Compacted structural domain PCR Polymerase chain reaction CT Chromosome territory Pds5 Precocious dissociation of sisters 5 (protein) CTCF CCCTC-binding factor pH3 Phospho-histone H3

I want to thank my family

ii Abstract Characterizing the 3D organization of holocentric chromosomes in Bombyx mori The genome's organization within the cell nucleus has been observed and described in various different organisms for over 140 years. Most of the organisms where this has been done are monocentric, or organisms that have chromosomes with a single centromere. Studies have shown that centromeres strongly constrain the architecture of chromosomes in interphase and during my thesis, I contributed to a review describing this phenomenon [START_REF] Muller | The Impact of Centromeres on Spatial Genome Architecture[END_REF]. However, across the eukaryotic tree of life you can find several examples of organisms that are holocentric, or organisms that have centromeres distributed along the entire length of their chromosomes.

To study the impact of centromeres on these types of chromosomes, we choose the silkworm, Bombyx mori, as our model organism. Although the holocentric chromosomes of B. mori have been subjects of studies describing the organization and formation of their centromeres and kinetochores (Cortes-Silva et al., 2020;[START_REF] Senaratne | Formation of the CenH3-Deficient Holocentromere in Lepidoptera Avoids Active Chromatin[END_REF], their genome organization has yet to be described. My thesis aims at characterizing the genome organization of B. mori with the use of sequencing-based techniques and bioinformatic approaches on two experimental systems.

In the first part of my thesis I used a combination of Hi-C and ChIP-Seq data from B. mori embryos to identify and characterize genome organizational features. Using Hi-C, I produced contact maps for all 28 chromosomes of the B. mori genome assembly and I was able to show that B. mori chromosomes make infrequent contacts between themselves resulting in strong chromosome territories. I then combined this Hi-C data with ChIP-Seq data sets corresponding to several B. mori embryo epigenetic marks in order to define and characterize chromosome compartments. This study revealed that B. mori chromosomes are organized into three genome-wide compartments: A, B and X. The A and B compartments in B. mori are reminiscent of those first described in human chromosomes. The X compartment is composed of highly compact, genepoor regions that do not interact with the other two compartments, nor with like compartments on the same chromosome. These findings are included in a study that I was a part of describing the genome organization of B. mori embryos (Muller et al., in progress).

In the second part of my thesis I turned to B. mori cell lines in order to determine some of the factors contributing to this genome organization. In order to do this, I used RNAi to perturb centromeres, cohesins and condensins, which have been shown to have an impact on genome organization in other organisms. I acquired Hi-C data and profiled different epigenetic marks in each condition and show that centromeres and SMC complexes play a role in B. mori genome organization with cohesin and condensin II having opposite effects in short-and long-range chromosome folding. To properly analyze this data, I developed bioinformatic tools to account for the holocentric nature of B. mori chromosomes. I even had the chance to contribute my bioinformatic skill set to a collaboration studying meiotic pairing of B. mori chromosomes (Rosin et al., 2021b). Taken together, using Hi-C and ChIP-Seq and performing the bioinformatic analysis of both, I was able to describe for the first time the genome organization of B. mori and characterize the roles of factors contributing to it.

iii Résumé Caractérisation de l'organisation tridimensionnelle des chromosomes chez l'organisme holocentrique Bombyx mori L'organisation du génome dans le noyau cellulaire a été observée et décrite dans différents organismes depuis plus de 140 ans. La plupart des organismes où cela a été fait sont monocentriques, c'est-à-dire dont les chromosomes n'ont qu'un seul centromère. Il a été montré que les centromères contraignent fortement l'architecture des chromosomes en interphase, et j'ai contribué lors de ma thèse à l'écriture d'une revue décrivant ce phénomène [START_REF] Muller | The Impact of Centromeres on Spatial Genome Architecture[END_REF]. Cependant, à travers l'Arbre de la Vie eucaryote, on peut trouver plusieurs exemples d'organismes qui sont holocentriques, c'est-à-dire qui ont plusieurs centromères distribués sur toute la longueur de leurs chromosomes. Pour étudier l'impact des centromères sur ce type de chromosomes, nous avons choisi comme organisme modèle le ver à soie, Bombyx mori. Bien que les chromosomes holocentriques de B. mori aient fait l'objet d'études décrivant l'organisation et la formation de leurs centromères et kinétochores (Cortes-Silva et al., 2020 ;[START_REF] Senaratne | Formation of the CenH3-Deficient Holocentromere in Lepidoptera Avoids Active Chromatin[END_REF], l'organisation de leur génome reste à décrire. Ma thèse vise à caractériser l'organisation du génome de B. mori en utilisant des techniques basées sur le séquençage et des approches bioinformatiques sur deux systèmes expérimentaux.

Dans la première partie de ma thèse, j'ai utilisé une combinaison de données Hi-C et ChIP-Seq d'embryons de B. mori pour identifier et caractériser les caractéristiques de l'organisation du génome. En utilisant les techniques Hi-C, j'ai produit des cartes de contact pour les 28 chromosomes de l'assemblage du génome de B. mori et j'ai pu montrer que les chromosomes de B. mori établissent des contacts peu fréquents entre eux, ce qui donne lieu à des territoires chromosomiques forts. J'ai ensuite combiné ces données Hi-C avec des ensembles de données ChIP-Seq correspondant à plusieurs marques épigénétiques de l'embryon de B. mori afin de définir et de caractériser les compartiments chromosomiques. Cette étude a révélé que les chromosomes de B. mori sont organisés en trois compartiments à l'échelle du génome : A, B et X. Les compartiments A et B de B. mori rappellent ceux décrits pour la première fois dans les chromosomes humains. Le compartiment X est composé de régions très compactes et pauvres en gènes qui n'interagissent ni avec les deux autres compartiments, ni avec des compartiments similaires sur le même chromosome. Ces résultats sont inclus dans une étude à laquelle j'ai participé et qui décrit l'organisation du génome des embryons de B. mori (Muller et al., en cours).

Dans la deuxième partie de ma thèse, je me suis tourné vers les lignées cellulaires de B. mori afin de déterminer certains des facteurs contribuant à cette organisation du génome. Pour ce faire, j'ai utilisé l'ARN interférence pour perturber les centromères, les cohésines et les condensines, dont l'impact sur l'organisation du génome a été démontré chez d'autres organismes. J'ai acquis des données Hi-C, profilé différentes marques épigénétiques dans chaques conditions, et montré que les centromères et les complexes SMC jouent un rôle dans l'organisation du génome de B. mori, la cohésine et la condensine II ayant des effets opposés sur le repliement des chromosomes à courte et longue distance. Pour analyser correctement ces données, j'ai développé des outils bioinformatiques pour tenir compte de la nature holocentrique des chromosomes de B. mori. J'ai également eu l'occasion de mettre mes compétences en bioinformatiques au service d'une collaboration portant sur l'appariement méiotique des chromosomes de B. mori (Rosin et al., 2021). Ensemble, en utilisant Hi-C et ChIP-Seq et en effectuant l'analyse bioinformatique des deux, j'ai pu décrire pour la première fois l'organisation du génome de B. mori et caractériser les rôles des facteurs qui y contribuent. The previous methods rely on a specific protein-DNA interaction, but the following sequencingbased methods do not. Split-Pool Recognition of Interactions by Tag Extension, or SPRITE, is also done in crosslinked conditions. The nuclei are then isolated and the chromatin is fragmented. The entire working sample is then "split" evenly into 96 wells where they are all ligated with unique barcodes. All 96 wells are then pooled and subsequently split again into 96 wells where another unique barcode is added. This is repeated several times [START_REF] Quinodoz | Higher-Order Interchromosomal Hubs Shape 3D Genome Organization in the Nucleus[END_REF]. If DNA fragments belong to the same cluster, then they should have the same sequence of unique barcodes because they should stay together after every "split". This allows you to determine spatial proximity of not only DNA-DNA multivalent interactions, but also DNA-RNA and RNA-RNA multivalent interactions. Genome Architecture Mapping (GAM) is another technique that allows you to recover genome-wide multivalent interactions. GAM uses fixed cells that are frozen and cut into ultrathin slices via cryosectioning. The slicing is done in random orientations, and the DNA is extracted and sequenced from each slice. Loci that are closer to each other in spatial proximity are sequenced together more frequently than those that are not [START_REF] Beagrie | Complex multi-enhancer contacts captured by genome architecture mapping[END_REF]. This information allows you to build an interaction map based on all of the DNA fragments that were sequenced together. GPSeq is another method that allows you to inferring distances and genome organization from the nuclear lamina to the nuclear radius, genome-wide. It does this by relying on time-dependent, gradual restriction digestion of chromatin from the nuclear lamina toward the nuclear center, followed by sequencing of the generated cut sites [START_REF] Girelli | GPSeq reveals the radial organization of chromatin in the cell nucleus[END_REF]). The result is information regarding the radial genome organization of the cell, which can also be paired with Hi-C to better model whole-genome structure.

The sequencing-based methods that have arguably been the most common and have almost become synonymous with studying spatial genome organization are chromosome conformation capture (3C)-based techniques. These methods will be discussed in detail in the following section.

Development of 3C Techniques

Chromosome conformation capture is a technique that was developed to elucidate the spatial interactions between two different loci. With the arrival and development of microscopy, we were able to see that the genome is spatially organized within the nucleus. We also knew that this 3D organization played a crucial role in gene regulation and genome stability. Due to the limitation in resolution of most visual methods at the time, we were unable to see interactions interactions between two know target sequences, but it had its limitations. The very nature of the method made it so you could only detect contacts between know sequences, and you could only look at two different loci at a time leading to the "one vs one" description. Also, there was a limit in the distance between the loci only up to a few hundred kilobases [START_REF] Simonis | An evaluation of 3C-based methods to capture DNA interactions[END_REF]. Another limitation in 3C, that was later overcome with Hi-C, was the inability to achieve sequencing deep enough to garner genome-wide information. Chromosome conformation capture-on-chip (4C) was a step up from 3C because it allowed you to look at all of the interactions being made with a single locus. Instead of the chimeric DNA strands being quantified immediately by PCR their ends are cleaved by another restriction enzyme and, at low concentrations, the fragments circularize. This is followed by inverse PCR with a primer based on a region of interest. The resulting fragments represented all the contacts made with that region and then these fragments were evaluated using a microarray chip, leading to the "on-chip" portion of the name [START_REF] Simonis | Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C)[END_REF]. The method later moved away from microarray and use high-throughput sequencing instead. This version of the method was called 4C-Seq [START_REF] Gheldof | Detecting Long-Range Chromatin Interactions Using the Chromosome Conformation Capture Sequencing (4C-seq) Method[END_REF]. Another variation of 4C that was also coupled with parallel sequencing was used to produce the first genome-wide contact map in yeast [START_REF] Duan | A three-dimensional model of the yeast genome[END_REF]. Chromosome conformation capture carbon copy (5C) might be a bit misleading in its name since one might think it builds off of the 4C method, but in fact it is analogous to 3C. 5C allowed the simultaneous detection of millions of interactions and this is due to the way the PCR primers are designed. This method also uses primers specific to regions of interest, except these primers also contain a universal sequence at their ends that allow for multiplexing [START_REF] Dostie | Chromosome Conformation Capture Carbon Copy (5C): A massively parallel solution for mapping interactions between genomic elements[END_REF]. Given that this method, in principal, allows for several 3C

experiments to be done at once, it carries many of the same limitations as 3C.

2.1.2. ChIA-PET and Capture Hi-C

Chromatin interaction analyses combined with paired-end tagging (ChIA-PET) was one of the first 3C-based methods developed that had a step to enrich interactions. In the case of ChIA-PET, the method incorporates a chromatin immunoprecipitation (ChIP) experiment to capture a protein of interest after the fixation and fragmentation step of a 3C-based protocol. This method allows you to recover only the contacts that were bound by your protein of interest. ChIA-PET brought the advantage of producing a library of relatively low complexity, meaning that you could validate more contacts with less reads [START_REF] Han | 3C and 3C-based techniques: the powerful tools for spatial genome organization deciphering[END_REF]. However, you still needed a large amount of cells in order to recover these contacts, making it very inefficient compared to later variations [START_REF] Mumbach | HiChIP: efficient and sensitive analysis of protein-directed genome architecture[END_REF]. Capture Hi-C also enriches for specific interactions, but not those specific to a protein of interest. Capture Hi-C is an adaptation to the Hi-C method, which will be discussed in detail in the following section. The Capture Hi-C method follows the Hi-C protocol almost exactly. Except, after the Hi-C library is amplified, an RNA bait based on a genomic region of interest is used to pull down the Hi-C fragments that are specific to that region.

Then, this subset of fragments is amplified, followed by high-throughput sequencing [START_REF] Mifsud | Mapping long-range promoter contacts in human cells with high-resolution capture Hi-C[END_REF].

Hi-C

Hi-C was the first 3C-based method that was used to identify spatial chromatin interactions genome-wide. The first steps of the method are similar to the rest of the 3C-based methods:

fixation, digestion and ligation. However, the ligation step is critical because the ends of the fragments are filled in with biotin and then they undergo blunt-end ligation. The fragments are then sonicated and de-crosslinked and pulled down via the biotin. This recovers all possible contacts that were biotinylated, genome-wide. These chimeric DNA fragments are then ligated to adapters and go through high-throughput sequencing (Figure 6). Having been established in 2009, the Hi-C protocol has undergone some optimizations and variations of the method have also been introduced claiming to improve upon Hi-C or make it more efficient in certain ways.

Here, I will discuss the original method and what came from it. I will also describe some of these optimizations followed by some alternate uses.

Early Studies

Originally, the Hi-C method was performed on human cell lines and generated over eight million uniquely mapping reads [START_REF] Lieberman-Aiden | Comprehensive Mapping of Long-Range Interactions Reveals Folding Principles of the Human Genome[END_REF]. This method was novel in the sense that it did not involve any primers selecting for ligation events within a specific region of interest.

Instead, you biotinylated the ligation junction, which allowed you to target the ligated fragments that encompass the information, rather than all sequences in between the restriction sites that are just pieces of the genome without any 3D information. Because this ligation process was done in diluted conditions you favored ligation events between fragments that were crosslinked near each other in spatial proximity. This new method boasted an unbiased approach to identify chromatin interactions genome-wide. Of course, the lack of bias was referring to the genomic regions past 3C-based methods were limited to, not to technical and experimental biases, which will be discussed in 3.4.

Optimizations and Adaptations

Since the introduction of Hi-C there have been variants of the technique that have been developed to improve the types of results you would expect to obtain from Hi-C. Each variant's protocol seems to be very similar, if not the same, as Hi-C with some exceptions at specific steps.

Here I will describe three of these variants, the differences between them and Hi-C and the results obtained from using them. 3.2.1. Tethered Conformation Capture (TCC) In 2011, Kalhor et al. published a method called TCC that was designed to be an improved alternative to Hi-C. By performing the ligation step on a solid substrate instead of in solution it was supposed to greatly enhance the signal-to-noise ratio. This was done by biotinylating the protein in the crosslinked DNA-protein complexes, followed by restriction enzyme digestion.

After this, the biotinylated proteins were immobilized on streptavidin-coated beads. This allowed you to essentially remove all non-crosslinked DNA fragments that could ligate to DNA fragments they are not crosslinked to. Another improvement was that the fragment overhangs left after restriction enzyme digestion were not only filled with a biotinylated nucleotide, but also with an α-thio-triphosphate-containing nucleotide that is supposed to be resistant to exonucleases. This method was directly compared to Hi-C under the same conditions and it was shown that, by reducing random intermolecular ligations, TCC had increased sensitivity for contact signal in the interchromosomal space. However, there was not much improvement in the intrachromosomal data between TCC and Hi-C.

In-situ Hi-C

In the case of in-situ Hi-C, it was the original developers of the Hi-C method that improved upon their own protocol. The method is the same step-by-step method as before, except in in-situ the chromatin is first crosslinked and then the nucleus is permeabilized while in the original protocol it's the other way around. By staying in a constrained environment like the nucleus this method is supposed to reduce the frequency of random ligation events, like TCC. This method also used a 4-cutter enzyme instead of a 6-cutter enzyme like the one used in the original protocol. By doing this it was able to achieve much higher resolution than before. This method was also faster to complete than the original, shortening the protocol from seven days to three days [START_REF] Rao | A 3D Map of the Human Genome at Kilobase Resolution Reveals Principles of Chromatin Looping[END_REF].

Micro-C

In contrast with the previous two methods, Micro-C is not meant to substitute the Hi-C protocol, but instead enhance the resolution at short genomic distances. Instead of digesting crosslinked chromatin with a restriction enzyme, Micro-C uses micrococcal nuclease to fragment chromatin.

This makes it possible to achieve nucleosome resolution, the highest of any Hi-C-based technique.

The method is very well-suited to study short-range contacts and nucleosome fiber folding.

However, the method falters when it comes to capturing long-range contacts. This is precisely why it is mentioned that this method can be used to compliment Hi-C, but not replace it [START_REF] Hsieh | Mapping Nucleosome Resolution Chromosome Folding in Yeast by Micro-C[END_REF].

Uses in Genome Assembly and Genome Assembly Correction

The result of a Hi-C experiment should be a 2D, visual representation of the 3D genome conformation in a cell population. Given the physical properties of chromatin, there are certain contact trends and signals you expect to see on a Hi-C contact map, especially in regards to contact decay. Because of this, genome misassemblies are visually obvious in Hi-C maps. Errors in a genome assembly create contact signatures on the Hi-C map that are physically impossible whenever the 1D and 3D signals agree. Very often, the misassemblies are easy to identify. Today, genome assembly not only requires a mixture of short-and long-reads to efficiently assemble scaffolds and contigs, but it also requires Hi-C reads to ensure that these scaffolds and contigs are ordered correctly based on the contact signatures between them [START_REF] Dudchenko | De novo assembly of the Aedes aegypti genome using Hi-C yields chromosome-length scaffolds[END_REF]. In the same way, Hi-C can be used to correct already publish genome assemblies.

Before Hi-C, long-read sequencing was the main option for resolving most assembly issues, but during this time it was still in development. MATE-pair sequencing was later designed to bridge gaps in shorter scale assemblies. However, with the development of 3C, which was very powerful in linking almost any contig and scaffold to a chromosome, MATE-pair sequencing was rapidly abandoned. With the emergence of 3C techniques came the development of tools that took advantage of the physical contacts experienced by a portion of DNA to determine its genomic position. The initial tools used clustering methods to scaffold genome data. Lachesis was an algorithm that clustered contigs into chromosome groups by using the contact probabilities form Hi-C data [START_REF] Burton | Chromosomescale scaffolding of de novo genome assemblies based on chromatin interactions[END_REF]. The program then orders and orients these scaffolds to form contiguous segments of DNA. There were other tools like GRAAL that ordered contigs based on probabilistic methods. GRAAL is another algorithm that takes contigs and breaks them into restriction fragments bins, then used Hi-C data to orient and reorient the bins to make contiguous

Hierarchical Genome Organization

The folding of genomes requires a hierarchy of organizational structures ranging from chromosome territories to DNA loops. Historically, microscopy approaches allowed us to study the structure and dynamics of this organization. In the past 20 years, with the development of 3C

technologies and methods, we now have an additional approach that permits us to study genome conformation by measuring the frequency of physical contacts between different genomic loci.

Here I will describe different levels of organization and what we have learned by studying them in different contexts and in different organisms.

Chromosome Territories Across Species

Chromosomes are often thought of as having a condensed and rod-like shape we know from most illustrations. In fact, this is how they were described in initial observations. However, this is the form chromosomes take on during cell division and we now know that they exist in a different conformation during interphase. During interphase, chromosomes decondense and for decades they were impossible to distinguish from one another in this state, especially in mammals. Before any direct observations could be made two different models were proposed to explain the organization of the decondensed state during interphase. As mentioned in 1.1, in 1885 Carl Rabl proposed the chromosome territory model were chromosomes occupied specific areas of the nucleus only overlapping with their neighbors in interphase (Rabl, 1885). A second, competing model colloquially known as the "spaghetti" model proposed that during interphase DNA fibers of uncondensed chromosomes spread throughout the nucleus, intermingling with those of other chromosomes. This was the widely accepted model during the 1970's and 1980's [START_REF] Cremer | Chromosome Territories[END_REF]. It was in 1982 that Thomas and Christoph Cremer provided experimental evidence supporting the chromosome territory theory. By using a laser to induce DNA damage the periphery, similar to what is seen in mammals [START_REF] Marshall | Specific interactions of chromatin with the nuclear envelope: positional determination within the nucleus in Drosophila melanogaster[END_REF]Rosin et al., 2018).

Caenorhabditis elegans

Whole-chromosome oligopaint FISH has been used to visualize all six chromosomes of Caenorhabditis elegans and show that they also occupy distinct, non-overlapping territories (Fields et al., 2019). Studies have shown that a possible factor contributing to territory formation is the perinuclear anchoring of chromosome arms via CEC-4, a H3K9me binding nuclear envelope protein. This anchoring results in C. elegans chromosomes taking on a tripartite conformation :

Left-arm, center, right-arm [START_REF] Bian | Histone H3K9 methylation promotes formation of genome compartments in Caenorhabditis elegans via chromosome compaction and perinuclear anchoring[END_REF][START_REF] Sawh | Lamina-Dependent Stretching and Unconventional Chromosome Compartments in Early C. elegans Embryos[END_REF].

Chromosome Compartments

Looking within chromosome territories we find that chromatin is spatially separated into compartments and sub-compartments. Compartments are made up of self-associating regions of chromatin that make long-range interactions with compartments of the same type and little to no interactions with compartments of different types. The compartment type can generally be defined by epigenetic and physical characteristics. Studies have shown that compartments can be further divided into sub-compartments that are composed of more specific epigenetic factors. In this section I will describe observations mainly seen in humans. For compartment descriptions in other organisms see 3.1.2.

Establishment of A/B Compartments

Originally it was stated that the human genome was spatially segregated into two genome-wide compartments, which were labelled A and B. The A compartment was made up of active and more open chromatin. The B compartment was composed of more closed chromatin. As mentioned in 3.1.1, the first signs of chromosome compartmentalization came from the plaid pattern created by the normalized Hi-C interactions in intrachromosomal contact maps.

Converting the contact map into a Pearson correlation map further strengthened this pattern and made it easier to distinguish between the two compartments. In order to define precise locations for the different regions that make up each compartment, PCA was used to produce a twodimensional track based on the three-dimensional data. The alternating positive and negative signal of the eigenvector clearly corresponded to compartment interaction signals in the plaid pattern of most chromosomes [START_REF] Lieberman-Aiden | Comprehensive Mapping of Long-Range Interactions Reveals Folding Principles of the Human Genome[END_REF]. This relationship justified the use of a principal component eigenvector to define compartments in Hi-C data, at least preliminarily. profiling, but no epigenetic description was provided for this compartment. However, an interesting observation that was made was that interactions between B compartments were consistently higher than interactions between A compartments. From this it was suggested that compartment B is more densely packed than A [START_REF] Lieberman-Aiden | Comprehensive Mapping of Long-Range Interactions Reveals Folding Principles of the Human Genome[END_REF].

Over time the definitions of A and B compartments have changed slightly. With improvements in the Hi-C protocol allowing for higher resolution, and integration of several more genetic and epigenetic data sets, A and B compartments became more clearly defined. In addition to active epigenetic marks, the new data showed that A compartments are also associated to early replication timing regions, which is consistent with how it has been described before. The new data also showed that the B compartment is correlated with several inactive chromatin marks, late replication timing regions, lamina and nucleolus associated domains [START_REF] Rao | A 3D Map of the Human Genome at Kilobase Resolution Reveals Principles of Chromatin Looping[END_REF]. The higher resolution Hi-C data also revealed new contact patterns that could not previously be seen at lower resolution. Analysis of the contacts within each compartment showed that these patterns can actually be define by sub-compartments, which will be discussed in the next section.

Identification of Sub-Compartments

Originally, the A and B compartments were identified and defined by studying long-range contact patterns on Hi-C data at 1 Mbp resolution in human cell lines. New Hi-C studies were able to achieve 25 kb resolution. At this resolution, long-range contact patterns provided evidence for six sub-compartments within the two previously defined compartments (Figure 12). Of these six sub-compartments, two of them were in the A compartment and four of them were in the B compartment [START_REF] Rao | A 3D Map of the Human Genome at Kilobase Resolution Reveals Principles of Chromatin Looping[END_REF].

2012; [START_REF] Nora | Spatial partitioning of the regulatory landscape of the X-inactivation centre[END_REF]. It was also shown that TAD positioning remains conserved between different cells types, during cell differentiation and, for syntenic regions, between different mammals [START_REF] Dixon | Topological domains in mammalian genomes identified by analysis of chromatin interactions[END_REF][START_REF] Nora | Spatial partitioning of the regulatory landscape of the X-inactivation centre[END_REF]. However, perturbing the boundaries between neighboring TADs exhibited an alteration in TAD structure [START_REF] Nora | Spatial partitioning of the regulatory landscape of the X-inactivation centre[END_REF].

When looking at the cis-regulatory landscape of the X chromosome inactivation center in mice (Xic), it was shown that the Xist-Tsix-Xite region was located between two TADs and the deletion of this region resulted in an increase of contacts between them. The overall structure of the two TADs remained the same, however this increase in contacts demonstrated that key players behind TAD separation lie at their boundaries [START_REF] Nora | Spatial partitioning of the regulatory landscape of the X-inactivation centre[END_REF]. Aside from this perturbation study, it was also shown that CCCTC-binding factor (CTCF) was found at TAD boundaries, however, at the time it was not recognized to play a major role in TAD formation. This was because only a small portion of CTCF binding sites were found at TAD boundaries and the rest were found within TADs

Later studies showed that CTCF plays a much bigger role in TAD formation than previously seen.

This was likely missed in earlier studies due to a lack of resolution and a deeper inspection of CTCF binding site orientation. In human cell lines, over 9,448 peaks corresponding to chromatin loops were identified. These peaks are associated to 12,903 peak loci and 86% of these loci were shown to be bound by CTCF. Furthermore, it was also shown that CTCF binding sites located at peak loci are found in convergent orientation 92% of the time and that 39% of TADs were composed of peaks with this convergent orientation [START_REF] Rao | A 3D Map of the Human Genome at Kilobase Resolution Reveals Principles of Chromatin Looping[END_REF].

Mutating specific CTCF binding sites at TAD boundaries alters the boundary location, while mutating the same type of binding sites between two TAD leads to the TADs merging [START_REF] Sanborn | Chromatin extrusion explains key features of loop and domain formation in wild-type and engineered genomes[END_REF]. Depletion of CTCF genome-wide leads to a loss of TAD insulation [START_REF] Nora | Targeted Degradation of CTCF Decouples Local Insulation of Chromosome Domains from Genomic Compartmentalization[END_REF]. CTCF-associated TAD boundary disruptions have also been shown to lead to gene misregulation that can result in pathogenic phenotypes, different forms of cancer and alterations in cell state [START_REF] Flavahan | Insulator dysfunction and oncogene activation in IDH mutant gliomas[END_REF][START_REF] Hnisz | Activation of proto-oncogenes by disruption of chromosome neighborhoods[END_REF][START_REF] Lupiáñez | Disruptions of Topological Chromatin Domains Cause Pathogenic Rewiring of Gene-Enhancer Interactions[END_REF][START_REF] Narendra | CTCF establishes discrete functional chromatin domains at the Hox clusters during differentiation[END_REF][START_REF] Van Bemmel | The bipartite TAD organization of the X-inactivation center ensures opposing developmental regulation of Tsix and Xist[END_REF]. All of these obsevations suggest that CTCF is crucial to TAD formation in mammals.

CTCF is one of the main TAD boundary insulators in mammals, but does not seem to play a similar role in other organisms like D. melanogaster [START_REF] Cubeñas-Potts | Different enhancer classes in Drosophila bind distinct architectural proteins and mediate unique chromatin interactions and 3D architecture[END_REF][START_REF] Rowley | Evolutionarily Conserved Principles Predict 3D Chromatin Organization[END_REF][START_REF] Ulianov | Active chromatin and transcription play a key role in chromosome partitioning into topologically associating domains[END_REF]. TAD boundaries in D. melanogaster have been shown to be demarcated by BEAF-32 either paired with CP190 or Chromator and only marginally enriched in CTCF [START_REF] Wang | Sub-kb Hi-C in D . melanogaster reveals conserved characteristics of TADs between insect and mammalian cells[END_REF]. Early studies in D. melanogaster also showed that TAD formation was highly linked with epigenetic and transcriptional state, contrary to what was seen in humans [START_REF] Rowley | Evolutionarily Conserved Principles Predict 3D Chromatin Organization[END_REF][START_REF] Sexton | Three-Dimensional Folding and Functional Organization Principles of the Drosophila Genome[END_REF][START_REF] Ulianov | Active chromatin and transcription play a key role in chromosome partitioning into topologically associating domains[END_REF][START_REF] Wang | Sub-kb Hi-C in D . melanogaster reveals conserved characteristics of TADs between insect and mammalian cells[END_REF]. A more recent study showed that condensin II and cohesin play counteracting roles in TAD formation in active chromatin [START_REF] Rowley | Condensin II Counteracts Cohesin and RNA Polymerase II in the Establishment of 3D Chromatin Organization[END_REF].

Architectural Proteins

For decades it has been shown that insulators play a large role in genome organization. Here I will discuss the insulator proteins and insulator complexes that have been shown to be involve in genome organization in both mammals and D. melanogaster. Since the onset of their study, structural maintenance of chromosome (SMC) complexes have been known to play a role in genome organization. In this section I will also discuss the role of two SMC complexes: cohesin and condensin II.

Insulator Proteins

DNA-protein complexes that play roles in either blocking enhancer-promotor interactions or inhibiting the spread of epigenetic marks are known as insulators [START_REF] Yang | Chromatin Insulators: A Role in Nuclear Organization and Gene Expression[END_REF]. The structure of insulator complexes usually involves one DNA-binding protein that is accompanied by other proteins, which determine the insulator type and its function. There are a variety of insulator proteins and protein complexes in mammals and, mainly, in D. melanogaster (Figure 14).

Here I will outline some of the major insulators that have been studied.

CTCF

Before being recognized as one of the major players in genome architecture, and especially in the TAD formation, CTCF has been caraterized with several other functions. CTCF serves many purposes in the genome and is essential for the viability of mammalian cells (11,19,[START_REF] Albert | Whole-chromosome paints in maize reveal rearrangements, nuclear domains, and chromosomal relationships[END_REF]. As a transcription factor it has been shown to have a variety of roles in both repressing [START_REF] Lobanenkov | A novel sequencespecific DNA binding protein which interacts with three regularly spaced direct repeats of the CCCTC-motif in the 50-flanking sequence of the chicken c-myc gene[END_REF] and activating [START_REF] Filippova | An exceptionally conserved transcriptional repressor, CTCF, employs different combinations of zinc fingers to bind diverged promoter sequences of avian and mammalian c-myc oncogenes[END_REF][START_REF] Klenova | CTCF, a conserved nuclear factor required for optimal transcriptional activity of the chicken c-myc gene, is an 11-Zn-finger protein differentially expressed in multiple forms[END_REF] transcription. It has also been shown to have insulator activity and was one of the first proteins to be shown to be involved in chromatin looping between binding sites [START_REF] Handoko | CTCFmediated functional chromatin interactome in pluripotent cells[END_REF][START_REF] Splinter | CTCF mediates long-range chromatin looping and local histone modification in the β-globin locus[END_REF]. Initially, CTCF was known for its insulator roles at the b-globin and H19/Igf2 loci. In chicken, CTCF was found to be responsible for the insulator activity of the 5'HS4 site in the b-globin that separates it from neighboring chromatin [START_REF] Bell | The Protein CTCF Is Required for the Enhancer Blocking Activity of Vertebrate Insulators[END_REF]. CTCF is also involved in the b-globin locus of mammals [START_REF] Bulger | A complex chromatin landscape revealed by patterns of nuclease sensitivity and histone modification within the mouse beta-globin locus[END_REF][START_REF] Farrell | Conserved CTCF insulator elements flank the mouse and human beta-globin loci[END_REF], but here, it is found to encompass the locus in a large chromatin loop, essentially promoting spatial interactions within it [START_REF] Holwerda | CTCF: the protein, the binding partners, the binding sites and their chromatin loops[END_REF][START_REF] Palstra | The betaglobin nuclear compartment in development and erythroid differentiation[END_REF][START_REF] Splinter | CTCF mediates long-range chromatin looping and local histone modification in the β-globin locus[END_REF]. At the H19/Igf2 locus, CTCF is responsible for the controlled activation of the H19 gene on the maternal allele and Igf2 on the paternal allele by binding to the imprinting control region (ICR) in a methylation dependent manner. By binding to the unmethylated ICR of the maternal allele it insulates the region between the H19 enhancers and the Igf2 gene so they cannot reach. On the paternal allele the ICR is methylated and therefore CTCF cannot bind [START_REF] Bell | Methylation of a CTCF-dependent boundary controls imprinted expression of the Igf2 gene[END_REF][START_REF] Hark | CTCF mediates methylation-sensitive enhancer-blocking activity at the H19/Igf2 locus[END_REF][START_REF] Kurukuti | CTCF binding at the H19 imprinting control region mediates maternally inherited higher-order chromatin conformation to restrict enhancer access to Igf2[END_REF][START_REF] Murrell | Interaction between differentially methylated regions partitions the imprinted genes Igf2 and H19 into parent-specific chromatin loops[END_REF]. These early studies showed that CTCF can interfere with promoter-enhancer communication and form chromatin loops between its binding sites [START_REF] Holwerda | CTCF: the protein, the binding partners, the binding sites and their chromatin loops[END_REF]. Revisiting these roles with Hi-C data has shown that CTCF's insulator roles play a role in TAD boundary formation in mammals and, together with cohesin, is responsible for local genome organization. For more on the role CTCF plays in TAD boundaries, see 4.3. For more on the role of CTCF with cohesin, see 5.2.1. CTCF is also found in D. melanogaster, but has been shown to not play a similar in organizing the genome as in mammals [START_REF] Cubeñas-Potts | Different enhancer classes in Drosophila bind distinct architectural proteins and mediate unique chromatin interactions and 3D architecture[END_REF][START_REF] Rowley | Evolutionarily Conserved Principles Predict 3D Chromatin Organization[END_REF][START_REF] Ulianov | Active chromatin and transcription play a key role in chromosome partitioning into topologically associating domains[END_REF].

There are also far less CTCF binding sites in D. melanogaster than in humans [START_REF] Pugacheva | CTCF mediates chromatin looping via N-terminal domain-dependent cohesin retention[END_REF]. However, a recent study has shown that although CTCF is dispensable for D. melanogaster early development [START_REF] Gambetta | The Insulator Protein CTCF Is Required for Correct Hox Gene Expression, but Not for Embryonic Development in Drosophila[END_REF], its expression is critical for the nervous system.

The same study also showed that CTCF does in fact contribute to contact boundary formation, but only a small fraction (<10%) [START_REF] Kaushal | CTCF loss has limited effects on global genome architecture in Drosophila despite critical regulatory functions[END_REF]. Many contact domain boundaries in D.

melanogaster are also occupied by other insulator proteins, which I will describe in the following sections.

BEAF-32/CP190

D. melanogaster boundary element-associated factors (BEAF) can be found as two different DNAbinding proteins, BEAF-32A and BEAF-32B and were discovered to interact with scs' boundary element sequences on polytene chromosomes [START_REF] Zhao | Visualization of chromosomal domains with boundary element-associated factor BEAF-32[END_REF]. Perturbations of BEAF-32

were shown to interfere with polytene chromosome structure and lead to developmental defects [START_REF] Gilbert | The Drosophila Boundary Element-Associated Factors BEAF-32A and BEAF-32B Affect Chromatin Structure[END_REF]. BEAF was found to colocalize with centrosomal protein 190 (CP190) and together mediate long-range chromatin interactions [START_REF] Vogelmann | Chromatin Insulator Factors Involved in Long-Range DNA Interactions and Their Role in the Folding of the Drosophila Genome[END_REF]. They have also other instances of gene activation, happen via chromatin conformation alteration in the form of DNA looping [START_REF] Kurukuti | CTCF binding at the H19 imprinting control region mediates maternally inherited higher-order chromatin conformation to restrict enhancer access to Igf2[END_REF][START_REF] Majumder | The insulator factor CTCF controls MHC class II gene expression and is required for the formation of long-distance chromatin interactions[END_REF][START_REF] Splinter | CTCF mediates long-range chromatin looping and local histone modification in the β-globin locus[END_REF]. For years, the mechanism behind CTCF's long-range interactions was not known. This was until it was shown that, in humans, cohesin binding sites colocalize with CTCF binding sites via ChIP-chip and ChIP-qPCR [START_REF] Wendt | Cohesin mediates transcriptional insulation by CCCTC-binding factor[END_REF]. The same study reported that cohesin was required for CTCF's insulator function. Now that a relationship between cohesin and CTCF was establish, future studies looked to investigate this relationship in regards to the long-range chromosomal interactions both were previously reported to make. In a study looking specifically at this relationship, it was found that cohesin is required for CTCF to maintain long-range interactions at the IFNG locus in human CCR5 + T H 1 cells [START_REF] Hadjur | Cohesins form chromosomal cis-interactions at the developmentally regulated IFNG locus[END_REF].

There are many studies that have establish cohesin's non-canonical roles outside of sister chromatid cohesion. These roles mainly pertain to enhancer-promotor interactions and gene regulation. It was shown that the mechanism behind these roles involves cohesin's ability to enclose two separate chromatin fibers to form loops. This was demonstrated early on with 3C, but with the development of Hi-C in 2009, we were able to detect how this function effects genome organization at a greater 3D scale. In a large Hi-C study involving nine human cell lines, over 10,000 loop signatures or "peaks" were identified. Out of these peaks, 86% of them were reported to be bound by CTCF and cohesin subunit, RAD21, although this was consistent with what was previously reported about the relationship between CTCF, cohesin and loop formation, what was new was that these proteins were also concentrated at TAD boundaries, something that could only be seen with Hi-C [START_REF] Rao | A 3D Map of the Human Genome at Kilobase Resolution Reveals Principles of Chromatin Looping[END_REF].

In order to study the effects of cohesin on spatial genome architecture in humans, studies have perturbed cohesin, typically by depleting one of its core subunits. In the case of depleting RAD21 (Scc1), cohesin is rendered unable to associate with DNA. Via Hi-C, it was shown that this removal of cohesin leads to the loss of all looping events and the loss of all contact domains that were formed by CTCF-cohesin DNA loops (Figure 16). It also resulted in stronger genome compartmentalization. This loss of cohesin did not affect CTCF localization, nor did it affect histone modification patterns [START_REF] Rao | Cohesin Loss Eliminates All Loop Domains[END_REF][START_REF] Wutz | Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins[END_REF][START_REF] Zuin | Cohesin and CTCF differentially affect chromatin architecture and gene expression in human cells[END_REF]. Two other factors that can influence the effects of cohesin are its loader, NIPBL (Scc2), and its remover, WAPL. Cohesin is loaded onto DNA by NIPBL-MAU2 (Scc2/Scc4) complex, often just referred to as NIPBL [START_REF] Ciosk | Cohesin's Binding to Chromosomes Depends on a Separate Complex Consisting of Scc2 and Scc4 Proteins[END_REF]. Depleting NIPBL reduces the amount of cohesin loaded onto DNA and, like depleting RAD21, reduces the amount of CTCF-cohesin loops and strengthens the segregation between compartments while not affecting CTCF binding [START_REF] Haarhuis | The Cohesin Release Factor WAPL Restricts Chromatin Loop Extension[END_REF].

The factor responsible for removing cohesin from DNA is it's antagonist, WAPL [START_REF] Gandhi | Human Wapl Is a Cohesin-Binding Protein that Promotes Sister-Chromatid Resolution in Mitotic Prophase[END_REF][START_REF] Kueng | Wapl Controls the Dynamic Association of Cohesin with Chromatin[END_REF]. Depletion of WAPL has practically the opposite effect as depleting NIPBL, it results in longer CTCF-cohesin loops and a reduction of compartmental interactions [START_REF] Haarhuis | The Cohesin Release Factor WAPL Restricts Chromatin Loop Extension[END_REF][START_REF] Rao | Cohesin Loss Eliminates All Loop Domains[END_REF][START_REF] Wutz | Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins[END_REF].

In mouse, another mammal, the loss of cohesin is very similar to what we see in human. Depletion of RAD21 results in loss of contact domains and strengthens the segregation between active and inactive chromatin [START_REF] Gassler | A mechanism of cohesin-dependent loop extrusion organizes zygotic genome architecture[END_REF][START_REF] Seitan | Cohesin-based chromatin interactions enable regulated gene expression within preexisting architectural compartments[END_REF]. Loss of NIPBL also reduces the amount of cohesin loaded onto DNA, thus leading to the disappearance of contact domains and CTCF-cohesin loops along with the enhancement of compartmentalization, all without altering CTCF binding [START_REF] Schwarzer | Two independent modes of chromatin organization revealed by cohesin removal[END_REF]. Deletion of WAPL strengthens contact domains and loops, opposite to the effect of NIPBL loss [START_REF] Gassler | A mechanism of cohesin-dependent loop extrusion organizes zygotic genome architecture[END_REF].

Moving away from mammals, we see that cohesin also plays a role in genome organization and gene regulation, but differently from what we know in human and mouse. In D. melanogaster, cohesin has not been shown to associate, with CTCF. In fact, it has been shown to associate with RNA PolII and play a role in mediating active chromatin organization [START_REF] Misulovin | Association of cohesin and Nipped-B with transcriptionally active regions of the Drosophila melanogaster genome[END_REF][START_REF] Rowley | Condensin II Counteracts Cohesin and RNA Polymerase II in the Establishment of 3D Chromatin Organization[END_REF]. Cohesin loss via RAD21 knockdown reveals a reduction of intragenic contacts. Additionally, it also results in a decrease in homotypic A-compartment interactions, which is different from the effect cohesin has at the compartment level in mammals [START_REF] Rowley | Condensin II Counteracts Cohesin and RNA Polymerase II in the Establishment of 3D Chromatin Organization[END_REF]. The D. melanogaster ortholog of NIPBL, Nipped-B, is also known for loading cohesin on to DNA. Knocking down Nipped-B also leads to a reduction of cohesin binding, sister chromatid cohesion defects and the misregulation of certain genes [START_REF] Dorsett | Effects of sister chromatid cohesion proteins on cut gene expression during wing development in Drosophila[END_REF][START_REF] Gause | Dosage-Sensitive Regulation of Cohesin Chromosome Binding and Dynamics by Nipped-B, Pds5, and Wapl[END_REF][START_REF] Misulovin | Association of cohesin and Nipped-B with transcriptionally active regions of the Drosophila melanogaster genome[END_REF][START_REF] Rollins | Drosophila Nipped-B Protein Supports Sister Chromatid Cohesion and Opposes the Stromalin/Scc3 Cohesion Factor To Facilitate Long-Range Activation of the cut Gene[END_REF], but the effects on genome architecture via Hi-C have not been shown. Also similar to mammals, Wapl is a cohesin antagonist and removes it from DNA [START_REF] Gause | Dosage-Sensitive Regulation of Cohesin Chromosome Binding and Dynamics by Nipped-B, Pds5, and Wapl[END_REF]. Mutations in Wapl were shown to prevent sister chromatid separation and increase cohesin stability on chromatin [START_REF] Cunningham | Wapl antagonizes cohesin binding and promotes Polycomb-group silencing in Drosophila[END_REF][START_REF] Gause | Dosage-Sensitive Regulation of Cohesin Chromosome Binding and Dynamics by Nipped-B, Pds5, and Wapl[END_REF][START_REF] Vernì | Genetic and Molecular Analysis of wings apart-like (wapl), a Gene Controlling Heterochromatin Organization in Drosophila melanogaster[END_REF]. However, just like Nipped-B, the effects of Wapl depletions on genome architecture via Hi-C have not been shown in D. melanogaster.

In C. elegans, like in other organisms, the depletion of cohesin subunits lead to defects in cell division [START_REF] Chambeyron | Chromatin decondensation is sufficient to alter nuclear organization in embryonic stem cells[END_REF][START_REF] Mito | Distinct Developmental Function of Two Caenorhabditis elegans Homologs of the Cohesin Subunit Scc1/Rad21[END_REF][START_REF] Wang | Caenorhabditis elegans EVL-14/PDS-5 and SCC-3 Are Essential for Sister Chromatid Cohesion in Meiosis and Mitosis[END_REF]. However, there are four different homologs for the Rad21 subunit (COH-1, SCC-1/COH-2, COH-3, and REC-8) and depletion of different ones can lead to different defects [START_REF] Das | Condensin I folds the C. elegans genome[END_REF][START_REF] Mito | Distinct Developmental Function of Two Caenorhabditis elegans Homologs of the Cohesin Subunit Scc1/Rad21[END_REF]. As far as the role of cohesin in genome organization goes, it has been reported that after cohesin depletion via SCC-1 cleavage only a mild loss of short distance contacts can be seen via Hi-C, unlike in mammals and D. melanogaster. The same study shows that it is in fact condensin I that is responsible for large scale genome folding in C. elegans [START_REF] Das | Condensin I folds the C. elegans genome[END_REF]. This is further discussed in 5.1.2.1. 5.2.2. Condensins In most eukaryotes, two classes of condensins exist, condensin I and II. They differ only in their non-SMC subunits, and they mainly have nonoverlapping functions [START_REF] Ono | Differential Contributions of Condensin I and Condensin II to Mitotic Chromosome Architecture in Vertebrate Cells[END_REF]. In order to properly understand the role of condensins in genome organization, it is important to specify between the two. In vertebrates, although both condensins play essential roles in mitosis, it occurs at different parts of the cell cycle. Condensin I is the cytoplasmic condensin that only gains access to chromatin after nuclear envelope breakdown and contributes to chromosome condensation between prometaphase and telophase [START_REF] Ono | Spatial and Temporal Regulation of Condensins I and II in Mitotic Chromosome Assembly in Human Cells[END_REF]. Upon entry to the nucleus, condensin I contributes to the dissociation of cohesin from chromosome arms and chromosome shortening [START_REF] Hirota | Distinct functions of condensin I and II in mitotic chromosome assembly[END_REF]. Studies have shown that condensin I may not play a crucial role in chromosome compaction during cell division, but is more-so tasked with maintaining already condensed chromatin in a rigid state to facilitate spindle attachment [START_REF] Gerlich | Condensin I Stabilizes Chromosomes Mechanically through a Dynamic Interaction in Live Cells[END_REF]. Other studies support this claim and have shown that condensin I may only act in maintaining condensed chromatin loops, rather than playing a role in the initial condensation [START_REF] Green | Contrasting roles of condensin I and condensin II in mitotic chromosome formation[END_REF]. A recent study has shown more specifically, that condensin I is responsible for the formation of short, 60-80 kb, inner loops in prometaphase cells [START_REF] Gibcus | A pathway for mitotic chromosome formation[END_REF]. These studies give us insight into the role of condensin I during cell division, however, there have not been any major studies showing a role for it in organizing the genome during interphase, which is likely due to it being absent in the nucleus during this stage. Although, some studies have shown that some condensin I subunits are associated to gene regulation in interphase [START_REF] Li | Condensin I and II Complexes License Full Estrogen Receptor α-Dependent Enhancer Activation[END_REF][START_REF] Zhang | Condensin I and II behaviour in interphase nuclei and cells undergoing premature chromosome condensation[END_REF].

Condensin I also plays a vital role in D. melanogaster mitosis, being necessary for not only proper formation and segregation of mitotic chromosomes but also for maintaining the integrity of centromere heterochromatin [START_REF] Dej | Mutations in the Drosophila Condensin Subunit dCAP-G: Defining the Role of Condensin for Chromosome Condensation in Mitosis and Gene Expression in Interphase[END_REF][START_REF] Oliveira | The Condensin I Subunit Barren/CAP-H Is Essential for the Structural Integrity of Centromeric Heterochromatin during Mitosis[END_REF][START_REF] Savvidou | Drosophila CAP-D2 is required for condensin complex stability and resolution of sister chromatids[END_REF]. It has also been shown to be necessary for topoisomerase II localization to sister chromatids for proper condensation [START_REF] Coelho | Condensin-dependent localisation of topoisomerase II to an axial chromosomal structure is required for sister chromatid resolution during mitosis[END_REF]. There have not been any studies showing a role for condensin I in genome organization in interphase, which is likely because it is condensin II that has the significant role there, which will be discussed in the next section.

There have been extensive studies showing that condensins are crucial for chromosome dynamics and genome organization in interphase in C. elegans. The X-chromosome specific condensin I, or condensin I DC , has been shown to be crucial for dosage compensation. The dosage compensated X chromosome also exhibits contact domains similar to what is seen on mammalian chromosomes, which makes it distinct from the autosomes [START_REF] Crane | Condensin-driven remodelling of X chromosome topology during dosage compensation[END_REF][START_REF] Das | Condensin I folds the C. elegans genome[END_REF][START_REF] Rowley | Analysis of Hi-C data using SIP effectively identifies loops in organisms from C. elegans to mammals[END_REF]. This distinct structure is thought to be linked with the chromosome-wide reduction in gene expression that contributes to balancing expression levels between hermaphrodites (XX) and males (X0) [START_REF] Crane | Condensin-driven remodelling of X chromosome topology during dosage compensation[END_REF][START_REF] Jans | A condensin-like dosage compensation complex acts at a distance to control expression throughout the genome[END_REF].Condensin I has also been shown to be responsible for long-range genome folding on C. elegans autosomes. Loss of condensin I leads to the loss of contact domains, reinforcement of epigenetic compartments and genome-wide decompaction [START_REF] Das | Condensin I folds the C. elegans genome[END_REF]. Similar to the loss of cohesin in mammals, which was described in the previous section.

Condensin II

In vertebrates, condensin II can be found in the nucleus of a cell in interphase and concentrates on chromosomes in prophase to participate in the early condensation stage of chromosomes prior to cell division [START_REF] Ono | Differential Contributions of Condensin I and Condensin II to Mitotic Chromosome Architecture in Vertebrate Cells[END_REF]. Condensin II has been shown to mediate long-range DNA interactions and establish the initial chromatin axis on condensed chromosomes and localize at the inner kinetochore plate. Depleting condensin II leads to largely decondensed chromosome that contain anaphase bridges, and lose longitudinal shortening and helical winding [START_REF] Gibcus | A pathway for mitotic chromosome formation[END_REF][START_REF] Green | Contrasting roles of condensin I and condensin II in mitotic chromosome formation[END_REF][START_REF] Ono | Spatial and Temporal Regulation of Condensins I and II in Mitotic Chromosome Assembly in Human Cells[END_REF]. Given that condensin II is located in the nucleus during interphase, there has been a study to see if it contributes to genome organization. What was reported was that condensin II does not seem to play a role in genome organization in interphase when studied via Hi-C. Depletion of the condensin II subunit CapH2, which is essential for its function, resulted in no changes at any level of genome organization or gene expression when compared to the wild-type [START_REF] Abdennur | Condensin II inactivation in interphase does not affect chromatin folding or gene expression[END_REF].

Centromere Impact on Genome Organization

Centromeres are mostly known for the crucial functions they play during mitosis and meiosis, however that is not their only role in the genome as centromeres are not inert during interphase.

In this section I will describe the impact centromeres have on genome organization in interphase.

Basic Characteristics of Centromeres

As mentioned in 1.2, chromosomes were first observed with a microscope and during these initial observations the centromere was identified and initially called the "primary constriction" that spindle fibers attached to in order to ensure chromosome segregation (Flemming, 1882). Since then centromeres have lied at the center of studies revolving around cell division.

Centromeres are regions of specialized chromatin found on each chromosome. The DNA at centromeres is often characterized as AT-rich, gene-poor and transcribed at low levels [START_REF] Talbert | What makes a centromere?[END_REF]. In most organisms the centormeres are epigenetically defined by the presence of centromeric nucleosomes. Centromeric nucleosomes are specialized in that they contain a histone-H3 variant known as CenH3, or CENP-A in mammals [START_REF] Earnshaw | Identification of a family of human centromere proteins using autoimmune sera from patients with scleroderma[END_REF][START_REF] Palmer | Purification of the centromere-specific protein CENP-A and demonstration that it is a distinctive histone[END_REF]. For certain varieties of centromeres, the DNA can also be composed of specific centromere sequences and/or tandem repeats [START_REF] Steiner | Diversity in the organization of centromeric chromatin[END_REF]. One example of a centromere specific sequence are CENP-B boxes, which can be found at human centromeres. CENP-B boxes are 17-bp long sequences that are binding sites for CENP-B, which is a protein that helps position centromeric nucleosomes [START_REF] Masumoto | The role of CENP-B and alpha-satellite DNA: de novo assembly and epigenetic maintenance of human centromeres[END_REF]. Most centromeres can be characterized by the presence of centromeric nucleosomes, specific sequences and repetitive elements, however, exceptions can be found for each one of these features within the large centromere diversity among eukaryotes.

The primary role of a centromere is to provide the foundation for kinetochore assembly. The kinetochore is a macromolecular protein complex and is responsible for mediating the attachement of spindle fibers to centromeres during cell division [START_REF] Fukagawa | The Centromere: Chromatin Foundation for the Kinetochore Machinery[END_REF].

The proteins making up the kinetochore can be subdivided into two major groups: those making up the inner-kinetochore and those making up the outer-kinetochore. The inner-kinetochore is the part of the kinetochore that is in contact and most proximal to the centromere. Within the inner-kinetochore is a set of proteins known as the constitutive centromere-associated network, or CCAN, which is present at the centromere throughout the cell cycle. During cell division, the evolutionary (Drinnenberg et al., 2014;[START_REF] Melters | Comparative analysis of tandem repeats from hundreds of species reveals unique insights into centromere evolution[END_REF]Melters et al., , 2012)), centromere and kinetochore studies (Cortes-Silva et al., 2020;[START_REF] Gassmann | An inverse relationship to germline transcription defines centromeric chromatin in C. elegans[END_REF][START_REF] Hofstatter | Repeat-based holocentromeres influence genome architecture and karyotype evolution[END_REF][START_REF] Marques | Holocentromeres in Rhynchospora are associated with genomewide centromere-specific repeat arrays interspersed among euchromatin[END_REF][START_REF] Senaratne | Formation of the CenH3-Deficient Holocentromere in Lepidoptera Avoids Active Chromatin[END_REF][START_REF] Steiner | Diversity in the organization of centromeric chromatin[END_REF].

Holocentric organisms have been identified among insects, flowing plants, green alga and most famously in nematodes (Drinnenberg et al., 2014;Melters et al., 2012;[START_REF] Zedek | Holocentric chromosomes: from tolerance to fragmentation to colonization of the land[END_REF].

In all cases they are found evolutionarily within monocentric groups, which supports the theory that holocentric organisms evolved from monocentric ancestors. There is currently no evidence to suggest a transition from holocentricity to monocentricity, which implies that holocentricity is a stable trait. Like monocentromeres, holocentromeres are diverse when it comes to the composition of their underlying DNA, as holocentromeres can not be soley defined by the presence of specific sequences or sequence repeats or even centromeric nucleosomes. Their unity comes in the structure they assume during cell division as they all follow the criteria previously mentioned in the first paragraph (Melters et al., 2012). It is these same criteria that sets monocentric chromosomes and holocentric chromosomes apart.

Several models have been proposed to explain how holocentricity emerged [START_REF] Senaratne | Mesoderm differentiation in vertebrate development and regenerative medicine by Fiona Wardle / Special Issue:Genomic and epigenetic stability and inheritance by[END_REF]. One such model is that centromeric satellite repeats may have spread across the length of the chromosome instead of remaining restricted to one site, effectively giving kinetochores the ability to assembly along the entire chromosome [START_REF] Senaratne | Mesoderm differentiation in vertebrate development and regenerative medicine by Fiona Wardle / Special Issue:Genomic and epigenetic stability and inheritance by[END_REF]. This model implies that kinetochores or CenH3 recognize specific sequence features, thus supporting a genetic definition of centromeres. Indeed, some holocentric plants assemble CenH3 and kinetochores on repetitive DNA (Rynchospora). Support for this has been found in the holocentric plant Luzula nivea [START_REF] Neumann | Stretching the Rules: Monocentric Chromosomes with Multiple Centromere Domains[END_REF]. Centromeric sites of other holocentric organisms are however not associated with repeats. Here a "loss of centromere identity" (i.e. disruption of the epigenetic loop that maintains CenH3 at centromeres) could have lead to the ectopic incorporation of CenH3 and thus the holocentric architecture [START_REF] Senaratne | Mesoderm differentiation in vertebrate development and regenerative medicine by Fiona Wardle / Special Issue:Genomic and epigenetic stability and inheritance by[END_REF]. Since holocentricity emerged independantly several times and holocentric architectures are diverse, it is possible that they all did not emerge in the same way.

Another difference that arises with this centromere organization is a hightened tolerance to DNA damage. For example, in the case of chromosomal fragmentation, a fragmented holocentric chromosome can still propogate to the next cell cycle entirely because the entire length has kinetochore attachement sites [START_REF] Zedek | Holocentric chromosomes: from tolerance to fragmentation to colonization of the land[END_REF]. In monocentric chromosomes, chromosomal fragments without a centromere are lost. Due to characteristics such as this and the lack of evidence for a holocentric to monocentric transition, it would appear that holocentricity does presents some advantages over monocentricity. Why we do not see more holocentricity occuring in evolution is a question still open in the field.

Thesis Motivation

As far as holocentric genome organization goes, there is only one well-studied example, C. elegans.

Studies have shown that in C. elegans, the transcriptional state of the chromatin not only plays a part in compartmentalization, but in interchromosomal interactions as well. Looking at the genome-wide contact map, the noticeable interactions are between the active chromosome centers and the inactive chromosome arms, which is mainly due to the anchoring of the inactive arms to the nuclear envelope by H3K9me-binding protein [START_REF] Sawh | Lamina-Dependent Stretching and Unconventional Chromosome Compartments in Early C. elegans Embryos[END_REF][START_REF] Bian | Histone H3K9 methylation promotes formation of genome compartments in Caenorhabditis elegans via chromosome compaction and perinuclear anchoring[END_REF].

Studies have also shown that, in C. elegans, condensin I is the main culprit responsible for spatial genome organization [START_REF] Das | Condensin I folds the C. elegans genome[END_REF]. Being that this is the only reference we have to holocentric genome organization; a wide gap of knowledge remains. In my thesis I explored another group of holocentric animals, insects, focusing on the silkworm, Bombyx mori.

Although B. mori and C. elegans are both holocentric, it is known that their holocentricity evolved independently of each other (Melters et al., 2012;Drinnenberg et al., 2014). The holocentric chromosomes of B. mori have been subjects of studies describing the linear organization and formation of their centromeres [START_REF] Senaratne | Formation of the CenH3-Deficient Holocentromere in Lepidoptera Avoids Active Chromatin[END_REF], along with the kinetochore formation on them (Cortes-Silva et al. 2020). However, the 3D organization of holocentric chromosomes in B.

mori has yet to be described.

The overall aim of my thesis was to provide a general description of the hierarchical genome organization of B. mori holocentric chromosomes and determine the factors that contribute to it.

In order to do this, I took advantage of two experimental systems available for this organism.

The first system is the p50 silkworm strain from which post-diapause embryos were used to perform Hi-C and ChIP-Seq experiments. We selected this strain because it was used to build the first B. mori genome assembly in 2004 (Mita et al., 2004;Xia et al., 2004). The assembly was later updated in 2008 (International Silkworm Genome Consortium, 2008) and in 2019 the genome was reassembled using deep sequencing of PacBio and Illumina reads (Kawamoto et al., 2019). Using this strain, we expect to get the best read-alignment results for both Hi-C and ChIP-Seq. This is crucial to perform accurate analysis and provide the best description of the genome as possible.

Aim I: Describe the different hierarchical levels of B. mori genome organization.

Part I: Analysis of interchromosomal contacts.

Result I: B. mori embryos exhibit strong chromosome territories.

Part II: Analysis of intrachromosomal contacts.

Result II: Compartment analysis reveals three genome-wide contact patterns.

Result III. The X-compartment is composed of gene-poor and highly compact, self-interacting domains.

The second system is BmN4-SID1, an RNAi competent cell line derived from B. mori ovaries [START_REF] Kobayashi | SID-1 protein of Caenorhabditis elegans mediates uptake of dsRNA into Bombyx cells[END_REF][START_REF] Maeda | Production of human α -interferon in silkworm using a baculovirus vector[END_REF]. This system is used to facilitate the use of molecular biology techniques such as introducing plasmid constructs via transfections and genomic perturbations via RNAi depletion of target mRNAs. This system is also useful because the cells can readily be grown and maintained. The BmN4-SID1 cell line provides experimental capabilities that are vital to determine the key players involved in organizing the B. mori genome.

Aim II: Investigate what factors contribute to B. mori genome organization.

Part III: RNAi depletion of cohesin and condensin II subunits.

Result IV: Cohesin and condensin II have counteracting roles in maintaining genome organization.

Result V: Centromere localization is not affected by cohesin or condensin II depletion.

Part IV: RNAi depletion of essential centromere protein.

Result VI: Centromere region perturbations do not impact short-range genome organization in interphase.

Results

compartment B, while the other ~50% of the genome is either compartment A (~30%), X (~16%) or unmapped regions (Figure 29C).

Next, I investigated how the contact frequency changes over increasing genomic distances for each compartment. For this, I looked at both homotypic and heterotypic contacts between all combinations of compartments A, B and X. What I found was that A-A, B-B and A-B P(s) curves confirm observations made in other genomes, namely that homotypic contacts are always stronger than heterotypic. Additionally, B-B contacts are stronger at short distances (<300 kb) while A-A contacts become stronger at longer distances. We then plotted X-X contacts together with X-A and X-B contacts. We confirm the pronounced short-range contacts within X that drop dramatically at a distance of around 300-500 kb to meet the range of A-B contact frequencies at distances of 1-2 Mbp. At all size ranges, X-X contacts are higher than X-A or X-B with X-A and X-B contact decay curves overlapping, which confirms the individuality of the X compartment (Figure 29D).

Lastly, having identified the regions of the genome pertaining to each compartment I wanted to know how the different compartments interact in the interchromosomal space. Using the same observed/expected data as in Result 1, I found that interchromosomal contacts mainly come from interactions between A compartments and not between B compartments (Figure 29E).

Surprisingly, interchromosomal contacts are also enriched for X compartments interactions, in proportions similar to A's (Figure 29E). The apparent lack of long-range contacts raised the question of where X domains are located with respect to their CT. To address this question, we collaborated with Leah Rosin from the NIH to label portions of single A, B and X compartment domains with oligopaint FISH probes. We found that X compartment domains are more likely to be located in the periphery of the CT compared to A or B compartment domains. It is thus possible that the preferential positioning of X compartment domains on the outside of CTs places them in a favorable location for interchromosomal contacts despite their high levels of compaction and very poor propension to make intrachromosomal long-range contacts (Supp. Figure 1).

Taken together, we conclude that B. mori chromosomes segregate into three genome-wide compartments. Reminiscent of other organisms, we have identified A and B compartments in B.

mori that represent epigenetically defined active and inactive regions of the genome. The newly identified compartment X exhibits mixed epigenetic features with only weak enrichments.

Compartment X is comprised of about one sixth of the genome and has distinct short-and longrange contact profiles.

Result III. The X-compartment is composed of gene-poor and highly compact selfinteracting domains.

In order to get a more general idea of the intrachromosomal interaction patterns in B. mori chromosomes I took a closer look at the genome-wide P(s) curve. The trend, or rate of contact decay, of the curve can be further studied by plotting it's derivative, or the change in slope of the curve as a function of genomic distance. Plotting the genome-wide P(s) curve together with its derivative shows that the rate of contact decay is slightly decreased between 10 kb and 50 kb and then increases between 50 kb and 300 kb (Figure 30A, top). This change in rate of contact decay appears in the derivative as a bump between 10 kb and 300 kb (Figure 30A, bottom). The initial decrease in rate of decay represented by a shallow slope indicates local compaction of chromatin and this region of the P(s) curve is often referred to as a "shoulder". The local chromatin compaction that creates this shoulder in the P(s) curve is thought to be created by loop extruding factors, mainly cohesin [START_REF] Fudenberg | Emerging Evidence of Chromosome Folding by Loop Extrusion[END_REF]. At larger genomic distances there is another, smaller shoulder that creates another bump in the derivative at ~3 Mbp. Although not shown experimentally in interphase, this shoulder is thought to be a result of higher-level chromatin folding by condensin during cell division [START_REF] Abdennur | Condensin II inactivation in interphase does not affect chromatin folding or gene expression[END_REF][START_REF] Gibcus | A pathway for mitotic chromosome formation[END_REF].

With this in mind, we then wondered whether levels of compaction or loop extrusion processes were different in each compartment. To address this question, I generated intradomain P(s) curves for each of the three compartments (Figure 30B, top). These plots revealed distinct rates of contact decay inside A, B and X. The P(s) curve of X domains consistently shows higher intradomain interaction frequencies compared to A and B. X domains thus appear to be more compact than A and B domains. Plotting the derivatives of intradomain P(s) curves for each compartment revealed a strong bump at ~40 kb for the X compartment. The P(s) curve derivatives for A and B also show bumps around 40 kb and another around ~80-100 kb but those are much lower compared to X. This indicates that the shoulder in the genome-wide P(s) curve (Figure 30A, top) and the bump at 50 kb in the derivative (Figure 30A, bottom) are mainly that it is higher than the interchromosomal mean of the WT. This effect could indicate a slight decompaction of the chromosome (or regions of the chromosome) that may lead to an increase of trans-chromosomal contacts. However, it is difficult to assess interchromosomal contact patterns in the cell line data due to extensive aneuploidy and rearrangements, which will be further addressed in Discussion and Future Directions. Along with the Scc1 RNAi I also performed an RNAi depletion of CapH2, condensin II's kleisin subunit, by treating BmN4-SID1 cells with CapH2 dsRNA for 3 days, I harvested the cells for Hi-C, ChIP-Seq, IF to measure mitotic index and also for RNA-Seq to verify the depletion (Figure 32). Condensin II is another SMC complex that has been shown to extrude loops of DNA [START_REF] Ganji | Real-time imaging of DNA loop extrusion by condensin[END_REF][START_REF] Terakawa | The condensin complex is a mechanochemical motor that translocates along DNA[END_REF]. Although it has been suggested to not play a role in interphase genome organization in mammals, it has been shown to counteract the role of cohesin in genome organization in D. melanogaster [START_REF] Rowley | Condensin II Counteracts Cohesin and RNA Polymerase II in the Establishment of 3D Chromatin Organization[END_REF]. With this in mind I was motivated to see if condensin II played a role in B. mori genome organization. Performing the same analyses as with Scc1 RNAi , I first made a visual inspection of the normalized contact maps for CapH2 RNAi and compared them to the WT condition. In this case, the contact domains seen in WT are better defined in CapH2 RNAi . The domains in CapH2 RNAi are easily distinguished from the interdomain regions (Figure 33A). Looking at the insulation scores for the plotted region you can see that the interdomain insulation scores of CapH2 RNAi are in the same range as that of WT, but the intradomain scores are higher. Comparing the insulation scores between CapH2 RNAi and WT at the genome-wide scale bares an increase in the range of insulation scores. This signifies that, overall, the intradomain contact frequency is higher in CapH2 RNAi than in WT, while the interdomain contact frequency remains mostly unchanged.

As with Scc1 RNAi I also calculated and plotted the P(s) curve for CapH2 RNAi . I found that at distances under 1 Mbp, CapH2 RNAi has a higher contact frequency than WT. At distances between 1 Mbp and 3 Mb the contact frequency between CapH2 RNAi and WT are similar and at distances greater than 3 Mbp CapH2 RNAi has a higher contact frequency than WT. I also looked at the effect of CapH2 RNAi on interchromosomal contacts and found that, like Scc1 RNAi , there is an overall higher frequency of contacts between chromosomes than in WT. Although it has been shown in D.

melanogaster that CapH2 depletion leads to increased intermixing between interphase chromosomes (Rosin et al., 2018), it is difficult to draw the same conclusion in this condition because of the aforementioned reason.

This observation led me to wonder if the factors contributing to the formation of contact domains might play a role, if any, in the enrichment of CenpT within these regions. In order to partly address this question, I also performed CenpT ChIP-Seq experiments with Scc1 RNAi and CapH2 RNAi cells. What I found was that depleting either cohesin or condensin II results in marginal differences in CenpT enrichment throughout the genome. Visual inspection of ChIP-Seq enrichment and annotated domains for CenpT in both WT and RNAi conditions exhibits that, overall, CenpT enrichment in the RNAi conditions stays within the same contact domains as in the WT, regardless of a loss or gain in intradomain contact frequencies (Figure 34A). In both RNAi conditions CenpT does not seem to be enriched in regions where it was not previously found. Correlating the CenpT ChIP-Seq data from the two RNAi conditions with the CenpT ChIP-Seq data from WT shows that both conditions are highly correlated with the WT (Figure 34B). Comparing the sizes of the annotated CenpT domains between conditions shows that CenpT domains are larger in the RNAi conditions as compared to the WT (Figure 34C). We suspect that this is likely due to the increase in enrichment rather than the spread of CenpT to previously depleted regions.

In summary, we find that depletion of cohesin and condensin II have opposing effects on interphase genome architecture in the B. mori cell line. As seen in other organisms, depleting Scc1 leads to a reduction of intradomain contact frequency and, in cases of strong depletion, a loss of contact domains all together. In this case, a 3-day RNAi depletion of Scc1 was enough to reduce intradomain contacts globally, but not enough to abolish contact domains. The loss of intradomain contacts after cohesin depletion is likely due to the loss of loop extrusion. With the loss of cohesin, the chromatin within the insulated boundaries of the contact domains in WT are no longer held together by cohesin's loop extrusion processivity, thus rendering them less likely to make spatial contacts. We also see an increase in long-range contacts in Scc1 RNAi as well. In contrast, what we see in CapH2 RNAi is an increase in intradomain contact frequency. This increase in intradomain contacts is interesting because it shows that cohesin and condensin II have counteracting roles in local chromatin compaction in B. mori interphase chromosomes. We also find that centromere regions are localized within contact domains and remain there despite cohesin or condensin II depletion.

experiments I will consider using spike-in controls to test for a global effect of the depletion on the transcriptome.

Comparing the CenpT RNAi normalized contact map with the WT normalized contact map shows no noticeable difference between them (Figure 34A). Plotting the insulation scores for both conditions in the same region of the genome shows an almost complete overlap between the two sets of insulation scores (Figure 34A). Indicating that there is no change in intra-or interdomain contact frequency between the CenpT RNAi and WT in this region. Looking at the insulation scores genome-wide confirms the previous observation and shows that the distribution of insulation scores between CenpT RNAi and WT are in the same range (Figure 34B).

To determine if there are any changes in contact frequency at increasing genomic distances I calculated and plotted the P(s) curve for CenpT RNAi and compared it to WT. The P(s) curve confirms that the short-range contacts frequencies are similar between CenpT RNAi and WT.

However, as genomic distance increases we find that there is also an increase in contact frequency in CenpT RNAi as compared to WT (Figure 34D). The interchromosomal contact mean is also higher in CenpT RNAi (Figure 34C). To see if centromere regions have any barrier-like characteristics I calculated and plotted the P(s) curves for CenpT and non-CenpT regions for both WT and CenpT RNAi . I found that contact frequency is higher in non-CenpT regions than in CenpT regions in both conditions at short distances. However, there is no change in contact frequency for either curve between CenpT RNAi and WT as genomic distance increases beyond contact domains.

(Figure 34E).

The results of this study show that depletion of CenpT does not affect local genome organization, but does seem to increase contact frequency at longer intrachromosomal distances. Preliminary investigation into whether or not centromere regions act as barriers between neighboring regions shows that contact frequencies between centromere and non-centromere regions remain the same after depletion of CenpT. More analysis will need to be done to validate this observation.

A note on BmN4-SID1 genome organization

The BmN4-SID1 cell line has extensive aneuploidy and rearrangments. So extensive, that we are not fully aware of how many copies of each chromosome exist. We assume based on microscopy images (data not shown) that each chromosome exists beyond a diploid set of complete and, During my thesis, I aimed at describing the genome organization in a group of organisms where it has previously not been described, holocentric insects. The results of my work bring new insights into CT's and chromosome compartment types within these organisms. Along with a hint of some factors involved in folding the genome.

B. mori interchromosomal contacts

To start, I took a top-down approach on hierarchical genome organization. A standard first step in studying genome organization via Hi-C is to plot the genome-wide contact map. This is to say, a contact map that displays all intra-and interchromosomal contacts within the genome assembly.

By doing this you can observe contacts being made between chromosomes and compare them to the relative frequency of contacts being made within chromosomes. A common observation is for the chromosomes to be distinguishable on the contact map as squares on the diagonal.

Distinguishable squares are also a sign of CT's since they indicate that intrachromosomal contacts are higher than interchromosomal contacts at all lengths of the chromosomes. Due to this characteristic, comparing the intrachromosomal contact mean to the interchromosomal contact mean is a good indicator of CT strength (Figure 25C).

Doing this for the B. mori embryo data produced a contact map with 28 distinguishable squares making up the diagonal, and no noticeable interchromosomal contacts or contact patterns (Figure 24B). Enlarging the image to a view of five chromosomes and their interchromosomal contacts reveals that there are colored bins present in between chromosomes representing contacts, but these bins are sparse and low in contact frequency. Comparing the P(s) curve to the interchromosomal contact mean shows that the lowest intrachromosomal mean (at the longest intrachromosomal distances) is almost two orders of magnitude higher. Analyzing this data for B.

mori indicated that the CT's in this organism are strong, so I compared this data to other organisms to get a better idea of how strong, or if this level of CT strength was not uncommon. I decided to analyze CT strength in C. elegans and D. melanogaster in the same way. I selected these organisms to compare with for a few reasons. C. elegans is another holocentric organism in which the genome organization has been studied and D. melanogaster is another well studied insect.

These two organisms are appropriate for a comparison with B. mori, a holocentric insect. I also selected these organisms because the chromosome sizes (or chromosome arm sizes) are comparable and the genome sizes are all within the same order of magnitude. After comparing the organisms in various ways (Figure 25) I concluded that B. mori displayed the strongest CTs of the three. Perhaps the most convincing analysis is the comparison of the P(s) curve with the mean interchromosomal contact frequency, which showed that the weakest intrachromosomal contacts in B. mori are still considerably higher than the interchromosomal contact mean than in the other organisms.

Our Hi-C analysis of CTs was further supported by a collaborator via microscopy techniques.

Using whole chromosome oligopaint FISH for six chromosomes in B. mori, the study revealed that B. mori has spatially distinct CTs with low levels of intermixing. The study also showed that the volume of B. mori nuclei is similar to that of D. melanogaster, but the B. mori genome is over three times as large. The study also looks into chromosome compaction and found that B. mori autosomes are tightly folded (Rosin et al., 2021a). Perhaps this characteristic can explain the lack of contacts between chromosomes. In order to fit more genomic material without increasing nuclear volume the chromosomes need to be more compact and more compact chromosomes make less interchromosomal contacts due to less intermixing. It is also interesting to speculate on the roles of factors that contribute to CT formation, like condensin II, and if these factors are more abundant, active or efficient in B. mori.

Nevertheless, I still did an analysis on the interchromosomal contacts to see if there were any preferential contacts between specific chromosomes. I found that, in general, higher frequencies of interchromosomal contacts are made between smaller chromosomes as compared to the larger ones. I also found that the two smallest chromosomes are not restricted to only making contacts with the other small chromosomes as they make more contacts than expected with the large ones as well (Figure 26). In humans, small gene-rich chromosomes have been shown to preferentially interact with each other, which is consistent with those localizing close to each other in nuclear space [START_REF] Boyle | The spatial organization of human chromosomes within the nuclei of normal and emerin-mutant cells[END_REF][START_REF] Lieberman-Aiden | Comprehensive Mapping of Long-Range Interactions Reveals Folding Principles of the Human Genome[END_REF][START_REF] Tanabe | Non-random radial arrangements of interphase chromosome territories: evolutionary considerations and functional implications[END_REF]. In an effort to explain the interchromosomal interactions between chromosomes, we made several correlations between the chromosomes for different genetic and epigenetic factors. Calculating the gene density for each chromosome shows that gene content varies only slightly between chromosomes (Supp. Figure 2). Thus, the interchromosomal contact preferences is likely not due to higher gene densities. We also noticed that the average interchromosomal contacts of chromosomes are positively correlated with their GC content and overall enrichments in H3K27me3, H3K9me3 and H3K36me3 but not H3K4me3 histone marks (Supp. Figure 3). It is thus hard to hypothesize what are the main factors involved in these interaction preferences.

One could speculate that perhaps the lack of heterogeneity between these factors could lead to an overall more homogeneous level of interchromosomal interactions. In mouse, it has been shown that the more gene-rich chromosomes preferentially interact with each other as well, but not to the extent at which they do in humans. It was hypothesized that this could be partly due to the fact that mouse chromosomes are more similar in size than human chromosomes. Gene density is more homogenous between mouse chromosomes as well [START_REF] Mayer | Common themes and cell type specific variations of higher order chromatin arrangements in the mouse[END_REF]. In B.

mori, the smallest chromosome is about half the size of the largest and our analysis on gene density shows that it varies only slightly. All in all, perhaps this indicates that when there is no striking difference between the genetic or epigenetic characteristics of chromosomes their interchromosomal interactions are determined by factors relating to their size.

B. mori intrachromosomal contacts

Moving from CTs down the genome organizational hierarchy, I next looked to investigate the contact patterns within chromosomes. Plotting the intrachromosomal contact map for any chromosome in the B. mori genome assembly will reveal signs of compartmentalization throughout the whole chromosome. In mammals, along with normalized contact maps it is common to also plot Pearson correlation maps. Calculating the Pearson correlations of the contacts is useful to "sharpen" or better define the contact pattern. This step is not totally necessary for B. mori contact maps because the compartmentalization is quite clear. This classic checkerboarding pattern indicating a segregation of contacts is visible along the length of the chromosomes and is occasionally interrupted by blank regions indicating areas where no (or few) contacts are made. Taking a closer look at these blank regions shows that there are areas of the chromosomes that do not participate in the checkerboard pattern. These areas also seem to make strong cis-interactions and can vary in size (Figure 29B). From visual inspection we assumed that the checkerboard pattern was a result of homotypic interactions between two distinct compartments. Initially we hesitated referring to this third, introverted region as a compartment.

One of the key characteristics of compartments was the high frequency of homotypic interactions and low frequency for heterotypic interactions. This third region seems to have neither.

Regardless, in order to define these regions, we used PCA data as input for a k-means clustering analysis and defined k=3. The resulting clusters corresponded to the three previously identified regions with high accuracy. We then characterized these regions physically, genetically and epigenetically and found that one of the regions involved in the checkerboard pattern is gene-rich and highly enriched in active epigenetic marks. The other checkerboarding region is depleted in active marks and in genes as compared to the genome. Given these characteristics and what has been found in other organisms we labelled these compartments A and B, respectively, like the ones originally defined in humans [START_REF] Lieberman-Aiden | Comprehensive Mapping of Long-Range Interactions Reveals Folding Principles of the Human Genome[END_REF]. Given the fact that a third compartment such as the one we have identified has not been extensively described in literature we arbitrarily labeled it compartment "X".

Our studies showed that compartment X is composed of some of the most gene-poor regions of the genome and that it is relatively enriched in H4K20me1. However, this enrichment is not consistent across all X compartment domains. H4K20me1 has a variety of roles in different organisms, but some studies have shown that this epigenetic mark may play a role in chromatin compaction [START_REF] Brejc | Dynamic Control of X Chromosome Conformation and Repression by a Histone H4K20 Demethylase[END_REF][START_REF] Tjalsma | H4K20me1 and H3K27me3 are concurrently loaded onto the inactive X chromosome but dispensable for inducing gene silencing[END_REF]. This is interesting because the compartment X P(s) curve does indicate that they are highly compact regions (Figure 29D,28B). We draw this conclusion from the P(s) curve because reports have shown that studying contact frequency as a function of genomic distance can reveal signs of local chromatin compaction. The main indicator of this compaction is when the slope of the P(s) curve becomes shallow at a specific, typically short, genomic distance. In addition, there is a pronounced bump indicating loop formation in the P(s) curve derivative. These indicators are present in B. mori P(s) curves and we have discovered that this local compaction mainly comes from the X compartment (Figure 30). These studies claim that this local compaction is a result of high levels of loop extrusion processivity done by cohesin [START_REF] Fudenberg | Emerging Evidence of Chromosome Folding by Loop Extrusion[END_REF][START_REF] Fudenberg | Formation of Chromosomal Domains by Loop Extrusion[END_REF]. Furthermore, it has also been shown experimentally that when you deplete cohesin you lose local compaction and the shoulder in the P(s) curve and the bump in the derivative are either reduced or disappear [START_REF] Schwarzer | Two independent modes of chromatin organization revealed by cohesin removal[END_REF][START_REF] Spracklin | Heterochromatin diversity modulates genome compartmentalization and loop extrusion barriers[END_REF], which is hypothesized to result from the loss of loop extrusion.

It would be interesting to investigate if there is a link between loop extrusion and the deposition of H4K20me1 given that they are both associated with chromatin compaction and may play a role in the existence of compartment X. A future perspective would be to use the B. mori cell line to map the enrichment profile of H4K20me1 via ChIP-Seq. Perhaps this characteristic can be used as a proxy to identify some X compartment regions in the cell line. Once these areas are identified one could repeat an RNAi depletion of Scc1 and see if H4K20me1 enrichment is reduced in these areas. If so, we could speculate that perhaps a component of loop extruder complexes is also involved in depositing the mark. This could explain the enrichment of the mark at compartment X. However, one can also not exclude the inverse. A situation in which chromatin enriched in H4K20me1 is in some way more permissive to loop extrusion. In this case we would have to identify the factor responsible for H4K20me1 deposition in B. mori, perform a knockdown or depletion of this factor and see how that effects local compaction of X compartments and/or enrichment of cohesin components at X compartment boundaries.

Although this is the first-time compartment X has been described in a holocentric insect, it is not et al., 2021). The B. mori X compartment is also enriched in H3K27me3, but differs from compartment I in contact frequency. The I compartment was reported to make contacts with both compartment A and B, when in our case with the X compartment, it makes contacts with neither. Similar to that characteristic, a compartment was identified in human HCT116 cells. In this study, this compartment is called "B0" and is described as having a neutral interaction preference. It is also described as 'poised heterochromatin' that can transition between active and inactive chromatin in different cell types [START_REF] Spracklin | Heterochromatin diversity modulates genome compartmentalization and loop extrusion barriers[END_REF]. Although this compartment visually resembles compartment X in B. mori, it was shown to have a slight enrichment in histone variant H2A.Z and ChIP-Seq for H2A.Z in B. mori does not show a preferential enrichment in X compartments (data not shown).

Our study on interchromosomal homotypic compartment contact frequency revealed that X compartments make more contacts between chromosomes than expected. Collaborators have shown that X compartments are positioned more peripherally in CTs than A and B compartments. This could not only explain why they make contacts interchromosomal, but also why they make so few contacts intrachromosomally (Supp. Figure 1). Efforts were also made to see if the X compartment was associated with centromere regions. Initial experiments to map the Cenp-T enrichment profile in the p50 embryos were met with technical issues as our Cenp-T antibody did not seem to work efficiently and we were not able to produce an informative ChIP profile. Given the similarity of short-range contacts between p50 and BmN4-SID1 Hi-C data, I aligned Cenp-T ChIP-Seq data from BmN4-SID1 to p50 Hi-C data and found that, visually, there is no correlation between Cenp-T enrichment and the X compartment. Still, it will be necessary to map centromere regions in the p50 embryos to validate this initial observation. Until then, we can not rule out a relationship between the X compartment and centromeric function at some point during the cell cycle. [START_REF] Gassler | A mechanism of cohesin-dependent loop extrusion organizes zygotic genome architecture[END_REF][START_REF] Haarhuis | The Cohesin Release Factor WAPL Restricts Chromatin Loop Extension[END_REF][START_REF] Rao | Cohesin Loss Eliminates All Loop Domains[END_REF][START_REF] Rowley | Condensin II Counteracts Cohesin and RNA Polymerase II in the Establishment of 3D Chromatin Organization[END_REF][START_REF] Schwarzer | Two independent modes of chromatin organization revealed by cohesin removal[END_REF][START_REF] Wutz | Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins[END_REF]. The insulation score analysis after cohesin depletion showed that intradomain contacts were reduced, but interdomain contacts remained mostly unchanged. This indicates a loss of extrusion and not of insulation,

which is what has come to be expected from cohesin depletion. One would expect to see a loss of insulation after the depletion of insulator proteins such as CTCF [START_REF] Kubo | Preservation of Chromatin Organization after Acute Loss of CTCF in Mouse Embryonic Stem Cells[END_REF][START_REF] Nora | Targeted Degradation of CTCF Decouples Local Insulation of Chromosome Domains from Genomic Compartmentalization[END_REF][START_REF] Wutz | Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins[END_REF]. Comparison of the Scc1 RNAi P(s) curve to WT indicates a reduction of local compaction, but it is difficult to say whether the loop extrusion "shoulder" is no longer present. The cell line Hi-C data that I've analyzed is at 20 kb resolution. The embryo data that I previously analyzed exhibiting the shoulder is at 2 kb resolution. The loop extrusion shoulder in this data is found between 10 kb and 50 kb, so I am unable to see it completely with the cell line Hi-C data.

To further characterize the role of cohesin in B. mori it is necessary to map its locations across the genome. Commercially available antibodies against cohesin components do not seem to efficiently target B. mori cohesin. To overcome this, we have cloned Scc1 fused to a 3X-FLAG peptide under an overexpressed promotor and have confirmed nuclear localization by immunofluorescence (data not shown). At this point we are in the process of cloning the same construct under the endogenous promotors. Once done, the next step will be to transfect the construct into the BmN4-SID1 cells and perform a ChIP-Seq experiment targeting the 3X-FLAG peptide, which we have an efficient antibody for. Based on the results from the cohesin depletion I would expect this experiment to render results showing an enrichment of Scc1 at contact domain boundaries.

In parallel to the Scc1 RNAi analysis I also analyzed data from condensin II depletion. I depleted condensin II by performing RNAi against its subunit, CapH2. In D. melanogaster, condensin II has been shown to contribute to genome organization in interphase cells (Rosin et al., 2018;[START_REF] Rowley | Condensin II Counteracts Cohesin and RNA Polymerase II in the Establishment of 3D Chromatin Organization[END_REF] assume the effects of cohesin and condensin II depletion to be at the global scale and not restricted to active sites. However, more analysis will have to be done in order to verify this.

Together with the Hi-C experiments done in the RNAi conditions I did ChIP-Seq against CenpT.

In the WT condition, CenpT localizes within contact domains (Figure 34A). Depleting either cohesin or condensin II does not seem to change the localization of CenpT enrichment (Figure 34A). This indicates that perhaps CenpT enrichment is insulated by insulator proteins and not promoted by an increase in contact frequency by loop extrusion. Along with the RNAi depletion of cohesin and condensin II components, I also performed an RNAi depletion of CenpT. I analyzed the results of CenpT RNAi in the same manner as for the SMC complexes. In this way, I found that the depletion of centromere regions does not have a local effect on genome organization (Figure 35A,B). It is possible that my analysis method is biased towards revealing the effects of loop extruders and perhaps is not appropriate to analyze the effect of centromere depletion. Alternative methods of analysis will have to be explored in order to conclude with certainty that centromere depletion has no effect. The CenpT RNAi P(s) curve did indicate a gain in long range contacts (Figure 35D). RNAi depletion of Scc1, CapH2 and CenpT also indicated an increase in mean interchromosomal contact frequency when compared to WT (Figure 33D, Figure 35C). However, it is difficult to assess the effects on long-range and interchromosomal contacts in the cell line for the following reason.

Methods

B. mori strains

All experiments have been carried out using B. mori embryos of the p50 reference strain from the Kyushu University facility (silkworm Genetic Resource Database https://shigen.nig.ac.jp/silkwormbase/topAction.do), except for oligopaint FISH (see dedicated paragraph). Upon arrival, diapaused eggs are stored in the cold (4°C) for more than 70 days before placed at RT (~25°C) to break diapause and restart the embryonic development. Late embryos (24h post-diapause release) are harvested to perform Hi-C and ChIP experiments.

BmN4-SID1 culture conditions BmN4-SID1(RRID:CVCL_Z091) cell lines were maintained in Sf-900 II SFM medium (GIBCO catalog # 10902-088) supplemented with 10% fetal bovine serum (Eurobio catalog # CVFSVF0001), antibiotic-antimycotic (GIBCO catalog # 15240-062) and 2mM L-glutamine (GIBCO catalog # 25030-024) at 27°C. Immunofluorescence BmN4-SID1 cells were fixed in 100% ice-cold MeOH and permeabilized using 0.3% Triton x-100 in PBS. Cells were then blocked in 3% BSA-PBS. Primary antibody incubations were done in blocking buffer overnight at 4°C. The following primary antibodies were used at 1:1000 dilution: anti-CenpT serum (rabbit polyclonal) (Cortes-Silva et al., 2020), anti-phospho-Histone H3 (Ser10) (mouse monoclonal, Merck Millipore Catalog # 05-806). The next day, cells were washed three times with 0.3% Triton x-100 in PBS and incubated for 1 hour at 4°C with secondary antibodies diluted to 1:1000 in blocking buffer. The following fluorescent-conjugated secondary antibodies were used: goat anti-rabbit IgG Alexa Fluor 488 (Thermo Fisher Scientific, catalog # A-11034, RRID AB_2576217), and goat anti-mouse IgG Alexa Fluor 568 (Thermo Fisher Scientific, catalog # A-11004, RRID AB_2534072). Cells were washed three times with 0.3% Triton x-100 in PBS and counterstained with DAPI for 3 min at room temperature (Sigma catalog # D9542) before washing again in 0.3% Triton x-100 in PBS and mounting samples in Vectashield Antifade Mounting Medium (Vector Laboratories catalog # H1000; RRID:AB_2336789).

Microscopy

Images of BmN4-SID1 cells in untreated or RNAi conditions were acquired on a Zeiss Axiovert Z1 light microscope. Images for phospho-Histone H3 (Ser10) positive cells were acquired using the 10X objective. Quantification of mitotic index was performed using the Fiji (ImageJ) software version 2.1.0/1.53c [START_REF] Schindelin | Fiji: an open-source platform for biological-image analysis[END_REF] on unprocessed TIFF images.

GC content analysis GC content for B. mori genome has been calculated using the BEDTools suite [START_REF] Quinlan | BEDTools: a flexible suite of utilities for comparing genomic features[END_REF]. We used the function "bedtools nuc" on 40 kb or 100 kb non-overlapping windows of the genome to extract the GC% for each window. For windows including "N" nucleotides and end of chromosome windows, the window size has been adjusted accordingly (considering only the size of "As", "Ts" "Cs" and "Gs") for accurate GC% calculation. The GC content distribution genome-wide and inside each compartment are shown as box plots and corresponds to the GC% per 40 kb windows of the genome or the compartment of interest, using the "bedtools intersect" function of the BEDTools suite.

B. mori repeat analysis

To determine the positions of transposable elements (TEs) in the B. mori chromosome of the genome assembly, we use the RepeatMasker software (Smit, AFA, Hubley, R & Green, P. RepeatMasker Open-4.0. 2013-2015 http://www.repeatmasker.org) on a custom B. mori transposon library (kindly provided to us by M. Kawamoto, University of Tokyo) using the "nolow" parameter. We then aggregated the output of defined TE coordinates along the genome using the "bedtools merge" function of BEDTools suite [START_REF] Quinlan | BEDTools: a flexible suite of utilities for comparing genomic features[END_REF]. This list allows us to confirm the overall genome-wide coverage of TE of 46% in B. mori. The list has been further used to determine the TE coverage per 40 kb windows genome-wide and inside each compartment using "bedtools makewindows", "bedtools coverage" and the "bedtools intersect" function of the BEDTools suite and shown as box plots.

Gene coverage analysis

The gene coverage has been plotted as box plot representing the distribution per 40 kb windows along the genome and inside each compartment and has been computed using the "bedtools makewindows" and "bedtools coverage" function of the BEDTools suite based on the gene list cleared from TEs previously generated.

Hi-C experiment and read processing For RNAi samples: cells were harvested and centrifuged for 5 min at 500 g, resuspended in 5 ml PBS and crosslinked with 2% final formaldehyde solution (Sigma-Aldrich F8775). Samples were inverted 10 times and left to incubate for 10 min at RT. Crosslinking was quenched with 200 mM glycine and inverted 10 times and left to incubate for 5 min at RT follow by an additional incubation for 15 min at 4°C. Samples were centrifuged for 5 min at 300 g and washed with 1 ml PBS and transfered to 1.5 ml tubes. Samples were then centrifuged for 3 min at 2000 g and stored at -80°C until Hi-C processing. For p50 embryos: Embryos have been fixed by poking chorions of ~30 eggs per experiment with a fine needle directly followed by immersion in 3 % final formaldehyde solution (Sigma-Aldrich F8775) in PBS for 30 min at RT. The reaction is then quenched by addition of glycine to a final concentration of 200 mM, 20 min at RT. Eggs are then washed in PBS and dissected using fine forceps while immerged in PBS. Embryos are transferred to 1.5 ml Eppendorf tubes, spined shortly to be pelleted. Then, residual PBS was removed and embryos froze at -80°C. After dissection, adult heads have been similarly dug in 3% formaldehyde, quenched in glycine 200 mM, washed in PBS and froze. All sampled have then been processed for Hi-C using the Arima-HiC+ kit from Arima Genomics following manufacturer's instructions and Illumina library have been prepared using the KAPA HyperPrep Kit from Roche (ref: KK8504) coupled to the Illumina TruSeq sequencing adapters (ref 20020590) following the modified version of the protocol by Arima. Hi-C library have been sequenced as PE100 on NovaSeq at the CurieCoreTech high-throughput sequencing platform. Pair-end reads have been processed using the HiCPro suite [START_REF] Servant | HiC-Pro: an optimized and flexible pipeline for Hi-C data processing[END_REF]. ".matrix" and "abs.bed" file from HiCPro are used to build Hi-C matrices in h5 format using the "hicConvertFormat" tool from HiCExplorer. For RNAi Hi-C data: matrix reads are then subsampled to the data set with smallest read count using "hicNormalize" and parameter "-normalize smallest". Subsampled Hi-C data is then normalized using "hicCorrectMatrix" and parameters "--correctionMethod ICE --filterThreshold -1.56 4" [START_REF] Ramírez | High-resolution TADs reveal DNA sequences underlying genome organization in flies[END_REF]. For comparision study: Hi-C reads from other species have been fetched from publicly available datasets and processed similarly as for p50 embryos (C. elegans: SRR14049318 [START_REF] Jimenez | Condensin DC spreads linearly and bidirectionally from recruitment sites to create loop-anchored TADs in C. elegans[END_REF]; D. melanogaster: SRR10485713 [START_REF] Zenk | HP1 drives de novo 3D genome reorganization in early Drosophila embryos[END_REF]). p50 B. mori embryo ChIP experiment and read processing ChIP-seq experiments have been performed on embryos 24h after diapause released. Approximately 200 eggs are gathered in a 1.5ml Eppendorf tube per ChIP experiment. 200 µl of methanol-free formaldehyde (FA) (ThermoFIsher Scientific # 28906) 1% in PBS is added to the tube before egg chorion are mechanically broken using a piston pellet (Dutcher # 45650) and FA volume adjusted to 1 ml. Embryos are fixed for 10 min at RT on wheel and then the reaction is quenched by addition of glycine to a final concentration of 0.125 M for 5 min at RT on wheel. Fixed embryonic cells in the supernatant are transferred to a clean 2 ml Eppendorf tube, and chorion are crushed with piston pellets and washed with 200 µl of PBS two to three times until only clear chorion pieces are visible and all embryonic cells are recovered in the 2 ml tube. Cells are then centrifugated 3 min at 600 g, 4°C, washed with 1 ml PSB and resuspended in 130 µl Lysis buffer w/o SDS (EDTA 10 mM,Tris-HCl (pH8.1) 50 mM + Protease Inhibitor (cOmplete™, EDTA-free Protease Inhibitor Cocktail, Roche #04693132001). Embryonic tissues are then ruptured using Covaris sonication with the following parameters on a E220Evolution apparatus: 60 sec, Duty 10%, Power 75 W, cycles/burst 1000, 7 °C. Cells are then lysed by addition of SDS to a final concentration of 1% and incubated for 30-60 min on ice. A this stage an aliquot of 10 µl is taken to serve as a quality control to check the non-degradation of the DNA. The remaining chromatin sample is sheared using Covaris sonication on a E220Evolution apparatus: 8 min (480 sec), Duty 10%, Power 105 W, cycles/burst 200, 7 °C. To allow for immunoprecipitation (IP), ChIP buffer (Triton X-100 1%, NaCl 150 mM, EDTA 2 mM, Tris-HCl 20 mM, pH 8.1 + Protease Inhibitor) is added to a final volume of 1500 µl and the sample is centrifugated for 10 min at 16000g, 4 °C. Cell debris are discarded and the supernatant containing soluble chromatin is transferred to a clean 2 ml Eppendorf tube. A sample of 50 µl of chromatin is kept as Input and the rest is added to the antibody (Ab) of choice (CenpT: serum (Cortes-Silva et al., 2020), H3K36me3: abcam ab9050, H3K4me3: diagenode C15200152, H3K27me3: cell signaling 9733S, H3K9me3: abcam ab8898, K4K20me1: abcam ab9051) bound to Dynabeads protein A (ThermoFisher Scientific # 10002D) according to manufacturer's instructions. The samples (Input and IP) are incubated on wheel at 4°C O/N. IP samples containing the Ab bound to Protein A and magnetic beads are then washed once in cold TSEI (SDS 0.1%, Triton X-100 1%, NaCl 150 mM, EDTA 2 mM, Tris-HCl 20 mM pH 8.1), 4 times in cold TSEII (SDS 0.1%, Triton X-100 1%, NaCl 500 mM, EDTA 2 mM, Tris-HCl 20 mM pH8.1) and three times in cold TE (Tris-HCl 10 mM pH8.1, EDTA 1 mM), incubated 2 min at RT on a wheel between each wash and recovered on a magnetic rack. Finally, DNA from all samples (QC, Input and IP) is extracted by addition of Extraction buffer (Tris-HCl 20 mM pH8.1, EDTA 10 mM, EGTA 5 mM, NaCl 300 mM, SDS 1%, RNase 0.1 mg/ml) to a final volume of 100 µl. After 15 min incubation at 37°C to allow for RNase activity, 3µl of Proteinase K (20 mg/ml, Qiagen # 19131) is added to each sample and incubation is continued at 65°C for at least 4h or O/N. DNA is purified using 1.2 x SPRIselect beads (Beckman Coulter #B23318) according to manufacturer's instructions and quality and quantity of samples are evaluated on a TapeStation instrument (Agilent 4200 TapeStation System). Input and IP samples are processed to make Illumina library using the KAPA HyperPrep Kit from Roche (ref: KK8504) coupled to the Illumina TruSeq sequencing adapters (ref 20020590) and sequenced as PE100 on NovaSeq at the CurieCoreTech high-throughput sequencing platform. Reads have been analyzed using in house pipeline written by the bioinformatics platform of the Institut Curie (Valentin Laroche, Nicolas Servant) aligning reads with Bowtie2 using default parameters and then filtered for a minimum mapping quality of 10 and non-ambiguously mapping reads (grep -v "XS:i"). The resulting alignment files for each ChIP experiment are normalized against their corresponding input using the "bamCompare" function of the DeepTools suite [START_REF] Ramírez | deepTools2: a next generation web server for deep-sequencing data analysis[END_REF] to produce log2 ratio of IP vs Input at various bin sizes that are further potted using pyGenomeTracks [START_REF] Lopez-Delisle | pyGenomeTracks: reproducible plots for multivariate genomic datasets[END_REF] or Matplotlib v3.5.2 [START_REF] Hunter | Matplotlib: A 2D Graphics Environment[END_REF].

BmN4-SID1 ChIP experiment and read processing Two confluent T75 flasks (Thermofisher, catalog # 156499) of BmN4-SID1 cells were used for each CenpT ChIP. Cells were cross-linked in freshly prepared 1% MeOH-free formaldehyde (Thermofisher catalog # 28906) for 10 min on rotation at RT. Cross-linking was quenched by adding glycine to 125 mM for 2 min at RT. Cells were then washed in ice-cold PBS and incubated for 10 min with 150 ml ice-cold lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris-HCl pH 8.1, 1 X cOmplete Protease Inhibitor Cocktail (Roche catalog # 11697498001)). To the cell lysates, 1350 ml ChIP buffer (1% Triton X-100, 150 mM NaCl, 2 mM EDTA, 20 mM Tris-HCl pH 8.1, 1 X Protease inhibitor cocktail) was added along with 4.5 ml CaCl2 1 M (3 mM final) and then pre-warmed for 2 min at 37C. Nuclei were then treated with 1 of MNase (Sigma, catalog # N3755-500UN) for 45 min, respectively at 37C with 700 rpm agitation. MNase reaction was stopped by adding a mix of 30 ml EDTA (0.5 M stock) and 60 ml EGTA (0.5 M stock). Each MNase-treated nuclei sample was then sonicated using a Covaris E220 sonicator under the following parameters: 150 s, Duty 10%, Power 75 W, cycles/burst 200, 7C. Sonicated chromatin was centrifuged 3 min at 16000 g and clear supernatant containing the solubilized chromatin was saved either as input or for ChIP. Anti-CenpT serum (rabbit polyclonal) (Cortes-Silva et al., 2020) diluted in PBS-T (0.02% tween-20) was incubated with Protein A Dynabeads (Thermofisher, catalog # 10001D) for 10 min at RT to allow for beads-antibody binding. Antibody-bound beads were washed with ChIP buffer and mixed with ChIP chromatin. All samples were incubated overnight at 4C. Chromatin-bound beads were collected the next day on a magnetic rack and washed with the following ice-cold buffers: once with low-salt TSE 1 (0.1% SDS, 1% Triton X-100, 150 mM NaCl, 2 mM EDTA, 20 mM Tris-HCl pH 8.1,); four times with high-salt TSE II (0.1% SDS, 1% Triton X-100, 500 mM NaCl, 2 mM EDTA, 20 mM Tris-HCl pH 8.1); and three times with 1x TE. DNA was directly extracted from chromatin-beads or input by adding DNA extraction buffer (20 mM Tris-HCl pH 8.1, 10 mM EDTA, 5 mM EGTA, 300 mM NaCl, 1% SDS) and incubating at 37C followed by reversing cross-links by addition of proteinase K (QIAGEN, catalog # 19131) and incubation overnight at 65C. DNA is purified using 1.2 x SPRIselect beads (Beckman Coulter #B23318) according to manufacturer's instructions and quality and quantity of samples are evaluated on a TapeStation instrument (Agilent 4200 TapeStation System). Input and IP samples are processed to make Illumina library using the KAPA HyperPrep Kit from Roche (ref: KK8504) coupled to the Illumina TruSeq sequencing adapters (ref 20020590) and sequenced as PE100 on NovaSeq at the CurieCoreTech high-throughput sequencing platform. Reads have been analyzed using in house pipeline written by the bioinformatics platform of the Institut Curie (Valentin Laroche, Nicolas Servant) aligning reads with Bowtie2 using default parameters and then duplicates are removed from the data using Picard "MarkDuplicates" with parameter "REMOVE_DUPLICATES=TRUE" (http://broadinstitute.github.io/picard/). The resulting alignment files for each ChIP experiment are normalized against their corresponding input using the "bamCompare" function of the DeepTools suite [START_REF] Ramírez | deepTools2: a next generation web server for deep-sequencing data analysis[END_REF] to produce log2 ratio of IP vs Input at various bin sizes.

Transcriptional profiling and data analysis Total RNA was isolated using Trizol reagent (Invitrogen, catalog # 15596018) following the manufacturer's instructions. RNA has been processed to Illumina library by the Stranded mRNA prep Ligation protocol performed and sequenced as PE100 on NovaSeq at the CurieCoreTech high-throughput sequencing platform. Adaptor-trimmed, paired-end reads of 100 bp length were mapped to the B. mori genome assembly using STAR version 2.7. [START_REF] Dobin | STAR: ultrafast universal RNA-seq aligner[END_REF]. The number of mapped RNA-seq reads in gene models (kindly provided to us by M. Kawamoto, University of Tokyo) or in genome-wide 10 kb windows were counted using ht-seq software [START_REF] Putri | Analysing high-throughput sequencing data in Python with HTSeq 2.0[END_REF]. Read counts were transformed into TPM (transcripts per million mapped transcripts) scores to evaluate normalized expression levels. 10 kb resolution TPM-normalized RNA-seq coverage tracks were further generated using the "bamCompare" function of the DeepTools suite [START_REF] Ramírez | deepTools2: a next generation web server for deep-sequencing data analysis[END_REF].

RNAi-mediated knock-down BmN4-SID1 cells were grown in T75 flasks (Thermofisher, catalog # 156499) and incubated with 250 pg/ml dsRNA for three days (Scc1, CapH2) or five days (CenpT). After three or five days, cells were fixed and processed for Hi-C, ChIP-Seq, RNA-Seq and IF as described.

Annotation of CenpT domains CenpT domains were annotated as previously described [START_REF] Senaratne | Formation of the CenH3-Deficient Holocentromere in Lepidoptera Avoids Active Chromatin[END_REF] with few modifications. CenpT ChIP-Seq signal originally in 10 kb windows was averaged over 20 kb using a sliding window method. The averaged windows with positive scores (log2-ratio > 0) within a genomic distance of 50 kb were merged using BEDTools suite [START_REF] Quinlan | BEDTools: a flexible suite of utilities for comparing genomic features[END_REF].

Compartment calling: k-mean clustering The 40 kb bin resolution matrix for p50 24h post-diapause embryos have been processed using the HiCExplorer utilities [START_REF] Ramírez | High-resolution TADs reveal DNA sequences underlying genome organization in flies[END_REF] to produce the corresponding Pearson correlation matrix. After normalization, PCA 1 to 3 have been extracted using Scikit-learn (Scikit-learn: Machine Learning in Python, Pedregosa et al., JMLR 12, pp. 2825-2830, 2011 ; API design for machine learning software: experiences from the scikit-learn project, Buitinck et al., 2013 (https://doi.org/10.48550/arXiv.1309.0238)). The explained variance ratio per PCA for each chromosome is provided in Table S3 (p50_D2_24h_Bmori-v4_40000_pearson_PCA_explained_variance_ratio.xlsx). Although the first two PCA were explaining most of the signal for all chromosomes, we kept the three first PCA to apply the K-Means clustering method of Scikit-learn on the Pearson correlation matrix for each chromosome of the genome to take more than 80 % of the signal in to account. We initially called three clusters per chromosomes, to fit our visual observation of the matrix. We realize that this was overcalling the cluster corresponding to the X compartment on chromosomes where it is sparse. Thus, for Chr15, 17, 21 and 28, we decided to call 5 clusters instead, keep one corresponding to the sparse X compartment, and fuse two corresponding to the A compartment and the last two the B (See Figure S7 for cluster representation along PCA1 and PCA2). As mentioned previously, Chr11 matrix is peculiar and split into two large non-interacting parts due to the presence of the rDNA locus. Thus, we had to treat the two parts (1:4479999 and 4480000:20440007) as if they were independent chromosomes to be able to have PCAs representing similar features than for the other chromosomes. Chr24 also has a peculiar matrix profile interrupted by a very large repeatrich and unmappable region. Thus, we also treated the two large mappable regions (1:11599999 and 14200000:17359173) as individual chromosomes. Clusters have been attributed to compartments A, B or X according to the positions of their centroids (See Figure S7). For most chromosomes, PCA1 was mainly used to distinguish between A and B compartments and PCA2 to isolate compartment X. For some chromosomes (Chr11: 1-4479999, Chr24: 14200000-17359173 and Chr27), PCA1 and PCA2 signals were inverted, so the compartment calling has been adjusted accordingly. All python scripts have been written using Python 3 in Jupyternotebook (Kluyver, T., Ragan-Kelley, B., Fernando P'erez, Granger, B., Bussonnier, M., Frederic, J., … Willing, C. (2016). Jupyter Notebooks -a publishing format for reproducible computational workflows. In F. Loizides & B. Schmidt (Eds.), Positioning and Power in Academic Publishing: Players, Agents and Agendas (pp. [87][88][89][90]). We provide the compartment coordinates for all chromosomes as files Sx_p50_D2-24h_Bmori-v4_40kb_comp_A_merge.bed Sy_p50_D2-24h_Bmori-v4_40kb_comp_B_merge.bed and Sz_p50_D2-24h_Bmori-v4_40kb_comp_X_merge.bed.

Genomic distance plot, P(s) curves and derivative For whole genome P(s) curves, all calculations were done for each chromosome individually and then averaged together at the end. The separation "s" between each pairwise contact was calculated by subtracting the start position of the first coordinate from the start position of the second. All contacts were grouped into logarithmically sized bins based on their "s". Logarithmic bins were determined by the range of "s" values for each chromosome. We calculated the mean of all contact probability values "P" within each logarithmic bin and plotted them on a log-log scale.

The derivative of the P(s) was determined by plotting the change of slope at each point for the P(s) curve. We do this in R (R Core Team (2021). R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. URL https://www.Rproject.org/) by performing the linear regression for each point in the P(s) curve with the following five points. We extract the slope of the line of regression using the coefficients() tool. Then we plot the slope for each "s" on an x-log scale. For compartmental domain P(s) curves, we use BEDTools "intersect" [START_REF] Quinlan | BEDTools: a flexible suite of utilities for comparing genomic features[END_REF] to extract the locus for each individual compartmental domain from the first and second coordinates of the Hi-C matrices. This results in a file containing only the pairwise contacts within that domain. We follow the previously described method to find the P(s) curve and derivative for each compartmental domain individually and then average them together at the end.

Interchromosomal contact analysis All intrachromosomal contact data was omitted from this analysis. The interchromosomal contact probability for each pairwise chromosome combination was calculated by dividing the observed sum of contacts between the two chromosomes by the expected sum. The observed value was calculated by taking the sum of all the contacts between each pairwise combination of chromosomes. The expected value was calculated by taking the sum of all the contacts between each pairwise combination of chromosomes from an "expected" matrix where each interchromosomal bin had a value equivalent to the mean of all interchomosomal values from the observed data. We determine the order of the chromosome pairs via hierarchical clustering of the observed/expected values. For compartment interchromosomal contact probability, we use BEDTools "intersect" [START_REF] Quinlan | BEDTools: a flexible suite of utilities for comparing genomic features[END_REF] to extract the loci for each individual compartment from the first and second coordinates of the observed and expected Hi-C matrices. This results in a file containing only the pairwise contacts within that compartment. We then follow the previously described method to find the observed sum and expected sum for each pairwise combination of chromosomes.

Interchromosomal contact mean All intrachromosomal contact bins are filtered out of the Hi-C data. The normalized contact frequency for each remaining bin is summed. This sum is divided by the total number of bins (mapped or unmapped) in the interchromosomal space, depending on the resolution.

ChIP enrichment in compartment analysis

For each ChIP-Seq data set we plot the IP/Input ratio as a track using deepTools "bamCompare" [START_REF] Ramírez | deepTools2: a next generation web server for deep-sequencing data analysis[END_REF]. Using BEDTools "intersect" we extract the regions of the ChIP-Seq track pertaining to the loci of each compartment. We calculate the median ChIP-Seq enrichment score (ratio of read counts) within each compartment. We then divide this score by the median ChIP-Seq enrichment score genome-wide. ChIP enrichment profiles per compartments for supplementary figure S8 have been computed using the "computeMatrix scale-regions", "plotProfile" and "plotHeatmap" of the deepTools v3.5.0 suite [START_REF] Ramírez | deepTools2: a next generation web server for deep-sequencing data analysis[END_REF] on the log2 ratio files.

Hi-C Insulation Scores Insulation scores were calculated using the TADtool "tads" tool with the "-a insulation" and "-v" parameters. Insulation scores were called using a window size of 100000 and a cutoff of 9. These window size and cutoff were defined using the TADtool "plot" tool on untreated BmN4-SID1 Hi-C data.

Copy Number Variation

Copy number variation track was produced using Control-FREEC [START_REF] Boeva | Control-FREEC: a tool for assessing copy number and allelic content using next-generation sequencing data[END_REF] software using an INPUT ChIP bam file as input and running the program with "BedGraphOutput=TRUE" parameter. Bedgraph file from the output was converted to bigwig using bedGraphToBigWig v2.8 from UCSC tools set [START_REF] Kent | BigWig and BigBed: enabling browsing of large distributed datasets[END_REF].

Oligopaint design, synthesis, DNA FISH, and image analysis Oligopaint libraries designed using a modified version of the Oligominer pipeline (Beliveau et al., 2018). CT paints were previously published [START_REF] Rosin | Dosage compensation in Bombyx mori is achieved by partial repression of both Z chromosomes in males[END_REF](Rosin et al., , 2021b)). Coordinates and all information for single domain paints can be found in Table 3. Oligopaints were synthesized as previously described by adding barcodes to each 70 bp oligo for PCR-based amplification (Nguyen and Joyce, 2019). FISH with Oligopaints was performed as previously described [START_REF] Rosin | Dosage compensation in Bombyx mori is achieved by partial repression of both Z chromosomes in males[END_REF]. Briefly, after fixation, cells on slides were washed 3x 5 min each in 0.1% Triton X-100 in PBS (PBS-T0.1%) at RT, then permeabilized with PBS-T0.5% for 15 min at RT. Cells were subsequently pre-denatured using the following washes: 1x 5 min in 2xSSCT (0.3 M NaCl, 0.03 M sodium citrate, 0.1% Tween-20) at RT, 1x 5 min 2xSSCT/50% formamide at RT, 1x 2.5 min in 2×SSCT/50% formamide at 92°C, 1x 20 min at 60°C in 2xSSCT/50% formamide. Primary Oligopaint probes* in hybridization buffer (10% dextran sulfate/2xSSCT/50% formamide/4% polyvinylsulfonic acid) were applied to slides and sealed under a 22x22 mm coverslip with rubber cement. Once rubber cement had dried, cells were then denatured at 92°C for 2.5 min and placed in a humidified chamber at 37°C overnight. The next day, slides were washed as follows: 2xSSCT at 60°C for 15 min, 2xSSCT at RT for 15 min, 0.2xSSC at RT for 10 min. Secondary probes (10 pmol/25 µL) containing fluorophores were added to slides, resuspended in hybridization buffer as described above, covered with 22x22 mm coverslip and sealed with rubber cement. Slides were incubated at 37°C in a humidified chamber for 2 h before repeating the above washes. Finally, some slides were stained with DAPI DNA stain in PBS for 5 min, followed by 2x 5 min washes in PBS-T0.1%. All slides were mounted in Prolong Diamond (Invitrogen) and left to cure overnight at RT before sealing with nail polish. * Primary probes were used for either 3-or 4-color FISH. For 3-color FISH, 2 domains plus the CT were painted and cells were stained with DAPI. For 4-color FISH, 3 domains plus the CT were painted. Data from these two experiment types were grouped for all analyses. A total of at least 3 biological replicates (10-12 pooled embryos) were performed for each FISH experiment measuring domain position in the CT. Images were acquired on a Leica DMi6000 widefield fluorescence microscope using an HCX PL APO 100x/1.40-0.70 Oil objective (Leica) and Leica DFC9000 sCMOS Monochrome Camera. DAPI, CY3, CY5, and FITC filter cubes were used for image acquisition for all experiments. All images were processed using the LasX software and Huygens deconvolution software, and tiffs were created in ImageJ. For quantification, deconvolved images were segmented and measured using a customized pipeline in the TANGO 3D-segmentation plug-in for ImageJ [START_REF] Ollion | TANGO: a generic tool for highthroughput 3D image analysis for studying nuclear organization[END_REF], using either the 'Hysteresis' or 'Spot Detector 3D' algorithms. Statistical analyses were performed using Prism 9 software by GraphPad. Figures were assembled in Adobe Illustrator.

Review

The Impact of Centromeres on Spatial Genome Architecture

Héloïse Muller, 1,2, * José Gil, Jr, 1, 2 and Ines Anna Drinnenberg 1,2 The development of new technologies and experimental techniques is enabling researchers to see what was once unable to be seen. For example, the centromere was first seen as the mediator between spindle fiber and chromosome during mitosis and meiosis. Although this continues to be its most prominent role, we now know that the centromere functions beyond cellular division with important roles in genome organization and chromatin regulation. Here we aim to share the structures and functions of centromeres in various organisms beginning with the diversity of their DNA sequence anatomies. We zoom out to describe their position in the nucleus and ultimately detail the different ways they contribute to genome organization and regulation at the spatial level.

Centromeres: Beyond Chromosome Segregation

Eukaryotic genomes are not randomly organized. Chromatin is organized into multiple domains juxtaposed one after the other along the length of a chromosome and positioned into particular layouts and in particular regions of the nuclear space [1,2]. Among these chromatin domains, centromeres are universal to all eukaryotic chromosomes. First cytologically described in 1882 by Walther Flemming as the primary constrictions on condensed chromosomes attaching to spindle fibers [3], centromeres are essential for cell division by ensuring the equal partitioning of DNA into daughter cells.

Although their crucial function comes into play during mitosis and meiosis, centromeres are not inert during interphase. Their importance in the genomes' 3D organization and regulation has only started to be recognized. In diverse eukaryotes, centromeres occupy particular domains in interphase nuclei. This nonrandom organization of centromeres in turn impacts chromatinbased processes such as transcription and replication. Here, we review centromere composition, localization, and interaction patterns across different eukaryotes. Simultaneously, we highlight conserved and divergent principles of linear and spatial centromeric architectures and their relationship with genome organization and function. For this, we survey observations from microscopy and more recent genome conformation studies to provide a systematic overview on spatial centromere organization in diverse eukaryotes. We also discuss the functional relevance of particular centromeric arrangements in interphase chromatin and emphasize the current limitations of and alternative strategies for genomic approaches in determining the spatial organization of centromeres.

Centromeres Are Enigmas of Many Genome Assemblies

Despite their essential function in mediating and controlling chromosome segregation, centromere architecture is remarkably diverse among different organisms. The length of centromeres ranges from as small as 120 bp to up to several megabases of DNA [START_REF] Agostinho | Sexual dimorphism in the width of the mouse synaptonemal complex[END_REF]. With some recent exceptions, sequence information is available only for shorter centromeres, whereas assemblies of larger centromeres are limited due to their length and repetitive sequence composition (Figure 1 and Box 1).

Highlights

Centromeres' primary role is to ensure chromosome segregation. In addition to this well-known function, centromeres also have an impact on genome architecture.

Microscopy studies have shown a propensity of centromere to cluster in many organisms. In addition, these clusters are not randomly localized in the nuclear space and tend to be found close to the nuclear periphery or around the nucleolus.

At a large scale, chromosome architecture is constrained by centromeric interactions and the position of the centromere along each chromosome.

Chromosome conformation capture experiments confirmed centromere clustering while shedding light onto unexpected consequences for genome spatial regulation: centromeres act as strong topological barriers preventing specific types of contacts between the two chromosomal arms.

Centromere sequences of many yeasts and fungal species have been characterized (Figure 1 and Box 1). Comparative genomic analyses show that centromere size and synteny (see Glossary) are mostly conserved within both the Saccharomycetaceae and Candida clades, pointing toward a common origin in each of the two clades. Nevertheless, centromere sequences have rapidly diverged, with very little or no detectable homology found between species [5][6][7][8][9][10]. In contrast to the budding yeasts, the well-studied fission yeast Schizosaccharomyces pombe has an incomplete genome assembly for two out its three centromeres, with the number of pericentromeric inverted repeats still unknown (Figure 1 and Box 1)(http://www.pombase.org). Compared with fungi, sequence assemblies of the megabase-long animal and plant centromeres, comprising satellite DNA interspersed with transposable elements, are even more limited and remain largely unresolved even after using long-read sequencing approaches [11][12][13]. Still, in some plant organisms, including Oryza sativa and Arabidopsis thaliana, the molecular structure of large centromeric regions for several chromosomes has been determined (Figure 1 and Box 1) [14][15][16][17][18].In addition, a recent study characterized the organization and sequence composition of all native Drosophila centromeres, confirming the structure unraveled by original mapping efforts on the Dp1230 minichromosome (Figure 1 and Box 1) [19,[START_REF] Albert | Whole-chromosome paints in maize reveal rearrangements, nuclear domains, and chromosomal relationships[END_REF]. Finally, the complete sequence of the first human centromere has also been released by assembling the higher-order-repeat (HOR) structure of the human chromosome Y satellite array (Figure 1 and Box 1) [21].

Note that some species, including chicken, horse, and potato, have repeat-based centromeres as well as repeat-free centromeres depending on the chromosome [22][23][24]. For instance, while most centromeres from the chicken macrochromosomes comprise chromosome-specifichomogeneous arrays of tandem repeats, chromosomes 5, 27, and Z comprise unique sequences.

Comparative genomic studies show that the latter are likely to represent evolutionarily more recent and 'immature' centromeres that could represent an intermediate state [25].Consistent with this, neocentromeres often form on sequences devoid of repetitive DNA [25] and tandem repeats have only recently been accumulating in some evolutionarily new centromeres. Regardless of their evolutionary framework, the nonrepetitive nature of these centromeres provides a unique opportunity allowing their unambiguous assembly.

Seeing Is Believing: Centromeres Follow a Distinct Nuclear Organization

While centromere assemblies are often limited, the location of centromeres can be visualized by the presence of the histone variant CenH3 (also called CENP-A in humans), which almost universally marks centromeric chromatin in eukaryotes [26]. Microscopy studies provided the first insights into centromere organization in interphase nuclei. In numerous fungal species, including Saccharomyces cerevisiae and S. pombe, centromeres cluster all along the cell cycle, which are visible as one large focus in the vicinity of the spindle pole body (SPB) and opposite the nucleolus (Figure 2) [27][28][29][30]. Nevertheless, this organization can vary in some fungi. In Cryptococcus neoformans, centromeres do not cluster in premeiotic cells and are instead separately positioned adjacent to the nuclear envelope (Figure 2). Only when cells progress toward mitosis do the centromeres gradually coalesce to a single cluster [START_REF] Rosin | Chromosome territory formation attenuates the translocation potential of cells[END_REF].

Chromosome organization in most plants has classically been divided into two main categories: Rabl and non-Rabl. The Rabl organization is characterized by: (i) all centromeres being restricted to one pole of the nucleus, opposite the nucleolus; and (ii) the telomeres being more dispersed in the opposite pole. This organization is observed in wheat, rye, barley, and oat root-tip cells (Figure 2) [32]. By contrast, sorghum and rice root cells adopt a non-Rabl conformation with centromeres and telomeres dispersed and intermingled in the nuclear area (Figure 2) [32]. Some species, like maize, have an intermediate conformation between Rabl and non-Rabl in nuclei from root-tip cells [32].Inaddition,l aterst udiesmodulated this original classification by reporting that chromosome organization can vary between different cell types. For example, most root Hi-C: variation of 3C in which biotin is incorporated at the ligation junction enabling it to be selected and to obtain genome-wide contact maps. Holocentric: referring to chromosomes that have multiple centromeres along their entire length, as opposed to monocentric where centromeres are restricted to one region on each chromosome.

Neocentromeres: new centromeres that form at an ectopic locus on the chromosome that is not centromeric. They typically arise due to the disruption of the endogenous centromere. Neocentromeres are often not associated with repetitive DNA but instead comprise unique sequences. Pericentromeric inversion: an inversion is a chromosome rearrangement in which a segment of a chromosome is reversed end to end. In a pericentric inversion, the segment includes the centromere. Synteny: conservation of the order of homologous genetic loci between two chromosomes.

nuclei of rice show non-Rabl organization, as described above, while in the xylem vessel cells centromeres and telomeres localize to opposite poles of the nuclei, consistent with a Rabl configuration [32,33].

Compared with the Rabl and non-Rabl conformations, A. thaliana has a distinct centromeric organization with (peri-)centromeric heterochromatin organized as clearly distinguishable chromocenters in the nucleus (Figure 2). These are not randomly positioned inside the nucleus and are found near the periphery or close to the nucleolus. Moreover, the number of chromocenters observed is often (in ~88% of observed nuclei) fewer than ten (corresponding to 2n =10 chromosomes), demonstrating a propensity to associate together [34].

Centromere clustering in interphase nuclei has also been observed in several animal model organisms. Counting the number of CENH3 CID -positive foci in interphase nuclei using immunofluorescence experiments demonstrated that centromeres cluster in flycells [35]. In particular, S2 aneuploid cells (13 stable chromosomes) show four to six clusters (Figure 2) and third-instar larval diploid (2n = 8) hemocytes show two or three clusters, which are localized close to the periphery of the nucleolus in both cell types.

Centromere clustering is also a common feature in human and mouse. Early studies on mouse B and T lymphocytes describe clustering of centromeres and their colocalization with

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Mus musculus

Homo sapiens

Saccharomyces cerevisiae

Candida albicans

Schizosaccharomyces pombe

Cryptococcus neoformans

Drosophila melanogaster When the centromere of a particular chromosome is depicted, its number is indicated after the species name and an approximate scale is shown on the right.

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chromocenters comprising pericentromeric heterochromatin (Figure 2) [36,37]. More recent analyses of nondividing/quiescent (G0) lymphocytes from mouse and humans revealed that centromeres cluster preferentially at the nuclear periphery, forming on average 13 or nine signals per human (2n = 46) or mouse (2n = 40) nucleus, respectively [38]. In the few cases of central localization, clusters are found to be associated with the nucleolus. Refining centromere locations within the nuclear region occupied by their corresponding chromosome [also called chromosome territory (CT)], centromeres are found at the periphery of CTs in 95% of studied cases and positioned toward the nuclear periphery. Other studies confirm and expand these results, showing that centromere clustering is a common feature observed to various extents in all cell types analyzed [39][40][41][42] but changes dynamically over the course of the cell cycle [41,42].The cell types analyzed include hematopoietic progenitors and mature blood cells (B and T lymphocytes, granulocytes, monocytes) as models for cell differentiation, as well as fibroblasts, which offer a comparison with a terminally differentiated cell from another lineage. On exit from mitosis, in early G1, very little to no clustering is observed, and the number of kinetochore signals decrease in late G1 and S phase indicating centromere clustering. Over the course of G2, centromeres separate out again as chromosomes start to condense in preparation for mitosis. In addition, peripheral localization and clustering of centromeres is less pronounced in the nuclei Box 1. Centromere Organization Is Diverse among Eukaryotes Saccharomyces cerevisiae 'point' centromeres are genetically defined by three genetic elements -CDEI, II, and IIIand incorporate a single centromere-specific histone H3 variant, CenH3 (Cse4) [99,100].

In the Candida clade, including Candida albicans, centromeres occur in large open reading frame (ORF)-free regions of 4-18 kb, which includes a CenH3-containing-nucleosome core region of 3-5 kb. They share no common sequence motif or repeats either within one organism or between them [5][6][7].

In Schizosaccharomyces pombe, the central core (cnt), of about 4 kb in length, assembles CenH3-containing-nucleosomes. It is surrounded by numerous alternating dg, dh, and cen253 elements forming innermost (imr) and outermost (otr) repeats, giving rise to centromeric regions of 35-120 kb [101]. A similar organization with a nonrepetitive mid-core Arabidopsis thaliana contains centromeres of several megabases with a central domain comprising mainly 180-bp satellite DNA and fragments of Athila retrotransposon elements, 106B. The flanking regions contain multiple families of LTR retrotransposons and 5SrDNA sequences [15,16,104].

Rice and maize contain SatDNA, named CentO (155 bp) and CentC (156 bp), respectively. Each also contains the centromere-specific retrotransposons CRM (maize) and CRR (rice), while in wheat CRW (wheat) is found but no SatDNA [14,17,18,[105][106][107][108].

Analysis of the Drosophila melanogaster centromere organization revealed that they comprise islands of complex DNA enriched in retrotransposons flanked by arrays of short satellite repeats (5-12 bp) [19,[START_REF] Albert | Whole-chromosome paints in maize reveal rearrangements, nuclear domains, and chromosomal relationships[END_REF]. Whether satellite or retrotransposon sequences are the functional parts of D. melanogaster centromeres based on CenH3 (Cid) enrichment remains, however, controversial [109].

Mouse chromosomes are all acrocentric, with centromeres comprising 120-bp minor satellites constituting the core centromeric region toward the telomere and the more abundant 234-bp major satellite forming pericentromeric heterochromatin [110,111].

The centromere of the human Y chromosome comprises 171-bp alpha-satellite monomers organized in a tandem fashion into large arrays of HOR [21]. While annotated alpha-satellite arrays are dominated by HORs on BAC clones conferring chromosome specificity [112], another study proposes that the core of human centromeres comprises two classes of highly homogeneous alpha-satellite dimers with HORs being pericentromeric [113].

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of cycling cells or progenitor cells compared with noncycling and terminally differentiated cells [41,42]. In particular, while centromere clustering is maintained in human embryonic stem cells, their peripheral localization is mainly lost, possibly due to rapid divisions of these cells [43]. Drosophila melanogaster [35]; Arabidopsis thaliana [34];

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Triticum aestivum and Sorghum bicolor [32]; Cryptococcus neoformans [START_REF] Rosin | Chromosome territory formation attenuates the translocation potential of cells[END_REF]; Candida albicans [30]; Schizosaccharomyces pombe [27]; Saccharomyces cerevisiae [115].Bar,5μm.

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In summary, while the insights gained from microscopy approaches are limited, these studies reveal that centromeres are often confined to or even clustered in specific nuclear locations in many organisms. Their organization within the nucleus, however, varies from one species to another.

Centromere Interactions Drive Genome 3D Organization and Regulation

Genome-wide chromosome architecture studies using high-throughput molecular biology techniques, like chromosome conformation capture (3C) and its derivatives (4C, Hi-C, etc.) offer new ways to study the importance of centromeres in genome organization. Early studies define the top of the genome organization hierarchy as the CTs [44]. In line with that, Hi-C data from various organisms demonstrate that intrachromosomal interactions are always more frequent than interchromosomal interactions. In many organisms, and especially in mammals, chromosomes can be further segregated into two compartments: the A compartment, which includes regions of the genome enriched in genes and histone modifications associated with active chromatin, and the B compartment, which is enriched in histone modifications associated with inactive chromatin [45]. Going down the chromosome organization to the scale of several hundred kilobases, one can find domains that interact more frequently within themselves than with neighboring regions, often referred to as topologically associated domains (TADs) [46]. In addition, other signatures of genome organization are visible on Hi-C maps and account for specific and strong landmarks of 3D genome organization. Here, the centromere is a classic example of a region with strong topological impact. Assembling data from many organisms published to date, we will discuss at least two implications of previously described centromere interactions visible on Hi-C matrices: (i) centromeres build a specific subcompartment inside the nucleus; and (ii) they form a barrier to intrachromosomal arm interactions.

Centromeres Form Strong Interacting Subcompartments 3D genome organization has been studied in several fungal species. The first S. cerevisiae full-genome contact map was published using a high-throughput 4C-derived experiment [47]. From this map, centromere clustering was reported to be the most striking feature of interchromosomal contacts, in accordance with the known Rabl-like organization of budding yeast chromosomes.

Besides that, centromeres engage in relatively few long-range intrachromosomal interactions (Figure 3A). These findings have been confirmed in later studies of other budding yeasts, filamentous fungi, and S. pombe [48][49][50][51][52]. Centromere clustering is such a prominent characteristic of yeast genome organization that the 3C signal has even been taken as a starting point to locate previously unidentified centromeres [53,54].

The remarkable plasticity of yeast genomes has allowed drastic remodeling of the location and number of centromeres. For example, selecting for neocentromeres in Candida albicans or manipulating the chromosome number by genome engineering in S. cerevisiae results in a drastic reorganization of genomic contacts [55][START_REF] Maeda | Chiasma Studies in the Silkworm, Bombyx mori L[END_REF][57]. In both cases, upon the loss of a centromere the strong interactions of flanking sequences with other centromeric regions disappear. In C. albicans, chromosomal regions near a neocentromere acquire new and strong interactions with all of the other seven centromeric regions. The impact of this reorganization on gene expression was also analyzed because coregulated genes often cluster along chromosome arms and in centromere-proximal regions [58,59]. In this case, however, the impact on gene expression upon centromere delocalization is marginal, at least under standard laboratory growth conditions. In addition, the reshuffling of centromere location might also interfere with DNA metabolism and recombination. Centromeres are among the earliest-replicated regions in the yeast genome [48,60,61] and negatively impact the rate of crossover during meiosis [62]. Future

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studies could therefore investigate the effects of centromere delocalization on other chromatin processes besides gene expression.

Centromeric and pericentromeric regions also drive specific physical contacts in interphase in other organisms as visualized on their respective Hi-C contact maps (Figure 3). In line with their Rabl or non-Rabl organization, two different types of contact map have been described in plants.

The contact map of barley shows a strong cross-shaped signal within and between chromosomes, consistent with their typical Rabl conformation [63]. By contrast, the contact map of rice, tomato, foxtail millet, and sorghum, which are described as non-Rabl, show tripartite segregation of their chromosomes, with centromeres and large pericentromeric heterochromatin grouped in a central B (inactive) compartment. Although interactions are enriched within all inactive compartments across the genome, they tend to be less involved in interchromosomal interactions than the two distal A (active) compartments [32,64]. Although maize was previously reported to have an intermediate organization, Hi-C analysis demonstrated a strong intra-and interchromosomal cross signal, indicating a Rabl organization similar to that in barley, at least in mesophyll protoplasts cells [32,64]. The genome-wide contact map of A. thaliana shows detectable interactions between all pairs of centromeric regions that make very few contacts with other genomic regions, except for the large heterochromatic locus on chromosome 4 called 'knob' (Figure 3B) [65]. Here, centromere clustering gives rise to a less-pronounced cross-shaped signal than observed in plants with Rabl conformation. Still, the strongest genome-wide Hi-C contacts of A. thaliana correspond to pericentromeric chromatin interactions, both within the same pericentromere and between different pericentromeres, consistent with the reported formation of heterochromatic chromocenters [65].

Centromere-centromere contacts are also visible on the Drosophila melanogaster Hi-C matrix, which shows strong interaction signals between the megabase-long heterochromatic pericentromeric regions (Figure 3C) [66]. To explain these interactions, as well as other general chromosomal organization in flies (see below), it is hypothesized that (peri)centromeric regions may assemble into their own nuclear compartment, again consistent with previous observations by microscopy [67].

Genome-wide Hi-C maps in vertebrates also show the presence of CTs, but centromere interactions are not as apparent (Figure 3D,E) [45,68]. The lack of strong centromere signals in vertebrate Hi-C contact maps is possibly due to weaker centromere interactions in these organisms. In addition, the lack of full centromere sequence assemblies might be a confounding factor. Satellite sequences are systematically excluded from most next-generation sequencing analyses due to their repetitive nature. Nevertheless, as the interaction signal spreads along neighboring chromatin fibers due to their physical properties, it is possible to recover the interaction encompassing repetitive regions by looking at adjacent nonambiguous regions. Using this principle to reanalyze human and mouse Hi-C data, it was found that decomposing the contact map signal into a series of genomic tracks, known as eigenvectors, enabled the determination of the contribution of particular genomic features to higher-order chromosomal organization [69]. They further determined that the first eigenvector relates to genomic sequences and local epigenetic chromatin states (as described before by the compartmentalization), while the second (or third in the case of the mouse) relates to the position along the chromosome arm and highlights centromere-centromere interactions. The association of all repetitive elements in the human and mouse genomes has also been studied using neighboring regions of ambiguous sequences as well as stringent and restrictive mapping parameters [70]. With these techniques, it was shown that some satellites significantly colocalize inside the nucleus, particularly alphasatellites in the human genome. Furthermore, by using higher-resolution maps of the human genome (down to 1-kb bins), the two original compartments that were described by the first eigenvector were refined and divided into five subcompartments, two in A and three in B [71]. Sixty-two percent of pericentromeric heterochromatin is found in subcompartment B2, which is associated with the nuclear lamina-and nucleolus-associated domains. Finally, superimposition of 3D genome structure determination from single-cell Hi-C data with microscopy visualizing CENP-A reveal a Rabl-conformation with centromeres and telomeres clustered at opposite sides of the nucleus in G1 mouse ES cells [72]. Taking these findings together, centromere clustering in vertebrates, although less apparent in Hi-C contact maps, still account for a prevalent class of contacts that have been demonstrated by many different means.

To overcome the problem of mapping repetitive sequences using a different strategy, centromere interactions were analyzed in chicken cells, taking advantage of the nonrepetitive nature of centromeres on chromosomes 5, 27, and Z, as well as engineered strains with neocentromeres at several positions on the Z chromosome [73]. As in humans, neocentromeres in the chicken are neither flanked by heterochromatin nor enriched for satellite repeat sequences [74][75][76].Using this setup, it has been shown that repetitive and nonrepetitive centromeres associate with one another in 4C analyses and that nonrepetitive centromeres and neocentromeres interact with heterochromatic regions of the genome.

Centromere clustering even extends to apicomplexans, single-celled organisms that include the human malaria parasite Plasmodium falciparum. Recent Hi-C analyses of several Plasmodium species and two additional apicomplexan parasites reveal varying degrees of centromere and telomere clustering in most developmental stages. However, centromere interactions appear to be lost in sporozoites, haploid cells that invade the insect salivary glands for transmission to the host [77,78].

To conclude, it is important to note that while the described features often arise from asynchronous cell populations, they are unlikely to be the result of interactions coming from the low fraction of mitotic cells as previously proposed in studies using synchronized cells [79][START_REF] Rasmussen | Chromosome interlocking during synapsis-a transient disorder[END_REF][81]. Mitotic chromosomes are devoid of compartments and TADs and enriched in intrachromosomal long-range contacts compared with interphase chromosomes. This particular folding makes them drastically different from interphase chromosomes that resemble and better recapitulate the chromosome conformation in exponentially growing cells. Taking these findings together, despite different linear genome architectures and centromere sequence compositions, centromere interactions appear to be a common feature in many organisms scattered throughout the eukaryotic phylogeny.

Centromeres Individualize Chromosome Arms

In addition to engaging in preferential interactions and building a specific subcompartment, centromeres tend to create a barrier within each chromosome resulting in less frequent contacts between the chromosome arms on either side of them compared with intra-arm contact frequencies (Figure 3F). This insulation property is visible on contact maps of species harboring a classical Rabl or Rabl-like chromosome conformation (Figure 3). Measurements in S. pombe, for instance, show more frequent intra-arm than interarm interactions, the latter being in the same range as interaction frequencies between chromosomes [50]. The barrier effect of centromeres also plays a role in species with other genome configurations. The first 3D studies in A. thaliana applying 4C on several viewpoints distributed across all five chromosomes demonstrate that chromosome arms are the main interaction units of the genome [82]. This result was later confirmed on genome-wide analysis showing a higher level of interactions between chromatin regions on the same side of a centromere than interactions on opposite sides [65].

Genomic rearrangements involving the centromere in mutant flies demonstrate the insulating properties of centromeres. Interactions between polycomb target genes in D. melanogaster wild type and in a mutant strain carrying a pericentromeric inversion on chromosome 3 has been analyzed using 4C. This demonstrated preferential interactions between these loci when present on the same chromosome arm. These findings imply strong rewiring of chromatin interactions in the rearranged strain consistent with a model of distinct territories formed by individual chromosome arms [67]. This model, however, has to be modulated depending on the developmental stage or tissue analyzed, because relative interactions between arms and chromosomes are reported to vary [66,83,84].

Analyses in human cells using a modified 3C technique called tethered conformation capture (TCC) revealed that the contact profile of inactive regions decreases abruptly when they are located on opposite sides of the centromere [85]. This effect, however, is not seen for active regions, which tend to be involved in long-range contacts regardless of the presence of the centromere [85]. These analyses demonstrate that the human centromere can also act as a contact barrier, at least for some categories of compartments.

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In addition, the position of the centromere on the chromosome constrains its overall architecture. This effect can be observed in S. cerevisiae, where the small and long chromosome arms tend to interact more frequently with one another, respectively [86,87]. Furthermore, the position of the centromere on the chromosome influences centromere-centromere interactions themselves.

For instance, the centromeric regions of human acrocentric chromosomes are more likely to contact each other than those of metacentric chromosomes [85].

Overall, the centromere influences interaction frequencies along the entire chromosome, preventing certain contacts between particular chromosomal regions. Variation in the position of the centromere can directly impact long-distance interactions and the general 3D conformation of chromosomes. It will be important to determine to what extend altered centromere locations can lead to perturbations in gene expression or other DNA-related metabolism in future studies.

Concluding Remarks and Future Perspectives

In this review, we provide a collective view on the spatial organization of centromeres in a range of distinct eukaryotes (Table 1, Key Table ). Despite their essential function, centromere architectures and sequences are highly diverse, yet in many organisms the spatial organization of centromeres appears to be conserved. Centromeres localize to distinct nuclear subcompartments or even cluster with one another in interphase nuclei. Given that the first layer of genome organization is the segregation of chromosomes into CTs with intrachromosomal interactions being the most frequent of all, centromere clustering across different chromosomes is not an intuitive result. This raises the questions of what leads to centromere clustering and whether there is a functional relevance to this organization (see Outstanding Questions).

Centromere clustering could be mediated by proteins that bind to centromeres or pericentromeres and stabilize long-range interactions between them. This mechanism could be similar to the formation of chromocenters in flies and mice that were proposed to be mediated by proteins bound to pericentromeric satellites and capable of bundling multiple DNA strands [88]. Future studies could aim to identify additional proteins involved in the formation of centromere clusters. The physical properties of centromere clusters could phase separate them into nuclear subcompartments as has been proposed to underlie A/B compartmentalization observed in Hi-C data [89] and the formation of heterochromatin foci bound by HP1 [90]. Kinetochore components are enriched in coiled-coil and disordered domains [START_REF] Westermann | Family matters: structural and functional conservation of centromere-associated proteins from yeast to humans[END_REF], which represent structural features frequently associated with protein assemblies that induce protein phase separation [START_REF] Woodruff | Organization and function of nondynamic biomolecular condensates[END_REF].

Several observations point toward a functional relevance of centromere clustering. Recent studies have hypothesized that the aggregation of chromosomes via chromocenters in interphase could prevent micronucleus formation and thus play a role in genome stability [88]. This is consistent with observations showing that the destabilization of spatial centromere positioning impairs silencing, which in turn leads to an increase in the transcription of transposable elements. This is associated with the accumulation of DNA double-strand breaks throughout the genome, leading to mitotic defects and genome instability [35,[START_REF] P E N G | .( 2 0 0 7 )H 3 K 9m e t h y l a t i o na n d RNA interference regulate nucleolar organization and repeated DNA stability[END_REF][START_REF] Mekhail | Role for perinuclear chromosome tethering in maintenance of genome stability[END_REF]. Additionally, centromere organization is subject to changes during the cell cycle and during various developmental stages including plant germination or floraison and mouse embryogenesis [START_REF] Simon | Structure and function of centromeric and pericentromeric heterochromatin in Arabidopsis thaliana[END_REF][START_REF] Martin | Genome restructuring in mouse embryos during reprogramming and early development[END_REF]. Several studies have also pointed out possible higher-order cluster organization in human cells, with specific combinations of centromere associations [39,40,85,[START_REF] Ioannou | A new model of sperm nuclear architecture following assessment of the organization of centromeres and telomeres in three-dimensions[END_REF][START_REF] Tjong | Population-based 3D genome structure analysis reveals driving forces in spatial genome organization[END_REF]. Hence, the question arises of whether such nonrandom centromere associations have functional implications for surrounding chromatin processes. However, given the dominant role of the cell cycle in determining genome architecture, temporal changes in cell-cycle stages will have to be taken into consideration when comparing centromere configurations across different organisms and developmental stages.

Outstanding Questions

How are centromeres organized and what is the identity of centromeric sequences?

What leads to centromere clustering and which factors are involved?

In particular, how is centromere clustering affected on the depletion of factors affecting chromatin organization and chromocenter formation, including DNA methyltransferases and methyl CpGbinding domain proteins, histone-modifying enzymes, and known architectural proteins?

How are centromeres localized within the nuclear space?

In particular, which factors enable centromere localization to specific nuclear compartments, including the nuclear lamina and the nucleolus?

What are the consequences of centromere interactions for genome architecture?

To what extent can the information on spatial genome architecture explain and predict the location of neocentromere formation?

Is there a functional relevance of centromere clustering for DNA metabolism, including gene expression or replication? How do changes in centromere organization or compaction (i.e. developmental, malignant, or evolutionary changes) lead to changes in 3D nuclear organization and what is the effect?

The position of the centromere on the chromosome together with its 3D organization also impacts the chromatin environment in its vicinity. It will therefore be interesting in future studies to investigate how genome architecture changes with centromere organization. This aspect could be addressed in organisms that have evolved neocentromeres or contain evolutionarily new centromeres by comparing their genetic environment with the ancestral state. Additionally, studies in organisms with drastically differing centromere organization, such as holocentric species, could also provide insights into the impact of centromeres on genome architecture.

Finally, a comprehensive picture on the conservation of 3D centromere conformation will require data from additional organisms, including non-model organisms. However, to do this we will have to overcome the challenges brought forth by the repetitive nature of many centromeres that result in incomplete genome assemblies. New and improved sequencing technologies will be required to obtain insights into centromere configurations in organisms with repetitive centromeres, including humans.

Key Table Table 1

Introduction

Precise homolog pairing and unpairing during meiosis is essential for genetic recombination and accurate chromosome segregation. Errors in chromosome segregation during meiosis can lead to reduced fertility, miscarriages, or chromosomal disorders in progeny, such as Down or Turner Syndromes [1]. Decades of research have gone into characterizing the synaptonemal complex (SC), a ubiquitous, proteinaceous structure that holds homologs together during meiotic prophase [2,3]. Yet how homologs find each other and come together in three dimensional (3D) space is still poorly understood. One of the main reasons that homolog pairing has remained such an enigma is the lack of cytological tools available for assaying chromosomeand locus-specific pairing dynamics during meiosis. Several recent studies have taken advantage of advances in super-resolution microscopy techniques, such as Structure Illumination Microscopy (SIM) and Stochastic Optical Reconstruction Microscopy (STORM), to visualize meiotic pairing in more detail than ever before [START_REF] Agostinho | Sexual dimorphism in the width of the mouse synaptonemal complex[END_REF][5][6][7][8][9]. However, these approaches have been limited to studying pairing genome-wide by fluorescently labeling elements of the SC [5,[7][8][9][10][11][12] or to visualizing small genomic loci by FISH [13][14][15][16].

Recent technological innovations in the design and synthesis of specialized DNA FISH probes called Oligopaints have made visualizing whole, individual chromosomes or complex sub-chromosomal loci in meiotic cells feasible. Unlike traditional BAC-based FISH probes, Oligopaints are computationally designed based on genome sequence data [17,18]. This approach allows for only unique, single copy sequences to be labeled, significantly increasing the specificity and resolution of FISH. Here, we leverage the flexibility of the Oligopaint design to add barcodes to label either whole chromosomes or different sub-chromosomal loci using the same set of oligos, as previously described [19]. This multiplexed approach allows for different highly specific FISH probes to be generated at low cost and high throughput. Oligopaints and related oligo-based FISH approaches have previously been used for karyotype analyses or characterization of interphase chromosome dynamics in Drosophila, C. elegans, mammals, and plants [16,[19][START_REF] Albert | Whole-chromosome paints in maize reveal rearrangements, nuclear domains, and chromosomal relationships[END_REF][21][22][23][24][25][26][27][28][29][30][START_REF] Rosin | Chromosome territory formation attenuates the translocation potential of cells[END_REF]. Recently, similar approaches have also been applied to the study of small chromosomal loci during meiosis [32,33], but Oligopaints have never before been used to characterize compaction and pairing of multiple, whole chromosomes during meiosis. Finally, this is the first study to use Oligopaints to visualize chromosomes in Lepidoptera (moths and butterflies).

Here, we combine Oligopaint DNA FISH with one of the first model systems used to study meiotic chromosomes, the silkworm moth. Bombyx mori are holocentric insects, with centromeres that form all along the chromosome during mitosis [34][35][36][37]. The holocentric mitotic configuration is also seen in many plants and nematodes, including C. elegans [38][39][40]. However, the holocentric chromosome configuration prevents accurate biorientation of bivalents formed after recombination and is therefore incompatible with canonical meiosis [40,41]. Instead, chromosomes in holocentric organisms often display "telokinetic" or "telokinetic-like" chromosomes during meiosis, where kinetochore activity is restricted toward telomere domains [40,[42][43][44][45][46][47]. In C. elegans, crossover position dictates which telomere faces poleward to connect to the spindle microtubules [42,45,[48][49][50]. A similar telokinetic mechanism for segregating meiotic chromosomes was also previously hypothesized to occur in B. mori [51][52][53] but has never before been directly observed. Furthermore, meiotic segregation in C. elegans occurs in the absence of the centromere-specifying factor Centromere Protein A (CENP-A) [50]. Instead, microtubules either run parallel to chromosomes to facilitate segregation or connect directly to cup-shaped kinetochores that form at chromosome ends [46,54]. Interestingly, CENP-A is entirely absent from the genomes of butterflies and moths [55]. How moths and butterflies segregate chromosomes during meiosis in the absence of CENP-A remains to be explored.

Unlike B. mori spermatogenesis, which has been reported to support crossovers and canonical pairing, oogenesis in B. mori is unconventional. Chiasmata are not observed in female meiosis in silkworms. Furthermore, the central elements of the SC break down just after pachytene (one of the sub-stages of meiotic prophase I), and the lateral elements of the SC are thought to be completely remodeled to form masses of protein (modified SC) between the two homologs [53,[START_REF] Maeda | Chiasma Studies in the Silkworm, Bombyx mori L[END_REF][57][58]. This modified SC is reported to be greater than one micron in width by the end of prophase I, thereby ultimately undoing tight end-to-end homolog pairing while still holding homologs together until anaphase I [58]. Thus, pairing along the entire length of the chromosomes is not expected after pachytene.

Our studies here illustrate that Oligopaints are robust and specific in germline cells and can be used to visualize chromosomes even in unconventional model systems with draft genomes. Using this FISH-based approach, we demonstrate that pairing in early meiotic prophase in the B. mori male germline is initiated at chromosome ends, with gene-rich chromosome ends more often initiating pairing. We further show that telomeric regions face poleward during metaphase I and act as localized kinetochores during male meiosis, and both telomeres on any given chromosome harbor the ability to act as local kinetochores. Additionally, using immunofluorescence combined with FISH, we reveal that the DNA-binding kinetochore protein Centromere Protein T (CENP-T) is present on meiotic chromosomes and likely facilitates microtubule attachments at metaphase I. Finally, using super-resolution microscopy combined with Oligopaints in the female germline, we show that homologs remain paired at telomere regions throughout meiotic prophase I even after remodeling of the SC. Overall, we provide the first extensive characterization of whole and sub-chromosome dynamics during meiosis in any species, thereby pioneering the use of Oligopaints as a tool for studying meiotic pairing and progression.

Results

B. mori Oligopaint design

To visualize chromosomes in the silkworm B. mori, we designed and generated Oligopaint libraries targeting six of the 27 autosomes and the Z sex chromosome. We chose chromosomes of varying size and gene density (Table 1) so we could explore whether these properties affect the dynamics of meiotic chromosomes. We were also interested in examining the sex chromosomes to determine whether they behave differently than autosomes in meiosis. Our Oligopaint libraries were designed using the Oligominer pipeline [18,59] based on the updated 2019 silkworm genome assembly [60]. Oligos were designed with 80 bp of homology and map to the genome only once (therefore only labeling unique, single copy sequences). As the W sex chromosome is largely composed of repetitive sequences, it was not suitable for the Oligopaint design strategy utilized here, and therefore we focused on autosomes and the Z chromosome. This Oligopaint design approach yielded a maximum probe density of approximately 3 oligos per kilobase (kb) of DNA. For most chromosomes in our analysis, we further reduced this density to 1 or 1.5 probes per kb (Table 1), a probe density that has been shown previously to be sufficient for whole chromosome paints [30]. The resulting oligos are fairly evenly distributed along each chromosome, with gaps in regions where repetitive sequences are more abundant (Fig 1A -1C). These oligo libraries were then multiplexed as described previously [19] with one or more barcode sequences to allow for the amplification of individual chromosomes, sub-chromosomal stripes, and/or active and inactive chromatin domains (Tables 1-5 andFigs 1B, 1C, andS1). In total, the libraries consisted of 191,536 oligos (designated as "primary oligos") up to 160 bp in length, which includes the 80 bp of homology, up to two unique 20 bp sub-chromosomal barcodes, and two 20 bp whole chromosome universal barcodes (Fig 1D). During the PCR amplification steps, secondary oligo binding sites are added to the primary oligos, to which fluorescently labeled secondary oligos anneal during the FISH protocol (Fig 1E; [21,30]). This method allows for increased flexibility when combining probes for multichannel imaging. Together, this multiplexed probe design combined with secondary oligo labeling both increases the efficiency of Oligopaint labeling and reduces the cost of Oligopaint synthesis.

B. mori Oligopaints are highly specific

As the B. mori genome used to design the Oligopaints is in a semi-draft state (assembled into chromosomes but with many unmapped contigs), we first tested the specificity of our B. mori chromosome paints using karyotype analysis. Due to the small, holocentric nature of silkworm chromosomes, mitotic chromosomes are highly compact, while chromosomes in meiotic prophase I (pachytene sub-stage; Fig 2A; reviewed in [61]) are more linear due to synapsis (S2 Fig) [62]. Therefore, meiotic chromosomes are better suited for karyotype analyses. As silkmoths have a very short adult lifespan (only 5-7 days), meiosis begins early in the larval stages [63]. Hence, meiotic chromosomes from late 4 th or early 5 th instar larval testes and ovaries were visualized using a squashing technique. This approach uses a hypotonic shock combined with physical squashing to burst cells and disperse nuclei onto the slide, retaining only a minimal amount of spatial and temporal information from the tissue. A detailed description of our meiotic squash protocol can be found in the Materials and Methods. Since homologs are paired during most sub-stages of meiotic prophase I (Fig 2A ), we expected a single fluorescence signal per chromosome for a given probe.

Using our whole chromosome paints, three chromosomes at a time were labeled on meiotic squashes from testes and ovaries. We were able to easily identify cells in the pachytene stage, when homologs are tightly paired end-to-end, using a combination of DAPI structure and chromosome morphology. Indeed, we observed singular and distinct fluorescence signals for S3, and S4).

In larval ovary squashes, we were also able to identify linear, paired pachytene chromosomes (Fig 2E). At this early pachytene stage, both oocyte-and nurse cell-destined germ cells pair their homologs. Subsequently during differentiation, nurse cell chromosomes begin to unpair. Post-pachytene nurse cells could be identified in our squashes by their unpaired but proximal, condensed chromosomes (S5 Fig; [57]). Interestingly, when we attempted to use these paints to characterize chromosome morphology in a B. mori ovary-derived cell line, BmN4, we found that our probes partially labeled multiple chromosomes, suggesting that the karyotype in these cells has undergone dramatic rearrangements and possible ploidy changes compared to the genome-derived strain (S6 Fig) . This observation further validated the specificity of our probes and their ability to detect translocations. Together, these data not only illustrate that our Oligopaint libraries are specific but also validate the B. mori genome assembly. PLOS GENETICS PLOS GENETICS

Detection of stripe and chromatin state sub-libraries

While the specificity of our whole chromosome paints indicated that the B. mori genome is accurately assembled at the chromosome level, we needed to validate the intra-chromosomal genome assembly and in turn, the specificity of our sub-chromosomal paints. For this, we again turned to pachytene squashes in the male germline, where the linear nature of chromosomes allowed us to verify the linear order of our probes. We started with the stripe sub-libraries, wherein selected chromosomes were sub-divided into 3 Mb or 1.5 Mb domains, depending on the chromosome (Table 2 andFig 1B). The first, middle, and last stripes were labeled with secondary oligos to visualize three stripes along each chromosome (Figs 1B and 3A-3E; telomere 1 (tel1), middle (mid), and telomere 2 (tel2), respectively). FISH with these stripe paints for ch15 in pachytene squashes from larval testes revealed a singular focus for each stripe, with the tel1, mid, and tel2 domains positioned in the predicted order along the linear chromosome (Fig 3A). This was also true for ch7, ch16, ch23, and chZ (Fig 3B -3E).

To test the specificity of our transcriptionally active and inactive chromatin domain paints, we labeled all three chromosomes with this barcode index (ch7, 15, and 16) at the same time on larval testes pachytene squashes. This analysis revealed three separate linear chromosomes with distinct banding patterns corresponding to the respective paint schematic (Figs 1 and3F). Together, these results indicate that the intra-chromosomal assembly for these chromosomes is highly accurate and our paints are specific for their target chromosomal domains.

Homolog pairing in early meiotic prophase is not synchronous

When examining the images of whole chromosome paints on meiotic squashes from testes, we noticed a wide variety of partially paired chromosome configurations (Figs 2F and 4A). We therefore attempted to quantify these configurations to determine the prevalence of each in order to ascertain whether they may be biologically relevant. We broadly classified the most common partially paired configurations as linear with loops, forked, circular, or lollipop (Fig 4A and 4B). Interestingly, the forked and circular configurations were observed most frequently for all analyzed chromosomes (Fig 4B). Furthermore, not all chromosomes within a single cell pair simultaneously, as many cells harbor some paired and some unpaired chromosomes (Fig 4C -4E). When analyzing three chromosomes at a time, less than 10% of cells show no pairing between any three examined chromosomes, while more than 40% show pairing between all three visualized homologous pairs (Fig 4D).

To determine if certain chromosomes more frequently initiate pairing earlier than others, we quantified which chromosome initiated pairing in the cells in which only one of the three labeled chromosome sets had begun to pair. This analysis revealed that large, gene-rich chromosomes (ch15, 17,[START_REF] Agostinho | Sexual dimorphism in the width of the mouse synaptonemal complex[END_REF]Fig 4E) more frequently initiate pairing before both small chromosomes (ch7 and ch16) and large, gene-poor chromosomes (ch23 and chZ). Notably, no cells were observed with chZ pairing before any given autosomal chromosome, suggesting sex chromosome pairing may be delayed compared to autosome pairing.

Next, we sought to use our stripe paints to better understand the circular and forked chromosome configurations. In addition to identifying pachytene cells (Fig 3), we were able to identify cells in all stages of meiosis I up to metaphase I as well as cells in interphase and mitosis using the stripe paints (Figs 5A and S7). By examining chromosome configurations in zygotene nuclei, we found that chromosomes in the forked configuration pair at a single telomere domain (Fig 5B), while chromosomes in the circular configuration harbor pairing at both Prophase I is typically subdivided into five distinct stages: leptotene, zygotene, pachytene, diplotene, and diakinesis. Briefly: in leptotene, replicated chromosomes are reorganized and compacted into a linear scaffold structure. In zygotene, synapsis begins between the homologous chromosomes. In pachytene, synapsis is complete (black dots represent the synaptonemal complex holding the homologs together). This stage is also when crossing over can occur. In diplotene, the homologs repulse, condense further, and the SC breaks down. The homologs remain attached via chiasma (crossovers). Finally, in diakinesis, chromosome condensation and cruciform bivalent formation is nearly complete as the cell prepares for metaphase I. The finding that both forked and circular chromosome pairing structures include paired telomere domains led us to predict that pairing may be initiated at chromosome ends. To test this hypothesis, we quantified the fraction of zygotene cells exhibiting pairing in only a single stripe domain (tel1, mid, or tel2) for four autosomes and chZ. In agreement with our prediction, we found a significant majority of cells harboring pairing at only one of the two telomere domains as opposed to pairing at only the middle domain (Fig 5F). Interestingly, some chromosomes showed a significant bias for pairing at one telomere domain versus the other, with ch15 pairing in the tel1 domain most often and ch23 pairing in the tel2 domain most frequently (Fig 5F). We investigated differences between the two chromosome ends on these chromosomes and found a significant bias in the distribution of genes along these chromosomes compared to the other chromosomes in our analyses, with gene-rich chromosome ends correlating with increased pairing initiation (Figs 5G and S8). This result, combined with our previous finding that large, gene rich chromosomes pair earlier, suggests that transcription may play an important role in pairing initiation in Bombyx. Interestingly, chZ does not follow this trend, as there is a slight bias in the distribution of genes toward the tel1 domain but a slight increase in pairing frequency of the tel2 domain. In agreement with our earlier finding that chZ pairs later than autosomes, this result suggests that sex chromosome pairing may be governed by different mechanisms than autosomal pairing, which may be transcription-based.

Telomeres face poleward at random and recruit CENP-T at metaphase I in larval testes

When analyzing pairing initiation with our stripe paints, we also noted that traditional cruciform bivalents are formed at metaphase I. These bivalents are highly reminiscent of those seen in meiosis in the nematode C. elegans, with one telomere remaining paired and the other telomere facing poleward (Fig 5H,reviewed in [44]). While this "telokinetic" chromosome configuration was previously hypothesized to occur in B. mori meiosis [51][52][53], it has never before been directly observed. In C. elegans, either telomere on any given chromosome can harbor kinetochore activity, and both do so with equal probability, depending on where crossovers form during meiotic prophase I [42,48]. Additionally, this telokinetic kinetochore activity is independent of the centromere-specifying histone CENP-A [46,50,54].

We first wanted to determine if both telomeres can also act as kinetochores at metaphase I in B. mori. One possibility is that the pairing bias we observed for prophase I persists through metaphase I such that gene-rich telomeres are more likely to remain paired and thus genepoor telomeres are more likely to be functional kinetochores. To test this hypothesis, we analyzed metaphase I bivalents in larval testes squashes using our stripe paints for ch7, 15, 16, 23, and Z. Quantification of telomere orientation at the metaphase plate revealed that approximately half of metaphase I cells harbor pairing in the tel1 domain and half harboring pairing . The unpaired telomere domain faces poleward, suggesting that like nematodes, B. mori telomere regions likely act as localized kinetochores during meiosis. This result indicates that, while pairing initiation shows a bias toward gene-rich telomeres, the orientation of chromosomes in the metaphase I bivalent is not biased, with both telomeres having an equal probability of either remaining paired or facing poleward.

Next, we repeated the experiment using ch7 stripe paints in whole mount late 5 th instar larval testes to look at metaphase I telomere orientation in the native cellular context. This less invasive whole mount approach also allowed us to maintain critical spatial and temporal information and to better visualize germline development as a whole. As predicted based on our above and previous findings [53], in addition to meiotic cells, 5 th instar larval testes also harbor mitotic cells that have unpaired homologs that highly resemble those seen in whole-mount embryos and testes squashes (Figs 6A-6C1, S7, and S9) and primary spermatocytes at all stages of meiosis I (Fig 6A -6D). Interestingly, B. mori and other Lepidopteran insects utilize two distinct spermatogenic pathways, ultimately resulting in apyrene sperm (without nuclei, which are thought to act as support cells for sperm migration [64]) and eupyrene sperm (with nuclei [64][65][66][67][68][69][70]). In whole-mount testes, we were able to clearly identify eupyrene secondary spermatocyte bundles (Fig 6D3 ) and mature eupyrene sperm (S10 and S11 Figs). Additionally, we identified secondary spermatocyte bundles apparently apyrene-destined, where some cells have no DNA, and the FISH signal is instead diffuse in the cytoplasm. This observation suggests that the cells in these bundles are beginning the process of nuclear degradation (Fig 6D4). Importantly, quantification of metaphase I bivalent formation in whole mount testes was completely in agreement with our findings from squashes, showing ch7 tel1 paired in 47% and ch7 tel2 paired in 53% of cells (Fig 6D2).

Finally, we wanted to interrogate how these bivalents attach to the meiotic spindle. While CENP-A is absent from the B. mori genome [55], mitotic kinetochore recruitment instead requires the DNA-binding protein CENP-T [34]. To determine if CENP-T is present on meiotic chromosomes or if microtubule attachment is independent of CENP-T, we performed coimmunofluorescence/FISH with anti-CENP-T and anti-Tubulin antibodies plus ch23 Oligopaint on dissociated cell cytospreads from late 4 th instar larval testes. This cytocentrifugebased approach allowed for clearer visualization of kinetochore/microtubule attachments, as chromosomes become more spaced during the centrifugation process (S12A and S12B Fig) .  FISH for a single chromosome enabled us to distinguish between meiotic and mitotic cells (one versus two FISH signals). This experiment revealed that indeed, CENP-T is present on meiotic chromosomes and therefore may facilitate attachment to microtubules. Interestingly, using super-resolution microscopy, we found that these kinetochores are more canonical than the cup-shaped kinetochores observed in C. elegans meiosis [50], with a single kinetochore structure forming at the microtubule-proximal end of each chromatid (S12C Fig) . Together, these findings suggest that B. mori chromosomes form traditional bivalent structures at metaphase I with localized CENP-T-based kinetochore activity restricted to one telomeric region at random. completely paired while ch16 remains unpaired. Dashed line approximates the nuclear edge. (D) Quantification of pairing initiation for three chromosomes at a time in cells with zygotene DAPI morphology and linear chromosome morphology. (E) Quantification of which chromosome is initiating pairing in nuclei where only one of the three visualized chromosomes is partially paired (while the other two chromosomes show no contact between homologs). Chromosomes are ordered smallest to largest. Gray outline indicates gene-poor chromosomes, while black outline indicates gene-rich chromosome. Statistics, Fisher's Exact Test comparing pairing initiated versus uninitiated. For all quantification, ��� p<0.0001, �� p<0.001, � p<0.01. All data were collected from n = 2-3 testes squashes per chromosome.

https://doi.org/10.1371/journal.pgen.1009700.g004

Homolog pairing at chromosome ends is persistent in B. mori female larval ovaries

In contrast to what we and others have observed in B. mori males, homolog pairing in females is reported to be unconventional, without chiasma formation and with the SC partially breaking down and transforming into a large proteinaceous mass greater than one micron in width between the homologs before metaphase I [53,[START_REF] Maeda | Chiasma Studies in the Silkworm, Bombyx mori L[END_REF][57][58]71]. Despite these previous observations, we found a significant number of nuclei with homologs paired end-to-end in late 4 th /early 5 th instar larval ovary squashes (Figs 2 andS4). This finding led us to wonder whether homolog pairing is more stable in B. mori female meiosis than previously appreciated.

To better visualize homolog pairing and the linear progression of female meiosis, we performed whole-mount DNA FISH with ch7 stripe paints in 5 th instar larval ovaries and used super-resolution microscopy to measure the distance between homologous copies. Like Drosophila, the B. mori larval ovary is composed of polytrophic meroistic ovarioles (where the oocyte is part of a group of inter-connected cells including supportive nurse cells). Each ovariole contains a linear arrays of developing egg chambers, with the tip (germarium) harboring germline stem cells that are mitotically dividing and the most mature chambers being the most distal from the stem cell niche ( [63,72,73], Figs 7A andS13). However, as moths have a much shorter adult lifespan than flies (only 5-7 days for silkmoths versus 2-3 months for Drosophila), the majority of oogenesis occurs in the larval and pupal rather than adult stage [63]. Therefore, 5 th instar larval ovaries harbor oocytes in all stages of pairing.

Interestingly, despite our diffraction-limited imaging of ovary squashes appearing to show tight homolog pairing in early pachytene and late pachytene (Figs 2, 7, and S4), super-resolution imaging of homologs in the zone of pairing in the germarium (the early stages of pachytene) revealed that homologous chromosome copies are approximately 0.15-0.20 μm apart at this stage (Fig 7B and7E). This configuration is a much looser pairing association than we observed in larval testes, which harbor closely paired homologs with no detectable separation even at super-resolution (S14 Fig) . By late pachytene, when nurse cells and the developing oocytes begin to differentiate, ch7 homologs are even further separated (0.25-0.30 μm, Fig 7C and7E). In the first egg chambers most proximal to the germarium, chromosomes are still pachytene-like in structure, but homologs are approximately 0. . The majority of oocytes in the developing egg chambers outside the germarium in 5 th instar larvae are arrested in late diakinesis [START_REF] Maeda | Chiasma Studies in the Silkworm, Bombyx mori L[END_REF], which should be after transformation of the SC at the end of pachytene. In agreement with this hypothesis, super-resolution imaging revealed that in oocytes in the most mature egg chambers most distal from the germarium, homologs remain linear but are approximately 0.45-0.55 μm apart, with the two homologous copies connected only at telomere domains, forming a large chromosomal loop . This finding suggests that in the absence of chiasma and after the partial breakdown and transformation of the SC, contacts between homologous chromosomes persist throughout meiotic prophase I in female B. mori. These contacts appear to be mediated by both telomeric pairing mechanisms possibly in addition to any remaining modified SC, revealing that homologs are indeed closer together than the predicted one micron separation.

Discussion

The silkworm, B. mori, was a model system for studying meiotic chromosomes for decades. Like Drosophila, silkworms are readily reared in a laboratory setting and amenable to genetic manipulation including RNAi and CRISPR. However, unlike fruit flies, silkworms are large in size, combining the increased ease of dissection and structure visualization commonly associated with mammalian models with the short generation time of an insect system. The large amount of tissue provided by B. mori increases the feasibility of genomics assays and other cell population-based approaches, which generally require a large number of cells. Importantly, the recent sequencing of the B. mori genome revealed that there is a high degree of sequence homology between silkworm genes and mammalian disease genes [60,[74][75][76]. Furthermore, B. mori harbor 28 chromosomes while humans have 23 (Drosophila have only 4), and our studies along with others have illustrated that, like in mammals, somatic homolog pairing is largely absent in B. mori [77]. This observation is in stark contrast to the high levels of somatic homolog pairing seen in Drosophila [78,79], making the study of B. mori genome dynamics more directly relevant to human biology.

Our studies examine pairing initiation in great detail in larval testes. We found that pairing is asynchronous in B. mori, with long, gene-rich chromosomes more frequently initiating pairing before their shorter or gene-poor counterparts. Interestingly, in zygotene nuclei, we observed a wide variety of partially paired chromosome configurations, many of which include (1, mitotic zone) and spermatocytes in all stages of meiosis up to mature sperm; progressing from right to left in the image as indicated: (2) meiosis I/primary spermatocytes, (3) meiosis II/secondary spermatocytes. Additionally, each lobe is surrounded by somatic cells in the sheath (4) and in the septae separating the lobes. (5)(6) prophase I: (5) leptotene/zygotene, (6) pachytene/diplotene, (7) metaphase/anaphase I, (8) prophase/metaphase II, (9) spermatids, (10) loop-like structures. We suspect that some of these loops (such as the "linear with loops" configuration) represent interlocked chromosomes as previously observed in Bombyx [START_REF] Rasmussen | Chromosome interlocking during synapsis-a transient disorder[END_REF]81], while others, like the highly abundant "circular" and "forked" configurations, represent pairing intermediates. It is also worth noting that "pairing" as measured by Oligopaint signal colocalization, even at approximately 140 nm x-y super-resolution, may not necessarily indicate SC formation and true synapsis (and we currently do not have SC antibodies for B. mori). We believe it is possible, and even likely, that there is a chromosome-wide "search" leading to transient colocalization of various chromosome regions (including the middle domain of chromosomes), but SC formation may always be initiated at telomeres, as in C. elegans [48,[82][83][84].

Importantly, we found that chromosome ends more frequently pair before the central domain of both autosomes and chZ. Additionally, we observed a bias toward gene-rich chromosome ends initiating pairing earlier than gene-poor chromosome ends of autosomes. Together, these data support a model in which transcription, the transcription machinery, or active histone modifications modulate pairing initiation in Bombyx. This study is not the first report linking transcription with homolog pairing. In fact, somatic pairing in Drosophila is largely influenced by transcription, and furthermore, transcription factories have been hypothesized to facilitate meiotic pairing in other species [85][86][87]. Still, whether or not telomeric genes are even transcribed in meiosis is unclear as meiotic transcriptome profiles for B. mori have yet to be established, and thus, the exact mechanism behind pairing initiation in Bombyx and beyond remains unclear.

Our FISH-based approach further reveals how chromosomes compact after prophase I and partially unpair when aligned at metaphase I. While crossing over has been reported in male meiosis in B. mori [52,[START_REF] Maeda | Chiasma Studies in the Silkworm, Bombyx mori L[END_REF], clear chiasmata were not apparent in post-pachytene spermatocytes (Figs 2,[START_REF] Agostinho | Sexual dimorphism in the width of the mouse synaptonemal complex[END_REF]and 5). We suspect this inability to detect chiasmata is likely due to the small size and compact nature of B. mori chromosomes in diplotene. We also show that mitotic chromosomes in B. mori, which are holocentric in structure, align parallel to the metaphase plate, with both telomeres aligned with the plate and homologs remaining unpaired. Our studies further illustrate that, like those in C. elegans, B. mori chromosomes do not retain the holocentric configuration in meiosis. Instead, meiotic chromosomes at metaphase I in spermatogenesis align perpendicular to the metaphase plate such that telomeric regions face the spindle poles, recruit CENP-T, and act as localized kinetochores. Super-resolution imaging of meiotic chromosomes at metaphase I revealed that these CENP-T-based kinetochores do not resemble the cupshaped kinetochores in C. elegans [50], but instead appear to be more canonical in structure, with a single kinetochore forming at the telomere of each chromatid.

Moreover, we demonstrate that both telomeres are equally likely to face poleward and harbor kinetochore activity. A similar telokinetic mechanism for meiosis has also been observed in the holocentric milkweed bug Oncopeltus fasciatus [88] and the kissing bug Triatoma infestans [89]. Whether crossover position dictates bivalent structure in B. mori or other holocentric insects, as in C. elegans [42,45,[48][49][50], remains to be explored. Interestingly, the broadly conserved centromere-specific histone H3 variant CENP-A is absent from the genome of Lepidopteran insects [55], suggesting that both mitotic and meiotic chromosome segregation in Bombyx occur independently of CENP-A.

We combined our Oligopaints with super-resolution microscopy to measure chromosomewide pairing in B. mori female meiotic prophase. We show that homologs are never paired as tightly in female meiosis as they are in male meiosis, in agreement with lack of chiasma formation in ovaries [53,[START_REF] Maeda | Chiasma Studies in the Silkworm, Bombyx mori L[END_REF]57]. Additionally, we find that after transformation of the SC at the end of pachytene, homologs appear to remain linked through telomere regions. Previous studies also suggest that modified SC remaining at this stage may play a role in keeping homologs together until anaphase I [58]. Yet, the exact mechanism facilitating and maintaining pairing in female meiosis in B. mori remains to be elucidated. Finally, the studies presented here illustrate that Oligopaints can be designed using, and act as validation for, draft genome assemblies, and demonstrate the feasibility of using Oligopaints to study meiotic chromosomes. Using this approach, we reveal novel insights into pairing initiation and meiotic chromosome segregation that may be conserved across species.

Materials and methods

B. mori strains and cell line

Embryos were obtained from Carolina Biological (Burlington, NC), Coastal Silkworms (Jacksonville, FL), Mulberry Farms (Fallbrook, CA), or were freshly laid in the lab by adults derived from embryos from these sources. Some larvae were obtained from Rainbow Mealworms (Compton, CA). Embryos were kept at 4˚C for less than 1 mo. For rearing, embryos were transferred to 28˚C, and larvae were fed fresh mulberry leaves or powdered mulberry chow (Carolina Biological or Rainbow Mealworms). BmN4 cells are commercially available from ATCC (Manassas, VA).

Oligopaint design and synthesis

Oligopaint libraries were designed as described in the main text. Active and inactive domains were determined primarily based on CENP-T depletion or enrichment. CENP-T ChIP-seq profiles were obtained from BmN4 cells, and domains were called as previously described [37] with the following modifications: CENP-T ChIP-seq signal originally in 10 kb windows was averaged over 50 kb. Subsequently, negative CENP-T domains were subtracted from positive CENP-T domains to obtain final CENP-T depleted domains. As previously observed, domains enriched for CENP-T were shown to strongly correlate with enrichment for the repressive histone mark H3K27me3, while domains depleted of CENP-T were shown to strongly correlate with enrichment of the active chromatin marks H3K4me3 and H3K36me3. All information regarding genomic coordinates for Oligopaints and probe density can be found in Tables 12345. Oligo pools were purchased from CustomArray/GenScript (Redmond, WA; ch 7,15,16) or Twist Biosciences (San Francisco,CA;ch 4,17,23,Z). Oligopaints were synthesized as previously described by adding barcodes to each oligo for PCR-based amplification [17,30,90].

Preparation of meiotic squashes and DNA FISH

For meiotic squashes, late 4 th instar or early 5 th instar larvae (approximately 3 in long) were sacrificed by decapitation. The caterpillars where then cut open anterior to posterior and fileted on a silicone dissecting dish using standard sewing needles. Gonads were harvested using forceps and placed into 1.5 mL tubes containing sf-900 (Gibco/ThermoFisher, Waltham, MA) tissue culture media. Gonads were then rinsed thrice in 1X PBS, then incubated in 0.5% sodium citrate for 8-10 min. Using forceps, gonads were then transferred to siliconized coverslips (1 gonad per coverslip) and covered with ~10 μL of 45% acetic acid/1% PFA/1X PBS and fixed for 6 min. Using a poly-L-lysine coated glass slide, gonads were then physically squashed and slide/coverslip were flash frozen in liquid nitrogen. After carefully removing slides from liquid nitrogen, coverslips were removed with a razor blade, and slides were postfixed in cold (pre-chilled to -20˚C) 3:1 methanol:glacial acetic acid for 10 min. After fixation, slides were washed thrice in 1X PBS and subjected to an ethanol row at -20˚C (70%, 90%, 100% ethanol, 5 min each) before drying completely at room temp. Slides were dried for 24-72 h.

FISH on meiotic squashes was performed as previously described for mitotic spreads [START_REF] Rosin | Chromosome territory formation attenuates the translocation potential of cells[END_REF]. Briefly, after drying slides, slides were denatured at 72˚C for 2.5 min in 2xSSCT/70% formamide before again drying with an ethanol row at -20˚C. Slides were then left to air dry for 10 min at room temperature. Primary Oligopaint probes were resuspended in hybridization buffer (10% dextran sulfate/2xSSCT/50% formamide/4% polyvinylsulfonic acid), placed on slides, covered with a coverslip, and sealed with rubber cement. Slides were denatured on a heat block in a water bath set to 92˚C for 2.5 min, after which slides were transferred to a humidified chamber and incubated at 37˚C overnight. The next day, coverslips were removed using a razor blade, and slides were washed as follows: 2×SSCT at 60˚C for 15 min, 2×SSCT at RT for 15 min, and 0.2×SSC at RT for 5 min. Fluorescently labeled secondary probes were then added to slides, again resuspended in hybridization buffer, covered with a coverslip, and sealed with rubber cement. Slides were incubated at 37˚C for 2 h in a humidified chamber before repeating the above washes. All slides were stained with DAPI and mounted in Prolong Diamond (Invitrogen/ThermoFisher, Waltham, MA). Slides were cured overnight before sealing with clear nail polish and imaging.

FISH on whole-mount gonads and embryos

For whole-mount DNA FISH in gonads, ovaries and testes from late 5 th instar larvae (after cessation of eating) were dissected in sf-900 cell culture media, washed thrice briefly with 1X PBS, and then fixed for 30 min in 4% PFA in PBS with 0.1% Triton-X-100 (0.1% PBS-T) at RT. Gonads were then washed again thrice in 1X PBS and permeabilized with 0.5% PBS-T for 15 min at RT. Gonads were pre-denatured by washing as follows: 2xSSCT for 10 min at RT, 2xSSCT/20% formamide for 10 min at RT, 2xSSCT/50% formamide for 10 min at RT, 2xSSCT/50% formamide for 3 h at 37˚C, 2xSSCT/50% formamide for 3 min at 92˚C, 2xSSCT/ 50% formamide for 20 min at 60˚C. To the 2xSSCT/50% formamide, 100 pmol of each probe was directly added. Gonads were then denatured for 3 min at 92˚C and incubated overnight at 37˚C. The next day, gonads were washed: 3x 30 min each in 2xSSCT/50% formamide at 37˚C, 1x 15 min in 2xSSCT at RT. 20 pmol of each secondary oligo was added with 50% formamide and incubated for 3 h at 37˚C. Final washes were performed (2x 30 min in 2xSSCT/50% formamide at 37˚C, 1x 10 min in 2xSSCT/50% formamide at RT, 1x 10 min in 2xSSCT/20% formamide at RT, 1x 10 min in 2xSSCT at RT), gonads were stained with DAPI, and mounted on slides with Prolong Diamond (Invitrogen/ThermoFisher, Waltham, MA).

For whole-mount embryo FISH, diapausing embryos were removed from 4˚C and kept at RT for 3-5 d. Chorions were weakened by soaking in 50% bleach for 15 min and then manually removed with forceps. Embryos were subsequently fixed for 30 min in 4% PFA in 0.1% PBS-T at RT, and FISH was performed as described above for whole-mount gonads.

IF-FISH on cytocentrifuge testes spreads

Testes were dissected as described above in sf-900 tissue culture media, followed by three quick washes in 1X PBS, then incubated in 500 μL 0.5% sodium citrate for 8-10 min in 1.5 mL tubes. Gonads were then gently crushed with a p1000 pipet tip, and cells were dissociated by gently pipetting up and down. 250 μl of the dissociated cells were added to a cytofunnel containing a poly-L lysine-coated slide. Cytocentrifugation (Shandon Cytospin 4, ThermoFisher, Waltham, MA) was performed for 5 min at 500 rpm with high acceleration. Slides were then fixed in 4% PFA for 10 min followed by three washes in 1X PBS. Cells were subsequently permeabilized in 100% methanol at -20˚C for 20 min, and then in 0.5% PBS-T for 15 min at RT. IF was then performed by blocking slides in 5% milk for 1 h at RT followed by incubation with primary antibodies diluted 1/500 in 5% milk overnight at 4˚C. Custom anti-CENP-T [34] and commercial anti-Tubulin antibodies were used (MilliporeSigma T6074, Burlington, MA).

The next day, slides were washed thrice in 0.1% PBS-T followed by incubation with secondary antibodies diluted 1/500 in 5% milk (goat anti-mouse alexa 546 and goat anti-rabbit alexa 488, ThermoFisher). After antibody incubation, slides were washed 3X in 0.1% PBS-T and post-fixed with 4% PFA for 10 min before proceeding with FISH as previously described [30].

Meiotic staging

Stages of meiosis were determined based largely on DAPI morphology, chromosome morphology, and/or cell position in whole-mount gonads. Leptotene/zygotene chromatin (DAPI) is a dense lampbrush structure, with chromosomes being linear but unpaired in leptotene and linear but partially paired in zygotene. Pachytene chromatin is a loose lampbrush with chromosomes being linear and completely paired from end to end. Diplotene/diakinesis chromatin is diffuse with chromosomes being paired or partially paired, nonlinear, and condensing. Metaphase I chromatin is highly condensed and aligned at the metaphase plate, with chromosomes being paired in a bivalent structure and highly condensed. Late 4 th -early 5 th instar male larvae were used for squashes as they only possess primary spermatocytes in meiosis I. In whole-mount testes, meiosis I and II were distinguished based on position in the gonad and based on the number of cells per bundle (with meiosis I bundles harboring approximately 64 cells and meiosis II bundles harboring approximately 128 cells).

Mitotic spreads from BmN4 cells

To induce mitotic arrest, approximately 1 × 10 5 cells were treated with 0.5 μg/mL Colcemid Solution (Gibco/ThermoFisher) for 2 h in a 28˚C heat block. Cells were then spun for 5 min at 600x g at RT to pellet and resuspended in hypotonic solution (500 mL of 0.5% sodium citrate). Cells were incubated in hypotonic solution for 8 min at RT. 100 μL of the cell suspension was then placed in a cytofunnel and spun at 1200 rpm for 5 min with high acceleration using a cytocentrifuge (Shandon Cytospin 4; ThermoFisher). For FISH, slides were then fixed in cold 3:1 methanol: acetic acid for 10 min and washed 3X 5 min in PBS-T (PBS with 0.1% Triton X-100). FISH was performed as described above for meiotic squashes.

Imaging, quantification, and data analysis

Widefield images of meiotic squashes and cytocentrifuge meiotic cells were acquired on a Leica DMi6000 wide-field inverted fluorescence microscope using an HCX PL APO 63x/1.40-0.60 Oil objective (Leica Biosystems, Buffalo Grove, IL), Leica DFC9000 sCMOS Monochrome Camera, and LasX software. Whole mount images were acquired using a Zeiss LSM 780 point scanning confocal (Zeiss Microscope Systems, Jena, Germany) with high sensitivity 32 anode Hybrid-GaAsP detectors. BmN4 mitotic spreads were acquired on a Zeiss AxioObserver Z1 wide-field inverted fluorescence microscope with 100x/1.4 oil Plan-APO objective, a Hamamatsu C13440 ORCA-Flash 4.0 V3 Digital CMOS camera, and ZEN blue software. Super-resolution imaging of cytospin meiotic spindles and CENP-T immunofluorescence was performed using a Nikon CSU-W1 SoRa spinning disk confocal microscope (Nikon Instruments, Inc, Melville, NY), controlled by Nikon Elements AR software ver 5.21.02. Images were collected using z-stacks of multi-channel fluorescence. The image detectors used were dual Hamamatsu Fusion BT (model C14440-20UP) cameras. SoRa super-resolution mode was used with objective lens Nikon CFI Apo TIRF 60x/1.49 Oil DIC N2, and intermediate SoRa magnification 4.0x, producing a pixel size of 27 nm. The main excitation/emission dichroic beamsplitter used was a Semrock Di01-T405/488/568/647 (IDEX Health & Science, LLC, Rochester, NY), and the image-splitter dichroic was 'DM A561 LP'. All emission filters were made by Chroma Technology Corporation (Bellows Falls, VT). Alexa Fluor 647 was excited by the 640 nm laser and emission filtered by ET655lp. Alexa Fluor 546 was excited by the 561 nm laser and emission filtered by ET605/52m. Alexa Fluor 488 was excited by the 488 nm laser and emission filtered by ET520/40m. DAPI was excited by the 405 nm laser and emission filtered by Semrock FF02-447/60. Post-processing of the images was performed in Nikon Elements AR software and included denoising via Denoise.ai and 3D deconvolution in automatic mode. Super-resolution imaging of Oligopaints in whole mount ovaries was performed using a Zeiss LSM 880 Airyscan with 63x/1.4 oil Plan-APO objective. Airyscan images were acquired in SR mode using the Zeiss 32 channel Airyscan detector, and images were processed under auto strength setting with Zen black software. Leica and Zeiss-acquired images were processed using Huygens deconvolution software (SVI, Hilversum, Netherlands). For all images, tiffs were created in ImageJ. All quantification was performed manually. All data are reported in S1 File.

  The compartments were characterized by comparing the Pearson correlation map, along with the eigenvector, to epigenetic, gene occupancy and DNase sensitivity data. From this you could see that the positive signal in the eigenvector, or compartment A, correlated with gene-rich and DNAse sensitive regions (open chromatin). With analyses at higher resolution, it was also shown that this compartment was correlated with H3K36me3 enrichment. Taken together, compartment A was described as open, accessible, actively transcribed chromatin. Compartment B was also described by comparing it to the same data as compartment A. What was gathered was that it was correlated with gene-poor and closed chromatin regions, based on DNase I

  the first time a third genome-wide compartment has been identified via Hi-C. Aside from the A and B compartment, a third compartment called the intermediate (I) compartment was identified when studying B-cell differentiation. It is called the I compartment because of its intermediate first eigenvector coefficient score, and it was also shown to be enriched in H3K27me3 (Vilarrasa-Blasi

Figure 1 .

 1 Figure1. Sequence Organization of Centromeres in Eukaryotes. For each species, the typical centromere organization is shown, with the size range for features of interest. When the centromere of a particular chromosome is depicted, its number is indicated after the species name and an approximate scale is shown on the right.

( 2 -

 2 5 kb) flanked by inverted repeats (2-5 kb) is found in the phylogenetically distant yeast Candida tropicalis [102]. In Cryptococcus species, centromeres are in syntenic positions in ORF-free, poorly transcribed regions featuring variable combinations of retrotransposons or their remnants. Centromeres span from 44 to 62 kb on average in C. neoformans and C. deneoformans, respectively, and about 14 kb in Candida deuterogatii [103].

Figure 2 .

 2 Figure 2. Schematic of Centromere Localization in Interphase. Blue, DAPI; red, centromere probe by immunofluorescence (IF) or fluorescence in situ hybridization (FISH) as indicated. Adapted from: Mus musculus [114];Drosophila melanogaster[35]; Arabidopsis thaliana[34]; Triticum aestivum and Sorghum bicolor[32]; Cryptococcus neoformans[START_REF] Rosin | Chromosome territory formation attenuates the translocation potential of cells[END_REF]; Candida albicans[30]; Schizosaccharomyces pombe[27]; Saccharomyces cerevisiae[115].Bar,5μm.

Figure 3 .

 3 Figure 3. Normalized Contact Maps. (A) Saccharomyces cerevisiae: full genome, resolution 5 kb, dataset from [86]. (B) Arabidopsis thaliana: full genome, resolution 20 kb, dataset from [65]. (C) Drosophila melanogaster: full genome, resolution 100 kb, dataset from [84]. (D) GM06990 human cell line: chromosome 3-5, resolution 1 Mb, dataset from [45]. (E) Chromosome 3 as in (D): top half, normalized contact map; bottom half, correlation matrix. Contact maps have been generated using the HiC-Pro suite, normalized by ICE using program default parameters [116] and drawn using HiCExplorer [117]. The correlation matrix has been generated by the HiTC R package [118]. (F) Schematic of a simplified Hi-C contact map displaying three chromosomes. In the intrachromosomal area (black-broken-line box), chromosome arms are individualized on each side of the centromere. Centromere-centromere interactions are the main features observed in the interchromosomal area (white-broken-line box).

Fig 1 .

 1 Fig 1. B. mori Oligopaint design. (A-C) Schematic of Oligopaints in B. mori. Whole chromosome Oligopaints are shown in A, stripe Oligopaints in B, and active/inactive Oligopaints in C. White regions indicate the absence of oligos (A) or regions not labeled by the respective barcode indices (B, C). (D) Schematic of primary probe design, showing whole chromosome barcodes and two unique barcodes (for stripes or active/inactive domains). (E) Schematic for Oligopaint DNA FISH assay with labeled secondary oligos. First, ordered oligos are amplified with primers containing barcode of interest and secondary oligo binding site, generating primary oligos. Primary oligos are then annealed to DNA and labeled with secondary oligos (shown in green). https://doi.org/10.1371/journal.pgen.1009700.g001

Fig 2 .

 2 Fig 2. Whole chromosome Oligopaints in B. mori 5 th instar germline squashes. (A) Schematic of early meiosis I (prophase I and metaphase I). One pair of homologous chromosomes is shown (red = paternal; blue = maternal).Prophase I is typically subdivided into five distinct stages: leptotene, zygotene, pachytene, diplotene, and diakinesis. Briefly: in leptotene, replicated chromosomes are reorganized and compacted into a linear scaffold structure. In zygotene, synapsis begins between the homologous chromosomes. In pachytene, synapsis is complete (black dots represent the synaptonemal complex holding the homologs together). This stage is also when crossing over can occur. In diplotene, the homologs repulse, condense further, and the SC breaks down. The homologs remain attached via chiasma (crossovers). Finally, in diakinesis, chromosome condensation and cruciform bivalent formation is nearly complete as the cell prepares for metaphase I. Arrows in metaphase I schematic indicate the assumed direction of spindle poles based on the direction of the metaphase plate. (B-E) Pachytene cells labeled with three whole chromosome Oligopaints, as indicated. B-D, larval testes. E, larval ovary. Scale bars = 10 μm. DAPI is shown in gray. F-H) Meiotic prophase I cells from larval testes squashes with whole chromosome paints for ch7 (magenta), ch15 (orange), and ch16 (green). Boxes indicate subsequent panels as indicated. DAPI is shown in gray. (F) Leptotene/zygotene cells, with unpaired, decondensed chromosomes and a dense lampbrush DNA stain. (G) Pachytene cells, with paired, linear, and relatively decondensed chromosomes and a loose lampbrush DNA stain. (H) Diplotene/diakinesis cells, with paired, less linear and more compact chromosomes and a diffuse DNA stain. (I) Quantification of prophase cell staging from late 4 th -early 5 th instar larval testes. n = 3 testes were analyzed. Error bars show mean and standard deviation between testes. (J) Metaphase I cells, with paired homologs condensed and aligned along the metaphase plate. Arrows indicate the direction of spindle poles. (K) Mitotic cells from larval testes, with chromosomes condensed and aligned along the metaphase plate but with unpaired homologs. Arrows indicate the direction of spindle poles.

  Fig 2. Whole chromosome Oligopaints in B. mori 5 th instar germline squashes. (A) Schematic of early meiosis I (prophase I and metaphase I). One pair of homologous chromosomes is shown (red = paternal; blue = maternal).Prophase I is typically subdivided into five distinct stages: leptotene, zygotene, pachytene, diplotene, and diakinesis. Briefly: in leptotene, replicated chromosomes are reorganized and compacted into a linear scaffold structure. In zygotene, synapsis begins between the homologous chromosomes. In pachytene, synapsis is complete (black dots represent the synaptonemal complex holding the homologs together). This stage is also when crossing over can occur. In diplotene, the homologs repulse, condense further, and the SC breaks down. The homologs remain attached via chiasma (crossovers). Finally, in diakinesis, chromosome condensation and cruciform bivalent formation is nearly complete as the cell prepares for metaphase I. Arrows in metaphase I schematic indicate the assumed direction of spindle poles based on the direction of the metaphase plate. (B-E) Pachytene cells labeled with three whole chromosome Oligopaints, as indicated. B-D, larval testes. E, larval ovary. Scale bars = 10 μm. DAPI is shown in gray. F-H) Meiotic prophase I cells from larval testes squashes with whole chromosome paints for ch7 (magenta), ch15 (orange), and ch16 (green). Boxes indicate subsequent panels as indicated. DAPI is shown in gray. (F) Leptotene/zygotene cells, with unpaired, decondensed chromosomes and a dense lampbrush DNA stain. (G) Pachytene cells, with paired, linear, and relatively decondensed chromosomes and a loose lampbrush DNA stain. (H) Diplotene/diakinesis cells, with paired, less linear and more compact chromosomes and a diffuse DNA stain. (I) Quantification of prophase cell staging from late 4 th -early 5 th instar larval testes. n = 3 testes were analyzed. Error bars show mean and standard deviation between testes. (J) Metaphase I cells, with paired homologs condensed and aligned along the metaphase plate. Arrows indicate the direction of spindle poles. (K) Mitotic cells from larval testes, with chromosomes condensed and aligned along the metaphase plate but with unpaired homologs. Arrows indicate the direction of spindle poles. https://doi.org/10.1371/journal.pgen.1009700.g002

Fig 3 .

 3 Fig 3. Stripe and active/inactive chromosome paints in B. mori 5 th instar testes squashes. (A) Left: Schematic of stripe paints for ch15, with tel1 in cyan, mid in yellow, and tel2 in magenta. Right: Pachytene cells labeled with ch15 stripe paints. (B-E) Left: Schematic of stripe paints for ch7, ch16, ch23 or chZ. Right: Representative pachytene nucleus labeled with stripe paints shown on left. (F) Left: Schematic of active/inactive paints for ch7, 15, and 16. Active domains are shown in green, and inactive domains are shown in red. Right: Representative pachytene nucleus labeled with paints for all 3 chromosomes. https://doi.org/10.1371/journal.pgen.1009700.g003

Fig 4 .

 4 Fig 4. Whole chromosome Oligopaints in zygotene-pachytene transition show a variety of folding and pairing configurations. (A) Oligopaints for ch16 (green) or ch15 (orange) in representative zygotene nuclei showing the most prevalent partially paired chromosome configurations during pairing initiation. Dashed line approximates the nuclear edge. (B) Frequency histogram showing the fraction of cells harboring chromosomes in each partially paired configuration for the indicated chromosome. Statistics, Mann-Whitney test with each chromosome as a replicate. (C) Oligopaints for ch16 (green) and ch15 (magenta) in representative zygotene nuclei showing asynchronous pairing initiation. Top: Ch16 has begun pairing while ch15 remains entirely unpaired. Bottom: Ch15 is nearly

Fig 5 .

 5 Fig 5. Analysis of pairing and metaphase I bivalent formation in 5 th instar larval testes squashes using stripe paints. (A) Left: Schematic of stripe paints for ch15, with tel1 in cyan, mid in yellow, and tel2 in magenta. Right: representative nuclei at the designated stages labeled with ch15 stripe paints. When cells enter meiosis, chromosomes begin to decondense (leptotene) and homologs pair (zygotene). Pairing is complete by pachytene, with complete synapsis for crossing over, and chromosomes are linear. Chromosomes begin to condense for segregation in diplotene and diakinesis. DAPI is shown in gray. (B) Top: Representative zygotene nucleus labeled with ch15 stripe paints showing a forked chromosome pairing configuration with tel1 paired. Bottom: Cartoon schematic of ch15 in above cell. (C) Top: Representative zygotene nucleus labeled with ch15 stripe paints showing a circular chromosome with a head-to-head pairing configuration. Bottom: Cartoon schematic of ch15 in above cell. (D) Top: Two representative zygotene nuclei labeled with ch15 stripe paints showing a circular chromosome with a head-to-tail pairing configuration. Bottom: Cartoon schematics of ch15 in above cells. (B-D) Dashed line indicates nuclear edge. (E) Quantification of circular chromosome pairing configurations in zygotene. Graph includes pooled data for ch15 and ch23, with n = 3 testes quantified for each. Error bars show standard deviation between chromosomes. �� p = 0.0082, unpaired t-test. (F) Quantification of stripe domain pairing in cells with only one stripe domain paired. n = 3 testes were quantified for each chromosome. For all chromosomes, pairing at both telomere domains is significantly enriched compared to the middle domain (p<0.0001, Fisher's exact test comparing tel1 vs mid and tel2 vs mid). Cyan asterisk = tel1 significantly enriched compared to tel2, magenta asterisk = tel2 significantly enriched compared to tel1 (Fisher's exact test, p<0.05). (G) Dot plot showing tel1: tel2 pairing ratio (X-axis) versus tel1:tel2 gene density ratio (Y-axis). Line of best fit calculated for autosome data only. Z chromosome data (gray) included for comparison. (H) Metaphase I bivalents labeled with ch15 stripe paints. Top: bivalent with pairing in tel1 domain. Bottom: bivalent with pairing in tel2 domain. Schematics of bivalents shown on the right. (I) Quantification of metaphase I orientation for ch7, 15, 16, 23, and Z. Ch7, n = 357 (48% tel 1 paired). Ch15, n = 182 (45% tel 1 paired). Ch16, n = 252 (50% tel1 paired). Ch23, n = 519 (47% tel1 paired). ChZ, n = 169 (48% tel1 paired). Each FISH assay was performed on n = 2-3 testes. https://doi.org/10.1371/journal.pgen.1009700.g005

Fig 6 .

 6 Fig 6.FISH with stripe chromosome paints in whole mount 5 th instar larval testes. (A) Cartoon schematic of 5 th instar larval testis. Mature 5 th instar larval testes are comprised of four testicular lobes, each of which harbors germline stem cells(1, mitotic zone) and spermatocytes in all stages of meiosis up to mature sperm; progressing from right to left in the image as indicated:(2) meiosis I/primary spermatocytes, (3) meiosis II/secondary spermatocytes. Additionally, each lobe is surrounded by somatic cells in the sheath[START_REF] Agostinho | Sexual dimorphism in the width of the mouse synaptonemal complex[END_REF] and in the septae separating the lobes.(5)(6) prophase I: (5) leptotene/zygotene, (6) pachytene/diplotene,(7) metaphase/anaphase I,(8) prophase/metaphase II, (9) spermatids,(10) mature sperm bundles. (B) Confocal image of 5 th instar larval testis stained with DAPI. Boxes indicate subsequently zoomed panels as indicated. Inset: ch7 stripe paints used in C-E. (C) Zoom of mitotic and pachytene region of larval testes as shown in A, labeled with ch7 stripe paints. Red boxes indicate zooms shown to the right. (C1) mitotic cells-chromosomes condensed, homologs unpaired, and aligned at the metaphase plate. Note how chromosomes are compacted perpendicular to the metaphase plate. (C2) leptotene cells, chromosomes are slightly decondensed and homologs are unpaired. (C3) pachytene cells, chromosomes are paired head-to-tail and linear. (D) Zoom of late prophase/metaphase I region of larval testis as shown in A, labeled with ch7 stripe paints. Red boxes indicate zooms shown to the right and below. Please note that some FISH signal background is visible within the DNA in this image. This is an artifact of the size of the larval gonad and the imaging depth. (D1) diplotene cells (diffuse stage), chromosomes are still paired and beginning to condense. Left: merged with DAPI in gray. Right: ch7 tel1 in cyan, mid in yellow, and tel2 in magenta. (D2) metaphase I cells labeled with ch7 stripe paints. White box indicates zoom shown in E. Left: merged with DAPI in gray. Right: tel1 (cyan) and tel2 (magenta) paints. Percent of cells with tel1 or tel2 paired indicated inside the panel. Cells from 1-2 bundles each from n = 3 testes were quantified for a total of n = 196 cells (47% tel1 paired). (D3) eupyrene-destined secondary spermatocyte bundle. Left: merged ch7 stripe paints with DAPI in gray. Right: ch7 stripe paints, tel1 (cyan) and tel2 (magenta). (D4) apyrene-destined secondary spermatocyte bundle. Left: merged ch7 stripe paints with DAPI in gray. Right: DAPI. Yellow arrowheads indicate spermatocytes that have already undergone nuclear degradation. Scale bar = 2 μm for all panels 1-4. (E) Zoom of metaphase I cell indicated in C2. Top: merged with DAPI in gray. Bottom: ch7 tel1 (cyan), mid (yellow), and tel2 (magenta) paints. Bivalent pairing is in tel2 domain in this cell. Note: zoomed fields for all panels may display a slightly different Z position than the larger field views for better clarity.

  Fig 6.FISH with stripe chromosome paints in whole mount 5 th instar larval testes. (A) Cartoon schematic of 5 th instar larval testis. Mature 5 th instar larval testes are comprised of four testicular lobes, each of which harbors germline stem cells(1, mitotic zone) and spermatocytes in all stages of meiosis up to mature sperm; progressing from right to left in the image as indicated:(2) meiosis I/primary spermatocytes, (3) meiosis II/secondary spermatocytes. Additionally, each lobe is surrounded by somatic cells in the sheath[START_REF] Agostinho | Sexual dimorphism in the width of the mouse synaptonemal complex[END_REF] and in the septae separating the lobes.(5)(6) prophase I: (5) leptotene/zygotene, (6) pachytene/diplotene,(7) metaphase/anaphase I,(8) prophase/metaphase II, (9) spermatids,(10) mature sperm bundles. (B) Confocal image of 5 th instar larval testis stained with DAPI. Boxes indicate subsequently zoomed panels as indicated. Inset: ch7 stripe paints used in C-E. (C) Zoom of mitotic and pachytene region of larval testes as shown in A, labeled with ch7 stripe paints. Red boxes indicate zooms shown to the right. (C1) mitotic cells-chromosomes condensed, homologs unpaired, and aligned at the metaphase plate. Note how chromosomes are compacted perpendicular to the metaphase plate. (C2) leptotene cells, chromosomes are slightly decondensed and homologs are unpaired. (C3) pachytene cells, chromosomes are paired head-to-tail and linear. (D) Zoom of late prophase/metaphase I region of larval testis as shown in A, labeled with ch7 stripe paints. Red boxes indicate zooms shown to the right and below. Please note that some FISH signal background is visible within the DNA in this image. This is an artifact of the size of the larval gonad and the imaging depth. (D1) diplotene cells (diffuse stage), chromosomes are still paired and beginning to condense. Left: merged with DAPI in gray. Right: ch7 tel1 in cyan, mid in yellow, and tel2 in magenta. (D2) metaphase I cells labeled with ch7 stripe paints. White box indicates zoom shown in E. Left: merged with DAPI in gray. Right: tel1 (cyan) and tel2 (magenta) paints. Percent of cells with tel1 or tel2 paired indicated inside the panel. Cells from 1-2 bundles each from n = 3 testes were quantified for a total of n = 196 cells (47% tel1 paired). (D3) eupyrene-destined secondary spermatocyte bundle. Left: merged ch7 stripe paints with DAPI in gray. Right: ch7 stripe paints, tel1 (cyan) and tel2 (magenta). (D4) apyrene-destined secondary spermatocyte bundle. Left: merged ch7 stripe paints with DAPI in gray. Right: DAPI. Yellow arrowheads indicate spermatocytes that have already undergone nuclear degradation. Scale bar = 2 μm for all panels 1-4. (E) Zoom of metaphase I cell indicated in C2. Top: merged with DAPI in gray. Bottom: ch7 tel1 (cyan), mid (yellow), and tel2 (magenta) paints. Bivalent pairing is in tel2 domain in this cell. Note: zoomed fields for all panels may display a slightly different Z position than the larger field views for better clarity. https://doi.org/10.1371/journal.pgen.1009700.g006

Fig 7 .

 7 Fig 7. FISH with stripe chromosome paints in whole mount 5 th instar larval ovary. (A) Schematic of ovariole from 5 th instar B. mori larval ovary. (B1-D1) Representative field of nuclei from larval ovary labeled with ch7 stripe paints imaged with diffraction-limited confocal microscopy. DAPI is shown in gray. (B) germarium (zone of pairing, region 1), (C) region of differentiation (region 2), (D) mature larval oocytes (region 3b). (C1-D1) yellow arrowheads indicate oocytes which were identified based on their unique chromosome morphology and weak DAPI stain. (B2-D5) Representative nuclei labeled with ch7 stripe paints and DAPI imaged with super-resolution microscopy. (B4-D4) zoom of ch7 shown in B2-D2. (D5) zoom of ch7 shown in D2, DAPI only. (D6) cartoon schematic of chromosomes in mature larval oocytes. (E) Measurement of distances between homologs obtained from super-resolution images. n = 3 ovaries were imaged and quantified. Measurements were taken from 3-5 chromosomes in 5-10 cells per ovary, resulting in 50-75 data points per region. ��� p<0.001, Mann-Whitney test. https://doi.org/10.1371/journal.pgen.1009700.g007

Table 2 :

 2 List of Hi-C data normalization methods.

	Method	Approach	Publication
	ICE	Iterative Correction and Eigenvector decomposition	Imakaev et al., 2012
	SCN	Sequential Component Normalization	Cournac et al., 2012 Venev et al., 2021
	HiCNorm	A generalized linear model approach	Hu et al., 2012
	KR	An implicit individual-sample normalization approach	Knight and Ruiz, 2013
	multiHiCcompare	A locally weighted regression approach	Stansfield et al., 2019

  , so I was curious to see what effect condensin II might have in B. mori cells. What I found was the condensin II depletion has the opposite effect that cohesin depletion has. Depleting condensin II results in an increase in intradomain contact frequency and an increase in local compaction. This was shown in the same manner as the effect for Scc1 RNAi . This effect is interesting because the results indicate that depleting CapH2 increases the loop extrusion that is likely done by cohesin. Meaning that CapH2 is possibly limiting loop extrusion in WT conditions. Another possibility is that condensin II could be limiting cohesin loop extrusion by promoting other

interactions in the genome. Depleting condensin II allows these regions to now be extruded by cohesin in its absence. In order to investigate how this may be occurring, it would also be interesting to map the locations of CapH2 in the B. mori genome. Knowing the regions of CapH2 enrichment could help in determining the mechanism by which condensin II counteracts cohesin.

Taken together, cohesin and condensin II seem to have opposing roles, similar to what has been reported in D. melanogaster

[START_REF] Rowley | Condensin II Counteracts Cohesin and RNA Polymerase II in the Establishment of 3D Chromatin Organization[END_REF]

. However, in D. melanogaster, this counteracting effect is associated to RNAPolII activity and intragenic regions. I have not done an experiment or analysis to verify if the effects I see are related to active chromatin regions, but I can report that the effects seen in B. mori also occur in regions known to be inactive. With this observation, I

Table 3 :

 3 Oligopaint coordinates

	Domain	paint start	paint stop	size (kb)	probes/kb	total # of
						oligos
	Chr04_set1_A	8800000	9000000	200	3
	Chr04_set2_A	13000000	13200000	200	2.9
	Chr04_set1_B	3360000	3560000	200	4.4
	Chr04_set2_B	7560000	7760000	200	4
	Chr04_set1_X	12100000	12300000	200	3
	Chr04_set2_X	16300000	16500000	200	3.3
	Chr17_A	5040000	5320000	280	2.3
	Chr17_B	8960000	9240000	280	2.4
	Chr17_X	12040000	12320000	280	2.4
	Chr23_A	15920000	16240000	320	2.4
	Chr23_B	9100000	9420000	320	2.4
	Chr23_X	4580000	4900000	320	2.4

Trends Trends in in Genetics Genetics

Rosin ID 1 *, Jose Gil, Jr. 2 , Ines A. Drinnenberg ID 2 , Elissa P. Lei ID 1

  . Centromere Organization in Eukaryotes

	Trends in Genetics
	Species	Number of	Type of centromere	Centromere imaging	Chromosome
		chromosomes			organization
	Saccharomyces cerevisiae	n =16	Point centromeres	One focus near SPB	Rabl-like
		2n =32
	Candida albicans	2n = 16	Regional without repetitive	One focus near SPB	Rabl-like
			sequence
	Schizosaccharomyces pombe	n =3	Regional with inverted repeats	One focus near SPB	Rabl-like
		2n =6
	Cryptococcus neoformans	n = 14	Regional with transposable	Individual foci at nuclear periphery	ND
			elements
	Arabidopsis thaliana	2n = 10	Regional with transposable	Clustered foci at chromocenters	Centromere
			elements and SatDNA		clustering
	Oryza sativa (rice)/ Sorghum	2n = 24/2n = 20	Regional with transposable	Dispersed foci in nucleus	Non-Rabl
	bicolor		elements and SatDNA
	Triticum aestivum (wheat)	2n = 6X = 42	ND	Clustered foci at one cell pole,	Rabl
				opposite nucleolus
	Drosophila melanogaster	2n = 8	Regional with transposable	Clustered foci around the	CT and arm territory
			elements and SatDNA	nucleolus
	Mus musculus	2n = 40	Regional with SatDNA	Clustered foci at chromocenters	CT
				localized at nuclear periphery
	Homo sapiens	2n = 46	Regional with SatDNA	Clustering at nuclear periphery	CT
	ND, not determined.

Trends in Genetics, August 2019, Vol. 35, No. 8 575 RESEARCH ARTICLE Oligopaint DNA FISH reveals telomere-based meiotic pairing dynamics in the silkworm, Bombyx mori Leah F.

Table 1 . Chromosome and chromosome paint information.

 1 Fig 2H) in which chromosomes are more condensed, paired, and have diffuse DAPI signal. Of cells in Prophase I in late 4 th -early 5 th instar larval testes, the majority are in pachytene (Fig 2I). Moreover, we were able to identify cells in which chromosomal bivalents are compacted and aligned at the metaphase plate (metaphase I, Fig 2J). Finally, in addition to meiotic cells, we observed interphase and mitotic cells in our testes squashes harboring two distinct, round fluorescence signals per chromosome, indicating that homologs are unpaired in the majority of non-meiotic cells in B. mori (Figs 2K,

	Chrom	chromosome size (bp)	size painted (bp)	paint start	paint stop	density (probes/kb)	total # of oligos	gene density (genes per Mb)
	4	18737234	18639239	282	18639521	1.5	26841	39.7
	7	13944894	13868845	35931	13904776	3	42625	36.0
	15	18440292	18354755	21089	18375844	1	17756	43.6
	16	14337292	14275583	27737	14303320	1.5	20190	43.7
	17	16840672	16806551	9415	16815966	1.5	23834	38.3
	23	21465692	21339065	123188	21462253	1.5	30506	35.8
	Z/1	20666287	20578020	37936	20615956	1.5	29784	32.2
	https://doi.org/10.1371/journal.pgen.1009700.t001

Table 2 . Stripe sub-library paint information. Chrom. and stripe paint start paint stop Stripe size (Mb)

 2

	7 tel1	35931	2809682	2.77
	7 mid	5587874	8357288	2.76
	7 tel2	11131025	13904776	2.77
	15 tel1	21089	3684775	3.66
	15 mid	7380341	11061106	3.68
	15 tel2	14748412	18375844	3.62
	16 tel1	27737	2865990	2.84
	16 mid	5732698	8598132	2.87
	16 tel2	11464265	14303320	2.84
	23 tel1	123188	1638259	1.51
	23 mid	9909172	11556867	1.65
	23 tel2	19815237	21462253	1.65
	Z tel1	37936	1589202	1.55
	Z mid	9538407	11119201	1.58
	Z tel2	19078122	20615956	1.53
	https://doi.org/10.1371/journal.pgen.1009700.t002

Table 3 . Ch7 Active and inactive chromatin domains paint information.

 3

	Domain	Start	Stop	Size (bp)
	inactive	35931	800000
	active	800001	900000
	inactive	1000001	1100000
	active	1100001	1250000
	inactive	1300001	1500000
	active	1500001	1600000
	inactive	1600001	1700000
	active	1700001	1850000
	inactive	1850001	1900000
	active	1950001	2000000
	inactive	2000001	2300000
	inactive	2400001	2600000
	active	2600001	2700000
	inactive	2700001	3350000
	active	3400001	3650000
	inactive	3650001	3900000
	active	3900001	4000000
	inactive	4000001	4550000
	active	4550001	4650000
	inactive	4650001	4950000
	active	4950001	5400000
	inactive	5450001	5800000
	active	5900001	6050000
	inactive	6050001	6250000
	inactive	6300001	6450000
	active	6450001	6550000
	inactive	6550001	6650000
	active	6700001	6800000
	active	6850001	7150000
	inactive	7150001	7400000
	active	7400001	7600000
	inactive	7600001	7900000
	active	7950001	8200000
	inactive	8250001	8750000
	inactive	8800001	9050000
	active	9050001	9300000
	inactive	9350001	9400000
	active	9400001	9550000
	inactive	9650001	10800000
	active	10850001	11100000
	active	11150001	11400000
	inactive	11400001	11650000
	active	11650001	11950000
	inactive	11950001	12300000
	active	12300001	12600000
	inactive	12600001	12700000
	active	12700001	12750000
				(Continued )

Table 3 .

 3 (Continued)

	Domain	Start	Stop	Size (bp)
	inactive	12800001	12950000
	active	12950001	13150000
	inactive	13150001	13250000
	active	13350001	13450000
	inactive	13500001	13650000
	active	13700001	13849955
	https://doi.org/10.1371/journal.pgen.1009700.t003

Table 4 . Ch15 Active and inactive chromatin domains paint information.

 4

	Domain	Start	Stop	Size (bp)
	inactive	21089	200000
	active	250001	850000
	active	900001	1000000
	inactive	1000001	1400000
	active	1400001	1950000
	inactive	1950001	2250000
	active	2250001	2550000
	inactive	2550001	2800000
	active	2850001	3200000
	inactive	3200001	3500000
	active	3550001	3850000
	inactive	3850001	3900000
	active	3950001	4100000
	inactive	4100001	4250000
	active	4250001	4350000
	inactive	4350001	4650000
	active	4650001	4750000
	inactive	4750001	4950000
	inactive	5000001	5150000
	active	5150001	5250000
	inactive	5300001	5350000
	active	5350001	6100000
	inactive	6100001	6200000
	active	6250001	6500000
	inactive	6500001	6700000
	active	6700001	6850000
	inactive	6900001	7100000
	active	7100001	7300000
	inactive	7300001	7550000
	active	7550001	7800000
	inactive	7800001	8100000
	active	8100001	8300000
	inactive	8400001	8650000
	active	8650001	8750000
	inactive	8750001	8800000
	active	8850001	8900000
				(Continued )

Table 4 .

 4 (Continued)

	Domain	Start	Stop	Size (bp)
	inactive	8900001	9100000
	active	9100001	9500000
	inactive	9500001	9550000
	active	9600001	9750000
	active	9800001	9950000
	inactive	10000001	10500000
	active	10500001	11200000
	active	11350001	11400000
	inactive	11400001	11500000
	active	11500001	11950000
	inactive	11950001	12000000
	active	12050001	12200000
	inactive	12200001	12250000
	active	12300001	12500000
	inactive	12500001	12600000
	active	12600001	12850000
	inactive	12850001	12950000
	active	12950001	13400000
	inactive	13450001	13600000
	active	13650001	13750000
	inactive	13750001	14150000
	inactive	14200001	14850000
	active	14900001	15050000
	inactive	15100001	15250000
	active	15300001	15500000
	inactive	15550001	15600000
	active	15600001	15750000
	inactive	15750001	16000000
	inactive	16050001	16350000
	inactive	16400001	17850000
	inactive	17900001	18100000
	active	18150001	18200000
	inactive	18200001	18250000
	active	18300001	18350000
	inactive	18350001	18375844
	https://doi.org/10.1371/journal.pgen.1009700.t004

Table 5 . Ch16 Active and inactive chromatin domains paint information.

 5

	Domain	Start	Stop	Size (bp)
	inactive	27737	50000
	active	50001	150000
	inactive	150001	250000
	active	250001	350000
	inactive	350001	650000
	active	700001	1350000
	inactive	1400001	1700000
	active	1700001	2300000
	inactive	2300001	2400000
	active	2400001	2700000
	inactive	2700001	3200000
	active	3200001	3600000
	inactive	3600001	4050000
	active	4050001	4200000
	inactive	4200001	4500000
	active	4500001	4600000
	active	4700001	4750000
	inactive	4750001	4800000
	active	4900001	4950000
	active	5000001	5250000
	inactive	5250001	5600000
	active	5600001	6000000
	inactive	6000001	6150000
	active	6200001	6250000
	active	6300001	6500000
	active	6550001	6650000
	inactive	6650001	6850000
	active	6850001	7000000
	inactive	7050001	7950000
	inactive	8000001	8500000
	active	8550001	8750000
	inactive	8800001	8900000
	active	8900001	9400000
	inactive	9400001	9450000
	active	9500001	10100000
	inactive	10100001	11450000
	active	11450001	11550000
	inactive	11550001	11900000
	active	11950001	12250000
	inactive	12250001	12400000
	active	12400001	12500000
	inactive	12500001	12700000
	active	12750001	13200000
	inactive	13250001	13800000
	active	13800001	14100000
	inactive	14100001	14303320
	https://doi.org/10.1371/journal.pgen.1009700.t005

Trends in Genetics, August 2019, Vol. 35, No. 8 567

Trends in Genetics, August 2019, Vol. 35, No. 8

Trends in Genetics, August 2019, Vol. 35, No. 8

Trends in Genetics, August 2019, Vol. 35, No. 8 571

Trends in Genetics, August 2019, Vol. 35, No. 8

Trends in Genetics, August 2019, Vol. 35, No. 8

Trends in Genetics, August 2019, Vol. 35, No. 8

PLOS Genetics | https://doi.org/10.1371/journal.pgen.1009700July 28, 2021

Acknowledgements

There are so many people that played a part in my journey. Some in big ways, some in small ways, but all of great importance. For all of those who helped me when I needed you, thank you. This one is for us. I want to thank my thesis supervisors, Ines Drinnenberg and Héloïse Muller, for the opportunity to work on such an interesting project and improve my scientific abilities. I also want to thank you for your time and effort. I gained so much from this experience. I hope you were able to gain something as well. Both of your have changed my life for the better, and for that I am forever grateful.

I want to thank my thesis advisory committee (Laurent Lacroix, Leonid Mirny and Leandro Quadrana), for all of the great advice and input to help me improve my thesis project. I especially want to thank Leandro. You taught me everything I knew about bioinformatics. You helped me overcome hurdles I thought impossible. Thank you for giving me the access to such a powerful toolset. Also, thank you for always looking out for me. I'd also like to thank all of the PI's of the UMR3664 (Geneviève Almouzni, Antoine Coulon, Nathalie Dostatni, Angela Taddei) for all of the advice, support and kindness.

I want to thank my team members for all of the help and the time we shared together. Gaetan, thank you for your unwaivering help. I hope I can become a Jedi now. Pablo, thank you for all of your help and all of the conversations. Messi > C. Ronaldo, and it's not even close. Jaime, thank you for all of the advice and for the conversations as well. Gané y lo sabes! Sundar, thank you for all of the help as well. I still think of how you were drying those samples. Amazing. Acknowledgment We thank Harmit Malik and all the members of the Drinnenberg lab for discussion. I.A.D. receives salary support from the CNRS. This work is supported by the Labex DEEP ANR-11-LABX-0044 part of the IDEX Idex PSL ANR-10-IDEX-0001-02 PSL, the Institut Curie and funds from the Atip Avenir 2015 program and the ERC (758757-CENEVO).

Acknowledgments

We would like to thank the members of the Lei and Drinnenberg labs, as well as Jamie Walters, Petr Nguyen, Martina Dalikova, Eric Joyce, and Son Nguyen for helpful discussion. We would also like to thank Catherine McManus, Jennifer Luppino, Stacie Hughes, Jean-Rene ´Huynh, and Anahi Molla-Herman for critical reading of the manuscript. Finally, we would like to thank Xufeng Wu and Jeff Reese in the NHLBI and NIDDK microscopy cores, respectively, for assistance with super-resolution microscopy.

This work was funded by the Intramural Program of the National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health (NIDDK; DK015602 to E.P.L.), the Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health (NICHD; 1K99HD104851 to L.F.R), LabEx DEEP grants from the Agence National de la Recherche (ANR-11-LABX-0044 and ANR-10-IDEX-0001-02 to I.A.D.), an ATIP-AVENIR research grant from CNRS/INSERM, the European Research Council (ERC; CENEVO-758757 to I.A.D.), and Institut Curie (I.A.D). NIDDK provides salary support for E.P.L. and L.F.R. Institute Curie and CNRS provide salary support for I.A.D. ERC provides salary support for J.G. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

All relevant data are within the manuscript and its Supporting Information files.

Abbreviations

Discussion and Future Directions Supplementary Figures Annex [START_REF] Muller | The Impact of Centromeres on Spatial Genome Architecture[END_REF]Rosin et al., 2021b During my thesis I was able to contribute to three publications. In Muller et al., 2019, I provided the introductory paragraph and I also contributed to writing the section titled "Centromere Interactions Drive Genome 3D Organization and Regulation". In this publication I provided Figure 3 and the definitions for the keywords in the glossary. These contributions earned me second author placement. Rosin et al., 2021b is the result of a collaboration between Leah F. Rosin and Elissa P. Lei from the National Institute of Health (NIH) in the United States of America and Ines A. Drinnenberg and I. I contributed to this publication by performing the bioinformatic aspects of the oligopaint designs. This contribution earned me a second author placement.

Gil Jr. et al., 2023 is in progess and is not included in this annex. To this publication I provided the analysis of observed/ expected interchromosomal contacts. I also provided the analysis for the P(s) curves of homotypic and heterotypic compartment contacts along with their respective derivatives. I provided the analysis for the epigenetic characterization of the compartments and I also provided the analysis for the compartment size distribution and genome coverage. In the supplementary data, I provided the analysis for interchromosomal, homotypic compartment contacts. I contributed to the design of figures 1, 2 and 4 and I also contributed to the discussion of the article. These contributions earned me first author placement. Writing -review & editing: Leah F. Rosin, Jose Gil, Jr., Ines A. Drinnenberg, Elissa P. Lei.

Supporting information