Escherichia coli. We find that histone proteins from the archaeon Methanothermus fervidus assemble on the E. coli chromosome in vivo and protect DNA from micrococcal nuclease digestion, allowing us to map binding footprints genome-wide. We show that higher nucleosome occupancy at promoters is associated with lower transcript levels, consistent with local repressive effects. Surprisingly, however, this sudden enforced chromatinization has only mild repercussions for growth unless cells experience topological stress. Our results suggest that histones can become established as ubiquitous chromatin proteins without interfering critically with key DNA-templated processes.Nucleosomes restrict DNA accessibility throughout eukaryotic genomes, with repercussions for replication, transcription, and other DNA-templated processes. How this globally restrictive organization emerged during evolution remains poorly understood. Here, to better understand the challenges associated with establishing globally restrictive chromatin, we express histones in a naive system that has not evolved to deal with nucleosomal structures:  All cellular systems face the dual challenge of protecting and compacting their resident genomes while making the underlying genetic information dynamically accessible. In eukaryotes, this challenge is solved, at a fundamental level, by nucleosomes,~147 bp of DNA wrapped around an octameric histone complex. Nucleosomes can act as platforms for the recruitment of transcriptional silencing factors such as heterochromatin protein 1 (HP1) in animals  and Sir Histones are not confined to eukaryotes, but are also common in archaea . They shThinking about the evolution of restrictive chromatin and its molecular underpinnings, we wondered how the presence of histones would affect a system that is normally devoid of nucleosomal structures. How would a cell that has neither dedicated nucleosome remodelers nor co-evolved sequence context cope with chromatinization? Could global chromatinization occur without fundamentally interfering with DNA-templated processes? How easy or hard is it to transition from a system without histones to one where histones are abundant? What are the key adaptations required, if any, to accommodate histones?Escherichia coli strains expressing histones from the hyperthermophilic archaeon Methanothermus fervidus (HMfA or HMfB), on which, thanks to the pioneering work of Reeve and co-workers, much of our foundational knowledge about archaeal histones is based. HMfA and HMfB are 85% identical at the amino acid level but differ with regard to their DNA binding affinity and expression across the M. fervidus growth cycle, with HMfB more prominent toward the latter stages of growth and able to provide greater DNA compaction in vitro  digestion, allowing us to map nucleosomes in E. coli in vivo. We present evidence for sequence-dependent nucleosome positioning and occupancy and consider how the presence of histones affects transcription on a genome-wide scale. Importantly, we find evidence for local repressive effects associated with histone occupancy yet only mild repercussions for growth and cell morphology, unless cells are forced to deal with excess levels of DNA damage or topological stress. Under favourable conditions, E. coli copes remarkably well with enforced chromatinization, despite evidence that histones disrupt the binding of native nucleoid-associated proteins (NAPs). Our findings have implications for how histones became established as global repressive regulators during the evolution of eukaryotes and for the evolvability of transcriptional ground states.Motivated by these questions, we built in vitro . We findE. coli K-12 MG1655 strain with plasmids carrying either hmfA or hmfB, codon-optimised for expression in E. coli and under the control of a rhamnose-inducible promoter  that are absent from Ec-EV, but present in M. fervidus and our histone-bearing E. coli strains.We carried out MNase digestion experiments using samples from late exponential and stationary phase, corresponding to 2 hr and 16–17 hr after induction, respectively . In response to a wide range of enzyme concentrations, MNase digestion of chromatin from Ec-hmfA/B  yields a ladder-like pattern of protection that is not observed in Ec-EV . Across cultures . Modal fcultures , respectcultures . HoweverE. coli genome, we find that binding is ubiquitous. On a coarse scale, coverage across the chromosome appears relatively even  previously shown to abolish DNA binding of HMfB  and then excluded genes from further analysis that were significantly up-regulated (or down-regulated) in both comparisons, reasoning that coincident patterns of change are not uniquely attributable to binding and might instead derive from systemic responses to heterologous expression. We then considered differential expression in Ec-hmfA/B versus Ec-EV for the remaining genes as a function of nucleosome occupancy.To address the first question, we generated two additional strains, Ec-hmfA of HMfB . MNase t of HMfB . Using Radj <0.05) down-regulated in histone-bearing strains display significantly higher nucleosome occupancy at TSSs than upregulated genes , oxidative stress (H2O2), DNA damage (UV), and supercoiling stress (novobiocin). To capture effects of histone occupancy during lag phase and ensure that stress responses are measured in cells where histones are established, we inoculated new cultures with cells that had already been expressing histone genes for 2 hr . When these pre-induced cells are re-inoculated, we observe a slightly prolonged lag phase  to >950 previously published differential expression profiles from a broad range of perturbations .The results above suggest that histones do not compromise dynamic responses to stress in general but that their presence is problematic when sensing or dealing with altered DNA topology or damage. To better understand the molecular basis of altered growth, we compared the transcriptome-wide signature of differential expression in Ec-hmfA , we find that correlations between expression profiles is modest (maximum ρ = 0.34), indicating that the transcriptional response to histone expression has a strong unique component. Histone-expressing strains are most similar to perturbations that are marked by transient growth arrest and induction of the stringent response . Most notably, genes previously identified as direct H-NS targets (green icons in h-ns is deleted (as one would expect), but also upon HMf expression. This might indicate that histones displace H-NS, but fail to provide similar silencing, leading to de-repression of H-NS target genes. In line with this hypothesis, we find that histone occupancy is not significantly reduced at known binding footprints of H-NS .We also find little, if any, evidence for competitive exclusion at known binding sites of other endogenous NAPs . In contE. coli are not associated with a strong growth phenotype, with the notable exception of the hupA/hupB double deletion (ΔΔHU) strain, which grows notably more slowly compared to its C600 wild-type progenitor , remains to be established. However, access regulation outside of a transcriptional context might well have provided the original evolutionary impetus for histones to spread across the genome, as genomes evolved to defend themselves against selfish elements that target the host genome for integration , previously shown to jointly abolish DNA binding of HMfB  at 37°C with agitation (170 rpm). Histone expression was induced by adding L-Rhamnose monohydrate to a final concentration of 15 mM at OD600 ~ 0.6. Cells were harvested after 2 hr or 16–17 hr following induction.HMf protein purification was performed as in Bacteria were harvested by centrifugation (4000 rpm for 15 min at 4°C), the supernatant discarded, and the pellet resuspended in a small volume of Histone Wash Buffer . Cell envelopes were disrupted using a Bioruptor Plus sonication system  for 10 cycles, 30 s on/off with power set to high. The soluble protein fraction was separated from cellular debris by centrifugation at 15,000 x g for 15 min at 4°C, while the insoluble fraction was obtained by re-suspending the pelleted debris in Histone Wash Buffer. The protein concentration in the cell lysate was quantified with a Pierce BCA Protein Assay Kit  using the provided albumin as standard. Protein fractions were separated by means of 16.5% Tris-tricine precast gels  and bands were revealed by colloidal Coomassie  staining. Histone-expressing strains showed a band close to the size expected for HMfA/B . This baOvernight pre-cultures were diluted 1:500 into LB medium plus kanamycin (50 μg/ml). Samples were plated in replicate into a flat bottom Nunc 96-well plate  and incubated at 37°C at 100 rpm for 30 min. OD measurement were performed using a high-throughput microplate reader  in which bacteria were grown at 37°C under continuous shaking . Optical density was measured at 600 nm every 30 min for 12.5 hr. For induction, the microplate reader was paused at cycle 6 and L-Rhamnose monohydrate added manually to the relevant wells to a final concentration of 15 mM. Results presented are from three biological replicates per strain, each averaged across six technical replicates.Bacterial cultures were harvested by centrifugation (4000 rpm for 15 min at 4°C), the supernatant discarded and the pelleted cells re-suspended in chilled 1x PBS . Cells were then fixed by adding a fixation solution  for 10 min at room temperature under slow rotation, after which fixation was quenched by adding 140 mM glycine. Following a further round of centrifugation (4000 rpm for 5 min at 4°C), bacteria were washed twice with 10 ml chilled 1x PBS and incubated in a lysozyme buffer  for 10 min at 37°C to generate protoplasts. Cells were pelleted (15000 rpm for 3 min at room temperature) and re-suspended in 500 μl of lysis buffer , transferred to a new microcentrifuge tube and incubated on ice for 20 min. Subsequently, the lysate was spun down and the pellet washed with 500 μl of -CA buffer  without re-suspending. The washed pellet was finally re-suspended in 500 μl of +CA buffer  to a uniform suspension. 50 μl of this suspension were digested with micrococcal nuclease  for 10 min (20 min for cells in stationary phase) at room temperature and finally blocked with a STOP solution containing calcium-chelating agents . Each sample was further diluted with -CA buffer and treated with 10% SDS and 150 ng/ml proteinase K overnight at 65°C with shaking at 500 rpm. Undigested DNA fragments were purified by two rounds of phenol:chloroform extraction separated by an RNase A digestion step . Finally, DNA fragments were precipitated in ethanol and re-suspended in 40 μl distilled water. The quality of the digest and the size of the retrieved fragments were assessed by agarose DNA electrophoresis (2.5% agarose gel in 1x TBE run at 150V for 30 min).M. fervidus harvested in late exponential and stationary phase were purchased from the Archaeenzentrum in Regensburg, Germany. We then followed the MNase protocol outlined above with the following modifications: first,~0.5 g of frozen pellet were thawed and re-suspended in 9 ml of 1x PBS before fixation. Second, due to differences in cell wall composition between M. fervidus and E. coli, the lysozyme digestion step was replaced by mechanical disruption with a French press: after the wash that follows fixation, the pellet was re-suspended in 20 ml of chilled 1x PBS, the cell suspension passaged twice through a TS Series French press (Constant Systems) at 15kpsi and then spun down at 4000 rpm for 15 min at 4°C before proceeding with cell lysis. Finally, the extracted chromatin was re-suspended in 250 μl of +CA buffer (instead of 500 μl). Digestion, fragment purification, sequencing and analysis were performed as for E.coli but with a micrococcal nuclease concentration of 100 U/ml.Frozen pellets of M. fervidus were analysed with an Agilent Bioanalyser DNA1000 chip. For each of these strains, three biological replicates were selected for sequencing. Twenty nanograms per sample were used for library construction with the NEBNext Ultra II DNA Library Prep Kit for Illumina and NEBNext Multiplex Oligos for Illumina. The output was then taken to 10 PCR cycles and purified using a 1.8x Ampure XP bead clean-up kit. Libraries were quantified via Qubit and quality assessment carried out on an Agilent Bioanalyser DNA 1000 chip. Libraries were then sequenced on an Illumina MiSeq sequencer using single-end 160 bp reads.Size distributions of the DNA fragments retrieved by MNase digestion of strains Ec-EV, Ec-hmfA, Ec-hmfB and Escherichia coli K-12 MG1655 genome (NC_000913.3) or the M. fervidus DSM2088 genome (NC_014658.1) using Bowtie2  to remove adapter sequences. This did not remove short remnant adapter sequences so that we submitted reads to a further round of trimming using Trimgalore v0.4.1 with default parameters. Trimmed reads were aligned, as appropriate, to either the  Bowtie2 . Only un Bowtie2 .Nucleosome peaks were called using the NucleR package in R as described previously . See SupE.coli genome (genomic positions 0–773608) and tested on the remainder of the genome.LASSO modeling was carried out for different footprint size classes  using empty vector-normalized coverage. Empty vector coverage was computed across fragment sizes and coverage across the genome uniformly increased by one to enable analysis of zero-coverage regions. K-mer counts  were computed using the R seqTools package over windows of three different sizes . Subsequent LASSO modeling was then carried out as described previously , with mohttp://regulondb.ccg.unam.mx/menu/download/datasets/files/High_throughput_transcription_initiation_mapping_with_5_tri_or_monophosphate_enrichment_v3.0.txt). The position inside each broad TSS associated with the most reads (column three in the file above) was defined as the TSS for downstream analysis. Promoter annotations were obtained from the same source (http://regulondb.ccg.unam.mx/menu/download/datasets/files/PromoterSet.txt).Experimentally defined transcriptional start sites were obtained from RegulonDB  , which aggregates differential transcriptional responses . Similarity between differential expression in Ec-hmfA versus Ec-hmfAnb and other pairwise comparisons . The supernatant was discarded and the pellet re-suspended in 100 μl of Y1 Buffer  and incubated at 37°C for 1 hr at 500 rpm. The cell suspension was added to 350 μl of RLT buffer, 250 μl 100% ethanol and loaded onto an RNeasy column from the RNeasy Kit . RNA was then washed and eluted following the manufacturer’s protocol. Eluted samples were incubated with DNase I  for 10 min at 37°C and then cleaned up with a second passage through the RNeasy column . Samples were finally eluted in 30 μl of RNase-free water and RNA quantified with Nanodrop. Quality assessment of the extracted RNA was carried out with an Agilent Bioanalyser RNAnano chip and five replicates per strain/condition were chosen for sequencing.For each replicate/strain/condition, 1.5 μg of total RNA were depleted of rRNA using the Ribo-Zero rRNA depletion kit (Illumina) and libraries constructed using a TrueSeq Stranded RNA LT Kit (Illumina). After 12 PCR cycles, library quality was assessed with an Agilent Bioanalyser HS-DNA chip and quantified by Qubit. No size selection was carried out and the samples were sequenced on a HiSeq 2500 machine using paired-end 100 bp reads.E. coli K-12 MG1655 genome (NC_000913.3) with Bowtie2 (--no-discordant --no-mixed). As a technical aside, we note that, despite the above filtering steps, some of the samples had an unusually low alignment rate (<30%). We found that most of the unaligned reads were perfect matches to rRNA sequences from Bacillus subtilis but not E. coli and had therefore eluded the above filter. As contamination at this scale is unlikely (no bacteria other than E. coli are grown or sequenced in the lab and a plain LB control was added to check for contamination when growing the samples for RNA extraction), we suspect these reads are the result of carrying over RiboZero oligos. The addition of a further round of filtering to discard reads that match non-coding RNA sequences from Bacillus subtilis increased the alignment rate to E. coli index up to ~90%.Using Bowtie2, reads were first aligned to all annotated non-coding RNA genes . Reads that mapped to any of these genes were discarded, even if they mapped to more than one location in the genome. We then used Trim Galore v0.4.1 with default parameters to trim adapters and low-quality terminal sequences. Trimmed reads were aligned to the By-gene read counts were computed from read alignments using the summarizeOverlaps function  from the GenomicAlignments package in BioConductor. Differential gene expression analysis was carried out using DESeq2 . Replicahttps://sils.fnwi.uva.nl/bcb/objectj/examples/Coli-Inspector/Coli-Inspector-MD/coli-inspector.html).Overnight pre-cultures of Ec-EV, Ec-hmfA, and Ec-hmfB were diluted in fresh LB medium plus antibiotic and grown as described above. ~300 μl of culture were harvested by centrifugation . Pellets were resuspended in 1% FA in PBS and fixed for 10 min at room temperature. Fixating agent was removed by spinning  and pellets were resuspended in 1 ml PBS. 5 μl of cellular suspension was spread onto an agarose pad, covered in VectaShield containing DAPI (Vector Laboratories) and the excess liquid removed. Slides were imaged using a Manual Leica DMRB with phase contrast and DIC for transmitted light illumination. For quantification, images from three independent experiments were analysed with MicrobeJ  to perfoGenomic regions bound by Fis and H-NS were obtained from The HMfB dodecamer unit cell was reconstructed from the crystal structure (PDB ID: 5t5k). Dimers were removed sequentially from the structure to build different sized HMfB polymers. We used the AMBER ff14SB forcefield  and solv2O2 were used at respective concentrations of 10 µg/mL, 0.31 ng/mL, and 1.25 mM. For UV stress, cells used as inoculum were grown as for chemical stress assay and 106 and 103 dilutions (in 1x PBS) were plated onto plates containing 15 mM rhamnose and kanamycin. Plates were irradiated with 10000µJ using Stratalinker 2400.To minimise the risk of suppressor mutations all cultures involving NAP deletions were inoculated straight from −80°C stocks. 5 mL LB cultures containing the antibiotic necessary for plasmid selection were inoculated in 50 mL tubes at OD = 0.1 from overnight LB cultures that contained antibiotics for both plasmid and mutant selection. After 2 hr, rhamnose was added to a final concentration of 15 mM to induce histone or YFP expression. Two hours after induction, 10 µL of cultures were used to inoculate 200 µL wells in Nunc 96-well microplates . Growth was monitored as described above. For chemical stress assays, novobiocin, rifampicin and Hhttps://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE127680).Datasets generated for this study have been deposited in the NCBI Gene Expression Omnibus under accession number GSE127680  the histone proteins assemble genome-wide into multimeric structures of varying sizes according to known intrinsic sequence preferences; (2) nucleosome displacement during transcription and replication; (3) local transcriptional repression due to nucleosome binding at promoters and gene bodies; (4) modest to inconclusive global effects on transcription; and (5) surprisingly, no major repercussions to cell growth.Rojec and colleagues present a simple but rather provocative experiment: would expressing archaeal histones from E. coli genome with M. fervidus histones. The experiments have been planned with great care. A nice and novel observation is that the size of nucleosomes bound at transcription start sites correlates with repression efficiency. The paper will be intriguing for different subfields because it suggests that early histone-fold like proteins may not have evolved solely to regulate transcription, although the other posited roles remain hypothetical. Nevertheless, the amount of novel information generated in relation to archaeal histone function is limited: the reported results largely confirm earlier observations. We ask that the authors add more discussion as to whether there are additional biological insights about hmfA/B. Furthermore, the manuscript poses questions in the Introduction, which are perhaps, eukaryotic-centric. We ask that the authors reframe these questions as current work on archaeal histones suggests many similarities between bacteria and archaeal DNA organizing proteins. Finally, there was consensus that the authors will need to further justify their assertion that the effect to E. coli "…is remarkably unremarkable" and provide more analyses on the effects to transcription.The manuscript is well-written and a pleasure to read. It reports a large body of convincing data on the neochromatinization of the Essential revisions:E. coli [with archaeal histones].… is remarkably unremarkable". Rich medium obviates the requirement for many E. coli genes for survival. Further, exponential growth has a relatively "fixed" transcriptome. In fact, the lagging phase  are likely where the nucleosomes have the largest effect. We would like to see a sampling of different types of stress: transcription , DNA damage , metabolic . This can be done in a reasonable length of time. A more detailed analysis of gene expression, including the data they excluded, may offer clues as to the type of defect, if any, the histone expressing cells may be experiencing. Perhaps they could even include a cell cycle analysis.1) All reviewers agreed that the exponential growth experiments in rich medium (LB) were inadequate to support the statement that "growth of 2) It is rather striking that the hmfA/B expressing strains have both larger cell size and greater size heterogeneity. This was only briefly mentioned in the Results but is rather striking. The fact that the SOS response is activated suggests that the histones are affecting more processes than suggested. Would growing the cells in conditions that cause DNA stress (UV for example) exacerbate the morphology? Are the cells overall bigger or just narrower and longer ? Could this be due to increased volume of chromatinized DNA? We would be satisfied with a more thorough discussion on this topic but would also like to see whether HfmA/B/nb expressing cells display additional morphological changes. It would also be useful if the authors could present additional information, if available, on cell-to-cell variability in terms of expression of the histone proteins. This might aid in making their point that hmfA/B have little impact on cell physiology.E. coli to adapt to histones regardless of whether they are bound to DNA? Finally, the correlation between GC content vs. histone occupancy is not formally quantified (only browser tracks are shown). What is the relationship genome-wide? Is it significant? Here we are only asking for additional analyses of the existing data.3) We generally found that the "locally repressive effects" was not robust and needed further exploration. The authors could partially address this by explicitly showing, examining, and categorizing by function the genes that are down- or up-regulated in hmfA/B/nb vs. EV, and determine the corresponding level of nucleosome signal in hmfA/B expressing cells. They reasonably exclude these as potentially generic responses to ectopic protein expression. But is that the case? Do these genes change expression in response to any ectopically expressed proteins or are there residual hmfAnb effects? The idea is to deconvolute whether histones incorporate more readily with already downregulated genes or cause their repression. Could these include genes that enable E. coli lacking one or more of its own DNA organizing proteins? We would like to see some straightforward estimates (using RT-qPCR) of a few stress-regulated genes under direct control of bacterial NAPs when grown under stress. We also ask whether there is phenotypic complementation of H-NS and HU knockout strains. In the case of HU this would make clear whether the similar structural properties of the two proteins are enough to have complementation. In the case of H-NS it could suggest that invasion of hmfA/B into large H-NS regulated regions indeed has only mild effects on global transcription. Such growth experiments carried out under a few conditions (see 1) should be fairly straightforward. What is the difference between AT-rich non-histone binding regions and AT-rich H-NS binding regions? What is the effect of histones in gene bodies; is some of the repression of spurious intragenic transcription by H-NS taken over by bound archaeal histones?4) We ask for additional analyses on bacterial organizational structure in the context of AT/GC compared to endogenous bacterial NAPs. What is conceptually the difference between over-expressing a bacterial DNA organizing protein, one from another bacterium, or an archaeal histone? Why do we not see a phenotype similar to that of 5) One of the most interesting questions put forward by the authors is 'how easy or hard is it to transition from a system without histones to one where histones are abundant?', but the answers to this question leaves something to be desired. Although the authors use an inducible system (2 h and 16 h), perhaps only subpopulations of the cells accommodate histones? We think it is important to rule out potential adaptive mechanisms especially by looking more deeply into the gene expression changes. It is quite possible that this occurs at the level of single-cells, which is beyond the scope of this study, but perhaps explains some of the variability. We again suggest simple experiments such as determining whether the SOS response is required to tolerate expression of histones or their binding to DNA. […] The amount of novel information generated in relation to archaeal histone function is limited: the reported results largely confirm earlier observations. We ask that the authors add more discussion as to whether there are additional biological insights about hmfA/B.M. fervidus and other archaea operate in their native cellular context. Even though not the focal point of the manuscript, it is worth highlighting two such leads:We share the reviewers’ desire to learn more about the native functions of archaeal histones. Given that we express archaeal histones in a heterologous system, we have been cautious in drawing firm conclusions about native functionality. That said, we do think that our experiments have revealed interesting leads that might help us better understand how the histones of E. coli in Figure 1C) or by varying the relative levels of different histone paralogs that are more or less likely to form stable longer oligomers.First, as highlighted by the editor, we provide some evidence – yet to be reported for any archaeon – that longer oligomers elicit stronger repression. We now also provide evidence from molecular dynamics simulations that longer oligomeric complexes are more stable . It is certainly conceivable that archaea make use of this property to dynamically regulate genome accessibility, either by changing absolute histone concentrations . There, we suggest that oligomer formation is subject to sequence-dependent nucleation-extension dynamics. We show that initial tetramer formation relies on a relatively high affinity site but that, from this nucleation point, oligomers can reach into less conducive sequence space. Importantly, this might enable repression of promoters that – by virtue of their sequence composition – disfavour nucleation. We anticipate that our heterologous Regarding insights into HMfA versus HmfB in particular  we note that differences between the two paralogs clearly do exist, as evident from MNase profiles , the transcriptomic response  and prior work by John Reeve’s group. However, these differences are comparatively subtle. As this issue is not central to the manuscript, we decided not to focus on these differences further. However, our data will allow a more focused exploration of this in the future.Furthermore, the manuscript poses questions in the Introduction, which are perhaps, eukaryotic-centric. We ask that the authors reframe these questions as current work on archaeal histones suggests many similarities between bacteria and archaeal DNA organizing proteins.We agree that there are important similarities between bacterial NAPs and archaeal histones. In fact, we stated this forcefully in the Discussion.“… care should be taken in projecting properties of eukaryotic histones onto those of archaea. In many instances, archaeal histones might be better understood with reference to bacterial NAPs, especially when considering how concentration drives opportunities for oligomerization, cooperativity, and bridging interactions with DNA.”It is therefore not because we are unaware of these similarities that we pose eukaryotic-centric questions in the Introduction. Rather, the framing is a considered choice. We believe that most readers will approach this manuscript from a eukaryote-based knowledge of histones and histone function. We aim to pick them up from their place of relative comfort and journey towards a better understanding of archaeal histones, ending up with a more nuanced understanding of the nature of archaeal histones – functionally somewhere between eukaryotic histones and bacterial nucleoid-associated proteins.Finally, there was consensus that the authors will need to further justify their assertion that the effect to E. coli "…is remarkably unremarkable" and provide more analyses on the effects to transcription.We appreciate that what is and is not remarkable is very much in the eye of the beholder. We have therefore characterized growth in a number of different stress conditions, as detailed below (point #5). We have also added a more rigorous comparative analysis of global and local transcriptional effects. To reflect that our astonishment is necessarily subjective we have rephrased the relevant sentence (subsection “Histone binding is associated with mild phenotypic effects under favourable conditions”). Whether the reviewers and readers share our astonishment is, of course, entirely up to them.Essential revisions:1) All reviewers agreed that the exponential growth experiments in rich medium (LB) were inadequate to support the statement that "growth of E. coli [with archaeal histones].… is remarkably unremarkable". Rich medium obviates the requirement for many E. coli genes for survival. Further, exponential growth has a relatively "fixed" transcriptome. In fact, the lagging phase  are likely where the nucleosomes have the largest effect. We would like to see a sampling of different types of stress: transcription , DNA damage , metabolic . This can be done in a reasonable length of time.2O2 (oxidative stress), UV (DNA damage), and novobiocin treatment (supercoiling stress). We added the latter condition to explore whether histones, known to constrain negative supercoils, exacerbate the effects of gyrase inhibition. To stress-test the cells, we slightly modified the growth protocol, inducing histone expression in exponentially growing cells with rhamnose, growing these cells for a further two hours in rhamnose, and then using them to inoculate a new culture that contains rhamnose as well as the drug of interest (rifampicin etc). In this way, we ensure that stress responses are measured in cells where histones are already established, and that we can adequately capture effects of histone occupancy during lag phase. We then tracked growth/survival of these strains by measuring optical density or, in the case of UV irradiation, colony formation. The results of these experiments are displayed in Figure 6.We share the reviewers’ curiosity to see how our histone-expressing strains respond to various stresses. In the revised manuscript we now present data on growth in response to rifampicin , HBriefly, the salient insights are as follows:- When pre-induced cells are transferred into new LB medium, there is a slightly prolonged lag phase. However, histone-expressing strains recover quickly and eventually catch up with non-binding/EV control strains.2O2. Again, histone-expressing strains recover well. Under these conditions, histones can therefore be said to have a mild bacteriostatic but no bacteriocidal effect.- Lag phase is extended further in strains treated with rifampicin or H- In contrast, the presence of histones matters in the face of UV and novobiocin treatment, where colony formation and growth, respectively, are severely affected. In novobiocin-treated cells, we observe strong morphological changes, as cells become conspicuously elongated . The presence of histones therefore seems to interfere with the cells’ ability to effectively deal with double strand breaks (UV) and altered supercoiling levels.E. coli”.Based on these results, which are complemented by new insights from transcriptomic analysis (see below), it would be reasonable to surmise that histones do not compromise dynamic responses to stress in general. However, their presence is problematic where the stress specifically relates to sensing or dealing with altered DNA topology or damage. Given what we know about histones, this is not unexpected. We present and discuss these results in the subsection “Systemic transcriptional responses to histone expression in A more detailed analysis of gene expression, including the data they excluded, may offer clues as to the type of defect, if any, the histone expressing cells may be experiencing. Perhaps they could even include a cell cycle analysis.). This allows us to place the effect of histone expression into a comparative framework and, by virtue of similarity to other conditions, suggest molecular pathways through which histones affect E. coli physiology.We have carried out substantial further analysis of our gene expression data, including uncensored data, and now provide a more in-depth comparison of our strains to other perturbations that had previously been assessed via global transcriptome profiling (https://genexpdb.okstate.edu/nb) and previous experiments that involved differential gene expression analysis.Figure 7A summarizes the key results of this analysis. It displays the similarity in transcriptional responses between histone expression (Ec-hmfA versus Ec-hmfAnb) to other experiments, several interesting patterns emerge:Comparing differential expression in exponential phase Ec-hmfA . Genes in the rpoS regulon are also upregulated in histone-expressing strains  while flagellar genes are downregulated, a response our strains share with the stringent response.- In terms of genes that are up-regulated, histone-expressing strains are similar to conditions associated with environments where carbon sources are scarce/exhausted  or altered (glucose-to-lactose shift).These transcriptional signatures of transient growth arrest and/or slowed growth are in line with the growth phenotypes we observe, which are characterized by an extended lag period . This suggests that histone expression slows initial growth until cells have had time to adjust. Such adaptive adjustments almost certainly involve the downregulation of gyrases, as previously noted. Histones wrap DNA in negatively constrained supercoils. Gyrases introduce negative (or relax positive) supercoils so counteracting the histone-associated built-up of negative supercoiling by downregulating gyrases makes sense. Downregulation of gyrases then likely renders cells more susceptible to novobiocin . This model is further consistent with the observation that expressing CcdB, a gyrase poison, leads to gene expression changes that have some similarity to histone expression .h-ns.We were further intrigued to see that, with regard to upregulated genes, the effect of histone expression is similar to the effects of deleting Finally, and importantly, we note that “similarity in expression” here is strictly relative. The strongest correlation to a dataset outside this study is modest (ρ=0.34). Thus, the transcriptional response to histone expression by no means precisely mimics any of the perturbations discussed above but has a strong unique component.2) It is rather striking that the hmfA/B expressing strains have both larger cell size and greater size heterogeneity. This was only briefly mentioned in the Results but is rather striking. The fact that the SOS response is activated suggests that the histones are affecting more processes than suggested. Would growing the cells in conditions that cause DNA stress (UV for example) exacerbate the morphology?We have considered how stress might exacerbate the impact of histone expression on cell morphology in the context of novobiocin treatment. Gyrase inhibition does indeed exacerbate morphology in histone-expressing cells, which become extremely elongated .Are the cells overall bigger or just narrower and longer ? Could this be due to increased volume of chromatinized DNA? We would be satisfied with a more thorough discussion on this topic but would also like to see whether HfmA/B/nb expressing cells display additional morphological changes.E. coli cells. As a potential starting point for future more in-depth analysis, we therefore decided to carry out a quantitative analysis of the relationship between nucleoid and cell volume based on DAPI-stained cells. As evident from Figure 5—figure supplement 1, we find no obvious changes to the nucleoid/cytoplasm ratio in histone-expressing cells and therefore no evidence for either compaction or decompaction. Ec-hmfA/B have a somewhat larger relative nucleoid volume than Ec-EV, but are indistinguishable from WT, suggesting that this is an Ec-EV-specific effect.We agree that it is interesting to explore changes to nucleoid morphology and nucleoid/cytoplasm ratio in histone-expressing It would also be useful if the authors could present additional information, if available, on cell-to-cell variability in terms of expression of the histone proteins. This might aid in making their point that hmfA/B have little impact on cell physiology.We initially considered tagging the histones with a fluorescent reporter, which would have allowed direct quantification. However, prior attempts at tagging by others were abandoned as even small tags were found to affect DNA binding . Tagging is much easier in eukaryotic histones, where the tag can be attached to the long flexible tails that are absent from archaeal histones. As an imperfect alternative, we sought to approximate heterogeneity by monitoring cell-to-cell variability of YFP expressed from the same promoter, either with or without HMfA simultaneously expressed (from a different plasmid but the same promoter) in the same cell. What we can conclude from 3) We generally found that the "locally repressive effects" was not robust and needed further exploration. The authors could partially address this by explicitly showing, examining, and categorizing by function the genes that are down- or up-regulated in hmfA/B/nb vs. EV, and determine the corresponding level of nucleosome signal in hmfA/B expressing cells. They reasonably exclude these as potentially generic responses to ectopic protein expression. But is that the case? Do these genes change expression in response to any ectopically expressed proteins or are there residual hmfAnb effects? The idea is to deconvolute whether histones incorporate more readily with already downregulated genes or cause their repression.We think the reviewers make several distinct points here. Below, we have tried to disaggregate these points as we understand them.a) The reviewers would like to see additional information to support the finding of locally repressive effects. They reasonably suggest that an alternative explanation might be true, namely that cause and effect are reversed, i.e. increased histone occupancy follows (rather than elicits) downregulation. There are two key pieces of evidence that strongly argue against this interpretation. First, as now highlighted in Figure 3F, unlike sometimes observed in eukaryotes, there is no negative global relationship between mRNA levels (as measured in Ec-EV to approximate the pre-histone state) and histone occupancy in our system. In fact, the relationship is slightly positive . Thus, lower expression per se is not associated with higher histone occupancy. Second, we showed previously  that the relationship between histone occupancy and expression, while present at the promoter, breaks down when considering a control window just downstream of the promoter. This is inconsistent with the alternative model where downregulation precedes histone deposition. If histones simply accumulated on the DNA because transcription was reduced or switched off, the same relief from transcriptional perturbation should be observed immediately downstream of the promoter. It is not.and the non-binding strain necessarily means that downregulation in the binding strain is really unrelated to DNA-binding. This is a good question. To begin with, we cannot and do not claim that coincident downregulation in both the binding and non-binding strain implies that downregulation in the binding strain is necessarily unrelated to binding. It is perfectly possible, for example, that histones exert a mild repressive effect at gene X and the same gene X in the non-binding histone strain also experiences downregulation, but for different reasons. We chose to focus on genes that are up- or down-regulated only in strains with a DNA-binding histones to minimise type I errors. Our overriding concern here is to isolate the effects of DNA-binding on gene expression. Naturally, in doing so we almost inevitably decrease sensitivity and miss genes at which histones exert a genuine effect via DNA binding and nucleosome formation. In the context of obtaining robust results on the direct effects of nucleosome formation, we believe this is a price worth paying.b) The reviewers also wonder, if we understand correctly, whether finding downregulation in the binding strain nb. However, we have shown experimentally that the non-binding mutants do not form nucleosomes and therefore would have to associate with DNA in a qualitatively distinct manner. Does that mean that overexpression of non-binding histones has impacts that are distinct from expressing other proteins? Yes. Is this a good thing? We would argue that it is, since it provides a more tailored control for other features of histones  that would not be controlled for by expressing a more inert gratuitous protein, and it is this well-matched control that allows us to home in on the specific effects of nucleosome formation along the E. coli chromosome.c) Finally, it is possible, that non-binding mutant histones still associate with DNA. The unexpectedly poor growth upon novobiocin treatment also hints at that possibility for HmfAWe struggled to follow the reviewers’ comment that categorizing genes by function would shed light on the issues above. There is no a priori reason to suspect that direct, locally mediated repressive effects of histone occupancy are divided along functional lines . One would expect functional structure for transcriptional responses launched to cope with the stress induced by histones (in terms of both heterologous expression and stress associated with binding), but not in terms of the initial targets of local repression. NB: we did carry out a classic gene set enrichment analysis (GSEA), which showed depletion of genes in COG M (caused by flagellar genes) and COG J (translation), very much in line with the findings from comparative transcriptomic analysis. We did not present those results for two reasons: first, we think the comparative transcriptomic analysis is more insightful. Second, we think that toolkit to carry out GSEA in prokaryotes is fundamentally incomplete. Tools inherited from eukaryotes do not take into account operons. This is problematic because genes cluster into operons by function. As a consequence, changes to the regulation of a single operon get amplified by the number of genes of the same function that operon contains. In light of this problem, it is typically unclear what statistical enrichments in prokaryotic GSEA actually mean.Could these include genes that enable E. coli to adapt to histones regardless of whether they are bound to DNA?Yes, absolutely, although it is arguably harder to pinpoint specific genes that react in this manner . Our experiments were not designed to address this specifically.Finally, the correlation between GC content vs. histone occupancy is not formally quantified (only browser tracks are shown). What is the relationship genome-wide? Is it significant? Here we are only asking for additional analyses of the existing data.-16).We already provided a quantitative assessment of the relationship between GC content and histone occupancy in Figure 3, in particular 3D. There are strong correlations in stationary phase between GC content and histone occupancy  We ask for additional analyses on bacterial organizational structure in the context of AT/GC compared to endogenous bacterial NAPs. What is conceptually the difference between over-expressing a bacterial DNA organizing protein, one from another bacterium, or an archaeal histone? Why do we not see a phenotype similar to that of E. coli lacking one or more of its own DNA organizing proteins? We would like to see some straightforward estimates (using RT-qPCR) of a few stress-regulated genes under direct control of bacterial NAPs when grown under stress. We also ask whether there is phenotypic complementation of H-NS and HU knockout strains. In the case of HU this would make clear whether the similar structural properties of the two proteins are enough to have complementation. In the case of H-NS it could suggest that invasion of hmfA/B into large H-NS regulated regions indeed has only mild effects on global transcription. Such growth experiments carried out under a few conditions (see 1) should be fairly straightforward. What is the difference between AT-rich non-histone binding regions and AT-rich H-NS binding regions? What is the effect of histones in gene bodies; is some of the repression of spurious intragenic transcription by H-NS taken over by bound archaeal histones?We agree with the reviewers that it would be interesting to understand in greater molecular detail how histone expression impacts nucleoid organization, how histones interact with endogenous NAPs, and whether histone expression poses a challenge distinct from overexpressing one of these NAPs.h-ns deletion strain . As direct targets are arguably better characterized for H-NS than any other NAP, we focused our investigation on H-NS. We began by comparing gene expression changes in our strain to those in a previously published h-ns deletion strain . Perhaps most notably, genes previously identified as direct H-NS targets , are amongst the most upregulated genes not only when h-ns is deleted (as one would expect), but also upon HMf expression. This might indicate, as suggested by the reviewers, that histones displace H-NS, but fail to provide similar silencing, leading to de-repression of H-NS target genes. This hypothesis is consistent with our previous analysis  showing that histones are not significantly excluded from previously determined H-NS binding sites, implying that they successfully compete for binding at those sites.We now present several new lines of investigation that shed light on these questions. We identified intriguing similarities in the global transcriptional response between hmfA/B-expressing strains and an h-ns deletion and the corresponding wildtype strain. In other words, we cannot detect what are considered typical effects of H-NS removal in this dataset. This might be because spurious transcripts are short and/or unstable and therefore not well represented in data from a typical RNA-Seq workflow . Irrespective of the underlying reason, this precludes an informed comparison of spurious transcription between this and our data. As the focus of this manuscript is not on H-NS, we did not pursue this further experimentally.We also attempted to look at spurious (intragenic and intergenic) transcription in this dataset. However, we found that the proportions of reads mapping to intergenes or antisense were effectively indistinguishable in the h-ns, we found no similarities of note when comparing our data to other NAP deletion mutants . As far as we are aware, transcriptome-wide data on the effects of NAP overexpression (vis-à-vis deletion) in a comparable genetic background are not available. They would also be intrinsically harder to compare and interpret given that a) effects will inexorably depend on the level to which the protein is overexpressed and b) phenotypes reflect expression burden as well as protein-specific effects unless suitable controls are in place. However, it is interesting to note that strong (>40-fold) overproduction of H-NS has previously been reported to lead to a transient  growth arrest after which cells resume growth . This situation, which the authors dubbed “artificial stationary phase”, is qualitatively reminiscent of the prolonged lag phase we observe in the presence of HMf (see point #5) and further underlines the transcriptional similarity between histone-expressing strains and E. coli growing in stationary-phase(-like) conditions. Why would histone expression resemble both h-ns deletion and h-ns overexpression? We suggest, based on the chain of events proposed above, that histone displacement of H-NS not only leads to dysregulation of target genes, but – like overexpression – also generates a pool of free H-NS that might go on to bind off-targets (including perhaps AT-rich promoters) or have toxic effects unrelated to DNA binding.In contrast to ΔIn contrast to H-NS, HU overexpression from the same promoter led to 1.8-fold slower growth but no growth arrest, again underlining the outlier status of H-NS.Our results, both old and new, indicate that histones likely invade genomic real estate normally occupied by NAPs. Histones also share some key properties with NAPs. Notably, they resemble HU in their ability to constrain negative supercoils. We therefore agree that it is interesting to ask if histones complement NAP deletion mutants. To address this question, we have assayed effects of HMfA expression on growth in a collection of NAP deletion strains. The results are summarized in Figure 8, considering differences in the time to maximum growth rate  and doubling time at the time of maximum growth as determined by curve fitting . We note the following:E. coli are not associated with a strong growth phenotypes, with two exceptions:- NAP deletions in fis deletion (Δfis) is associated with a reduced lag phase;hupA/hupB deletion (ΔΔHU) strain grows more slowly (increased doubling time) compared to its C600 WT progenitor strain.the fis (and – for unknown reasons – in M182 WT)- HMfA expression generally leads to an increase in lag phase duration, which is particularly pronounced in ΔhupA/hupB deletion and HMfA expression are not additive, suggesting that histone expression might partially alleviate defects associated with the absence of HU.- HMfA expression is associated with a relatively small but consistent increase in doubling time, but this effect does not appear to be compounded in NAP deletion strains. Interestingly, the growth retardation associated with Note that we use YFP here for historical reasons – we had already carried out some of these assays early on in the project, prior to constructing the non-binding histone control strains. YFP serves as a control for the burden imposed by gratuitous expression.E. coli, prefer binding to AT-rich DNA. This might be because they have an ancestral or extant function in the silencing of genomic invaders, which – for reasons that remain poorly understood – tend to have a GC content lower than the host genome. Histones, in contrast, prefer the GC-richer parts of the genome. This has some interesting implications. For example, as we suggest in the manuscript, histones might naturally have a lesser impact on sequences like promoters that are naturally AT-rich, reducing their impact on transcription. These divergent sequence preferences might also imply that, at low dosage, histones and NAPs such as H-NS might come to occupy different parts of the genome. If dosage is sufficiently high, however, histones, having saturated their preferred binding sites, will start to compete with NAPs for their preferred targets. Of course, these differences are not dogmatic. For example, we recently showed that a HU ortholog in Thermoplasma acidophilum prefers GC-rich sequences and behaves in many respects more like an archaeal histone than a “classic” bacterial HU . Arguably the biggest difference, however, is evolutionary context: the absence of molecular machinery that has co-evolved with histone proteins.Finally, we want to comment more generally on whether there are  “conceptual differences” between over-expressing a histone or an endogenous or heterologous bacterial NAP. We believe there are differences, but we would not call them conceptual. Rather, differences are grounded in the specific properties of histones vis-à-vis DNA binding proteins found in bacteria as well as in evolutionary context. What are these differences? To begin with, histones are arguably unusual in the way they wrap DNA and in the fact that they bind DNA strongly enough to provide protection from MNase. One might therefore suspect that they provide a stronger barrier than many other NAPs to the molecular machines that need to access DNA. In addition, many well-characterized bacterial NAPs, certainly those in 5) One of the most interesting questions put forward by the authors is 'how easy or hard is it to transition from a system without histones to one where histones are abundant?', but the answers to this question leaves something to be desired. Although the authors use an inducible system (2 h and 16 h), perhaps only subpopulations of the cells accommodate histones?The observation that, under standard conditions, histone expression has a mild bacteriostatic but no bacteriocidal effect argues strongly against the possibility that a significant fraction of cells drop out of the population because they cannot accommodate histones. That does not mean, of course, that there is no variability in histone expression and that some cells have an easier time than others, but note that across the population we observe histone:DNA ratios of up to ~0.7:1, so cells with a reduced burden are offset against those where the ratio exceeds 0.7:1.We think it is important to rule out potential adaptive mechanisms especially by looking more deeply into the gene expression changes. It is quite possible that this occurs at the level of single-cells, which is beyond the scope of this study, but perhaps explains some of the variability. We again suggest simple experiments such as determining whether the SOS response is required to tolerate expression of histones or their binding to DNA.E. coli cells to adapt to histone expression, we assayed growth in a recA deletion strain expressing histones. Histone expression leads to a significant reduction in growth compared to the control but is by no means lethal . This suggests that the SOS response facilitates adaptation to some aspect of the presence of histones, but is not critical for survival. Consistent with the SOS response being triggered following histone expression, we also show concerted upregulation of the rpoS regulon in our strains (see above).To understand the role of the SOS response in allowing E. coli does not respond to histone expression in an adaptive manner, i.e. by activating or repressing genes that help it cope with the presence of histones. In fact, we previously highlighted downregulation of gyrases as a likely adaptive response. The key, from an evolutionary point of view, is that E. coli copes through activation of generic stress responses rather than through a mechanism specifically evolved to allow the accommodation of histones. The cells survive without machinery dedicated to dealing with histones or adaptations to the transcriptional or replicative apparatus.On a more general note, we did not and do not argue that


Posttranslational modification of histone tails such as histone 3 lysine 27 acetylation (H3K27ac) is tightly coupled to epigenetic regulation of gene expression. To explore whether this is involved in placenta pathology, we probed genome-wide H3K27ac occupancy by chromatin immunoprecipitation sequencing (ChIP-seq) in healthy placentas and placentas from pathological pregnancies with fetal growth restriction (FGR). Furthermore, we related specific acetylation profiles of FGR placentas to gene expression changes.Analysis of H3K27ac occupancy in FGR compared to healthy placentas showed 970 differentially acetylated regions distributed throughout the genome. Principal component analysis and hierarchical clustering revealed complete segregation of the FGR and control group. Next, we identified 569 upregulated genes and 521 downregulated genes in FGR placentas by RNA sequencing. Differential gene transcription largely corresponded to expected direction based on H3K27ac status. Pathway analysis on upregulated transcripts originating from hyperacetylated sites revealed genes related to the HIF-1-alpha transcription factor network and several other genes with known involvement in placental pathology . Downregulated transcripts in the vicinity of hypoacetylated sites were related to the immune system and growth hormone receptor signaling. Additionally, we found enrichment of 141 transcription factor binding motifs within differentially acetylated regions. Of the corresponding transcription factors, four were upregulated, SP1, ARNT2, HEY2, and VDR, and two downregulated, FOSL and NR4A1.We demonstrate a key role for genome-wide alterations in H3K27ac in FGR placentas corresponding with changes in transcription profiles of regions relevant to placental function. Future studies on the role of H3K27ac in FGR and placental-fetal development may help to identify novel targets for therapy of this currently incurable disease.The online version of this article (10.1186/s13148-018-0508-x) contains supplementary material, which is available to authorized users. The dynamics of histone 3 lysine 27 acetylation (H3K27ac) in DNA regulatory regions is one of the components playing a fundamental role in the precise timing and level of gene transcription , 2. ConsFGR, a condition in which the fetus is unable to achieve its full growth potential through inadequate supply of nutrients and growth factors, occurs in approximately 5% of pregnancies  and progPrevious studies of placentas of FGR pregnancies have reported differences in gene transcription across many regions across the genome. Pathways associated with altered placental gene expression in FGR include angiogenesis, immune modulation, energy production, and growth signaling –15. MostIn this study, we mapped differential H3K27ac profiles in DNA regulatory regions in relation to disrupted development of the placenta, by exploring the presence of H3K27ac using chromatin immunoprecipitation sequencing (ChIP-seq) in FGR and control placentas. Next, we performed RNA-seq to examine whether differences in H3K27ac also reflects gene expression levels. With this approach, we identified previously unstudied alterations in promoter and enhancer activity related to placenta pathology.n = 4) and FGR pregnancies (n = 5) and identified 30,288 H3K27ac peaks that were present in at least two independent samples. Of these, 970 regions showed differential H3K27ac levels in FGR compared to controls with 366 being hyperacetylated and 604 regions being hypoacetylated in FGR . We annotated 368 genes corresponding to hyperacytelated sites and 313 genes associated with hypoacetylated sites  using AME . We deteIn this study, we used ChIP-seq and RNA-seq to perform an in-depth analysis of DNA regions with differential chromatin activity in healthy human placentas and pathological placentas of pregnancies affected by FGR. With this combined approach, we were able to identify H3K27 acetylation as a key additional layer involved in epigenetic regulation of gene expression in placental function which in turn might be related to impaired fetal growth.In summary, our findings confirm that, in FGR, the placenta exhibits substantial genome-wide alterations in H3K27 acetylation, with corresponding changes in transcription profiles in several regions pertinent to placental development and presumably function. The identified acetylation patterns show a clear distinction between FGR and healthy placentas, and the genes annotated to differentially acetylated regions are involved in pathways known to be affected in FGR . We confTo date, we are not aware of any previous reports on histone modification in FGR placentas. Other groups have focused attention on epigenetic regulation of placentas in FGR involving DNA methylation. Genome-wide methylation studies in placenta of FGR suggest involvement of pathways associated with lipid metabolism, transcription, and cadherin and Wnt signaling –22. OtheBarker hypothesis), i.e., the concept that exposure to intrauterine conditions may have long-lasting effects on fetal development, these epigenetic alterations might well be the result of changes in the intrauterine environment associated with FGR [Mechanisms involved in differential acetylation of histones within the human placenta have not been studied. In view of the “fetal origins of adult disease” hypothesis  or repressed chromatin (H3K27me3) . MoreoveWe demonstrate involvement of H3K27ac in key regions related to the placental function and placental pathology associating with impaired fetal growth. Our findings underscore that an approach using combined CHIP-seq and RNA-seq analyses facilitates discovery of novel genes involved in FGR, and identification of molecular pathways and processes associated with the placental function and early (disrupted) human growth and development. Future studies on unraveling the role of H3K27ac in FGR pregnancies and placental-fetal development may help to identify novel targets for therapy of this currently incurable disease.Placenta biopsies were collected immediately after birth from women with a FGR pregnancy and women with uncomplicated pregnancy. Each placenta was sampled at four random locations. For this study, we only included women undergoing a primary Cesarean section. We defined FGR as an estimated fetal weight <p3 and only included cases with pulsatility index of the uterine artery >p97.5 since we were particularly interested in cases with FGR based on placenta insufficiency. Controls were included when planned for Cesarean section because of either breech presentation or history of Caesarian section. Additional file The powdered samples, two scoops diluted in 700 μl sPBS, were used for chromatin isolation using the MAGnify™ Chromatin Immunoprecipitation System kit  according to manufacturer’s instructions. In brief, the tissue was crosslinked with 1% formaldehyde and the crosslinking was stopped by adding 1.25 M glycine. Cells were lysed using the kit-provided lysis buffer, and nuclei were sonicated using Covaris microTUBE . We aimed for DNA fragments of 200–2000 bp long. We continued with one fifth of the volume, and the sheared chromatin was diluted and then incubated with 1 μl anti-H3K27ac antibody  pre-coupled to magnetic beads for 2 h at 4 °C. Beads were extensively washed, and crosslinking was reversed by the kit-provided reverse crosslinking buffer with proteinase K. DNA was purified using ChIP DNA Clean & Concentrator kit (Zymo Research). Libraries were prepared using the NEXTflex™ Rapid DNA Sequencing Kit (Bioo Scientific). Samples were PCR-amplified and checked for the proper size range and for the absence of adaptor dimers on a 2% agarose gel, and barcoded libraries were sequenced 75 bp single-end on Illumina NextSeq500 sequencer (Utrecht Sequencing Facility).Sequencing reads were mapped against the reference genome  using BWA package (mem –t 7 –c 100 –M –R) . Multiplp < 0.05 by Wald test) [n = 5 FGR and n = 4 control samples.Regions with differential H3K27ac occupancy between FGR samples and controls were identified using DESeq2 (ld test)  and are To assign biological meaning to differentially acetylated peaks, we used three approaches. First, we annotated genes to the differentially acetylated sites in silico using a window of ± 20 kb from transcription start site (TSS). Second, GREAT software (Stanford) was used to assign biological meaning to a set of non-coding genomic regions by analyzing the annotations of genes flanking differentially acetylated regions . GREAT iTotal RNA was extracted from placental powder using RNeasy®  according to the manufacturer’s instructions by which all RNA molecules longer than 200 nucleotides are purified. Polyadenylated mRNA fraction was isolated using Poly(A) Beads , and sequencing libraries were constructed using the Rapid Directional RNA-seq kit . Libraries were sequenced on the Nextseq500 platform , producing single-end reads of 75 bp (Utrecht Sequencing Facility). Reads were aligned to the human reference genome GRCh37 using STAR version 2.4.2a. Picard’s AddOrReplaceReadGroups (v1.98) was used to add read groups to the BAM files, which were sorted with Sambamba v0.4.5, and transcript abundances were quantified with HTSeq-count version 0.6.1p1 using the union mode. Subsequently, reads per kilobase million reads sequenced (RPKMs) were calculated with edgeR’s RPKM function.p value calculation, FDR B&H correction, p value cutoff 0.05.Differentially expressed genes were identified using the DESeq2 package with standard settings. Genes with padj < 0.05 were considered as differentially expressed. Again, ToppFun was used for gene list enrichment analysis and candidate gene prioritization (accessed in March 207). The list of up- and downregulated genes were tested separately using probability density function To validate functional consequences of differentially acetylated regions, we overlapped the genes annotated to differentially hyperacetylated regions with upregulated genes derived from the mRNA-seq and genes annotated to differentially hypoacetylated regions with downregulated genes derived from the mRNA-seq. Functional pathway analysis with TOPPFUN were performed on both sets of overlapping genes. In addition, we searched the up- and downregulated genes for transcription factors (TF) with enriched TFBM within the differentially acetylated regions.t test or χ2 test.Data are shown as mean ± SD or median (range) and analyzed with Student’s Additional file 1:Table S1. Differentially acetylated regions in FGR compared to controls. Table S2. Genes annotated to hyper- and hypoacetylated sites in placenta of FGR using a 20-kb window. Table S3. Functional pathways related to hypo- and hyperacetylated in placenta of FGR identified by GREAT. Table S4. Up- and downregulated gene transcripts in placenta of FGR compared to controls. Table S5. RNA-seq pathways TOPPfunn. Table S6. Functional pathways related to overlapping genes in CHIP-seq and RNA-seq. Table S7. AME output: differentially acetylated TFBM’s in placenta of FGR vs. controls and functional pathways related to differentially acetylated TFBM (XLSX 694 kb)Additional file 2:Heatmap, MA plot, and V plot CHIP-seq. (DOCX 175 kb)Additional file 3:Heatmap, MA plot, and V plot RNA-seq. (DOCX 120 kb)Additional file 4:Pregnancy characteristics of the placenta samples. (DOCX 15 kb)


Cryptococcus neoformans, the transcription factor Znf2 controls yeast-to-hypha differentiation. Through a forward genetic screen, we identified the basidiomycete-specific factor Brf1. We discovered Brf1 works together with Snf5 in the SWI/SNF chromatin remodeling complex in concert with existent Znf2 to execute cellular differentiation. We demonstrated that SWI/SNF assists Znf2 in opening the promoter regions of hyphal specific genes, including the ZNF2 gene itself. This complex also supports Znf2 to fully associate with its target regions. Importantly, our findings revealed key differences in composition and biological function of the SWI/SNF complex in the two major phyla of Kingdom Fungi.Cellular differentiation is instructed by developmental regulators in coordination with chromatin remodeling complexes. Much information about their coordination comes from studies in the model ascomycetous yeasts. It is not clear, however, what kind of information that can be extrapolated to species of other phyla in Kingdom Fungi. In the basidiomycete  Cryptococcus neoformans. They show that the chromatin remodeling complex works with the transcription factor Znf2 in cellular differentiation.Jianfeng Lin et al. report the identification and characterization of the phylum specific SWI/SNF subunit Brf1 from  Through modulating gene expression, the SWI/SNF family of complexes are critical to a variety of cellular processes including stemness and differentiation.Cellular differentiation allows genetically identical cells to exhibit distinct phenotypes. Although functional and morphological differences are most pronounced in higher eukaryotes, cellular differentiation is a universal phenomenon, often directed by differential gene expression. In eukaryotes, nucleosomes serve as a general barrier preventing transcription and specific regulatory mechanisms make certain genomic DNA regions accessible to transcription factors and RNA polymerase II. The ATP-dependent SWItch/Sucrose Non-Fermentable (SWI/SNF) chromatin remodeling family mainly facilitates Saccharomyces cerevisiae and 11–15 subunits in humans3. However, most human SWI/SNF subunits have several isoforms, permitting dozens of combinatorial assemblies and a spectrum of related complexes4. It is, therefore, challenging to attribute observed phenotypes based on a mutation of a particular subunit to the function of a specific complex. Here, we use the term family when we discuss the SWI/SNF family of complexes. In S. cerevisiae and Schizosaccharomyces pombe, there are only two complexes in the SWI/SNF family: the SWI/SNF complex and the RSC (remodeling structure of chromatin) complex. These two complexes have their distinct catalytic and accessory subunits while sharing some common subunits6. The relative simplicity in these model yeasts has facilitated investigation of the molecular functions of SWI/SNF complexes and individual subunits. It is unclear, however, what functions and features of the SWI/SNF complexes established in these two model ascomycete yeasts can be extrapolated to other eukaryotic species, including other distantly related fungal species.The SWI/SNF complex is composed of 12 subunits in Cryptococcus neoformans belongs to a different phylum in Kingdom Fungi: Basidiomycota. Basidiomycetes diverged from ascomycetes about one billion years ago. They share key features with higher eukaryotes that are absent from the model yeasts. For instance, >90% of cryptococcal protein-coding genes contain multiple introns. Epigenetic regulation, such as RNAi and DNA methylation, plays important roles in cryptococcal biology10. C. neoformans can exist in multiple morphotypes and morphogenesis is associated with its pathogenicity11. For instance, yeasts and spores are infectious and virulent13; titan cells are proposed to be dormant and stress-resistant in hosts15; pseudohyphae and hyphae are attenuated in virulence in mammalian hosts16. In the environment, however, hyphae are an integral part of its life cycle and confer cryptococcal resistance to its natural predators like soil amoeba17.In contrast to these model yeasts, C. neoformans. During bisexual reproduction, a and α yeast partners conjugate after activating the pheromone-sensing cascade, and the resulting zygote differentiates to hyphae. In the absence of a compatible mating partner, the fungus can undergo similar cellular differentiation events18, with the exception that the non-self-recognition system is dispensable20. The zinc-finger transcription factor, Znf2, is an essential regulator for hyphal growth21. Deletion of ZNF2 confines cryptococcal cells to the yeast form and overexpression of ZNF2 drives filamentation regardless of growth conditions21. It is unknown whether chromatin remodeling factors coordinate with Znf2 to control the yeast-hypha differentiation in this basidiomycete.The yeast-to-hypha transition is the best-understood cellular differentiation process in Saccharomyces through genetic screens for mating-type switching or sucrose metabolism factors23. Here, through a forward genetic screen in a ZNF2 overexpression strain in C. neoformans, we identified two genes SNF5 and BRF1 that are essential for hyphal differentiation even when Znf2 protein is produced. Snf5 is a conserved core subunit in the SWI/SNF complex, while Brf5 is a novel basidiomycete-specific protein. We discovered that Brf1 works together with Snf5 in the SWI/SNF complex. We further demonstrated that Brf1 is essential for transcriptional induction of ZNF2 and is required for Znf2’s full association to the promoter regions of its downstream target genes, including the ZNF2 gene itself. Furthermore, the promoter region of ZNF2 and its downstream targets important for filamentation become transcriptionally inaccessible in the absence of BRF1 or SNF5. To our knowledge, this is the first identification and functional characterization of a phylum-specific subunit in the SWI/SNF complex in basidiomycetes. Our findings also revealed major differences in composition and biological functions of the SWI/SNF and the RSC complexes between the two major phyla in the Kingdom Fungi.The ATP-dependent chromatin remodeling SWI/SNF family complexes were initially discovered in ZNF2 in regulating yeast-to-hypha transition. Here, we employed a reporter strain to identify Znf2’s partners through a forward genetic screen. In this reporter strain, the native ZNF2 gene is deleted and an ectopic copy of mCherry-fused ZNF2 is expressed under the control of an inducible promotor of a copper transporter CTR4. This reporter strain switched from yeasts to hyphae when ZNF2 was induced in the presence of the copper chelator bathocuproinedisulfonic acid (BCS)  and visually screened 88,000 T-DNA insertional mutants made in this reporter strain for smooth yeast colonies on filamentation-inducing V8 agar medium containing BCS  Fig. , as expenal Fig. . Here weBCS Fig. . Eighty-BCS Fig. .Fig. 1BRFifty-eight mutants showed nuclear mCherry-Znf2 while 26 had altered subcellular distribution of the fluorescence signal. The mutants with mis-localized Znf2 signal might be defective in Znf2 trafficking, post-translational modification, and/or stability. Here, we set to identify factors that are required for nuclear-localized Znf2 protein to exert its function. Among these 58 mutants, 25 candidates showed only yeast cellular morphology and nuclear mCherry-Znf2 signal when cultured in YPD + BCS-inducing condition  to the wild-type congenic mating partner XL280a, micro-dissected basidiospores, and analyzed the phenotypes of the meiotic progeny. The genetic linkage analyses indicate that 8 out of the 25 mutants likely harbor a sole T-DNA insertion that is genetically linked to their non-filamentous phenotype on BCS media  is present , which encodes a general transcriptional co-repressor, reduced filamentation in the wild-type (WT) background , a core subunit of the conserved SWI/SNF complex. Snf5 is critical for cellular differentiation in all organisms tested, including ascomycetous and basidiomycetous fungi32. In C. neoformans, a bilateral snf5Δα × snf5Δa cross yielded no mating hyphae or fruiting body in the serotype A H99 strain background25. Consistently, we found that a bilateral snf5Δα × snf5Δa cross in the serotype D XL280 background yielded no mating hyphae . The predicted protein has an AT-rich interacting domain (ARID) but no other recognizable domains. Because it is basidiomycete specific .SNF5 or BRF1 abolished or nearly abolished self-filamentation  did not produce any aerial mating hyphae or sexual spores . Taken together, we propose that Brf1 functions together with Snf5.Given the striking resemblance of the ays Fig. . Constit36. C. neoformans carries three genes encoding ARID-containing proteins: RUM1α (CND05870), AVC1 (CNK00710), and BRF1. None of these three cryptococcal genes showed high sequence homology to either SWI1 or SOL1, the ARID-containing subunit in the SWI/SNF complex in S. cerevisiae or S. pombe, respectively. However, among the three proteins, Brf1 is more similar to ScSwi1 or SpSol1 in terms of domain architecture  or the RSC complex (sfh1Δ and rsc9Δ) on filamentation. Indeed, the RSC complex mutants, sfh1Δ and rsc9Δ, were still filamentous on V8 medium, in contrast to the yeast growth of the SWI/SNF complex mutants brf1Δ and snf5Δ , pheromone receptor STE3α (14× reduction), pheromone exporter STE6 (7× reduction), MAPK CPK1 (5.1× reduction), and pheromone transcription factor MAT2 (9× reduction)  had minimal impact on the transcript level of BRF1 (1.27× increase). The transcript of BRF1 remained at a low and steady level when cells were cultured either in YPD or on V8 medium. The steady level of BRF1 transcripts was also observed at different cell cycle stages  experiments using 3xFLAG-tagged Znf2 as the bait in the WT or the old Fig. . Thus, tZNF2 and CFL1 promoter regions, was dampened upon the disruption of Brf1 45 and compared genetic regions with open chromatin structures in the WT strain, grown under filamentation-repressing or -inducing conditions, as well as in the brf1Δ, znf2Δ, and snf5Δ strains cultured under filamentation-inducing conditions. We included the brf1Δ/BRF1oe strain as an additional control. As expected, accessible chromatin fell in the promoter regions, with expressed genes having a higher level of accessibility based on the relative enrichment of ATAC-seq reads  . Disruption of either Brf1, Snf5, or Znf2 abolished accessibility of the CFL1 promoter (ATAC-seq), where Znf2 binds to (ChIP-seq)  Fig. . CFL1 iseq) Fig. . Accordivel Fig. . SimilarCollectively, the data support the interdependence between the transcription factor Znf2 and the SWI/SNF complex in opening up chromatin to facilitate transcription of filamentation genes.S. cerevisiae and S. pombe have offered a relatively simple system for mechanistic studies. Basidiomycetous fungi resemble more of the higher eukaryotes in terms of genome structures, epigenetic regulation, and transcriptome complexity, but research on chromatin remodeling in this major phylum of the fungal kingdom is scarce.Chromatin remodeling plays critical roles in cellular differentiation in eukaryotes. The model ascomycetous yeasts C. neoformans. This complex remodels the chromatin structure of the promoter regions of filamentation genes to make them accessible for transcription . Our findings are consistent with published literature in other organisms, in which the SWI/SNF complex contributes to the DNA binding of the transcription factor47. The SWI/SNF complex has been shown being targeted to specific genetic loci by sequence-specific transcription factors and acetylation of histone tails49. In the ascomycetous fungus C. albicans, the transcription factor Efg1 recruits the histone acetyltransferase complex NuA4 to the promoters of hypha-associated genes, allowing, in turn, the recruitment of the SWI/SNF complex to activate their transcription50. We reason that Znf2 may recruit the SWI/SNF complex to its target sites given that accessibility at the CFL1 promoter region is lost in the absence of ZNF2 , Sfh1, and Rsc9 (refs. 41)  and SFH1 (RSC-specific) together slowed growth but did not cause lethality  between Ascomycota and Basidiomycota. In ity Fig. . ComparaC. neoformans and other basidiomycetes harbor two proteins with an Snf5 domain: SNF5 and SFH1 and they cluster closely with the Sfh1 clade in ascomycetes  likely belong to two different modules39 given the phenotypical differences of the snf5 and swi1 mutants52. However, Cryptococcus snf5 and brf1 mutants have nearly identical phenotypes. Furthermore, Snf5 proteins in basidiomycetes  are much larger in size compared to those in ascomycetes or in humans  , presumably one acting as the Snf5 subunit in the SWI/SNF complex and one acting as the Sfh1 subunit in the RSC complex Fig. . Based otes Fig. . The SWIaa) Fig. . Two intaa) Fig.  could po3. A search for the HSA domain (PF07529) and the BROMO domain (PF00439) revealed that some fungi, including ascomycete S. cerevisiae and zygomycete Mucor circinelloides, have two distinct Snf2 family ATPases, presumably one for the SWI/SNF complex and one for the RSC complex. Some other fungi, including ascomycete Aspergillus nidulans and basidiomycete C. neoformans, however, have only one Snf2 ATPase, presumably shared by SWI/SNF and RSC  complex and the PBAP (RSC) complex share the same catalytic ATPase subunit (BRM). Human BAF (mSNF/SWI) complex and PBAF (mRSC) complex can share the same ATPase BRG-1 subunit or use different ATPases (hBRM and BRG-1)3. Thus, the ATPase could be shared by or be unique to the SWI/SNF and the RSC complexes. The copy number variation of SNF2 might have driven such divergence independent of the evolutionary distance of the species.The SWI/SNF family employs Snf2 as the ATPase catalytic subunit, which carries an HSA and a BROMO domain (the DEXDc and HELICc regions)main PF0039 revealIn conclusion, we discovered the basidiomycete-specific subunit of SWI/SNF chromatin remodeling complex and revealed the major differences in the composition and biology of this complex between ascomycetes and basidiomycetes. Given the universal importance of the SWI/SNF complex in cellular differentiation in eukaryotes, our findings provide an important platform for future comparative functional analyses of this complex in diverse eukaryotic lineages.C. neoformans cells were cultured in YPD medium unless stated otherwise. For some assays, the defined minimal YNB medium  was used as specified in the figure legends. For mating or filamentation assays, V8 juice agar medium  was used.Strains used in this study are listed in Supplementary Table Agrobacterium-mediated cryptococcal transformation was carried out as described previously53. Briefly, engineered Agrobacterium cells that carry a Ti plasmid with the hygromycin-resistant marker were co-incubated with the recipient Cryptococcus cells  on the induction medium at 22 °C for 2 days. The Cryptococcus–Agrobacterium cocultures were collected, diluted, and transferred onto the selection medium (with 50 μM of BCS) that would induce ZNF2 expression. Cryptococcal transformants were cultured on this selective medium for 3–5 days at 22 °C in dark. Approximately 500 colonies grew on each selective plate and about 88,000 transformants were screened for smooth yeast colonies.The 54. The genomic DNA for each strain was then pooled together with equal molar concentration and the pooled DNA was sent for sequencing . The insertion sites in the mutants were identified via the AIMHII program developed by Esher et al.26 with default parameters.Genomic DNA of the selected mutants was prepared with the CTAB DNA extraction protocol as described previously56. Single-guide RNAs were designed with Eukaryotic Pathogen CRISPR guide RNA/DNA Design Tool (http://grna.ctegd.uga.edu/) for TRACE.The gene knockout constructs were constructed by fusing 5′ and 3′ homologous arms (each about 1 kb) with dominant drug marker via overlap PCRs. The constructs were introduced into cryptococcal cells by biolistic transformation or by TRACE (Transient CRISPR-Cas9 Coupled with Electroporation) as described previouslyFseI and AsiSI cutting sites was digested and ligated into the vectors where the ORF was placed downstream of the GPD1, the TEF1, or the CTR4-2 promoter, as we described previously42. A fluorescent tag (tdTomato or mNeonGreen) or a CBP -2×FLAG tag was placed immediately in-frame downstream of the AsiSI cutting site, which allows in frame tagging of the protein at the C-terminus. ZNF2 ORF was ligated into a vector where a 3xFLAG tag was downstream of the CTR4 promoter to allow tagging at the N-terminus. The functionality of the tagged proteins was confirmed by their ability to rescue/restore the phenotypes in the corresponding gene deletion strains. All constructed plasmids used in this study are listed in Supplementary Table To generate gene overexpression strains, the ORF amplicon with added 600 = 3). Three microliters of cell suspension and 10× serial dilutions were spotted onto YPD agar medium or YPD with the supplement of Congo Red (0.3–0.5%) and were incubated at 30 or 37 °C for 1–2 days. To test carbon source utilization, glucose in the YNB medium was replaced by either sucrose or raffinose .For phenotypic analyses, cells of the tested strains were cultured in YPD broth overnight at 30 °C with shaking at 230 r.p.m. Cells were collected by centrifugation, washed twice with sterile water, and then suspended with water to the same optical density at 600 nm  and incubated at 22 °C in dark for 1–7 days.To test the ability of these strains to undergo self-filamentation, 3 μL of cells with ODCTR4-2-inducible promoter to drive the expression of the examined genes, cells were maintained on YPD medium with 50 μM CuSO4 to suppress the gene expression. To induce the gene expression, the copper chelator BCS was supplemented to the medium with the final concentration of 200 μM in YPD and 50 μM in V8.For strains that use the 600 = 1. Fifty microliters of each strain were added to 950 μL of the indicated liquid medium in a 24-well plate . The growth of the cryptococcal strains (each with two replicates) was monitored at OD600 hourly in the Cytation 5 multi-mode reader (BioTek Instrument) with double orbit shaking (at 365 cpm) at 30 °C. The reads for each well were fitted to a preloaded logistic model Cryptococcal cells from overnight culture in YPD broth were centrifuged, washed twice with water, and resuspended in water. The cell concentration was normalized to the optical density of ODa mating partners were crossed on V8 juice agar medium (pH 5 or 7) and incubated at 22 °C in dark for 2–3 weeks until adequate spores were produced. Spores were micro-manipulated using a dissecting microscope. The mating type of the germinated spores was determined by successful mating with either JEC20a or JEC21α reference strain. Genetic linkage between the presence of the drug marker and the observed mutant phenotype was established by analyzing the dissected spores as we described previously57. If the insertional mutant harbors only one copy of the T-DNA, which carries the hygromycin selection marker HYG, we would expect a 1:1 ratio of hygromycin-sensitive (HYGS) and hygromycin-resistant (HYGR) progeny. If that single insertion caused the non-filamentous phenotype, then all HYGR progeny would be non-filamentous while all HYGS progeny would be filamentous under a Znf2-inducing condition. In comparison, when multiple T-DNAs inserted in the genome, the ratio between HYGS and HYGR meiotic progeny would be 1:3 (two unlinked insertions) or lower (>2 unlinked T-DNA insertions). For each genetic linkage assay, approximately 32 viable spores were analyzed, which gave a 97% confidence level of our analysis Strains of α and brf1::NAT/PTEF1-BRF1-CBP-2xFLAG-NEO (α) with znf2::NAT/PCTR4-2-3xFLAG-ZNF2-NEO (a) and examined the segregation of the genotypes (by diagnostic PCRs) and phenotypes in the progeny  and the control strain (prf1::NAT) were collected and suspended in water to the same optical density . Each mutant with NATR was mixed with the mating type a tester strain with NEOR marker (strain YSB133) with equal volume58. Three microliters of cell mixture were spotted onto V8 medium and incubated at 22 °C in dark for 24 h before they were collected, plated  onto YPD + NEO + NAT agar medium, and incubated at 30 °C for 3–5 days to select for fusion products.To determine the cell–cell fusion efficiency, mutants of the mating type α  was retrieved from the Ensembl Fungi database. Simple Modular Architecture Research Tool (SMART) online server was used to analyze the domain layouts for proteins. IBS online illustrator tool was used to illustrate all the protein domain layouts59.We used 60. HiSeq Rapid 175bp pair-end RNA sequencing was performed at the Georgia genome sequencing facility. The low-quality bases of the raw reads were trimmed with a custom perl script as published before60. Tophat2 was used to map the processed reads to the reference genome. The program HTSeq-count and DESeq2 were used to count the reads and identify the differentially expressed genes. The raw sequencing reads were deposited at NCBI with the BioProject accession number PRJNA534125 and SRA file numbers from SRR8947060 to SRR8947075.RNAs were extracted from cells cultured on V8 medium at 22 °C in dark for 24 h and prepared for sequencing as previously described® (Next Advance) without buffer for five cycles (60 s maximum blending followed by 90 s chilling on ice) and then with 1 mL of lysis buffer ) for another five cycles. Cell lysates were centrifuged at 500 × g at 4 °C for 5 min. The supernatant was collected and centrifuged again at 10,000 × g at 4 °C for 20 min. The supernatant was then incubated with pre-washed anti-FLAG M2 agarose beads  at 4 °C on a rotator overnight. For immunoprecipitation with mCherry or mNeonGreen-tagged proteins, RFP-Trap®_MA or mNeonGreen-Trap_MA beads from Chromotek (Germany) were used following the manufacturer’s instructions.Strains with tagged proteins were cultured in 15 mL YPD media overnight, washed twice with cold water, flash frozen in liquid nitrogen, and lyophilized. Lyophilized cells were broken into fine powder with silica beads in a Bullet Blender BlueAfter washing, immunoprecipitated protein samples were released from beads in the 2×SDS loading buffer  by boiling for 10 min.https://pams.uga.edu/) at the University of Georgia for identification. When analyzing the results, we applied the following criteria to exclude nonspecific proteins: proteins from the negative control, proteins not shared between samples of the two PTEF1-BRF1-CBP-2×FLAG strains, and non-nuclear proteins as Brf1 localizes to the nucleus.For Co-IP/MS, protein samples were then loaded into pre-casted 4–12% SurePAGE™ Bis-Tris Gel (GeneScript), ran for 5 min, and visualized by Coomassie Blue staining. The total pulled down protein samples were excised from the gel and sent to the proteomics and mass spectrometry facility  were run in an SDS-PAGE gel (30 μL of WCE and IP), transferred onto PVSF membrane, and western-blotted with anti-FLAG M2 antibodies .600 = 0.2) were incubated in 50mL YPD + BCS broth at 30 °C until the optical density reached OD600 = 1. Cells were then fixed in the medium with formaldehyde at 1% final concentration at 22 °C for 15 min with occasional swirling. Glycine with the final concentration of 0.125 M was added to quench the crosslinking at 22 °C for 5 min. The cells were then washed twice with cold sterile water and lyophilized. Lyophilized cells were broken as described above for Co-IP experiment with a different lysis buffer . The following steps were similar to what has been described previously63 and briefly, the cell lysates were centrifuged at 8000 r.p.m. at 4 °C for 10 min to enrich nuclei. The released nuclei were resuspended in 350 μL of lysis buffer. The nuclear suspensions were sheared by sonication in a Diagenode BioruptorTM for 25 cycles at 4 °C . The protein–chromatin complex was recovered from the supernatant after centrifugation at 14,000 r.p.m. at 4 °C for 10 min and the volume was brought to 1 mL. Fifty microliters of each protein–chromatin suspension was saved as Input DNA. The rest of the sample was added to 30 μL anti-FLAG M2 monoclonal antibodies on magnetic beads  to precipitate the FLAG-tagged protein. After incubation with antibodies at 4 °C overnight, the beads were washed twice with 1 mL ChIP lysis buffer, once in 1 mL ChIP lysis buffer containing 0.5 M NaCl, once in 1 mL ChIP wash buffer , and once in 1 mL TE , all at 4 °C. The immunoprecipitated Znf2-3xFLAG was eluted twice by adding 200 µL of TES buffer  and incubated at 75 °C for 15 min before the supernatant was transferred to a new tube. Twenty microliters of 5 M NaCl was added to each sample to de-crosslink at 65 °C for 4 h. Three hundred and fifty microliters of TES was added to each Input DNA sample before adding 20 µL of 5 M NaCl to reverse formaldehyde crosslinks. One microliter of RNAse A (10 µg/µL) was added and the sample was incubated at 37 °C for 30 min. Four microliters of Proteinase K (20 µg/µL) was then added to each sample and incubated at 45 °C for at least 1 h to digest all the proteins. Eventually, 2 µL of glycogen  was added to chromatin DNA samples. Ethanol of 2.5 volumes was added to each sample to precipitate the glycogen tangled DNA. The precipitated DNA sample was then suspended in 80 μL of nuclease-free water.FLAG-tagged Znf2 strains (strain JL653 and JL665)  where ΔCt [normalized ChIP] = Ct [ChIP]−(Ct [Input] −log2(input dilution factor)) as described previously63.For ChIP-qPCR, 3 μL of five times diluted DNA sample and 1 μL each from two primers (50 μM) were mixed with 5 μL of SYBR Green 2x qPCR premix reagents (Invitrogen). The qPCR reactions were carried out in a Realplex system (Eppendorf) with technical duplicates. The % Input is calculated as 2 65. ChIP-seq have been submitted to the GEO database (accession # GSE137248).The same ChIP samples for qPCRs were sent for DNA sequencing at Georgia Genomics Facility (UGA) on an Illumina NextSeq500 platform. Input from ChIP-seq experiments in WT strains was used as a background control for all samples. Differential peaks called using the findPeaks tool, part of them were annotated using ChipSeekerC. neoformans cells were cultured in 15 mL YPD liquid media at 30 °C overnight or on V8 medium (pH 7) at 22 °C in dark for 24 h. Cells were collected and washed two times with cold sterile water. Five hundred microliters of cold lysis buffer  β-me; 0.3% TrixtonX-100) and 200 μL (~1 PCR tube) of acid-washed glass beads  were added to the cell pellet of about 108 cells in 1.7 mL Eppendorf tubes. Cells were broken in a cell disruptor  at 4 °C with maximum speed for 2 min and then spun down at 50 × g at 4 °C for 2 min to remove the glass beads. The supernatant was transferred into a new Eppendorf tube and spun at 200 × g for another 2 min to remove most of the broken cells and debris. The enrichment of nuclei in the supernatant was verified by microscopic observation and the nuclei were collected for making the ATAC-seq library.ATAC-seq libraries were generated as follows. Briefly, about 0.2 million nuclei were incubated with Tn5 transposase preloaded with Illumina sequencing adapters at 37 °C for 30 min followed by purification of the DNA fragments by a reaction cleanup kit . Libraries were PCR amplified for 10 cycles with Phusion polymerase . Sequencing libraries were then cleaned with magnetic beads to remove free adapters and primer dimers . Libraries were mixed in equimolar ratios and pair-end sequenced by the Georgia Genomics Facility (UGA) on an Illumina NextSeq500 platform.https://github.com/FelixKrueger/TrimGalore). Duplicate reads were removed using Picard MarkDuplicates tool (http://broadinstitute.github.io/picard/). Reads were aligned to the JEC21 reference genome (Refseq assembly GCF_000091045.1) with HISAT2 using non-spliced alignment and a maximum fragment length of 2000 bp. Peak calling was performed with MACS2 using a q-value of 0.01, extension size of 73, and shifting reads by 37 bp to center on the insertion site. MACS2 was also used to call differential peaks66. Differential peak files were first sorted by fold change and then combined into a single file. Genes nearby Znf2 peaks were annotated using ChipSeeker65. ATAC-seq enrichment was calculated across Znf2 peaks or differential peaks between WT and brf1Δ using the annotatePeaks function, part of the HOMER (Hypergeometric Optimization of Motif EnRichment) suite of informatics tools, with parameters -size 3000 and -ghist67. Heatmaps were generated in R using the pheatmap package version 1.0.12. Differential peaks from ATAC-seq were compared to each other or Znf2 peaks or to genes identified as being up-regulated in WT V8 from RNA-seq analysis using BEDTools68 intersect to isolate these genes from the JEC21 annotation file for determining genes associated with differential peaks. ATAC-seq have been submitted to the GEO database (accession # GSE137248).ATAC-seq reads were filtered for low-quality and short reads with Trim Galore . More details can be referred to the corresponding sections.Further information on research design is available in the Supplementary InformationDescription of Additional Supplementary FilesSupplementary Data 1Supplementary Data 2Supplementary Data 3Supplementary Data 4Supplementary Data 5Supplementary Data 6Reporting SummaryPeer Review File