Comparative proteomics of eukaryotic histone modifications

We analyzed the phylogenetic distribution and evolutionary history of histone proteins. To this end, we surveyed the presence of histone-fold proteins across 172 eukaryotic and 4,226 archaeal taxa, using HMM searches (Fig. 1a,b andSupplementary Data 1). Histone proteins are found in all eukaryotic genomes. We clustered the identified 8,576 histone-encoding proteins using pairwise local alignments and then classified individual sequences in these clusters based on pairwise alignments to a reference database⁴⁸ (Fig. 1a andExtended Data Fig. 1a). This reveals four broad clusters corresponding to the four main eukaryotic histones (H2A, H2B, H3, and H4) and their variants (H2A.Z, macroH2A, and cenH3), as well as a fifth cluster composed of archaeal HMfB homologs. Finally, this classification also uncovers three large connected components composed of transcription factors with histone-like DNA binding domains, which are widely distributed in eukaryotes (POLE3, POLE4, DR1) and/or archaea (NFYB). Further analysis of the genomic distribution of these histone genes shows a frequent occurrence of H3-H4 and H2A-H2B pairs in head-to-head orientation (5' to 5'), strongly indicating co-regulation across eukaryotes (Extended Data Fig. 1b,c andSupplementary Data 2).

Next, we investigated the distribution and conservation of hPTMs across major eukaryotic groups and Archaea, including methylations, acetylations, crotonylations, phosphorylations, and ubiquitylations. To this end, histones from 19 different eukaryotic species were extracted, chemically derivatized⁴⁹ and analyzed by mass-spectrometry (Fig. 1c andSupplementary Data 3), adding to previously available hPTM proteomics data for additional seven species. Our extensive taxon sampling covers all major eukaryotic groups, as well as hitherto unsampled early-diverging eukaryotic lineages—such as the malawimonadGefionella okellyi, the discobanNaegleria gruberi, or the ancyromonadFabomonas tropica—, thus providing a comprehensive comparative framework for evolutionary inference.

We focused first on hPTMs present in canonical histones, as defined by their highly conservedN-terminal regions, phylogenetic analyses, and sequence similarity to curated reference canonical histones (Fig. 1d; seeMethods). hPTMs are detected in all canonical histones from all species. After correcting by sequence coverage, we observe that hPTMs are particularly abundant in H3 canonical histones (median = 23.5 hPTMs per species, mean = 24.3), compared with H2A, H2B and H4 (medians between 6.5 and 9, means between 9.5 and 13.4;Extended Data Fig. 2a). Holozoan canonical H2As (Homo sapiens, Sycon ciliatum andCapsaspora owczarzaki) represent an exception to this trend and contain similar number of modifications to H3s in these species. We also examined the reproducibility of hPTM detection across replicate samples, showing that the majority of hPTMs (87.5%) can be found in more than one sample (Extended Data Fig. 2b,c). Despite this, it is worth emphasizing that our data may contain false negatives, beyond the lack of coverage for particular residues that we systematically report. For example, some marks might be globally too scarce in the nucleosomes of a particular species, while other modifications like phosphorylations and ubiquitination are difficult to detect by mass-spectrometry without dedicated peptide-enrichment protocols.

Canonical H3 and H4N-terminal tails contain the majority of phylogenetically-conserved hPTMs, in stark contrast with the relative paucity of conserved hPTMs in canonical H2A and H2B. A striking example of paneukaryotic conservation comes from the acetylation of the H4 K5, K8, K12 and K16 residues (Fig. 1d, second panel), all of which mark gene expression-permissive chromatin environments in multiple eukaryotic species²². A similar conservation pattern is observed in the acetylation of a group ofN-terminal H3 lysines (K9, K14, K18, K23, K27) associated with similar functions, while other H3 acetylations are only found in a few species (e.g., residues K4, K56 and K79). While acetylations are highly conserved, only seven histone H3/H4 methylations are broadly conserved across eukaryotic lineages: H3K4me1/2/3, H3K9me1/2/3, H3K27me1/2/3, H3K36me1/2/3, H3K37me1/2/3 and, more sparsely, H3K79me1/2 and H4K20me1. Many of these broadly conserved marks have conserved roles in demarcating active (e.g., H3K4me) and repressive chromatin states (e.g., H3K9me and H3K27me)^22,42,50. The scarcity of conserved hPTMs in H2A and H2B canonical histones can partially explained by their higher degree of sequence divergence (Fig. 1e), which is reflected in many non-homologous lysine residues (Fig. 1d). But even among homologous positions, we found little evidence of conservation, with the exception of H2A K5ac (associated to active promoters⁵¹) and, in fewer species, methylation of H2A K5 and H2B K5. Finally, we were also able to identify phosphorylations in serine and threonine residues and a few instances of ubiquitylation. In general, these marks show more restricted phylogenetic distributions than lysine acetylation or methylation, even in the tightly conserved H3 and H4 histones. We can identify conserved phosphorylations in H2A T120 and S122, which are shared by most opisthokonts, and the ubiquitylation of H2A K119 only in some holozoan species.

Mass-spectrometry analysis detected histone variants in all species included in our study, suggesting that they are relatively abundant in the chromatin of these eukaryotes (Fig. 1e). Most of these variants are lineage-specific, with the exception of the paneukaryotic variants H2A.Z, H3/cenH3 and H3.3; and the macroH2A variant found in holozoans andMeteora sp. (belonging to an orphan eukaryotic lineage). Interestingly, we find hPTMs in the vast majority of detected variants, both conserved and lineage-specific, particularly acetylations and methylations (Fig. 1e andExtended Data Fig. 2d). Overall, our comparative proteomic analysis suggests the existence of a highly conserved set of canonical hPTMs of ancestral eukaryotic origin in H3 and H4, which co-exists with less conserved hPTMs in H2A, H2B, and lineage-specific modifications in variant histones.

Archaeal histone post-translational modifications

In contrast with the paneukaryotic distribution of histones, sequence searches show that only a fraction of archaeal genomes encode for histones (28.1% of the taxa here examined;Fig. 2a). Archaeal histones exhibit a patchy phylogenetic distribution, similar to other gene families shared with eukaryotes⁵². Among others, histones are present in Euryarchaeota, the TACK superphylum and Asgard archaea^12,53–56. Asgard are generally are considered to be the closest known archaeal relatives of eukaryotes^57,58, although this sister-group relationship has been challenged by some studies⁵⁹. Our extended sampling revealed that Asgard archaea histones, particularly in the Lokiarchaeota and Heimdallarchaeota clades⁵⁵, often have lysine-richN-terminal tails in the manner of eukaryotic histones (Fig. 2a-c). These Asgard histones appear to be conserved across multiple taxa, albeit without direct sequence similarity compared to canonical eukaryotic histones (Extended Data Fig. 1d). When compared against eukaryotic sequences classified in HistoneDB⁴⁸, these archaeal histones clearly cluster in a separate group and are most similar to either eukaryotic H4 or, to a lesser degree, H3 canonical histones, in line with previous findings^12,55,60.

To identify potential archaeal hPTMs, we performed proteomics analysis of histones in three Euryarchaeota (the MethanobacteriotaMethanobrevibacter cuticularis and the HalobacteriotaMethanospirillum stamsii andMethanosarcina spelaei) and one Thaumarchaeota species (Nitrososphaera viennensis;Fig. 2b). Mass-spectrometry detects histone proteins in all of them: 2-4 in the euryarchaeotes (with 27-90% protein coverage) and one in the thaumarchaeote (80% protein coverage), including homologs withN-terminal tails encoded by each of the three euryarchaeotes in our survey (22-40 aa, 0.09-28 lysines per residue;Fig. 2c). Moreover, this proteomics analysis finds evidence of hPTMs in archaeal histones. However, in comparison with eukaryotic histones, hPTMs are extremely scarce in archaeal histones. Specifically, we identify no hPTMs inN. viennensis andM. spelaei (one and two histones detected, respectively), three acetylations and one methylation inM. stamsii (in three out of four histones detected), and one acetylation and two methylations inM. cuticularis (in two out of four histones;Fig. 2b, top). Interestingly, we find conserved lysine residues with shared modifications inM. stamsii andM. cuticularis (methylation in K54 and acetylation in K57;Fig. 2b, bottom). This result indicates that highly-abundant hPTMs represent a eukaryotic innovation, likely linked to dynamic nucleosomal regulation in eukaryotes but not in Archaea.

Taxonomic distribution of chromatin-associated proteins

hPTMs are deposited and removed by specific modifying enzymes (‘writers’ and ‘erasers’), while ‘reader’ protein domains found in diverse proteins bind and recognize specific hPTMs. For example, Bromo and Chromo domains bind acetylated and methylated lysine residues, respectively. In addition, the control of histone loading/eviction from specific genomicloci is mediated by chromatin remodellers, like SNF2 proteins²⁷, and histone chaperones²⁶. To date, the classification and evolutionary analysis of this chromatin machinery has been based on biased, partial taxonomic samplings and has not employed phylogenetic methods⁶¹ (with rare exceptions^12,27), often resulting in inaccurate orthologous relationships and confounded classification and naming schemes.

We sought to obtain a systematic, phylogenetics-based classification of histone remodellers, chaperones, readers, and modifiers in order to understand the evolutionary history of eukaryotic chromatin (Fig. 3a). To this end, we (i) compiled a taxa-rich dataset of 172 eukaryotic genomes and transcriptomes, covering all major eukaryotic supergroups and devoting particular attention to early-branching, non-parasitic lineages (Supplementary Data 1), as well as genomic data from 4,226 Archaea, 24,886 Bacteria and 185,579 viral taxa; (ii) defined a protein structural domain as a proxy for each gene family (Supplementary Data 4) and retrieved all genes in these genomes that contained these domains; and (iii) inferred accurate orthology groups from phylogenetic analyses of each gene class (next section).

We examined the taxonomic distribution and abundance of the major gene classes (Fig. 3b,c). Many domains with chromatin-associated functions in eukaryotes are also present in Archaea and Bacteria, albeit with scattered phylogenetic distributions (Fig. 3b andExtended Data Fig. 3a,b). Families with prokaryotic homologs include mostly catalytic gene classes (writer, eraser and remodeller enzymes), whereas readers and histone chaperones are virtually absent from prokaryotes (Fig. 3b). Histone fold-encoding genes constitute a case in point for this patchy distribution of chromatin proteins in prokaryotes: they are present in most archaeal phyla, but are absent in about half of the sampled genomes within each (Fig. 3b). Yet, there is a qualitative difference between the phylogenetic distribution of archaeal and bacterial chromatin-associated gene classes: whereas archaeal histones tend to co-occur with chromatin-associated gene classes, the bacterial complement of writers and erasers is much less conserved and is uncorrelated with the extremely rare presence of histone-like genes (Fig. 3d).

Within eukaryotes, most gene structural classes associated with chromatin functions are ubiquitously distributed across all lineages here surveyed, supporting an early eukaryotic origin for the core chromatin machinery (Fig. 3b andExtended Data Fig. 3d). In fact, the total number of chromatin writer, eraser and remodeller enzymes remains remarkably stable across eukaryotes (Fig. 3e). The only exception is the marked increase in genes encoding reader domains observed in lineages exhibiting complex multicellularity: animals, streptophyte plants, and, to a lesser degree, phaeophyte brown algae (Stramenopila). This occurs partially due to the addition of new gene classes (e.g., SAWADEE in the Plantaes.l. + Cryptista lineage, or ADD_DNMT3 in bilaterians and cnidarians), but also via the expansion of ancient, widely-distributed reader gene classes (e.g., Tudor, PHD, Chromo or Bromo domains). These taxonomic patterns indicate that chromatin modifying and remodelling catalytic activities originated in prokaryotes, while reader and chaperone structural domains are eukaryotic innovations.

Phylogenetics of chromatin modifiers and remodellers

To gain detailed insights into the origin and evolution of chromatin gene families, we used phylogenetic analysis to define orthology groups from paneukaryotic gene trees. We surveyed 172 eukaryotic species and defined a total of 1,713 gene families (orthogroups), 95% of which were conserved in two or more high-ranking taxonomic groups (as listed inFig. 1a), and which included 51,426 genes in total (Supplementary Data 5). We annotated each gene family according to known members from eukaryotic model species. For simplicity, we use a human-based naming scheme throughout the present manuscript (unless otherwise stated), but we also provide a dictionary of orthologs in three additional model species (Arabidopsis thaliana,Saccharomyces cerevisiae andDrosophila melanogaster; seeSupplementary Data 5). This phylogenetic classification scheme of eukaryotic chromatin gene families, as well as the sequences and associated phylogenetic trees, can be explored and retrieved in an interactive database:https://sebe-lab.shinyapps.io/chromatin_evolution

We first investigated the potential pre-eukaryotic origins of these gene families/orthogroups by comparing their phylogenetic distance to prokaryotic sequences and to other eukaryotic orthogroups (Fig. 4a). Most eukaryotic gene families are more closely related to other eukaryotes than to prokaryotic sequences, supporting the idea that writers, erasers, remodellers and readers diversified within the eukaryotic lineage, as previously noted for histones¹². This analysis also reveals a substantial fraction of eukaryotic gene families with close orthogroups in Archaea and Bacteria, which pinpoints components that were (i) inherited from a prokaryotic ancestor during eukaryogenesis; (ii) laterally transferred between eukaryotes and prokaryotes at later stages; or (iii) a combination of both phenomena. For example, we identified a well-supported sister-group relationship between the eukaryotic SIRT7 deacetylase and a clade of Asgard archaea Sirtuin enzymes (Heimdallarchaeota and Lokiarchaeota), a topology compatible with an archaeal origin or ancient transfers to/from Asgard and eukaryotes⁶²; whereas SIRT6 appears nested within other eukaryotic sequences (Fig. 4b, left). Likewise, the KAT14 acetylase is more closely related to bacterial enzymes than to other eukaryotic acetylases (Fig. 4b, right).

Next, we mapped the phylogenetic distribution of orthogroups in order to infer the origin and diversification of individual chromatin gene families (Fig. 4c andExtended Data Fig. 4a). Using probabilistic inference of ancestral gene content, we reconstruct a rich Last Eukaryotic Common Ancestor (LECA) complement of chromatin-associated gene families: 65 acetylases (amongst which 61 were conserved in at least two of the most deeply sampled eukaryotic early-branching lineages, namely Amorphea, Diaphoretickes, and Discoba); 20 deacetylases (19 in these early-branching eukaryotic lineages); 59 methyltransferases (55); 42 demethylases (38); 33 remodellers (33); and 25 chaperones (18) (Fig. 4c andSupplementary Data 5). The subsequent evolution of these families is characterized by relative stasis, with few new orthologous families emerging in later-branching eukaryotic lineages. Notable exceptions include the origin of KAT5 deacetylases and KMT5B/C SET methyltransferases in Opisthokonta; KAT8 and SIRT7 in Holozoa; and Viridiplantae-specific deacetylases (homologs ofA. thaliana HDA7 and HDA14 deacetylases) and SETs (A. thaliana PTAC14); among others.

In spite of their broad distributions across eukaryotes, many chromatin modifier families exhibit variation in their protein domain architectures, likely conferring them functional properties such as distinct binding preferences (Extended Data Fig. 4b). For example, most CREBBP/EP300 acetylases consist of a catalytic HAT_KAT11 domain and two TAZ and ZZ zinc finger domains, but different lineages have acquired different reader domains: an acetylation-reading Bromo domain in holozoans and stramenopiles, PHD in plants and some stramenopiles, and no known reader domains in other lineages (e.g., in the fungal orthologs of theS. cerevisiae protein RTT109). A similar pattern is apparent in SET methyltransferase families sharing a core catalytic domain (SET) harboring variable DNA- and chromatin-interacting domains – animal SETDB1/2 homologs have MBD domains that bind CpG methylated DNA, while plants have SAD_SAR domains with the same function; and holozoan ASH1L homologs encode Bromo and BAH readers, whereas phaeophytes encode PHD domains (Extended Data Fig. 4b). Other architectures, however, are much more conserved, as exemplified by the presence of Tudor-knot and MYST zinc finger domains in most KAT5 deacetylases; or the ubiquitous co-occurrence of Helicase-C and SNF2_N domains in most remodellers (Extended Data Fig. 4b).

Specific examples of evolutionarily conserved chromatin gene families include the catalytic core and the subunits of well-studied chromatin complexes⁶³ like PRC1 (RING1/AB, PCGF), PRC2 (EZH1/2, SUZ12, EED, RBBP4/7) and Trithorax/MLL (MLL1/2/3/4, WRD5, ASH2L, RBBP5, DPY-30;Fig. 4d,e). However, when we compared the distribution of these complexes with the hPTMs they are related to, we found a generally poor co-occurrence (Fig. 4f-h). For example, organisms likeDictyostelium discoideum andCreolimax fragrantissima lack EZH1/2 orthologs, but we detected H3K27me3 in these species; whileThecamonas trahens andNaegleria gruberi lack Dot1 orthologs but have H3K79me marks. A poor correlation is also observed between the occurrence of H3K9me and that of SUV39H1 orthologs. An exception to this pattern is the ubiquitous distribution of H4K16ac and the acetylase family KAT5/8⁶⁴ (Fig. 4h). These patterns suggest that the specificity between hPTMs and their writers might not be completely conserved across eukaryotes, with distinct members of the same gene classes (e.g., methyltransferases) performing similar roles. In this context, reading domains present in writing/erasing enzymes (directly in the same protein or as part of multi-protein complexes) are likely to play a major role in the re-purposing of chromatin catalytic activities.

Evolutionary expansion of chromatin readers

Multiple protein structural domains have been involved in the recognition of hPTMs, such as Bromo domains binding to acetylated lysines or Chromo, MBT and Tudor domains binding to methylated lysines^23,24. These are generally small domains and can be found both as stand-alone proteins as well as in combination with other domains, often catalytic activities such as hPTM writers, erasers and remodellers. Thus, they are central in the establishment of functional connections between chromatin states. To understand the contribution of these reading domains to the evolutionary diversification of chromatin networks, we studied in detail the phylogeny and protein architecture of reader domains across eukaryotes.

We quantified the co-occurrence frequency of reader and catalytic domains, finding (i) that most reader domains are present in genes without writer, eraser or remodeller domains (87%,Fig. 5a); and (ii) that most cases of reader-catalytic co-occurrence involve PHD, Chromo and Bromo domains (Extended Data Fig. 5a). For example, the conserved architecture of the paneukaryotic CHD3/4/5 re-modellers includes Chromo readers in most species and PHD domains specifically in animals and plants (Extended Data Fig. 4b). Likewise, PHD domains are often present in the KMT2A/B and KMT2C/D SET methyltransfrase; and the ASH1L family has recruited Bromo and BAH domains in holozoans, and PHD in multicellular stramenopiles (Extended Data Fig. 4b). In spite of these redundancies, reader families typically have independent evolutionary histories, as illustrated by the fact that most reader domain-containing genes encode only one such domain (92%,Extended Data Fig. 5b).

We next performed phylogenetic analyses of individual reader domains and reconstructed the gains and losses of these reader gene families/orthogroups (Fig. 5a). Compared to the relative stasis of catalytic enzyme families, this reader-centric analysis revealed a strikingly different evolutionary pattern of lineage-specific bursts of innovation, particularly amongst PHD, Chromo and Bromo genes, as well as Tudor in animals (Fig. 5a andExtended Data Fig. 5c). PHD, Chromo and Bromo families also appeared as the most abundant in the reconstructed LECA reader domain repertoire, which amounted to 89 gene families (Fig. 5a, left). The distribution of gene family ages in extant species also corroborates that more readers have emerged at evolutionarily more recent nodes of the tree of life than catalytic gene families (Fig. 5b).

Co-option of chromatin machinery by transposable elements

Further examination of the domain co-occurrence networks of readers revealed that Chromo and PHD domains are often present together with protein domains found in transposable elements (TEs;Fig. 5c andSupplementary Data 6), including retrotransposons (e.g., retrotranscriptases and integrases; orange modules inFig. 5c) and DNA transposons (e.g., DNA binding domains and transposases; red modules). It is known that some TEs show insertion-preferences associated to specific chromatin states⁶⁵, often mediated by direct chromatin tethering mechanisms⁶⁶. For example, the Chromo domain of the MAGGY gypsy retrotransposon of the fungusMagnaporthe grisea targets H3K9me regions⁶⁷. Reciprocally, some protein domains of TE origin, often DNA-binding domains, have been co-opted into chromatin and transcriptional regulators⁶⁸. Thus, we decided to explore in detail the occurrence of chromatin-associated domain (readers, but also catalytic domains) linked to TEs in the 172 eukaryotic genomes in our dataset (Fig. 5d). Moreover, we used available RNA-seq datasets in many of these species to validate some of these TE fusions (Fig. 5d-e). A fully validated fusion gene would (i) come from a non-discontinuous gene model in the original assembly, and (ii) have evidence of expression, with reads mapping along the entire region between the TE-associated domain and the chromatin-associated domain (Extended Data Fig. S6).

We identified 823 predicted gene models containing both chromatin- and TE-associated domains (Fig. 5d). Whilst these TE fusions were not exclusive of reader domains, most such fusions involved PHD and Chromo-encoding genes; followed by SNF2_N remodellers, SET methyltransferases, and others. An homology search against a database of eukaryotic TEs revealed that most of these candidate TE fusions could be aligned to known retrotransposons or DNA transposons. For example, by way of validation, our analysis identifies the SETMAR human gene, a previously-described fusion between a SET methyltranferase and a Mariner-class DNA transposon⁶⁹. Overall, 31% of the candidate fusion genes were supported by valid gene models according to our stringent criteria (Fig. 5d). Interestingly, we find very few cases of hypothetical fusions between TEs and Bromo domains, which recognize K acetylations and are otherwise highly abundant across eukaryotes, and none of them is validated by RNA-seq data. This could be explained by the detrimental effect of targeting TE insertions to sites of active chromatin demarcated by histone acetylations, such as promoter and enhancer elements.

Some of these validated fusions have a broad phylogenetic distribution (Fig. 5e), such as a Gypsy-ERV retrotransposon with aC-terminal Chromo domain (Unk. Chromo 2.1 inFig. 5e) that is widely distributed in animals and various microbial eukaryotes, and contains dozens of paralogs in vertebrateDanio rerio or the charophyteChara braunii, many of which are expressed. Another widespread Gypsy-ERV retrotransposon with a Chromo domain is present in multiple expressed and highly similar copies in the fungusRhizopus delemar (Fig. 5f,e), suggesting a successful colonization of this genome by this TE. By contrast, other TE fusions are taxonomically restricted to one or few related species, such as the fusion of hAT activator DNA transposons with Chromo CBX and CDY readers in the spongeEphydatia muelleri; or multiple instances of fusions with Chromo and PHD readers in cnidarians. A common fusion in cnidarians involves different retrotransposon classes with PHD domains orthologous to the PYGO1/2 protein (Fig. 5e), which is known to recognize specifically H3K4me⁷⁰. Globally, this analysis reveals that recruitment of chromatin reading and even modifying domains by TE has occurred in many eukaryotic species, in a way that might facilitate the evasion from suppressing mechanisms in the host genomes as suggested by the expansion of Chromo-fused TEs in the genomes ofChara braunii (Viridiplantae),Chromera velia (Alveolata) andRhizopus delemar (Fungi).

Chromatin components in viral genomes

In addition to TEs, chromatin is also involved in the suppression of another type of genomic parasites: viruses. Some chromatin-related genes, including histones, have been found in viral genomes, especially among the nucleocytoplasmic large DNA viruses – also known as giant viruses. Eukaryotic core histones have been even hypothesized to have evolved from giant virus homologs, after the discovery that certain Marseilleviridae genomes encoded deeply-diverging orthologs of the four canonical histones⁷¹. These viral histones have been recently shown to form nucleosome-like particles that package viral DNA^72,73.

We analyzed the distribution and abundance of chromatin-related protein domains among viruses, including data from 1,816 giant virus genomes. Based on structural domain searches, we identified 2,163 viral chromatin-related proteins (Fig. 5g andSupplementary Data 6). The majority of these proteins are encoded by giant viruses (55%), followed by Caudovirales (37%). Among these two groups, only giant virus genomes encode histones – specifically, the Iridoviridae, Marseilleviridae, Mimiviridae, Pithoviridae, and Phycodnaviridae families. Concordantly with previous studies⁷⁴, we also identify remodellers in all giant virus families; as well as less abundant components of the chromatin writer/eraser/reader toolkit (Fig. 5g).

We then investigated the phylogenetic affinities of these viral chromatin proteins, starting with histones (Fig. 5h). Our analysis recovers the phylogenetic affinity of Marseilleviridae histones with specific eukaryotic histone families⁷¹, and makes this pattern extensive to Mimiviridae, Iridoviridae, and Pithoviridae giant viruses (Fig. 5h), with the caveat of the ambiguous clustering of the H4-like viral histones with either H4 eukaryotic or archaeal HMfB genes. In all these lineages, we identify genes encoding two histone-fold domains orthologous to H2B + H2A (inset table inFig. 5h), whereas the H4 + H3 histone doublet genes appears to be exclusive to Marseilleviridae. By contrast, histone homologs in Phycodnaviridae, Pandoraviridae (also giant viruses), and Polydnaviridae (incertae sedis) are never found as either doublets or as early-branching homologs of eukaryotic histones, suggesting recent acquisition from eukaryotes.

Unlike histones, most of the viral chromatin-associated genes exhibited a mixture of prokaryotic and eukaryotic phylogenetic affinities and often lack affinity to any specific eukaryotic gene family (Fig. 5i andExtended Data Fig. 7). Viral readers, on the other hand, are often embedded within eukaryotic clades in gene trees and are similar tobona fide eukaryotic families, exhibiting topologies consistent with recent, secondary acquisitions. This is the case of BIRC2/3/XIAP readers widespread in the Baculoviridae, which encode BIR domains that are often hijacked from their hosts⁷⁵. We also find a number of viral Chromo-encoding genes, which fall in two main taxonomic categories: (i) giant virus homologs of the eukaryotic CBX1/3/5 family (present in Mimiviridae, Iridoviridae and Phycodnaviridae); and (ii) homologs from various Adintoviridae, which are closely related to animal Chromo genes encodingrve integrase domains⁷⁶ (Fig. 5i). Finally, we also identify a handful of eukaryotic-like viral genes with deep-branching positions relative to core eukaryotic gene families, as seen in histones (Fig. 5h). This includes Mimiviridae homologs of the eukaryotic methyltransferases SMYD1-5 and DOT1 (Extended Data Fig. 7d,e), as well as SNF remodeller families with homologs in distinct giant virus clades (HLTF/TTF2 in Phycodnaviridae, Mimiviridae and Iridoviridae). These results indicate that cases of horizontal transfer from eukaryotes to viruses are common in different chromatin-related gene families, including histones. Therefore, it is likely that basally-branching giant virus histones were similarly acquired from a stem eukaryotic lineage and this would explain the observed histone tree topology with extant eukaryotic species. In any case, most of the eukaryotic chromatin machinery appears to have cellular roots.

Eukaryotic cell culture and tissue sources

Capsaspora owczarzaki strain ATCC30864 filopodial cells were grown axenically in 5 ml flasks with ATCC medium 1034 (modified PYNFH medium) in an incubator at 23ºC (Sebé-Pedrós et al., 2013a).

Corallochytrium limacisporum strain India was axenically grown in Difco Marine Broth medium at 23ºC,Creolimax fragrantissima strain CH2 was axenically grown in Difco Marine Broth medium at 12ºC,Spizellomyces punctatus strain DAOM BR117 was axenically grown in (0,5% yeast extract, 3% glycerol,1g/L K₂HPO₄, 0,5% EtOH) medium at 17ºC,Thecamonas trahens strain ATCC50062 was grown in ATCC medium: 1525 Seawater 802 medium,Chlamydomonas reinhardtii strain CC-503 cw92 mt+ was axenically grown in Gibco TAP medium at 29ºC,Guillardia theta strain CCMP2712 was axenically grown in L1+500uM NH₄Cl medium at 18ºC,Emiliania huxleyi strain CCMP1516 was grown in L1-Si medium at 18ºC,Thalassiosira pseudonana strain CCMP1335 was axenically grown in L1 medium at 18ºC,Bigelowiella natans strain CCMP2755 was axenically grown in L1-Si medium at 23ºC,Naegleria gruberi strain ATCC30224 was axenically grown in ATCC medium 1034 (modified PYNFH medium) at 29ºC,Gefionella okellyi strain 249 was grown in 15% Water Complete Cereal Grass Media (WC-CGM3) at 18ºC and Fabomonas tropica strain NYK3C was grown in L1 + YT medium at 18ºC. All cells were grown in 250 ml culture flasks.

In addition, we used frozen tissues/cells from the following species:Homo sapiens (ES cells, courtesy of Cecilia Ballaré, CRG),Physcomitrella patens (strain Gransden 2004, vegetative stage, courtesy of Josep Casacuberta, CRAG-CSIC),Sycon ciliatum (adult sponges sampled from Bergen, Norway, courtesy of Maja Adamska, ANU) andPhytophthora infestans (strain T30-4, courtesy of Harold J.G.Meijer, Wageningen University).

Archaeal cell culture

Cultures ofMethanobrevibacter cuticularis DSM 11139,Methanospirillum stamsii DSM 26304 andMethanosarcina spelaei DSM 26047 were purchased from the Deutsche Stammsammlung von Mikroorganismen und Zellkulturen GmbH (DSMZ), Braunschweig, Germany. Cultures were grown in closed batch in 50mL of defined media in 120mL serum bottles (La-Pha-Pack, Langerwehe, Germany). Growth was monitored as OD (600 nm; Analytik Jena, Specord 200 plus).Methanobrevibacter-cuticularis was grown in modifiedMethanobrevibacter cuticularis medium DSMZ 734a (DSMZ 2014) omitting bovine rumen fluid, yeast extract and Na-resazurin at 1.5 bar overpressure H₂CO₂ (20 vol.-% CO2 in H₂) at 37°C. As soon as a change in OD was observed, a constant agitation at 90rpm was applied.Methanospirillum stamsii was grown in modifiedMethanobacterium medium DSMZ 119 (DSMZ 2017) omitting sludge fluid, yeast extract and Na-resazurin at 1 bar overpressure H₂CO₂ (20 vol.-% CO2 in H₂) at 29°C, under constant agitation at 90rpm.Methanosarcina spelaei was grown in modifiedMethanosarcina barkeri medium DSMZ 120a (DSMZ 2014) omitting yeast extract and Na-resazurin at 1.5 bar overpressure H₂CO₂ (20 vol.-% CO2 in H₂) at 33°C, under constant agitation at 90rpm. All gases were obtained from Air Liquide GmbH, Schwechat, Austria.Nitrososphaera viennensis EN76 was grown in continuous culture in a bioreactor as previously described¹⁰⁰.

Cells were harvested via centrifugation at 21,000xg 4°C 1h (Thermo scientific, Sorvall Lynx 4000 centrifuge), the supernatant discarded and the resulting pellet resuspended in 1ml of spent medium, followed by another round of centrifugation at 21,000xg 4°C for 1h (Eppendorf, Centrifuge 5424R). Pellets were stored at -70°C. All archaeal histones were extracted as described below.

Histone acid extraction

Starting material was a pellet of 50-100M cells (washed once with cold PBS) or a flash-frozen tissue homogenate in liquid nitrogen using a ceramic mortar grinder. Cells were washed first in 10ml of buffer I (10 mM TrisHCl pH 8, 10 mM MgCl₂, 0.4M Sucrose). After 5min incubation, samples were centrifuged at 8.000g for 20min at 4°C and supernatant was removed. The resulting pellet was resuspended in 1.5ml of Buffer II (10 mM TrisHCl pH 8, 10 mM MgCl₂, 0.25M Sucrose, 1% Triton X-100, 1% Igepal Ca-630) and incubated 15min on ice. In specific cases, cells at this stage were broken using a 2ml Dounce homogenizer (with Pestle B) or with a 20G syringe. Then samples were centrifuged at 15.000g for 10min at 4°C and supernatant was removed. The resulting pellet was then slowly resuspended in 300μL of Buffer III (10 mM TrisHCl pH 8, 2 mM MgCl₂, 1.7M Sucrose, 1% Triton X-100) and then resulting resuspended nuclei were layered on top of another 300μL of Buffer III. Sample was centrifuged at 20.000g for 1h at 4°C and supernatant was removed, resulting in a nuclear pellet ready for acid histone extraction. All buffers were supplemented with spermidine (1:1000), beta-mercaptoethanol (1:1000), protease inhibitors (1x cOmplete cocktail Roche #11697498001, 1mM PMSF, 1:2000 Pepstatin), phosphatase inhibitors (1x phoSTOP cocktail Roche #4906845001) and deacetylase inhibitors (10mM Sodium butyrate).

For samples processed using a high-salt + HCl extraction protocol^101,102, the pellet was resuspended in 500μL of High Salt Extraction Buffer (20 mM TrisHCl pH 7.4, CaCl₂ 1M and protease, phosphatase and deacetylase inhibitors, same as above). Sample was incubated on ice for 30min and then pure HCl has added to a final 0.3N concentration (12.82μL to the initial 500μL). Samples were incubated for at least 2h on a rotor at 4°C and then centrifuged at 16.000g for 10min at 4°C to remove cellular/nuclear debris. The resulting supernatant containing solubilized histones was transferred to a clean 1.5ml tube and Trichloroacetic Acid (TCA) was added drop-wise to 25% final concentration (171μL TCA to an approximate initial 513μL sample) and left overnight at 4°C to precipitate histones. Samples were then centrifuged at 20.000g for 30min at 4°C and the supernatant removed. The pellet was then washed twice with 500μL of cold acetone and then dried for 20min at room temperature. Finally, clean histone pellets were resuspended in 30-50μL of ultrapure water. Protein concentration in the sample was measured using BCA and extraction was examined using an SDS-PAGE protein gel with Coomassie staining.

For samples processed using H₂SO₄¹⁰², the protocol was exactly the same except that 400μL 0.4N H₂SO₄ (freshly diluted) was used instead, with a similar incubation time of at least 2h at 4°C.

Histone chemical derivatization

Histones samples were quantified by the BCA method and 10 μg of each sample were derivatized with propionic anhydride, digested with trypsin and derivatized again with phenylisocyanate as previously described⁴⁹. Briefly, samples were dissolved in 9 μL of H2O and 1 μL of triethyl ammonium bicarbonate was added to bring the pH to 8.5. The propionic anhydride was prepared by adding 1 μL of propionic anhydride to 99 μL of H2O and 1 μL of propionic anhydride solution was added immediately to the samples with vortexing and incubation for 2 minutes. The reaction was quenched with 1 μL of 80mM hydroxylamine and samples were incubated at room temperature for 20 minutes. Tryptic digestion was performed for 3 h with 0.1 μg trypsin (Promega Sequencing Grade; Madison, WI) per sample. A 1% v/v solution of phenyl isocyanate (PIC) in acetonitrile was freshly prepared and 3 μl added to each sample (17 mM final concentration) and incubated for 60 min at 37 °C. Samples were acidified by adding 50 μL of 5% formic acid, vacuum dried and desalted with C18 ultramicrospin columns (The Nest Group, Inc, Southborough, MA).

Liquid Chromatography-Tandem Mass Spectrometry Sample Acquisition

A 2-μg aliquot of the peptide mixture was analyzed using a LTQ-Orbitrap Fusion Lumos mass spec-trometer (Thermo Fisher Scientific, San Jose, CA) coupled to an EASY-nLC 1000 (Thermo Fisher Scientific, San Jose, CA) with both collision induced dissociation (CID) and high energy collision dissociation (HCD) fragmentation.

Peptides were loaded directly onto the analytical column and were separated by reversed-phase chromatography using a 50-cm column with an inner diameter of 75 μm, packed with 2 μm C18 particles spectrometer (Thermo Scientific, San Jose, CA, USA) with a 90 min chromatographic gradient. The mass spectrometer was operated in positive ionization mode using a data dependent acquisition method. The “Top Speed” acquisition algorithm determined the number of selected precursor ions for fragmentation.

Mass-spectrometry Data Analysis

Acquired data were analyzed using the Proteome Discoverer software suite (v2.0, Thermo Fisher Scientific), and the Mascot search engine (v2.6, Matrix Science¹⁰³) was used for peptide identification using a double-search strategy. First, data were searched against each organism protein database plus the most common contaminants considering Propionylation onN-terminal, Propionylation on Lysines and Phenylisocyanate onN-terminal as variable modifications. Then a new database was generated with the proteins identified in the first search,, and a second search was done considering Propionylation onN-terminal, Propionylation on Lysines, Phenylisocyanate onN-terminal, Dimethyl lysine, trimethyl lysine, propionyl + methyl lysine, acetyl lysine, crotonyl lysine as variable modifications. Precursor ion mass tolerance of 7 ppm at the MS1 level was used, and up to 5 missed cleavages for trypsin were allowed. False discovery rate (FDR) in peptide identification was set to a maximum of 5%. The identified peptides were filtered by mascot ion score higher than 20 and only PTMs with a localization score ptmRS¹⁰⁴ higher than 45 were considered. The raw proteomics data have been deposited to the PRIDE¹⁰⁵ repository with the dataset identifier PXD031991.

Analysis of hPTM conservation

Comparative genomics analysis of chromatin-associated proteins

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Supplementary Materials

Data Availability Statement

The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD031991.

Code for reproducing the analysis is available in our lab Github repository (https://github.com/sebepedroslab/chromatin-evolution-analysis).