A key maxim in modernist architecture is that ‘form follows function'. While modernist buildings are hopefully the product of intelligent design, the architectures of chromosomes have been sculpted by the forces of evolution over many thousands of generations. In the following, I will describe recent advances in our understanding of chromosome architecture in the archaeal domain of life. Although much remains to be learned about the mechanistic details of archaeal chromosome organization, some general principles have emerged. At the 10–100 kb level, archaeal chromosomes have a conserved local organization reminiscent of bacterial genomes. In contrast, lineage-specific innovations appear to have imposed distinct large-scale architectural features. The ultimate functions of genomes are to store and to express genetic information. Gene expression profiles have been shown to influence chromosome architecture, thus their form follows function. However, local changes to chromosome conformation can also influence gene expression and therefore, in these instances, function follows form.

The seminal studies of Woese and Fox revealed Archaea to be a distinct domain of life from Bacteria and Eukaryotes [1]. More recent advances in phylogenetic methods, coupled with massively increased taxonomic sampling have revealed deep divisions between archaeal phyla and have provided support for the emergence of eukaryotes from within the archaeal domain of life [2]. Metagenomic studies have provided a wealth of primary sequence information on archaeal species yet comparatively few organisms have been successfully grown as pure cultures, and, of these cultivated species, only a handful have proven amenable to genetic manipulation. Much of what we currently know about archaeal molecular genetics and chromosome biology is therefore derived from a limited number of species, most notably, members of the Sulfolobales order of the crenarchaea and the Halobacteriales and Thermococcales orders of the euryarchaea.

Thus far, all archaeal genomes are encoded on circular chromosomes and, like bacterial genomes, encode polycistronic units. As in bacteria, it is believed that transcription and translation are coupled in archaea [3]. Echoing the deep phyletic divide between cren- and euryarchaea, there are some fundamental differences in cell cycle logic and DNA replication mode between Sulfolobus and the euryarchaeal Haloferax and Thermococcus (Table 1). Sulfolobus species, and other crenarchaea, have a cell cycle in which a new-born cell has a single copy of its chromosome [4]. After a brief G1 period DNA replication initiates. Sulfolobus chromosomes possesses three replication origins, of which two (oriC1 and oriC3) fire at the same time thereby defining the start of S-phase and the third origin fires a few minutes into S-phase [5]. Ultimately, all cells fire all three replication origins. Following the completion of S-phase, approximately half the cell cycle is spent with two copies of the chromosome and fluorescence in situ hybridization (FISH) of a limited number of loci around the genome provided evidence for the persistence of sister chromatid cohesion for the majority of the G2 period [6]. This cohesion correlated with the existence of hemi-catenane junctions between the daughter molecules. At the end of G2, genomes compact concomitant with their segregation, in a process that likely involves Sulfolobus homologs of a bacterial ParA protein, termed SegA and its DNA binding partner, SegB [7]. Sulfolobus cells then divide utilizing the ESCRT machinery [8,9].

In contrast with the tightly organized 1C–2C ploidy control and regimented cell cycle of Sulfolobus, the euryarchaea are typically polyploid [10]. For example, Haloferax volcanii possesses ∼20 copies of its main chromosome. Studies in synchronized halophilic archaea that followed BrdU incorporation to monitor DNA replication or utilized GFP fused to the single-stranded DNA binding protein, provided evidence to suggest that DNA replication occurs throughout the cell cycle with no defined pre- and post-replicative phases. Indeed, replication was even observed in cells undergoing division. Quantitation of the SSB-GFP foci indicated that only 1–3 chromosomes replicate at a given time (for a recent review see [11]). Thus, for the halophiles, at least in regard to replication, there is a non-equivalence between chromosomes. Whether all chromosomes in a given cell have the same transcription profile and/or have the same conformation are currently unresolved issues. Another intriguing difference between the model euryarchaea and the crenarchaeote Sulfolobus lies in DNA replication mode. Sulfolobus has three DNA replication origins and the activity of at least one is required for viability [12]. In contrast, while Haloferax volcanii typically uses DNA replication origins during normal growth, it is possible to delete all active origins and retain viability [13]. Remarkably, origin-lacking cells even double more rapidly than their wild-type counterparts under laboratory growth conditions. Replication in these cells is likely mediated via homologous recombination as the cells are highly sensitive to levels of the RadA protein, the ortholog of bacterial RecA and eukaryotic Rad51 [13]. In Thermococcus kodakarensis, studies in exponentially growing cells failed to provide any evidence for replication origin firing or defined replication initiation sites [14]. However, genetic studies implicated origin function in surviving prolonged starvation. It was suggested that during nutrient deprivation, ongoing cell division could eventually reduce the cells to single chromosome content. In a 1C state, with no homologs present, replication origin function, rather than homologous recombination, would presumably be required to facilitate the restart of chromosome replication.

Our understanding of the conformation of bacterial chromosomes has been established by a combination of microscopy approaches, using FISH, fluorescent repressor/operator systems (FROS), a FROS-related technology utilizing an orthogonal fluorescent-ParB/parS system and through the application of proximity ligation chromosome conformation capture (3C)-based methodologies [15]. While FISH has been employed on a limited number of loci in Sulfolobus, it has not been extensively used in archaea. Nor have FROS-like technologies been employed in any archaeal species to date. Archaeal chromosome conformation studies have therefore focused on the application of 3C technologies to archaeal cultures [16–19]. The results of these studies of asynchronous populations report on the dominant and persistent architectural features. A further caveat must be borne in mind with the polyploid euryarchaea, as it is possible that the various copies of the chromosome have distinct and non-equivalent architectures [11]. In Sulfolobus, the existence of sister chromatid cohesion during the majority of the G2 phase of the cell cycle gives some confidence that 3C methods do indeed capture the conformation of both chromosomes in a given cell [6]. A clear goal for future work will be to apply 3C methods to single archaeal cells. It will also be of considerable interest to perform 3C on synchronized cultures to determine the modulation of chromosome architecture during the course of the cell cycle.

High-resolution 3C studies on both euryarchaeal and crenarchaeal species have revealed the presence of self-interacting domains throughout the genomes [17,18]. The size of these domains range from 25–570 kb in Haloferax volcanii and 14–196 kb in Sulfolobus species. In both species there is a statistically significant correlation between the location of domain boundaries and highly expressed genes. This combination of scale and boundary definition suggests these structures are analogous to ‘chromosomal interaction domains' (CIDs) first characterized in the bacterium Caulobacter crescentus and which are now known to be a universal feature of bacterial chromosome organization [20,21]. Studies in Sulfolobus confirmed that, as in bacteria, gene expression plays an active role is establishing boundaries [18]. Global inhibition of transcription using the DNA intercalator Actinomycin D reduced boundary definition and genetic approaches to either delete or insert highly transcribed genes resulted in loss or gain of CID boundaries, respectively. It has been proposed that highly expressed genes may act as CID boundaries by serving as barriers to the diffusion of plectonemic supercoiling in bacteria. It seems highly plausible that a similar mechanism may be at play in archaeal chromosomes. It is worthwhile emphasizing that this proposed dependence of CIDs on a fundamental topological feature of DNA readily accounts for the similarities in CIDs across a broad range of organisms both within and across domains of life even though their transcription machineries and chromatin compaction systems are highly distinct.

In addition to CIDs, the high-resolution studies have provided support for the existence of defined loop structures between disparate loci in archaeal chromosomes. In both Sulfolobus and Haloferax ∼70 loops of varying length were detected throughout the genome [17,18]. Notably, loops could encompass multiple CIDs, suggesting CIDs and loops are distinct organizational features of the chromosome. Analysis of the identity of the genes in the vicinity of loop anchors in Sulfolobus revealed a significant enrichment of genes for protein and RNA components of the ribosome [18]. While the 3C method employed reports on binary interactions, the observation that, within the population, a single anchor point could interact with a number of discrete loci suggests that a hub-like organization could exist, allowing the spatial colocalization of genes involved in ribosome biogenesis. In both Haloferax and Sulfolobus, loops appear to be dependent upon transcription, as Actinomycin D treatment led to a loss of loops in both species [17,18]. Additionally, entry into stationary phase reduced loops, and, in Sulfolobus, the ‘ribosomal gene’ loops were particularly impacted. The mechanistic basis of loop formation is currently not fully resolved. One explanation could be long-range bridging events between transcription factors bound at the anchor loci. Another possibility, at least for the Sulfolobus ribosomal loops, is that coordinated translation and ribosomal assembly could provide an attractive force that creates a nexus for ribosome biogenesis. Finally, as will be discussed below, in Haloferax, deletion of a gene for a SMC-superfamily protein had an impact on loop formation, albeit less of an effect than observed upon Actinomycin D treatment or entry into stationary phase [17].

While loops and CIDs are shared features of the crenarchaea and euryarchaeal studied thus far, work in Sulfolobus has revealed an additional architectural level, compartmentalization that, thus far, has not been observed in euryarchaeal species [16]. In addition to the ∼100 kb CIDs, Sulfolobus chromosomes possess an additional level of larger scale self-interacting segments. These segments form discrete higher-order interactions, generating a checker board-like plaid pattern in the Hi-C contact maps. The strong bias of these interactions defines two distinct compartments. Strikingly, this compartmentalized architecture has also been observed in mammalian cells, where the compartments are referred to as A and B type [22]. The same A/B nomenclature has been adopted for Sulfolobus. As with the mammalian A compartment, the Sulfolobus A compartment possesses the majority of the highly expressed genes in the genome. The B compartment in Sulfolobus is more lowly expressed, contains non-essential genes and also is enriched for transposable elements [16]. Thus, at the functional level, the Sulfolobus A and B compartments are reminiscent of the phenomena of euchromatin and heterochromatin, respectively, in eukaryotes. It is important to stress that while there is a conceptual and functional analogy, the mechanistic bases of definition of the A/B compartments in Sulfolobus and of eukaryotic euchromatin and heterochromatin are entirely distinct.

Notably, the three replication origins in the Sulfolobus chromosome are located in the A compartment. However, inactivation of the origins by deletion of the cognate initiator protein gene did not affect compartmentalization. Thus, ongoing replication is not required for the maintenance of A compartment identity [16]. However, as discussed below, it seems likely that over multi-generational, evolutionary timescales the replication program will have contributed to sculpting the conformation of the chromosome. Compartment identity, like CIDs and loops, is dependent on the transcriptional profile of cells. Inhibition of transcription with Actinomycin D abrogated compartments. Entry into stationary phase, during which a relative increase in transcription occurs within the B compartment, also triggers a loss of compartmentalization. Interestingly, disruption of the genes for the extremely abundant cell surface S-layer subunits also resulted in a loss of compartmentalization [18]. This is an extremely pleiotropic mutation with cells exhibiting altered morphology, size and ploidy [23]. However, this observation could perhaps hint at a contribution of cell shape to underlying chromosome organization. It is important to stress that compartmentalization is distinct from CID formation. CIDs are found in both A and B compartment and even under conditions where compartmentalization is lost, CIDs persist. In contrast with the independence of compartments and CIDs, the overall number of loop structures was reduced in conditions where compartments were weakened, suggesting a causal interplay between these two phenomena [18]. Given that 80% of loops are intra-compartmental, one possibility is that loops could act to reinforce the compartmentalized architecture.

SMC proteins are member of an extended family of ATPases, possessing a bipartite ATPase head domain, a coiled-coil domain and, at the opposite end of the coiled-coil from the ATPase domain, a hinge domain that serves as a dimerization module. Depending on the species and protein complex in question, SMC proteins form homo- or heterodimers and further associate with additional partner proteins. For excellent overviews of the phylogeny of the SMC superfamily and the closely related RAD50 family of proteins, the reader is directed to recent articles by Yoshinaga and Inagaki [24,25]. In eukaryotes, the SMC-containing complexes, cohesin and condensin are well documented to play key roles in establishing and maintaining chromosome tertiary structure. While cohesin is a eukaryotic innovation, condensins are near universal, being found in eukaryotes, bacteria and many, but not all, archaea [26]. In bacteria, condensin facilitates juxtaposition of chromosome arms and data support a loop-extrusion model to generate the juxtaposition [27]. While helping to establish long-range architecture in bacteria, there is currently no evidence to indicate a role for bacterial condensins in establishing or maintaining CIDs [28]. In eukaryotes, cohesin functions to establish CID-like domains, the Topologically Associating Domains (TADs). Cohesin is enriched at TAD boundaries and thus a TAD could be viewed as a loop extruded by cohesin [29–31]. In mammals, the oriented DNA binding of the protein CTCF provides the directional cue to limit cohesion movement and thus CTCF and cohesin enrichment are hallmarks of mammalian TAD boundaries [32–34].

While high resolution protein structures have been determined for archaeal condensin components, both for the SMC subunit and the partner proteins ScpA and ScpB [35–37], very little is known about the physiological roles of the archaeal SMC family proteins. Genetic studies of a condensin SMC homolog in the euryarchaeon Methanococcus voltae revealed that disruption of the gene resulted in aberrant chromosome segregation and cell shape, engendering a ‘titan cell' phenotype [38]. Another SMC-superfamily member, Sph1, of Halobacterium salinarum was implicated in events late in chromosome replication and segregation [39]. More recently, the high-resolution 3C studies in Haloferax volcanii tested the contributions of SMC-superfamily proteins to chromosome conformation [17]. The gene for the condensin SMC subunit was non-essential for viability with similar growth rate, morphology and DNA content to wild-type cells. The cells however, displayed a loss of 9 domain boundaries across the genome. Importantly, this was not due to transcriptional changes in the boundary-proximal genes, suggesting a direct architectural role for the protein in boundary definition. Whether this is due to physical localization of the condensin complex at these loci is currently unknown. A modest but significant reduction in loops was also observed in the mutant cells. Deletion mutants of genes for two further SMC-superfamily proteins, RAD50 and Sph4, did not show any significant reduction in boundaries or loops but, like the strain lacking the condensin SMC, showed some reduction in short-range contacts [17].

Condensin is found throughout the archaeal lineages, with the exception of the crenarchaeal phylum, of which Sulfolobus is a member. Examination of the genome sequence of Sulfolobus identified open-reading frame encoding a signature SMC-related protein that we have termed coalescin, or ClsN [16,40]. ClsN is a small SMC-family protein of 68 kDa with ∼350 amino acids of coiled-coil. Interestingly, the dimerization domain of ClsN appears to be a Zinc-hook motif, a feature previously identified in the RAD50 family of proteins [41].

ChIP-Seq revealed ClsN to be enriched in the B compartment of Sulfolobus and over-expression of the clsN gene led to a strengthening of compartmentalization with a concomitant further reduction in the already low gene expression in the B compartment. Furthermore, ClsN occupancy, as adjudged by ChIP-Seq, displayed a general anti-correlation with transcription levels of loci with which it associated. Importantly, induction of transcription correlated with loss of ClsN at the induced loci [16]. Thus, there appears to be a mutually antagonistic interplay between gene transcription and coalescin binding. In metazoan, the self-interacting TADs are demarcated by the enrichment of cohesin and the DNA binding protein CTCF at boundaries between TADs. The available data strongly support cohesin acting to extrude loops of DNA with CTCF acting as a polar barrier to cohesin movement [30,31,33,34]. In contrast, in Sulfolobus there is no detectable enrichment of coalescin at boundaries between the self-interacting CID domains [18]. Rather, coalescin shows elevated enrichment within the body of B compartment CIDs. Thus, within the B compartment, CIDs seem to be defined by locally high gene expression levels at the boundaries and ClsN enrichment within the CID body. The mechanism of ClsN action is unresolved at this time but the current observations of its localization are compatible with either local dynamic DNA loop extrusion by the protein or a more static binding and second-strand capture models. Many additional aspects of ClsN function remain to be determined: whether dedicated ClsN loaders and unloaders exist, whether ClsN interacts with additional partner proteins and how ClsN localization is modulated during the cell cycle are all key topics requiring resolution.

The compartmentalization of Sulfolobus chromosomes with its partitioning into more and less transcriptionally active domains prompts comparisons with eukaryotic euchromatin and heterochromatin. However, the basis of the evolution of this modular nature may have mechanistic parallels with bacterial chromosomes. It is well established that many bacterial species show gradients of gene expression from their single DNA replication origins to the termination sites. This is interpreted as a mechanism arising over evolutionary timescales that allows selection for elevated expression of important growth-related genes, in essence by ensuring coupling of gene dosage with early chromosome replication [42]. A similar principle could be at play in Sulfolobus with the additional complexity that with three replication origins, three distinct regions can now provide safe harbors for the genes undergoing selection. As alluded to above, looping between regulatory elements that govern coordinate expression of these origin-proximal genes and/or attractive forces mediated by coupled transcription/translation of multi-protein assemblies, such as ribosomes, could provide a mechanism for consolidating the 3D juxtaposition of origin proximal loci. In the case of Sulfolobus, the general antagonism between ClsN and transcription results in enrichment of ClsN in transcriptionally quiescent regions. Through loop extrusion and/or second strand capture, ClsN can lead to the coalescence of these less highly expressed genes. Thus, the combination of selection over evolutionary time scales for origin-proximal localization for highly expressed genes, their regulatory looping and the action of ClsN to generate coalesced loci of low transcriptional activity could drive the formation of the compartmentalized structure seen in present-day Sulfolobus genomes (Figure 1). It is interesting to note that the definition of compartmentalization [as adjudged by principal component (PC) analyses] is weakest in the vicinity of oriC2 [19]. It is notable, therefore, that oriC2 is a Sulfolobus-specific replication origin and, presumably, the most recently acquired in this lineage. The heterogeneity in the PC signals in the vicinity of oriC2 may reflect the ongoing evolutionary streamlining of gene order in the vicinity of this locus.

Finally, the advent of compartmentalization has profound implications for the ongoing evolution of Sulfolobus chromosomes. It has been established that Sulfolobus chromosomes display uneven rates of single nucleotide polymorphisms (SNPs) [43]. Remarkably, regions of low and high SNP accumulation correspond to the A and B compartments, respectively [19]. Similarly, gene synteny is also more conserved within the A compartment than in the B compartment. Interestingly, proximity to replication termination sites, rather than distance from replication origins, may have a causal role in SNP accumulation in the B compartment [19]. In principle, the different levels of transcription of A and B compartments could contribute to this bias via distinct levels of transcription-coupled repair (TCR) in the two compartments. However, current evidence suggests there is no TCR pathway in Sulfolobus [44]. Thus, the mechanistic basis of this differential accumulation of SNPs remains undetermined at this time. It is notable that data derived from Assay for Transposase-Accessible Chromatin (ATAC)-Seq demonstrated that the B compartment is less accessible to transposition events than the A compartment [19]. A key implication of this result is that the B compartment may have a denser, more highly compacted organization than the A compartment. The denser, more restrictive nature of the B compartment may result in slower kinetics of DNA-repair associated events, perhaps explaining the higher mutation rate of this compartment.

Our knowledge of archaeal chromosome architecture and its modulation and regulation is still at a very rudimentary level. Much remains to be determined about the identity and molecular roles of the key players that effect chromosome organization and how these players are regulated throughout the cell-cycle and during culture growth. It is clear that transcription plays a vital role in sculpting the domain organization of archaeal genomes, an illustration of function defining form. Yet, beyond speculation about insulating plectoneme diffusion, how transcriptional activity leads to CID boundaries is unresolved. Indeed, we know essentially nothing about the topological landscape of archaeal chromosomes. While extensive biochemical and structural studies have been performed on archaeal topoisomerases [45], how supercoiling is distributed on archaeal chromosomes and how this distribution could impact on higher-order organization remains unknown. We also know very little at this point about the fine-level chromatin landscape of archaeal cells. A variety of chromatin or nucleoid-associated proteins have been characterized in archaeal species [46]. These include orthologs of histones in essentially all archaeal phyla apart from the crenarchaea. How these various protein interact with, and are distributed on, archaeal genomes and how they contribute to higher order organization remain exciting topics for future work. Understanding the basis of locus-locus loop formation is another key goal. The observation that the evolutionary trajectory of a given locus is influenced by its location in B or A compartment reveals how function can follow form. It is exciting to note that this context-dependent bias in mutation rate is not restricted to archaea, similar epigenome-driven phenomena can be observed in Arabidopsis and in human cancer cells [47,48].

  • The position of archaea in the tree of life gives unique insights into the mechanisms and evolution of chromosome organization

  • Studies of archaeal chromosome conformation reveal an intriguing blend of features previously associated with bacterial and eukaryotic chromosome organization

  • Ongoing studies will reveal the mechanistic basis of chromosome organization via the interplay of DNA topology, gene transcription, chromatin proteins and SMC complexes.

The author declares that there are no competing interests associated with this manuscript.

Work in my lab is funded by the College of Arts and Sciences, Indiana University and by National Institutes of Health grants R01GM125579 and R01GM135178.

S.D.B. wrote this article.

I thank Rachel Samson for her insightful comments on this manuscript and members of my lab for their work on this project.

CIDs

chromosomal interaction domains

FISH

fluorescence in situ hybridization

FROS

fluorescent repressor/operator systems

PC

principal component

SNPs

single nucleotide polymorphisms

TADs

topologically associating domains

TCR

transcription-coupled repair

1
Woese
,
C.R.
and
Fox
,
G.E.
(
1977
)
Phylogenetic structure of the prokaryotic domain: the primary kingdoms
.
Proc. Natl Acad. Sci. U.S.A.
74
,
5088
5090
2
Spang
,
A.
,
Caceres
,
E.F.
and
Ettema
,
T.J.G.
(
2017
)
Genomic exploration of the diversity, ecology, and evolution of the archaeal domain of life
.
Science
357
,
eaaf3883
3
French
,
S.L.
,
Santangelo
,
T.J.
,
Beyer
,
A.L.
and
Reeve
,
J.N.
(
2007
)
Transcription and translation are coupled in Archaea
.
Mol. Biol. Evol.
24
,
893
895
4
Bernander
,
R.
and
Poplawski
,
A.
(
1997
)
Cell cycle characteristics of thermophilic archaea
.
J. Bacteriol.
179
,
4963
4969
5
Duggin
,
I.G.
,
McCallum
,
S.A.
and
Bell
,
S.D.
(
2008
)
Chromosome replication dynamics in the archaeon Sulfolobus acidocaldarius
.
Proc. Natl Acad. Sci. U.S.A.
105
,
16737
16742
6
Robinson
,
N.P.
,
Blood
,
K.A.
,
McCallum
,
S.A.
,
Edwards
,
P.A.
and
Bell
,
S.D.
(
2007
)
Sister chromatid junctions in the hyperthermophilic archaeon Sulfolobus solfataricus
.
EMBO J.
26
,
816
824
7
Kalliomaa-Sanford
,
A.K.
,
Rodriguez-Castaneda
,
F.A.
,
McLeod
,
B.N.
,
Latorre-Rosello
,
V.
,
Smith
,
J.H.
,
Reimann
,
J.
et al (
2012
)
Chromosome segregation in Archaea mediated by a hybrid DNA partition machine
.
Proc. Natl Acad. Sci. U.S.A.
109
,
3754
3759
8
Samson
,
R.Y.
,
Obita
,
T.
,
Freund
,
S.M.
,
Williams
,
R.L.
and
Bell
,
S.D.
(
2008
)
A role for the ESCRT system in cell division in archaea
.
Science
322
,
1710
1713
9
Lindas
,
A.C.
,
Karlsson
,
E.A.
,
Lindgren
,
M.T.
,
Ettema
,
T.J.
and
Bernander
,
R.
(
2008
)
A unique cell division machinery in the Archaea
.
Proc. Natl Acad. Sci. U.S.A.
105
,
18942
6
10
Soppa
,
J.
(
2011
)
Ploidy and gene conversion in Archaea
.
Biochem. Soc. Trans.
39
,
150
154
11
Soppa
,
J.
(
2022
)
Non-equivalent genomes in polyploid prokaryotes
.
Nat. Microbiol.
7
,
186
188
12
Samson
,
R.Y.
,
Xu
,
Y.
,
Gadelha
,
C.
,
Stone
,
T.A.
,
Faqiri
,
J.N.
,
Li
,
D.
et al (
2013
)
Specificity and function of archaeal DNA replication initiator proteins
.
Cell Rep.
3
,
485
496
13
Hawkins
,
M.
,
Malla
,
S.
,
Blythe
,
M.J.
,
Nieduszynski
,
C.A.
and
Allers
,
T.
(
2013
)
Accelerated growth in the absence of DNA replication origins
.
Nature
503
,
544
547
14
Gehring
,
A.M.
,
Astling
,
D.P.
,
Matsumi
,
R.
,
Burkhart
,
B.W.
,
Kelman
,
Z.
,
Reeve
,
J.N.
et al (
2017
)
Genome replication in Thermococcus kodakarensis independent of Cdc6 and an origin of replication
.
Front. Microbiol.
8
,
2084
15
Badrinarayanan
,
A.
,
Le
,
T.B.
and
Laub
,
M.T.
(
2015
)
Bacterial chromosome organization and segregation
.
Annu. Rev. Cell Dev. Biol.
31
,
171
199
16
Takemata
,
N.
,
Samson
,
R.Y.
and
Bell
,
S.D.
(
2019
)
Physical and functional compartmentalization of archaeal chromosomes
.
Cell
179
,
165
79.e18
17
Cockram
,
C.
,
Thierry
,
A.
,
Gorlas
,
A.
,
Lestini
,
R.
and
Koszul
,
R.
(
2021
)
Euryarchaeal genomes are folded into SMC-dependent loops and domains, but lack transcription-mediated compartmentalization
.
Mol. Cell
81
,
459
72.e10
18
Takemata
,
N.
and
Bell
,
S.D.
(
2021
)
Multi-scale architecture of archaeal chromosomes
.
Mol. Cell
81
,
473
87.e6
19
Badel
,
C.
,
Samson
,
R.Y.
and
Bell
,
S.D.
(
2022
)
Chromosome organization affects genome evolution in Sulfolobus archaea
.
Nat. Microbiol.
7
,
820
830
20
Le
,
T.B.
and
Laub
,
M.T.
(
2014
)
New approaches to understanding the spatial organization of bacterial genomes
.
Curr. Opin. Microbiol.
22
,
15
21
21
Le
,
T.B.
and
Laub
,
M.T.
(
2016
)
Transcription rate and transcript length drive formation of chromosomal interaction domain boundaries
.
EMBO J.
35
,
1582
1595
22
Lieberman-Aiden
,
E.
,
van Berkum
,
N.L.
,
Williams
,
L.
,
Imakaev
,
M.
,
Ragoczy
,
T.
,
Telling
,
A.
et al (
2009
)
Comprehensive mapping of long-range interactions reveals folding principles of the human genome
.
Science
326
,
289
293
23
Zhang
,
C.
,
Wipfler
,
R.L.
,
Li
,
Y.
,
Wang
,
Z.
,
Hallett
,
E.N.
and
Whitaker
,
R.J.
(
2019
)
Cell structure changes in the hyperthermophilic crenarchaeon Sulfolobus islandicus lacking the S-layer
.
mBio
10
,
e01589-19
24
Yoshinaga
,
M.
and
Inagaki
,
Y.
(
2021
)
Ubiquity and origins of structural maintenance of chromosomes (SMC) proteins in eukaryotes
.
Genome Biol. Evol.
13
,
evab256
25
Yoshinaga
,
M.
,
Nakayama
,
T.
and
Inagaki
,
Y.
(
2022
)
A novel structural maintenance of chromosomes (SMC)-related protein family specific to archaea
.
Front. Microbiol.
13
,
913088
26
Hirano
,
T.
(
2016
)
Condensin-based chromosome organization from bacteria to vertebrates
.
Cell
164
,
847
857
27
Wang
,
X.
,
Brandao
,
H.B.
,
Le
,
T.B.
,
Laub
,
M.T.
and
Rudner
,
D.Z.
(
2017
)
Bacillus subtilis SMC complexes juxtapose chromosome arms as they travel from origin to terminus
.
Science
355
,
524
527
28
Le
,
T.B.
,
Imakaev
,
M.V.
,
Mirny
,
L.A.
and
Laub
,
M.T.
(
2013
)
High-resolution mapping of the spatial organization of a bacterial chromosome
.
Science
342
,
731
734
29
Nasmyth
,
K.
(
2001
)
Disseminating the genome: joining, resolving, and separating sister chromatids during mitosis and meiosis
.
Annu. Rev. Genet.
35
,
673
745
30
Sanborn
,
A.L.
,
Rao
,
S.S.
,
Huang
,
S.C.
,
Durand
,
N.C.
,
Huntley
,
M.H.
,
Jewett
,
A.I.
et al (
2015
)
Chromatin extrusion explains key features of loop and domain formation in wild-type and engineered genomes
.
Proc. Natl Acad. Sci. U.S.A.
112
,
E6456
E6465
31
Fudenberg
,
G.
,
Abdennur
,
N.
,
Imakaev
,
M.
,
Goloborodko
,
A.
and
Mirny
,
L.A.
(
2017
)
Emerging evidence of chromosome folding by loop extrusion
.
Cold Spring Harb. Symp. Quant. Biol.
82
,
45
55
32
Nora
,
E.P.
,
Goloborodko
,
A.
,
Valton
,
A.L.
,
Gibcus
,
J.H.
,
Uebersohn
,
A.
,
Abdennur
,
N.
et al (
2017
)
Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization
.
Cell
169
,
930
44.e22
33
Rao
,
S.S.P.
,
Huang
,
S.C.
,
Glenn St Hilaire
,
B.
,
Engreitz
,
J.M.
,
Perez
,
E.M.
,
Kieffer-Kwon
,
K.R.
et al (
2017
)
Cohesin loss eliminates all loop domains
.
Cell
171
,
305
20.e24
34
Wutz
,
G.
,
Varnai
,
C.
,
Nagasaka
,
K.
,
Cisneros
,
D.A.
,
Stocsits
,
R.R.
,
Tang
,
W.
et al (
2017
)
Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins
.
EMBO J.
36
,
3573
3599
35
Burmann
,
F.
,
Shin
,
H.C.
,
Basquin
,
J.
,
Soh
,
Y.M.
,
Gimenez-Oya
,
V.
,
Kim
,
Y.G.
et al (
2013
)
An asymmetric SMC-kleisin bridge in prokaryotic condensin
.
Nat. Struct. Mol. Biol.
20
,
371
379
36
Diebold-Durand
,
M.L.
,
Lee
,
H.
,
Ruiz Avila
,
L.B.
,
Noh
,
H.
,
Shin
,
H.C.
,
Im
,
H.
et al (
2017
)
Structure of full-length SMC and rearrangements required for chromosome organization
.
Mol. Cell
67
,
334
47.e5
37
Jeon
,
J.H.
,
Lee
,
H.S.
,
Shin
,
H.C.
,
Kwak
,
M.J.
,
Kim
,
Y.G.
,
Gruber
,
S.
et al (
2020
)
Evidence for binary Smc complexes lacking kite subunits in archaea
.
IUCrJ
7
,
193
206
38
Long
,
S.W.
and
Faguy
,
D.M.
(
2004
)
Anucleate and titan cell phenotypes caused by insertional inactivation of the structural maintenance of chromosomes (smc) gene in the archaeon Methanococcus voltae
.
Mol. Microbiol.
52
,
1567
1577
39
Herrmann
,
U.
and
Soppa
,
J.
(
2002
)
Cell cycle-dependent expression of an essential SMC-like protein and dynamic chromosome localization in the archaeon Halobacterium salinarum
.
Mol. Microbiol.
46
,
395
409
40
Takemata
,
N.
and
Bell
,
S.D.
(
2020
)
Emerging views of genome organization in Archaea
.
J. Cell Sci.
133
,
jcs243782
41
Hopfner
,
K.P.
,
Craig
,
L.
,
Moncalian
,
G.
,
Zinkel
,
R.A.
,
Usui
,
T.
,
Owen
,
B.A.
et al (
2002
)
The Rad50 zinc-hook is a structure joining Mre11 complexes in DNA recombination and repair
.
Nature
418
,
562
566
42
Rocha
,
E.P.C.
(
2004
)
The replication-related organization of bacterial genomes
.
Microbiology (Reading)
150
,
1609
1627
43
Krause
,
D.J.
,
Didelot
,
X.
,
Cadillo-Quiroz
,
H.
and
Whitaker
,
R.J.
(
2014
)
Recombination shapes genome architecture in an organism from the archaeal domain
.
Genome Biol. Evol.
6
,
170
178
44
Dorazi
,
R.
,
Gotz
,
D.
,
Munro
,
S.
,
Bernander
,
R.
and
White
,
M.F.
(
2007
)
Equal rates of repair of DNA photoproducts in transcribed and non-transcribed strands in sulfolobus solfataricus
.
Mol. Microbiol.
63
,
521
529
45
Garnier
,
F.
,
Couturier
,
M.
,
Debat
,
H.
and
Nadal
,
M.
(
2021
)
Archaea: a gold mine for topoisomerase diversity
.
Front. Microbiol.
12
,
661411
46
Laursen
,
S.P.
,
Bowerman
,
S.
and
Luger
,
K.
(
2021
)
Archaea: the final frontier of chromatin
.
J. Mol. Biol.
433
,
166791
47
Schuster-Bockler
,
B.
and
Lehner
,
B.
(
2012
)
Chromatin organization is a major influence on regional mutation rates in human cancer cells
.
Nature
488
,
504
507
48
Monroe
,
J.G.
,
Srikant
,
T.
,
Carbonell-Bejerano
,
P.
,
Becker
,
C.
,
Lensink
,
M.
,
Exposito-Alonso
,
M.
et al (
2022
)
Mutation bias reflects natural selection in Arabidopsis thaliana
.
Nature
602
,
101
105
49
Zerulla
,
K.
,
Chimileski
,
S.
,
Nather
,
D.
,
Gophna
,
U.
,
Papke
,
R.T.
and
Soppa
,
J.
(
2014
)
DNA as a phosphate storage polymer and the alternative advantages of polyploidy for growth or survival
.
PLoS ONE
9
,
e94819
50
Ausiannikava
,
D.
,
Mitchell
,
L.
,
Marriott
,
H.
,
Smith
,
V.
,
Hawkins
,
M.
,
Makarova
,
K.S.
et al (
2018
)
Evolution of genome architecture in archaea: spontaneous generation of a new chromosome in Haloferax volcanii
.
Mol. Biol. Evol.
35
,
1855
1868
This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution License 4.0 (CC BY).