Nature Reviews Microbiology | AOP, published online 6 May 2015; doi:10.1038/nrmicro3467

PROGRESS The interplay between nucleoid organization and transcription in archaeal genomes Eveline Peeters, Rosalie P. C. Driessen, Finn Werner and Remus T. Dame

Abstract | The archaeal genome is organized by either eukaryotic-like histone proteins or bacterial-like nucleoid-associated proteins. Recent studies have revealed novel insights into chromatin dynamics and their effect on gene expression in archaeal model organisms. In this Progress article, we discuss the interplay between chromatin proteins, such as histones and Alba, and components of the basal transcription machinery, as well as between chromatin structure and gene-specific transcription factors in archaea. Such an interplay suggests that chromatin might have a role in regulating gene expression on both a global and a gene-specific level. Moreover, several archaeal transcription factors combine a global gene regulatory role with an architectural role, thus contributing to chromatin organization and compaction, as well as gene expression. We describe the emerging principles underlying how these factors cooperate in nucleoid structuring and gene regulation. In all domains of life, genome organization occurs in a functional and dynamic manner, packaging the genome into the nucleus (in eukaryotes) or into the cell (in bacteria or archaea) and, at the same time, efficiently accommodating DNA-based processes such as transcription, replication and repair. Histone proteins are the prototypical organizers of eukaryotic DNA, wrapping the DNA around an octameric histone protein core into nucleosomes, which imposes the first level of organization1. Bacteria lack histone proteins and instead harbour nucleoidassociated proteins (NAPs) that reduce the effective volume of the genome by local DNA deformation (bending or wrapping) or by stabilizing long-distance contacts along the genome and forming loops (bridging). Proteins involved in compactly folding and organizing the genome (that is, eukaryotic histones or bacterial NAPs) are collectively termed ‘chromatin proteins’. In addition to genome compaction, both eukaryotes and bacteria have mechanisms that divide the genome into functional domains, including the formation of chromatin loops2–4 and the

attachment of chromatin to the nuclear lamina in eukaryotes5 or the cell membrane in bacteria6. Cell differentiation processes and cellular adaptation to varying physiological conditions are controlled by the regulation of distinct genes, which is partly mediated by local and global chromatin remodelling in both eukaryotes and bacteria. In eukaryotes, the exact positioning of nucleo­somes along the genome is crucial in global gene regulation, and histone modifications constitute an important mechanism to modulate genome accessibility 7. Chromatin remodelling is an active process involving large and complex ATP-consuming complexes8,9. In bacteria, chromatin remodelling seems to be a passive process that relies on changes in the relative expression levels of NAPs with different architectural properties10,11. By contrast, in archaea, the interplay between chromatin organization and transcription processes still remains enigmatic. Archaea harbour various chromatin proteins; whereas Euryarchaeota mainly have homologues of eukaryotic histone proteins, Crenarchaeota generally lack histone

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homologues and only rely on a set of other small proteins that are functional homologues of bacterial NAPs12. In each archaeal organism, gene expression processes are also characterized by a bacterial–eukaryoti­c hybrid nature of features. The archaeal basal transcription machinery shows strong similarities with its eukaryotic RNA polymerase II counterpart 13, but archaeal gene organization (for example, the presence of operons) resembles that of bacteria, and most archaeal regulatory transcription factors are similar to typical bacterial transcription factors14 (BOX 1). In this Progress article, we discuss emerging concepts of archaeal nucleoid organization in the context of gene expression. The phylogenetic heterogeneity in archaeal nucleoid components suggests that the interplay between chromatin organization and transcription does not fit either a bacterial or a eukaryotic paradigm but is unique in many aspects. Furthermore, we emphasize the fact that it is not always straightforward to classify archaeal DNA-binding proteins as either dedicated nucleoid-structuring proteins or conventional transcription factors. We first discuss chromatin organization in archaea and the key factors involved, followed by a review of our current understanding of how histones and the chromatin protein Alba affect basal transcription. Finally, we discuss recent insights into the interplay between histones and gene-specific transcription factors in regulating gene expression and chromatin structure. Chromatin organization in archaea Each archaeal genome harbours at least two different types of chromatin proteins with distinct architectural properties (FIG. 1) and often multiple paralogues of each type with functional overlap. So far, several distinct chromatin protein families that show an uneven phylogenetic distribution have been characterized (TABLE 1). The best-studied chromatin proteins belong to the Alba superfamily, which is widely distributed and almost universally present in archaea15. Alba seems to have an ancient evolutionary history and considerable functional plasticity 16. Most Alba proteins interact with RNA in addition ADVANCE ONLINE PUBLICATION | 1

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PROGRESS Box 1 | Basal and regulated transcription in archaea The archaeal transcription machinery is closely related to the RNA polymerase II system in terms of subunit composition, structure, use of general factors and molecular mechanisms86,87. Basal factors form a pre-initiation complex (PIC) with RNA polymerase (RNAP) and guide this enzyme through initiation and elongation by modulating its DNA- and RNA-binding, and catalytic properties13. Binding of TATA-binding protein (TBP) to the promoter induces a recruitment cascade that leads to transcription initiation; transcription factor B (TFB) binds to the TBP– DNA complex, and the DNA–TBP–TFB complex subsequently recruits RNAP and transcription factor E (TFE) (see the figure, part a). TFB facilitates the loading of the template strand into the catalytic centre of RNAP and stimulates early catalysis88, whereas TFE aids DNA melting and stabilizes the ‘open’ complex46,89,90. TBP, TFB and TFE interact with promoter DNA in distinct ways: TBP and TFB bind in a sequence-specific manner to the double-stranded TATA and B‑recognition element (BRE) motifs of the promoter, respectively, whereas TFE interacts with the non-template strand, probably in a sequence-independent manner. An additional cis-element named initiator (Inr) includes the transcription start site and influences the strength of some promoters91. In contrast to the basal transcription machinery, gene-specific signal-responsive transcription regulation is mediated by small bacterial-like

a Transcription initiation PIC

transcription factors14,92,93. Transcription factors can regulate transcription by binding close to or overlapping with the promoter, thereby enhancing or repressing the sequential assembly of the PIC94,95. In many instances, it has been shown that cooperative binding of multiple protein molecules leads to regulation (see the figure, part b). Whether a transcription factor affects transcription initiation in a positive or negative manner is mainly determined by the relative position of the transcription factor-binding site (or sites) with respect to the TATA box, BRE and Inr elements. Activators usually bind to DNA upstream of the BRE and function in the initial steps of PIC assembly by stimulating the recruitment of TBP and/or TFB to the promoter through protein–protein interactions96 (see the figure, part b). By contrast, repressors bind to promoter-overlapping sequences, thereby preventing access of either the basal transcription factors TBP and TFB (if the operator overlaps with the TATA box and BRE; see the figure, part b) or of RNAP in a later stage of PIC assembly (if the operator directly precedes or overlaps with the Inr)97. Archaeal transcription factors are not always dedicated activators or repressors but can sometimes alternate between both roles at different promoters, depending on operator positioning80,98,99, or at the same promoter, depending on operator occupancy94.

b Transcription activation or repression

RNAP

TBP and TFB recruitment

TFB

TBP

RNAP

BRE

TATA

TFE

No TBP and TFB recruitment

TFB

TBP

BRE

TATA

+1

Activating transcription factor

TFE

TFB TBP BRE

TATA +1

Repressing transcription factor

+1 Nature Reviews | Microbiology

to binding to double-stranded DNA (dsDNA)17,18 and have been suggested to function in RNA metabolism16. In euryarchaeal methanogenic archaea, Alba proteins are low-abundance, sequence-specific dsDNA-binding proteins19, whereas in crenarchaeal organisms, it was shown that Alba is a highly abundant cellular protein that binds to dsDNA without apparent sequence specificity 15,20. Alba assembles into dimers, which are homodimeric or hetero­ dimeric depending on whether paralogues are encoded and on their relative amounts. These dimers show dimer–dimer interactions and bind to DNA in cis or trans (FIG. 1c). As binding of Alba proteins in euryarchaea is sequence-specific, it is expected that binding in trans — yielding loops — occurs between two specific sites (FIG. 1a,b). Electron microscopy and atomic force microscopy (AFM) studies of Alba–DNA complexes revealed that crenarchaeal Alba shows a bimodal DNA-binding behaviour depending on the protein concentration21–23. At low and intermediate concentrations, binding of Alba results in DNA compaction as a result of the formation of looped structures (FIG. 1c). At high protein concentrations, it fully coats

DNA as a result of cooperative binding, giving rise to stiff filaments. The co‑crystal structure of the DNA-bound Alba homologue from Aeropyrum pernix revealed that dimer–dimer interactions of Alba enable the bridging of two DNA duplexes24, which is consistent with the microscopy studies showing loop formation21. Moreover, a study using Alba paralogues from Sulfolobus solfa‑ taricus and mutants confirmed the importance of the dimer–dimer interface and highlighted the crucial role of the Phe60 residue in both DNA bridging and cooperative binding along DNA23. In the euryarchaeal phylum, histones have evolved as the most abundant chromatin proteins25 (FIG. 1a,b). Most eury­archaeal species encode two homologues of the eukaryotic core histones H3 and H4, which form wrapped nucleoprotein structures26. The prevailing view is that these nucleo­somes are tetrameric structures composed of either heterodimers or homodimers27 with around 60 bp of DNA wrapped around the surface of the tetramer 26,28,29, which is in contrast to the 150 bp associated with eukaryotic octameric nucleosomes1. Visualization of protein–DNA complexes formed with the

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histone proteins HMf (from Methanothermus fervidus), MkaH (from Methanopyrus kandleri) and TmkH (from Thermococcus kodakarensis) revealed sparsely decorated ‘beads‑on‑a‑string’ structures similar to eukaryotic chromatin fibres30–33. Haloferax volcanii has a uniform chromatin architecture with regularly spaced tetrasomes34 (FIG. 1a). By contrast, T. kodakarensis and Methanothermobacter thermautotrophicus histones can assemble into chromatin particles of variable sizes that occupy DNA stretches ranging from 30 bp to more than 450 bp, with 30 bp of DNA added for complexes of increasing size29,35 (FIG. 1b). The 30 bp unit size suggests that histones bind to DNA as dimers that — analogous to the assembly of eukaryotic octasomes — can associate side‑by‑side and assemble into large multimers. Consistent with this, in addition to forming tetrameric nucleosome-like structures, HMf proteins can also associate with DNA as dimers and induce DNA bending 36. The determinants of forming larger multi­mers along DNA are currently unclear; histone dimers possibly bind to repeating sequence elements that position the dimeric units favourably for dimer–dimer interactions37. www.nature.com/reviews/micro

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PROGRESS a Euryarchaeal nucleoid

b Euryarchaeal nucleoid

H. volcanii

T. kodakarensis

60 bp

Histone tetramer

120 bp

60 bp 30 bp

180 bp

150 bp

Alba tetramer

c Crenarchaeal nucleoid Sulfolobus spp. Alba1 homodimer cis Alba1–Alba2 heterodimer

Cren7 or Sul7

Figure 1 | Chromatin organization in archaea.  The structure of the archaeal nucleoid varies among different archaeal species depending on the chromatin proteins they express. a,b | The euryarchaeal nucleoid is mainly organized by histone proteins that bend or wrap DNA, as well as by Alba that binds to DNA as a homodimer or a heterodimer and that forms looped structures by bridging two DNA duplexes. In Haloferax volcanii, histone proteins form tetrameric nucleoprotein structures that wrap about 60 bp of DNA around their surface (part a). These nucleosomes form a regular ‘beads‑on‑a‑string’ structure similar to eukaryotic chromatin. In Thermococcus kodakarensis, histone proteins assemble into multimeric forms that cover variable sizes of DNA ranging from 30 bp (indicative of a

Crenarchaeota have, in addition to Alba, several 7 kDa-sized architectural proteins such as Cren7 and Sul7 (REFS 38–40) (FIG. 1c; TABLE 1). The lack of conservation of archaeal chromatin proteins within the archaeal phylum is further corroborated by the ongoing discovery of novel NAPs in crenarchaea: examples are Sso7c and Sso10a in S. solfataricus 41,42 and the crenarchaeal chromatin protein 1 (CC1) that is found in Thermoproteus tenax, Pyrobaculum aero‑ philum and A. pernix 43,44 (TABLE 1). Several of these proteins have DNA-binding folds that are typical of bacterial and archaeal transcription factors32,41,42. These structural similarities show that it is problematic to deduct function from sequence or structure alone and suggest that these factors can have dual functions in chromatin structuring and transcriptional regulation. Chromatin and basal transcription Despite differences on a sequence and a structure level, most archaeal chromatin proteins have in common that they are small (7–10 kDa), highly abundant in the cell and typically bind to dsDNA with no or very low

trans

trans cis

dimer binding) to 450 bp (part b). c | The crenarchaeal nucleoid is organNature Reviews | Microbiology ized by proteins that bend DNA (for example, Cren7 and Sul7 in Sulfolobus spp.), as well as by Alba that either forms looped structures by bridging two DNA duplexes or forms stiff filaments by binding cooperatively side by side. The bimodal DNA-binding behaviour of Alba depends on protein concentration: bridges are formed at low and intermediate concentrations, and stiff filaments are formed at high concentrations. In Sulfolobus spp., cis and trans binding of Alba is affected by the formation of either homodimers or heterodimers. Alba homodimers bind cooperatively and have a tendency to bind in cis, whereas heterodimers bind non-cooperatively and are therefore more likely to bind in trans (see inset).

sequence specificity. Continued research efforts have yielded detailed mechanistic insights into how most of these proteins interact with DNA; however, the effect of these proteins on the basal transcription machinery has only been mechanistically unravelled for archaeal histones and Alba. Histone binding to DNA represses transcription. Accumulating evidence suggests that nucleosomal organization in archaea is interlinked with transcription processes (FIG. 2). As in eukaryotic chromatin, nucleosome-depleted regions are found immediately upstream and downstream of transcription units, comprising promoters and terminators, respectively 28,29,34, and are surrounded by an upstream and a downstream nucleosome-enriched region (–1 and +1 nucleosomes)28,34. Furthermore, histone binding is decreased throughout highly transcribed genes such as ribosomal DNA genes and the mcr operon in methanogenic archaea26,34. This suggests that there is a functional link between the organization of nucleosomes and transcription activity; however, it remains to be determined

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whether active transcription is primarily the outcome of a histone-free DNA template, whether it causes the depletion of histones or whether it is a combination of both28,29. Similarly to their eukaryotic homologues, archaeal histones have a slight sequence bias towards alternating GC- and AT‑rich dinucleotides and trinucleotides that are off-set by half a helical turn28,29,34. Chromatin that has been reconstituted in vitro from recombinant histones and purified genomic DNA has a near-identical histone occupancy profile compared with native chromatin. As reconstitution occurs in the absence of ongoing transcription, the sequence bias of histone–DNA interactions seems to have a major role in nucleosome deposition. Several studies have shown archaeal histone-mediated gene silencing in vitro. Histones can repress transcription by preventing the binding of the initiation factors TATA-binding protein (TBP) and transcription factor B (TFB) to the TATA box and B-recognition element (BRE) promoter elements, respectively (FIG. 2a), or by blocking RNA polymerase (RNAP) binding to the template35,45. Using a Methanocaldococcus ADVANCE ONLINE PUBLICATION | 3

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PROGRESS Table 1 | Distribution* of archaeal chromatin proteins and families of archaeal transcription factors among different archaeal species

Sso7c

MC1

7kMK

TrK

Lrp

TrmB

ArsR

HTH_3

–

–

✓

✓

✓

✓

–

✓

–

–

–

–

✓

✓

✓

✓

✓

✓

–

✓

–

–

–

✓

✓

✓

✓

–

✓

✓

–

✓

–

–

–

✓

✓

✓

✓

–

–

✓

✓

–

✓

–

–

–

✓

✓

✓

✓

–

–

✓

–

–

–

–

–

–

✓

✓

✓

✓

✓

–

✓

–

–

–

–

–

–

✓

✓

✓

✓

✓

–

✓

✓

–

–

–

–

–

✓

✓

✓

✓

–

–

–

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✓

–

–

–

–

–

✓

✓

✓

✓

✓

–

–

–

✓

–

–

–

–

–

–

✓

✓

✓

✓

Pyrobaculum

✓

–

✓

–

✓

✓

–

–

–

–

–

✓

✓

✓

✓

Thermoproteus

✓

–

✓

–

✓

✓

–

–

–

–

–

✓

✓

✓

✓

Vulcanisaeta

✓

–

–

–

✓

✓

✓

–

–

–

–

✓

✓

✓

✓

Caldivirga

✓

–

–

–

✓

✓

✓

–

–

–

–

✓

✓

✓

✓

Thermofilum

✓

–

–

–

–

–

✓

–

–

–

–

✓

✓

✓

✓

Korarchaeota

‘Candidatus Korarchaeum’

✓

–

–

–

–

–

✓

–

–

–

–

✓

✓

✓

✓

Euryarchaeota

Archaeoglobus

✓

–

–

–

–

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✓

–

–

–

–

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✓

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Methanosarcina

–

–

–

–

–

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–

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–

–

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Halobacterium

–

–

–

–

–

–

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–

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Methanobacterium

✓

–

–

–

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–

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–

–

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Methanococcus

✓

–

–

–

–

–

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–

–

–

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Methanosphaerula

✓

–

–

–

–

–

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–

–

–

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Methanopyrus

✓

–

–

–

–

–

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–

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Pyrococcus

✓

–

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Thermoplasma

✓

–

–

✓

–

–

–

–

–

–

–

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Nanoarchaeum

✓

–

–

–

–

–

✓

–

–

–

✓

✓

✓

–

–

Cren7

–

HU

–

CC1

✓

Sul7

–

Genus

Alba

Histone

Transcription regulators Sso10a

Chromatin proteins

Thaumarchaeota

Cenarchaeum

✓

–

–

–

–

Aigarchaeota

‘Candidatus Caldiarchaeum’

✓

–

–

–

–

Sulfolobus

✓

✓

–

–

Metallosphaera

✓

✓

–

Acidianus

✓

✓

Ignicoccus

✓

–

Desulfurococcus

✓

–

Aeropyrum

✓

–

Ignisphaera

✓

Acidilobus

Phylum

Crenarchaeota

Order

Sulfolobales

Desulfurococcales

Thermoproteales

Thermococcus Nanoarchaeota

*Protein distributions were obtained by carrying out standard protein BLAST . 100

jannaschii in vitro transcription system, it was shown that the formation of tetrameric nucleosomes in the promoter region efficiently represses transcription, and the inhibition level correlates with the histone concentration45. Similarly, histones are also capable of inhibiting transcription initiation when initially bound in coding sequences, which are more common nucleosome localization sites compared with promoter regions. For example, when HMtA2 histones of M. thermautotrophicus bind downstream

of the transcription start site (TSS), they facilitate the formation of a filament that extends upstream and that covers the promoter elements, thus inhibiting transcription initiation35. Unfortunately, the effect of transcription factor E (TFE), a third initiation factor, has not been established in these experiments. As TFE stabilizes the transcription initiation complex 46, it might counteract the repressive effects of histones. In addition to interfering with the initiation phase of the transcription cycle, nucleosomes can

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function as a barrier to transcription elongation complexes (FIG. 2b). In vitro transcription elongation assays showed that although HMtA2 does not block RNAP progression, it reduces the average transcription elongation rate by threefold35. DNA-bound histones induce pausing of transcription elongation complexes at multiple locations but predominantly directly upstream of the HMtA2 tetrasome assembled at a high-affinity histone-binding site, which is located about 125 bp downstream of the TSS. www.nature.com/reviews/micro

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PROGRESS Future in vitro experiments are required to determine RNAP elongation rates on chromatinized templates in the presence of transcription elongation factors such as the cleavage factor TFS47,48 and particularly the processivity factor Spt4/5. TFS is crucial in reactivating backtracked elongation complexes, whereas Spt4/5 improves RNAP processivity and increases its elongation rate. All of these factors thereby alter the properties of RNAP and are likely to assist the elongation complex in overcoming physical obstacles such as histone-based nucleosomes and other chromatin structures49–51. Methanococcus voltae and T. kodakarensis strains lacking single histone proteins (HstA and HtkB, respectively) are viable but show some growth defects52,53. Substantial transcriptomic and proteomic differences were observed in these mutant strains compared with wild-type strains, which is in agreement with a role for histones in gene expression. How histone deletion affects growth remains to be determined, but the largest upregulation was observed in the expression of proteins that have a role in purine metabolism53. Interestingly, in each of these mutant strains, genes were not only upregulated but also downregulated, which suggests that histones are also capable of activating transcription either directly or indirectly 28,52,53. Alba protein levels decreased upon deletion of a histone protein in T. kodakarensis53. However, the nucleosome occupancy profiles immediately upstream of T. kodakarensis genes that were differentially expressed in the absence of either HtkA or HtkB did not reveal a consistent correlation between histone binding and transcriptional repression or activation28. Thus, the relationship between nucleosomal organization and gene expression in vivo seems to be more complex than that implied by the in vitro studies. Moreover, as the DNAbinding affinity of histone tetramers depends on the subunit composition54, histone-based chromatin structure and thus gene expression could also be modulated by differential expression of histone paralogues55,56. According to the eukaryotic paradigm, gene regulation is mediated by post-translational modification (PTM; such as methylation, acetylation and phosphorylation) of histones at the histone tails protruding from the nucleosome, thereby modulating chromatin structure57,58. Interestingly, euryarchaeal histones lack the amino- and carboxy-terminal tails that are modified in their eukaryotic counterparts and also do not seem to be post-translationally modified at other sites59. Nevertheless, archaeal genomes encode bona fide homologues of

eukaryotic chromatin remodellers, including prefoldin, which is a co‑chaperone that was recently shown to have a role in eukaryotic chromatin dynamics60–62. Whether these homologues modify histone-based chromatin and reorganize the archaeal chromatin structure, and whether this has any effect on transcription, remains to be determined.

a Competitive binding

Histones

TFB TBP BRE TATA

b Decreasing RNAP elongation rates RNAP

Alba differentially regulates gene expression. Although Alba has evolved into a chromatin protein in the Crenarchaeota, this is not true for all homologues in other archaeal lineages. In mesophilic methanococci, Alba proteins seem to have a specific role in transcription regulation19. As an example, the Methanococcus maripaludis Alba protein, Mma10b, is expressed at lower levels than its orthologue in Sulfolobus shi‑ batae (0.01% versus 1.6% of the total cellular protein)19 and binds to DNA in a sequencespecific manner — two characteristics that are typical of transcription factors and not of chromatin proteins. Deletion of Mma10b or its homologue in M. voltae has been shown to result in upregulation of several genes involved in carbon dioxide assimilation, including the gene encoding pyruvate ferredoxin oxido­reductase19,52, which suggests that this Alba protein has a specific gene regulatory role in autotrophic growth. In support of this, autotrophic growth is impaired in the M. maripaludis mma10b mutant 19. Sulfolobus spp. Alba1 is acetylated by protein acetyltransferase (Pat) at Lys16 on the DNA-binding surface17, and silent information regulator 2 (Sir2) deacetylates this residue15. In vitro DNA-binding experiments have shown that Lys16 acetylation reduces the DNA-binding affinity of Alba1 (REF. 15) (FIG. 2c). The functional modulation of Sir2 via NAD63 suggests that the acetylation state and thus DNA binding of Alba1 might be affected by NAD levels in vivo. Moreover, a study using a reconstituted in vitro transcription system that included the strong Sulfolobus spp. spindle-shaped virus 1 (SSV1) T6 promoter, TBP, TFB and RNAP showed that transcription was repressed by recombinant (non-acetylated) Alba1 purified from Escherichia coli, whereas native (acetylated) Alba1 purified from S. solfa‑ taricus had no effect on transcription15. As high concentrations of non-acetylated Alba1 repress transcription in vitro, its acetylation state potentially regulates transcription in vivo15,64 (FIG. 2c). The Alba1 Lys16 residue that is subject to acetylation is not conserved in all archaeal species19, which suggests that this mode of regulation is not universal.

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c Chromatin protein modulation Alba1

TFB TBP BRE TATA K16ac

Figure 2 | Putative transcription regulatory Nature Reviews | Microbiology mechanisms of chromatin proteins in archaea.  a | Histone proteins compete for DNA binding with the transcription initiation factors TATAbinding protein (TBP) and transcription factor B (TFB) to their respective promoter elements, the TATA box and B-recognition element (BRE). Depending on many factors (for example, expression levels of chromatin proteins and their DNAbinding affinity), either histones or the initiation factors bind to the DNA, and transcription initiation is repressed or permitted, respectively. b | During transcription elongation, DNA-bound histones reduce RNA polymerase (RNAP) elongation rates, but elongation is not completely abrogated. c | Acetylation of the Lys16 residue (K16ac) of Alba1 reduces its DNA-binding affinity. It has been suggested that this post-translational modification has a role in global gene regulation by permitting access of transcription factors to the promoter site.

It remains to be determined whether PTMs of Alba1 regulate gene expression in vivo and whether these phenomena are specific for subsets of genes, individual genes or whether they affect transcription on a global level. Chromatin and transcription factors The action of chromatin proteins affects gene expression not only by modulating the basal transcription machinery but also through the interplay with gene-specific transcription factors. In addition to their main role as gene regulators, transcription factors can contribute to chromatin struc­ turing. Thus, chromatin and transcription factors have a bilateral relationship.

Nucleosomes affect transcription factor binding. Archaea have small transcription factors that typically bind to inverted repeat ADVANCE ONLINE PUBLICATION | 5

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PROGRESS a

Histone TATA

UAS

No rb2 expression +1

b TFB BRE

TBP TATA

Low rb2 expression level

+1

c Ptr2 TFB BRE

TBP TATA

+1

High rb2 expression level

Figure 3 | Interplay between transcription factors and histones modulates transcriptional regu­ Nature Reviews | Microbiology lation.  In the euryarchaeon Methanocaldococcus jannaschii, the upstream activating site (UAS) is located directly upstream of the rubredoxin (rb2) promoter. Both histones and the Lrp-type regulator Ptr2 have been shown to bind to this region. a | In the absence of Ptr2, the histone tetramers bind to the UAS as well as to upstream and downstream regions, which results in transcriptional repression in vitro. b | TATA-binding protein (TBP) and transcription factor B (TFB) binding can overcome transcriptional repression mediated by the histones. c | At high Ptr2 concentrations, the transcription factor binds to the UAS and activates transcription by recruiting TBP to the promoter, which leads to high expression levels.

sequences as dimers or higher-order dimer oligomers in accordance with the bacterial paradigm of transcription factor–DNA interactions. This feature has been exploited in a comparative analysis of histone-binding regions and inverted repeat-harbouring sequences in the genome of T. kodakarensis, which shows lower nucleosome occupancy around putative transcription factor-binding sites29. Nucleosome depletion in genomic regions associated with regulatory transcription factors is also observed in eukaryotic chromatin65; however, the causal relationship between nucleosome positioning and transcription factor binding is unclear and highly debated in the eukaryotic field66. The similarity between in vitro and in vivo nucleo­some occupancy profiles28 suggests that sequence-directed nucleosome positioning enables preferential binding of transcription factors in chromatin-free regions rather than that transcription factor binding poses a barrier for binding of histones and determines the positioning around resulting chromatin-free regions. The anti-correlation pattern between histone-binding sites and transcription factor-binding sites is not universal, as shown by the observation of a direct interplay between histones and a regulatory transcription factor in M. jannaschii 45. In

this euryarchaeon, the transcription factor Ptr2 — a member of the bacterial–archaeal Lrp (also known as AsnC) family of transcription factors — activates the fdxA and rb2 genes, which encode a ferredoxin and a rubredoxin, respectively 67,68. In both target promoter regions, Ptr2 binds to an upstream activating site (UAS) composed of two helical aligned 15 bp Ptr2‑binding sites located directly upstream of the BRE. Ptr2 stimulates the recruitment of TBP, thus activating transcription67. The UAS in the rb2 promoter region has a dual function: in addition to being a high-affinity binding site for Ptr2, it also contains binding sites for histone tetramers45. In an in vitro transcription system it has been shown that, although histones silence the rb2 promoter compared with a non-chromatinized template (FIG. 3a), TBP and TFB are still capable of competing to some extent for access to the promoter elements45 (FIG. 3b). However, when present at high levels, Ptr2 outcompetes histone binding and carries out transcriptional activation (FIG. 3c). Notably, the level of activation (comparing transcription output in the absence or the presence of Ptr2) is dependent on histone concentration, and sufficiently high histone levels have been shown to induce maximal activation levels. Overall, Ptr2‑mediated activation levels are

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higher with a histone-bound template than with a non-chromatinized template45. Thus, histones have the capability to increase the potency of transcriptional activators, which would be an advantage in the case of genes with intermediate to high intrinsic promoter strengths. In conclusion, this case study yields a contradictory view from the above-mentioned genome-wide analysis in T. kodakarensis and shows that histones and transcription factors are not always binding in an independent manner from each other. Transcription factors modulate chromatin structure. Some of the best-studied archaeal transcription factor families contain members that combine a global gene regulatory role with an architectural role. It is therefore plausible that these regulators contribute to chromatin organization and compaction, and regulate physiological processes (for example, biofilm formation). The Lrp transcription factor family is widespread in archaea and is also found in bacteria69. The prototypic Lrp protein in E. coli is not only a transcription regulator of genes involved in amino acid metabolism but also a DNA-organizing NAP70,71. In archaea, several Lrp-type transcription factors have been characterized and were shown to regulate amino acid metabolism, as well as central carbon and energy metabolism69. Although most of these characterized proteins are specific local transcription regulators that control only one or a few target genes (for example, Ss‑LrpB in S. sol‑ fataricus activates the expression of a pyruvate ferredoxin oxidoreductase operon and two permease genes)68,72,73, some Lrp-type transcription factors were shown to regulate the expression of many genes (for example, Lrp in Halobacterium salinarum regulates many genes involved in amino acid synthesis, central carbon metabolism, transport processes and transcription regulation)73,74. Sa‑Lrp of Sulfolobus acidocaldarius, which is highly conserved in Sulfolobales, has been implicated in stress-induced cell aggregation on the basis of the diminished cellular aggregation characteristics of an Sa‑lrp deletion strain and the observed upregulation and downregulation of several genes encoding pili in this strain. It has been suggested that Sa‑Lrp has a global gene regulatory role as well as an architectural role instead of being a specific transcription factor. Besides the observation that genes encoding amino acid biosynthetic enzymes and other transcription factors were differentially regulated in the Sa‑lrp deletion strain, it was also observed that Sa‑Lrp is expressed at high www.nature.com/reviews/micro

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PROGRESS intracellular concentrations and that it binds to DNA with low sequence specificity; these two features are typical characteristics of chromatin proteins. Furthermore, AFM imaging of Sa‑Lrp–DNA complexes showed Sa‑Lrp-induced DNA wrapping. These findings support the idea that Sa‑Lrp contributes to nucleoid architecture, which is analogous to E. coli Lrp75. The archaea-specific Lrs14 family is another example of gene-regulating transcription factors that also function as NAPs. Lrs14‑like proteins were isolated from cellular extracts of S. solfataricus 76–78, and in vitro transcription assays have shown that Lrs14‑like proteins can either repress or activate transcription depending on the target gene77,78, which supports their role in gene regulation. In addition, genetic analysis of lrs14‑like genes in S. acidocaldarius showed that these proteins have a role in biofilm formation and cellular motility 79. The Lrs14‑like Abfr1 in S. acidocaldarius and its orthologue in S. solfataricus (named Smj12) bind to DNA with high affinity but very low sequence specificity and induce strong DNA deformations76,79. Furthermore, Smj12 has been shown to induce positive super­coiling 76. This suggests a dual role of Abfr1 and Smj12 in the regulation of gene expression as well as in the modulation of chromatin structure. Finally, archaeal TrmB-like proteins are specific transcription factors involved in the regulation of sugar metabolism in response to the direct binding of sugar ligands80. However, the TrmB-like protein TrmBL2, which belongs to the same protein family as these sugar-responsive regulatory proteins, was identified as a structural component of T. kodakarensis chromatin. This 32 kDa protein is abundantly present in histone-free regions in Thermococci in vivo and forms thick stiff filaments when bound to DNA in vitro30, which suggests that TrmBL2 has an important role in chromatin organization30. Conclusions and perspectives Pioneering research has revealed a diverse range of DNA-binding proteins that organize the archaeal genome on a structural level akin to bacterial NAPs. In addition, euryarchaeal histones form nucleo­protein complexes homologous to eukaryotic nucleo­somes29,34. However, although archaea use chromatin proteins that are similar to bacterial and eukaryotic prototypes, functional details can be distinct. For example, archaeal nucleosomes are smaller and more dynamic than their eukaryotic counterparts, and they seem to represent

an ancestral state of eukaryotic chromatin. Euryarchaeal nucleosomes can negatively affect the initiation and elongation stages of transcription35,45. Negative interference by promoter occlusion involves a competition for DNA binding between chromatin proteins (histones and Alba) and transcription factors, including both basal and gene-specific transcription factors. This suggests that chromatin might have a role in regulating gene expression on a global as well as a gene-specific level. Furthermore, although nucleosome positioning is directed by sequence motifs in the DNA, similarly to eukaryotic histone binding 28,29, archaea do not use an orthodox equivalent of the eukaryotic ‘histone code’ whereby histone PTMs in eukaryotic nucleosomes guide chromatin remodelling and gene regulation. Phosphorylation, acetylation and/or methylation have been reported for RNAP sub­units, transcription factors and chromatin proteins in Sulfolobus spp.15,81–85; however, any putative regulatory function of these PTMs are mostly unknown. For example, the reversible acetylation of Alba by Sir2 and Pat modulates its DNA-binding properties in vitro15, but whether this regulates gene expression in vivo remains unclear. The knowledge of which proteins are involved in modulating chromatin structure in archaeal organisms is incomplete, and the views on the interplay between chromatin proteins and transcription are only fragmentary. To push the frontiers of this field, several fundamental questions need to be addressed. Considering the variety of archaeal chromatin proteins, is there a correlation between genome size, complexity and positive or negative supercoiling on the one hand and the presence of distinct classes of architectural chromatin proteins on the other? What is the effect of reverse gyrase-mediated positive supercoiling that is prevalent in hyperthermophilic archaea on chromatin structure and dynamics? What are the underlying principles of nucleosome depositioning across the genome — is this process passive or active, and is it facilitated by factors such as the chaperone prefoldin and the acetyltransferase Elp3? In future studies, an integrated approach is highly desirable that combines in vitro analyses with in vivo studies such as whole-genome occupancy profiling of the transcription machinery, histones, NAPs and architectural transcription factors. We need to apply genetic tools to study the perturbation of chromatin by mutations introduced in all of these factors in order to establish causal connections between chromatin structure

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and ongoing transcription. Would the downregulation or deletion of specific histone variants unmask binding sites of transcription factors that would in turn activate transcription of specific subsets of genes? Would a dechromatinized genome provide less of a barrier for transcription elongation complexes and alter the genome-wide occupancy profile of RNAP? Conversely, would mutations in RNAP and transcription processivity factors such as Spt4/5, and TFS alter the chromatin landscape of the genome? To complete the description of chromatin and gene expression, we need to develop and apply in vivo proximity analyses such as chromosome conformation capture (3C) and its variants, as well as 3D visualization technologies, including super-resolution microscopy and electron cryotomography. Addressing the key issues outlined above will markedly increase our understanding of the relationship between chromatin dynamics and gene expression in all three domains of life. Eveline Peeters is at the Research Group of Microbiology, Department of Bio-engineering Sciences, Vrije Universiteit Brussel, Pleinlaan 2, B-1050 Brussels, Belgium. Rosalie P. C. Driessen and Remus T. Dame are at the Leiden Institute of Chemistry, Leiden University, Einsteinweg 55, 2333 CC Leiden, The Netherlands. Finn Werner is at the Institute for Structural and Molecular Biology, Division of Biosciences, University College London, Darwin Building, Gower Street, London WC1E 6BT, UK. E.P. and R.P.C.D. contributed equally to this work. Correspondence to R.T.D.  e‑mail: [email protected] doi:10.1038/nrmicro3467 Published online 6 May 2015 Luger, K., Mader, A. W., Richmond, R. K., Sargent, D. F. & Richmond, T. J. Crystal structure of the nucleosome core particle at 2.8 Å resolution. Nature 389, 251–260 (1997). 2. Papantonis, A. & Cook, P. R. Transcription factories: genome organization and gene regulation. Chem. Rev. 113, 8683–8705 (2013). 3. van der Valk, R. A., Vreede, J., Crémazy, F. & Dame, R. T. Genomic looping: a key principle of chromatin organization J. Mol. Microbiol. Biotechnol. 24, 344–359 (2014). 4. Merkenschlager, M. & Odom, D. T. CTCF and cohesin: linking gene regulatory elements with their targets. Cell 152, 1285–1297 (2013). 5. Bickmore, W. A. & van Steensel, B. Genome architecture: domain organization of interphase chromosomes. Cell 152, 1270–1284 (2013). 6. Woldringh, C. L. The role of co‑transcriptional translation and protein translocation (transertion) in bacterial chromosome segregation. Mol. Microbiol. 45, 17–29 (2002). 7. Bannister, A. J. & Kouzarides, T. Regulation of chromatin by histone modifications. Cell Res. 21, 381–395 (2011). 8. Vignali, M., Hassan, A. H., Neely, K. E. & Workman, J. L. ATP-dependent chromatin-remodeling complexes. Mol. Cell. Biol. 20, 1899–1910 (2000). 9. Narlikar, G. J., Sundaramoorthy, R. & Owen-Hughes, T. Mechanisms and functions of ATP-dependent chromatin-remodeling enzymes. Cell 154, 490–503 (2013). 10. Ali Azam, T., Iwata, A., Nishimura, A., Ueda, S. & Ishihama, A. Growth phase-dependent variation in protein composition of the Escherichia coli nucleoid. J. Bacteriol. 181, 6361–6370 (1999). 1.

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Acknowledgements

Research in the laboratory of R.T.D. is supported by grants from the Netherlands Organization for Scientific Research (864.08.001), High Tech Systems & Materials NanoNextNL program 8B, the FOM Foundation for Fundamental Research on Matter program ‘Crowd management: the physics of genome processing in complex environments’ and the Human Frontier Science Program (RGP0014/2014). E.P. is supported by the Research Foundation Flanders (FWO-Vlaanderen) and by the Research Council of Vrije Universiteit Brussel.

Competing interests statement

The authors declare no competing interests.

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