Next Article in Journal / Special Issue
Transposon and Transgene Tribulations in Mosquitoes: A Perspective of piRNA Proportions
Previous Article in Journal / Special Issue
Chromatin Organization during C. elegans Early Development
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
DNA
Volume 4
Issue 1
10.3390/dna4010005
dna-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

How Chromatin Motor Complexes Influence the Nuclear Architecture: A Review of Chromatin Organization, Cohesins, and Condensins with a Focus on C. elegans

by
Bahaar Chawla
and
Györgyi Csankovszki
*
Department of Molecular, Cellular, and Developmental Biology, University of Michigan, Ann Arbor, MI 48109-1085, USA
*
Author to whom correspondence should be addressed.
DNA 2024, 4(1), 84-103; https://doi.org/10.3390/dna4010005
Submission received: 29 December 2023 / Revised: 2 February 2024 / Accepted: 5 March 2024 / Published: 11 March 2024
(This article belongs to the Special Issue DNA Organization in Model Organisms)
Browse Figures
Review Reports Versions Notes

Abstract

:
Chromatin is the complex of DNA and associated proteins found in the nuclei of living organisms. How it is organized is a major research field as it has implications for replication, repair, and gene expression. This review summarizes the current state of the chromatin organization field, with a special focus on chromatin motor complexes cohesin and condensin. Containing the highly conserved SMC proteins, these complexes are responsible for organizing chromatin during cell division. Additionally, research has demonstrated that condensin and cohesin also have important functions during interphase to shape the organization of chromatin and regulate expression of genes. Using the model organism C. elegans, the authors review the current knowledge of how these complexes perform such diverse roles and what open questions still exist in the field.

Graphical Abstract

1. Introduction: Chromatin Organization

The word chromatin comes from the Greek word for color, khroma. In the late 1880s, Walter Flemming used this term to describe the dark stained material found in the cell preparations he treated with basophilic dyes [1]. Chromatin is now defined as the complex of DNA and associated proteins found in the nuclei of living organisms. The most discrete unit of eukaryotic chromatin is a nucleosome [2]. Discovered in the 1970s, nucleosomes are structures made up of an octamer of histone proteins and approximately 160 base pairs of DNA wrapped around the octamer, like thread on a spool [3,4]. As the cell is preparing to undergo mitosis, these nucleosomes are tightly condensed until they become the stereotypical chromosomes that can be seen under a microscope. Once cell division is completed, the chromosomes lose this degree of compaction [5].
However, that does not mean that interphase nuclei have no structure or organization. Instead, we now understand that how the chromatin is organized has a major impact on gene expression, tissue specification, and cell homeostasis [6,7,8,9]. In fact, long-range communication is known to be essential for gene expression, where a necessary enhancer may be many kilobases away from the promoter and must be brought in contact with the RNA transcription machinery for transcription to proceed [10,11]. One way in which this can be accomplished is through the recruitment of enhancer-binding proteins, such as CCAAT/the Enhancer-Binding Protein Family. Members of this family use their highly conserved basic-leucine zipper domain to both bind DNA and dimerize with other members of the same family or even other transcription factors to bring segments of DNA together by bending the DNA strands physically [12]. These chromatin-organizing events are often transient [13]. Some transcription factors, like those in the WNT signaling family are sequestered in the cytoplasm until circumstances are reached that promote them to translocate into the nucleus [14]. Therefore, transcription factors and enhancer-binding proteins alone are not the sole drivers of chromatin organization [15].
Instead, there are specific orders of chromatin organization, each accomplished through a variety of drivers. At the lowest level, the positioning of nucleosomes is specific. Nucleosomes are unlikely to be present at transcriptional start sites of actively transcribed genes, but instead, they are found in well-characterized patterns along the rest of the gene body [16]. The next level of organization is the clustering of nucleosomes. Known as “clutches”, these are defined as a group of more than three nucleosomes that are distinctly separated from other groups of nucleosomes by a distance more than 20 nm [17,18]. Clutches are often of variable number and density, the latter correlated with cell potency [19]. In somatic cells, clutches tend to have more nucleosomes that are packed in tighter, resulting in higher density. This contrasts with pluripotent cells, where the clutches are smaller and less dense, resulting in more open chromatin [17]. Open chromatin, or euchromatin, is also defined as accessible DNA, where the histone tails of the nucleosomes are modified to prevent tight packaging. This contrasts with heterochromatin, or inactive chromatin, where the nucleosomes can be packed very tightly, facilitated by different post-translational modifications on the histone tails [20].
The next level of chromatin organization is the formation of topologically associated domains, TADs. TADs are higher-order structures that are caused by the increased interaction of DNA within a genomic region with itself compared to interactions with DNA in a different genomic region [21,22]. Genes within a TAD are generally of the same transcriptional state, resulting in enrichment of histone modifications reflecting that state in the TAD [23,24]. TADs have a well-defined size, with the median in mouse cells at 880 kb and similar sizes in non-mammalian cells [21]. TADs are thought to promote gene expression through the close-range organization of enhancers and promoters while also insulating these from other DNA sequences [25]. This has been demonstrated through the removal of a TAD boundary, where the lack of a boundary resulted in new promoter–enhancer interactions and misregulated gene expression [26,27,28]. TAD boundaries are also very stereotypical and surprisingly well conserved between species [29,30]. However, plants have been shown to not utilize TADs as a method of genome organization [31]. In fact, even in species that do utilize TADs, it has been demonstrated that a large-scale disruption of TADs by depleting the key proteins responsible results in little disruption of gene expression. These results demonstrate that TADs are not the primary way to regulate gene expression, making the goal of understanding the link between chromatin organization and gene regulation still a work in progress [32,33].
The next order of chromatin organization is controversial. Initial studies suggested that chromatin at this level is separated into two compartments, A and B. The A compartment is made up of active chromatin, while the B compartment is made up of inactive chromatin [34,35]. DNA is classified into these compartments based on calculating principal eigenvectors of the frequent contacts in a Hi-C map [36,37]. However, recent work in human lymphoblastoid cells argues that the compartments are much smaller than what was previously defined. The newer smaller size puts compartments at the same resolution as TADs or even smaller, but unlike TADs, their boundaries do not line up with the same protein-enriched loops [38].
The final and highest order of chromatin are chromosome territories. Each chromosome is found in a discrete region of nucleus, and the only instances of overlapping between chromosomes is at the boundaries of their territories [39,40]. Generally, these territories are spherical, as in many mammalian and Arabidopsis cells [41,42]. How these chromosome territories are positioned in the nucleus has been found to be non-random, with euchromatin-rich chromosomes more central and heterochromatin-heavy chromosomes located at the nuclear periphery [43,44]. This organization results in a very complex nuclear architecture, one that is still being dissected to understand the nuances and implications.
This review will focus on chromatin organization and its impact on gene expression. While the focus of the review is condensins in Caenorhabditis elegans, this review will also summarize the literature on cohesins and the role of both complexes in chromatin organization in C. elegans as well as other organisms to provide broader context.

2. Structural Maintenance of Chromosome Proteins Are the Backbone of Chromatin Motor Complexes

Structural Maintenance of Chromosome (SMC) proteins are a family of highly conserved proteins found from bacteria to eukaryotes [45]. SMC proteins share a characteristic tertiary structure that allows them to accomplish their various functions [46,47,48]. Each SMC protein has two globular domains at each terminus that are separated by an alpha-helical region, then a “hinge”, and another alpha helix. The SMC proteins fold on their hinges, and the two alpha helices come together as coiled-coil (Figure 1). The coiled-coil brings the two globular domains in contact with each other, where they make up the ATPase “head” [49]. The ATPase is a key functional domain, as studies have shown that preventing ATP hydrolysis will also abolish loop extrusion and other functions of the SMC proteins [46,47].
In eukaryotes, there are generally six SMC proteins, named SMC1-6 [50]. These are found in three chromatin motor complexes, with differing functions [50]. SMC-1 and SMC-3 are the SMC protein pair found in cohesin [49]. SMC-5 and SMC-6 are found with a few other proteins in the SMC-5/6 complex, which functions in double-stranded break repair, restarting a stalled replication fork, and maintaining telomeres [51]. The final pair, SMC-2 (known as MIX-1 in C. elegans) and SMC-4 are found with three other proteins in complexes known as condensins [50,52]. All SMC proteins function in pairs, as their ATPase head domains require dimerization [46,53]. This dimerization is carried out through specific residues found in the hinge domain. Each hinge domain will interact with the hinge domain of the partnering SMC protein [49,54]. All three SMC protein-containing chromatin motor complexes can loop-extrude DNA. However, they perform very different functions from each other, not all of which contribute to chromatin architecture [45,55].
Along with the SMC proteins, another protein found in chromatin motor complexes is the kleisin. Kleisin comes from the Greek word for closure, kleisimo [56]. Kleisins are a superfamily of proteins, but all members are generally known to be SMC partners. There are three eukaryotic families and a single bacterial family. Despite the wide variety in these families, the main motifs that allow the kleisin to bind an SMC head at each terminus are well conserved [57].
The two chromatin motor complexes capable of modifying chromatin architecture, cohesin and condensin, comprise a pair of SMC proteins and a kleisin, along with different regulatory subunits. Cohesin resembles a “ring”, through which DNA is loop-extruded (Figure 2A). In C. elegans, this ring is made up of SMC-1; SMC-3; the kleisin subunit SCC-1 (sometimes referred to as COH-2); and a HEAT-repeat domain containing regulatory protein, SCC-3 [58]. There are homologues for all cohesin subunits found throughout eukaryotic organisms (see Table 1), with some eukaryotes containing multiple homologues of SCC-1 and SCC-3 [59,60,61]. These homologues allow for differential regulation and loading of cohesin complexes, which is especially important given the many roles cohesin is responsible for (discussed later in Section 3).
The full condensin complex (now called condensin I in eukaryotes) was originally found in Xenopus extracts, where co-immunoprecipitation using antibodies against the SMC proteins was used to pull down the non-SMC protein members of the complex [52]. These non-SMC proteins are now identified as CAP-D2, CAP-G, and CAP-H (DPY-28, CAPG-1, and DPY-26 in C. elegans). CAP-D2 and CAP-G are both HEAT-repeat domain containing proteins, whereas CAP-H is the kleisin [62]. The binding of the kleisin to the SMC proteins directly regulates the ATPase and loop-extruding function of the condensin. The current predicted mechanism suggests that the N-terminal tail of CAP-H relaxes its hold on SMC-2, which allows the SMC heads to work together to bind and hydrolyze ATP, opening the condensin ring to load onto DNA [63,64].
While the SMC/kleisin proteins are highly conserved from bacteria to eukaryotes, the non-SMC/kleisin regulatory proteins are not. A key difference between bacterial and eukaryotic condensins is that eukaryotic condensins have these HEAT-repeat domain containing proteins. Bacterial condensins instead have Kite (kleisin interacting winged-helix tandem elements) dimers to finish out their condensin rings with the SMC proteins and kleisin, and these Kites do not contain heat repeats [65]. Instead, the HEAT-repeat domain containing proteins found in present-day eukaryotic condensins and cohesins are now considered part of a unique subgroup of HEAT-repeat proteins; HAWKs (HEAT proteins associated with kleisins) [66]. Interestingly, these HAWKs are assumed to have replaced Kites in the condensin inherited from the last common eukaryotic ancestor [66].
Unlike cohesins that have multiple paralogous subunits to differentiate between functions, there are just two distinct condensin complexes with different functions in most eukaryotes, known as condensin I and condensin II [67]. Condensin I and II share the same SMC proteins, SMC2/4, but contain different non-SMC proteins (Figure 2B). Condensin I consists of CAP-D2, CAP-G, and CAP-H (see Table 2). Condensin II replaces those with CAP-D3, CAP-G2, and CAP H2 (see Table 3). One main difference in the two sets of non-SMC proteins is how they regulate condensin function. As mentioned previously, the ring-opening of condensin I is controlled by the kleisin CAP-H. Moreover, mitotic loading of condensin I onto DNA has been shown to be regulated by phosphorylating the N-terminal tail of CAP-H [63]. In contrast, CAP-H2 of condensin II does not have an N-terminal tail. Instead, the unique regulatory elements are found in one of the HAWK proteins, CAP-D3. CAP-D3 contains an additional helical HEAT “docker”, not found in CAP-D2, that can be phosphorylated [68].
C. elegans contain these condensins with their key structures, but unlike other eukaryotes, C. elegans contain three condensins. Condensin I is made up of the SMC protein pair MIX-1 (the commonly used name for C. elegans SMC-2) and SMC-4, along with the kleisin DPY-26 and HAWKs CAPG-1 and DPY-28. Condensin II has the same SMC pair, but with the kleisin KLE-2 and HAWKs CAPG-2 and HCP-6 [69]. The third condensin, condensin IDC, is identical to condensin I, but for one subunit (Figure 2B). Instead of having the SMC pair of MIX-1 and SMC-4, it has MIX-1 and a unique SMC protein DPY-27, and it is involved in dosage compensation, which will be discussed in detail later [69].
The final SMC-protein-containing complex is a not a chromatin architecture-modifying complex. Therefore, we will only discuss it briefly here. The SMC-5/6 complex is made up of the final SMC proteins SMC-5 and SMC-6, a kleisin NSE-4, and up to five regulatory proteins (see Table 4) [70,71]. Generally, the SMC-5/6 complex functions in DNA replication, DNA repair, and silencing of extra-chromosomal DNA [72,73]. In C. elegans, it has been shown that the SMC-5/6 complex processes double-stranded breaks in meiosis and prevents ectopic recombination [74]. The SMC-5/6 complex does not have a ring shape and, instead, is more analogous to a “boomerang” (Figure 2C) [72]. Interestingly, some of the regulatory subunits (NSE-1-3) are Kites, which could mean that the SMC-5/6 complex diverged from other SMC complexes very early [66].

3. The Unique Chromatin Architecture of C. elegans and the Role of Chromatin Motor Complexes in Cell Division

If chromosomes of model organisms were to be compared, most model organisms have monocentric chromosomes [75,76,77]. Each chromosome has only a single centromere that holds the sister chromatids together. The positioning of the centromere along the chromosome can be varied, but the centromeric DNA and its surroundings are usually condensed into heterochromatin during interphase [78]. The presence of multiple centromeres is typically a result of chromosomal rearrangements and can lead to defects in chromosome segregation [75]. However, the nematode C. elegans has holocentric chromosomes [79]. These holocentric chromosomes lack a single point centromere and instead have kinetochore attachments along the entire length of the chromosomes [79]. Despite this difference, C. elegans chromosomes follow principles of chromatin organization during interphase seen in other species [80]. Therefore, this section will highlight chromatin organization during cell division in the context of C. elegans while also identifying those mechanisms unique to C. elegans.
Holocentric chromosomes have evolved multiple times independently throughout the different phyla [79]. One hypothesis is that holocentric chromosomes having microtubule attachments along their lengths allows for the correct segregation of fragments, fusion, or translocated chromosomes [81]. However, this same physical characteristic is also the source of a meiotic conundrum. When the bivalent is formed in meiosis I, the holocentric chromosome can have spindle attachments that would pull the bivalents in multiple different directions, leading to problems in segregation [79]. Therefore, there must be an adaptation to overcome this potential pitfall. C. elegans accomplishes this through two approaches. The first is that C. elegans bivalents have only one chiasma that is closer to one end of the chromosome than the other [82,83,84]. This results in a cruciform-shaped bivalent, like monocentric bivalents (Figure 3B). The second adaptation is that the bivalents condense very tightly so that the short arms of the cruciform are no longer visible and the entire bivalent becomes a capsule that is pulled along the spindle [85]. Now resolved, meiosis proceeds as expected.
What roles do the chromatin motor complexes play? Biochemical (or single-molecule) experiments with purified condensins from other organisms suggest that the mechanism that both chromatin motor complexes employ to create higher-order structures is loop extrusion [86,87]. One description of this mechanism hypothesizes that the base of the loop is captured by the ATPase heads of the SMC proteins, while the hinge grabs a further segment of DNA. Then, the hinge is “scrunched” back and transfers that DNA to the heads to create a loop [88]. Another hypothetical mechanism is that the kleisin moves and allows DNA to enter the ring of the chromatin motor complex [89], or that the DNA is pushed into the ring by an ATP-dependent “stroke” [64,90]. A final hypothetical mechanism called “reel and seal” argues that chromatin motor complexes do not hold DNA within the ring, but instead, they hold DNA at two points, the interface of the kleisin and each HAWK and that the SMC proteins “reel” more DNA and feed it into the loop [91]. While still debated, the final product of loop extrusion are those same darkly stained bodies that Walter Flemming saw in his microscope centuries ago.
After DNA replication, the two sister chromatids must be held together until metaphase where they are oriented correctly for proper migration to opposing spindle poles [61,92]. Sister chromatid cohesion is accomplished by cohesin [49,58,93,94,95]. The sister chromatids are contained within the cohesin ring until metaphase, when separase, a protease, cleaves the SCC-1 subunit of cohesin and releases the sister chromatids [96,97].
In meiosis, the sister chromatids must be held together, but this differs greatly from mitosis [98]. In prophase I, homologous recombination occurs, which results in crossover of non-sister chromatid arms. Once the cell reaches anaphase I, the homologues must divide. During this event, cohesin is removed from the chromatid arms distal to the chiasma but maintained at the rest of the homologues. Cohesin is still present but only maintains cohesion between the sister chromatids. Then, in metaphase II, cohesin will be completely removed from the chromatids [99]. This diversity of cohesin’s roles in meiosis is accomplished through the multiple paralogues of each subunit. In C. elegans, the meiotic specific kleisins REC-8 and COH-3/4 replace SCC-1 [60,100,101]. Cohesins with REC-8 are recruited through a replication-dependent mechanism to the chromosomes, while cohesins with COH-3/4 are recruited during homologous recombination [60]. This is not unique to C. elegans. In fission yeast cohesins, the meiosis-specific kleisin Rec8 replaces Scc1, while the meiosis-specific HAWK Rec11 selectively replaces Psc3 [102,103]. Cohesin containing Psc3 is found at centromeres, but not at chromatids arms. In contrast, Rec11 forms a specific complex with Rec8 that is required for homologous recombination. Therefore, the many paralogues of cohesin subunits work together to maintain chromosome integrity during meiosis.
The condensin I and II complexes display different associations with DNA throughout the cell cycle. For example, during mitosis in mammalian cells, condensin II associates with DNA during prophase, while condensin I does not become DNA-localized until after nuclear envelope breakdown in prometaphase [104]. This is also true for C. elegans, where condensin II is present in the nucleus prior to nuclear envelope breakdown [105]. Metaphase onwards, condensin I and II are found in a mutually distinct pattern along the length of mammalian chromosomes [106]. In C. elegans, condensin I has been shown to “coat” mitotic chromosomes, whereas condensin II localizes near centromeric proteins which face the spindle poles (Figure 3A) [69,92]. Interestingly, condensin I partially shifts localization to the spindle midzone in anaphase where it delays cytokinesis to resolve chromatin bridges [107]. Somewhat contrary to the understanding that condensins are major contributors to the condensation of DNA for cell division [5,83,92,108], it has been shown in C. elegans that in condensin mutants, chromosomes are still able to condense, but the lack of condensin results in segregation defects [83,92].
Condensin localization becomes even more specific in meiosis [109]. Using mammalian meiosis as a paradigm, condensin I can first be visualized in metaphase I near centromeric regions. Then, in anaphase I, condensin I localization spreads also to chromosome arms, where it stays until metaphase II. In contrast, condensin II is visible from interphase onwards. From metaphase I through metaphase II, condensin II is localized to the meiotic chromosomes, except near centromeres. These distinct localizations have been shown to be required for specific events and the successful progression of meiosis in mammals, plants, and more [109,110,111]. In C. elegans, the pattern of condensin I and II localization is similar to other organisms, where condensin II is present in prophase, but condensin I only begins to localize to the meiotic chromosomes after nuclear envelope breakdown (Figure 3B) [69,105,109]. Interestingly, in C. elegans, condensin I also plays a unique role in protecting cohesin from premature degradation during meiosis [112].

4. Chromatin Motor Complexes Shape Chromatin Architecture during Interphase

While chromosome territories allow for an overall view of genomic dynamics in interphase, TADs are a common unit of organization both at the broader genomic level and in more discrete single gene–enhancer interactions [113,114]. Unlike the higher strata, TADs have distinct physical features. Specifically, TADs are defined by the enrichment of cohesin and CTCF at their boundaries [21]. Unlike cell division where condesin plays a major role, it is cohesin that is responsible for the interphase chromatin architecture [108]. In mammals and Drosophila, cohesin continuously moves along DNA, extruding loops, until it runs into CTCF [115]. CTCF is one of the most well-known chromatin organization drivers. A transcription factor, it is expressed ubiquitously through many different tissue types. Along with other transcription factors, CTCF creates strong boundaries that cohesin cannot pass, resulting in TAD boundaries [116].
One of the largest differences in how chromatin is organized between C. elegans and other species is the role of cohesion and CTCF. Unlike other organisms, where cohesin and CTCF play a large role in defining chromatin architecture in interphase, this is not seen in C. elegans. First, a CTCF homolog has not been identified in C. elegans [117]. Second, there do not seem to be cohesin-defined TAD boundaries in the autosomal chromosomes [118]. Instead, C. elegans autosomal chromosomes have areas of high-density heterochromatin that are located at the nuclear periphery [119]. These arms are tethered by interactions between histone modifications, chromodomain reader proteins, and lamina proteins [119,120,121,122]. Regions of the chromosomes attached to the nuclear lamina are referred to as lamina-associated domains (LADs), and they have also been observed in mammalian cells, plants, and others [123,124]. Using Hi-C, it has been shown that these heterochromatin-containing autosomal arms are organized in TADs, but they are smaller than in other organisms [118]. These smaller TADs are defined by an enrichment of histone modifications. Inactive domains are enriched for H3K27me3 with a median size of 18 kb. Active domains are enriched for a variety of “active” modifications, such as H3K36me3, and have a median size of 13 kb [24,125]. These domains also interact preferentially with like-domains, reminiscent of A/B compartments [24]. Taken together, it is apparent that C. elegans genomic architecture utilizes familiar strategies, even without CTCF.
As for cohesin’s roles in C. elegans, not much is known about its interphase functions. Like other eukaryotes, there are multiple SCC-1 homologs in C. elegans. SCC-1/COH-2 is the mitotic kleisin and SCC-1-containing cohesins are present in interphase [58], but their interphase roles have not been investigated. However, there is an SCC-1 homolog, COH-1, that plays a role in development that is not related to cell division. This protein has been observed in somatic nuclei and is consistently localized to DNA, regardless of cell cycle. Defects in COH-1-deficient worms result in muscular movement impairment, but not mitosis [58]. Whether this somatic COH-1 is found with members of cohesin and whether that complex is involved in gene regulation are still under investigation.
The interphase localization and function of each condensin is even more varied by species and even cell type. In Drosophila melanogaster, it has been found that condensin I is required for proper gene expression in differentiated neurons [126]. In contrast, condensin II is required for proper chromosome territory formation in D. melanogaster ovarian nurse cells [127]. In human interphase cells, condensin II is found in the nucleus where it has been linked to gene regulation and cellular senescence [128], while condensin I is not nuclear [104]. In fact, condensin II has been postulated to be the main determinant of spherical chromosome territories in chordates. A study compared the chromosomal architecture of 24 eukaryotic species, representing all subphyla of chordates, and found two distinct groups of chromosomal architecture [129]. One group had spherical chromosome territories while the other had what is known as the Rabl configuration. The Rabl configuration is characterized by the clustering of all centromeres at one pole and all the telomeres at the opposite pole, resulting in chromosome arms with lengths equal to the nuclear diameter [130]. Strikingly, all the species with Rabl configuration lacked the homologs for condensin II subunits [129]. While studies seem to agree on the requirement of condensin II during interphase, the wide variety of functions, such as spatial organization between centromeres and rDNA in plants or silencing retrotransposons in D. melanogaster, makes it difficult to group these functions as conserved or unique [131,132].
In C. elegans, condensin I does not associate with chromosomes during interphase [69,105], and while condensin II is present and binds some chromosomal sites [133], its function in interphase has not been determined. However, the third condensin, condensin IDC, is present. Condensin IDC gets its name because it is uniquely functional in dosage compensation, where it is responsible for the unique chromatin architecture that separates the X chromosomes from the rest of the genome.

5. TADs in C. elegans Are a Result of Dosage Compensation

The lack of CTCF does not mean that C. elegans do not have any strongly defined TADs. Instead, strongly defined TADs are uniquely found on the X chromosomes [114,118]. C. elegans have two sexes, hermaphrodites and males, which are determined chromosomally. There are two X chromosomes in hermaphrodites, but only a single X chromosome in males [134]. This difference in chromosome copy, however, leads to a dosage imbalance between the sexes.
Imbalance in gene dosage due to sex chromosomes is common among species that have chromosomally determined sex [135,136,137]. In the XY system, females have two X chromosomes and are therefore homogametic, while males have an X and Y chromosome apiece and are heterogametic. Mammals, flies, some plants, and some fish use the XY system [135,136,138,139]. In the ZW system, males are the homogametic sex with two Z chromosomes, while females are the heterogametic sex with a Z and W chromosome apiece. The ZW system is used in birds, amphibians, reptiles, and others [140,141].
To overcome the gene imbalance between homogametic and heterogametic sexes, many species have evolved an adaptation known as dosage compensation. However, while the end goal is the same, the actual dosage compensation mechanisms are incredibly diverse [142]. For example, in mammals, dosage compensation is achieved through X-inactivation, where one of the two X chromosomes in XX cells is silenced [135]. In contrast, flies, despite also using the XY system, upregulate the expression of a single male X so that the gene products produced are equal to the total gene products produced in XX cells [136].
C. elegans have a completely different dosage compensation mechanism [143,144]. The dosage compensation complex (DCC) in C. elegans is a multi-protein complex that binds to both hermaphrodite X chromosomes, downregulating each so that the total gene products in the hermaphrodite are equal to the gene products in the male. The DCC is made up of 10 proteins (Figure 4). The first five are those of condensin IDC; the SMC proteins MIX-1 and DPY-27, the kleisin DPY-26, and the regulatory HAWKs DPY-28 and CAPG-1 [69,145,146,147]. Then, there are three proteins, SDC-1, SDC-2, and SDC-3, which get their names from their dual roles in dosage compensation and sex determination [148,149,150]. The final two proteins are DPY-21 and DPY-30. DPY-21 is a jumonji-domain-containing demethylase [151,152], while DPY-30 is an essential regulator of H3K4 methylation [153,154,155], although its role in dosage compensation appears to be independent of its role in H3K4 methylation [156].
The localization of condensin IDC is not subject to the cell cycle like the other condensins. Instead, it is subject to the establishment of dosage compensation. Prior to dosage compensation establishment, the members of condensin IDC are not DNA-associated. Dosage compensation in C. elegans begins at the 30-cell stage, when SDC-2 expression begins in hermaphrodite embryos [157,158]. DCC members bind to the X chromosomes in a sequential pattern, beginning with the localization of SDC-2 to the X chromosomes [158]. It is followed by SDC-3 and DPY-30, then condensin IDC, and finally SDC-1 and DPY-21 [144,145,151,153]. Once dosage compensation is established, condensin IDC will be localized to the X chromosomes, regardless of cell cycle stage [69].
Localization of the DCC to the X chromosomes is dependent on rex sites. rex, or recruitment elements on X, sites are sequences of variable length that contain clusters of motifs that must act in combination to recruit the DCC [159,160,161]. These rex sites are distributed along the length of the X chromosomes in a non-random pattern that includes strong recruiting, weak recruiting, and non-recruiting regions based on DCC member occupancy using ChIP-seq [162]. How the DCC localizes across the X chromosomes is debated. One model is that the DCC binds initially to rex sites and then “spreads” to coat non-rex chromatin [160,163,164,165]. This model is supported by a ChIP-seq analysis of a fusion of the X chromosome with the fifth autosome (X:V) that demonstrates that the DCC can “spread” from the X chromosome into chromosome V [164]. However, an alternative model suggests that the rex sites are not recruitment elements, but they are two-sided blocks to prevent condensin IDC from moving past them [166,167]. This model proposes that condensin IDC is capable of loop-extruding randomly like other chromatin motor complexes and that the rex sites where SDC-2 binds and prevents condensin IDC from proceeding are analogous to CTCF-binding sites where CTCF prevents cohesin from looping further. Which model gives a more accurate understanding of the true mechanism is still unclear and therefore an area of ongoing work.

6. C. elegans Dosage Compensation as a Paradigm for Chromosome-Wide Gene Regulation

As our knowledge of chromatin organization has increased, so have developed theories of how gene regulation is no longer just about a single gene, but how the whole genome is coordinated in the context of tissue type, developmental stage, or as a response to external stressors [168,169,170]. Dosage compensation in C. elegans is an elegant paradigm for dissecting the mechanisms involved in chromosome-wide gene regulation, as the DCC is specific to X chromosomes only while also utilizing a highly conserved condensin so that the insights made in this system can be applicable broadly. Therefore, this section will highlight the many mechanisms used by the DCC to downregulate the X chromosomes.
At the gene level, binding of the DCC across the X chromosomes has been found to localize to transcriptional start sites, presumably to antagonize transcriptional machinery binding [163,171]. It has been demonstrated in many other systems that occupying the promoter region prevents the transcriptional machinery from binding and assembling [172,173].
At the nucleosome level, the DCC facilitates the post-translational modification of histone tails in multiple different ways. First, DPY-21, a member of the DCC, is responsible for the enrichment of H4K20me1 on nucleosomes of the X chromosomes [152,174,175]. This pattern is set up in a cell-cycle specific manner. First, the entire genome is stripped of H4K20me1 during DNA replication [176,177]. During mitosis, SET-1 will add H4K20me1 to the entire genome. After mitosis, that mark will be converted to H4K20me2/3 by SET-4 [174]. However, DPY-21 and the DCC will specifically convert H4K20me2 back to H4K20me1 on the X chromosome nucleosomes [143,152]. In fact, this strategy is not unique to C. elegans. In mammalian nuclei, the inactivated X chromosome is also enriched for H4K20me1 [178]. The other histone modification is H4K16ac, which is depleted on dosage-compensated X chromosomes and found to be associated with open chromatin [175]. A third modification which influences dosage compensation, H3K9me3, is not unique to the X chromosomes, as it is associated with any heterochromatin [121]. However, a lamina-associated chromatin reader, CEC-4, binds to H3K9me3-associated chromatin and, in cooperation with the DCC, tethers the X chromosomes to the nuclear lamina [122,179]. Tethering of heterochromatin is not unique to dosage compensation in C. elegans [121,122,180], but in conjunction with the DCC, it has a unique impact on the compaction and subnuclear localization of the X chromosomes. It has also been demonstrated that the nuclear RNAi machinery is also involved in X chromosome compaction and repression, but whether this effect is direct or indirect is not yet known [181]. A similar pathway involving Argonautes to establish heterochromatin has been observed in D. melanogaster [182]. Finally, it has been shown that one of the HAWKs, DPY-28, can interact with the tails of histone 3 and 4 [183], and this function may be a conserved one [184].
At higher-order chromatin levels, condensin IDC restructures the X chromosome into strongly defined TADs, whose boundaries are demarcated by rex sites and DCC occupancy [118,164]. Many studies have demonstrated how condensins in other organisms are capable of DNA-dependent loop extrusion [86,185,186,187]. In C. elegans, there is also evidence of how condensin IDC works in cooperation with topoisomerase I and II to create TADs [188]. Again, similar interactions between topoisomerases and chromatin motor complexes have been noted in other models [189]. At the chromosomal level, the X chromosomes in dosage compensated somatic cells are tightly condensed and located at the nuclear periphery [143,179], and the gene expression of both X chromosomes equal that of the male X [190]. The compaction and nuclear localization of the X chromosomes can be seen as a model of heterochromatic sequestering to the nuclear lamina [182].
Despite all these recent advances, there are still open questions in the field. While condensin IDC is structurally equivalent to mitotic condensins, we do not have the evidence to claim that DPY-27 is a true SMC protein. Instead, there is a possibility that condensin IDC serves only as a scaffold for the rest of the DCC. If so, how does the DCC loop the X chromosomes into the distinct TADs seen in Hi-C maps? Secondly, the developmental necessity of the DCC has been demonstrated. However, whether maintenance of dosage compensation requires active function of the DCC is not known. The recognition of DPY-21 as a demethylase and its function to enrich H4K20me1 are settled. However, it has been shown that DPY-21 has a role that is independent of its jumonji domain [183]. What that role entails is not known. Finally, mutating the DCC does not cause the nuclear occupancy of the X chromosomes in hermaphrodites equal that of the male X [191]. Therefore, there may still be other mechanisms unknown, waiting to be discovered and to be added to our understanding of C. elegans dosage compensation. These future studies in C. elegans will hopefully provide insights into our larger knowledge of chromatin organization and chromosome-wide gene regulation.

Author Contributions

Conceptualization, B.C. and G.C.; writing—original draft preparation, B.C.; writing—review and editing, B.C. and G.C.; supervision, G.C.; project administration, G.C.; funding acquisition, G.C. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the National Institutes of Health, grant numbers R01GM13385801 (to G.C.) and R35GM149543 (to G.C.), and by the National Science Foundation, grant number MCB 1923206 (to G.C.).

Conflicts of Interest

The authors declare no competing financial interests.

References

  1. Paweletz, N. Walther Flemming: Pioneer of mitosis research. Nat. Rev. Mol. Cell Biol. 2001, 2, 72–75. [Google Scholar] [CrossRef] [PubMed]
  2. Lee, J.Y.; Orr-Weaver, T.L. Chromatin. In Encyclopedia of Genetics; Brenner, S., Miller, J.H., Eds.; Academic Press: New York, NY, USA, 2001; pp. 340–343. [Google Scholar]
  3. Richmond, T.J.; Finch, J.T.; Rushton, B.; Rhodes, D.; Klug, A. Structure of the nucleosome core particle at 7 Å resolution. Nature 1984, 311, 532–537. [Google Scholar] [CrossRef] [PubMed]
  4. Kornberg, R.D. Chromatin Structure: A Repeating Unit of Histones and DNA. Science 1974, 184, 868–871. [Google Scholar] [CrossRef] [PubMed]
  5. Antonin, W.; Neumann, H. Chromosome condensation and decondensation during mitosis. Curr. Opin. Cell Biol. 2016, 40, 15–22. [Google Scholar] [CrossRef] [PubMed]
  6. Merkenschlager, M.; Nora, E.P. CTCF and Cohesin in Genome Folding and Transcriptional Gene Regulation. Annu. Rev. Genom. Hum. Genet. 2016, 17, 17–43. [Google Scholar] [CrossRef]
  7. Han, H.J.; Russo, J.; Kohwi, Y.; Kohwi-Shigematsu, T. SATB1 reprogrammes gene expression to promote breast tumour growth and metastasis. Nature 2008, 452, 187–193. [Google Scholar] [CrossRef]
  8. Tang, Z.; Luo, O.J.; Li, X.; Zheng, M.; Zhu, J.J.; Szalaj, P.; Trzaskoma, P.; Magalska, A.; Wlodarczyk, J.; Ruszczycki, B.; et al. CTCF-Mediated Human 3D Genome Architecture Reveals Chromatin Topology for Transcription. Cell 2015, 163, 1611–1627. [Google Scholar] [CrossRef]
  9. Phillips-Cremins, J.E.; Sauria, M.E.G.; Sanyal, A.; Gerasimova, T.I.; Lajoie, B.R.; Bell, J.S.K.; Ong, C.-T.; Hookway, T.A.; Guo, C.; Sun, Y.; et al. Architectural Protein Subclasses Shape 3D Organization of Genomes during Lineage Commitment. Cell 2013, 153, 1281–1295. [Google Scholar] [CrossRef]
  10. Spitz, F.; Furlong, E.E.M. Transcription factors: From enhancer binding to developmental control. Nat. Rev. Genet. 2012, 13, 613–626. [Google Scholar] [CrossRef]
  11. Furlong, E.E.M.; Levine, M. Developmental enhancers and chromosome topology. Science 2018, 361, 1341–1345. [Google Scholar] [CrossRef]
  12. Renfro, Z.; White, B.E.; Stephens, K.E. CCAAT enhancer binding protein gamma (C/EBP-γ): An understudied transcription factor. Adv. Biol. Regul. 2022, 84, 100861. [Google Scholar] [CrossRef]
  13. Heslop, J.A.; Pournasr, B.; Duncan, S.A. Chromatin remodeling is restricted by transient GATA6 binding during iPSC differentiation to definitive endoderm. iScience 2022, 25, 104300. [Google Scholar] [CrossRef] [PubMed]
  14. Liu, J.; Xiao, Q.; Xiao, J.; Niu, C.; Li, Y.; Zhang, X.; Zhou, Z.; Shu, G.; Yin, G. Wnt/β-catenin signalling: Function, biological mechanisms, and therapeutic opportunities. Signal Transduct. Target. Ther. 2022, 7, 3. [Google Scholar] [CrossRef]
  15. Swinstead, E.E.; Paakinaho, V.; Presman, D.M.; Hager, G.L. Pioneer factors and ATP-dependent chromatin remodeling factors interact dynamically: A new perspective: Multiple transcription factors can effect chromatin pioneer functions through dynamic interactions with ATP-dependent chromatin remodeling factors. Bioessays 2016, 38, 1150–1157. [Google Scholar] [CrossRef] [PubMed]
  16. Radman-Livaja, M.; Rando, O.J. Nucleosome positioning: How is it established, and why does it matter? Dev. Biol. 2010, 339, 258–266. [Google Scholar] [CrossRef] [PubMed]
  17. Ricci, M.A.; Manzo, C.; García-Parajo, M.F.; Lakadamyali, M.; Cosma, M.P. Chromatin fibers are formed by heterogeneous groups of nucleosomes in vivo. Cell 2015, 160, 1145–1158. [Google Scholar] [CrossRef]
  18. Portillo-Ledesma, S.; Tsao, L.H.; Wagley, M.; Lakadamyali, M.; Cosma, M.P.; Schlick, T. Nucleosome Clutches are Regulated by Chromatin Internal Parameters. J. Mol. Biol. 2021, 433, 166701. [Google Scholar] [CrossRef]
  19. Zentgraf, H.; Franke, W.W. Differences of supranucleosomal organization in different kinds of chromatin: Cell type-specific globular subunits containing different numbers of nucleosomes. J. Cell Biol. 1984, 99, 272–286. [Google Scholar] [CrossRef]
  20. Bannister, A.J.; Kouzarides, T. Regulation of chromatin by histone modifications. Cell Res. 2011, 21, 381–395. [Google Scholar] [CrossRef]
  21. Dixon, J.R.; Selvaraj, S.; Yue, F.; Kim, A.; Li, Y.; Shen, Y.; Hu, M.; Liu, J.S.; Ren, B. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 2012, 485, 376–380. [Google Scholar] [CrossRef]
  22. Nora, E.P.; Lajoie, B.R.; Schulz, E.G.; Giorgetti, L.; Okamoto, I.; Servant, N.; Piolot, T.; van Berkum, N.L.; Meisig, J.; Sedat, J.; et al. Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 2012, 485, 381–385. [Google Scholar] [CrossRef] [PubMed]
  23. Rowley, M.J.; Nichols, M.H.; Lyu, X.; Ando-Kuri, M.; Rivera, I.S.M.; Hermetz, K.; Wang, P.; Ruan, Y.; Corces, V.G. Evolutionarily Conserved Principles Predict 3D Chromatin Organization. Mol. Cell 2017, 67, 837–852.e837. [Google Scholar] [CrossRef] [PubMed]
  24. Huang, N.; Seow, W.Q.; Appert, A.; Dong, Y.; Stempor, P.; Ahringer, J. Accessible Region Conformation Capture (ARC-C) gives high-resolution insights into genome architecture and regulation. Genome Res. 2022, 32, 357–366. [Google Scholar] [CrossRef] [PubMed]
  25. Ghavi-Helm, Y.; Klein, F.A.; Pakozdi, T.; Ciglar, L.; Noordermeer, D.; Huber, W.; Furlong, E.E.M. Enhancer loops appear stable during development and are associated with paused polymerase. Nature 2014, 512, 96–100. [Google Scholar] [CrossRef] [PubMed]
  26. Guo, Y.; Xu, Q.; Canzio, D.; Shou, J.; Li, J.; Gorkin, D.U.; Jung, I.; Wu, H.; Zhai, Y.; Tang, Y.; et al. CRISPR Inversion of CTCF Sites Alters Genome Topology and Enhancer/Promoter Function. Cell 2015, 162, 900–910. [Google Scholar] [CrossRef] [PubMed]
  27. Lupiáñez, D.G.; Kraft, K.; Heinrich, V.; Krawitz, P.; Brancati, F.; Klopocki, E.; Horn, D.; Kayserili, H.; Opitz, J.M.; Laxova, R.; et al. Disruptions of Topological Chromatin Domains Cause Pathogenic Rewiring of Gene-Enhancer Interactions. Cell 2015, 161, 1012–1025. [Google Scholar] [CrossRef]
  28. Franke, M.; Ibrahim, D.M.; Andrey, G.; Schwarzer, W.; Heinrich, V.; Schöpflin, R.; Kraft, K.; Kempfer, R.; Jerković, I.; Chan, W.-L.; et al. Formation of new chromatin domains determines pathogenicity of genomic duplications. Nature 2016, 538, 265–269. [Google Scholar] [CrossRef]
  29. Yu, M.; Ren, B. The Three-Dimensional Organization of Mammalian Genomes. Annu. Rev. Cell Dev. Biol. 2017, 33, 265–289. [Google Scholar] [CrossRef]
  30. Li, D.; He, M.; Tang, Q.; Tian, S.; Zhang, J.; Li, Y.; Wang, D.; Jin, L.; Ning, C.; Zhu, W.; et al. Comparative 3D genome architecture in vertebrates. BMC Biol. 2022, 20, 99. [Google Scholar] [CrossRef]
  31. Wang, C.; Liu, C.; Roqueiro, D.; Grimm, D.; Schwab, R.; Becker, C.; Lanz, C.; Weigel, D. Genome-wide analysis of local chromatin packing in Arabidopsis thaliana. Genome Res. 2015, 25, 246–256. [Google Scholar] [CrossRef]
  32. Rao, S.S.P.; Huang, S.C.; Glenn St Hilaire, B.; Engreitz, J.M.; Perez, E.M.; Kieffer-Kwon, K.R.; Sanborn, A.L.; Johnstone, S.E.; Bascom, G.D.; Bochkov, I.D.; et al. Cohesin Loss Eliminates All Loop Domains. Cell 2017, 171, 305–320.e324. [Google Scholar] [CrossRef]
  33. Nora, E.P.; Goloborodko, A.; Valton, A.L.; Gibcus, J.H.; Uebersohn, A.; Abdennur, N.; Dekker, J.; Mirny, L.A.; Bruneau, B.G. Targeted Degradation of CTCF Decouples Local Insulation of Chromosome Domains from Genomic Compartmentalization. Cell 2017, 169, 930–944.e922. [Google Scholar] [CrossRef]
  34. Lieberman-Aiden, E.; van Berkum, N.L.; Williams, L.; Imakaev, M.; Ragoczy, T.; Telling, A.; Amit, I.; Lajoie, B.R.; Sabo, P.J.; Dorschner, M.O.; et al. Comprehensive Mapping of Long-Range Interactions Reveals Folding Principles of the Human Genome. Science 2009, 326, 289–293. [Google Scholar] [CrossRef] [PubMed]
  35. Sexton, T.; Yaffe, E.; Kenigsberg, E.; Bantignies, F.; Leblanc, B.; Hoichman, M.; Parrinello, H.; Tanay, A.; Cavalli, G. Three-Dimensional Folding and Functional Organization Principles of the Drosophila Genome. Cell 2012, 148, 458–472. [Google Scholar] [CrossRef] [PubMed]
  36. Krietenstein, N.; Abraham, S.; Venev, S.V.; Abdennur, N.; Gibcus, J.; Hsieh, T.-H.S.; Parsi, K.M.; Yang, L.; Maehr, R.; Mirny, L.A.; et al. Ultrastructural Details of Mammalian Chromosome Architecture. Mol. Cell 2020, 78, 554–565.e557. [Google Scholar] [CrossRef]
  37. Hsieh, T.-H.S.; Cattoglio, C.; Slobodyanyuk, E.; Hansen, A.S.; Rando, O.J.; Tjian, R.; Darzacq, X. Resolving the 3D Landscape of Transcription-Linked Mammalian Chromatin Folding. Mol. Cell 2020, 78, 539–553.e538. [Google Scholar] [CrossRef]
  38. Harris, H.L.; Gu, H.; Olshansky, M.; Wang, A.; Farabella, I.; Eliaz, Y.; Kalluchi, A.; Krishna, A.; Jacobs, M.; Cauer, G.; et al. Chromatin alternates between A and B compartments at kilobase scale for subgenic organization. Nat. Commun. 2023, 14, 3303. [Google Scholar] [CrossRef]
  39. Cremer, T.; Cremer, M. Chromosome territories. Cold Spring Harb. Perspect. Biol. 2010, 2, a003889. [Google Scholar] [CrossRef]
  40. Branco, M.R.; Pombo, A. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biol. 2006, 4, e138. [Google Scholar] [CrossRef] [PubMed]
  41. Pecinka, A.; Schubert, V.; Meister, A.; Kreth, G.; Klatte, M.; Lysak, M.A.; Fuchs, J.; Schubert, I. Chromosome territory arrangement and homologous pairing in nuclei of Arabidopsis thaliana are predominantly random except for NOR-bearing chromosomes. Chromosoma 2004, 113, 258–269. [Google Scholar] [CrossRef]
  42. Zink, D.; Cremer, T.; Saffrich, R.; Fischer, R.; Trendelenburg, M.F.; Ansorge, W.; Stelzer, E.H. Structure and dynamics of human interphase chromosome territories in vivo. Hum. Genet. 1998, 102, 241–251. [Google Scholar] [CrossRef]
  43. Croft, J.A.; Bridger, J.M.; Boyle, S.; Perry, P.; Teague, P.; Bickmore, W.A. Differences in the localization and morphology of chromosomes in the human nucleus. J. Cell Biol. 1999, 145, 1119–1131. [Google Scholar] [CrossRef]
  44. Fritz, A.J.; Sehgal, N.; Pliss, A.; Xu, J.; Berezney, R. Chromosome territories and the global regulation of the genome. Genes Chromosomes Cancer 2019, 58, 407–426. [Google Scholar] [CrossRef]
  45. Harvey, S.H.; Krien, M.J.; O’Connell, M.J. Structural maintenance of chromosomes (SMC) proteins, a family of conserved ATPases. Genome Biol. 2002, 3, Reviews3003. [Google Scholar] [CrossRef]
  46. Lammens, A.; Schele, A.; Hopfner, K.P. Structural biochemistry of ATP-driven dimerization and DNA-stimulated activation of SMC ATPases. Curr. Biol. 2004, 14, 1778–1782. [Google Scholar] [CrossRef]
  47. Palou, R.; Dhanaraman, T.; Marrakchi, R.; Pascariu, M.; Tyers, M.; D’Amours, D. Condensin ATPase motifs contribute differentially to the maintenance of chromosome morphology and genome stability. PLoS Biol. 2018, 16, e2003980. [Google Scholar] [CrossRef]
  48. Abdul, F.; Diman, A.; Baechler, B.; Ramakrishnan, D.; Kornyeyev, D.; Beran, R.K.; Fletcher, S.P.; Strubin, M. Smc5/6 silences episomal transcription by a three-step function. Nat. Struct. Mol. Biol. 2022, 29, 922–931. [Google Scholar] [CrossRef]
  49. Haering, C.H.; Lowe, J.; Hochwagen, A.; Nasmyth, K. Molecular Architecture of SMC Proteins and the Yeast Cohesin Complex. Mol. Cell 2002, 9, 773–788. [Google Scholar] [CrossRef] [PubMed]
  50. Jessberger, R. The many functions of smc proteins in chromosome dynamics. Nat. Rev. Mol. Cell Biol. 2002, 3, 767–778. [Google Scholar] [CrossRef] [PubMed]
  51. Duan, X.; Yang, Y.; Chen, Y.H.; Arenz, J.; Rangi, G.K.; Zhao, X.; Ye, H. Architecture of the Smc5/6 Complex of Saccharomyces cerevisiae Reveals a Unique Interaction between the Nse5-6 Subcomplex and the Hinge Regions of Smc5 and Smc6. J. Biol. Chem. 2009, 284, 8507–8515. [Google Scholar] [CrossRef] [PubMed]
  52. Hirano, T.; Kobayashi, R.; Hirano, M. Condensins, Chromosome Condensation Protein Complexes Containing XCAP-C, XCAP-E and a Xenopus Homolog of the Drosophila Barren Protein. Cell 1997, 89, 511–521. [Google Scholar] [CrossRef]
  53. Hassler, M.; Shaltiel, I.A.; Kschonsak, M.; Simon, B.; Merkel, F.; Tharichen, L.; Bailey, H.J.; Macosek, J.; Bravo, S.; Metz, J.; et al. Structural Basis of an Asymmetric Condensin ATPase Cycle. Mol. Cell 2019, 74, 1175–1188.e9. [Google Scholar] [CrossRef]
  54. Hirano, M.; Hirano, T. Opening closed arms: Long-distance activation of SMC ATPase by hinge-DNA interactions. Mol. Cell 2006, 21, 175–186. [Google Scholar] [CrossRef]
  55. Pradhan, B.; Kanno, T.; Umeda Igarashi, M.; Loke, M.S.; Baaske, M.D.; Wong, J.S.K.; Jeppsson, K.; Björkegren, C.; Kim, E. The Smc5/6 complex is a DNA loop-extruding motor. Nature 2023, 616, 843–848. [Google Scholar] [CrossRef]
  56. Cheng, H.; Zhang, N.; Pati, D. Cohesin subunit RAD21: From biology to disease. Gene 2020, 758, 144966. [Google Scholar] [CrossRef]
  57. Schleiffer, A.; Kaitna, S.; Maurer-Stroh, S.; Glotzer, M.; Nasmyth, K.; Eisenhaber, F. Kleisins: A Superfamily of Bacterial and Eukaryotic SMC Protein Partners. Mol. Cell 2003, 11, 571–575. [Google Scholar] [CrossRef]
  58. Mito, Y.; Sugimoto, A.; Yamamoto, M. Distinct developmental function of two Caenorhabditis elegans homologs of the cohesin subunit Scc1/Rad21. Mol. Biol. Cell 2003, 14, 2399–2409. [Google Scholar] [CrossRef] [PubMed]
  59. Severson, A.F.; Ling, L.; van Zuylen, V.; Meyer, B.J. The axial element protein HTP-3 promotes cohesin loading and meiotic axis assembly in C. elegans to implement the meiotic program of chromosome segregation. Genes. Dev. 2009, 23, 1763–1778. [Google Scholar] [CrossRef]
  60. Severson, A.F.; Meyer, B.J. Divergent kleisin subunits of cohesin specify mechanisms to tether and release meiotic chromosomes. eLife 2014, 3, e03467. [Google Scholar] [CrossRef] [PubMed]
  61. Brooker, A.S.; Berkowitz, K.M. The roles of cohesins in mitosis, meiosis, and human health and disease. Methods Mol. Biol. 2014, 1170, 229–266. [Google Scholar] [CrossRef] [PubMed]
  62. Cutts, E.E.; Vannini, A. Condensin complexes: Understanding loop extrusion one conformational change at a time. Biochem. Soc. Trans. 2020, 48, 2089–2100. [Google Scholar] [CrossRef] [PubMed]
  63. Tane, S.; Shintomi, K.; Kinoshita, K.; Tsubota, Y.; Yoshida, M.M.; Nishiyama, T.; Hirano, T. Cell cycle-specific loading of condensin I is regulated by the N-terminal tail of its kleisin subunit. eLife 2022, 11, e84694. [Google Scholar] [CrossRef] [PubMed]
  64. Shaltiel, I.A.; Datta, S.; Lecomte, L.; Hassler, M.; Kschonsak, M.; Bravo, S.; Stober, C.; Ormanns, J.; Eustermann, S.; Haering, C.H. A hold-and-feed mechanism drives directional DNA loop extrusion by condensin. Science 2022, 376, 1087–1094. [Google Scholar] [CrossRef]
  65. Palecek, J.J.; Gruber, S. Kite Proteins: A Superfamily of SMC/Kleisin Partners Conserved Across Bacteria, Archaea, and Eukaryotes. Structure 2015, 23, 2183–2190. [Google Scholar] [CrossRef] [PubMed]
  66. Wells, J.N.; Gligoris, T.G.; Nasmyth, K.A.; Marsh, J.A. Evolution of condensin and cohesin complexes driven by replacement of Kite by Hawk proteins. Curr. Biol. 2017, 27, R17–R18. [Google Scholar] [CrossRef] [PubMed]
  67. Ono, T.; Losada, A.; Hirano, M.; Myers, M.P.; Neuwald, A.F.; Hirano, T. Differential contributions of condensin I and condensin II to mitotic chromosome architecture in vertebrate cells. Cell 2003, 115, 109–121. [Google Scholar] [CrossRef]
  68. Yoshida, M.M.; Kinoshita, K.; Shintomi, K.; Aizawa, Y.; Hirano, T. Regulation of condensin II by self-suppression and release mechanisms. Mol. Biol. Cell 2023, 35, ar21. [Google Scholar] [CrossRef]
  69. Csankovszki, G.; Collette, K.; Spahl, K.; Carey, J.; Snyder, M.; Petty, E.; Patel, U.; Tabuchi, T.; Liu, H.; McLeod, I.; et al. Three distinct condensin complexes control C. elegans chromosome dynamics. Curr. Biol. 2009, 19, 9–19. [Google Scholar] [CrossRef]
  70. Solé-Soler, R.; Torres-Rosell, J. Smc5/6, an atypical SMC complex with two RING-type subunits. Biochem. Soc. Trans. 2020, 48, 2159–2171. [Google Scholar] [CrossRef] [PubMed]
  71. Pebernard, S.; Wohlschlegel, J.; McDonald, W.H.; Yates, J.R., 3rd; Boddy, M.N. The Nse5-Nse6 dimer mediates DNA repair roles of the Smc5-Smc6 complex. Mol. Cell Biol. 2006, 26, 1617–1630. [Google Scholar] [CrossRef] [PubMed]
  72. Peng, X.P.; Zhao, X. The multi-functional Smc5/6 complex in genome protection and disease. Nat. Struct. Mol. Biol. 2023, 30, 724–734. [Google Scholar] [CrossRef]
  73. Odiba, A.S.; Ezechukwu, C.S.; Liao, G.; Li, S.; Chen, Z.; Liu, X.; Fang, W.; Jin, C.; Wang, B. Loss of NSE-4 Perturbs Genome Stability and DNA Repair in Caenorhabditis elegans. Int. J. Mol. Sci. 2022, 23, 7202. [Google Scholar] [CrossRef]
  74. Hong, Y.; Sonneville, R.; Agostinho, A.; Meier, B.; Wang, B.; Blow, J.J.; Gartner, A. The SMC-5/6 Complex and the HIM-6 (BLM) Helicase Synergistically Promote Meiotic Recombination Intermediate Processing and Chromosome Maturation during Caenorhabditis elegans Meiosis. PLoS Genet. 2016, 12, e1005872. [Google Scholar] [CrossRef]
  75. Barra, V.; Fachinetti, D. The dark side of centromeres: Types, causes and consequences of structural abnormalities implicating centromeric DNA. Nat. Commun. 2018, 9, 4340. [Google Scholar] [CrossRef]
  76. Kyriacou, E.; Heun, P. Centromere structure and function: Lessons from Drosophila. Genetics 2023, 225, iyad170. [Google Scholar] [CrossRef]
  77. Simon, L.; Voisin, M.; Tatout, C.; Probst, A.V. Structure and Function of Centromeric and Pericentromeric Heterochromatin in Arabidopsis thaliana. Front. Plant Sci. 2015, 6, 1049. [Google Scholar] [CrossRef]
  78. Xu, R.; Pan, Z.; Nakagawa, T. Gross Chromosomal Rearrangement at Centromeres. Biomolecules 2023, 14, 28. [Google Scholar] [CrossRef]
  79. Melters, D.P.; Paliulis, L.V.; Korf, I.F.; Chan, S.W.L. Holocentric chromosomes: Convergent evolution, meiotic adaptations, and genomic analysis. Chromosome Res. 2012, 20, 579–593. [Google Scholar] [CrossRef]
  80. Emerson, F.J.; Lee, S.S. Chromatin: The old and young of it. Front. Mol. Biosci. 2023, 10, 1270285. [Google Scholar] [CrossRef]
  81. Márquez-Corro, J.I.; Escudero, M.; Luceño, M. Do holocentric chromosomes represent an evolutionary advantage? A study of paired analyses of diversification rates of lineages with holocentric chromosomes and their monocentric closest relatives. Chromosome Res. 2018, 26, 139–152. [Google Scholar] [CrossRef]
  82. Nabeshima, K.; Villeneuve, A.M.; Colaiácovo, M.P. Crossing over is coupled to late meiotic prophase bivalent differentiation through asymmetric disassembly of the SC. J. Cell Biol. 2005, 168, 683–689. [Google Scholar] [CrossRef]
  83. Chan, R.C.; Severson, A.F.; Meyer, B.J. Condensin restructures chromosomes in preparation for meiotic divisions. J. Cell Biol. 2004, 167, 613–625. [Google Scholar] [CrossRef]
  84. Hillers, K.J.; Villeneuve, A.M. Chromosome-Wide Control of Meiotic Crossing over in C. elegans. Curr. Biol. 2003, 13, 1641–1647. [Google Scholar] [CrossRef]
  85. Schvarzstein, M.; Wignall, S.M.; Villeneuve, A.M. Coordinating cohesion, co-orientation, and congression during meiosis: Lessons from holocentric chromosomes. Genes. Dev. 2010, 24, 219–228. [Google Scholar] [CrossRef]
  86. Ganji, M.; Shaltiel, I.A.; Bisht, S.; Kim, E.; Kalichava, A.; Haering, C.H.; Dekker, C. Real-time imaging ofDNA loop extrusion by condensin. Science 2018, 360, 102–105. [Google Scholar] [CrossRef]
  87. Kim, Y.; Shi, Z.; Zhang, H.; Finkelstein, I.J.; Yu, H. Human cohesin compacts DNA by loop extrusion. Science 2019, 366, 1345–1349. [Google Scholar] [CrossRef]
  88. Higashi, T.L.; Pobegalov, G.; Tang, M.; Molodtsov, M.I.; Uhlmann, F. A Brownian ratchet model for DNA loop extrusion by the cohesin complex. eLife 2021, 10, e67530. [Google Scholar] [CrossRef]
  89. Marko, J.F.; De Los Rios, P.; Barducci, A.; Gruber, S. DNA-segment-capture model for loop extrusion by structural maintenance of chromosome (SMC) protein complexes. Nucleic Acids Res. 2019, 47, 6956–6972. [Google Scholar] [CrossRef]
  90. Nomidis, S.K.; Carlon, E.; Gruber, S.; Marko, J.F. DNA tension-modulated translocation and loop extrusion by SMC complexes revealed by molecular dynamics simulations. Nucleic Acids Res. 2022, 50, 4974–4987. [Google Scholar] [CrossRef]
  91. Dekker, C.; Haering, C.H.; Peters, J.-M.; Rowland, B.D. How do molecular motors fold the genome? Science 2023, 382, 646–648. [Google Scholar] [CrossRef]
  92. Hagstrom, K.A.; Holmes, V.F.; Cozzarelli, N.R.; Meyer, B.J. C. elegans condensin promotes mitotic chromosome architecture, centromere organization, and sister chromatid segregation during mitosis and meiosis. Genes. Dev. 2002, 16, 729–742. [Google Scholar] [CrossRef]
  93. Uhlmann, F.; Nasmyth, K. Cohesion between sister chromatids must be established during DNA replication. Curr. Biol. 1998, 8, 1095–1102. [Google Scholar] [CrossRef] [PubMed]
  94. Chan, R.C.; Chan, A.; Jeon, M.; Wu, T.F.; Pasqualone, D.; Rougvie, A.E.; Meyer, B.J. Chromosome cohesion is regulated by a clock gene paralogue TIM-1. Nature 2003, 423, 1002–1009. [Google Scholar] [CrossRef]
  95. Pasierbek, P.; Födermayr, M.; Jantsch, V.; Jantsch, M.; Schweizer, D.; Loidl, J. The Caenorhabditis elegans SCC-3 homologue is required for meiotic synapsis and for proper chromosome disjunction in mitosis and meiosis. Exp. Cell Res. 2003, 289, 245–255. [Google Scholar] [CrossRef]
  96. Siomos, M.F.; Badrinath, A.; Pasierbek, P.; Livingstone, D.; White, J.; Glotzer, M.; Nasmyth, K. Separase is required for chromosome segregation during meiosis I in Caenorhabditis elegans. Curr. Biol. 2001, 11, 1825–1835. [Google Scholar] [CrossRef]
  97. Luo, S.; Tong, L. Structure and Function of the Separase-Securin Complex. Subcell. Biochem. 2021, 96, 217–232. [Google Scholar] [CrossRef]
  98. Michaelis, C.; Ciosk, R.; Nasmyth, K. Cohesins: Chromosomal proteins that prevent premature separation of sister chromatids. Cell 1997, 91, 35–45. [Google Scholar] [CrossRef]
  99. Ishiguro, K.I. The cohesin complex in mammalian meiosis. Genes. Cells 2019, 24, 6–30. [Google Scholar] [CrossRef]
  100. Pasierbek, P.; Jantsch, M.; Melcher, M.; Schleiffer, A.; Schweizer, D.; Loidl, J. A Caenorhabditis elegans cohesion protein with functions in meiotic chromosome pairing and disjunction. Genes. Dev. 2001, 15, 1349–1360. [Google Scholar] [CrossRef]
  101. Castellano-Pozo, M.; Sioutas, G.; Barroso, C.; Prince, J.P.; Lopez-Jimenez, P.; Davy, J.; Jaso-Tamame, A.-L.; Crawley, O.; Shao, N.; Page, J.; et al. The kleisin subunit controls the function of C. elegans meiotic cohesins by determining the mode of DNA binding and differential regulation by SCC-2 and WAPL-1. eLife 2023, 12, e84138. [Google Scholar] [CrossRef]
  102. Klein, F.; Mahr, P.; Galova, M.; Buonomo, S.B.; Michaelis, C.; Nairz, K.; Nasmyth, K. A central role for cohesins in sister chromatid cohesion, formation of axial elements, and recombination during yeast meiosis. Cell 1999, 98, 91–103. [Google Scholar] [CrossRef]
  103. Kitajima, T.S.; Yokobayashi, S.; Yamamoto, M.; Watanabe, Y. Distinct cohesin complexes organize meiotic chromosome domains. Science 2003, 300, 1152–1155. [Google Scholar] [CrossRef]
  104. Ono, T.; Fang, Y.; Spector, D.L.; Hirano, T. Spatial and temporal regulation of Condensins I and II in mitotic chromosome assembly in human cells. Mol. Biol. Cell 2004, 15, 3296–3308. [Google Scholar] [CrossRef] [PubMed]
  105. Collette, K.S.; Petty, E.L.; Golenberg, N.; Bembenek, J.N.; Csankovszki, G. Different roles for Aurora B in condensin targeting during mitosis and meiosis. J. Cell Sci. 2011, 124, 3684–3694. [Google Scholar] [CrossRef]
  106. Shintomi, K.; Hirano, T. The relative ratio of condensin I to II determines chromosome shapes. Genes. Dev. 2011, 25, 1464–1469. [Google Scholar] [CrossRef]
  107. Bembenek, J.N.; Verbrugghe, K.J.; Khanikar, J.; Csankovszki, G.; Chan, R.C. Condensin and the spindle midzone prevent cytokinesis failure induced by chromatin bridges in C. elegans embryos. Curr. Biol. 2013, 23, 937–946. [Google Scholar] [CrossRef]
  108. Golfier, S.; Quail, T.; Kimura, H.; Brugues, J. Cohesin and condensin extrude DNA loops in a cell cycle-dependent manner. Elife 2020, 9, e53885. [Google Scholar] [CrossRef]
  109. Lee, J.; Ogushi, S.; Saitou, M.; Hirano, T. Condensins I and II are essential for construction of bivalent chromosomes in mouse oocytes. Mol. Biol. Cell 2011, 22, 3465–3477. [Google Scholar] [CrossRef] [PubMed]
  110. Smith, S.J.; Osman, K.; Franklin, F.C. The condensin complexes play distinct roles to ensure normal chromosome morphogenesis during meiotic division in Arabidopsis. Plant J. 2014, 80, 255–268. [Google Scholar] [CrossRef] [PubMed]
  111. Resnick, T.D.; Dej, K.J.; Xiang, Y.; Hawley, R.S.; Ahn, C.; Orr-Weaver, T.L. Mutations in the chromosomal passenger complex and the condensin complex differentially affect synaptonemal complex disassembly and metaphase I configuration in Drosophila female meiosis. Genetics 2009, 181, 875–887. [Google Scholar] [CrossRef]
  112. Hernandez, M.R.; Davis, M.B.; Jiang, J.; Brouhard, E.A.; Severson, A.F.; Csankovszki, G. Condensin I protects meiotic cohesin from WAPL-1 mediated removal. PLoS Genet. 2018, 14, e1007382. [Google Scholar] [CrossRef]
  113. Dixon, J.R.; Gorkin, D.U.; Ren, B. Chromatin Domains: The Unit of Chromosome Organization. Mol. Cell 2016, 62, 668–680. [Google Scholar] [CrossRef]
  114. Dekker, J.; Heard, E. Structural and functional diversity of Topologically Associating Domains. FEBS Lett. 2015, 589, 2877–2884. [Google Scholar] [CrossRef]
  115. Makrantoni, V.; Marston, A.L. Cohesin and chromosome segregation. Curr. Biol. 2018, 28, R688–R693. [Google Scholar] [CrossRef]
  116. Holwerda, S.J.; de Laat, W. CTCF: The protein, the binding partners, the binding sites and their chromatin loops. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2013, 368, 20120369. [Google Scholar] [CrossRef] [PubMed]
  117. Ong, C.-T.; Corces, V.G. CTCF: An architectural protein bridging genome topology and function. Nat. Rev. Genet. 2014, 15, 234–246. [Google Scholar] [CrossRef] [PubMed]
  118. Crane, E.; Bian, Q.; McCord, R.P.; Lajoie, B.R.; Wheeler, B.S.; Ralston, E.J.; Uzawa, S.; Dekker, J.; Meyer, B.J. Condensin-driven remodelling of X chromosome topology during dosage compensation. Nature 2015, 523, 240–244. [Google Scholar] [CrossRef] [PubMed]
  119. Ikegami, K.; Egelhofer, T.A.; Strome, S.; Lieb, J.D. Caenorhabditis elegans chromosome arms are anchored to the nuclear membrane via discontinuous association with LEM-2. Genome Biol. 2010, 11, R120. [Google Scholar] [CrossRef]
  120. Cabianca, D.S.; Muñoz-Jiménez, C.; Kalck, V.; Gaidatzis, D.; Padeken, J.; Seeber, A.; Askjaer, P.; Gasser, S.M. Active chromatin marks drive spatial sequestration of heterochromatin in C. elegans nuclei. Nature 2019, 569, 734–739. [Google Scholar] [CrossRef]
  121. Towbin, B.D.; González-Aguilera, C.; Sack, R.; Gaidatzis, D.; Kalck, V.; Meister, P.; Askjaer, P.; Gasser, S.M. Step-wise methylation of histone H3K9 positions heterochromatin at the nuclear periphery. Cell 2012, 150, 934–947. [Google Scholar] [CrossRef]
  122. Gonzalez-Sandoval, A.; Towbin, B.D.; Kalck, V.; Cabianca, D.S.; Gaidatzis, D.; Hauer, M.H.; Geng, L.; Wang, L.; Yang, T.; Wang, X.; et al. Perinuclear Anchoring of H3K9-Methylated Chromatin Stabilizes Induced Cell Fate in C. elegans Embryos. Cell 2015, 163, 1333–1347. [Google Scholar] [CrossRef]
  123. Alagna, N.S.; Thomas, T.I.; Wilson, K.L.; Reddy, K.L. Choreography of lamina-associated domains: Structure meets dynamics. FEBS Lett. 2023, 597, 2806–2822. [Google Scholar] [CrossRef]
  124. Wang, N.; Wang, Z.; Tzourtzou, S.; Wang, X.; Bi, X.; Leimeister, J.; Xu, L.; Sakamoto, T.; Matsunaga, S.; Schaller, A.; et al. The plant nuclear lamina disassembles to regulate genome folding in stress conditions. Nat. Plants 2023, 9, 1081–1093. [Google Scholar] [CrossRef]
  125. Liu, T.; Rechtsteiner, A.; Egelhofer, T.A.; Vielle, A.; Latorre, I.; Cheung, M.S.; Ercan, S.; Ikegami, K.; Jensen, M.; Kolasinska-Zwierz, P.; et al. Broad chromosomal domains of histone modification patterns in C. elegans. Genome Res. 2011, 21, 227–236. [Google Scholar] [CrossRef]
  126. Hassan, A.; Rodriguez, P.A.; Heidmann, S.K.; Walmsley, E.L.; Aughey, G.N.; Southall, T.D. Condensin I subunit Cap-G is essential for proper gene expression during the maturation of post-mitotic neurons. eLife 2020, 9, e55159. [Google Scholar] [CrossRef]
  127. Bauer, C.R.; Hartl, T.A.; Bosco, G. Condensin II promotes the formation of chromosome territories by inducing axial compaction of polyploid interphase chromosomes. PLoS Genet. 2012, 8, e1002873. [Google Scholar] [CrossRef]
  128. Iwasaki, O.; Tanizawa, H.; Kim, K.D.; Kossenkov, A.; Nacarelli, T.; Tashiro, S.; Majumdar, S.; Showe, L.C.; Zhang, R.; Noma, K.I. Involvement of condensin in cellular senescence through gene regulation and compartmental reorganization. Nat. Commun. 2019, 10, 5688. [Google Scholar] [CrossRef]
  129. Hoencamp, C.; Dudchenko, O.; Elbatsh, A.M.O.; Brahmachari, S.; Raaijmakers, J.A.; van Schaik, T.; Sedeño Cacciatore, Á.; Contessoto, V.G.; van Heesbeen, R.; van den Broek, B.; et al. 3D genomics across the tree of life reveals condensin II as a determinant of architecture type. Science 2021, 372, 984–989. [Google Scholar] [CrossRef]
  130. Jin, Q.W.; Fuchs, J.; Loidl, J. Centromere clustering is a major determinant of yeast interphase nuclear organization. J. Cell Sci. 2000, 113, 1903–1912. [Google Scholar] [CrossRef]
  131. Sakamoto, T.; Sugiyama, T.; Yamashita, T.; Matsunaga, S. Plant condensin II is required for the correct spatial relationship between centromeres and rDNA arrays. Nucleus 2019, 10, 116–125. [Google Scholar] [CrossRef]
  132. Schuster, A.T.; Sarvepalli, K.; Murphy, E.A.; Longworth, M.S. Condensin II subunit dCAP-D3 restricts retrotransposon mobilization in Drosophila somatic cells. PLoS Genet. 2013, 9, e1003879. [Google Scholar] [CrossRef]
  133. Kranz, A.L.; Jiao, C.Y.; Winterkorn, L.H.; Albritton, S.E.; Kramer, M.; Ercan, S. Genome-wide analysis of condensin binding in Caenorhabditis elegans. Genome Biol. 2013, 14, R112. [Google Scholar] [CrossRef] [PubMed]
  134. Madl, J.E.; Herman, R.K. Polyploids and Sex Determination in Caenorhabditis elegans. Genetics 1979, 93, 393–402. [Google Scholar] [CrossRef] [PubMed]
  135. Brockdorff, N.; Turner, B.M. Dosage compensation in mammals. Cold Spring Harb. Perspect. Biol. 2015, 7, a019406. [Google Scholar] [CrossRef] [PubMed]
  136. Lucchesi, J.C.; Kuroda, M.I. Dosage compensation in Drosophila. Cold Spring Harb. Perspect. Biol. 2015, 7, a019398. [Google Scholar] [CrossRef]
  137. Lau, A.C.; Csankovszki, G. Balancing up and downregulation of the C. elegans X chromosomes. Curr. Opin. Genet. Dev. 2015, 31, 50–56. [Google Scholar] [CrossRef]
  138. Sanderson, B.J.; Feng, G.; Hu, N.; Carlson, C.H.; Smart, L.B.; Keefover-Ring, K.; Yin, T.; Ma, T.; Liu, J.; DiFazio, S.P.; et al. Sex determination through X-Y heterogamety in Salix nigra. Heredity 2021, 126, 630–639. [Google Scholar] [CrossRef] [PubMed]
  139. Hayman, E.S.; Fairgrieve, W.T.; Luckenbach, J.A. Molecular and morphological sex differentiation in sablefish (Anoplopoma fimbria), a marine teleost with XX/XY sex determination. Gene 2021, 764, 145093. [Google Scholar] [CrossRef]
  140. Nakamura, M. Sex determination in amphibians. Semin. Cell Dev. Biol. 2009, 20, 271–282. [Google Scholar] [CrossRef]
  141. Irwin, D.E. Sex chromosomes and speciation in birds and other ZW systems. Mol. Ecol. 2018, 27, 3831–3851. [Google Scholar] [CrossRef]
  142. Chandler, C.H. When and why does sex chromosome dosage compensation evolve? Ann. N. Y. Acad. Sci. 2017, 1389, 37–51. [Google Scholar] [CrossRef] [PubMed]
  143. Lau, A.C.; Nabeshima, K.; Csankovszki, G. The C. elegans dosage compensation complex mediates interphase X chromosome compaction. Epigenetics Chromatin 2014, 7, 31. [Google Scholar] [CrossRef] [PubMed]
  144. Meyer, B.J. Mechanisms of sex determination and X-chromosome dosage compensation. Genetics 2022, 220, iyab197. [Google Scholar] [CrossRef] [PubMed]
  145. Chuang, P.-T.; Lieb, J.D.; Meyer, B.J. Sex-Specific Assembly of a Dosage Compensation Complex on the Nematode X Chromosome. Science 1996, 274, 1736–1739. [Google Scholar] [CrossRef] [PubMed]
  146. Lieb, J.D.; Albrecht, M.R.; Chuang, P.T.; Meyer, B.J. MIX-1: An essential component of the C. elegans mitotic machinery executes X chromosome dosage compensation. Cell 1998, 92, 265–277. [Google Scholar] [CrossRef] [PubMed]
  147. Lieb, J.D.; Capowski, E.E.; Meneely, P.; Meyer, B.J. DPY-26, a link between dosage compensation and meiotic chromosome segregation in the nematode. Science 1996, 274, 1732–1736. [Google Scholar] [CrossRef] [PubMed]
  148. Nusbaum, C.; Meyer, B.J. The Caenorhabditis elegans gene sdc-2 controls sex determination and dosage compensation in XX animals. Genetics 1989, 122, 579–593. [Google Scholar] [CrossRef] [PubMed]
  149. Villeneuve, A.M.; Meyer, B.J. sdc-1: A link between sex determination and dosage compensation in C. elegans. Cell 1987, 48, 25–37. [Google Scholar] [CrossRef]
  150. DeLong, L.; Plenefisch, J.D.; Klein, R.D.; Meyer, B.J. Feedback control of sex determination by dosage compensation revealed through Caenorhabditis elegans sdc-3 mutations. Genetics 1993, 133, 875–896. [Google Scholar] [CrossRef]
  151. Yonker, S.A.; Meyer, B.J. Recruitment of C. elegans dosage compensation proteins for gene-specific versus chromosome-wide repression. Development 2003, 130, 6519–6532. [Google Scholar] [CrossRef]
  152. Brejc, K.; Bian, Q.; Uzawa, S.; Wheeler, B.S.; Anderson, E.C.; King, D.S.; Kranzusch, P.J.; Preston, C.G.; Meyer, B.J. Dynamic Control of X Chromosome Conformation and Repression by a Histone H4K20 Demethylase. Cell 2017, 171, 85–102.e23. [Google Scholar] [CrossRef]
  153. Pferdehirt, R.R.; Kruesi, W.S.; Meyer, B.J. An MLL/COMPASS subunit functions in the C. elegans dosage compensation complex to target X chromosomes for transcriptional regulation of gene expression. Genes. Dev. 2011, 25, 499–515. [Google Scholar] [CrossRef]
  154. Dong, X.; Peng, Y.; Peng, Y.; Xu, F.; He, X.; Wang, F.; Peng, X.; Qiang, B.; Yuan, J.; Rao, Z. Characterization and crystallization of human DPY-30-like protein, an essential component of dosage compensation complex. Biochim. Biophys. Acta (BBA) Proteins Proteom. 2005, 1753, 257–262. [Google Scholar] [CrossRef] [PubMed]
  155. Tremblay, V.; Zhang, P.; Chaturvedi, C.P.; Thornton, J.; Brunzelle, J.S.; Skiniotis, G.; Shilatifard, A.; Brand, M.; Couture, J.F. Molecular basis for DPY-30 association to COMPASS-like and NURF complexes. Structure 2014, 22, 1821–1830. [Google Scholar] [CrossRef] [PubMed]
  156. Petty, E.; Laughlin, E.; Csankovszki, G. Regulation of DCC localization by HTZ-1/H2A.Z and DPY-30 does not correlate with H3K4 methylation levels. PLoS ONE 2011, 6, e25973. [Google Scholar] [CrossRef]
  157. Custer, L.M.; Snyder, M.J.; Flegel, K.; Csankovszki, G. The onset of C. elegans dosage compensation is linked to the loss of developmental plasticity. Dev. Biol. 2014, 385, 279–290. [Google Scholar] [CrossRef]
  158. Dawes, H.E.; Berlin, D.S.; Lapidus, D.M.; Nusbaum, C.; Davis, T.L.; Meyer, B.J. Dosage Compensation Proteins Targeted to X Chromosomes by a Determinant of Hermaphrodite Fate. Science 1999, 284, 1800–1804. [Google Scholar] [CrossRef]
  159. McDonel, P.; Jans, J.; Peterson, B.K.; Meyer, B.J. Clustered DNA motifs mark X chromosomes for repression by a dosage compensation complex. Nature 2006, 444, 614–618. [Google Scholar] [CrossRef]
  160. Jans, J.; Gladden, J.M.; Ralston, E.J.; Pickle, C.S.; Michel, A.H.; Pferdehirt, R.R.; Eisen, M.B.; Meyer, B.J. A condensin-like dosage compensation complex acts at a distance to control expression throughout the genome. Genes. Dev. 2009, 23, 602–618. [Google Scholar] [CrossRef] [PubMed]
  161. Fuda, N.J.; Brejc, K.; Kruesi, W.S.; Ralston, E.J.; Bigley, R.; Shin, A.; Okada, M.; Meyer, B.J. Combinatorial clustering of distinct DNA motifs directs synergistic binding of Caenorhabditis elegans dosage compensation complex to X chromosomes. Proc. Natl. Acad. Sci. USA 2022, 119, e2211642119. [Google Scholar] [CrossRef]
  162. Albritton, S.E.; Kranz, A.L.; Winterkorn, L.H.; Street, L.A.; Ercan, S. Cooperation between a hierarchical set of recruitment sites targets the X chromosome for dosage compensation. eLife 2017, 6, e23645. [Google Scholar] [CrossRef] [PubMed]
  163. Ercan, S.; Giresi, P.G.; Whittle, C.M.; Zhang, X.; Green, R.D.; Lieb, J.D. X chromosome repression by localization of the C. elegans dosage compensation machinery to sites of transcription initiation. Nat. Genet. 2007, 39, 403–408. [Google Scholar] [CrossRef]
  164. Kim, J.; Jimenez, D.S.; Ragipani, B.; Zhang, B.; Street, L.A.; Kramer, M.; Albritton, S.E.; Winterkorn, L.; Morao, A.; Ercan, S. Condensin DC loads and spreads from recruitment sites to create loop- anchored TADs in C. elegans. eLife 2022, 11, e68745. [Google Scholar] [CrossRef]
  165. Csankovszki, G.; McDonel, P.; Meyer, B.J. Recruitment and spreading of the C. elegans dosage compensation complex along X chromosomes. Science 2004, 303, 1182–1185. [Google Scholar] [CrossRef]
  166. Anderson, E.C.; Frankino, P.A.; Higuchi-Sanabria, R.; Yang, Q.; Bian, Q.; Podshivalova, K.; Shin, A.; Kenyon, C.; Dillin, A.; Meyer, B.J. X Chromosome Domain Architecture Regulates Caenorhabditis elegans Lifespan but Not Dosage Compensation. Dev. Cell 2019, 51, 192–207.e196. [Google Scholar] [CrossRef]
  167. Rowley, M.J.; Poulet, A.; Nichols, M.H.; Bixler, B.J.; Sanborn, A.L.; Brouhard, E.A.; Hermetz, K.; Linsenbaum, H.; Csankovszki, G.; Lieberman Aiden, E.; et al. Analysis of Hi-C data using SIP effectively identifies loops in organisms from C. elegans to mammals. Genome Res. 2020, 30, 447–458. [Google Scholar] [CrossRef] [PubMed]
  168. Petit, F.; Sears, K.E.; Ahituv, N. Limb development: A paradigm of gene regulation. Nat. Rev. Genet. 2017, 18, 245–258. [Google Scholar] [CrossRef] [PubMed]
  169. Arvanitis, M.; Tayeb, K.; Strober, B.J.; Battle, A. Redefining tissue specificity of genetic regulation of gene expression in the presence of allelic heterogeneity. Am. J. Hum. Genet. 2022, 109, 223–239. [Google Scholar] [CrossRef]
  170. Chang, Y.N.; Zhu, C.; Jiang, J.; Zhang, H.; Zhu, J.K.; Duan, C.G. Epigenetic regulation in plant abiotic stress responses. J. Integr. Plant Biol. 2020, 62, 563–580. [Google Scholar] [CrossRef]
  171. Kruesi, W.S.; Core, L.J.; Waters, C.T.; Lis, J.T.; Meyer, B.J. Condensin controls recruitment of RNA polymerase II to achieve nematode X-chromosome dosage compensation. eLife 2013, 2, e00808. [Google Scholar] [CrossRef]
  172. Tzatsos, A.; Paskaleva, P.; Ferrari, F.; Deshpande, V.; Stoykova, S.; Contino, G.; Wong, K.K.; Lan, F.; Trojer, P.; Park, P.J.; et al. KDM2B promotes pancreatic cancer via Polycomb-dependent and -independent transcriptional programs. J. Clin. Investig. 2013, 123, 727–739. [Google Scholar] [CrossRef]
  173. Ward, J.R.; Vasu, K.; Deutschman, E.; Halawani, D.; Larson, P.A.; Zhang, D.; Willard, B.; Fox, P.L.; Moran, J.V.; Longworth, M.S. Condensin II and GAIT complexes cooperate to restrict LINE-1 retrotransposition in epithelial cells. PLoS Genet. 2017, 13, e1007051. [Google Scholar] [CrossRef] [PubMed]
  174. Vielle, A.; Lang, J.; Dong, Y.; Ercan, S.; Kotwaliwale, C.; Rechtsteiner, A.; Appert, A.; Chen, Q.B.; Dose, A.; Egelhofer, T.; et al. H4K20me1 contributes to downregulation of X-linked genes for C. elegans dosage compensation. PLoS Genet. 2012, 8, e1002933. [Google Scholar] [CrossRef] [PubMed]
  175. Wells, M.B.; Snyder, M.J.; Custer, L.M.; Csankovszki, G. Caenorhabditis elegans dosage compensation regulates histone H4 chromatin state on X chromosomes. Mol. Cell Biol. 2012, 32, 1710–1719. [Google Scholar] [CrossRef] [PubMed]
  176. Budhavarapu, V.N.; Chavez, M.; Tyler, J.K. How is epigenetic information maintained through DNA replication? Epigenetics Chromatin 2013, 6, 32. [Google Scholar] [CrossRef] [PubMed]
  177. Corvalan, A.Z.; Coller, H.A. Methylation of histone 4’s lysine 20: A critical analysis of the state of the field. Physiol. Genom. 2021, 53, 22–32. [Google Scholar] [CrossRef] [PubMed]
  178. Oda, H.; Okamoto, I.; Murphy, N.; Chu, J.; Price, S.M.; Shen, M.M.; Torres-Padilla, M.E.; Heard, E.; Reinberg, D. Monomethylation of histone H4-lysine 20 is involved in chromosome structure and stability and is essential for mouse development. Mol. Cell Biol. 2009, 29, 2278–2295. [Google Scholar] [CrossRef] [PubMed]
  179. Snyder, M.J.; Lau, A.C.; Brouhard, E.A.; Davis, M.B.; Jiang, J.; Sifuentes, M.H.; Csankovszki, G. Anchoring of Heterochromatin to the Nuclear Lamina Reinforces Dosage Compensation-Mediated Gene Repression. PLoS Genet. 2016, 12, e1006341. [Google Scholar] [CrossRef] [PubMed]
  180. Poleshko, A.; Katz, R.A. Specifying peripheral heterochromatin during nuclear lamina reassembly. Nucleus 2014, 5, 32–39. [Google Scholar] [CrossRef]
  181. Davis, M.B.; Jash, E.; Chawla, B.; Haines, R.A.; Tushman, L.E.; Troll, R.; Csankovszki, G. Dual roles for nuclear RNAi Argonautes in Caenorhabditis elegans dosage compensation. Genetics 2022, 221, iyac033. [Google Scholar] [CrossRef]
  182. Pal-Bhadra, M.; Leibovitch, B.A.; Gandhi, S.G.; Chikka, M.R.; Bhadra, U.; Birchler, J.A.; Elgin, S.C. Heterochromatic silencing and HP1 localization in Drosophila are dependent on the RNAi machinery. Science 2004, 303, 669–672. [Google Scholar] [CrossRef]
  183. Breimann, L.; Morao, A.K.; Kim, J.; Jimenez, D.S.; Maryn, N.; Bikkasani, K.; Carrozza, M.J.; Albritton, S.E.; Kramer, M.; Street, L.A.; et al. The histone H4 lysine 20 demethylase DPY-21 regulates thedynamics of condensin DC binding. J. Cell Sci. 2022, 135, jcs258818. [Google Scholar] [CrossRef]
  184. Kschonsak, M.; Merkel, F.; Bisht, S.; Metz, J.; Rybin, V.; Hassler, M.; Haering, C.H. Structural Basis for a Safety-Belt Mechanism That Anchors Condensin to Chromosomes. Cell 2017, 171, 588–600.e524. [Google Scholar] [CrossRef]
  185. Terakawa, T.; Bisht, S.; Eeftens, J.M.; Dekker, C.; Haering, C.H.; Greene, E.C. The condensin complex is a mechanochemical motor that translocates along DNA. Science 2017, 358, 672–676. [Google Scholar] [CrossRef] [PubMed]
  186. Kong, M.; Cutts, E.; Pan, D.; Beuron, F.; Kaliyappan, T.; Xue, C.; Morris, E.; Musacchio, A.; Vannini, A.; Greene, E.C. Human Condensin I and II Drive Extensive ATP-Dependent Compaction of Nucleosome-Bound DNA. Mol. Cell 2020, 79, 99–114. [Google Scholar] [CrossRef] [PubMed]
  187. Eeftens, J.M.; Bisht, S.; Kerssemakers, J.; Kschonsak, M.; Haering, C.H.; Dekker, C. Real-time detection of condensin-driven DNA compaction reveals a multistep binding mechanism. EMBO J. 2017, 36, 3448–3457. [Google Scholar] [CrossRef] [PubMed]
  188. Morao, A.K.; Kim, J.; Obaji, D.; Sun, S.; Ercan, S. Topoisomerases I and II facilitate condensin DC translocation to organize and repress X chromosomes in C. elegans. Mol. Cell 2022, 82, 4202–4217.e4205. [Google Scholar] [CrossRef] [PubMed]
  189. Uusküla-Reimand, L.; Hou, H.; Samavarchi-Tehrani, P.; Rudan, M.V.; Liang, M.; Medina-Rivera, A.; Mohammed, H.; Schmidt, D.; Schwalie, P.; Young, E.J.; et al. Topoisomerase II beta interacts with cohesin and CTCF at topological domain borders. Genome Biol. 2016, 17, 182. [Google Scholar] [CrossRef] [PubMed]
  190. Meyer, B.J.; Casson, L.P. Caenorhabditis elegans compensates for the difference in X chromosome dosage between the sexes by regulating transcript levels. Cell 1986, 47, 871–881. [Google Scholar] [CrossRef]
  191. Lau, A.C.; Zhu, K.P.; Brouhard, E.A.; Davis, M.B.; Csankovszki, G. An H4K16 histone acetyltransferase mediates decondensation of the X chromosome in C. elegans males. Epigenetics Chromatin 2016, 9, 44. [Google Scholar] [CrossRef]
Dna 04 00005 g001
Figure 1. SMC protein folding. An SMC protein is shown at the top with globular N and C terminal domains that are separated by a pair of alpha-helices on either side of a “hinge” domain. Once the hinge folds on itself, the two alpha-helices make up a coiled-coil, which allows for the N and C terminal domains to interact as the ATPase “head”. The coiled-coil is sometimes referred to as an “arm” with an “elbow”, where flexible bends can occur.
Figure 1. SMC protein folding. An SMC protein is shown at the top with globular N and C terminal domains that are separated by a pair of alpha-helices on either side of a “hinge” domain. Once the hinge folds on itself, the two alpha-helices make up a coiled-coil, which allows for the N and C terminal domains to interact as the ATPase “head”. The coiled-coil is sometimes referred to as an “arm” with an “elbow”, where flexible bends can occur.
Dna 04 00005 g001
Dna 04 00005 g002
Figure 2. Chromatin motor complexes in Caenorhabditis elegans. SMC proteins that interact with the N-terminus of the kleisin are in pink or purple, while SMC proteins that interact with the C-terminus of the kleisin are in orange or yellow. Kleisins are in shades of blue. HAWKs are in shades of green, and Kites are in shades of red. (A) The 4 subunits of cohesin in the closed ring conformation, shown here with the C. elegans subunit names. (B) Condensin I, condensin II, and condensin IDC are shown in the closed ring conformation, shown here with the C. elegans subunit names. (C) The SMC-5/6 complex is shown here with the characteristic “boomerang” shape and C. elegans subunit names.
Figure 2. Chromatin motor complexes in Caenorhabditis elegans. SMC proteins that interact with the N-terminus of the kleisin are in pink or purple, while SMC proteins that interact with the C-terminus of the kleisin are in orange or yellow. Kleisins are in shades of blue. HAWKs are in shades of green, and Kites are in shades of red. (A) The 4 subunits of cohesin in the closed ring conformation, shown here with the C. elegans subunit names. (B) Condensin I, condensin II, and condensin IDC are shown in the closed ring conformation, shown here with the C. elegans subunit names. (C) The SMC-5/6 complex is shown here with the characteristic “boomerang” shape and C. elegans subunit names.
Dna 04 00005 g002
Dna 04 00005 g003
Figure 3. Chromatin motor complex localization during C. elegans cell division. (A) In mitosis, cohesin is already present on the chromosomes before S-phase. After replication, cohesin is responsible for cohesion between the sister chromatids. Condensin II localizes to the chromosomes during prophase, while condensin I localizes to the chromosomes after nuclear envelope breakdown in prometaphase. In anaphase, cohesin is removed and a portion of condensin I moves to the spindle midzone. (B) In meiosis, cohesin is present along the chromosomes. After homologous recombination, condensin II localizes to the chromosomes at the end of prophase I. Condensin I localizes after nuclear envelope breakdown, specifically between the bivalents in metaphase I. In anaphase I, cohesin is removed from the bivalent short arms to allow for homolog separation and condensin I remains in the spindle midzone. Condensin II and cohesin remain on the long arm of the bivalent to keep sister chromatids attached.
Figure 3. Chromatin motor complex localization during C. elegans cell division. (A) In mitosis, cohesin is already present on the chromosomes before S-phase. After replication, cohesin is responsible for cohesion between the sister chromatids. Condensin II localizes to the chromosomes during prophase, while condensin I localizes to the chromosomes after nuclear envelope breakdown in prometaphase. In anaphase, cohesin is removed and a portion of condensin I moves to the spindle midzone. (B) In meiosis, cohesin is present along the chromosomes. After homologous recombination, condensin II localizes to the chromosomes at the end of prophase I. Condensin I localizes after nuclear envelope breakdown, specifically between the bivalents in metaphase I. In anaphase I, cohesin is removed from the bivalent short arms to allow for homolog separation and condensin I remains in the spindle midzone. Condensin II and cohesin remain on the long arm of the bivalent to keep sister chromatids attached.
Dna 04 00005 g003
Dna 04 00005 g004
Figure 4. The C. elegans Dosage Compensation Complex (DCC). Condensin IDC is shown with the other members of the DCC. Functions of the other members are highlighted in corresponding colors. The SDC proteins SDC-1, SDC-2, and SDC-3 are colored similarly to highlight their shared function in sex determination.
Figure 4. The C. elegans Dosage Compensation Complex (DCC). Condensin IDC is shown with the other members of the DCC. Functions of the other members are highlighted in corresponding colors. The SDC proteins SDC-1, SDC-2, and SDC-3 are colored similarly to highlight their shared function in sex determination.
Dna 04 00005 g004
Table 1. Cohesin mitotic and meiotic homologues, and cohesin regulator CTCF.
Table 1. Cohesin mitotic and meiotic homologues, and cohesin regulator CTCF.
OrganismC. elegansSaccharomyces cerevisiaeSaccharomyces pombeMammalsDrosophila melanogasterXenopus
SMC1SMC-1Smc1Psm1SMC1α, SMC1β *DmSMC1/Smc1XSMC1a/Smc1a
XSMC1b/Smc1b
SMC3SMC-3Smc3Psm3SMC3Cap/Smc3XSMC3/Smc3
KleisinSCC-1/COH-2
COH-1
REC-8 *
COH-3 *
COH-4 *		Scc1/Mcd1
Rec8 *		Rad21
Rec8 *		RAD21
RAD21L *
REC8 *		DRAD21/Rad21
c(2)M *		XRAD21/Rad21
XREC8/Rec8 *
HAWKSCC-3Scc3/Irr1Psc3
Rec11 *		SA1/STAG1
SA2/STAG2
SA3/STAG3 *		DSA1/Sa1
SA-2/Sa2 *		XSA1/Sa1
XSA2/Sa2
CTCF---CTCFDmCtcf/CtcfXCTCF/Ctcf
* indicates paralogues functional only during meiosis.
Table 2. Condensin I eukaryotic homologues.
Table 2. Condensin I eukaryotic homologues.
OrganismC. elegansS. cerevisiaeS. pombeMammalsD. melanogasterXenopus
SMC2SMC-2Smc2Cut14SMC2DmSmc2/Smc2XCAP-E/Smc2
SMC4SMC-4
DPY-27 *		Smc4Cut3SMC4DmSmc4/Smc4XCAP-C/Smc4
KleisinDPY-26Brn1Cnd2CAP-HCap-H/BarrenXCAP-H/Cap-H
HAWKsDPY-28
CAP-G1		Ysc4
Ysc1		Cnd1
Cnd3		CAP-D2
CAP-G		dCap-D2/Cap-D2
dcap-g/Cap-G		XCAP-D2/Cap-D2
XCAP-G/Cap-G
* indicates paralogues functional only in dosage compensation.
Table 3. Condensin II homologues in eukaryotes.
Table 3. Condensin II homologues in eukaryotes.
OrganismC. elegansMammalsD. melanogasterXenopus
SMC2SMC-2SMC2DmSMC2/Smc2XCAP-E/Smc2
SMC4SMC-4SMC4Glu/SMC4XCAP-C/Smc3
KleisinKLE-2CAP-H2dCap-H2/Cap-H2XCAP-H2/Cap-H2
HAWKsHCP-6
CAP-G2		CAP-D3
CAP-G2		dCAP-D3/Cap-D3
-1		XCAP-D3/Cap-D3
XCAP-G2/Cap-G2
1 A paralog for CAP-G2 has not been found in D. melanogaster.
Table 4. SMC5/6 complex eukaryotic homologues.
Table 4. SMC5/6 complex eukaryotic homologues.
OrganismC. elegansS. cerevisiaeS. pombeMammalsD. melanogasterXenopus
SMC5SMC-5Smc5Smc5/Spr18SMC5Smc5XSMC5/Smc5
SMC6SMC-6Smc6Smc6/Rad18SMC6Smc6XSMC6/Smc6
KleisinNSE-4Nse4Nse4/Rad62NSE4/NSMCE4Nse4Nse4
KitesNSE-1
NSE-2
NSE-3		Nse1
Nse2/Mms21
Nse3		Nse1
Nse2
Nse3		NSE1/NSMCE1
NSE2/NSMCE2
NSE3/NSMCE3		Nse1
Qjt/Nse2
MAGE/Nse3		Nse1
Nse2/Nsmce2/Mms21
Nse3
Other 1-
-		Nse5
Nse6/Kre29		Nse5
Nse6		SIMC1, SLF1
SLF2		-
-		Slf1
Slf2/Fam178a
1 Proteins in this category are not related to proteins in other SMC complexes, and homologues have not been found in all species.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Chawla, B.; Csankovszki, G. How Chromatin Motor Complexes Influence the Nuclear Architecture: A Review of Chromatin Organization, Cohesins, and Condensins with a Focus on C. elegans. DNA 2024, 4, 84-103. https://doi.org/10.3390/dna4010005

AMA Style

Chawla B, Csankovszki G. How Chromatin Motor Complexes Influence the Nuclear Architecture: A Review of Chromatin Organization, Cohesins, and Condensins with a Focus on C. elegans. DNA. 2024; 4(1):84-103. https://doi.org/10.3390/dna4010005

Chicago/Turabian Style

Chawla, Bahaar, and Györgyi Csankovszki. 2024. "How Chromatin Motor Complexes Influence the Nuclear Architecture: A Review of Chromatin Organization, Cohesins, and Condensins with a Focus on C. elegans" DNA 4, no. 1: 84-103. https://doi.org/10.3390/dna4010005

Article Metrics

Back to TopTop