Lena K Oesterlin
email: lena.oesterlin@curie.fr

3,4)P 2 : Phosphatidylinositol 3,4-biphosphate PI(3,4,5)P 3 : Phosphatidylinositol 3,4,5-triphosphate PI(4)P: Phosphatidylinositol 4-phosphate PI(4,5)P 2 : Phosphatidylinositol 4,5-biphosphate

Keywords: Amphipathic helix Biotin carboxyl carrier protein BMP: Bis(monoacylglycero)phosphate 1, 2-Dihexadecanoyl-sn-glycero-3-phosphoethanolamine (triethylammonium Protein-lipid interaction by fluorescence PMSF: Phenylmethylsulfonyl fluoride Photomultiplier tube Membrane order, curvature sensing, lipid packing defects, prenylation, geranylgeranyl

Vesicular trafficking between intracellular compartments is regulated by many proteins such as RAB GTPases. Deciphering the molecular mechanisms governing RAB localization is thus critical to understand intracellular transport. We focused our study on the role of the physicochemical properties of membranes in the specific recruitment of RAB proteins. Using in vitro reconstitution, we demonstrate that a balance between electrostatic interactions and hydrophobic insertion of the RAB C-terminal prenyl group into lipid packing defects controls the recruitment of RAB proteins.

famous pharmaceutical company (my initials might help to figure out which one it is…). It was not an easy decision to make. I made this famous list of pros and cons, discussed with many people, but I was still not able to make up my mind. One conversation that I will never forget is the one that I had with my parents. They made me realize that I had been talking about doing a PhD for months, even years, and that when hesitating between two alternatives, it is often better to go with the first one. I did not have the feeling that my parents were pushing me into doing anything, they would've been happy for me either way. But these words made all the difference and on the 3 rd of March 2014 I started in Bruno's lab as an engineer, and a couple months later started my PhD.

First, I would like to thank Bruno Goud for welcoming me in his lab and for his supervision during the last three years and a half. Bruno was always available to answer my questions, to discuss about the project and to share his knowledge. He greatly contributed to the improvement of my scientific skills and made me (I think) a better scientist.

I also greatly acknowledge Lena Oesterlin for her supervision and support. In the past years we had our differences, did not always agree on everything, but I am confident that it made me stronger as a scientist and as a person. In the end, I think we can agree that what we accomplished was very exciting and successful.

I would like to deeply acknowledge the members of my thesis committee Sandrine Etienne-Manneville, Aymelt Itzen and Jean-Claude Guillemot for fruitful discussions and great guidance.

Many thanks to my thesis Jury, Joelle Sobczack-Thepot, Jacqueline Cherfils, Cathy Jackson, Bruno Antonny and Aurélien Roux for the time they have spent reading my manuscript and for the very interesting discussion that (I hope) we will have on December 13 th . Doing a PhD is a very rewarding experience not only because you learn a lot as a scientist but also because you get to meet and interact with many people. I would like to thank Jean-Baptiste M., for all the help over the past years. I learned a lot from him concerning membranes, microscopy and many other things. He was always available to answer my many questions, to have a look at the thesis manuscript and to help with the scientific article.

I would also like to thank Stephanie (do not call her "Steph", she hates it), not only for the critical reading of this manuscript, but also for being who she is. Stephanie and I have had many scientific and less scientific discussions. Surprisingly, Star wars and Game of thrones were recurrent subjects. I hope I will find someone up to the task to debrief the last season (damn need to wait until 2019!).

Great thanks to Sabine, the pillar of the lab (what am I saying, of the whole floor!), the one person that makes everything hold. I do not even know where to start... Thank you for all your advices, your support, for being here in good but also bad moments. You are a great person, full of humanity and respect.

All the other members of the Goud team have had such a good and positive influence on me.

Thank you all Kristine, Anahi, Samuel, Surya, Charlotte, Azumi and all the members of the Coudrier team for the nice atmosphere, and great exchanges especially during lab meetings.

I would like to take some time to point out some unexpected encounters with people who were colleagues in the beginning and now so much more. Bruno and Hugo, my buddies, thank you both for all the amazing moments playing shuffleboard and belote; and also for other let's say less serious moments... I think we can agree that shuffleboard is not my thing and that Bruno might need to improve his belote skills (maybe stop drinking while playing?). Hugo on the other hand will tell you that he is the 1992 world champion; still trying to figure out if this is true... Thanks to Hugo for all the protein purifications, none of this work would've been possible without your involvement. Thanks to Bruno and Laure for taking care of Tomy (it's my cat) when I was away, I am sure he will miss you guys! When arriving to this point, I can bet that some people will complain because I still did not acknowledge them. Girls, you brought me so much during the past years that it would just be impossible to not mention you! Imene, Alex, Melissa, Camilla and Lea, thank you all for your friendship and for supporting me during these last years. I still remember when I arrived at Curie and I would see Melissa and Lea taking a break, I did not know them but still thought "wow they look cool". It is only a couple months later, at the young scientist retreat in Aachen (thanks ADIC), that we really started talking and that I realized that the two of them would play an important role during my curie experience. I can now say that it has indeed been the case, especially Melissa who has been such a great and important support during good but also bad moments. Camilla was already in the lab when I arrived and it did not take me much time to realize what a great, selfless and strong person she is, always smiling and joyful. I am pretty sure she will always think about me as long as the "Dyson" brand exists and when going to the Spa… When I arrived in the lab, I was appointed an office which was supposed to be provisional. What I did not know at this time is that I would never leave it. I met Imene (aka "the dragon"), my break companion, with whom the vibe felt right straight from the beginning. You taught me so much and always pushed me to go further so thank you! People came and went through the office, and it will be difficult to acknowledge them all. I will, however, remember the 18 th of January 2016, the day when Alexandra arrived and sat on the desk just next to mine. This date marks the beginning of a very intense (maybe a bit unhealthy?) relationship. My grandmother often tells me "qui aime bien châtie bien" and I think this sentence is in this case very appropriate. Alexandra has a very honest and definite opinion about things, and this is a quality that I admire. Thanks Alex for all your advices, your help and for being so patient with me while I was writing this manuscript.

Again, thank you girls for all the awesome moments that we shared together, weekends, parties and all the ADIC (association des doctorants, post-doctorants et jeunes chercheurs de l'institut Curie) events. It's been great! Many other people greatly contributed to the good and effective working environment. Thanks to all the Tran/Paoletti team and especially those with whom I shared the office. Ana and Lara, thank you for all the laughter, helping me with the abstract shortening, making sure that I drink enough water and for always offering food.

I am also grateful to Aurelie and Edwige for all their help with administrative matters and their kindness. I would also like to thank the laverie, and especially Chedly for his good mood and for making sure the lab functioned properly.

Over my time at Curie, I had the opportunity and the pleasure to meet many people from other departments. I spent a great amount of time in the Bassereau lab mostly preparing vesicles; and this allowed me to exchange a lot with them scientifically but also personally. I would like to thank Patricia Bassereau for welcoming me in her lab, for stimulating discussions and for inviting me to her legendary parties; it made me feel like a true member of the Bassereau team. Big thanks to Julien, Ajay, Thibaut, Nicola, Alex, Kodjo, Stephanie and Alicia for the great atmosphere and laughter. I would especially like to acknowledge Maryam and Feng. I don't even know how many times I annoyed you with my questions, but you were always inclined to help me and to teach me everything there is to know about tube pulling. As I like to tell them, "physicists rock!!". It would not have been possible to keep a sane mind throughout this PhD, without the presence of let's say non-scientific friends. I would like to thank my high school friends the "Croustibat" crew, Mike, François, Simon, Romain, Samy, Lucas and Dimitry; and also the girls, Laura, Cha, Alex, Camille and Loulou. I feel so lucky to have such amazing and lasting friends. It is great to see that things don't change and that the distance does not prevent us from regularly seeing each other.

One other thing that hasn't changed in the past years is that I still am a bad loser. I am sure Fanny and Thibaut will agree with this! Thank you both for the great times that we spent together, playing board games, the treasure hunts in Paris and going out. I will definitely miss you once I leave Paris.

Over the years, I realized that work is not all (I think most people will agree with me), and that it is important to get involved in other activities. Thursday night was football night with A Pucchiacca international and it was a bad idea to miss it unless you wanted to suffer Jean-Baptiste's wrath! Thank you guys, even though we have lost many games (probably more losses than victories), we never gave up. Sweat, blood and eternal glory! My involvement in the ADIC also represented an important part of my life as a PhD student. I would like to acknowledge all the past and current members of the association for the great and sometimes complicated times that we spent trying to make things work! Great thanks to my family, and especially my parents. They have been so supportive not only throughout this PhD, but during my whole life. Big up to my two brothers Matthieu and François and their respective wives (Flore do not freak out) just because they are the best. Particular thoughts for my goddaughter who is the cutest child ever! I would also like to address special thanks to my godfather and godmother for inviting us over for dinner so many times.

Finally, it is said that behind every (great) man there is a woman, and I am no exception. I am sure that Louise will want to read what I wrote and for this reason I will continue in French. En 2014, lorsqu'il a fallu que je déménage à Paris tu n'as pas hésité à me suivre. Tu travaillais depuis à peine trois mois, nous n'avions jamais habité ensemble et pourtant, sans aucun doute, tu as foncé.

Comme j'aime te le dire tu as été mon roc pendant ces dernières années, un soutien crucial dans des moments de stress et de doutes, et d'une grande patience surtout lors de l'écriture de ce manuscrit. Pour tout cela je te remercie.

From cell discovery to membrane binding properties of RAB GTPases

The cell is known as the smallest functional unit of all living organisms. It was first discovered in 1665 by Robert Hooke who called it "cell" from the Latin word cellula, meaning a small room. It was only in the middle of the 19 th century that the "cell theory" was developed by Theodor Schwann, Jakob Schleiden and Rudolf Virchow, stating that all organisms are composed of at least one cell and that cells originate from preexisting ones. In the late 19 th century, the discovery of the Golgi apparatus and the endosomes by Camillo Golgi in 1898 and Ilya Metchnikoff in 1887, respectively, led to the finding that cells are divided into sub-cellular compartments. Other intracellular compartments such as lysosomes, mitochondria and the endoplasmic reticulum (ER) were discovered by Christian de Duve, George Palade and Albert Claude in the middle of the 20 th century with the emergence and development of new techniques such as ultra-centrifugation and electron microscopy. The isolation of these different compartments demonstrated that they have different lipid and protein compositions, therefore suggesting that they also have different functions. A new view of intracellular architecture had emerged: eukaryotic cells are constituted of multiple intracellular compartments called organelles. These different organelles are delimited by membranes and each of them has a specific and unique composition and function, thereby allowing the compartmentalization of specific reactions and an increased complexity as compared to prokaryotic cells.

Even though cells are well compartmentalized, the different intracellular organelles are not isolated structures fulfilling independent functions. They are, on the contrary, constantly communicating between each other through exchange of intracellular components such as proteins and lipids. The first pathway to be described was the secretory pathway: Proteins are synthetized in the ER, enter the Golgi on its cis side and are finally released from the Golgi in secretory vesicles or granules that are then released out of the cell [START_REF] Duve | Exploring cells with a centrifuge[END_REF][START_REF] Palade | Intracellular aspects of the process of protein synthesis[END_REF]. Up to date, many other trafficking routes have been identified. Endocytosis was first discovered by Ilya Metchnikoff in 1883 (at the time known as the "phagocytosis theory"), but was only described in more detail decades later as the process by which molecules can get internalized into the cell (see [START_REF] Schmid | Endocytosis: Past, present, and future[END_REF] for extensive review). Other retrograde pathways from the plasma membrane to the Golgi and to the ER have been described as well [START_REF] Johannes | Tracing the retrograde route in protein trafficking[END_REF].

Intracellular trafficking was proposed, based on electron microscopy studies [START_REF] Palade | Intracellular aspects of the process of protein synthesis[END_REF][START_REF] Roth | Yolk Protein Uptake in the Oocyte of the Mosquito Aedes Aegypti[END_REF] to occur by encapsulation of secretory proteins into small vesicles and their transport from a donor compartment (fission process) to an acceptor compartment (fusion process).

Among others, James Rothman, Randy Sheckman and Thomas Südhof, who were awarded the 2013 Nobel Prize in Physiology or Medicine, spent decades trying to unravel the molecular mechanisms of intracellular transport. Their studies led to a deep understanding of vesicular transport with the identification of coat proteins involved in vesicle budding and specific selection of cargos and the discovery of SNARE proteins that allow vesicle fusion with the acceptor compartment and the following release of the cargos [START_REF] Bonifacino | Vesicular transport earns a Nobel[END_REF].

Intracellular trafficking consists in the sequential action of multiple sets of proteins to allow the exchange of molecules from one compartment to another. Small GTPases such as RAB proteins are key components of this process. The first RAB gene YPT1 (in Saccharomyces cerevisiae) was discovered in 1983 by Gallwitz and coworkers [START_REF] Gallwitz | A yeast gene encoding a protein homologous to the human c-has/bas proto-oncogene product[END_REF], and many other RAB genes were identified in the following decade with the completion of the yeast and human genomessequencing projects. The RAB family of proteins is now composed of over 60 members. RAB proteins were rapidly described to localize to the cytosolic face of most intracellular organelles (compartmental membranes but also transport vesicles) and to be implicated in the secretory and endocytic vesicular transport pathways [START_REF] Martinez | Rab proteins[END_REF]. RABs were shown to be physically attached to intracellular compartments through the insertion of their prenyl group, a lipid moiety at their C-terminus consisting of one or two geranylgeranyl groups [START_REF] Casey | Protein prenyltransferases[END_REF].

RAB proteins have been shown to localize to specific compartments, thereby defining organelle identity [START_REF] Zerial | Rab proteins as membrane organizers[END_REF][START_REF] Zhen | Cellular functions of Rab GTPases at a glance[END_REF]; and this is thought to be crucial for the control and directionality of vesicular trafficking. The mechanisms regulating specific RAB membrane targeting and localization are thus of great interest to understand intracellular transport events. The question of RAB protein targeting to specific membranes was first addressed 25 years ago and many teams have tried to answer this question. Originally, the Cterminal hypervariable region of RAB proteins was suggested to contain specific targeting information [START_REF] Chavrier | Hypervariable C-terminal domain of rab proteins acts as a targeting signal[END_REF]. Other studies later revised this statement by suggesting that multiple sequence motifs of RAB proteins, including this hypervariable region, can contribute to their specific membrane targeting (Ali et al., 2004;[START_REF] Beranger | The effector domain of Rab6, plus a highly hydrophobic C terminus, is required for Golgi apparatus localization[END_REF]. These motifs were mostly described to mediate RAB specific membrane localization via their interaction with RAB interacting proteins such as RAB GEFs (Blumer et al., 2013) or effector proteins [START_REF] Aivazian | TIP47 is a key effector for Rab9 localization[END_REF][START_REF] Wu | Structural basis for recruitment of RILP by small GTPase Rab7[END_REF].

Most studies so far have focused on the role of RAB interacting proteins in the mechanisms allowing the targeting of RAB proteins to specific membranes; but the role of the physicochemical properties of the membranes themselves has remained largely unexplored. It is however now well established that intracellular membranes are fluid environments that display different compositions and structures. These divergent properties can be exemplified by the formation of nanodomains of lipids or by membrane curvature, important features for the recruitment of some peripheral membrane proteins. Membrane charge, which is specific of endocytic compartments and the plasma membrane, is also a key feature regulating the specific recruitment of Cterminally positively charged proteins [START_REF] Heo | PI(3,4,5)P3 and PI(4,5)P2 lipids target proteins with polybasic clusters to the plasma membrane[END_REF].

I thus focused my study on the physicochemical membrane properties regulating the recruitment of RAB proteins. This issue being difficult to assess inside the cell, I have used an in vitro approach consisting of purified and prenylated RAB proteins and giant unilamellar vesicles (GUVs) as model membranes.

I started my study with RAB1 and RAB6 which associate with pre-Golgi and Golgi/trans Golgi network membranes respectively, RAB5 which is present on early endosomes and RAB35 which localizes to recycling endosomes and to the plasma membrane (Chapter 4).

RAB4 and RAB11 were shown to localize to tubular structures on endosomes (Sonnichsen et al., 2000) suggesting that curvature might play an important role in their specific localization. Their specific membrane recruitment however proved to be more complex and this will be addressed in Chapter 5.

Finally, previous studies in the lab had led to the unexpected observation that RAB6 incorporation into GUVs can induce membrane tethering. I also got to investigate this process during my PhD and detailed information about the possible mechanism behind this effect will be found in Chapter 6.

Chapter 1

1 Biology and physics of membranes The first part of this chapter will focus on presenting membranes, how they are formed and how the very heterogeneous compositions of cellular membranes can lead to differences in their physicochemical properties. Because membranes are complex systems composed of many different molecules (proteins, lipids), their study in cells remains very challenging. Thus, in the second part of this chapter I will present multiple in vitro experimental model membranes that can be used to mimic in a more simplistic way the different properties of membranes.

The Lipid bilayer

Eukaryotic cells are divided into compartments. These compartments, as well as cells themselves, are delimited by membranes. In the middle of the 20 th century, the membrane was described as a two dimensional fluid structure (the lipid bilayer) which led to the fluid mosaic model (Singer and Nicolson, 1972). In this model, the membrane is a fluid matrix of lipids in which peripheral and integral proteins diffuse. Since then, this model has been refined and it appears that lipids are not immobile structures holding proteins together. Membranes are crowded and heterogeneous environments with lipids and proteins diffusing laterally allowing the formation of regions which vary in thickness and composition (Engelman, 2005).

From a lipid molecule to a bilayer

The basic components of membranes are lipids. Lipid molecules are amphiphilic and are thus composed of a hydrophilic part, the head, and a hydrophobic part, the tail. The hydrophilic head of the lipid defines the lipid type and can be neutral or charged. The hydrophobic tail is most often made of two aliphatic chains (can vary from 1 to 4 chains) of various length and degree of unsaturation. Due to their amphiphilic structure, lipids have the property to self-assemble in such a way that the heads are accessible to the solvent, whereas the tails (more hydrophobic) are buried in the core of the membrane. This spontaneous rearrangement of the lipids is of entropic origin and the result of a competition between the hydrophobic attraction (also called hydrophobic effect), which tends to aggregate the molecules together therefore reducing the interfacial area; and the repulsion of the hydrophilic head groups which tends to increase the interfacial area [START_REF] Israelachvili | Intermolecular and surface forces[END_REF].

This competition between hydrophobic attraction and headgroup repulsion results in a constant area per lipid. Depending on this parameter as well as on the geometrical shapes of lipids, lipid assemblies will exhibit different morphologies (Figure 1 It is important to mention that the shape (also called packing parameter) of a particular lipid is not fixed. External parameters such as hydration, temperature and pH can modulate the headgroup effective area as well as the apparent chain volume [START_REF] Pomorski | Model cell membranes: discerning lipid and protein contributions in shaping the cell[END_REF].

Different classes of lipids

The Polar lipids from the glycerophospholipid family are the main eukaryotic membrane lipids. They are based on a diacylglycerol backbone supplemented by a phosphate group (cone-shaped phosphatidic acid, PA) (Figure 1.2). Phosphatidylcholine (PC), the major species of this family is formed by the addition of a choline on top of the PA. PC lipids are zwitterionic (contain both a positive and a negative charge at physiological conditions) and usually have a rather cylindrical shape. Other groups can be added instead of the choline to form a phosphatidylethanolamine (PE), a species with a conical shape due to the small size of its headgroup. Phosphatidylserine (PS) is a negatively charged and cylinder-shaped phospholipid mainly concentrated in the inner (cytosolic) leaflet of the plasma membrane. Phosphatidylinositol lipids (PI) are present in smaller amounts and bear a negative charge. PI lipids are known to be phosphorylated by several different kinases and their derivatives are known to be involved in a multitude of signaling processes, mainly at the plasma membrane and at the endocytic compartments (Di Paolo and De Camilli, 2006;van Meer et al., 2008).

Sphingolipids have a ceramide (Cer) backbone (sphingosine base amid-linked to a fatty acid) (Figure 1.2). Sphingomyelin (SM) is the most abundant species of this family of lipids, and is composed of a phosphate-choline headgroup. Another important class of ceramide based lipids is the Glycosphingolipids (GSLs), consisting of a ceramide molecule attached to monosaccharides or polysaccharides. SM and GSL are found on the non-cytosolic (extracellular or luminal) leaflet of the plasma membrane (van Meer et al., 2008).

Sterols constitute another major class of lipids present in cellular membranes. The presence of the OH group on the lipid headgroup suggests that they are slightly polar and their specific structure suggests that they are non-bilayer forming molecules (even though they are able to insert in membranes). Cholesterol (Figure 1.2) is the major sterol present in mammalian cells and its level increases from the ER to the plasma membrane [START_REF] Ikonen | Cellular cholesterol trafficking and compartmentalization[END_REF][START_REF] Mesmin | Intracellular sterol dynamics[END_REF].

Lipid synthesis and distribution in cells

Lipids are not homogeneously distributed across cellular membranes (Figure 1.3). Additionally, the two leaflets of the Golgi, endosomal and plasma membranes all exhibit asymmetric lipid compositions [START_REF] Devaux | Static and dynamic lipid asymmetry in cell membranes[END_REF][START_REF] Verkleij | The asymmetric distribution of phospholipids in the human red cell membrane. A combined study using phospholipases and freeze-etch electron microscopy[END_REF][START_REF] Wood | Cholesterol asymmetry in synaptic plasma membranes[END_REF].

The endoplasmic reticulum (ER) is the main lipid and protein biosynthetic organelle [START_REF] Bell | Lipid topogenesis[END_REF]. The presence of ribosome complexes, which gives its rough aspect to the ER, is responsible for protein synthesis. This compartment is mostly composed of Glycerophospholipids (PC and PE). The ER produces the bulk of structural phospholipids and cholesterol. Ceramide, the precursor of complex sphingolipids is also produced at the ER. Even though sterols and complex sphingolipid precursors are synthesized in the ER, these products are rapidly leaving this compartment and are transported to other organelles via vesicular transport but also via a nonvesicular route involving membrane contact sites [START_REF] Jackson | Lipids and Their Trafficking: An Integral Part of Cellular Organization[END_REF] (also discussed in chapter 2). The Golgi apparatus is a lipid-based sorting station. It is composed of several compartments called cisternae. The cisterna on the ER side is called the cis-Golgi and the cisterna on the plasma membrane side is called the trans-Golgi. The main processes that take place at the Golgi apparatus are the post-translational modifications of newly synthesized proteins coming from the ER (known as protein maturation). The Golgi is the place where significant level of lipid synthesis occurs. This compartment is specialized in the synthesis of complex sphingolipids (SM, GSLs) [START_REF] Futerman | The ins and outs of sphingolipid synthesis[END_REF]. PC and PE synthesis can also take place at the Golgi apparatus.

Cholesterol levels are higher as compared to the ER membrane. The PI derivative PI(4)P, which acts as a signaling lipid, is enriched at the trans-Golgi network.

The plasma membrane has very different lipid content as compared to the other intracellular membranes (Figure 1.3). It is enriched in sphingolipids and sterols which are packed at higher density than glycerophospholipids and can resist mechanical stress. The plasma membrane bilayer is highly asymmetric as the outer leaflet mainly contains SM and PC while the inner leaflet contains mostly PE, PS but also the PI derivative PI(4,5)P 2 (van Meer et al., 2008). The inner leaflet is thus highly negatively charged.

The plasma membrane is not involved in the autonomous synthesis of structural lipids but numerous PI lipid derivatives (PI(3,4)P 2 , PI(4,5)P 2 , PI (3,4,5)P 3 , PI(4)P), involved in signaling cascades, were shown to be synthesized or degraded there (Di Paolo and De Camilli, 2006). SM synthesis was also shown to take place at the plasma membrane [START_REF] Tafesse | Both sphingomyelin synthases SMS1 and SMS2 are required for sphingomyelin homeostasis and growth in human HeLa cells[END_REF].

While the composition of early endosomes is very similar to that of the plasma membrane, late endosomes have a quite different composition as the amounts of sterols and PS decrease whereas the levels of bis(monoacylglycero)phosphate (BMP) increase [START_REF] Kobayashi | Separation and characterization of late endosomal membrane domains[END_REF]. BMP is a cone-shaped and negatively charged lipid which was shown to act in multivesicular body generation (endosome containing internal vesicles that originate from inward budding), fusion processes and sphingolipid hydrolysis [START_REF] Gallala | Biological function of the cellular lipid BMP-BMP as a key activator for cholesterol sorting and membrane digestion[END_REF][START_REF] Matsuo | Role of LBPA and Alix in multivesicular liposome formation and endosome organization[END_REF]. The endocytic compartments recruit specific sets of kinases and phosphatases that allow the regulation of phosphoinositide content (Di Paolo and De Camilli, 2006). Thus, early endosomes are composed of PI(3)P, whereas PI(3,5)P 2 is mainly found on late endosomes.

As a general important comment, the PI lipid derivatives previously mentioned act as signaling lipids which allow membrane identification and subsequently the recruitment of cytosolic proteins involved in vesicular transport (Di Paolo and De Camilli, 2006).

Membrane domain formation 1.2.1 Different states of membranes

As previously mentioned, cellular membranes are composed of mixtures of many different lipid species which display various geometrical shapes. This lipid structure is dependent on their physical properties (size of the headgroup, degree of unsaturation, aliphatic chain length) but also on external parameters such as temperature or hydration [START_REF] Pomorski | Model cell membranes: discerning lipid and protein contributions in shaping the cell[END_REF]. As any physical system, lipid bilayers can exist in different phases depending on the overall structures of their lipid content.

The membrane is fluid at high temperature and in different liquid-crystal phases at lower temperatures [START_REF] Los | Membrane fluidity and its roles in the perception of environmental signals[END_REF]. The two most extreme phases are the gel or solid ordered phase (So) and the liquid disordered phase (Ld) (Figure 1.4). In the So phase, the lipid acyl chains can undergo trans-isomerization which leads to their extension and more Van der Waals interactions. Stronger interactions lead to more ordered lipid packing which prevents any lateral lipid diffusion [START_REF] Seu | Influence of lipid chemistry on membrane fluidity: tail and headgroup interactions[END_REF]. The Ld phase is usually characterized by the presence of unsaturated lipids (one or more double bonds in the acyl chain) and their irregular packing.

Unsaturation of the acyl chains leads to kinks in their structure which reduces the surface area accessible to other lipids and thus weakens Van der Waals interactions. Consequently, the Ld phase is a highly fluid state in which individual lipids can freely diffuse [START_REF] Seu | Influence of lipid chemistry on membrane fluidity: tail and headgroup interactions[END_REF].

Under physiological conditions, intracellular bilayers tend to exist in a fluid phase and can undergo phase transition under correct environmental conditions. The temperature at which a membrane lipid can undergo phase transition from the gel to the liquid state is the melting temperature (Tm). The Tm can vary between lipids due to their different structural properties (acyl chain length and degree of unsaturation) [START_REF] Ceccato | How membrane chain-melting phase-transition temperature is affected by the lipid chain asymmetry and degree of unsaturation: an effective chain-length model[END_REF]. Lipids that exhibit longer acyl chains will have higher surface areas as compared to lipids exhibiting smaller chain length, resulting in stronger Van der Waals interactions between aliphatic chains and thus increased Tm. As previously mentioned, increasing unsaturation leads to weaker Van der Waals interactions and thus to lower Tm of the lipid. As the Tm strongly depends on the amount of unsaturation and on the length of the acyl chains, one can thus make the distinction between high Tm lipids that will be in a solid state at physiological conditions, and low Tm lipids that will be in a liquid state under the same conditions.

As an example, sphingolipids usually carry saturated or trans-unsaturated (linear) aliphatic chains whereas the acyl chains of glycerophospholipids are often unsaturated. Membranes composed of sphingolipids therefore adopt a more tightly packed structure (solid-like phase) as compared to glycerophospholipids containing membranes that form less ordered domains (L d phase). Cholesterol is another key component of eukaryotic cellular membranes and was shown to drastically affect the physical properties of membranes, through lateral order disruption of the gel phase and ordering of the Ld phase [START_REF] Henriksen | Universal behavior of membranes with sterols[END_REF][START_REF] Ipsen | Phase equilibria in the phosphatidylcholine-cholesterol system[END_REF]. The presence of cholesterol in membranes can lead to the appearance of an intermediate state between gel phase and Ld, the liquid ordered (Lo) phase (Figure 1.4). This Lo phase is characterized by a tight lipid packing, as in the gel solid-ordered phase, but also by a rapid lateral diffusion rate, as in Ld membranes (London, 2002;[START_REF] 'baye | Liquid ordered and gel phases of lipid bilayers: fluorescent probes reveal close fluidity but different hydration[END_REF]. Lo membranes are usually thicker, stiffer and less permeable than Ld membranes [START_REF] Rawicz | Elasticity, strength, and water permeability of bilayers that contain raft microdomain-forming lipids[END_REF]. The lateral diffusion coefficient is 2-3 fold less in the Lo phase as compared to the Ld phase [START_REF] Veatch | Seeing spots: complex phase behavior in simple membranes[END_REF].

Phase state of cellular membranes

Cellular membranes display inhomogeneous lipid distribution at different levels (van Meer et al., 2008). The lateral heterogeneity in cellular membranes can be described by the lipid raft hypothesis [START_REF] Simons | Functional rafts in cell membranes[END_REF] which has been a matter of great debate in the last 20 years. In biomembranes, lipid rafts, which are characterized by a tight packing of saturated lipids and cholesterol, likely exist in a Lo state and behave like islands floating in a sea of loosely-packed Ld domains of unsaturated glycerophospholipids [START_REF] 'baye | Liquid ordered and gel phases of lipid bilayers: fluorescent probes reveal close fluidity but different hydration[END_REF]. Lipid rafts have received a huge attention as they are believed to be involved in many cellular processes such as signal transduction, lipid trafficking and regulation of membrane protein activity [START_REF] Jacobson | Lipid rafts: at a crossroad between cell biology and physics[END_REF]Lingwood and Simons, 2010;[START_REF] Owen | The lipid raft hypothesis revisited--new insights on raft composition and function from super-resolution fluorescence microscopy[END_REF].

Another less described heterogeneity is related to the differential composition and organization of the plasma membrane as compared to intracellular membranes. Cholesterol and sphingolipid levels were found to be increased from the ER to the plasma membrane and this seems to be due to the directed anterograde transport and the absence of retrograde transport of these lipids [START_REF] Brugger | Evidence for segregation of sphingomyelin and cholesterol during formation of COPI-coated vesicles[END_REF][START_REF] Klemm | Segregation of sphingolipids and sterols during formation of secretory vesicles at the trans-Golgi network[END_REF]. As previously mentioned, even though raft components are synthesized in the ER and the Golgi complex, they are quickly leaving these compartments and are transported towards the plasma membrane. As an example, sphingomyelin is mainly found at the plasma membrane even though ceramide, the hydrophobic backbone of sphingolipids, is synthesized in the ER and the assembly of the sphingolipids headgroups to ceramide takes place at the Golgi complex [START_REF] Van Meer | Sphingolipid transport: rafts and translocators[END_REF]. Thus, lipid raft components are moved toward the plasma membrane where they concentrate but also spread into the endocytic recycling pathways [START_REF] Mukherjee | Role of membrane organization and membrane domains in endocytic lipid trafficking[END_REF]. This seems to be consistent with studies showing that more than 60% of all cellular cholesterol is located to the plasma membrane whereas intracellular membranes such as the ER or the Golgi membranes exhibit low levels of cholesterol [START_REF] Ikonen | Cellular cholesterol trafficking and compartmentalization[END_REF][START_REF] Mesmin | Intracellular sterol dynamics[END_REF].

As cholesterol is known to promote phase separation [START_REF] Ohvo-Rekila | Cholesterol interactions with phospholipids in membranes[END_REF][START_REF] Silvius | Cholesterol at different bilayer concentrations can promote or antagonize lateral segregation of phospholipids of differing acyl chain length[END_REF], this process is thought to mainly occur at the cholesterol-rich plasma membrane. Multiple studies, in particular the ones using fluorescent probes such as Laurdan [START_REF] Bagatolli | To see or not to see: lateral organization of biological membranes and fluorescence microscopy[END_REF][START_REF] Owen | The lipid raft hypothesis revisited--new insights on raft composition and function from super-resolution fluorescence microscopy[END_REF] and its derivatives [START_REF] Kim | Two-photon fluorescent turn-on probe for lipid rafts in live cell and tissue[END_REF][START_REF] Sezgin | Measuring lipid packing of model and cellular membranes with environment sensitive probes[END_REF] have focused on lipid raft organization in model and cell membranes. These probes have the ability to change fluorescence color and intensity in response to changes in membrane hydration and solvent relaxation [START_REF] Demchenko | Monitoring biophysical properties of lipid membranes by environment-sensitive fluorescent probes[END_REF], two parameters linked to lipid order.

Niko and coworkers recently developed a new probe based on pyrene which demonstrates enhanced photophysical properties over Laurdan (Niko et al., 2016). Their study confirms clear differences in lipid order among the different cellular membranes with the plasma membrane mainly composed of Lo domains while intracellular membranes are much closer to Ld phases. This is consistent with the higher amount of sphingomyelin and cholesterol, both responsible for the formation of lipid rafts, at the plasma membrane.

Membrane deformations

Biological membranes are two-dimensional surfaces with two principal curvatures C 1 = 1/R 1 and C 2 = 1/R 2 (with R 1 and R 2 referred to as the principal radii of curvature) along two perpendicular directions (Zimmerberg and Kozlov, 2006) (Figure 1.5). The total curvature of the membrane is

C = C 1 +C 2 .
In the case of a spherical vesicle of radius R, the membrane deforms equally in both directions leading to C 1 = C 2 = 1/R and a total curvature C v = 2/R. In the case of a cylindrical tube of radius R, which is curved only in one direction and flat in the other, C 1 > 0 and C 2 = 0 yielding a total curvature C t = 1/R. When proteins interact with the membrane, it is said that a protein can sense positive curvature if it senses the convex side of the membrane whereas the curvature is negative if it senses its concave side.

Lipid bilayers have an average thickness in the order of 4-5 nm [START_REF] Marquardt | Lipid bilayer thickness determines cholesterol's location in model membranes[END_REF]. Logically enough, this value also corresponds to the lower limit of radii to which a bilayer can be bent.

Thus, in the perspective of a protein, if a membrane has a curvature radius in the same range (10-50 nm) it will be considered as highly curved whereas if the radius is superior to 50 nm it will be considered as weakly curved.

Membrane curvature in cells

Due to their heterogeneous lipid and protein compositions, intracellular compartments exhibit membrane regions with both high and low curvature (Figure 1.6). For example, the ER and the budding processes, which displays two positive curvatures on the spherical vesicle part but also a negative curvature at the neck of the membrane bud. Adapted from (Zimmerberg and Kozlov, 2006).

Golgi form complex networks of interconnected flat sheets and highly curved tubules (30-60 nm diameter) (Shibata et al., 2009;Voeltz and Prinz, 2007). Mitochondria also display many curved invaginations (30 nm in diameter), called cristae, which are directed towards the mitochondrial matrix and thus greatly increase the total surface area necessary for chemical reactions (Voeltz and Prinz, 2007). Endosomes were also shown to exhibit tubular regions of high curvature and globular regions of low curvature (Sonnichsen et al., 2000).

These tubular structures are often highly dynamic. They can undergo continuous fission and fusion events and get continuously rearranged by moving along the actin or microtubule cytoskeleton. Particular membrane budding can also take place during the formation of multivesicular bodies (MVBs) [START_REF] Hurley | Membrane budding[END_REF] (Figure 1.6). MVBs are late endosomes containing internal vesicles that sort membrane proteins destined for degradation into these vesicles. These internal vesicles are formed by invagination of the endosomal membrane followed by scission of the buds. In contrast to clathrin and COP-coated vesicles formation which consists in budding towards the cytosol (implying positive curvature for cytosolic proteins), the generation of MVBs consists in the membrane budding away from the cytosol which thus has an opposite negative curvature.

Enveloped viruses release is also occurring away from the cytosol which also implies negative curvature. ESCRT (endosomal sorting complexes required for transport) proteins were described to be deeply involved in both MVB formation and virus budding processes [START_REF] Babst | MVB vesicle formation: ESCRT-dependent, ESCRT-independent and everything in between[END_REF][START_REF] Votteler | Virus budding and the ESCRT pathway[END_REF].

Intracellular trafficking also involves curved membranes (Figure 1.6) as it consists in the transport of highly curved transport vesicles of 40-80 nm in diameter but also in membrane deformation events allowing fission of these vesicles from the donor compartment and their subsequent fusion with the acceptor compartment [START_REF] Takamori | Molecular anatomy of a trafficking organelle[END_REF][START_REF] Bonifacino | The mechanisms of vesicle budding and fusion[END_REF]. This vesicle transport process will be further discussed in Chapter 2.

Mechanisms of membrane deformation

Membrane deformation can be achieved through the alteration of membrane properties in an asymmetric manner or by applying unilateral constraints to it. In other words, membrane curvature can be dynamically regulated not only by changes in lipid composition but also through the action of external factors such as the cytoskeleton or proteins that can directly get inserted in membranes and act as wedges or that can form scaffolds around the membrane [START_REF] Mcmahon | Membrane curvature and mechanisms of dynamic cell membrane remodelling[END_REF]. Figure 1.7 gives an overview of biologically relevant ways to deform membranes and each of them will be further discussed in the next sections.

Lipid dependent membrane deformations

As a general mechanism, bilayers that have symmetrical lipid content between the two leaflets and similar interacting environments should remain flat. However, if some degree of asymmetry is introduced into the system (a difference in composition between the leaflets), the bilayer acquires a spontaneous curvature.

As previously mentioned, depending on their geometrical properties, lipids will self-assemble into different structures. Even if they cannot form bilayers on their own, non-cylinder shaped lipids (conical or inverted-conical) can still be incorporated in these structures. In this case they will induce a spontaneous curvature of the monolayer they are embedded in (Figure 1.8). However, if these lipids are homogeneously distributed between the two monolayers (symmetrical bilayer) the spontaneous curvature of both leaflets will cancel each other (Zimmerberg and Kozlov, 2006).

In order for spontaneous curvature to actually happen, the enrichment of a specific lipid in one of the monolayers is required. Adapted from [START_REF] Mcmahon | Membrane curvature and mechanisms of dynamic cell membrane remodelling[END_REF].

Membrane asymmetry can be caused by a directional transfer of lipids between the two membrane leaflets (also called transbilayer flip-flop) or by lipid modifying enzymes that can modify the chemical properties of lipids.

Lipid transfer between two monolayers can be spontaneous in the case of cholesterol or lipids exhibiting either a small or a non-polar headgroup such as DAG and PA [START_REF] Sprong | How proteins move lipids and lipids move proteins[END_REF]; and can also be assisted by proteins such as flippases [START_REF] Devaux | How lipid flippases can modulate membrane structure[END_REF].

Phospholipid flippases, such as Type IV-type ATPases (P4-ATPase), control the translocation of phospholipid molecules across the membrane [START_REF] Muthusamy | Linking phospholipid flippases to vesicle-mediated protein transport[END_REF]. More specifically, they have been shown to establish and maintain plasma membrane asymmetry by transferring PS and PE from the external leaflet to the cytosolic leaflet of the bilayer and to be involved in sterol trafficking and metabolism, and in vesicle-mediated protein transport [START_REF] Muthusamy | Linking phospholipid flippases to vesicle-mediated protein transport[END_REF]. Lipid modifying enzymes also have the ability to asymmetrically change the lipid composition of one monolayer and induce membrane curvature. Phospholipase A (PLA) hydrolyses the acyl chain of a phospholipid to produce lysophospholipid (Figure 1.8: red lipids) known to build positively curved membranes. PLA 2 for example has been shown to be involved in the formation of tubules from the Golgi and endosomes [START_REF] Brown | Phospholipase A2 (PLA2) enzymes in membrane trafficking: mediators of membrane shape and function[END_REF]. Sphingomyelinases release the phosphocholine headgroup of sphingomyelin (known to have no spontaneous curvature) and produce the ceramide lipid known to build negative curvature. This process was suggested to be used by cells to mediate exosome formation [START_REF] Trajkovic | Ceramide triggers budding of exosome vesicles into multivesicular endosomes[END_REF]. In contrast, phospholipase D (PLD) modifies the polar headgroup to yield cone-shaped PA, while phospholipase C (PLC) produces cone-shaped DAG (Figure 1.8: green lipids). These phospholipases were also shown to induce membrane deformations [START_REF] Holopainen | Macroscopic consequences of the action of phospholipase C on giant unilamellar liposomes[END_REF][START_REF] Inaba | Phospholipase Cbeta1 induces membrane tubulation and is involved in caveolae formation[END_REF].

However even though spontaneous curvature is known to exist in cells, it is not believed to play a major role in the generation of membrane curvature (Kozlov et al., 2014;Shibata et al., 2009).

Flippases for example are more effective if they localize to a small membrane area. If they act only on a small portion of a larger surface, such as the plasma membrane, their effect will be equilibrated on the entire surface and the resulting influence on membrane curvature would be negligible.

Membrane deformation was also observed in artificial membranes exhibiting distinct lipid domains (such as Lo and Ld). The boundary separating these domains has an energy cost, due to line tension, that the system would like to minimize. This is achieved with the Ostwald ripening [START_REF] Lifshitz | The kinetics of precipitation from supersaturated solid solution[END_REF], a process by which the size and shape of membrane domains is adjusted to minimize the energy associated to the surface tension. Over time, lipids diffuse and domains coalesce so that the number of domains constantly decreases whereas the average domain size constantly increases, ultimately resulting in a more energetically favorable complete phase separation (fully separated vesicle with two domains). The shape of such a vesicle is no longer spherical but is the result of a competition the bending energy and the line tension of the boundary separating the two domains. This effect was shown to lead to domain budding (Baumgart et al., 2003) and even to membrane fission (Roux et al., 2005).

Deformation induced by external objects

In the previous section, I focused on models where the membrane shape is determined by its lipid composition. However, other types of deformations exist in a biological context and consist in the interaction of membranes with external objects. For example, the generation of membrane carriers requires close and independent collaboration between the membrane itself and the cytoskeleton (filament polymerization), motor proteins and membrane shaping proteins (coat proteins for example) [START_REF] Anitei | Bridging membrane and cytoskeleton dynamics in the secretory and endocytic pathways[END_REF].

Cytoskeleton

The cytoskeleton is composed of networks of polymers and filaments involved in various biological processes. The actin and microtubule filaments, which are the best characterized, can generate pushing forces on cellular membranes (Figure 1.9A). It is well described that cell membranes can exhibit very thin extensions called filopodia or extended flat lamellipodia which are formed by different modes of actin polymerization and play a major role in cell migration. The formation of these structures requires multiple proteins such as proteins nucleating and elongating the filaments, protecting them from capping, and bundling them together [START_REF] Mattila | Filopodia: molecular architecture and cellular functions[END_REF]. It has been shown in vitro that actin filament and microtubule polymerization forces are able to tubulate membranes [START_REF] Limozin | On the organization of self-assembled actin networks in giant vesicles[END_REF][START_REF] Nomura | Morphological and topological transformation of membrane vesicles[END_REF], and actin polymerization has been suggested to be involved in the generation of tubular transport intermediates in the secretory and endocytic pathways [START_REF] Anitei | Bridging membrane and cytoskeleton dynamics in the secretory and endocytic pathways[END_REF]. filaments. Adapted from [START_REF] Gurel | Connecting the cytoskeleton to the endoplasmic reticulum and Golgi[END_REF].

Motor proteins that have the ability to "walk" on these filaments (myosins on actin filaments; and kinesins and dyneins on microtubules) can also play a role in the biogenesis of intracellular carriers at the ER or at the Golgi [START_REF] Gurel | Connecting the cytoskeleton to the endoplasmic reticulum and Golgi[END_REF]. Membrane-bound motor proteins have the possibility to induce membrane deformations by moving along the cytoskeleton tracks (Figure

1.9B and Figure 1.9C).

In vitro studies have also shown that microtubule-associated kinesins [START_REF] Koster | Membrane tube formation from giant vesicles by dynamic association of motor proteins[END_REF][START_REF] Leduc | Cooperative extraction of membrane nanotubes by molecular motors[END_REF][START_REF] Roux | A minimal system allowing tubulation with molecular motors pulling on giant liposomes[END_REF] and actin-associated Myosin II [START_REF] Carvalho | Actin polymerization or myosin contraction: two ways to build up cortical tension for symmetry breaking[END_REF] have the ability to deform liposomes by applying a pulling force on them.

Hydrophobic insertion or wedging

Certain peripheral or transmembrane proteins also have the ability to mold biological membranes through the insertion of hydrophobic or amphipathic domains. These protein insertions can be separated in two classes: shallow insertions, which are only incorporated in the external part of a monolayer and integral insertions that occupy the whole membrane space. Shallow insertions have been demonstrated to be much more powerful generators of membrane curvature [START_REF] Campelo | The hydrophobic insertion mechanism of membrane curvature generation by proteins[END_REF]. These proteins can generate membrane asymmetry through the hydrophobic insertion of hydrophobic or amphipathic domains (helix) into one monolayer. This process known as "wedging" was mostly described as the incorporation of a helix to about 30-40% of the monolayer thickness [START_REF] Campelo | The hydrophobic insertion mechanism of membrane curvature generation by proteins[END_REF]. The region of the lipid headgroups is expanded while the region of the acyl chains remains undisturbed. This leads to a strong membrane asymmetry and thus to the generation of positive membrane curvature.

To this class belong the amphipathic helices (AH) of the N-BAR domains of various proteins [START_REF] Frost | The BAR domain superfamily: membrane-molding macromolecules[END_REF][START_REF] Gallop | Mechanism of endophilin N-BAR domain-mediated membrane curvature[END_REF], of epsin N-terminal homology (ENTH) domains [START_REF] Ford | Curvature of clathrin-coated pits driven by epsin[END_REF] and of small G-proteins such as ARF1 and SAR1 [START_REF] Lee | Sar1p N-terminal helix initiates membrane curvature and completes the fission of a COPII vesicle[END_REF][START_REF] Lundmark | Arf family GTP loading is activated by, and generates, positive membrane curvature[END_REF].

N-BAR domain proteins such as amphiphysin or endophilin possess an AH at their N-terminal extremity and have been shown to be involved in membrane tubulation [START_REF] Gallop | Mechanism of endophilin N-BAR domain-mediated membrane curvature[END_REF][START_REF] Masuda | Endophilin BAR domain drives membrane curvature by two newly identified structure-based mechanisms[END_REF].

The ENTH domain of Epsin folds into an AH upon binding to PI(4,5)P 2 and was shown to tubulate synthetic vesicles [START_REF] Ford | Curvature of clathrin-coated pits driven by epsin[END_REF]. It is also known to play a role in clathrin-dependent endocytosis [START_REF] Boucrot | Membrane fission is promoted by insertion of amphipathic helices and is restricted by crescent BAR domains[END_REF][START_REF] Itoh | BAR, F-BAR (EFC) and ENTH/ANTH domains in the regulation of membrane-cytosol interfaces and membrane curvature[END_REF].

The small GTPases ARF1 and SAR1 are recruited to the cytosolic leaflet of membranes through the exposure of an AH. This process, induced by GTP binding, consists in the displacement of the Nterminal domain from the core of the protein and the subsequent availability of the AH for membrane binding. AHs of SAR1 and ARF1 were suggested to be important for vesicle budding, through the respective induction of COPII [START_REF] Lee | Sar1p N-terminal helix initiates membrane curvature and completes the fission of a COPII vesicle[END_REF] and COPI-coated vesicle curvature [START_REF] Beck | Membrane curvature induced by Arf1-GTP is essential for vesicle formation[END_REF].

Interestingly, a study from Stachowiak and coworkers demonstrates that membrane bending and tubulation can still occur when the AH of the ENTH domain of Epsin is removed and that moderate protein coverage is able to induce membrane bending independently of the wedge mechanism [START_REF] Stachowiak | Membrane bending by protein-protein crowding[END_REF]. Membrane curvature was thus suggested to be induced by hydrophilic proteins through a crowding mechanism. This process consists in the thermally induced lateral diffusion of proteins which collide with each other. The generated steric effect leads to a lateral pressure on the membrane which can be released through the formation of tubules.

Scaffolding

The binding of hydrophilic protein domains to the surface of one monolayer can also lead to membrane asymmetry and subsequently to membrane curvature.

The scaffolding mechanism consists in the transfer of curvature from the intrinsically curved interaction site of the protein to the underlying membrane. This curved interface can be preexisting in one protein monomer or be the result of protein oligomerization. Protein scaffolds can impose their curvature on membranes only if they can overcome the rigidity of the bilayer. Coat proteins such as COPI, COPII and clathrin are well documented protein coats that polymerize around nascent vesicles into a cage-like shell and induce membrane curvature [START_REF] Faini | Vesicle coats: structure, function, and general principles of assembly[END_REF].

COPII and COPI coats are respectively implicated in the anterograde pathway (export from the ER) and in the intra-Golgi and retrograde (Golgi to ER) pathways. Clathrin-mediated membrane deformations were shown to be involved in endocytosis (McMahon and Boucrot, 2011) but also in trafficking events at the TGN [START_REF] Antonny | Membrane deformation by protein coats[END_REF].

Reticulons and Caveolins are membrane-embedded proteins that were described to generate curvature through the formation of wedge-shaped insertions and oligomeric scaffolds [START_REF] Shibata | The reticulon and DP1/Yop1p proteins form immobile oligomers in the tubular endoplasmic reticulum[END_REF][START_REF] Walser | Constitutive formation of caveolae in a bacterium[END_REF]. Reticulons and Caveolins were respectively shown to be involved in the formation of tubules at the ER [START_REF] Voeltz | A class of membrane proteins shaping the tubular endoplasmic reticulum[END_REF] and the creation of plasma membrane invaginations [START_REF] Walser | Constitutive formation of caveolae in a bacterium[END_REF]. ESCRT proteins were also described to drive membrane deformation during MVB generation by polymerizing at the neck of budding vesicles [START_REF] Mccullough | Structure and membrane remodeling activity of ESCRT-III helical polymers[END_REF][START_REF] Wollert | Molecular mechanism of multivesicular body biogenesis by ESCRT complexes[END_REF]. Dynamin [START_REF] Ferguson | Dynamin, a membrane-remodelling GTPase[END_REF] and BAR domain proteins [START_REF] Masuda | Structural characteristics of BAR domain superfamily to sculpt the membrane[END_REF] also have the ability to deform membranes into tubules by acting as cylindrical oligomeric scaffolds.

Finally, shallow hydrophobic insertion and intrinsically curved protein scaffolds appear as the most effective generators of membrane curvature. However, even though these two mechanisms are different in their way to generate curvature, it is thought that they can still be complementary to each other. Indeed, as certain proteins, such as N-BAR, Reticulons or Caveolins, are involved in both scaffolding mechanism and hydrophobic insertion, membrane curvature generation could very well be the result of a combination of these two effects. Studies show that mutation of the SAR1 AH compromises COPII vesicle budding even though COPII was still able to deform liposomes [START_REF] Lee | Sar1p N-terminal helix initiates membrane curvature and completes the fission of a COPII vesicle[END_REF]. This suggests that the COPII mediated scaffolding mechanism is not sufficient to drive vesicle budding. Similar studies about ARF and its involvement in COPI-coated vesicle budding [START_REF] Beck | Membrane curvature induced by Arf1-GTP is essential for vesicle formation[END_REF][START_REF] Krauss | Arf1-GTP-induced tubule formation suggests a function of Arf family proteins in curvature acquisition at sites of vesicle budding[END_REF] show that inhibition of ARF1 dimerization prevents vesicle budding without disrupting COPI membrane interaction. Therefore, the intracellular generation of membrane curvature is probably the result of a concerted action of several protein-based mechanisms.

Protein curvature sensing

Membrane curvature is not only a consequence of the mechanical or enzymatic actions of some proteins; it can also serve as spatial information for the recruitment and docking of proteins.

Curvature sensing is thought to be important for vesicular trafficking processes (Antonny, 2011;[START_REF] Liu | Mechanochemical crosstalk during endocytic vesicle formation[END_REF].

The regulation of protein membrane binding by curvature was first suggested for ARFGAP1 which is involved in the uncoating of COPI vesicles [START_REF] Bigay | Lipid packing sensed by ArfGAP1 couples COPI coat disassembly to membrane bilayer curvature[END_REF]. Vesicle budding from the Golgi membrane is mediated by the assembly of COPI coat proteins that are attached to the membrane via the activated (GTP-bound) adaptor protein ARF1. Subsequent disassembly of the coat requires ARF1 inactivation by ARFGAP1. Bigay and coworkers demonstrated that ARF1 inactivation and subsequent release of coat proteins was faster at the surface of small liposomes suggesting that ARFGAP1 recruitment was enhanced in presence of curved membranes [START_REF] Bigay | Lipid packing sensed by ArfGAP1 couples COPI coat disassembly to membrane bilayer curvature[END_REF].

Logically enough all mentioned curvature generators are also able to detect membrane curvature.

This was mostly described for BAR domain proteins (Bhatia et al., 2009;[START_REF] Gallop | Mechanism of endophilin N-BAR domain-mediated membrane curvature[END_REF]Peter et al., 2004), Epsin [START_REF] Capraro | Curvature sensing by the epsin N-terminal homology domain measured on cylindrical lipid membrane tethers[END_REF] and Dynamin [START_REF] Roux | Membrane curvature controls dynamin polymerization[END_REF]. This double function is thought to be dependent on their membrane-bound density (Antonny, 2011;Sorre et al., 2012) with curvature generation and sensing respectively representing the high and low density behavior of the same protein.

Because the observed curvature generation activity of these proteins was assessed either by overexpression in cells or by high concentration on liposomes, it might not be physiologically relevant in vivo. The curvature generation and sensing activities can however be related in the way that if a protein accumulates on a curved surface in the cell, it might reach a high enough concentration which is sufficient to induce further deformations.

Two major mechanisms of curvature sensing have been described: the sensing of surface geometry and the sensing of lipid packing defects (Figure 1.10). N-BAR domain proteins such as Amphiphysin or Endophilin are curvature sensors that have been suggested to use both mechanisms.

Sensing of surface geometry and charge

BAR domains are extended dimers containing cationic residues and exhibiting an arc-shaped structure that will preferentially bind curved and negatively charged membranes (Peter et al., 2004) (Figure 1.10A). In the case of BAR domain curvature sensing appears intuitive. If the membrane displays a curvature that is closer to that of the BAR domain, this will lead to more electrostatic interactions between the negatively charged membrane and the positively charged domain, thus leading to a higher gain of energy. Of note, classical BAR domains preferentially sense highly positively curved membranes whereas F-BAR (extended FCH domain) domains sense low positive curvature and I-BAR (Inverse BAR) domains bind to negatively curved membrane surfaces [START_REF] Mim | Membrane curvature and its generation by BAR proteins[END_REF].

Dynamin is a curvature sensor that, contrary to BAR domain proteins, does not display a curved interface [START_REF] Ferguson | Dynamin, a membrane-remodelling GTPase[END_REF]. It was shown to be preferentially recruited to highly curved tubes and this was suggested to be related to its polymerization activity (formation of a curved dynamin coat) on curved membranes [START_REF] Roux | Membrane curvature controls dynamin polymerization[END_REF]. Dynamin also possess a hydrophobic loop, emerging from its pleckstrin homology domain, which displays curvature sensing activity [START_REF] Vallis | Importance of the pleckstrin homology domain of dynamin in clathrin-mediated endocytosis[END_REF], possibly by the second curvature sensing mechanism: the sensing of lipid packing defects.

Hydrophobic insertion into lipid packing defects

Proteins can also sense curvature, not through the sensing of membrane surface geometry but through the sensing of lipid packing defects induced by membrane curvature [START_REF] Bigay | ArfGAP1 responds to membrane curvature through the folding of a lipid packing sensor motif[END_REF][START_REF] Drin | A general amphipathic alpha-helical motif for sensing membrane curvature[END_REF]. When a membrane is curved, the surface area of the lipid headgroup region of one monolayer is expanded whereas the hydrophobic region remains undisturbed thus leading to the formation of lipid packing defects (Figure 1.10B). Of note, defects in the arrangement of lipids can also be related to the specific membrane compositions and consequently to membrane only to curved membranes via the insertion of hydrophobic residues (yellow squares). In contrast, α-Synuclein has small and poorly hydrophobic residues (yellow circles) and a zwiterrionic polar face (positive and negative charges) and will bind only to curved and negatively charged (red lipid headgroups) membranes. (C) Specific lipid compositions (conical-shaped lipids) can also lead to the formation of lipid packing defects. Mono-unsaturated lipids promote the formation of deep defects, which are preferential binding sites for ALPS motifs, whereas poly-unsaturated lipids promote the formation of shallow defects which allow α-Synuclein membrane insertion. Adapted from [START_REF] Drin | A general amphipathic alpha-helical motif for sensing membrane curvature[END_REF].

order (Antonny, 2011). Indeed, cone-shaped lipids can promote the formation of packing defects (Figure 1.10C), and consequently the formation of a liquid-disordered phase.

This type of curvature sensing is mediated by the insertion of hydrophobic or amphipathic protein domains.

Amphipathic helix (AH) membrane insertion into curved membranes was shown to be dependent on the presence of higher amounts of lipid packing defects as compared to flat membranes [START_REF] Vanni | Amphipathic lipid packing sensor motifs: probing bilayer defects with hydrophobic residues[END_REF]. AHs are well described curvature sensors that can be found in many different proteins, involved for example in vesicular trafficking processes. Proteins such as Epsin are involved in coat assembly [START_REF] Ford | Curvature of clathrin-coated pits driven by epsin[END_REF] whereas ARFGAP1 is involved in coat disassembly [START_REF] Bigay | Lipid packing sensed by ArfGAP1 couples COPI coat disassembly to membrane bilayer curvature[END_REF]. AHs are thought to fold upon contact with membranes and insert their hydrophobic face in the lipid bilayer in contact with the lipid hydrophobic tails often with the help of positively charged residues on the helix that will interact with the lipid polar head group [START_REF] Drin | A general amphipathic alpha-helical motif for sensing membrane curvature[END_REF].

More recently, lipidated proteins such as trimeric G proteins [START_REF] Hatzakis | How curved membranes recruit amphipathic helices and protein anchoring motifs[END_REF] or RAS proteins (Larsen et al., 2015;Larsen et al., 2017) were also shown to sense curvature. The membrane recruitment of N-RAS proteins, which bear a palmitoyl and a farnesyl lipid group, was enhanced on highly curved liposomes and an increase in membrane curvature was shown to shift the preferential N-RAS recruitment from Ld membranes to Lo membranes (Larsen et al., 2015).

N-RAS membrane binding was subsequently shown to be dependent on the amount of lipid packing defects induced by curvature and lipid geometrical shape (Larsen et al., 2017).

Due to the fact that, not only AHs, but also alkyl hydrophobic chain motifs can sense lipid packing defects induced by curvature, it was suggested that this type of membrane curvature sensing is not dependent on the affinity of the hydrophobic domain for the membrane but simply on the ability of a curved membrane to host more proteins [START_REF] Hatzakis | How curved membranes recruit amphipathic helices and protein anchoring motifs[END_REF].

However, the "Velcro model" (Antonny, 2011) suggests that the AH's global and effective interaction with curved membranes is a result of many weak and identical interactions of different AH hydrophobic regions with the membrane. Thus, the binding of shorter AHs displaying a good balance between polar and non-polar faces will require less packing defects as compared to longer and strongly imbalanced (larger hydrophobic side) AHs.

Because of these variations in their chemical properties, AH motifs can be recruited to different cellular and curved membranes (Antonny, 2011;Bigay and Antonny, 2012). For example, membranes of the early secretory pathway display loose packing and low surface charges which will favor curvature recognition by AHs composed of a larger hydrophobic side and a small polar face, such as the ALPS motif found in ARFGAP1 (Figure 1.10B) [START_REF] Mesmin | Two lipid-packing sensor motifs contribute to the sensitivity of ArfGAP1 to membrane curvature[END_REF]. In contrast to this, α-Synuclein, which displays a smaller hydrophobic side and more charged residues, will preferentially recognize membrane curvature in bilayers exhibiting tighter lipid packing and higher amounts of negatively charged lipids (Figure 1.10B), mainly found in the late secretory pathway [START_REF] Jao | Structure of membrane-bound alphasynuclein from site-directed spin labeling and computational refinement[END_REF].

Thus, one can differentiate between deep lipid packing defects, preferentially recognized by the ALPS motif and shallow lipid packing defects sensed by α-Synuclein [START_REF] Pranke | alpha-Synuclein and ALPS motifs are membrane curvature sensors whose contrasting chemistry mediates selective vesicle binding[END_REF] (Figure 1.10C).

In vitro experimental approaches 1.4.1 Model membranes for in vitro experiments

Due to the high complexity of cellular membranes, their study remains very challenging. Thus, many model membrane systems of controlled lipid composition are now available to study membrane related events in vitro [START_REF] Bagatolli | Phase behavior of multicomponent membranes: Experimental and computational techniques[END_REF][START_REF] Sezgin | Model membrane platforms to study protein-membrane interactions[END_REF].

Depending on the study and more specifically on the relevant membrane physical parameters, model membranes with various geometries can be used (Figure 1.11): planar shape bilayers like supported lipid bilayers or spherical shape bilayers like vesicles. Spherical vesicles with different sizes can be obtained; from the nanometer range (small unilamellar vesicles, SUVs) to the micrometer range (giant unilamellar vesicles, GUVs). Some of the existing model membrane systems are summarized in Figure 1.11 as well as the methods to produce them.

Multilamellar vesicles can be synthesized through the hydration of dry lipid films. Subsequent sonication of these vesicles leads to the formation of SUVs (30 to 50 nm in diameter) whereas their extrusion through polycarbonate membranes of varying pore sizes leads to the formation of large unilamellar vesicles (LUVs) with diameters ranging between 100 nm and 1 µm (Figure 1.11).

GUVs are unilamellar vesicles of larger sizes (1 to 100 µm in diameter) that are produced by the hydration of dry lipid films and subsequent electroformation under an alternating current (AC) electric field (Angelova et al., 1992).

Unilamellar vesicles with asymmetric membranes can also be obtained using the inverted emulsion technique [START_REF] Pautot | Production of unilamellar vesicles using an inverted emulsion[END_REF], which consists in the transfer of inverted emulsion droplets covered with a lipid monolayer through a second monolayer at an oil-water interface.

This method can also be used to encapsulate proteins inside GUVs [START_REF] Pontani | Reconstitution of an actin cortex inside a liposome[END_REF].

Vesicles, called giant plasma membrane vesicles (GPMVs), can also be obtained from the cell plasma membrane by inducing cell blebbing [START_REF] Levental | Giant plasma membrane vesicles: models for understanding membrane organization[END_REF].

The study that is being presented in this manuscript mostly focuses on the use of GUVs as their micrometer size allowed suitable visualization under a light microscope. They can also be deformed and their tension can be adjusted using micromanipulation techniques thereby allowing the study of the membrane mechanical properties [START_REF] Henriksen | Measurement of membrane elasticity by micro-pipette aspiration[END_REF][START_REF] Rawicz | Effect of chain length and unsaturation on elasticity of lipid bilayers[END_REF][START_REF] Rawicz | Elasticity, strength, and water permeability of bilayers that contain raft microdomain-forming lipids[END_REF][START_REF] Tierney | Elasticity and phase behavior of DPPC membrane modulated by cholesterol, ergosterol, and ethanol[END_REF]. GUVs have also been widely used to study demixing behaviors of lipid mixtures [START_REF] Scherfeld | Lipid dynamics and domain formation in model membranes composed of ternary mixtures of unsaturated and saturated phosphatidylcholines and cholesterol[END_REF][START_REF] Veatch | Separation of liquid phases in giant vesicles of ternary mixtures of phospholipids and cholesterol[END_REF][START_REF] Wesolowska | Giant unilamellar vesicles -a perfect tool to visualize phase separation and lipid rafts in model systems[END_REF] and protein interaction and sorting (Ambroggio et al., 2010;[START_REF] Saarikangas | Molecular mechanisms of membrane deformation by I-BAR domain proteins[END_REF]Sorre et al., 2012). A detailed description of giant vesicle applications can be found in (Walde et al., 2010).

One particular disadvantage of using the electroformation method is that only growth buffers containing low levels of salt can be used [START_REF] Bucher | Giant vesicles as biochemical compartments: the use of microinjection techniques[END_REF][START_REF] Dimova | A practical guide to giant vesicles. Probing the membrane nanoregime via optical microscopy[END_REF]. This problem can be circumvented using an optimized electroformation method consisting of platinum electrodes [START_REF] Meleard | Giant unilamellar vesicle electroformation from lipid mixtures to native membranes under physiological conditions[END_REF].

Another new method, exhibiting higher yield of GUV unilamellarity and faster GUV growth, has more recently been proposed. As compared to the original swelling method where lipids were spread onto glass, lipids are in this case spread on top of a polymer gel consisting of either agarose [START_REF] Horger | Films of agarose enable rapid formation of giant liposomes in solutions of physiologic ionic strength[END_REF] or polyvinyl alcohol [START_REF] Weinberger | Gel-assisted formation of giant unilamellar vesicles[END_REF]. This method also allows the use of aqueous buffer of physiological ionic strength.

Phase separation from living cells to model membranes

The most famous method used to study lipid rafts and their association with membrane proteins consists in the observation of detergent-resistant membranes. However, this method was shown not to be optimal as the use of detergents is thought to potentially induce phase separation and to affect the partitioning of membrane proteins to a specific phase [START_REF] Heerklotz | Triton promotes domain formation in lipid raft mixtures[END_REF][START_REF] Lichtenberg | Detergent-resistant membranes should not be identified with membrane rafts[END_REF]. Other techniques have thus been developed in order to study native membranes in situ [START_REF] Klymchenko | Fluorescent probes for lipid rafts: from model membranes to living cells[END_REF], such as mass spectrometry to monitor the chemical composition of cell membranes [START_REF] Lozano | Colocalization of the ganglioside G(M1) and cholesterol detected by secondary ion mass spectrometry[END_REF] or atomic force microscopy (AFM) which allows nanoscopic resolution and clear distinction between Lo and Ld domains [START_REF] Goksu | AFM for structure and dynamics of biomembranes[END_REF].

Lipid rafts in live cells can only be studied and visualized using optical techniques and fluorescent probes. However, even though optical microscopy improved in the last decades now allowing imaging at tens of nanometers resolution, the study of lipid rafts in cells is still very limited considering their small size, thought to be in the nanometer range [START_REF] Pike | Rafts defined: a report on the Keystone Symposium on Lipid Rafts and Cell Function[END_REF]. Additionally, due to their high complexity in composition and structure, the study of membrane rafts in live cells is very challenging. Biophysicists have thus been trying in the last decades to reproduce the complexity of rafts with model systems exhibiting phase separation, such as planar supported lipid bilayers [START_REF] Longo | Stability and liquid-liquid phase separation in mixed saturated lipid bilayers[END_REF][START_REF] Mulligan | AFM investigations of phase separation in supported membranes of binary mixtures of POPC and an eicosanyl-based bisphosphocholine bolalipid[END_REF][START_REF] Szmodis | Thermally induced phase separation in supported bilayers of glycosphingolipid and phospholipid mixtures[END_REF] and GUVs (Roux et al., 2005;[START_REF] Veatch | Separation of liquid phases in giant vesicles of ternary mixtures of phospholipids and cholesterol[END_REF], whose physical properties can be controlled and varied.

In single component systems, the transition to a certain phase occurs at a well-defined melting temperature. Coexistence between the So and Ld phases can be achieved in a 2-component system of high and low Tm lipids. As previously mentioned, cholesterol is key to the formation of the Lo phase. Thus, Lo and Ld domains can coexist in ternary lipid mixtures consisting of high and low Tm lipids and cholesterol. At equilibrium conditions and at a given temperature and composition, the nature of the thermodynamic phases can be predicted and referenced with phase diagrams (Figure 1.12A). These representations can be obtained using the Gibbs triangle [START_REF] Veatch | Seeing spots: complex phase behavior in simple membranes[END_REF]) and many can be found in the literature [START_REF] Feigenson | Phase behavior of lipid mixtures[END_REF][START_REF] Goni | Phase diagrams of lipid mixtures relevant to the study of membrane rafts[END_REF]. From (Baumgart et al., 2003).

It is important to note that this description of ternary mixture phase behavior is very general and details will vary from one system to another depending on the lipids used and their physical properties (acyl chain length, degree of unsaturation, charge). Lipid and protein interactions have also been shown to have an influence on phase diagrams. One of the best characterized examples is the interaction between Cholera toxin and GM1 lipids. Cholera toxin interacts with GM1 lipids and induces lipid clustering which thus leads to phase separation [START_REF] Hammond | Crosslinking a lipid raft component triggers liquid ordered-liquid disordered phase separation in model plasma membranes[END_REF].

In the case of Lo/Ld phase separation (that was used in this study), model membranes exhibit areas having Lo properties which are connected to Ld domains. It is possible to identify the different phases by using fluorescent lipids or lipophilic probes that will preferentially sort into one of the two phases [START_REF] Baumgart | Fluorescence probe partitioning between Lo/Ld phases in lipid membranes[END_REF][START_REF] Klymchenko | Fluorescent probes for lipid rafts: from model membranes to living cells[END_REF] (Figure 1.12B).

Curvature sensing on model membranes

Different model membrane systems and protocols exist to detect protein curvature sensing (Figure 1.13).

Wavy supported lipid bilayers are a very interesting system as they display a continuum of positively and negatively curved regions [START_REF] Cheney | Single Lipid Molecule Dynamics on Supported Lipid Bilayers with Membrane Curvature[END_REF][START_REF] Hsieh | Curvature sorting of peripheral proteins on solid-supported wavy membranes[END_REF]. However this method is very limited as the displayed curvature values are relatively small (radius in the few hundred nanometer range) (Figure 1.13A). A good alternative system that has been recently described is the use of supported membrane tubes [START_REF] Dar | Use of the supported membrane tube assay system for real-time analysis of membrane fission reactions[END_REF]. This system has the advantage, contrary to the previously mentioned system, to display highly curved regions (between 10 and 40 nm tube radii) (Figure 1.13B).

Curvature sensing can also be assessed by using liposomes of decreasing diameters, which can, as previously mentioned, be generated by vesicle extrusion through pores of decreasing sizes

(Figure 1.13C).
Vesicles are incubated with the proteins and centrifuged. Subsequent SDS PAGE analysis of the pellet and the supernatant fractions will allow the assessment of protein binding (Peter et al., 2004). An alternative and more effective way to study liposome-protein interactions is by flotation assay, where vesicles are centrifuged in a sucrose gradient of higher density than the liposome preparation buffer. Vesicles are thus always recovered in the top fraction. All fractions are analyzed by SDS PAGE to assess membrane localization [START_REF] Bigay | ArfGAP1 responds to membrane curvature through the folding of a lipid packing sensor motif[END_REF]. The efficiency of protein binding to vesicles can also be studied using a lipid-sensitive probe covalently attached to the protein [START_REF] Pranke | alpha-Synuclein and ALPS motifs are membrane curvature sensors whose contrasting chemistry mediates selective vesicle binding[END_REF].

The single liposome curvature assay (SLiC) is another method used to assess liposome curvature sensing and, in contrast to the previous mentioned methods, allows the assessment of protein binding to individual vesicles with diameters ranging from 30 nm to 1 µm [START_REF] Jensen | Membrane curvature sensing by amphipathic helices: a single liposome study using alphasynuclein and annexin B12[END_REF]Larsen et al., 2015). Liposomes are immobilized on a passivated glass surface through biotinstreptavidin interactions (Figure 1.13D). Binding of fluorescent proteins as well as fluorescent liposome signal are then assessed by confocal microscopy. The great advantage of this method is that because SLiC works at the single-liposome level, and each frame is comprised of hundreds of vesicles exhibiting various curvatures, it thus permits to probe a wide range of curvatures in the same experiment (Larsen et al., 2015). Finally, the setup that has been used in this study consists in the pulling of membrane nanotubes from GUVs (Figure 1.13E). Tube formation occurs through the aspiration of the GUV with a micropipette on one side and the trapping (using an optical tweezer setup) of a bead tethered to the vesicle on the other side. This technique allows the quantitative study of protein enrichment on a highly curved membrane region (the tube) as compared to a flat membrane region (the GUV) (Ambroggio et al., 2010;[START_REF] Prevost | IRSp53 senses negative membrane curvature and phase separates along membrane tubules[END_REF]Sorre et al., 2012).

The micropipette aspiration technique, which was first introduced by Evans on red blood cells [START_REF] Waugh | Thermoelasticity of red blood cell membrane[END_REF] and later on large vesicles [START_REF] Kwok | Thermoelasticity of large lecithin bilayer vesicles[END_REF], allows the control of membrane tension and was thus originally meant to assess elastic properties of membranes. I used it for two purposes: holding the GUV and controlling membrane tension (and consequently tube curvature).

Tube extraction was first achieved by applying a flow on red blood cells attached to a glass surface [START_REF] Hochmuth | Measurement of the elastic modulus for red cell membrane using a fluid mechanical technique[END_REF]. Later on, cells were held with a micropipette and tubes were pulled using a bead immobilized with a second micropipette [START_REF] Hochmuth | Extensional flow of erythrocyte membrane from cell body to elastic tether. II. Experiment[END_REF].

Finally, tubes were extracted using optically or magnetically trapped beads. While holding the vesicle with a micropipette, these trapped beads are used to apply a point force on the membrane and consequently extract the tubes [START_REF] Cuvelier | Coalescence of membrane tethers: experiments, theory, and applications[END_REF][START_REF] Heinrich | A piconewton force transducer and its application to measurement of the bending stiffness of phospholipid membranes[END_REF] (Figure

1.13E).

Optical tweezers allow the micromanipulation of micron-size particles. The optical trap is created by tightly focusing a laser beam with an objective of high numerical opening. The dielectric and trapped particle experiences two forces: a scattering force which tends to push the particle in the direction of the propagation of light; and a gradient force, which acts in the direction of the intensity gradient and will thus tend to bring the particle back to the focus (Figure 1.14) [START_REF] Neuman | Optical trapping[END_REF]. Thus, for an object to be efficiently trapped, the gradient force must exceed the scattering force. This is the reason why the laser beam must be tightly focused (higher focusing results in a greater intensity gradient).

Of note, other similar tube-pulling methods have been developed, such as the use of forces generated by kinesin motors walking on microtubules [START_REF] Roux | A minimal system allowing tubulation with molecular motors pulling on giant liposomes[END_REF] (Figure 1.13F). Briefly, biotinylated kinesins interact with membranes composed of biotinylated lipids, in presence of streptavidin beads. Subsequent ATP addition will activate the kinesins and allow them to walk on the polymerized microtubules and consequently to pull membrane tubes. A detailed experimental protocol will be described in Chapter 3. of same intensity and opposite direction is then imparted to the bead, resulting in the force depicted by the gray arrows. The thicker ray carries more intensity and therefore transmits a greater momentum to the bead. As a result, the net force on the bead is towards the high intensity (and slightly downward). B: To form a stable trap, the light has to be focused. To do so, the bead is illuminated by a focused beam of light with a radial intensity gradient (highest intensity is depicted in black). In this case, the two lateral rays have the same intensity and their refraction leads to a transfer of momentum resulting in lateral forces that balance each other. Therefore only the axial force remains. This axial force is balanced by the scattering force (not shown), resulting in the axial stability of the bead. If the bead is moved away from the center of the beam, the radial gradient force will bring it back to the equilibrium position. From [START_REF] Neuman | Optical trapping[END_REF].

Light intensity profile Light intensity profile

A B

Chapter 2

2 RAB GTPases GTPases. This superfamily is comprised of more than 160 members and is divided into five major families [START_REF] Rojas | The Ras protein superfamily: evolutionary tree and role of conserved amino acids[END_REF]. RAS proteins couple extracellular signals to intracellular pathways and thus play a critical role in cellular processes such as proliferation, apoptosis, differentiation, motility and transcription [START_REF] Karnoub | Ras oncogenes: split personalities[END_REF]. The RHO (RAS homology) family of proteins regulates various cellular processes including cell polarity, cytoskeletal organization, cell cycle and transcriptomal dynamics [START_REF] Bustelo | GTP-binding proteins of the Rho/Rac family: regulation, effectors and functions in vivo[END_REF] and the RAN (RAS-related nuclear protein) family mediates nuclear transport processes and mitosis [START_REF] Melchior | Ran GTPase cycle: oOne mechanism --two functions[END_REF]. The members of the ARF (ADP-ribosylation factor) family of proteins are involved in the recruitment of coat proteins, the regulation of phospholipid metabolism and in the modulation of actin structure at the surface of membranes; and thereby regulate membrane trafficking events and organelle structure [START_REF] Donaldson | ARF family G proteins and their regulators: roles in membrane transport, development and disease[END_REF][START_REF] Jackson | Arfs at a glance[END_REF]. Finally, RAB proteins represent the largest family of small GTPases with over 60 RAB genes identified in humans. These proteins actively regulate membrane trafficking events in the cell and localize to distinct compartments, thereby defining organelle identity [START_REF] Zhen | Cellular functions of Rab GTPases at a glance[END_REF].

As this study focuses on the membrane binding properties of RAB proteins, a detailed description of RAB structure, localization and function will be provided in this second chapter and a particular attention will be set on the different RAB proteins used in this study.

RAB discovery and evolution

The first two RAB genes, YPT1 and SEC4, were discovered in yeast [START_REF] Gallwitz | A yeast gene encoding a protein homologous to the human c-has/bas proto-oncogene product[END_REF][START_REF] Salminen | A ras-like protein is required for a post-Golgi event in yeast secretion[END_REF]. At this time, these RAS-like proteins were thought, like all G proteins, to act in the transduction of signals through the plasma membrane [START_REF] Barbacid | ras genes[END_REF][START_REF] Gilman | G proteins: transducers of receptor-generated signals[END_REF]. The observation that SEC4 mutations confer late secretory defects suggested a role for SEC4 in intracellular trafficking [START_REF] Salminen | A ras-like protein is required for a post-Golgi event in yeast secretion[END_REF]. A year later, YPT1 was shown to localize to the yeast Golgi and YPT1 mutant cells were shown to exhibit defects in the transfer of material between the ER and the Golgi [START_REF] Segev | The yeast GTP-binding YPT1 protein and a mammalian counterpart are associated with the secretion machinery[END_REF]. YPT1 became the first GTPase shown to function inside cells and to regulate intracellular trafficking. The term RAB for RAS-related in Brain came from the isolation of four RAS genes, homologous to the yeast YPT proteins, using a screen of a rat brain cDNA library [START_REF] Touchot | Four additional members of the ras gene superfamily isolated by an oligonucleotide strategy: molecular cloning of YPT-related cDNAs from a rat brain library[END_REF].

The conservation of YPTs in evolution was first shown for YPT1. Its closest human homolog, RAB1 was shown to not only share 71% amino acid sequence identity [START_REF] Touchot | Four additional members of the ras gene superfamily isolated by an oligonucleotide strategy: molecular cloning of YPT-related cDNAs from a rat brain library[END_REF] but also to localize to the Golgi [START_REF] Segev | The yeast GTP-binding YPT1 protein and a mammalian counterpart are associated with the secretion machinery[END_REF]. The subsequent discovery that mouse RAB1 could functionally replace YPT1 in yeast led to the assumption that membrane trafficking events were regulated by conserved machineries [START_REF] Haubruck | The ras-related mouse ypt1 protein can functionally replace the YPT1 gene product in yeast[END_REF]. More YPTs and RABs were then discovered by searching for homologs of YPT1 and SEC4 [START_REF] Zahraoui | The human Rab genes encode a family of GTP-binding proteins related to yeast YPT1 and SEC4 products involved in secretion[END_REF] and even more were identified when the yeast and human genome-sequencing projects were completed [START_REF] Bock | A genomic perspective on membrane compartment organization[END_REF][START_REF] Lazar | Vesicular transport: how many Ypt/Rab-GTPases make a eukaryotic cell?[END_REF]. Currently, eleven yeast YPTs and more than 60 human RABs have been identified; and YPT1, YPT51, YPT6, YPT7, SEC4 and YPT31/YPT32 have proved to be very homologous to RAB1, RAB5, RAB6, RAB7, RAB8 and RAB11 respectively in mammalian cells (Figure 2.1). Interestingly, the YPT/RAB family expanded significantly from yeast to multicellular organisms, such as worms and flies, and then to mammals [START_REF] Bock | A genomic perspective on membrane compartment organization[END_REF], implying that trafficking pathways in mammals are much more tightly regulated, but also tissue specific (some RAB proteins are not ubiquitously expressed). The YPT/RAB family has significantly expanded from unicellular to multicellular organisms and is now composed of more than 60 members in humans (see (Hutagalung and Novick, 2011) for detailed review about existing RAB proteins). Sequences were retrieved from UniProt and aligned using ClustalW.

RAB sequence and structure

Extensive sequence analysis and crystallographic studies led to the identification of several RAB sequence regions and motifs which are critical for function and specific membrane recruitment.

G-domain

RAB proteins are composed of only one structural domain, common to all members of the RAS superfamily, called the G domain. This domain, which structurally consists of a central β-sheet with six strands that is flanked by five α-helices on both sides, serves as the binding and hydrolysis site of guanine nucleotides, and switches conformation upon GTP or GDP binding (Figure 2.2A) [START_REF] Dumas | Structural basis of activation and GTP hydrolysis in Rab proteins[END_REF][START_REF] Ostermeier | Structural basis of Rab effector specificity: crystal structure of the small G protein Rab3A complexed with the effector domain of rabphilin-3A[END_REF]. The specific regions that can sense the nature of the bound nucleotides and are involved in GTP hydrolysis are the switch I and switch II regions. The respective contributions of these two regions vary among the members of the RAS superfamily. Both switch regions were shown to interact with the γ-phosphate of GTP whereas GDP interaction (through its guanine base) is only mediated by the Switch I region [START_REF] Cherfils | Regulation of small GTPases by GEFs, GAPs, and GDIs[END_REF][START_REF] Vetter | The guanine nucleotide-binding switch in three dimensions[END_REF].

Both tend to be disordered in the GDP-bound form and undergo major changes to adopt a structurally ordered state upon GTP binding (Figure 2.2B and Figure 2.2C) [START_REF] Lee | Structural mechanisms for regulation of membrane traffic by rab GTPases[END_REF].

When comparing the switch region crystal structures of several GTP-bound RAB proteins, significant resemblance can be observed in terms of overall length and boundaries but structural differences between RAB subfamilies can also be detected (Figure 2.2C), specifically in their switch and complementary determining (CDRs) regions [START_REF] Pfeffer | Structural clues to Rab GTPase functional diversity[END_REF].

RAB specific sequence motifs

Extensive sequence analyses of different RAB proteins have shown the existence of specific motifs (Figure 2.3). Pereira-Leal and Seabra (Pereira-Leal and Seabra, 2000) have identified five RAB family regions (RABF) that distinguish RAB proteins from the other members of the RAS superfamily and four subfamily regions (RABSF) that stand to differentiate each RAB subfamily (RAB1 from RAB6 for example). To note, the previously mentioned CDRs correspond approximately to the RABSF motifs (RABSF1, RABSF3, RABSF4) [START_REF] Pfeffer | Structural clues to Rab GTPase functional diversity[END_REF]. Different combinations of mutations of these motifs led to mislocalization of the RAB proteins suggesting that membrane specificity is also determined by these specific RAB sequence domains (Ali et al., 2004).

RAB C-terminal region

The C-terminal extremity of RAB proteins, and more specifically the last 35 to 40 amino acid region, is unstructured and highly variable among the different RAB proteins. It was initially proposed that RAB specific membrane targeting was only dependent on the RAB hypervariable Cterminal sequence [START_REF] Chavrier | Hypervariable C-terminal domain of rab proteins acts as a targeting signal[END_REF]. This hypothesis was however later questioned when the specific localization of RAB proteins was shown to not always be dependent on their C-terminal region but instead to be dependent on motifs distributed throughout their primary sequence (Ali et al., 2004;[START_REF] Beranger | The effector domain of Rab6, plus a highly hydrophobic C terminus, is required for Golgi apparatus localization[END_REF]. An even more recent study demonstrated that when most of the hypervariable region was replaced with a polyethylene glycol linker, RAB1 and RAB5 were still found to localize to the correct intracellular compartment (Li et al., 2014); thus also going against the prediction that the C-terminus of RAB proteins is critical for RAB targeting.

Some RAB proteins however do possess C-terminal residues involved in specific membrane targeting. It was shown that the replacement of the polybasic C-terminal sequence of RAB35 with uncharged elements led to the absence of proper plasma membrane localization (Li et al., 2014).

It was thus suggested that the specific recruitment of RAB35 to the plasma membrane (which is negatively charged) is mediated by electrostatic interactions. RAB7 specific localization to late endosomes / lysosomes was also described to be dependent on its hypervariable region and more specifically on the C-terminal region involved in the binding with the RAB-interacting lysosomal protein (RILP) (Li et al., 2014).

The C-terminal extremity of RAB proteins is also composed of a mono or di-cysteine motif which allows the covalent attachment of respectively one or two geranylgeranyl lipid moieties necessary for RAB membrane insertion [START_REF] Desnoyers | Geranylgeranylation of Rab proteins[END_REF]. This will be further discussed in the next section.

RAB posttranslational modifications 2.3.1 RAB activation cycle

Like all GTPases, YPTs/RABs cycle between an active GTP-bound form and an inactive GDPbound form (Figure 2.4). Upon membrane incorporation, RAB proteins are activated by a guanine nucleotide exchange factor (GEF) and bound GDP is replaced by the approximately 10fold more abundant GTP [START_REF] Barr | Rab GEFs and GAPs[END_REF][START_REF] Cherfils | Regulation of small GTPases by GEFs, GAPs, and GDIs[END_REF]. It is important to note that GEF proteins can catalyze the exchange reaction in both directions but directionality is dictated by the higher concentration of GTP over GDP [START_REF] Goody | Exchange factors, effectors, GAPs and motor proteins: common thermodynamic and kinetic principles for different functions[END_REF]. In such a conformation, RABs can fulfill their functions in the trafficking processes through their interaction with effector proteins. RABs then undergo an inactivation process which is mediated by GTPase-activating proteins (GAPs) and consists in the hydrolysis of GTP to GDP [START_REF] Barr | Rab GEFs and GAPs[END_REF][START_REF] Cherfils | Regulation of small GTPases by GEFs, GAPs, and GDIs[END_REF].

GEFs and GAPs regulate RAB activation in a spatiotemporally controlled manner. So far, many

RABGEFs have been identified including the two large families Vps9-domain containing proteins (specific for the RAB5 family) [START_REF] Carney | Vps9 domain-containing proteins: activators of Rab5 GTPases from yeast to neurons[END_REF][START_REF] Horiuchi | A novel Rab5 GDP/GTP exchange factor complexed to Rabaptin-5 links nucleotide exchange to effector recruitment and function[END_REF] and DENN domain GEFs which act on several different RABs [START_REF] Marat | DENN domain proteins: regulators of Rab GTPases[END_REF]. Many other structurally different GEFs, that are not part of these two families, have also been identified (see [START_REF] Muller | Molecular control of Rab activity by GEFs, GAPs and GDI[END_REF] for extensive review). These different RABGEF proteins exhibit very low structural and sequence homology to each other, which explains why the corresponding GEFs of some RABs are yet to be identified. which allows their interaction with effector proteins. RABs subsequently interact with GTPase activating proteins (GAPs) which catalyze GTP to GDP hydrolysis and GDP-bound RABs can get recycled back to the donor compartment by the GDP dissociation inhibitor (GDI). Adapted from (Hutagalung and Novick, 2011).

Several GEFs of small GTPases, such as RASGEFs SOS and ARFGEFs Cytohesins, have been reported to be controlled by feedback loops, in which the GTP-bound GTPases or their effectors bind to their cognate GEFs and modify their basal nucleotide exchange rate [START_REF] Cherfils | Regulation of small GTPases by GEFs, GAPs, and GDIs[END_REF]. Interestingly, some RABGEFs have been suggested to be regulated by similar feedback loops.

RAB5GEF Rabex-5 is the best characterized example as it was described to interact with Rabaptin-5, a RAB5 effector [START_REF] Horiuchi | A novel Rab5 GDP/GTP exchange factor complexed to Rabaptin-5 links nucleotide exchange to effector recruitment and function[END_REF]. Briefly, RAB5 is activated by Rabex-5 upon membrane recruitment and is then able to interact with its various effectors, including Rabaptin-5. Rabaptin-5 then interacts with Rabex-5 and allows the increase of nucleotide exchange activity of Rabex-5 towards RAB5 (feedback effect). These interactions generate a feedback loop which prevents RAB5 inactivation by its cognate GAP, and GDI-mediated membrane extraction [START_REF] Lippe | Functional synergy between Rab5 effector Rabaptin-5 and exchange factor Rabex-5 when physically associated in a complex[END_REF]. This feedback mechanism was however recently challenged with the finding that Rabaptin-5 recruitment to early endosomes is independent of RAB5 and relies on its interaction with Rabex-5 and RAB4 [START_REF] Kalin | Rabaptin5 is recruited to endosomes by Rab4 and Rabex5 to regulate endosome maturation[END_REF]. A similar feedback mechanism was suggested to occur in yeast where SEC4 and its cognate GEF SEC2 both interact with SEC15, a subunit of the exocyst complex [START_REF] Medkova | The rab exchange factor Sec2p reversibly associates with the exocyst[END_REF].

A key feature of small GTPases is their ability to intrinsically hydrolyze GTP, but this activity is very low. Thus, as mentioned above, RAB proteins get more rapidly inactivated through GAP mediated GTP hydrolysis. In contrast to RAB GEFs, human RAB GAPs consist of one major family, the TBC domain containing proteins with more than 40 members in humans [START_REF] Fukuda | TBC proteins: GAPs for mammalian small GTPase Rab?[END_REF]). An exception to this is found with the RAB3 subfamily-specific GAP complex which consists of 2 different proteins [START_REF] Nagano | Molecular cloning and characterization of the noncatalytic subunit of the Rab3 subfamily-specific GTPase-activating protein[END_REF].

GAP proteins but also GEFs were shown to be involved in RAB cascades, where RABs recruit effectors that act as GAPs for upstream RAB protein or as GEFs for RABs acting further down the pathway. YPT32 was shown to recruit SEC2, the GEF of the later acting SEC4 [START_REF] Ortiz | Ypt32 recruits the Sec4p guanine nucleotide exchange factor, Sec2p, to secretory vesicles; evidence for a Rab cascade in yeast[END_REF], and also to recruit GYP1 and GYP6, the two respective GAPs of the previous acting YPT1 and YPT6

(Rivera-Molina and [START_REF] Rivera-Molina | A Rab GAP cascade defines the boundary between two Rab GTPases on the secretory pathway[END_REF][START_REF] Suda | Rab GAP cascade regulates dynamics of Ypt6 in the Golgi traffic[END_REF]. A similar cascade was described for endocytic RABs; RAB5 recruits the class C VPS/HOPS complex, a known GEF for RAB7 [START_REF] Rink | Rab conversion as a mechanism of progression from early to late endosomes[END_REF]. These RAB cascades were suggested to keep RAB domains distinct through the removal of a RAB from a domain in which it does not belong [START_REF] Nottingham | Defining the boundaries: Rab GEFs and GAPs[END_REF].

RAB membrane insertion and extraction

RAB proteins cycle between a cytosolic and a membrane bound form (Figure 2.4). In order to reversibly bind to membranes, RABs, but also RAS and RHO/RAC proteins, need to be prenylated at their C-terminal cysteine residues (Figure 2.5) [START_REF] Zhang | Protein prenylation: molecular mechanisms and functional consequences[END_REF]. Prenylation is a posttranslational modification that was discovered in the late 1970s but was only described for RABs more than 10 years later [START_REF] Farnsworth | C terminus of the small GTP-binding protein smg p25A contains two geranylgeranylated cysteine residues and a methyl ester[END_REF][START_REF] Khosravi-Far | Ras (CXXX) and Rab (CC/CXC) prenylation signal sequences are unique and functionally distinct[END_REF].

Prenylation consists in the addition of either a farnesyl (Figure 2.5A) or geranylgeranyl group (Figure 2.5B) at the C-terminus of target proteins and is catalyzed by 3 different enzymes.

Farnesyltransferase (FTase) and geranylgeranyltransferase 1 (GGTase 1) both recognize C-terminal CAAX motifs (C is a Cystein, A is an aliphatic amino acid and X is any amino acid) and attach a single farnesyl or geranylgeranyl group to the cysteine via a thioester linkage [START_REF] Seabra | Protein farnesyltransferase and geranylgeranyltransferase share a common alpha subunit[END_REF].

If X is a methionine, serine, glutamine, or alanine, the substrate is farnesylated whereas if it is a leucine or phenylalanine, the protein is geranylgeranylated. Conversely, RAB geranylgeranyl transferase (RABGGTase, also called GGTase II) catalyzes the addition of usually two geranylgeranyl groups on a di-cysteine motif (CC, CXC or CCXX) at the C-terminal extremity of RAB proteins (Figure 2.5B) [START_REF] Seabra | Rab geranylgeranyl transferase. A multisubunit enzyme that prenylates GTP-binding proteins terminating in Cys-X-Cys or Cys-Cys[END_REF].

After their synthesis, RAB proteins first bind the RAB escort protein (REP) which then presents the RAB to the RABGGTase for geranylgeranylation (Figure 2.4) (Alexandrov et al., 1999;[START_REF] Andres | cDNA cloning of component A of Rab geranylgeranyl transferase and demonstration of its role as a Rab escort protein[END_REF]. The subsequent binding of a new geranylgeranylpyrophosphate (GGpp) substrate molecule to the active site of the RABGGTase leads to RAB dissociation from the REP and delivery to the target membrane (Thoma et al., 2001a;Thoma et al., 2001b). It is important to note that some RABs, such as RAB8 and RAB13, display a mono-cysteine CAAX motif at their C-terminus and are modified by a single geranylgeranyl moiety [START_REF] Joberty | Isoprenylation of Rab proteins possessing a C-terminal CaaX motif[END_REF]. This monoprenylation reaction seems to still be mediated by REP and RABGGTase (Gomes et al., 2003;[START_REF] Wilson | Prenylation of Rab8 GTPase by type I and type II geranylgeranyl transferases[END_REF]. It has also been suggested that the mono-and diprenylation motifs are important for RAB specific membrane targeting possibly because of the higher hydrophobicity of the double lipid modification or the preferential recognition of double prenylated RABs by regulatory proteins [START_REF] Calero | Dual prenylation is required for Rab protein localization and function[END_REF]Gomes et al., 2003). Once membrane bound RAB proteins have fulfilled their functions, they need to be extracted from membranes and recycled back to the donor compartment (Figure 2.4). The protein responsible for this was originally identified as a RAB3 interacting protein that inhibited the dissociation of GDP and was thus names GDP dissociation inhibitor (GDI) [START_REF] Sasaki | Purification and characterization from bovine brain cytosol of a protein that inhibits the dissociation of GDP from and the subsequent binding of GTP to smg p25A, a ras p21-like GTP-binding protein[END_REF].

Interestingly, RABGDI and REP were shown to be structurally very homologous to each other (Alory and Balch, 2001). However, even though both REP and GDI were shown to bind GDPbound RABs with higher affinity as compared to GTP (Wu et al., 2010), REP displayed higher affinity for unprenylated RABs whereas GDI displayed higher affinity for the prenylated form [START_REF] Wu | Interaction analysis of prenylated Rab GTPase with Rab escort protein and GDP dissociation inhibitor explains the need for both regulators[END_REF]. This ensures that RAB interaction with GDI only occurs once their trafficking cycle is completed and they have been inactivated.

Due to the high affinity of the RAB:GDI complex, the intrinsic rate of GDI dissociation is very low which hence limits the rate of RAB recycling and consequently of active RAB generation [START_REF] Shapiro | Quantitative analysis of the interactions between prenyl Rab9, GDP dissociation inhibitor-alpha, and guanine nucleotides[END_REF]. This led to the assumption that possible mechanisms exist to accelerate this process.

Some studies suggested that the YIP3/PRA1 protein might function as a GDI dissociation factor (GDF) that is able to dissociate RAB proteins from high affinity RAB:GDI complexes to catalyze the final step of their recycling to the donor compartment [START_REF] Dirac-Svejstrup | Identification of a GDI displacement factor that releases endosomal Rab GTPases from Rab-GDI[END_REF]Sivars et al., 2003). YIP proteins have consequently been considered as potential GDFs for various RAB proteins (Barrowman and Novick, 2003). The absolute requirement for GDFs has however been disputed as the GEF activity of the Legionella pneumophila DrrA/SidM was shown to be sufficient to displace GDI from the RAB1:GDI complex [START_REF] Schoebel | RabGDI displacement by DrrA from Legionella is a consequence of its guanine nucleotide exchange activity[END_REF]. Additional studies demonstrated that deletion of YIP proteins did not alter the localization of YPT7 [START_REF] Cabrera | Guanine nucleotide exchange factors (GEFs) have a critical but not exclusive role in organelle localization of Rab GTPases[END_REF], again arguing against the role of YIP proteins as potential GDFs.

Other studies showed that, after spontaneous dissociation of RAB:GDI complexes, phosphocholination and adenylylation respectively mediated by AnkX and DrrA/SidM from Legionella pneumophila, can inhibit the reformation of the RAB:GDI complexes (Oesterlin et al., 2012). It was at that time thought that human cells could naturally use similar mechanisms to displace RABs from GDI. This was recently confirmed when phosphorylation of RAB1 at the switch II region by TGF-β activated kinase 1 was shown to preferentially occur when RAB1 is GDPbound, and to consequently disrupt RAB-GDI interaction (but not interactions with GEF or GAP proteins) [START_REF] Levin | Innate immunity kinase TAK1 phosphorylates Rab1 on a hotspot for posttranslational modifications by host and pathogen[END_REF].

Other RAB phosphorylations at different positions of their primary sequence have been reported.

RAB4 was shown to be phosphorylated at Ser 204 near to the C-terminal cysteines and this was described to regulate its localization during cell cycle progression [START_REF] Bailly | Phosphorylation of two small GTP-binding proteins of the Rab family by p34cdc2[END_REF]van der Sluijs et al., 1992a). RAB8 and RAB13 were also shown to be phosphorylated at Ser 111 by PTEN-induced kinase 1. Phosphorylation of RAB8 was at the same time shown to negatively regulate its activation by its cognate GEF Rabin8 [START_REF] Lai | Phosphoproteomic screening identifies Rab GTPases as novel downstream targets of PINK1[END_REF]. Interestingly, phosphorylation events seem to rather negatively regulate the membrane insertion of some RAB proteins.

Some RAB proteins can undergo additional C-terminal modifications following geranylgeranylation, such as proteolysis and/or carboxyl methylation, depending on their prenylation motif. Similarly to RAS and RHO GTPases, CAAX motifs-containing RAB proteins undergo proteolytic cleavage of the AAX tripeptide which allows the exposure of the C-terminal cysteine (Leung et al., 2007). Following proteolysis, RAB proteins exhibiting CAAX but also CXC motifs at their C-terminal extremity are methylated (Leung et al., 2007;Smeland et al., 1994).

RAB carboxyl methylation, which consists in the addition of a carboxyl group to the exposed prenylated cysteine, was shown to enhance the hydrophobicity of the C-terminus and subsequently to increase membrane affinity [START_REF] Michaelson | Postprenylation CAAX processing is required for proper localization of Ras but not Rho GTPases[END_REF][START_REF] Silvius | Fluorimetric evaluation of the affinities of isoprenylated peptides for lipid bilayers[END_REF].

Methylation was subsequently shown to be required for RAS [START_REF] Michaelson | Postprenylation CAAX processing is required for proper localization of Ras but not Rho GTPases[END_REF] but not for RAB (Leung et al., 2007) proper localization. Both studies also observed an increase of protein cytosolic pool in the absence of methylation, which appears to be related to increased RHO/RAC:GDI and RAB:GDI affinities. membrane targeting was thought to be mediated by the RAB hypervariable C-terminal domain [START_REF] Chavrier | Hypervariable C-terminal domain of rab proteins acts as a targeting signal[END_REF]. Although this region was confirmed to be important for the specific localization of some RAB proteins (Li et al., 2014), other studies demonstrated that this is not Specific localization and function of the other RABs will be detailed here. RAB2 localizes to the ER-Golgi intermediate compartment and is involved in Golgi-ER trafficking. RAB33 and RAB40 localize to the Golgi and regulate intra-Golgi transport (RAB33 also involved in autophagosome formation with RAB24). RAB8 is found on the plasma membrane and mediates trans -Golgi network to plasma membrane transport, ciliogenesis (together with RAB17 and RAB23) and GLUT4 vesicle translocation (together with RAB10 and RAB14). RAB3 (secretory vesicles, plasma membrane), RAB26 (secretory granules), RAB27 (melanosomes) and RAB37 (secretory granules) are all involved in exocytic events. RAB32 and RAB38 both localize to melanosomes and are involved in their biogenesis; RAB32 is also found on mitochondria and was shown to play a role in mitochondria fission. RAB13 is involved in tight junction assembly between cells. RAB18 regulates lipid droplet formation. RAB21 and RAB22 are both found on early endosomes; and respectively mediate integrin endocytosis and TGN-early endosomes transport / early phagosome maturation (together with RAB5 and RAB14). RAB15 localizes to early and recycling endosomes and is involved in trafficking between these two compartments and to the plasma membrane. RAB17 mediates the transport between recycling endosomes and the plasma membrane. RAB7 mostly localizes to late endosomes and lysosomes and is involved in late endosome / phagosome maturation. From (Stenmark, 2009).

Membrane targeting of RAB GTPases

always the case and that additional sequence motifs of RAB proteins also play a key role (Ali et al., 2004;[START_REF] Beranger | The effector domain of Rab6, plus a highly hydrophobic C terminus, is required for Golgi apparatus localization[END_REF].

In relation to this, studies suggested that RAB protein interaction with some effectors could also mediate RAB specific localization. As an example, mutations of RAB7 specific RABSF1 and RABSF4 domains led to RAB7 mislocalization. This was suggested to be due to the absence of interaction with the effector RILP, which is known to partially interact with these parts of the RAB7 sequence [START_REF] Wu | Structural basis for recruitment of RILP by small GTPase Rab7[END_REF]. Another study has shown that a chimeric construct consisting of RAB5 with the C-terminal hypervariable domain of RAB9 was re-localized to late endosomes (where RAB9 is originally found) [START_REF] Aivazian | TIP47 is a key effector for Rab9 localization[END_REF]. This has been suggested to be due to the interaction of the RAB9 C-terminus with its effector TIP47, and was confirmed when the abrogation of RAB9-TIP47 interaction (through TIP47 mutation) failed to localize the chimeric RAB5/RAB9 protein to late endosomes [START_REF] Aivazian | TIP47 is a key effector for Rab9 localization[END_REF].

A role in membrane targeting was also suggested for GEFs, since they can catalyze the activation of RAB proteins and consequently stabilize them on membranes (Blumer et al., 2013;[START_REF] Schoebel | RabGDI displacement by DrrA from Legionella is a consequence of its guanine nucleotide exchange activity[END_REF]. Indeed, the Legionella pneumophila protein DrrA, a GEF for RAB1 was shown to displace GDI from RAB1:GDI complexes and to subsequently activate RAB1 [START_REF] Schoebel | RabGDI displacement by DrrA from Legionella is a consequence of its guanine nucleotide exchange activity[END_REF]; and mistargeting of DrrA, Rabex-5 and Rabin-8 (known GEFS for RAB1, RAB5 and RAB8 respectively) to mitochondria led to RAB1A, RAB5A and RAB8A mislocalization to the same compartment (Blumer et al., 2013).

As already mentioned, membrane localized GDFs are thought to allow the dissociation of GDIs from RABs and thus to contribute to membrane specific localization [START_REF] Dirac-Svejstrup | Identification of a GDI displacement factor that releases endosomal Rab GTPases from Rab-GDI[END_REF]Sivars et al., 2003) but this matter is still debated as it was later suggested that GDFs are not absolutely required for RAB membrane targeting [START_REF] Cabrera | Guanine nucleotide exchange factors (GEFs) have a critical but not exclusive role in organelle localization of Rab GTPases[END_REF][START_REF] Schoebel | RabGDI displacement by DrrA from Legionella is a consequence of its guanine nucleotide exchange activity[END_REF].

In conclusion, RAB specific membrane targeting seems to be mediated by a plethora of factors, such as GDFs, GEFs, effector proteins and RAB C-terminal hypervariable region.

Most studies so far have focused on protein factors, but one cannot forget that RAB proteins are peripheral membrane-bound proteins that localize to specific membranes exhibiting specific composition and physical properties. In relation to this, there is strong evidence that membranebound molecules are not randomly distributed in the membrane bilayer but are enriched in membrane domains of varying lipid compositions [START_REF] Mukherjee | Role of membrane organization and membrane domains in endocytic lipid trafficking[END_REF]. Good examples are the glycosylphosphatidyl-inositol anchor proteins which were shown to preferentially localize to lipid rafts at the plasma membrane [START_REF] Paulick | The glycosylphosphatidylinositol anchor: a complex membraneanchoring structure for proteins[END_REF] but also other lipidated proteins such as N-RAS, H-RAS and K-RAS that were respectively described to preferentially bind to lipid packing defects (Larsen et al., 2015;Larsen et al., 2017), lipid rafts and disordered membranes in a GTP-dependent manner [START_REF] Prior | GTP-dependent segregation of H-ras from lipid rafts is required for biological activity[END_REF]; and negatively charged membranes [START_REF] Guet | Plasma membrane phosphatidylinositol 4-phosphate and 4,5-bisphosphate determine the distribution and function of K-Ras4B but not H-Ras proteins. The Journal of biological chemistry[END_REF]Jang et al., 2015).

Interestingly, clear compartmentalization of RAB5, RAB4 and RAB11 within the endosomal membrane has also been observed (Sonnichsen et al., 2000). In this study, three main populations of endosomes could be distinguished: one that is only composed of RAB5, another one that contains both RAB5 and RAB4 and a third one that harbours RAB4 and RAB11. Although it was proposed that effector protein recruitment and clustering might mediate the formation of RAB membrane domains through simultaneous effector interaction with RABs and specific lipids [START_REF] Zerial | Rab proteins as membrane organizers[END_REF], another possible explanation could consist in the direct binding of RAB proteins to membrane domains exhibiting differential geometry and physicochemical properties (curvature, order, charge). The study from Sonnichsen also demonstrates that on the same endosomal membrane, RAB5 localizes to globular structures whereas RAB4 is found on tubular (and thus more curved) structures (Sonnichsen et al., 2000), thus suggesting that some RAB proteins might preferentially bind to curved membranes.

RAB GTPases and vesicular transport 2.5.1 General mechanism of intracellular transport

Intracellular membrane trafficking, which allows the exchange of material between the different cellular organelles, can broadly be divided into two distinct pathways. The secretion pathway, which was the first one to be described [START_REF] Duve | Exploring cells with a centrifuge[END_REF][START_REF] Palade | Intracellular aspects of the process of protein synthesis[END_REF], consists in the transport of newly synthetized secretory proteins from the endoplasmic reticulum (ER) to and through the Golgi complex, release from the Golgi to secretory vesicles and final delivery to the extracellular space [START_REF] Bonifacino | The mechanisms of vesicle budding and fusion[END_REF]. In contrast to this directed transport toward the cell exterior, the endocytic pathway can be described as an inward-bound pathway from the cell exterior to the endosomal and lysosomal compartments. The transport of proteins or lipids from the ER towards the plasma membrane, endosomes or lysosomes is referred to as the anterograde pathway. In contrast to this, proteins and lipids can also be recycled from the cell surface/endosomes back to intracellular compartments through the retrograde pathway [START_REF] Bonifacino | Retrograde transport from endosomes to the trans-Golgi network[END_REF][START_REF] Johannes | Tracing the retrograde route in protein trafficking[END_REF]. Intracellular membrane transport has been extensively studied in the last decades and led to the identification of a complex variety of lipids and proteins, including RABs, involved in the regulation of this process (Cai et al., 2007a;Hutagalung and Novick, 2011;Stenmark, 2009).

The transport of vesicles from one compartment to the other occurs through different steps, all of which are to some extent regulated by RAB GTPases (Figure 2.7).

The recruitment and assembly of coat proteins (Clathrin, COPI, COPII) leads to membrane distortion and consequently to vesicle budding from the donor compartment. The selective incorporation of cargoes into the forming vesicles is mediated by adaptor proteins and their activation can be mediated by RAB proteins (Figure 2.7A). As an example, in the recycling of mannose-6-phosphate receptors from late endosomes to the trans-Golgi network, the cytosolic tail of the receptor is recognized by the sorting adaptor TIP47, a known RAB9 effector [START_REF] Carroll | Role of Rab9 GTPase in facilitating receptor recruitment by TIP47[END_REF].

After vesicle budding, partial vesicle uncoating occurs. Membrane dissociation of COPI coats is thought to mostly be mediated by ARF1 GTP hydrolysis [START_REF] Bigay | Lipid packing sensed by ArfGAP1 couples COPI coat disassembly to membrane bilayer curvature[END_REF][START_REF] Tanigawa | Hydrolysis of bound GTP by ARF protein triggers uncoating of Golgi-derived COP-coated vesicles[END_REF] whereas COPII dissociation is achieved through SAR1 GTP hydrolysis [START_REF] Aridor | Sequential coupling between COPII and COPI vesicle coats in endoplasmic reticulum to Golgi transport[END_REF][START_REF] Yoshihisa | Requirement for a GTPase-activating protein in vesicle budding from the endoplasmic reticulum[END_REF]. RAB proteins have also been proposed to play a role in this process (Figure 2.7B) through the recruitment of PI kinases or phosphatases [START_REF] Christoforidis | Phosphatidylinositol-3-OH kinases are Rab5 effectors[END_REF][START_REF] Shin | An enzymatic cascade of Rab5 effectors regulates phosphoinositide turnover in the endocytic pathway[END_REF] which alter the vesicle composition and can consequently cause the dissociation of PI-binding protein coats [START_REF] Haucke | Phosphoinositide regulation of clathrin-mediated endocytosis[END_REF].

Cargo containing vesicles are then transported to their final destination by diffusion or by motormediated transport along cytoskeletal tracks. RAB proteins are also involved in this process through recruitment of motor adaptor proteins or direct interaction with motors (Figure 2.7C) [START_REF] Hammer | Rabs grab motors: defining the connections between Rab GTPases and motor proteins[END_REF][START_REF] Seabra | Rab GTPases and myosin motors in organelle motility[END_REF].

Dyneins and kinesins are microtubule-based motors which move towards the minus end and either the plus or minus end of microtubules, respectively. The best described example of RABkinesin direct interaction is between RAB6 and KIF20A (originally called Rabkinesin-6) [START_REF] Echard | Interaction of a Golgi-associated kinesin-like protein with Rab6[END_REF] which mediates transport from the Golgi and was shown to be required for successful cleavage furrow and cytokinesis [START_REF] Hill | The Rab6-binding kinesin, Rab6-KIFL, is required for cytokinesis[END_REF]. Many studies however indicate that RAB proteins, such as RAB 3, 6, 9, 11 and 27 mostly interact indirectly with motors via adaptor proteins [START_REF] Horgan | Rab GTPases and microtubule motors[END_REF]. Another type of indirect interaction can be illustrated with RAB5 and KIF16B. RAB5 recruits VPS34, a phosphoinositide 3-kinase, to early endosomes. VPS34 locally synthesizes PI(3)P and consequently recruits KIF16B [START_REF] Hoepfner | Modulation of receptor recycling and degradation by the endosomal kinesin KIF16B[END_REF].

While some RAB proteins, such as endosomal RAB4, were shown to directly interact with dynein motors [START_REF] Bielli | The small GTPase Rab4A interacts with the central region of cytoplasmic dynein light intermediate chain-1[END_REF], most RAB-dynein interactions are indirect. RAB11 associates with dynein-1 through the binding of its effector FIP3 to the LIC1 and LIC2 (light intermediate chain)

subunits [START_REF] Horgan | Rab11-FIP3 links the Rab11 GTPase and cytoplasmic dynein to mediate transport to the endosomal-recycling compartment[END_REF]. RAB7 associates with the cytoplasmic dynein-1-dynactin motor complex through the binding of its effector RILP to the dynactin p150 Glued subunit [START_REF] Jordens | The Rab7 effector protein RILP controls lysosomal transport by inducing the recruitment of dynein-dynactin motors[END_REF]. RAB6 was also shown to interact with the p150 Glued subunit of the dynactin complex and with the dynactin-interacting protein Bicaudal D2 [START_REF] Short | The Rab6 GTPase regulates recruitment of the dynactin complex to Golgi membranes[END_REF].

Myosins are actin-based motors which, with the exception of Myosin VI, move towards the barbed (+) end of actin filaments. The best characterized RAB-Myosin interactions involve mainly class V myosins. RAB27 and Myosin Va interact with each other through Melanophilin and MyRIP (Myosin and RAB interacting protein), which allows the regulation of melanosome transport in melanocytes [START_REF] El-Amraoui | MyRIP, a novel Rab effector, enables myosin VIIa recruitment to retinal melanosomes[END_REF][START_REF] Matesic | Mutations in Mlph, encoding a member of the Rab effector family, cause the melanosome transport defects observed in leaden mice[END_REF]. Later on, Myosin Va was shown to directly associate with RAB3A [START_REF] Wollert | Myosin5a tail associates directly with Rab3A-containing compartments in neurons[END_REF], RAB8A/RAB10 [START_REF] Roland | Alternative splicing in class V myosins determines association with Rab10[END_REF], and to multiple other RABs, including RAB6, RAB14 and RAB39B, via three distinct RAB-binding domains [START_REF] Lindsay | Identification and characterization of multiple novel Rab-myosin Va interactions[END_REF]. Additionally, RAB6 regulates the fission of transport vesicles from the Golgi through its direct interaction with Myosin II [START_REF] Miserey-Lenkei | Rab and actomyosin-dependent fission of transport vesicles at the Golgi complex[END_REF].

Finally, vesicle fusion with the acceptor compartment is mediated by the pairing of SNAREs (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) (Figure 2.7E) (see [START_REF] Jahn | SNAREs--engines for membrane fusion[END_REF] for extensive review about SNAREs). A SNARE on a transport vesicle (v-SNARE) interacts with its cognate SNARE-binding partner (t-SNARE) and forms a complex. The free energy release, derived from the SNARE complex assembly, is able to overcome the repulsive forces between opposing bilayers and to subsequently drive the fusion of the two opposite membranes [START_REF] Hanson | Structure and conformational changes in NSF and its membrane receptor complexes visualized by quick-freeze/deep-etch electron microscopy[END_REF][START_REF] Jena | Assembly and disassembly of SNAREs in membrane fusion[END_REF]. RAB proteins are also involved in this process through interaction with proteins that regulate SNARE function [START_REF] Novick | Interactions between Rabs, tethers, SNAREs and their regulators in exocytosis[END_REF][START_REF] Ohya | Reconstitution of Rab-and SNARE-dependent membrane fusion by synthetic endosomes[END_REF]. As an example, two RAB5 effector proteins EEA1 (early endosome antigen 1) and Rabenosyn-5 respectively interact with the SNARE proteins Syntaxin-6 and Syntaxin- 13 (McBride et al., 1999;[START_REF] Simonsen | The Rab5 effector EEA1 interacts directly with syntaxin-6[END_REF], and VPS45, a member of the SEC1 family of SNARE regulators [START_REF] Morrison | Regulation of early endosomal entry by the Drosophila tumor suppressors Rabenosyn and Vps45[END_REF][START_REF] Nielsen | Rabenosyn-5, a novel Rab5 effector, is complexed with hVPS45 and recruited to endosomes through a FYVE finger domain[END_REF].

RABs and membrane tethering

Prior to vesicle fusion, the interaction between the vesicle and its target membrane needs to be established and this is referred to as membrane tethering (Figure 2.7D). Proteins and protein complexes called tethers or tethering factors are key to this process [START_REF] Brocker | Multisubunit tethering complexes and their role in membrane fusion[END_REF][START_REF] Sztul | Role of tethering factors in secretory membrane traffic[END_REF]. All tethering factors can broadly be divided into two categories: coiled-coil tethers and multi-subunit tethers. Tethering is mostly achieved through tether interactions with RAB GTPases (Hutagalung and Novick, 2011;Stenmark, 2009) and coat proteins (Figure 2.8C) (Cai et al., 2007a;[START_REF] Trahey | Transport vesicle uncoating: it's later than you think[END_REF]. A large number of tethers are RAB effectors and some have been described to also act as RABGEFs.

Most coiled-coil tethers have been found in the Golgi and are now referred to as Golgins [START_REF] Short | Golgins and GTPases, giving identity and structure to the Golgi apparatus[END_REF]. Due to their large size and elongated structure (up to 3000 residues) they are able to bridge distances of more than 200 nm. They also exhibit multiple RAB binding sites in their structure (Figure 2.8A). Because of these properties Golgins are thought to potentially serve as scaffolds to recruit RAB containing vesicles to the correct side of the Golgi [START_REF] Sinka | Golgi coiled-coil proteins contain multiple binding sites for Rab family G proteins[END_REF].

The previously mentioned RAB5 effectors EEA1 and Rabenosyn-5 involved in SNARE-mediated membrane fusion were also identified as coiled-coil tethering factors [START_REF] Nielsen | Rabenosyn-5, a novel Rab5 effector, is complexed with hVPS45 and recruited to endosomes through a FYVE finger domain[END_REF][START_REF] Simonsen | EEA1 links PI(3)K function to Rab5 regulation of endosome fusion[END_REF]. Both effectors possess a FYVE domain that binds PI(3)P which is usually found on early endosomes. Specific RAB effectors can therefore connect membranes and simultaneously recruit specific SNARE proteins that mediate fusion [START_REF] Dubuke | The Secret Life of Tethers: The Role of Tethering Factors in SNARE Complex Regulation[END_REF]. Multi-subunit tethering complexes can also mediate vesicle tethering via their interaction with RAB (B) or coat (C) proteins. Some tethering factors might also recruit SNAREs or their regulators and thereby can also regulate vesicle fusion. Adapted from [START_REF] Brocker | Multisubunit tethering complexes and their role in membrane fusion[END_REF].

Multi-subunit tethering complexes can be divided into two general groups: those required for membrane fusion with organelles of the secretory pathway (Dsl1p, COG, GARP, exocyst) and those of the endo-lysosomal pathway (CORVET and HOPS). Each of these protein complexes, except Dsl1p, were shown to act as RAB protein effectors (Figure 2.8B) (see [START_REF] Brocker | Multisubunit tethering complexes and their role in membrane fusion[END_REF] for extensive review) and they all seem to couple the recognition of membranes via RAB GTPases with the subsequent SNARE-mediated membrane fusion, again demonstrating that membrane tethering and fusion are closely related processes [START_REF] Dubuke | The Secret Life of Tethers: The Role of Tethering Factors in SNARE Complex Regulation[END_REF].

The TRAPP complexes, known to act as GEFs for RAB1/YPT1 and YPT31/YPT32 [START_REF] Jones | The TRAPP complex is a nucleotide exchanger for Ypt1 and Ypt31/32[END_REF][START_REF] Wang | TRAPP stimulates guanine nucleotide exchange on Ypt1p[END_REF] were shown to promote membrane tethering through their interaction with COPII proteins (Cai et al., 2007b;[START_REF] Sacher | TRAPP I implicated in the specificity of tethering in ER-to-Golgi transport[END_REF] and can thus also be referred to as tethering factors.

Thus, abundant evidence indicates that RAB proteins and their effectors promote membrane tethering. However, tethering activity was originally mostly assessed in vivo, or by using cell extracts. Due to the large amount of molecules in these systems, it remains very challenging to identify their precise mode of action; some might directly mediate tethering while others only act as upstream regulators that promote tethering but do not directly mediate it. As a consequence, more and more biological processes were reconstituted in vitro using minimal reaction systems of purified components, in order to explore more precisely their mechanisms of action [START_REF] Cheung | Protein flexibility is required for vesicle tethering at the Golgi[END_REF][START_REF] Drin | Asymmetric tethering of flat and curved lipid membranes by a golgin[END_REF][START_REF] Lurick | Multivalent Rab interactions determine tether-mediated membrane fusion[END_REF][START_REF] Murray | An endosomal tether undergoes an entropic collapse to bring vesicles together[END_REF].

Interestingly, by using this type of chemically defined system, the yeast RAB5 ortholog VPS21 as This additional study also suggests that RAB proteins could potentially directly drive membrane tethering during vesicle transport events, possibly via a RAB-RAB interaction. It is important to note that membrane tethering events (with no subsequent fusion) can also mediate transient contact sites mostly between the ER and other organelles such as Golgi, Mitochondria, endosomes, plasma membrane and lipid droplets. These membrane contact sites have been described to be crucial for the exchange of material such as lipids (especially cholesterol) and Ca 2+ between compartments (see [START_REF] Jackson | Lipids and Their Trafficking: An Integral Part of Cellular Organization[END_REF][START_REF] Phillips | Structure and function of ER membrane contact sites with other organelles[END_REF] for extensive reviews). RAB7 was shown to promote ER-late endosome contact sites through its interaction with Protrudin, an ER localized protein [START_REF] Raiborg | Repeated ER-endosome contacts promote endosome translocation and neurite outgrowth[END_REF]. It has been suggested that these contact sites may be regulated by activated RAB7 since Protrudin-RAB7 interaction only occurs when RAB7 is GTP-bound and expression of dominant-active RAB7 (RAB7Q67L mutant) leads to an increasing amount of contact sites [START_REF] Raiborg | Repeated ER-endosome contacts promote endosome translocation and neurite outgrowth[END_REF].

As membrane tethering was also shown to occur in vitro in the unique presence of RAB proteins [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF][START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF], the formation of contact sites between organelles could also be driven by heterotypic interactions of RAB proteins from two opposite membranes.

Focus on the RAB proteins used in this study

RAB1 and the ER-Golgi intermediate compartment

RAB1 has been highly conserved through evolution. It was shown to be a functional homolog of YPT1 [START_REF] Haubruck | The ras-related mouse ypt1 protein can functionally replace the YPT1 gene product in yeast[END_REF] and exists as two isoforms RAB1A and RAB1B which share 93% amino acid identity. RAB1 was firstly shown to localize to the Golgi [START_REF] Preuss | Characterization of the Saccharomyces Golgi complex through the cell cycle by immunoelectron microscopy[END_REF][START_REF] Segev | The yeast GTP-binding YPT1 protein and a mammalian counterpart are associated with the secretion machinery[END_REF] and has also later been shown to localize to the pre-Golgi intermediate compartment [START_REF] Saraste | Localization of the small GTP-binding protein rab1p to early compartments of the secretory pathway[END_REF]. RAB1 was first suggested to regulate the anterograde transport of cargoes between the ER and the cis-Golgi [START_REF] Plutner | Rab1b regulates vesicular transport between the endoplasmic reticulum and successive Golgi compartments[END_REF][START_REF] Tisdale | GTP-binding mutants of rab1 and rab2 are potent inhibitors of vesicular transport from the endoplasmic reticulum to the Golgi complex[END_REF] but subsequent studies have shown that RAB1 is also required for retrograde transport in Saccharomyces cerevisiae [START_REF] Kamena | Ypt1p is essential for retrograde Golgi-ER transport and for Golgi maintenance in S. cerevisiae[END_REF] and seems to play a role in autophagy [START_REF] Huang | Antibacterial autophagy occurs at PI(3)P-enriched domains of the endoplasmic reticulum and requires Rab1 GTPase[END_REF][START_REF] Zoppino | Autophagosome formation depends on the small GTPase Rab1 and functional ER exit sites[END_REF].

Interestingly, RAB1 is also important for Golgi biogenesis and structure maintenance [START_REF] Galea | A high-content screening microscopy approach to dissect the role of Rab proteins in Golgi-to-ER retrograde trafficking[END_REF], as well as for the extension of the intermediate compartment in the pericentriolar region [START_REF] Marie | The function of the intermediate compartment in pre-Golgi trafficking involves its stable connection with the centrosome[END_REF]. Most RAB1 effectors were identified as tethering factors (COG complex, Golgin-84, etc.) [START_REF] Satoh | Golgin-84 is a rab1 binding partner involved in Golgi structure[END_REF][START_REF] Suvorova | The Sec34/Sec35p complex, a Ypt1p effector required for retrograde intra-Golgi trafficking, interacts with Golgi SNAREs and COPI vesicle coat proteins[END_REF], therefore highly suggesting that RAB1 is mostly involved in the regulation of vesicle-compartment tethering during transport.

RAB6 and the Golgi

RAB6 is another RAB gene conserved from yeast to humans (Pereira-Leal and [START_REF] Pereira-Leal | Evolution of the Rab family of small GTP-binding proteins[END_REF]. In human, the RAB6 subfamily is composed of the splice variants RAB6A and RAB6A' [START_REF] Echard | Alternative splicing of the human Rab6A gene generates two close but functionally different isoforms[END_REF] which differ by only 3 amino acids but were shown to play non-overlapping roles (Figure 2.10) (Del [START_REF] Nery | Rab6A and Rab6A' GTPases play non-overlapping roles in membrane trafficking[END_REF]; and RAB6B which displays 91% identity to RAB6A (most amino acid differences located at the C-terminal hypervariable region) and is preferentially expressed in neuronal cells [START_REF] Opdam | The small GTPase Rab6B, a novel Rab6 subfamily member, is cell-type specifically expressed and localised to the Golgi apparatus[END_REF]. A divergent RAB6 isoform named RAB6C has also been identified [START_REF] Shan | WTH3, a new member of the Rab6 gene family, and multidrug resistance[END_REF][START_REF] Simpson | Systematic subcellular localization of novel proteins identified by large-scale cDNA sequencing[END_REF]. RAB6C, which is very homologous to RAB6A' (97%) but possess an additional C-terminal extension of 46 amino acids, was reported to be a primate-specific retrogene derived from a RAB6A' transcript [START_REF] Young | RAB6C is a retrogene that encodes a centrosomal protein involved in cell cycle progression[END_REF]. RAB6C, which was shown to localize to the centrosome and was suggested to play a role in cell cycle progression [START_REF] Young | RAB6C is a retrogene that encodes a centrosomal protein involved in cell cycle progression[END_REF], is less abundant and less stable than RAB6A', does not localize to membranes and exhibits a reduced GTP-binding affinity.

All RAB6A/A' and B localize to the medial and trans Golgi cisternae and trans-Golgi network (TGN) as well as on dynamic tubulovesicular carriers that move along microtubules [START_REF] Nizak | Recombinant antibodies to the small GTPase Rab6 as conformation sensors[END_REF]. RAB6 was shown to regulate trafficking from endosomes to the Golgi (Figure 2.10) [START_REF] Mallard | Early/recycling endosomes-to-TGN transport involves two SNARE complexes and a Rab6 isoform[END_REF][START_REF] Young | Regulation of microtubule-dependent recycling at the trans-Golgi network by Rab6A and Rab6A[END_REF]. This is thought to partially be mediated by the involvement of RAB6 in the targeting and tethering of endosome derived vesicles to the TGN membrane, through its interaction with the multi-subunit tether GARP [START_REF] Liewen | Characterization of the human GARP (Golgi associated retrograde protein) complex[END_REF] Perez-Victoria and [START_REF] Perez-Victoria | Dual roles of the mammalian GARP complex in tethering and SNARE complex assembly at the trans-golgi network[END_REF].

The major pool of RAB6 is located on Golgi membranes and RAB6 has been suggested to play a key role in intra-Golgi transport and Golgi homeostasis (Figure 2.10). RAB6 associates with several golgins [START_REF] Short | Golgins and GTPases, giving identity and structure to the Golgi apparatus[END_REF] and other tethering factors such as the COG complex [START_REF] Fukuda | Large scale screening for novel rab effectors reveals unexpected broad Rab binding specificity[END_REF], and thereby drives intra-Golgi membrane tethering.

Finally, RAB6 was also suggested to be implicated in Golgi-to-ER [START_REF] White | Rab6 coordinates a novel Golgi to ER retrograde transport pathway in live cells[END_REF] and Golgito-plasma membrane [START_REF] Grigoriev | Rab6 regulates transport and targeting of exocytotic carriers[END_REF] transport (Figure 2.10) where it can trigger myosin II and actin -dependent fission of transport carriers [START_REF] Miserey-Lenkei | Rab and actomyosin-dependent fission of transport vesicles at the Golgi complex[END_REF] but also act in cooperation with its effector Bicaudal D2 to recruit the dynamin/dynactin motor complex necessary for vesicle translocation [START_REF] Matanis | Bicaudal-D regulates COPI-independent Golgi-ER transport by recruiting the dynein-dynactin motor complex[END_REF][START_REF] Short | The Rab6 GTPase regulates recruitment of the dynactin complex to Golgi membranes[END_REF]. Finally, RAB6 was also shown to be involved in mitosis and cytokinesis [START_REF] Bardin | Phenotypic characterisation of RAB6A knockout mouse embryonic fibroblasts[END_REF][START_REF] Miserey-Lenkei | A role for the Rab6A' GTPase in the inactivation of the Mad2spindle checkpoint[END_REF].

RAB4 / RAB5 / RAB11 and the endosomal system

As a very general picture, the entry of cargoes from the plasma membrane to early endosomes (EEs) is mediated by RAB5 (Figure 2.11). Cargoes can eventually be recycled back to the cell surface by a fast RAB4-mediated pathway and a slow RAB11-mediated process via late recycling endosomes. Of note, RAB5 can also mediate cargo degradation from EEs to multivesicular bodies and subsequently to lysosomes via RAB7; and RAB9 controls the retrograde transport of cargoes from late endosomes to the TGN. All these endosomal RAB proteins were described to segregate into specific domains on endosomes (Figure 2.11) [START_REF] Barbero | Visualization of Rab9-mediated vesicle transport from endosomes to the trans-Golgi in living cells[END_REF]Sonnichsen et al., 2000). RAB5 is a key component of the early endocytic pathway that localizes to the cytosolic side of the plasma membrane, endocytic vesicles and EEs [START_REF] Chavrier | Localization of low molecular weight GTP binding proteins to exocytic and endocytic compartments[END_REF]. RAB5 has three ubiquitously expressed isoforms; RAB5A, RAB5B and RAB5C that share over 90% sequence homology (the main differences are located at the N-terminal extremity and the C-terminal hypervariable region). These isoforms appear to be functionally redundant in endocytic trafficking events [START_REF] Bucci | Co-operative regulation of endocytosis by three Rab5 isoforms[END_REF][START_REF] Gurkan | Large-scale profiling of Rab GTPase trafficking networks: the membrome[END_REF], although RAB5C was recently described to selectively modulate the growth factor-mediated activation of RAC1 and consequently cell motility [START_REF] Chen | Rab5 isoforms orchestrate a "division of labor" in the endocytic network; Rab5C modulates Rac-mediated cell motility[END_REF].

RAB5 regulates the transport of cargo from the plasma membrane to EEs [START_REF] Bucci | The small GTPase rab5 functions as a regulatory factor in the early endocytic pathway[END_REF], the generation of PI(3)P lipids on EEs [START_REF] Christoforidis | Phosphatidylinositol-3-OH kinases are Rab5 effectors[END_REF], EE fusion [START_REF] Barbieri | Rab5 regulates the dynamics of early endosome fusion[END_REF] and EE motility on cytoskeletal tracks through its interaction with effector proteins and motors [START_REF] Nielsen | Rab5 regulates motility of early endosomes on microtubules[END_REF]. RAB5 was also shown to activate signaling pathways from EEs [START_REF] Benmerah | Endocytosis: signaling from endocytic membranes to the nucleus[END_REF][START_REF] Miaczynska | Not just a sink: endosomes in control of signal transduction[END_REF]. As previously mentioned, RAB5 can promote the generation of PI(3)P lipids on EEs via its ability to recruit PI(3)P-kinase [START_REF] Christoforidis | Phosphatidylinositol-3-OH kinases are Rab5 effectors[END_REF], and regulate the levels of PI(4,5)P 2 through the recruitment of 5-phosphatases [START_REF] Hyvola | Membrane targeting and activation of the Lowe syndrome protein OCRL1 by rab GTPases[END_REF]. This change in membrane composition allows the recruitment of RAB5 effectors, such as EEA1 and Rabenosyn-5 that can act as tethering factors, and also mediates the direct or indirect assembly of SNARE proteins, thus enabling EE fusion [START_REF] Mills | Relationships between EEA1 binding partners and their role in endosome fusion[END_REF][START_REF] Morrison | Regulation of early endosomal entry by the Drosophila tumor suppressors Rabenosyn and Vps45[END_REF].

The RAB4 subfamily of proteins is comprised of two isoforms in human RAB4A and RAB4B which display 93% sequence homology and are ubiquitously expressed, even though RAB4A is highly expressed in the brain [START_REF] Hoogenraad | Neuron specific Rab4 effector GRASP-1 coordinates membrane specialization and maturation of recycling endosomes[END_REF] and RAB4B is the predominant isoform in B cells [START_REF] Krawczyk | Expression of RAB4B, a protein governing endocytic recycling, is co-regulated with MHC class II genes[END_REF]. Both isoforms are thought to have redundant functions in recycling [START_REF] Krawczyk | Expression of RAB4B, a protein governing endocytic recycling, is co-regulated with MHC class II genes[END_REF]. RAB4 was shown to localize to EEs [START_REF] Van Der Sluijs | The small GTP-binding protein rab4 is associated with early endosomes[END_REF], as well as to RAB11-positive recycling endosomes [START_REF] Trischler | Biochemical analysis of distinct Rab5-and Rab11positive endosomes along the transferrin pathway[END_REF]. In contrast to RAB5, RAB4

mediates the recycling of cargo from EEs directly back to the plasma membrane (fast recycling route) [START_REF] Sheff | The receptor recycling pathway contains two distinct populations of early endosomes with different sorting functions[END_REF]van der Sluijs et al., 1992b) or directs the sorting of cargoes to the endosomal recycling compartment (ERC) via a "slow recycling" route [START_REF] Deneka | Rab'ing up endosomal membrane transport[END_REF].

RAB4 is also involved in regulating other more specialized trafficking pathways, such as the transport of the glucose transporter GLUT4 in adipocytes [START_REF] Cormont | Potential role of Rab4 in the regulation of subcellular localization of Glut4 in adipocytes[END_REF] or the processing of a receptor-mediated antigen in B lymphocytes [START_REF] Lazzarino | The monomeric guanosine triphosphatase rab4 controls an essential step on the pathway of receptor-mediated antigen processing in B cells[END_REF]. Interestingly, Rabenosyn-5, in addition to be a RAB5 effector, has also been shown to bind GTP-bound RAB4

and has been proposed to act as a linker between the RAB4-RAB5 domains on EEs and to coordinate the sorting of cargoes from EEs (de [START_REF] De Renzis | Divalent Rab effectors regulate the subcompartmental organization and sorting of early endosomes[END_REF].

Alternatively, cargoes can be recycled back to the plasma membrane via a late endocytic recycling compartment (ERC) on which RAB4 but also RAB11 are present and form distinct domains (Figure 2.11) (Sonnichsen et al., 2000). The RAB11 protein family is composed of three isoforms:

RAB11A and RAB11B which have redundant functions and are ubiquitously expressed, although RAB11B is more abundant in heart, brain and testes [START_REF] Lai | Molecular analysis of mouse Rab11b: a new type of mammalian YPT/Rab protein[END_REF]; and RAB25 (also known as RAB11C) is expressed in polarized epithelial cells derived from colon, lung and kidney tissues [START_REF] Goldenring | Identification of a small GTP-binding protein, Rab25, expressed in the gastrointestinal mucosa, kidney, and lung[END_REF]. RAB11A is the best studied and characterized member of this subfamily and has been found to localize to the ERC [START_REF] Ullrich | Rab11 regulates recycling through the pericentriolar recycling endosome[END_REF] and to be implicated in the transport of internalized receptor from ERC to the plasma membrane [START_REF] Maxfield | Endocytic recycling[END_REF]. The recycling of cargos from the ERC to the plasma membrane was shown to be highly dependent on RAB11 and its effectors, which mediate transport along actin filaments [START_REF] Hales | Rab11 family interacting protein 2 associates with Myosin Vb and regulates plasma membrane recycling[END_REF] and subsequent tethering and fusion [START_REF] Takahashi | Rab11 regulates exocytosis of recycling vesicles at the plasma membrane[END_REF].

Additionally, RAB11 and his yeast homolog YPT31/YPT32 were also shown to localize to the TGN [START_REF] Urbe | Rab11, a small GTPase associated with both constitutive and regulated secretory pathways in PC12 cells[END_REF] and to regulate transport from the Golgi to the plasma membrane [START_REF] Satoh | Rab11 mediates post-Golgi trafficking of rhodopsin to the photosensitive apical membrane of Drosophila photoreceptors[END_REF] and the recycling of plasma membrane proteins back to the Golgi [START_REF] Chen | Ypt31/32 GTPases and their novel Fbox effector protein Rcy1 regulate protein recycling[END_REF].

RAB35 and the plasma membrane

RAB35 was initially called RAB1C due to its high sequence similarity with RAB1A and RAB1B and for this reason only started to mobilize attention in the last 10 years. RAB35 was shown to localize to the plasma membrane and to endosomes, and to be involved in endocytic recycling, cytokinesis processes [START_REF] Dambournet | Rab35 GTPase and OCRL phosphatase remodel lipids and F-actin for successful cytokinesis[END_REF]Kouranti et al., 2006) and exosome secretion [START_REF] Hsu | Regulation of exosome secretion by Rab35 and its GTPase-activating proteins TBC1D10A-C[END_REF]. RAB35 plasma membrane localization was suggested to be dependent on its highly conserved polybasic C-terminal extremity, through its direct interaction with negatively charged lipids, PI(4,5)P 2 and PI(3,4,5)P 3 [START_REF] Heo | PI(3,4,5)P3 and PI(4,5)P2 lipids target proteins with polybasic clusters to the plasma membrane[END_REF]Li et al., 2014). RAB35 was also described to regulate PI membrane composition through its interaction with OCRL (Oculo-Cerebro-Renal syndrome of Lowe) [START_REF] Cauvin | Rab35 GTPase Triggers Switch-like Recruitment of the Lowe Syndrome Lipid Phosphatase OCRL on Newborn Endosomes[END_REF][START_REF] Dambournet | Rab35 GTPase and OCRL phosphatase remodel lipids and F-actin for successful cytokinesis[END_REF]. OCRL, which is also an effector for RAB5 and RAB6 [START_REF] Hyvola | Membrane targeting and activation of the Lowe syndrome protein OCRL1 by rab GTPases[END_REF], was shown to selectively hydrolyze PI(4,5)P 2 into PI(4)P through its phosphatase domain [START_REF] Lowe | Structure and function of the Lowe syndrome protein OCRL1[END_REF] and to thereby regulate PI(4,5)P 2 levels on endosomes. RAB35 also plays a fundamental role in actin-based events at the plasma membrane leading to its subsequent involvement in a large variety of cellular functions, such as endosomal trafficking, phagocytosis, cell migration and others [START_REF] Klinkert | Rab35 GTPase: A Central Regulator of Phosphoinositides and F-actin in Endocytic Recycling and Beyond[END_REF].

Chapter 3

3 Materials and Methods Proteins, and more specifically RAB5A, were purified using the following method.

Expression: RAB5A expression and purification were performed as previously described (Oesterlin, 2012). RAB5A was expressed as a cleavable His6-MBP-fusion construct. Expression was performed in BL21(DE3) cells (Novagen/Merck) and was induced with 0.2 mM IPTG at 37°C for 5h.

The following protein purification was performed in several steps. First cell lysis was achieved by re-suspending them in 150 mL lysis buffer consisting of 50 mM Hepes pH7.5, 500 mM LiCl, 2 mM β-Mercaptoethanol, 1 mM MgCl 2 , 10 µM GDP, 5% Glycerol supplemented with 1 protease inhibitor tablet (complete EDTA free, Roche) per 50 mL lysis buffer, 2 µg/mL Pepstatin and 1 mM PMSF. The cell suspension was incubated for 30 min on ice under continuous stirring and cells were passed through a cell disruptor (Constant Systems Ltd) at 2.5 kBar. As an alternative to the cell disruptor, cells can also be sonicated. The cell lysate was centrifuged at 20000 rpm at 4°C for 45 minutes and supernatant was filtered with a 0.22 µm pore size filter (Millipore Express PLUS).

Protein purification / affinity column n°1: RAB5A was then isolated by affinity chromatography using 2 x 5 mL HiTrap TM Chelating HP columns (GE Healthcare) charged with

Ni 2+ mounted on an AKTA purifier system (Amersham Biosciences). As an alternative, Protino Ni-NTA Agarose beads (Macherey-Nagel) packed on an empty column can also be used. Two buffers A and B were used; both the same as the lysis buffer (without protease inhibitors) and buffer B supplemented with 500 mM Imidazole. The cell extract was loaded on the column and sequential washes with buffer A, 2% buffer B and 4 % buffer B were performed to get rid of contaminants which bind unspecifically to the column (Figure 3.1). Elution was performed with 200 mL continuous gradient of buffer B (4% to 100%) and 3 mL fractions were collected (Figure 3.1). In the case of Ni-NTA Agarose beads, elution was performed with 5%, 10% and 15% buffer B.

Protein purification / digestion: In the case of RAB5A, the protein was expressed using an Nterminal MBP tag that now has to be removed. The MBP tag was followed by a Tobacco etch virus Protein samples at each step were ran on a 12% SDS-PAGE gel to identify the RAB5A positive fractions and pool them together. Protein concentration was usually determined at this step using a Bradford protein assay (Bio-Rad).

Protein purification / affinity column n°2: This second protein isolation, meant to separate the His6-MBP tag from the digested RAB5A protein, was performed the same way as before except that washes were only done with buffer A, 4% buffer B and 8% buffer B. At this step, the protein of interest is normally collected in the "flow-through" (when the dialysis product is loaded on the column) while the His6 tag is retained on the column. However, in this particular case, digested RAB5A was found to unspecifically interact with His6-MBP and thus most of it was only collected in the 4% buffer B wash.

The protein was then concentrated to a 3 mL final volume using Amicon Ultra 15 MWCO 10.000 Da concentration tube.

Protein purification / Gel filtration column:

To further isolate RAB5A from other protein contaminants, a gel filtration (or size-exclusion) chromatography can be performed using a HiLoad 16/600 Superdex 75 column (GE Healthcare) mounted on the AKTA system with a 4 mL loop. The Gel Filtration column was equilibrated and performed using a buffer consisting of 25 mM Hepes pH7.5, 200 mM NaCl, 1 mM MgCl 2 , 2 mM DTE, 10 µM GDP, 5% Glycerol. The first 30 mL were not collected as it corresponds to the dead volume of the column. 60 fractions of 1.5 mL were then collected and loaded on a 12% SDS-PAGE gel. The fractions of interest [START_REF] Hutagalung | Role of Rab GTPases in membrane traffic and cell physiology[END_REF][START_REF] Baumgart | Imaging coexisting fluid domains in biomembrane models coupling curvature and line tension[END_REF][START_REF] Roux | Role of curvature and phase transition in lipid sorting and fission of membrane tubules[END_REF][START_REF] Pechlivanis | Hydrophobic modifications of Ras proteins by isoprenoid groups and fatty acids--More than just membrane anchoring[END_REF][START_REF] Gomes | Membrane targeting of Rab GTPases is influenced by the prenylation motif[END_REF][START_REF] Wu | Membrane targeting mechanism of Rab GTPases elucidated by semisynthetic protein probes[END_REF][START_REF] Stenmark | Rab GTPases as coordinators of vesicle traffic[END_REF][START_REF] Smeland | Geranylgeranylated Rab proteins terminating in Cys-Ala-Cys, but not Cys-Cys, are carboxyl-methylated by bovine brain membranes in vitro[END_REF][START_REF] Leung | Rab GTPases containing a CAAX motif are processed post-geranylgeranylation by proteolysis and methylation[END_REF][START_REF] Jang | Mechanisms of membrane binding of small GTPase K-Ras4B farnesylated hypervariable region[END_REF] in this case) were then pooled together, the protein concentration was measured by Bradford assay and proteins were aliquoted and stored at -80°C until further use.

Many other proteins were used for this study. Expression and purification of these proteins were performed very similarly to that of RAB5 and varying details can be found in Table 3.1.

In vitro modifications of RAB and GST proteins

As previously mentioned in chapter 2, following their synthesis RAB proteins undergo posttranslational modifications (addition of the prenyl group) and get activated/deactivated. In our in vitro model, purified RAB proteins were also activated/deactivated and prenylated.

Nucleotide exchange reaction

During the purification process, most buffers used contain GTP or GDP nucleotides which are required for protein stabilization. However, nucleotide exchange still needs to be performed to make sure that all proteins bind a given nucleotide.

Nucleotide exchange was induced chemically. Briefly, RAB proteins are incubated with a 20 fold molar excess of the desired nucleotide and a 5 fold molar excess of EDTA (over MgCl 2 in buffer), to destabilize the nucleotide binding pocket, for 3 h at 25°C. For protein activation, RAB6 was exchanged to GTP (Sigma Aldrich) and RAB1, RAB4, RAB5, RAB11, RAB35 were exchanged to GppNHp (Jena Bioscience), a non-hydrolysable analog of GTP. Following incubation, a 2 fold molar excess of MgCl 2 (over EDTA) is added to re-stabilize the binding pocket and the protein sample is run on a NAP-5 / NAP-10 size exclusion column (GE Healthcare) to remove the excess of nucleotide. In order to measure the nucleotide bound state of the protein, an HPLC ion-pair reverse phase chromatography was performed as previously described [START_REF] Eberth | In vitro GEF and GAP assays[END_REF]

Prenylation reaction

The prenylation reaction consists in the addition of one or two C20 geranylgeranyl moieties Monoprenylation reactions were performed at 25°C for 1.5 h with a molar ratio of 0.5:1:5 GGTase1, RAB and GGpp. Molar ratios for the diprenylation reaction were: 1:5:0.5:0.75 RABGGTase:GGpp:RAB:REP and the reaction was performed at 25°C for 4 h. Of note, diprenylation can only be achieved for GDP-bound RABs (REP only recognizes the GDP-bound form) whereas monoprenylation can be performed with both GTP-and GDP-bound RAB proteins.

To control efficient protein prenylation, NBD-Farnesyl pyrophosphate (NBD-Fpp, Jena Bioscience), a C15 fluorescent analog of GeranylGeranyl pyrophosphate, was used as previously described (Dursina et al., 2006). The prenylation reactions with NBD-Fpp were performed using the same molar ratios as for monoprenylation with GGpp. 15 µL of each reaction was loaded on a 15% SDS-polyacrylamide gel and gel fluorescence was visualized using the Ethidium Bromide program of a ChemiDoc imaging system (Bio-Rad) (Figure 3.4).

Labeling reaction

Prior to prenylation, proteins could be labeled using Alexa Fluor TM 488 Sulfodichlorophenol ester (Alexa488 5-SDP ester, Molecular Probes / Life Technologies) or Alexa Fluor TM 568 Succinimidyl ester (Alexa568 NHS ester, Molecular Probes / Life Technologies) and by following the associated protocol. To ensure that the label does not interfere with the binding of some effectors or antibodies, proteins were only labeled on their N-terminal amine group. As the pKa of the terminal amine is lower than that of the amine group containing lysines, specific N-terminal labeling could be achieved using a buffer close to neutral pH (in our case proteins are already at pH 7.5).

The protein was incubated with a 4 fold molar excess of Alexa dye for 2 h at 25°C. A 250 fold molar excess of freshly prepared Hydroxylamine was added and the reaction was incubated for 1h at 25°C. A NAP column (GE Healthcare), pre-equilibrated with the protein buffer, was then run to remove excess unbound fluorophore. Protein concentration was determined by a Bradford assay.

The efficiency of labeling (n * ) which corresponds to the ratio of moles of dye per mole of protein was determined after absorption measurements at the nanodrop (protein and label program, ThermoFisher Scientific) and using the following formula:

𝑛 * = 𝐴 𝑚𝑎𝑥 Ɛ ×𝑝𝑟𝑜𝑡𝑒𝑖𝑛 𝑐𝑜𝑛𝑐𝑒𝑛𝑡𝑟𝑎𝑡𝑖𝑜𝑛 (𝑀) × 𝑑𝑖𝑙𝑢𝑡𝑖𝑜𝑛 𝑓𝑎𝑐𝑡𝑜𝑟 [1]
Where A max is the absorbance of the fluorescent protein at the wavelength maximum of the dye (λ max = 488 nm for Alexa488 and 568 nm for Alexa568) and Ɛ is the fluorescent dye molar extinction coefficient (71,000 M -1 cm -1 and 91,300 M -1 cm -1 for Alexa488 and Alexa568 respectively).

Experimental studies with GUVs

Electroformation with ITO coated slides

GUVs were grown on conductive indium tin oxide(ITO) coated glass slides using the electroformation (EF) technique (Angelova et al., 1992) (Figure 3.5A). 15 µL of a 0.5 mg/mL lipid mix was dried on pre-washed (water, ethanol, water, chloroform) ITO coated slides for a few minutes at 50°C and subsequently under vacuum for at least 2 h. The chamber is then assembled.

Sealing wax (Vitrex, Denmark) is applied around the dried lipid films of two opposing ITO glass slides. The entire chamber is immobilized using two paper clips while 4 layers of Teflon tape on each side are used as a spacer (Figure 3.5A). The dried lipid films were then rehydrated in a sucrose solution (osmolarity between 100 and 430 mOsm, depending on the osmolarity of the protein buffer used for the experiments) and GUVs were grown for 3 h under a sinusoidal voltage (1.1 V, 10 Hz).

GUV growth was most of the time performed at room temperature except in the case of phase separation where it was performed at 50°C. In the latter, the sealing wax which is normally used to construct and isolate the chamber starts to melt which results in leakage of the sucrose solution. For this reason chambers were constructed using a hand-made PDMS chamber which is resistant to such temperatures.

Electroformation with platinum wires

The previously described EF method has the disadvantage to be restricted to growth buffers containing low levels of salt. In the case of lipid mixtures containing specific charged lipids (PI(4,5)P 2 , PI(3)P)), growth needed to be performed in presence of salt for stabilization purposes and the original EF method was not adapted. We thus used a more recently described method suitable for the use of buffers with higher concentrations of salt [START_REF] Meleard | Giant unilamellar vesicle electroformation from lipid mixtures to native membranes under physiological conditions[END_REF].

The EF chamber is made out of a block of Teflon of 5 mm thickness and 40 mm length, with three regularly spaced square wells of 8 x 8 mm (Figure 3.5B). The chamber is also perforated throughout its length by two narrow holes which are 3 mm apart from each other. Two platinum wires of 0.5 mm diameter (LS413074 Goodfellow, UK) are inserted into these holes, cross all three wells and are inserted in the hole on the opposite side of the chamber. These wires come out of the chamber by a few mm which allows us to later connect them to a function generator and to consequently build up an electric field inside the wells. Around 5 µL of a 3 mg/mL lipid mixture was applied drop-by-drop on pre-washed wires (5 min.

sonication steps in acetone, ethanol and water) and subsequent drying under high vacuum was performed.

The next step consists in the mounting of the EF chamber (Figure 3.5B). Sealing wax (Vitrex, Denmark) was applied on the side of the chamber to prevent leakage from the holes (from which the platinum wires come out). The bottom side of the chamber was closed using a glass coverslip fixed to the chamber by the anterior addition of vacuum grease (Dow Corning, USA). All the wells were filled with growth buffer and the chamber was completely sealed with a second glass coverslip. The chamber was connected to a generator through the extended platinum wires on the side and growth was performed at 4 °C overnight under sinusoidal voltage (0.35 V, 500 Hz).

GUVs are then collected by pipetting directly next to the wires.

Generalities of the experimental approach

Experimental setup

As already mentioned, all experiments were performed in vitro using GUVs as controlled model systems and purified Golgi membranes. The experimental setup, which allowed us to perform the measurements (optical tweezer, necessary for tube pulling experiments) and optical acquisition at the same time, was originally developed by P. Bassereau's team at Institut Curie (Sorre, 2010;[START_REF] Sorre | Curvature-driven lipid sorting needs proximity to a demixing point and is aided by proteins[END_REF] and was later adapted in our lab by David Guet, a previous PhD student [START_REF] Guet | Plasma membrane phosphatidylinositol 4-phosphate and 4,5-bisphosphate determine the distribution and function of K-Ras4B but not H-Ras proteins. The Journal of biological chemistry[END_REF]) (Figure 3.6). The setup is originally based on a commercial Nikon eclipse Ti inverted microscope which was modified with the optional stage riser (Nikon), in order to create an extra port for epifluorescence microscopy (port #2). The confocal head consists in an A1R confocal system equipped with four laser lines (405 nm, 488 nm, 561 nm and 640 nm) and has two acquisition modes: Galvano and Resonant. Briefly, the first one allows an optimal signal to noise ratio and is more suited for fluorescence intensity measurements whereas the second one displays a higher scanning speed and thus allows the user to scan at a higher frame rate. In the scope of our experiments we always used the Galvano mode.

We also took advantage of an added DIC bright field visualization module (Figure 3 Because the three modules (confocal microscopy, optical tweezers and DIC) are not usually compatible, the light spectrum was thereby split in three separate channels. Confocal fluorescence microscopy was performed in the visible range (400 nm < λ < 750 nm), optical trapping in the infra-red channel (λ > 900 nm) and DIC was used in the near infra-red channel (750 nm < λ < 900 nm). Bright field imaging video was visualized using a Labview based custom software. Additional details about the setup can be found in D. Guet's thesis [START_REF] Guet | Plasma membrane phosphatidylinositol 4-phosphate and 4,5-bisphosphate determine the distribution and function of K-Ras4B but not H-Ras proteins. The Journal of biological chemistry[END_REF].

Measurement of fluorescence intensity

Many of the performed experiments required measurements of fluorescence intensities (Figure

3.7).

In In the case of chapter 6, fluorescence intensities were measured all over the vesicle with the Fiji software using the oval profile plugin (Figure 3.7B). An oval selection was drawn around the vesicle and the integrated oval profile plugin was set to measure the pixel of highest intensity for each degree (360° in total) of this oval selection. Because some experiments required the assessment of protein membrane binding, each intensity value was subtracted by the background intensity value outside the vesicle.

Measurement of protein density on the membrane

A standard measurement of fluorescence intensities is a good tool to assess and compare the binding or sorting of one labeled protein in different experimental GUV conditions. However, because the different proteins were labeled with various efficiencies, their binding to membranes could not be compared simply by using fluorescence intensity values.

Fluorescence intensities were thus converted into protein surface densities to overcome this issue and also to obtain more representative values of protein binding.

In order to convert fluorescence intensities into protein densities, we needed a calibration standard. In other words, we needed a fluorescent species that we could incorporate in GUVs at known densities. Fluorescence was thus calibrated using GUVs made of BodipyFL-C5-HPC (bodipyFL), a green fluorescent lipid. We synthetized pure EggPC GUVs with various molar ratios of bodipyFL and measured the fluorescence intensities on the membane. BodipyFL surface density (Φ bodipyFL ) and the resulting fluorescence intensity on the membrane (I bodipyFL ) are related by:

Φ 𝑏𝑜𝑑𝑖𝑝𝑦𝐹𝐿 = 𝐴 𝑔𝑎𝑖𝑛 𝐼 𝑏𝑜𝑑𝑖𝑝𝑦𝐹𝐿 (𝑔𝑎𝑖𝑛) [2]
Where A gain is the calibration coefficient at a given confocal photomultiplier tube detector (PMT) gain (same as the one used for protein experiments).

The bodipyFL surface density (Φ bodipyFL ) at each molar ratio was calculated by assuming that all the dye of the initial lipid mix was incorporated into the GUVs. The area per EggPC lipid is approximately 0.7 nm 2 (Nagle, 2000) which results in a number of lipid per µm 2 of 2 ×10 6 0.7 = 2.86 10 6 (where the factor 2 is to take both leaflets into account). The bodipyFL surface density was thus defined as: 𝛷 𝑏𝑜𝑑𝑖𝑝𝑦𝐹𝐿 = 2.86 10 6 𝑥 𝑏𝑜𝑑𝑖𝑝𝑦𝐹𝐿 (x bodipyFL representing the molar ratio of the lipid dye).

Finally A gain was determined using Eq. F is given by the ratio of left and right slopes.

Finally, as the bulk experiments were performed with Alexa488 alone, the correction factor needed to be adjusted with the number of fluorophores per protein. The protein labeling efficiency was thus taken into account by calculating the degree of labeling (n * ) of the protein using Eq. 1.

In the end, the protein density on the GUV membrane (Φ v prot ) was defined by:

Φ 𝑣 𝑝𝑟𝑜𝑡 = 𝐴 𝑔𝑎𝑖𝑛 𝐹 𝑥 𝑛 * × 𝐼 𝑣 𝑝𝑟𝑜𝑡 (𝑔𝑎𝑖𝑛) [3]

Lipid dependent protein recruitment experiments

In the simple case of protein recruitment to differently composed GUVs, a hand-made metal insert (gift from Phong Tran, Institut Curie, crafted by the university of Philadelphia) was used (Figure 3.9). A hand-made PDMS chamber with a 5 mm diameter hole in the middle was fixed to a pre-washed (water, ethanol, water) 22 x 40 mm glass coverslip, inserted and immobilized into the metal insert using two screws. The chamber was incubated with 35 µL of 0.5 mg/mL β-Casein (Sigma) for a couple of minutes to prevent adhesion of the GUV to the glass. The solution was removed and the chamber was washed using 50 µL protein buffer supplemented with 0.1 mg/mL β-Casein. After removal of the protein buffer, 1 or 2 µL of GUV solution and 2 µM proteins are added to the chamber in a total volume of 20 µL (completing to 20 µL with the previously used protein buffer).

Curvature sensing experiments with GUVs

The study of protein curvature sensing was performed by pulling a highly curved tube from a GUV and measuring the enrichment of the protein in the tube as compared to the GUV. In order to do so we used a setup originally developed by the Bassereau team (Institut Curie) and adapted in our lab by David Guet, a previous PhD student [START_REF] Guet | Plasma membrane phosphatidylinositol 4-phosphate and 4,5-bisphosphate determine the distribution and function of K-Ras4B but not H-Ras proteins. The Journal of biological chemistry[END_REF]. Biotinylated GUVs are held on one side by a micropipette which is inserted into the experimental chamber. On their other side these biotinylated GUVs interact with streptavidin coated beads which are immobilized using an optical trap. By moving the micropipette away from the optical trap, the Biotin-streptavidin interaction allows to pull a highly curved tube. Subsequently, tube radius can be modulated by controlling the membrane tension through micropipette aspiration of the GUV (see "micropipette aspiration" section). The sorting (or distribution) ratio of the protein between the highly curved tube and the flat GUV membrane can thus be measured for decreasing tube radii (or increasing tube curvatures). In the next sections the different components of the setup will be presented as well as the overall experiment and data analyses.

Experimental chamber

A 22 x 40 glass slide was cut in its length in two parts (1/3 and 2/3 of the original glass slide) using a Retractable Diamond Scriber (Electron Microscopy Sciences). The two glass parts were washed The chamber was incubated with 200 µL of 0.5 mg/mL β-Casein (Sigma) for a couple of minutes to prevent adhesion of the GUV to the glass. The solution was removed and the chamber was washed using 200 µL protein buffer supplemented with 0.1 mg/mL β-Casein. After removal of the protein buffer, 1 or 2 µL of GUV solution and 100-300 nM proteins were added to the chamber in a total volume of 200 µL (completing to 200 µL with the previously used protein buffer).

Micropipette aspiration

The micropipette aspiration technique is usually used to study the thermoelastic and mechanical properties of cells or synthetic vesicles. The micropipette is a key component of the tube pulling experimental setup as it allowed us to hold and manipulate GUVs and to control membrane tension (and consequently modulate the radius of the tube).

Micropipettes were made of borosilicate capillaries of 1 mm outer diameter and 0.58 mm inner diameter (Harvard Apparatus, UK). The forging of micropipettes is performed in two steps. First, an elongated tip is created on the capillary using a micropipette puller (P-2000, Sutter

Instrument, USA). The latter pulls on both ends of the capillary while heating in the middle with a laser beam leading to the fabrication of two micropipettes with elongated tips that are closed at their end by the merged glass walls. Then, a smooth opening of 5-10 µm is generated at the tip of the micropipette using a microforge (MF-830, Narishige, Japan). A streptavidin coated bead is immobilized using an optical trap and a membrane tube is formed by the biotin-streptavidin interaction. Lowering of the water tank will lead to an increase in membrane tension and consequently a decrease of the tube radius. (Not to scale).

In order to control membrane tension, the micropipette, filled with protein buffer supplemented with 0.1 mg/mL β-Casein (to prevent adhesion of GUVs to the pipette), is connected to a water circuit which ends with a water reservoir of adjustable height (Figure 3.11).

The pipette is fixed to a mechanical micromanipulator which enables us to move it in three directions (x, y, z). The GUV is aspirated by decreasing the pressure inside the micropipette (by lowering the height of the water reservoir). Membrane tension is then proportional to the pressure variation (ΔP) between the micropipette interior and exterior (the chamber) and can be calculated using the Young-Laplace equation [START_REF] Evans | Entropy-driven tension and bending elasticity in condensed-fluid membranes[END_REF]:

𝜎 = 𝛥𝑃𝑅 𝑝 2(1- 𝑅 𝑝 𝑅 𝑣 ) [4]
Where R p is the radius of the pipette and R v is the radius of the vesicle Consequently, in the case of tube pulling experiments, tube radius can be decreased by increasing membrane tension through stronger GUV aspiration.

Optical tweezers

Optical tweezers are micromanipulation tools used to manipulate micron-size particles very precisely. The optical trap was created by tightly focusing a laser beam with an objective of high numerical aperture. The trapped particle experiences a force in the piconewton range, referred to as a gradient force, which pushes it towards the laser focus where the light intensity is the highest. Optical trapping uses infrared light (1063 nm) which is less-invasive and thus has the advantage of causing limited damage to the sample.

An optical tweezer can function as a Hook spring which applies an elastic force on the bead. This force (f) is proportional to the displacement of the bead (x) from its equilibrium position at the trap center (x 0 ) and can thus be defined by:

𝑓 = 𝑘(𝑥 -𝑥 0 ) [5]
Where k is the trap stiffness.

In our experiments, the stiffness of the optical trap (k) at a given laser power needed to be measured (see next section). Charlotte Alibert, another PhD student in the lab, took care of this calibration.

Briefly, the chamber is filled with water containing micron-sized beads (around 3 µm diameter).

One bead is trapped by the optical tweezers and the chamber is moved at a controlled velocity using a piezo-electric actuator (Mad City Labs, MCL-S02456).

The force applied on the bead can be calculated using Stokes' law:

𝑓 = 6𝜋𝜂𝑟𝑣 [6]
Where η is the viscosity of the fluid, r the radius of the bead and v the velocity of the fluid.

The displacement of the bead was assessed with the confocal microscope software (NIS Elements, Nikon) and analyzed using a home-made Matlab code to track the center of the bead.

The trap stiffness (k) at a given laser power was measured using Eq. 5. The force exerted by the trap is plotted as a function of the bead displacement and the resulting slope of the linear fit gives the stiffness value (at a given laser power).

Trap stiffness for other laser power values were also measured the same way. The trap stiffness was plotted as a function of the laser power and the slope of the linear fit gave the value of the trap stiffness per unit power: K = 216.2 ± 6 pN/µm/W.

Calibration and measurements of tube radii

As previously mentioned, GUV tension can be modulated through micropipette aspiration and consequently the tube radius can be varied (Figure 3.11). The tube radius can be measured in two ways.

First, the radius can be measured using the previously described equilibrium relation between the tube radius (R tube ), the force required to hold the tube (f) and the tension of the membrane (σ) [START_REF] Derenyi | Formation and interaction of membrane tubes[END_REF]:

𝑅 𝑡𝑢𝑏𝑒 = 𝑓 4𝜋𝜎 [7]
With f and σ that can respectively be deduced from Eq. 5 and Eq. 4.

However, when tube pulling experiments were performed in presence of proteins, the validity of this equation can be questioned, as the binding of peripheral proteins can change the properties of the membrane (such as membrane tension). Therefore, in the presence of proteins, the tube radius value was assessed with a second method, by measuring the intensity of lipid dye in the tube. Because the studied proteins were all labeled in green with Alexa488 or GFP dyes, we used a red lipid dye consisting of TexasRed-DHPE (Molecular Probes).

Fluorescence in the tube is proportional to the number of fluorophores per unit length and the number of fluorophores is logically proportional to the size of the tube. Thus the fluorescence of the tube is also proportional to its radius (R tube ).

The fluorescence of the tube at a given radius can however vary from one experiment to another and even from one GUV to another if the dye is not homogeneously distributed. The tube fluorescence will also be dependent on the type of fluorescent lipid used. For these reasons the fluorescence of the tube was normalized to that on the vesicle (I ves ) and tube radius was defined to be proportional to

𝐼 𝑡𝑢𝑏𝑒 𝐼 𝑣𝑒𝑠

:

𝑅 𝑡𝑢𝑏𝑒 = 𝑃 𝑐 𝐼 𝑡𝑢𝑏𝑒 𝐼 𝑣𝑒𝑠 [8]
Where P c is the unknown proportionality constant (in nm).

P c was calibrated and determined experimentally by using Eq. 8 in the absence of proteins. In this case, R tube could be measured using Eq. 7 and the P c value was consequently measured by plotting R tube as a function of the 𝐼 𝑡𝑢𝑏𝑒 𝐼 𝑣𝑒𝑠 ratio and taking the slope of the resulting linear fit (Figure 3.12).

Measurements of tube radius and ratio of tube to vesicle fluorescence values were performed with several vesicles and P c was determined to be 239 ± 10 nm. Tube radius is measured using Eq. 7 and plotted as a function of its fluorescence intensity. Each point represents a GUV at a given membrane tension (N = 10 GUVs in total). P c is determined using Eq. 8 by taking the slope of the resulting linear fit.

When looking at the calibration graph, the data seems very dispersed at high tube radii. This could be the result of two things: The GUV bilayer is asymmetrical (due to differences in inside and outside buffers) which can eventually lead to a spontaneous curvature (C 0 ) of the membrane.

The impact of the spontaneous curvature will be higher on large tube radii and Eq. 7 would need to be modified by taking C 0 into account.

Another explanation is that this calibration is less reliable when the diameter of the tube gets closer to the thickness of the confocal volume.

Measurement of sorting ratio

In order to quantify protein sorting to the tube, the fluorescence intensity of the Alexa488 labeled protein (I protein ) was normalized by the intensity of the fluorescent lipid (TexasRed-DHPE, I lipid ) at each tension step increase. The sorting ratio S corresponds to the ratio between the normalized protein intensity on the tube and the same normalized intensity on the GUV (Fig. S4B):

𝑆 = (𝐼 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 /𝐼 𝑙𝑖𝑝𝑖𝑑 ) 𝑡𝑢𝑏𝑒 (𝐼 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 /𝐼 𝑙𝑖𝑝𝑖𝑑 ) 𝑣𝑒𝑠𝑖𝑐𝑙𝑒 [9]
3.3 Experimental studies with purified Golgi membranes

Purification of Rat Liver Golgi stacks

Purified Golgi membranes were obtained by Hugo Bousquet (engineer in the lab) and Lena

Oesterlin by following a previously described protocol [START_REF] Slusarewicz | Isolation of a matrix that binds medial Golgi enzymes[END_REF]. It is important to note, that the purified membrane fraction is only enriched in Golgi membranes and does not consist of pure Golgi membranes. Final buffer consisted of 100 mM Potassium Phosphate pH 6.7, 5 mM MgCl 2 , 250 mM Sucrose.

Experimental chamber

The chamber used for experiments with purified Golgi membranes was handmade (Figure 3.13). The mounted chamber was heated for 5 seconds on a heating and agitating device, which allowed the parafilm to melt and consequently to fix the coverslip to the glass.

Pulling tubes with kinesins

When using purified Golgi stacks as model membranes, protein sorting to tubular structures was monitored. Single tubes could be pulled out of immobilized Golgi membranes using the previously described optical tweezer setup. However, I took advantage of a previously described method [START_REF] Roux | A minimal system allowing tubulation with molecular motors pulling on giant liposomes[END_REF] consisting in Golgi membrane tubulation (can also be performed with

Microtubule preparation

Microtubules were prepared from a tubulin solution (given to us by Jean-Baptiste Manneville, a researcher in the lab). A 50 µL aliquot of tubulin solution was incubated for 15 minutes at 37°C. 2 µL of 1 mM Taxol solution was added in order to stabilize the microtubules and the mix was incubated for 15 minutes at 37°C. The microtubule solution was then centrifuged for 17 minutes at 90000 rpm (TLA 120.2 rotor pre-heated at 37°C) and 37°C; and the pellet was resuspended in supplemented with 30 µM Taxol. Microtubules were stored at room temperature and could be used one or two days after.

Experimental protocol

The experimental chamber was incubated with 5 µL of the previously prepared microtubule solution for 15 minutes in a humid chamber (with coverslip on the bottom so that microtubules can attach to the glass). To note, the direction of injection was always marked to make sure that all injections were performed on the same side of the chamber. The chamber was then incubated for 15 minutes with 5 µL solution of Imi Casein buffer (7 mg/mL β-Casein in 50 mM Imidazole pH 6.7, 50 mM NaCl, 2 mM EGTA, 1 mM MgCl 2 ), supplemented with 30 µM Taxol, to passivate the glass surface. The chamber is then washed with 5 µL solution of 5 µM DTT, 30 µM Taxol in Imi buffer.

On the side, 15 µL of 1.8 mg/mL purified biotinylated kinesin (Kinesin-11401 -BCCP, see protein purification section) was incubated with 5 µL of 1 mg/mL Streptavidin solution (Sigma) on ice for at least 5 minutes. The chamber was then incubated with 5 µL of this kinesin / Streptavidin mix for 15 minutes in the humid chamber (coverslip down).

In parallel , purified Golgi membranes (4 µL) were incubated with 2 µM green or red labeled RAB protein and with an oppositely labeled lipid marker (red Bodipy TR C 5 -ceramide complexed to BSA or green Bodipy FL C 5 -ceramide complexed to BSA, both from Molecular Probes) for 20-30 minutes on ice (5 µL total reaction volume). Subsequently, 7.5 µL of motility buffer (In 100 µL total volume: 5 µM DTT, 30 µM Taxol, 2 mM ATP, 25 mM Glucose, 50 µL 2X Golgi membrane buffer, 3 µL Oxygen Scavenger*) and 1.25 µL of 0.2 mg/mL biotinylated lipids (Biotin-CAP-PE, Avanti Polar lipids) were added to the previous Golgi membrane mix.

Following its incubation with the kinesin / Streptavidin mix, the chamber was then injected with 5 µL of the Golgi membrane mix directly after motility buffer and biotinylated lipid addition.

After membranes have settled down to the bottom of the chamber and have interacted with microtubule-binding kinesins, they were visualized under the confocal microscope.

*Oxygen Scavenger: 50X stock with 9 mg Catalase (C9322, Sigma) and 18.5 mg Glucose Oxidase (G2133) in 100 µL Imi buffer.

Immunofluorescence on Golgi membranes

Golgi stacks were in reality only enriched in Golgi membranes but also contained other membrane types. In the course of our study, it became important to identify these different membranes. In order to do so, I adapted and modified the immunofluorescence protocol, usually used to identify different compartments of a cell.

First Golgi membranes needed to be immobilized at the bottom of the experimental chamber.

The bottom side of the chamber was coated with Streptavidin by incubation with 0.25 mg/mL Streptavidin solution (1/4 dilution of a 1 mg/mL stock solution in Golgi membrane buffer) for 1 hour in a humid chamber (coverslip down). The chamber was then washed with Golgi membrane buffer (all injections are performed on the same side) to remove the excess unfixed Streptavidin, and incubated with a Golgi membrane -biotinylated lipid mix (5 µL Golgi membrane solution + 1.5 µL Biotin-CAP-PE from Avanti Polar lipids) for 30 minutes.

The chamber was subsequently filled with 2 µM RAB proteins (dilution in Golgi membrane buffer) and washed 30 minutes later three times with Golgi membrane buffer. Fixation was then performed for 15 minutes using 4% paraformaldehyde (PFA, 16% stock diluted in Golgi membrane buffer) and the chamber was again washed two times with Golgi membrane buffer.

The chamber was incubated with Golgi membrane buffer supplemented with 0.2% (w/v) BSA for 15 minutes and finally injected with the primary antibody and, 1 hour later, with the secondary antibody (see Table 3.2 for list of antibodies used). Several washing steps were performed after both incubations with antibodies using Golgi membrane buffer.

Introduction

RAB proteins are small GTPases of the RAS superfamily that are involved in many steps of transport inside the cell. There are over 60 RAB proteins in humans and they all localize to distinct membrane compartments. RAB proteins which oscillate between an active form (GTP-bound) and an inactive form (GDP-bound) can bind to membranes with the help of their prenyl group (geranylgeranyl group), a post-translational lipid modification at their C-terminal extremity. The RAB Escort Protein (REP), known to be involved in RAB prenylation, and the GDP Dissociation Inhibitor (GDI), known to bind to soluble RABs, are known to play key roles in both the delivery and the recycling of RAB proteins to and from membranes (1,2) but cannot account for their specific intracellular localization. Until now, multiple studies have suggested that RAB specific membrane targeting could be mediated by protein factors such as Guanine nucleotide Exchange Factors (GEF), originally known to activate RABs by nucleotide exchange (3) , and GDI Displacement Factors (GDF), thought to influence the release of prenylated RAB proteins from GDI (4,5) . Extensive sequence analysis, domain swapping and mutagenesis studies of different RAB proteins have shown that specific domains are involved in RAB targeting to membranes. Pereira-Leal and coworkers (6) identified five RAB family regions that distinguish RAB proteins from the other members of the RAS superfamily and four subfamily regions that stand to differentiate each RAB subfamily. Different combinations of mutations of these domains led to mislocalization of the RAB proteins suggesting that membrane specificity is also determined by specific RAB sequences (7) . The hypervariable region of RAB35 has also been shown to be determinant for proper membrane targeting (8) . While protein-protein interaction has been widely studied to explain RAB specific membrane targeting, very little is known about the influence of the membrane itself. Diverging from the initial fluid mosaic model (9) , it is now known that membranes are crowded and heterogeneous environments with lipids and proteins diffusing laterally allowing the formation of regions which vary in thickness and composition (10) . Due to specific lipid metabolism and selective transport, cellular membranes have heterogeneous lipid compositions with asymmetrical lipid compositions between the two leaflets (11) . Because of their diversity, lipid membranes exhibit different physicochemical properties such as lipid order, bending rigidity or curvature. Lipid nanodomains have been shown to exist in biological membranes (12) in a so-called 'raft phase' (or Liquid ordered Lo phase) enriched in saturated lipids, and in a non-raft phase (or Liquid disordered Ld phase), enriched in unsaturated lipids. Membrane curvature is also a key feature of intracellular membranes as most cellular organelles display regions of both low and high curvature. For example, the endoplasmic reticulum is formed of a complex network of interconnected flat sheets and highly curved tubules (13,14) , and endosomes display globular (low curvature) and tubular regions (15) . In cellular membranes, specific membrane compositions or membrane curvature can induce the formation of defects in the arrangement of lipids. Indeed, membranes containing conical shaped lipids, such as phosphatidylethanolamine (PE) or diacylglycerols (DAG) will be less prone to have an ordered structure as compared to phosphatidylcholine (PC) membranes (16-18) . External constraints applied for instance by the cytoskeleton, protein coats, or insertion of amphipathic protein domains can force a lipid bilayer to bend (17-19) . In such a conformation the surface area of the lipid headgroup region of one monolayer is expanded whereas the hydrophobic region remains undisturbed. Biologically, lipid packing defects can be beneficial as they were shown to facilitate the folding of some transmembrane proteins and to be essential for binding to membranes of some peripheral proteins (20,21) . The varying amounts of packing defects could thus represent a key feature explaining why some membrane proteins are inserted into specific membrane areas.

In this work, we investigate the role of the physicochemical properties of membranes in the binding of RAB proteins using in vitro assays consisting of purified RAB proteins and giant unilamellar vesicles (GUVs) as model membranes of controlled lipid composition [START_REF] Walde | Giant vesicles: preparations and applications[END_REF] .

Results and discussion

Four RAB proteins, that localize to distinct membranes in cells, were chosen for our study: RAB1 and RAB6 which associate with pre-Golgi and Golgi/trans Golgi network membranes respectively, RAB5 which is present on early endosomes and RAB35 which mainly localizes to the plasma membrane (23) (see SI Text).

RAB6 specifically localizes to the Ld phase independently of its activation and prenylation state.

To test whether RAB proteins show specific recruitment to a given lipid phase, we investigated the recruitment of purified RAB proteins to GUVs exhibiting phase separation between Lo and Ld domains (24) . GUVs were formed using a lipid mixture consisting of brain sphingomyelin (BSM), cholesterol and 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) (3:1:3 molar ratio) (25) . No binding of unprenylated RAB proteins was observed on these membranes (Fig. S1) which is in good agreement with the commonly accepted view that RAB proteins are incorporated into biological membranes through their C-terminal geranylgeranyl groups (26) . Most RAB GTPases are diprenylated in the cell with the addition of two geranylgeranyl moieties on the two C-terminal cysteines (27) . The use of diprenylated proteins (Fig. S2A) is technically challenging due to the high affinity of the GDP-bound RAB for the REP (28) , which prevents binding to GUV membranes. Recruitment of diprenylated RAB6 to membranes exhibiting phase separation was however achieved through the use of additional protein factors (following a protocol detailed in Fig. S3) and clear segregation to the Ld phase was observed (Fig. S3A). Similar experiments were performed with monoprenylated GDP-bound RAB6 (Fig. S2B) and specific binding to Ld domains could also be observed in presence but also in absence of additional protein factors (Fig. S3B and Fig. 1A). To confirm these results, experiments were performed with GUVs composed of pure Ld phase (DOPC and cholesterol in a 1:1 molar ratio) or to GUVs composed of pure Lo phase (BSM and cholesterol in a 1:1 molar ratio) (25) . GDP-bound RAB6, independently of its mono-diprenylation status, was only recruited to Ld GUVs but not to Lo GUVs (Fig. S3 and Fig. 1B). Thus, RAB6 displayed similar membrane binding behavior on GUVs independently of its mono-or diprenylation status. Diprenylated RABs can only be activated by nucleotide exchange in the presence of membrane to avoid precipitation of the proteins; therefore preventing any measurement of the amount of active GTP-bound RABs in our system. For this reason we focused our study on monoprenylated RAB proteins.

The previous experiments were performed using GDP-bound (inactive) RAB6. However RAB proteins localize in the cytosol in their GDP-bound inactivated form and get activated (GTP-bound) by GEFs upon membrane incorporation [START_REF] Stenmark | Rab GTPases as coordinators of vesicle traffic[END_REF] . As membrane bound RAB proteins are mostly active, we investigated the binding of RAB proteins in their GTPbound form. Similarly to its GDP-bound form, monoprenylated GTP-bound RAB6 was only recruited to Ld domains on GUVs displaying phase separation (Fig. 1A) and recruitment was only observed on Ld vesicles but not on Lo vesicles (Fig. 1B); indicating that RAB6 specific binding was independent of its activation state. It should be noted that a quantitative comparison of fluorescence intensities is not feasible due to different prenylation and labeling efficiencies. As membrane-bound RAB proteins are mostly loaded with GTP in the cell and are known to fulfill their functions as such, only activated (GTP-or GppNHpbound) RABs were used in the following experiments.

Some RAB proteins can undergo additional C-terminal modifications following geranylgeranylation, such as proteolysis and/or carboxyl methylation, depending on their prenylation motif (30,31) . RAB carboxyl methylation, which consists in the addition of a carboxyl group to the exposed prenylated cysteine, was shown to enhance the hydrophobicity of the C-terminus and subsequently to increase membrane affinity (31) . However, since the absence of methylation was shown to only affect the cycle of RAB membrane/cytosol partitioning, but not their specific membrane localization (31) , we did not investigate the potential effects of RAB carboxyl methylation in our in vitro experiments.

RAB proteins specifically localize to the Ld phase through their geranylgeranyl group.

Similarly to RAB6, monoprenylated and activated RAB1 and RAB5 segregated specifically to Ld domains on GUVs displaying phase separation (Fig. 1A) and recruitment was only observed on Ld vesicles but not on Lo vesicles (Fig. 1B).

We next investigated whether the prenyl group plays a direct role in the specific recruitment of RAB proteins to the Ld phase. For that purpose, we looked at the recruitment of glutathione S-transferase (GST) to which a CVIL prenylation motif was added at its C-terminus. The purified and fluorescently labeled protein was enzymatically monoprenylated using the same protocol than for the RAB proteins. As shown in Fig. 1, monoprenylated GST also specifically segregates to Ld domains. On the other hand, no recruitment of unprenylated GST could either be detected on Lo or Ld domains (Fig. S1), confirming that the prenyl group is required and sufficient for GST membrane insertion.

Altogether, the above results suggest that the recruitment of RAB proteins to Ld membranes is mediated by the geranylgeranyl moiety. The chemical structure of this C20 isoprenoid chain shows a high degree of unsaturation ("kinks" in the prenyl chain) (Fig. S2C), which might lead to preferential insertion of the geranylgeranyl moiety into Ld membranes. In agreement with this, it was recently shown that the unsaturated C15 isoprenoid farnesyl group, linked to the C-terminus of K-RAS4B spontaneously inserts into loosely packed bilayers consisting of unsaturated lipids (Ld phase) [START_REF] Jang | Mechanisms of membrane binding of small GTPase K-Ras4B farnesylated hypervariable region[END_REF] . Preferential partitioning to flat Ld membranes was also observed for N-RAS proteins (33) . In contrast, the addition of a saturated C16 hydrocarbon chain palmitoyl group to transmembrane proteins mediates their dynamic targeting to raft-like Lo phases (34) . Thus, a likely hypothesis is that the high degree of unsaturation of the prenyl groups favors their insertion into Ld membranes.

RAB35 membrane recruitment is driven by both the charged hypervariable region and the prenyl group.

The great majority of RAB GTPases, including the previously tested RAB1, RAB5 and RAB6, are found associated with intracellular membranes (23) . RAB35 on the other hand, was shown to localize to intracellular endocytic compartments but also to the plasma membrane (35) . We thus wondered whether RAB35 membrane binding was governed by a similar mechanism. We first tested the recruitment of monoprenylated RAB35 to Lo and Ld GUVs. Unexpectedly, RAB35 was not recruited to either of these membranes (Fig. 2A), indicating that the prenyl group is not sufficient to drive RAB35 membrane insertion.

Endosomal and plasma membranes are known to be negatively charged due to the large amount of phosphoinositides and phosphatidyl-serine (11) , anionic lipids known to play major roles in signaling processes and membrane dynamics [START_REF] Paolo | Phosphoinositides in cell regulation and membrane dynamics[END_REF][START_REF] Raghupathy | Transbilayer lipid interactions mediate nanoclustering of lipid-anchored proteins[END_REF] . RAB35 contains stretches of positively charged residues at its C-terminal region, the last 20 amino acid region being the most charged as compared to that of the other RABs (Table S1). In cellulo studies have shown that this polybasic region is essential for targeting RAB35 to the plasma membrane (8) indicating that RAB35 localization depends on electrostatic interactions between the negative charge of the inner leaflet of the plasma membrane and the positive charges of the RAB35 C-terminal region. To address the role of electrostatic interactions, we monitored the recruitment of RAB35 on negatively charged GUVs. The experiments were performed with anionic Lo and Ld GUVs by replacing cholesterol with negatively charged sulfate cholesterol (See SI Text). RAB35 binding was now observed on both Lo and Ld vesicles (Fig. 2A). These results confirm that RAB35 membrane recruitment is mediated by electrostatic interactions and clearly demonstrate that, in contrast to the other RAB proteins that we tested, RAB35 can also bind to Lo domains.

We then investigated whether the prenyl group is required for RAB35 membrane binding to negatively charged vesicles. No binding of unprenylated RAB35 to charged vesicles was observed (Fig. 2A), suggesting that both the electrostatic interactions and the prenyl group are necessary for RAB35 recruitment, but also that prenyl groups are able in some cases to interact with Lo membranes. Additionally, we quantified the area density of prenylated RAB35 (Φv) using Eq. 1 and observed a threefold increase in recruitment to Ld vesicles as compared to Lo vesicles (Fig. 2B). This is consistent with the previous observations that RAB proteins preferentially bind to Ld domains. Taken together, our results suggest that the membrane recruitment of C-terminally charged and prenylated RAB proteins is primarily dependent on the presence of anionic lipids. This specificity for negatively charged membranes gives the ability to RAB35 to overcome the exclusive binding of the prenyl group to Ld domains. This charge dependency is crucial for RAB35 interaction with negatively charged endosomal and plasma membranes. Interestingly, when comparing the charge of the last 20 amino acids of all human RAB proteins, we found that RAB23 and RAB35 display the highest positive charge (Table S1). RAB23 has also been shown to localize to the plasma membrane (38) suggesting that its specific recruitment to the plasma membrane might also be mediated by electrostatic interactions. Furthermore, in good agreement with this, previous studies demonstrated that the recruitment of proteins of the RAS family (K-RAS4B and RND3) to the plasma membrane can be modulated by electrostatic interactions between the positively charged C-terminus and anionic phospholipid headgroups [START_REF] Gomez | Electrical properties of plasma membrane modulate subcellular distribution of K-Ras[END_REF][START_REF] Madigan | Regulation of Rnd3 localization and function by protein kinase C alpha-mediated phosphorylation[END_REF] .

RAB proteins can sense membrane curvature through their prenyl group.

Most RAB proteins, for instance RAB1 or RAB6, are present on transport vesicles which typically have a diameter of 40-60 nm (41,42) and can thus be regarded as curved membranes. We therefore investigated the influence of membrane curvature on RAB membrane recruitment. As a model, we used an optical tweezer setup to pull membrane tubes from EggPC GUVs (with additional 0.1% mol of TexasRed-DHPE lipids) (43,44) (Fig. S4A). RAB protein relative enrichment (or sorting) between the highly curved tube and the flat GUV membrane was imaged by confocal microscopy. Tuning membrane tension through micropipette aspiration of the GUV allows us to modulate the tube radius and to measure protein sorting for increasing curvature (up to 1/15 nm -1 ). Biological membranes are two-dimensional surfaces with two principal curvatures C1 = 1/R1 and C2 = 1/R2 (with R1 and R2 referred to as the principal radii of curvature) along two perpendicular directions (45) . The total curvature of the membrane is C = C1+C2. In the case of a spherical vesicle of radius R, the membrane deforms equally in both directions leading to C1 = C2 = 1/R and a total curvature Cv = 2/R. In the case of a cylindrical tube of radius R, which is curved only in one direction and flat in the other, C1 > 0 and C2 = 0 yielding a total curvature Ct = 1/R (45) . A 15 nm radius tube will thus have the same curvature as a 30 nm radius intracellular transport vesicle (42) , indicating that the typical curvatures in our experiments are biologically relevant.

Curvature sensing was assessed by calculating the sorting ratio (S) defined as the protein/lipid signal ratio on the tube divided by that observed on the GUV (43,44) (Eq. 2 and Fig. S4B). When the tube radius was decreased (i.e. the curvature was increased), a clear enrichment of the proteins was detected in the tube region (Fig. 3A). Sorting was different among the studied RAB proteins, with a ratio increasing up to 5.5, 3 and 2.5 times at a 15 nm tube radius for RAB5, RAB6 and RAB1, respectively.

Because curvature sensing depends on the protein area density (Φv) (44) , sorting values cannot be directly compared among RAB proteins. The protein density is coupled to membrane curvature through a protein curvature coupling coefficient (also called protein spontaneous curvature, Cp) [START_REF] Sorre | Nature of curvature coupling of amphiphysin with membranes depends on its bound density[END_REF] . To quantitatively compare the sorting of RAB proteins with that of other proteins, we used the theoretical model previously developed by Sorre and coworkers (44) and the resulting equation: S = 1 + 1/(RtCpϕv) where S is the sorting ratio, Rt is the tube radius, Cp is the effective spontaneous curvature of the protein and ϕv is the protein area fraction which is related to Φv by Φv = ρ ϕv (ρ is the inverse of the area per protein). The intrinsic curvature radius of the protein Cp -1 can be determined by plotting (S-1) ϕv as a function of curvature (1/Rt) and taking the resulting slope of the linear fit (Fig. 3B). Φv was assessed using Eq. 1 and ρ was estimated by assuming that RAB proteins are spherical proteins of around 25 kDa with a corresponding average radius of 2 nm (ρ = 1/12.6 nm -2 ) [START_REF] Erickson | Size and shape of protein molecules at the nanometer level determined by sedimentation, gel filtration, and electron microscopy[END_REF] . Cp -1 values were respectively 2.1 ± 0.2 nm, 2.6 ± 0.1 nm and 1.5 ± 0.1 nm for RAB1, RAB5 and RAB6. The three RAB proteins display similar spontaneous curvatures, in the same range as what was obtained for Amphiphysin (1.9 ± 0.4 nm) [START_REF] Sorre | Nature of curvature coupling of amphiphysin with membranes depends on its bound density[END_REF] , a known curvature sensor (47) . However, Amphiphysin senses curvature both through its highly curved Bin/Amphiphysin/Rvs (BAR) domain and its Nterminal amphipathic helix (47,48) . Rab proteins interact with membranes through the hydrophobic insertion of their prenyl group into the bilayer while a few amino acid residues close to the prenylation site will be in proximity to the lipid headgroups. Thus, the geometry of the inserted domain may be comparable to that of lipids with inverted conical shapes, such as lysophosphatidic acids (LPAs). LPAs generate local positive curvature and display a spontaneous curvature radius of 2 nm (45) , a value close to those we obtained for RAB proteins.

We next investigated the contribution of the prenyl group to the sensitivity of RAB proteins for curved membranes using monoprenylated GST. As shown in Fig. 3A, a clear enrichment of monoprenylated GST in membrane tubes pulled from GUVs was observed (4 fold increase on tubes of 15 nm radius). The spontaneous curvature radius of monoprenylated GST was measured using the same model as above. We found Cp -1 = 2.8 ± 0.1 nm, which is very similar to the values obtained for RAB proteins (Fig. 3B). This suggests that RAB curvature sensing is independent of the tertiary structure of the protein but, similarly to their preference for Ld membranes, depends on the geranylgeranyl moiety.

Prenylated proteins show preferences for lipid packing defects.

The Ld phase is characterized by the assembly of unsaturated lipids which are known to promote lipid packing defects (49) . Membrane curvature was also shown to lead to the appearance of defects in the arrangement of lipids (49) . To explain the preferential binding of RAB proteins to Ld membranes and their sensitivity to curvature, we hypothesized that RAB membrane recruitment is dependent on the presence of lipid packing defects in the bilayer.

To test this hypothesis, we performed recruitment experiments with GUVs containing 15% mol 1-2-dioleoyl-sn-glycerol (DOG), a conical-shaped lipid that was shown to induce the formation of packing defects similar to those found on positively curved membranes (50) . Control GUVs containing lower amounts of lipid packing defects were composed of 15% mol DOPC cylindrical lipids (See SI Text).

The membrane recruitment of all monoprenylated proteins was significantly increased in the presence of DOG (Fig. 4), i.e. in presence of higher amounts of lipid packing defects.

Unlike DOG, polyunsaturated fatty acids (PUFAs), such as 1-stearoyl-2-docosahexaenoyl-snglycero-3-phosphoethanolamine (PUFA PE), were shown to decrease the amount of lipid packing defects, especially in curved membranes [START_REF] Pinot | Lipid cell biology. Polyunsaturated phospholipids facilitate membrane deformation and fission by endocytic proteins[END_REF] . We measured RAB and GST binding on GUVs composed of 30% mol PUFA PE and used as a control GUVs containing higher amounts of packing defects and composed of 30% mol 1-palmitoyl-2-oleoyl-snglycero-3-phosphoethanolamine (POPE) (See SI Text). We found that the membrane recruitment of geranylgeranylated proteins signify-cantly decreases in the presence of PUFA PE (Fig. 5) i.e. when the amount of packing defects is decreased.

Altogether, the above results suggest that lipid packing defects are drivers of RAB membrane recruitment and that this lipid packing defect sensing is mediated by the C-terminal prenyl group.

Conclusions

It has been known for a long time that prenyl groups act as non-specific membrane anchors but our results, together with recently published data [START_REF] Larsen | Membrane curvature enables N-Ras lipid anchor sorting to liquid-ordered membrane phases[END_REF][START_REF] Larsen | Membrane Curvature and Lipid Composition Synergize To Regulate N-Ras Anchor Recruitment[END_REF] , highlight a role for prenyl groups (farnesyl and geranylgeranyl) in specific membrane domain targeting. Similarly to what we found with geranylgeranylated RAB proteins, farnesylated N-RAS preferentially binds to Ld domains on flat membranes and its differential membrane recruitment was shown to rely on the presence of lipid packing defects induced by curvature and specific lipid geometrical shapes (33,52) . A likely explanation is that prenyl groups are largely unsaturated and have a kinked structure allowing them to get preferentially inserted into membranes containing packing defects such as Ld or curved membranes. This lipid-driven membrane binding mechanism sheds new light on how RAB GTPases could bind to membranes. Intracellular membranes are mainly composed of Ld phases (53) , and many RAB proteins associate with highly curved transport vesicles (41,42) . Our hypothesis is that the addition of one or two geranylgeranyl moieties on all RAB proteins serves as a core mechanism to bind them to specific membrane domains displaying lipid packing defects, the specificity for a given compartment (ER, Golgi, endosomes) relying then on other mechanisms such as the presence of specific GEFs (3) . An interesting variation to this theme is given by RAB35 which has a positively charged C-terminus and is mainly found at steady state associated with the plasma membrane and the endocytic compartments [START_REF] Kouranti | Rab35 regulates an endocytic recycling pathway essential for the terminal steps of cytokinesis[END_REF] . We showed that RAB35 membrane recruitment is primarily dependent on the presence of negatively charged lipids which are also predominantly found on endosomal and plasma membranes (11) . Even though lipid packing defects enhance RAB35 membrane affinity, they are not essential for membrane binding.

In conclusion, our work illustrates that the physicochemical properties of membranes, such as charge distribution and lipid packing defects, could be prime determinants of the localization of RAB proteins to cellular membranes.

Material and Methods

In vitro monoprenylation and diprenylation. The prenylation reaction consists in the addition of one or two C20 geranylgeranyl moieties (Geranylgeranyl pyrophosphate, GGpp, Sigma) at the C-terminal extremity of the proteins. Prenylation was achieved either through monoprenylation (addition of one geranylgeranyl group) using purified Geranylgeranyl Transferase type 1 (GGTase1) (Fig. S2B) or diprenylation (addition of two geranylgeranyl groups) using the native prenylation machinery consisting of purified RAB Geranylgeranyl Transferase (RABGGTase or GGTase2) and RAB Escort Protein (REP) (Fig. S2A). Monoprenylation reactions were performed at 25°C for 1.5 h with a molar ratio of 0.5:1:5 GGTase1, RAB and GGpp. Molar ratios for the diprenylation reaction were: 1:5:0.5:0.75 RABGGTase:GGpp: RAB:REP and the reaction was performed at 25°C for 4 h. To control efficient protein prenylation, NBD-Farnesyl pyrophosphate (Jena Bioscience), a C15 fluorescent analog of GeranylGeranyl pyro-phosphate, was used as described in (54) .

Giant Unilamellar Vesicles. GUVs were grown on indium tin oxide coated (ITO) glass slides using the electroformation technique [START_REF] Angelova | Preparation of giant vesicles by external AC electric fields. Kinetics and applications[END_REF] . 15 µL of a 0.5 mg/mL lipid mix was dried on ITO coated slides for a few minutes at 50°C and subsequently under vacuum for at least 2 h. The dried lipid film was then rehydrated in a sucrose solution (osmolarity between 100 and 430 mOsm, depending on the osmolarity of the protein buffer used for the experiments) and GUVs were grown for 3 h under a sinusoïdal voltage (1.1 V,10 Hz). GUV growth was most of the time performed at room temperature except in the case of phase separation (see SI Text) where it was performed at 50°C.

Membrane tube pulling by optical tweezers.

A highly curved membrane tube was pulled out from an EggPC containing GUV, aspirated in a micropipette to control its membrane tension, using 3.2 µm diameter beads trapped in an optical tweezer as previously described in (43) (Fig. S4A). The membrane tension was increased in a stepwise fashion to decrease the tube radius and hence increase membrane curvature (43) . The reaction buffer used was the one specific of the studied protein and was supplemented with 0.1 mg/mL β-Casein to prevent adhesion of the GUV to the glass. Membrane binding was studied using 100-300 nM final concentration of protein.

Measurement of protein density on the membrane.

Protein density was assessed as previously described (44) . Briefly, fluorescence was calibrated using GUVs made of EggPC lipids and BodipyFL-C5-HPC (HPC), a green fluorescent lipid, at various concentrations. The HPC area density on the GUV (Φv HPC ) can be calculated by assuming that the average area occupied by a single PC molecule is 0.7 nm 2 [START_REF] Nagle | Structure of lipid bilayers[END_REF] . The fluorescent intensity of this lipid on the GUV membrane was measured (Iv HPC at a given confocal photomultiplier tube detector gain) for each area density. A linear fit of the fluorescence vs. area density plot gave the conversion constant (Again) (Φv HPC = Again X Iv HPC ). Proteins were labeled with the Alexa488 fluorophore and lipids with BodipyFL-C5-HPC, two fluorophore exhibiting different spectral properties. Thus, we measured the correction factor F = Ibulk A488 / Ibulk HPC , i.e. the ratio of fluorescence intensities of Alexa488 and HPC respectively at a given concentration in solution. Both fluorescent signals in bulk scaled linearly with their concentration and F is defined as the ratio between the slopes of the Alexa488 linear fit and that of HPC. The protein labeling efficiency was taken into account by calculating the degree of labeling (n * ) of the protein using Eq. S1. Protein density on the GUV membrane (Φv prot ) was thus given by:

Φ 𝑣 𝑝𝑟𝑜𝑡 = 𝐴 𝑔𝑎𝑖𝑛 𝐹 𝑥 𝑛 * × 𝐼 𝑣 𝑝𝑟𝑜𝑡 (𝑔𝑎𝑖𝑛) [1]
Measurement of sorting ratio. In order to quantify protein sorting to the tube, the fluorescence intensity of the Alexa488 labeled protein (Iprotein) was normalized by the intensity of the fluorescent lipid (TexasRed-DHPE, Ilipid) at each tension step increase.

The sorting ratio S corresponds to the ratio between the normalized protein intensity on the tube and the same normalized intensity on the GUV (Fig. S4B):

𝑆 = (𝐼 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 /𝐼 𝑙𝑖𝑝𝑖𝑑 ) 𝑡𝑢𝑏𝑒 (𝐼 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 /𝐼 𝑙𝑖𝑝𝑖𝑑 ) 𝑣𝑒𝑠𝑖𝑐𝑙𝑒 [2]
ACKNOWLEDGMENTS. We thank Roger Goody for providing the plasmids to express the prenylation machinery. Ahmed El Marjou is acknowledged for the purification of the REP protein. We thank Aymelt Itzen for critical reading of the manuscript. We thank Bruno Antonny, Sandrine Etienne-Manneville and Jean-Claude Guillemot for stimulating discussions. We thank Ernesto Ambroggio, Maryam Alqabandi and Feng-Ching Tsai for their help with the fluorescence quantifications. We thank Mathieu Pinot for his technical assistance with the optical tweezer setup. Fig. 3. Prenylated proteins can sense membrane curvature. A highly curved membrane tube was pulled with optical tweezers from an EggPC GUV containing the fluorescent lipid marker TexasRed-DHPE (red) in the presence of 100-300 nM Alexa488 labeled monoprenylated proteins (RAB1, RAB5, RAB6 and GST). (A) The plots show the protein sorting ratios (Eq. 1) as a function of tube curvature (1/Rtube). Data was obtained from 10 (RAB1, GST) or 7 (RAB5, RAB6) independent experiments. Each dot represents one sorting measurement at a given tube radius. Each plot was fitted with a linear regression (black line). For all prenylated proteins, sorting increases when the curvature is increased (i.e. when the tube radius is decreased). (B) Same data as in A plotted as (S-1) ϕv as a function of curvature (1/Rt). Following the theoretical model from Sorre and coworkers (44) , the sorting ratio S is given by S = 1 + 1/(RtCpϕv) where Rt is the tube radius, Cp is the effective spontaneous curvature of the protein and ϕv is the protein area fraction. (S-1) ϕv is thus predicted to scale linearly with the tube curvature (1/Rt) with a slope Cp -1 . Each plot was fitted with a linear regression (black line) to measure the intrinsic curvature radius of the protein Cp -1 (in nm).

Fig. 4. Increasing amounts of lipid packing defects enhance RAB membrane binding. RAB1, RAB5, RAB6 and GST were labeled using Alexa488, monoprenylated and incubated with GUVs at 2 µM final concentration. DOG containing GUVs with a high density of lipid packing defects were formed using an 85% EggPC: 15% DOG (mol/mol) mix. In control GUVs, DOPC replaced DOG. The right panel shows the quantification of protein density on the membrane (Φv) in both DOPC and DOG containing vesicles. We observed a significant increase in protein recruitment on GUVs with higher levels of lipid packing defects. (Scale bar: 10 µm; *** = t-test, p-value ˂ 0.0001; ** = t-test, p-value = 0.0006).

Fig. 5. Decreasing amounts of lipid packing defects reduce RAB membrane binding. RAB1, RAB5, RAB6 and GST were labeled using Alexa488, monoprenylated and incubated with GUVs at 2 µM final concentration. PUFA PE containing GUVs with a low density of lipid packing defects were formed using a 55% EggPC : 15% DOG : 30% PUFA PE (mol/mol) mix. In control GUVs POPE replaced PUFA PE. The right panel represents the quantification of protein density on the membrane (Φv) in both POPE and PUFA PE containing vesicles. We observed a significant decrease in protein recruitment on GUVs with lower levels of lipid packing defects. (Scale bar: 10 µm; *** = t-test, pvalue ˂ 0.0001). )

Ф v (10 3
.μm -2

)

Ф v (10 3 .μm -2
)

Supplementary text

Protein purification. Expression and purification of the prenyltransferases GGTase1 and RABGGTase were performed as described previously (1) . REP-1 was expressed in insect cells using a pFastBac vector and purified as in (2) . RAB5A and codon optimized RAB1B were C-terminally modified, expressed and purified as described in (3) . RAB6A, RAB35 and GST were also C-terminally modified with a CVIL motif (recognized by GGTase1 for monoprenylation) and were respectively expressed as a cleavable His6fusion, His-GFP-fusion and GST-fusion constructs. RAB protein purification was performed at pH7.5 as previously described for RAB1B. GST purification was performed using Protino Glutathione Agarose 4B beads (Macherey Nagel). For RAB6A, residual Histag was removed using a HisTrap HP column (GE Healthcare) and His6-GFP-RAB35 was isolated as a fusion protein (no protease cleavage and no second affinity chromatography was required). Wild-Type RAB6A protein (RAB6A-WT) was purified the same way as the C-terminally modified RAB6A. His6-mCherry-LidAaa201-583 was expressed and purified as previously described (4) .

In vitro nucleotide exchange. Nucleotide exchange was induced chemically as described in (5) . RAB6 was exchanged to GTP and GDP (Sigma Aldrich) and RAB1, RAB5, RAB35 were exchanged to GppNHp (Jena Bioscience), a non-hydrolysable analog of GTP. In order to measure the nucleotide bound state of the protein, an ion-paired reverse phase chromatography was performed as described in (6) .

In vitro N-terminal labeling. The N-terminal labeling of the protein was performed using Alexa Fluor TM 488 Sulfodichlorophenol ester (Alexa488 5-SDP ester, Molecular Probes). To ensure specific N-terminal labeling the reaction was performed at pH 7.5. The protein was incubated with a 4 fold molar excess of Alexa488 for 2 h at 25°C. A 250 fold molar excess of freshly prepared Hydroxylamine was added, for 1h at 25°C, to stop the reaction. A NAP column (GE Healthcare), pre-equilibrated with the protein buffer, was then run to remove excess unbound fluorophore. Protein concentration was determined by a Bradford assay. The efficiency of labeling (n * ) which corresponds to the ratio of moles of dye per mole of protein was determined after absorption measurements using the following formula:

𝑛 * = 𝐴 𝑚𝑎𝑥 𝑜𝑓 𝑡ℎ𝑒 𝑙𝑎𝑏𝑒𝑙𝑒𝑑 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 Ɛ ×𝑝𝑟𝑜𝑡𝑒𝑖𝑛 𝑐𝑜𝑛𝑐𝑒𝑛𝑡𝑟𝑎𝑡𝑖𝑜𝑛 (𝑀) × 𝑑𝑖𝑙𝑢𝑡𝑖𝑜𝑛 𝑓𝑎𝑐𝑡𝑜𝑟 [S1]
Where Amax is the absorbance of the fluorescent protein at the wavelength maximum of the dye (λmax = 488 nm for Alexa488) and Ɛ is the fluorescent dye molar extinction coefficient (71,000 M -1 cm -1 for Alexa488).

Giant Unilamellar Vesicles: preparation and reagents. All lipids were from Avanti Polar Lipids. Phase separated GUVs contained Brain Sphingomyelin (BSM), Cholesterol and 1,2-dioleoyl-snglycero-3-phosphocholine (DOPC) at a molar ratio of 3:1:3 respectively (7) and were electroformed at 50°C

to allow lipid mixing. The other GUVs tested were electroformed at room temperature. Lo vesicles and Ld vesicles were respectively composed of BSM : Cholesterol and DOPC : Cholesterol at 1:1 molar ratios. The charged versions of Lo and Ld vesicles were composed of Cholesterol 3-sulfate instead of Cholesterol. To induce lipid packing defects, a 85% Lα-phospha-tidylcholine (Egg, Chicken) (EggPC) : 15% 1-2-dioleoyl-sn-glycerol (DOG) (mol/mol) mix was used. In control experiments, DOG was replaced by DOPC. In order to decrease the amount of lipid packing defects 30% (mol/mol) of 1-stearoyl-2docosahexaenoyl-sn-glycero-3-phosphoethanolamine (PUFA PE) was added to the previously mentioned DOG-containing mix, at the expense of EggPC. Control experiments were performed using 30% (mol/mol) of 1-palmitoyl-2-oleoyl-sn-glycero-3phosphoethanolamine (POPE) instead of PUFA PE.

Tube pulling experiments were performed using EggPC vesicles. To achieve adhesion between the GUV membrane and the streptavidin-coated beads, 0.035% (mol/mol) of 1,2-distearoyl-snglycero-3-phosphoethanolamine-N-[biotinyl(polyethyleneglycol)-2000] (ammonium salt) (DSPE-PEG(2000)-Biotin) was added to the lipid mix (see also 'Membrane tube pulling by optical tweezers' section).

GUVs were made fluorescent by adding 0.1 (mol/mol) % of the red-emitting dye TexasRed TM 1,2-Dihexadecanoyl-sn-glycero-3-phosphoethanola-mine (triethylammonium salt) (TexasRed-DHPE) from Molecular Probes.

Confocal microscopy.

Confocal images were taken on a Nikon A1R microscope using a x100 objective. Green fluorescent dyes were excited with a 488 nm laser and red fluorescent dyes were excited with a 561 nm laser. Experiments were performed at room temperature (22°C). Images were quantified using the Fiji software (8) .

Measurement of fluorescence intensities. Fluorescent intensities were measured using a rectangular selection including either the horizontal tube or the GUV membrane and subsequent averaging along vertical lines. All intensities were calculated after subtracting the noise level (intensity inside the vesicle) from the maximum of the fluorescence peak. charged whereas the red colored amino acids are negatively charged. RAB23 and RAB35 both display a highly positive C-terminus as compared to the other RAB proteins. On the other hand, RAB4 and RAB11 preferentially localize to curved tubular structures, whereas RAB5 is predominantly found on globular and flatter structures (Sonnichsen et al., 2000). This raises the possibility that the segregation of these RAB proteins into distinct domains could also be mediated by the physicochemical properties of membranes.

Description of the in vitro approach

In order to investigate the RAB4/RAB11 membrane binding requirements, I used an in vitro approach consisting of purified and prenylated RAB proteins, and giant unilamellar vesicles (GUVs) or purified Golgi fractions as model membranes.

Following purification, RAB proteins were exchanged to GTP/GppNHp (non-hydrolysable analog of GTP) or GDP using a chemically-induced reaction. Nucleotide exchange efficiency was then assessed by reverse-phase ion-pair chromatography. Of note, as membrane-bound RAB proteins are mostly loaded with GTP in the cell and are known to fulfill their functions as such, most experiments were performed using activated GTP/GppNHp-bound RABs.

RAB proteins were then fluorescently labeled at their N-terminus using Alexa fluorophores, and prenylated. Most RAB proteins, including RAB4 and RAB11 (but also RAB1, RAB5 and RAB6), are diprenylated (di-cysteine motif) in the cell by the joint action of the RAB escort protein (REP) and the RAB geranylgeranyl transferase (RABGGTase) (Alexandrov et al., 1994;[START_REF] Seabra | Rab geranylgeranyl transferase. A multisubunit enzyme that prenylates GTP-binding proteins terminating in Cys-X-Cys or Cys-Cys[END_REF].

Diprenylation of RAB proteins could be achieved in vitro using purified REP and RABGGTase and geranylgeranyl pyrophosphate as a substrate. The particularity of this reaction is that REP only recognizes the GDP-bound form of RAB proteins [START_REF] Seabra | Nucleotide dependence of Rab geranylgeranylation. Rab escort protein interacts preferentially with GDP-bound Rab[END_REF]. Therefore, the nucleotide exchange to GTP can only be performed once proteins are diprenylated. Activation of diprenylated RAB proteins leads to the dissociation of the REP-RAB complex and to the subsequent exposure of the hydrophobic prenyl groups to the solvent, thereby causing protein precipitation. Protein precipitation can be avoided by activating diprenylated RABs in the presence of membranes, which allows stabilization of the hydrophobic prenyl groups; but consequently prevent any measurement of the amount of active GTP-bound RABs in the system.

In order to overcome this issue, RAB proteins were monoprenylated using purified Geranylgeranyl transferase I (GGTaseI), which can catalyze the prenylation reaction in a nucleotide independent manner. AS GGTaseI recognizes the C-terminal mono-cysteine CAAX motif, RAB proteins were C-terminally modified to a CVIL motif.

More detailed information about this in vitro approach can be found in Chapter 3.

RAB4 and RAB11 recruitment to GUV membranes

RAB4 and RAB11 are not recruited to PC-containing membranes

Following RAB protein nucleotide exchange to GTP/GppNHp, N-terminal labeling with Alexa488 fluorophore, and monoprenylation, RAB recruitment to membranes was monitored using pure EggPC GUVs. Of note, this entire study started with the use of RAB1B, RAB5A and RAB6A (Chapter 4), which were all found to be effectively recruited to these EggPC-containing membranes. 2 µM of RAB4A or RAB11A were incubated with pure EggPC vesicles (fluorescently labeled with 0.1% (mol) TexasRed-DHPE lipids). Surprisingly, no protein fluorescence was observed on the membrane in presence of either of these proteins (Figure 5.1A).

A 10% and 6.5% degree of labeling was quantified for Alexa488-RAB4A:GppNHp and Alexa488-RAB11A:GppNHp, respectively, which is in the same range as what was obtained for the effectively recruited Alexa488-RAB1B:GppNHp (5%). These measurements indicate that the absence of RAB4A/RAB11A membrane fluorescence intensity is not due to poor labeling efficiency, but probably the result of a lack of protein recruitment to the membrane.

To exclude the possibility that RAB proteins are not recruited because of ineffective prenylation, the efficiency of the GGTase1-mediated prenylation reaction was assessed using NBD-Farnesyl pyrophosphate, a C15 fluorescent analog of geranylgeranyl pyrophosphate (Chapter 3). The exposure of a polyacrylamide gel, loaded with RAB1B, RAB4A and RAB11A prenylation reactions, under a ChemiDoc imaging system, demonstrate that all these proteins are prenylated with comparable efficiencies (Figure 5.1B). This observation suggests that the absence of RAB4A / RAB11A membrane recruitment is not due to the absence of the prenyl group but rather due to the physicochemical properties of the membrane. These EggPC-containing GUV membranes might not display the appropriate characteristics necessary for RAB4A and RAB11A recruitment. The proteins on the gel are then revealed using coomassie blue staining. The two upper bands correspond to the two GGTaseI subunits and the lower band corresponds to the RAB proteins.

RAB4 and RAB11 are not recruited to GUVs of various lipid composition

The previous observations suggest that RAB4A and RAB11A membrane binding might require In order to verify that the RAB4A and RAB11A absence of recruitment is not due to vesicle defects, I checked the binding of 2 µM monoprenylated Alexa488-labeled GppNHp-bound RAB5A to the same vesicles. RAB5A was found to be effectively recruited to all membrane types (Figure 5.2), thereby confirming that vesicles were in good shape. This clearly suggests that, in contrast to RAB5A, RAB4A and RAB11A require more demanding membrane parameters for their effective recruitment.

Membrane curvature has no effect on the recruitment of RAB4 and RAB11

RAB4 and RAB11 localize to tubular endosomal structures (Sonnichsen et al., 2000). I thus thought that the absence of RAB4A and RAB11A recruitment to GUVs could reflect a high sensitivity to membrane curvature (GUVs have a 10 to 20 µm diameter and can therefore be considered as flat). As an example, ARFGAP is a known curvature sensor which was described to only bind to curved membranes displaying radii lower than 35nm (Ambroggio et al., 2010). I thus wondered whether RAB4A and RAB11A might only be able to bind curved membranes.

In order to answer this question, I studied the recruitment of RAB4A and RAB11A to curved membrane structures by pulling tubes from an EggPC-containing GUV (labeled with 0.1% TexasRed-DHPE lipids), using the previously described optical tweezer setup (see Chapter 3) (Figure 5.3) (Ambroggio et al., 2010;Sorre et al., 2012).

This method allows the assessment of protein enrichment to the curved tube as compared to the "flat" vesicle. The holding of the vesicle with a micropipette connected to a water tank allows the control and variation of membrane tension, and consequently of tube curvature. As RAB4A and RAB11A were found not to be recruited to any of the tested GUVs, and as additional fluorescence studies using differently composed GUVs would have most certainly been time consuming and tedious, I made the decision to change the membrane model and to investigate the binding of these proteins to purified rat liver Golgi fractions. Electron microscopy analyses of these membrane fractions (purified in the lab by Hugo Bousquet and Lena Oesterlin)

were performed by Ilse Hurbain (Graça Raposo lab, Institut Curie) and showed that the purified Golgi fractions were enriched in Golgi membranes but still contained other membrane types (Figure 5.4). Purified Golgi fractions were identified by the addition of Rhodamine labeled WGA (wheat germ agglutinin, Vector labs) which interacts with N-acetylglucosamine sugars on the membrane.

Logically, I first checked the recruitment of RAB6A, a known Golgi-localized RAB protein [START_REF] Antony | The small GTPbinding protein rab6p is distributed from medial Golgi to the trans-Golgi network as determined by a confocal microscopic approach[END_REF][START_REF] Goud | Small GTP-binding protein associated with Golgi cisternae[END_REF], in order to verify that RAB proteins can effectively be recruited to purified Golgi fractions. 2 µM monoprenylated Alexa488-labeled GTP-bound RAB6A was incubated in the presence of purified Rhodamine-WGA-labeled Golgi fractions (Figure 5.5A).

Clear membrane recruitment of monoprenylated RAB6A was observed. I then investigated the binding of monoprenylated Alexa488-labeled and GppNHp-bound RAB4A and RAB11A (Figure

5.5A).

Similarly to monoprenylated RAB6A, both monoprenylated RAB4A and RAB11A were found to be recruited to these Rhodamine-WGA-labeled membranes (Figure 5.5A).

In contrast, no binding to purified Golgi fractions was observed for unprenylated RAB6A, RAB4A and RAB11A (Figure 5.5B) which is in good agreement with the commonly accepted view that RAB proteins are incorporated into biological membranes through their C-terminal prenyl groups (Pechlivanis and Kuhlmann, 2006). The above observations highlight the fact that the prenyl group is absolutely required for membrane binding and that the previously observed absence of recruitment to GUV membranes was probably due to inadequate membrane composition (membrane lipids and/or proteins) and/or structure. were incubated with purified Golgi fractions (labeled with Rhodamine WGA (Wheat Germ Agglutinin)).

Unprenylated and active RAB proteins were not recruited to these membranes. (Scale bar = 10 µm).

RAB4/RAB11 membrane recruitment does not depend on the presence of effector proteins

The previous observations indicate that the prenyl group is required for RAB4A / RAB11A membrane binding but does not exclude the possibility that other protein factors might be needed. One could imagine that membrane-bound proteins present on purified Golgi membranes, might also interact with prenylated RABs, therefore allowing their recruitment and stabilization on membranes. To address this point, I investigated the binding of inactive GDPbound RAB proteins that cannot interact with effector proteins (Figure 5.6). RAB4A and RAB11A

were both exchanged to GDP, prior to monoprenylation, and the nucleotide exchange efficiency was assessed by reverse phase ion-pair chromatography (see Chapter 3). Both proteins were exchanged to GDP with a 100% efficiency (not shown). 2 µM monoprenylated Alexa488-labeled GDP-bound RAB4A and RAB11A were then incubated with purified Rhodamine-WGA-labeled Golgi fractions (Figure 5.6A). Both monoprenylated inactive RABs were found to be effectively recruited to these membranes. Consistently with results from the last section (5.3.1), no binding of unprenylated GDP-bound RAB4A and RAB11A proteins was observed on purified Golgi fractions (Figure 5.6B).

The effective membrane recruitment of these prenylated GDP-bound proteins suggests that RAB4A and RAB11A membrane binding does not require protein factors and is mostly dependent on prenyl group incorporation into membranes exhibiting specific lipid composition and/or structure. While assessing the recruitment of monoprenylated Alexa488-labeled and GTP/GppNHp-bound RAB6A, RAB4A and RAB11A to purified Golgi fractions (Figure 5.5A), I noticed that it was not homogeneous, meaning that RAB4A and RAB11A seemed to preferentially localize to some membrane structures. This was not very surprising, as these purified membrane fractions consisted of a mix of Golgi membranes and other membrane types. The three RAB proteins tested localize to specific compartments in the cell. RAB6A localizes to medial-Golgi, trans-Golgi and trans-Golgi network membranes [START_REF] Antony | The small GTPbinding protein rab6p is distributed from medial Golgi to the trans-Golgi network as determined by a confocal microscopic approach[END_REF][START_REF] Goud | Small GTP-binding protein associated with Golgi cisternae[END_REF]; while RAB4A localizes to early endosomes [START_REF] Van Der Sluijs | The small GTP-binding protein rab4 is associated with early endosomes[END_REF] and recycling endosomes [START_REF] Trischler | Biochemical analysis of distinct Rab5-and Rab11positive endosomes along the transferrin pathway[END_REF]; and

Monoprenylated

RAB11A is found on recycling endosomes [START_REF] Ullrich | Rab11 regulates recycling through the pericentriolar recycling endosome[END_REF] and trans-Golgi network membranes [START_REF] Urbe | Rab11, a small GTPase associated with both constitutive and regulated secretory pathways in PC12 cells[END_REF].

However, an interesting observation was made when 2 µM monoprenylated Alexa568-RAB4A:GppNHp and 2 µM monoprenylated Alexa488-RAB11A:GppNHp were added together to purified membrane fractions. Both proteins were found to specifically co-localize to the same membrane structures (Figure 5.7A). Similar experiments were performed in presence of 2 µM monoprenylated Alexa488-RAB6A:GTP and Alexa568-labeled GppNHp-bound RAB4A (Figure 5.7B) or RAB11A (Figure 5.7C). Surprisingly, RAB6A was also found to co-localize with both RAB4A and RAB11A. The above results suggest that, when using purified Golgi fractions as model membranes, monoprenylated RAB proteins might be localizing by default to the same membrane structures. This absence of specific localization could be due to the artificial system used. Purified

Golgi fractions are missing soluble proteins, and probably also some membrane-bound proteins, which might be important for RAB specific membrane targeting. Another important particularity of this system consists in the use of mono-geranylgeranylated RAB proteins instead of the digeranylgeranylated forms, which are usually found in cells. This will be further discussed in the last section of this chapter.

Monoprenylated RAB proteins do not localize to Golgi or recycling endosomal structures

The identification of these RAB4A, RAB6A and RAB11A positive membrane structures became important for two reasons: to identify the membrane type on which monoprenylated proteins seem to bind by default and to further characterize the RAB4A and RAB11A membrane binding requirements.

As RAB6A is known to specifically localize to medial-Golgi trans-Golgi and trans-Golgi network membranes in cells [START_REF] Antony | The small GTPbinding protein rab6p is distributed from medial Golgi to the trans-Golgi network as determined by a confocal microscopic approach[END_REF][START_REF] Goud | Small GTP-binding protein associated with Golgi cisternae[END_REF], I started by investigating whether monoprenylated RAB proteins localize by default to Golgi structures. I started by labeling specifically trans-Golgi and trans-Golgi network membranes using ATTO488-labeled αRAB6:GTP antibody (Adipogen). Simultaneously to the addition of αRAB6:GTP, 2 µM monoprenylated Alexa568-labeled GppNHp-bound RAB4A or RAB11A were added. The membrane regions where both proteins were found to preferentially localize were negative for the Golgi marker αRAB6:GTP (Figure 5.8A). This was expected since RAB4A and RAB11A preferentially localize to endocytic compartments. However, because RAB6A was previously found to localize to the same membrane structures as RAB4A and RAB11A, the above results suggest that monoprenylated RAB6A does not localize to Golgi membranes in vitro and is therefore mislocalized.

Follow-up immunofluorescence experiments were performed using the cis-Golgi marker α-GM130

(anti-Golgi matrix protein 130) and monoprenylated Alexa488-labeled GppNHp-bound RAB11A (Figure 5.8B). Monoprenylated RAB11A was not found to co-localize with α-GM130 suggesting that monoprenylated proteins are not recruited to cis-Golgi membranes.

As RAB4A and RAB11A localize to recycling endosomes in cells [START_REF] Trischler | Biochemical analysis of distinct Rab5-and Rab11positive endosomes along the transferrin pathway[END_REF][START_REF] Ullrich | Rab11 regulates recycling through the pericentriolar recycling endosome[END_REF], I then checked whether monoprenylated RAB proteins might localize by default to endosomal structures.

By immunofluorescence, I labeled specifically recycling endosomal structures using an α-RAB11

antibody. Monoprenylated Alexa488-labeled GTP-bound RAB6A and α-RAB11 were added simultaneously (Figure 5.8C). Monoprenylated RAB6A did not seem to co-localize with the α-RAB11 antibody, indicating that monoprenylated RAB6A, and consequently monoprenylated RAB4A and RAB11A do not localize to recycling endosomal membranes. Monoprenylated RAB4A and RAB11A are therefore also mislocalized.

Other immunofluorescence studies were performed using an antibody targeted against the early endosomal marker EEA1 and an antibody recognizing the ER. Unfortunately, poor membrane labeling was achieved when using these antibodies and no conclusive observations could be made.

Discussion

In this chapter I showed that RAB4A and RAB11A were not directly recruited to PC-containing membranes. This led me to assume that the membrane binding requirements of these two proteins were much more specific. However, RAB4A and RAB11A were also not recruited to GUVS exhibiting distinct Lo/Ld phases and pulling of highly curved tubes did also not allow RAB4A/RAB11A membrane binding. The above results suggest that, in contrast to what was observed for RAB1B, RAB5A and RAB6A (Chapter 4), lipid packing defects, promoted by unsaturated lipids mainly found in Ld membranes and by membrane curvature (Bigay and Antonny, 2012) are not sufficient to drive RAB4A and RAB11A membrane recruitment.

Although RAB4A/RAB11A and phosphoinositides were described to work synergistically through the recruitment of many common effectors [START_REF] Campa | Rab11 and phosphoinositides: A synergy of signal transducers in the control of vesicular trafficking[END_REF][START_REF] Jean | Coordination between RAB GTPase and phosphoinositide regulation and functions[END_REF], RAB4A/RAB11A membrane insertion has never been shown, to my knowledge, to be primarily dependent on the presence of phosphoinositides. In addition, both proteins partially localize to recycling endosomes which exhibit only minor levels of PI (3,4,5)P 3 and PI(4,5)P 2 [START_REF] Fields | Phosphatidylinositol 3,4,5-trisphosphate localization in recycling endosomes is necessary for AP-1B-dependent sorting in polarized epithelial cells[END_REF][START_REF] Thapa | Phosphoinositide signaling regulates the exocyst complex and polarized integrin trafficking in directionally migrating cells[END_REF]. Finally, RAB4A and RAB11A display low levels of positive charges at their C-terminal extremity (see Figure S1, Chapter 4) as compared to RAB35, which was shown to only bind to negatively charged membranes (Chapter 4). However, in order not to exclude the possibility of charge dependency, I investigated the recruitment of RAB4A/RAB11A to PI(4,5)P2 and to PI(3)P-containing GUVs. The observed absence of recruitment of these proteins suggest that negatively charged lipids are not sufficient to drive RAB4A/RAB11A membrane binding.

Interestingly, RAB4A and RAB11A were found to be recruited to purified Golgi fractions. This recruitment was shown to be both prenyl group dependent and nucleotide independent, thus confirming the commonly accepted view that the prenyl group is required for membrane binding (Pechlivanis and Kuhlmann, 2006) but also indicating that membrane binding does not require interactions with effector proteins. These findings suggest that RAB4A and RAB11A membrane incorporation is mainly dependent on the physicochemical properties of the membranes themselves and that recruitment to GUV membranes could be achieved if the appropriate membrane requirements are met.

I reasoned that RAB4A/RAB11A membrane recruitment could very well be due to a combination of multiple specific membrane features. As an example, α-Synuclein, a known curvature sensor, was described to only bind to curved and negatively charged membranes [START_REF] Pranke | alpha-Synuclein and ALPS motifs are membrane curvature sensors whose contrasting chemistry mediates selective vesicle binding[END_REF]. These Golgi membranes seemed to gather the necessary requirements for RAB4A/RAB11A binding. However, the direct assessment of the geometry (curvature) of RAB4A/RAB11A positive membranes proved to be very difficult due to the fact that these membranes tend to aggregate. The effect of membrane curvature on RAB recruitment was therefore investigated by using a previously described tube pulling approach (see Chapter 3) [START_REF] Roux | A minimal system allowing tubulation with molecular motors pulling on giant liposomes[END_REF]. In this method, multiple tubes are pulled from Golgi membranes by using the force exerted by kinesin motors walking on microtubules; and relative protein enrichment to the tube can be estimated. These tube pulling experiments were performed in the presence of monoprenylated Alexa568-labeled RAB11A (Figure 5.9A) and Alexa488-labeled RAB6A (Figure 5.9B) but did not lead to any conclusive observations as the background protein fluorescence was very high. When performing such experiments, pulled tubes are visualized under the microscope at a very close distance (2-3 µm) from the coverslip. This high protein intensity background could have thus been the result of non-specific interactions of the proteins with the solid glass surface.

Although the coating of the glass with β-Casein was supposed to prevent such interactions, it proved not to be sufficient and clear background fluorescence could still be observed. In order to overcome this issue, the use of a polymer cushion, such as Polyethyleneglycol (PEG) [START_REF] Wagner | Tethered polymer-supported planar lipid bilayers for reconstitution of integral membrane proteins: silane-polyethyleneglycol-lipid as a cushion and covalent linker[END_REF], should be considered. Polymer cushions were originally used to increase the mobility of integral membrane proteins in supported planar lipid bilayers by preventing the nonspecific interaction of the substrate-exposed protein domain with the hydrophilic substrate.

PEGylated polymer cushions, which can prevent the non-specific binding of proteins to surfaces [START_REF] Du | Grafted poly-(ethylene glycol) on lipid surfaces inhibits protein adsorption and cell adhesion[END_REF] may therefore also be used in the presence of purified Golgi membranes.

The use of synthetic vesicles, that allow a controlled lipid composition, seems however to be more favorable for such a study, as the properties of purified Golgi fractions are difficult to assess.

Multiple combinations of membrane specific features (charge, curvature, etc.) could be achieved using synthetic vesicles and subsequent fluorescence studies by confocal microscopy. However, when visualizing protein binding by confocal microscopy, only one condition can be investigated at a time. Microscope studies of protein binding to GUVs exhibiting various physiochemical properties would therefore be time consuming and tedious.

In order to more rapidly assess the RAB4/RAB11 membrane binding properties, the use of the recently described high-throughput PLIF (protein-lipid interaction by fluorescence) method [START_REF] Ceccato | How membrane chain-melting phase-transition temperature is affected by the lipid chain asymmetry and degree of unsaturation: an effective chain-length model[END_REF] should be considered. This technique was originally developed to investigate the binding of proteins to liposomes containing specific phosphoinositide derivatives. I thought to adapt this method to my study, in order to probe the binding of RAB4A and RAB11A to liposomes exhibiting a wide range of different lipid compositions (Figure 5.10). This experimental approach consists as the following: GFP-tagged RAB proteins are incubated on anti-GFP antibody-coated 96-well plates with specific fluorescent liposome preparations. The level of lipids is then measured with a microplate reader by assessing the fluorescent signal intensity of bound liposomes. This sensitive experimental procedure, which is currently being optimized, would therefore allow the rapid screening of RAB-membrane interactions. Of note, influence of curvature could simultaneously be investigated by probing the binding to liposomes of decreasing sizes. The plate is then washed several times using the liposome buffer, in order to remove unbound liposomes, and incubated for 5 minutes under agitation with 1% triton solution (in PBS) to lyse the liposome solution.

Levels of lipids are finally measured by assessing the fluorescent signal intensity of bound liposomes using a microplate reader. Adapted from Ceccato, 2016.

While performing recruitment experiments using purified Golgi fractions, I observed that both monoprenylated RAB4A and RAB11A, but also monoprenylated RAB6A, localized to the same membrane structures. These membrane structures were found to be negative for cis-Golgi, medial-Golgi, trans-Golgi, trans-Golgi network and recycling endosomal markers. Using purified membrane fractions, monoprenylated RAB4A, RAB11A and RAB6A are therefore all mislocalized to the same membrane structures.

Previous studies have shown that interactions with effectors could regulate the specific membrane targeting of RAB proteins [START_REF] Aivazian | TIP47 is a key effector for Rab9 localization[END_REF]Li et al., 2014). When using purified Golgi fractions, one has to remember that this system does not contain soluble proteins and that some membrane-bound proteins might not be present as well due to the purification procedure. I therefore assumed that the absence of proper localization of RAB proteins in this in vitro system might be due to the absence of specific targeting factors.

The mislocalization of RAB proteins could also implicate the prenyl group. All my experiments were performed using monoprenylated (one geranylgeranyl group) RAB proteins, whereas most RABs, including RAB4, RAB6 and RAB11, were found to be diprenylated (two geranylgeranyl groups) in cells [START_REF] Leung | Thematic review series: lipid posttranslational modifications. geranylgeranylation of Rab GTPases[END_REF]. Studies showed that RAB5A and RAB27A mutants containing only one prenylatable cysteine were mistargeted to the ER and non-functional (Gomes et al., 2003). This could be explained by the fact that decreasing the amount of prenyl groups might result in a loss of specific targeting information or in a decreased affinity for the target membrane.

Studies showed that YPT1 and SEC4 were mislocalized and not functional when mutated from a di-cysteine to a mono-cysteine motif [START_REF] Calero | Dual prenylation is required for Rab protein localization and function[END_REF]. In this same study, the interaction between mono-cysteine motif-containing YPT1 / SEC4 and YIP1, a protein known to only bind prenylated RAB proteins, was shown to be impaired; and loss of functional YIP1 was described to impact YPT1 localization to the Golgi [START_REF] Calero | Dual prenylation is required for Rab protein localization and function[END_REF]. It was thus suggested that mislocalization of monoprenylated RAB proteins might be the result of an absence of interaction with specific targeting factors. However, contradicting results showed that inactivation or depletion of YIP1 does not change YPT1 membrane association and localization (Barrowman et al., 2003;[START_REF] Heidtman | A role for Yip1p in COPII vesicle biogenesis[END_REF], therefore implying that mislocalization of monoprenylated RAB proteins might result from other mechanisms.

Membrane targeting of mono-and diprenylated RABs could also be regulated by the lipid environment, for instance if monogeranylgeranylated RAB proteins exhibit lower affinity or are less stable in the target membranes than diprenylated RABs. In vitro studies using synthetic vesicles have indeed suggested that proteins that possess two geranylgeranyl modifications have a half-life in the order of hours [START_REF] Shahinian | Doubly-lipid-modified protein sequence motifs exhibit long-lived anchorage to lipid bilayer membranes[END_REF] whereas monoprenylated proteins usually exhibit a half-life in the order of a second or less [START_REF] Schroeder | S-Acylation and plasma membrane targeting of the farnesylated carboxyl-terminal peptide of N-ras in mammalian fibroblasts[END_REF]. More recent in vitro studies demonstrated that, at a given membrane composition, membrane dissociation constant values of monofarnesylated N-RAS are generally higher than that of difarnesylated N-RAS proteins [START_REF] Gohlke | Influence of the lipid anchor motif of N-ras on the interaction with lipid membranes: a surface plasmon resonance study[END_REF]. Monoprenylated RAB proteins could therefore dissociate faster from their target membranes and might consequently re-incorporate themselves into the most abundant cellular membrane, the ER.

Nevertheless, additional immunofluorescence experiments should be performed to confirm the identity of the RAB4A/RAB6A/RAB11A positive membrane structures. The use of a good anti-ER antibody now seems necessary in order to assess whether monoprenylated localize by default to ER membranes. Further investigations using the diprenylated versions of these RAB proteins should also be carried out in order to determine whether, when using purified Golgi fractions as model membranes, mono and diprenylated RABs localize differently.

Chapter 6

6 RAB6-induced membrane tethering As already mentioned in the previous chapter, RAB proteins were exchanged to GTP/GppNHp or GDP using a chemically-induced reaction, N-terminally labeled with Alexa fluorophores and monoprenylated using purified geranylgeranyl transferase I (GGTaseI). In contrast to the previous studies, the understanding of this particular tethering mechanism did not require the use of different membrane types but rather the use of different purified proteins. All experiments were thus performed using only EggPC GUVs.

Specificities of RAB6-induced membrane tethering 6.1.1 Vesicle tethering is a RAB6-specific effect

We have observed, by DIC and confocal microscope, vesicle tethering upon the addition of 2 µM monoprenylated Alexa488-RAB6:GTP (RAB6A and also RAB6A') (Figure 6.1A). In contrast, no vesicle tethering was observed in presence of identical concentrations of GppNHp-bound and monoprenylated Alexa488-RAB1B and Alexa488-RAB5A (Figure 6.1B), thus suggesting that vesicle tethering is a RAB6 specific effect.

This observation is partially consistent with previous studies [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF][START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF] in the sense that RAB proteins can mediate liposome aggregation in vitro (these studies are discussed in detail in chapter 2). However, our results are also in conflict with these two studies as VPS21 [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF], the yeast RAB5 ortholog, and RAB5A [START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF] but not RAB6A [START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF] were shown to induce liposome tethering. An important difference between our experimental setups consists in the way of recruiting the proteins to the membranes. In the case of my study and similarly to what occurs in the cell, RAB proteins bound to membranes with their prenyl group whereas in their studies proteins were anchored to membranes using Histidine tagged proteins and Nickel-NTA containing liposomes.

One could hypothesize that His-tagged and prenylated RAB proteins might display structural differences which could account for the differential tethering effects.

Vesicle tethering is nucleotide and concentration dependent

The tethering process seems to preferentially occur when the protein is in its active, GTP bound state but not when GDP-bound (Figure 6.2A). Additionally, a concentration threshold was roughly estimated simply by observing the frontier at which vesicles start to tether. I have identified a concentration threshold of 1.6 µM (Figure 6.2B). This concentration threshold is however not very representative as it might vary depending on the protein membrane density and thus on the protein prenylation efficiency but also on the affinity of the protein for the membrane. The measurement of a protein density threshold would therefore be more relevant and this should be further investigated. In relation to this, one could argue that the absence of vesicle tethering with GDP-bound RAB6A could possibly simply be due to a decrease in membrane recruitment of the protein. The study from Lo and coworkers showed that VPS21, the yeast RAB5 ortholog, is also able to tether vesicles [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF]. This study demonstrates that tethering is GTP dependent and that it can consequently be regulated by GEFs and GAPs. In the case of RAB6, the impact of the nucleotide-bound state could be further investigated in a similar manner with the addition of known RAB6A GEFs or GAPs. Ric1-Rgp1 was described to act as a RAB6 GEF [START_REF] Pusapati | Ric1-Rgp1 complex is a guanine nucleotide exchange factor for the late Golgi Rab6A GTPase and an effector of the medial Golgi Rab33B GTPase[END_REF][START_REF] Siniossoglou | Ric1p and Rgp1p form a complex that catalyses nucleotide exchange on Ypt6p[END_REF] but no attempt was made to test it in our assay. On the other hand, no RAB6 GAPs have been clearly identified so far. Originally, TBC1D11/GAPCenA was proposed to act as a GAP for RAB6 [START_REF] Cuif | Characterization of GAPCenA, a GTPase activating protein for Rab6, part of which associates with the centrosome[END_REF] but this was later questioned by Barr and colleagues when GAP activity was only detected towards RAB4 but not RAB6 [START_REF] Fuchs | Specific Rab GTPaseactivating proteins define the Shiga toxin and epidermal growth factor uptake pathways[END_REF].

A comparison of membrane recruitment of GDP-bound versus GTP-bound RAB6 (and other RABs) could nevertheless also be of interest for future studies. This will be further discussed in the last section of this chapter.

RAB6-induced vesicle tethering is mediated by a RAB-RAB dimerization in trans

The observation of vesicle tethering in confocal microscopy using Alexa488-labeled Rab6A (Figure 6.1A) gave the impression that the protein gets concentrated in the site of vesicle interaction. To further investigate this point, tethering of TexasRed-DHPE-labeled vesicles was induced by the addition of 2 µM monoprenylated Alexa488-RAB6A:GTP; and lipid and protein enrichments at the interaction sites (as compared to the non-interacting regions) were assessed (Figure 6.3A). Quantification of fluorescence intensities of tethered vesicles showed that, whereas TexasRed DHPE lipids concentrate 2 times at the interaction site (because of the contact between two membranes), a 5 fold increase in Alexa488-RAB6A:GTP fluorescence intensity could be detected in the same region (Figure 6.3A). Cryo-electron microscopy analysis that were performed in collaboration with Daniel Levy's team (UMR168, Institut Curie) showed clear protein densities of 5 ± 1 nm thickness at the interaction sites of the tethered vesicles whereas no protein densities could be observed in non-interacting regions (Figure 6.3B). Similarly to what was previously suggested with VPS21 [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF], we suggest a model in which the RAB6 proteins interact with the membrane through their C-terminal geranylgeranyl moiety and one RAB protein from one membrane would interact with one RAB from another membrane (Figure

6.3C).

In this model, based on the analysis of vesicle tethering by Cryo-EM, the average distance between the two opposing membranes is 10-12 nm with protein densities (RAB proteins) having a thickness of 5 ± 1 nm; and spaces between protein densities and vesicular membranes (corresponding to the length of the extended RAB6 C-termini) of 3.5 ± 0.5 nm. The structure of the YPT1/GDI complex (Figure 6.4) [START_REF] Pylypenko | Structure of doubly prenylated Ypt1:GDI complex and the mechanism of GDI-mediated Rab recycling[END_REF] indicates that, when in complex with GDI, YPT1 has a core size of around 4.6 nm and a partially unresolved extended C-terminal tail that seems to be able to bridge distances of at least 4.3 nm; thus fitting nicely with my model. The reason why the C-terminal tail is partially unresolved resides in the fact that many C-terminal residues are flexible. However, the Cryo-EM analysis of vesicle tethering (Figure 6.3B)

demonstrates that the 3.5 ± 0.5 nm space between protein densities and vesicular membranes is quite stable and conserved, thereby suggesting that, in such a configuration, the RAB6 C-terminal tail might no longer be flexible.

The RAB-RAB interaction is dynamic

Protein densities visualized by Cryo-EM seemed to display a solid-like structure, as they were continuous and did not exhibit any apparent gaps. This suggested that strong and stable contacts are established between the dimerizing RAB6 proteins. I thus wondered how stable this interaction was. In order to investigate this point, I induced EggPC vesicle tethering with the addition of 2 µM red labeled RAB6A (monoprenylated Alexa568-RAB6A:GTP) and subsequently to the formation of vesicle aggregates, an extra 2 µM of green labeled RAB6A (monoprenylated Alexa488-RAB6A:GTP) was added. The membrane recruitment of the green labeled RAB6 protein was assessed over time (Figure 6.5A). Interestingly, even though vesicle tethering had already occurred, the second RAB6 protein was rapidly recruited to the membrane and to the interaction sites (Figure 6.5A); and was also found to concentrate there (together with the first RAB6 protein) (Figure 6.5B). This suggests that the RAB6-RAB6 interaction is highly dynamic, therefore allowing a fast mixing of the proteins at the interface between two membranes. Vesicle tethering could still be observed in presence of mutated RAB6A; which suggests that the Y35 and D49 residues might not be involved in RAB6-RAB6 dimerization (Figure 6.7A). An interesting observation was however made during the purification procedure, and more specifically during the gel filtration step. In the case of RAB5 and RAB6 but not of RAB1, clear protein dimerization could be observed when performing size-exclusion chromatography.

However, during RAB6A-Y35R-D49R purification, no dimer complexes were obtained, suggesting that the double mutation might have prevented the formation of this complex in solution. The observation that RAB5 can aggregate in solution but not when bound to membranes and that conversely RAB6-Y35R-D49R induces vesicle tethering but does not aggregate in solution strongly suggests that these two types of oligomerization are not related and that RAB proteins might be able to oligomerize in different ways. This could partially account for the conflicting results with previously published data [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF][START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF].

Unprenylated RAB6A does not interact with membranebound RAB6A

The already mentioned nucleotide specificity of vesicle tethering suggests that this interaction might involve the Switch regions. I therefore reasoned that, if the interaction involves the Switch regions, it would be possible to inhibit membrane tethering by out-competing the RAB-RAB interaction through the recruitment of unprenylated RAB6 or effector proteins. Alexa568-labeled RAB6A was found to be recruited to the vesicular membrane and to concentrate at the interaction sites, Alexa488-labeled RAB6A was not detected in any of these regions. (Scale bar = 10 µm). According to this model, bivalent αRAB6:GTP should be expected to promote vesicle tethering.

This hypothesis was confirmed with the observation that the addition of 40 µM bivalent anti-RAB1:GTP antibody (ATTO488-αRAB1:GTP, courtesy of Adipogen) was able to promote vesicle tethering in presence of 2 µM monoprenylated RAB1B:GppNHp (Figure 6.9D left). Additionally, ATTO488-αRAB1:GTP was also found to concentrate at the interaction sites (Figure 6.9D right)

Interestingly, many RAB effector proteins are dimers that can interact with two RAB proteins [START_REF] Oesterlin | Effectors of Rab GTPases: Rab binding specificity and their role in coordination of Rab functionand localization[END_REF]. A tempting hypothesis would be that these effector dimers could promote, to some extent, tethering between two opposing membranes.

Effect of monovalent RAB6 effector proteins

As bivalent effectors seem to promote vesicle tethering, I then thought to inhibit the tethering by using monovalent effector proteins.

In relation to the previous section, I investigated the impact of the single-chain variable fragment of the αRAB6:GTP antibody (scFv). Addition of a 5 fold molar excess of purified αRAB6:GTP antibody (scFv) prior (Figure 6.10A), but also after RAB6-induced vesicle tethering, did not inhibit vesicle aggregation. When equimolar amounts (2 µM) of Alexa568-RAB6A:GTP and Alexa488-αRAB6:GTP (scFv) were added simultaneously, RAB6A was still found to concentrate around 5 times at the interaction site while αRAB6:GTP (scFv) was concentrated up to 2 times (Figure 6.10B). This suggests that the affinity of αRAB6:GTP (scFv) for RAB6A might be high enough to allow the formation of αRAB6:GTP (scFv)-RAB6A complexes at the interaction site but not sufficient to out-compete the RAB6-RAB6 dimerization. (scFv) at the interaction sites were quantified as in Figure 6.3A. RAB6A:GTP was found to concentrate around 5 times in these regions while only a two fold increase in recruitment was detected for αRAB6:GTP (scFv). (Scale bar = 10 µm).

LidA from Legionella pneumophila is known to have a very high (picomolar) affinity for RAB6A, but also for RAB1B and RAB8A (Schoebel et al., 2011). When equimolar amounts ( 2 This suggests that monovalent effectors with a high affinity for RAB proteins are be able to compete with the dimerization process. The OCRL protein (oculocerebrorenal syndrome of Lowe protein) is a known effector of several RAB GTPases, including RAB6 [START_REF] Hyvola | Membrane targeting and activation of the Lowe syndrome protein OCRL1 by rab GTPases[END_REF] were both found to concentrate at the interaction sites up to 7 and 2 times, respectively. (B) Same experiment as in (A) with Alexa568-labeled RAB6A:GTP and Alexa488-labeled OCRL 538-901 . RAB6A:GTP was found to concentrate up to 4 times at the interaction sites whereas Alexa488-labeled OCRL 538-901 was found to be excluded from these regions. (Scale bar = 10 µm).

These results suggest that Alexa488-OCRL 538-901 exhibits lower affinity towards RAB6A than mCherry-OCRL 538-901 and probably unlabeled OCRL 538-901 . Due to its lower affinity for RAB6A, Alexa488-labeled OCRL 538-901 is not able to compete with the RAB-RAB interaction and to inhibit vesicle tethering. This decreased affinity for RAB6 might be due to the labeling reaction. This reaction consists in the covalent attachment of the Alexa488 fluorophore to the amine groups of the protein. We usually perform it at pH7.5 which should only allow labeling of the N-terminal amine group and not of the amine group-containing lysisnes (lower pKa of the N-terminal amine group). One possibility would be that the RAB-binding domain of OCRL (OCRL 538-901 ) was also labeled on some lysine residues, which led to a decreased affinity for RAB6.

Nevertheless, an excess of mcherry-tagged and untagged OCRL 538-901 was found to moderately inhibit vesicle tethering.

The effect of OCRL 538-901 on RAB6-induced vesicle tethering was found to be less strong than that of LidA 201-583 . As already mentioned, LidA was shown to display a very high affinity (30 pM) towards RAB6 (Schoebel et al., 2011) which is higher than what can be found for usual RAB effectors, including OCRL whose affinity for RAB6 was shown to be in the micromolar range [START_REF] Hou | A structural basis for Lowe syndrome caused by mutations in the Rab-binding domain of OCRL1[END_REF]. LidA is therefore expected to out-compete the RAB6-RAB6 interaction much more effectively than OCRL.

Discussion

I have shown that activated GTP-bound RAB6 (A and A'), but not RAB1B or RAB5A, is able to tether vesicles in vitro without the need of any additional factors (such as tethering factors). We suggest that this tethering effect is mediated by RAB6-RAB6 dimerization in trans. This is partially consistent with previous studies showing that some RAB proteins can induce vesicle aggregation [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF][START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF] and that such an effect is modulated by RAB-RAB interactions in their GTP-bound state [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF]. However, in contrast to my work, these studies indicated that VPS21/RAB5A but not RAB6A are able to induce vesicle tethering. A difference between these studies and mine is the mean by which RAB proteins are anchored to membranes. RAB binding was previously achieved through the interaction of histidine-tagged proteins with Nickel-NTA-containing liposomes [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF][START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF] whereas in my study RAB proteins were incorporated into membranes through their geranylgeranyl group.

The observation that unprenylated RAB6 was not membrane localized in the presence of monoprenylated RAB6 indicate that the prenylated and non-prenylated proteins do not interact with each other. We therefore suggest that monoprenylated RAB6 undergoes conformational changes upon membrane insertion of its prenyl group, which only allows it's interaction with another monoprenylated and membrane-bound RAB6 protein. As the previously mentioned studies used RAB proteins anchored to membranes through a histidine tag, we hypothesize that their proteins might not undergo similar changes in conformation and thereby that the RAB-RAB interactions possibly involve different sequence regions.

One can argue that my study mostly relies on qualitative measurements of vesicle tethering by confocal microscopy. More quantitative analyses of liposome aggregation, using dynamic light scattering [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF] or turbidity assays [START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF], should be performed in order to confirm these observations. RAB proteins are thought to drive membrane tethering through the recruitment of tethering factors (coiled-coil tethers and multisubunit tethering complexes) [START_REF] Brocker | Multisubunit tethering complexes and their role in membrane fusion[END_REF]Cai et al., 2007a). Our results suggest that some RAB proteins, including RAB6, might also operate in concert with these conventional effector proteins to directly drive membrane recognition.

As mentioned in chapter 2, tethering events also occur during the formation of membrane contact sites (MCSs) allowing the transfer of material, such as lipids, between organelles [START_REF] Jackson | Lipids and Their Trafficking: An Integral Part of Cellular Organization[END_REF][START_REF] Phillips | Structure and function of ER membrane contact sites with other organelles[END_REF]. ER-Golgi contact sites have been described to be mostly promoted by a variety of lipid transfer proteins which contain a pleckstrin homology (PH) domain, allowing their binding to Golgi-localized PI(4)P lipids, and an FFAT motif (diphenylalanine in an acidic tract) that can interact with the ER-localized VAP-A protein (vesicle-associated membrane protein-associated protein A). Among them, the oxysterol-binding protein (OSBP), which mediates the transfer of both cholesterol and PI(4)P between the ER and the Golgi, was also shown to bind ARF1 at the Golgi membrane [START_REF] Mesmin | A four-step cycle driven by PI(4)P hydrolysis directs sterol/PI(4)P exchange by the ER-Golgi tether OSBP[END_REF]. Of note, a construct consisting of the PH domain, the coiled-coil region and the FFAT motif of OSBP (PH-FFAT) was sufficient to promote and stabilize ER-Golgi contact sites in HeLa cells [START_REF] Mesmin | A four-step cycle driven by PI(4)P hydrolysis directs sterol/PI(4)P exchange by the ER-Golgi tether OSBP[END_REF].

Prior to my arrival, it was found by Cathy Jackson's team (Institut Jacques Monod) that GBF1 (Golgi-specific brefeldin A-resistance guanine nucleotide exchange factor 1), a known cis-Golgi localized GEF for ARF1 [START_REF] Zhao | Localization of large ADP-ribosylation factor-guanine nucleotide exchange factors to different Golgi compartments: evidence for distinct functions in protein traffic[END_REF], interacts with RAB6. This interaction was first assessed by co-immunoprecipitation/mass spectrometry analyses and subsequently confirmed by

Stephanie Miserey-Lenkei (Bruno Goud's lab, Institut Curie) by yeast two hybrid. Later on, Cathy

Jackson's team also identified VAP-A as a potential GBF1 interacting protein by immunoprecipitation. They subsequently observed, using a proximity ligation assay, that both proteins co-localize at PH-FFAT stabilized contact sites in Hela cells. Altogether, the above results raise the interesting possibility that RAB6 might also localize to ER-Golgi contact sites and might participate in membrane tethering. Further experiments are needed to test this hypothesis.

Part of my work consisted in investigating the effect of RAB6 effector recruitment on RAB6induced vesicle tethering. Because this effect seemed to be dependent on the nucleotide bound state of the protein, I assumed that effector proteins might be able to out-compete the RAB6-RAB6 interaction. Experiments performed in the presence of bivalent αRAB:GTP antibodies strongly suggest that, instead of impairing vesicle aggregation, they can promote vesicle tethering through their interaction with two RAB proteins from distinct membranes. This suggests that effector proteins containing two or more RAB binding sites might be able to promote tethering between two membranes. In relation to this, one could make a distinction between dimeric RAB effector proteins [START_REF] Oesterlin | Effectors of Rab GTPases: Rab binding specificity and their role in coordination of Rab functionand localization[END_REF] that would rather promote homotypic fusion (interaction with two proteins of the same RAB subfamily) and effector proteins that have the ability to interact with different RAB proteins and thereby would rather drive heterotypic fusion. A striking example of such an effector is the Golgi-localized coiled-coil tethering factor GCC185 which associates as a dimer and can therefore bind two RAB proteins [START_REF] Oesterlin | Effectors of Rab GTPases: Rab binding specificity and their role in coordination of Rab functionand localization[END_REF]. Although a direct interaction between GCC185 and RAB6 is controversial [START_REF] Burguete | Rab and Arl GTPase family members cooperate in the localization of the golgin GCC185[END_REF][START_REF] Houghton | The localization of the Golgin GCC185 is independent of Rab6A/A' and Arl1[END_REF], GCC185 also contains multiple other RAB binding-sites which have been suggested to be necessary for the maintenance of Golgi stack morphology [START_REF] Hayes | Multiple Rab GTPase binding sites in GCC185 suggest a model for vesicle tethering at the trans-Golgi[END_REF]. Another tethering factor Rabenosyn-5, which possesses separate RAB4 and RAB5 binding sites, was suggested to connect distinct domains in cis on early endosomes [START_REF] De Renzis | Divalent Rab effectors regulate the subcompartmental organization and sorting of early endosomes[END_REF] and could possibly also mediate to some extent membrane interaction in trans.

In contrast, monomeric effector proteins containing only one RAB6 binding site did not promote vesicle aggregation but, as expected, seemed to rather compete with the RAB-RAB interaction. This competition was more or less efficient, likely depending on their affinities for RAB6. Both αRAB6:GTP (scFv) and OCRL were found to localize at the interaction sites, and OCRL was able to moderately impair vesicle tethering. Their effective recruitment to the interaction sites suggests that both effectors exhibit an affinity towards RAB6 that is high enough to take advantage of the high dynamics of the RAB-RAB interaction but not sufficient to completely impair dimerization. An increased concentration of OCRL might eventually result in a complete inhibition of the vesicle tethering process. However, if the required concentration is too high, impairment of this process might not be experimentally achievable. These results suggest that the RAB6-RAB6 affinity might be higher than that of OCRL-RAB6 which was previously described to be in the low micromolar range (3.7 µM) [START_REF] Hou | A structural basis for Lowe syndrome caused by mutations in the Rab-binding domain of OCRL1[END_REF].

Many human RAB6 effectors were described to have low micromolar to high nanomolar affinities towards RAB6 [START_REF] Oesterlin | Effectors of Rab GTPases: Rab binding specificity and their role in coordination of Rab functionand localization[END_REF], suggesting that they might not always be able to compete with RAB6-RAB6 dimerization. I thus investigated the effect of LidA, a protein translocated by Legionella pneumophila into the host cytosol at the beginning of infection which was shown to interact with RAB1, RAB6 and RAB8 [START_REF] Machner | Targeting of host Rab GTPase function by the intravacuolar pathogen Legionella pneumophila[END_REF] with an extremely high (picomolar) affinity (Schoebel et al., 2011). Interestingly, no vesicle aggregation was observed after simultaneous addition of RAB6 and LidA; and the addition of LidA to previously tethered vesicles seemed to impair the stability of vesicle aggregates. This suggests that LidA, due to its high affinity for RAB6, might out-compete the RAB6-RAB6 interaction, thereby inhibiting homotypic membrane tethering. LidA was previously shown to interfere with the early steps of the secretory pathway [START_REF] Derre | LidA, a translocated substrate of the Legionella pneumophila type IV secretion system, interferes with the early secretory pathway[END_REF]. This was suggested to be mediated by its interaction with GTP but also GDP-bound RAB1 [START_REF] Machner | Targeting of host Rab GTPase function by the intravacuolar pathogen Legionella pneumophila[END_REF]. In addition, LidA was shown to exhibit similar properties towards GTP-and GDP-bound RAB6 (Schoebel et al., 2011), which strongly suggests that it can also interfere with RAB6-mediated trafficking events, including RAB6-induced membrane tethering.

Our study raised the question of the influence of the nucleotide bound state of RAB proteins on their recruitment to membranes. All members of the RAS superfamily of proteins are activated by GEFs upon membrane binding. GDP to GTP exchange was extensively described to be crucial for interaction with effector proteins. In the case of ARF1, GTP binding was also shown to allow the displacement of its N-terminal amphipathic from the core of the protein and its subsequent availability for membrane anchorage [START_REF] Antonny | N-terminal hydrophobic residues of the G-protein ADP-ribosylation factor-1 insert into membrane phospholipids upon GDP to GTP exchange[END_REF]. Additionally, H-RAS was shown to be first targeted to lipid rafts at the plasma membrane and then laterally redistributed to disordered membranes upon activation [START_REF] Prior | GTP-dependent segregation of H-ras from lipid rafts is required for biological activity[END_REF]. This process was suggested to rely on conformational changes of the H-RAS N-terminal domain, coincident with GTP binding, which could be transmitted to the lipid anchor. To my knowledge, no such mechanisms were described for RAB proteins. RABs do not possess N-terminal amphipathic helices and were observed to preferentially localize to disordered membranes independently of their nucleotide-bound state (see Chapter 4). This point should however be further investigated.

another plasma membrane-localized RAB protein (Evans et al., 2003), also displays a C-terminal polybasic hypervariable region, thereby suggesting that its specific recruitment to the plasma membrane might also be mediated by electrostatic interactions. As many other intracellular compartments are enriched in specific phosphoinositide derivatives, known to be involved in various cellular signaling processes, electrostatic interactions might also mediate the recruitment of other RAB proteins.

RAB4 and RAB11 were effectively recruited to purified Golgi fractions in an effectorindependent manner. However membrane charges, but also lipid packing defects, promoted by unsaturated lipids or membrane curvature, were not sufficient to promote RAB4 and RAB11 recruitment to synthetic vesicles. Altogether, this suggests that RAB4 and RAB11 require more demanding membrane physicochemical properties. Possibly, RAB4 and RAB11 effective membrane binding might require a combination of multiple membrane features, such as charge and curvature. This could be rapidly investigated using a "protein-lipid interaction by fluorescence" assay [START_REF] Ceccato | How membrane chain-melting phase-transition temperature is affected by the lipid chain asymmetry and degree of unsaturation: an effective chain-length model[END_REF].

Most studies so far have identified protein factors, such as GEFs, as being responsible for the targeting of RAB proteins to specific intracellular membranes. This study sheds new light on the important role of the physicochemical properties of membranes in mediating the recruitment of RAB proteins. An important question however remains. What is the relative involvement of each of these mechanisms? One could argue that because RAB1, RAB5 and RAB6 can all sense lipid packing defects via their prenyl group, the specificity of localization to a given compartment would then rather rely on protein-mediated mechanisms. However, RAB35 only binds to negatively charged membranes and RAB4/RAB11 membrane binding appears much more specific, suggesting that protein factors and properties of membranes are equally important for the regulation of RAB specific membrane targeting. This matter generates an exciting area for future

investigations. An increased complexity of the in vitro system, through the addition of other membrane-bound proteins such as GEFs or lipid-binding effector proteins, could offer an interesting new way to better understand how RAB proteins are specifically localized.

Abstract

RAB GTPases are major regulators of vesicular trafficking and localize to specific compartments. Deciphering the molecular mechanisms governing RAB localization is thus critical to understand intracellular transport processes. We have managed, for the first time, to incorporate purified and prenylated RABs into artificial membranes. By doing so, we observed that RAB6, but not RAB1 or RAB5, is able to promote by itself vesicle tethering. We believe that RAB6 is able to interact in trans with itself and to consequently drive homotypic membrane tethering. In the main part of this study, we investigated the physicochemical membrane requirements necessary for RAB recruitment. RAB1, RAB5 and RAB6 were all found to only localize to disordered membrane domains and to preferentially bind to curved membranes. We demonstrated that this specific recruitment of RAB1, RAB5 and RAB6 is primarily dependent on the hydrophobic insertion of their prenyl group into lipid packing defects. In contrast, RAB35 recruitment was primarily dependent on the presence of negatively charged lipids and was found to be modulated, to a lesser extent, by lipid packing defects. Although RAB4 and RAB11 were effectively recruited to purified Golgi fractions, in an effector-independent manner, membrane charges and lipid packing defects were not sufficient to promote their recruitment to synthetic vesicles; suggesting that RAB4 and RAB11 require more demanding membrane physicochemical properties. Our work demonstrates that the properties of membranes are critical for the regulation of RAB specific membrane targeting.

Résumé

Les RAB GTPases sont des régulateurs majeurs du trafic vésiculaire et sont localisées sur des compartiments spécifiques. L'identification des processus moléculaires régulant la localisation des RAB est donc cruciale afin de comprendre les mécanismes de transport intracellulaire.

RND3:

  RHO family GTPase 3 SAR: Secretion-associated RAS-related protein scFv: Single-chain variable fragment SDS-PAGE: Sodium dodecyl sulfate polyacrylamide gel electrophoresis SLB: Supported planar lipid bilayer SLiC: Single liposome curvature assay SM: Sphingomyelin SNARE: Soluble N-ethylmalemeide-sensitive factor attachment protein receptor So: Solid-ordered SUV: Small unilamellar vesicle TBC: Tre2/Bub2/Cdc16 TEV: Tobacco etch virus TGF: Transforming growth factor TGN: Trans-Golgi network TIP47: 47-kDa tail-interacting protein Tm: Melting temperature TRAPP: Transport protein particle t-SNARE: Target-SNARE VAP-A: Vesicle-associated membrane protein-associated protein A VPS: Vacuolar protein sorting v-SNARE: Vesicle-SNARE WGA: Wheat germ agglutinin WT: Wild-type YIP: YPT-interacting protein YPT: Yeast proximal tubulin List of figures

Figure 1 . 1 :

 11 Figure 1.1: Lipids can self-assemble into different structures depending on their molecular shapes.. .................................................................................................................................................

Figure 1 . 2 :

 12 Figure 1.2: Membrane lipids ..............................................................................................................

Figure 1 . 3 :

 13 Figure 1.3: Lipid synthesis and distribution in cells .........................................................................

Figure 1 . 4 :

 14 Figure 1.4: Mechanism of lipid domain formation ...........................................................................

Figure 1 . 5 :

 15 Figure 1.5: Membrane shapes and related curvatures ......................................................................

Figure 1 . 6 :

 16 Figure 1.6: Membrane curvature in cells ...........................................................................................

Figure 1 . 7 :

 17 Figure 1.7: Mechanisms of membrane deformation ..........................................................................

Figure 1 . 8 :

 18 Figure 1.8: Deformation induced by changes in lipid composition ................................................

Figure 1 . 9 :

 19 Figure 1.9: Mechanisms of membrane deformation by the cytoskeleton .......................................

Figure 1 . 10 :

 110 Figure 1.10: Mechanisms of curvature sensing ..................................................................................

Figure 1 . 11 :

 111 Figure 1.11: Schematic of the different available methods to produce model membranes .............

Figure 1 . 12 :

 112 Figure 1.12: Phase separation in model membranes.........................................................................

Figure 1 . 13 :

 113 Figure 1.13: In vitro experimental setups for the study of curvature sensing ..................................

Figure 1 . 14 :

 114 Figure 1.14: Description of the gradient force ...................................................................................

Figure 2 . 1 :

 21 Figure 2.1: Alignment of all YPTs (in red) and their human RAB homologues (in black) .............

Figure 2 . 2 :

 22 Figure 2.2: Structural representation of the different members of the RAB family .......................

Figure 2 . 3 :

 23 Figure 2.3: Schematic representation of RAB protein motifs ..........................................................

Figure 2 . 4 :

 24 Figure 2.4: RAB prenylation and activation cycle ............................................................................

Figure 2 . 5 :

 25 Figure 2.5: Mechanisms of protein prenylation ...............................................................................

Figure 2 . 6 :

 26 Figure 2.6: Subcellular localization of RAB proteins .......................................................................

Figure 2 . 7 :

 27 Figure 2.7: RAB proteins are involved in all steps of vesicular transport ........................................

Figure 2 . 8 :

 28 Figure 2.8: Mechanisms of vesicle recognition and tethering .........................................................

Figure 2 . 9 :

 29 Figure 2.9: RAB proteins can directly drive vesicle tethering in vitro .............................................

Figure 2 . 10 :Figure 2 . 11 :

 210211 Figure 2.10: RAB6 regulates multiple trafficking pathways to and from the Golgi .........................77 Figure 2.11: Mechanism of cargo endocytosis mediated by specific RAB domains ........................

Figure 3 . 1 :

 31 Figure 3.1: RAB5A purification on affinity column using the ÄKTA purifier system .....................

Figure 3 . 2 :

 32 Figure 3.2: Measurement of nucleotide exchange efficiency by HPLC ion-paired reverse phase chromatography using the ÄKTA system ..........................................................................................

Figure 3 . 3 :

 33 Figure 3.3: Mechanisms of RAB prenylation ....................................................................................

Figure 3 . 4 :

 34 Figure 3.4: RAB1 and RAB4 prenylation tests ...................................................................................

Figure 3 . 5 :

 35 Figure 3.5: Schematic of GUV electroformation chambers .............................................................

Figure 3 . 6 :

 36 Figure 3.6: Schematic of the microscope experimental setup .........................................................

Figure 3 . 7 :

 37 Figure 3.7: Measurements of fluorescence intensities .....................................................................

Figure 3 . 8 :

 38 Figure 3.8: Green fluorescence calibration .......................................................................................

Figure 3 . 9 :

 39 Figure 3.9: Hand-made PDMS chamber in metal insert ..................................................................

Figure 3 . 10 :

 310 Figure 3.10: Experimental chamber for tube pulling experiments ..................................................

Figure 3 . 11 :

 311 Figure 3.11: Tube pulling experimental setup .................................................................................

Figure 3 . 12 :

 312 Figure 3.12: Determination of the proportionality constant between tube radius and fluorescence ............................................................................................................................................................

Figure 3 . 13 :

 313 Figure 3.13: Experimental chamber for Golgi membranes studies .................................................

Figure 3 . 14 :

 314 Figure 3.14: Tubulation of Golgi membranes mediated by kinesin motors ...................................

Figure 5 . 1 :

 51 Figure 5.1: Monoprenylated RAB4A and RAB11A are not recruited to PC-containing vesicles .....

Figure 5 . 2 :

 52 Figure 5.2: Monoprenylated RAB4A and RAB11A are not recruited to GUV membranes of various lipid composition ...............................................................................................................................

Figure 5 . 3 :

 53 Figure 5.3: Monoprenylated RAB4A and RAB11A are not recruited to curved membranes ...........

Figure 5 . 4 :

 54 Figure 5.4: Electron microscopy analyses of the purified Golgi fractions ......................................

Figure 5 . 5 :

 55 Figure 5.5: Monoprenylated and active RAB6A, RAB4A and RAB11A are recruited to purified Golgi fractions through their prenyl group ................................................................................................

Figure 5 . 6 :

 56 Figure 5.6: Monoprenylated and inactive RAB4A and RAB11A are recruited to purified Golgi fractions through their prenyl group ................................................................................................

Figure 5 . 7 :

 57 Figure 5.7: Monoprenylated RAB4A, RAB11A and RAB6A localize to the same membrane structures ...........................................................................................................................................

Figure 5 . 8 :

 58 Figure 5.8: Monoprenylated RAB4A, RAB11A and RAB6A do not localize to Golgi or recycling endosomal structures ........................................................................................................................

Figure 5 . 9 :

 59 Figure 5.9: Study of RAB4A/RAB11A curvature sensing using purified Golgi fractions and kinesin motors .................................................................................................................................................

Figure 5 . 10 :

 510 Figure 5.10: Description of the PLIF (protein-lipid interaction by fluorescence) assay adapted to my study .............................................................................................................................................

Figure 6 . 1 :

 61 Figure 6.1: Vesicle tethering is a RAB6-specific effect ....................................................................

Figure 6 . 2 :

 62 Figure 6.2: Vesicle tethering is nucleotide and concentration dependent ....................................

Figure 6 . 3 :

 63 Figure 6.3: Vesicle tethering is induced by a RAB6-RAB6 dimerization in trans ..........................

Figure 6 . 4 :

 64 Figure 6.4: Structure of the YPT1:GDI complex ..............................................................................

Figure 6 . 5 :

 65 Figure 6.5: The RAB6-RAB6 interaction is dynamic .......................................................................

Figure 6 . 6 :

 66 Figure 6.6: RAB6 dimer crystal structure ........................................................................................

Figure 6 . 7 :

 67 Figure 6.7: Vesicle tether in presence of RAB6A-Y35R-D49R ........................................................

Figure 6 . 8 :

 68 Figure 6.8: Unprenylated RAB6A does not compete with membrane-bound RAB6A..................

Figure 6 . 9 :

 69 Figure 6.9: Bivalent αRAB:GTP antibodies promote vesicle tethering ..........................................

Figure 6 . 10 :

 610 Figure 6.10: αRAB6:GTP (scFv) can access the interaction sites but does not inhibit vesicle tethering ..............................................................................................................................................

Figure 6 . 11 :

 611 Figure 6.11: Inhibition of vesicle tethering by LidA 201-583 ................................................................

Figure 6 . 12 :

 612 Figure 6.12: Effect of OCRL 538-901 on vesicle tethering ....................................................................

. 1 )

 1 . Most common lipid assemblies in water are micelles (also called hexagonal phase) (Figure 1.1A) and bilayers (lamellar phase) (Figure 1.1B), which are respectively mainly composed of inverted cone-shaped and cylindershaped lipids. Micelles are globular structures where the lipid head groups form a spherical shell protecting the lipid tails (Figure 1.1A). Bilayers are composed of two monolayers in which the lipids are parallel to each other and the tails of each monolayer are facing each other in the core of the membrane (Figure 1.1B). On average bilayers have a thickness of around 5 nm with an area per lipid of around 0.7 nm 2 (this will of course depend on the type of lipid). Lipids with a conical shape have the possibility to self-assemble into an inverted micelle structure (also called inverted hexagonal phase) (Figure 1.1C).

Figure 1 . 1 :

 11 Figure 1.1: Lipids can self-assemble into different structures depending on their molecular shapes. (A) Inverted-conical lipids will tend to self-assemble into micelles, (B) cylinder-shaped lipids into bilayers and (C) cone-shaped lipids into inverted micelles. *Cardiolipins (or Diphosphatidylglycerol) contain 4 acyl chains and consequently display a high conical shape.

Figure 1 . 2 :

 12 Figure 1.2: Membrane lipids. Phospholipids comprise two subclasses of lipids: glycerophospholipids containing a diacylglyerol backbone and sphingolipids which contain a ceramide backbone. These subclasses are further divided into lipid species that display different hedgroups. Here we show the four types of glycerophospholipids (Phosphatidylcholine, Phosphatidylethanolamine, Phosphatidylserine,Phosphatidylinositol), and one type of sphingolipid (Sphingomyelin) used in the study. Glucosylceramide contains a ceramide backbone and is the key precursor for most Glycosphingolipids. Cholesterol is the major sterol found in eukaryotic cells. Of note glycerophospholipids are mostly composed of unsaturated acyl chains whereas sphingolipids mostly exhibit saturated tails. Adapted from(van Meer et al., 2008).

Figure 1 . 3 :

 13 Figure 1.3: Lipid synthesis and distribution in cells. Lipid composition is shown as graphs and expressed as the percentage of total phospholipid content (blue for mammals and light blue for yeast). The figure shows the sites of synthesis of the major phospholipids (in blue) and that of signaling lipids involved in membrane recognition (in red). From (van Meer et al., 2008).

Figure 1 . 4 :

 14 Figure 1.4: Mechanism of lipid domain formation. Membranes can exist as a fluid state or as a solidstate at high and low temperature, respectively. The temperature at which a lipid can transition from one phase to another is the melting temperature (Tm). The Tm is dependent of the specific structure of the lipid (acyl chain length, degree of unsaturation). At physiological conditions, high Tm lipids (long saturated acyl chains) will localize to the solid phase whereas low Tm lipids (short unsaturated acyl chains) will transition to the liquid-disordered state (Ld). Cholesterol can induce the formation of an intermediate liquid-ordered phase(Lo).

Figure 1 . 5 :

 15 Figure 1.5: Membrane shapes and related curvatures. Cellular membranes can be found harboring different shapes. (A) A flat or plane shape, such as basic compartmental membranes, which displays no curvature (C 1 = C 2 = 0). (B) A spherical shape, like intracellular vesicles, which is similarly positively curved in two directions (C 1 = C 2 > 0). (C) A tubular shape, such as tubules emanating from the Golgi, which is only deformed in one direction (C 1 > 0, C 2 = 0). (D) A saddle shape, which can be observed during vesicle

Figure 1 . 6 :

 16 Figure 1.6: Membrane curvature in cells. Almost all cellular membranes display regions of high curvature. Regions of high positive curvature are highlighted in reed. From (McMahon and Gallop, 2005).

Figure 1 . 7 :

 17 Figure 1.7: Mechanisms of membrane deformation. Membranes can be deformed by internal (changes in lipid composition) but also external constraints (protein scaffolding, insertion of hydrophobic domains in one monolayer, cytoskeleton polymerization and movement of motor proteins on cytoskeletal tracks).

Figure 1 . 8 :

 18 Figure 1.8: Deformation induced by changes in lipid composition. Asymmetric bilayers can undergo some deformations. Inverted cone-shaped lipids such as lysophospholipids (in red) tend to induce positive curvature whereas cone-shaped lipids (in green) induce negative curvature.

Figure 1 . 9 :

 19 Figure 1.9: Mechanisms of membrane deformation by the cytoskeleton. (A) Membranes can be pushed by the addition of new G-actin or α-β-tubulin subunits at the + end of actin filaments and microtubules, respectively. (B) Membranes can be pulled via the movement of motor proteins on cytoskeletal tracks. Kinesin and Dynein motors move on microtubules (mostly towards the + and -end, respectively); and Myosins move on actin filaments (mostly towards the barbed (or +) end). (C) Motor proteins such as non-muscle Myosin II can induce the constriction of membranes attached to actin

Figure 1 . 10 :

 110 Figure 1.10: Mechanisms of curvature sensing. (A) Proteins, such as BAR domain proteins, can sense the geometry of curved membranes through their curved BAR domain and electrostatic interactions with negatively charged lipids (red lipid headgroups). (B) Membrane curvature induces the formation of lipid packing defects (expansion of the headgroup region of one monolayer) which allows the preferential insertion of protein hydrophobic domains (amphipathic helices or lipidated groups). The ALPS motif binds

Figure 1 . 11 :

 111 Figure 1.11: Schematic of the different available methods to produce model membranes. MLVs: multilamellar vesicles, SUVs: small unilamellar vesicles, GUVs: giant unilamellar vesicles, LUVs: large unilamellar vesicles, SLBs: supported-planar lipid bilayers. Adapted from[START_REF] Bagatolli | Phase behavior of multicomponent membranes: Experimental and computational techniques[END_REF].

Figure 1 .--

 1 12A shows typical phase diagrams for binary (high and low Tm lipids) and ternary lipid mixtures system (high Tm lipid (A)/Cholesterol (C)/low Tm lipid (B)). Many different phases can be distinguished with ternary lipid mixtures. Pure homogeneous Ld phase when the low Tm lipid (Figure 1.12A (B)) is in excess. -Pure homogeneous Lo phase when the high Tm lipid (Figure 1.12A (A)) is in excess with moderate cholesterol (Figure 1.12A (C)) amount. Coexistence of multiple phases: the three components are in comparable amounts (central area of the phase diagram). Two specific regions can be distinguished: (1) Lo/Ld and (2) So/Lo/Ld phase coexistence.

Figure 1 . 12 :

 112 Figure 1.12: Phase separation in model membranes. (A) Representation of the phase behavior of lipid mixtures using phase diagrams. Phase coexistence can be observed with binary lipid mixtures of different Tm (left diagram) and also with ternary lipid mixtures containing additional cholesterol (right diagram). A: High Tm lipid, B: Low Tm lipid, C: Choleterol. Adapted from[START_REF] Bagatolli | Phase behavior of multicomponent membranes: Experimental and computational techniques[END_REF]. (B) Phase separation on giant vesicles visualized using a rhodamine -1,2-Bis(diphenyl-phosphoino)ethane red fluorescent probe for Ld domains and perylene to mark the Lo phase (blue). From(Baumgart et al., 2003).

Figure 1 . 13 :

 113 Figure 1.13: In vitro experimental setups for the study of curvature sensing. (A) Wavy supported lipid bilayers. Adapted from[START_REF] Hsieh | Curvature sorting of peripheral proteins on solid-supported wavy membranes[END_REF]. (B) Supported membrane tubes. Adapted from[START_REF] Dar | Use of the supported membrane tube assay system for real-time analysis of membrane fission reactions[END_REF].(C) Generation of curved liposomes by extrusion. Extruder image from Avanti polar lipids. (D) Single liposome curvature assay. Adapted from[START_REF] Jensen | Membrane curvature sensing by amphipathic helices: a single liposome study using alphasynuclein and annexin B12[END_REF]. (E) Tube pulled from a GUV with optical tweezers. (F) Tube pulled from a GUV with kinesin motors. Adapted from[START_REF] Van Meer | Membrane curvature sorts lipids. Stabilized lipid rafts in membrane transport[END_REF].

Figure 1 . 14 :

 114 Figure 1.14: Description of the gradient force. A: a micrometric transparent bead is illuminated by a parallel beam of light with an intensity gradient increasing from left to right (represented by a white (low intensity) to black (high intensity) gradient). Two example rays of different intensities are shown in black (black lines of different thickness). These rays are refracted, and thus change direction, when entering and exiting the bead, resulting in a similar directional change of the momentum of the photons. A momentum

Figure 2 . 1 :

 21 Figure 2.1: Alignment of all YPTs (in red) and their human RAB homologues (in black). The

Figure 2 . 2 :

 22 Figure 2.2: Structural representation of the different members of the RAB family. (A) Structural representation of RAB3 with its different functional regions. (B) Comparison of inactivated GDP-bound RAB proteins. The switch regions display a poorly ordered structure. (C) Superposition of GTP-boundRABs. Switch regions are more ordered than with GDP-bound RABs and Switch II displays great conformational differences between RABs. From[START_REF] Lee | Structural mechanisms for regulation of membrane traffic by rab GTPases[END_REF].

Figure 2 . 3 :

 23 Figure 2.3: Schematic representation of RAB protein motifs. RAB family motifs (F1 to F5) differentiate the RAB proteins from the other members of the RAS superfamily while RAB subfamily motifs (SF1 to SF4)are variable among RAB proteins. The C-terminus of most small GTPases, including RAB proteins, is composed of a hypervariable region (suggested to be important for RAB specific membrane targeting) and a mono or di-cysteine motif on which can be added one or two geranylgeranyl groups. The black bars show the location of the phosphate-loop, the switch regions and the hypervariable domain. Adapted from[START_REF] Ali | Targeting of Rab GTPases to cellular membranes[END_REF].

Figure 2 . 4 :

 24 Figure 2.4: RAB prenylation and activation cycle. The RAB escort protein (REP) interacts with the GDPbound RAB protein and directs it towards the RAB geranylgeranyltransferase (RABGGTase) which catalyzes the transfer of one or two (mostly two) prenyl groups (geranylgeranyl moieties) onto the C-terminal monoor di-cysteine motif of the RAB protein. Once they are prenylated, RAB proteins can bind to membranes and get subsequently activated (GDP to GTP exchange) by guanine nucleotide exchange factors (GEF)

Figure 2 . 5 :

 25 Figure 2.5: Mechanisms of protein prenylation. Most members of the RAS superfamily (RAS, RHO and RAB proteins) are prenylated, at their C-terminus, by prenyl transferases. Prenylation consists in the addition of a (A) farnesyl group (substrate is farnesylpyrophosphate) by a farnesyl transferase, specific toRAS proteins, or in the addition of (B) one or two geranylgeranylgroups (substrate is geranylgeranylpyrophosphate) by geranylgeranyl transferases for RHO and RAB proteins. Farnesyl transferase (FTase) and geranylgeranyl transferase I (GGTase I) recognize the CAAX (C is a cysteine, A is an aliphatic amino acid and X is any amino acid) motif at the protein C-terminus. In contrast, the RAB geranylgeranyl transferase (RABGGTase) does not recognize any specific motif. RAB proteins interact with the RAB escort protein (REP) by forming two binding interfaces: the first is between the RAB-binding platform and effectors loops of the RAB protein; and the second is between the C-terminal-binding region (CBR) and the CBR interacting motif (CIM). The REP then recruits the RABGGTase for prenylation and subsequently directs the RAB to the target membrane. GDP-bound RHO and RAB proteins can get extracted from membranes and recycled back to their original compartment by GDP dissociation inhibitors (RHOGDI and RABGDI respectively). Adapted from(Guo et al., 2008).

  RAB proteins have been shown to localize to specific compartments, thereby defining organelle identity[START_REF] Zerial | Rab proteins as membrane organizers[END_REF][START_REF] Zhen | Cellular functions of Rab GTPases at a glance[END_REF] (Figure 2.6); and this is thought to be crucial for the control and directionality of vesicular trafficking. The mechanisms regulating specific RAB membrane targeting and localization are thus of great interest to understand intracellular transport events. The question of RAB protein targeting to specific membranes was initially probed 25 years ago and many have tried to answer this question. Originally, RAB

Figure 2 . 6 :

 26 Figure 2.6: Subcellular localization of RAB proteins. All RAB GTPases localize to specific membranes and are involved in specific transport pathways. Detailed information about the different RAB proteins used in this study (RAB1, RAB4, RAB5, RAB6, RAB11 and RAB35) can be found in the last part of this chapter.

Figure 2 . 7 :

 27 Figure 2.7: RAB proteins are involved in all steps of vesicular transport. (A) During vesicle budding, RAB proteins can activate sorting adaptors to selectively incorporate cargoes into the vesicle. (B) After vesicle fission, coat proteins are partially dissociated from the vesicle. RAB proteins can recruit phosphoinositide (PI) kinases or phosphatases leading to the dissociation of PI binding coat proteins. (C) RAB proteins can mediate vesicle movement on cytoskeletal tracks either by interacting with motor adaptors or directly with motor proteins. (D) Membrane recognition and vesicle tethering can be mediated by RAB proteins through their interaction with tethering factors. (E) These tethering factors can simultaneously recruit SNARE proteins (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) and allow SNARE complex assembly and subsequent vesicle fusion with the acceptor compartment. Adapted from(Stenmark, 2009).

(Figure 2 .

 2 8A and Figure 2.8B)

Figure 2 . 8 :

 28 Figure 2.8: Mechanisms of vesicle recognition and tethering. Tethering is mostly achieved through tether interaction with RAB proteins (A and B) or with coat proteins. (C). (A) RAB proteins can recognize coiled coil tethering factors which exhibit multiple RAB binding sites and are able to bridge long distances.

  well as other endosomal RAB proteins from Saccharomyces cerevisiae (YPT10 and YPT53) were shown to undergo GTP-regulated RAB-RAB interactions that directly drive membrane tethering in vitro, without the need of any effectors (Figure2.9A and Figure2.9B)[START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF]. VPS21mediated vesicle tethering could be observed by using a liposome-based in vitro system (His-tag VPS21 proteins interact with Nickel-NTA-DOGS-containing liposome and subsequent liposome tethering is observed by quasi-elastic light scattering) (Figure2.9A). This tethering effect could only be observed when VPS21 was preloaded with GTP, and was shown to be positively regulated by VPS9 (GEF for VPS21) and inhibited in presence of GYP1 (VPS21 GAP protein). Additionally, by using a bead-liposome assay, vesicle tethering was shown to occur in a symmetric manner, with VPS21 anchored to two opposite membranes and subsequent vesicle tethering mediated by VPS21 dimerization or oligomerization in trans (Figure2.9B). Finally, heterotypic vesicle tethering was shown to occur as well through VPS21 interaction with two other endosomal RABs, YPT10 and YPT53.The VPS21 intrinsic tethering capacity was finally suggested to act in concert with other effector tethers and SNAREs to mediate membrane recognition, tethering and fusion events during endosomal transport.Furthermore, during the course of my thesis, a study has demonstrated that human RAB5A, but also RAB2A (ER/Golgi localized) and RAB7A (late endosome/lysosome localized) also drive vesicle tethering in vitro without the need of any additional components (Figure2.9C and Figure2.9D)[START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF]. Vesicle tethering was observed using three different methods consisting of streptavidin-bead assays (Figure2.9C), turbidity assays and fluorescence microscopy (Figure2.9D). The nucleotide dependency of this effect was not extensively assessed using RABs preloaded with GTP or GDP or using specific GEFs and GAPs and thus no conclusion was made on this point.

Figure 2 . 9 :

 29 Figure 2.9: RAB proteins can directly drive vesicle tethering in vitro. (A) Poly-histidine tagged VPS21 was incorporated into liposomal membranes containing Nickel-NTA-DOGS lipids and liposome tethering was observed by quasi-elastic light scattering in presence of GTP-bound VPS21 only. Adapted from (Lo et al., 2011). (B) Using epifluorescence microscopy, GST tagged VPS21 immobilized on glutathione coated beads were shown to interact with the previously mentioned VPS21 containing liposomes (made fluorescent with TexasRed DPPE) thereby suggesting that vesicle tethering is mediated by VPS21 dimerization/

Figure 2 . 10 :

 210 Figure 2.10: RAB6 regulates multiple trafficking pathways to and from the Golgi. RAB6A and RAB6A'are involved in retrograde trafficking from the endosomal compartments to the Golgi and from the Golgi to the ER. RAB6 is also implicated in intra-Golgi transport and Golgi to plasma membrane transport. RAB6 mediates fission of transport carriers through its direct interaction with Myosin II. RAB6 regulates vesicle motility in cooperation with Bicaudal D2 and the dynamin/dynactin complex. Finally, RAB6 also plays a role in vesicle docking and fusion with the acceptor compartment through its interaction with tethering factors (GARP, COG, Golgins) and direct or indirect recruitment of SNARE proteins. Adapted from[START_REF] Valente | Rab6 and myosin II at the cutting edge of membrane fission[END_REF].

Figure 2 . 11 :

 211 Figure 2.11: Mechanism of cargo endocytosis mediated by specific RAB domains. RAB5 mediates the transfer of cargoes from the plasma membrane to early/sorting endosomes (EEs). EEs display distinct RAB4 and RAB5 domains involved in endocytic recycling and endosome fusion, respectively. RAB4 and RAB11segregate into specific domains on recycling endosomes and are respectively implicated in vesicular transport from EEs and towards the plasma membrane. Late endosomes, also called multivesicular bodies, are composed of two distinct RAB7 and RAB9 domains which regulate cargo degradation via lysosomes and transport towards the trans-Golgi network, respectively. Adapted from(Stenmark, 2009).

3. 3 . 1 1

 311 Purification of Rat Liver Golgi stacks ............................................................................ 104 3.3.2 Experimental chamber ................................................................................................... 104 3.3.3 Pulling tubes with kinesins ............................................................................................ 105 3.3.4 Immunofluorescence on Golgi membranes .................................................................. 107 Protein expression and purificationThis study uses many different proteins. Many DNA constructs were given to us by Roger Goody's lab (Max Planck Institute in Dortmund) and some of the proteins were expressed and purified by them (Table3.1). Others were constructed using standard cloning techniques (amplification, digestion, and ligation). REP-1 was expressed and purified by Ahmed El Marjou (Recombinant Protein Facility, Institut Curie). All monoprenylated proteins were C-terminally modified to contain the CVIL amino acid sequence, a CAAX box recognized by the Geranylgeranyl transferase I. Modifications of the C-terminal recognition site were performed using a simple cloning strategy consisting of oligonucleotides containing the 12 bases coding for the CVIL motif. All constructs were amplified using OmniMAX competent cells (Invitrogen) and the NucleoSpin Plasmid QuickPure kit (Macherey-Nagel); and sequenced by the Institut Curie sequencing platform.

(

  TEV) protease cleavage site which allows digestion and separation of the MBP tag from the rest of the protein. RAB5A was incubated with TEV protease (1 mg protease for 50 mg protein) and dialyzed overnight at 4°C against 5 L of 50 mM Hepes pH 7.5, 200 mM NaCl, 2 mM β-Mercaptoethanol, 1 mM MgCl 2 , 10 µM GDP in 4 spectra/Por Dialysis membrane (MWCO 12.-14.000, Vol/length: 6.4 mL/cm, Spectrum Laboratories).

Figure 3 . 1 :

 31 Figure 3.1: RAB5A purification on affinity column using the ÄKTA purifier system. Protein absorbance at 280 nm (blue) and 254 nm (red) at every washing step. The green line represents the percentage of buffer B. His-tag proteins interact with the nickel column and were only eluted with increasing concentrations of Imidazole (when fractionation occurs).

  using the ÄKTA purification system. The stationary phase of a Protonsil 120-3-C18 column (Bischoff chromatography) is composed of C18 modified silica. The column is equilibrated with the mobile phase containing 50 mM KPi pH 6.7, 10 mM Tetrabutylammonium bromide, 15-25% (v/v) acetonitrile. 100 µL of the 50-100 µM GDP/GTP/GppNHp-bound protein is injected on the column and retention volumes are compared to previously injected GTP/GDP/GppNHp standards (Figure 3.2A). Absorption of the nucleotides is measured at 254 nm. If the protein was exchanged to GTP, the nucleotide exchange efficiency can be measured by calculating the ratio of the GTP integrated peak area over the sum of all nucleotide integrated peak areas (Figure 3.2B).

Figure 3 . 2 :

 32 Figure 3.2: Measurement of nucleotide exchange efficiency by HPLC ion-pair reverse phase chromatography using the ÄKTA system. (A) Absorbance and retention volumes of pure 50 µM GTP and GDP standards at 280 nm (blue) and 254 nm (red). (B) Absorbance and retention volumes of the newly exchanged RAB6 (exchanged to GTP) at 280 nm (blue) and 254 (nm) red. The efficiency of nucleotide exchange is measured at 254 nm, by calculating the GTP peak area over the sum of both nucleotide peak areas.

(

  Geranylgeranyl pyrophosphate, GGpp, Sigma) to the C-terminal extremity of the proteins (Figure 3.3A). Prenylation was achieved either through monoprenylation (addition of one geranylgeranyl group) using purified geranylgeranyl transferase type I (GGTaseI) (Figure 3.3B) or diprenylation (addition of two geranylgeranyl groups) using the native prenylation machinery consisting of purified RAB geranylgeranyl transferase (RABGGTase or GGTaseII) and RAB escort protein (REP-1) (Figure 3.3C). Of note, because GGTaseI only recognizes the C-terminal CAAX motif, monoprenylated proteins were mutated to a CVIL motif at their C-terminus.

Figure 3 . 3 :

 33 Figure 3.3: Mechanisms of RAB prenylation. (A) Native diprenylation machinery consisting of a GDPbound RAB protein with a double cysteine motif at its C-terminal extremity, the RAB geranylgeranyl transferase (RABGGTase, with two subunits α and β) and the RAB escort protein (REP). The RABGGTase transfers two geranylgeranyl groups on the double cysteine motif of the RAB protein, with the REP acting as a chaperone. (B) Monoprenylation of a RAB protein mediated by the Geranylgeranyl Transferase type 1 (two subunits α and β). The GGTase1 recognizes the CVIL motif at the C-terminus of the RAB protein (CAAX box) and transfers one geranylgeranyl group on the cysteine.

Figure 3

 3 Figure 3.4: RAB1 and RAB4 prenylation tests for (A) 1 h (B) 1 h 45 minutes and (C) 2 h 30 minutes incubation times. The right panel shows the NDB-Fpp fluorescence (detected using a ChemiDoc imaging system) and the left panel shows the coomassie blue staining.

3 (

 3 All lipids were from Avanti Polar Lipids. Phase separated GUVs contained Brain Sphingomyelin (BSM), Cholesterol and 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) at a molar ratio of 3:1:Roux et al., 2005) and were electroformed at 50°C to allow lipid mixing. The other GUVs tested were electroformed at room temperature. Lo vesicles and Ld vesicles were respectively composed of BSM : Cholesterol and DOPC : Cholesterol at 1:1 molar ratios(Roux et al., 2005). The charged versions of Lo and Ld vesicles were composed of Cholesterol 3-sulfate instead of Cholesterol. To induce lipid packing defects, a 85% L-α-phosphatidylcholine (Egg, Chicken) (EggPC) : 15% 1-2dioleoyl-sn-glycerol (DOG) (mol) mix was used. In control experiments, DOG was replaced by DOPC. In order to decrease the amount of lipid packing defects 30% (mol) of 1-stearoyl-2docosahexaenoyl-sn-glycero-3-phosphoethanolamine (PUFA PE) was added to the previously mentioned DOG-containing mix, at the expense of EggPC. Control experiments were performed using 30% (mol) of 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE) instead of PUFA PE. Tube pulling experiments were performed using EggPC vesicles. To achieve adhesion between the GUV membrane and the streptavidin-coated beads, 0.035% (mol) of 1,2-distearoylsnglycero-3-phosphoethanolamine-N-[biotinyl(polyethyleneglycol)-2000] PEG(2000)-Biotin) was added to the lipid mix. GUVs were made fluorescent by adding 0.1% (mol) of the red-emitting dye TexasRed TM 1,2-Dihexadecanoyl-sn-glycero-3-phosphoethanolamine (triethylammonium salt) (TexasRed-DHPE) from Molecular Probes. The green-emitting dye β-BODIPY™ FL C 5 -HPC (2-(4,4-Difluoro-5,7-Dimethyl-4-Bora-3a,4a-Diaza-s-Indacene-3-Pentanoyl)-1-Hexadecanoylsn-Glycero-3-Phosphocholine) from Molecular Probes was also used; mostly for calibration measurements (see 3.2.2.3).

Figure 3 . 5 :

 35 Figure 3.5: Schematic of GUV electroformation chambers. (A) Original electroformation using ITO coated slides. Two slides with dried lipid films are assembled a bit shifted from each other while sealing wax (Vitrex) or PDMS is used to isolate the chamber. 4 layers of Teflon tape on each side act as spacers and the chamber is immobilized using paper clips. (B) Electroformation under physiological conditions (high levels of salt). The chamber is made out of a block of Teflon with three identical wells that are sealed, at the top and the bottom of the chamber, by two glass coverslips. Lipids are applied on two platinum wires that cross the whole chamber and are connected to the generator on both extremities.

. 6 )

 6 which allowed us to perform force measurements (calibration for tube pulling experiments) and helped us to easily find model membranes in the different experimental chambers (avoiding the use of the confocal module and fluorescence).

Figure 3 . 6 :

 36 Figure 3.6: Schematic of the microscope experimental setup. The setup developed in the lab by D.Guet is composed of a confocal head which uses visible light (rainbow beam with simple arrow, port n°3), an optical tweezer which functions at 1063 nm (red beam, port n°1), a bright field visualization module which uses near infrared light (brown beam, port n°2) and finally epifluorescence mode (rainbow beam with triple arrows, port n°2). From[START_REF] Guet | Plasma membrane phosphatidylinositol 4-phosphate and 4,5-bisphosphate determine the distribution and function of K-Ras4B but not H-Ras proteins. The Journal of biological chemistry[END_REF].

  chapter 4 and chapter 5, these were measured with the Fiji software using a rectangular selection (10 pixel width) including either the horizontal tube (for tube pulling experiments) or the GUV membrane and subsequent averaging along vertical lines (Figure 3.7A). All intensities were calculated after subtracting the noise level (intensity inside the vesicle) from the maximum of the fluorescence peak.

Figure 3 . 7 :

 37 Figure 3.7: Measurements of fluorescence intensities. (A) The left images represent typical images obtained with tube pulling experiments. Each vertical line of the rectangular selection gives a plot of fluorescence intensity as a function of the pixel number along the line. The right panel of the figure shows the average plot of the rectangular selection. The maximum of the fluorescence peak is subtracted with the noise level (intensity inside the vesicle). (B) An oval selection is drawn around the vesicle and the oval profile plugin measures the pixel of highest intensity for each degree (360 in total). The plot on the right shows the fluorescence intensity values that were subtracted by the background (outside the vesicle) intensity value. (Scale bar: 10 µm).

2 .

 2 A linear fit of bodipyFL fluorescence vs. bodipyFL area density plot gave the conversion constant (A gain ) (Figure 3.8A). Proteins were labeled with the Alexa488 fluorophore and lipids with BodipyFL-C5-HPC, two fluorophores exhibiting different spectral properties. The calibration coefficient A gain could thus not directly be used for the calculation of protein densities. To overcome this issue, the coefficient was corrected with the correction factor F = I bulk A488 / I bulk HPC , i.e. the ratio of fluorescence intensities of Alexa488 and bodipyFL respectively at a given concentration in solution. Both fluorescent signals in bulk scaled linearly with their concentration and F is defined as the ratio between the slopes of the Alexa488 linear fit and that of HPC (Figure 3.8B).

Figure 3 . 8 :

 38 Figure 3.8: Green fluorescence calibration. (A) Determination of the proportionality constant A gain by plotting the green lipid (here BodipyFL) surface density as a function of its fluorescence intensity (at a given PMT gain) and taking the slope of the resulting linear fit. Each point represents the average BodipyFL membrane fluorescent intensity of N = 40 GUVs at a given surface density. (B) Correction factor F determination. Fluorescent intensities of both green dyes are measured in solution at given concentrations.

Figure 3 . 9 :

 39 Figure 3.9: Hand-made PDMS chamber in metal insert. Studies of protein recruitment to GUVs of different compositions were always performed with this device.

(

  water, ethanol, water) and assembled on the microscope insert (Figure 3.10). More specifically, the larger glass part was fixed on the bottom side of two protruding StarFrost glass slides (themselves already fixed to the microscope insert) (Knittel Glass), using vacuum grease (Dow Corning, USA), while the smaller glass part is fixed on the upper part of the StarFrost glass slides.

Figure 3 . 10 :

 310 Figure 3.10: Experimental chamber for tube pulling experiments. This hand-made chamber has the particularity to be opened on the side allowing the insertion of the micropipette.

Figure 3 . 11 :

 311 Figure 3.11: Tube pulling experimental setup. A biotinylated lipid-containing GUV is aspirated into a micropipette connected to a water tank of adjustable height which allows the control of membrane tension.

Figure 3 . 12 :

 312 Figure 3.12: Determination of the proportionality constant between tube radius and fluorescence.

Figure 3 . 13 :

 313 Figure 3.13: Experimental chamber for Golgi membranes studies. A glass coverslip is fixed to aStarfrost glass slide with melted parafilm in between. The parafilm acts as a spacer and also allows isolation of the chamber. This hand-made chamber has the advantage of working with small volumes (5 µL).

  GUVs) using purified kinesin motors (Figure3.14). Briefly, biotinylated kinesin molecules are attached to biotin containing Golgi membranes through streptavidin (previously, small streptavidin coated beads were used) and tube formation occurs in presence of microtubules and ATP.

Figure 3 . 14 :

 314 Figure 3.14: Tubulation of Golgi membranes mediated by kinesin motors. The chamber is coated with microtubules. Biotinylated kinesins and biotinylated Golgi membranes interact in presence of Streptavidin and ATP addition allows kinesins to move along microtubules and the subsequent pulling of membrane tubes.

  Paris Sciences et Lettres Research University, CNRS, UMR144, F-75005 Paris, France b Sorbonne Universités, Université Pierre et Marie Curie, University Paris 06, CNRS, UMR 144, F-75005 Paris, France

Fig. 1 .

 1 Figure legends:Fig.1. Monoprenylated proteins specifically bind to liquid disordered domains independently of their activation/inactivation status. Prenylated proteins (RAB1, RAB5, RAB6 and GST) were labeled with Alexa488 fluorophore and monoprenylated. All images show GTP-or GppNHp-bound RAB proteins except for RAB6 which was also GDP-bound. GUVs were incubated with 2 µM protein. (A) GUV phase separation was achieved using a 3:1:3 (molar ratios) BSM:Cholesterol:DOPC lipid mixture. The Ld phase is marked with 0.1% (mol/mol) TexasRed-DHPE lipids whereas the Lo phase composed of saturated lipids is unlabeled. All tested proteins localize specifically to the Ld phase as shown by the merge images. (B) Lo and Ld GUVs were respectively composed of 1:1 (molar ratios) BSM:Cholesterol and DOPC:Cholesterol. Prenylated proteins were recruited to Ld vesicles but not to Lo vesicles. (Scale bar: 10 μm).

Fig. 2 .

 2 Fig. 2. Negative charges allow binding of RAB35 to both Ld and Lo membranes with a significant preference for Ld domains. GUVs were incubated with 2 µM GFP-tagged RAB35. Neutral Lo and Ld vesicles were formed using 1:1 (molar ratios) of BSM:Cholesterol and DOPC:Cholesterol respectively. Negatively charged Lo and Ld vesicles were formed by replacing Cholesterol with Cholesterol Sulfate. (A) Monoprenylated RAB35 was only recruited to charged Lo and Ld vesicles but not to neutral Ld vesicles. A clear preference for the Ld negatively charged membrane can be observed. Unprenylated RAB35 is not recruited to charged vesicles. (B) Quantification of GFP-RAB35 protein density (Φv) on charged Lo and Ld membranes. We observed a threefold increase in RAB35 recruitment on disordered membranes. (Scale bar: 10 μm; *** = t-test, p-value ˂ 0.0001).

  Figure 1

  Figure S1

Figure 5 . 1 :

 51 Figure 5.1: Monoprenylated RAB4A and RAB11A are not recruited to PC-containing vesicles. (A) 2 µM monoprenylated Alexa488-labeled and GppNHp-bound RAB4A or RAB11A were incubated with EggPCcontaining GUVs (labeled with 0.1% TexasRed-DHPE lipids). Proteins were not recruited to these membranes. (Scale bar = 10 µm). (B) Prenylation efficiencies were assessed by performing monoprenylation reactions in the presence of NBD-Farnesyl pyrophosphate substrate (fluorescent analog of geranylgeranyl pyrophosphate). Monoprenylation reactions were performed on 5 µg GppNHp-bound RAB4A, RAB11A and RAB1B (control) using GGTaseI and the NBD-Farnesyl pyrophosphate substrate. After 1h30 minutes, the prenylation reactions were loaded on an acrylamide gel and analyzed under a ChemiDoc imaging system. The observed fluorescent bands (right panel) correspond to proteins that underwent effective prenylation.

  additional membrane factors (different lipid composition and/or structure) than RAB1B, RAB5A and RAB6A. I thus tested RAB4A and RAB11A recruitment to GUVs made of different lipid compositions. I investigated the binding of 2 µM monoprenylated, Alexa488-labeled, GppNHpbound RAB4A and RAB11A to negatively charged GUVs, endosomal-like GUVs with PI(3)P, brain total lipid extract-containing GUVs supplemented with PI(4,5)P 2 , and vesicles exhibiting Lo and Ld phases. RAB4A and RAB11A were not recruited to any of these membranes (Figure5.2).

Figure 5 . 2 :

 52 Figure 5.2: Monoprenylated RAB4A and RAB11A are not recruited to GUV membranes of various lipid composition. 2 µM monoprenylated Alexa488-labeled and GppNHp-bound RAB4A, RAB11A or RAB5A (control) were incubated with charged GUVs, endosomal-like + PI(3)P GUVs, brain total lipid extract PI(4,5)P 2 -containing GUVs and GUVs exhibiting Lo/Ld domains. In contrast to RAB5A, RAB4A and RAB11A were not recruited to these GUVs of different lipid composition. Charged lipid mix: 10% (mol) brain

  2 µM monoprenylated Alexa488-labeled GppNHp-bound RAB4A or RAB11A were incubated with TexasRed-DHPE-labeled EggPC GUVS. Membrane tubes were pulled and tube curvature was increased until rupture. Regardless of tube curvature, RAB4A and RAB11A were not recruited (Figure 5.3).

Figure 5 . 3 :

 53 Figure 5.3: Monoprenylated RAB4A and RAB11A are not recruited to curved membranes. 2 µM monoprenylated Alexa488-labeled and GppNHp-bound RAB4A or RAB11A were incubated with EggPCcontaining GUVs (labeled with 0.1% TexasRed-DHPE lipids and containing an extra 0.035% DSPE-PEG(2000)-Biotin) and 3.2 µm streptavidin-coated beads. Highly curved tubes were pulled from micropipette-aspirated GUVs using an optical tweezer setup. Protein enrichment to the tube was assessed for increasing tube curvature values. The displayed images show protein (green) and lipid (red) signals at 20 nm tube radius. RAB4A and RAB11A were not recruited to highly curved tubes. (Scale bar = 10 µm).

Figure 5 . 4 :

 54 Figure 5.4: Electron microscopy analyses of the purified Golgi fractions. Red arrows indicate the Golgi stacks. From Ilse Hurbain (Graça Raposo's team, Institut Curie).

Figure 5 . 5 :

 55 Figure 5.5: Monoprenylated and active RAB6A, RAB4A and RAB11A are recruited to purified Golgi fractions through their prenyl group. (A) 2 µM monoprenylated Alexa488-labeled and GTP/GppNHpbound RAB6A, RAB4A or RAB11A were incubated with purified Golgi fractions (labeled with Rhodamine WGA (Wheat Germ Agglutinin)). All monoprenylated and active RAB proteins were recruited to these membranes. (B) 2 µM unprenylated Alexa488-labeled and GTP/GppNHp-bound RAB6A, RAB4A or RAB11A

Figure 5 . 6 :

 56 Figure 5.6: Monoprenylated and inactive RAB4A and RAB11A are recruited to purified Golgi fractions through their prenyl group. (A) 2 µM monoprenylated Alexa488-labeled or GDP-bound RAB4A and RAB11A were incubated with purified Golgi fractions (labeled with Rhodamine WGA (Wheat Germ Agglutinin)). All monoprenylated and inactive RAB proteins were recruited to these membranes. (B) 2 µM unprenylated Alexa488-labeled or GDP-bound RAB4A and RAB11A were incubated with purified Golgi fractions (labeled with Rhodamine WGA (Wheat Germ Agglutinin)). Unprenylated and inactive RAB proteins were not recruited to these membranes. (Scale bar = 10 µm).

Figure 5 . 7 :

 57 Figure 5.7: Monoprenylated RAB4A, RAB11A and RAB6A localize to the same membrane structures. (A) 2 µM monoprenylated GppNHp-bound Alexa488-RAB11A and Alexa568-RAB4A were incubated together with purified Golgi fractions. Both monoprenylated and active RAB proteins were found to colocalize to the same membrane structures. (B) 2 µM monoprenylated Alexa488-RAB6A:GTP and Alexa568-RAB4A:GppNHp were incubated together with purified Golgi fractions. Both monoprenylated and active RAB proteins were found to co-localize to the same membrane structures. (C) 2 µM monoprenylated Alexa488-RAB6A:GTP and Alexa568-RAB11A:GppNHp were incubated together with purified Golgi fractions. Both monoprenylated and active RAB proteins were found to co-localize to the same membrane structures. (Scale bar = 10 µm).

Figure 5 . 8 :

 58 Figure 5.8: Monoprenylated RAB4A, RAB11A and RAB6A do not localize to Golgi or recycling endosomal structures. (A) 2 µM monoprenylated Alexa568-labeled and GppNHp-bound RAB4A or RAB11A were incubated with the medial-trans-Golgi and trans-Golgi network marker Atto488-labeled αRAB6:GTP antibody in the presence of purified Golgi fractions. Both monoprenylated and active RAB proteins were found to preferentially localize to αRAB6:GTP antibody negative structures.(B) Immunofluorescence experiments were performed in presence of 2 µM monoprenylated Alexa488-labeled and GppNHp-bound RAB11A and an antibody targeted against the cis-Golgi marker GM130 in the presence of purified Golgi fractions. Monoprenylated and active RAB11A was found to preferentially localize to α-GM130 antibody negative structures. (C) Immunofluorescence experiments were performed in presence of 2 µM monoprenylated Alexa488-labeled and GTP-bound RAB6A and an antibody targeted against the recycling endosomal marker RAB11 in the presence of purified Golgi fractions. Monoprenylated and active RAB6A was found to preferentially localize to α-RAB11 antibody negative structures. (Scale bar = 10 µm).

Figure 5 . 9 :

 59 Figure 5.9: Study of RAB4A/RAB11A curvature sensing using purified Golgi fractions and kinesin motors. These experiments were performed using microtubule-coated chambers. Biotinylated kinesins interact with biotinylated Golgi membranes in the presence of streptavidin. Addition of an ATP-containing buffer allows kinesins to move on microtubules and to subsequently pull multiple tubes from Golgi membranes. (A) 2 µM monoprenylated, Alexa568-labeled and GppNHp-bound RAB11A was incubated with purified Golgi fractions (labeled with Bodipy Fluorescein Ceramide) (B) 2 µM monoprenylated, Alexa488labeled and GTP-bound RAB6A was incubated with purified Golgi fractions (labeled with Bodipy TexasRed Ceramide). (Scale bar = 10 µm).

Figure 5 . 10 :

 510 Figure 5.10: Description of the PLIF (protein-lipid interaction by fluorescence) assay adapted to my study. GFP-tagged RAB proteins are incubated on anti-GFP antibody-coated 96-well plates. Redfluorescent liposomes are added in the wells with each well containing liposomes of specific size and/or lipid composition. The plate is put under agitation for 15 to 20 minutes to allow RAB-liposome interaction.

Figure 6 . 1 :

 61 Figure 6.1: Vesicle tethering is a RAB6-specific effect. (A) Vesicle tethering was assessed over time (during 30 minutes), using a confocal microscope, in presence of 2 µM monoprenylated Alexa488-RAB6A:GTP. RAB6 was added to the EggPC vesicle mix at t = 00:00. The same observation was made in the presence of the RAB6A' isoform (not shown) (B) The same experiments were carried out in presence of monoprenylated Alexa488-RAB5A:GppNHp or monoprenylated Alexa488-RAB1B:GppNHp and no vesicle tethering was observed. (Scale bar = 10 µm).

Figure 6 . 2 :

 62 Figure 6.2: Vesicle tethering is nucleotide and concentration dependent. (A) Vesicle (pure EggPC) aggregation was assessed, using a confocal microscope, 26 minutes and 40 seconds after addition of 2 µM GTP-bound (left) and GDP-bound (right) monoprenylated Alexa488-RAB6A. No vesicle tethering was observed in presence of inactivated (GDP-bound) RAB6A. (B) Vesicle (pure EggPC) aggregation was assessed, using a DIC microscopy, 30 minutes after addition of various concentrations of monoprenylated RAB6A:GTP. Vesicle tethering was observed in presence of 1.6 µM RAB6A (right) but not after addition of 1.5 µM RAB6A (left). (Scale bar = 10 µM).

Figure 6 . 3 :

 63 Figure 6.3: Vesicle tethering is induced by a RAB6-RAB6 dimerization in trans. (A) Quantification of fluorescence intensities of tethered vesicles. Vesicle (pure EggPC with 0.1% (mol/mol) TexasRed-DHPE lipids) tethering was induced with 2 µM monoprenylated Alexa488-RAB6A:GTP. Fluorescence intensities of the protein (green channel) and lipids (red channel) were measured using the oval profile method (see Chapter 3). In order to assess the enrichment of the protein at the sites of vesicle interaction (plot on the right), each fluorescence intensity value was subtracted by the background and normalized to the average fluorescence intensity value of non-interacting regions. The normalized fluorescence intensity of the noninteracting regions therefore has approximately a value of 1. TexasRed DHPE lipids were found to concentrate 2 times at the interaction site (because of the contact between two membranes) whereas a 5

Figure 6 . 4 :

 64 Figure 6.4: Structure of the YPT1:GDI complex. Ribbon representation of YPT1 (light blue) bound to GDI (green). The geranylgeranyl lipid anchor is shown. Distances are indicated in ångström. Adapted from[START_REF] Pylypenko | Structure of doubly prenylated Ypt1:GDI complex and the mechanism of GDI-mediated Rab recycling[END_REF].

Figure 6 . 5 :

 65 Figure 6.5: The RAB6-RAB6 interaction is dynamic. (A) Effect of the membrane incorporation of 2 µM monoprenylated Alexa488-RAB6A:GTP after vesicle tethering has occurred (achieved with previous addition of 2 µM monoprenylated Alexa568-RAB6A:GTP) was assessed over time (during 30 minutes), by confocal microscopy. Alexa488-RAB6A was added to the tethered EggPC vesicle / Alexa568-RAB6A mix at t = 00:00. (B) Respective enrichments of both labeled proteins at the interaction sites were quantified as in Figure 6.3A. Both proteins were found to concentrate between 3 and 5 times in these regions. (Scale bar = 10 µm).

Figure 6 . 7 :

 67 Figure 6.7: Vesicle tether in presence of RAB6A-Y35R-D49R. Vesicle tethering was assessed over time (during 42 minutes), using a confocal microscope, in presence of 2 µM monoprenylated Alexa488-RAB6A-Y35R-D49R:GTP. Mutated RAB6 was added to the EggPC vesicle mix at t = 00:00. (Scale bar = 10 µm).

First, I tried

  to out-compete this interaction by adding an excess of C-terminally truncated RAB6A (RAB6A ) which cannot be prenylated. I started by monitoring whether RAB6A 8-195 could inhibit vesicle tethering. I induced EggPC vesicle tethering with the addition of 2 µM monoprenylated GTP-bound RAB6A and subsequently to the formation of vesicle aggregates, I added a 10 fold molar excess of GTP-bound RAB6A . Surprisingly, vesicles were still found to tether (not shown). In another experiment, 2 µM monoprenylated Alexa488-labeled GTP-bound RAB6A and a 10 fold molar excess of GTP-bound RAB6A 8-195 were added simultaneously. Vesicles were still aggregating and RAB6 was still found to concentrate at the interaction sites (Figure6.8A), thereby indicating that addition of unprenylated RAB6A does not impede vesicle tethering.

Figure 6 . 8 :

 68 Figure 6.8: Unprenylated RAB6A does not compete with membrane-bound RAB6A. (A) 2 µM monoprenylated Alexa488-RAB6A:GTP and a 10 fold molar excess of GTP-bound C-terminally truncated RAB6A (RAB6A 8-195 ) were added simultaneously (at t = 00:00). Vesicle aggregation was assessed over time (during 30 minutes), by confocal microscopy. RAB6A 8-195 :GTP did not prevent RAB6A-induced vesicle tethering. (B) Equimolar amounts (2 µM) of monoprenylated Alexa568-labeled GTP-bound RAB6A and Alexa488-labeled GTP-bound RAB6A 8-195 were added simultaneously. The membrane fluorescence intensities of each labeled protein were quantified, after vesicle tethering, using the oval profile method (see Chapter 3), and the obtained intensity values were subtracted by that of the background. Although

Figure 6 . 9 :Figure 6 .

 696 Figure 6.9: Bivalent αRAB:GTP antibodies promote vesicle tethering. (A) Effect of the addition of 40 µM ATTO488-αRAB6:GTP after vesicle tethering has occurred (achieved with previous addition of 2 µM monoprenylated Alexa568-RAB6A:GTP) was monitored over time (during 10 minutes), by confocal microscopy. ATTO488-αRAB6:GTP was added to the tethered EggPC vesicle / Alexa568-RAB6A mix at t = 00:00. (Scale bar = 10 µm). (B) Respective enrichments of Alexa568-RAB6A:GTP and ATTO488-αRAB6:GTP at the interaction sites were quantified as in Figure 6.3A. Both were found to concentrate

Figure 6 .

 6 Figure 6.10: αRAB6:GTP (scFv) can access the interaction sites but does not inhibit vesicle tethering. 2 µM monoprenylated Alexa568-RAB6A:GTP and a 5 fold molar excess of αRAB6:GTP (scFv) were added simultaneously (at t = 00:00). Vesicle aggregation was monitored over time (during 24 minutes), by confocal microscopy. αRAB6:GTP (scFv) did not prevent RAB6A-induced vesicle tethering. (B) Equimolar amounts (2 µM) of monoprenylated Alexa568-RAB6A:GTP and Alexa488-αRAB6:GTP (scFv) were added simultaneously. Respective enrichments of Alexa568-RAB6A:GTP and Alexa488-αRAB6:GTP

  µM) of mCherry-LidA 201-583 (amino acids 201 to 583) and monoprenylated Alexa488-labeled GTP-bound RAB6A were added simultaneously, no vesicle tethering was observed (not shown). In a separate experiment, I induced EggPC vesicle tethering with the addition of 2 µM monoprenylated GTPbound RAB6A and subsequently to the formation of vesicle aggregates added a 2.5 fold molar excess of mCherry-LidA 201-583 . Consistently with the previous experiment, the addition of LidA 201- 583 led to a complete destruction of the tethering complex (Figure 6.11A), and to the homogeneous redistribution of Alexa488-RA6A:GTP over the GUV membrane (Figure 6.11B).

Figure 6 . 11 :

 611 Figure 6.11: Inhibition of vesicle tethering by LidA 201-583 . (A) Effect of the addition of 5 µM mCherry-LidA 201-583 after vesicle tethering has occurred (achieved with previous addition of 2 µM monoprenylated Alexa488-RAB6A:GTP) was monitored over time (during 10 minutes), by confocal microscopy. mCherry-LidA 201-583 was added to the tethered EggPC vesicle / Alexa488-RAB6A mix at t = 00:00. (B) Enrichment of Alexa488-RAB6A:GTP at the interaction sites, before (t = 00:00, green) and after (t = 10:16, light green) mCherry-LidA 201-583 addition, were quantified as in Figure 6.3A. Alexa488-RAB6A:GTP was found to concentrate less at the interaction sites after the addition of LidA 201-583 . (Scale bar = 10 µm).

  . The study of the effect of OCRL on vesicle tethering gave rise to conflicting observations. I induced EggPC vesicle tethering with the addition of 2 µM monoprenylated GTP-bound RAB6A. Subsequently to the formation of vesicle aggregates, I added a 10 fold molar excess (20 µM) of either the mCherry-tagged RAB binding domain of OCRL (mCherry-OCRL 538-901 ) or the Alexa488-labeled OCRL 538-901 . Although addition of mCherry-OCRL 538-901 seemed to partially impede the tethering, no alteration was observed after addition of Alexa488-labeled OCRL 538-901 . When performing the same experiment using an untagged version of OCRL 538-901 , the resulting effect was very similar to that obtained with mcherry-tagged OCRL 538-901 . A likely explanation is that the Alexa488 fluorophore might somehow decrease the affinity of OCRL 538-901 for RAB6 thereby making it more difficult to inhibit vesicle tethering. In relation to this, when equimolar amounts (2 µM) of monoprenylated Alexa488-labeled GTP-bound RAB6A and mCherry-OCRL 538-901 were added simultaneously, RAB6A was still found to concentrate around 5 times at the interaction site while mCherry-OCRL 538-901 concentrated up to 2 times (Figure6.12A). Conversely, Alexa488-labeled OCRL 538-901 , when added in equimolar amount, was found to be excluded from the interaction sites (Figure6.12B).

Figure 6 . 12 :

 612 Figure 6.12: Effect of OCRL 538-901 on vesicle tethering. (A) RAB6-induced vesicle tethering was monitored after simultaneous addition of equimolar amounts (2 µM) of monoprenylated Alexa488-labeled RAB6A:GTP and mCherry-OCRL 538-901 , by confocal microscopy. Respective protein enrichments at the interaction sites were quantified as in Figure 6.3A. Alexa488-labeled RAB6A:GTP and mCherry-OCRL 538-901

  1.1 The Lipid bilayer ..................................................................................................................... 20 1.1.1 From a lipid molecule to a bilayer .................................................................................. 20 1.1.2 Different classes of lipids ................................................................................................. 21 1.1.3 Lipid synthesis and distribution in cells ........................................................................ 23

1.2 Membrane domain formation ............................................................................................ 26 1.2.1 Different states of membranes ....................................................................................... 26 1.2.2 Phase state of cellular membranes ................................................................................. 28 1.3 Membrane deformations .................................................................................................... 29 1.3.1 Membrane curvature in cells .......................................................................................... 29 1.3.2 Mechanisms of membrane deformation ........................................................................ 32 1.3.3 Protein curvature sensing ............................................................................................... 39 1.4 In vitro experimental approaches ....................................................................................... 43 1.4.1 Model membranes for in vitro experiments .................................................................. 43 1.4.2 Phase separation from living cells to model membranes .............................................. 45 1.4.3 Curvature sensing on model membranes ...................................................................... 48

  2.1 RAB discovery and evolution .............................................................................................. 54 2.2 RAB sequence and structure ............................................................................................... 55 2.2.1 G-domain......................................................................................................................... 56 2.2.2 RAB specific sequence motifs ......................................................................................... 57

2.2.3 RAB C-terminal region ................................................................................................... 57 2.3 RAB posttranslational modifications .................................................................................. 58 2.3.1 RAB activation cycle ....................................................................................................... 58 2.3.2 RAB membrane insertion and extraction ....................................................................... 61 2.4 Membrane targeting of RAB GTPases ................................................................................ 64 2.5 RAB GTPases and vesicular transport ................................................................................ 68 2.5.1 General mechanism of intracellular transport .............................................................. 68 2.5.2 RABs and membrane tethering ....................................................................................... 71 2.6 Focus on the RAB proteins used in this study ................................................................... 75 2.6.1 RAB1 and the ER-Golgi intermediate compartment ..................................................... 75 2.6.2 RAB6 and the Golgi......................................................................................................... 76 2.6.3 RAB4 / RAB5 / RAB11 and the endosomal system ..........................................................77 2.6.4 RAB35 and the plasma membrane ................................................................................. 80 RAB (Ras-like in brain) proteins are members of the RAS (Rat Sarcoma) superfamily of small

  Protein synthesis and modification ..................................................................................... 81 3.1.1 Protein expression and purification ................................................................................ 81 3.1.2 In vitro modifications of RAB and GST proteins ........................................................... 84 3.2 Experimental studies with GUVs ........................................................................................ 90 3.2.1 Synthesis of giant unilamellar vesicles ........................................................................... 90 3.2.2 Generalities of the experimental approach .................................................................... 93 3.2.3 Curvature sensing experiments with GUVs ................................................................... 98

	3.1

3.3

Experimental studies with purified Golgi membranes ..................................................... 104

Table 3 .1: List of proteins used in this study and how they were obtained

 3 . BL21(DE3) and BL21-CodonPlus(DE3)-RIL cells were respectively obtained from Novagen/Merck and Stratagene/Agilent Technologies. Protino Ni-NTA agarose beads and Protino glutathione agarose 4B beads were both obtained from Macherey-Nagel. The TEV (tobacco etch virus) protease was purified in the lab and the PreScission protease (human rhinovirus 3C protease and GST fusion protein) was purified by Ahmed El Marjou (Recombinant Protein Facility, Institut Curie).

	Protein	Tag	Expression	Affinity column / Beads	Digestion	Final Buffer	Previously described in	Construct from
	Codon optimized RAB1B-CVIL	His6-MBP	BL21(DE3) 0.2 mM IPTG 20h at 20°C	Ni	2+	Yes with TEV	20 mM Hepes DTE, 10 μM GDP pH7.5, 50 mM NaCl, 1 mM MgCl2, 2 mM	Oesterlin 2012	Goody lab, MPI, Dortmund
	RAB4A-CVIL	His6-EGFP	BL21(DE3) 0.3 mM IPTG 20h at 20°C	Ni	2+	Yes with PreScission	50 mM HEPES pH 7,5, 150 mM NaCl, 2 mM DTE, 5 mM MgCl2, 5% Glycerol		Cloning in a pOPINN vector (Addgene)
							25 mM Hepes
	RAB5A-CVIL	His6-MBP	BL21(DE3) 0.2 mM IPTG 5h at 37°C	Ni	2+	Yes with TEV	pH7.5, 200 mM NaCl, 1 mM MgCl2, 2 mM DTE, 10 µM	Oesterlin 2012	Goody lab, MPI, Dortmund
							GDP, 5% Glycerol
	RAB6A-CVIL	His6	BL21-CodonPlus(DE3)-RIL 0.2 mM IPTG 5h at 37°C	Ni 2+	Yes with TEV	25 mM Hepes pH 7.5, 50 mM NaCl, 1 mM MgCl2, 2 mM DTE, 10 μM GTP		Modification of RAB6A-WT plasmid via standard cloning strategy
	RAB6A-WT	His6	BL21-CodonPlus(DE3)-RIL 5h at 37°C 0.2 mM IPTG	Ni	2+	Yes with TEV	20 mM Hepes pH MgCl2, 10 μM GTP 7.5, 50 mM NaCl, 2 mM DTE, 1 mM		Goody lab, MPI, Dortmund
	RAB11A-CVIL	His6-EGFP	BL21(DE3) 0.3 mM IPTG 20h at 20°C	Ni 2+	Yes with PreScission	25 mM HEPES pH 7,5, 200 mM NaCl, 2 mM DTE, 2 mM MgCl2, 10 µM GDP		Cloning in a pOPINN vector (Addgene)
	RAB35-CVIL	His6-EGFP	BL21-CodonPlus(DE3)-RIL 0.2 mM IPTG 20h at 20°C	Ni 2+	No	25 mM Hepes pH 7,5, 50 mM NaCl, 2 mM DTE, 1 mM MgCl2, 10 μM GDP		Cloning in a pOPINN vector (Addgene)
	GST-CVIL	GST	BL21(DE3) 0.3 mM IPTG 20h at 20°C	Glutathione beads	No	50 mM Hepes pH 7.5, 150 mM NaCl, 1 mM DTE, 5% Glycerol		Cloning in a pGEX-6P1 vector (Addgene)
									Goody lab, MPI,
									Dortmund.
	REP-1	His6	Insect cells (Sf9)	Ni 2+	No	25 mM HEPES pH 7.2, 40 mM NaCl, 10 mM DTT	Alexandrov 1999	Purified by Ahmed El Marjhou (Recombinant
									Protein Facility
									of Institut Curie)
			BL21-
	RABGGTase (α and β subunits)	α : His6-GST β : His6	CodonPlus(DE3)-RIL electrcompetent 0.1 mM IPTG	Ni	2+	Yes with TEV	20 mM Hepes pH 7.5, 50 mM NaCl, 2 mM DTE	Dursina 2006	Goody lab, MPI, Dortmund
			20h at 16°C

* BCCP: Biotin carboxyl carrier protein.

Table 3 .2: List of primary and secondary antibodies used in this study.

 3

	Primary antibody	Species	Dilution	From	Secondary antibody	Fluorescent dye	Dilution
	Anti-RAB11	Rabbit	1/100	Invitrogen	Anti-Rabbit	Far red Alexa647	1/400
				Recombinant
	Anti-RAB1 (ROF7)	Human	1/100	antibodies platform,	Anti-Human	Red Cy3	1/400
				Institut Curie
	Anti-GM130	Mouse	1/1000	BD Biosciences	Anti-Mouse	Far red Alexa647	1/400
	Anti-EEA1	Goat	1/200	Santa Cruz Biotechnology	Anti-Goat	Red Cy3	1/400
	Anti-ER	Mouse	1/1000	Perez team, Institut Curie	Anti-Mouse	Red Cy3	1/400

packing defects and membrane charge control RAB GTPase recruitment Running title: RAB proteins bind lipid packing defects

	Guillaume Kulakowski	a,b , Hugo Bousquet

  Specificities of RAB6-induced membrane tethering ........................................................ 158 6.1.1 Vesicle tethering is a RAB6-specific effect .................................................................... 158 6.1.2 Vesicle tethering is nucleotide and concentration dependent .................................... 158 6.1.3 RAB6-induced vesicle tethering is mediated by a RAB-RAB dimerization in trans .... 161 6.1.4 The RAB-RAB interaction is dynamic ........................................................................... 163 6.2 Involvement of the Switch regions .................................................................................... 165 6.2.1 RAB6A mutant induces vesicle tethering ..................................................................... 165

	6.1

6.2.2 Unprenylated RAB6A does not interact with membrane-bound RAB6A ................... 166 6.2.3 Bivalent αRAB:GTP antibodies promote vesicle tethering .......................................... 168 6.2.4 Effect of monovalent RAB6 effector proteins ............................................................... 170 6.3 Discussion ........................................................................................................................... 174

Prior to my arrival to the lab, similar in vitro studies, using purified and prenylated RAB proteins and synthetic vesicles as model membranes, had led to the observation that vesicles tether in presence of monoprenylated and GTP-bound RAB6A. Part of my project consisted in investigating this process in greater detail.

  Nous sommes parvenus, pour la première fois, à incorporer des protéines RAB purifiées et prénylées dans des membranes artificielles. Nous avons tout d'abord observé que RAB6 est capable de promouvoir une agrégation de vésicules, phénomène qui n'est pas observé avec RAB1 et RAB5. Nous suggérons un modèle dans lequel RAB6 interagit en trans avec lui-même et par conséquent induit un accolement de membranes. La partie principale de cette étude consistait à identifier les propriétés physicochimiques des membranes requises pour le recrutement des protéines RAB. Nous avons observé que RAB1, RAB5 et RAB6 se lient préférentiellement à des membranes désordonnées et courbées, phénomène qui s'explique par l'insertion du groupement prenyl hydrophobe au niveau de défauts d'agencement de lipides. En revanche, le recrutement de RAB35 requiert la présence de lipides chargés négativement et peut être modulé, dans une moindre mesure, par les défauts d'agencement lipidique. Bien que RAB4 et RAB11 soient recrutées sur des fractions de Golgi purifiées, les charges membranaires et les défauts d'agencement lipidique ne sont pas suffisants pour permettre leur recrutement sur des vésicules synthétiques. Cela suggère que le recrutement de RAB4 et RAB11 nécessite des propriétés membranaires plus complexes. Nos travaux démontrent que les propriétés membranaires sont cruciales pour la localisation spécifique des protéines RAB.

Acknowledgements ................................................................................................................... Abbreviations ...........................................................................................................................

Acknowledgements

Acknowledgements

When I first encountered Bruno Goud, it was to start as an engineer, and eventually start a few months later a PhD. At the same time, I had a very interesting offer for a 2 year-position in a Acknowledgements Acknowledgements

Abbreviations

List of supplementary information Supplementary text: Material and methods not necessary to understand the overall message of the paper, but required to reproduce the experiments.

Figure S2

: Graphic illustration of the different prenylation reactions performed in this study. This figure is mentioned at the beginning of the results and discussion section and can also be associated with the main material and methods section. It should allow the reader to better understand the protein modification reactions performed.

Figure S3: Control experiment to demonstrate why our study focused on the use of monoprenylated proteins rather than diprenylated proteins, which are most commonly found in the cell. This figure is mentioned in the first part of the results and discussion section and demonstrates that mono-and diprenylated RAB6 proteins are recruited to the same specific membrane domains.

Figure S4:

Graphic illustration of the optical tweezer setup used to pull tubes from GUVs and of the quantification method used to assess protein enrichment to the tube. This figure is mentioned in the RAB curvature sensing section and can also be associated with the main material and methods section. It should help the reader to better understand the overall method used to study protein curvature sensing. showing the charge properties of the last 20 amino acids of all human RAB proteins. This figure is mentioned in the third part of the results and discussion section (on the study of RAB35 membrane recruitment). It will allow the reader to assess more easily the charge variations among the hypervariable domains of all RAB proteins and more specifically the higher positive charge of RAB35 and RAB23 as compared to the other RABs.

Table S1: Table

Supplementary figure and table legends

Fig. S1. Membrane recruitment of RAB proteins depends on the presence of the prenyl group. GUV phase separation was achieved by using a 3:1:3 (molar ratios) BSM:Cholesterol:DOPC lipid mixture. The Ld phase (made of of unsaturated lipids) was labelled with 0.1% (mol/mol) TexasRed-DHPE lipids (red) whereas the Lo phase (made of saturated lipids) was unlabeled. GUVs were incubated with 2 µM Alexa488-labeled and unprenylated proteins. No membrane binding was observed with unprenylated proteins. (Scale bar: 10 μm).

Fig. S2. Mechanisms of RAB prenylation.

(A) Native diprenylation machinery consisting of a RAB protein with a double cysteine motif at its C-terminal extremity, the RAB Geranylgeranyl Transferase (RABGGTase, with two subunits α and β) and the RAB Escort Protein (REP). The RABGGTase transfers two geranylgeranyl groups on the double cysteine motif of the RAB protein, with the REP acting as a chaperone. (B) Monoprenylation of a RAB protein mediated by the Geranylgeranyl Transferase type 1 (two subunits α and β). The GGTase1 recognizes the CVIL motif at the C-terminus of the RAB protein (CAAX box) and transfers one geranylgeranyl group on the cysteine. (C) Structure of the geranylgeranyl pyrophosphate group. This lipid moiety has a C20 isoprenoid structure containing four unsaturations.

Fig. S3. Diprenylated and monoprenylated GDP-bound RAB6 are specifically recruited to disordered membranes. GUV phase separation was achieved by using a 3:1:3 (molar ratios) BSM:Cholesterol:DOPC lipid mixture. The Ld phase (made of of unsaturated lipids) was labelled with 0.1% (mol/mol) TexasRed-DHPE lipids (red) whereas the Lo phase (made of saturated lipids) was unlabeled. Lo and Ld GUVs were respectively composed of 1:1 (molar ratios) BSM:Cholesterol and DOPC:Cholesterol. Alexa488 labeled RAB6 proteins were used at a 2 µM final concentration. In vitro diprenylation of RAB6 WT by both REP and the RABGGTase can only be performed if the RAB is inactive (i.e. GDP-bound). REP interacts with the RAB protein and then binds the RABGGTase in the presence of geranylgeranyl pyrophosphate. As the affinity of the RAB protein for the REP is very high (low nanomolar range) (9) , the RAB remains tightly bound to the complex even in the presence of membranes, thus preventing its recruitment. In order to overcome this issue and allow RAB recruitment, we added to the mixture a truncated version of the RAB6 supereffector from Legionella pneumophila LidA (LidA201-583), which as an unusually high affinity for the RAB proteins in their GDP and GTP bound forms (respectively 4 nM and 30 pM affinities). As it was shown that LidA201-583 covers an unusually large surface area of some RAB proteins (4) and therefore shows overlapping binding sites with REP, it is possible to outcompete REP binding through the addition of this effector protein. By adding an equimolar concentration of LidA201-583 and RAB:REP:RABGGTase (2 µM) in the presence of membranes, we observed RAB membrane recruitment, confirming that LidA201-583 outcompetes the RAB:REP:RABGGTase interaction. However, binding of RAB6 was also observed on Lo GUVs under these conditions. This is probably due to the dissociation of the RAB from the complex which forces the newly added hydrophobic prenyl groups to be in contact with the solvent. Therefore, because it is more energetically favorable, the RAB incorporates its prenyl group into the hydrophobic and more stable membrane environment. To prove this theory, we added GGTase1 to the experimental chamber (same molar ratios as for the monoprenylation reaction, see material and methods). Once LidA201-583 has outcompeted the RAB:REP:RABGGTase interaction, GGTase1 (which does not have overlapping RAB binding sites with LidA201-583) will interact with the prenyl group and shield it from the solvent. (A) In the presence of both LidA201-583 and GGTase1, diprenylated RAB6 localizes specifically to the Ld phase on phase separated GUVs and to Ld vesicles. No recruitment is observed on Lo GUVs. (B) Monoprenylated GDP-bound RAB6 exhibits the same recruitment behavior in the presence of LidA201-583 and GGTase1 as diprenylated RAB6. In the absence of LidA201-583, monoprenylated GDP-bound RAB6 was also shown to bind specifically to Ld domains (Fig. 1) which demonstrates that this effector does not have any influence on the specific localization of RAB6 proteins. We conclude that RAB6 specifically binds to Ld domains independently of the quantity of prenyl groups. (Scale bar: 10 μm).

Fig. S4

. Pulling a tube from a GUV using an optical tweezer setup. (A) A biotin containing GUV was aspirated with a micropipette connected to a water reservoir which allows to control membrane tension. A bead coated with streptavidin was immobilized using an optical trap (highly focused infra-red laser beam) (10,11) . After addition of 100-300 nM of protein, a tube was pulled from the GUV through the interaction of the biotinylated lipids with the immobilized streptavidin coated bead. GUV tension was increased in a stepwise manner by decreasing the height of the water tank. 1 minute waiting time was required at each tension step increase to allow protein redistribution and sorting. (B) Vesicles were formed using EggPC lipids. Lipids were labelled using 0.1% (mol/mol) TexasRed-DHPE and 0.035% of DSPE-PEG(2000)-Biotin was added to the lipid mix to achieve adhesion between the GUV membrane and the streptavidin coated beads. At each tension step, confocal images of the green (protein) and red (lipid) channels were obtained and both fluorescence intensities were measured on the tube and on the GUV. The graphs represent the plots of intensities for both channels on the GUV and on the tube. Fluorescence intensities were calculated by subtracting the background noise (inside the vesicle) from the maximum of the fluorescence peak. The equation on the right defines the protein sorting ratio S as the protein/lipid signal ratio on the tube divided by that observed on the GUV. (Scale bar: 10 µm).

Table S1. Charges at the RAB protein C-termini (last 20 amino acids).

The charges of the last 20 amino acids of each RAB protein were calculated at a physiological pH (7.4) using a protein calculator tool (Protein calculator.

[online] Available at: http://protcalc.sourceforge.net/). The amino acid residues colored in green are positively

Involvement of the Switch regions 6.2.1 RAB6A mutant induces vesicle tethering

To further understand the mechanism regulating RAB6-RAB6 dimerization, we searched for published crystal structures in which RAB6 is a dimer. One crystal structure of RAB6 from Drosophila melanogaster (DOI: 10.2210/pdb2y8e/pdb) was found to be crystalized as a dimer (Figure 6.6) [START_REF] Walden | Structure of the Drosophila melanogaster Rab6 GTPase at 1.4 A resolution[END_REF]. The analysis of the dimer crystal structure by Olena Pylypenko (Anne Houdusse's lab, Institut Curie) revealed that the RAB6-RAB6 interaction might occur via βsheet-β-sheet interactions (hydrogen bonds) and/or an aromatic ring interaction between tyrosines (at position 35) (Figure 6.6). To test this hypothesis, the aspartic acid (D) 49 residue, localized to the β-sheet structure and known to not be involved in interaction with effectors, and the tyrosine (Y) 35 residue were both mutated to arginine (R), a positively charged bulky residue which was expected to impede dimerization through repulsive electrostatic interactions. The mutated RAB6A protein (RAB6A-Y35R-D49R) was purified, exchanged to GTP, labeled and monoprenylated in a similar manner as non-mutated RAB6A. 2 µM monoprenylated Alexa488labeled GTP-bound RAB6A-Y35R-D49R was incubated in presence of EggPC-containing GUVs.

In order to precisely investigate RAB6A was fluorescently labeled at its N-terminus with an Alexa488 fluorophore. Equimolar amounts (2 µM) of monoprenylated Alexa568-labeled GTP-bound RAB6A and Alexa488-labeled GTP-bound RAB6A 8-195 were added simultaneously. Interestingly, RAB6A was positively recruited to the vesicles and concentrated at the interaction sites whereas RAB6A was found to be completely excluded from all of these regions (Figure 6.8B); thereby indicating that RAB6A and RAB6A 8-195 cannot interact. The absence of interaction logically explains why vesicle tethering was not impaired but also strongly suggests that monoprenylated RAB6A might undergo conformational changes upon membrane binding, thereby preventing its interaction with unprenylated RAB6A. Therefore, the previously described absence of liposome aggregation in presence of RAB6A [START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF]) could be explained by the fact that RAB6A membrane binding is mediated via a His tag-Nickel-NTA interaction which might not cause conformational changes necessary for RAB6 dimerization. The above results also imply that the RAB-mediated tethering effects previously described [START_REF] Lo | Insights into lipid raft structure and formation from experiments in model membranes[END_REF][START_REF] Tamura | Membrane-anchored human Rab GTPases directly mediate membrane tethering in vitro[END_REF] and the RAB6 specific tethering effect described here might be a result of different mechanisms involving differently folded proteins.

Bivalent αRAB:GTP antibodies promote vesicle tethering

In order to out compete the RAB-RAB interaction, I investigated the effect of a green labeled anti-RAB6:GTP antibody (ATTO488-αRAB6:GTP, from Adipogen). The addition of a 20 fold molar excess ATTO488-αRAB6:GTP prior to, but also after, RAB6-induced vesicle tethering did not prevent or inhibit vesicle aggregation (Figure 6.9A). Interestingly, Alexa568-RAB6A and ATTO488-αRAB6:GTP were both found to concentrate, between 4 and 6 times, at the interaction sites (Figure 6.9B). The effect of αRAB6:GTP on vesicle tethering was also monitored by cryoelectron microscopy (again in collaboration with Daniel Levy's team) and clear protein densities could be detected at the interface between tethered vesicles (Figure 6.9C). The space between interacting membranes was also increased three times as compared to when αRAB6:GTP is not added to the GUV/RAB6 mix (Figure 6.9C left and Figure 6.3B right). This could be explained by the fact that the 110 kDa αRAB6:GTP is a bivalent (dimeric) antibody and can bind two RABs from different membranes and its recruitment to the interaction site would increase the space between the interacting vesicles (Figure 6.9C right).

Concluding remarks

In this study, we have managed for the first time to incorporate prenylated RABs into artificial membranes. My PhD project started with the study of the RAB6-induced membrane tethering, a phenomenon which had just been observed before my arrival to the lab. I have confirmed that RAB6, but not RAB1 or RAB5, is able to promote by itself, and in a GTP-dependent manner, vesicle tethering. We believe that prenylated RAB6 is able to interact in trans with itself and thereby promotes homotypic membrane tethering. This suggests that RAB6 might regulate, in concert with tethering factors or lipid transfer proteins, membrane recognition and tethering events during lipid and protein transport processes.

The main part of my project consisted in investigating the membrane binding properties necessary for the effective recruitment of prenylated RAB GTPases. RAB1, RAB5 and RAB6 were all found to only localize to Ld membrane domains and to preferentially bind to curved membranes; and this seemed to occur independently of the nucleotide-bound state and the number of geranylgeranyl groups. We demonstrated that the specific recruitment of RAB1, RAB5 and RAB6 is primarily dependent on the hydrophobic insertion of the prenyl group into lipid packing defects. We therefore showed that, in accordance with recently published data on farnesylated N-RAS proteins (Larsen et al., 2015;Larsen et al., 2017), prenyl groups do not only act as non-specific anchors but are also involved in specific membrane targeting. This lipid-driven mechanism might partially explain how RAB proteins could bind to intracellular membranes which are mainly found in a disordered state (Niko et al., 2016), and associate with highly curved transport carriers.

Although the membrane incorporation of the prenyl group was shown to be sufficient for RAB1, RAB5 and RAB6 membrane binding, it was proven not to be the case for other RABs.

The membrane recruitment of RAB35, known to localize in cells to the endocytic compartments and the plasma membrane (Kouranti et al., 2006), was shown to be primarily dependent on the presence of negatively charged lipids. This observation is consistent with previous studies [START_REF] Heo | PI(3,4,5)P3 and PI(4,5)P2 lipids target proteins with polybasic clusters to the plasma membrane[END_REF]Li et al., 2014) demonstrating that RAB35 plasma membrane localization is regulated by the interaction of its C-terminal positively charged hypervariable region with negatively charged phosphoinositides. Although lipid packing defects were shown to enhance RAB35 membrane affinity, they were not essential for membrane binding. RAB23,