Dušan Živković

Angelique Sanchez

Thomas Menneteau

Adrien Schahl

Sandrine Lise

Christine Kervarrec

Ana Toste Rêgo

Paula C A Da Fonseca

Matthieu Chavent

Charles Pineau

Odile Burlet-Schiltz

Julien Marcoux
email: julien.marcoux@ipbs.fr

Marie-Pierre Bousquet

-PAGE: sodium dodecyl sulphatepolyacrylamide gel electrophoresis SEC: size exclusion chromatography SPE: solid phase extraction TD(P): Top-Down (proteomics) TOF: time-of-flight UVPD: ultraviolet photodissociation XL-MS: cross-linking -mass spectrometry

Keywords: affinity purification AR, axial elements BU collision-induced dissociation CO, crossing over capillary zone electrophoresis ECD, electron capture dissociation EM, electron microscopy ESI, electrospray ionization ETD, electron transfer dissociation FACS, fluorescence activated cell sorting FT, Fourier transform GELFrEE, gel-eluted liquid-fraction entrapment electrophoresis GFP, Green Fluorescent Protein ion cyclotron resonance IEC, ion exchange chromatography IEM, ion evaporation model IM, ion mobility IRM, ion routing multipole iSCAMS (MP), interferometric scattering mass spectrometry (mass photometry) mass photometry (iSCAMS) quadrupole RPLC, reverse phase liquid chromatography SC, synaptonemal complex SDS

Now, here, you see, it takes all the running you can do, to keep in the same place. If you want to get somewhere else, you must run at least twice as fast as that!"

The Red Queen in Through the Looking-Glass, by Carroll, Lewis "Gradually it was disclosed to me that the line separating good and evil passes not through states, nor between classes, nor between political parties either-but right through every human heart-and through all human hearts. This line shifts. Inside us, it oscillates with the years. And even within hearts overwhelmed by evil, one small bridgehead of good is retained. And even in the best of all hearts, there remains… an unuprooted small corner of evil. "

The Gulag Archipelago, by Solzhenitsyn, Alexander "The moment you follow someone you cease to follow Truth." Krishnamurti, Jiddu "For those who follow nature everything is easy and straightforward, whereas for those who fight against her life is just like rowing against the stream."

Letters, by Seneca my project. Thank you David for your hospitality and warm welcome, great conversations, scientific or not and everything you taught me about proteomics.

Thank you Anne for your discussions and insights. They helped to obtain a lot of perspective on my work and proteomics in general.

Great many thanks to all the other members of the team, Emma, Renaud, Marlene and Karima (never separated those two, not even here), Alex, Manue and Carine, for your tutorship and for all the help you provided with enthusiasm and sincere will to teach me. I thank to Elenore and Lucie for the for their pleasant company. I thank Mylene for sharing the stories of her travels and for welcoming me warmly to the team.

I thank all my dear friends from the lab, who made the days of hard work feel like we're in this together. Giulia, you were a true Sun's ray and you bring joy wherever you go! Laurie, thank you for all the good laughs we had. You were the example to me of how a PhD should be handled and how an aspiring scientist should set his goals, and I thank you for that as well. Thank you, Neil, for being comrade in arms during these though four years. Thank you Angelique, for showing me what perseverance is and for teaching me the true value of modesty. Thank you Rim, for teaching me what it means to take oneself seriously and how powerful independent thinking is. Thank you Frances, for showing me what enthusiasm for science means. Thank you Xime, for your infinite care and love. Your support means a

Mass spectrometry -Swiss Army knife to dissect the spermatoproteasome structure and function

La spectrométrie de masse -un couteau suisse pour disséquer la structure et la fonction du spermatoprotéasome

Abstract

The proteasome is the main enzymatic complex for targeted proteolysis in the cell. Its core complex (20S) consists of four stacked heptameric rings and requires activator complexes: 19S, PA28αβ, PA28γ and PA200, which regulate 20S activity and substrate specificity. Alternative 20S subunits exist to further modulate the proteasome activity. Spermatogenesis is a process of male germ cell differentiation, where spermatogonia (SPG) transform through spermatocyte (SPC), then spermatid (SPT) stages, to become spermatozoa. This process requires intense proteolysis. The spermatoproteasome (spt20S) is specific to the developing gametes and essential for spermatogenesis. It differs from the standard proteasome (std20S) by only one subunit -α4s, which replaces the constitutive α4 subunit. Together with PA200, the spt20S plays an important role in meiosis progression, however, the mechanisms that make it different compared to std20S remain unknown.

We established complementary proteomic pipelines for characterisation of proteasome complexes in the testes, combining immunopurification (IP) and mass spectrometry (MS) analysis. Our Top-Down analysis of purified proteasome, showed for the first time that both α4 and α4s carry the same PTMs. Using Bottom-Up proteomics we compared the interactome of total proteasomes with that of the spt20S only, obtained using a specific antibody we developed for this purpose. We established that α4 and α4s do not co-exist in the same 20S, although they are almost equally abundant in the testes. We also measured that 19S and PA200 regulators were bound in higher ratios to the spt20S compared to std20S. Among other preferentiallyassociated proteins of spt20S were PI31 and Fbxo7, both shown to be crucial for fertility. They mediate proteasome transport and docking of E3 ligases -both processes that could be crucial for spt20S function.

We then obtained germ cells at different differentiation stages and performed a proteomics analysis of both lysates and IP-ed proteasome complexes, to establish a dynamic image of the proteasome throughout spermatogenesis. We observed a complete shift from std20S to spt20S between pre-meiotic SPG and meiotic and post-meiotic SPC and SPT cells. We explained this by a shift in expression, rather than preferential incorporation of α4s. Upon entering meiosis, the PA200 association with core proteasome increased 7-fold, marking its importance in gamete development. Although PA200 was represented in literature as the main spt20S interactor, we show that 19S was undoubtedly stoichiometrically dominant, occupying 60% of the existing 20S in SPCs -an unprecedented proteasome activation. Identified spt20S-interacting proteins largely correlated with previous interactome analysis on the whole testes, showing robustness of our methods. We identified synaptonemal proteins bound exclusively to spt20S and numerous proteins involved in ubiquitylation, cell cycle and meiotic progression as well as cellular transport, which fits the current model of spt20S role, proposed by earlier work.

The shift from α4 to α4s in meiosis was shown to be crucial, but what is the molecular basis for this transition? The hydrogen-deuterium exchange experiment coupled to MS helped us to show for the first time that the two proteasomes exhibit different binding interfaces: α4s contains regions that are more flexible compared to α4. We further supported this finding with pull-down assays, which showed that 19S binds more strongly to the spt20S than to std20S, which would explain the increase in proteolytic activity required during meiosis. The spt20S showed a higher tryptic activity compared to the std20S in vitro, which might reflect a particular need for histone degradation.

Altogether, our data reveal a more complex process of spt20S regulation than previously suggested and set the basis for structural and functional differences between the spt20S and std20S.

Résumé

Le protéasome est le complexe enzymatique protéolytique principal de la cellule. Son coeur catalytique (20S) est formé de quatre anneaux heptamériques. Son activité et sa spécificité de substrat peuvent être régulées par les complexes 19S, PA28αβ, PA28γ et PA200 ainsi que par des sous-unités 20S alternatives. La spermatogenèse est un processus de différenciation des cellules germinales mâles: les spermatogonies (SPG) se transforment en spermatocytes (SPC), en spermatides (SPT) puis en spermatozoïdes. Ce processus requière une protéolyse intense. Le spermatoprotéasome (spt20S) est spécifique des gamètes en développement et essentiel à la spermatogenèse. Il diffère du protéasome standard (std20S) par la sous-unité α4s qui remplace la sous-unité constitutive α4. Le spt20S joue un rôle important avec PA200 dans la progression de la méiose, mais les mécanismes qui le rendent différent du std20S restent inconnus.

Nous avons établi des stratégies protéomiques complémentaires pour caractériser les complexes du protéasome immunopurifiés à partir de testicules. L'analyse Top-Down de protéasome purifié, nous a permis de montrer pour la première fois qu'α4s et α4 portent les mêmes MPTs. La protéomique Bottom-Up, nous a permis de comparer les immunopurifications (IPs) de protéasomes totaux avec celles obtenues avec un anticorps spécifique du stp20S que nous avons développé. Nous avons établi qu'α4 et α4s ne coexistent pas dans le même 20S, bien qu'ils soient presque également abondants dans les testicules. Nous avons également trouvé plus de 19S et de PA200 liés au spt20S qu'au std20S. Les autres protéines préférentiellement associées au spt20S incluent PI31 et Fbxo7 qui sont cruciales pour la fertilité et d'importants médiateurs du transport du protéasome et de l'ancrage des E3 ligases -deux processus qui semblent cruciaux pour la fonction du spt20S pendant la spermatogenèse.

Nous avons ensuite obtenu des cellules germinales à différents stades de la différenciation et l'analyse protéomique des lysats ainsi que des IPs, nous a permis d'établir un interactome dynamique du protéasome tout au long de la spermatogenèse. Nous avons observé un changement total du std20S au spt20S entre les SPG pré-méiotiques et les SPC/SPT méiotiques et post-méiotiques. Un changement d'expression semble responsable, plutôt qu'une incorporation préférentielle d'α4s. En entrant dans la méiose, l'association de PA200 avec le protéasome a augmenté 7 fois, confirmant son importance dans le développement des gamètes. Bien que PA200 soit d'après la littérature le principal activateur du spt20S, nous montrons que le 19S est est en réalité majoritaire, lié à 60% des 20S dans les SPC -une activation du protéasome sans précédent. De nombreux partenaires du spt20S sont identifiés à la fois dans les cellules germinales et dans les testicules entiers, montrant la robustesse de nos méthodes. Ceux-ci incluent des protéines synaptonémales, de nombreuses protéines impliquées dans l'ubiquitylation, le cycle cellulaire et la progression méiotique ainsi que le transport cellulaire, en accord avec les fonctions du spt20S proposées dans la littérature.

Le passage d'α4 à α4s semble crucial pour la méiose, mais quelles sont les bases moléculaires de cette transition ? L'échange hydrogène-deutérium nous a permis de montrer pour la première fois que les deux protéasomes présentent des interfaces d'interaction différentes : α4s contient des régions plus flexibles qu'α4. Cette découverte est confirmée par des pull-down in vitro montrant que le 19S se lie plus fortement au spt20S qu'au std20S, expliquant la hausse d'activité protéolytique pendant la méiose. L'activité trypsique du spt20S est plus élevée que celle du std20S in vitro, ce qui pourrait refléter la nécessité de dégradation des histones.

Globalement, nos données révèlent un processus de régulation du spt20S qui est plus complexe que ce qui avait été suggéré précédemment et jettent les bases des différences structurales et fonctionnelles entre le spt20S et le std20S. Table 1. Brief summary of structural MS techniques …….……………………………………………………. Table 2. Theoretical masses of the proteasome complexes and their regulators. ……………... Table 3. Gene Ontology term enrichment for biological process -search results. ……………..

Introduction

2.1 Spermatogenesis

The role of germ cells

Germ cells are unique in their role of transmitting genetic information from parents to the offspring. Not only do germ cells transmit the blueprint for the development of the new organism, but they provide the constituents of the first cell, out of which the new organism develops.

Morphology of the testes

To understand the process of spermatogenesis, we must first understand the organisation of the organ that facilitates it. The human testes are a pair of organs oval in shape, measuring 2.5 x 4 cm and held by a connective tissue capsule called tunica albuginea. The testis is divided into 370 lobules containing seminiferous tubules and intertubular tissue. While the intertubular tissue contains Leydig cells that secrete hormones, the seminiferous tubules contain the differentiating germ cells and the Sertoli cells that support and modulate germ cell development [START_REF] Holstein | Understanding spermatogenesis is a prerequisite for treatment[END_REF]. The seminiferous tubules are very coiled in shape and both of their ends connect to the rete testis -a sort of canal that helps pooling and transporting differentiated germ cells into the epididymis (see Figure 1.A). The epididymis is a meandering canal that accumulates the differentiated germ cells and facilitates its final maturation stages, during which the spermatozoa obtains motility and capacity for fertilization [START_REF] James | The role of the epididymis and the contribution of epididymosomes to mammalian reproduction[END_REF]. The seminiferous tubules are around 180 µm in diameter and are made up of germinal epithelium and peritubular tissue that surrounds it and holds it together in a tube shape. The germinal epithelium is where the germ cell differentiation takes place. The stem cells perpetually dividing from puberty onwards, are found at the base -outward part of the cross-section of the seminiferous tubule (see Figure 1.B) and as they enter differentiation, migrate towards the middle, until they are released into the lumen. As the germ cells develop, they are enveloped in the Sertoli cell membranes that support them by providing structural scaffold, nutrients and growth factors. They also prevent immature cells from drifting into the lumen [START_REF] Stukenborg | Ontogenesis of Testis Development and Function in Humans[END_REF]). Among themselves, Sertoli cells establish special connections called tight junctions that separate the basal and adluminal compartments of the germinal epithelium and creates the bloodtestis barrier. This barrier is crossed by the differentiating cells and is considered a protected zone from potential harmful influence, such as pathogens and drugs or toxins.

Goniogenesis

The germ cell line is continuously renewed in males throughout life, originating from the primordial germ cells. The exact origin of the human primordial germ cell line is not exactly known, due to ethical constraints of the studies. Between the 4 th and 6 th week of gestation, the primordial gonial cells (PGC) migrate to the gonads in development, from which point they are referred to as gonocytes (see Figure 2.A). The sex of the gonocytes is not determined by themselves, but rather by the surrounding cells -in males that would be the hormonal control of Leydig, Sertoli and mesenchymal cells. Unlike their female counterparts, the gonocytes do not enter meiosis, but start proliferating mitotically and six months postnatally, they migrate to the basal membrane of the seminiferous tubules and become spermatogonia [START_REF] Stukenborg | Ontogenesis of Testis Development and Function in Humans[END_REF]. Two main types of spermatogonia exist: A and B. The A type is dividing mitotically to preserve the germ stem cell fund. A certain part of this population will enter the path of differentiation and thus become spermatogonia B. Morphology of the human testis. A: Cross-section of the human testes. The organ is divided in lobules, each containing highly coiled seminiferous tubules, eluting its secretions from both ends into the rete testis, which transfers the differentiated germ cells into the efferent ducts of the epididymis. As they travel through the epididymis, the germ cells reach full maturation and are stored in the epididymis. B: Cross-section micrograph of a histological preparation of a seminiferous tubule. The pool stem cells called spermatogonia renews itself by division, while a part of the population begins to differentiate. As they do so, germ cells move towards the middle of the seminiferous tubules, until they are released into the lumen. Throughout the differentiation process, germ cells are immersed in invaginations on the surface of Sertoli cells that serve as support and quality control.

Differentiation

Spermatogenesis encompasses three distinct phases: mitotic proliferation and development of spermatogonia; meiotic division of the spermatocytes to yield cells containing only one recombined chromosome copy and chromatin condensation and polarization of the spermatids, which ends in spermiogenesis -development of motile spermatozoa.

Spermatogonia B go through two mitotic divisions to give four copies, while maintaining the cytoplasmic bridges, without fully separating from each other through meiosis, all the way until late spermatid stages. These bridges are called syncytial connections and are thought to play an important role in the synchronisation of the sperm cell development.

The whole process takes more than two months to complete in humans and at a single spot, it is not initiated constantly, but in spiral-shaped waves. This means that as they develop, the cells migrate longitudinally, along the length of the tubule and towards the lumen. Since the dividing cells stay connected for the good part of the differentiation process, cells in seminiferous tubule crosssections can be observed in groups of the same development stage, with previous or next wave of differentiation of neighbouring cells being visible on the same cross-section. The schematic representation of the spermatogenesis wave can be seen on Figure 2.B. Note that we do not usually see all stages of differentiation in a single histological preparation where a seminiferous tubule is perpendicularly cut, due to the spatial organisation of the differentiation process. Germ cell differentiation overview. A: Before birth, the primordial gonial cells migrate to the gonads, where they become gonocytes. During foetal development, gonocytes settle in the basal layer of the seminiferous tubules, thus becoming spermatogonia. The type A spermatogonia divide and a subpopulation of them enter the differentiation pathway by becoming type B spermatogonia. They further divide mitotically and upon entering spermatocyte stage, they initiate meiosis. After meiosis, the cells enter the spermatid stage, in which they contain recombined genetic material and reduced number of chromosomes. The polarisation of the cells follows: spermatids form a head and a tail, discard excess cytoplasmic material and become fully separated. Upon maturation, they are released by the Sertoli cells into the lumen. The letter markings C and N stand for number of copies and ploidy, respectively. B: Spermatogenesis is initiated in waves. Three neighbouring waves of differentiation are given as an example (marked as I, II and III), with stages of differentiation marked on the ribbons: L-leptonema, Z-zygonema, P-pachynema. As the differentiation progresses, cells migrate along the seminiferous tubule and towards the lumen (see lower diagram representing the migration of the neighbouring waves inside of a seminiferous tubule). In a cross-section we would be able to distinguish neighbouring waves and different stages -later ones towards the middle and the earlier ones towards the peritubular tissue. Adapted from A: (Bolcun-Filas and Handel, 2018); B: [START_REF] Hirsh | The anatomical preparations of the human testis and epididymis in the Glasgow Hunterian Anatomical Collection[END_REF].

Meiosis

Meiotic division at the spermatocyte stage is a crucial step in gametogenesis, since it results in genetic shuffling and is thus responsible for genetic variability. The process occurs through meiosis I and meiosis II. Meiosis I is responsible for duplication of the genetic material and shuffling parts of the chromosomes by breaking and cross-joining the DNA strands of homologous chromosomesmaternal and paternal, through a process called recombination, which is followed by cell division. Meiosis II follows, whereby single copies of homologous chromosomes are divided between haploid sister cells.

Somatic cells have two sets of chromosomes -one maternal and one paternal, also called diploidy; it can thus be said that they also possess two copies of each chromosome. The state of the genetic material in a somatic cell can thus be described as 2N2C, where N is the number of sets of chromosomes and the C is the number of copies of each chromosome. For an overview of the germ cell differentiation, the reader is referred to Figure 2. The spermatocytes I (primary spermatocytes) initiate their S phase, where they double the amount of DNA, effectively reaching 2N4C state -two copies of each chromosome, from both maternal and paternal sets -four copies in total. Here, they enter the prolonged prophase of meiosis I, where the recombination takes place (Figure 3). It is a complex, highly coordinated process lasting 24 days in humans. The prophase I is divided into five differentiable states: leptonema, zygonema, pachynema, diplonema, diakinesis. Progression through metaphase, anaphase and telophase take only a fraction of the time, whereby primary spermatocytes divide into spermatocytes II (secondary spermatocytes); their state can be described as 1N2C: they have two copies of one recombined set of chromosomes, where each copy is a mix from both parents. It is important to note that each of these two copies of each chromosome are unique in their combination of maternal and paternal genes. Meiosis II is essentially a cell cycle of the secondary spermatocytes, where the number of copies of chromosomes is reduced, resulting in haploid cells, and only takes a few hours to complete. The theoretical result of meiosis I and II are four haploid spermatids originating from each primary spermatocyte, each containing a unique combination of maternal and paternal genes. Spermatids then proceed into spermiogenesis -a process that transforms them into spermatozoa (Bolcun-Filas and Handel, 2018).

As mentioned before, this is the basis of the genetic variability among offspring -the genes of the individual's parents are uniquely recombined to give half of its offspring's genome. The male haploid cell, combined with the female haploid cell, result in an individual carrying two sets of chromosomes, each set being a unique combination of their grandparents' genes.

Events of the prophase I

The prophase I is marked by two major events: pairing of the homologous chromosomes and their recombination. To facilitate the pairing and recombination process, specific protein structures arise to harness the DNA. The cohesin directly interacts with the chromatin -the cohesin structures maintain the sister chromatids together throughout meiosis I, until their separation in anaphase II (see Figure 4). Meanwhile, they also support the integrity of the chromatids after crossover events and serve as a platform for synaptonemal complexes formation. Furthermore, sandwiched plates called synaptonemal complex -SC, connect two homolog chromosomes together. Outer layers of the SCs are composed of SYCP2/3 proteins and are called axial elements (AE), which organize the DNA for the recombination event. The inner layer of the "sandwich" is called central element (CE), flanked on both sides by transverse filaments, composed of the SYCP1 protein. In these sites, various proteins are recruited to perform different steps of recombination. The SC is a dynamic structure, which alters its size and constituents throughout meiosis. Strand Breaks (DSBs) of one of the two homologous chromosomes. Single thread of cleaved DNA is inserted into the homologous DNA ladder that unwinds to accommodate the invading strand. This complex may be stabilized or not and therefore a non-crossover (NCO) or a crossover (CO) scenario can occur. In a crossover scenario, single strands of DNA are exchanged between the homologs forming double Holliday junction. Upon resolving the junction by nickase and ligase activity, two novel recombined double DNA strands emerge. Adapted from (Marston and Amon, 2004).

The recombination is initiated by a homology search and pairing of DNA from maternal and paternal homologous chromosomes (Figure 3). The double-stranded DNA break is induced on one chromosome and only one DNA strand from that break is introduced to the DNA ladder of the other homologous chromosome ("strand invasion"). The homologous DNA ladder unwinds and opens to accommodate the invading broken strand that binds to a complementary sequence. A recognisable D-loop is formed and the events that follow can take two courses. In a non-crossover (NCO) scenario (Figure 3, left), the DNA does not undergo further conformational changes and the invading strand is ejected. In a crossover (CO) scenario (Figure 3, right), DNA undergoes conformational changes and in turn, one of the "accommodating" strands breaks off to pair with the chromosome of the invading strand. Missing parts of the DNA ladders are filled in by polymerase and ligase activity. At this stage, there are two points at which DNA crosses between the chromosomes, called Holliday Junctions. These junctions are resolved by nicking the DNA in a manner that allows for the exchange of large portions of DNA between the homologous chromosomes. The coordinated action of several enzymes is necessary to facilitate these events. The sites of the crossovers seem to be conserved between species and have been mapped in mice. Around fifty crossovers are typical for human spermatocytes (Eichenlaub-Ritter, 2014).

Leptonema is marked by chromatin condensation, whereby two sister chromatids become visible and are connected via cohesins. Chromosomes also begin to pair in leptotene, where the telomeres (chromosome ends) attach to the inner surface of the nuclear membrane at the same spot. Cohesins will fetch and hold together the DNA of the sister chromatids, with loops protruding along the AE Arrays of AE spread along the deposited cohesin protein that directly interact with the DNA of the chromatids (in red in Figure 4). On some of the DNA loops, additional protein complexes form, made of PRDM9 and associated proteins that activate the region for recombination. SPO-11 complexes are then recruited to induce DSBs -this is a signal for recombination machinery to be employed (Eichenlaub-Ritter, 2014). Chronological order of events at the synaptonemal complex. Cohesin core maintains the integrity of the chromosomes and keeps sister chromatids together (left scheme). In leptonema, the axial elements bind to the cohesin and protruding DNA loops bind protein complexes that induce DSBs. In zygonema, the pairing of the homologs continues by the formation of the first central elements, and the first recombination nodules are formed. The left schema represents the way SC holds the homologs together. In pachynema, the pairing of the homologs is complete and the recombination events are brought to the end. The sex body contains X and Y chromosomes (yellow) that only partially participate in the recombination. In diplonema, the SCs are dissolved and the homologs are connected only by the chiasmata. What follows is the separation of recombined homologs. In the legend, key markers and the regulators of the events are listed, as well as the constituents of the SC. Adapted from (Bolcun-Filas and Handel, 2018).

During zygonema, the pairing of the chromosomes continues: the transversal elements (in light green in Figure 4) interact with each other to form the central element (containing SYCE1, 2 and 3, TEX12 and Six6OS1; in dark green in Figure 4) and the SC pairing proceeds in a zipper-like manner along the entire AE-covered regions. Early recombination nodules are formed, containing proteins such as RAD51 and DMC1. Formation of the CE is coordinated at the same time as the HORMAD protein removal coordinating the DSB-related events. Transitional recombination nodules mediate the beginning of the crossover events, marked by proteins such as RPA, BLM and RNF212 (Hunter, 2015).

In pachynema, the homologous chromosomes are fully paired, along their entire length. In males, a distinct structure is formed between X and Y chromosomes, called the sex body (yellow blob in Figure 4, top). The X and Y chromosome are transcriptionally inactive and only participate partially in recombination, via the homologous region called pseudoautosomal region. Main crossover events take place -late recombination nodules appear and facilitate final DSB repair, resolving the DSBs, either as a CO or an NCO event.

In diplonema, homolog chromosomes detach along the entire length, while the sister chromatids of each chromosome still remain together. HORMAD proteins colonize again the cohesin surface as the SC is dissolved (Figure 4, top right). The only thing then keeping the homolog chromosomes together are the sites where crossing-over occurred -the chiasmata, which are basically threads of DNA that are now exchanged between the two homologous chromosomes.

Diakinesis describes the events following dissolution of the nuclear membrane and involves the separation of the recombined homologous chromosomes (Eichenlaub-Ritter, 2014).

Events after the prophase I

At the end of prophase I, the homologous chromosomes are still attached to each other by one or several chiasmas. After the dissolution of the nuclear membrane, a bipolar spindle is formed (Figure 5). Two pairs of centrioles, barrel-shaped organelles composed of microtubules, position themselves on the opposite sides of the cell and form the so-called meiotic spindle -the scaffold permitting the separation of homologous chromosomes and their migration to the separate poles of the cell. The microtubules attach themselves to the centromeres of both sister chromatids, while pairs of homolog chromosomes are set to separate into different poles of the cell. Resulting tension between pairs of chromosomes, still connected by the chiasma, leads to their alignment to the equator of the cell. The centromeres recruit spindle assembly checkpoint proteins (SAC) that ensure proper attachment of the chromosomes. Anaphase-promoting complex (APC) is rendered inactive by the SAC, as long as it senses irregularities in chromosome binding to the meiotic spindle. At anaphase I, homolog chromosomes migrate to the poles of the cell that will separate into two daughter cells.

Nucleus is formed very briefly and chromosomes re-condense to initiate prometaphase II -a short stage in which sister chromatids are organised at the equator for separation. Anaphase II takes place, whereby sister chromatids are separated and migrate to what is to become separate (haploid) cells (Eichenlaub-Ritter, 2014; Hunter, 2015). Stages in meiotic division. On the left, microscopy of the distinct stages of prophase I are displayed in line with the scheme of the molecular events at the chromatin level. The copies of DNA are duplicated in the S phase giving sister chromatids. Homolog pairing begins in leptonema. The synapsis forms in zygonema and is fully achieved in pachynema. Crossover begins in zygonema but is fully complete in pachynema. In diplonema, synapses disappear and the only thing holding the homologs together are chiasmata -the strands of DNA marking the spot of the crossover. The meiotic spindle now forms and the recombined homologs are separated into two cells. Another division separates the recombined sister chromatids into haploid cells. The theoretical outcome of the process is four haploid spermatids from one primary spermatocyte. Adapted from (Hunter, 2015).

Spermiogenesis and spermiation

The process of spermiogenesis correspond to the development of the spermatids into spermatozoa. Key events taking place during this process are as follows (Figure 6). Nuclear chromatin is condensed to one tenth of its initial volume by removing histones and replacing them with protamines. This process is a highly organised succession of histone post-translational modifications [START_REF] Sheng | The Role of Histone Ubiquitination during Spermatogenesis[END_REF]Wang et al., 2019) and histone replacements by transitory DNA-binding proteins [START_REF] Rathke | Chromatin dynamics during spermiogenesis[END_REF]. The process seems to be at least in part coordinated by a set of enzymes that are part of the Ubiquitin-Proteasome System that will be explained in detail in the following chapter. Acrosome cap is formed, which is an organelle that digests the proteoglycan envelope of the egg during fertilization (the event known as the acrosomal reaction). The acrosome originates from the Golgi apparatus, is filled with enzymes and it is located at the top of the nucleus. Flagellum is developed and attached to the nucleus. Besides a portion of mitochondria, most of the organelles are discarded with the cytoplasm. Upon finishing this process, the process called spermiation takes place, whereby spermatozoa are released into the lumen and further removal of cytoplasm and its contents occurs. The cytoplasmic residue is phagocytised by Sertoli cells. The spermatozoids are not yet motile nor are they capable of acrosomal reaction and are transported by peristaltic movements into the rete testis, floating in the secretions of the Sertoli cells. The sperm maturation is far from over and will require additional stay in the epididymis (O'Donnell et al., 2011). Figure 6. Spermiogenesis and spermiation. Spermatid polarizes by reorganising its contents. The Golgi apparatus develops into an acrosome, spermatid compactifies the nucleus, grows a tail and sheds most of its cytoplasm. Mitochondria are placed at the base of the nucleus and help power the motors of the flagellum. In the process of spermiation, spermatozoa are released from the grip of the Sertoli cells and additionally shed cytoplasm. However, the process of maturation is not done. The spermatozoa are neither motile nor capable of acrosomal reaction. They will acquire this capability only in the epididymis. Adapted from [START_REF] Schatten | Sperm Centrosomes: Kiss Your Asterless Goodbye, for Fertility's Sake[END_REF].

The process of spermiogenesis is far from perfect -high occurrence of malformed spermatozoids is expected. These malformations come in many various forms, some of which are a lack or malformation of the tail, a double tail, multinucleated sperm cells, absence of acrosome, etc. The whole process of spermatogenesis is very inefficient. Only 25% of the cells reach full maturity and more than half of them are malformed. The remaining 75% are lost in apoptosis or degradation. Even though, the rate of daily production of spermatozoa per gram of testicular tissue in men is 3 to 4 million [START_REF] Holstein | Understanding spermatogenesis is a prerequisite for treatment[END_REF]. Every lost cell or cytoplasmic vesicle is phagocytized and reused again as a source of energy or a building block for the new gamete cells. In a tissue with such a tremendous protein turnover, the role of protease systems, which recycle the cellular material, is paramount.

Proteasome

Proteolytic systems of the cell

In order to maintain homeostasis, the cell must respond to changes in its internal and external environment. On another level, a tight regulation of cellular behaviour is necessary for functioning of the organism. In both of these concepts, protein degradation is as essential as the protein synthesis. At the cellular level, unnecessary or damaged proteins need to be degraded, while others are degraded as a part of signal transduction or regulation of their activity. Vast number of proteases operating within the cells exert their activity on one or very few substrates. However, there are two major systems of broader substrate spectrum that degrade proteins down to short peptides: the lysosome and the proteasome. Lysosomes are specialized intracellular organelles which maintain low pH environment and contain a variety of proteases used to degrade any introduced material. This may be another entire organelle marked for recycling, as a part of a damage response [START_REF] Youle | Mechanisms of mitophagy[END_REF], engulfed cytoplasmic material (starvation response) [START_REF] Shang | Nutrient starvation elicits an acute autophagic response mediated by Ulk1 dephosphorylation and its subsequent dissociation from AMPK[END_REF] or a targeted protein imported into lysosome [START_REF] Bandyopadhyay | The Chaperone-Mediated Autophagy Receptor Organizes in Dynamic Protein Complexes at the Lysosomal Membrane[END_REF]. While lysosomal proteolysis is a tightly regulated process, the enzymes involved are not substrate-specific. The proteasome is the primary system for targeted degradation of proteins and its activity can be regulated in many ways.

The 20S proteasome

The proteasome is a multiprotease complex responsible for the targeted degradation of most proteins in the cell. It does so through a complex system of enzymes in charge of substrate tagging and recognition. It consists of the core particle which houses the catalytic sites and associates with regulators, which modulate its activity and substrate specificity. The core particle, also called the 20S proteasome, is a barrel-like 28-mer consisting of four stacked hetereoheptameric rings, 715 kDa in size. Each of the two inner rings (β rings) have three catalytic sites located on three different catalytic subunits. The two β rings are identical and are facing each other, forming a cavity (Figure 7). From each side, the two β rings are capped by two other identical rings (called α rings), effectively closing the catalytic cavity and forming a gated pore for substrate introduction. The yeast and mammalian proteasomes have 14 different subunits forming the core, seven per each type of ring, while the bacterial and archaeal proteasomes are much simpler -α and β rings are homoheptameric in archaea, while bacteria have two types subunits per ring (Kniepert and Groettrup, 2014). However, in this work we will focus on the more evolved mammalian systems. The proteasome complexes are often named by their Svedberg sedimentation coefficient -for the core proteasome particle, that would be 20S.

The 20S proteasome itself has relatively low specificity and low capacity for protein digestion, since the channel for the protein introduction is mostly closed by the protruding N-termini of the α ring subunits (Osmulski, Hochstrasser and Gaczynska, 2009). The channel is opened by the association of the 20S with one of its regulators. In the case of the 19S, its major regulator, the unlocking of the channel is induced by the introduction of protruding loops of the 19S into the so-called α-pocketsthe pits formed at the interface of neighbouring α ring subunits (Smith et al., 2007). Most of the proteasome substrates require energy in the form of ATP to be unfolded and subsequently degraded. To achieve specificity, proteins are modified by Ubiquitin (Ub) -a small protein tag which marks them for degradation. As it will be discussed later, this is not always the case: many proteins can be degraded without ATP-hydrolysis, notably denatured proteins and intrinsically-disordered proteins. The proteasome is a threonine protease: the cavity containing the active sites is around 100 Å in length and 60 Å in width (see Figure 7, right). In this space, six threonine residues act as nucleophiles to catalyse the hydrolysis of the peptide backbone. The threonine residues come from three different subunits of the two inner rings (β1, β2, and β5), marked in red on the sliced view of the proteasome complex in Figure 7 (right). Specific type of activity is commonly associated to each of the catalytic subunit, depending on the exact nature of the catalytic site that is being formed upon proteasome assembly: the caspase-like activity is attributed to β1, trypsin-like activity to β2 and chymotrypsin-like activity to β5 subunit, however, the cleavage preferences go beyond this simple division [START_REF] Nussbaum | Cleavage motifs of the yeast 20S proteasome subunits deduced from digests of enolase 1[END_REF]. The unfolded protein finds itself in a tightly confined space, with the peptide bonds exposed to the highly effective concentration of catalytic sites (∼50 mM), ensuring efficient digestion into peptides [START_REF] Nussbaum | Cleavage motifs of the yeast 20S proteasome subunits deduced from digests of enolase 1[END_REF].

Figure 7. Structure of the eukaryotic 20S proteasome. Left: the 20S proteasome is composed of four stacked heteroheptameric rings in αββα configuration. It is a symmetric complex, with two β rings facing each other and two α rings flanking them. On the right, longitudinal cross section of the complex reveals how the two β rings form a catalytic chamber and the α rings create gated pores which protects from unregulated proteolysis. The N-terminal threonine catalytic sites are marked in red. Adapted from (Finley, 2009).

Proteins degraded through the proteasome system yield peptides of 8 amino acid residues on average, which can then be further degraded by peptidases to amino acids. In higher organisms, the proteolytic peptides can become part of the innate immune system, via the major histocompatibility class I (MHC-I) complex [START_REF] Rock | Protein degradation and the generation of MHC class I-presented peptides[END_REF], which presents surface antigens on the cells, in order for immune cells to differentiate the self from non-self and to be able to detect abnormal cells. There are several ways the proteasome activity can be regulated: 1) by incorporation of the alternative, tissuespecific subunits that alter its activity and substrate specificity; 2) post-translational modification of its subunits; 3) association with proteins and protein complexes which activate it and regulate its substrate specificity, also known as proteasome activators/regulators 4), or other proteasomeinteracting proteins (PIPs) that further modify the activity of the proteasome or proteasome-activator dyad; 5) through differential expression of proteasome subunit-coding genes. These systems of regulation give a powerful platform for substrate targeting, making the proteasome a hugely versatile system involved in almost all cellular processes: cell division (Min and Lindon, 2012), differentiation (T cell - [START_REF] Widjaja | Proteasome activity regulates CD8+ T lymphocyte metabolism and fate specification[END_REF], sperm cell -(Q. [START_REF] Zhang | Meiosis I progression in spermatogenesis requires a type of testis-specific 20S core proteasome[END_REF], apoptosis [START_REF] Sohn | The Proteasome Is Required for Rapid Initiation of Death Receptor-Induced Apoptosis[END_REF], heat shock response [START_REF] Aristizábal | Heat stress reveals high molecular mass proteasomes in Arabidopsis thaliana suspension cells cultures[END_REF], DNA repair [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF], immunity (Kloetzel, 2004) and many others. The summary of principal dimensions of the control of the proteasome activity can be seen on Figure 8.

Finally, the proteasome itself can become the target of the degradation machinery. It has been observed that inactive proteasomes (e.g. inhibited by a drug) are Ub-ylated and transported to the autophagosome, where they get degraded by lysosomal enzymes and low pH [START_REF] Hoeller | How the proteasome is degraded[END_REF]. In order to properly maintain homeostasis, the proteasome is regulated on multiple levels: 1) new 20S complexes can be syntesized by controlling the gene expression and 2) PTMs can be introduced to modulate 20S activity or even mark it for degradation; 3) alternative, tissue-specific subunits exist, which upon incorporation modulate the peptide product profile or 4) the interaction with proteasome-associating proteins. Many regulatory particles can bind to the 20S, altering its substrate specificity, the prevalent being 19S, which makes up the 26S complex together with the proteasome core. Proteasome-dependant proteolysis is further regulated by substrate-interacting proteins, which can help select or shuttle targets for degradation or proteasome-interacting proteins, which can inhibit or activate an existing proteasome complex. Finally, the proteasome can be translocated directly to needed location, within or without the cell. Adapted from (Ben-Nissan and Sharon, 2014).

The 20S proteasome assembly

The detailed schematic representation of the assembly is presented in the Figure 9: Two heterodimeric chaperone complexes facilitate the assembly of the α ring -PAC1/2 and PAC3/4, which ensure that all the subunits are inserted in the proper place and no premature regulator binding can occur [START_REF] Hirano | A heterodimeric complex that promotes the assembly of mammalian 20S proteasomes[END_REF]Le Tallec et al., 2007). The Ump1 protein (or POMP) then binds to the α ring to facilitate the binding of the β subunits in a specific order [START_REF] Ramos | Ump1p Is Required for Proper Maturation of the 20S Proteasome and Becomes Its Substrate upon Completion of the Assembly[END_REF]: β2, β3 then β4. Upon binding of the β3, PAC3/4 is displaced and β5, β6, β1 and β7 bind to form the hemiproteasome (15S complex). Upon β7 binding, hemiproteasome dimerizes into preholoproteasome -an immature proteasome core particle. The N-terminal propeptides of the β1, β2, β5, β6 and β7 subunits are then auto-cleaved to expose the active sites, which degrade the Ump1 chaperone, which is degraded in the process, yielding a mature 20S proteasome particle. An excellent review on the proteasome assembly was published by Bertolotti and Rousseau [START_REF] Rousseau | Regulation of proteasome assembly and activity in health and disease[END_REF].

Tissue-specific 20S proteasomes

As already mentioned in the previous chapter, the proteasome core particle subunits have alternative isoforms expressed only in specific tissues or under certain conditions. The incorporation of these alternative isoforms modulates proteasome behaviour: the proteasome activity, substrate specificity, types of peptides produced and regulator binding. For an overview of the principal proteasome types, refer to Figure 10.

The immunoproteasome and its intermediates are constitutively expressed by the cells of the immune system, however, their expression can be induced in almost any tissue by proinflammatory factors (Akiyama et al., 1994;[START_REF] Früh | Displacement of housekeeping proteasome subunits by MHC-encoded LMPs: a newly discovered mechanism for modulating the multicatalytic proteinase complex[END_REF]. In the tissues in course of infection, it can replace up to 90% of the standard proteasome (Khan et al., 2001). In the immunoproteasome, the catalytic subunits are replaced by the inducible isoforms: β1i, β2i and β5i. Intermediary forms also exist, where only one or two standard catalytic subunits are replaced [START_REF] Guillaume | Two abundant proteasome subtypes that uniquely process some antigens presented by HLA class I molecules[END_REF]. The peptides generated by the immunoproteasome can serve as antigens, presented by the MHC class-I complex at the surface of the cell. The T lymphocytes use these complexes to differentiate the healthy from abnormal or non-self cells. In order for a peptide to be recruited into the complex, it must fulfil two prerequisites: its length must be 8-9 amino acid residues and it must have C-terminal anchor residues (in humans, they are basic or hydrophobic in nature). The immune-subunits alter the proteasome peptide profile: the resulting peptides have C-termini that are not acidic and are more hydrophobic, resulting in a greater number of successfully-presented antigens (Kincaid et al., 2012). The role of the immunoproteasome is thus to modulate the antigen presentation in cases of infection or abnormal cells, thereby improving the immune response. Beyond antigen presentation, the immunoproteasome is essential in maintaining the proliferation of T cells in response to infection [START_REF] Moebius | Immunoproteasomes are essential for survival and expansion of T cells in virus-infected mice[END_REF] and oxidative stress (Pickering et al., 2010;Abi Habib et al., 2020) .

The thymoproteasome is only expressed in the epithelial cells of the thymic cortex and plays a role in T-cell education. The thymus is the lymphoid organ in which immature T-cells are positively Proteasome and negatively selected to differentiate self from non-self proteins. The thymus cortical and medullar cells express MHC complexes presenting many self-antigens. In the positive selection round, immature T-cells must be able to bind to the MHC, otherwise, they will undergo apoptosis. The fate of those that bind will be determined by the strength of the interaction: stronger binding induces differentiation to CD4 + (helper T-cells), while weak interaction induces CD8 + fate (killer T-cells). The negative selection round serves to eliminate the cells that would cause an autoimmune reaction [START_REF] Klein | Antigen presentation in the thymus for positive selection and central tolerance induction[END_REF]. In the thymoproteasome, the β5 subunit is replaced by the β5t isoform, while the β1 and β2 subunits are replaced by the β1i and β2i isoforms [START_REF] Murata | Regulation of CD8+ T cell development by thymus-specific proteasomes[END_REF] which induces a reduction in chymotrypsin-like activity of the proteasome, while the other two remain unaffected compared to the immunoproteasome [START_REF] Murata | Regulation of CD8+ T cell development by thymus-specific proteasomes[END_REF]. Lower chymotrypsin-like activity yields weaker ligand peptides of the MHC complex, since fewer peptides with hydrophobic C-termini are produced. Murata et al. postulated that this leads to lower-affinity interactions between the thymocytes and the T-cells, which is a prerequisite for successful T-cell selection [START_REF] Murata | Thymoproteasome: probable role in generating positively selecting peptides[END_REF]. As per Murata's model, 80% fewer CD8 + T-cells are produced in β5t knock out (KO) models, confirming this theory [START_REF] Murata | Regulation of CD8+ T cell development by thymus-specific proteasomes[END_REF].

The spermatoproteasome is a proteasome subtype expressed only in gonadal tissue. It has proven to be vital for germ cell development, however, no apparent role has been determined for it in females. Due to its central position in the subject of the study, the spermatoproteasome will be presented in detail in a subsequent chapter.

The proteasome regulators

The best described proteasome regulator is the 19S regulatory particle, which is capable of recognising and unfolding proteins marked for degradation via an Ub tag, then feed the unfolded protein thread into the 20S proteasome. The 19S can attach to the 20S core from either of its two sides, resulting in a 26S or 30S proteasome complex. Although imprecise, the term 26S proteasome is often used in the literature to generally refer to both the singly-capped and doubly-capped 19S-20S complexes. The 19S is the only proteasome regulator that uses ATP and is the largest. It is the most represented in literature, due to its involvement in the ubiquitin-proteasome system -the great targeted protein-recycling system of the cell. Other proteasome activators exist, such as PA28αβ, PA28γ and PA200, which, together with the 19S, can bind in different hybrid combinations to the 20S [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF]. The PA28αβ, PA28γ and the PA200 do not use ATP and have their own set of specific targets and activity-modulating properties, as we will see later.

19S

Unveiling the structure and function of the proteasome took 50 years of research, from the first negative stain electron microscopy to the near-atomic resolution structures of today [START_REF] Harris | Release of a macromolecular protein component from human erythrocyte ghosts[END_REF]Dong et al., 2019) Unlike the conformationally-stable 20S proteasome, the 19S (therefore the 26S as well), possesses a wide range of conformations, necessary for substrate processing (Dong et al., 2019).

Binding of the 19S does not immediately induce 20S α-ring pore opening; this is only the last step in the process of the substrate introduction to the catalytic sites [START_REF] Chen | Structural basis for dynamic regulation of the human 26S proteasome[END_REF][START_REF] Bard | Structure and Function of the 26S Proteasome[END_REF][START_REF] Bard | The 26S proteasome utilizes a kinetic gateway to prioritize substrate degradation[END_REF].

The proteasome regulatory particle, known as RP or 19S, is a regulatory complex consisting of at least 18 subunits in eukaryotes. Structurally, it is made up of two parts: the base and the lid, which may assemble separately. The base subcomplex consists of six different subunits (RPT1, RPT2, RPT3, RPT4, RPT5, RPT6 in S. cerevisiae) belonging to the ATPases Associated with diverse cellular Activities (AAA) family, which form a heterohexameric ring capable of binding to the 20S core through their Cterminal hydrophobic-tyrosine-X (HbYX) motif and which hydrolyse ATP to generate mechanical motion, similar to other AAA motors [START_REF] Esaki | Cdc48 AAA ATPase Regulates Protein Dynamics and Turnover in Mitochondria[END_REF]. The N-termini of the six base subunits make three α-helix bundle protrusions, onto which the lid subunits bind (see Figure 11, middle panel).

Two Ub/ Ub-like (UBL) receptors bind directly to the ATPase ring: RPN1 and RPN10, whereby RPN1 binds almost independently of the rest of the lid. The lid is composed of non-ATPase subunits, six of which are very similar in structure (RPN3, RPN5, RPN6, RPN7, RPN9 and RPN12) and two subunits form a dimer (RPN8 and RPN11). They come together to form a horseshoe-like complex, interacting via bundled C-terminal α-helices; bound to these eight subunits is RPN15 (X. There are two ways the 26S proteasome can assemble. The first way is the assembly of the lid and the base separately, which then unite into the 19S, to associate with the 20S. This pathway is more probable to be the accurate description of the 19S assembly in mammals. Recently, a protein responsible for the 19S-20S association has been identified -p28, which recruits the 19S and favours a specific 19S conformation necessary for the interaction to occur (Y. Lu et al., 2017). Besides p28, several other assembly chaperones are necessary for the 19S assembly, such as S5b, p27 and PAAF1 [START_REF] Krzywda | The Crystal Structure of Gankyrin, an Oncoprotein Found in Complexes with Cyclin-dependent Kinase 4, a 19 S Proteasomal ATPase Regulator, and the Tumor Suppressors Rb and p53[END_REF][START_REF] Roelofs | Chaperone-mediated pathway of proteasome regulatory particle assembly[END_REF][START_REF] Park | Assembly manual for the proteasome regulatory particle: the first draft[END_REF]. The second way of assembly is by using the 20S as the scaffold for the assembly of the AAA base, upon which the lid is constructed. The intermediates of this pathway have been observed in yeast (Park et al., 2013).

The components of the 19S regulator, with minor variations in number, are present throughout the eukaryotic branch of the tree of life. Furthermore, true orthologs and functional homologs are found throughout the archaea, showing that, from an evolutionary standpoint, the ancestors of the 19S are nearly as old as the proteasome [START_REF] Fort | Evolution of Proteasome Regulators in Eukaryotes[END_REF].

The first step in substrate digestion is the Ub recognition, mediated by the Ub receptors in the base -RPN1 [START_REF] Shi | Rpn1 provides adjacent receptor sites for substrate binding and deubiquitination by the proteasome[END_REF], RPN10 [START_REF] Van Nocker | The multiubiquitin-chain-binding protein Mcb1 is a component of the 26S proteasome in Saccharomyces cerevisiae and plays a nonessential, substrate-specific role in protein turnover[END_REF] and RPN13, bound via RPN2 [START_REF] Husnjak | Proteasome subunit Rpn13 is a novel ubiquitin receptor[END_REF]. In addition, the substrate can be delivered to the 26S via shuttle proteins which are capable of binding to the Ub via Ub-associated (UBA) domains and can be recognized by the 19S via their Ub-like (UBL) domains (Finley, 2009;[START_REF] Paraskevopoulos | Dss1 Is a 26S Proteasome Ubiquitin Receptor[END_REF]. The substrate N-or Cterminus [START_REF] Berko | The Direction of Protein Entry into the Proteasome Determines the Variety of Products and Depends on the Force Needed to Unfold Its Two Termini[END_REF] is then fed to the pore of the RPT engine (Dong et al., 2019). It is a prerequisite for the substrate to have a disordered N-terminus, long enough to reach the RPT pore, in order to be efficiently primed and degraded [START_REF] Fishbain | Sequence composition of disordered regions fine-tunes protein half-life[END_REF], or at least an internal disordered loop region that satisfies the same criteria [START_REF] Piwko | Proteasome-mediated protein processing by bidirectional degradation initiated from an internal site[END_REF]. In a sense, this is a regulation mechanism, whereby the proteins without disordered termini are protected from proteasomal degradation -their half-life is therefore longer than that of most proteins with a disordered N-or Cterminus. The Ub now must be cleaved from the substrate to trigger its unfolding. This is mediated by the DUB activity of RPN11 (Verma, 2002) or by an exterior DUB that does not belong to the 26S complex, such as USP14 (B.-H. Lee et al., 2016) or a DUB which interacts with the proteasome in a transitory manner, through 19S components, such as UCHL5 [START_REF] Deol | Proteasome-Bound UCH37/UCHL5 Debranches Ubiquitin Chains to Promote Degradation[END_REF]. In the resting state, only a few C-termini of the 19S base actually dock into the 20S α ring. Upon Ub-cleavage, several more of the C-termini of the 19S base subunits bury into the α-α pockets and induce the opening of the 20S pore, which in turn allows the unfolding of the substrate by the 19S to continue. Furthermore, the axes of the pores of the 19S and the 20S, which are not aligned in the resting state, now align, to allow for substrate passage (Matyskiela, Lander and Martin, 2013;Dong et al., 2019).

There are six ATP-binding pockets on the 19S base, one at every AAA ring subunit and they can exist in three different states: ATP-bound, ADP-bound and apo-like state (Dong et al., 2019) (Figure 11, right panel). Without the substrate, the AAA domain is in its most compact form, hydrolysing minimal amounts of ATP, about 27 ATPs per complex [START_REF] Hoffman | Nucleotidase Activities of the 26 S Proteasome and Its Regulatory Complex[END_REF]. Binding of the ATP, hydrolysis and then release of the ADP + P is coupled with the motion of the hinge of the AAA subunit along the substrate chain. The six subunits all work in concert, alternating between the three mentioned states, which effectively moves the peptide through the 20S proteasome pore and into the catalytic chamber (Dong et al., 2019). The proposed mechanism for ATPase ring operation is schematically presented on Figure 11, right panel. Interestingly, large side chain groups improve AAA subunit hinge annealing to the substrate backbone, while shorter sidechains on the substrate thread do not allow for a tight grip of the AAA hinges, rendering the translocation less efficient. This can serve as a mechanism of early substrate release and it has been observed to serve as a mechanism to activate a certain transcription factor, by removing the inhibiting propeptide sequence [START_REF] Tian | A conserved processing mechanism regulates the activity of transcription factors Cubitus interruptus and NF-κB', Nature Structural & of the 26S proteasome[END_REF].

It is considered that the majority of the proteasome-mediated protein degradation occurs via 26S and 30S proteasomes. Since the proteasomal degradation is involved in most cellular processes, this system, along with its diverse, far-reaching network of Ub ligases and other adjuvant proteins have been studied in-depth and deserve their own chapter.

PA28αβ

PA28αβ is a heteroheptamer with an α4β3 topology (Huber and Groll, 2017). The α and β subunits form a cone shaped ring (Figure 12, left) which adapts to the 20S proteasome and regulates substrate access. The protruding loops dock into the α-α pockets and unlock the 20S particle pore, in a manner that is similar, but not the same as with the 19S. PA28αβ can also form hybrid complexes with singlycapped 26S proteasomes [START_REF] Cascio | Preparation of hybrid (19S-20S-PA28) proteasome complexes and analysis of peptides generated during protein degradation[END_REF]. Its expression can be induced by the IFN-γ, a cytokine important in immune response, secreted by immune cells and induces immunoproteasome and macrophage activation [START_REF] Boehm | CELLULAR RESPONSES TO INTERFERON-γ[END_REF]. PA28αβ association with the immunoproteasome modulates the proteolytic peptide profile of the proteasome, namely by creating shorter peptides which are more acidic on their C-termini, and thereby augmenting the antigen presentation. Although the involvement of PA28αβ in the MHC presentation is evident, it is not yet clear how the different peptide profile contributes to the improved immune response [START_REF] Raule | Enhanced rate of degradation of basic proteins by 26S immunoproteasomes[END_REF]. Although both are induced in the immune response, the immunoproteasome and PA28αβ are able to function independently in the supply of antigenic peptides for MHC complexes [START_REF] Schwarz | The proteasome regulator PA28α/β can enhance antigen presentation without affecting 20S proteasome subunit composition[END_REF]de Graaf et al., 2011). From these observations, a question arises whether PA28αβ binds more strongly to the immunoproteasome compared to the standard proteasome. Although at first disputed [START_REF] Schmidtke | The 20S immunoproteasome and constitutive proteasome bind with the same affinity to PA28αβ and equally degrade FAT10[END_REF], the evidence of this preferential interaction was shown using protein correlation profiling, and affinity purification coupled to mass spectrometry [START_REF] Fabre | Deciphering preferential interactions within supramolecular protein complexes: the proteasome case[END_REF]. Together with the immunoproteasome, PA28αβ plays an important role in response to the oxidative stress (Pickering et al., 2010;Abi Habib et al., 2020), where it activates the 20S proteasome and helps degrade in an ATPindependent manner the unfolded proteins which accumulate as a result of the oxidative damage (Pickering and Davies, 2012). Evolutionarily, it appeared more recently than PA28γ, probably to specifically play a role in the immune system [START_REF] Fort | Evolution of Proteasome Regulators in Eukaryotes[END_REF].

PA28γ

This regulator is also a part of the PA28 regulator family. PA28γ is a homoheptamer (Figure 12, center) and appears to be evolutionarily older than the PA28αβ, with its sequence and structure closely resembling the orthologs found in other branches of life [START_REF] Fort | Evolution of Proteasome Regulators in Eukaryotes[END_REF]. The orthologs of PA28 have been identified in protozoans and showed similar behaviour (Xie et al., 2019). Like PA28αβ, PA28γ is a dome-like structure which caps the 20S particle by binding to the α ring interface, and unlocks the 20S pore via an activation loop able to docks into the α-α pockets. The inside of the PA28 regulators is lined with polar residues, which was proposed to play a role in substrate retention (Xie et al., 2019). Besides simply opening the pore of the 20S, PA28 activators induce allosteric changes throughout the 20S [START_REF] Chen | Cryo-EM of mammalian PA28αβ-iCP immunoproteasome reveals a distinct mechanism of proteasome activation by PA28αβ[END_REF]. Unlike PA28αβ, PA28γ does not seem to modify the profile of peptide produced by the 20S. It is postulated however, that the differences in the highly-flexible loops on the apical sides of the complex may play a role in the substrate selection [START_REF] Cascio | PA28γ: New Insights on an Ancient Proteasome Activator[END_REF]. PA28γ is not induced by the IFNγ, like PA28αβ or the immunoproteasome, but it seems to have a rather distinct set of roles.

The PA28γ is rather uniformly distributed across tissues [START_REF] Kim | A draft map of the human proteome[END_REF], but it is, however, predominantly localized in the nucleus [START_REF] Masson | Identification and Characterization of a DrosophilaNuclear Proteasome Regulator[END_REF], where it plays a major role in the organisation of the genetic material (Baldin et al., 2008;[START_REF] Zannini | REGγ/PA28γ proteasome activator interacts with PML and Chk2 and affects PML nuclear bodies number[END_REF]. Its association with the 20S proteasome in physiological state seems to be limited [START_REF] Fabre | Label-Free Quantitative Proteomics Reveals the Dynamics of Proteasome Complexes Composition and Stoichiometry in a Wide Range of Human Cell Lines[END_REF] and it rather serves as a stock for urgent response (Welk et al., 2016). Indeed, PA28γ association seems to be induced in case of oxidative stress (Abi Habib et al., 2020), where it improves the 20S's ability to degrade unfolded proteins (Pickering and Davies, 2012). PA28γ is a very important effector in the process of DNA repair, where it helps recruit proteasomes that remove proteins at the damage site (Levy- [START_REF] Levy-Barda | Involvement of the nuclear proteasome activator PA28γ in the cellular response to DNA double-strand breaks[END_REF]. It has been shown that PA28γ is an important regulator of the cell cycle [START_REF] Murata | Growth Retardation in Mice Lacking the Proteasome Activator PA28γ[END_REF][START_REF] Zannini | REGγ proteasome activator is involved in the maintenance of chromosomal stability[END_REF], by targeting the cell cycle regulators for degradation [START_REF] Li | Ubiquitin-and ATP-Independent Proteolytic Turnover of p21 by the REGγ-Proteasome Pathway[END_REF]. It also targets signalling proteins important in apoptosis (S. Liu et al., 2018) and consequentially plays an important role in many cancers. Indeed, PA28γ is involved in oncogenesis through several different pathways [START_REF] He | REGγ is associated with multiple oncogenic pathways in human cancers[END_REF]Ali et al., 2013;[START_REF] Moncsek | Evidence for anti-apoptotic roles of proteasome activator 28γ via inhibiting caspase activity[END_REF].

Together with PA200, PA28γ plays a role in spermatogenesis: double KO of PA200 and PA28γ was shown to be fully infertile, with greatly impaired motility of spermatozoa. Interestingly enough, the KO germ cells did suffer from high levels of oxidative damage, indicative of, but not limited to the role of PA28γ in removing the oxidation-damaged proteins (L. Huang et al., 2016). Recent work by Gao et al. described however that the PA28γ KO mice model is sub-fertile [START_REF] Gao | The REGγ-Proteasome Regulates Spermatogenesis Partially by P53-PLZF Signaling[END_REF], due to a reduced number of spermatogonia. They further unveiled a PA28γ -p53 -PLZF axis of cell proliferation control. The PLZF is a transcription repressor responsible for self-renewal of gonial cells. The p53 is a proteasome substrate, specifically targeted by the PA28γ and is a suppressor of the PLZF gene. Among others, it also regulates apoptosis. The absence of PA28γ thus results in p53 accumulation, which represses the PLZF expression, which in turn leads to spermatogonial apoptosis.

PA200

The Proteasome Activator 200 (PA200) is a monomer of 200kDa, asymmetric in shape and capable of binding to the 20S proteasome and opening the substrate channel. The majority of its sequence is composed of HEAT repeats (Kajava et al., 2004), forming an α-solenoid winding on itself and forming a pore in its centre (Figure 12, right). The N-terminus is opposite of the 20S binding face, which is followed by the ensemble of the HEAT repeats, while the C-terminus facing the α-ring of the 20S is structured and highly conserved throughout species [START_REF] Fort | Evolution of Proteasome Regulators in Eukaryotes[END_REF]. The C-terminus indeed contains a HbYX (hydrophobic-tyrosine-other) motif that binds into the α-α pockets and another α-α pocket anchoring point is formed by a protruding loop of atypical sequence. Two recent structural studies identified two inositol-binding pores/openings, with yet-unconfirmed purpose (Toste Rêgo and da Fonseca, 2019; Guan et al., 2020). PA200 contains atypical bromodomain-like domain (Guan et al., 2020), which has been shown to recognize acetylated proteins -a role well described in acetylated histone removal [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF]. Docking of the PA200 regulator induces both pore opening and a structural rearrangement throughout the 20S core (Toste Rêgo and da Fonseca, 2019), however, functional implications of these changes are not yet clear.

PA200 is an ATP-independent regulator involved in many cellular processes. Besides, it is involved in the process of mitochondrial fission [START_REF] Tar | Proteasomes Associated with the Blm10 Activator Protein Antagonize Mitochondrial Fission through Degradation of the Fission Protein Dnm1[END_REF] and oxidative stress response (Pickering and Davies, 2012). Of note are its roles in DNA damage repair and in spermatogenesis. The presence of a bromodomain-like domain in PA200 enables the recognition of acetylated histones. Indeed, PA200 has been shown to enhance the degradation of acetylated, but not Ub-ylated histones in vitro [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF]. This leads to an important Ub-independent degradation pathway, which seems to be activated in two cases: upon DNA damage response and for chromatin condensation in spermatogenesis [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF][START_REF] Mandemaker | DNA damage-induced replication stress results in PA200proteasome-mediated degradation of acetylated histones[END_REF]. What these two biological processes have in common is the need to remove histones from the DNA, which hinder the access of other intervening proteins, such as DNA repair machinery, transcription complexes or transitional histone isoforms. In spermatogenesis there are two particular events where the PA200 is recruited for such activity: the DSB repair on recombination sites during prophase I and the histone removal after meiosis. Histones are then to be replaced by transitional histone isoforms and in the end, by protamines, which compactify the chromatin (Wang et al., 2019). Failure to express PA200 leads to genetic instability, manifested as a reduced viability of the cells upon IR light-induced DNA damage. The IR light exposure also induced the formation of the hybrid 19S-20S-PA200 complex and its accumulation on the chromatin [START_REF] Blickwedehl | Role for proteasome activator PA200 and postglutamyl proteasome activity in genomic stability[END_REF]. The same study reports that a similar phenotype to PA200 loss can be induced by inhibition of caspase-like activity of the 20S. PA200 is predominantly localized in the nucleus, although during mitosis it can be found throughout the cell (Ustrell, 2002). Any kind of DNA damage (oxidative, UV or γ-rays) seems to induce PA200 to form nuclear foci -a localization pattern typical of DNA-repair proteins (Ustrell, 2002). Although the oxidative damage induces a typical DNA-repair response, PA200 itself does not seems to improve the degradation of oxidized proteins, suggesting that it responds indirectly, by acting via other proteins, to favour DNA repair (Pickering and Davies, 2012). Although PA200 expression and re-localization is a part of response to induced DNA damage, the physiologically-induced DNA double-strand breaks, such as lymphocyte development and immunoglobin class switching in B-cells, do not seem to require PA200 to occur properly [START_REF] Khor | Proteasome Activator PA200 Is Required for Normal Spermatogenesis[END_REF] -this further supports the thesis that the PA200 acts indirectly to favour DNA repair, through degradation of proteins at the DNA damage site.

PA200 is expressed across many different tissues in human, however its expression peaks in male and female gonads [START_REF] Kim | A draft map of the human proteome[END_REF]. The KO model for PA200 did not show any apparent phenotype for female mice, while males were subfertile. Among other problems, the male KOs display a lower sperm count and exhibit motility defects. On the histological level, KOs display normal spermatogonia, however, remarkable defects are observed in cells entering meiosis. Namely, pachytene spermatocytes were either abnormal or undergoing apoptosis or necrosis, while the spermatids were being massively phagocytised and reduced in number [START_REF] Khor | Proteasome Activator PA200 Is Required for Normal Spermatogenesis[END_REF]. It was then demonstrated that the PA200 deficient males showed inability to remove acetylated histones, which are targeted by PA200 for proteasomal degradation [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF]. In another study, double KOs for PA28γ and PA200 were completely infertile, principally due to lack of motility, but were able to complete the acrosomal reaction. Single KO of each of these proteins, although it had significant effects, did not induce complete infertility. Upon further investigation, it was concluded that the double KO germ cells suffered from substantial oxidative damage and displayed accumulation of Ubylated proteins. However, the PA200 and PA28γ can compensate for each other's absence, probably due to somewhat overlapping roles (L. Huang et al., 2016).

The association with the 20S (but not the expression) of both PA28γ and PA200 increases in response to 20S proteasome inhibition. Reversely, the specific inhibition of 26S induced both increased association and expression of PA200 (Welk et al., 2016). This observation shows how both PA28γ and PA200 regulators are involved in cellular response to stress and behave as a sort of intracellular protein stock of proteasome activators. Altogether, these two ATP-independent proteasome regulators seem to have indispensable, non-redundant roles in proteasome-mediated regulation of cellular processes.

Figure 12. The ATP-independent proteasome regulators. Left: the PA28αβ regulator is a heteroheptamer in α4β3 configuration. PA28γ (centre) is composed of seven identical subunits and is evolutionarily related to PA28αβ. The PA200 regulator is a 200 kDa monomer, which forms an α-solenoid winding on itself and forming a pore in its centre (potentially even two pores). It is markedly asymmetric. All three representations were made in UCSF ChimeraX, using 6KWX PDB entry for PA200 and the models of PA28αβ and PA28γ generated in [START_REF] Lesne | Conformational maps of human 20S proteasomes reveal PA28-and immuno-dependent inter-ring crosstalks[END_REF].

There seems to be a relation between the roles of PA200 and the spermatoproteasome in spermatogenesis. Namely, the PSMA8 KO mouse model (gene coding for α4s), besides being infertile, showed inability to degrade acetylated histones in the developing germ cells. Moreover, it was shown that in this model, unlike in the wild type, PA200 was completely absent from the surface of the chromosomes in pachytene due to drastically lower PA200 abundance on the protein level, suggesting that somehow the spermatoproteasome regulates the PA200 expression [START_REF]The PSMA8 subunit of the spermatoproteasome is essential for proper meiotic exit and mouse fertility[END_REF]. Another recent study of the PSMA8 KO finally confirmed the mutual role of the spermatoproteasome and PA200 in acetylated histone degradation, while also showing that the spermatoproteasome-PA200 complex is more efficient at degrading acetylated histones compared to its standard 20S-PA200 equivalent. Beyond this role, and in accord with the previously described roles of PA200, deletion of α4s suppressed the repair of the DNA double-strand breaks in metaphase I of spermatogenesisanother indicative of their shared role [START_REF] Zhang | Proteasome subunit α4s is essential for formation of spermatoproteasomes and histone degradation during meiotic DNA repair in spermatocytes[END_REF].

The ubiquitin-proteasome system (UPS)

As mentioned earlier, the proteasome is the main machinery for targeted proteolytic degradation. As such, it must have a large, hierarchically organized network of regulator proteins which spans across the proteome and is highly regulated at multiple levels. Protein half-life is defined in great part by the compactness of its structure and the presence of a disordered N-terminus or a specific N-terminal amino acid. Structurally-disordered proteins can be even degraded in an ATPindependent manner. Most proteins, however, are targeted for degradation via Ub (or Ub-like protein) tags. Ub is a 76-amino acid protein sequence, highly conserved and ubiquitously present in eukaryotes [START_REF] Goldstein | Isolation of a polypeptide that has lymphocyte-differentiating properties and is probably represented universally in living cells[END_REF]. Its C-terminus carries a glycine residue, which can be engaged in a covalent bond with the amine group of virtually any exposed lysine sidechain, or less commonly through Nterminal amine residue, hydroxy group of a serine or threonine, or thiol group of the serine [START_REF] Stewart | E2 enzymes: more than just middle men[END_REF]. Chains of multiple Ub can form to promote the degradation.

The ubiquitination system

The target protein is covalently modified with Ub via a cascade of three groups of enzymes: E1-3 (Figure 13). There are very few E1 enzymes [START_REF] Schulman | Ubiquitin-like protein activation by E1 enzymes: the apex for downstream signalling pathways[END_REF] in mammals and they are the Ub-activating enzymes that recruit the Ub moiety by covalently binding to it in an ATPdependant process. Besides Ub, there are several other Ub-like proteins that can be used by Ub-ligases in the process of signalling (see Figure 14). The E2 (ub-conjugating) enzymes are more numerous (around 40 in humans; [START_REF] Stewart | E2 enzymes: more than just middle men[END_REF]) and each E1 enzyme transfers the Ub over to a specific set of E2 enzymes. The E3 Ub ligases are able to recognise a specific set of substrates as well as a specific set of E2 ligases and transfer the Ub from the E2 ligases to its final recipient -the target substrate. They count hundreds of proteins [START_REF] Jackson | CRL4s: the CUL4-RING E3 ubiquitin ligases[END_REF] that can be divided in HECT and RING families, with a third hybrid family called RING-IBR-RING (RBR) [START_REF] Zheng | Ubiquitin ligases: Structure, function, and regulation[END_REF].

In the case or RING family of E3 ligases, the Ub tag is transferred directly from the E2 ligase to the substrate and the E3 complex is there to serve as a scaffold and to promote the transfer. The substrate polyUb-ylation is promoted through sequential Ub -transfers onto the existing Ub elements, where the residue through which the linking is occurring depends strongly on the E2 ligase specificity [START_REF] Schulman | Ubiquitin-like protein activation by E1 enzymes: the apex for downstream signalling pathways[END_REF]). In the case of HECT E3 ligases, a thioester intermediary of covalently bound Ub-E3 complex forms, before the transfer of Ub to the substrate [START_REF] Pickart | Mechanisms Underlying Ubiquitination[END_REF]. Certain types of E2 ligases can even transfer the Ub to the substrate on their own [START_REF] David | The E2 Ubiquitin-conjugating Enzymes Direct Polyubiquitination to Preferred Lysines[END_REF].

Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ubylation enzymes. E1 Ub-activating enzymes are very small in number, with each member targeting a specific group of E2 Ub-conjugating enzymes, which in turn have their own preferences for E3 ligases, which are very numerous -it is estimated that over 1000 distinct E3 ligase exist in humans [START_REF] Jackson | CRL4s: the CUL4-RING E3 ubiquitin ligases[END_REF]. The Ub moieties are transferred down the hierarchy to the E3 ligases -where each E3 type complex can incorporate dozens of different substrate-recognizing adaptors. After the first Ub is transferred to the substrate, the chain can be elongated by the E2-E3 ligase complex, to amplify the degradation signal or further modified with various other complexes. This process is balanced with the activity of the de-ubiquitination enzymes (DUBs), which can shorten or completely remove the Ub chains. Not all Ub modification is a degradation signal -activity of many enzymes is regulated in this way or a protein affinity for a certain interactor can be altered. The substrate is then recognized by the 26S or 30S proteasome. On the schema, the 26S proteasome is presented. The 20S core can be capped from the other side by any other proteasome regulator. The substrate recognition can be aided by a shuttle protein. The Ub is cleaved away from the substrate by an intrinsic 19S DUB activity and recycled for further use by the E1 ligases, while the protein is degraded into peptides.

Two most studied groups of E3 complexes (and most relevant in the context of this thesis) come from a RING E3 ligase family: the cullin-RING ligase (CRLs) and the anaphase-promoting complex/cyclosome (APC/C). These two complexes introduce Ub chains on many cell cycle regulators and mark them for degradation, such as Aurora A and B, cyclin A, B, and E, Cdk-inhibitory kinase Wee1, Cdc25 phosphatase and many other cell division regulators [START_REF] Nakayama | Ubiquitin ligases: cell-cycle control and cancer[END_REF][START_REF] Craney | Dynamic regulation of ubiquitin-dependent cell cycle control[END_REF]. Thereby, these two E3 ligase complexes, very similar in structure, are crucial for cell cycle regulation. They serve as platforms which are able to host numerous different proteins that are responsible for substrate specificity (example on the Figure 13). The CRL family's receptor fund is huge and enables targeting of vast number of substrates: CRL1 has 69 human F-box proteins, CRL3 -circa 100 BTB proteins and CRL4 -circa 60 WD40-containing DCAFs. Based on the need of the cell, these proteins are exchanged dynamically, through a very clever system of disassembly protein that has high affinity for assembled complex but a low affinity for a disassembled CRL [START_REF] Craney | Dynamic regulation of ubiquitin-dependent cell cycle control[END_REF]. The APC is crucial for metaphase to anaphase transition -a complex network of signalling molecules called spindle assembly checkpoint, inhibits the APC from operation, until all chromosomes have achieved attachment [START_REF] Craney | Dynamic regulation of ubiquitin-dependent cell cycle control[END_REF]. Interestingly, these two complexes exert their activity in the separate times of the cell cycle: APC from early M to the end of the G1 phase (cell division and quiescent period), while a particular CRL (SKP1-CUL1-F-box) complex has prominent substrates from the end of G1 phase, through the S phase (DNA replication), to the end of G2 phase (chromosome condensation) [START_REF] Nakayama | Ubiquitin ligases: cell-cycle control and cancer[END_REF]).

The Ubiquitin as a signal

Many substrates are Ub-ylated on several sites and Ub itself can be conjugated with other Ub's via several exposed lysine sites (K6, K11, K27, K29, K33, K48, K63) yielding Ub chains of different structural properties or even branched chains [START_REF] Yau | Assembly and Function of Heterotypic Ubiquitin Chains in Cell-Cycle and Protein Quality Control[END_REF]. In parallel to the polymerisation, the DUBs may exert their activity to trim the Ub chains (Figure 13) [START_REF] Komander | Breaking the chains: structure and function of the deubiquitinases[END_REF]. The positions and number of Ub on the protein, the length and branchedness of the poly-Ub chains represent a code which is interpreted by the ensemble of enzymes (Figure 14) [START_REF] Oh | Principles of Ubiquitin-Dependent Signaling[END_REF]. The interpretation of the code will depend on the topology of the Ub chain (binding sites) and its structural features, such as flexibility or compactness [START_REF] Komander | The Ubiquitin Code[END_REF]. The K48 link chains are the most compact and also are crucial for proteasomal degradation of Ub-ylated substrates, however, other linkages can be recognized by the proteasome, such as K11, though to a lesser degree. The K11 linkage type chains were found to be particularly important for degradation of cell cycle regulators, notably the APC substrates [START_REF] Matsumoto | K11-Linked Polyubiquitination in Cell Cycle Control Revealed by a K11 Linkage-Specific Antibody[END_REF]. It was demonstrated that the lysosomal degradation of a substrate can be triggered through the K63-linked chains -an interesting example on non-proteasomal, Ub-dependent proteolysis [START_REF] Mukhopadhyay | Proteasome-Independent Functions of Ubiquitin in Endocytosis and Signaling[END_REF]. Finally, branched chains are yet poorly understood, but for now seem to amplify the signal for degradation through proteasome [START_REF] Meyer | Enhanced Protein Degradation by Branched Ubiquitin Chains[END_REF].

However, the Ub tags do not necessarily lead to degradation. Often, they serve a purpose of protein-protein interaction modulation, enzymatic activity regulation, or even the regulation of localization in the cell [START_REF] Welchman | Ubiquitin and ubiquitin-like proteins as multifunctional signals[END_REF]. Roles in numerous processes were assigned to the Ub chains that do not induce proteasome recruitment: the Methionine1-linked Ub, together with the K63-linked Ub, is involved in NF-κB signalling through mixed type chains (which can be branched), the K6-linked chains in mitochondrial homeostasis and the K33 linked Ub chains -in intracellular protein trafficking [START_REF] Swatek | Ubiquitin modifications[END_REF]).

An entire family of Ub-like molecules exist, very similar in structure to the Ub itself, which can be conjugated with the Ub-chains or on their own, through Ub ligase system (Dohmen, Huibregtse and Scheffner, 2016; Cappadocia and Lima, 2018), among which the SUMO1-4 proteins, NEDD-8, ISG15, LC3 (ATG8) and ATG12 are the best characterized. In that context, it is more appropriate to speak about the forementioned systems as the Ub-like regulation, rather than Ub regulation.

Other proteins involved in the UPS network

Many proteins are involved in substrate delivery and processing, which are neither part of the ubiquitination system nor the intrinsic DUBs. Among them are the shuttle proteins that are capable of recognizing the substrates and being recognized by the constitutive 19S subunits. As already mentioned, this is achieved via the Ub-associating (UBA) and Ub-like (UBL) domains. Many such proteins exist in humans: HR23A and B (orthologs of the RAD23A and B in yeast), UBQLN1, 2, 3 and 4, DDI1 and 2, UBAC1, UB7 and NUB1L. These proteins have different affinities towards the 19S intrinsic Ub receptors, moreover, the shuttles themselves allow the proteasome to target a greater variety of Ub branches and linkages. Thereby, the shuttles greatly expand the field of potential substrates that may not be recognized by the 26S proteasome alone. Many proteins are being identified that cannot be simply put in the UBA-UBL shuttle group, but are able to act as shuttle factors or even recognize intrinsically disordered proteins that are not Ub-ylated and introduce them to the proteasome. Others (such as RAD23) even act as protectors from DUB activity, by binding to Ub-ylated proteins, and ""escorting them to the proteasome [START_REF] Richly | A Series of Ubiquitin Binding Factors Connects CDC48/p97 to Substrate Multiubiquitylation and Proteasomal Targeting[END_REF]Tsuchiya et al., 2017). Many extrinsic DUBs are capable of binding reversibly to the proteasome, improving the Ub-cleavage, and regulating substrate specificity: e.g. USP14 binds to RPN1 [START_REF] Borodovsky | A novel active site-directed probe specific for deubiquitylating enzymes reveals proteasome association of USP14[END_REF] and UCH37 to RPN13 [START_REF] Sahtoe | Mechanism of UCH-L5 Activation and Inhibition by DEUBAD Domains in RPN13 and INO80G[END_REF].

UPS in health and disease

The importance of proper UPS functioning for cell (and organism-level) homeostasis cannot be overstated. Many diseases have been described that involve UPS impairment, such as neurodegenerative diseases [START_REF] Hegde | Perturbations of Ubiquitin-Proteasome-Mediated Proteolysis in Aging and Alzheimer's Disease[END_REF] and many types of cancer [START_REF] Baloghova | Ubiquitin Ligases Involved in the Regulation of Wnt, TGF-β, and Notch Signaling Pathways and Their Roles in Mouse Development and Homeostasis[END_REF]Wang et al., 2020). As already mentioned, the 26S proteasome regulates the NF-κB maturation by cleavage and mediates the degradation of the transcription activators of this pathway (Kravtsova-Ivantsiv, Cohen and Ciechanover, 2009) that regulates many cellular processes, such as adaptive immunity, inflammation, stress response and B-cell development. Aggregation of proteins in neurons leads to dysfunction of the cells and finally their death. Thus, maintaining proper clearance of the misfolded proteins is imperative for neuronal function. There is accumulating evidence that defects in UPS can lead to development of neurodegenerative diseases [START_REF] Hegde | Perturbations of Ubiquitin-Proteasome-Mediated Proteolysis in Aging and Alzheimer's Disease[END_REF]. Moreover, it has been found that the UPS has an important role in neuronal plasticity and memory [START_REF] Djakovic | Phosphorylation of Rpt6 Regulates Synaptic Strength in Hippocampal Neurons[END_REF]. Reduced proteasomal activity is known to lead to age-related diseases. Likewise, processes that induce autophagy and induce proteasome activation in general are known to slow down the onset of age-related symptoms [START_REF] Saez | The Mechanistic Links Between Proteasome Activity, Aging and Agerelated Diseases[END_REF]VerPlank et al., 2020). Many cyclins are under the control of the UPS and their dysregulation is known to induce malignancies. Moreover, many cancer cells rely on high protein turnover through the UPS. It is thus no surprise that the UPS is a common target of cancer therapy, not only via the 20S, but also the 19S subunits (Anchoori et al., 2018) and shuttle proteins such as UCH37 (X. Lu et al., 2017). Several 20S-targetting drugs have been developed for cancer treatment, such as bortezomib, carfilzomib and ixazomib mostly used to treat multiple myeloma (Scott et al., 2016;[START_REF] Richardson | Ixazomib for the treatment of multiple myeloma[END_REF][START_REF] Groen | Carfilzomib for relapsed and refractory multiple myeloma[END_REF]. Although revolutionary in multiple myeloma treatment, the issue with these drugs in that they target the entire UPS system and essentially block all proteasome-related activity. Although this treatment has proved effective in anti-proliferative activity, the side effects are not negligible, since UPS inhibition leads to toxic accumulation of misfolded protein [START_REF] Hou | Bortezomib treatment causes long-term testicular dysfunction in young male mice[END_REF]Tundo et al., 2020). Thus, a more targeted approach is envisioned, whereby a particular Ub ligase or a subset of ligases is selectively targeted, in an effort to reduce adverse effects when treating cancers [START_REF] Bielskienė | E3 ubiquitin ligases as drug targets and prognostic biomarkers in melanoma[END_REF]. Literature on proteasome involvement in disease and as a potential drug target for diseases treatment is vast and goes beyond the scope of this thesis; for more information on this topic, the reader is referred to a great review by Tundo et al. (Tundo et al., 2020).

UPS in spermatogenesis

Spermatogenesis is a process by which permanently dividing stem cells called spermatogonia differentiate into spermatocytes, then spermatids, which then further develop to become fully functional spermatozoids [START_REF] Holstein | Understanding spermatogenesis is a prerequisite for treatment[END_REF]. This process will initially require the cell to go through meiosis, whereby recombination and equal distribution of genetic material must occur, followed by a tremendous reorganization of the cellular morphology and genome. Then the differentiating cell goes through spermiogenesis, whereby it becomes polarized, with a head and a tail, losing most of its cytoplasmic material. Throughout this process, the genetic material becomes more densely packed and less transcriptionally active, with bulky histones being replaced by smaller protamines, allowing a tighter DNA packaging [START_REF] Rathke | Chromatin dynamics during spermiogenesis[END_REF]Wang et al., 2019). For this to occur successfully, many new proteins must be synthetized, while the others must be removed, namely the ones that served their purpose or are damaged. The translational process must be constantly regulated by histone modification, until finally the histones themselves are removed. Although it has been clear for a long time that the UPS is essential in the coordination of this process, the exact particularities of the UPS pathways by which the differentiation is controlled are only beginning to be unveiled. Throughout human tissues, the expression of the 26S proteasome genes are highest in the gonads [START_REF] Kim | A draft map of the human proteome[END_REF], suggestive of the high necessity for efficient protein degradation.

UPS is involved in differentiation signalling

The spermatogonia expresses Stra8 -a marker of differentiation, when stimulated by retinoic acid [START_REF] Oulad-Abdelghani | Characterization of a premeiotic germ cell-specific cytoplasmic protein encoded by Stra8, a novel retinoic acid-responsive gene[END_REF]. If, however, the proteasome activity is inhibited, this differentiation signal will lack [START_REF] Manku | Expression of the Ubiquitin Proteasome System in Neonatal Rat Gonocytes and Spermatogonia: Role in Gonocyte Differentiation1[END_REF]. Another candidate for a germ cell differentiation regulator is Uba6 -an E1 ligase highly expressed in germ cells, exactly at the point of transition from mitosis to meiosis [START_REF] Hogarth | Identification and Expression of Potential Regulators of the Mammalian Mitotic-to-Meiotic Transition1[END_REF], however, its exact role remains to be elucidated. Another Ub ligase has been identified as crucial for germ cell differentiation -βTrCP (which is one of the substrate recognition proteins for Skp1-Cul1-F-box (SCF) ubiquitin ligase family -see "The Ubiquitination system" section on cullin-RING ligases). The loss of this protein stops spermatogenesis; however, this defect was reversed upon removal of Snail1-a substrate of SCF βTrCP -a fine example of the importance of the specific action of Ub ligases (Kanarek et al., 2010). The cell cycle, including meiosis [START_REF] Futcher | Cyclins in Meiosis: Lost in Translation[END_REF], is regulated by the signalling molecules called cyclins. Their degradation is regulated through ubiquitination, orchestrated by the APC/C complex (Golan, Yudkovsky and Hershko, 2002; Meyer and Rape, 2014) and is required for the cell cycle transitions.

DNA repair and histone removal

For the DSB to occur and a proper recombination event to take place, it is necessary for the histones to be modified or removed and entire ensemble of the DNA repair machinery to be assembled to the DSB site. This process seems to be, at least in part, mediated by the UPS. The 26S has been shown to be recruited to the sites of chemically induced DSBs and it is possible that the 26S responds in the same way for DSBs induced in other biological contexts -physiological or nonphysiological [START_REF] Krogan | Proteasome Involvement in the Repair of DNA Double-Strand Breaks[END_REF]. The deletion of one of the poly-Ub genes -Ubb in mice, arrests the spermatogenesis at the prophase of meiosis I, indicating that this is a particularly Ub-demanding step. In this KO model, there was a marked lack of γH2AX modification on histones -a mark of the DSBs and homologous recombination [START_REF] Rogakou | DNA Double-stranded Breaks Induce Histone H2AX Phosphorylation on Serine 139[END_REF]. The H2A histone was, however, Ub-ylated, although its abundance was far lower than in wild type mice [START_REF] Ryu | The Mouse Polyubiquitin Gene Ubb Is Essential for Meiotic Progression[END_REF]. The role of proteasome in meiotic recombination was demonstrated with studies on two orthologous Ub ligases, Rad6 and Hr6a/Hr6b, in yeast and mammals, respectively. Mutation or Rad6 in yeast leads to impaired meiosis I and a decreased number of DSBs in these cells [START_REF] Game | The role of RAD6 in recombinational repair, checkpoints and meiosis via histone modification[END_REF]). It appears that Rad6 works in accordance with Bre1 -another Ub-ligase, to mediate the Ub-ylation of H2B histones [START_REF] Yamashita | Rad6-Bre1-mediated histone H2B ubiquitylation modulates the formation of double-strand breaks during meiosis[END_REF]. In mice, Hr6b deletion has induced longer synaptonemal complexes -the connections between chromosomes in course of recombination. The number of recombination events in this KO model was remarkably increased (Baarends et al., 2003). Recent paper by Jasvinder et al. showed that the 26S proteasome is indeed recruited to the synaptonemal complex during meiosis and is essential for recombination in yeast. The recruitment was found to be mediated by an E3 ligase called Zip1 -an orthologue of the mammalian RNF212 [START_REF] Ahuja | Control of meiotic pairing and recombination by chromosomally tethered 26S proteasome[END_REF].

Related to DSBs, in the spermatogenesis context, is the formation of the synaptonemal complexes, since they facilitate the crossovers that are the consequence of the physiologically induced DSBs (see Chapter 2.1.4.1.1 Events of the prophase I). Recent study showed proteasome localization on these complexes (Gómez-H et al., 2019), which correlates interestingly with another study, which described that a loss of the E3 ligase called Ubr2 leads to a complete absence or heavy impairment of the synaptonemal complexes in spermatocytes, while the spermatogonia seemed intact [START_REF] Kwon | Female Lethality and Apoptosis of Spermatocytes in Mice Lacking the UBR2 Ubiquitin Ligase of the N-End Rule Pathway[END_REF].

UBC4 is an E2 Ub-ligase expressed particularly in round and elongated spermatids, where it was found to be particularly active [START_REF] Rajapurohitam | Activation of a UBC4-Dependent Pathway of Ubiquitin Conjugation during Postnatal Development of the Rat Testis[END_REF] and another isoform of this protein -UBC8A, is specific to the testis [START_REF] Wing | A novel rat homolog of the Saccharomyces cerevisiae ubiquitinconjugating enzymes UBC4 and UBC5 with distinct biochemical features is induced during spermatogenesis[END_REF]. An E3 ligase called Huwe1 was identified to interact specifically with the UBC4 E2 ligase, whereby its activity is entirely dependent on UBC4 and its isoforms. Most importantly, this HECT domain-containing E3 ligase is able to Ub-ylate all the histone subtypes in vitro [START_REF] Liu | Characterization of E3Histone, a novel testis ubiquitin protein ligase which ubiquitinates histones[END_REF]. Huwe1 is expressed in the cytoplasm throughout tissues, however, it is highly expressed in the nuclei of spermatogonia and mid-pachytene spermatocytes. Its presence was not detected in the spermatids, where the histone removal takes place, indicating that this ligase may have a role in chromatin remodelling in early differentiation stages of the male germ cells [START_REF] Liu | Regulated expression of the ubiquitin protein ligase, E3Histone/LASU1/Mule/ARF-BP1/HUWE1, during spermatogenesis[END_REF]. For the correct sperm development, DNA repackaging must occur, whereby histones are replaced, first by the intermediary proteins, then by protamines (Wang et al., 2019). An initial step in this process seems to be carried out by the E3 ligase called Rnf8 that mediates histone Ub-ylation. The KO model for Rnf8 is unable to remove histones during spermatogenesis [START_REF] Lu | RNF8-Dependent Histone Modifications Regulate Nucleosome Removal during Spermatogenesis[END_REF], which fits well together with the known roles of the Rnf8 -namely the coordination of DNA repair through multiple pathways [START_REF] Kolas | Orchestration of the DNA-Damage Response by the RNF8 Ubiquitin Ligase[END_REF]. UPS activity may be mixed with the PA200 -an ATP-independent proteasome regulator, described in the previous section. It was already mentioned earlier that the alternative proteasome regulator PA200 plays a great role in DNA repair and spermatogenesis (Ustrell, 2002; [START_REF] Khor | Proteasome Activator PA200 Is Required for Normal Spermatogenesis[END_REF][START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF]. It is uncertain, however, if the Ub-mediated proteasomal pathways act separately from the alternative proteasome activators in this process. Some evidence for the existence of mixed complexes (20S core with 19S and PA200 activator on each side) was put forward [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF][START_REF] Zhang | Proteasome subunit α4s is essential for formation of spermatoproteasomes and histone degradation during meiotic DNA repair in spermatocytes[END_REF] and the implications of this discovery have yet to be elucidated.

Taken together, these studies begin to reveal the extent of the intricate UPS network that coordinates germ cell differentiation. It appears that the hotspot of proteasomal activity revolves around the recombination event, which employs great amounts of Ub and recruits proteasome complexes. The implications of the spermatoproteasome -a testis-specific proteasome (see Chapter 2.2.6) in these events is yet to be revealed and will be further discussed.

PI31

PI31 is a proteasome-interacting protein that is not an activator per se, but seems to modulate differently the proteasome activity based on the biochemical/biological context. It is a 30 kDa protein consisting of principally two domains (Figure 15A). The N-terminal side contains an FP domain capable of homo-and hetero-dimerization ("FP" stands for "Fbxo7-PI31" domain) [START_REF] Kirk | Structure of a Conserved Dimerization Domain within the F-box Protein Fbxo7 and the PI31 Proteasome Inhibitor[END_REF][START_REF] Shang | Structure of the FP domain of Fbxo7 reveals a novel mode of proteinprotein interaction[END_REF]. On the C-terminal side, the proline-rich (PR) domain seems to be less structured and interacts with the proteasome via multiple regions. On the very C-terminus of PI31 there is an HbYX motif, similar to the motifs found in several proteasome activators, capable of binding into the α-α pockets and activating the proteasome (Figure 15B). Although the motif itself indeed activates the 20S, PI31 inhibits 20S activity and it does so via its PR domain, even in the absence of the HbYX motif, indicative of multiple regions of interaction with the 20S [START_REF] Bader | A conserved F box regulatory complex controls proteasome activity in Drosophila[END_REF][START_REF] Li | Molecular and Cellular Roles of PI31 (PSMF1) Protein in Regulation of Proteasome Function[END_REF][START_REF] Yashiroda | N-Terminal α7 Deletion of the Proteasome 20S Core Particle Substitutes for Yeast PI31 Function[END_REF]. Furthermore, PI31 seems to inhibit the assembly of the 26S in vitro [START_REF] Li | Molecular and Cellular Roles of PI31 (PSMF1) Protein in Regulation of Proteasome Function[END_REF] as well as the 20S-PA28 association [START_REF] Mccutchen-Maloney | cDNA Cloning, Expression, and Functional Characterization of PI31, a Proline-rich Inhibitor of the Proteasome[END_REF]. Interestingly, this behaviour can be regulated by ADP-ribosylation: this PTM prevents PI31 from inhibiting the 20S activity and seems to promote 26S assembly in vivo (Cho-Park and Steller, 2013), showing intricate modes of regulation of PI31 activity. On the other hand, both PI31 and its HbYX motif alone were shown to increase the 26S activity [START_REF] Bader | A conserved F box regulatory complex controls proteasome activity in Drosophila[END_REF]; however, this observation seems to be replicated with moderate success [START_REF] Li | Molecular and Cellular Roles of PI31 (PSMF1) Protein in Regulation of Proteasome Function[END_REF][START_REF] Yashiroda | N-Terminal α7 Deletion of the Proteasome 20S Core Particle Substitutes for Yeast PI31 Function[END_REF].

The role of the PI31 is not an easy one to study, since the KO models exhibit lethality in early development. Mice die perinatally, while the PI31 KOs of D. melanogaster die as larvae [START_REF] Bader | A conserved F box regulatory complex controls proteasome activity in Drosophila[END_REF]Minis et al., 2019). Conditional KO permitted a study in mice and showed that PI31 was crucial in maintaining the neuronal homeostasis, more particularly in regulating the proteasome transport in axons and dendrites, effectively managing proteotoxic stress in the neurons (Minis et al., 2019). It was further shown that PI31 serves as an adaptor protein for proteasome that is recruited by the dynein light chain proteins (dDYNLL1/2) and transported along the axons. The abrogation of PI31 results in failure to recruit proteasome and toxic accumulation of protein aggregates. It appears that the signal triggering PI31 binding to dDYNLL1/2 is a phosphorylation at a single serine residue (Ser168 in mice and Ser153 in human) (K. Liu et al., 2019).

The FP domain of the N-terminal side of PI31 dimerizes with a very similar domain found in Fbxo7 -part of the SCF E3 ligase complex (Kirk et 15A). Although very similar in structure, the FP domains from these two proteins seem to each have a unique way of binding into a homodimer, but have a specific way of heterodimerization [START_REF] Shang | The FP domains of PI31 and Fbxo7 have the same protein fold but very different modes of protein-protein interaction[END_REF], which was shown to increase PI31 stability in vivo [START_REF] Bader | A conserved F box regulatory complex controls proteasome activity in Drosophila[END_REF][START_REF] Rathje | A Conserved Requirement for Fbxo7 During Male Germ Cell Cytoplasmic Remodeling[END_REF]. Both of these proteins play crucial roles during spermatogenesis [START_REF] Bader | A conserved F box regulatory complex controls proteasome activity in Drosophila[END_REF][START_REF] Rathje | A Conserved Requirement for Fbxo7 During Male Germ Cell Cytoplasmic Remodeling[END_REF]. In D. melanogaster, PI31 seems to be highly expressed during the individualization process of germ cells, whereby the excess cytoplasm and its contents are ejected to enable germ cells to reach full maturity. PI31 colocalizes with Fbxo7 in the cellular structures whose role is to manage the excess cytoplasm contents, indicative of their potential mutual role in this process. Tissue-specific KO model for PI31 seemed unable to undergo meiosis. Proteins important in cell cycle, which are normally degraded by the proteasome, such as cyclin B, accumulated in the KO germ cells. Moreover, aggregation of Ub-ylated proteins occurred, indicating strong proteasome activity impairment caused by the PI31 KO [START_REF] Bader | A conserved F box regulatory complex controls proteasome activity in Drosophila[END_REF].

Fbxo7 is a subunit of the SCF Ub E3 ligase complex (Figure 15B) and plays a role in substrate recognition. As such, it has a broad spectrum of functions, including cell cycle regulation [START_REF] Laman | Transforming activity of Fbxo7 is mediated specifically through regulation of cyclin D/cdk6[END_REF]. The KO model for Fbxo7 is completely sterile, as the gamete development arrests at round spermatid stage. As already mentioned, Fbxo7 ablation leads to reduction in PI31 protein, but not mRNA levels, indicative of structure stabilization between these two proteins [START_REF] Rathje | A Conserved Requirement for Fbxo7 During Male Germ Cell Cytoplasmic Remodeling[END_REF], meaning that the Fbxo7 protects PI31 from degradation. The relation between Fbxo7 and PI31 in proteasome regulation remained elusive until recent in vivo studies, probably because PI31 and Fbxo7 interact strictly in context-dependent ways and thus cannot be easily studied together in cell culture, as suggested in [START_REF] Nelson | Beyond ubiquitination: the atypical functions of Fbxo7 and other F-box proteins[END_REF]. The question arises, whether this behaviour is regulated by PTMs, as observed earlier with PI31. These two proteins seem to be evolutionary cousins, sharing two FP and PR domains (Figure 15A). However, Fbxo7 has several more domains in charge of complex formation, namely, F-box region that binds Skp1 (an E3 ligase component), a cdk-binding domain (responsible for its interaction with cell-cycle regulators) and an Ub-like domain with yet unclear role, but potentially involved in proteasome regulation [START_REF] Swatek | Ubiquitin modifications[END_REF]. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. A: domain organisation of PI31 and Fbxo7. The two proteins dimerize via their FP domain, while the proline rich domain in PI31 binds to the 20S proteasome. Fbxo7 also contains domains that enable it to bind to other proteins: the F box domain enables it to participate in an E3 ligase complex, cdk domain which interacts with cell cycle regulators and a Ub-like domain. Adapted from [START_REF] Shang | The FP domains of PI31 and Fbxo7 have the same protein fold but very different modes of protein-protein interaction[END_REF]. B: Proposed model of PI31/Fbxo7mediated proteasome regulation. PI31 acts as a linker between the 26S proteasome and Fbxo7, which is a part of an E3 ligase. Together, they may make up a system for degradation of proteins crucial for the progression of spermatogenesis. Adapted from [START_REF] Bader | A conserved F box regulatory complex controls proteasome activity in Drosophila[END_REF].

Spermatoproteasome

The discovery of the spermatoproteasome began with the identification of duplicated proteasome genes in Drosophila, observed to be expressed only in testes. It was established that 10 additional 20S proteasome subunits and 6 additional 19S subunits existed and were expressed in the testes [START_REF] Belote | Duplicated proteasome subunit genes in Drosophila and their roles in spermatogenesis[END_REF]. Upon further investigation it was established that α6T is essential for sperm development in Drosophila [START_REF] Zhong | The testis-specific proteasome subunit Pros 6T of D. melanogaster is required for individualization and nuclear maturation during spermatogenesis[END_REF]. Namely, the α6T would become prominent in the spermatids after meiosis and persist in the mature sperm. The KO model for α6T was completely infertile with irregularities in sperm individualization and nuclear maturation.

The first identification of an ortholog in mammals was detected in bovine seminiferous tubules and the 20S particle showed impaired ability to degrade Ub-ylated substrate, when compared to muscle or spleen proteasome [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF]. Detailed investigation of the testis-specific proteasome in mice by Uechi et al. determined that an alternative proteasome subunit, similar in sequence to α4 was expressed in the testes, particularly in gamete cells, from early pachytene, onwards into the spermatid stage and they named it α4s (Uechi, Hamazaki and Murata, 2014). It was then established that α4s replaces α4 in the 20S proteasome complex and that their existence is mutually exclusive. Sequence comparison of the two subunit isoforms in humans and mice revealed high homology, with differences concentrated in the C-terminal region (Figure 15, right), which, in α4, is found at the surface of the proteasome, oriented sideways, towards the solvent. Then proposed structural model of α4s can be seen on Figure 15, left panel. Since α4s is part of the α ring, with differences concentrated to the C-terminal region, it was not expected that it would alter the catalytic activity of the proteasome core, but would rather modify its interaction with regulators. Indeed, the activity assays on the recombinantly produced tagged, standard and spermato-20S particles showed no apparent difference in performance. Interestingly, the Ub-ylated protein removal was shown to be spermatoproteasome-dependent, indicating a probable involvement of the 26S in the process (Uechi, Hamazaki and Murata, 2014).

Figure 16. The spermatoproteasome sequence and model. Left: the alternative proteasome subunit α4s (yellow and red) is modelled onto the existing standard proteasome structure (gray). The differences between the standard proteasome and the spermatoproteasome-specific isoform are concentrated in the two outwardprotruding helices, here coloured in red. Model was generated using Swiss-Model platform, based on the 1IRU PDB entry and displayed in UCSF ChimeraX. Right: the sequence alignment of the bovine α4 and α4s show that the differences between the two are confined to two small regions towards the C-terminal. Alignment was done using ClustalW algorithm and Uniprot database canonical protein sequence entries.

Following the initial study by Uechi et al., it was established that α4s was necessary for proper germ cell maturation in mice: the double KO of the PSMA8 gene, encoding for α4s, renders the male mice infertile, while the female mice seem to be unaffected (Gómez-H et al., 2019; Q. [START_REF] Zhang | Meiosis I progression in spermatogenesis requires a type of testis-specific 20S core proteasome[END_REF]. Moreover, the spermatoproteasome was shown to localize along the synaptonemal complex and cells with deficient synaptonemal complex also seem to have a dispersed spermatoproteasome profile. Unexpectedly, the PSMA8 KO seems to be proficient in the homologous recombination and in the ability to assemble and disassemble the synapsis. It appears that the residual activity of the standard proteasome is somewhat able to compensate for the loss of α4s. However, developing germ cells go into apoptosis at either metaphase I or II (see Chapter 2.1.4.1.2. Events after the prophase I, section of the Spermatogenesis chapter). It was further shown that the spermatoproteasome interacts with PA200 and that without α4s, the PA200 almost completely lost association with the 20S in germ cells and even decreased in expression. Furthermore, there was a great accumulation of acetylated histones upon PSMA8 KO. These findings taken together show that the spermatoproteasome and PA200 work together to target histones, thereby modulating the chromatin organisation in developing germ cells. The effect of the spermatoproteasome and PA200 on histone removal could not be assessed, since germ cells go into apoptosis well before the programmed histone replacement (Gómez-H et al., 2019; Q. [START_REF] Zhang | Meiosis I progression in spermatogenesis requires a type of testis-specific 20S core proteasome[END_REF]. However, the role of the spermatoproteasome does not seem to stop there.

Besides PA200, the spermatoproteasome seems to interact with the SYCP3 and SYCP1 synaptonemal proteins and various cell cycle-related proteins. It is expected that without spermatoproteasome, accumulation of the signal proteins CDK1 and TRIP13 inhibits the transition from metaphase to anaphase, thus arresting cell progression (Gómez-H et al., 2019). Indeed, some proteins are stabilized upon α4s deletion, as it is the case with Rad51 -a homologous recombination factor, as confirmed in a recent study by Gómez-H et al. Furthermore, the PA28γ was shown to be highly expressed in the testes [START_REF] Kim | A draft map of the human proteome[END_REF] and also very important in male fertility (L. Huang et al., 2016); mainly by regulating cell cycle proteins through proteasome activity [START_REF] Gao | The REGγ-Proteasome Regulates Spermatogenesis Partially by P53-PLZF Signaling[END_REF]. PA28αβ seems to be completely absent from the spermatoproteasome complexes [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF], as expected, due to its particular role in immunity.

Although Gómez-H et al. did not detect impairments in the recombination performance, there was a noticeable impairment in the DSB repair in metaphase I of the α4s KO, further pointing to the suspected role of the spermatoproteasome in this process. It was further confirmed by in vitro assays that the spermatoproteasome was more efficient at degrading acetylated histones when activated by PA200, compared to its standard counterpart [START_REF] Zhang | Proteasome subunit α4s is essential for formation of spermatoproteasomes and histone degradation during meiotic DNA repair in spermatocytes[END_REF].

Finally, we can summarize that three distinct roles of the spermatoproteasome have been identified so far: 1) the degradation of cell cycle-mediating proteins, through the UPS network or via alternative proteasome regulators [START_REF] Gao | The REGγ-Proteasome Regulates Spermatogenesis Partially by P53-PLZF Signaling[END_REF]; 2) the mediation of DSBs during meiosis [START_REF] Ahuja | Control of meiotic pairing and recombination by chromosomally tethered 26S proteasome[END_REF][START_REF] Zhang | Proteasome subunit α4s is essential for formation of spermatoproteasomes and histone degradation during meiotic DNA repair in spermatocytes[END_REF] by affecting signalling molecules and acting upon the histones and 3) the degradation of acetylated histones through association with PA200 [START_REF] Qian | Acetylation-Mediated Proteasomal Degradation of Core Histones during DNA Repair and Spermatogenesis[END_REF][START_REF] Zhang | Proteasome subunit α4s is essential for formation of spermatoproteasomes and histone degradation during meiotic DNA repair in spermatocytes[END_REF]). It appears that spermatoproteasome plays a vital and non-redundant role in coordinating spermatogenesis, through UPS and association with alternative regulators such as PA200 and PA28γ. It appears that there are necessary functions in this process that could not simply be performed just by increasing standard proteasome activity, reflected by the presence of alternative testis proteasome in evolutionarily distant cousins such as Drosophila melanogaster [START_REF] Belote | Duplicated proteasome subunit genes in Drosophila and their roles in spermatogenesis[END_REF]. The exact reasons for this particular necessity remain to be elucidated through structural and functional studies of the spermatoproteasome complex.

Characterization of protein complexes by mass spectrometry 2.3.1 Introduction to mass spectrometry

MS is a technique that enables measurement of mass-to-charge ratio (m/z value) of charged particles, which can be used to determine their mass. By precisely measuring the mass of ions, and some a priori knowledge about the analyte, we can chemically identify the ion species. As schematically represented on the Figure 17, a mass spectrometer consists of an ion source, an analyser and a detector. In the source, the sample molecules are first ionized and dispersed, which makes them fly down a voltage gradient, established along the instrument, inside of which, different levels of vacuum are maintained. In the analyser section, different electric and/or magnetic fields are applied so as to select only the ions of a certain m/z value. The remaining ions that correspond to the desired m/z value are usually then passed into the detector -a section of the instrument capable of detecting and quantifying the ions. The analyser can then select ions of an incrementally different m/z value, which are then detected and quantified -that way the analyser can perform a scan of different m/z values. Modern instruments can perform this scan really quickly, depending on the desired resolution, sensitivity and the size of the range of m/z values. Primary output of the analysis is a mass spectrum -a plot of different m/z values of detected ions versus their relative intensity.

There are several desired properties of a mass spectrometer that are considered to be important parameters when describing its performance. Resolution of an analyser is its capability to separate ions of different m/z. It can be expressed as the lowest mass difference between two peaks the analyser is able to distinguish. Resolution is calculated form a single peak of an ion in a mass spectrum, as the ratio between the m/z of the ion and the width of the peak at a certain height, commonly at the middle of the peak (a.k.a. full width at half maximum -FWHM). Resolution is a unitless parameter, usually a big number and it is always defined for a given m/z. Sensitivity of an instrument is determined by its capacity to create ions, efficiency of the transmission and the capacity to detect and convert them to signal. It is important that the instrument maintains linearity of the signal response to ion abundance, across the broadest range of quantities possible. The sensitivity of a system is usually defined as the lowest molar quantity of an analyte it can detect, nowadays commonly expressed in picomoles, attomoles or femtomoles. Inherent to its mechanism of ions separation and geometry, an MS instrument will have a determined mass range of ions it can detect, that is, a determined mass limit. Some typical measurement terms can be introduced, such as the mass precision and the mass accuracy. Accuracy is defined as the proximity of the measurement to the true value. In mass spectrometry, this is often termed as mass measurement accuracy (or error) and is often expressed in parts per million (ppm), milli mass units (mmu) or root mean square (RMS). Milli mass unit is defined as 0.001Da and the error can be expressed as the difference between the measured mass and the theoretical mass of the ion. The error in ppm is expressed as the mass difference in mmu divided by the mass of the measured ion, multiplied by one million -it thus changes with the mass of the ion for an error of a given mmu. Precision is defined as the repeatability of a measurement as a consequence of random errors -repeated measurements of a value manifest as a distribution of values around an average, and the less dispersed the values are, the more precise the measuring technique is. This value can be expressed quantitatively as the standard deviation of a measurement [START_REF] Brenton | Accurate Mass Measurement: Terminology and Treatment of Data[END_REF]. The analytes can be introduced to the mass spectrometer in various ways. Ionization enables the analytes to be manipulated by the electric or magnetic fields of the mass analyser and later to be detected. Signal from the detector is then processed by a computer into a mass spectrum -representation of mass-to-charge versus intensity of the ions.

Ion sources

Numerous techniques have been developed to ionize sample molecules, that is, to provide relevant analytes with a charge and introduce them as individual desolvated molecules to the analyser of the mass spectrometer. Depending on the amount of transferred energy, the analysed molecules can fragment or stay largely unchanged -in the so called quasi-molecular ion state (IUPAC Gold book-(Gold, 2019)), which divides the techniques in two distinct groups-hard ionisation, which induces partial sample degradation and soft ionization techniques, which produce quasi-molecular ions. Relevant for the study of biomolecules are the soft ionisation techniques -of which the most widelyadopted are matrix-assisted laser desorption ionisation (MALDI) and electrospray ionisation (ESI).

The MALDI approach was developed by Karas et al. (Karas et al., 1987) and Tanaka received a part of the Nobel Prize in 2002 for the developments in the field [START_REF] Tanaka | Protein and polymer analyses up tom/z 100 000 by laser ionization time-of-flight mass spectrometry[END_REF]. The analyte is mixed into a matrix of a small organic compound and spread onto a surface with the help of a volatile solvent. The matrix is capable of absorbing light energy and transferring a part of it to the molecules around it, causing them to ionize. When a UV laser is discharged into the matrix, the matrix absorbs the radiation, which causes the analyte molecules to ionize and desorb from the surface into the gas phase. This process mostly produces singly-charged molecules.

A great problem with MALDI is resolving the complexity of the sample, when working with complex mixes. It remains very useful in the analysis of biomarkers and its capability to couple with in situ tissue analysis (whereby the tissue is fixed and prepared with the ionizing matrix). Since the analyte is released in pulses/packages, MALDI has been very successfully coupled to TOF analysers (see following section) who operate in the same (pulsed) manner. Some attempts to couple it to liquid chromatography (LC) separation have been made, [START_REF] Pereira | A Nano LC-MALDI Mass Spectrometry Droplet Interface for the Analysis of Complex Protein Samples[END_REF] to resolve the sample complexity issue.

The ESI technique was invented by Malcolm Dole [START_REF] Dole | Molecular Beams of Macroions[END_REF]) and it proved to be so significant for the characterization of large molecules, that a part of the 2002 Nobel Prize for Chemistry was awarded to John B. Fenn, "for the development of methods for identification and structure analyses of biological macromolecules" [START_REF] Fenn | Electrospray wings for molecular elephants (Nobel lecture)[END_REF]. The functioning of ESI is depicted on Figure 18 and is based on the following principles: when high voltage (few kV order) is applied to a conductive capillary containing liquid, positive (or negative) ions accumulate near the meniscus of the liquid at the end of the capillary, which destabilizes the meniscus and leads to the formation of the Taylor cone, culminating in the ejection of droplets into the gas phase. The ejected droplets start losing volume due to solvent evaporation, which reduces their surface, causing the charges on the surface to concentrate. At some point, (Rayleigh limit, defined by the maximum amount of charge that can be on the droplet) the repelling Coulomb forces overcome the surface tension of the droplet, which causes the droplet to fission, into smaller droplets, re-distributing charges. The process repeats itself until there is mostly analyte molecules carrying charges (Kebarle and Verkerk, 2009). The polarity of this process can be inversed, but most protein studies are done on positive ions. Unlike MALDI, it is common for ESI to produce multiply-charged ions, which enabled for the first time the study of bigger molecules, due to a shift in the m/z into a range that was more manageable by mass spectrometers. The fact that ESI is based on emission of particles from droplets emanating from a capillary is what unlocked the possibilities of analysis complex mixtures of non-volatile molecules like peptides and proteins, since the MS systems could now be coupled to liquid chromatography (LC).

The resulting quasi-molecular ions of the positive mode ESI can usually be described via the general formula of [M+nH] n+ , where M is mass of the original molecule, and n is the number of charges received in the form of protons. Apart from protons, the ionisation can occur via adduct formation with other cations, e.g. [M+nNa] n+ .

To facilitate evaporation from the droplet surface, volatile organic solvents are usually added to the water-based mixes, such as methanol or acetonitrile. Small amount of volatile acid (formic acid or trifluoro acetic acid) is most commonly added to the solution, to improve conductivity and to produce pH under which most analyte ions are protonated, when positive mode analysis is employed. In the analysis of native proteins under non-covalent (non-denaturing) conditions (vide infra), addition of ammonium acetate is practiced, which is a volatile salt that helps maintain a physiological pH and suppresses the formation of undesirable salt adducts. There are two prevalent theories on the exact ion formation mechanism: ion evaporation model (IEM) and charge residue model (CRM). The IEM proposes that the charged particles are ejected from the solvent droplet, carrying the charge with them and leaving the solvent droplet behind, which seems to be in accordance with experimental data on formation of smaller ions. The CRM proposes that after several fissions, the multiply-charged ions are formed when all of the solvent evaporates away [START_REF] Winger | Observation and implications of high mass-to-charge ratio ions from electrospray ionization mass spectrometry[END_REF].

With ESI, most of the analyte is lost at the interface with the MS, so in order to analyse smaller amounts of analyte, there was a need for smaller flows at the capillary, thus the nanoESI was developed in 1995 [START_REF] Wilm | Analytical Properties of the Nanoelectrospray Ion Source[END_REF]. In nanoESI (~10nL/min), there is a smaller, more fissile droplet formation, better ionisation yields and therefore better sensitivity. Moreover, nanoESI does not necessarily require positive pressure across the capillary, but the liquid is continuously pulled towards the Taylor cone end as a result of the electrostatic and capillary forces [START_REF] Wilm | Analytical Properties of the Nanoelectrospray Ion Source[END_REF]. Due to lower flow-rates nanoESI eliminated the need for heated source and sheath gas and drastically reduced the amount of volatile solvents used in the analysis.

Great importance of ESI lies in its capacity to couple MS with LC systems. This enables the analysis of complex mixtures, by separating the analytes based on their retention time. This is particularly important in proteomics, where sample complexity is a major point of consideration in experiments.

Mass analysers

The way an ion will behave in an electric field in a vacuum depends on several parameters, but mainly on its mass and charge-the more massive an ion is, the higher inertia it carries while in motion and it is thus more difficult for an electric field to change its direction. The more charges it carries, the more susceptible it becomes to the electrical field forces and voltage differentials. It is exactly these properties of the ions that the analysers use to manipulate them. Nowadays there are several common types of analysers in use, each having inherent advantages and disadvantages in the design: time-of-flight (TOF), quadrupole (Q), ion trap (IT) and Fourier-transform analysers (FT) -ion cyclotron resonance (ICR) and Orbitrap.

TOF analysers operate on a very simple principle: they measure the time it takes ions, accelerated by initial voltage differential, to travel a fixed distance in a vacuum tube, free of electric field. This "time of flight" is correlated with the mass-to-charge ratio of an ion: energy uptake of the ions by the initially applied voltage is directly proportional to their charge, but the speed is inversely proportional to their mass. Thus, the ions of the same charge will gain the same kinetic energy in the electric field, but their speed will depend on their mass, due to a simple relation: Ek=½(mV 2 ), whereby Ek is the kinetic energy, and V is the speed of an ion with a certain mass m. The TOF analysers offer very large mass range and speed of analysis, however, variability of velocities of the same ion species can broaden peaks measured by the detector, which limits the resolution and they are known to take up a lot of space. In order to extend the ion path across the analyser, and thereby increasing its nominal resolution, set of reflectron lenses is introduced into its geometry, giving a V-shaped ion path, across the same TOF space (See Figure 19A). Introduction of a reflectron has an added benefit of focusing the ions incoming to the detector into narrower peaks. TOF analysers offer resolution which in theory can be extended by increasing the TOF distance, however, in practice, the resolution does not go above 50-80,000. On the other hand, their mass range is practically unlimited (up to 1,000,000 m/z in rapifleX series from Bruker) Q analysers are very compact systems -they consist of four parallel electrodes, placed at equal distance from the axis of ion passage. Perpendicular electrodes are connected to the same DC polarity, while adjacent electrodes have opposite DC polarities (Figure 19B). These DC voltages offset the radio frequency that is applied between the two pairs of electrodes. These voltages are manipulated in such a way that only ions of certain m/z value maintain a stable trajectory through the analyser, while the others fall out of path -creating what is called a mass filter. This mode of operation is also called a selected ion monitoring mode or SIM. The quadrupole can be operated in a scan mode, where a scan of a certain m/z range can be performed at a speed of up to 15 000 amu/s (current Shimadzu instruments), thus permitting the ions of different m/z to be quantified in a short span of time. The Q analysers offer a simplicity of design and the capability to select a really narrow band of m/z, but in the attempt to more precisely select an exact m/z, they lose in sensitivity, since most of the introduced ions are not perfectly centred along the axis of transmission and are thus lost during filtering process and on their own, they do not offer great resolutive power. The Q analysers are very useful as filter analysers and are found in various configurations in tandem mass spectrometersinstrument setups composed of more than one analyser. One of the first configurations still in use to date are the triple quadrupoles, consisting of two linearly connected analyser Qs bridged by a third one that serves as a collision cell (QqQ). Other common configurations are Q-Orbitrap and Q-TOF. Tandem mass spectrometers and their modes of operation will be explained in detail in the following section.

IT analysers are based on the same principle of operation as quadrupoles, except that they are designed to confine the incoming ions oscillating inside defined space over a period of time. There are two types of designs: linear ion trap (LIT) and ring ion trap (presented on the Figure 19C). LIT confines the ions via linear electrodes along the axis of entry, akin to Q analysers, while the end-cap potential is applied to repel the ions and prevent them from escaping at the LIT terminals. Ring ion trap confines the ions inside a 3D space via one ring and two hyperbolic end-cap electrodes. In both cases, the ions can be accumulated over time, then ejected into the detector one by one, by gradually augmenting the current that maintains the confined ions. The voltage used for ion ejection can be directly related to their m/z value. The advantages of ion traps are that they are cheap and easy to maintain and can accumulate the ions over time, which gives them a higher sensitivity, however the capacity of an ion trap is limited, as well as the possible resolution. The capacity to confine ions gives tremendous options for experimentation, since the ions can be accumulated, then selected for fragmentation. The fragments themselves can be then further fragmented, enabling MS n experiments, which will be explained in the following chapter.

The Orbitrap analyser belongs to the family of FT mass analysers but is also a type of ion trap [START_REF] Makarov | Electrostatic Axially Harmonic Orbital Trapping: A High-Performance Technique of Mass Analysis[END_REF]. The principle of operation consists in trapping the ions of a certain m/z range inside a hollow electrode on a trajectory around an axial electrode (Figure 19D). As a result of the electrostatic confinement and the initial kinetic energy, the ions will maintain a complex path around the axial electrode, oscillating in multiple ways, thereby having a rotational radial and axial frequency. The motion of ions along the axial electrode defines their axial frequency, which is the only one that will not depend on the spatial spread of ions or their energy, but is directly related to the square root of their m/z ratio. In this analyser, all the ions are detected simultaneously. The oscillating ions inside the trap induce current in the surrounding electrode, that is then measured. The resulting signal is a function of complex harmonic oscillations of numerous ions and cannot be read as such. A mathematical procedure called Fourier transform decomplexifies the recorded overlayed periodic functions into a set of m/z values and intensities. Huge advantage of FT analysers is that the resolution of the acquired spectra increases with the time of acquisition, which gives them great resolutive power (480 000 at 200 m/z for Thermo Scientific's Exploris 480 model).

FT-ICR analysers are similar to the Orbitraps in the regard that they also trap ions that perform very complex motion inside the confinement area and consequently induce current in nearby detection plates. The signal produced from this motion also has to be deconvoluted using Fourier transform. Unlike Orbitraps, which use electrode geometry and electrostatic field to trap the ions, FT-ICR uses a magnetic field. When in a magnetic field, ions undergo circular motion, perpendicular to the direction of magnetic field. The frequency of oscillation will depend on the m/z of the ions. In order to keep the ions of the same m/z in a package and to introduce enough kinetic energy to the system, so that the ion motion can be reliably measured, an electric RF field perpendicular to the direction of the magnetic field is applied, in a process called excitation. The same principle regarding mass resolution as for the Orbitraps, applies here -the longer the detection time, the better the resolution. To achieve longer detection times, the ions must maintain their energy and trajectory. For this, an ultra-high vacuum and a strong, homogenous magnetic field must be applied, which is very costly and equipment is rather bulky, however, ICRs remain the best analysers in terms of resolution and mass accuracy [START_REF] Marshall | Fourier transform ion cyclotron resonance mass spectrometry: A primer[END_REF]. Current instrumentation constructed by Bruker, which employs magnets of up to 15T (solariX series), is able to achieve the resolving power beyond 10,000,000 and accuracy below 250ppb. In Q analysers, the RF field maintains the ions on a stable trajectory through the cavity along the electrodes. Applying offset DC voltage can destabilize the trajectories of ions of certain m/z, effectively creating a mass filter. C: Cross-section of a ring IT: the ions are maintained in constant motion via one ring-shaped electrode, capped by two hyperbolic electrodes. Ions can be filtered through a similar mechanism as in quadrupoles, collided with a gas to induce fragmentation and their fragments can be further analysed in the same IT. D: Orbitraps confine the ions in a complex rotating motion around the axial electrode. The ions induce current in the outer electrode, which is recoded as signal. Complex waveform of the signal is deconvoluted through mathematical transformation into mass spectra. E. ICR uses strong magnetic field to confine ions in a circular motion. Electrical RF field is used to add energy to the system. Moving ions induce current in the nearby plates, which is recorded as signal. ICR, like the Orbitraps, are part of the FT-family of mass analysers, meaning that the signal is decomplexified through Fourier transformations in order to obtain information on the m/z of the ions. Adapted from [START_REF] Haag | Mass Analyzers and Mass Spectrometers[END_REF].

Ion mobility cells

Ion mobility spectrometry is a technique of separation of ions in a gas phase using electric fields. Unlike mass spectrometry, where the separation occurs under vacuum, the ions here are submitted to non-destructive collisions with an inert gas, which makes them separate based on their size and shape as well. In a simplest setup, molecules of analyte are ionized and flown into a tube having a certain pressure of inert gas, across which a constant electric field is applied to maintain the kinetic energy to the ions. The measured parameter for these ions is the time it takes to cross the tube -the drift time. The friction forces of the gas environment (the drift gas) will slow down the drift of the ions from one side of the tube to another. The extent to which an ion is slowed down is directly proportional to its collisional cross section (CSS) -the area amenable for collisions with the gas atoms. It can be thus said that the IMS measures size-to-charge ratio, making it an orthogonal ion separation technique to the MS. Groups of ions of the same m/z will not necessarily have the same drift timethey may be composed of several different ions or a same compound in different structural conformations, e.g. a peptide folded in two different ways, having two different CCSs. Using only m/z, to characterize molecules becomes increasingly difficult as the size of the molecule increases. Even the identification of a simple molecule such as a monosaccharide can be difficult if we use only its exact mass, due to structural isomers and usually a lot of a priori knowledge about the sample is needed. Extremely useful approach in MS is to introduce energy to the analyte ions, thereby dissociating them into fragments, which may or may not be charged; the mass of the ionized fragmentation products is then studied to either infer the structure or exact molecular composition of the initial ion, or a signature product ion (or a set of ions) is used to confirm the identity of the analyte in case of some a priori biochemical knowledge. This approach, where the precursor ions and the fragmentation (also called "ion activation") products of the said ions are studied is called tandem MS, or MS n , whereby the exponent "n" represents the number of generations of ions originating from the first precursor ion. It is important to note that the m/z window is usually left wide enough to encompass several isotopes of an ion, for the sake of ion abundance/ signal intensity.

The MS n experiments can be separated by space or time. The separation in space is done when the selection of the precursor, fragmentation and the analysis of the fragment ions is done in separate analyzers, connected by ion-routing elements. The MS n separation in time is a characteristic of ITs, due to their capacity to store ions -meaning that selection, fragmentation and m/z analysis can be done inside one trap. This way, an IT is capable of making several generations of ions, giving us MS 3 and so on, but the trap's ion capacity is the limiting factor.

Collision-induced fragmentation (CID)

In CID, analyte ions are subjected to collisions with the molecules of neutral and inert gas (usually N2, He or Ar), while confined inside a collision cell. The resulting transfer of energy from the gas molecules to the analyte ions causes them to fragment. Depending on the exact conditions of the reaction (gas pressure, temperature, kinetic energy of the analyte ions), different fragmentation patterns can occur. Relevant for the study of peptides and proteins are the low-energy collisions (1-100 eV), usually done inside the Q or LIT. They suggest several successive lesser-energy collisions with the gas, before the molecule has enough energy to dissociate into products [START_REF] Sleno | Ion activation methods for tandem mass spectrometry[END_REF]). Variation on this approach is the higher-energy C-trap dissociation (or higher-energy collisional dissociation, HCD) developed by Olsen et al. [START_REF] Olsen | Higher-energy C-trap dissociation for peptide modification analysis[END_REF]. In HCD approach, the collision is done in a type of IT with adjusted higher capacity and extended m/z range for ion trapping, useful for small reporter ion detection. Another variation on the CID is the SID -surface-induced dissociation. Unlike CID or HCD, where the ions undergo multiple collisions with molecules of gas, in SID the ions collide against a solid surface. This technique has been used with great success in native MS, where it enables the dissociation of protein quaternary structures, without denaturation/unfolding of the subunits [START_REF] Stiving | Surface-Induced Dissociation: An Effective Method for Characterization of Protein Quaternary Structure[END_REF].

Electron-capture dissociation (ECD) and electron transfer dissociation (ETD)

In ECD, low-energy electrons are emitted by a filament and captured by multiply charged ions, which causes charge reduction and fragmentation of the ion. When applied to the study of peptides and proteins, ECD is known to preserve labile PTMs such as glycosylation or phosphorylation, but rather fragment peptide backbones themselves. However, in order to manipulate both cations and electrons, an FT ICR device is needed, which is a costly solution [START_REF] Zubarev | Electron capture dissociation of multiply charged protein cations. A nonergodic process[END_REF]. Similar method has been proposed, where anions capable of transferring electrons are introduced into a LIT containing the analyte cations. The result of the electron transfer would be cation dissociation and the reaction is thus called electron transfer dissociation (ETD) [START_REF] Syka | Peptide and protein sequence analysis by electron transfer dissociation mass spectrometry[END_REF]. Due to its lower cost and propensity to preserve labile chains in the fragments, ETD is a very convenient way to study the PTMs of peptides and proteins and is popular in (Native) Top-Down Proteomics [START_REF] Lermyte | Characterization of top-down ETD in a travelling-wave ion guide[END_REF] -an approach that will be detailed in the native MS chapter (2.4.3.2).

Ultraviolet photo-dissociation (UVPD)

Besides collision or chemical reactions, another way to increase internal energy of a molecule is by light absorption. The chemical bonds in an analyte molecule can be targeted for fragmentation using light wavelength corresponding to maxima in their absorption spectra. When the UV light is absorbed by the chemical bond, two events commonly take place: the electrons partaking in the bond formation are moved into dissociative orbitals or they "fall" back into the previous state while releasing vibrational energy (R. Julian, 2017). In the first case transitions happen at a time scale which does not allow for other relaxation pathways to take place and the absorbed energy is spent directly on dissociation of the said bond. The second pathway results in a hot molecule, alike to the state after CID. These two pathways come together to yield unique dissociation patterns, with good sequence coverage for large ions, useful for intact protein studies (Greer and Brodbelt, 2018; Gomes et al., 2020).

Peptide fragmentation

When analyzing peptides, it is possible to determine their exact sequence via MS 2 experiment. The peptide ion is first selected as a precursor, activated by one of the previously described methods and the fragments are then analyzed. In a typical MS 2 experiment of a peptide, depending on which bond in the peptide backbone is broken, several different ions can occur as a result of the precursor peptide activation. Typical products of fragmentation are illustrated on the Figure 20A: breakage of the peptide bond will result in the formation of b-and y-ions -typical of CID and HCD, while the breakage of the Cα -N bond results in the formation of z-and c-ions. The ETD is known to produce many c-, z-and y-ions. Breaking of the C -C bond will produce x-and a-ions. The convention of naming the ions has been proposed by Roepstorff and Fohlman in 1984 (Roepstorff, Fohlman and P. [START_REF] Roepstorff | Letter to the editors[END_REF].

Upon activation, in a large pool of identical peptide ions, it will happen that, e.g., CID yields a series of b-and y-ions of different sizes. The mass difference between "adjacent" ions of the same series (y-or b-) will correspond to the mass of a particular amino acid residue. For example, in the Figure 20A, the mass difference between the y2 and y3 ion will be the mass of the R2-containing amino acid residue. By interpreting the mass spectra of the fragments, we can thus deduce the sequence of the precursor peptide. Knowing the mass of the precursor will help us fill the information gap to determine its full sequence. An example of sequence determination of a peptide precursor, based on the b-and y-ion series of its fragment spectra is illustrated on Figure 20B. In a common Bottom-Up proteomic experiment, this procedure is somewhat different and automatized, whereby the ions in the MS2 spectra are assigned by comparison of their masses to the masses of library entries of fragment ions produced in silico. The precursor ions are often found in the MS 2 spectra, along with the fragments. Here, the precursor ion with the formula MH + can be found on the far right of the spectra. Mass difference between the precursor and different fragment ions correspond to the mass of the sequentially lost amino acid residues. This permits to reconstruct the sequence of the peptide. Adapted from (Baars and Perlman, 2016).

Tandem MS in protein analysis

As explained earlier, tandem MS permits us to more precisely characterize the structure/composition of the analyte. In protein analysis (especially in shotgun proteomics) we rarely deal with samples simple enough to comprehensively analyse as such. It is thus very common to separate the analytes via LC system coupled directly to the MS. Following method considerations will take this assumption by default. Depending on the goal of the analysis, there are several different scanning modes of MS n experiments: Product Ion Scan, Precursor Ion Scan, Neutral Loss Scan, Single Reaction Monitoring, Multiple Reaction Monitoring, Parallel Reaction Monitoring. For the sake of simplicity, let us consider MS 2 experiments involving two analysers and the collision cell, such as the QqQ configuration mentioned earlier in the analysers section.

Product Ion Scan: The first analyser filters only a narrow m/z range and the ions are fragmented in the collision cell. The fragments are then analysed by the second analyser. Each fragment spectrum (MS/MS or MS 2 spectrum) is associated with the m/z of the precursor and can now serve to identify it. The more narrowly we define the m/z value of a precursor, the less ions we get due to the imperfections of the selection process -thus a compromise must be made between sensitivity and precise precursor selection. This approach is commonly used to determine structure of molecules and to identify peptide/protein sequences in proteomics. Two strategies exist to obtain the MS 2 spectra in this type of experiment: data-dependent acquisition (DDA) and data-independent acquisition (DIA). The schematic comparison of the DDA and DIA approach are represented on the Figure 21.

In DDA approach, a first MS survey scan is done (~1s), to quantify all the present ion species and spot the most intense precursors to fragment. The 1 st analyser voltages are set to let all ions reach the 2 nd analyser that is used to scan and analyse the ions. The instrument then compiles a list of all the detected m/z values and sequentially selects N most intense ions as precursors for fragmentation in the following MS 2 scan. Ion selection (where ideally only one ion species is selected via narrow m/z band), fragmentation and analysis of the fragments is done for each precursor individually (~50 ms) until a defined number N of MS 2 scans is reached or there is no more ions to analyse, after which a new cycle of analysis starts with another MS survey scan. Instead of defining the N of MS 2 scans, cycle time can be defined instead. Practical considerations for the cycle time length in proteomics are following. An average chromatographic peak in nanoLC setups lasts around 20 to 30 s. By convention, we consider a minimum number of measurement points per peak to be 8 to 10 for a sufficient LC resolution. It thus follows that the cycle time should not exceed 2.5 to 3 s. In order to avoid fragmenting the same ion several times, a certain m/z can be excluded from precursors for a certain period -this is called dynamic exclusion and can greatly improve performance in complex samples, however, there is a risk that the MS 2 spectra will not be obtained at the apex of the peak, thus losing in intensity. The DDA approach suffers from a limited dynamic range for ion identification (as only the most intense precursors are usually fragmented) and results in missing values in quantification results. This is more pronounced in complex samples, where several analytes elute from the LC at the same retention time. It follows that these limitations can be diminished by reducing the co-eluting analytes. Strategies to achieve this will be covered in detail in the following chapters. Nonetheless, DDA developed marvellously the field of proteomics, where in Bottom-Up approach, up to 11500 proteins can be quantified, with the help of various sample fractionation strategies (Azimifar et al., 2014).

The DIA approach does not employ a narrow m/z band selection in the first analyser. On the contrary, the strategy of simultaneous multiple ion fragmentation is employed, which yields more complex MS 2 spectra. Special bioinformatic treatment of the said spectra must be done to assign the fragments to a certain precursor, in an attempt to identify it. The family of DIA is diverse [START_REF] Zhang | Data-Independent Acquisition Mass Spectrometry-Based Proteomics and Software Tools: A Glimpse in 2020[END_REF], but two representative strategies will be described: SWATH and MS E . In the SWATH approach, in order to cover a defined scan range (e.g. 400 to 1500 m/z), sequential non-overlapping (or partially overlapping) bands of 20 -30 m/z in width are selected for transmission in the first analyser, then fragmented and MS 2 spectra is obtained. Here, a cycle time is the time required to scan the full m/z range by fragmenting these neighbouring bands, one by one. The name "swath" is an ensemble of fragment ion spectra for one band, across a defined chromatographic time. In Bottom-Up proteomics, the bioinformatic analysis of DIA data usually requires pre-existing spectral libraries acquired in DDA mode. These libraries contain information about the retention time, charge, precursor m/z, fragment type and fragment m/z values. The libraries are needed because the fragments cannot be directly associated to a precursor, since a broad range of m/z is used to select multiple precursors to be fragmented at the same time. To create these spectral libraries, a sample of the same type (and species) must be used, usually the mixture of aliquots of all biological samples that are to be analysed individually by DIA, to ensure covering the whole proteome in the created database. This also permits that a sufficient quantity of biological material (mg) is available for a first fractionation step. The sample mix is fractionated by a separation method orthogonal to the one used in the LC-MS system in order to improve the proteome coverage. Signals obtained from individually injected biological samples will finally be bioinformatically interpreted by matching m/z and retention times of DIA fragments in the spectral library. Since this is very time and resource-consuming, comprehensive community-driven spectral libraries are made for several organisms and are constantly improved (SWATHAtlas.org), thus eliminating the need for in-house project-specific library to be made, while improving cross-laboratory experiment reproducibility [START_REF] Rosenberger | A repository of assays to quantify 10,000 human proteins by SWATH-MS[END_REF].

Very innovative approach to DIA, called the scanning SWATH has recently been developed, whereby instead of selecting for fragmentation the increments of defined m/z range, one at a time, a continuous sweep of m/z is performed in the precursor selection process. As the m/z sweep is performed, the fragment intensities will appear in a distribution, centred around the m/z of its precursor -this facilitates the association of the MS2 spectra with its corresponding precursor [START_REF] Messner | Ultra-fast proteomics with Scanning SWATH[END_REF].

The MS E is a technique patented by Waters (Manchester, UK) [START_REF] Silva | Absolute Quantification of Proteins by LCMSE[END_REF]. Here, the full m/z range of ions is transmitted and alternating low and high collision energy is used in the MS spectra acquisition. When the collision energy is low (~4V), no fragmentation occurs and the MS spectra gathers all the parent-ions. When the collision energy is high, all ions are fragmented simultaneously. By alternating these two modes, a pairing can be done between the precursors in the low-energy mode and the fragments in the high-energy mode. The algorithm for spectra interpretation is therefore somewhat more complex and involves aligning the extracted ion chromatograms of the precursors and fragments to create pseudo MS 2 spectra. Compared to DDA, the dynamic range of DIA is only limited by the detector sensitivity and the chromatographic separation, making it better for the deep proteome studies. However, the complexity of the MS 2 spectra and the pre-requirement of spectral libraries in most cases limited the use of DIA approach and for a long time DDA has been the standard of analysis. Recent improvements in DIA methods have put this approach on par and even better than DDA, both in terms of identification and quantification of proteins [START_REF] Dowell | Benchmarking Quantitative Performance in Label-Free Proteomics[END_REF]. Another major advantage of DIA over DDA is a better reproducibility of MS 2 spectra -due to the stochastic nature of precursor selection in DDA, a lot of MS 2 missing values are generated. For most DIA approaches, the need for spectral libraries remains a limitation. However, recent efforts in applying deep learning show that this problem may be circumvented all together by the creation of in silico spectral libraries [START_REF] Yang | In silico spectral libraries by deep learning facilitate data-independent acquisition proteomics[END_REF]. In Precursor Ion Scan mode, the first analyser scans for precursors and successively selects them for fragmentation. However, the second analyser will only filter for a very specific m/zexpecting a very specific fragment mass. This approach is used in the study of PTMs in proteomics, where a fragment ion of a certain mass can be a reporter for a PTM (such as phosphorylated peptides with +80 Da shift or N-/O-Glycosylated peptides with +162 Da mass whift).

Neutral Loss Scan is a mode in which the two analysers scan at the same speed, but two different m/z ranges, offset by a certain mass difference. This difference corresponds to an expected neutral mass loss for a known fragmentation pathway (e.g. fragmenting a phosphorylated peptide can give a neutral loss of 80 or 98 Da).

In Single Reaction Monitoring (SRM), the first analyser selects a narrow m/z range of ions, which are then fragmented. The second analyser also selects a very narrow m/z, thereby monitoring for a specific precursor ion -fragment ion transition. This type of monitoring is used for accurate identification and quantification of biomarkers, metabolites, toxins in the environment, etc. It is necessary to study in advance the fragmentation of the molecule in question.

Multiple Reaction Monitoring (MRM) is based on the same principle as SRM, except that several parent ion -fragment ion transitions are observed simultaneously. Very useful for identification and quantification of several known analytes, MRM gives us better dynamic range and sensitivity, compared to DDA, however, only a few transitions can be studied and the interferences even in a narrow m/z band are possible.

In Parallel Reaction Monitoring (PRM), precursors are selected for fragmentation in the first analyser (like in SRM/MRM) and all of the fragmentation products are observed simultaneously in the second analyser. For such experiment, a high-resolution second analyser is required. This approach benefits from parallel observation of multiple fragment ions in that it has higher specificity and lower interferences from non-targeted ions.

Not all the instrument configurations have the capability to perform all the modes of analysis listed above. Product Ion Scan can be implemented on all types of instruments but not the other scanning modes. Triple quadruple-like instruments (QqQ & QqTrap configurations) can perform all scanning modes except PRM (which requires high resolutive power) and Q-Orbitrap / Q-TOF configurations are more commonly used for Product Ion Scan (no a priori knowledge of proteins) or PRM (targeted analyses).

Instrument geometries

To illustrate the possibilities for the combination of analyzers into a single integrated MS architecture, two examples will be given, namely the ones used throughout the experimental sections of this work.

The Synapt G2Si is an instrument commercialized by Waters (Manchester, UK). It is a Q-TOF with an IM cell integrated between the Q and the TOF. The analytes are ionized at atmospheric pressure and introduced to the system via a set of ion guides. By displacing the ion route out of line of sight of the first analyzer (StepWave TM ), the introduction of neutral ions further into the system is limited. As presented by the scheme on the Figure 22, next in the path of the ions is the quadrupole, followed by the TriWave cell which consists of three segments -an ion mobility separation element sandwiched between two linear ion traps (Trap and Transfer). Finally, the latter trap connects the IM cell to the TOF analyzer.

Each of the elements offers a set of possibilities, which combined give a broad range of possibilities for different MS experiments. In a simplest, MS-only method, the Q is in transmission mode, while the TOF measures the m/z of the transmitted ions. The internal energy of the ions can be increased by collisions in either of the two collision cells, giving possibility of ion fragmentation before and/or after the ion mobility cell. In a common MS/MS method, the Q can select a specific precursor ion, which is fragmented in one or both cells and the fragments analyzed in TOF. The IM cell can be activated to separate the ions in an orthogonal manner for any of the mentioned modes. The analyte can be characterized via drift time and m/z values in IM-TOF mode, or a specific m/z can be selected in Q for subsequent separation in IM, with or without fragmentation in any of the two cells. In the TOF analyzer, the path of the ions can be extended from a V-shaped path to a W-shape, thus increasing the resolution of the analyzer, at the expense of the sensitivity. The Q-IM-TOF configuration is convenient for the study of intact proteins, since the ion mobility cell offers the separation of different conformations and the TOF analyzer has a mass range only limited by the transmission capacity of the quadrupole.

The MS/MS experiment can be done in two different modes: DDA approach, where the Q isolates the most intense ions of a specific m/z, which are fragmented in one of the traps adjacent to the IM cell; second approach is the before-mentioned MS E , where Q is constantly in transmission mode, while the trap shifts periodically between low-and high-energy mode, fragmenting all the available ions. The Orbitrap Fusion Tribrid is a Q-orbitrap instrument with a linear ion trap, made by Thermo Scientific (CA, USA). In an MS-only experiment, the ions are introduced into the system and focused by the S-Lens, while the neutral particles are eliminated via the geometry of the Active Beam Guide (see Figure 23), to reduce the noise and improve the signal-to-noise ratio. The ions are transferred through the Q and into the Ion-Routing Multipole (IRM) that acts as a trap for the incoming ions. After a certain quantity threshold (regulated by the automated gain control -AGC), the ions are introduced from the IRM into the orbitrap (via the so-called C-trap), where the spectra are derived from the FT signal. In an MS 2 experiment, the ions of certain m/z are filtered through the Q, accumulated in the IRM and passed into the Linear Ion Trap (LIT), where they are fragmented. The ions can also be fragmented in the IRM, via CID-type fragmentation called HCD, described in the previous chapter. The fragmentation products can then be analyzed in the LIT (equipped with a pair of detectors) or in the Orbitrap, if higher resolution is desired. This configuration is capable of performing simultaneous MS and MS 2 acquisitions: while the MS spectra is acquired in the orbitrap, the Q, IRM and LIT can work in concert to accumulate, select, fragment the ions and analyze the fragments. This is particularly important in DDA acquisition of MS 2 spectra because the parallel functioning of these different modules helps the selection and fragmentation of a higher number of ions for the same cycle time. Moreover, parallel acquisition leads to higher overall sensitivity of the analysis, since it allows the Orbitrap cell to have a longer time for scanning, which results in the higher resolution of analysis. For more than a century of its existence, MS was used to answer questions across many different fields, from initial discovery of isotopes and preparative application for uranium, over the analysis of small organic molecules and peptides, to the study of native protein complexes in the MDa range. The development of tandem MS and the introduction of soft ionization techniques enabled the study of fragile molecules such as peptides and proteins, which upon decades of incremental improvements in the analytic technology allowed for the development of the entirely new fields of proteomics and structural MS.

Proteomics

The development of molecular tools has revolutionised biology research in recent decades. Veritable explosion of genetic study and the emergence of fields such as genomics has occurred due to the development of polymerase chain reaction (PCR) as a tool to amplify DNA [START_REF] Mullis | Specific Enzymatic Amplification of DNA In Vitro: The Polymerase Chain Reaction[END_REF] and several generations of DNA sequencing technologies [START_REF] Sanger | DNA sequencing with chain-terminating inhibitors[END_REF][START_REF] Bentley | Accurate whole human genome sequencing using reversible terminator chemistry[END_REF]. Subsequent discovery and implementation of the reverse transcriptase as a molecular biology tool, which translates mRNA into cDNA, has enabled the study of the transcriptome (Aatsinki et al., 1994).

It would be convenient to introduce here the term proteoform: distinct protein forms arising from alternative splicing and/or various PTMs, but nonetheless transcribed from the same gene. Necessity for a specific term arose from confusion over the term "protein isoform" which IUPAC refers to only for genetic isoforms. Disambiguation of various terms appearing in the literature can be found here: [START_REF] Smith | Proteoform: a single term describing protein complexity[END_REF].

For a flow of information from genes to messenger RNA to proteins, there is an everincreasing number of possible proteoforms. The number of possibilities of alternative mRNA splicing multiplied by the number of possible post-translational modifications of the protein give a huge number of potential products. Only a fraction of these possibilities occurs, of course. For about 21 300 human protein-coding genes [START_REF] Pertea | Thousands of large-scale RNA sequencing experiments yield a comprehensive new human gene list and reveal extensive transcriptional noise[END_REF] it is estimated to exist at least 70 000 proteoforms (Aebersold et al., 2018), however, this number is very difficult to estimate or exactly determine.

Proteomics aims at identifying and quantifying proteins from a generally complex system (tissue, cell, cellular compartment, biological fluid, etc.). There is no PCR equivalent (amplification) for protein study, which is why sensitivity has limited proteomics studies for a very long time. However, recent analytical method developments in sample preparation, chromatography MS instrumentation, and bioinformatics enabled the development of this field. Nowadays, the field of proteomics is completely identified with the MS analysis for identification of proteins. General proteomics workflow consists in extracting the protein fraction from a sample, followed by separation of protein species into simpler mixtures to facilitate identification by tandem MS. Two general approaches to proteomics study exist: Bottom-Up (BU) and Top-Down (TD). Essentially, in a typical BU approach the proteins are first enzymatically digested into peptides, then the peptides are separated via different biochemical methods to be subsequently analysed via MS. Prior to enzymatic digestion, protein mixture can be simplified by fractionation, by LC, SDS-PAGE or another technique. In TD approach, entire proteins are separated and analysed as such. Another chimeric approach exists, called Middle-Down, which involves partial digestion of proteins and subsequent study of intermediate fragments. TD and Middle-Down approaches can be considered to be structural techniques, since they enable the determination of the primary protein structure, by sequence and PTM identification (and thereby an exact proteoform determination). In a dynamic and rapidly growing area such as protein MS, the boundaries of definition are often being moved around. Recent expansion to the TD field is the native TD, where in a single MS experiment, the components of the noncovalent complexes are isolated and fragmented in order to identify them as a discrete proteoform [START_REF] Ro | Native top-down mass spectrometry provides insights into the copper centers of membrane-bound methane monooxygenase[END_REF].

Two great challenges in proteomics are the large dynamic range and the complexity of the sample. There is indeed a great discrepancy in abundance between different protein species in the same sample -while myosin and albumin may constitute up to 15% of protein mass in a muscle tissue sample, cyclins may be virtually undetectable, due to their low copy number in a cell. Haemoglobin constitutes most of the protein mass of a red blood cell. Many approaches have been commercially developed to deplete these abundant proteins from the samples and to enrich the low-abundant species [START_REF] Polaskova | High-abundance protein depletion: Comparison of methods for human plasma biomarker discovery[END_REF]de Jesus et al., 2017) and thereby decrease the dynamic range.

To reduce sample complexity, proteins or peptides are separated via chromatography, either in 1D or 2D. The most common way of separation is reverse-phase (RP) liquid chromatography on C18based columns (for peptides) or C4/C8 columns (for proteins), coupled directly to the mass spectrometer. Thereby, the retention time of an analyte can greatly help in its identification. In order to get a deeper proteome, offline fractionation can be performed prior to the RP separation. A first step can be gel filtration, gel electrophoresis, ion exchange, isoelectric focusing, HILIC, or basic RP, whereby the eluting analytes are pooled into several fractions, which are then analysed one by one via acidic RP LC-MS. Alternatively, 2D LC separation can couple two orthogonal chromatographic techniques (Swearingen and Moritz, 2012).

Principles of Bottom-Up Proteomics

As mentioned earlier, in BU proteomics the purified proteins are enzymatically digested into peptides before LC-MSMS analysis. The approach relies on the assumption that every protein will have at least one, but preferably several proteospecific peptides -that is, peptides unique to the said protein. A pre-requisite for shotgun proteomics is the existence of a genomic database (with a characterized transcriptome), which enables to predict theoretical protein sequences. Comprehensive proteome database for a large number of organisms can be found in Swiss-Prot database (>42 000 entries for human, >564 000 entries in total), which is a curated and up-to-date collection of experimentally confirmed proteins. Swiss-Prot is a subgroup of a much larger TrEMBL database -a transcriptome-based proteome database made up of theoretically predicted proteins (more than >174 000 entries for human, >207 800 000 entries in total). Both of these can be accessed via online platforms such as Uniprot. First, an in silico digestion of the proteins is done based on a protein sequence database and possible peptides are predicted, as well as their theoretical MS 2 spectra. These theoretical spectra are then compared to the experimental spectra obtained in the LC-MSMS run. Matching spectra are assigned to peptides, which are then assigned to proteins. Different quantitation approaches can be employed, which will be discussed later on.

Why would one resort to such strategy? Although at first counter-intuitive, the BU approach for now yields far superior results in terms of proteome coverage and quantitation compared to TD for several reasons. The BU approach is more sensitive and is not limited by the size of the protein. Furthermore, it is less susceptible to ionization competition of the coeluting proteins, whereby the smaller proteins will have better propensity to ionize and thus "steal" the charges form the bigger coeluting proteins. Finally, the bigger unfolded proteins will ionize with higher charge states which manifest themselves as denser charge state envelope, which requires higher instrument resolving power to properly separate.

The Bottom-Up Workflow Sample preparation

For the overview of the workflow, see the Figure 24. First, the proteins need to be extracted from the sample. Many sample-specific considerations need to be made, since each sample type (e.g. muscle, brain, bone, milk, cell culture, fruit, leaf, soil sample, archaeological artefact or a painting) represents a different matrix from which the proteins need to be extracted and has its own set of constraints. Overall, sample is homogenized enough to release proteins. The proteic fraction is biochemically separated from the rest and the proteins are denatured as to enable more efficient enzymatic digestion. Prior to denaturation and digestion, the proteins can be separated into fractions to decomplexify the analysed mixture. Let us begin by considering the choice of enzyme for protein digestion. Trypsin is the most widely adopted enzyme in BU proteomics for several reasons. It is a serine protease which cleaves at the carboxyl side, after arginine and lysine residues, with high specificity. The distribution of these two amino acids throughout the protein sequences across proteome stays relatively uniform and is within the convenient size range for MS analysers. Moreover, lysine and arginine have basic sidechains that can be protonated, making them ideal for ionization in positive mode. The tryptic peptides contain at least one basic site at both N-term and C-term ends, which greatly enhances fragmentation efficiency. Moreover, trypsin is a sufficiently small enzyme to penetrate PAGE gels, which makes it convenient for in-gel digestion of proteins. Wild type trypsin is prone to autolysis, but this issue has been fixed in commercially available enzymes. One pertinent problem is the incompleteness of digestion. When the proteins are very structured, the cleavage sites are not available for digestion. The presence of negatively charged residues near the cutting sites inhibits cleavage, especially after lysine. To address this issue, Trypsin/LysC mix can be used, since LysC is highly specific for C-terminal of Lys residues. Chymotrypsin covers protein sequences in an orthogonal manner to trypsin, because of its preference for hydrophobic amino acids, however it misses a lot of cleavage sites. Pepsin is a very small and fast enzyme, with broad specificity. It is very active in low pH and low temperature. All of these properties make it ideal for hydrogen/deuterium exchange MS studies, more of which will be covered later. Good comparison of different enzymes in BU proteomics has been done by Giansanti et al. [START_REF] Giansanti | Six alternative proteases for mass spectrometry-based proteomics beyond trypsin[END_REF].

LC-MS 2 acquisition

The peptides are then separated and analysed via LC-MS 2 . By far the most common chromatography for peptides is RP nanoLC chromatography with C18-functionalized silica, either packed or monolithic. With current instrumentation, mode than 5 000 protein identifications from a single-shot 30-minute analysis are possible, using miniscule amounts of peptide sample, on the order of 0.5 to 1µg (Kelstrup et al., 2018). In case a deeper proteome is needed, fractionation of the sample in an orthogonal manner is required: ion exchange or high pH RP fractionation.

The MS instrument methods depend on the geometry and the capabilities of the instrument, but in any case MS and MS 2 spectra should be obtained for peptide identification. The two DDA and DIA strategies employed for MS 2 have already been described in detail earlier.

Peptide identification and protein inferrence

Depending on the strategy used, different bioinformatics approaches are available for peptide identification. In DDA strategy, a software will create in-silico the theoretical tandem mass spectra, based on the provided proteome database (containing sequences and protein identities and related data) and a set of given parameters, such as the enzyme used for digestion, possible PTMs, fragmentation type, number or mis-cleavages, maxiumum shift allowed between experimental m/z and theoretical one. These in silico-created spectra are essentially lists of potential m/z values for all possible peptide fragments, based on a database. These libraries are then used to search the experimental MS 2 spectra for spectral peptide matches (PSMs)-within the user-defined tolerances. The PSMs are scored by their quality in various ways and are different depending on the search engine used. The work in this thesis was done using Mascot search engine (Matrix Science Limited, UK), which uses a probabilistic model of scoring PSMs -meaning that it tries to estimate how probable it is that the PSM happened at random. Another very important way of verifying the fidelity of identifications is the False Discovery Rate (FDR): an artificial database of random protein sequences is created (or reverse order sequences compared to the actual database used) and another decoy search is ran against it. The FDR is estimated as a percentage of decoy hits in the results. The search result stringency is usually set based on the FDR value. After peptide identification and filtering, protein identity is inferred. Peptides can either be proteospecific, or shared between multiple proteins and this association step poses a great problem. One of the partial solutions is assigning peptides to protein groups rather than individual proteins when there is ambiguity. Peptide abundances can be measured based on the area of the eXtracted Ion Chromatograms (XICs) of the theoretically most intense isotope and from these the protein intensity can be inferred in various ways.

Protein quantitation using Bottom-Up proteomics

The BU approach offers a plethora of different ways for quantitating proteins. Accurate analyte quantitation faces several challenges: ionization efficiency depends on peptide sequence and liquid phase conditions, thus it is difficult to compare different peptides; the dependence of signal response may not be linear to peptide quantity -depends on the sample complexity and the instrument properties; signal suppression may occur in co-eluting peptides. General strategies to combat these difficulties are to only compare identical peptides, in not-too-different amounts, to compare samples of similar composition and to separate well the peptides. The set of quantitation tools can be divided in absolute quantitation and relative quantitation techniques. As the name implies, in absolute quantitation techniques the MS intensities of the identified peptides are related to absolute quantity values (e.g. [ng protein] / [g of sample]), while in relative quantitation, the signal arising from the same species are matched across injected samples and their intensities compared. Most quantitation datasets published in the literature are relative as variations between a reference sample and others are usually enough to answer the biological question, while the absolute quantification is more challenging from an analytical point of view, especially so if a high accuracy quantification is expected. But "semi-absolute" quantification of proteins in a sample can be obtained from the label free quantification to estimate the abundance of a protein relative to the others in the same sample.

Absolute quantification strategies

The absolute approaches usually rely on the addition at a known concentration of isotopicallylabelled standards, which are chemically identical to some of the analytes. Examples of its use are the investigation of the protein complex stoichiometry [START_REF] Menneteau | Mass Spectrometrybased Absolute Quantification of 20S Proteasome Status for Controlled Ex-vivo Expansion of Human Adipose-derived Mesenchymal Stromal/Stem Cells[END_REF], or biomarker quantification for clinical purposes [START_REF] Brun | Isotope-labeled Protein Standards[END_REF]. Several methods are employed: intact isotope labelled protein standard absolute quantification (PSAQ or absolute SILAC), biologically synthesized artificial quantification concatemer peptides (QconCAT) and synthetic stable isotope labelled peptides (AQUA) [START_REF] Brun | Isotope dilution strategies for absolute quantitative proteomics[END_REF]. AQUA are synthetic peptides added to the digested sample, where the identical light-isotope peptide to be measured is expected to be found in the sample. In QconCAT, a recombinant and chimeric protein is designed and produced, that yields a defined set of tryptic peptides (arising from different proteins), matching the peptides one wants to study. It is added before trypsin digestion. For both QconCAT and AQUA, any additional sample treatment (such as fractionation) should be avoided, since it may lead to inaccuracies -the synthetic proteins or peptides do not behave identically as the species in the sample. Moreover, the tryptic cleavage is not performed in the same conditions in the natural protein than in the QconCAT (different sequence) or in the AQUA approach (no cleavage), leading to variations in enzymatic cleavage efficiency and quantification biases. For PSAQ, heavy standard recombinant proteins are produced in vivo, by supplementing a cell culture with stable heavy isotope-labelled amino acids. Produced, purified, and accurately quantified heavy protein can be introduced in the beginning of the sample treatment and will behave identically as the protein originating from the sample, which makes it a very reliable standard. The before mentioned strategies can be both used for targeted methods such as SRM/MRM and for global analysis (non-targeted analysis in product ion scan mode; see Chapter 2.3.1.4.6).

Relative quantification strategies

Relative quantification strategies are divided in several groups: metabolic isotope labelling, chemical labelling, enzyme labelling and label-free quantification (LFQ) [START_REF] Zhang | Protein Analysis by Shotgun/Bottom-up Proteomics[END_REF]. Here, a different kind of comparison is made, between two or more samples. Metabolic labelling is done in vivo, using nutrient sources with heavy isotopes. A great example of the technique is stable isotope labelling with amino acids in cell culture (SILAC): cell culture medium made with heavy isotopelabelled amino acids ( 15 N-, 13 C-or 2 H) is used to feed a cell culture. After several passages, most of the protein will be made up of "heavy" amino acids. The protein from such culture can be spiked and used as internal control for experimental groups made with the same cell line. Interestingly, the same concept can be used to produce SILAC mice, by feeding them with heavy isotope-labelled food. The elution times will be the same for both heavy and light peptides of the same sequence, while the MS and MS 2 spectra of heavy peptides will be shifted by a certain increment compared to the light ones of the same sequence. Although having the advantage of reliable global protein comparison, this approach suffers from increased sample complexity and is limited to cell culture and small animals, moreover, maximum of three samples can be compared at the same time [START_REF] Hilger | Triple SILAC to Determine Stimulus Specific Interactions in the Wnt Pathway[END_REF].

Chemical labelling can be done on protein (iCAT) or peptide level (iTRAQ, TMT) [START_REF] Moulder | Analysis of the plasma proteome using iTRAQ and TMT-based Isobaric labeling[END_REF]. Reagents that react with functional groups of proteins or peptides are used. The iCAT approach uses a set of cysteine-binding reagents -a light and a heavy version, to tag and compare two different samples. The iTRAQ and TMT are both based on amine-reacting compounds. These sets of compounds are developed in such a way to be isobaric, but upon fragmentation they give a reporter ion of different mass. For iTRAQ system there are 8 different compounds, while TMT has 16 isobaric amine-reacting tags, giving 8 or 16 comparison "channels", respectively. These systems allow samples to be compared by tagging them separately, each with a different isobaric compound, then mixing the samples and analysing them together. The iTRAQ and TMT allow for reliable relative quantification of proteins across samples, however they have couple of issues.

Enzymatic labelling is based on reactions with the protein sample, where a heavy isotopelabelled group is introduced to the protein/peptide via an enzyme. Elegant example is the tryptic digestion of protein in presence of H2 18 O [START_REF] Ye | 18O Stable Isotope Labeling in MS-based Proteomics[END_REF]. Upon what is effectively a catalysed hydrolysis of the peptide bond, the heavy oxygen binds to the carboxyl side of the peptide bond in the form of a hydroxyl group. Another heavy oxygen can be exchanged via reaction with one of the resonant structure of the carboxyl group. The sample treated in this way can then be compared to a trypsin-digested sample in regular water by mixing them together. Although it is a very cheap and convenient way to label peptides, a drawback of this reaction is the reversibility of the reaction. Label-Free Quantification Due to unique advantages every one of them offers, each of the labelling techniques has found its niche use. However, often incomplete labelling, resulting complexity of the sample/ complexity of the spectra, additional sample preparation steps needed, lack of high throughput implementation and often high price of the material, a quantitation technique that requires no labelling or standard addition to the sample would be preferable -the LFQ approach. Gradual improvements in quantitation strategies have led to several different software solutions that give reliable and precise quantification results. Two basic strategies are employed to calculate relative intensity value for a protein: spectral counting and ion intensity-based. The spectral counting approach relates the number of times a peptide of a protein has been identified during analysis as a measure of abundance. In the ion intensity approach, the area of the chromatographic elution peak for a certain peptide is used for quantitation. All the global proteomic analyses presented in this thesis employ the LFQ approach. Proline software was used [START_REF] Bouyssié | Proline: an efficient and user-friendly software suite for large-scale proteomics[END_REF] and the data-processing pipeline is as follows.

As already described, the acquired spectra must first be searched against a database of theoretical spectra defined by a user-provided protein sequence database (see Figure 25 for the full workflow). The search results, which consist of PSMs and suggested protein hits, submitted spectra and search parameters are then imported into the quantification software.

Validation step can be performed based on scoring systems or user-defined criteria, in an attempt to eliminate false positives. Validated peptide sequences must now be clustered into proteins. The peptides are clustered hierarchically and in protein groups, in such a way that each peptide is classified as belonging to either one or multiple indistinguishable proteins. The protein inference is done in such a way that the least possible number of protein groups is used to describe the existing list of peptides -this approach is known in logic as the Occam's razor (law of parsimony).

Proline counts the MS 2 spectra both at the peptide and at the protein level to make the relative protein group comparison. The quantitation can be based on proteospecific peptides only, or on shared peptides as well. The way the shared peptides can be used is by calculating a weighing factor as to how to distribute counts based on the count of the proteospecific peptides. The algorithm will associate validated PSMs with the chromatographic peaks by identifying the m/z value of the precursor in the MS spectra, its elution time, and associating them with the chromatographic peak m/z. Then the abundance of each ion is estimated based on the peak area of its extracted ion chromatogram. Once the peptide is identified via MS 2 events, its retention times are aligned across all LC-MS runs to recover some of the missing values, by re-extracting the intensities on MS level and trying to associate the "orphan" MS 2 spectra at the putative retention time and get a more complete set of MS 2 identifications for each peptide.

The protein abundances can now be calculated based on the peptide abundances. There are several different approaches to do this, ranging from simple functions such as sum, median or average of all peptides, the average of several most intense peptides or a set of more complex functions for protein abundance estimation such as median ratio fitting (MRF). For more details on exact mathematical procedures, one can refer to the original paper by Bouyssié et al. (10.1093/bioinformatics/btaa118). The quantitation data can now be exported for further treatment or visualization using the same tool.

Although BU improvements lead to tremendous depth of analysis, it does have great shortcomings. Commonly, the sequence coverage in most large BU datasets for most proteins is low. Besides commonly stressed points (chromatographic peak capacity and dynamic range of the instrument), there are problems inherent to the nature of the technique. Upon trypsin digestion, irregularly interspaced lysine and arginine residues in some proteins will yield peptides that are either too long or too short for analysis. Differences in ionization yields of co-eluting peptides may cause some of them not to be detected at all. Short peptides are less likely to be proteospecific, while shared peptides cannot be used for exact identification of a protein, but rather indicate the presence of at least one of several proteins. Since the information about a protein is collected in pieces, it is sometimes not possible to infer exact proteoforms and their relative abundance if several are present. An identified PTM shows that some of the protein is modified, but an absence of modified peptides does not necessarily mean that the identified protein is not modified. Even though it faces some difficulties, BU proteomics has proved itself to be a valuable tool in many fields of biology, answering increasingly complex questions (Aebersold and Mann, 2016), with constant incremental improvements being made in sample preparation, instrumentation and signal processing, making this approach increasingly robust, sensitive and precise.

Interactome study via Bottom-Up Proteomics

Knowledge of the way proteins interact to form complexes gives valuable insights about their role in the studied systems. There are several different ways that the protein-protein interactions can be studied through proteomics: affinity purification (AP) either through an affinity tag or through immunopurification (IP), enzymatic labelling and protein correlation profiling (PCP) (see Figure 26).

Two most conventional ways to study an interactome are through co-immunopurification (CoIP), using an antibody directed against the target protein or a recombinantly expressed protein with an affinity tag. The principle is to affinity purify a protein (the bait) and observe the co-purified proteins using proteomics. The interactors of a protein will appear enriched in the eluate compared to a negative control AP eluate. Specificity towards a single epitope has an inherent bias towards the protein complexes The antibody must be able to recognize the native protein, which also depends on the peptide used as an epitope. Finally, whole protein can be used for immunisation, however this approach is often impractical and has many issues. Since antibodies very commonly underperform and are expensive to produce/develop, recombinant expression of a target-affinity tag chimera can also be considered. Most systems of this type use two tags separated by a cleavage site, in order to improve AP specificity. The tags are not naturally-occurring in these proteins, so they could thus affect the interactome. Furthermore, this approach is limited to model-systems and cannot be used with some samples such as patient samples, tissues or biological fluids and can generate biases due to recombinant expression.

Two common problems in AP-MS are false positives and false negatives. False positives come from non-specific interactions with some component of the purification system -antibody or affinity matrix and these can be attenuated using a proper negative control. In the case of CoIP, negative control consists of an IP of a sample that has an identical (or similar) background/proteome, but lacks the target protein; an alternative to this would be an IP of the same sample using a mock antibody, which does not carry affinity for any protein in the analysed sample. In the case of AP of an affinitytagged protein, the proper control AP is to use the recombinant expression system expressing only the tag itself. False negatives come from unstable protein-protein interactions which fall apart during the purification process. Fairly common way of improving protein interactor recovery is through the use of cross-linking (XL). Crosslinkers are chemical agents capable of binding covalently to two functional groups on a protein, to stabilize transient protein-protein interactions [START_REF] Klockenbusch | Optimization of Formaldehyde Cross-Linking for Protein Interaction Analysis of Non-Tagged Integrinβ1[END_REF]. Formaldehyde is a fairly common XL agent and it is a reversible crosslinker, which makes it convenient to adapt to BU proteomic workflows. One obtains the information on interactors, but no exact topologically-relevant information or exact interaction network. One approach to study this is again through XL-MS, which is a structural MS technique explained in the section 2.3.3.5.

Another approach to study protein interactome is by enzymatic labelling of interacting proteins within a certain radius of the target protein. Namely, a set of techniques exist (BioID, TurboID) that are based on in-vivo biotin labelling -a recombinant chimera of a promiscuous biotin ligase and the protein of interest is expressed in either a cell line or an organism; as a consequence, native interactors of the target protein will bind to the recombinant protein version and get biotinylated due to proximity of the ligase. The interactors can be identified through proteomic analysis as having biotin modifications on their peptides or can be enriched upon biotin-streptavidin based enrichment step. This method can help improve recovery of less abundant or transient interactors [START_REF] Roux | BioID: A Screen for Protein-Protein Interactions[END_REF][START_REF] May | Comparative Application of BioID and TurboID for Protein-Proximity Biotinylation[END_REF].

PCP is based on the following principle: if two or more proteins in a sample form a complex, and are subjected to non-denaturing chromatography or some other form of non-denaturing sample fractionation, their abundance profiles will directly correlate across different fractions (Andersen et al., 2003;[START_REF] Fabre | Deciphering preferential interactions within supramolecular protein complexes: the proteasome case[END_REF][START_REF] Larance | Global Membrane Protein Interactome Analysis using In vivo Crosslinking and Mass Spectrometry-based Protein Correlation Profiling[END_REF]. It consists in fractionating the different protein complexes, usually cross-linked, via gel filtration or a density gradient. Multiple fractions are collected and the proteins analysed in each fraction. The protein signal is compared across fractions and a dynamic image of protein abundance versus elution volume/fraction number is obtained. Proteins belonging to the same complex will have relative abundance profiles with a high correlation coefficient, meaning that they elute similarly across fractions. PCP helps in resolving the heterogeneity of big protein complexes, when one protein (or a set of proteins) participates in multiple complexes of different sizes, which cannot be discerned using a simple CoIP.

Structural mass spectrometry

Introduction

Besides identifying and quantifying proteins/proteoforms in a sample, mass spectrometry experiments can be used to study the structure of proteins and protein complexes (protein-protein, protein-DNA, -lipid, -carbohydrates, -ligand complexes). To do this, the structural MS toolbox contains a plethora of different approaches that give complementary information to other biophysical techniques. Moreover, advancements in instrumentalization enable more sophisticated MS experiments to be performed, which combine multiple approaches into a single experiment.

The structural MS techniques can be roughly categorized into several groups. On the one hand, we have techniques that operate on the protein level: native MS (nMS) and TD proteomics, and since recently native TD proteomics. On the other hand, we have the techniques that operate on peptide level and include some form of labelling: cross-linking MS (XL-MS; covalent labelling) and Hydrogen-Deuterium eXchange (HDX; non-covalent labelling). The Middle-down proteomics is most closely related to TD proteomics, and employs the study of partially digested proteins.

None of the before mentioned techniques provide us with exact protein structure per se, the way X-ray crystallography, electron microscopy (EM) or nuclear magnetic resonance (NMR) do. However, each of them provides unique type of information not available to the classical structural techniques, circumventing many of the classical structural techniques' limitations, such as sample purity, sample quantity, protein size, analysis throughput, instrument availability, etc. Review of mentioned techniques is given in the Figure 27. Along with the figure, Table 1 provides the short summary of advantages and disadvantages of each described technique, compared to traditional structural techniques.

Hydrogen-Deuterium exchange -MS

Requires less sample compared to conventional structural techniques.

Limited by sequence coverage of the method. Data interpretation can be complicated (especially without an available structure/model).

Native mass spectrometry

nMS is the study of proteins and protein complexes in their native form. Using ESI, proteins are dispersed from the liquid phase at physiological pH, ionized and desolvated using heat and dry gas, to then be analysed in the vacuum of the instrument, while maintaining (at least partially) noncovalent interactions. Whether the proteins truly stay in their native conformation(s) in this transition to gas phase has been the subject of debate [START_REF] Kitova | Reliable Determinations of Protein-Ligand Interactions by Direct ESI-MS Measurements. Are We There Yet?[END_REF], however numerous studies indicate this to be case-specific behaviour. Labile protein quaternary structures and interactions with other biomolecules can indeed be maintained. Using nMS we can determine the oligomeric state of a protein complex and determine the ratios at which proteins interact, also if multiple types of complexes are formed. The strength of these interactions can be measured, and ligand binding affinity can also be determined (Kd), although this is not yet a routine practice (El-Hawiet et al., 2012), nonetheless, its potential in drug discovery is just being unveiled [START_REF] Gavriilidou | Application of Native ESI-MS to Characterize Interactions between Compounds Derived from Fragment-Based Discovery Campaigns and Two Pharmaceutically Relevant Proteins[END_REF]. Membrane proteins can be studied, although the presence of detergents needed to maintain these structures poses a problem, due to peak broadening and ion suppression [START_REF] Ilag | Drug Binding Revealed by Tandem Mass Spectrometry of a Protein-Micelle Complex[END_REF], however, many developments in MS-compatible detergents, protein entrapment methods and specific instrument settings enabled this field to develop (Keener, Zhang and Marty, 2021), being able to study some proteins ejected even directly from native membranes [START_REF] Chorev | Protein assemblies ejected directly from native membranes yield complexes for mass spectrometry[END_REF]

Top-Down Proteomics

TD proteomics enables to identify exact splicing variant of the protein (isoform) as well as to identify PTMs and their exact position, not only for one or few proteins at a time, but for relatively complex samples. It is a protein-centric approach, providing information on actual proteoforms in their integrity, as they are in the biological system, with sequence coverage much greater than BU. It circumvents the problem of protein inference and information loss regarding actual proteome complexity, which BU has. The exact number of existing human proteoforms is hard to estimate and is at least several fold greater than the number of protein-coding genes (Aebersold et al., 2018). TDP enables simultaneous and unambiguous identification of differently-modified protein variants, making it irreplaceable in the study of heavily-modified proteins, such as histones [START_REF] Greer | Top-Down Characterization of Heavily Modified Histones Using 193 nm Ultraviolet Photodissociation Mass Spectrometry[END_REF]. To illustrate the explosion of complexity between genomics and proteomics, it is convenient to use the histone example. Typical single-shot (no-replicates, no fractionation) BU proteomic LC-MS 2 analysis will nowadays result in identification of ~5000 protein entries, via peptides which may or may not be modified, using spectral search based on transcriptome-level information. One TDP study of an affinity-enriched histone fraction alone resulted in the identification of more than 500 unique proteoforms [START_REF] Greer | Top-Down Characterization of Heavily Modified Histones Using 193 nm Ultraviolet Photodissociation Mass Spectrometry[END_REF]. Another great example is the work of Tran et al. (Tran et al., 2011), showing tremendous ratio of proteoforms (>3000) versus gene products (1043). Another advantage of entire protein study is the ability to detect intraprotein and interprotein disulphide bonds, which can be very structurally informative. Although the disulphide bridge presence usually complicates MS 2 identification, there have been developments in successful, descriptive fragmentation techniques [START_REF] Rush | Top-Down Characterization of Proteins with Intact Disulfide Bonds Using Activated-Ion Electron Transfer Dissociation[END_REF]. Disulphide bridge can be connecting two regions of the same protein, two different proteins -homodimers or heterodimers, originating from one or multiple peptide chains. Usually, the MS analysis is done on denatured proteins, however as we approach high MWs, it becomes difficult or impossible. Here we enter the territory of the hybrid approach of native TD, which was mentioned earlier. It should also be noted that TD is very often used as a targeted analysis technique, wherein is its original application, while the global proteomic TD analysis approach has only begun to be popularized since recently.

For all its theoretical advantages, TDP has its own set of technical hurdles which impede it from being on par with BU, all associated with the difficulties of analysing entire proteins. The set of problems can be divided in several categories: the protein separation, ionisation, MS detection, identification/ data treatment.

Coeluting proteins are often difficult to analyse at the same time, mainly due to the ionization competition between proteins, whereby the smaller proteins dominate the large ones in ionization efficiency. Since no single biochemical property would be sufficient to be used as a parameter for their separation, it is common for global TDP experiments of very complex protein mixtures, such as lysates, to employ multi-dimensional separation [START_REF] Cai | Top-Down Proteomics of Large Proteins up to 223 kDa Enabled by Serial Size Exclusion Chromatography Strategy[END_REF][START_REF] Mccool | Deep Top-Down Proteomics Using Capillary Zone Electrophoresis-Tandem Mass Spectrometry: Identification of 5700 Proteoforms from the Escherichia coli Proteome[END_REF]Tucholski et al., 2019), where orthogonal techniques are used together, such as SEC and RPLC, IEC (ion exchange chromatography) and RPLC, or electrophoresis techniques (gel-eluted liquid-fraction entrapment electrophoresis -GELFrEE, capillary zone electrophoresis -CZE) and RPLC. First separation is done offline and can be done on native proteins (SEC, native GELFrEE) or denatured proteins (GELFrEE) and several fractions are collected, each having a certain mass range. This is followed by an on-line RP separation and protein identification. Several different separation modes can be employed to reduce the dynamic range of the sample and improve proteome depth, example being the work of McCool et al. who identified 5700 proteoforms from E. coli lysate by pre-fractionation via sequential SEC and RPLC to analyse fractions on CZE-MS [START_REF] Mccool | Deep Top-Down Proteomics Using Capillary Zone Electrophoresis-Tandem Mass Spectrometry: Identification of 5700 Proteoforms from the Escherichia coli Proteome[END_REF]. Another interesting example are several works from the group of Ying Ge, who employ serially-connected SEC columns of different porosity (sSEC) in off-line fractionation combined with on-line RPLC [START_REF] Cai | Top-Down Proteomics of Large Proteins up to 223 kDa Enabled by Serial Size Exclusion Chromatography Strategy[END_REF]Tucholski et al., 2019). Finally, it has been proven that a single-dimension separation can be successfully achieved for small proteins if long gradients and >1m columns are used [START_REF] Shen | High-resolution ultrahigh-pressure long column reversed-phase liquid chromatography for top-down proteomics[END_REF].

Ionization efficiency and ionization competition play a determinant role in the detection of proteins. Although ESI is very efficient at creating protein ions, proteoform signal intensity is divided across all the different charge states it exists in. Besides ionization competition, this signal loss due to spreading is one of the culprits behind the decrease in signal to noise ratio for big proteins and is more pronounced in denatured proteins as they exist in a higher number of charge states than in nMS. For this reason, best ionisation results are obtained for 10 to 50 kDa proteins, with the upper limit on the analysis of denatured proteins at ~200 kDa, using a Q-TOF instrument [START_REF] Cai | Top-Down Proteomics of Large Proteins up to 223 kDa Enabled by Serial Size Exclusion Chromatography Strategy[END_REF]. This problem has been addressed with the Proton Transfer Charge Reduction technique (PTCR), available in the Orbitrap Eclipse Tribrid mass spectrometer, which is used to reduce charge states of the select m/z ranges of high MW proteins [START_REF] Huguet | Proton Transfer Charge Reduction Enables High-Throughput Top-Down Analysis of Large Proteoforms[END_REF]. The signal suppression due to presence of salts and ionisable detergents represents a great issue for TDP experiments and special care must be taken to eliminate them. Usage of RPLC coupled to the MS helps separate such ions from the eluting proteins.

For improved resolution in order to interpret complex spectra and better high-MW protein signal recovery, FT-ICR instruments are very often the norm for deep TDP exploration (Anderson et al., 2017; Tucholski et al., 2019). However, adapting data acquisition strategies can help improve the quality of the acquired data and the number of identified proteoforms for a Q-Orbitrap system [START_REF] Fornelli | Advancing Top-down Analysis of the Human Proteome Using a Benchtop Quadrupole-Orbitrap Mass Spectrometer[END_REF].

Fragmentation techniques used in TDP are varied, however, to obtain better sequence coverage while maintaining labile PTMs, the most worthy have proven ECD and ETD techniques, although the fragmentation yield is lower compared to CID techniques, especially HCD. Hybrid technique exists, which combines these two technologies, termed EThcD and ETciD. Some particular studies of disulphide bridge-containing proteins employed activated-ion ETD (AI-ETD) to efficiently fragment and localize the highly-linked regions and localize these bonds [START_REF] Rush | Top-Down Characterization of Proteins with Intact Disulfide Bonds Using Activated-Ion Electron Transfer Dissociation[END_REF]. This technique uses infrared photons to improve the fragmentation efficiency of the ETD. Another novel technique which does not disrupt the labile PTMs but gives great fragmentation yield is ultraviolet photodissociation (UVPD), which uses high-energy photons to disrupt the peptide bonds [START_REF] Shaw | Complete Protein Characterization Using Top-Down Mass Spectrometry and Ultraviolet Photodissociation[END_REF] and is becoming widely accepted [START_REF] Shaw | Complete Protein Characterization Using Top-Down Mass Spectrometry and Ultraviolet Photodissociation[END_REF][START_REF] Cannon | Ultraviolet Photodissociation for Characterization of Whole Proteins on a Chromatographic Time Scale[END_REF][START_REF] Greer | Top-Down Characterization of Heavily Modified Histones Using 193 nm Ultraviolet Photodissociation Mass Spectrometry[END_REF].

Integral membrane proteins usually demand the use of detergents, which rendered them incompatible with the TDP. However, developments in the area of MS-compatible detergents such as Dodecyl-β-D-maltoside (DDM), enabled the membrane protein study. Some studies indicate that many proteins can be recovered without the special use of detergents [START_REF] Catherman | Large-scale top-down proteomics of the human proteome: membrane proteins, mitochondria, and senescence[END_REF] Many technical aspects have drastically improved over the recent years and the TDP has grown from a niche approach to a vibrant community of researchers with established guidelines [START_REF] Donnelly | Best practices and benchmarks for intact protein analysis for topdown mass spectrometry[END_REF].

Middle-Down proteomics

Middle-Down proteomics is an approach which employs partial enzymatic digestion of a protein and subsequent proteomic analysis. The digestion yields longer peptides (50-60 amino acid residues), which makes it convenient to more precisely determine exact proteoforms, when many PTMs/ splicing variants are anticipated. The MDP has particularly found its niche in histone code study [START_REF] Sidoli | Middle-down proteomics: a still unexploited resource for chromatin biology[END_REF]. Namely, histones are regulated via their terminal regions or "tails", which are heavily post-translationally modified in a dynamic manner. Bottom-Up yields short peptides which cannot be properly associated with a single proteoform and isomeric PTM-peptides cannot be separated properly to generate unambiguous MS 2 spectrum, while MDP is able to unambiguously attribute PTMs to their original protein. The techniques often used are HILIC for peptide separation, ECD or UVPD for peptide fragmentation and even IM [START_REF] Shliaha | Middle-Down Proteomic Analyses with Ion Mobility Separations of Endogenous Isomeric Proteoforms[END_REF]. Another great field of application of MDP is the study of mAb and Antibody-Drug Conjugates, whereby the Fc and Fab fragments can be cleaved by specific enzymes such as IdeS or Papaïn in order to facilitate their MS analysis [START_REF] Beck | Characterization of Therapeutic Antibodies and Related Products[END_REF].

Cross-linking -mass spectrometry (XL-MS)

XL-MS is a structural MS approach where purified protein complexes are chemically labelled with cross-linkers, which are able to react with two different protein sites to create covalent bonds across neighbouring protein structures. These cross-links can occur within the same protein, across two proteins or be "dead-ended", meaning the reagent only bound to one site, while the other side remains unbound/neutralized. The cross-linked protein complex is then digested into peptides, separated and analysed using conventional BU techniques. The data is however, interpreted differently, in attempt to find and sequence cross-linked peptides, as to determine between which sites was the cross-link established. This way, one learns which proteins/ protein regions are in immediate vicinity to each other. The cross-linking agents, depending on their length, can be used as molecular rulers, since the physical length of the molecule determines the distance across which they can react. Great improvement in the XL-MS workflow was made with the introduction of MS-cleavable cross-linkers, which asymmetrically fragment upon ion activation into two fragments, leaving recognisable signature on the peptide backbone, thereby simplifying the peptide identification from MS 2 spectra. The reader is referred to a great review by Rappsilber and O'Reilly for further information on this topic (O'Reilly and Rappsilber, 2018).

Hydrogen-Deuterium eXchange -Mass Spectrometry

HDX-MS experiments are based on the capacity of protein backbone amide groups to exchange their only hydrogen for deuterium, when exposed to deuterated water. Propensity of an amide group to participate in H/D exchange will depend on the exposedness to the solvent, flexibility of the region within it is located and whether it participates in any hydrogen bonds (stabilisation of secondary structures). The deuterium uptake can be measured as a shift in mass compared to the non-deuterated state. By locally determining the relative deuterium uptake, we can infer structural properties about a protein or a protein -ligand complex, e.g. if a binding of a ligand stabilizes a protein or induces a conformational change, which would manifest itself in changed deuterium exchange profile along the protein sequence.

HDX experiment begins by deuterium incorporation: a protein or a protein complex is diluted in a D2O-based buffer (see Figure 28). The rate of exchange will depend on several parameters: the D2O diffusion rate, the involvement of the hydrogen in a bond, local electrostatic properties of the environment, the pD of the solution (pH equivalent for deuterated systems) and temperature (Bai et al., 1993). If the specific protein region is buried inside the protein core or is facing an interaction surface with another protein, or is intrinsically very compact, the D2O access will be limited and thus the exchange rate kinetics will be very slow. Thus, one can expect loop regions to show high exchange rates, which is the case, with few exceptions. These exceptions are attributed to the existence of the locally-ordered water clusters, forming hydrogen bonds with an amide group. When it comes to hydrogen bonds, involvement of hydrogen in stable secondary structures will greatly impede any exchange. It is important to note that the deuterium uptake can occur in places other than amide group, however these reactions happen at a time scale either too short (lateral chains of amino-acids) or too long (C-H) for practical study by MS. This reaction is reversible -as soon as the protein is in H2O-based solution, the back exchange occurs. To prevent this, a sample must be cooled down to 0-4°C and be acidified -optimal pH to minimize back exchange has been found to be 2.5 [START_REF] Berry | Hydrogen Deuterium Exchange Mass Spectrometry of Oxygen Sensitive Proteins[END_REF]. Although it is common to study the deuterium uptake, its release kinetics can also be studied, by first fully deuterating a protein, then measuring the back exchange [START_REF] Woods | High resolution, high-throughput amide deuterium exchange-mass spectrometry (DXMS) determination of protein binding site structure and dynamics: Utility in pharmaceutical design[END_REF], but this approach remains limited by incomplete protein deuteration.

In order to get better spatial resolution on different regions of the protein, the deuterium uptake must now be studied locally. To do this, there are two ways: BU and TD. As one might guess, the bottom-up implies the proteins to be digested, and is more common. The TD approach employs the fragmentation of intact proteins inside the instrument [START_REF] Pan | Hydrogen/Deuterium Exchange Mass Spectrometry with Top-Down Electron Capture Dissociation for Characterizing Structural Transitions of a 17 kDa Protein[END_REF]. Collision-induced fragmentation induces the so-called hydrogen/deuterium scrambling: when the high-energy transfer occurs between the analyte and the collision gas, the hydrogen atoms re-arrange themselves, thus scrambling the deuteration rate information. However, employing radical-based techniques, such as ECD or ETD, no re-arrangement occurs, due to less energy transfer and faster fragmentation events [START_REF] Rand | Measuring the Hydrogen/Deuterium Exchange of Proteins at High Spatial Resolution by Mass Spectrometry: Overcoming Gas-Phase Hydrogen/Deuterium Scrambling[END_REF]. Although TD results in less back exchange compared to bottom-up, due to shorter sample manipulation, it is difficult to obtain good sequence coverage for larger proteins [START_REF] Pan | Hydrogen/Deuterium Exchange Mass Spectrometry with Top-Down Electron Capture Dissociation for Characterizing Structural Transitions of a 17 kDa Protein[END_REF] and synthetic control peptides should be used to check for potential scrambling.

For bottom-up, the deuterated protein is put in low temperature and pH of 2.5, in order to quench the reaction and prevent back exchange. Usually, the protein is denatured and reduced at the same time, using additives in the buffer (guanidinium chloride, TCEP). This is followed by protein digestion -to do this efficiently, a temperature of around 10°C is needed and enzymes compatible with acidic conditions are used. The most commonly used enzyme in HDX is pepsin, since it cleaves very rapidly even in low temperature, is stable and operates well at low pH conditions. Pepsin has lower cleavage specificity, however, resulting peptides are reproducible for the same proteins. Other enzymatic options are being investigated (Ahn et al., 2013). After digestion, the peptides are analysed using RPLC-MS. Nowadays, the entire HDX-MS workflow can be automatized via programmable robot sample manipulation and on-line protein digestion, directly coupled to LC-MS [START_REF] Espada | A Decoupled Automation Platform for Hydrogen/Deuterium Exchange Mass Spectrometry Experiments[END_REF]Watson et al., 2021). Data treatment following the HDX-MS analysis is relatively difficult: MS E spectra obtained on a pepsin digest are more difficult to interpret than the ones based on tryptic peptides (since no cleavage specificity can be used). Furthermore, after deuteration, isotopes from two different peptides can overlap, since short LC gradients, which are necessary to reduce back exchange, result in more complex spectra due to co-elution. Usually, manual curation of spectra is required and obtaining full-sequence coverage can be difficult, which limits the HDX use for large protein complexes. Recent work from our group [START_REF] Lesne | Conformational maps of human 20S proteasomes reveal PA28-and immuno-dependent inter-ring crosstalks[END_REF] on the HDX-MS study of proteasome complexes, has raised the bar in terms of protein size to be successfully studied by this technique and has brought new structural insights on proteasome-regulator complexes above 900 kDa in size. As mentioned before, deuterium uptake is usually compared to the non-deuterated reference, and is most effectively presented using a protein sequence-associated heatmap (see Figure 28). This heatmap can also be plotted onto a model or a structure obtained via different technique, for more insightful data visualisation (Bouyssié et al., 2019).

This technique is successfully applied to membrane protein studies, using many innovative approaches (Mehmood et Great advantage of HDX-MS is the completely native approach of the isotopic labelling in solution. Besides, it consumes less sample than traditional structural approaches (typically ~1000 pmol for a complete kinetics in triplicate injections), while providing useful complementary structural information for a great mass range of protein complexes. and after certain time of deuteration, the reaction is quenched and the protein is digested into peptides. Exposed or flexible peptides will manifest a positive shift in mass due deuteration, which can be presented in a form of a heatmap or visualized onto the 3D structure of a protein or a protein complex.

Mass Photometry

Mass Photometry (MP), also known as interferometric scattering mass spectrometry (iSCAMS), is a very recent technique, only first published in 2018 [START_REF] Young | Quantitative mass imaging of single biological macromolecules[END_REF]) that allows the determination of the molecular weight at the single-molecule level. When a protein in solution is placed on the glass coverslip, the molecules will begin to adhere to the glass and locally displace the water on the solution-glass interface, causing the local refractive index to change (Figure 29.A). The technique is based on illuminating the interface between the glass and the sample solution and measuring the contrast produced in light interferences, which is directly proportional to the mass of the molecule that caused the event. The contrast-to-MW dependence is fairly linear for most proteins of globular shape (~2% for 50-800 kDa range) and deviates only for filamentous proteins (Sonn-Segev et al., 2020).

The mass photometer records a video of binding events and then sums the binding events into a histogram for a recorded contrast range (Figure 29.B). By using a mix of standards, the mass photometer can be calibrated to produce a histogram of the heterogeneity of particles of different masses in the sample. The advantages of this technique are numerous. The measurement is done in solution, on native molecules, without any prior labelling. Moreover, it is compatible with a broad range of detergents, buffers and additives. It consumes very little sample (3-10 μl of 100 pM-100 nM solution) and actually counts individual molecules. It has a huge mass range: between 40kDa and several 5MDa. The downside of this technique is its low mass resolution (full width at half-maximum between 20 and 100 kDa, depending on the size of the protein).

All of these traits make it suitable for the study of oligomeric states of large proteins and the formation of large protein complexes, such as the proteasome. Moreover, the formation of these complexes can be observed in real time, opening a whole range of possibilities for experimentation. Very recently, Olerinyova et al. showed that it is feasible to use this technique to study membrane proteins [START_REF] Olerinyova | Mass Photometry of Membrane Proteins[END_REF].

The study of proteasome complexes using mass spectrometry

The proteasome has a long track record of being studied by mass spectrometry, at first only for the study of its core composition by proteomics and, as the techniques got developed, for the structural study as well. Since its discovery in 1978 [START_REF] Ciehanover | A heat-stable polypeptide component of an ATP-dependent proteolytic system from reticulocytes[END_REF], the proteasome has been the object of intense study, because of its involvement in a great number of processes, as described in the Chapter 2.2. Because of this polyvalence of the proteasome, there are many different contexts of proteasome study: cell cycle regulation and cancer, antigen peptide processing and immunity, cell cycle regulation and cell differentiation, protein catabolism and fasting, failure to degrade proteins and diseases related to protein aggregation, such as Alzheimer's disease, just being some of the main examples (Coux, Zieba and Meiners, 2020). For this reason, there are many different questions to be answered.

The BU proteomics approach has been used on many different occasions to identify proteasome subunits and answer following questions. What is the 20S proteasome composition / what proteasome types can be found in a sample and in which ratios Moreover, the BU proteomics, often in association with two-dimensional gel electrophoresis, which helps the separation of isoforms, has for long been used to identify PTMs in the proteasome subunits or its partners [START_REF] Zong | Two-dimensional electrophoresis-based characterization of posttranslational modifications of mammalian 20S proteasome complexes[END_REF]. More recently, the TD approach has been developed. It is the best suited technique for this task because it is able not only of identifying individual PTMs, but characterizing exact proteoforms (exact sequence and all the PTMs), even when multiple exist, as shown is several occasions [START_REF] Gersch | A Mass Spectrometry Platform for a Streamlined Investigation of Proteasome Integrity, Posttranslational Modifications, and Inhibitor Binding[END_REF]. Next, although the ELISA approach has been widely used to determine the absolute quantity of the proteasome in a sample [START_REF] Majetschak | Immunological methods to quantify and characterize proteasome complexes: Development and application[END_REF], BU proteomics-based approaches using addition of heavy isotope-labelled standards and targeted analysis approaches have been developed to measure all proteasome subtypes in a single high throughput analysis [START_REF] Menneteau | Mass Spectrometrybased Absolute Quantification of 20S Proteasome Status for Controlled Ex-vivo Expansion of Human Adipose-derived Mesenchymal Stromal/Stem Cells[END_REF]. The team of Yifat Merbl published a novel way of identifying proteasome substrates, through analysis of the peptides processed by the proteasome (Wolf- [START_REF] Wolf-Levy | Revealing the cellular degradome by mass spectrometry analysis of proteasome-cleaved peptides[END_REF]. This methodology holds many promises for the study of the proteasome role -inferred de facto from its substrates.

Structural biology techniques brought highly valuable information on the proteasome structure, by determining the 20S core structure first by negative staining EM, then by crystallography. X-ray crystallography also provided high-resolution structures of proteasomes alone or in the presence of covalent inhibitors [START_REF] Huber | Immuno-and constitutive proteasome crystal structures reveal differences in substrate and inhibitor specificity[END_REF] and regulators (Knowlton et al., 1997;Huber and Groll, 2017).

Recent cryoEM studies determined the structure of the 20S with several different regulators (Toste Rêgo and da Fonseca, 2019; Xie et al., 2019). Exceptional effort has been made to determine the 26S structure and try to reconstruct the catalytic intermediate-states of the 26S (X. Huang et al., 2016;[START_REF] Mao | Structure, Dynamics and Function of the 26S Proteasome[END_REF]. The combination of cryo-EM and covalent cross-linking coupled to mass spectrometry (MS) allowed to refine the structure of these different catalytic intermediates (Beck et al., 2012;[START_REF] Wehmer | Structural insights into the functional cycle of the ATPase module of the 26S proteasome[END_REF].

Despite such great achievements, these high-resolution methods are still limited by either the size, the heterogeneity, or the dynamics of the purified complexes. This is evidenced by the small number of structures of the 20S bound to regulators and by the numerous missing regions (too dynamic) in these resolved structures. The emergence of native MS contributed to a better understanding of the 20S assembly mechanism [START_REF] Sharon | Mass Spectrometry Reveals the Missing Links in the Assembly Pathway of the Bacterial 20 S Proteasome[END_REF] and of the architecture of the 19S lid [START_REF] Taverner | Subunit Architecture of Intact Protein Complexes from Mass Spectrometry and Homology Modeling[END_REF] as well as the identification of new partners of the 20S proteasome extracted from rat liver (Moscovitz et al., 2015). Structural MS is thus not limited by the same factors as the traditional structural biology approaches, such as flexibility of the domains (X-ray crystallography, cryoEM), or complex size and multiple subunits (such is the case with NMR).

Context and objectives

This thesis project was realized in the Proteomics and Mass Spectrometry of Biomolecules team, led by Dr Odile Schiltz, at the IPBS, Toulouse, France. On the one hand, the team specializes in the analysis of proteins from multiple aspects: BU and TD proteomics, targeted protein analysis and absolute quantification and structural MS (nMS and HDX-MS). On the other hand, the team specializes in proteasome biology. For the realization of this project, we established a collaboration with the team of Charles Pineau at the Irset, Rennes, who specializes in the biology of spermatogenesis. The Pineau team provided us with precious samples and worked with us to interpret our findings in their biological context, thus complementing our competences.

In a broader sense, the goal of the project was to elucidate the role of the spermatoproteasome upon germ cell differentiation. More precisely, we wanted to understand what makes spermatoproteasome unique from a structural and functional point of view and how this relates mechanistically to its role. How does the proteasome complex composition change throughout spermatogenesis? Does the incorporation of an alternative, tissue-specific α4s subunit into the 20S proteasome core lead to structural changes? If so, what are the different structural features and how does this relate to changes in catalytic behavior and regulator binding? What are the substrates in spermatogenesis that are specific to the spermatoproteasome?

To answer these questions, a set of experiments was conceived, with the attempt to maximally benefit from the team's expertise in MS and their 15 years-long experience in the MS-based study of the proteasome. While conducting the principal project, I had the opportunity to participate in several international collaborations established with the team, concerning MS-based proteasome analyses

Results

The manuscript of the article that is entirely concerned with the work done on this thesis project and which represents a cohesive set of experiments and their insights is presented in the Chapter 4.1.2. The said article is submitted to the Proceedings of the National Academy of Sciences (PNAS) and is currently under review (03/10/2021). The entirety of the work in the Chapter 4.1.2 was done by me (Dušan Živković), except for the molecular dynamics analyses.

Additional experiments were conducted and yielded comprehensive results, but due to their incompleteness or lack of cohesion with the principal narrative of the article in preparation, they are included in a separate chapter named Additional experiments.

Besides the study of the spermatoproteasome, I was involved in several related projects that resulted in peer-reviewed publications. The objective, results and my contribution to these studies are briefly described in a separate chapter and the articles themselves are presented in the Annex chapter.

Besides the work presented in the Results section, great efforts were put in two aspects of the project, which never reached their goals: the recombinant production of the PA200 regulator and the production of the PSMA8 KO mouse model. Two consecutive papers were published by other groups on the study of both of these aspects: first, on the structural characterization of the PA200-20S complex by da Fonseca team, Cambridge, UK (Toste Rêgo and da Fonseca, 2019); and second on the phenotype study of the mouse PSMA8 KO model, by the Pendas team in Salamanca, Spain (Gómez-H et al., 2019). Both of these publications greatly diminished the impact of the two aspects of the project, which were then terminated due to high cost of development. However, collaborations were established with both groups. The da Fonseca team provided us with the PA200 material for the publication in preparation (Chapter 4.1.2), while a new project with the Pendas team was established, concerning the in-depth proteomic characterization of the PSMA8 KO model.

Proteasome complexes experience profound structural and functional

rearrangements throughout mammalian spermatogenesis

Objectives

The study was divided in two sections, as presented in the Figure 30, below. We first studied the spermatoproteasome interactome. More precisely, we studied how the spermatoproteasome behaves throughout germ cell differentiation, in terms of total amount in the cell, the types of interactors it associates with and their relative abundance. This was achieved through BU proteomics approaches, combined with CoIPs. We used a whole organ CoIP approach for the interactome study, due to the availability of the material in large quantities. However, separating the germ cells by stages of development enabled us to obtain a dynamic image of the proteasome composition. The comparison between the proteasome CoIP samples and the lysates shows us whether the proteasome components that are expressed in the cell are indeed incorporated into the complexes and what proportion exists non-associated in the cytosol. The high sensitivity of BU proteomics enables the detection and quantitation of low-abundant interactors.

In the second part of the study, we wanted to compare the structural feature of the standard and spermatoproteasome and see the implications of those differences in the way they interact with a subset of interactors that we identified to be relevant in the spermatogenesis context (based on the findings from the first section).

To first compare the structural feature of the two proteasomes, we employed TD proteomics to characterize and relatively quantify the exact proteasome proteoforms in the testis, including those of α4s and α4. Differences in the primary structure (such as PTMs or truncations) could, at least partially explain differences in behavior/function. We further employed HDX-MS to compare the structural features of α4s and α4, more precisely, their stability and exposedness to the solvent (complementary to traditional structural techniques, yet very informative). These findings were further supported by molecular dynamics simulations.

We then wanted to see if these differences in structure convey differences in the way proteasome interacts with relevant regulators (using Pull-Down assays), and/or if they convey differences in proteasome activity.

Article under review in PNAS (2021)

Proteasome complexes experience profound structural and functional rearrangements throughout mammalian spermatogenesis

Introduction

Spermatogenesis is a process of cell differentiation, whereby a part of the population of stem cells called spermatogonia (SPG) enter the differentiation pathway and develop into spermatids, which then enter the spermiation process to become fully developed spermatozoa. Spermiogenesis is the differentiation of haploid round spermatids into elongated spermatids. During this maturation, a number of cell remodeling events are completed, including several critical cell-specific remodeling processes such as DNA condensation, mitochondrial reorganization, production of the flagellum and cytoplasm removal (1). SPG first divide mitotically and develop into spermatocytes (SPC), which enter meiosis. Meiosis requires duplication of the genetic material, its condensation, recombination between the maternal and paternal homologue chromosomes and re-distribution of the recombined chromosomes into separate cells. This is immediately followed by another division resulting in daughter cells called spermatids (SPT) with only one recombined copy of each chromosome. This entire process is facilitated by the Sertoli cells (SER) that serve to support developing gametes (2). Recombination is a particularly lengthy process requiring the formation of special bridges across paired chromosomes, called synaptonemal complexes (SCs), which enable the exchange of parts of chromosomes (3). Overall, spermatogenesis and spermiogenesis are both proteolysis-intensive processes with high protein turnover that require intense engagement of the primary proteolytic machinery of the cell, the proteasome (4).

The proteasome is a macromolecular proteolytic machinery in charge of controlled degradation of proteins (5). The proteasome core, also known as the 20S proteasome, is a symmetric, barrel-like structure that consists of a catalytic chamber formed by β subunits, which is sealed on both ends by a ring of α subunits (5)(6)(7). The 20S associates with various proteins and protein complexes, the most notable being the proteasome activator complexes 19S regulatory particle (to form the 26S proteasome), PA28αβ, PA28γ and PA200. These regulators modify the 20S activity and substrate specificity in order to regulate various processes such as cell division, differentiation, heat shock response, DNA repair, immune response, apoptosis, and many others (8)(9)(10)(11)(12)(13)(14)(15). Moreover, the proteasome complexity is further increased by the presence of several proteasome subtypes where one or several subunits of the constitutive complex are replaced by alternative isoforms. Thus far, in addition to the constitutive 20S proteasome (c20S), immuno-(i20S), thymo-(t20S), and spermatoproteasomes (s20S) have been identified and described in the literature (16).

The s20S is a proteasome subtype in which the standard α4 subunit (PSMA7 gene) is replaced by α4s (PSMA8 gene) that is expressed exclusively in gamete cells (17). Previous studies in PSMA8-/-KO mouse models have established that the α4s subunit is essential for spermatogenesis (14,18). Functionally, mammalian s20S has been shown to degrade acetylated histones via association with PA200 and play a role in DNA damage repair in spermatocytes and the maturation of spermatids (11,14,19). Double strand break (DSB) repair was shown to be dependent on s20S-mediated degradation of non-histone substrates (19,20). Cell cycle-mediating proteins were also reported as substrates of s20S (21,22). However, despite these reports, s20S remains understudied and there is currently a lack on basic information on the exact nature and relative stoichiometry of the regulators binding to the s20S during spermatogenesis. In this context, revealing the dynamics of the 20S proteasome composition and partners throughout spermatogenesis could help explain how the s20S functions are conveyed, and further elucidate the underlying molecular mechanisms.

Here we use advanced mass spectrometry (MS)-based approaches to map interactomes of mammalian s20S during male germ differentiation and compare them with those of c20S. Our analyses in whole testes and in isolated male germ cells revealed specific s20S partners and profound changes in the dominant 20S proteasome subtypes present throughout differentiation. Incorporation of α4s into s20S was strikingly correlated with an increased association with 19S, PA200 and PI31 regulators, and with an overall proteasome activation, especially regarding its trypsin-and chymotrypsin-like activities. Moreover, using hydrogen-deuterium exchange (HDX)-MS and molecular dynamics simulations we identified conformational differences between α4 and α4s, providing a molecular rationale for the observed differences between c20S and s20S.

Results

Spermatoproteasome represents a major fraction of total 20S proteasome in mammalian testes.

To examine differential expression patterns of s20S and c20S, as well as other proteasome-associated factors, we analyzed tissue-specific human proteome maps (17) and observed that the expression pattern of the PA200 and 19S regulators closely follows that of α4s (Fig. S1A). We noted that both male and female gonads express s20S (Fig. S1A), but given that the only reported role for s20S is in male germ cells we decided to focus our analysis on whole testes and purified male germ cells. This does not exclude that the s20S may play a still unknown role in females.

To map the relative abundance of s20S, we used freshly frozen bovine testis lysate where the s20S is highly expressed. We immunopurified (IP-ed) 20S proteasome using the MCP21 antibody that recognizes the α2 proteasome subunit (present in all 20S proteasome subtypes), as described previously (23). The 20S proteasome was then further separated from its regulators by an additional Size Exclusion Chromatography (SEC) step (24). After IP and SEC, the purified 20S proteasomes were analyzed on a LC-MS system, using a previously optimized method (25). We were able to identify not only the proteasome subunits that are conserved among the different 20S subtypes, but also the specific catalytic subunits of the c20S (β1, β2, β5) and the i20S (β1i, β2i, β5i), as well as the testes-specific α4s subunit of the s20S (Fig. 1A; Table S1). Taking the β5 subunit abundance as reference, the results show that most of the 20S proteasomes in testis contain exclusively the three constitutive catalytic subunits (64% ± 1 %), while some complexes with different combinations of constitutive and immune subunits are also present (26,27) (Table S1). Additionally, we could quantify the proportion of s20S using the MS signals of α4 vs. α4s subunits, by assuming that they have similar ionization yields based on their high sequence identities (Fig. S1B). In this way we established that the s20S constitutes 48 ± 1 % of the total testis immunopurified proteasomes, which will thereafter be called "testes 20S pool". In addition to quantifying different proteasome species, we were able to observe various PTMs (Table S2), that were in accordance with published data (25,29,30). Interestingly, we noticed that α4s harbors the same PTMs as the α4 proteoform, i.e. loss of the initial methionine and N-terminal acetylation. Additionally, we observed that repeated freeze-thaw cycles led to formation of truncated versions of α4 and α4s subunits that lack the last two amino acid residues (Fig. 1B-C), which were later confirmed by LC-MS/MS Top-Down (TD) sequencing (Fig. 1D). A similar loss in mass, corresponding to a truncated version of α4, was recently described in rat and rabbit (30). Given that our results indicate that these lighter α4/α4s isoforms are produced upon storage, most likely via proteolytic cleavage of their solvent accessible C-termini, we consider them unlikely to be biologically relevant. We also noted +73 Da and -13 Da mass differences observed for β6 and β2, respectively (Fig. S2). TD sequencing confirmed that in our sample these subunits differ from the official (curated) Swiss-Prot entries by 1 amino-acid (G233E for β6 and N252T for β2) but are in agreement with predicted TrEMBL entries. These mutations are barely visible using bottom-up proteomics where proteins are commonly identified with only a few peptides, illustrating the benefits of TD-MS for proper proteoform characterization and database curation. Taken together, our TD-MS analysis of the immunopurified proteasome pool from bovine testes revealed that s20S represents a major fraction of the total proteasome, suggesting a key functional role.

The s20S has its specific set of interactors in bovine testes.

To further characterize the s20S in gamete cells, we mapped its interactome using an IP strategy. We performed two IPs from bovine testes lysates using either an anti-α4s-specific antibody generated according to (31), and validated as shown in Fig. S3), which targets only the s20S, or an anti-α2 antibody, which recognizes all proteasome types, and compared the relative abundances of coimmunopurified proteins. In a first set of experiments, lists of proteasome interacting proteins (PIPs) specific to α2 and α4s were obtained by comparing the anti-α2 or anti-α4s IPs against a control IP (antibody directed against rat CD8-OX8, as used previously (32)). 3867 proteins were validated and quantified in the anti-α2 and anti-α4s IPs, with a total of 1177 proteins exhibiting over a 2-fold change (FC) enrichment (significance threshold at p<0.05) compared to the control (Fig. S4,S5). To highlight putative specific or enriched components in c20S and s20S interactomes, we compared the relative abundances of each of these proteins in the anti-α2 IP (total 20S) vs. anti-α4s IP (s20S). To do so, protein abundances were normalized based on the 20S content in each IP (estimated as the average of the abundances of the non-catalytic 20S subunits), then the fold changes (anti-α4s IP / anti-α2 IP) and their significance (p-value) were calculated and represented as a Volcano plot (Fig. 2A). Here an unequal distribution of enriched proteins is observed, skewed towards the anti-ɑ4s IP. One possible explanation for this observation is that the anti-α4s antibody might fetch 20S interactors that appear below the detection threshold when analyzing the whole 20S interactome (i.e., with the anti-α2 antibody only). However, we also acknowledge that the anti-α4s polyclonal antibodies used may display more non-specific bindings, compared to monoclonal antibodies used in the anti-α2 IP. We therefore looked specifically at proteins that might be of interest, namely proteasome-and ubiquitin-related proteins.

The distribution displayed on Fig. 2A highlights the specific composition features of c20S and s20S complexes. Only proteins that were enriched (FC>2, p-value <0.05) in anti-ɑ2 or in anti-ɑ4s IPs compared to the control IPs are displayed in this comparison. We observed that the non-catalytic 20S subunits (except α4 and α4s -PSMA7 and PSMA8 genes, respectively) are found tightly centered around the y-axis (mean FC = 1.0 +/-0.2), showing that the normalization of the dataset is well done and that there are no major variations in their relative abundance. As expected, the bait of the anti-α4s IP is found on the right (α4s protein -PSMA8, FC = 6.4, p-value = 0.016) whereas the subunit replacing it in the c20S is found on the left (α4 protein -PSMA7 gene, FC = 0.16, p-value = 12.E-4). We then used the relative abundances of the α4 (PSMA7) and α4s proteins (PSMA8) in the anti-α2 IP to estimate the part of s20S in the total pool of 20S proteasome in bovine testis (42 ± 7%; Fig. 2B), which is in good agreement with the results obtained by the TD approach. On the other hand, the anti-α4s IP contains 95 ± 9% of s20S (Fig. 2B), confirming the exclusive nature of the α4s subunit incorporation into the s20S (14,18). Taken together, our data provide evidence that the α4s and α4 subunits do not co-exist within a hybrid 20S proteasome. This subunit exclusivity is in agreement with what has been previously observed for some i20S catalytic subunits (33,34), and could result from exclusive expression of either the α4 or the α4s or from chaperone-mediated preferential incorporation of α4s in 20S proteasome. We observed that the three i20S-specific catalytic subunits (PSMB8-10 proteins, Fig. 2A) were more abundant in the anti-α2 IP and were practically absent from the anti-α4s IP, implying that little or no immuno-subunits constitute the s20S (FC of immuno subunits in anti-α4s IP = 0.006, p-value = 7.E-5, Fig. 2C), as previously observed using an orthogonal immunodetection approach (18). This interesting result is also in agreement with the 6.1 fold decrease, in the anti-α4s IP, of PA28αβ (p-value = 0.01), known to interact preferentially with the i20S (23). Moreover, while the four 20S Proteasome Assembly Chaperones (PAC) are equally distributed in the α4-and α4s-containing 20S (Fig. 2A, Table S3), the POMP maturation protein is 5.3 fold increased in the α4-containing 20S (p-value = 0.02, Fig. 2C, Table S3). POMP is known to preferentially promote the assembly of i20S catalytic subunits over c20S ones (33,35,36), which agrees with our observation that immuno-catalytic subunits are low abundant in the anti-α4s IP.

Our analysis of main 20S interactors revealed that all subunits of the 19S particle could be quantified and their very tight distribution in the Volcano plot (i.e. very close FCs and pvalues, Fig. 2A) emphasizes the quality of this dataset. Interestingly, we could measure an average increase of 2.13 ± 0.15 of 19S subunits in the anti-α4s IP compared to the anti-α2 IP. Since the anti-α4s antibody purifies almost exclusively α4s-containing 20S (95 ± 9%) and the anti-α2 IP contains a mix of proteasomes (42% of α4s-containing 20S), we can estimate that in whole testes, the abundance of the 19S bound to the s20S is approximately 15 times higher than the one associated with the c20S (p-value = 0.02) (Fig. 2C and Table S3). Interestingly, other known 19S interactors were also found to be increased in the anti-α4s IP compared to the anti-α2 IP. This is the case for the two main 19S-associated deubiquitinases USP14 and UCHL5 (FC = 3.2 and 3.9, p-values = 0.02 and 0.05, respectively), ADRM1 protein, a known 19S receptor of polyubiquitin chains (FC = 1.8, p-value = 0.01), as well as RPS27, the main cellular precursor of ubiquitin (FC = 1.9, p-value = 0.03). The K48 polyubiquitin chains identified by the LIFAGK(GG)QLEDGR peptide (Døskeland, 2006) were increased by a factor of 3.0 (p-value = 0.001) in the anti-α4s IP compared to the anti-α2 IP. Altogether this dataset suggests a higher loading of polyubiquitinated substrates onto s20S compared to total proteasome complexes. Accordingly, among the 40 ubiquitin-related enzymes (conjugating enzymes, ligases and deubiquitinylases) that are found significantly regulated between the two IPs, all were increased in the s20S interactome except TRIM21, an E3 ligase that is mainly involved in immune response (Table S4).

Another PIP found to be highly regulated in this dataset is PI31, a 20S interactor of controversial function (37). PI31 displayed a 20-fold enrichment in the anti-α4s IP compared to the anti-α2 IP (Fig. 2A, p-value = 0.01) and seems to be a preferred partner of the s20S (Fig. 2C and Table S3). Strikingly, a known heterodimerization partner of PI31, Fbxo7 (38), is also significantly more abundant in the s20S IP (FC 72; p-value = 0.007). Recent KO studies of Fbxo7 and Nutcracker, its heterolog in drosophila, indicate important roles of both proteins during spermatogenesis (39,40). Due to the established role of PA200 in spermatogenesis (11), a complementary IP directed against PA200 was conducted and also showed that PI31 and Fbxo7 proteins were highly enriched in the anti-α4s IP compared to the PA200 IP (FCs = 12.2 and 4.1, p-values = 1E-04 and 0.05, respectively, Fig. S6,S7), suggesting that these proteins are not bound to the s20S through an interaction with PA200.

We found that the synaptonemal proteins SYCP3 and SYCE1 are specifically enriched in the anti-α2 and anti-α4s IPs, compared to their respective control IP (Fig. S4,S5), and significantly more abundant in the s20S interactome compared to the other α4-containing 20S isoforms (FCs = 294 and 3.8, respectively; p-values = 8E-03 and 0.05, respectively (Fig. 2A, 2C and Table S3). Previous imaging studies have suggested that these proteins interact directly with the s20S (18), and our dataset further demonstrates such specific interactions of the s20S with the SC.

Despite their proposed role in spermatogenesis (41), the nuclear PA28γ and PA200 regulators of the 20S core were either not (FC = 0.8; p-value = 0.4) or only slightly, but still significantly (FC = 2.0; p-value = 0.03), enriched in the s20S compared to the α4-containing 20S subtype (Fig. 2C and Table S3). Moreover, when comparing the relative abundances of α4 and α4s co-purified in the anti-α2 and in the anti-PA200 IPs, we observed similar values (Fig. 2B). Altogether, these results suggest that PA200 may not interact preferentially with the s20S, contrary to what was previously speculated (16). However, if the α4 and α4s isoforms are expressed in different cell types, as previously reported (14), they would not have to compete for PA200 association. Thus, higher association observed between PA200 and the s20S could be due to a higher PSMA8 gene expression in these cells.

Taken together, our IP strategy revealed that the s20S has an interactome distinct from that of the c20S. The s20S interactome we mapped here is enriched for the 19S particle, as well as different components of the UPS and synaptonemal proteins. The differences between the c20S and s20S interactomes suggest distinct functions of these proteasome variants.

Proteasome dynamics analysis shows profound rearrangements throughout

spermatogenesis. An important limitation of our whole testis analysis is the lack of information on the dynamic interactions that are frequently lost and/or rearranged upon tissue lysis (42), and information on proteasome dynamics throughout spermatogenesis. To provide a detailed map of the proteasome interactome during spermatogenesis, we initiated a study of individual germ cell populations. We harvested testes from young adult or prepubescent rats, and immediately dissected and digested them into individual cells. Rat germ cells were purified and separated either by sedimentation at unit gravity (SPG) or centrifugal elutriation (SPC and SPT), in order to obtain highly enriched cell populations. SER were also purified, since they are support cells of the germ cell lineage and might carry relevant information. Proteasome complexes from these purified fractions of cells were in vivo crosslinked to maintain protein-protein interactions (43) and then co-immunopurified using the antibody directed against the α2 proteasome subunit (23). Purified complexes were digested with trypsin and the resulting peptides analyzed on a nanoLC-MS/MS system, followed by protein identification and relative quantification. In parallel, lysates of individual groups of cells were directly analyzed without the IP step to obtain complementary information on protein expression and to validate the cell purification protocol. Among the 5750 proteins validated and quantified in the lysates using label-free MS, a pool of 12 proteins were unambiguously identified as specific markers of each cell type (Fig. S8). Proteasome complexes immunopurified from the different groups of cells were then analyzed using the same approach to observe changes in 20S proteasome composition throughout spermatogenesis (Fig. 3). None of the three i20S catalytic subunits could be detected in the four different cell types studied, suggesting that the i20S subunits previously detected in the whole bovine testis sample are likely derived from infiltrated immune cells. Another striking observation was that the α4s isoform almost completely replaced α4 at the SPC and remained the main isoform throughout spermatogenesis (Fig. 3A). This trend was also observed in cell lysates (Fig. 3B), and may be driven by changes in the protein expression or stability of the two isoforms.

Our analysis of proteasome activators (19S, PA28αβ, PA28γ, and PA200) indicated that the 19S is the predominant regulator associated with the 20S particles, whatever the cell type analyzed, and was bound from around 30% (in SER and SPG) up to 60% (in SPC and SPT) of the total 20S pool (Fig. 3C). The fraction of 20S-19S complexes thus significantly increases 1.7fold (p-value = 0.01) when SPG proceeds into the SPC stage (Fig. 3D), and, as previously seen in bovine testes, the K48 polyubiquitin chains identified by the LIFAGK(GG)QLEDGR peptide were increased by a factor of 2.9 (p-value = 1E-03), highlighting a high demand for cellular ubiquitin-dependent proteolysis. While the nuclear PA28γ activator levels do not significantly change through spermatogenesis, its cytoplasmic counterpart PA28αβ significantly decreased (7.2-fold) from the SPG to SPC stages (p-value = 0.005) (Fig. 3D). The abundance of PA200 bound to the 20S core particle increases 7.5-fold in SPC (p-value = 0.002) and 6.6-fold in SPT (p-value = 0.01) compared to SPG, in agreement with the proposed role of PA200 in spermatogenesis (11,44).

We also noted that PI31 has a dramatic increase (11-fold) in 20S core particle association in both SPC and SPT stages, compared to non-differentiated SPG cells (p-values of 2E-03 and 8E-04, respectively) (Fig. 3D). Accordingly, its known interactor Fbxo7 was only detected in proteasome complexes purified from SPC and SPT.

Given that 20S-associated regulators display different substrate specificities and subcellular localization (45), the observed variations in the panel of proteasome activators suggest profound changes in the biological function and/or subcellular localization of proteasomes throughout spermatogenesis. Overall, a large proportion of the 20S proteasome pool was bound to activators, reaching almost 75% in SPC cells, which represents a specific feature of testis cells as the proportion of activated 20S core is usually around 20 to 60% in other tissues (27). Moreover, all three germ cell types contain slightly but significantly higher amounts of total 20S compared to SER (Fig. 3E). Chymotrypsin, trypsin, and caspase-like proteolytic activities were significantly higher in meiotic and post-meiotic germ cells (SPC & SPT) compared to undifferentiated SPG (Fig. 3F), which is most probably the consequence of two major events, i.e. the complete replacement of α4 by α4s isoform and the increase in activator-associated 20S, in particular the higher loading of 19S and PA200 particles. No significant variation of 20S-associated regulators can be observed from SER to SPG cells which also display similar 20S proteasome composition, in particular their high content of α4. The observed variations in proteasome proteolytic activities and 20S-associated regulators in premeiotic (SPG) vs. meiotic and post-meiotic cell types (SPC & SPT) thus suggest either a possible increased ability of the s20S to interact with its activators, compared to the c20S, or a transcriptionally-driven increase in the quantities of these regulators in post-meiotic stages.

To answer this crucial question, we analyzed variations of expression of these different proteins in the different cell lysates. Changes from SPG (containing exclusively c20S) to SPC and SPT (containing 85-90% s20S, Fig. 3A) were thus compared both in the immunopurified proteasome samples and in the lysates (Fig. S9).

Concerning α4s, its increase in lysates from the SPG to SPC stages is approx. two-fold higher than its increase in immunopurified proteasomes (p-value = 0.007, Fig. S9). This may be due to a chaperone-mediated resistance to α4s incorporation into functional 20S proteasomes, or to the lag time between synthesis of α4s and its incorporation in newlyassembled s20S. Increase in proteasome bound PA200 appears to be mainly the consequence of increased expression (Fig. S9), which is in accordance with the existing literature (19). However, given that we did not observe increased binding of PA200 to s20S in the whole testis interactome analysis, it remains difficult to draw conclusions about specific PA200 interactions at this stage. Finally, the increase in the association of 19S and PI31 with 20S proteasomes from SPG to SPC is, to a certain extent but not completely, explained by higher cellular abundances, since their increase in the immunopurified proteasomes are significantly higher than in full lysates (FC = 1.4 and 2.7; p-value = 0.03 and 0.006, respectively, Fig. S9).

Altogether this dataset indicates that changes in the composition and activation of 20S proteasomes from the SPC stage onwards may, at least partly, result from transcriptional regulation; however, at this stage we can't exclude the possibility that other mechanisms and factors may be involved.

Pull-Down assays show a preferential binding of the s20S to the 19S in vitro.

In order to examine preferential interactions between α4s and PI31, 19S and PA200 (compared to α4), we took advantage of the testes 20S pool immunopurified in mg quantities from bovine testes. After incubation of these three partners of the 20S with either pure c20S (purified from bovine muscle) or the testes 20S pool (purified from bovine testes), we pulled-down complexes using anti-PI31, anti-19S or anti-PA200 antibodies-grafted beads, and estimated the relative amounts of α4 and α4s in the eluates using LC-MS/MS label free quantification. A control pull-down was run concurrently with the MCP21 antibody that equally recognizes the c20S and s20S. After normalization of the signal with the MCP21 IP, we found that PI31 was interacting more with proteasomes with α4 than with α4s (Fig. 4A) (p-value = 0.018). However, the 19S pull-down (anti-PSMC2) was much more efficient in co-purifying α4s than α4 (p-value = 0.0002) (Fig. 4B). Finally, the anti-PA200 IP showed no significant difference between α4 and α4s pulldowns (Fig. 4C). Altogether, these in vitro binary interaction experiments suggest that the c20S and s20S preferentially interact with PI31 and the 19S, respectively; while there is no clear evidence that PA200 interacts preferentially for any of these 20S subtypes. Testis and muscle proteasomes show distinct peptidase activities. We tested whether ɑ4s incorporation into the 20S core induces any change in proteasome activity on its own or in complex with a regulator. We compared the testes 20S pool purified from bovine testes, which contains approximately 50% of s20S, and the c20S purified from bovine muscle in their ability to degrade fluorogenic substrates. The substrates assayed were boc-LRR-AMC, suc-LLVY-AMC and z-LLE-AMC, which probe the trypsin-like, chymotrypsin-like and caspaselike activity, respectively. The two proteasome samples were also activated with a PA200:20S molar ratio of 8:1. When comparing the basal activity of the proteasome, the s20S mix showed greater trypsin-and chymotrypsin-like activity, while the caspase-like activity was more or less the same (Fig. 4D). The addition of PA200 did activate the three protease activities of both 20S proteasomes. Although trypsin-like activity change upon PA200 binding is greater in c20S, the overall tryptic activation of the complex remains significantly greater in the s20S. The activated complex's chymotrypsin-like activity is also greater in the s20S. The activation of the caspase-like activity is somewhat more pronounced in the c20S compared to the s20S. Overall, these results show that s20S seems to have higher trypsin and chymotrypsin-like activities than c20S, both at basal state and when activated with PA200.

HDX-MS rationalizes the spermatoproteasome structural singularity.

We recently implemented HDX-MS, a technique which reports on solvent accessibility and/or flexibility of the backbone amide protons, on the c20S and i20S proteasome complexes, in order to decipher their structural rearrangements upon replacement of their catalytic subunits and binding to PA28 regulators (46). Building on this expertise, we investigated the conformational differences due to the replacement of α4 by α4s. The relatively large amount (~180 µg per condition) of pure 20S required to perform such structural studies led us to use the testes 20S pool purified from bovine testes. This prevented us from comparing the deuteration of any peptides common to the c20S and s20S. However, 33 and 27 of the 53 and 47 peptides obtained upon pepsin digestion of α4 and α4s, respectively, were proteospecific (Fig. S10). We manually compared the deuteration heatmaps obtained for both α4 and α4s after alignment (Fig. 5A) and clearly identified two regions, encompassing residues 180-189 and 225-250, that were more readily deuterated in α4s compared to α4 (Fig. 5A-C). Interestingly, these two regions face each other on the outer surface of the α-ring, based on the c20S structure (Fig. 5B). To explore possible impacts of these differences on structure and dynamics of α4 and α4s, we employed 1 µs all-atom molecular dynamics simulations. We could detect ~64% more hydrogen bonds (H-bonds) between the NH groups of the backbone and water molecules for α4s than for α4 (Fig. S11A) in line with our HDX-MS results. Furthermore, the distribution of the crossing angle between the two last α-helices was broader in α4s compared to α4 (Fig. S11B, standard deviation = 8.3° and 6.2° for α4s and α4, respectively), indicating that the motion between these two helices is more important in α4s. Looking in more detail, we observed that these two helices were stabilized by H-bonds between three pairs of side chain residues (E235/K193, E231/K189 and Y228/D185 for α4 and E235/R193, E231/K189 and Q228/N185 for α4s) (Fig. S11C). Interestingly, the frequency of these H-bonds was found much higher in α4 than in α4s, especially between residues 235 and 189 (Fig. S11,S12A), explaining the higher motion of these helices, the larger number of Hbonds between its backbone and water molecules and the overall faster deuteration in α4s. In line with these results, the average Accessible Surface Area (ASA) of residues 184-198 and 225-241 was higher in α4s than in α4 (Fig. S12B). Altogether, our results indicate more dynamic movements of both 180-189 and 225-250 regions in the s20S compared to the c20S, potentially creating a different binding interface and resulting in the differences in the interactomes we mapped above.

Discussion

Despite being recognized as important for mammalian male germ cell differentiation and spermatogenesis, the testis-specific proteasome isoform s20S remains incompletely characterized. Here, we combined state-of-the-art MS-based proteomic analyses with in vitro studies to conduct a comparison between s20S and c20S, using α4s and α4 as markers for each proteasome variant, respectively. We found that α4s represents ~42-45% of all α4 variants found in whole bovine testis, suggesting that the s20S is a major proteasome form in testicular tissue. We also showed that α4 and α4s do not co-exist within assembled proteasomes, in line with immunofluorescence imaging experiments (14). This exclusive incorporation of α4 or α4s subunits into proteasomes may be either due to a selective subunit incorporation or to a sudden change in protein expression levels. To address this question, as well as map dynamic proteasome changes that accompany spermatogenesis, we analyzed isolated populations of rat germ cells at different differentiation stages (SPG, SPC, SPT) and their supportive cells (SER). This demonstrated that SPG 20S proteasomes contained almost exclusively α4 (>98%), whereas pachytene SPC and SPT predominantly had α4s-containing proteasomes (>82%-92% α4s). This sudden change in the abundance of α4s vs. α4 may be driven by rapid changes in transcription or in protein turnover resulting in major differences in protein levels.

The shift of c20S to s20S observed from pre-meiotic SPG to meiotic SPC was accompanied by profound changes in the major 20S-associated regulators, in particular an increase in PA200 and 19S activators, concomitantly with a decrease in PA28αβ. Surprisingly, no change in PA28γ could be observed, although this 20S activator was previously shown to be relevant to male fertility (22,41). It has been recently proposed that PA200 constitutes a major component of the s20S complex (11,19) and that α4s may help the formation of PA200capped complexes containing only standard catalytic subunits (19). Accordingly, our results indicate that PA200 is an important player in s20S function, as its level of incorporation into assembled proteasomes in SPC and SPT (around 8-10%) is at least 10-fold higher than in SPG and previously measured for several other cell types (27). However, our results also demonstrate that the 19S is the major activator of 20S core particles in germ cells, and in particular in meiotic SPC where α4s completely replaces α4. Indeed, the 19S is bound to around 30-40% of the total 20S pool in SER and undifferentiated germ cells and, strikingly, this proportion increases up to 60% in SPC and SPT cells, making meiotic and post-meiotic proteasomes the most activated proteasome complexes we have analyzed so far. This agrees with the increased proteolytic activities in SPC and SPT, compared to SPG and SER, in line with previous observations using whole testis lysate compared to muscle tissue (11). Another study also showed high levels of 19S-containing species in post-meiotic germ cells (18) and in whole testes (11,19). Thus, contrary to what we expected from previous studies (11,19), we clearly demonstrated that PA200 is far from being the sole activator of s20S during germ cell differentiation. The increased association between s20S and its 19S activator is probably aligned with high ubiquitin-dependent proteolytic requirements at the SPC stage, as suggested by increased polyubiquitin chains at the s20S complex in meiotic SPC compared to pre-meiotic SPG. In particular, meiosis I progression would require above all the ubiquitinproteasome system (40,47,48), and, to a lesser extent, PA200 (14) which was shown to be rather involved in later acetylated histone turnover events (19).

We observed that PI31, previously described as an in vitro 20S proteasome inhibitor (37), but also a physiological 26S proteasome activator (40,49) or assembly factor (50), together with its binding and stabilizing partner Fbxo7 (38), were both enriched in s20S interactomes compared to c20S but also PA200 IPs. Strikingly, both PI31 and Fbxo7 are essential for proper spermatogenesis in Drosophila (40) and mice (39), respectively. To our knowledge, our work is the first establishing a direct link between the α4s specific subunit of the s20S and the PI31/Fbxo7 axis. PI31 was shown to mediate proteasome transport in axons and dendrites in mice, by regulating the loading of proteasomes onto microtubule-dependent molecular motors (51). Accordingly, here we identified several microtubule-related proteins interacting with the s20S, which could facilitate such transport of s20S complexes. On the other hand, as a part of its E3 ligase activity, Fbxo7 targets proteins involved in cell cycle regulation (52). Thus, we speculate that PI31 and Fbxo7 might act as shuttle proteins for the s20S, targeting cyclins and other spermatogenesis-specific substrates; however, mechanistic details of s20S-PI31-Fbxo7 remain to be established.

Another class of s20S-specific interaction partners we identified are SYCE1 and SYCP3, components of synaptonemal complexes. These findings further support the proposed mechanism whereby s20S binds to the synapses along the axes of the meiotic chromosomes, to regulate this process through the degradation of specific proteins (18,47). Although previous work suggested that SYCP3 could be a substrate of the s20S (18,47), our data indicate high sequence coverages of SYCP3 and SYCE1 in our interactome study (25% and 34%, with 4 and 9 peptides, respectively), which is typical of true interactors. Moreover, recent work on s20S KO mouse models did not replicate the SYCP3 accumulation (14), further strengthening the argument for SYCP3 as a s20S interactor. Taken together, our quantitative interactome data indicate extensive differences between c20S and s20S, and show that spermatogenesis is accompanied by a major shift from c20S as the major proteolytic machinery towards s20S.

Given that the only difference between c20S and s20S is the presence of ɑ4 or ɑ4s isoforms, we employed HDX-MS to examine whether they display any differences in structure and dynamics. Our analysis revealed pronounced differences in flexibility of the C-terminal regions of the two isoforms, corresponding to the two last ɑ-helices located on the outer, solvent-exposed side of both ɑ4 and ɑ4s structures. Our HDX-MS data showed that these two helices are more rigid (or stabilized) in ɑ4 than in ɑ4s. Molecular dynamics simulations provide a rationale for these observations, by predicting more stable hydrogen bonds between the two C-terminal helices in ɑ4. In order to understand functional consequences of these differences, we performed pull-down assays with ɑ4-and ɑ4s-containing proteasome and immobilized proteasome regulators, PI31, PA200 and 19S. We observed differences in pull-down efficiency for PI31, which showed significantly greater affinity towards the ɑ4containing proteasome (c20S), and 19S, which had a higher affinity towards the s20S. However, we did not observe any differences in affinity of PA200. Finally, we also tested proteolytic activity of proteasomes purified from bovine testes (~50% s20S) and from the muscle (100% c20S) in two different conditions: alone and when activated by PA200. We measured that s20S has higher basal trypsin-and chymotrypsin-like activity, and this trend held upon PA200 activation. On the other hand, caspase-like activity for 20S alone was either the same or very similar, while the c20S-PA200 complex was more active than the s20S-PA200. Interestingly, upon PA200 binding, the fold-change in caspase-like activity is the most pronounced, confirming previous observations (53). Increased basal level of tryptic activity can lead to improved degradation of substrates that are highly positively charged, such as histones, in agreement with the report that showed histones to be the targets of the PA200-20S complex in the context of spermatogenesis (11). Overall, these activity measurements indicate significant difference in behavior of the ɑ4s-containing proteasome, compared to the ɑ4-containing proteasome.

Taken together, the large amount of data analyzed and presented in the context of this study highlight some key differences between c20S and s20S. Our results imply a more complex process of s20S regulation than previously suggested. Based on these, we can speculate that the structural differences between the s20S and c20S proteasome variants trigger the recruitment of specific partners and ubiquitin-related enzymatic modulators that are key for proteasome cellular relocalization to the SC and for the degradation of important meiotic players, respectively.

Experimental Procedures

Proteome Repository Search. For data on relative expression at the protein level of the proteasome and proteasome-related genes we searched The Human Proteome Map portal (17) by querying the list of relevant genes.

Reagents. Unless stated otherwise, all reagents were purchased from Euromedex. The c20S, i20S, PA28γ and PA28αβ were purchased from Enzo Life Science. The mouse IgG1 anti-ɑ2 antibody was produced from the hybridoma cell line MCP21 (European Collection of Cell Cultures). Antibody development. Anti-α4s antibodies were produced by Biotem (Apprieu, France) using procedures described in SI Experimental Procedures.

Preparation of separated germ cells. Cells were obtained from rats using procedures described in SI Experimental Procedures.

LC-MSMS analysis. Bottom-up and top-down proteomics experiments were performed on an Orbitrap Fusion instrument coupled to an Ultimate 3000 chromatography system. Acquisition parameters and data analysis are detailed in SI Experimental Procedures. HDX-MS and MD simulations. HDX-MS experiments were performed on a Synapt-G2Si (Waters Scientific, Manchester, UK) coupled to a Twin HTS PAL dispensing and labelling robot (Trajan Scientific, Milton Keynes, UK) via a NanoAcquity system with HDX technology (Waters, Manchester, UK), as described previously (46). More details, as well as MD simulation workflow can be found in SI Experimental Procedures.

In vitro assays. The activity assay, based on fluorogenic peptide degradation, and the pull-down assays are further detailed in SI Experimental Procedures.

Supporting Information

SI Experimental Procedures

Antibody development Two peptides were synthesized, C+ILNPEEIEKYVAEIEKEKEENEKKKQKKAS -corresponding to the ɑ4-specific C-terminal sequence (P1 peptide) and C+MFSAKEIELQVNEIEKEKEEAEKKKSKKTT -corresponding to the ɑ4s-specific C-terminal sequence (P2 peptide). Two New Zealand White rabbits, 20 to, 25 weeks old, were immunized with a KLH-P2 conjugate (Day 0), by subcutaneous injection containing incomplete Freund's adjuvant. Immunisations were repeated on Day 7, Day 14 and Day 35. The rabbits were exsanguinated on Day 42 after the immunisation and the antibodies were affinity-purified from the blood serum using the immobilized P1 peptide, followed by depletion step using immobilized P2 peptide, in order to remove the ɑ4-reactive antibodies. The obtained anti-α4s polyclonal antibodies exhibited an EC50 of 0.005 µg/mL against the α4s epitope and of 2.12 µg/mL against its α4 counterpart (Fig. S4B). Moreover, the specificity of the newly-produced anti-α4s antibody was verified by western-blot assay (Fig. S4C).

Preparation of separated germ cells from rats Spermatogonia (SPG) were isolated from testes obtained from thirty 8-day-old Sprague Dawley (SD) rat testes with a purity of greater than 90% as previously described (1). Seminiferous epithelial cells were dispersed by enzyme treatment and separated. Briefly, the cells were separated by sedimentation velocity at unit gravity at 4 °C, using a 2-4% BSA gradient in Ham'F12/DMEM in an SP-120 chamber (STAPUT). After 2.5 h of sedimentation, 35 fractions were collected. Cell fractions 16 to 21 were pooled, washed 3 times with Phosphate Buffered Saline pH 7.4 and cells were immediately subjected to in vivo cross-linking to stabilize proteasome interactors, as previously described (2). Cross-linked cells were stored at -80°C. Pachytene spermatocytes (SPC) and early spermatids (SPT) were isolated from testes obtained from eight 90-day-old SD rat and prepared by centrifugal elutriation with a purity greater than 90% according to a previously described method (3), except that enzymatic dissociation of cells was replaced by mechanical dispersion. Upon isolation, cells were washed 3 times with Phosphate Buffered Saline pH 7.4 and cells were immediately subjected to in vivo cross-linking as previously described (2). Cross-linked cells were stored at -80°C. Sertoli cells (SER) were isolated from testes obtained from eight 20-day-old SD rats according to previously described methods (4,5). The cell suspensions were seeded at a density of approximately 1x10^6 cells/mL in 75 cm2 tissue culture flasks (NUNC, Copenhagen, Denmark). The cells were then incubated at 32°C in a humidified atmosphere of 5% CO2 and 95% air in Ham's F12/ DMEM supplemented with insulin (10 mg/mL), Human transferrin (5 mg/mL) and gentamycin (10 mg/mL), all from Life Technologies (Eragny, France). Culture medium was changed daily until the end of the experiment. On the second day of culture, the cells were exposed to a hypotonic treatment in order to eliminate the contaminating germ cells. The degree of purity of the isolated Sertoli cells was > 98%. On day 5 of culture, cells were recovered, washed 3 times with Phosphate Buffered Saline pH 7.4 and were immediately subjected to in vivo cross-linking as previously described (2). Cross-linked cells were stored at -80°C. Proteasome Purification The proteasome complexes were immunopurified from tissues or cell lysates using the MCP21 antibody as described before ( 6), the anti-α4s or anti-PA200 antibodies as fully described in SI Experimental Procedures. Protein extract were trypsin digested using S-trap micro cartridges (Protifi, Farmingdale NY USA) Eight mg of MCP21 antibody was grafted to one gram of CNBr-activated Sepharose beads. Frozen bovine testis (-80 °C) from an adult individual were broken into pieces using a hammer and chisel. Lysis and coIP were then done at 4 °C. Ten grams of tissue were diced and placed into a blender with the lysis buffer (20 mM Tris HCl, 0.25% TritonX-100, 100 mM NaCl, 10 mM ATP, 5 mM MgCl2, 1 tablet of inhibitors/50 mL (cOmplete™ ULTRA Tablets Mini EDTA-free, Merck, Darmstadt, Germany), pH 7.6), at 1.6 weight to volume ratio. The lysate was passed through a Dounce homogenizer and then sonicated in a VibraCell (Bioblock Scientific, France) (power = 2, 60% active cycle, 10 cycles, 30 sec per cycle). The lysates were centrifuged for 30 min at 4 °C, 4500 g and the greasy supernatant was aspirated from the top of the lysate. The solid pellet was also discarded while the remaining liquid was again centrifuged at 4 °C and 16000 g, for 30 min. The supernatant was filtered through a 0.22 µm membrane. The filtered lysate was then mixed with the Sepharose grafted with antibodies and incubated at 4°C overnight. The next day, the Sepharose was washed with 50 mL of Equilibration buffer (20 mM Tris HCl, 1 mM EDTA, 10% glycerol, 100 mM NaCl, pH 7.6) and eluted with Equilibration buffer supplemented with 3 M NaCl. The eluate was concentrated on Amicon filters of 100 kDa cutoff (Millipore) and separated using gel filtration chromatography. The chromatography was performed on a Superose 6 10/300 GL column, using TSDG buffer (10 mM Tris-HCl pH 7.0, 1 M KCl, 10 mM NaCl, 5.5 mM MgCl2, 0.1 mM EDTA, 1 mM DTT, 10% Glycerol). The fractions were assayed by SDS-PAGE and 20S proteasome fractions were pooled together.

Proteasome immunopurification from rat germ cells and sample preparation for LC-MS Immunopurification of proteasome complexes from rat germs cells was done using the MCP21 clone of the anti-α2 antibody and sample preparation was done by in-gel digestion, as described in (7). For this protocol, pools of cells from different rats, 8-10x10^6 SER, 20-25x10^6 of SPGs, 8x10^6 of SPCs and 20x10^6 of SPTs per replicate were used. Three independent experiments were run for each type of cell.

Co-immunopurification for interactomics from bovine testis and sample preparation for LC-MS/MS

Four biological replicates of frozen bovine testes (-80 °C) from adult individuals were broken into pieces using a hammer and chisel. Lysis and coIP were then done at 4 °C. Ten grams of tissue were diced and placed into a blender with the lysis buffer (20 mM Tris HCl, 0.25% TritonX-100, 100 mM NaCl, 10 mM ATP, 5 mM MgCl2, 1 tablet of protease inhibitors/50 mL (cOmplete™ ULTRA Tablets Mini EDTA-free, Merck, Darmstadt, Germany), pH 7.6), 1.6 weight to volume ratio. The lysate was passed through a Dounce homogenizer and then sonicated in a VibraCell (Bioblock Scientific, France) (power=2, 60% active cycle, 10 cycles, 30 sec per cycle). The lysates were centrifuged for 30 min at 4 °C, 4500 g and the greasy supernatant was aspirated from the top of the lysate. The solid pellet was also discarded while the remaining liquid was again centrifuged at 4°C and 16,000 g for 30 min. The supernatant was filtered through a 0.22 µm membrane. The co-immunopurification of the proteasome complexes was done using ProteinG coupled to the magnetic beads (MagBeads, Cat. No. L00274, GenScript USA Inc. Piscataway, NJ USA). The protocol proposed by the supplier was followed with the following modifications. The amount of beads used per sample was 100 µL of suspension. The amount of antibodies used per replicate was 100 µg of anti-α4s, 50 µg of anti-α2, 10 µg of anti-PA200 and 50 µg of OX8 control antibody. One mL of lysate of each replicate was incubated with ProteinG-MagBeads, coupled with different antibodies, overnight at 4 °C on a rocker. Samples were then washed four times with Equilibration buffer (20 mM Tris HCl, 1 mM EDTA, 10% glycerol, 100 mM NaCl, 2 mM ATP, 5 mM MgCl2, pH 7.6), then eluted with 25 µL of S-trap Elution Buffer (5% SDS, 50 mM TEAB, pH 7.55). In case of anti-ɑ4s, proteasome complexes were eluted via competitive elution method, for improved signal/noise ratio. Namely, the peptide used for immunization of the animals and production of antibody was used to elute the antibody from the MagBeads. Peptide was diluted in the Equilibration Buffer to 160 µg/mL and 100 µL of the peptide solution were added per replicate. The beads were incubated for 1h at room temperature, while mixing. The supernatant was recovered and mixed with SDS solution to final 5% SDS. Control IP was also done with the OX8 control antibody and the peptide elution procedure. Preparation for LC-MS analysis was done using S-trap micro cartridges (Protifi, Farmingdale NY USA), by default protocol of the manufacturer. Around 2 µg of peptides per sample per run were used for LC-MSMS analysis.

Bottom-up LC-MSMS Analysis was performed on an Orbitrap Fusion instrument, with nano ESI source, coupled to the Ultimate 3000 chromatography system using 90 min runs. Peptides were loaded onto a precolumn and washed (97.95% water, 2% ACN,0.05% trifluoroacetic acid). Upon loading onto the column (made in-house, 75µm ID, 50cm column, packed with Reprosil C18, 3 µm phase, Cluzeau), the elution gradient is initiated at 300 µL/min, by 100% phase A (0.2% formic acid) for 3min, then a slope 10-30% phase B (80% acetonitrile, 0.2% formic acid) for 50min, followed by 30-45% B gradient for 10min, to finish with 45-80% B during 1min. The 80% B was maintained for 10 to wash the column, followed by re-equilibration with the 100% phase A during 15min. The nano-ion source was at +1900 V, with ion transfer tube temperature at Comparison between the anti-α2 and anti-α4s was done in the following way:

(1) Datasheet from the Proline export file, containing all the protein set entries with their relative abundance, called "Protein sets" was copied for treatment and the empty fields were replaced with zeros. (2) Logical tests for every protein entry were performed, to determine if it was detected in at least 4 out of 8 analysis runs of at least one group of samples (4 samples times 2 technical replicates), then that it was determined by specific peptides, then filtered only the entries that satisfy the said tests. The logic tests were done on raw intensities, not the normalized ones.

(3)

The technical replicates were averaged and the fifth percentile for every group was calculated. (4) Number of missing values for each IP sample were calculated and the missing values were replaced by the fifth percentile for protein entries with two or more missing values. (5) The iBAQ values were calculated by dividing the intensities with the number of observable peptides for each entry. (6) Mean proteasome abundance was calculated as average of iBAQs for non-catalytic subunits of the proteasome, excluding the ɑ4 and ɑ4s. (7) The iBAQ values were normalized in all the groups with mean proteasome value for each group. (8) The one-sided paired T-test and fold change were calculated between the anti-α2 and anti-α4s groups. The comparison was done only for proteins significantly enriched between the experimental and control groups (see the end of previous paragraph). Estimation of the regulator stoichiometry of the ɑ4-and ɑ4s-containing proteasome : The composition of these two complexes can be estimated for known interactors based on their intensities in the two CoIPs that have a different ratio of enrichment of ɑ4 and ɑ4s, which is the case in our experiments (anti-ɑ2 and anti-ɑ4s CoIP). Firstly, we need to know the ratio of ɑ4 versus ɑ4s in both CoIPs. The intensity of an interactor of a particular complex (ɑ4-or ɑ4s-containing proteasome) can be expressed via the following formulas:

Experimental Design and Statistical Rationale

Label-free MS Analyses: All statistical analyses were performed on at least three independent biological replicates. For each biological replicate, results from at least two injection replicates were averaged. The analyses on bovine testes were done in four biological replicates. Probability values (p) were determined by one-way paired Student's test for groups of equal variance. Differences were statistically significant at confidence levels of 95% (*), 99% (**), or 99.9% (***). The Pull-Down assays were done in three technical replicates, with p values determined by two-sided, non-paired Student's test for groups of equal variance. The "rat germ cells" dataset contains MS results from the analysis of the IPs and lysates of 4 cell types (Sertoli cells, spermatogonia, spermatocytes, spermatids) in biological triplicates (pools of cells from different individuals for each group), analysed in three injection replicates, for a total of 72 raw files. The "bovine testis CoIP" dataset contains MS results for 4 types of IPs (anti-PA200, anti-α2, anti-α4s, and OX-8 control) from four biological replicates, two technical injections per biological replicate, and lysates for each biological sample, for a total of 40 .raw files. The quantitative comparison of proteins was done based on normalized protein intensities from the "Protein groups" sheet in the Proline output file.

Top-down LC-MSMS Nano-LC-MS analyses of commercial or immunopurified 20S were performed on a nanoRS UHPLC system (Dionex) coupled to an LTQ-Orbitrap Fusion Tribrid mass spectrometer (Thermo Fisher Scientific), as previously described (9). Briefly, a total of 5 μL of sample at 0.5 μM was loaded onto a reverse-phase C4-precolumn (300 μm i.d. × 5 mm; Thermo Fisher Scientific) at 20 μL/min in 2% acetonitrile (ACN) and 0.2% formic acid (FA). After 5 minutes of desalting, the precolumn was switched online with an analytical C4 nanocolumn (75 μm i.d. × 15 cm; in-house packed with C4 Reprosil) equilibrated in 95% solvent A (5% ACN, 0.2% FA) and 5% solvent B (0.2% FA in ACN). Proteins were eluted using a binary gradient ranging from 5% to 40% (5 min) and then 40% to 99% (33 min) of solvent B at a flow rate of 300 nL/min. The Fusion Tribrid (Thermo Fisher Scientific) was operated in single MS acquisition mode with the Xcalibur software (Thermo Fisher Scientific). The spray voltage was set to 1900 V, the ion transfer tube temperature to 350°C, the RF lens to 60%, and no in-source dissociation was applied. The MS scans were acquired in the 700 to 2000 m/z range with the resolution set to 15 000 and using 10 μscans for averaging. LC-MS .raw files were automatically deconvoluted with the rolling window deconvolution software RoWinPro (10) and the proteoform footprints were visualized with VisioProt-MS v2.0 (11).

For MSMS acquisition, 3 second cycles were used, with the following MS parameters: MS scan range between 1000 and 2000 m/z in Orbitrap at 7,500 resolution, 10 µscans for averaging. MSMS activation (HCD) was done with 25% activation energy, with spectra acquisition in the Orbitrap at 60,000 resolution. The AGC target was set to 1e6. Dynamic exclusion for 600 s was used within 60 s to prevent repetitive selection of the same precursor (selection tolerance: ±10ppm) and improve the number of identified proteins. The following source settings were used: Spray voltage 1900V, RF lens 60%, no in-source fragmentation was used. The MSMS analysis of the degraded 20S proteasome used the same parameters, except for the activation method, whereby EThcD was used instead of HCD: ETD activation time of 5ms, ETD reagent target intensity set at 7e5, maximum ETD reagent injection time set at 200ms, with supplemental EThcD activation at 25% energy. The spectra identification was done in Proteome Discoverer (Thermo Fisher Scientific) v.2.3, using an Absolute Mass Search with the following parameters: precursor mass tolerance -1000 Da, fragment mass tolerance -15ppm. Mutations and truncations of the proteoforms were then manually curated. The search database was generated from the ensemble of the available Swiss-Prot and TrEMBL entries for bovin 20S proteasome, bovin proteasome regulators (19S, PA28 subunits, and PA200) and their isoforms.

HDX-MS and MS simulations HDX-MS experiments were performed on a Synapt-G2Si (Waters Scientific, Manchester, UK) coupled to a Twin HTS PAL dispensing and labelling robot (Trajan Scientific, Milton Keynes, UK) via a NanoAcquity system with HDX technology (Waters, Manchester, UK). Data were collected with MassLynX v.4.1 (Waters Scientific (Manchester, UK) and the robot was controlled via HDX Director v.1.0.3.9 (LEAP Technologies, Carborro, NC, USA). Each step was optimized to minimize sample loss and work with such a heterogeneous sample, as described previously (12). Briefly, 5.7 µL of bovin 20S proteasome at 2.3 µM were aspirated and 5.2 µL were diluted in 98.8 µL of protonated (peptide mapping) or deuterated buffer (20 mM Tris pH/pD 7.4, 1 mM EDTA, 1 mM DTT) and incubated at 20 °C for 0, 0.5, 1, 5, 10, and 30 min. The final D2O percentage was 95% in the labelled samples. 99 µL were then transferred to vials containing 11 µL of precooled quenching solution (500 mM glycine at pH 2.3). After 30 s. of quenching, 105 µL were injected into a 100 µL loop. Proteins were digested online with a 2.1 × 30 mm Poros Immobilized Pepsin column (Life Technologies/Applied Biosystems, Carlsbad, CA, USA). The temperature of the digestion room was set at 15 °C. Peptides were desalted for 3 min on a C18 pre-column (Acquity UPLC BEH 1.7 µm, VANGUARD) and separated on a C18 column (Acquity UPLC BEH 1.7 µm, 1.0 × 100 mm) by the following gradient: 5-35% buffer B (100% acetonitrile, 0.2% formic acid) for 12 min, 35-40% for 1 min, 40-95% for 1 min, 2 min at 95% followed by 2 cycles of 5-95% for 2 min and a final equilibration at 5% buffer A (5% acetonitrile, 0.2% formic acid) for 2 min. The total runtime was 25 min. The temperature of the chromatographic module was set at 4 °C. Experiments were run in triplicates and the protonated buffer was injected between each triplicate to wash the column and avoid cross-over contamination. The acquisitions were performed in positive and resolution mode in the m/z range 50-2000 Th. The sample cone and capillary voltages were set at 30 and 3 kV, respectively. The analysis cycles for nondeuterated samples alternated between a 0.3 s low energy scan (Trap and Transfer collision energies set to 4 V and 2 V, respectively), a 0.3 sec high energy scan (Ramp Trap and Transfer collision energies set to 18 to 40 V and 2 to 2 V, respectively) and a 0.3 sec lockspray scan (0.1 µM [Glu1]-Fibrinopeptide in 50% acetonitrile, 50% water and 0.2% formic acid infused at 10 µL/min). The lockspray trap collision energy was set at 32 V and a GFP scan of 0.3 sec is acquired every min. Deuterated samples were acquired only with the low energy and lockspray functions. Peptide identification was performed with ProteinLynx Global SERVER v.3.0.2 (PLGS, Waters, Manchester, UK) based on the MSE data acquired on the nondeuterated samples. The MSMS spectra were searched against a home-made database containing sequences of the 18 c20S, i20S and s20S subunits, all the 19S subunits, as well as the entries for PA28ɑꞵ, PA28γ and PA200 regulators, as well as pepsin from Sus scrofa. Peptides were filtered in DynamX v.3.0 from Waters Scientific (Manchester, UK) with the following parameters: peptides identified in at least two replicates, 0.2 fragments per amino-acid, intensity threshold 1000. The RDUs were not corrected for back exchange. These data were then used for structural data mining with HDX-Viewer (13). Molecular representations of Fig. 5 were generated in UCSF ChimeraX v.0.9 (14) and Visual Molecular Dynamics (15), respectively.

Molecular Dynamics simulations and analysis

For molecular dynamics (MD simulations), we used the α4 model to perform an homology model of the bovine α4s subunit using the modeler program (16) (83.2 % sequence identity between α4 and α4s). For each structure, we then performed 1 µs of molecular dynamics (MD) simulations using GROMACS 2020 (17) in combination with AMBER14SB force field (18).

We first embedded each model in a waterbox using TIP3P water model (19). Cl ions were added to neutralize the system. Energy minimization was performed using the steepest descent algorithm and each system was equilibrated with a constant temperature (canonical ensemble, NVT, 300 K) ensemble for 100 picoseconds (ps), followed by a 100 ps equilibration at constant pressure (isothermal-isobaric, NPT, 1 bar). For equilibration steps, the protein backbone was kept constrained. For equilibration and production runs, we applied the velocity-rescaling thermostat (20) on protein and solvent, coupled with the Parrinello-Rahman barostat (21), with a time constant of 2 ps and compressibility of 4.5x10-5 bar-1. Long-range electrostatics were modeled using the Particle-Mesh Ewald method (22,23). All bonds were treated using the LINCS algorithm (24). For both α4 and α4s models, we performed 1 µs of simulation at 300 K and ambient pressure of 1 bar with an integration time step of 2 femto-seconds. We then analyzed the dynamics of the two last α-helices. We analyzed the creation of hydrogen bonds between water molecules and helices NH groups. To do so, we used VMD (15) Hbonds plugin (https://www.ks.uiuc.edu/Research/vmd/plugins/hbonds/) to count hydrogen bonds formed every 100 ps of the trajectory using a donor-acceptor distance of 3.5 Å and a cutoff angle of 30°. We then calculated the Solvent Accessible Surface Area (SASA) using VMD functions (https://www.ks.uiuc.edu/Research/vmd/vmd-1.8.3/ug/node121.html). To be comparable with HDX-MS data, only the NH groups of residues 180 to 189 and 225 to 241 were used for these two analyses. We calculated hydrogen bonds formed between the two helices (respectively residues 185 to 198 and 224 to 241) as well as the inter-helix angle θ every 100 ps. We used VMD Hbonds plugin and Tcl functions to calculate respectively interhelix hydrogen bonds and interhelix angle.

Activity assays

The activity assay was performed in 96-well black plates (Greiner Bio-One, UK). For assays on purified proteins, 11 µL of purified 20S proteasome sample at 0.57 µM (either c20S or s20S) was mixed with 17.75 µL of recombinant PA200 regulator at 2.84 µM and left to interact for 30 min at 37°C. After they were allowed to interact, the complexes were diluted with activity buffer (20 mM Tris HCl, 5 mM MgCl2, 2 mM ATP, 0.5 mM DTT) to a final volume of 425 µL. As a control, 11 µL of purified 20S proteasome sample at 0.57 µM (either c20S or s20S) was also diluted to a final volume of 425 µL. All the mixes were distributed into wells in aliquots of 50 µL (except for the assays of LLVY, where 25 µL of sample + 25 µL of Activity buffer were used) to which 50 µL of Activity buffer was added, supplemented with 100 µM fluorogenic peptide substrate: Suc-LLVY-AMC, Boc-LRR-AMC or z-LLE-AMC, to probe for chymotrypsin-, trypsinor caspase-like activities, respectively. The kinetic assays were performed at 37 °C in a FLX-800 spectrofluorometer (BIOTEK) over 60 min with one reading every 5 min, at 360 nm for excitation and 460 nm for emission. The slope of the kinetic assay (increase in fluorescence intensity over time) and the quantity of proteasome were used to determine the proteasome's specific activity.

Pull-Down Assays 20S proteasomes were purified from bovine testes as already described above. It is established that the 20S pool contains ~50% s20S and ~50% α4-containing 20S. Purified recombinant human 19S complex was bought from R&D Systems (Minneapolis, USA), recombinant, FLAG-tagged human PI31 protein was bought from OriGene (Rockville, USA) and PA200 protein was prepared as described in (Toste Rêgo and da Fonseca, 2019). The PI31 and 20S pool were mixed at a 10 to 1 ratio in Equilibration buffer 1 (20 mM Tris HCl, 1 mM EDTA, 10% glycerol, 100 mM NaCl, 2 mM ATP, 5 mM MgCl2, pH 7.6) in a total volume of 100µL, at final concentrations of 280nM 20S and 2.8µM PI31. The mix was incubated for 3,5h at 4°C with either the anti-FLAG resin or the proteinG MagBeads containing 100µg of anti-α2 antibody. The resin or the beads were washed with 5mL Equilibration buffer. The elution was done with 2x volume of the slurry of Glycine buffer (Glycine pH3,5 0,1 M) and the samples were prepared for LC-MS analysis on mini-S-trap cartridges (Protifi, Farmingdale NY USA) as recommended by the supplier. For FLAG-tag IP, 280 µL of 50% M2 anti-FLAG affinity resin slurry was used (Merck, Darmstadt, Germany). For anti-α2 IP, 100 µL of 25% MagBeads slurry were used (GenScript, Piscataway, NJ USA). The PA200 of 19S were separately mixed with 20S pool in a 1:2 ratio (10 µg of 19S with 20 µg of 20S proteasome and 2.94 µg PA200 with 20 µg 20S proteasome) in the Equilibration buffer 2 (50 mM Tris pH 7.6, 1 mM MgCl2, 200 µM ATP, 0.5 M DTT) in a total volume of 24 µL. Both mixes were incubated for 30 min at 37 °C, then split in two halves. One half was mixed with the experimental group antibody (10 µg of anti-PSMC2 for 19S-20S mix or anti-PA200 for PA200-20S mix) or control group antibody (10 µg of anti-α2 antibody) and incubated for 30 min at 37°C. Each of the samples was then mixed with ProteinG MagBeads (50 µL 25% slurry per sample) and incubated overnight at 4 °C. The elution of complexes was done with 50 µL of S-Trap buffer and the samples were prepared for LC-MS analysis on mini-S-trap cartridges (Protifi, Farmingdale NY USA) as recommended by the supplier.

Additional

Figure S1. (A)

Expression of all the proteasome subunits and proteasome regulators in different tissues. Of particular interest here are the α4 (PSMA7) subunit and its isoform -α4s (PSMA8) subunit, specific to s20S and proteasome regulators -19S (PSMC, PSMD), PA28αß (PSME1 & 2), PA28γ (PSME3), and PA200 (PSME4). For the sake of simplicity, gene names are used to represent proteins. Abundances at the protein level are presented across different tissues in a form of a heatmap, generated on Human Proteome Map [START_REF] Kim | A draft map of the human proteome[END_REF]. We can see the shared pattern of expression for the PA200 and 19S regulators and the α4s (PSMA8) subunit, suggesting a mutual role. (B) Differences between α4 and α4s isoforms. Sequence identity between these two separate isoforms is very high; however, the differences are concentrated in two regions at the C-terminus of only about 30 amino acid residues in total. In α4, this region is completely on the outer surface, exposed to the solvent, and not involved in the α ring interface. Left: α4s (yellow) was modelled on the structure of the c20S. The regions of pronounced differences are marked in red, notably the two outwardfacing helices. Right: Sequence alignment of bovine α4 and α4s. The sequences are almost identical, except for the regions marked by the two red squares. and the different concentration of polyclonal antibody produced against peptide 2 were used for the readout. This resulted in EC50 of 0.005 µg/mL and 2.12 µg/mL for coating peptides 2 and 1, respectively. (C) Western-blot showing the specificity of the antibody produced against bovine α4s. c20S proteasome were purified from bovine muscle tissue and s20S (mixed with c20S) was purified from bovine testes. The anti-α2 antibody was obtained from the MCP21 hybridoma. Anti-α4s Anti-PA200 spermatogonia (SPG), spermatocytes (SPC), and spermatids (SPT). Among 5750 proteins validated and quantified in the lysates dataset obtained by LC-MS/MS, a pool of 12 protein markers of each separated cell type were used to validate the cell purification protocol.

Additional experiments

Mass photometry to study proteasome complex formation

Introduction and rationale

We wanted to compare the way spermatoproteasome or standard proteasome bind to different regulators, namely, PA200, PA28γ and PI31. This assay would help us better understand the functional implications of α4s incorporation into the proteasome and the structural differences observed in the previous chapter (4.1.2 Article in preparation), more precisely, whether it changes the way the 20S proteasome interacts with its regulators. To study the formation of these complexes, there are several advantages to using mass photometry, when compared to other native techniques such as isothermal titration calorimetry (ITC) or native MS. Firstly, the required concentration of the sample is relatively low -1 to 10 nM; the quantity of the samples consumed is low (~10 µL) and although it is not the practice -the sample can be recovered, since the technique is non-destructive. Next, the system is compatible with most biological buffer formulations (with few exceptions, like high glycerol content) and it allows to relatively quantify complexes of very different masses in a reliable way (40 kDa -5 MDa). However, the mass resolution of the technique is quite limited (25 kDa at 60 kDa) and better suited for the study of large protein complexes, which is the case of the proteasome and its regulators.

After running feasibility tests, we observed that the resolution of the mass photometer was not sufficient to efficiently distinguish between the PI31-bound (746 kDa) and non-bound form (716 kDa) of the 20S proteasome. The study was also performed with PA200 and PA28γ. The access to the MP instrument was limited and was done at Refeyn headquarters in Oxford, UK. We consider the results as preliminary, yet nonetheless very informative.

Experimental section

The different proteasome complexes (bovine) and PA200 (human) were purified, as previously described in the manuscript of the unpublished article (see Chapter 4.1.2). The PA28γ complex was bought from Enzo Life Sciences (Villeurbanne, France). Buffer used in the experiments was the same used in the proteasome purification -(20 mM Tris HCl, 1 mM EDTA, 10% glycerol, 100 mM NaCl, pH 7.6). Proteasome samples were deposited on the microscope slides at final concentrations between 1 and 2 nM, by diluting the initial stock at 10 nM, directly on the slides placed inside a RefeynOne MP instrument (Refeyn, Oxford, UK). The data was collected for 100 frames on average. The regulators were mixed at different ratios to the 10 nM proteasome and then left to interact for 1 minute on ice, followed by dilution on the microscope slide.

The ratios of different observed complexes, as well as the molar ratios in which the interactors were mixed in, were calculated based on the observed/counted molecules in the mass histograms and not based on the initialy assumed concentration (that we based on Bradford assay or a 280nm absorbance). This way we ensure the fidelity of the ratios and avoid bias from wrongly assumed initial concentrations of the interactors.

Results

In the Figure 31, the mass photometry histograms for testis proteasome incubated with PA200 are presented, mixed at two different ratios. For the facility of interpretation, the Table 2 presents the theoretical masses of the proteasome, proteasome regulators and their complexes. On the left panel of Figure 31, a 2:1 PA200:20S ratio was used and on the right this is the 10:1 ratio. On both panels, we can see the normal distribution of masses around ~212-215 kDa, forming a very narrow peak, corresponding to PA200's theoretical molecular weight. An unidentified peak is visible at 70kDa, resembling more to a "smear" of masses, indicative of degradation, possibly an artefact of the proteasome purification or of freeze-thaw cycle. Comparing the left and right panels, it becomes obvious that the smear originates from the proteasome sample and to our knowledge, it did not Additional experiments 123 interfere with the experiments. Next is the ~427-431 kDa peak corresponding to the theoretical mass of a dimer of PA200). Next to the right, we see the free 20S proteasome itself at 714 kDa (theoretical MW= 715 kDa), but only on the left panel corresponding to the lower PA200:20S ratio. Additional peaks correspond to singly-capped (~0.9 MDa) and doubly-capped (~1.1 MDa) proteasomes. The right panel of the Figure 31 shows only the doubly-capped proteasome complex. At a 2:1 (PA200:20S) ratio (left histogram), both singly-capped and doubly-capped complexes form, in approximately the same amount. Upon increase of PA200 concentration (right histogram), the particles of certain masses completely disappear, as the 20S and singly capped proteasome get completely occupied by PA200. As expected, free PA200, largely in excess compared to the proteasome dominates in intensity, especially at the 10:1 ratio. Surprisingly, the two proteasome subtypes interacted identically, both with PA200 and PA28γ. 29 817 Equivalent experiments were run to study the interaction between 20S proteasomes from muscle and testes and PA28γ. Next, we decided to study the distribution of different proteasome complexes as the molar excess of the regulators was increased. For that purpose, we observed the way the amount of occupied proteasome change compared to the total proteasome, as more regulator is mixed with the 20S proteasome. To do that, the exact ratios are re-calculated based on the observed count of individual complexes, for each MP acquisition (Figure 32). On the left-hand side, the percentage of occupied 20S is presented, which is the sum of the percentages of the singly and doubly capped proteasome. The singly capped complexes decrease as the regulator is added, while the share of doubly capped complexes increases (can be said with certainty only for PA200). The measurements for the two regulators, however, were not done in the same range of ratios, nor was it the case for the two proteasome types. We seem to observe what may be a sigmoidal curve dependency in PA200 doubly capped complex formation (lower row, right-hand side).

Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. Both muscle proteasome and the testis proteasome were assayed with two regulators, PA28γ (upper panels) and the PA200 (lower panels), in different ratios. The two proteasome samples seem to behave the same, however, the two regulators seem to bind at a somewhat different affinity.

Discussion

As the regulators are allowed to interact, different complexes form: singly and doubly capped proteasomes. In lower abundance of the regulator, the proteasome can still exist in the free form. Upon further addition of the regulator (increase in the regulator:20S ratio), the proteasome complexes quickly saturate, leaving the doubly-capped proteasome as the only proteasome species. PA200 exhibits a propensity to form a dimer -behaviour also observed for PA28γ, which can hardly be due to non-specific interactions, given the very low concentrations used (10-20 nM) (data not shown).

The two proteasome subtypes seem to behave identically with the two tested regulators, suggesting that there is no preferred interaction towards any of the two proteasome types. The 20S is 100% occupied at around 2:1 ratio for PA200, while the PA28γ plot lacks data points at the same range of ratios, or until the plateau of saturation is achieved. The share of singly-capped 20S begins to drop faster with PA200 compared to PA28γ , as the regulator abundance increases. Furthermore, the share of doubly-capped 20S seems to follow a sigmoidal function, which could be indicative of a positive cooperativity for PA200 proteasome binding. However, no conclusion can be made at this stage, since the dataset is incomplete.

Conclusion and perspectives

Overall, mass photometry proves to be a very useful tool to study the formation of native complexes and their dissociation coefficients, particularly for proteasome complexes. The discovery of cooperativity for the PA200 proteasome binding would be a novel insight for the proteasome function, however, the dataset is incomplete and no conclusions can be made until several issues are addressed: 1) the lack of higher ratio measurements for the PA28γ regulator and 2) higher ratio measurement for the muscle (standard) proteasome in general; 3) the ratios used in the experiments need to be more evenly distributed across the range of studied ratios, in order to improve the predictability of the models. These results can thus be considered as preliminary and insufficient to make any firm conclusions. With a complete dataset, the coefficient of dissociation between the 20S and its regulators could be obtained using this method, and a set of binding assays for 19S regulator could also be done.

Mass photometry is well-suited for the study of protein-protein interactions in native conditions and can be used to determine the oligomeric state of the protein complexes or to determine the dissociation constant for pairs of proteins. However, these proteins need to be above certain size for the MP to be able to distinguish between them or even detect them. For binding of small proteins, and for the study of ligand binding, the nMS is well suited (although conditions for analysis need to be optimized for each protein complex).

Native MS

Introduction

The idea of studying proteasome by nMS is not new per se (Ben-Nissan et al., 2019), however, it remains a challenge to observe multiple species of very different masses at the same time. The goal of this experiment was to investigate if we could use nMS to compare the association of the 20S standard and spermatoproteasome complexes with different regulators.

Experimental section

The standard bovine proteasome was mixed with the recombinant PA200 regulator in the ratio of 1:10 (PA200:std20S) in the Equilibration buffer (IP -see Article in preparation chapter) and left to interact for 5 minutes on ice. The sample was then desalted on Micro Bio-Spin columns (Biorad) into 200 mM ammonium acetate pH 7.4. The complex was then analyzed on the Synapt G2-Si mass spectrometer coupled to a TriVersa NanoMate chip-based nanoESI system (Advion, NY USA). Besides the PA200:20S complex, the PA200 and the 20S alone were also analyzed. MS conditions for PA200 analysis: Sampling Cone Voltage = 150V; Capillary Voltage = adjusted by TriVersa NanoMate; Mass Range = 1-10 000 Th; Trap Collision Energy = 4eV; Trap Fas Flow = 4mL/min. The parameters that were different for 20S and 20S:PA200 analysis were: Mass Range = 2-15 000 Th; Trap Fas Flow = 8mL/min. Spectra were manually extracted and analysed with UniDec [START_REF] Marty | Bayesian Deconvolution of Mass and Ion Mobility Spectra: From Binary Interactions to Polydisperse Ensembles[END_REF].

Results

Figure 33 displays acquired spectra of the different protein complexes. On the left-hand side, the processed mass spectra are displayed, with smoothed peaks and subtracted baseline. Different charge states of the same complex are marked by the same symbol (circle, triangle, square or rhomboid of different colours). These mass spectra were deconvoluted with UniDec and are presented on the right-hand side of Figure 33. Calculated masses are given in the coloured tables on the right.

Analysis of the PA200 sample alone (Figure 33.A) reveals the existence of two distinct species, separated by 1338 Da.

Next, we analysed the standard bovine proteasome alone and measured two distinct species, at 724.6 and 754.7 kDa, which roughly correspond to the theoretical mass of the 20S proteasome (715.8 kDa). We were not able to explain so far the mass difference measured for the higher MW signal.

The mass spectrum of the 20S-PA200 mix contains additional peaks corresponding to singlycapped proteasome at ~900 kDa. Another interesting mass can be observed at ~430 kDa (Figure 33.C), which seems to correspond to a dimer of PA200. Finally, we remark that the PA200:20S complex requires slightly higher cone voltage and collision gas pressure than the PA200 alone.

Discussion

Recent work by Toste Rego et al. (Toste Rêgo and da Fonseca, 2019) on the structure of the PA200:20S complex revealed the existence of two extra-densities corresponding to two inositol hexakisphosphate (InsP6) molecules bound at two distinct sites. The average molecular weight of InsP6 is 660 Da, meaning that the two InsP6 molecules bound to PA200 would correspond to the mass shift observed here (2x660=1320; 18 Da can be attributed to one molecule of water or simply mass measurement error).

Two distinct mass species in native spectra of the 20S proteasome, separated by 6 kDa, have been previously observed by us and others (unpublished data). However, the origin of the higher-mass peak has not yet been confirmed. Since TD-MS revealed that the molecular weight of each alpha and beta subunits were corresponding to the expected ones, we think that this mass shift could either come to some non-covalent binding of one or multiple ligands to be identified, or to a large covalent modification that was not visible in our proteoform footprints. This issue could be addressed by Native Top-Down using a UHMR instrument capable of selecting and fragmenting high MW species.

The MW of the singly-capped proteasome complex (~900 kDa) does not fully correspond to the theoretical mass (927 kDa) but is within the mass precision range for such a large complex. Better transmission of such high m/z species could be achieved using a 32k quadrupole (instead of an 8k quadrupole) or on a UHMR instrument.

Finally, the species observed at ~430 kDa, seem to correspond to a dimer of PA200, as already observed by mass photometry. Native mass spectra of the PA200:20S complex formation. On the right are the deconvoluted spectra along with the calculated masses of the detected mass species. A: PA200 spectra B: bovine standard 20S proteasome spectra C: PA00:20S proteasome mix, with a ~900 kDa peak corresponding to the singly-capped proteasome complex.

Conclusion and perspectives

We show how the proteasome regulator binding event can be observed using nMS. If the experimental conditions are further developed to improve the resolution of higher mass species, we could use nMS for relative affinity assessment of a regulator towards different 20S types. More precisely/reliably determined masses may tell us more about the different proteoform binding behaviour, e.g. the strength of interaction of modified vs non-modified regulators, such as PI31, which upon phosphorylation binds to dDYNLL1/2, to facilitate proteasome transport (K. Liu et al., 2018). It would be interesting to attempt to fragment the higher MW species identified in 20S-only sample, in order to determine which proteasome subunit may carry a PTM or non-covalent ligand(s). However, for such an experiment, an instrument capable of surface-induced dissociation would be best suited, since it would provide richer MSMS spectra [START_REF] Vimer | Comparative Structural Analysis of 20S Proteasome Ortholog Protein Complexes by Native Mass Spectrometry[END_REF].

spt20S substrate experiment

Introduction and rationale

Substrate specificity is a key aspect of proteasome function. Knowing the substrates specific to the spermatoproteasome would bring key information about the whereabouts of its requirement in germ cells. To study such a vital complex in a process which is not replicable in vitro, that is, spermatogenesis, carries many hurdles. However, substrate degradation could be studied in a closely related system. For this purpose, we selected an immortalized mouse spermatocyte cell line -GC-2spd, treated by transfection of large tumor antigen protein to the developing germ cells. As a result, the cell line differentiation arrests in the beginning of the spermatocyte stage [START_REF] Hofmann | Immortalized germ cells undergo meiosis in vitro[END_REF]. This cell line on its own does not express α4s, but only the α4 and its proteome is expected to be somewhat similar to the one of native spermatocytes. If α4s was to be recombinantly expressed in this cell line, we could observe the changes at the proteome level, to better understand the physiological implications of the spermatoproteasome expression. We thus set out to recombinantly express α4s and α4 in two separate cell cultures, whereby the α4s-expressing cell line is the experimental group and the α4-expressing cell line is the control group.

Experimental section

Firstly, we created a pCDNA3.1 plasmid construct, expressing the GFP-P2A-PSMA8 or the GFP-P2A-PSMA7, whereby the GFP serves as a reporter of proper expression at the protein level and the P2A self-cleaving peptide sequence ensures proper and efficient cleavage into two separate proteins. The GC-2spd cell line was grown until 60% confluence, then transfected with GFP-P2A-PSMA7 or GFP-P2A-PSMA8 plasmid constructs, using polyethyleneimine transfection agent, as described in [START_REF] Merolle | Overexpression of the mitochondrial Mg channel MRS2 increases total cellular Mg concentration and influences sensitivity to apoptosis[END_REF], then grown in 15 cm culture plates at 37°C for 24h, as recommended by the supplier (ATCC Cell lines). The cells were sorted on a Becton Dickinson LSRII flow cytometer in three groups -GFP-negative, GFP-feeble and GFP-intense, based on the GFP fluorescence signal, as shown in the Figure 34. For each group, one million cells were sorted. Besides the two transfected cell cultures, two additional controls were introduced: regular 37°C cell culture and a cell culture treated only with the transfection agent (no DNA). The cells were lysed and prepared for BU proteomics analysis using the STrap protocol, as described in the manuscript of the unpublished article (see Chapter 4.1.2). The identification and quantification of proteins, then data analysis (protein fold enrichment and statistical significance calculation) were all performed in the Proline software [START_REF] Bouyssié | Proline: an efficient and user-friendly software suite for large-scale proteomics[END_REF]. The Gene Ontology term enrichment was done using GO PANTHER search engine, using the list of augmented proteins between the two experimental groups, applying Fisher's test and Bonferroni correction for multiple testing. As a reference background, we used the list of proteins identified in the experiment.

Besides the main experimental procedure, one GC2spd cell culture plate (10cm petri dish) of each transfection (with GFP-P2A-PSMA7 or GFP-P2A-PSMA8 construct) was lysed with 2mL of Lysis buffer (CoIP) and the lysate was clarified. Then, the 20S proteasome was CoIP-ed from the lysates, by incubation with 50µL MagBeads that carry 50µg of anti-α2 Ab. Then the beads were washed and complexes eluted with 500µL of 0.1M Glycine at pH3.5 (5 min incubation) and the flowthrough (FT) was kept for analysis. The samples were diluted in Laemmli (50µL). Bovine testis lysate was used for the positive control (the GC-2spd is a murine cell line). The anti-α4s Western blot was performed in order to establish if the recombinantly-expressed α4s is indeed incorporated into the proteasome core.

Results

The typical cell sorting plots represent the fluorescent probe intensity versus the forwardscattered light, directly related to the cell volume. Figure 34.A displays the parameters of each sorting group. On the panel B, the fluorescence intensities versus cell count are displayed. We can see that the majority of the cells display no fluorescence signal (background noise, intensity <500 RFU) and very few of them express GFP fluorescence (intensity >500 RFU). On the panel C, the table is indicative of relative count estimates for each cell sorting group. Nonetheless, one million cells expressing our GFP-construct (with RFU>500) were sorted for each group.

As described in the Experimental section, both PSMA7-expressing and PSMA8-expressing cell cultures were lysed and the 20S proteasomes purified, in order to verify by Western blot the incorporation of the α4s subunit into the proteasome core (see Figure 35). The same amount of cells and the same amount of beads were used for both cell culture IPs. The anti-α4s staining shows the strongest signal in positive control (bovine testis lysate) and in the purified 20S from α4s (PSMA8)transfected cell line. Then follows the flowthrough (FT) of the IP. Some signal is however present in the 20S purified from the α4-transfected cell line, probably due to the non-specific binding of the antibody to highly-concentrated α4 subunit. Another interesting observation are the 50 kDa bands present only in the FT of the IP. We very commonly observe these bands in IP samples and we attribute them to the antibody fragments. Namely, some of the Protein G-tethered antibody remains in the FT after overnight incubation. Our initial doubt was that some of the recombinant protein would remain non-cleaved, however, this chimera would be at least 61 kDa in size. Interestingly, the bands from the IPs appear somewhat at a higher MW than that of bovine positive control. For both constructs: PSMA7-expressing and PSMA8-expressing, three cell groups were analysed in BU proteomics (GFP-feeble, GFP-negative, GFP-intense). Besides these cells, just a regular cell culture and the cell culture treated with the PEI transfection agent were used as two additional control groups. The GFP-intense groups were selected for further analysis, since they were the only one to overexpress α4 or α4s. The table is the witness of the low transfection rate and presumably high cell mortality. We compared the proteomes of the PSMA8-expressing (α4s) and PMSA7-expressing (α4) cells first to the control groups (normal cell culture, transfecting agent-treated cell culture), then between themselves, in order to narrow down the number of proteins to observe. Namely, we considered relevant for observation only the proteins that change in expression when the PSMA7 or PSMA8 gene are expressed. We then shortlisted the proteins into four groups:

 overexpressed in PSMA8  underexpressed in PSMA8  overexpressed in PSMA7  underexpressed in PSMA7,
which represent the differentially expressed proteins (two-sided homoscedastic T test, (p value) ≤ 0.05) in the two respective groups of cells, compared to the control groups. The protein lists were searched for Gene Ontology term enrichment, against a reference list of all identified proteins in the experiment. Only one of the protein groups, namely, overexpressed proteins in PSMA8-expressing cells, yielded results, as presented in Table 3, while all the others returned no statistically significant enrichment results in GO term enrichment for biological process. Interestingly, all of the GO IDs were relevant to gene expression or chromatin remodelling, as referred in Table 3. The "fold enrichment" column refers to the enrichment in the GO term occurrence in the overexpressed protein list compared to the reference list, and not to protein abundance fold change. The p-value column is the best descriptor of the significance of a certain GO term hit, since it reflects the (im)probability that the hit occurred by chance.

To better describe the GO term enrichment query output, we performed visual regrouping of the GO terms, using REVIGO engine [START_REF] Supek | REVIGO Summarizes and Visualizes Long Lists of Gene Ontology Terms[END_REF], as presented in Figure 36. The colour intensity represents the corrected p-value -more red means more significant. The size of the circle corresponds to the "generalness" of the GO term -the bigger the circle, the more frequent the term is in the database and the smaller it is -the more specific process it describes. The regrouping is performed in semantic space -meaning it has no inherent quantitative meaning. It is there to visually separate the terms that are less semantically related and collate together more related terms. Table 3. Gene Ontology term enrichment for biological process -search results for proteins increased in PSMA8-expressing group vs. non-transfected cells, along with relevant parameters. The corrected p-value and fold enrichment were calculated based on the reference list of all the proteins identified in the proteomic analysis of the GC-2spd cell line. Although the initial goal of the experiment was to identify novel substrates of the spermatoproteasome, in the PSMA8-expressing group, no proteins that are significant in the biological context of spermatogenesis were found to be specifically underexpressed. Among reduced proteins that could potentially be interesting is CDK4 -cyclin-dependent kinase 4, reduced by 60% in PSMA8 versus control but not significantly augmented in PSMA7 versus control.

We next wanted to observe how these relevantly augmented proteins (two-sided homoscedastic T test, (p value) ≤ 0.05) compare between the PSMA7-expressing and PSMA8expressing groups, as presented in Figure 37. The gene names are used for corresponding proteins, for the sake of clarity. Proteins that are more abundant in the PSMA7-expressing cells are found on the left side, while on the right side the proteins are found more abundant in the PSMA8-expressing cells. The points in blue represent the proteins which cross the significance threshold of 0.05 for pvalue and of at least 2-fold change. On the far right we see PSMA8, in red, while PSMA7 is slightly on the left, below both significance thresholds. There is no way of distinguishing the intrinsically compared to the extrachromosomally-expressed α4 levels, other than comparing their abundances to those of other non-catalytic proteasome subunits or to the intensities in the control groups. Both cell lines have intrinsic α4 protein levels, which explains why the extrachromosomally-expressed α4 is not as clearly distributed as α4s. In orange are the points that represent proteins enriched under similar GO terms for chromatin re-organisation, in the right-hand side of Figure 37. These proteins are more expressed in the PSMA8-expressing cells.

Discussion

Although this experiment was initially designed to identify the substrates of the spermatoproteasome, to our understanding, it did not provide any unambiguous identification of such substrates. In the few proteins that are underexpressed in the PSMA8-expressing cells, none correspond to the expected protein groups. Since the context of PSMA8 expression is spermatogenesis, we were expecting to see the disappearance of cyclins or cyclin-related proteins, a decrease in Ub-ylated proteins or histones, none of which is the case. It is possible that the cellular context in which PSMA8 was expressed does not correspond to the native function of the spermatoproteasome, even though the GC-2spd cell line originates from spermatocytes immortalized in the beginning of the meiosis and should have a somewhat similar proteome compared to the original cells. Another possibility is that the spermatoproteasome requires other relevant proteins to be expressed in order to function properly, such as PA200 or 19S regulators, or PI31 and Fbxo7, which could serve as adaptor proteins for substrate targeting. Moreover, there exists a possibility that a good portion of the recombinantly expressed, 20S-incorporated α4 or α4s is being Ub-ylated, since the bands of the IP groups in the Figure 35 appear somewhat higher than the positive control -around 6kDa, which corresponds to one Ub moiety. This could bias the findings and more investigation would be needed to confirm this.

On the other hand, we unexpectedly observed that chromosome reorganization-related genes are being specifically overexpressed in the PSMA8-expressing cells (Figure 37 and Table 3). Indeed, chromatin reorganization occurs throughout spermatogenesis [START_REF] Rathke | Chromatin dynamics during spermiogenesis[END_REF]Wang et al., 2019), in order to condense the chromatin and activate meiosis-specific genes, however, how the spermatoproteasome could trigger this shift towards a specific biological process is not clear. Potential candidate for spermatoproteasome substrate, whose degradation could maybe help to induce such changes is the cyclin-related protein CDK4, reduced by 60% in PSMA8 versus control but not significantly augmented in PSMA7 versus control; furthermore, it is significantly decreased in PSMA8 versus PSMA7-expressing group (data not shown).

Conclusions and perspectives

The experiment does not seem to be able to identify direct spermatoproteasome-specific substrates. However, we did observe an interesting induction of genes responsible for chromatin remodeling, including several types of histones, in the cells expressing the spermatoproteasome, although it is not clear why, since to our knowledge, we do not identify any negative transcription factor or any inhibitor of a positive transcription factor to be specifically degraded by the spermatoproteasome. An experimental design including several cell lines could show whether this gene induction is consistent across cell lines and an inherent property of the spermatoproteasome.

In order to study spermatoproteasome substrates, several other approaches are possible. First, the MS analysis of proteolytic peptides (MAPP) can also be applied to study the spermatoproteasome. This method permits to study the proteasome substrates by analysing the peptides trapped inside the cross-linked proteasome (Wolf-Levy et al., 2018). Namely, a cross-linker is used to enzymatically inactivate the proteasomes by covalently binding them to their substrates in an ex vivo setting. The proteasomes are then IP-ed along with their substrates and the cross-links are reversed. The complexes are then loaded on a Solid Phase Extraction column, and in a two-step elution, the substrate peptides are eluted, followed by the elution of proteasome complexes. The peptide elution fraction can now be used to identify the substrates by LC-MSMS. Applied to the study of the spermatoproteasome directly in testes, this approach would have the advantage of a native environment, with the possibility to target exclusively the spermatoproteasome, using our α4sspecific antibody, as well as the total proteasome using the anti-α2 antibody.

Second, proteome analysis of the testes from the mouse PSMA8 KO model can be examined. The absence of PSMA8 in the gamete cells could potentially lead to the accumulation of spermatoproteasome substrates. In gene knock out animal models, the cells often undergo metabolic and proteome rearrangement atypical of their lineage. It is thus crucial to study the PSMA8 KO model organs at the age when the PSMA8 is supposed to begin be expressed (19 days postpartum), to avoid that the adaptive mechanisms of gamete cells interfere with the proteome study. Another possibility would be the development of a conditional KO model, whereby the effect of this gene on the proteome could be studied on the already developed cells that are otherwise not allowed to develop, due to detrimental consequences of the constitutive KO for their lineage. A collaboration is already established with the group of Alberto Pendas (University of Salamanca, Spain), that developed this KO model [START_REF]The PSMA8 subunit of the spermatoproteasome is essential for proper meiotic exit and mouse fertility[END_REF] and the material production is in progress.

Among the interactors identified in the spermatoproteasome interactome experiment (Chapter 4.1.2), the Fbxo7 -part of an E3 ligase, seems crucial in this biological context [START_REF] Rathje | A Conserved Requirement for Fbxo7 During Male Germ Cell Cytoplasmic Remodeling[END_REF]. Thus, identification of Fbxo7 targets in this context would seem to somewhat overlap with the study of the spermatoproteasome substrates. The KO model for Fbxo7 has already been developed [START_REF] Rathje | A Conserved Requirement for Fbxo7 During Male Germ Cell Cytoplasmic Remodeling[END_REF] and the differential study of its proteome compared to wild type might be complementary to the spermatoproteasome substrate experiments.

Collaborations on articles

During the realization of the work presented in this thesis, I (the author) published several articles as a result of the collaboration in subjects related to its principal subject, namely, the study of proteasome complexes using mass spectrometry approaches. Three of the collaborative papers will be listed in this chapter, specifying my contribution to each paper.

Publication 1

The proteasome generates the peptides involved in MHC class I antigen presentation and is therefore involved in the way these cells are recognized by T-cells. In the case of autoimmune diseases, such as type 1 diabetes, the β-cells are recognized as non-self and destroyed. In this work, which can be read in the Annex chapter (Publication 1), we studied the proteasome composition of pancreatic β-cells in order to better understand the context of the autoimmune diseases in which these cells are involved.

We showed that the immunocatalytic subunits are constitutively expressed in the β-cells in moderate amounts, and participate in the 20S proteasome formation. They can be further induced by the introduction of proinflammatory signals, such as IL-1β and IFN-γ, which in turn increased all three types of the proteasome peptidase activity, in cell lines and both human and mouse islets. Using BU proteomics approach, we identified the β5i subunit as the constituent of the intermediate immunoproteasome in the INS-1E cells, but no immunoproteasome (as the β2i subunit was not detected). The purified proteasome mix was found to interact mostly with the 19S regulator, and to a smaller extent with PA28αβ.

This study laid the basis for further investigation of the genesis and treatment of the autoimmune response in pancreas, by identifying the intermediate immunoproteasomes that are constitutively expressed in the islet cells and whose expression can be increased in β-cells by small quantities (pg/mL -level) of IL-1β.

My contribution to this work consisted in performing the BU proteomics analysis of the INS-1E cell line (model of pancreatic β-cells) to assess their proteasome composition. Briefly, the CoIP of the formaldehyde cross-linked proteasome complexes using anti-α2 antibody was performed and the LC-MSMS analysis was done as previously described [START_REF] Fabre | Comparison of label-free quantification methods for the determination of protein complexes subunits stoichiometry[END_REF][START_REF] Fabre | Label-Free Quantitative Proteomics Reveals the Dynamics of Proteasome Complexes Composition and Stoichiometry in a Wide Range of Human Cell Lines[END_REF]. Collected raw data was analysed using the Proline software [START_REF] Bouyssié | Proline: an efficient and user-friendly software suite for large-scale proteomics[END_REF], for the proteasome composition, through the label-free relative quantification approach described in detail in the Materials and Methods section of Chapter 4.1.2.

Publication 2

Similar contributions of relative quantification of the proteasome and its associated regulators through proteomics were published in another collaboration (Abi Habib et al., 2020), which can be found in the Annex chapter (Publication 2).

Here, we dealt with the efficiency of different proteasome subtypes to degrade Ub-ylated or oxidized substrates, which has broad physiological implications, such as to maintain the protein homeostasis and degrade potentially toxic, denatured proteins during oxidative stress damage, or to signal the immune system through changes in the MHC-bound peptide repertoire. Four different proteasome types were studied: standard proteasome, immunoproteasome and two intermediate immunoproteasome types of the following catalytic subunit composition: β1-β2-β5i and β1i-β2-β5i.

No difference in the capacity to degrade Ub-ylated substrates was observed for the 26S complexes of each proteasome type. However, the 20S were shown to be more efficient at degrading the oxidized or intrinsically disordered proteins (IDPs) than the 26S complexes. Among the different 20S types, the immunoproteasome and its intermediate 20S complexes were the most efficient in oxidized or IDPs. Upon further investigation, it was confirmed that the β5i subunit (found in the immunoproteasome and all of its intermediates) plays a crucial role in stimulating this type of activity. The degradation signal was shown to be related to the disordered protein structure, rather than the exposedness of hydrophobic patches or oxidized residues. Proteomics analysis revealed that upon oxidative stimuli, the 26S partially dissociates to release 20S, evidently, in order to more efficiently degrade the denatured proteins, that do not require the ATP-ase dependant unfolding by the 19S regulator. Moreover, the oxidative stress induces the PA28γ and PA28αβ regulators to bind to the 20S, although their expression levels were not increased in the lysates, indicating active regulator recruitment.

In conclusion, upon oxidative stress induction in these cell line models, the 26S proteasome complexes partially dissociate to enable more efficient disordered protein degradation by the 20S proteasome and the β5i subunit enhances this type of proteasome activity. This work is in accordance with previous findings on a different cell line, that the oxidative stress induces 26S dissociation (Wang et al., 2017).

Publication 3

Finally, modest contributions were made to the work of Lesne et al. [START_REF] Lesne | Conformational maps of human 20S proteasomes reveal PA28-and immuno-dependent inter-ring crosstalks[END_REF] in the comparative study of regulator binding to the standard proteasome versus immunoproteasome, by measuring the peptidase activity of the proteasome complexes via fluorogenic peptide activity assays.

In this study, HDX-MS was used to probe the structural features of the two aforementioned proteasome subtypes and their interaction with the PA28αβ and PA28γ regulators. More precisely, we investigated what are the specific structural changes induced upon regulator binding for each 20S-PA28 pair. Can we specify the differences between the standard and the immunoproteasome in the way they interact with the activators, due to the allosteric changes in the binding surface of the α ring induced by the immuno subunit incorporation? Firstly, we observed inner-to-outer conformational change in proteasome rings induced by the immuno subunits being incorporated into the 20S core. Notably, the rearrangement of the α3 Cterminus, difficult to study in this context via other techniques due to its flexibility. The central pore of the 20S and the β interface of the α ring were more flexible in the immunoproteasome compared to the standard proteasome, especially the α5 subunit facing β5i.

Next, we observed a general behavior for PA28 activator binding, whereby the interaction of the PA28 activation loops and the C-termini induce allosteric changes on the other side of the regulators. Loops missing in any PA28 structure were observed to be stabilized upon PA28 binding to the standard proteasome but not the immunoproteasome. PA28αβ seems to be more protected than PA28γ from the H/D exchange upon binding to the proteasome, which did not convey different proteasome activation. Nonetheless, upon binding to the 20S, different deuteration patterns seem to be induced in the proteasome between PA28αβ and PA28γ, indicating different activation mechanisms.

Overall, we observed outer-to-inner allosteric changes upon PA28 binding for every 20S-PA28 pair, with overall 20S core destabilization upon binding of regulators, while the opening of the central 20S pore seems to occur in a manner specific to each 20S-PA28 pair.

The HDX offers great performance across a very broad range of masses, while at the same time studies the complexes in their native state. However, complexes of such size and heterogeneity were never studied by HDX-MS and in addition to unveiling the allosteric changes upon regulator binding to the 20S and the differences between the two proteasome types in this process, the achieved developments presented in this work represent a technological achievement. This article also sets the basis for the HDX-MS analysis of the spermatoproteasome vs. standard proteasome presented in Chapter 4.1.2.

Conclusions and Perspectives

By establishing complementary BU proteomic approaches to study proteasome complexes, we managed to characterize a major shift in the proteasome composition throughout spermatogenesis, both in the composition of the 20S core and in the panel of proteasome activators. Namely, we observed that from the pachytene spermatocyte stage onwards, the spermatoproteasome-specific subunit, α4s, completely replaces the standard subunit α4 in the 20S core and does not coexist with immuno-subunits within the same 20S complex. This shift seems to be mainly transcriptionally driven. Under physiological conditions, the spermatoproteasome is highly activated by the 19S and PA200 that are the major regulators and this correlates with higher in vitro tryptic and chymotryptic activities in α4s-containing cells compared to those bearing α4. Among other proteins that interact with the spermatoproteasome, we found PI31 and Fbxo7, previously identified to be important for spermatogenesis. Moreover, we identified several synaptonemal proteins as specific interactors of the spermatoproteasome, confirming recent observations and suggesting that the shift of α4 to α4s might be important for proteasome complexes cellular relocalization.

Using TD proteomics, we managed to identify the exact proteoforms of the proteasome subunits present in mammalian testis and to relatively quantify them. Through structural MS study, we identified differences in flexibility between the α4 and α4s subunits, whereby α4s is more flexible at its C-terminal region, a domain formed by two helices protruding outwards from the proteasome core. These findings were further confirmed by molecular dynamics simulations, which predict higher number of hydrogen bonds between these two helices in the α4 compared to α4s, and reversely, higher interaction with water for α4s.

We then wanted to understand the implications of these structural differences by studying the way the two proteasome subtypes interact with the regulators we previously identified as relevant in the spermatogenesis context. Pull-down assays with 19S, PA200 and PI31 showed higher affinity of 19S towards α4s-containing proteasome compared to α4-containing proteasome. On the contrary, PI31 showed greater affinity towards the α4-containing proteasome. To further understand the implications of these structural differences, we compared the peptidase activity of the proteasome from testis and from muscle, both at the basal stage and upon activation by PA200. The two proteasome subtypes had the most pronounced differences in the trypsin-like activity, while PA200 strongly stimulated the caspase-like activity in both proteasomes.

Higher affinity of spermatoproteasome towards 19S can be explained by the need of the cell for high protein turnover during differentiation. The cell thus requires more active proteasomes. During spermatogenesis, histones are sequentially replaced by transitional histone types, until protamines take their place. Moreover, high transcriptional activity implies high rate of nucleosome disassembly. For both of these reasons, higher tryptic activity of the spermatoproteasome could thus be justified, since the histones are very positively charged, arginine-rich proteins and trypsin-like activity is defined as preference for cleaving after the arginine or lysine amino acid residues. However, the exact implications of differences in the behavior of the two proteasome subtypes remain to be explained in the physiological context.

More complete comparative study of the spermatoproteasome enzymatic activity should be put in place, including other regulators, such as 19S and PI31 and further in vitro testing of potential substrates, such as Ub-ylated or acetylated histones. This could greatly broaden our understanding of the enzymatic behavior of the spermatoproteasome. Knowing the spermatoproteasome substrates, especially spermatoproteasome-specific substrates, would be crucial for understanding its role. The current experimental setup, using recombinantly expressed α4s did not produce a comprehensive set of candidates. However, several different approaches would enable a study in more physiologically relevant conditions. Firstly, the MS analysis of proteolytic peptides (MAPP) can be applied to study the proteasome in the testis, by cross-linking it in its native environment along with the peptide residues of its substates, combined with an immunoenrichment (Wolf-Levy et al., 2018). Another approach consists in the comparative proteomic analysis of the testis of the PSMA8 KO mouse model and the wild type mice (already developed by our collaborators). The constitutive KO does not permit the full gamete differentiation and thus the role of the PSMA8 gene cannot be studied in these later stages. Thus, to improve the relevance of this study, an inducible KO could be developed in the future.

Mass photometry is a novel approach to study protein complexes in native conditions and has proven to be very useful in the study of proteasome-regulator interactions. However, we would need to obtain a more complete dataset to study the binding dynamic, determine the dissociation constants of 20S:PA200 and 20S:PA28γ complexes (potentially even 19S) and compare them between different subtypes of proteasomes.

Since mass photometry is limited in terms of resolution, nMS can be used to study the formation of these complexes in greater detail and potentially characterize the exact Multi-Proteoform-Complexes that participate in the binding. PI31 is a great candidate for the nMS study, since it is too small to study its binding by mass photometry and nMS would allow us if certain phosphorylated state of PI31 has a better affinity for the proteasome.

HDX-MS can be employed to answer many interesting spermatoproteasome questions. First, does the α4s incorporation into the 20S result in structural rearrangement in other parts of the proteasome, particularly in the β rings? To answer this, instead of ~50% spermatoproteasome mix from testis, pure spermatoproteasome would be required for such analysis.

Next, the interaction of PI31 and 20S would be interesting to study, since the binding site of PI31 on the proteasome is yet unknown and multiple binding mechanisms were described [START_REF] Li | Molecular and Cellular Roles of PI31 (PSMF1) Protein in Regulation of Proteasome Function[END_REF]. Besides the determination of the binding sites, a comparative study of the bound versus nonbound state for PI31:20S complex could provide insights into the mechanism of proteasome inhibition by PI31.

The proline-rich domain of PI31 has not yet been structurally characterized, probably due to its disordered structure [START_REF] Kirk | Structure of a Conserved Dimerization Domain within the F-box Protein Fbxo7 and the PI31 Proteasome Inhibitor[END_REF]. The HDX-MS study of PI31 could provide first insights into the structural features of this domain.

Akin to the study by Lesne et al. [START_REF] Lesne | Conformational maps of human 20S proteasomes reveal PA28-and immuno-dependent inter-ring crosstalks[END_REF], the differences in PI31-induced structural rearrangement in two different proteasomes (standard and spermatoproteasome) could be studied by HDX-MS. So could the effect of PA200 binding, however, PA200 being a 200 kDa monomer, raises the bar for methodological improvement very high and so far, we were not able to obtain satisfactory sequence coverage (or spectral quality) for the 20S:PA200 complex (data not shown). The use of ionmobility to increase the sequence coverage of such large complexes is currently being investigated in the team and shows great promises, at the expense of a longer manual curation time.

Finally, we are left with numerous questions. What is the functional implication of the observed differences in behavior between the two proteasomes? Through which E3 ligases does the spermatoproteasome realize its function? Is the spermatoproteasome involved in cell cycle regulation, via an E3 ligase context or any other? Here we explored the potential of the mass spectrometry toolbox to characterize the structure and function of this novel proteasome type and its complexes. Furthermore, we gave rationale for the continuation of the MS-guided spermatoproteasome study, in combination with other biochemical and molecular biology techniques. In a broader context, we provide a rich, highly descriptive dataset to the proteasome community that can build upon our findings, by investigating the matter from other methodological standpoints.

Introduction

The proteasome is a multi-subunit complex essential for the proteolytic degradation of cellular proteins and in the generation of specific sets of bioactive peptides [1] influencing a variety of cellular processes e.g. transcriptional regulation, signaling and the regulation of the cell cycle progression [2][3][4][5].

Proteasomal activity is executed by the proteolytic core, known as the 20S proteasome. It consists of a stack of four heptameric rings: two outer α and two inner β rings [1]. The β rings are composed of catalytically active subunits (β1, β2 and β5) that cleave peptide bonds at the C-terminal side of proteins [6] with caspase-, trypsin-and chymotrypsin-like activities, respectively [7,8]. The standard 26S proteasome contains a 19S regulatory cap that binds the polyubiquitin chain, denatures the protein, and feeds it into the proteolytic core of the proteasome [9].

Standard proteasomes (s-proteasome) assembled with β1, β2 and β5 subunits are ubiquitously expressed, but specialized proteasomes also exist and are constitutively expressed by e.g. immune cells [9], where they represent the dominant form. Formation of the proteolytic core of these specialized proteasomes involves substitution of the constitutively expressed catalytic β1, β2 and β5 subunits with the interferon (IFN)-γ-inducible β1i, β2i and β5i subunits (alternatively termed Psmb9/LMP2, Psmb10/MECL-1/LMP10 and Psmb8/LMP7, respectively) [6,10,11]. The immune-proteasome (i-proteasome) has an alternative 20S catalytic core where all βsubunits are replaced by IFN-γ inducible β-subunits and where the 20S-associated 19S can be replaced by the 11S (also termed PA28αβ) proteasome regulator [9,12,13].

When standard and inducible subunits are present in cells, the latter are preferentially incorporated into newly produced 20S proteasomes [14,15]. Interestingly, co-expression of standard and inducible β subunits enables cells to assemble a variety of distinct 20S complexes, collectively referred to as intermediate proteasomes (int-proteasomes) [9]. The two most common int-proteasomes are composed of two inner rings containing either β1/β2/β5i or β1i/β2/ β5i. These int-proteasomes are not exclusive, as other combinations have been observed, including 20S proteasome with one constitutive (β1/ β2/ β5) and one immune (β1i/ β2i/ β5i) inner ring (also called asymmetric proteasomes, [16][17][18]).

Immune cells permanently and many other cells under conditions of oxidative stress, inflammation, cytokine stimulation, or viral and bacterial infection express and assemble iand int-proteasomes [9,19]. Recently, induction of expression of such proteasomes upon exposure of human pancreatic islets and rat and mouse insulinoma cells to INFγ and β but not high concentrations of IL-1β, was reported [18,20]. Furthermore, int-proteasomes (but not iproteasomes) are constitutively expressed in various cells, including liver, heart, kidney, lung or colon [16,[21][22][23][24]. They constitute between 1% (heart) to 50% (liver) of the total proteasome pool [16,21,23,24].

The proteasomal composition in cells has broad implications, as proteasomes exhibit diverse substrate specificities. This affects the peptide repertoire generated for presentation on major histocompatibility complex (MHC) class I molecules [13,19,25], signal transduction via e.g nuclear factor kappa-light-chain-enhancer of activated B cells (NFκB) [26] and protein degradation e.g. of proinsulin [27].

The s-proteasome is known to improve glucose-stimulated insulin secretion [28], regulate intracellular proinsulin levels [27] or protect against lipotoxic endoplasmic reticulum stress [29]. However, the functions of i-and int-proteasomes are poorly defined. Importantly, constitutive expression of inducible proteasome subunits in pancreatic β cells has not been described, but their induction upon INFγ and β treatment has been suggested to play a protective role against cytokine-induced apoptosis [20] and during antiviral responses [18]. Of special interest to type 1 and 2 diabetes pathogenesis is the constitutive profile of the β cell proteasomes and their regulation. Type 1 diabetes (T1D) is an autoimmune disease, in which tolerance to β cells is broken, with proinsulin serving as a major autoantigen. T1D is histologically characterized by pancreatic islet inflammation with increased levels of cytokines i.e. IL-1β, INF-γ/β and TNF-α, in the islet microenvironment [30]. Type 2 diabetes (T2D) arises when insulin secretion fails to meet demands mainly due to impaired insulin sensitivity, with β-cell oxidative and endoplasmic reticulum stress, lipotoxicity and glucotoxicity as consequences causing progressive loss of β cell functional mass [31]. All these cellular stresses induce an inflammatory response or are exacerbated by or associated with low-grade systemic inflammation via production of interleukin 1β (IL-1β) and IL-6 and recruitment and activation of innate immune cells [32,33]. As i-and int-proteasomes can modify e.g. signal transduction and MHC I peptide presentation, their constitutive and/or induced expression in β cells by inflammatory cytokines is of high interest and therapeutic potential.

Here, we hypothesized that β cells constitutively express active non-standard proteasomes and that the expression is upregulated by innate inflammatory signals at low levels. We therefore set out to analyze the composition of proteasomes in human and mouse islets as well as in the commonly used β-cell model INS-1E cell line in non-stimulated or cytokine-stimulated conditions. We report constitutive transcription and translation of inducible proteasome subunits (β1i/ β2i/ β5i) in β-cells, albeit with lower expression levels compared to immune celllines. Of the inducible subunits, β5i is incorporated into active proteasomes in non-stimulated INS-1E cells, forming intermediate proteasomes that constitute 14% of total proteasomes in these cells. Furthermore, mRNA and protein expression of inducible subunits is upregulated by low concentrations of IL-1β. β5i and β1i subunits were induced in all tested cellular models while β2i was induced in mouse (but not human) islets and INS-1E cells. Consequently the composition and both constitutive and stimulated activity of proteasomes in β cells has to be considered when investigating degradation mechanisms and antigen presentation on MHC I molecules of proinsulin and other β-cell proteins.

Materials and methods

Cell culture

The rat insulinoma INS-1E cell line, a gift from Claes Wollheim and Pierre Maechler, University Medical Center, Geneva, Switzerland, was maintained as previously described [11]. The mouse insulinoma MIN6 cell line, was cultured in DMEM (Life Technologies, Naerum, Denmark) with 25 mM glucose, supplemented with 10% FBS, 0.1% Penicillin/Streptomycin (P/S), 50 uM β-mercaptoethanol and 2 mM L-glutamine. The mouse lymphocyte cell line A20, donated by Prof. Søren Buus, Department of Immunology and Microbiology, University of Copenhagen, Denmark, was cultured in RMPI-1640 (Life Technologies, Naerum, Denmark), containing 10% FBS, 1% P/S, 10 mM HEPES, 50 uM β-mercaptoethanol and 4.5 g/L D-glucose. The human T lymphocyte cell line Jurkat, also from Prof. Buus, was cultured in RPMI-1640 with 10% FBS and 1% P/S. All cells were maintained at 37 o C with 5% CO 2 . All cell-lines were Mycoplasma negative.

Animal Care

B6 C57BL/6NRJ mice were housed, handled and sacrificed according to Danish legislation for animal experimentation and with prior approval from the local animal ethics committee, issued by the Department of Experimental Medicine, University of Copenhagen. Animal handling and procedures were conducted by researchers with FELASA certification and supervised by veterinarians.

Islet isolation and culture

Mouse islets were isolated by injection of LiberaseTM TL (Roche1, Hvidovre, Denmark) through the common bile duct to digest exocrine tissue. Islets were handpicked and either lysed immediately or cultured for 3-5 days in RPMI-1640 supplemented with 10% FBS and 1% P/S, at 37 o C and 5% CO 2 . All data points represent separate islet collections (tested in technical triplicates) and thus denote biological variability.

Human islets were isolated from healthy, heart-beating donors by the European Consortium for Islet Transplantation (ECIT) in Milan, Italy, with local ethical approval. The obtained islets were ~90% pure and no apparent difference in their quality was observed. Details on islet donors are included in the Table 1. Islets were cultured as previously described in [11].

Cytokine exposure

INS-1E cells were exposed to 10 ng/mL rat IFN-γ (R&D, Minnesota, USA) or 15 or150 pg/mL rat IL-1β (BD Bioscience, Lyngby, Denmark) or control medium for 24h. Human islets were exposed to 10 ng/ml human IFN-γ (BD Bioscience, New Jersey, USA) or 30 or 300 pg/ml rat IL-1β, while mouse islets were exposed to either 10 ng/mL rat IFN-γ or 50 or 300 pg/mL rat IL-1β or control medium, both for 24 hours prior to experiments.

Western blotting

Prior to experiments cells or islets were lysed in lysis buffer, consisting of 100 mM Tris (pH 8.0), 30 mM NaCl, 10 mM KCl, 10 mM MgCl 2 , 2% NP-40, 20 mM iodoacetamide and protease inhibitor cocktail (Life Technologies, Naerum, Denmark). Protein concentrations were measured using Bio-Rad Protein Assay Dye Reagent (Bio-Rad, Copenhagen, Denmark). Indicated amounts of proteins were loaded on Nu-Page 4-12% bis-tris gels (Thermo Fisher Scientific, Hvidovre, Denmark), and proteins were separated by SDS-PAGE. Gels were transferred to PVDF membranes using the iBLOT2 system (Thermo Fisher Scientific, Hvidovre, Denmark). Membranes were cut prior to incubation with primary antibodies (Table 2) overnight. Primary antibodies were diluted in 2% BSA in TBST (50 mM Tris pH 8, 150 mM NaCl, 0.1% Tween). Membranes were blotted with appropriate secondary antibodies for 1 hour. Blots were developed using chemiluminescence and captured using the Azure1Saphire Biomolecular Imager. Western blots were quantified using ImageJ software (v. 1.52a, [34]).

Proteasome activity

INS-1E, A-20 and Jurkat cells, human and mouse islets were plated in duplicates or triplicates in 96-well plates and treated with 50 nM ONX-0914, a selective inhibitor of theβ5i subunit activity (Selleck Chemicals, Rungsted, Denmark, IC50: ~10 nM for β5i, [35]) or 2 µM MG132, a broad proteasome inhibitor (Sigma-Aldrich, Søborg, Denmark) or control medium for 2 hours prior to experiments. Chymotrypsin-, trypsin-and caspase-like activity was measured through luminescent assay using commercially available Proteasome-Glo TM Assay (Promega, Nacka, Sweden) according to the manufacturer's protocol. Depicted data are averages of either technical duplicates or triplicates as indicated. The added trypsin-like, chymotrypsin-like and caspase-like activity is referred to as total proteasome activity.

Bulk mouse islet RNA Sequencing

Five hundred mouse islets were plated and exposed to IL-1β (50 pg/mL) for 10 days or left non-exposed for 10 days. Total mRNA was extracted from the islets by employing RNeasy 1 Micro Kit (Qiagen, Vedbaek, Denmark). Single-stranded, single-end sequencing libraries were generated using 35 ng of extracted RNA by means of TruSeq1 Stranded mRNA Library Prep (Illumina1, Copenhagen, Denmark), and library sequencing was done with the HiSeq 4000 System (Illumina1, Copenhagen, Denmark). Sequence files were drawn to the UCSC mouse genome NCB137/mm9. Further technical and analysis details in [36] and RNA-seq raw data are accessible here: https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE110691. In brief, expression levels of all genes were estimated by Cufflink (cufflinks v2.2.1,-p 6 -G $gtf_filemax-bundle-frags 1000000000) using only the reads with exact matches. Since specific mRNA levels were analyzed no correction for multiple testing was done. Results (RPKM) for the specific genes of 3 independent experiments were analyzed by Student's paired t-test, n = 3. RPKM for each gene is provided in Table 3.

Single-Cell RNA Sequencing of Pancreatic Islets

Each single-cell transcriptome was sequenced to *750,000 reads, sufficient for cell-type classification. Islet cell subpopulations were analyzed for PSMB8, PSMB9 and PSMB10 genes expression using published human islet single-cell sequencing data [37]. FastQ files were downloaded from ArrayExpress (accession: E-MTAB-5061). Data was analyzed with bcbionextgen (https://github.com/chapmanb/bcbio-nextgen), using the hisat2 algorithm [38] to align sequence reads to human genome version hg38 and uniquely aligned reads within RefSeq gene annotations were used to quantify gene expression with the Salmon algorithm [39]. Data is then expressed as log2 of counts per million (CPM). Only cells that passed the quality control in the original study [37] were maintained for further analysis, and the cell type classification from the original study was also maintained.

Mass spectrometry for proteasome composition analysis

INS-1E cells were grown to 90% confluence in T175 flasks. The cells were washed with HBSS before incubation with pre-warmed culture media complemented with 0.1% formaldehyde for cross-linking for 15 minutes. Next, 125 mM glycine was added for 10 minutes at 37˚C to quench the formaldehyde. The advantages of live cell cross-linking vs non-crosslinking step has been evaluated in [40]. The cells were then washed three times with HBSS and centrifuged, and pellets were stored at -80˚C for later proteasome composition analysis. Immuno-purification of the proteasomes from the in-vivo cross-linked lysates, was performed as previously described [41]. Briefly, proteasomes were purified by incubating the lysates with CNBr sepharose beads (GE Healthcare) covalently bound to the antibody specific for the α2 subunit of the proteasome (MCP21) (100 mg of beads for 0.8 mg antibody), using 150 million cells per 50 mg of grafted beads. The supernatant was collected, and the beads were washed three times with 40 bead volumes of washing buffer (20 mM Tris-HCl pH 7.6, 1 mM EDTA, 10% glycerol, 150 mM NaCl, 0.1% NP-40, 2 mM ATP and 5 mM MgCl2). Finally, proteins were eluted with 0.5 ml of elution buffer (20 mM Tris-HCl pH 7.6, 1 mM EDTA, 10% glycerol, 3 M NaCl, 2 mM ATP and 5 mM MgCl 2 ). Two additional cycles of purification were conducted, reincubating the collected supernatant with antibody-grafted beads. All fractions were pooled. LC-MS/MS analysis was performed as previously described [21,42]. Briefly, immuno-purified proteasome samples were precipitated with 20% trichloroacetic acid (TCA), washed with cold acetone and then denatured by boiling at 95˚C for 30 min in the Laemmli buffer, also reversing the crosslinks [41]. Proteins were alkylated and concentrated on 12% acrylamide SDS-PAGE gel as a single band, which was cut and washed. Trypsin digestion was then performed overnight at 37˚C and the peptides were extracted from the gel. The digestion mixture was then dried in a Speed-Vac and resuspended with 2% acetonitrile, 0.05% trifluoroacetic acid. The peptide Upregulation of inducible proteasome subunits upon prolonged, low-dose exposure to IL1-β. Five hundred mouse islets were cultured and exposed to IL-1β (50 pg/mL) for 10 days. Total mRNA was extracted, and bulk (whole pancreatic islets) sequenced and genes identified using the UCSC mouse genome NCB137/mm9. mRNA levels of PSMB8, PSMB9 and PSMB10 for inducible subunits β5i, β1i and β2i, respectively, were significantly (p = <0.0005 each) upregulated by IL-1β exposure while PSMB5, PSMB6 and PSMB7 for corresponding standard proteasome subunits β5, β1 and β2 remained unchanged. Data presented as RPMK for individual islet collection (biological replicates) in the respective conditions and their means. Results were analyzed by Student's paired t-test, n = 3.

https://doi.org/10.1371/journal.pone.0222432.t003

mixture was then analyzed by nano-LC-MS/MS using an UltiMate3000 system (Dionex) coupled to Orbitrap Fusion mass spectrometer (Thermo Fisher Scientific, Bremen, Germany). Proteins identification, validation and relative quantification were performed as previously published [41].

Statistical analysis

All samples were selected without bias and represent biological not technical variations. Distribution of islets, specifically, were randomized and independent of e.g. size and shape. As a result, samples should be homogenous and represent biological variation, and both protein expression and activity is therefore assumed to be normally distributed [43,44]. Furthermore, normality of all expression data was tested with a Shapiro-Wilk test and found normally distributed and tested using a student t-test. Meanwhile proteasome activity and cell viability each data point is represented by a mean value of technical replicates, and as such should be normally distributed according to the central limits theorem [45]. Differences between two groups were assessed by a two-tailed Student's t-test. All statistical analyses were done using GraphPad Prism (v. 6, La Jolla, CA). Data is represented as means ± SD or SEM. P-values of �0.05 were considered significant.

Results

Inducible proteasome subunits are constitutively expressed in pancreatic islets and β-cell lines

To investigate whether proteasome inducible β subunits are expressed in non-stimulated β cells we re-analyzed RNA-sequencing data of single-cells dispersed from pancreatic islets from healthy individuals [37] and found that between 3.5 to 40% of β, α and δ cells express constitutively mRNA of all inducible subunits (Fig 1A and1B). Next, we lysed human and mouse islets, INS-1E (β-cell insulinoma), A20 (B cell lymphoma) and Jurkat (T cell leukemia) cells and analyzed their protein contents by SDS-PAGE and Western blotting. As expected, immune cell lines (A20 and Jurkat) showed high expression of the inducible subunits (Fig 1C -1E). Interestingly, we detected relatively low but consistent expression of all three inducible subunits (β1i, β2i and β5i) in human and mouse islets and INS-1E cells (Fig 1C -1E).

INS-1E cells contain two major types of proteasomes

We next investigated the composition of proteasomes in INS-1E cells through immunoprecipitation (IP) of the 20S α2 proteasome subunit that is an obligatory member of all types of assembled proteasomes [46].

In vivo gently cross-linked INS-1E cells proteasomes were immunoprecipitated and samples analyzed by liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS). About 86% of the total proteasomes were found to be s-proteasomes with enzymatically active rings composed of β1-β2-β5 subunits (Fig 2). However, almost 14% of the precipitated proteasomes contained β5i replacing the standard β5 subunit, forming an intermediate proteasome β1-β2-β5i. We found neither β1i nor β2i in protein complexes precipitated with α2 subunit.

Finally, our MS data indicated the presence of proteasomal regulatory particles 19S and 11S (PA28αγ) within α2 proteasome complexes at the rate of 55.8% and 5.1%, respectively (Fig 2). However, with our experimental approach we cannot assign proteasome types to the detected specific regulatory particles. The remaining 39.1% of 20S corresponds to free (unactivated) proteasome [21,41].

The β5i-selective small-molecule inhibitor ONX-0914 reduces chymotrypsin-like activity of the β cell proteasome

We next examined the profile of proteasome proteolytic activities: chymotrypsin-, trypsinand caspase-like. The proteolytic activities were tested in unstimulated live cells by addition of specific substrates to the medium (Cell-Based Proteasome-Glo™ Assay). As shown in Fig 3A- 3C intact human and mouse islets, INS-1E, MIN6 and Jurkat cells exhibited strikingly similar profiles. Under basal conditions, chymotrypsin-like activity constituted 50-60% of total proteasome activity, with the remaining 40-50% of activity almost equally divided between trypsin-and caspase-like activities. In A20 cells, trypsin-and caspase-like activities constituted a larger part of the proteasome activity than did chymotrypsin-like activity, despite the fact that they expressed the highest amounts of β5i and β1i (both subunits have chymotrypsin-like activities).

We next probed what portion of the observed proteasome chymotrypsin-like activity can be attributed to the inducible subunit β5i. We took an advantage of a selective and potent β5i subunit inhibitor, ONX-0914. Pretreatment of human and mouse islets and INS-1E cells with 50 nM ONX-0914 for 4 hours reduced chymotrypsin-like activity by 40%, indicating that β5i is enzymatically active in those islets and cells (Fig 3D -3F). At the same time, trypsin-and caspase-like activities were not affected by ONX-0914. As expected two µM of MG-132 treatment (broad proteasome inhibitor) almost completely blocked all three types of enzymatic activities in all tested cells and islets.

Low concentrations of IL-1β upregulate β1i, β2i and β5i subunit expression in β cells

Previous work has shown that the β1i, β2i and β5i proteasome subunits are expressed in response to IFN-γ/β in cells other than that of hematopoietic origin including β-cells [20,21,41] but their expression was not regulated by the high concentrations of IL-1β treatment [20].

Here we asked, if a similar to IFN-γ/β expression upregulation can be achieved by mimicking low-grade inflammation with the application of a low stimulatory concentration of IL-1β [36].

Mouse islets exposed to 50 pg/ml of IL-1β for 10 days exhibited a significantly higher mRNA expression of β5i (Psmb8), β2i (Psmb9) and β1i (Psmb10) compared to the untreated islets (Table 3). The mRNA levels for β5 (Psmb5), β2 (Psmb6) and β1 (Psmb7) genes that encode standard subunits remained unchanged after the same exposure.

Furthermore, exposure of human and mouse islets, as well as INS-1E cells, to low concentration of IL-1β for 24 h (15 pg/ml for INS-1E, 30 pg/ml for human islets and 50 pg/ml for mouse islets) induced expression of β1i, β2i and β5i (Fig 4A -4C) with the exception of β2i in human islets. The high concentration of IL-1β (150 pg/ml for INS-1E and 300 pg/ml for human/mouse islets) further induced β1i and β5i expression (but not β2i) in INS-1E cells, but failed to induce upregulation in the subunit expression in human and mouse islets (Fig 4A and4B). As expected, low IFN-γ treatment for 24 h induced expression of all inducible subunits (Fig 4A -4C). Concentrations of IL-1β in the low range are known to increase insulin biosynthesis [47,48]. Interestingly, induction of inducible proteasome expression by low concentrations of IL-1β or IFN-γ was associated with increased proinsulin expression levels in INS-1E cells while high concentrations of IL-1β diminished proinsulin expression (S1A and S1BFig). Furthermore, low concentration of IL-1β did not decrease the viability of mouse islets and INS1-E cells over the 24 h exposure to the cytokine (S1C and S1D Fig).

Low concentrations of IL-1β increase proteasome activity in β cells

INS-1E cells exposed to either low concentrations of IL-1β or IFN-γ showed significant increase in all three proteasome catalytic activities. Furthermore, high concentration of IL-1β increased chymotrypsin-like and trypsin-like activity compared to controls, although the increase was less significant than that observed for low concentration exposure (Fig 5A). Mouse and human islets showed a similar pattern of increasing chymotrypsin-, trypsin-and caspase-like activity, when exposed to low concentrations of IL-1β or IFN-γ (Fig 5B and5C).

A high concentration of IL-1β did not have a significant effect on any of the proteasome-based catalytic activities in mouse and human islets.

Discussion

The present work shows that 1) inducible proteasome subunits are constitutively expressed in human and rodent islets and a β-cell line, 2) β5i is incorporated into an active proteasome, forming int-proteasomes and 3) inducible subunit expression is upregulated by low IL-1β concentrations. The cellular composition of proteasomes and their expressional regulation is of particular interest, because different types of proteasomes degrade proteins and peptides with different efficiency and specificity [49] influencing a variety of cellular processes including antigen presentation and thereby maintenance of peripheral tolerance or induction of autoimmunity [8,13].

The presence of int-proteasomes as normal constituents in different tissues has been established before as they have been reported to constitute up to 50% of the total proteasome pool, depending on the tissue [16,24], but a comprehensive investigation of proteasome composition in primary β cells or β cell models in non-stimulated conditions has not been performed. The Human Protein Atlas RNA-seq data indicated constitutive expression of the inducible β1i, β2i and β5i subunits in human pancreas, with 10% of the RNA sequencing reads originating from the islets of Langerhans and 75% coming from exocrine glandular cells [50]. Immunostaining of the islets for specific inducible subunits detected β5i protein by one of two employed antibodies, while β1i and β2i proteins were not detected [50]. To clarify this issue, we first investigated the expression of inducible proteasome subunits in unstimulated human dispersed islet cells. Re-analysis of the previously published data set of single-cell RNA sequencing [37] uncovered substantial subpopulations of α-, βand δ-cells that constitutively express all inducible subunits (Fig 1A and1B) placing human islets on par with other tissues that express those subunits constitutively [9,16]. When testing β cell models, as well as human and mouse islets, we found that all three inducible β-subunits were detectable at the protein level without the need for cytokine stimulation (Fig 1C -1E), although the expression levels of each subunit varied substantially between tested groups. β cells can therefore potentially assemble i-and int-proteasomes containing one, two or three inducible subunits without immune-stimulation. It is however important to stress that the number of β1i-positive β cells is low in human islets (Fig 1B ) thus substantially limiting the possibility to assemble i-proteasome. This observation should be taken into account while investigating proteasome function and composition in pancreatic β cells.

The similar profile of expression of inducible β subunits in β, α and δ cells indicates that those subunits play parallel roles in degradation of the hormones abundantly handled by each cell type ER: insulin, glucagon and somatostatin, respectively. The cellular localization of the iand int-proteasomes may also play a role in that process, as β5i and β1i subunits are found in close proximity to the ER while the s-proteasomes are homogenously distributed in both nucleus and cytoplasm [51].

Next, we used mass spectrometry to identify the proteasome subtypes in β cells. By immunoprecipitating the α2 subunit from INS-1E cell lysates, we purified active proteasomes. Eighty-six % of total proteasomes contained standard β subunits forming s-proteasomes (Fig 2). The remaining 14% contained only β5i subunit, whereas β1i and β2i could not be detected. Therefore, INS-1E cells constitutively express two types of proteasomes, the s-proteasome and an int-proteasome, where at least one βring contains a β5i subunit. We neither detected iproteasomes nor int-proteasomes with incorporation of inducible subunits other than β5i, despite their expression in unstimulated INS-1E cells. According to the rules of cooperative assembly, β1i cannot be incorporated without β5i but the opposite is feasible [52] and thus it is theoretically possible but biologically less plausible that β1i and β2i are expressed but do not participate in the formation of a pool of active proteasomes. Alternatively, sensitivity of the antibody used for the detection of β1i may be high relative to the sensitivity of the other antibodies used for subunits detection, distorting the evaluation of the intracellular stoichiometry of the inducible subunits. The MS data that failed to detect β1i and β2i incorporation into proteasomes may thus be a more valid measure of the actual subunit stoichiometry in INS-1E cells. Furthermore, according to the rules of proteasome assembly, the lack of incorporation of β1i would prevent the incorporation of β2i subunit [52]. However, we cannot rule out that β1i and β2i are incorporated but constitute the minor portion of active proteasomes, below the detection limit of our MS method.

The enzymatically active proteasomes are generally capped on one or both ends of the central 20S proteasomal core by regulatory particles 19S or 11S, but the method employed in our study does not distinguish which type of proteasome is associated with a given regulatory particle. We have found that 56% of proteasomes in INS-1E cells contained the 19S particle known to associate with all types of proteasomes [21,46,53,54], while 5.1% proteasomes contained 11S that preferentially associates with int-and i-proteasomes [12,21,46,55]. This would indicate that about one third of all INS-1E int-proteasomes are bound to 11S particles while the other two third is associated with 19S particle, (hybrid int-proteasome) or not associated with any regulatory particle and thus presumably not active. Regulatory particles dictate substrate availability and specificity with 19S recognizing client proteins marked by polyubiquitin chains and 11S being involved in the degradation of short and non-ubiquitinated peptides and antigen processing for MHC I presentation [56]. As a result, their presence within assembled and active proteasomes demonstrates that unstimulated β cells contain specialized and mixed populations of proteasomes, possibly reflecting functional specificity.

When profiles of proteasome substrate-specific activities were analyzed, we found that the islets and cell lines (with the exception of A20) all showed similar proteolytic profiles. Chymotrypsin-like activity constituted between 50 and 60% of the total proteasome activity, while trypsin-and caspase-like activities were responsible for the remaining 40-50% (Fig 3A -3C). Immune cell lines have generally been reported to express a higher basal level of inducible subunits, and the i-proteasome constitutes a dominant form of their proteasomes [8]. The fact that INS-1E cells and islets share a similar proteasomal catalytic activity profile indicates that inducible subunits codetermine the activity profiles not only in immune cells.

Off note, we have observed clear differences in the proteasomal catalytic activities in two tested immune cells models, Jurkat and A20. The latter cell profile indicates persistent if not dominant incorporation of β1 subunit (with caspase-like activity) but not β1i (chymotrypsinlike activity) and diminished activity of β5 and/or β5i subunits (chymotrypsin-like activity) that are obligatory part of active proteasomes [56]. The reasons for observed differences are not known but may indicate cancer-cell-specific adjustments, human (Jurkat) vs mouse (A20) divergence or reflect more physiologically important differences between T (Jurkat) and B (A20) cells. Finally, it is plausible that proteasome activity in A20 cells is additionally modified by e.g. post-translational modifications or altered transcription of proteasomal activators, as reviewed in [57].

We next pretreated islets and cells with a β5i selective small-molecule inhibitor, ONX-0194, and found a 30 to 50% reduction in chymotrypsin-like activity, further indicating that the β5i subunit is proteolytically active in β cells (Fig 3D -3F).

Interpretation of the pathophysiological consequences of the proteasomes diversity in β cells requires better understanding of factors influencing its expression and composition. The human genes coding for β5i and β1i map to chromosome 6 precisely between the DNA sequences coding for human leucocyte antigen (HLA)-DQ, HLA-DM and Transporter 1 ATP Binding Cassette Subfamily B Member (TAP) 1 and 2 (S2 Fig), genes known to be major determinants of antigen presentation and predisposing to autoimmune diseases, including type 1 diabetes [58]. The promoter region of β5i contains binding sites for the NFκB transcription factor (S2 Fig), but high concentrations of IL-1β, a strong inducer of NFκB [59], do not increase β5i expression [20]. Accordingly, studies in neurons have shown that high concentration of IL-1β induces Early growth response 1 protein (Egr1) that strongly inhibits transcriptional activity at the β5i promoter [60]. At the same time, Freudenburg et al. speculated that since viral infections induce IL-1β synthesis, iNOS expression and nitric oxide production impeding on mitochondrial function, the resulting reduction in ATP levels would trigger iproteasome activation and generation of altered peptides that may be immunogenic and enable killing of infected target cells as an appropriate host antiviral response [18]. How can these apparently disparate IL-1β functions be reconciled? We suggest that cytokine concentration and/or duration of exposure are a key determinants of cell fate. It has been reported that low IL-1β concentrations (0.01-0.1 ng/ml) are stimulating and protective for β cells e.g. they improve insulin biosynthesis and secretion and increase β cell proliferation, while higher concentrations (5-20 ng/ml) can induce cell apoptosis and necrosis through e.g. induction of endoplasmic reticulum and mitochondrial stress [47,48]. These two outcomes employ different cellular pathways, the stimulatory pathway depending on PKC and phospholipases and the toxic pathway on NFκB signaling. We therefore used 10-100 fold lower IL-1β concentrations in our experiments compared to previous publications [20] as well as, in case of mouse islet used for bulk sequencing (Table 3), we extended the islet exposure to IL-1β up to 10 days in order to better mimic long-term low-grade inflammation. Indeed, we found that treatment of human and mouse islets and INS-1E cells with low concentrations of IL-1β increased the mRNA (Table 3) and protein expression of all inducible proteasome β subunits (except of β2i in human islets, Fig 4), while it had no impact on mRNA levels of standard subunits (Table 3) and increased all substrate-specific proteolytic activities in human and mouse islets and INS-1E cells (Fig 5). Off note, 10 day mouse islets exposure to low concentrations of IL-1β did diminished their glucose induced insulin secretion but did not reduced islets insulin content nor induced endoplasmic reticulum stress or cell death as reported in Ibarra et al. Mol Cell Endocrinol. 2019.

Our results indicate that cytokine concentration is critical when evaluating the regulatory role of cytokines in proteasome expression and activity.

Proteasomes process proteins of both endogenous and exogenous origin and produce peptides that are complexed with MHC I. The shift in composition of proteasomes towards i-proteasomes, changes the peptide repertoire from non-immunogenic to immunogenic [18,61] and can contribute to the progression towards autoimmune diabetes [18,20,62,63]. Importantly, the observed differences in cytokine action may reflect changing conditions in the islet microenvironment during inflammatory or metabolic stress. IL-1β is a central promoter of low-grade inflammation and protection against certain viral infections, including influenza [64]. One of the possible host protective mechanisms engaged by this cytokine could involve expression and assembly of int-and i-proteasomes that would result in an increased presentation of viral antigens and/or modified self-antigens, thereby enabling T effector-cell dependent eradication of infected cells. Similarly, low grade inflammation and local IL-1β production in the islet microenvironment, could facilitate neoepitope presentation by β cells through preferential incorporation of inducible subunits to form int-or i-proteasomes. Interestingly, cells deficient in β5i show lower MHC I expression and peptide presentation, and β5i pharmacological inhibition slows disease progression in mouse models of inflammatory diseases such as arthritis and lupus [35,65]. Furthermore, β5i has also been implicated in type 1 diabetes, and its inhibition has been shown to have a protective effect [35]. This could reflect the fact that the MHC I peptides repertoire is at least in part dependent upon the activity of β5i subunit in β cells expressing int-or i-proteasomes.

The role of immune-and, especially, intermediate proteasomes in β cell pathophysiology remains to be uncovered in detail. The perspective that differential proteasome subunit expression dictates the repertoire of β-cell neoepitopes presented by MHC I deserves future investigation. Discoveries in this field could lead to targeted proteasome inhibition as treatment options in diseases with an autoimmune component. In this study, we lay the groundwork for such future investigations. For the first time, we show that int-proteasomes are constitutively expressed and active in β cells and that inducible proteasome subunits can be upregulated in β cells in response to stimulatory low concentrations of IL-1β along with increases in total proteasome activities. The turnover of intracellular proteins is essential to maintain protein homeostasis in cells. It can occur by autophagy through the delivery of proteins to lysosomes, or by proteolysis by the proteasome, a tightly regulated complex that selectively degrades undesirable or damaged proteins 1, 2 and generates small peptides to be presented on major histocompatibility complex (MHC) class I molecules and recognized by cytolytic T lymphocytes in the process of immune surveillance 3 . The proteasome is a large multi-catalytic protease complex. Its basal structure, called 20S proteasome, is composed of two copies of fourteen different subunits assembled into four hetero-heptameric rings that delimit a cylindrical complex inside which proteins are degraded. The two identical outer rings are composed of α1-α7 subunits and form the gate, which regulates the entry of intracellular proteins inside the catalytic chamber of the 20S proteasome. The two inner rings are each composed of seven β-subunits (β1-β7), three of which bear a catalytic N-terminal threonine that initiates peptide bond hydrolysis 4 . In the standard proteasome (SP), which is the most abundant proteasome subtype in most cell types at steady state, the three constitutively expressed catalytic open 1 Ludwig Institute for Cancer Research, Brussels 1200, Belgium. 2 de Duve Institute, UCLouvain, Brussels 1200, Belgium. 3 WELBIO (Walloon Excellence in Life Sciences and Biotechnology), Brussels 1200, Belgium. 4 Institut de Pharmacologie et Biologie Structurale, IPBS, Université de Toulouse, CNRS, UPS, Toulouse, France. 5 Centre hospitalier de Jolimont, service de biochimie médicale, La Louvière, Belgium. 6 Structural Biology Research Center, VIB, 1050 Brussels, Belgium. 7 Structural Biology Brussels Laboratory, Vrije Universiteit Brussel, 1050 Brussels, Belgium. 8 Brussels Center for Redox Biology, 1050 Brussels, Belgium. 9 Medizinische Klinik III, Hämatologie, Onkologie und Tumorimmunologie Charité -Universitätsmedizin Berlin, corporate member of Freie Universität Berlin, Humboldt-Universität Zu Berlin, and Berlin Institute of Health, Campus Benjamin Franklin, Hindenburgdamm 30, 12203 Berlin, Germany. 10 Partner Site Berlin and German Cancer Research Center (DKFZ), German Cancer Consortium (DKTK), Heidelberg, Germany. 11 These authors contributed equally: Nathalie Vigneron and Benoit J. Van den Eynde. * email: nathalie.vigneron@bru.licr.org; benoit.vandeneynde@bru.licr.org subunits are β1 (PSMB6), β2 (PSMB7) and β5 (PSMB5). In immune cells and in cells exposed to inflammatory cytokines, these three catalytic subunits are replaced by their inducible counterparts β1i (PSMB9, LMP2), β2i (PSMB10, MECL1) and β5i (PSMB8, LMP7), forming a second subtype of proteasome called the immunoproteasome (IP) 5 . Two additional proteasome subtypes expressing a mix of constitutive and inducible subunits are also found in normal tissues and in some human cancer cell lines: the intermediate proteasome β5i (hereafter called SIP for single intermediate proteasome), which contains β1, β2, β5i, and the intermediate proteasome β1i-β5i (called DIP for double intermediate proteasome), which contains β1i and β5i along with β2 6 .

The different proteasome subtypes are found either as free 20S particles or associated with regulatory particles (RP) that interact with the N-terminal tails of the α-subunits to facilitate opening of the gate 7 . These RP include PA28αβ (or 11S RP), PA28γ, PA200 and the 19S RP (or PA700), which associates with 20S proteasomes to form 26S proteasomes, which operate the ATP-and ubiquitin-dependent degradation of intracellular proteins. Ubiquitinated substrates are targeted to 26S proteasomes through interactions between ubiquitin and the different ubiquitin receptors of the 19S regulator. This step is followed by conformational modifications that favor translocation of the protein substrate coupled with its unfolding, deubiquitination and the opening of the proteasome gate 8-11 .

The catalytic properties of each proteasome subtype are determined by its combination of catalytic subunits. Each catalytic subunit has a different cleavage specificity dictated by its structure and the nature of amino acids that line the catalytic pocket. Three major catalytic activities were described: the caspase-like, the trypsin-like and the chymotrypsin-like activities, which respectively cleave after acidic, basic and hydrophobic residues. The β1 subunit has caspase-like activity, whereas β1i rather cleaves after small hydrophobic or branched-chain amino acids, with the so-called branched amino acid preferring (BrAAP) activity. Subunits β2i and β2 both display trypsin-like activity, β2i cleaving more efficiently than β2. Finally, the two homologous subunits β5 and β5i display chymotrypsin-like activity, β5i being more efficient than β5 6, 12, 13 . As a result, each proteasome subtype produces a unique repertoire of antigenic peptides 6, 14-16 . Because the IP has a higher propensity to cleave after hydrophobic and basic residues, it was predicted to produce peptides that bind more efficiently to MHC class I 15, 17, 18 . Aside from the production of MHC class I peptides, several other proposed functions of the IP, such as activation of the NF κB pathway 19, 20 or selective turnover of ubiquitinated proteins 21 , remain controversial 15, 22-24 .

Seifert et al. showed that the IP was more efficient than the SP at degrading ubiquitinated proteins and suggested a specific role of the IP to rapidly degrade newly synthesized defective proteins that accumulate following IFNγ exposure and are both ubiquinated and oxidized 21 . These results were soon challenged by Nathan et al. who showed in a very similar set of experiments that the SP and the IP did not differ in their capacity to degrade ubiquitinated proteins 23 . Because of these discrepancies, it is unclear whether the IP is more efficient than the SP in the ubiquitin-dependent degradation of proteins. Furthermore, the role of the intermediate proteasomes in this matter has never been explored. In order to address these issues, we first set up an experimental approach that is independent of the use of IFNγ, to study the degradation of ubiquitinated proteins by 26S SP, IP and intermediate proteasomes. We then studied how the four proteasome subtypes degrade oxidized proteins. Because in our hands the turnover of oxidized proteins could not be reliably assessed in cells, and because free 20S proteasomes were shown to degrade non-ubiquitinated oxidized proteins that accumulate upon oxidative stress 25-27 , we set up an in vitro approach to study the ATP-and ubiquitin-independent degradation of oxidized proteins by the four 20S proteasome subtypes. Finally, we explored the rules that dictate this unconventional degradation of oxidized proteins.

Results

Ubiquitinated proteins are degraded at similar rates by the four 26S proteasome subtypes. To examine the relative efficiency of the four proteasome subtypes to degrade ubiquitinated proteins, we used human HEK293-EBNA cell lines (hereafter called 293) expressing exclusively either the standard proteasome (SP), the immunoproteasome (IP), the intermediate proteasome β5i (SIP) or the intermediate proteasome β1i-β5i (DIP) 6 . These cell lines were previously obtained following successive transfection of a parental 293 cell line, which exclusively expresses the SP (hereafter called 293 SP), with strong expression vectors encoding the different inducible subunits, generating the 293 SIP, DIP and IP. These three cell lines maintained similar levels of transcription of the constitutive subunits PSMB6 (β1), PSMB7 (β2) and PSMB5 (β5) as the parental 293 SP as shown by RNA-seq analysis (Fig. S1a). However, the stable overexpression of the inducible subunits PSMB8 (β5i), PSMB9 (β1i) and PSMB10 (β2i) (Fig. S1a,b), together with their preferential incorporation into nascent proteasomes 28-30 , ensured a complete replacement of SP by SIP in β5i-transfected cells, by DIP in β1i-β5i-transfected cells, or by IP in β1i-β2i-β5i-transfected cells. Indeed, LC-MS/MS analysis of proteasomes purified from these cell lines using a sucrose gradient based technique adapted from Schmidtke et al. 31 , which is not specific for a given proteasome subtype, showed that 293 SP, SIP, DIP and IP cell lines contained exclusively SP, SIP, DIP and IP respectively (Fig. S5b). The lack of incorporation of constitutive subunits in the proteasomes of 293 SIP, DIP and IP cell lines appears to trigger their degradation, as the western blot analysis of the total lysates did not show any trace of unincoroporated subunits (Fig. S1b). Because these cell lines have the same background and only differ in their composition of proteasome catalytic subunits, they are ideally suited to compare the intracellular degradation of ubiquitinated proteins by 26S SP, IP and intermediate proteasomes.

As a first step, we monitored the half-life of the pool of ubiquitinated proteins in the four 293 cells treated with MLN7243, an inhibitor of the ubiquitin-activating enzyme 32 . The decay of ubiquitinated proteins was similar in the four cell lines, and was prevented to a similar extent by the proteasome inhibitor MG132, suggesting equal degradation of ubiquitinated proteins by the four proteasome subtypes (Fig. S2). Of note, MG132 did not fully prevent the decay of ubiquitinated proteins, which also resulted from the action of deubiquitinases, whose inhibition by PR619 increased the amount of ubiquitinated proteins (Fig. S2a). We then studied the degradation of two specific protein substrates: the cell cycle regulator p21 (CIP1/WAF1) and the transcription factor c-myc. We first verified that the degradation of these two proteins was proteasome and ubiquitin-dependent in our 293 cells: it was efficiently blocked by proteasome inhibitors MG132 and bortezomib, and by ubiquitin-activating enzyme inhibitor MLN7243 (Fig. 1a,b). Ubiquitination of these two proteins was confirmed by isolating ubiquitinated proteins using the tandem ubiquitin binding entity (TUBE)-pull down assay (Fig. S3a) 33 and detecting p21 or c -myc by western blotting: ubiquitinated p21 and c-myc were detected and they were stabilized after treatment with proteasome inhibitor MG132 (Fig. S3b,c). Finally, we also checked that the four 293 cell lines did not differ in their expression of E1 ubiquitin-activating enzymes, E2 ubiquitin-conjugating enzymes and E3 ubiquitin ligases. RNA-seq data showed similar levels of transcripts for the E1 and E2 enzymes that are associated with the ubiquitin proteasome system (Fig. S4). Likewise, the transcripts associated with the E3 ubiquitin ligases responsible for ubiquitination of p21 and c -myc 34-39 were similarly expressed in the four 293 cell lines (Fig. S4). These cell lines therefore appear as a reliable model system to determine whether the four 26S proteasome subtypes differ in their ability to degrade ubiquitinated proteins, using p21 and c -myc as paradigms.

We treated the four 293 cell lines expressing SP, SIP, DIP or IP with cycloheximide, and monitored p21 and c-myc degradation by western blot. We observed a similar rate of degradation of ubiquitinated p21 and c-myc in the four cell lines (Fig. 1c-f). Thus, SP and IP are equally efficient at degrading ubiquitinated proteins, confirming the findings of Nathan et al. 23 . Furthermore, the two intermediate proteasomes degrade ubiquitinated proteins at a rate similar to SP and IP. teins. However, the intracellular assays available to quantify oxidized proteins in cells, such as the oxyblots, proved unreliable in our hands.

Immuno and intermediate

During oxidative stress, proteasome-mediated degradation of oxidized proteins possibly occurs independently from the 19S RP and the ubiquitination system 25-27 . Moreover, previous work indicated that, under oxidative stress, 26S proteasomes are unstable and dissociate into 20S proteasomes 40-42 . Using quantification by label-free nano LC-MS/MS of proteasome-interacting proteins 43-45 , we also observed an increased proportion of 20S proteasomes with an increased binding to ECM29, a proteasome-interacting particle suggested to promote 26S proteasome dissociation (Fig. 2) 41, 42 . Accordingly, we observed a concomitant decrease in 26S proteasome (Fig. 2). These results confirm that 26S proteasomes are unstable upon oxidative stress and dissociate into free 20S proteasomes. This, together with the notion that 20S proteasomes can degrade oxidized proteins in an ubiquitinindependent manner 25, 27 prompted us to use purified 20S proteasomes for in vitro digestion of oxidized proteins.

20S proteasomes were purified from the four 293 lines using a sucrose gradient based technique adapted from Schmidtke et al. 31 . We used LC-MS/MS and silver staining of polyacrylamide gels to evaluate the purity of our preparations and confirm that they contained almost exclusively 20S proteasome subunits (Fig. S5a). LC -MS/MS also showed that each proteasome preparation solely comprises the expected proteasome subtype (Fig. S5b). Using the small peptide GTPEGLYL, which is derived from the RPT2 subunit of the 19S RP and was shown to open the proteasome gate, we observed that the gate of our 20S proteasomes displayed a closed conformation (Fig. S5c) 46 . Finally, we used fluorogenic substrates to confirm that each proteasome preparation displayed the expected activities (Fig. S5d) 6, 13 . We then incubated native and H 2 O 2 -oxidized calmodulin with these 20S proteasomes and evaluated the degradation of calmodulin on immunoblots. As expected, native calmodulin was not degraded (Fig. 3a). Indeed, full-length, folded proteins are generally targeted to 26S proteasomes in an ubiquitin-dependent manner and are therefore resistant to degradation by 20S proteasomes. However, H 2 O 2 -oxidized calmodulin was degraded by all four 20S proteasome subtypes, with intermediate proteasomes and the IP degrading oxidized calmodulin faster than the SP (Fig. 3b,c). We confirmed this observation when digesting hemoglobin with purified 20S proteasomes. Here, we followed the degradation of 3 H-labeled hemoglobin by measuring the release of small radioactive peptides, resistant to trichloroacetic acid precipitation (Fig. 3d,e). Like oxidized calmodulin, oxidized hemoglobin was more rapidly degraded by β5i -containing 20S proteasomes (Fig. 3e). In this case, however, we also observed a partial degradation of unoxidized hemoglobin by 20S proteasomes (Fig. 3d), probably because hemoglobin is highly susceptible to spontaneous oxidation, through conversion of the ferrous form (Fe 2+ ) of the heme to its ferric form (Fe 3+ ) leading to reactive oxygen species production and autoxidation of the protein 47 . Overall, our results suggest that IP and intermediate 20S proteasomes more rapidly degrade oxidized proteins.

Because some reports described degradation of non-ubiquitinated proteins by the 26S proteasome 48, 49 , we also purified 26S proteasomes from the four 293 lines and monitored their ability to degrade oxidized calmodulin (Fig. S6a-c). After 8 h of digestion, we observed no degradation of oxidized calmodulin by any of the 26S proteasome subtypes (Fig. S6c), despite their high activity on fluorogenic substrates (Fig. S6b). In the same conditions, 20S DIP efficiently degraded oxidized calmodulin (Fig. S6c right panel). These results support our conclusion that ubiquitin-independent degradation of oxidized proteins is a selective attribute of 20S proteasomes.

Alteration of protein structure targets oxidized proteins to the 20S proteasome. We next investigated whether the preferential targeting of oxidized proteins to the 20S proteasome resulted from the modification of protein conformation. In particular, the exposure of hydrophobic patches by oxidized proteins was suggested to facilitate opening of the α-gate of the 20S proteasome to favor degradation of oxidized proteins 50 .

Using circular dichroism analysis, we confirmed that oxidation of calmodulin induced a dramatic change in secondary structure (Fig. 4a), which correlated with an increased degradation by the 20S proteasome (Fig. 4b left panel). Moreover, the addition of calcium chloride to oxidized calmodulin partially restored its secondary structure (Fig. 4a) and decreased its degradation by the IP, whose intrinsic activity was not affected by the addition of calcium ions (Fig. 4b). These results confirmed the observations of Ferrington et al. 51 and suggested that the loss of protein structure and not the presence of oxidized residues targets oxidized proteins to the catalytic chamber of the 20S proteasomes. Accordingly, when we analyzed the secondary structure of oxidized and native hemoglobin by circular dichroism, we also observed a disruption of secondary structure of oxidized hemoglobin that correlated with increased degradation by the 20S IP (Fig. 4c,d).

Oxidized hemoglobin also exposed more hydrophobic patches, as measured with Nile Red (Fig. 4f). However, hydrophobic patches did not seem involved in targeting oxidized proteins to 20S proteasomes since calmodulin oxidation actually decreased exposure of hydrophobic patches, and CaCl 2 did not affect the level of exposed hydrophobic patches (Fig. 4e). We therefore conclude that disruption of protein structure is the main factor that targets oxidized proteins to the 20S proteasome.

Degradation rate is independent of the presence of methionine sulfoxides. Even though oxidized residues do not target oxidized proteins to the proteasome, their presence could explain the better degradation of oxidized proteins by β5i-containing 20S proteasomes. To test this hypothesis, we generated four 8-amino acid long precursor peptides corresponding to sequences found either in calmodulin, in the β-chain of hemoglobin or in antigenic proteins such as NY-ESO 1 54 and the LCMV nucleoprotein 55 . All four precursor peptides contained either a methionine residue (unoxidized precursor) or a methionine sulfoxide at the corresponding position (oxidized precursor). Each precursor peptide also had its C-terminal residue coupled to a quencher (EDDnp) that absorbs the fluorescent signal emitted by the fluorescent group (Abz) bound to the N-terminus. Upon peptide cleavage, the quencher is released from the fluorescent probe and fluorescence can be detected (Fig. 5). Comparing the degradation of unoxidized versus oxidized precursors, we observed that the presence of oxidized methionine either increased (Fig. 5a,d), reduced (Fig. 5c) or did not modify (Fig. 5b) cleavage of the fluorogenic peptides by the four proteasome subtypes. Interestingly, the hierarchy of cleavage of the oxidized peptides by the four proteasome subtypes did not reflect the one observed for the degradation of oxidized proteins: the SP was even in some cases more efficient than the other subtypes (Fig. 5c,d). We therefore conclude that the presence of oxidized methionines inside the substrate protein does not explain the differential degradation of oxidized proteins by the four proteasome subtypes.

The β5i catalytic subunit is crucial for degradation of oxidized proteins. We then tested whether the catalytic subunits present in 20S IP and intermediate proteasomes could explain their increased ability to degrade oxidized proteins. Since they share the β5i catalytic subunit, we hypothesized that this subunit might be responsible. To test this, we digested oxidized calmodulin and hemoglobin with purified 20S proteasomes, in the presence of the β5i-specific inhibitor PR-957. The concentration of PR-957 was chosen to block the chymotrypsin-like activity of the IP and intermediate proteasomes without affecting that of the SP. Indeed, at higher concentrations, PR-957 is known to display off-target activity on the β5 subunit (Fig. S7) 56 . We observed that PR -957 slowed down the degradation of oxidized calmodulin and hemoglobin by both IP and DIP (Fig. 6a,b). Degradation by SIP was also affected by PR -957 (Fig. 6a,b), but to a lesser extent for oxidized calmodulin (Fig. 6a). Increasing the concentration of PR-957 up to 10 µM slightly improved the inhibition of oxidized calmodulin degradation by SIP (Fig. 6c). To ascertain that the effect of PR-957 resulted from β5i inhibition and not β1i, we repeated digestions with IP and DIP in the presence of the β1i-specific inhibitor calpeptin. Calpeptin did not block the degradation of oxidized calmodulin by IP and DIP at concentrations that inhibited hydrolysis of the β1i-specific fluorogenic substrate (Fig. S8). All in all, in β5i-containing 20S proteasomes, the catalytic activity of subunit β5i plays a critical role in the degradation of oxidized proteins.

Degradation of intrinsically disordered protein tau by β5i-containing 20S proteasomes.

Our observation that ubiquitin-independent degradation of oxidized proteins by 20S proteasomes was triggered by disruption of their structure prompted us to investigate whether intrinsically disordered proteins were also degraded by 20S proteasomes. A few reports support the contention that such proteins can be degraded by 20S proteasomes in an ubiquitin-independent manner 57-59 . It is the case of tau, a protein that accumulates in Alzheimer disease and exhibits little or no secondary structure. We compared the degradation of recombinant tau incubated with purifed 20S or 26S proteasomes. We observed that 20S proteasomes degraded tau (Fig. 7a,b), while 26S proteasomes did not (Fig. 7c). This result confirmed that tau was degraded in an ATP-ubiquitinindependent manner 59 , and suggested that the disordered structure of tau targeted this protein for degradation by 20S proteasomes, in a manner analogous to oxidized proteins. Accordingly, we observed that, like oxidized proteins, tau was degraded faster by β5i -containing 20S proteasomes. Overall, these results support a model in which proteins that are disordered, either intrinsically or as a result of oxidation, are efficiently degraded in an ubiquitin-independent manner by the 20S proteasomes that contain subunit β5i, i.e. the 20S immunoproteasome and the two 20S intermediate proteasomes.

Discussion

The proteasome is key to maintaining cell homeostasis and establishing the MHC class I immune repertoire. In particular, the immunoproteasome (IP), which is expressed in immune cells and induced under inflammatory conditions, is essential for the production of a high affinity MHC class I repertoire 15, 17, 18 . In addition to the standard proteasome (SP) and the IP, two additional proteasome subtypes [intermediate proteasomes β5i (SIP) and β1i-β5i (DIP)], containing a mix of constitutive and inducible subunits, are found in healthy tissues (such as kidney, small bowel or liver) and in tumors 6, 60 . Although these intermediate proteasomes constitute around 30-50% of the total proteasome content found in healthy tissues, their specific function, aside from altering the antigenic peptide repertoire, was not yet elucidated. We therefore studied the potential role of intermediate proteasomes in protein homeostasis notably in the degradation of ubiquitinated and oxidatively damaged proteins. The controversy regarding better clearance of ubiquitinated proteins by the IP and the lack of insights about the role of the intermediate proteasomes in this process, led us to compare the four proteasome subtypes for their ability to degrade ubiquitinated proteins. We used a system composed of four cell lines of the same origin that only differ by their proteasome content. We used them to compare the four proteasome subtypes for their ability to clear the pool of ubiquitinated proteins. We observed no difference in the clearance of total ubiquitinated proteins in the four cell lines. However, the interpretation of this result was complicated by the involvement of deubiquitinases, which also play an important role in the removal of ubiquitinated proteins. We therefore studied the degradation of two specific substrates, p21 and c-myc, for which we first ascertained that the degradation was ubiquitin and proteasome-dependent. In particular, the decay of p21 and c-myc was fully prevented by inhibition of ubiquitination or proteasome activity. We monitored the degradation of these substrates by western blot using Unoxidized precursor peptides contained a methionine residue which was replaced by methionine sulfoxide (Mox) in the corresponding oxidized precursor. These precursor peptides were incubated with the four proteasome subtypes, and their degradation was monitored by measuring fluorescence at 420 nm. RFU at 420 nm of each sample was reported to the RFU at 420 nm of the unoxidized precursor degraded by the SP. The absolute RFU values at 420 nm of the unoxidized precursors degraded by the four proteasome subtypes are indicated on the histogram. All values are mean + SEM of three independent experiments. specific antibodies, bypassing thereby any involvement of deubiquitinases. We observed no difference in the degradation of p21 and c-myc in the four cell lines. By showing that the SP and IP did not differ in their capacity to degrade ubiquitinated proteins, our results differ from those published by Seifert et al. 21 , but confirm those obtained by Nathan et al. 23 . Our results also show that intermediate proteasomes SIP and DIP degrade ubiquitinated proteins with the same efficiency as SP and IP. Previous studies showed that the rate of degradation by the 26S proteasome was limited by the 19S RP 61-63 , which controls recognition and removal of ubiquitin moieties, unfolding of the protein substrate and its progression into the 20S core. This may explain why the difference in catalytic subunits of the proteasome subtypes does not affect the degradation of ubiquitinated proteins, as their impact is likely masked by the rate-limiting effect of the 19S RP. This implies, however, that the 19S RP functions similarly in the four proteasome subtypes. This is supported by crystallographic studies of SP and IP, which revealed a high structural similarity between their outer α-rings, which comprise the binding interface between the 19S RP and the 20S proteasome 12 . Moreover, no difference in 19S subunit content was observed between the four proteasome subtypes 64 . In vivo cross-linking experiments performed on the four 293 cell lines, followed by LC-MS/MS analysis of immunopurified proteasomes, confirmed that the amount of 19S RP bound to 20S proteasomes was similar for all four proteasome subtypes 64 . Taken together, our results combined with previous studies suggest that the lack of difference in the degradation of 26S substrates by the four proteasome subtypes is caused by a similar functioning of the 19S RP in the four proteasome subtypes. Although in normal conditions the 26S proteasome is responsible for the degradation of most cellular proteins 3 , its role in degrading oxidized proteins that can accumulate under oxidative stress conditions is unclear. Contradictory results regarding the protective role of the 26S proteasome under oxidative stress conditions have made the field extremely controversial. On the one hand, several studies suggested that degradation of oxidized proteins was ensured by the 26S proteasome 65-67 , showing that cells with compromised 26S proteasomes accumulated oxidized proteins under oxidative stress, and that increasing 26S proteasome activity by blocking deubiquitinase Usp14 increased the degradation of oxidized proteins under oxidative stress 65, 67 . Moreover, Manohar et al. showed that after oxidative stress, half of the oxidized proteins were conjugated with K48 ubiquitin, which targets them for proteasomal degradation 66 . On the other hand, other studies suggested that degradation of oxidized proteins was mediated by 20S more than 26S proteasomes 25, 27, 40-42, 68-70 . They showed that oxidized proteins were better degraded by 20S proteasomes 25, 68 , and that cells with compromised 26S proteasomes or with a compromised ubiquitinated system were not more sensitive to oxidative stress than wild type cells 27, 70 . Moreover, they showed that the ubiquitination system was sensitive to oxidative stress 69 and that the 26S proteasome disassembled under oxidative conditions with the help of ECM29, thus increasing the abundance of 20S proteasomes 41, 42 . Finally, the 20S role in degradation of oxidized proteins was further supported by studies suggesting that oxidative stress was associated with depletion of ATP, which is needed for 26S proteasome functioning 71-73 .

As a first step, we evaluated the effect of oxidative stress on proteasome composition. Our mass spectrometry analysis of regulators bound to 20S proteasomes confirmed an increased proportion of free 20S proteasome in oxidant-damaged cells. This increased abundance of 20S correlated with a decreased level of 26S and with a recruitment of ECM29, which triggers disassembly of the 26S proteasome 41, 42 . This confirms the observations by Wang et al. and suggests that the 26S proteasome is sensitive to oxidative stress because it partially dissociates into 20S proteasome 41, 42 . Because degradation of oxidized proteins is possibly independent of the 26S proteasome and the ubiquitin system, we separated the study of degradation of oxidized proteins from that of ubiquitinated proteins. Because in our hands degradation of oxidized proteins could not be reliably assessed in cells, we analyzed their ubiquitin-independent degradation in vitro using the oxidant-resistant 20S proteasomes. Our in vitro experiments confirmed that 26S proteasomes could not degrade oxidized proteins in an ubiquitin-independent manner, while 20S proteasomes could. When comparing the in vitro degradation of oxidized calmodulin and hemoglobin by the four 20S proteasome subtypes, we observed that their turnover was faster with the IP and the two intermediate proteasomes when compared to the SP. Our data suggest that, aside from its role in the generation of MHC class I-restricted peptides, one of the functions of the IP might be to degrade oxidized proteins induced in inflammatory conditions, i.e. when IP expression is induced. Because SIP and DIP are both expressed in healthy tissues in the absence of obvious inflammation 6, 60 , their higher propensity to degrade oxidized proteins suggests that they could protect healthy tissues from the accumulation of oxidized proteins in non-inflammatory oxidative stress conditions, in which IP is not induced. The remaining 26S proteasomes that did not dissociate in oxidative stress could also contribute to the elimination of oxidized proteins, but these proteins should then be ubiquitinated and therefore equally degraded by the four 26S proteasome subtypes, because of the rate-limiting effect of the 19S RP, as discussed above. The absence of 19S RP in 20S proteasomes eliminates this effect, thereby unmasking the effect of the catalytic subunits of the proteasome subtypes. In sum, while oxidized proteins that are ubiquitinated are expected to be similarily degraded by the four 26S proteasome subtypes, oxidized proteins that are eliminated in an ubiquitin and ATP-independent manner by 20S proteasomes appear to be better degraded by IP and the intermediate proteasomes than by the standard proteasome.

Structural studies performed on 20S proteasomes reveal that the central pores formed by the α-rings are obstructed by the N-termini of their α-subunits 74 . The gates of our purified 20S proteasomes were predominantly closed (Fig. S5c). Closed-gated proteasomes restrict the entry of proteins to their catalytic chambers. The opening of their gates requires the help of the different regulatory particles, or can be induced in vitro using hydrophobic peptides or using low concentrations of SDS 74, 75 . Our in vitro assays proved that oxidized proteins were able to enter the catalytic chamber of closed-gated 20S proteasomes without the help of any regulatory particle or any gate-opening treatment, while native proteins were unable to do so. This unregulated entry of oxidized proteins into 20S proteasomes led us to evaluate whether a change in protein structure could explain the targeting of oxidized proteins for degradation by 20S proteasomes. Our circular dichroism data clearly showed that oxidation caused drastic modifications in the secondary structure of calmodulin and hemoglobin. Moreover, using Ca 2+ to restore the structure of oxidized calmodulin, we confirmed that the entry into the closed 20S proteasome was triggered by the disrupted structure of oxidized proteins rather than the increased exposure of hydrophobic patches or the presence of oxidized residues. This notion that the lack of structure targets proteins for degradation by the 20S proteasome was fully confirmed when we tested degradation of tau, an intrinsically disordered protein. We found that 20S proteasomes efficiently degraded tau in an ubiquitin-independent manner, while 26S proteasomes did not. Although our circular dichroism data show that degradation of oxidized proteins by 20S proteasomes correlates with the secondary structure alteration, it is unclear whether disruption of only the tertiary structure would suffice for ubiquitin-independent degradation by 20S proteasomes. In future studies, it will be interesting to determine the exact role of secondary and tertiary structure modifications in the unregulated degradation by 20S proteasomes.

Our study also revealed that the degradation of oxidized proteins was dependent on the nature of the catalytic subunits of 20S proteasomes. Using the β5i-specific inhibitor PR -957, we showed that the β5i catalytic subunit was essential for the degradation of oxidized calmodulin and hemoglobin. Interestingly, β5i was the main subunit involved in the degradation of oxidized calmodulin by DIP and IP. This result contrasts with the notion that all catalytic subunits contribute to the degradation of proteins by the 26S proteasome 76 . This divergence likely reflects the fact that the ubiquitin-dependent degradation of 26S protein substrates is different from the ubiquitin-independent degradation of oxidized proteins by the 20S proteasome. Unlike 26S protein substrates, which are unfolded by the 19S RP before entering the catalytic chamber, oxidized proteins probably access the catalytic chamber of the 20S proteasome through their disordered regions, which we found to be essential for degradation by 20S proteasomes. The improved degradation of oxidized proteins in the presence of β5i could relate to an increased sensitivity of disordered regions to the β5i catalytic activity. Another possibility could be that disordered regions more readily access the β5i catalytic site through narrow side openings at the interface between the α and β-rings, which were previously suggested to allow the entry of disordered proteins and polypeptides 4 . This proposed functional link between disordered protein regions and β5i catalytic activity is strikingly supported by our observation that tau, an intrinsically disordered protein, was better degraded by β5i-containing 20S proteasomes, even though it was not oxidized.

Although our results showed that the β5i catalytic activity was central to the degradation of oxidized proteins by 20S proteasomes, we observed that IP was more efficient than DIP at degrading oxidized proteins. This suggests that the β2i catalytic subunit, which is exclusively present in IP, might also be involved. The presence of β2i could slightly alter the proteasome structure, rendering β5i more active. On the other hand, degradation of oxidized proteins by SIP was less sensitive to the β5i inhibitor PR-957 when compared to IP or DIP. This suggests that, in SIP, either β5i is not the only subunit involved in degradation of oxidized proteins, or the presence of β1 in place of β1i (which is found in both IP and DIP) alters the structure of β5i preventing full inhibition by PR-957. The latter hypothesis is supported by the fact that the PR-957 dose required to fully block the chymotrypsinlike activity of SIP is higher than for IP and DIP (Fig. S7). Solving the crystal structure of the four proteasome subtypes bound to PR-957 should help to understand this differential effect of the inhibitor.

Besides confirming the partial dissociation of 26S proteasome upon oxidative stress, our mass spectrometry approach, which relied on affinity purification with an anti-α2 antibody, allowed us to have a broad view of all proteins associated with 20S proteasomes. We observed a significant recruitment of the PA28γ and the IFNγ-induced PA28α/β regulators, highlighting their potential function in oxidative stress (Fig. 2). Because the increased binding of PA28α/β and PA28γ to the 20S proteasome did not correlate with an increased expression in the total lysate (Fig. S9), these regulators seem to be actively recruited to the 20S proteasome upon oxidative stress. An activator function was already inferred for PA28α/β in the degradation of oxidized proteins 77-79 . Since PA28α/β is expressed, like IP, in inflammatory conditions, its increased binding to proteasomes upon oxidative stress could reinforce the activity of the IP towards oxidized proteins. In future studies, it will be interesting to explore whether proteasomes associated to PA28γ are more efficient than 20S proteasomes in degrading oxidized proteins, and to investigate how this regulatory particle could affect the function of the four proteasome subtypes.

In conclusion, we show here that the SP, the IP and the intermediate proteasomes all degrade ubiquitinated proteins at similar rates. On the other hand, we confirm that oxidative stress partially dissociates the 19S regulator from the 26S proteasome, resulting in more 20S in the cell. 20S proteasomes can degrade oxidized -but not nativeproteins and this is triggered by secondary structure alterations that occur during oxidation. Accordingly, 20S proteasomes also degrade intrinsically disordered proteins, like tau. Catalytic subunit β5i is key to the turnover of oxidized and disordered proteins, which are both better degraded by 20S IP, DIP and SIP as compared to 20S SP. Therefore, β5i-containing proteasomes seem to play a key role in the clearance of disordered and oxidatively damaged proteins.

Materials and methods

Degradation of ubiquitinated proteins. Four HEK293-EBNA cell lines, each expressing one proteasome subtype 6 , were used to study the kinetics of degradation of ubiquitinated p21 and c-myc. For the kinetic analysis, 1 × 10 6 cells of the four 293 lines were treated with 50 µg/ml of cycloheximide (Cell Signaling) and collected for lysis every hour and this for 5 h.

To confirm that p21 and c-myc were degraded in a proteasome and ubiquitin-dependent manner, 1 × 10 6 293 cells expressing standard proteasome were treated either with DMSO, 20 µM of MG132 (Selleck Chemicals), 5 µM of MLN7243 (Active Biochem) or 1 µM of Bortezomib (Santa-Cruz sc-217785) for 30 min. Cycloheximide was then added at a concentration of 50 µg/ml for 5 h.

Cells were then collected and lysed in lysis buffer containing 0.1% SDS, 1% deoxycholic acid, 0.5% NP40 supplemented with protease and phosphatase inhibitors (Roche). Proteins were quantified using the BCA protein assay (from Thermo Fisher Scientific) and analyzed by western blot.

Induction of oxidative stress in 293 cells and in vivo cross-linking. 1 × 10 8 293 SP cells were treated or not with 2 mM of H 2 O 2 for 30 min. They were then cross-linked for 15 min with 0.1% of formaldehyde and the cross-linking reaction was finally quenched using 125 mM of glycine. Cells were washed three times with PBS and then lysed in lysis buffer (10 mM Hepes pH 7.9, 10 mM KCl, 5 mM MgCl 2 , 10% glycerol, 10 mM ATP, 1% NP-40, protease and phosphatase inhibitors; Roche) for 15 min at 4 °C. The lysate was sonicated, centrifuged and the protein concentration was quantified using a detergent-compatible assay (DC assay; Bio-Rad).

Immuno-purification of proteasomes and LC-MS/MS analysis.

Immuno-purification of the proteasomes from the in vivo cross-linked lysates was performed as previously described 44 . Briefly, proteasomes were purified by incubating the lysates with 100 mg of CNBr sepharose beads (GE Healthcare) covalently-bound to 0.8 mg of the antibody specific for the α2 subunit of the proteasome (MCP21). The supernatant was collected, and the beads were washed three times with 40 bead volumes of washing buffer (20 mM Tris-HCl pH 7.6, 1 mM EDTA, 10% glycerol, 150 mM NaCl, 0.1% NP-40, 2 mM ATP and 5 mM MgCl 2 ). Finally, proteins were eluted with 0.5 ml of elution buffer (20 mM Tris-HCl pH 7.6, 1 mM EDTA, 10% glycerol, 3 M NaCl, 2 mM ATP and 5 mM MgCl 2 ). Two additional cycles of purification were conducted using the collected supernatant. All fractions were pooled. 20S proteasome quantification by sandwich ELISA assay was performed as previously described 44 .

LC-MS/MS analysis was performed as previously described 43, 45 . Briefly, immuno-purified proteasome samples were precipitated with 20% TCA, washed with acetone and then denatured by boiling at 95 °C for 30 min in the Laemmli buffer. Proteins were alkylated and concentrated on 12% acrylamide SDS-PAGE gel as a single band, which was cut and washed. Trypsin digestion was then performed overnight at 37 °C and the peptides were extracted from the gel. The digestion mixture was then dried in a Speed-Vac and resuspended with 2% acetonitrile, 0.05% trifluoroacetic acid. 5 µL of each peptide sample corresponding to 2.5 µg of 20S proteasome (estimated by ELISA) were then analyzed by nano-LC-MS/MS using an UltiMate3000 system (Dionex) coupled to LTQ-Orbitrap Velos mass spectrometers (Thermo Fisher Scientific). The Mascot Daemon software (version 2.3.2; Matrix Science, London, UK) was used for database search. Protein identification and validation were performed as previously published 43, 45 .

Relative quantification of free 20S proteasome and 20S associated regulators. In order to quantify free 20S proteasome and proteasome-bound regulators, we used a mass spectrometry-based approach as previously described 45 . For each of the subunits of the complex, a Protein Abundance Index (PAI) was calculated, which is defined as the average of extracts ion chromatograms (XIC) area values corresponding to the three most intense reference tryptic peptides identified from the protein. If only one or two peptides were identified (for example in the case of low abundance proteins), then the PAI was calculated on the basis of these two XIC area values. All PAIs were normalized (NPAIs) with the total amount of 20S proteasome, which was defined as the mean of the PAIs of 20S proteasome non-catalytic subunits (subunits α1 to α7, β3, β4, and β7) divided by two because each core proteasome subunit is integrated twice in each 20S core complex.

Five types of complexes were considered: the free 20S complex, the 19S-associated, the PA28αβ-associated, the PA28γ-associated, and the PA200-associated proteasomes. The 19S-associated proteasome fraction was calculated as the mean of the normalized NPAIs of the 19S subunits. The fraction of 20S proteasome in association with PA28αβ was calculated from the normalized NPAIs of each PA28α and PA28β subunits, taking into account the stoichiometry of both subunits in the final α4β3 heptameric complex. Similarly, for the PA28γ-associated 20S proteasome, the homo-heptameric structure of the PA28γ regulator complex was taken into account. The fraction of 20S proteasome that interacts with PA200 was obtained directly from the normalized NPAI of this large monomeric protein. As each regulator can be associated with the 20S complex in a stoichiometry of one or two regulators per 20S core, we approximated the fraction of regulator-associated proteasome by considering that the overall (20S proteasome: regulator) stoichiometry was (1:1.5). Thus, the fractions obtained were divided by 1.5 to take this stoichiometry into account. Using this approach, hybrid forms of the proteasome, corresponding to a 20S proteasome core interacting with one type of regulator, such as 19S, on one side, and another type of regulator, such as PA28, on the other side, could not be considered. Finally, the free 20S proteasome fraction was considered to correspond to the remaining 20S entities, by subtracting the regulator-associated 20S forms from the total 20S complexes 45 .

Purification of the four 20S proteasome subtypes. Purification of 20S proteasome was performed as previously described 31 using frozen pellets of 1 × 10 9 293 cells. Cell pellets were lysed using 20 ml of buffer A (100 mM KCl, 5 mM MgCl 2 , 10 mM HEPES pH 7.2) supplemented with 0.1% NP-40. The suspension was passed 3 times through a 23-gauge needle then 2 times through a 25-gauge needle and finally through a 27-gauge needle. The lysate was centrifuged at 30,000 g for 30 min at 4 °C with the SW32Ti rotor and the supernatant loaded into 10 ml packed volume of DEAE-Sephacel (Amersham Pharmacia Biotech). The column was washed using 5 bead volumes of buffer A and beads were eluted in 500 mM KCl, 5 mM MgCl 2 , 10 mM Hepes, pH 7.2. All proteins that were bound to the column were collected and precipitated with 35% of ammonium sulfate overnight under agitation at 4 °C. The precipitated proteins were discarded by centrifugation at 17,000 g for 20 min at 4 °C and the supernatant was loaded onto a phenyl-Sepharose column. Elution was performed with a gradient of 35-0% of ammonium sulfate in buffer A. Fractions of 1 ml were collected, analyzed by sandwich ELISA as previously described 80 and positive fractions containing the proteasome were pooled and precipitated with 80% ammonium sulfate overnight under agitation at 4 °C. Proteins that precipitated were collected by centrifugation at 12,000 g for 10 min at 4 °C, resuspended with 4 ml of buffer A and loaded into a gradient of 15-40% of sucrose in buffer A. The gradient was centrifuged at 29,200 rpm for 36 h at 4 °C using the rotor SW32Ti. Fractions of 0.8 ml were collected, tested by sandwich ELISA to identify the fractions containing the proteasome, pooled and diluted 5 times in buffer A. Pooled fractions were then concentrated using the Vivaspin 6 (cutoff 10,000 Da). A volume of 2 ml was collected and injected into a Superose-6 10-300 GL column. Purified proteasomes were collected from this size-exclusion column at 11-13 ml and then concentrated using the Vivaspin 6. Proteasome concentration was measured by performing three independent sandwich ELISA and the proteasome purity was assessed by migration in a PAGE gel and silver staining as well as by liquid chromatography coupled to LC-MS/ MS. Proteasome activity was tested using the fluorogenic substrates Suc-LLVY-AMC, Z-LLE-AMC or Boc-LRR-AMC (Enzo Life Sciences). Each of the four proteasome subtypes was purified independently three times and the in vitro kinetic assays were performed at least twice with each batch of proteasome.

Oxidation of calmodulin with H 2 o 2 . Recombinant bovine calmodulin (Merck Millipore Cat# or calmodulin purified from bovine brain (Merck Millipore Cat#208690-1MG) were resuspended in 20 mM of Tris-HCl pH 7.0 to a concentration of 2 mg/ml (similar results were obtained with both calmodulin sources). Half of the resuspended protein was oxidized by incubating 1 mg/ml of calmodulin with 50 mM Pipes pH 6.2, 100 mM KCl, 0.1 mM CaCl 2 , 1 mM MgCl 2 , in the presence of 50 mM H 2 O 2 at room temperature for 24 h. Oxidized calmodulin was then dialyzed 3 times against 20 mM Tris-HCl, pH 7.0 and its concentration was measured using BCA protein assay (from Thermo Fisher Scientific). Oxidation of the protein was verified by separating native and oxidized calmodulin (1.2 µg) in a denaturing 12% polyacrylamide gel, which was stained with Page blue protein staining solution (Thermo Fisher Scientific). As expected, the oxidized calmodulin migrated slower than its native counterpart demonstrating that the protein was efficiently oxidized (Fig. S11). Further verification was performed by analyzing the native and the oxidized calmodulin by HPLC-MS. This showed that more than 80% of the calmodulin bore 9 methionine sulfoxides and the remaining 20% contained at least 6 methionine sulfoxides.

Degradation of calmodulin by the 20S proteasome. Native or oxidized Calmodulin (2.9 µg) were mixed with purified 20S proteasomes (2.1 µg) in a buffer containing 5 mM Tris-HCl pH 7.4, 1 mM MgCl 2 , 10 mM KCl and 0.01 mM EGTA in a final volume of 40 µl and the reaction was incubated at 37 °C. Samples (1.6 µl) were collected at different time points and the reaction was stopped by freezing the samples on dry ice. These samples were then analyzed by western blot. Where indicated, the proteolysis buffer was supplemented with 0.5 mM of CaCl 2 and incubated with the oxidized calmodulin for 30 min at 37 °C prior to the addition of the 20S immunoproteasome. In some experiments, PR-957 (S7172-5MG SelleckChem) was incubated with purified proteasome for 30 min at 37 °C prior to the addition of the oxidized calmodulin.

Labeling and oxidation of bovine hemoglobin. Reductive methylation of bovine hemoglobin (Sigma-Aldrich Cat#H2500) was carried out in 1 ml reaction volumes containing 1 mg protein, 10 nmoles 3 H formaldehyde (Isobio (Fleurus) 10 Ci/mmole Cat#ART-0311-1), 20 mM NaCNBH 3 (Thermo Fisher Scientific Cat#44892) and 100 mM Hepes, pH 7.5, according to the previously described protocol by Jentoft et al. 81 . The mixture was incubated at room temperature for 24 h. The tritium-incorporation yield measured after 10% TCA precipitation varied between 7-20%. One-half of the sample was then removed and incubated with 50 mM H 2 O 2 for 24 h, which caused discoloration of the light brown hemoglobin solution. The labeled hemoglobin treated or not with hydrogen peroxide was then dialyzed 3 times against 20 mM Tris-HCl, pH 7.5.

Samples (1.5 µg) of native or oxidized hemoglobin were separated in a denaturing 4-12% polyacrylamide gel and stained with the SilverQuest Silver Staining kit (Thermo Fisher Scientific Cat#LC6070). For each of the 2 conditions, a band corresponding to the 15 kDa hemoglobin of the two monomers was clearly visible and the intensities of the bands were similar.

Degradation of labeled hemoglobin by the 20S proteasome. Labeled hemoglobin (1.5 µg) was mixed with purified 20S proteasome (2 µg) in a final volume of 20 µl of buffer containing 5 mM Tris.HCl pH 7.4, 1 mM MgCl 2 , 10 mM KCl and 0.01 mM EGTA. The final reaction was then incubated at 37 °C and samples (2 µl) were collected after 0, 2, 6 and 24 h. The reaction was stopped by adding 23 µl H 2 O, 25 µl BSA (40 mg/ml) and 800 µl ice-cold 10% trichloroacetic acid (TCA). After 30 min on ice, the supernatant was directly transferred to a counting vial. Radioactivity of the supernatant was measured in 12 ml scintillation cocktail (Ultima Gold, Perkin Elmer Cat#6013329) using a Beckman LS6000 IC liquid scintillation counter. Where indicated, 1 µM of PR-957 was added with the digestion mixture.

Purification of the four 26S proteasome subtypes. The four 26S proteasomes were purified from the four 293 cell lines by the Ubl-affinity method 82 following the instructions of the kit (rapid 26S purification Kit-L Cat# J4320). Frozen pellets of approximately 300 × 10 6 cells of the four 293 cells were used for the purification. Protein content was measured using Bradford assay. The concentration of the purified 26S proteasomes was estimated by western blot densitometry study. The levels of α2 subunit and 20S subunits of the four 26S preparation was compared to the levels of the same subunits of a 20S proteasome preparation of known concentration. Western blot. 20 µg of protein lysates or samples collected from the degradation of calmodulin by the 20S proteasomes were separated on a denaturing 4-12% polyacrylamide gel and then transferred to a nitrocellulose (Thermo Fisher Scientific Cat#IB23001) membrane using the iBlot 2 transferring apparatus (Thermo Fisher Sci-entific). Membranes were blocked for 1 h at room temperature in PBS containing 5% of dry milk and 0.1% Tween 20, washed three times in the washing buffer (PBS, 0.1% Tween) and incubated with the primary antibody. The mouse antibody directed against calmodulin (Millipore Cat#05-173, RRID:AB_309644) was diluted at 1/5,000 and incubated 1 h at room temperature. The rabbit p21 antibody (Cell Signaling Technology Cat#2947S, RRID:AB_823586) and the rabbit c-myc antibody (Cell Signaling Technology Cat#13987, RRID: AB_2631168) were diluted at 1/1,000 and the mouse vinculin antibody (Sigma-Aldrich Cat#V9131,RRID_AB:477629) was diluted at 1/2,000. All these antibodies were diluted in the washing buffer supplemented with 5% of bovine serum and 0.02% azide. The rabbit polyclonal antibody B19 that recognizes all isoforms of tau 83, 84 (kindly provided by Jean-Pierre Brion, Université libre de Bruxelles) was diluted at 1/2,000 in PBS supplemented with 2% BSA and 0.08% azide. Diluted antibodies were incubated with the membranes overnight under agitation at 4 °C. The membranes were washed three times with the washing buffer and then incubated with the secondary antibody for 1 h at room temperature under agitation. Goat anti-mouse HRP-conjugated IgG (R&D Systems Cat#HAF007, RRID:AB_357234) and goat anti-rabbit HRP-linked IgG (Cell Signaling Technology Cat#7074, RRID:AB_2099233) were both diluted 1/2,000 in the washing buffer supplemented with 5% of non-fat dry milk. Finally western blot signals were revealed with the chemiluminescent substrate West Pico SuperSignal (Thermo Fisher Scientific). Images were acquired by Fusion Fx camera from Vilber Lourmat. Quantification was performed with the Bio-1D (version 15.06b) software from Vilber Lourmat and with Image J.

Degradation of tau

Proteasome activity in the presence of CaCl 2 . 100 µM of the fluorogenic substrate Suc-LLVY-AMC were mixed with 10 nM of purified 20S immunoproteasome in a proteolysis buffer containing 5 mM Tris-HCl pH 7.4, 1 mM MgCl 2 , 10 mM KCl, 0.01 mM EGTA supplemented or not with 0.5 mM CaCl 2 . The final reaction volume (100 µL) was incubated at 37 °C for 2 h. The fluorescence intensity was measured for the digested peptides using the Glo-Max discover microplate reader (Promega Fitchburg) (Excitation 390 nm, Emission 415-445 nm).

Circular dichroism. Native and oxidized calmodulin or hemoglobin were initially dissolved in 20 mM Tris-HCl pH 7.0 buffer. To study their circular dichroism spectra, the Tris-HCl buffer was replaced by 10 mM sodium phosphate buffer pH 7.0 by three rounds of dialysis. The concentration of the proteins were then evaluated using BCA protein assay (Thermo Fisher Scientific) and diluted in the phosphate buffer to reach a protein concentration of 0.2 mg/ml. Circular dichroism measurements were performed using a Jasco J-715 circular dichroism spectropolarimeter at 25 °C in a quartz cuvette with a 1-mm path length (#110-1-40; Hellma Analytics). Far UV spectra (190 to 250 nm) were recorded at a scan speed of 50 nm/min, 1 nm bandwith. Five spectra were measured and averaged. The percentage of α-helical and β-strand content was determined by sending the records to the Dichroweb server.

Nile Red hydrophobicity assay. Nile Red (Sigma-Aldrich Cat#7385-67-3) was dissolved in DMSO to 0.25 mM. 0.2 µM of Nile Red was mixed in 40 mM HEPES-KOH pH 7.4 with either 10 µM of native or oxidized calmodulin or 5 µM of native or oxidized hemoglobin. Where indicated, oxidized calmodulin was pre-incubated 30 min with CaCl 2 . The fluorescence intensity was measured using the Glo-Max discover microplate reader (Promega Fitchburg) and measurement parameters were set to 520 nm excitation and 580-640 nm emission.

Synthesis and degradation of the fluorescent precursors. The FRET-fluorescent precursor peptides, Abz-peptidyl-EDDnp, were synthesized by solid-phase peptide synthesis (SPPS) using conventional fluorenylmethoxycarbonyl (Fmoc) chemistry on a Symphony synthesizer (Protein Technologies Inc). Dnp Nova Tag resin was used as the support for peptide synthesis to couple the EDDnp (2,4-dinitrophenyl ethylenediamine) (NovaBiochem, Merck Cat#855053) quencher group to the C-terminal carboxyl residue of the peptide. Fmoc-Abz-OH (Bachem Cat#B-3260) was used as the building block to couple the N-terminal residue to the fluorescent group Abz (ortho-aminobenzoic). All of the peptides obtained were purified by reverse-phase HPLC to > 95% purity and characterized by mass spectrometry. Finally the lyophilized peptides were dissolved in DMSO to a concentration of 10 mM.

Unoxidized or oxidized precursors (100 µM) were mixed with purified 20S proteasomes (14 nM) in a proteolysis buffer containing 5 mM Tris-HCl pH 7.4, 1 mM MgCl 2 , 10 mM KCl and 0.01 mM EGTA. The final reaction (50 µL) was incubated at 37 °C for 2 h. The fluorescence intensity was measured using the SpectraMax 190 Microplate Reader (Excitation 320 nm, Emission 420 nm).

Statistical method.

For the analysis of the effect of oxidative stress on the proteasome-interacting proteins and for the analysis of the effect of the PR-957 inhibitor on the degradation of oxidized hemoglobin an unpaired student t-test was performed. The p values are annotated as follows: *p < 0.05; **p < 0.01; ***p < 0.005; **** p < 0.0001. For the analysis of the degradation of oxidized proteins by the four 20S proteasome subtypes, for the analysis of the degradation of p21 and c-myc by the four proteasome subtypes, for the analysis of the levels of hydrophobic patches present in native and oxidized calmodulin and hemoglobin, and for the analysis of the degradation of tau by the four 20S proteasome subtypes a one-way ANOVA followed by Bonferroni posthoc comparison was performed. The p-values are annotated as follows: *p < 0.0332; **p < 0.0021; ***p < 0.0002; ****p < 0.0001. Hydrogen-Deuterium eXchange coupled to Mass Spectrometry (HDX-MS) is now common practice in structural biology. However, it is most of the time applied to rather small oligomeric complexes. Here, we report on the use of HDX-MS to investigate conformational differences between the human standard 20S (std20S) and immuno 20S (i20s) proteasomes alone or in complex with PA28αβ or PA28γ activators. Their solvent accessibility is analyzed through a dedicated bioinformatic pipeline including stringent statistical analysis and 3D visualization. These data confirm the existence of allosteric differences between the std20S and i20S at the surface of the α-ring triggered from inside the catalytic β-ring. Additionally, binding of the PA28 regulators to the 20S proteasomes modify solvent accessibility due to conformational changes of the β-rings. This work is not only a proof-of-concept that HDX-MS can be used to get structural insights on large multi-protein complexes in solution, it also demonstrates that the binding of the std20S or i20S subtype to any of its PA28 activator triggers allosteric changes that are specific to this 20S/PA28 pair. 1 he ubiquitin proteasome system (UPS) is central to proteostasis. Its most downstream element is the 26S proteasome that clears the cells of abnormal, denatured, or damaged proteins and regulates degradation of short-lived proteins, in most cases conjugated to ubiquitin. The 26S proteasome is a highly conserved compartmentalized multicatalytic protease composed of more than 30 subunits constituting the 20S catalytic core and the 19S regulatory complex 1 . Its activity is directly involved in many cytokines and hub proteins intracellular concentration 2 , and regulates immunogenic peptide production 3 . It is the focus of intense regulation and dynamically localizes within cells in response to physiological and external perturbations 4,5 and its genetic polymorphism is directly responsible for numerous pathologies including cancer, heart disease, and type 2 diabetes to name a few 6,7 .

The main mechanism regulating the proteasome activity is the substitution of the catalytic subunits β1, β2, and β5, respectively harboring caspase-like, trypsin-like, and chymotrypsin-like activities, and constituting the standard 20S (std20S) with other subunits. For example, the immunoproteasome (i20S) contains the immuno-subunits β1i, β2i, and β5i that have different cleavage specificities and generate immunogenic peptides 8 . It is strongly expressed in immune cells and can be induced in most tissues by interferon-γ.

The proteolytic activity of the 20S is also regulated by interacting proteins, the most common being the ATP-dependent 19S activator involved in the UPS degradation pathway. However, less abundant ubiquitin-independent regulators such as PA28αβ, PA28γ, or PA200 also activate the 20S and modify its substrate specificity. In addition, a vast collection of Proteasome Interacting Proteins (PIPs) regulate or assist proteasomal functions 9-16 .

Besides its subunit composition and association with regulators and/or PIPs, the proteasome is subjected to a wide variety of posttranslational modifications (PTMs) affecting its subunits activity 17-19 . The combination of these three levels of regulation (catalytic subunits, regulators, PTMs) implies a wide proteasome subtype heterogeneity, which probably results in specialized functions that can adapt protein degradation pathways to changing conditions in the cell. One of today's main challenges is to understand how different proteasome subtypes influence its proteolytic activity and the cell function.

The proteasome has been intensely studied from a structural and functional point of view since its discovery in 1988 20 . Electron microscopy (EM) allowed to observe the 20S in complex with different regulators 21-26 together with certain catalytic intermediate-states of the 26S 27 . Covalent cross-linking coupled to mass spectrometry (MS) helped refining 26S structures 28,29 and generating structural models involving different partners, including the 19S 23 , Ecm29 15 , Ubp6 30 , and other PIPs 15 . X-ray crystallography also provided high-resolution structures of proteasomes alone or in the presence of covalent inhibitors 31 , regulators 32,33 , and complexes of proteasome with their associated regulators 15,31,32 .

The complex size is obviously a limit for its analysis by Nuclear Magnetic Resonance (NMR). However, the structure of the eukaryotic proteasome from Thermoplasma acidophilum, containing only one type of subunit α and β, has been resolved. It showed evidence of a remodelling of the catalytic sites located in the center of the proteasome upon binding of a regulatory particle on its surface, which can be described as an outer-to-inner allosteric change 34 . A reverse inner-to-outer change was also observed at the binding interface when modifying the catalytic site, but this was not confirmed by the comparison of the std20S X-ray structure 35 with the recent i20S X-ray 36 and cryo-EM 37 structures (RMSD = 0.392 and 0.480 Å, respectively). However, the outer-to-inner mechanism was recently shown in the human 20S cryo-EM structure upon PA200 binding 25 : conformational changes were found not only at the binding interface (opening of the pore), but also down to the catalytic sites.

Despite these achievements, high-resolution methods are still limited by either the size, the heterogeneity, or the dynamics of the purified complexes. This is evidenced by the small number of structures of the 20S bound to regulators and their numerous unresolved regions.

Here, we optimized the emerging method Hydrogen-Deuterium eXchange coupled to MS (HDX-MS) to investigate conformational changes occurring upon binding of the std/i20S to PA28 regulators. This method, developed in the 90 s, enables the detection of conformational differences between two protein samples. Briefly, the proteins or protein complexes of interest are diluted in a deuterated buffer and hydrogen atoms from the peptide bonds are exchanged with the deuterium atoms of the solution. The rate of exchange of each amide hydrogen depends on its solvent accessibility and involvement in the stabilization of secondary structures: regions that are relatively more solventaccessible or flexible present a higher deuteration rate than the ones that are hidden in the core of the protein or rigid. This expanding method can thus be used to study protein dynamics 38 , compare protein conformations (in different buffer conditions, after mutation, ligand binding) 39 , or to identify protein-protein or protein-ligand binding interfaces 40,41 . We present the HDX-MS analysis of the entire human std and i20S core particles as well as their PA28αβ and PA28γ regulators, mapping solvent accessibility/dynamics for each complex. The conformational maps of the std and i20S present significant differences, and their differential analysis with and without regulators provide a molecular rationale for their distinct functions. Our data provide evidence for the human 20S inner-to-outer allosteric change upon incorporation of the immuno-subunits and the reverse outer-toinner transduction signal upon PA28 binding, illustrating the interplay between the different proteasome regulation pathways. Altogether, this work highlights the potential of HDX-MS to generate low resolution but informative structural information on large hetero-oligomeric complexes. It opens the door to many other applications, including identifying PIPs binding surfaces and the stabilizing/destabilizing effects of other regulators of the 20S activity, including small molecules.

Results

Dynamic interfaces uniting the four rings of the 20S proteasome. HDX consists in the exchange of backbone amide hydrogens (H) with deuteriums (D) in proteins in solution. The exchange rate depends on the solvent accessibility and/or the flexibility of a given region, thereby providing low resolution but crucial information on protein conformation. We performed two distinct analyses (Fig. 1a): (i) identification of the most accessible and/or dynamic regions of the proteins in solution, and (ii) differential analysis of each protein alone or in complex. We monitored the incorporation of deuterium for each sample from 0 to 30 min in triplicate experiments (Fig. 1b). For differential analyses, we compared the peptide relative deuterium uptakes (RDU) of the proteins alone vs. in presence of their binding partner. The peptides were considered significantly different when they presented a P value of the ANOVA ≤ 0.01 after correction for multiple testing (Benjamini-Hochberg) and three successive time points with an absolute difference between the two conditions >4 times the experimental standard deviation observed in the dataset (Fig. 1c) (see Methods for more detailed information). The peptide-level RDUs or differences of RDU were mapped to the protein sequences as described in Fig. 1d: when several peptides covered the same region of a protein we Finally, we mapped the RDUs to the protein 3D structures in order to visualize the regions the most accessible to solvent (Fig. 1e, left panel), or presenting significant differences of deuteration (Fig. 1e, right panel). For differential analysis, we colored the 3D structures with the sum of RDU differences between the two conditions (the values per timepoint are available in Supplementary Data 1-3). In this context, modification of solvent accessibility upon complex formation can be due to the presence of a binding interface or to allosteric changes. This approach was employed to identify the conformational differences between the std20S and i20S, and to compare their structural changes upon Fig. 1 HDX-MS workflow applied to the analysis of the 20S proteasome and its regulators. a Overview of the workflows applied to the analysis of proteins alone (left) or differential analysis of proteins alone vs. in complex (right). The different steps are described in more detail in b-e as referred to on the right side. b Sample preparation and MS data acquisition. Proteins (alone or in complex) were subjected to hydrogen (blue points) to deuterium (red points) exchange in triplicate experiments for 0, 0.5, 1, 5, 10, and 30 min before being subjected to pepsin digestion. Peptides were then separated by liquid chromatography (UPLC) and analysed by MS. Spectra were curated using DynamX before statistical analysis and/or mapping of their RDU to the protein sequence (consolidation). c Statistical comparison of proteins alone vs. in complex: kinetics of solvent accessibility were compared for each peptide using ANOVA. Peptides with a Benjamini-Hochberg corrected P value ≤ 0.01 and 3 successive time points presenting an absolute difference of RDU > 4 times the mean standard deviation between replicates in the entire dataset were considered regulated and colored in the final figures. Otherwise, peptides were colored in gray in the heatmaps and 3D figures of differential analyses. d RDUs of the smallest peptides were mapped to the amino-acid portion they cover on the protein sequence. These were then presented as color gradients on heatmaps: light green and dark blue correspond to low and high RDU, respectively. The origin of RDU reported on the heatmap is indicated above the protein sequence with peptides represented as solid and dashed lines when their RDUs were reported or ignored, respectively. Residue numbers are indicated on the top. e 3D visualization of the RDU (left panel) or significant differences of RDU (right panel) measured in this study: blue and red correspond to a reduction or an increase of RDU in the complex, respectively. The mapping of peptide-level data to protein sequence was performed as described in d with the sum of RDU per timepoint (sum(mean(RDU))) or the sum of the mean of difference of RDU per timepoint (excepted T0) (sum(mean(diff. RDU))). H hydrogen, D deuterium, diff. difference, RDU relative deuterium uptake.

binding to PA28αβ or PA28γ. For this, we purchased commercial i20S purified from human spleen previously used for in vitro proteolytic assays 42-44 . We obtained 89% subunit sequence coverage on average for the 17 subunits of the std/i20S (Supplementary Fig. 1, Supplementary Data 4 and 5, Supplementary Table 1). HDX-MS results were mapped on the 3D structure of the human std20S (PDB: 5LE5, Fig. 2a,b). As expected, we found a faster and higher deuteration of the solvent-facing α-ring interface of the std20S compared to any other ring interfaces (Fig. 2c). This observation benchmarks our approach by confirming that high RDUs directly indicate regions of the proteasome that are more dynamic and potentially accessible for interaction with regulators or PIPs. The N-terminal (N-ter) part of the α-subunits constituting the α-ring were highly deuterated (2-22 in α1, 2-20 in α2, 14-18 in α3, 2-8 in α4, 4-13 in α5, 2-9 in α6, and 2-11 in α7, Fig. 2c). These N-ter are located around the proteasome pore entry, and their high RDU strongly supports the hypothesis of a flexible pore entrance in the std20S that fluctuates between open and closed states 45-47 . Furthermore, the α-ring presented very dynamic patches consisting of noncontiguous peptides from the same subunit in α3 (176-186/ 238-261), α4 (44-54/197-206), and α5 (49-63/167-181) (Fig. 2c). These may interact with many different PIPs. Our analysis also identified dynamic patches at the interface between different subunits like α1 (2-22/165-178) and α2 (194-202), α3 (14-18) and α4 (44-54), and α5 (167-181) and α6 (48-56) that correspond to the 1-2, 3-4, and 5-6 α-pockets 48 (Fig. 2c, circled in white) known to accommodate the C-terminal (C-ter) tails of Rpt3, Rpt2, and Rpt5, respectively 23,48 . We also identified flexible patches on the interface of the α-ring facing the β-ring, where amino-acid stretches of α1 (146-152), α4 (91-102), α5 (102-107), and α7 (140-148) were slightly more flexible than the remaining of the interface (Fig. 2d and Supplementary Data 3,6).

The β-ring was globally less deuterated than the α-ring (Fig. 2e, f), which was expected since it does not present a large solvent interface. However, after 30 min, some α-facing bulges were moderately deuterated in all β subunits but β7: these include the loops between α-helices 1 and 2 of β1 (66-82), β2 (67-79), β3 (75-87), β4 (68-79), β5 (67-77), and β6 (74-91) (red circles in Fig. 2e, and Supplementary Data 6). Interestingly, some of these dynamic loops face the highly deuterated regions of the α-ring described above and define flexible regions of the α-ring/β-ring interface. The most dynamic regions of the β-ring were located on its outer surface and were constituted of the C-ter of β1 (178-203), β2 (186-234), β4 (195-201), β5 (185-204), β6 (151-168), and β7 (200-214) (Fig. 2f in red and Supplementary Data 6). These regions are particularly interesting since they can potentially interact with the numerous PIPs described in the literature.

An inner-to-outer conformational change upon substitution of standard to immuno-subunits. Although mouse i20S and std20S possess very similar β2 and β2i substrate binding channels, there are structural differences between their standard and immunocatalytic subunits β1 and β5. The S1 and S3 substrate pockets are smaller in β1i vs. β1, β5i possesses smaller S2 and S3 pockets, and the S1 pocket is larger in β5i than in β5 31 . However, very little structural difference is found between the α and noncatalytic β subunits of the i20S and std20S (Root-Mean Square Deviation of the Cα < 0.72 Å). The same is true when comparing the recent structure of the human i20S 36 with the std20S 35 (RMSD of the Cα < 0.67 Å). The std20S catalytic chamber-localized at the β-ring/β-ring interface-was locally very flexible in our data (Fig. 2f and Fig. 3). The residues forming the S1 and S2 pockets 31 of β1 were highly deuterated, together with the T23 of the S3 pocket (Fig. 3b, see Supplementary Data 4 for full kinetics). The catalytic residues of β2 were poorly deuterated compared to the remainder of this subunit (Fig. 3c), with the exception of 45-53 constituting its S1 and S2 pockets. Conversely, most of the residues forming β5 catalytic site and substrate pockets were highly deuterated (Fig. 3d). The relatively poor sequence coverage that we obtained for β2i limits its analysis and we cannot comment on the accessibility of its S1, S2, and S3 pockets (Fig. 3b and Supplementary Table 1). The direct comparison of RDUs between the standard and immuno-catalytic subunits (β1/β1i, β2/β2i, and β5/β5i) was not possible due to their sequence differences. Nevertheless, we could acquire the deuteration profiles of the peptides ADSRATAGAY (16-25) of β5 and AVDSRASAGSY (15)(16)(17)(18)(19)(20)(21)(22)(23)(24)(25) of β5i that cover the same portion of the two subunits. Their difference of deuteration profiles indicate that these residues are more flexible in std20S than in the i20S (Fig. 3d and Supplementary Data 4).

Allosteric differences beyond the catalytic subunits were shown in the prokaryotic std and i20S analysed by NMR 34 . These were not observed in mouse crystallographic structures 31 , may be due to crystal packing. In order to confirm or infirm this inner-toouter allosteric change in human, we compared the RDUs of all the noncatalytic subunits of the i20S (Supplementary Data 4-6) with those of the std20S (as presented Fig. 1).

The introduction of immuno-catalytic subunits resulted in significant conformational changes on the noncatalytic 20S subunits. The results of our statistical analysis are presented in Fig. 4, and Supplementary Data 1, 4, 7-10. Overall, the noncatalytic β-ring subunits were more dynamic in the i20S than in the std20S, (>50 peptides significantly more deuterated, Fig. 4a-b in red, Supplementary Data 1, 7). The β-ring/β-ring interface and the channel were particularly more dynamic/ accessible in the i20S vs. std20S (Fig. 4a/e and Supplementary Data 1), together with two α-facing bulges of β6 and β7 (Fig. 4b, red circles). Four β-facing bulges of α2/3/5/6 were also significantly more dynamic in the i20S vs. std20S (Fig. 4c, red circles, corrected P-values and RDU differences are available in Supplementary Data 9 and 10). The pore entrance on the α-ring solvent-accessible surface (α subunits N-ter) was significantly more dynamic/accessible upon replacement of β1/2/5 by the immuno-subunits (Fig. 4d red circles, Supplementary Data 1, 7). This suggests a long-range mechanism enabling the i20S pore to dwell longer in the open-state, and could explain its generally higher activity 49 as well as the various regions from the antechambers that were more accessible in the i20S vs. std20S (98-99 in α2; 102-109 in α3; 128-142 in α5; 100-108 in α6; 95-105 in β3; 94-102 in β4, Fig. 4e, corrected P-values and RDU differences are available in Supplementary Data 9 and 10). Conversely, three solvent-facing regions of the α-ring were significantly more dynamic/accessible in the std20S than in the i20S (Fig. 4d in blue, Supplementary Data 4, 7). Since these constitute potential binding sites for the proteasome regulators, this could explain the preferential association of specific regulators to different 20S subtypes. Interestingly, these three subunits also contain regions that are more accessible in the std20S on the β-ring facing interface (Fig. 4c), suggesting that this dynamic behavior can be related to the presence of the standard catalytic subunits within the β-ring. More precisely, the α1 102-114 (corrected P value = 5.87.10 -11 , sum of difference = -12%) and 146-152 (corrected P value = 2.54.10 -7 , sum of difference = -12%), which were more deuterated in the std20S vs. i20S, are in direct contact with a portion of β2/β2i (69-72) also more accessible in the std20S (Supplementary Data 1). Altogether, these results confirm the inner-to-outer allosteric effect following incorporation of the immuno-β subunits that lead to αring remodeling: more dynamic pore entrance but protected external anchor regions (α3 C-ter).

The activation loop of PA28β is less dynamic/accessible than PA28α/γ's. There is no structure yet for PA28γ but those of the human homoheptameric PA28α (nonphysiological) and murine heteroheptameric PA28α 4 β 3 were resolved in 1997 32 and 2017 33 , respectively. These monomers are composed of a four-helix bundle, assembled as a barrel-shaped heptamer (Fig. 5) that controls proteasome catalytic activity through a conserved activation loop 50,51 .

The structures of PA28α, PA28β, and PA28αβ miss a region of 15-31 residues bridging the α-helices 1 and 2, although they were present in the recombinant constructs (Fig. 5a-c and Supplementary Data 11 in red). These apical loops, most likely flexible, sit on top of the regulator and might thereby control the substrate access to the proteasome. The human PA28γ chain has a similar loop (43 residues), and no information on its structure has ever been reported. Since the sequence identity of human PA28γ is closer to PA28α (40%) than PA28β (35%) or Plasmodium falciparum (Pf) PA28 (34%) (Supplementary Table 2), we generated a homology model of human PA28γ (Fig. 5e) using the structure of human PA28α 33 as a template with the SWISS-MODEL server 52 . The sequence of Pf is used as a comparison here because it is part of the very few structures of PA28 regulator homologues available in the PDB with close homology to PA28γ and whose structure was recently solved in complex with the 20S 24 .

The sequence coverages obtained upon digestion were above 90% for all three PA28 subunits across all conditions (Supplementary Fig. 1, Supplementary Data 11, 12 and Supplementary Table 1). Their deuteration pattern revealed similar dynamics for both PA28αβ and PA28γ with a very strong protection of most residues inside the channel (Fig. 5d, e and Supplementary Data 11). The missing apical loops (Supplementary Data 11, red rectangles) were very quickly deuterated, a sign of high accessibility/flexibility, that could explain their absence in known structures. The proteasome-facing interface (unstructured N-and C-termini and activation loops) (Fig. 5d, e, right) and the entrance of the channel (Fig. 5d, e, left) were also strongly deuterated. Remarkably, the regions involved in the binding (Cter) and activation of the proteasome were very accessible/ dynamic when both regulators were alone in solution. However, the activation loop was less deuterated in PA28β. The reason why eukaryoric homoheptameric PA28 regulators have evolved towards more complex heteroheptameric PA28αβ is not well understood. Given the overall similar tridimensional structures of PA28αβ and PA28γ, the poorer flexibility observed on PA28β activation loop could, at least partially, explain their different functions.

Allosteric changes between PA28γ and PA28αβ upon 20S binding. Before comparing the PA28 regulators alone and in complex with the std20S, we confirmed their ability to increase the proteasome activity in vitro. In our hands, a 2-fold molar excess of any of the PA28 regulators doubled the std20S chymotrypsin-like activity (P value of a paired two-sided t-test: 0.0007 and 0.0013 for PA28γ and PA28αβ, respectively; Supplementary Fig. 2 and Supplementary Table 3). As expected, the PA28 proteasome-facing interfaces were highly protected in the complex compared to the regulators alone (Fig. 6a, b right, in blue; Supplementary Data 2, 12, Interestingly, other regions were less dynamic/accessible upon complex formation. The kinks of the first α-helix of PA28α (25-32; corrected P value = 2.93.10 -15 , sum of difference = -61%) and PA28γ (35-40; corrected P value = 6.55.10 -13 , sum of difference = -30%) (Fig. 5a and Fig. 6a, b, red ovals) were strongly protected upon std20S binding. The PA28β N-ter (5-16) that faces PA28α's kink, was also significantly protected upon std20S interaction (corrected P value = 9.19.10 -8 , sum of difference = -30%) (Supplementary Data 2). We thus hypothesize that PA28α/PA28β interaction might be strengthened and/or structurally rearranged upon binding to the std20S. The N-ter (6-10) of PA28γ facing the kink was slightly but not significantly protected (corrected P value = 3.77.10 -3 , sum of difference = -6.4%) (Fig. 6b and Supplementary Data 2).

The opposite side of the heptamer was also partially protected upon std20S binding: the loops between helices 3 and 4 of PA28α (179-197), PA28β (166-184), and PA28γ (203-208) (Fig. 6a,b) as well as the flexible loop (60-101) of PA28γ (Fig. 5b). These regions include the constriction sites at the top of PA28αβ (K187 and K190 in PA28α, K177 and K180 in PA28β) that were significantly protected upon std20S binding (Fig. 6a and Supplementary Data 2). In PA28γ, another constriction site, located underneath was protected upon std20S binding (R181, Fig. 6b and Supplementary Data 2).

We then compared the impact of std20S vs. i20S binding on PA28 solvent accessibility using the statistical strategy presented in Fig. 1c (comparison referred to as PA28 + std20S vs. PA28 + i20S in Supplementary Data 2, 9, 10). Most of the reduction in accessibility/flexibility that was observed on the PA28 chains upon binding to the std20S were dampened upon interaction with the i20S (Fig. 6c,d

and Supplementary Data 2).

A PA28-driven outer-to-inner allosteric change of the 20S proteasome. Atomic force spectroscopy showed that the 20S can alternate between open and closed states, possibly through allosteric regulation 45,46 . Our data indicate that PA28αβ and PA28γ do not present the same conformational rearrangements upon binding to the 20S, so they might allosterically affect the 20S pore entrance and/or its catalytic sites in a regulator-specific outer-to-inner mechanism, as suggested recently 25 . We compared the std/i20S RDUs in presence or absence of PA28αβ or PA28γ (Fig. 7, Supplementary Data 3, 14, and 15). Strikingly, very few regions of the α-ring in contact with the regulators were significantly protected in the std20S upon interaction with PA28αβ or PA28γ, and none in the i20S. The same C-ter region of α3 (238-261) that was found to be more accessible in the std20S vs. i20S (corrected P value = 1.55.10 -7 , sum of difference = -24%) (Fig. 4d) was protected in the std20S upon interaction with PA28αβ (corrected P value = 1.62.10 -5 , sum of difference = -28%), and to a lesser extent with PA28γ (corrected P value = 2.85.10 -4 , sum of difference = -16%) (Fig. 7a, f, red circle on α3). Other regions of the std20S that were protected when binding PA28αβ form two patches on the same solvent-exposed interface (Fig. 7a, red circles on α4, α5). These could be the main anchorage sites of PA28αβ at the surface of the std20S. The remainder of the std20S solvent-exposed surface was globally more dynamic upon PA28αβ binding, especially near the pore entrance (N-ter of α1/2/3/4/5/7), which can be interpreted as an opening of the pore (Fig. 7a, large red circle). Another external patch at the α1/ α2 interface was also more dynamic upon binding of PA28αβ (Fig. 7a, red circle on α1, α2).

On the β-facing α-ring, PA28αβ binding mainly destabilized α2 (136-145: corrected P value = 2.48.10 -5 , sum of difference = 10%) and the α5 bulge (102-107: corrected P value = 1.56.10 -5 , sum of difference = 19%) (Fig. 7b, red circles on α2, α5). Interestingly, this bulge faces the 109-135 β6 stretch encompassing 4 peptides more deuterated upon PA28αβ binding and is in close vicinity to the β5 interface, which is also more dynamic (Fig. 7c, red circles on β5, β6, corrected P values and RDU differences are available in Supplementary Data 9-10). Although not close enough to make hydrogen bonds, α2 (136-145) (Fig. 7b, red circle on α2) faces the β2/β3 interface that was more flexible/ accessible upon PA28αβ binding, especially in the 186-202 C-ter region of β2 extending towards β3 (Fig. 7d, circle on β2). The only peptides of the β-ring that were less dynamic upon PA28αβ binding are located on β1 (Fig. 7c, red circle on β1). We compared the effect of PA28αβ binding to the std20S (Fig. 7a-e) with the other conditions (Fig. 7f-t). Although the destabilization of the α-ring N-ter by PA28αβ was almost complete in the std20S (except α6) (Fig. 7a, large red circle), it was restricted to α2/3/7 in the i20S (Fig. 7k, large red circle), suggesting a partial opening of the pore. Binding of PA28γ did not affect any of the std20S α subunit N-ter (Fig. 7f) and only the i20S α2 and α7 N-ter (Fig. 7p, large red circle), indicating less PA28γ-driven opening of the std20S than the i20S pore.

Overall, PA28 binding to the std/i20S increased their RDU, especially at the pore entrance, at the α1/α2 interface (Fig. 7a,k,p, red circles on α1, α2), and the β2/β2i C-ter arm extending towards β3 (Fig. 7d, e, n, o, s, t). Despite these similarities, our data show subtle proteasome-as well as regulator-specificdifferences in the allosteric motion triggered upon complex formation, as suggested recently 25 . The std20S α3 C-ter was protected upon binding of PA28αβ and PA28γ (Fig. 7a,f). This seemed to induce the PA28γ-specific destabilization of α2 , sum of difference = 13%) and β7 (152-167) (Fig. 7i, red circles on β1, β6, β7; see Supplementary Data 8 for the corrected P values and RDU differences corresponding to the protein portions covered with multiple peptides). According to our statistical thresholds, the α3 C-ter protection upon PA28 binding was std20S-specific (Fig. 7k,p), but the remodeling of the α-ring occurred in both 20S (although differently) and was propagated to the β-ring of the i20S via α1/β3/β5i and α4/β3/β5i bulges for PA28αβ (Fig. 7l, m, red circles on α1/β3/β5i) and PA28γ, respectively (Fig. 7q, r, red circles on α4/β3/β5i). It resulted in remodeling of the β-ring/βring interface eventually affecting β1i/2i/7 or β1i/2i/5i/6/7 with PA28αβ (Fig. 7n, red circles on β1i/2i/7) and PA28γ (Fig. 7s, red circles on β1i/2i/5i/6/7), respectively. The limited sequence coverage around the catalytic sites prevented us from drawing exhaustive conclusions on the differential impact of regulator/20S complex formation on the proteasome active sites. However, β1 (10-25) (corrected P value = 9.49.10 -9 , sum of difference = 22%), β2 (16-42) (corrected P value = 9.54.10 -5 , sum of difference = 5.2%) and β5 (29-36) (corrected P value = 8.84.10 -3 , sum of difference = 8%) active sites were partially but significantly more dynamic/accessible upon PA28αβ binding (Fig. 7d, orange circles and Supplementary Data 3 and 14), in-line with an increased proteolytic activity. Binding of PA28αβ also increased the solvent accessibility of β1i active site (10-17) (corrected P value = 3.89.10 -3 , sum of difference = 14%) (Supplementary Data 3 and 14), which is in good agreement with previous work showing that the caspase-like activity of the i20S is enhanced to a higher extent by PA28αβ than PA28γ 53 . PA28γ was reported to have little or no effect on the i20S 44 , whereas it is known to increase all three std20S activities 54 . Our data do not indicate any major change in the std20S active site solvent accessibility upon interaction with the regulator (Fig. 7i). However, it revealed a significant decrease of β2i N-ter (2-6) (corrected P value = 3.80.10 -3 , sum of difference = -13%) solvent accessibility upon PA28γ interaction (Supplementary Data 3). Thus, the catalytic activation observed upon PA28γ/std20S binding may be driven by changes occurring further away from the active sites.

Discussion

This structural dataset on the human i20S compared to the std20S alone or bound to both PA28 regulators rationalizes from a mechanistic point of view previous observations that replacement 49 , modification 34 or ligand binding 49,55 to the catalytic subunits can allosterically modify the 20S core particle structure and alter its binding to potential PIPs. A main advantage of HDX-MS is that it provides information on very flexible loops, unlike X-ray crystallography or cryo-EM. Also, it is not theoretically limited by the complex size since the proteins are digested into peptides after deuteration: it can be applied to very small proteins (unlike cryo-EM) or very large complexes (unlike NMR). Large oligomeric complexes containing a reduced number of different monomers (limited number of chemical shifts) have already been analyzed by NMR, such as the prokaryotic 20S from T. acidophilum constituted of 14 identical α and 14 identical β subunits 34 . With the recent exception of the DNA-PKc analysis 56 (469 kDa monomer), HDX-MS is usually performed either on rather small and simple systems 39 or on homo-oligomeric complexes 38,57 , due to some technical limitations in protein digestion, peptide separation and data analysis. In comparison, our study encompasses 16 different ~25 kDa monomers. Despite this analytical challenge, we obtained an average sequence coverage per subunit of 89% for the 20S alone and 81% when in complex with PA28 regulators. It is important to note that, except for a few specific cases 58,59 , HDX-MS does not inform on the presence of multiple conformational states in solution since it is based on the measured average of their RDUs. Other approaches such as cryo-EM-based classification would be needed to better characterise intermediate or minor states. Nevertheless, comparing the same sample (like std20S) in presence or absence of regulator and performing thorough statistical analysis identified a wealth of information on the proteasome dynamics, binding interfaces and allosteric changes upon incorporation of the immuno-subunits or activation by PA28 regulators (Fig. 8).

These conformational maps revealed the subtle rearrangement of the std/i20S α3 C-ter. More precisely, the residues 238-261 were less dynamic/accessible upon incorporation of the immunocatalytic subunits (in blue Fig. 8a). The principal component analysis of the 20S high-resolution structures available in the Protein Data Bank (PDB) in 2014 (46 structures from yeast, 4 murine and 1 bovine) revealed that the 220-230 stretch of the mouse std20S and i20S α3 clustered with the yeast apo and peptide-bound forms, respectively 49 . We believe that the allosteric change occurring on α3 C-ter upon ligand binding to β5 in the yeast 20S could be similar to the change in solvent accessibility/ dynamics observed between the std20S and the i20S conformational maps. The highly charged C-ter of α3 points towards the outer surface of the α-ring and has also been suggested to be the Insulin-Degrading-Enzyme binding site 60 . It could be involved in the binding of PA28αβ/γ to the std20S, or in its subsequent activation. Interestingly, the very end of α3 C-ter is absent from most human 20S proteasome structures, which confirms its high flexibility but also impedes any modeling of 20S/PA28 interaction. Unfortunately, this protruding C-ter α helix is not present in the cryo-EM structure of the Pf 20S bound to its PA28 regulator, since it is nine residues shorter than its human counterpart 36 . It could be characteristic of more evolved 20S proteasomes and tune their binding with different regulators.

As commented before, the differential analysis of the catalytic subunits is impossible due to their sequence differences (the peptides resulting from the pepsin digestion are different). The comparison of their deuteration profiles was further hindered by the limited sequence coverage of the i20S. This was partly due to the presence of residual traces of std20S present in this sample 61 . Nevertheless, it was possible to compare the RDUs of the noncatalytic subunits of the two core particles. The central pore of the 20S was more flexible in the i20S vs. std20S (Fig. 4d and in red Fig. 8a), as was the β-facing interface of the α-ring, especially α5 that directly faces the β5 (std20S) or β5i (i20S) catalytic subunit (Fig. 4c). We propose that the conformational differences observed between the standard and immuno-α3 C-ter (red arrows Fig. 8a) could be transduced from the inner β-ring to the outer αring via α5.

Interestingly, upon binding of any regulator to the 20S, we observed a similar destabilization of the α5 region that faces β5/5i, suggesting that the exact opposite mechanism (from the outer αring to the inner β-ring) can also take place upon 20S/PA28 interaction, as suggested earlier 34 .

Overall, our data indicate a general mechanism of PA28 activation by the 20S core proteasome. More precisely, the interaction of PA28 activation loops and C-ter with the 20S seems to trigger a long-range allosteric change at the top of the regulator (loops between helices 1/2 and/or 3/4) via a rearrangement of the N-ter kink of PA28α and PA28γ (Fig. 8b). Although present in the three PA28 subtypes, the activation loop protection was stronger in PA28α and PA28γ vs. PA28β; most probably explaining why "the affinity of the proteasome towards the PA28αβ complex is about two orders of magnitude higher than towards the homomeric PA28α and PA28β complexes" 62 . Indeed, several studies showed that both PA28α and PA28β were required for an optimum activation of the 20S by PA28αβ 32,33 .

We also provide crucial information on the loops located between helices 1/2, missing in every PA28 high-resolution structure or model. Since these are located at the entrance/exit of the activator, any change in their flexibility could modify the nature of both substrates and/or products of the proteasome complex. Indeed, it has been shown in PA28α to constrain substrates within the catalytic chamber of the core particle, thereby generating shorter peptides 63 . We show that these flexible loops, together with PA28 constriction sites (conserved lysine between alpha helices 3 and 4) are allosterically stabilized upon binding of the std20S but not the i20S (Fig. 8c). This could explain how the binding of different PA28 to different 20S subtypes alters the length/nature of the generated peptides. Our data also suggest that PA28 might modify the 20S products by inducing conformational changes in the proteasomal active sites. Indeed, our results show distinctive changes in dynamics detected down to the β-ring and around the catalytic site and/or substrate pockets upon regulator binding (Fig. 8c).

We observed a stronger protection of PA28αβ than PA28γ upon std20S binding (Fig. 6 and Fig. 8c). Globally, both PA28 were more protected upon interaction with the std20S than with the i20S (Fig. 8c and Supplementary Data 2 and 13), suggesting a stronger interaction of the regulators with the std20S than the i20S (Supplementary Data 2). This was mirrored by regions on the i20S α-ring that were less protected than in the std20S upon incubation with both PA28, whereas previous studies showed that PA28αβ interaction was stronger with the i20S than with the std20S 64 or equivalent 21 . In this context, we think that protection to deuteration cannot be always correlated to relative binding

7. 1 . 1 Figure 1 .

 111 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 2 .Figure 3 .

 23 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 4 .

 4 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 5 .

 5 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 6 .

 6 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 7 .

 7 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 8 .

 8 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 9 .Figure 10 .Figure 11 .Figure 12 .Figure 13 .

 910111213 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 14 .Figure 15 .Figure 16 .Figure 17 .Figure 18 .Figure 19 .

 141516171819 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 20 .Figure 21 .Figure 22 .Figure 23 .

 20212223 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 24 .Figure 25 .

 2425 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 26 .Figure 27 .Figure 28 .Figure 29 .Figure 30 .Figure 31 .Figure 32 .

 26272829303132 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 33 .Figure 34 .

 3334 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 35 .

 35 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 36 .Figure 37 .

 3637 Figure 1. Morphology of the human testis. .............................................................................Figure 2. Germ cell differentiation overview. .......................................................................... Figure 3. Model of the recombination process. .....................................................................Figure 4. Chronological order of events at the synaptonemal complex. ................................Figure 5. Stages in meiotic division. .........................................................................................Figure 6. Spermiogenesis and spermiation. ............................................................................ Figure 7. Structure of the eukaryotic 20S proteasome. ..........................................................Figure 8. Regulation of the 20S proteasome activity and function. .......................................Figure 9. The assembly of the 20S proteasome. ..................................................................... Figure 10. Schematic representation of the composition of tissue-specific proteasomes. ....Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. ............. Figure 12. The ATP-independent proteasome regulators. ...................................................... Figure 13. The UPS. Proteins are targeted for degradation via hierarchically organised system of Ub-ylation enzymes. .........................................................................................................................Figure 14. Diversity of Ub and Ub-like signals. ........................................................................ Figure 15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. ............. Figure 16. The spermatoproteasome sequence and model. ................................................... Figure 17. Schematic representation of an MS system. .......................................................... Figure 18. ESI formation of suspended ions. ........................................................................... Figure 19. Schematic representations of different analysers. ................................................. Figure 20. A: The types of ions that can originate from peptide fragmentation .................... Figure 21. The schematic comparison of DDA and DIA modes of operation. ......................... Figure 22. Schematic overview of the Synapt G2Si from Waters. ...........................................Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid from Thermo Fisher Scientific. ....................................................................................................................................Figure 24. The Bottom-Up proteomics workflow. ................................................................... Figure 25. Label-Free Quantification using Bottom-Up proteomics. ......................................Figure 26. Different ways to study an interactome through BU proteomics. ......................... Figure 27. The overview of different structural MS techniques. ............................................. Figure 28. The workflow of the Bottom-Up HDX-MS experiment. .......................................... Figure 29. Principles of mass photometry. .............................................................................. Figure 30. Diagram of the study undertaken in the article manuscript. ................................. Figure 31. Mass photometry histograms: The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. ...........................................................................Figure 32. Plot of regulator/20S ratio versus the share (in %) of a particular complex in the total 20S population. ..........................................................................................................................Figure 33. Native mass spectra of the PA200:20S complex formation. ............................... Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4s-coding gene (PSMA8), with GFP fluorescence as a marker. ...................................Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4transfected cultures. ..........................................................................................................................Figure 36. Visual regrouping of the GO term enrichment results. ....................................... Figure 37. Volcano plot of α4s versus α4-expressing cell line. ..............................................

Figure 1 .

 1 Figure1. Morphology of the human testis. A: Cross-section of the human testes. The organ is divided in lobules, each containing highly coiled seminiferous tubules, eluting its secretions from both ends into the rete testis, which transfers the differentiated germ cells into the efferent ducts of the epididymis. As they travel through the epididymis, the germ cells reach full maturation and are stored in the epididymis. B: Cross-section micrograph of a histological preparation of a seminiferous tubule. The pool stem cells called spermatogonia renews itself by division, while a part of the population begins to differentiate. As they do so, germ cells move towards the middle of the seminiferous tubules, until they are released into the lumen. Throughout the differentiation process, germ cells are immersed in invaginations on the surface of Sertoli cells that serve as support and quality control.

Figure 2 .

 2 Figure 2. Germ cell differentiation overview. A: Before birth, the primordial gonial cells migrate to the gonads, where they become gonocytes. During foetal development, gonocytes settle in the basal layer of the seminiferous tubules, thus becoming spermatogonia. The type A spermatogonia divide and a subpopulation of them enter the differentiation pathway by becoming type B spermatogonia. They further divide mitotically and upon entering spermatocyte stage, they initiate meiosis. After meiosis, the cells enter the spermatid stage, in which they contain recombined genetic material and reduced number of chromosomes. The polarisation of the cells follows: spermatids form a head and a tail, discard excess cytoplasmic material and become fully separated.Upon maturation, they are released by the Sertoli cells into the lumen. The letter markings C and N stand for number of copies and ploidy, respectively. B: Spermatogenesis is initiated in waves. Three neighbouring waves of differentiation are given as an example (marked as I, II and III), with stages of differentiation marked on the ribbons: L-leptonema, Z-zygonema, P-pachynema. As the differentiation progresses, cells migrate along the seminiferous tubule and towards the lumen (see lower diagram representing the migration of the neighbouring waves inside of a seminiferous tubule). In a cross-section we would be able to distinguish neighbouring waves and different stages -later ones towards the middle and the earlier ones towards the peritubular tissue. Adapted from A: (Bolcun-Filas and Handel, 2018); B:[START_REF] Hirsh | The anatomical preparations of the human testis and epididymis in the Glasgow Hunterian Anatomical Collection[END_REF].

Figure 3 .

 3 Figure 3. Model of the recombination process. The recombination is initiated by introduction of Double Strand Breaks (DSBs) of one of the two homologous chromosomes. Single thread of cleaved DNA is inserted into the homologous DNA ladder that unwinds to accommodate the invading strand. This complex may be stabilized or not and therefore a non-crossover (NCO) or a crossover (CO) scenario can occur. In a crossover scenario, single strands of DNA are exchanged between the homologs forming double Holliday junction. Upon resolving the junction by nickase and ligase activity, two novel recombined double DNA strands emerge. Adapted from (Marston and Amon, 2004).

Figure 4 .

 4 Figure 4. Chronological order of events at the synaptonemal complex. Cohesin core maintains the integrity of the chromosomes and keeps sister chromatids together (left scheme). In leptonema, the axial elements bind to the cohesin and protruding DNA loops bind protein complexes that induce DSBs. In zygonema, the pairing of the homologs continues by the formation of the first central elements, and the first recombination nodules are formed. The left schema represents the way SC holds the homologs together. In pachynema, the pairing of the homologs is complete and the recombination events are brought to the end. The sex body contains X and Y chromosomes (yellow) that only partially participate in the recombination. In diplonema, the SCs are dissolved and the homologs are connected only by the chiasmata. What follows is the separation of recombined homologs. In the legend, key markers and the regulators of the events are listed, as well as the constituents of the SC. Adapted from (Bolcun-Filas and Handel, 2018).

Figure 5 .

 5 Figure 5.Stages in meiotic division. On the left, microscopy of the distinct stages of prophase I are displayed in line with the scheme of the molecular events at the chromatin level. The copies of DNA are duplicated in the S phase giving sister chromatids. Homolog pairing begins in leptonema. The synapsis forms in zygonema and is fully achieved in pachynema. Crossover begins in zygonema but is fully complete in pachynema. In diplonema, synapses disappear and the only thing holding the homologs together are chiasmata -the strands of DNA marking the spot of the crossover. The meiotic spindle now forms and the recombined homologs are separated into two cells. Another division separates the recombined sister chromatids into haploid cells. The theoretical outcome of the process is four haploid spermatids from one primary spermatocyte. Adapted from(Hunter, 2015).

Figure 8 .

 8 Figure 8. Regulation of the 20S proteasome activity and function. In order to properly maintain homeostasis, the proteasome is regulated on multiple levels: 1) new 20S complexes can be syntesized by controlling the gene expression and 2) PTMs can be introduced to modulate 20S activity or even mark it for degradation; 3) alternative, tissue-specific subunits exist, which upon incorporation modulate the peptide product profile or 4) the interaction with proteasome-associating proteins. Many regulatory particles can bind to the 20S, altering its substrate specificity, the prevalent being 19S, which makes up the 26S complex together with the proteasome core. Proteasome-dependant proteolysis is further regulated by substrate-interacting proteins, which can help select or shuttle targets for degradation or proteasome-interacting proteins, which can inhibit or activate an existing proteasome complex. Finally, the proteasome can be translocated directly to needed location, within or without the cell. Adapted from (Ben-Nissan and Sharon, 2014).

Figure 9 .

 9 Figure 9. The assembly of the 20S proteasome. Two heterodimers, PAC3/4 and PAC1/2 aid the oligomerisation of the α ring. Ump1/POMP aids in recruitment of the β-subunits to the α ring, which serves as a scaffold. Upon assembly of two hemiproteasomes, the propeptides of the β-subunits are cleaved and the remaining assembly chaperones are digested to give a mature 20S proteasome unit. Adapted from (Mao, 2021).

Figure 10 .

 10 Figure 10. Schematic representation of the composition of tissue-specific proteasomes. In the immunoproteasome, one or several catalytic subunits are replaced by an isoform typical for (but not exclusive to) the cells of the immune system and plays a role in antigen presentation. The thymoproteasome exists only in the epithelial cells of the thymic cortex and plays a role in T-cell education. In the spermatoproteasome, the α4 subunit is replaced by the gamete-specific α4s. This subunit proved to be vital for germ cell development (Gómez-H et al., 2019; Z.-H. Zhang et al., 2019).

Figure 11 .

 11 Figure 11. 26S proteasome structure and scheme of the base ring ATPase activity. Left: the 26S proteasome consists of the 20S proteasome catalytic core and 19S regulator, which in turn consists of the base (red) and the lid (yellow and orange). The lid is in charge of substrate recognition via the Ub-and UBL-recognising subunits (in orange) and the removal of the Ub tag. Centre: the base of the 19S couples ATP hydrolysis with unfolding of the substrate. Three protruding N-terminal helix bundles facilitate lid subunit binding. Right: the substrate is fed into the 20S via coordinated action of the six ATPase subunits. ADP + P release provides energy for the hinge motion that moves the polypeptide chain. Motion is coordinated among subunits to drive the polypeptide chain through the 19S base pore. Left and centre panel made in UCSF ChimeraX, using 5MPB PDB entry. Right panel adapted from (Dong et al., 2019).

Figure 14 .

 14 Figure 14. Diversity of Ub and Ub-like signals. The Ub signal sends a different message depending on the length, branchedness, type of linkage or PTMs. The type of linkage between the Ub moieties determines the structural properties of the chain and its topology (e.g. K48 linking is the most compact, while the K63-linked chains are looser). As a consequence, different chain types are recognized by different enzymes or by the same enzyme with different affinities. The K48 and K11 are most common signals for proteasomal degradation. Other types of linkages have a role in cell signalling, enzyme activity regulation or in other degradation pathways (lysosomal degradation through K63-linked chains; (Mukhopadhyay and Riezman, 2007)). Other Ub-like proteins also take part in the signalling, such as NEDD-8 or the SUMO family (Cappadocia and Lima, 2018).

Figure 15 .

 15 Figure15. PI31 and Fbxo7 are partners that seem to regulate proteasome activity. A: domain organisation of PI31 and Fbxo7. The two proteins dimerize via their FP domain, while the proline rich domain in PI31 binds to the 20S proteasome. Fbxo7 also contains domains that enable it to bind to other proteins: the F box domain enables it to participate in an E3 ligase complex, cdk domain which interacts with cell cycle regulators and a Ub-like domain. Adapted from[START_REF] Shang | The FP domains of PI31 and Fbxo7 have the same protein fold but very different modes of protein-protein interaction[END_REF]. B: Proposed model of PI31/Fbxo7mediated proteasome regulation. PI31 acts as a linker between the 26S proteasome and Fbxo7, which is a part of an E3 ligase. Together, they may make up a system for degradation of proteins crucial for the progression of spermatogenesis. Adapted from[START_REF] Bader | A conserved F box regulatory complex controls proteasome activity in Drosophila[END_REF].

Figure 17 .

 17 Figure 17. Schematic representation of an MS system. The analytes can be introduced to the mass spectrometer in various ways. Ionization enables the analytes to be manipulated by the electric or magnetic fields of the mass analyser and later to be detected. Signal from the detector is then processed by a computer into a mass spectrum -representation of mass-to-charge versus intensity of the ions.

Figure 18 .

 18 Figure 18. ESI formation of suspended ions. When voltage potential is applied between the capillary that carries solubilized analyte and an electrode, the ions will accumulate at the meniscus, which results in the formation of the Taylor cone and the emission of droplets. Upon continued evaporation of the solvent, the droplets continue to fission due to the repelling forces of the ions. Result of the repeated fissions are chargecarrying analytes that are completely or partially desolvated. Adapted from (Kebarle and Verkerk, 2009).

Figure 19 .

 19 Figure 19. Schematic representations of different analysers. A: In a TOF analyser, the m/z values are related to the time needed for ions to travel a certain distance. B: In Q analysers, the RF field maintains the ions on a stable trajectory through the cavity along the electrodes. Applying offset DC voltage can destabilize the trajectories of ions of certain m/z, effectively creating a mass filter. C: Cross-section of a ring IT: the ions are maintained in constant motion via one ring-shaped electrode, capped by two hyperbolic electrodes.Ions can be filtered through a similar mechanism as in quadrupoles, collided with a gas to induce fragmentation and their fragments can be further analysed in the same IT. D: Orbitraps confine the ions in a complex rotating motion around the axial electrode. The ions induce current in the outer electrode, which is recoded as signal. Complex waveform of the signal is deconvoluted through mathematical transformation into mass spectra. E. ICR uses strong magnetic field to confine ions in a circular motion. Electrical RF field is used to add energy to the system. Moving ions induce current in the nearby plates, which is recorded as signal. ICR, like the Orbitraps, are part of the FT-family of mass analysers, meaning that the signal is decomplexified through Fourier transformations in order to obtain information on the m/z of the ions. Adapted from[START_REF] Haag | Mass Analyzers and Mass Spectrometers[END_REF].

Figure 20 .

 20 Figure20. A: The types of ions that can originate from peptide fragmentation[START_REF] Hao | Current Methods for the Characterization of Posttranslational Modifications in Therapeutic Proteins Using Orbitrap Mass Spectrometry[END_REF]. B: Theoretical MS 2 spectrum of b-and y-ion series of a peptide Ser-Phe-Ala-Glu, including the loss of H2O and NH3. The precursor ions are often found in the MS 2 spectra, along with the fragments. Here, the precursor ion with the formula MH + can be found on the far right of the spectra. Mass difference between the precursor and different fragment ions correspond to the mass of the sequentially lost amino acid residues. This permits to reconstruct the sequence of the peptide. Adapted from (Baars and Perlman, 2016).

Figure 21 .

 21 Figure 21. The schematic comparison of DDA and DIA modes of operation. On the horizontal axis is the m/z range to be scanned, while the y axis represents the retention time. Horizontal bars represent the cycle time. A: Individual points represent MS 2 scans. Ideally, in a single MS 2 event, the isotopes of individual peptide are selected with a narrow 2 Th range (lower schema). B: The small bars represent increments of m/z range selected for MS2. Several peptides are selected for fragmentation at the same time (lower schema). Adapted from (Hu, Noble and Wolf-Yadlin, 2016).

Figure 22 .

 22 Figure 22. Schematic overview of the Synapt G2Si from Waters. The instrument has a Q-TOF configuration, with an IM cell between the two mass analysers, which can serve as a collision chamber (Image source -Waters).

Figure 23 .

 23 Figure 23. Schematic geometry representation of the Orbitrap Fusion Tribrid, from Thermo Fisher Scientific. (Image source-Planet Orbitrap)

Figure 24 .

 24 Figure 24. The Bottom-Up proteomics workflow. The affinity purification (AP) of the protein complexes is an option which enables the interactome study (see Chapter 2.3.3.4).

Figure 25 .

 25 Figure 25. Label-Free Quantification using Bottom-Up proteomics. The peptides are identified from the MS 2 spectra, using a database of candidates and a set of user-set constraints. The simplest possible set of proteins is inferred from the peptide list. Then the proteins are quantified based on the intensities of their peptides. Adapted from (10.1021/cr3003533).

Figure 26 .

 26 Figure 26. Different ways to study an interactome through BU proteomics. Left to right: affinity purification, either through an affinity tag or based on an antibody-antigen interaction; correlation of the abundance/elution profile of proteins from native chromatography/gel/density gradient can indicate they belong to the same complexes; BioID -expression of a chimera carrying a biotin ligase will tag any nearby interactor, which can then be affinity enriched and identified by MS, or identified by MS directly.

Figure 27 .

 27 Figure 27. The overview of different structural MS techniques. The diagrams represent basic mode of operation for each approach, with a short description of the information each technique provides. Adapted from (Allison and Bechara, 2019).

Figure 28 .

 28 Figure 28.The workflow of the Bottom-Up HDX-MS experiment. The native protein is diluted in D2O and after certain time of deuteration, the reaction is quenched and the protein is digested into peptides. Exposed or flexible peptides will manifest a positive shift in mass due deuteration, which can be presented in a form of a heatmap or visualized onto the 3D structure of a protein or a protein complex.

Figure 29 .

 29 Figure 29. Principles of mass photometry. A: Adherence of large molecules from solution to the glass surface, results in the interference of the scattered light. The intensity of the interference contrast is directly proportional to the mass of the molecule. The instrument records multiple adhering events over time (i, i+1, i+2) and sums them up in the form of a histogram (panel B), which represents discrete number of particles for a defined increment of contrast values. On the panel B, the example is given for analysis of a mixture containing three species, whereby the biggest is the most abundant one (far right). The contrast can be translated to mass using calibration standards of known masses. Figure adapted from (Sonn-Segev et al., 2020).

  (Bousquet-Dubouch et al., 2011; Khilji et al., 2020)? In combination with affinity purification techniques, it allows for the study of the proteasome interactome, and thus the identification (and relative quantification) of the proteasomebound activators (Fabre, Lambour, Garrigues, et al., 2014; Abi Habib et al., 2020).

Figure 30 .

 30 Figure 30. Diagram of the study undertaken in the article manuscript. The study was split in two parts -identifying the spermatoproteasome interactors and the study of the structural versus functional implications of the α4s incorporation into the proteasome. In blue squares are the techniques employed for the study and in orange are the questions they aim at responding to. Several types of material were used for the experiments: whole organs, purified germ cells or purified protein complexes. (spt20S -spermatoproteasome; std20Sstandard proteasome).

Figure 1 .

 1 Figure 1. (A) 3D proteoform footprint of the LC-MS trace of 20S subunits immunopurified from bovine testis, generated with VisioProt-MS (28). The names of the c20S subunits are indicated next to the corresponding signal. (B) Close-up view of the 3D proteoform footprint of 20S subunits immunopurified from bovine testes after repeated freeze-thaw cycles, resulting in truncated proteoforms of α4 and α4s. (C) The truncated α4 and α4s to the fulllength proteins ratio increase with repeated freeze-thaw cycles. (D) Top-down LC-MS/MS of α4 identifies the degradation as the loss of the last two amino-acids.

Figure 2 .

 2 Figure 2. (A) Analysis of proteasome composition and interacting proteins in bovine testes. Distribution of enriched proteins between two IPs against anti-α2 (left hand side) and anti-α4s (right hand side). Only proteins that were enriched (FC>2, p-value <0.05) in the anti-ɑ2 or anti-ɑ4s IPs compared to the control IPs are displayed here. For clarity, gene names are used as protein identifiers. 20S non-catalytic and 19S subunits are circled in red and blue, respectively. (B) Estimated percentages of α4 and α4s in the different IPs. The protein intensities were normalized based on the 20S content and the relative abundances of α4 and α4s estimated for the different IPs. (C) FCs of 20S-associated regulators in the s20S (α4scontaining proteasome) relative to α4-containing proteasomes. The relative quantities of α4 and α4s in the anti-α4s and anti-α2 IPs enables to estimate the fraction of 20S-associated regulator in the α4s-and α4-containing proteasomes. Finally, FCs in α4s-vs. α4-containing proteasomes were calculated, as detailed in the experimental section. Graphics represent mean and standard deviations. All values are the means of three independent experiments. Stars indicate significance. * : p<0.05; **: p<0.01; ***: p<0.001.

Figure 3 .

 3 Figure 3. Composition and proteolytic activity of proteasome complexes purified from rat SER, SPG, SPC and SPT. Cells obtained from rat testes were separated and then analyzed using label free proteomics. (A-B) Proportions of α4 and α4s proteins in (A) immunopurified proteasome complexes and (B) cell lysates. (C) The distribution of the different proteasome complexes was estimated for each cell type by label-free MS (27). We considered the free 20S complexes and the ones associated with 19S, PA28αβ, PA28γ, and PA200 regulators. Error bars indicate standard deviation across three biological replicates. (D) Major changes in the composition of proteasome complexes throughout spermatogenesis. The variations of major 20S-associated regulators were measured throughout spermatogenesis using label-free quantitative proteomics. The SPG stage was used as a reference and stars indicate significance between SPC vs. SPG or SPT vs. SPG. (E) Changes in the amount of total 20S proteasomes throughout spermatogenesis. The variations of 20S proteasome subunits (all non-catalytic subunits except α4 and α4s) were measured throughout spermatogenesis using label-free quantitative proteomics and normalized with the MS signal of total protein. Stars indicate significance between SPG vs. SER or SPC vs. SER or SPT vs. SER. (F) Proteasome chymotrypsinlike (ChT), trypsin-like and caspase-like specific activities, expressed in pmol AMC released per minute and per mg of total protein, have been measured in the four cell types lysates throughout spermatogenesis. Stars indicate significance between SPC vs. SPG or SPT vs. SPG. Graphics represent mean and standard deviations. All values are the means of three independent experiments. * : p<0.05; **: p<0.01; ***: p<0.001.

Figure 4 .

 4 Figure 4. In vitro assays. Bovine c20S or testes 20S pool were incubated with (A) PI31, (B) the 19S or (C) PA200, and complexes were purified with sepharose beads grafted with their corresponding antibodies. After washes and elution, α4 and α4s subunits were identified and quantified by LC MS/MS label-free quantification and their abundances in anti-PI31 IP, anti-PSMC2 IP, and anti-PA200 IP were normalized with their abundances in the corresponding control IPs. Anti-α2 subunit control IP (for total 20S) was performed on the same reaction mix for each experiment. Graphics represent mean and standard deviations. All values are the means of three independent experiments. Stars indicate significance * : p<0.05; **: p<0.01; ***: p<0.001. The vertical axis represents the intensity of a protein in a pull-down experiment relative to its intensity in a control IP (in the case of 19S -average intensity of a set of 19S base proteins), when normalized for 20S abundance. (D) Comparison of three peptidase activities in proteasomes purified from bovine testis (s20S) or muscle (c20S). Activation of the proteasomes by PA200 was assayed at a PA200:20S molar ratio of 8:1. The bars are grouped by the peptide used to probe each activity: boc-LRR-AMC, suc-LLVY-

Figure 5 .

 5 Figure 5. HDX-MS of the s20S vs. c20S. (A) Heatmap showing the relative deuterium uptake from 0% (blue) to 60% (red) of α4 and α4s at 0.5 min, 1 min, 5 min, 10 min and 30 min. (B) Cartoon representation of a homology model of α4s (left) and the structure of α4 (right) within the 20S bovine proteasome (PDB: 1IRU), viewed from the top of the α-ring. Residues of α4s and α4 are color coded with the relative deuterium uptake at 15 min. (C) Kinetics of

  x=(A(ɑ2)*d -A(ɑ4s)*b)/(a*d -b*c) y=(A(ɑ2)*c -A(ɑ4s)*a)/(b*c -a*d) if starting from statements: A(ɑ2)=a*x+b*y and A(ɑ4s)=c*x+d*y for a,b,c,d being the percentages in 0 to 1 format (e.g. 0.15 for 15%), where x is the intensity of a protein P bound to ɑ4-containing P20S and y is the intensity of a protein P bound to ɑ4scontaining P20S. A(ɑ2) is the experimentally measured intensity of the protein P in the anti-ɑ2 CoIP, while A(ɑ4s) is the experimentally measured intensity of the protein P in the anti-ɑ4s CoIP. Additionally: since a=1-b and c=1-d then: x=(b*A(ɑ4s) -d*A(ɑ2))/(b-d) y=((1-d)*A(ɑ2) + (b-1)*A(ɑ4s))/(b-d) (conditions: b≠d, bd≠d) The estimations are easily derived when the two formulae are solved together, once the intensity values of a protein P for each CoIP are plugged in, together with the ɑ4s percentage in each CoIP (b and d factors, 42% and 96% for anti-ɑ2 and anti-ɑ4s IP, respectively).

Figure S2 .

 S2 Figure S2. The Top-Down analysis of 20S proteasome from bovine testes highlighted errors in the Swiss-Prot entries of (A) β6 (Q2TBX6) and (C) β2 (Q2TBPO), whereas the TrEMBL counterparts of (B) β6 (G5E589: Arg233Glu) and β2 (D) F1MBI1 Asn252Thr) provided more fragments and higher identification scores.

Figure

  Figure S3. (A) Sequence alignment of bovine PSMA7 and PSMA8 proteins showing the Cterminal sequence chosen for the immunization strategy (Peptide 1 : ILNPEEIEKYVAEIEKEKEENEKKKQKKAS and Peptide 2 : MFSAKEIELQVNEIEKEKEEAEKKKSKKTT) (B) ELISA assay (plates were coated with peptide 1 (corresponding to the 219-248 C-term part of the α4 protein) or peptide 2 (corresponding to the 221-250 C-term part of the α4s protein)and the different concentration of polyclonal antibody produced against peptide 2 were used for the readout. This resulted in EC50 of 0.005 µg/mL and 2.12 µg/mL for coating peptides 2 and 1, respectively. (C) Western-blot showing the specificity of the antibody produced against bovine α4s. c20S proteasome were purified from bovine muscle tissue and s20S (mixed with c20S) was purified from bovine testes. The anti-α2 antibody was obtained from the MCP21 hybridoma.

Figure S4 .

 S4 Figure S4. Anti-α2 immunopurification (A) Volcano plot volcano presenting the statistical significance distribution against the log2-transformed fold change between anti-α2 IP and unrelated control OX8 IP. (B) Close-up view of proteins that were significantly enriched (FC>2, p-value <0.05) in anti-ɑ2 vs. OX8 IP. For the sake of clarity, gene names are used to represent the proteins.

Figure S5 .

 S5 Figure S5. Anti-α4s immunopurification (A) Volcano plot volcano presenting the statistical significance distribution against the log2-transformed fold change between anti-α4s IP and unrelated control OX8 IP. (B) Close-up view of the most significantly enriched proteins in anti-ɑ4s vs. OX8 control IP. For the sake of clarity, gene names are used to represent the proteins.

Figure S6 .

 S6 Figure S6. Anti-PA200 immunopurification Volcano plot volcano presenting the statistical significance distribution against the log2-transformed fold change between anti-PA200 IP and unrelated control OX8 IP. (B) Close-up view of proteins that were significantly enriched (FC>2, p-value <0.05) in anti-PA200 vs. OX8 IP. For the sake of clarity, gene names are used to represent the proteins.

Figure S7 .

 S7 Figure S7. Volcano plot volcano presenting the statistical significance distribution against the log2-transformed fold change between anti-α4s (left hand side) and anti-PA200 (right hand side).

Figure S8 .

 S8 Figure S8. Label-free MS quantification of protein markers of sertoli cells (SER),

Figure S9 .

 S9 Figure S9.Comparison of protein abundances changes from SPG to SPC stages in the lysate and in the immunopurified proteasome. For each protein or protein complex, the change in quantity from SPG to SPC (= ratio SPC/SPG) was calculated both in the lysate and in the immunopurified proteasome sample. Then, these ratios were normalized with the corresponding ratios in the lysates. Stars indicate significance between the lysate and its corresponding immunopurified proteasome sample (IP).

Figure S10 .

 S10 Figure S10. Sequence coverage obtained upon pepsin digestion. (A) 20 peptides are found in both α4 and α4s. Proteospecific peptides obtained for (B) α4 and (C) α4s.

Figure S11 .

 S11 Figure S11. Molecular dynamics simulations of α4 and α4s. (A) Number of H-bonds observed for the amide protons of residues 184-198 and 225-241 of α4s (orange line) and α4 (blue line) during the 1 µs simulation. (B) Distribution of crossing angles between the two last α-helices of α4s (yellow) and α4 (green). (C) Close-up view of the two last α-helices of α4 (left) and α4s (right). (D) Normalized frequency of H-bonds observed between the side-chain residues of the two last α-helices of α4 (left) and α4s (right).

Figure S12 .

 S12 Figure S12. (A) Normalized frequency of H-bonds observed between the side-chain residues of the two last α-helices of α4 (left) and α4s (right). (B) Distributions of Accessible Surface Areas (Ų) observed for the residues 184-198 and 225-241 α4 (blue) and α4s (red)

Figure 31 .

 31 Figure31. The testis proteasome is mixed in different ratios with the PA200 regulator and allowed to interact. Mass photometry histograms plot the masses of particles in solution versus their count. Here, mass photometry histograms show different complexes that arise as a result of the 20S-PA200 interaction.

Figure 33 .

 33 Figure33. Native mass spectra of the PA200:20S complex formation. On the right are the deconvoluted spectra along with the calculated masses of the detected mass species. A: PA200 spectra B: bovine standard 20S proteasome spectra C: PA00:20S proteasome mix, with a ~900 kDa peak corresponding to the singly-capped proteasome complex.

Figure 34 .

 34 Figure 34. Results of FACS performed on GC-2spd cell line, transfected with either α4-(PSMA7) or α4scoding gene (PSMA8), with GFP fluorescence as a marker. A: Cells were sorted in three different groups, based on the GFP intensity. Different sorting groups are represented by different rectangles. B: The count of cells versus the GFP fluorescence intensity -logarithmic scale. The majority of the cells do not express GFP and only show background fluorescence signal. C: Percentage of sorted cells to total sample cell count (two samples: PSMA7expressing and PSMA8-expressing).

Figure 35 .

 35 Figure 35. Western blot (anti-α4s) of 20S proteasomes CoIP-ed from the α4s-or α4-transfected cultures. We can see the strong signal at ~30 kDa coming from the α4s-transfected samples. The α4-transfected samples also display some amount of signal, probably caused by non-specific binding.

Figure 36 .

 36 Figure 36. Visual regrouping of the GO term enrichment results. The circles represent a single GO-term. The size represents generalness of the term and the colour represents the p value of the each GO term. Regrouping produced in REVIGO engine (Supek et al., 2011) and plotted in R Studio.

Figure 37 .

 37 Figure 37. Volcano plot of α4s versus α4-expressing cell line. Only the proteins that were differentially expressed in either of the cell lines compared to a control cell line are displayed. For clarity, gene names were used to represent proteins. In blue are all the statistically significant differentially expressed proteins ((p value) ≤ 0.005; (fold change) ≥ 2). The orange points represent a subgroup of significantly expressed proteins involved in chromatin re-organisation.

Fig 1 .

 1 Fig 1. Constitutive expression of proteasome inducible subunits in islets and cell lines. (A-B) Single cell RNA sequencing analysis of β1i, β2i and β5i gene expression in human pancreatic islet alpha, beta and delta cells from healthy individuals (n = 6). The data is shown as means with SEM. (B) presents the percentage of cells with detectable levels of inducible subunit mRNA. (C-D) SDS-PAGE and Western blot analysis of basal expression of proteasome inducible subunits in immune cell lines A20 and Jurkat, in insulinoma beta cell line INS-1E, and human islets (H-islets) and mouse islets (M-islets). Values on top of the Western blots show the amount of protein loaded. C and D are representative blots of n = 3. (E) Quantification of relative expression levels of inducible proteasome subunits normalized to tubulin (C) or actin (D) in tested cell lines/islets (n = 3, biological replicates). The data is shown as means with SD. https://doi.org/10.1371/journal.pone.0222432.g001

Fig 2 .Fig 3 .

 23 Fig 2. Identification of intermediate proteasomes in INS-1E cell line. Four x10 8 cells were cross-linked, lysed, their proteasomes immunoprecipitated with mAb MCP21 and analyzed by LC-MS/MS. The absolute quantities of each of the six catalytic subunits measured by the LC-MS/MS method were computed to calculate the stoichiometry of 20S proteasome subtypes and the fractions of regulatory particles associated with the 20S core particle, as detailed in Experimental Procedures. INS-1E cells were cultured at standard conditions and four biological replicates were analyzed. https://doi.org/10.1371/journal.pone.0222432.g002

Fig 4 .

 4 Fig 4. Cytokines induce upregulation of inducible proteasome subunits in islets/cells. Human (A, n = 4) and mouse islets (B, n = 3) and INS-1E cells (C, n = 4) were exposed for 24 h to IL-1β at low (50 pg/ml for mouse islets and 15 pg/ml for INS-1E) or high dose (300 pg/ml for human/mouse islets and 150 pg/ml for INS-1E) or IFN-γ (10 ng/ml). Islets/cells were lysed and protein content analyzed by SDS-PAGE and Western blotting. Representative blots of four independent experiments (biological replicates) are shown (left) and quantification of inducible subunit bands relative to the tubulin (A and C) or actin (B) is presented (right). Statistical analysis was performed by paired t-tests of treatments versus control. Experiments done on individual islet donors (A and B) or biological cell replicates (C) are connected by lines. https://doi.org/10.1371/journal.pone.0222432.g004

Fig 5 .

 5 Fig 5. Basal and cytokine induced activity of proteasome subunits in (A) INS-1E, (B) mouse and (C) human islets (all n = 3). Cells and islets were exposed to low IL-1β dose (30 pg/ml for human islets, 50 pg/ml for mouse islets and 15 pg/ml for INS-1E for 24 hours), high IL-1β dose (300 pg/ml for human islets, 300 pg/ml for mouse islets and150 pg/ml for INS-1E for 24 hours), INF-γ (10 ng/ ml for human islets, 10 ng/ml for mouse islets and INS-1E for 24 hours) or control media. Statistical analysis was performed by paired t-tests of treatments versus control. Experiments done on individual islet donors (B and C) or biological cell replicates (A) are connected by lines. https://doi.org/10.1371/journal.pone.0222432.g005

10 ,

 10 from ENCODE-track option. For clarity and in relevance to the current publication, only some transcription factors are presented. (TIF) 7.1.3 Article: Conformational maps of human 20S proteasomes reveal PA28-and immunodependent inter-ring crosstalk Nathalie Vigneron 1,2,3,11* & Benoit J. Van den Eynde 1,2,3,11* The proteasome is responsible for selective degradation of proteins. It exists in mammalian cells under four main subtypes, which differ by the combination of their catalytic subunits: the standard proteasome (β1-β2-β5), the immunoproteasome (β1i-β2i-β5i) and the two intermediate proteasomes (β1-β2-β5i and β1i-β2-β5i). The efficiency of the four proteasome subtypes to degrade ubiquitinated or oxidized proteins remains unclear. Using cells expressing exclusively one proteasome subtype, we observed that ubiquitinated p21 and c-myc were degraded at similar rates, indicating that the four 26S proteasomes degrade ubiquitinated proteins equally well. Under oxidative stress, we observed a partial dissociation of 26S into 20S proteasomes, which can degrade non-ubiquitinated oxidized proteins. Oxidized calmodulin and hemoglobin were best degraded in vitro by the three β5icontaining 20S proteasomes, while their native forms were not degraded. Circular dichroism analyses indicated that ubiquitin-independent recognition of oxidized proteins by 20S proteasomes was triggered by the disruption of their structure. Accordingly, β5i-containing 20S proteasomes degraded unoxidized naturally disordered protein tau, while 26S proteasomes did not. Our results suggest that the three β5i-containing 20S proteasomes, namely the immunoproteasome and the two intermediate proteasomes, might help cells to eliminate proteins containing disordered domains, including those induced by oxidative stress.

Figure 1 .

 1 Figure 1. Degradation of ubiquitinated proteins by the standard proteasome (SP), the intermediate proteasome β5i (SIP), the intermediate proteasome β1i-β5i (DIP) and the immunoproteasome (IP). (a) p21 and (b) c-myc are degraded in a proteasome and ubiquitin-dependent manner. Western blot analysis of lysates of 293 SP cells treated or not with cycloheximide (CHX) alone or in combination with MLN7243, an inhibitor of the ubiquitinactivating enzyme, or with MG132 or bortezomib, two proteasome inhibitors. Cells were treated with MLN7243, MG132 or Bortezomib 30 min prior to the 5 h treatment with cycloheximide. (c, d) Western blot analysis of the kinetics of degradation of (c) p21 and (d) c-myc in 293 cell lines expressing the different proteasome subtypes. (e, f) Densitometric evaluation of the kinetics of the degradation of (e) p21 and (f) c-myc in the four different cell lines. All values (+ SD) are collected from three independent experiments. Full-length images for (a-d) are presented in Fig. S10.

Figure 2 .

 2 Figure 2. Oxidative stress dissociates 26S proteasomes into free 20S proteasomes. (a) Relative abundance of the different proteasome regulators and ECM29 bound to 20S proteasomes under oxidative stress induced by H 2 O 2 . 293 SP cells were treated with or without 2 mM of H 2 O 2 for 30 min, proteins were cross-linked in vivo with formaldehyde, proteasomes were immuno-purified using the anti-α2 antibody (MCP21) and the different proteasome regulators were analyzed using label-free quantitative MS along with ECM29. The abundance of 20S interactors were measured in control and oxidized conditions, normalized to the abundance of 20S, and then the oxidized condition was normalized to the control condition. All values are means of three independent experiments + SD. (b) Pie chart showing the proportion of free 20S proteasome and of the different regulators associated to 20S proteasomes under control conditions (left panel) and oxidative stress (right panel).

Figure 3 .

 3 Figure 3. More efficient degradation of oxidized proteins by the 20S immunoproteasome (IP) and the two 20S intermediate proteasomes β5i (SIP) and β1i -β5i (DIP). (a-c) Western blot analysis of the kinetics of degradation of (a) native and (b) oxidized calmodulin by the four 20S proteasome subtypes. (c) Densitometric evaluation of the kinetics of the degradation of the oxidized calmodulin by the four proteasome subtypes. All values (+ SEM) are collected from seven independent experiments using three different batches of purified proteasomes.(d, e) Kinetics of degradation of tritium-labeled (d) native and (e) oxidized hemoglobin. These two forms of hemoglobin were incubated with the four 20S proteasome subtypes. To monitor protein degradation, samples were collected at different time points, precipitated with trichloroacetic acid (TCA) and the radioactivity present in the supernatant was measured. Remaining hemoglobin was quantified by subtracting the radioactivity measured in the supernatant from the total radioactivity. Proteins were oxidized by 24 h incubation in 50 mM H 2 O 2 . All values are means (+ SEM) of seven independent experiments using three different batches of purified proteasomes. Full-length images for (a-b) are presented in Fig.S11.

Figure 4 .

 4 Figure 4. Degradation of oxidized proteins is triggered by disruption of their structure. (a) Analysis by circular dichroism of the secondary structure of native 52, 53 calmodulin, oxidized calmodulin and oxidized calmodulin treated with 0.5 mM CaCl 2 . The top panel shows the circular dichroism spectra and the bottom panel shows percentages of the different secondary structures obtained by analyzing the circular dichroism spectra using the Contin-LL program. (b) The left panel illustrates a western blot showing that CaCl 2 treatment of oxidized calmodulin reduced its degradation by 20S immunoproteasomes. These results are representative of three independent experiments. The right panel shows that CaCl 2 did not affect the activity of 20S immunoproteasomes, as measured with fluorogenic substrate Suc-LLVY-AMC. (c) Analysis by circular dichroism of the secondary structure of native hemoglobin and oxidized hemoglobin. The top panel shows the circular dichroism spectra and the bottom panel shows percentages of the different secondary structures. (d) Kinetics of degradation of labeled unoxidized and oxidized hemoglobin. These two forms of hemoglobin were incubated with the 20S IP, and protein degradation was monitored as described in Fig. 3e. Similar results were obtained from seven independent experiments. (e, f) Assessing the level of surface exposed hydrophobic patches in the three forms of calmodulin (e) and in the two forms of hemoglobin (f) using the Nile Red probe. Nile Red was incubated with the different samples and its fluorescence at 630 nm, which increases in a nonpolar environment, was measured. Relative fluorescence units (RFU) at 630 nm of each sample was reported to the RFU at 630 nm of the corresponding native protein, and the absolute RFU values at 630 nm of the native proteins are indicated on the histogram. All values are mean + SEM of three independent experiments. Full-length images for (b) are presented in Fig. S12.

Calmodulin

  Unoxidized precursor: Abz-GTVMRSLG-EDDnp Oxidized precursor: Abz-GTVMoxRSLG-EDDnp Hemoglobin Unoxidized precursor: Abz-DAVMNNPK-EDDnp Oxidized precursor: Abz-DAVMoxNNPK-EDDnp b a NY-ESO1 Unoxidized precursor: Abz-ATPMEAEL-EDDnp Oxidized precursor: Abz-ATPMoxEAEL-EDDnp LCMV nucleoprotein Unoxidized precursor: Abz-GVYMGNTL-EDDnp Oxidized precursor: Abz-GVYMoxGNTL-EDDnp

Figure 5 .

 5 Figure5. The presence of oxidized residues does not explain the differential degradation of oxidized proteins by the four 20S proteasome subtypes. Assay comparing the fluorescence emitted following the degradation of oxidized and unoxidized FRET (Fluorescence resonance energy transfer)-peptides by the four proteasome subtypes. Oxidized and unoxidized precursor peptides were derived from (a) calmodulin, (b) hemoglobin, (c) NY-ESO1 and (d) LCMV nucleoprotein. Unoxidized precursor peptides contained a methionine residue which was replaced by methionine sulfoxide (Mox) in the corresponding oxidized precursor. These precursor peptides were incubated with the four proteasome subtypes, and their degradation was monitored by measuring fluorescence at 420 nm. RFU at 420 nm of each sample was reported to the RFU at 420 nm of the unoxidized precursor degraded by the SP. The absolute RFU values at 420 nm of the unoxidized precursors degraded by the four proteasome subtypes are indicated on the histogram. All values are mean + SEM of three independent experiments.

Figure 6 .Figure 7 .

 67 Figure 6. Catalytic immuno-subunit β5i plays an important role in the degradation of oxidized proteins by β5i-containing 20S proteasomes. (a) Western blot analysis of the kinetics of degradation of oxidized calmodulin by the four proteasome subtypes in the presence or absence of the β5i-specific inhibitor PR-957 (1 μM). These results are representative of three independent experiments. (b) Assessing the degradation of the radiolabeled oxidized hemoglobin by the four proteasome subtypes in the presence or in the absence of the β5i-specific inhibitor PR-957 after 24 h of digestion. The percentage of degraded hemoglobin was reported in each case to the percentage of degraded hemoglobin by the SP in the absence of PR-957, and the absolute percentages of degraded hemoglobin in the vehicle condition are indicated on the histogram. Error bars are SEM of three independent experiments. (c) Western blot analysis of the kinetics of degradation of oxidized calmodulin by the intermediate proteasome β5i treated with increased concentrations of the PR-957 inhibitor. These results are representative of three independent experiments. Full-length images for (a) and (c) are presented in Fig. S13.

  by 20S proteasomes. Recombinant tau 2N4R (8.3 µg) (Anaspec Cat#AS-55556-50) was mixed with purified 20S proteasomes (2.1 µg) in a buffer containing 5 mM Tris-HCl pH 7.4, 1 mM MgCl 2 , 10 mM KCl and 0.01 mM EGTA in a final volume of 40 µl and the reaction was incubated at 37 °C. Samples (1.6 µl) were collected at different time points and the reaction was stopped by freezing the samples on dry ice. These samples were then analyzed by western blot.Degradation of tau by 26S proteasomes.Recombinant tau 2N4R (1.35 µg) was mixed with purified 26S proteasomes (2.6 µg) in a buffer containing 5 mM Tris-HCl pH 7.4, 1 mM MgCl 2 , 10 mM KCl, 0.01 mM EGTA and 2 mM of ATP in a final volume of 20 µl and the reaction was incubated at 37 °C. Samples (1.6 µl) were collected at different time points and the reaction was stopped by freezing the samples on dry ice. These samples were then analyzed by western blot.

7. 1 . 2

 12 Article: Efficiency of the four proteasome subtypes to degrade ubiquitinated or oxidized proteins ARTICLE Conformational maps of human 20S proteasomes reveal PA28-and immuno-dependent inter-ring crosstalks Jean Lesne 1,3,5 , Marie Locard-Paulet 1,2,5 , Julien Parra 1 , Dušan Zivković 1 , Thomas Menneteau 1,4 , Marie-Pierre Bousquet 1 , Odile Burlet-Schiltz 1 & Julien Marcoux 1 ✉

  ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-020-19934-z 2 NATURE COMMUNICATIONS | (2020) 11:6140 | https://doi.org/10.1038/s41467-020-19934-z | www.nature.com/naturecommunications kept only the RDU (or difference of RDU) of the shortest one.

  (diff. RDU)) in complex alonePeptide detected RDU color-coded in heatmap

Fig. 2

 2 Fig. 2 HDX-MS of the std20S reveals a dynamic pore entrance and inter-ring interface. a, b Structure of the human std20S proteasome (PDB: 5LE5) 35 showing the four stacked αββα heptameric rings colored by subunit (a) or relative deuterium uptake (RDU) (b). c-f Their solvent accessibility is represented on each face of the α-ring (solvent interface (c) and β-ring interface (d)) and β-ring (α-ring interface (e) and β-ring interface (f)) as the sum of consolidated relative deuterium uptakes between 0.5 and 30 min (described in Fig. 1b, c). The α-pockets (docking sites for the C-termini of PA28 regulators) are circled in white in c. Regions discussed in the text are indicated with red arrows and the moderately flexible/accessible bulges at the interface between the αand β-ring are circled in red in e. The corresponding residues are indicated in red around the structures. Active sites are circled in orange (f). The ring faces visualized in d-f are indicated by arrows on the right of the structure in a.

Fig. 3 6 NATURE

 36 Fig.3Comparison of the catalytic-subunit solvent accessibilities. a Representation of the three catalytic subunits β1 (left), β2 (middle), and β5 (right) of the std20S (PDB: 5LE535 ). The residues forming the active site and the substrate-specific pockets are indicated and color-coded (active site, S1, S2, S3, and S' are represented in red, green, blue, purple, and yellow, respectively). b-d Deuteration heatmaps showing the relative deuterium uptake (RDU) for each timepoint of the kinetics, along the protein sequences of β1/β1i (b), β2/β2i (c), and β5/β5i (d). The peptide sequence coverage is presented above the heatmaps as explained in Fig.1d. For the sake of clarity, only the sequence stretches containing residues of the active sites and substrate pockets are represented here.

Fig. 4

 4 Fig. 4 Differential analysis of std20S and i20S. a-d Sum of differential relative uptakes between the i20S and the std20S (Δ(RDU) = RDU i20S -RDU std20S ): β-ring/β-ring interface (a); β-ring facing the α-ring (b); α-ring facing the β-ring (c); solvent-facing α-ring (d); inside view showing the antechambers and catalytic pockets (e). Color-coded regions were significantly more accessible/dynamic in the std20S (blue = -30%) or i20S (red = +30%) and regions of interest are indicated with arrows. Regions that are not covered in one or both conditions, or do not pass the statistical threshold are represented in black and grey, respectively (see Fig. 1 for more details). RDU relative deuterium uptake.

Fig. 5 PA28 8 NATURE

 58 Fig. 5 PA28 regulators present flexible loops on both ends of the channel. a, b Structure of monomeric PA28α, β, or γ showing the 4-helix-bundle colored based on amino-acid numbering (a) or the sum of hydrogen-deuterium relative uptake accumulated over 30 min in PA28γ or PA28α 4 β 3 in solution (b). The residues corresponding to the missing apical loops (dashed lines) are indicated in b. c Structure of the heptameric complex PA28α 4 β 3 (PDB: 5MX5 33 ). Only four apical loops out of seven are represented, for the sake of clarity. d, e Surface accessibility of the heptameric complexes PA28α 4 β 3 (d PDB: 5MX5) and PA28γ 7 (e homology model) viewed from the top (left), front and inside (middle), and bottom (right). Regions that were not detected are represented in black (described in Fig.1b, c). RDU relative deuterium uptake.

Fig. 6 Fig. 7

 67 Fig. 6 Activation of PA28αβ and PA28γ by the 20S involves a rearrangement of the N-ter kink and an allosteric change. The sum of differential RDUs accumulated from 0.5 to 30 min deuteration of the PA28 regulators alone or in presence of the 20S proteasome (Δ(RDU) = RDU PA28/20S -RDU PA28 ) is color coded from blue (less accessible in the complex) to red (more accessible in the complex) and ranges from -70%/+70% for PA28αβ + std20S (a) and -30%/+30% for PA28γ + std20S (b) PA28αβ + i20S (c) and PA28γ + i20S (d). The heptameric complexes PA28α 4 β 3 (a and c PDB: 5MX5) and PA28γ 7 (b and d homology model) are viewed from the top (left), front and inside (middle), and bottom (right).Residues of PA28 that present a significant decrease of deuteration upon binding of the 20S proteasome are located around the N-ter kink (red ovals), the activation loop and 20S interface and the entry of the channel (left). Residues in gray were not considered significantly different according to the statistical thresholds presented in Fig.1c. Residues with no RDU information in one or both conditions are in black. Δ(RDU) sum of differences of relative deuterium uptake.

  Huang et al., 2016; Dong et al., 2019). Another Ub receptor -RPN13 binds in substoichiometric ratios to the 19S, via the RPN2 lid subunit (Berko et al., 2014; Z. Liu et al., 2019) (see Figure 11, left). The 19S is highly dynamic, both in terms of structure and composition: besides its canonical set of subunits, it may interact with many other proteins in charge of substrate recognition or deubiquitylation, such as the deubiquitylation enzymes (DUBs). Moreover, 19S function is regulated by a number of PTMs (VerPlank et al., 2019; Liu et al., 2020).

Table 1 .

 1 Brief summary of structural MS techniques, with their advantages and disadvantages.

	Technique	Advantages	Disadvantages
	Native MS	Requires less sample compared	Buffer compatibility issues.
		to conventional structural	Potential gas phase dissociation
		techniques. Can identify more	/ not very good for
		than one analyte at a time.	quantification of different
		Quick analysis time.	oligomeric complexes intra
			spectrum (different ionization
			yields)
	Top-down proteomics	Can handle complex samples.	Limited by protein size (in
		Identify proteoforms and	complex mixtures). Loss of low-
		relatively quantify them. Can	abundant species. Requires
		handle relatively complex	fractionation (GELLFrEE, SEC, IEF,
		samples (IPs, secretomes)	etc...

) for deeper proteome analysis. Middle-down proteomics Can characterize proteins with very complex and heterogeneous PTMs. Less limited by protein size. Limited to few proteins, due to complexity limitations. Cross-linking proteomics Gives distance (molecular ruler) information for many interactors at once. Complex data analysis. Requires condition optimization.

  . Different instrument configurations give different flavours to nMS: combining the ion mobility cells with TOF to study gasphase protein unfolding (Vallejo et al., 2019), employing extended mass range analysers to observe MDa assemblies (van de Waterbeemd et al., 2016)or various fragmentation techniques for native TD (Lermyte et al., 2015; Greisch et al., 2019). As mentioned earlier, IM provides us with values directly relatable to theoretical CCS of proteins (Jurneczko andBarran, 2011) and enables to separate conformers or better resolve the spectra of co-eluting species (Uetrecht et al., 2011). Native TD enables us to isolate native complexes while inside the mass spectrometer and fragment them for identification, which is useful to identify specifically-interacting ligands or proteoforms[START_REF] Belov | From Protein Complexes to Subunit Backbone Fragments: A Multistage Approach to Native Mass Spectrometry[END_REF][START_REF] Gupta | The role of interfacial lipids in stabilizing membrane protein oligomers[END_REF].

  . Recent efforts of Brown et al. showed how many membrane proteins can be identified using TDP, combining smart precipitation techniques with MS-compatible SEC (Brown et al., 2020) fractionation. TDP is not yet widely adopted in clinical research, however, several studies prove the value of this approach (Molnar et al., 2009; Mazur et al., 2010; Ntai et al., 2016, 2018) . Particularly interesting study by Dong et al. from Ge group identified a phosphorylation site on a cardiac muscle proteintroponin I, responsible of hypertension (Dong et al., 2012).

  al., 2012; Reading et al., 2017; Kopcho, Chang and Komives, 2019). Protein-DNA studies (Zheng et al., 2017) and even protein-small ligand binding studies employ HDX. Small ligand binding indeed diverges into many possibilities. If the binding induces sufficient stabilization or conformational change -it can be observed by HDX: enzyme-inhibitor binding studies (S. Lee et al., 2016), ligand screening in drug discovery (Masson, Maslen and Williams, 2017) or even metal ion binding (Baños-Mateos et al., 2017). However, if the ligand purely interacts via sidechains and does not induce backbone changes, its binding remains invisible to HDX.

  MS analysis was done in the Orbitrap at 60k resolution, scan range between 350 and 1400 m/z, with RF lens value set to 60%, AGC target set to 4e5, maximum injection time to 50 ms, with 1 microscan performed in profile mode. MSMS was done in data-dependant mode, in the Orbitrap, at resolution of 15k (first mass at 100 m/z). Monoisotopic peak determination was set for peptides, intensity threshold of 2.5e4 was used and only charge states from 2 to 6 were used as precursors. Dynamic exclusion was used with the following parameters: exclude the ion after "1" times, during 30 sec, including isotopes, with mass tolerance, high and low, 10 ppm each. Ion isolation was done in the quadrupole, with an isolation window of 1.7 m/z. The activation was done by HCD at 28% collision energy. AGC was set to 5e4, ions were injected for all available parallelizable time, with maximum injection time of 22 ms. A total of 1 microscans per cycle was used and recorded in centroid mode.

	Proteasome complexes experience profound structural and functional rearrangements throughout
	mammalian spermatogenesis
	type: Intensity; Intensity threshold: 0). Normalization and statistics were done using the following parameters: Global options: (Apply profile clustering: false (method: QUANT_PROFILE); Use only specific peptides: true; Discard miscleaved peptides: false (method: KEEP_MOST_ABUNDANT_FORM); Discard oxidized peptides: false (method: KEEP_MOST_ABUNDANT_FORM); Abundance summarizer method: LFQ based on QUANT_PEPTIDE_IONS); Protein options: (Apply normalization: true; Apply missing values inference: false; Apply variance correction: false; Apply T-test: true (p-value: 0.01); Apply Z-test: true (p-value: 0.4)); Peptide options: Apply normalization: true; Apply missing values inference: false; Apply variance correction: false; Apply T-test: true (p-value: 0.01); Apply Z-test: true (p-value: 0.4). The p-value for a non-paired, one-sided T-test and fold change were calculated between the experimental IP groups and the control IP groups (IP with the OX8 antibody). Proteins enriched over 2-fold with a p-value below 0.05 were considered 275 °C. using following significantly enriched.
	parameters: PSM filters: Pretty Rank: 1.0, Peptide Seq Length: 7.0; PSM validation: expecting
	FDR of 1.0% on Mascot Adjusted E-Value; Validation of the protein sets was done using the
	following parameters: Min. specific peptide count: 1; Enable protein FDR (false discovery rate)
	filter: true; Min protein set FDR (%): 1.0; Peptide set scoring method: Mascot Modified
	MudPIT; Dataset merging mode: After validation (merge identification summaries). Label-free
	quantitation was done using following parameters: Quantitation type: label_free;
	Quantitation method: Label free based on the extraction of feature abundance; Extraction
	m/z tolerance: 5 ppm; Mapping m/z tolerance: 5 ppm; Min. peakel duration: 15 sec;
	Alignment computation method: (Iterative; Max number of iterations: 3; m/z tolerance: 5
	99

Bottom-Up data treatment Identification and quantitation parameters. The identification was done with Mascot search engine, using following search parameters: Peptide mass tolerance : 10 ppm; Peptide charge : 2+ and 3+; Cleavage enzyme: Trypsin/P; Max. missed cleavages : 2; Decoy : true; Mass : Monoisotopic; Fixed modifications : Carbamidomethyl (C); Variable modifications : Acetyl (Protein N-term),Oxidation (M); MS/MS IONS SEARCH : true; Error tolerant : false; MS/MS tolerance : 20 mmu; Data format : Mascot generic; Quantitation : None; Instrument : ESI FTMS HCD. The UniProt entries (SwissProt & TrEMBL) for Bos Taurus were used as a protein entry database, including isoforms. In the case of rat germ cells, the search was performed with Rattus norvegicus Uniprot database, including isoforms. The result files were then imported into Proline -software for Bottom-Up proteomics (8), using following parameters: Peaklist software: Proline; Generate intermediate mzIdentML file: false; Instrument configuration: ORBITRAP FUSION (A1=FTMS F=HCD A2=FTMS); Decoy strategy: Concatenated Decoy Database; Protein match decoy rule: SearchGUI REVERSED; Ion score cutoff: None; Subset threshold: None; Replace peptide matches scores by PEP values (Posterior Error Probabilities): false; Automatically associate raw files to identification results (DAT, OMX): true. Validation of the peptide IDs in Proline was done ppm; Time tolerance: 5 sec); Alignment smoothing: (Method: Landmark Range; Number of landmarks: 50; Sliding window overlap: 50); Features cross assignment: (Mapping m/z tolerance: 5 ppm; Time tolerance: 60 sec; Normalization method: None; Master feature filter

Table S1 .

 S1 Estimation of the relative abundances of the immuno-catalytic subunits in whole bovine testes (See Top-Down MS experimental part for detailed calculations).

						Standard
		Run 1	Run 2	Run 3	Average	deviation
	β5i (%)	35.0	36.3	36.8	36.0	0.9
	β1i (%)	32.0	29.6	30.5	30.7	1.2
	β2i (%)	17.5	7.5	9.9	11.7	5.2

Table S2 .

 S2 Theoretical

	PA28γ	0.831	0.363
	PA200	1.969	0.0270
	PI31	1451.3	0.002
	Fbxo7	71.58	0.007
	PAC1-4	0.887	0.136
	POMP	0.187	0.020
	SYCE1	294.31	0.009
	SYCP3 (rep1 excluded)	125.65	0.013
		ɑ4s/ɑ4	p-value (T-test)
	non-catalytic 20S	1.030	0.234
	catalytic c20S	3.133	0.002
	catalytic i20S	0.006	6.8E-05
	19S	14.555	0.025
	PA28αβ	0.164	0.012

and observed MWs of the bovine proteasome subunits, along with their corresponding PTMs *nomenclature as in

(25)

.

Table S3 .

 S3 Estimation of the abundance ratio of different proteins between ɑ4s and ɑ4 containing proteasomes in whole bovine testes (See experimental part for detailed calculations).

	UBE3A	3.40E-03	6.20E-04	7.98	1.11E-02
	NPLOC4	8.87E-03	1.79E-03	7.68	2.27E-02
	USP26	2.46E-04	3.40E-05	7.61	1.83E-03
	USP34	1.81E-04	3.68E-05	7.57	1.92E-02
	PELI2	8.44E-02	1.20E-02	7.52	3.60E-04
	UBR1	9.39E-04	1.26E-04	7.12	2.54E-02
	USP11	1.51E-04	2.38E-05	7.00	3.17E-02
	ARIH2	2.18E-04	4.67E-05	6.85	4.59E-02
	USP5	1.58E-02	2.30E-03	6.66	4.05E-04
	USP47	3.92E-04	1.02E-04	6.47	3.59E-02
	USP12	5.17E-04	9.10E-05	6.42	2.36E-03
	UFD1	1.46E-02	3.58E-03	5.27	1.28E-02
	Gene name RNF114	averaged normalized 3.42E-04	averaged normalized 7.91E-05	averaged FC (anti-α4s vs. 4.56	t test (anti-α4s vs. anti-α2) 2.37E-03
	UBE4B	iBAQ_IP anti-α4s 6.28E-05	iBAQ_IP anti-α2 1.61E-05	anti-α2) 4.51	1.55E-02
	USP10 UBAP2L	5.86E-03 3.25E-03	4.07E-05 1.33E-03	148.49 4.49	4.29E-05 1.73E-02
	RNF40 KCMF1	3.15E-02 2.61E-04	4.51E-04 7.17E-05	103.05 4.45	8.83E-04 2.17E-02
	RNF20 UBE2N	3.03E-02 7.15E-04	4.40E-04 2.67E-04	84.31 4.27	4.90E-04 4.67E-02
	UBE2A UCHL5	1.41E-02 9.81E-03	2.74E-03 3.96E-03	35.36 3.87	4.59E-02 4.65E-02
	DTX3L ITCH	1.80E-03 1.00E-04	7.59E-05 2.66E-05	33.26 3.48	9.08E-03 4.48E-03
	HUWE1 USP14	1.94E-03 5.68E-03	1.10E-04 2.19E-03	26.55 3.24	5.39E-03 1.48E-02
	HERC4 BRCC3	7.85E-04 1.82E-04	7.88E-05 7.70E-05	19.83 2.54	1.77E-02 4.33E-02
	USP16 RPS27A	6.28E-04 4.98E-02	6.48E-05 2.88E-02	18.89 1.89	7.17E-03 3.09E-02
	UFM1 ADRM1	1.04E-02 2.19E-02	5.81E-04 1.22E-02	18.88 1.83	4.96E-03 1.04E-02
	HERC5 HERC2	1.22E-03 2.55E-06	2.86E-04 5.69E-06	17.12 0.49	4.78E-02 2.82E-02
	USP40 TRIM21	2.53E-03 2.68E-03	3.42E-04 6.71E-02	16.04 0.04	1.44E-02 3.58E-05
	USP15	4.96E-04	1.10E-04	11.45	4.71E-02
	OTUD4	2.04E-04	2.52E-05	11.45	8.78E-03
	UFC1	1.05E-03	9.83E-05	10.71	5.12E-03
	UBR5	3.99E-03	6.87E-04	9.60	2.09E-02
	OTUB1	1.13E-02	2.15E-03	9.05	4.01E-02

Table S4 .

 S4 Averaged normalized IBAQ values and FC enrichment in anti-ɑ4s vs. anti ɑ2 IP for selected ubiquitin-related proteins.

Table 2 .

 2 Theoretical masses of the proteasome complexes and their regulators

	Protein complex	Theoretical MW [Da]
	standard proteasome (bovine)	715 750
	spermatoproteasome (bovine)	715 842
	PA200 (human)	211 334
	PA28γ (human)	216 126
	PI31 (human)

  The intermediate proteasome expression in pancreatic beta cells PLOS ONE | https://doi.org/10.1371/journal.pone.0222432 February 13, 2020 2 / 21 Funding: This study was funded by The Punjab Educational Endowment Fund https://www.peef. org.pk/ (M.S.K), the Department of Biomedical Sciences at the University of Copenhagen https:// bmi.ku.dk/ (T.D. and M.T.M.); the Augustinus

	Foundation https://augustinusfonden.dk/ (T.D. and
	M.T.M.); EFSD/JDRF/Lilly European Programme in
	Type 1 Diabetes http://www.europeandiabetes
	foundation.org/, Vissing Fonden http://www.vissing
	fonden.dk/, Bjarne Jensen Fonden http://www.
	bjarnejensensfond.dk/, Poul og Erna Sehested
	Hansens Fond, no website, Eva og Hans Carl
	Holms Mindelegat https://haldguttenberg.dk/1-
	april/(M.T.M). The funders had no role in study
	design, data collection and analysis, decision to
	publish, or preparation of the manuscript.
	Competing interests: The authors have declared
	that no competing interests exist.
	Abbreviations: i-proteasome, Immunoproteasome;
	int-proteasome, Intermediate proteasome;
	s-proteasome, Standard proteasome; β1, beta
	subunit 1; β2, beta subunit 2; β5, beta subunit 5;
	β1i, inducible beta subunit 1; β2i, inducible beta
	subunit 2; β5i, inducible beta subunit 5; PSMB8,
	Proteasome subunit beta type-8 = β5i; PSMB9,
	Proteasome subunit beta type-9 = β1i; PSMB10,
	Proteasome subunit beta type-10 = β2i; HBSS,
	Hank's balanced salt solution; IL-1β, Interleukin 1
	β; INF, Interferon; MHC, Major histocompatibility
	complex.

Table 1 . Human islets donors information and islet preparations used during the investigation. Islet donors information

 1

	Donor	1	2	3	4
	Age	63	62	58	20
	Gender (M/F)	F	F	M	M
	BMI	19.5	29.3	27.8	21.8
	blood group	A+	A+	O+	B+
	HLA (A:B)	2,11 : 18,57	26,29 : 7,18	2,26 : 35,55	11,24 : 18,51
	HLA (DR)	11,17	4,15	14,16	1,11
	Cold ischemia time (h)	9	5	8.5	8
	Islets culture duration (h) �	16	14	20	42
	Cause of death	Cerebral bleeding	Cerebral bleeding	Cerebral bleeding	Anoxia
	Source of islets	ECIT	ECIT	ECIT	ECIT
	Estimated viability (%)	95	95	95	95
	Estimated purity (%)	90	90	90	90
	Any additional note	EBV positive			EBV positive

� time from islets isolation to shipment https://doi.org/10.1371/journal.pone.0222432.t001

Table 2 . Primary and secondary antibodies used during the investigation.

 2

	Antibody target	Company	Cat#	Dilution
	β1i	Abcam	ab243556	1:10.000
	β5i	Abcam	ab3329	1:5.000
	β2i	Abcam	ab183506	1:1.000
	Tubulin	Sigma	T6074	1:10.000
	Actin	Thermofisher Scientific	MA5-11869	1:15.000
	Insulin	Cell signaling	8138S	1:5.000
	Anti-mouse secondary	Cell signaling	7076S	1:10.000
	Anti-rabbit secondary	Cell signaling	7074S	1:10.000
	https://doi.org/10.1371/journal.pone.0222432.t002

Table 3 . Low concentrations of IL-1β induce β subunit mRNA expression in mouse islets.

 3

	Gene	Ctrl 1	Ctrl 2	Ctrl 3	IL-1B	IL-1B	IL-1B	Ctrl	IL-1β treatment	P value
					1	3	3	mean	mean
	β5i (PSMB8)	9.38	8.54	9.03	28.13	24.57	26	8.98	26.23	0.0021
	β1i (PSMB9)	6.70	5.41	6.59	13.31	16.02	15.79	6.23	15.04	0.0172
	β2i (PSMB10)	12	11.75	9.57	33.99	36.21	32.18	11.11	34.13	0.001
	β5	34.53	37	38.27	42.61	42.78	34.31	36.6	39.9	0.4657
	(PSMB5)
	β1	102.84	93.47	90.64	93.09	109.07	98.63	95.65	100.26	0.6015
	(PSMB6)
	β2	79.63	76.68	79.96	85.41	83.28	72.34	78.76	80.34	0.7634
	(PSMB7)

Деда Томи, који увек верује у мене.

Proteasome complexes experience profound structural and functional rearrangements throughout mammalian spermatogenesis

PLOS ONE | https://doi.org/10.1371/journal.pone.0222432February 13, 2020

Scientific RepoRtS | (2020) 10:15765 | https://doi.org/10.1038/s41598-020-71550-5

NATURE COMMUNICATIONS | (2020) 11:6140 | https://doi.org/10.1038/s41467-020-19934-z | www.nature.com/naturecommunications

Acknowledgements

I give my most sincere thanks to my two mentors, Marie-Pierre and Julien, for their huge support throughout these four years. It is a privilege and an honor to work with such dedicated people. Without your engagement and care, this would have been an impossible task.

I thank sincerely to the reviewers Olivier Coux, Silke Meiners and Frank Sobott, for their good will to participate in this endeavor. I am looking forward to reading your advices and talking to you about this subject. I thank Christel Lutz for her participation as the member of the jury. Your contributions are much appreciated.

I thank Odile Schiltz for the given opportunity to work with the Proteomics and Mass Spectrometry team and for providing me with the means to realize the project.

My sincere thanks to the Doctoral School for the trust they put in me and for the funding of Acknowledgements. This work was supported by the French Ministry of Research (ANR-ProteasoRegMS to JM, Investissements d'Avenir Program, Proteomics French Infrastructure, ANR-10-INBS-08 to OB-S), University of Toulouse (grant to DZ), the Fonds Européens de Développement Régional Toulouse Métropole and the Région Midi-Pyrénées (OB-S). PCAdF and ATR were funded by the Medical Research Council (MC_UP_1201/5 grant to PCAdF).

Acknowledgments

The authors are thankful to JDRF award 31-2008-416 (ECIT Islet for Basic Research program) for providing human islets for this study. The authors also acknowledge C. Wollheim and P. Maechler (University Medical Centre, Geneva, Switzerland) for providing INS-1E cell line. Soren Buus, (Department of International Health, Immunology and Microbiology, Copenhagen, Denmark) generously provided the A20 and Jurkat cells. We would like to thank Nils Billestrup for sharing and discussing his islet sequencing data that is presented in this manuscript.

Acknowledgements

We are grateful to Ms Auriane Sibille for her precious help in the preparation of this manuscript. We thank M. Gyrd-Hansen (Ludwig Cancer Research institute, Oxford Branch) for providing the control GST and the GST-UBA fusion protein used for the TUBE-pulldown assay. We thank M. Jean-Pierre Brion (Université Libre de Bruxelles) for providing the B19 anti-tau antibody. We thank Dr. C. Uyttenhove for generation of the hybridomas producing the monoclonal antibodies targeting the β1i and β2i subunits of proteasome. We thank Therèse Aerts and Rui Cheng for their precious technical assistance. JAH was supported by a fellowship from the Fonds National de la Recherche Scientifique (TELEVIE Grant No. 7455115F). JM was supported by a grant from the V.I.B, Belgium. The work was supported by grants from the Fonds National de la Recherche Scientifique (F.N.R.S) and from the Fonds Wetenschappelijk Onderzoek-Vlaanderen (F.W.O) (ID EOS : 30837538 and PDR T.0091.18). This work was also supported by grants from the Walloon excellence in life sciences and biotechnology (WEL-BIO), from the Fondation contre le Cancer, the Fonds Maisin, Belgium and the Région Midi-Pyrénées, European funds (Fonds Européens de Développement Régional, FEDER), Toulouse Métropole, and by the French Ministry of Research with the Investissement d' Avenir Infrastructures Nationales en Biologie et Santé program (ProFI, Proteomics French Infrastructure project, ANR-10-INBS-08). Acknowledgements This work was supported by the French Ministry of Research (ANR-ProteasoRegMS to J. M. and Investissements d'Avenir Program, Proteomics French Infrastructure, ANR-10-INBS-08 to O.B.-S.), the 631 Fonds Européens de Développement Régional Toulouse Métropole and the Région Midi-Pyrénées (O.B.-S.), and the Novo Nordisk Foundation (NNF14CC0001).

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Data availability. The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE (54) partner repository with the dataset identifier PXD027436. Scientific RepoRtS | (2020) 10:15765 | https://doi.org/10.1038/s41598-020-71550-5

Data availability

The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE 69 partner repository with the dataset identifier PXD018921. All other data are available with this paper online.

Code availability

The R scripts of the analysis, with the fasta files and output tables and cxc files are freely available at zenodo.org [https://zenodo.org] with the https://doi.org/10.5281/ zenodo.3769174 under the Creative Commons Attribution 4.0 International licence.

Abbreviations

Competing interests

The authors declare no competing interests. the differences between the std20S-and i20S-specific subunits β1/1i, β2/2i and β5/5i located inside the complex trigger conformational changes at the pore entrance (aperture in the i20S: +++) and external surface of the proteasome, including the C-ter of α3 (inner-to-outer change represented as an arrow). The subunits that are more accessible/flexible in the std20S and the i20S are presented in blue and red, respectively. b PA28 solvent accessibilities analysed alone indicate similarities in their dynamic regions: their apical loops, kinks and activation loops were more deuterated (indicated with+). c 20Sand PA28-specific reorganizations occurring upon complex formation. This schematics presents the results of the comparison of HDX-MS data from the PA28 and std/i20S alone vs. in complex. The subunits in red and blue were globally more and less deuterated in the complex than in the protein alone, respectively. More localised changes are indicated by half-circles. Opening of the 20S pore entrance is indicated with + and +++ (for minor and more pronounced increase of deuteration) as well as the reorganisation of the C-ter of β2 and β2i upon complex formation.indicates the area of the proteins that were protected upon binding (apical loops, kinks and activation loops of the regulators; C-ter of α3 in the std20S). affinities. The very limited decrease of RDU on the α-ring interface interacting with PA28 regulators can be explained by the resulting gate opening (RDU increase) known to induce its increased proteolytic activity. Gate opening and binding interface protection are expected to have opposite impacts on RDU, so their combination in the same protein region would result in no significant changes of deuteration rates. We showed that both PA28 regulators activate the chymotrypsin-like activity of the std20S to a similar extent in our in vitro assay (Supplementary Fig. 2). However, the N-ter portions of the std20S α subunits (entrance of the channel) were significantly more deuterated when interacting with PA28αβ whereas this increase was not significant with PA28γ (Fig. 7a, f and Supplementary Data 3), suggesting that their allosteric mechanisms of activation are different. This could alternatively be explained by a shorter dwelltime at the surface of the std20S or by differences in binding stoichiometry. More experiments are needed to fully understand these allosteric mechanisms of activation.

Although not similarly distributed, a global core proteasome destabilization upon complex formation was found in all four conditions tested (Fig. 7), so we focused on the comparison of the most affected regions. We observed that each pair of PA28/20S complex underwent long-range conformational rearrangements in specific regions, as suggested recently 25 . Different sets of α Nter were destabilized depending on the PA28/20S pair (in red Fig. 8c), suggesting a gradual interaction-specific opening of the central pore, in-line with a recent cryo-EM structure that showed a partial opening of the bovine i20S gate in complex with human PA28αβ 65 . The β2 C-ter extending towards β3 was destabilized in all the conditions, except std20S/PA28γ. On the contrary, the region just before the α3 C-ter (217-236) was protected upon PA28 binding in all cases, except std20S/PA28αβ. Interestingly, this stretch is close to β3 74-86, which was also protected in all conditions tested except std20S/PA28αβ. These observations could provide a basis to understand how regulator binding propagates from the αto the β-ring depending on the activator/ 20S pair.

Altogether, these data provide a unique resource informing on the allosteric activation of the 20S proteasome complexes that should soon be completed by upcoming structural and functional studies. We think that HDX-MS could be applied to characterize, not only the binding sites of many new proteasome inhibitors, but also to identify their potential allosteric effects on the 20S core complex. The same is true for the many PIPs that regulate proteasome function and are still poorly characterized from a structural point of view 66 . More broadly, this work demonstrates the ability of HDX-MS to investigate dynamic events on megadalton assemblies including ribosomal particles, inflammasomes, and nucleosomes, to name a few.

Methods

Reagents. Unless stated otherwise, all reagents were purchased from Sigma-Aldrich. The std20S (BML-PW8720-0050), i20S (BML-PW9645-0050), PA28γ (BML-PW9875-0100), and PA28αβ (BML-PW9420-0025) were purchased from Enzo Life Science. Deuterium oxide (D215H) and sodium deuteroxide (D076Y) solutions were from Euriso-top and deuterium chloride (543047-50 G) was from Sigma-Aldrich. The fluorogenic peptide substrate (4011369.0025) for proteasome activity measurement was from Bachem.

Proteasome activity test. The assay was performed in black 96-well plates (Fluotrac 200-GREINER BIO-ONE). 5 μL of each sample (corresponding to 50 ng of 20S and/or 28 ng of PA28) were added in three independent experiments to 45 μL of Tris-HCl 100 mM pH 8 and 50 μL of Suc-LLVY-AMC (for chymotrypsin-like activity) substrate in 200 mM Tris-HCl, pH 8 at a final concentration of 400 μM/ well. The kinetic assays were performed at 37 °C in a FLX-800 spectrofluorimeter (BIOTEK, Winooski, VT, U.S.A.) over 90 min with one reading every 5 min, at 360 nm for excitation and 460 nm for emission. Each proteasome specific activity was obtained by dividing the activity by the proteasome quantity (50 ng). A two-sided ttest was used between the std20S alone and in presence of PA28.

Development of a HDX-MS pipeline dedicated to the comparative analysis of 20S proteasome/regulator complexes. In a classical HDX-MS workflow, the proteins of interest are digested online and the generated peptides are separated on a reverse phase column before MS analysis to monitor deuterium incorporation rates. The main challenges encountered when analysing heterogeneous complexes such as the proteasome by HDX-MS are threefold. First, the high number of peptides generated after digestion entailed specific optimization of their chromatographic separation. Second, we optimized the quenching step and injection parameters to reduce dead volumes, as well as the acquisition method, in order to handle samples at low concentration (<1 µM). Finally, we developed a computational pipeline dedicated to the multidimensional analysis of HDX-MS analysis of large complexes. The resulting data were mapped to available 3D structures using our recently developed open-source web application HDX-Viewer v.1.2 67 .

Automated Hydrogen-Deuterium eXchange coupled to Mass Spectrometry (HDX-MS). HDX-MS experiments were performed on a Synapt-G2Si (Waters Scientific, Manchester, UK) coupled to a Twin HTS PAL dispensing and labelling robot (LEAP Technologies, Carborro, NC, USA) via a NanoAcquity system with HDX technology (Waters, Manchester, UK). Data were collected with MassLynX v.4.1 from Waters Scientific (Manchester, UK) and the robot was controlled via HDx Director v.1.0.3.9 (LEAP Technologies, Carborro, NC, USA). Each step was optimized to minimize sample loss and work with such a heterogeneous and diluted sample:

Method in HDxDirector: The method recommended by Waters (HDX System Suitability Test) injects only 25% of the sample that is aspirated from the protein vial. In order to reduce sample loss, we (1) used a sample loop of 100 µl instead of 50 µl, (2) carefully reduced all the dead volumes, and (3) used 500 mM glycine pH 2.3 instead of 50 mM K 2 HPO 4 , 50 mM KH 2 PO 4 , pH 2.3 in order to optimize the ratio of quenching volume (10% instead of 50%). With this workflow, we increased by more than threefold the amount of starting material injected (79%). 20S proteasomes were incubated alone or with a 2-fold molar excess of PA28, with final concentrations of 0.4 and 0.8 µM, respectively. 5.7 µL of protein were aspirated and 5.2 µL were diluted in 98.8 µL of protonated (peptide mapping) or deuterated buffer (20 mM Tris pH/pD 7.4, 1 mM EDTA, 1 mM DTT) and incubated at 20 °C for 0, 0.5, 1, 5, 10, and 30 min. The final D 2 O percentage was 95% in the labelled samples. 99 µL were then transferred to vials containing 11 µL of precooled quenching solution (500 mM glycine at pH 2.3). For experiments involving PA28αβ, the quenching buffer was supplemented with 250 mM tris-(2carboxyethyl) phosphine (TCEP) in order to reduce the disulphide bridge between Cys21 of chain α and Cys3 of chain β. After 30 s. of quenching, 105 µL were injected into a 100 µL loop. Proteins were digested online with a 2.1 × 30 mm Poros Immobilized Pepsin column (Life Technologies/Applied Biosystems, Carlsbad, CA, USA). The temperature of the digestion room was set at 15 °C.

Chromatographic run: In order to cope with the unusual sample heterogeneity, the runtime of the chromatographic separation (12 min) was doubled compared to the one used for smaller protein complexes (6 min). Peptides were desalted for 3 min on a C18 pre-column (Acquity UPLC BEH 1.7 µm, VANGUARD) and separated on a C18 column (Acquity UPLC BEH 1.7 µm, 1.0 × 100 mm) by the following gradient: 5-35% buffer B (100% acetonitrile, 0.2% formic acid) for 12 min, 35-40% for 1 min, 40-95% for 1 min, 2 min at 95% followed by 2 cycles of 5-95% for 2 min and a final equilibration at 5% buffer A (5% acetonitrile, 0.2% formic acid) for 2 min. The total runtime was 25 min. The temperature of the chromatographic module was set at 4 °C. Experiments were run in triplicates and the protonated buffer was injected between each triplicate to wash the column and avoid cross-over contamination.

MS acquisition: The acquisitions were performed in positive and resolution mode in the m/z range 50-2000 Th. The sample cone and capillary voltages were set at 30 and 3 kV, respectively. The analysis cycles for nondeuterated samples alternated between a 0.3 s low energy scan (Trap and Transfer collision energies set to 4 V and 2 V, respectively), a 0.3 sec high energy scan (Ramp Trap and Transfer collision energies set to 18 to 40 V and 2 to 2 V, respectively) and a 0.3 sec lockspray scan (0.1 µM [Glu1]-Fibrinopeptide in 50% acetonitrile, 50% water and 0.2% formic acid infused at 10 µL/min). The lockspray trap collision energy was set at 32 V and a GFP scan of 0.3 sec is acquired every min. In order to double the signal intensity of deuterated peptides, deuterated samples were acquired only with the low energy and lockspray functions.

Data analysis: Peptide identification was performed with ProteinLynx Global SERVER v.3.0.2 (PLGS, Waters, Manchester, UK) based on the MS E data acquired on the nondeuterated samples. The MSMS spectra were searched against a homemade database containing sequences from the 17 std20S and i20S subunits, PA28α, PA28β, PA28γ, and pepsin from Sus scrofa. Peptides were filtered in DynamX v.3.0 from Waters Scientific (Manchester, UK) with the following parameters: peptides identified in at least two replicates, 0.2 fragments per amino-acid, intensity threshold 1000. The quantitative analysis of deuteration kinetics was performed using the statistical package R (R Development Core Team, 2012; http://www.Rproject.org/) on the corresponding MS intensities. The deuterium uptakes of each considering that all amino-acids (except proline residues and the first amino-acid or each peptide) were deuterated, and then averaged (weight = intensity) to get a value of relative deuterium uptake (RDU) per peptide sequence/condition/ timepoint (see Supplementary Data 4,5,12,14,16). The RDUs were not corrected for back exchange. To identify the protein regions that presented conformational changes in complex vs. alone, we performed an ANOVA (Anova(), type = III, singular.ok = T) followed by Benjamini-Hochberg correction of the P value on all quantified peptides. For each comparison, we considered significantly regulated the peptides with a corrected P value ≤ 0.01 and an absolute difference of RDU above 1.57% (four times the mean absolute difference of RDU in the entire dataset) for three successive time points (Supplementary Data 9-10). The corresponding volcano plots are presented in Supplementary Data 3, 7, 13 and all the regions statistically regulated in all the differential HDX-MS analysis are listed in Supplementary Data 8. The RDU and differences of RDU (for protein alones or comparison between conditions, respectively) were consolidated (RDU mapping from peptide-level to protein sequence) using the values of the smallest peptide to increase the spatial resolution (see consolidation heatmaps in Supplementary Data 1-3, 6, 11). These data (per timepoint or sum of all time points) were then used for structural data mining with the recently developed open-source web application called HDX-Viewer [https://masstools.ipbs.fr/hdx-viewer] 67 that allows to directly plot and visualize the whole dataset (14 different subunits per proteasome core particle) on the proteasome 3D structure. The scripts corresponding to the quality control and statistical analysis of this dataset are available at zenodo.org [https://zenodo.org] with the https://doi.org/10.5281/ zenodo.3769174 under the Creative Commons Attribution 4.0 International licence. All molecular representations were generated in UCSF ChimeraX v.0.9 (2019-06-06) 68 .

Modeling: The PA28γ model was generated using the structure of human PA28α (PDB:1AV0) as a template with the SWISS-MODEL server 52 . This model misses the first 3 and the last 7 amino-acids, as well as the 64-106 stretch.

Reporting summary. Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Author contributions

All authors contributed to editing the manuscript and figures. J.L. performed the HDX-MS optimization and data acquisition, model building and data analysis. J.P. performed the HDX-MS optimization, data acquisition, and data analysis. D.Z. and T.M. performed experiments. M.L.P. analyzed the data and wrote the paper. O.B.-S. provided critical input. M.P.B. provided critical expertise on the proteasome. J.M. designed, performed and supervised the experiments, data and structural analysis, and wrote the paper.

Competing interests

The authors declare no competing interests.

Additional information

Supplementary information is available for this paper at https://doi.org/10.1038/s41467-020-19934-z.

Correspondence and requests for materials should be addressed to J.M.

Peer review information Nature Communications thanks Frank Sobott and other, anonymous, reviewers for their contributions to the peer review of this work. Peer review reports are available.

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