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. 2024 Feb 25;123(2):101–108. doi: 10.1111/mmi.15245

Capturing chromosome conformation in Crenarchaea

Elyza Pilatowski‐Herzing 1,2, Rachel Y Samson 1,2, Naomichi Takemata 1,2,3, Catherine Badel 1,2,4, Peter B Bohall 1,2, Stephen D Bell 1,2,✉
PMCID: PMC11344861  PMID: 38404013

Abstract

While there is a considerable body of knowledge regarding the molecular and structural biology and biochemistry of archaeal information processing machineries, far less is known about the nature of the substrate for these machineries—the archaeal nucleoid. In this article, we will describe recent advances in our understanding of the three‐dimensional organization of the chromosomes of model organisms in the crenarchaeal phylum.

Keywords: Archaea, condensin, evolution, genome, SMC, Sulfolobus

To fit into cells, chromosomes must be folded. Here we describe how members of the Archaea effect this folding process and how chromosome architecture both influences and is influenced by gene expression.

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The archaea are arguably best known for species that thrive in a range of extreme environments. However, organisms in the archaeal domain of life are ubiquitous and abundant components of the biosphere that play key roles in global biogeochemical cycles (Teske et al., 2021). Pioneering work by Carl Woese and George Fox in the 1970s first recognized Archaea as a domain of life, distinct from bacteria and eukaryotes (Woese & Fox, 1977). More recently, advances in metagenomic sampling resulting in broader taxonomic representation and improvements in phylogenetic reconstruction methodologies have illuminated the diversity of the archaeal domain with the identification of a number of key archaeal phyla and super‐phyla (Spang et al., 2017). Foremost among these are the Euryarchaea, Asgard, DPANN (named from the first letters of the five phyla characterized within this superphylum: Diapherotrites, Parvarchaeota, Aenigmarchaeota, Nanoarchaeota, and Nanohaloarchaeota), and TACK groupings. The TACK superphylum is comprised of the Thaumarchaea, Aigarchaea, Crenarchaea, and Korarchaea. More recently, analyses have supported a revised model of the tree of life, suggesting that the eukaryote domain emerged from within the Archaea, most likely from within the Asgard phylum (Figure 1) (Eme et al., 2023).

FIGURE 1.

FIGURE 1

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Schematic representation of the two‐domain of life tree with eukaryotes emerging from within the Asgard grouping. Major phyla or superphyla of Archaea are indicated in purple (acronyms are explained in the text) eukaryotes in blue and bacteria in gray.

Despite the abundance of archaeal organisms and their ecological and evolutionary significance, in comparison with the wealth of information available for bacteria and eukaryotes, comparatively little is known about the basics of archaeal cell biology, molecular genetics, and chromosome biology (Bell, 2022; Blombach et al., 2019; Greci & Bell, 2020; van Wolferen et al., 2022). A major impediment to studying the molecular biology of archaea is that only a few archaeal strains can be cultured and grown under laboratory conditions and, of these species, only a limited subset can be genetically manipulated. Thus, much of our understanding of archaeal molecular biology stems from detailed analyses of a few species, most notably the Sulfolobales of the crenarchaeal phylum and the Halobacteriales, Methanococcales, and Thermococcales orders from the euryarchaeal phylum (Leigh et al., 2011).

The phylogenetic divergence between crenarchaea and euryarchaea is manifested in key differences in the fundamental basis of cell cycle logic and information processing machineries; for reviews see, Samson and Bell (2011), Bell (2022), and Soppa (2022). Briefly, while the euryarchaea are typically highly polyploid and appear to have concurrent rounds of DNA replication, chromosome segregation, and cell division, crenarchaea such as Sulfolobus have more overtly structured cell cycles. These hyperthermophilic acidophiles have single circular chromosomes, ranging from 2.2 to 3 megabases in size. The Sulfolobus cell cycle begins with a brief G1 growth period and transitions into DNA replication (Bernander & Poplawski, 1997). The start of replication, or S‐phase, is defined by the simultaneous firing of two of the three replication origins (oriC1 and oriC3), with the third (oriC2) firing shortly into S‐phase (Duggin et al., 2008). All three replication origins fire once per cell cycle. Upon completing replication, cells then enter G2 phase, a growth period where cells spend approximately half their cell cycle. During this period, daughter chromosomes are cohesed and hemi‐catenane junctions are detectable between the sister chromatids (Robinson et al., 2007). Late in G2, chromosomes segregate in a process that is believed to be dependent on an ortholog of bacterial ParA proteins, termed SegA in Sulfolobus, that functions with a non‐canonical partner protein, termed SegB (Kalliomaa‐Sanford et al., 2012). Segregation is rapidly followed by cell division in a process that utilizes orthologs of the eukaryotic ESCRT‐III and Vps4 proteins (Lindas et al., 2008; Samson et al., 2008).

A combination of light microscopy approaches and chromosome conformation capture techniques has demonstrated that the chromosomes of many bacteria (with the notable exception of E. coli) organize their chromosomes by pairing the chromosome arms symmetrically from loci normally in the vicinity of the origin of DNA replication (Badrinarayanan et al., 2015; Lioy et al., 2021; Reyes‐Lamothe & Sherratt, 2019; Sherratt, 2017). The emerging rule is that canonical bacterial ParB proteins bind to origin‐proximal parS sites and, upon DNA binding, ParB loads the condensin holocomplex composed of a structural maintenance of chromosome (SMC) protein and accessory proteins. As replication proceeds, condensin acts to zip up the two chromosome arms, facilitating the coupling of chromosome segregation with DNA replication. Bacteria chromosomes also have a second, more local, level of chromosome organization in the form of self‐interacting chromatin interaction domains (CIDs) (Le et al., 2013). CIDs are typically on the scale of tens of kilobases, and their boundaries are defined by, and dependent upon, highly transcribed genes or operons. Metazoan chromosomes are also organized into self‐associating local domains, termed topological associating domains (TADs) (Lieberman‐Aiden et al., 2009). TADs are thought to be defined principally by the action of the loop‐extruding cohesin SMC complex, with directional barriers imposed to loop extrusion by polar interactions with the boundary‐defining, DNA‐bound CTCF factor; for a recent review, see da Costa‐Nunes and Noordermeer (2023). Thus, in both bacteria and eukaryotes, SMC proteins are key players in mediating chromosome organization (Hirano, 2016; Yatskevich et al., 2019). A typical SMC protein possesses a hinge domain that mediates the dimerization between two proteins to form a homodimeric or heterodimeric SMC complex. This hinge domain is flanked by alpha‐helical regions that fold onto one another to form an anti‐parallel coiled‐coil region. The extreme N‐ and C‐terminal regions of the SMC protein fold together to reconstitute an ATPase head domain. The ATPase head domains engage or disengage during the ATP binding, hydrolysis, and release catalytic cycle. Additional accessory proteins interact with head and head‐proximal regions of the coiled‐coil of the SMC to form the holocomplex.

1. CHROMOSOME ORGANIZATION IN THE SULFOLOBALES

While condensin is nearly universal, genes for its subunits are notably absent from the crenarchaea. It has, therefore, been of considerable interest to determine how crenarchaeal chromosomes are organized and how this organization is brought about. Thus far, the majority of studies have focused on species within the Sulfolobales (Figure 2) (Badel et al., 2022; Takemata et al., 2019; Takemata & Bell, 2021c). Chromosome conformation capture (3C) techniques have shown Sulfolobales chromosomes to be structured on multiple levels. As in bacteria and the few euryarchaea studied thus far (Cockram et al., 2021; Lioy et al., 2021), Sulfolobales chromosomes possess the tens of kilobase‐scale CIDs, and as in bacteria, locally high transcription plays a key role in establishing inter‐CID boundaries (Takemata & Bell, 2021c). Sulfolobales chromosomes also have numerous anchored loops throughout the chromosome with long‐range loops frequently anchored at loci encoding ribosomal proteins or rRNA. The Sulfolobales chromosomes also show an additional higher‐order level of organization in the form of A and B compartments, superficially reminiscent of the organization of metazoan chromosomes (Takemata & Bell, 2020, 2021c). As in metazoans, the Sulfolobales A compartment is more transcriptionally active and ATAC‐Seq reveals it to have a more accessible or open organization (Badel et al., 2022). The A compartment contains the majority of the essential genes and also houses the three origins of chromosome replication. While origin activity is not required for the ongoing maintenance of compartmentalization, we have previously speculated the location of the origins may have contributed to the evolution of the A compartment's gene repertoire (Badel et al., 2022; Bell, 2022; Takemata et al., 2019). In comparison, the B compartment is less transcriptionally active, less accessible and has an enrichment of non‐essential genes and transposons. As stated above, crenarchaeal phylum members have lost canonical condensin. However, Sulfolobales encode a novel SMC protein, termed coalescin (ClsN) (Takemata et al., 2019). In addition to being a signature protein for the Sulfolobales, ClsN is a member of the “Archaea‐specific SMC‐related proteins”—ASRPs (Yoshinaga et al., 2022). The ASRPs show a broad but patchy distribution across archaeal phyla, a pattern suggestive of derivation from an extra‐chromosomal element (see below). Although clsN is presumably, in evolutionary terms, a relatively recent acquisition in the Sulfolobales, a genome‐wide transposon mutagenesis study has shown clsN to be an essential gene for Sulfolobus islandicus viability (Zhang et al., 2018). ChIP‐Seq studies demonstrated ClsN is enriched in the less transcriptionally active B compartment of the chromosome. A previously unpublished analysis of the compartmentalized organization of the Saccharolobus solfataricus P2 (SSO) genome is shown in Figure 3a–d along with the ChIP‐Seq profile of SSO ClsN. As can be seen in Figure 3d, there is a statistically significant (two‐sided Wilcoxon test p < 2 × 10−16) enrichment of ClsN in the B compartment. Genome‐wide ClsN is most commonly localized in intergenic regions. Both across the genome as a whole and within the compartments, there is an anticorrelation between ClsN occupancy of genes and their transcription level (Takemata et al., 2019). While clsN is an essential gene, experiments to over‐express ClsN resulted in intensification of the B compartment and further repression of the already low levels of transcription of genes in the vicinity of its binding sites. Conversely, induction of transcription of previously silent and ClsN‐occupied loci leads to eviction of ClsN, concomitant with increased transcriptional activity. Overall, the available evidence to date supports ClsN playing a key role in structuring the genomes of members of the Sulfolobales, and the demonstrated mutual antagonism of transcription and ClsN occupancy suggests a potential role for ClsN as global regulator of transcription (Takemata et al., 2019). Notably, there is a general induction of genes within the B compartment as cells enter stationary phase. While the total amount of ClsN associated with chromosomal DNA is modestly enhanced in stationary phase, there is a redistribution of the ClsN protein on the chromosome, with a reduction of the ClsN occupancy in the loci located in the exponential phase B compartment. This eviction of ClsN occurs in parallel with an overall loss of integrity of the B compartment and transcriptional induction of these loci (Takemata et al., 2019). Whether the eviction of ClsN is a bystander effect of the enhanced transcription of these loci or a determining factor that facilitates transcriptional induction remains to be determined.

FIGURE 2.

FIGURE 2

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Chromosome conformation capture contact maps of chromosomes of a range of Sulfolobales species (left column). Sis—Sulfolobus islandicus REY15A: Sto—Sulfurisphaera (formerly Sulfolobus) tokodaii: Sac—Sulfolobus acidocaldarius DSM639 Ste—Sulfuracidifex tepidarius. The scale (CS) indicated contact score × 10−3. Pearson correlation coefficient heat maps are also shown in the middle column, and the scale (PC) indicates the Pearson correlation coefficient and on the right are principal component plots indicating the localization of loci falling into A or B compartment. Figures are generated from previously published 3C experiments (Badel et al., 2022; Takemata et al., 2019; Takemata & Bell, 2021c). Data are available at the Gene Expression Omnibus (GEO) Accession code GSE128063 and GES159537 and NIH SRA Project numbers PRJNA814106 (Ste and Sto).

FIGURE 3.

FIGURE 3

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(a) Pearson correlation coefficient heat map of contacts within the Saccharolobus solfataricus P2 chromosome. (b) Principal component analysis of the data shown in part (a). Regions of the genome corresponding to A and B compartments are shown above and below the midline, respectively. (c) ChIP‐Seq analyses of the localization of ClsN on the Saccharolobus solfataricus chromosome. The positions of A and B compartments are shown with white and gray backgrounds, respectively. (d) Violin plot of the binding of ClsN to A and B compartment loci. Data were analyzed using a Wilcoxon test (two sided) and the p‐value indicated. Data will be deposited at the SRA. Note that these data are previously unpublished and have been deposited at the Sequence Read Archive, BioProject Accession PRJNA1074953.

2. LIFE WITHOUT SMCs

The genetic essentiality of ClsN, its role in defining the B compartment, and its apparently intimate association with gene expression reveal it to be a key component of chromosomal organization in the Sulfolobales. Thus, even though it is an atypical SMC superfamily protein, it can be seen to fulfill functions associated with canonical SMCs in other organisms. At first glance, this might be seen as representing a non‐orthologous gene displacement event. Such an evolutionary step already has precedent within the bacterial domain. E. coli and some other members of the gamma proteobacteria encode the MukBEF complex in place of the canonical SMC‐ScpAB bacterial condensin (Hiraga et al., 1989). However, examination of Figure 4 suggests a more complex evolutionary derivation of ClsN. It is apparent that canonical condensin is nearly universal in archaea, with the exception of the crenarchaea. Thus, the genes for this complex were lost at or near the root of the crenarchaeal phylum. While some crenarchaeal species have acquired genes for lineage‐specific novel SMC‐related proteins, for example, ClsN in the Sulfolobales or the distantly related Rkd4 in the Thermoproteales, other lineages, such as the Desulfurococcales, have no SMC‐like proteins other than the Rad50 DNA repair protein. We have recently investigated how one such species, Aeropyrum pernix, organizes its chromosome in the absence of SMC proteins (Badel & Bell, 2024).

FIGURE 4.

FIGURE 4

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Distribution of condensin and ClsN SMC superfamily proteins in the TACK superphylum. Black circles indicate the presence of orthologs, and gray circles indicate non‐ubiquitous presence of orthologs in a lineage. Rkd4 is a distant paralog of ClsN found in the Thermoproteales.

Using a candidate locus approach, we previously identified two origins of DNA replication in the A. pernix chromosome (Robinson & Bell, 2007). Using a genome‐wide origin mapping approach, we confirmed that these two origins are active and that no further origin exists in the A. pernix genome. In agreement with all other archaea examined to date, admittedly a limited subset of the Sulfolobales and euryarchaea (Cockram et al., 2021, Takemata & Bell, 2021c), 3C experiments revealed that the chromosome of A. pernix possesses CIDs. However, unlike the situation with Sulfolobales species, we could not detect any association of CID boundaries with highly expressed genes. Treating cells with the translation inhibitor chloramphenicol led to weakening of CID boundaries. Intriguingly, however, the DNA intercalating agent, actinomycin D, while reducing transcription globally, led to a strengthening of CID insulation. Actinomycin D also had the unanticipated effect of inducing the expression of a subset of genes, probably involved in stress response. The Pearson correlation coefficient heat map of the A. pernix chromosome did not reveal the clear plaid pattern seen with Sulfolobales species. However, there was a distinct region, highlighted in a dashed box in Figure 5a, that showed reduced long‐range interactions with the majority of the chromosome, with the exception of a number of small regions, Figure 5a. Therefore, it appears that there is a small, insulated domain formed in the A. pernix chromosome. Interestingly, this compartment was enriched for highly expressed genes and also contained a statistically significant level of loop structures with both anchors present within the domain. Thus, this domain, which we termed “High Expression Insulated Domain” or HEID, is reminiscent of the A compartment seen in Sulfolobus chromosomes. Interestingly, the reprogramming of transcription upon actinomycin D administration led to the generation of a novel HEID' domain, based on the novel sites of elevated transcription. In contrast, the rest of the chromosome “ROC” had lower levels of transcription but otherwise largely lacked defining features and we suspect it is principally defined by its exclusion from the HEID. Thus, A. pernix possesses a rudimentary proto‐compartmentalized architecture that presumably represents an evolutionary forerunner of the more organized compartmentalization seen in Sulfolobales species. Interestingly, comparison of the replicon architecture of the A. pernix and Sulfolobales chromosomes provides hints regarding this transition to more structured compartments in Sulfolobales. As mentioned above, A. pernix has two replication origins, in contrast to the three seen in Sulfolobales. Two of the Sulfolobales replication origins (oriC1 and oriC3) are related to those of Aeropyrum, indicating that the unique Sulfolobales origin of DNA replication, oriC2, was likely acquired more recently. In this regard, it is notable that Sulfolobales oriC2 is located within 50 kb of clsN in a broad range of Sulfolobales species. It seems plausible, therefore, that ClsN was encoded on the extrachromosomal element that integrated into an A. pernix‐like, progenitor Sulfolobales chromosome, donating its origin and the clsN gene in the process. ClsN, with its affinity for lowly expressed regions of the genome, would have helped orchestrate formation of these regions into the fully coalesced B compartment seen in present‐day Sulfolobales chromosomes (Figure 5b). It would be of considerable interest to determine the consequences of introducing the clsN gene into A. pernix, with respect to chromosome organization and gene expression in that organism. Regrettably, however, there is currently no system for the genetic manipulation of this organism.

FIGURE 5.

FIGURE 5

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(a) Pearson correlation coefficient heat map arising from 3C analyses of the Aeropyrum pernix chromosome. The main region corresponding to the HEID domain is highlighted in a dashed yellow box. Additional constituent regions contributing to HEID are indicated with purple triangles. The principal component analysis is shown below with HEID domain in purple and the rest of the chromosome in blue. These figures were generated from data deposited in NCBI Sequence Read Archive (SRA). Submission ID: SUB13894161. BioProject ID: PRJNA1027590 (Badel & Bell, 2024). (b) Schematic proposing a pathway for the evolution of the compartmentalized organization of the Sulfolobales chromosomes. Orthologous DNA replication origins are color‐coded to overcome the rather unfortunate current nomenclature. We suggest that the integration of an extrachromosomal element that was replicated via the green replication origin and additionally encoded clsN led to the generation of a three‐replication origin, clsN‐encoding ancestral Sulfolobus chromosome. ClsN with its preference for the transcriptionally more‐quiescent regions of the chromosome would lead to coalescence of loci into a B compartment.

Finally, it is deeply intriguing that the genes for canonical condensin appear to have been lost at the root of the crenarchaeal phylum. In most bacteria, condensin plays a key role in facilitating the coupling of chromosome segregation with DNA replication. Notably, the crenarchaea have a fundamentally distinct logic of cell cycle progression with replication and segregation temporally separated. Indeed, studies in Saccharolobus solfatarics P2 provided evidence that sister chromatids remain cohesed for a considerable portion of the post‐replicative period, prior to segregation (Robinson et al., 2007). There is currently no evidence for ClsN playing a role in facilitating this cohesion. It has been noted that the period of cohesion correlates with the presence of hemi‐catenane junctions between the newly replicated sister chromatids. How hemi‐catenanes are resolved prior to segregation and the process of segregation itself remain poorly understood phenomena. For condensin‐containing, non‐crenarchaeal archaea, very little is known about how cell cycles are regulated, with the exception of some studies of halophiles and members of the Thermococcales. These organisms are highly polyploid and appear to lack the regimented cell cycle parameters observed in crenarchaea; for a recent review, see Bell (2022). Instead, concurrent rounds of replication and cell division appear to occur and this could suggest a more bacterial‐like mode of coordinated DNA replication and segregation. If this is indeed the case, it seems possible that the loss of condensin in the crenarchaea may have helped trigger the transition into a cell cycle with temporally separated DNA replication and chromosome segregation seen in these species. This cell cycle logic is, of course, superficially reminiscent of that seen in present‐day eukaryotes. It should be emphasized, however, that current analyses place the root of eukaryotes within the Asgard archaea and the Asgards encode condensin (Teske et al., 2021). Undoubtedly, more studies of cell cycle mode and chromosome conformation are needed before we can draw firm conclusions about the relationships between SMC content, ploidy, and cell cycle mode. In particular, entire phyla of archaea, including the Asgards, remain entirely unstudied at these levels. Expanding analyses into this unexplored phylogenetic space is a key goal for future work and will undoubtedly give fundamental insights into the evolution of both archaeal and eukaryotic chromosome dynamics.

3. METHODS

3.1. Strains, media, and growth conditions

The Saccharolobus solfataricus P2 strain was grown in Brock's medium (Brock et al., 1972) containing 0.1% tryptone, pH 3.2 at 78°C.

3.2. ChIP‐Seq

Chromatin immunoprecipitation (ChIP) was performed as described in Samson et al. (2013). Cultures of S. solfataricus P2 were crosslinked with 1% formaldehyde for 20 min. After quenching with 125 mM glycine, the cells were pelleted and washed with 1X PBS. The pellets were resuspended in TBS‐TT (20 mM Tris, 150 mM NaCl, 0.1% Tween‐20, 0.1% Triton X‐100, pH 7.5), passed through a French press three times at 20,000 psi, and sonicated using a Diagenode Bioruptor to generate DNA fragments ranging from 200 to 1000 bp. The extract was then clarified by centrifugation. 10 μg of extract (based on protein concentration) was used in each 100 μL ChIP reaction. Samples were rotated for 2–3 h with 3 μL of antiserum at 4°C. 25 μL of a 50% slurry of protein A sepharose was then added, and the samples were rotated for another hour at 4°C. Each ChIP reaction was then washed five times at room temperature with TBS‐TT, once with TBS‐TT containing 500 mM NaCl, and once with TBS‐TT containing 0.5% Tween‐20 and 0.5% Triton X‐100. Protein–DNA complexes were eluted from the protein A sepharose in 20 mM Tris, 10 mM EDTA, 0.5% SDS, pH 7.8 at 65°C for 30 min. Crosslinking was reversed and protein was digested by incubating the samples with 10 ng/μL proteinase K for 6 h at 65°C followed by 10 h at 37°C. The samples were extracted with phenol/chloroform/isoamyl alcohol first and then chloroform alone, and the DNA was precipitated in 100% ethanol containing 20 μg of glycogen. After washing with 70% ethanol and air drying, the DNA was resuspended in 50 μL TE buffer.

ChIP reactions were performed in triplicate and pooled. DNA libraries were prepared and sequenced on an Illumina HiSeq 2000 platform at the University of Wisconsin. Single‐end sequence reads were mapped to the S. solfataricus P2 genome using Geneious v 2022.0.2 (www.geneious.com). Data were exported as .bam files and analyzed using SeqMonk (www.bioinformatics.babraham.ac.uk/projects/seqmonk/). Read counts were quantified in 1 kb windows and were normalized to ChIP input DNA.

3.3. 3C‐seq

3C‐seq for S. solfataricus P2 was carried out essentially as described for our previous analyses of Sulfolobus islandicus—for detailed procedures, see Takemata and Bell (2021a); Takemata and Bell (2021b). Briefly, 20 mL of cell culture was crosslinked with 6% formaldehyde, quenched, washed, and stored at −80°C.

For library preparation, a frozen cell pellet was thawed, lysed, and digested with AluI. The reaction was then quenched with SDS prior to ligation with T4 DNA ligase (NEB) in the presence of Triton X‐100. Crosslinked were reversed by heat treatment, and the reaction was then treated with proteinase K. DNA was recovered by organic extraction followed by ethanol precipitation prior to resuspension in 40 μL of 1 × NEBuffer 2 containing 0.1 mg/mL RNase A. The success of proximity ligation was confirmed by running 10 μL of the DNA on a gel. The remaining DNA was further extracted with phenol:chloroform:isoamyl alcohol and ethanol‐precipitated. The DNA then resuspended in Buffer EB (QIAGEN) and sheared with a Bioruptor (Diagenode). The sheared DNA was used for library construction with NEBNext Ultra DNA Library Prep Kit for Illumina and NEBNext Multiplex Oligos for Illumina (NEB) according to the manufacturer's instructions. DNA was purified using AMPure XP Beads (Beckman Coulter) to enrich adaptor‐ligated DNA molecules of 400–500 bp. Eight or nine cycles of PCR were conducted for library amplification. DNA libraries were paired‐end sequenced (43 bp × 2) on the Illumina NextSeq platform at the Center for Genomics and Bioinformatics at Indiana University.

3.3.1. Generation of 3C‐seq matrices

3C‐seq reads were processed using HiC‐Pro version 2.9.0 (Servant et al., 2015) as described (Takemata et al., 2019). Genome sequences were then binned at different resolutions to generate raw contact matrices. Valid read pairs from biological replicates were pooled and used for iterative correction implemented in HiC‐Pro. Bins with extremely low coverage were filtered out by setting the FILTER_LOW_COUNT_PER parameter to 0.01. The low coverage over the filtered‐out bins was due to a large number of repeat sequences (transposons, etc.) in the S. islandicus genome, whereas the number of such repeats is very small in the S. acidocaldarius genome. After the iterative correction, all values were multiplied to make the sum of interaction scores equal to 1000 for each row and column.

Distance‐normalized contact matrices, Pearson correlation matrices, and compartment index plots were generated as described previously using HiTC (Servant et al., 2012; Takemata et al., 2019).

3.4. NGS data

The 3C‐Seq and ChIP‐Seq data have been deposited at the Sequence Read Archive, BioProject accession PRJNA1074953 https://www.ncbi.nlm.nih.gov/sra/?term=PRJNA1074953.

AUTHOR CONTRIBUTIONS

Stephen D. Bell: Conceptualization; funding acquisition; writing – original draft; supervision; project administration. Elyza Pilatowski‐Herzing: Writing – original draft. Rachel Y. Samson: Writing – review and editing; investigation; conceptualization. Naomichi Takemata: Conceptualization; investigation; writing – review and editing; software. Catherine Badel: Conceptualization; writing – review and editing; software. Peter B. Bohall: Writing – review and editing; visualization.

CONFLICT OF INTEREST STATEMENT

The authors declare no conflict of interests.

ETHICS STATEMENT

No human or animal subjects were used in this work.

ACKNOWLEDGMENTS

We thank the DNA Sequencing Facility at the Biotechnology Center, University of Wisconsin–Madison, and the Center for Genomics and Bioinformatics at Indiana University, Bloomington, for DNA sequencing. This work in SDB's lab has been funded by NIH grant R01GM135178 and is currently funded by R35GM152171.

Pilatowski‐Herzing, E. , Samson, R.Y. , Takemata, N. , Badel, C. , Bohall, P.B. & Bell, S.D. (2025) Capturing chromosome conformation in Crenarchaea. Molecular Microbiology, 123, 101–108. Available from: 10.1111/mmi.15245

DATA AVAILABILITY STATEMENT

The data that support the findings of this study are openly available in NIH Sequence Read Archive at https://www.ncbi.nlm.nih.gov/sra/?term=PRJNA1074953, reference number PRJNA1074953.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Data Availability Statement

The data that support the findings of this study are openly available in NIH Sequence Read Archive at https://www.ncbi.nlm.nih.gov/sra/?term=PRJNA1074953, reference number PRJNA1074953.

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