Histone post-translational modifications — cause and consequence of genome function

Depósito de investigación de la Universidad de Sevilla https://idus.us.es/ “This version of the article has been accepted for publication, after peer review (when applicable) and is subject to Springer Nature’s AM terms of use, but is not the Version of Record and does not reflect post-acceptance improvements, or any corrections. The Version of Record is available online at: https://doi.org/10.1038/s41576-022-00468-7 ” The give and take of histone modifications in 1 genome function 2 3 Gonzalo Millán-Zambrano1,2,7, Adam Burton3,7, Andrew J. Bannister4† and Robert 4 Schneider5,6† 5 6 1Centro Andaluz de Biología Molecular y Medicina Regenerativa CABIMER, 7 Universidad de Sevilla-CSIC-Universidad Pablo de Olavide, Seville, Spain 8 2Departamento de Genética, Facultad de Biología, Universidad de Sevilla, Seville, 9 Spain 10 3Institute for Epigenetics and Stem Cells, Helmholtz Center Munich, Germany 11 4Gurdon Institute and Department of Pathology, University of Cambridge, UK 12 5Institute for Functional Epigenetics, Helmholtz Center Munich, Germany 13 6Faculty of Biology LMU Munich, Germany 14 7These authors contributed equally: Gonzalo Millán-Zambrano, Adam Burton 15 †email: [email protected]; [email protected] 16 17 Abstract 18 Over the past decades, much has been learned about histone post-translational 19 modifications (PTMs) and how they affect DNA-templated processes at the molecular 20 level. This understanding has been bolstered recently by the identification of new types 21 of histone PTMs, the advent of new genome-wide mapping approaches, and methods 22 to locally and acutely deposit or remove PTMs. Now, with the availability of vast 23 amounts of data across various biological systems, the functional role of PTMs in key 24 biological processes, such as transcription, recombination, replication, repair and 25 genome architecture, is slowly emerging. Here, we review recent advances in the field, 26 which have taught us that histone PTMs can both have direct effects, with a more 27 causative function, as well as indirect effects, often acting as reinforcers and 28 bookmarkers of epigenetic events in a complex regulatory network. 29 30 Introduction 31 Virtually every cell in the human body contains the same genetic information encoded 32 within approximately 2 m of linear DNA. This presents a huge organizational problem; 33 how to package this length of DNA within a relatively small nuclear volume, whilst 34 maintaining its accessibility in a spatially and temporally coordinated manner. The 35 solution is achieved by the DNA associating with proteins, predominantly conserved 36 histone proteins, to form a complex structure referred to as chromatin. The generally 37 accepted view is that chromatinised DNA is ultimately folded into higher-order relatively 38 stable chromosomal structures, which need to be decondensed to facilitate DNA 39 templated processes. However, the actual long range spatial and temporal organization 40 of chromatin in vivo appears far more complex and variable than this simple vision, with 41 recent data pointing towards a continuously changing architectural landscape in which 42 different chromatin states are continuously morphing and inter-converting (reviewed in 43 1). Thus, chromatin represents much more than a mere inert packaging structure; it is a 44 dynamic scaffold capable of responding to various cues to regulate DNA accessibility to 45 different cellular machinery. 46 The fundamental unit of chromatin is the nucleosome, consisting of a central 47 histone octamer (two each of histones H2A, H2B, H3 and H4) around which 48 approximately 1.75 left-handed superhelical turns of DNA are wound2. The histones are 49 decorated by a plethora of post-translational modifications (PTMs) (Fig. 1). These PTMs 50 are often referred to as ‘epigenetic marks’ and serve to regulate chromatin structure and 51 hence DNA-templated processes, such as transcription, recombination, replication or 52 repair. It has been suggested that they serve as a ‘code’, in which “all have their own 53 message to convey”3. 54 The histone modification landscape is laid down, maintained and reset by 55 numerous interconnected signalling pathways, involving enzymes that catalyse specific 56 modifications (the ‘writers’), proteins recognizing them via specific domains (the 57 ‘readers’), and enzymes removing them (the ‘erasers’). Many of the enzymes involved 58 rely upon cellular cofactors, intimately linking their activity with cellular metabolic state4, 59 5. Adding to this complexity, numerous variants of histones exist, depending on the 60 species, which can also be differentially modified6. Their importance is underscored by 61 the discovery that modified residues in histones are mutated in cancer, where they can 62 act as oncogenic drivers7. 63 Histone PTMs exert their effects via a multitude of mechanisms (Fig. 2), either 64 directly, such as by local, structural perturbation, or indirectly, by recruitment of effector 65 complexes (Fig. 2a). It was long thought that some histone modifications, such as lysine 66 acetylation, are highly dynamic, whereas others, such as lysine methylation, are more 67 stable8. Nowadays, it is appreciated that most, if not all, histone modifications are 68 reversible, at least to some extent; in many cases, both a writer and an eraser of the 69 PTM have been identified. Indeed, it is now evident that the balance of activity (i.e. the 70 equilibrium) between writers and erasers is critical in establishing biological output 71 (reviewed in 9). 72 Fundamentally, the histone modification landscape is established and maintained 73 by highly regulated, spatially and temporally coordinated, recruitment of the various 74 enzymes to specific regions of the genome (Fig. 2b). Technological advances (Table 1), 75 such as chromatin immunoprecipitation followed by sequencing (ChIP–seq), have been 76 instrumental in demonstrating that specific functional genomic regions, such as active 77 genes and enhancers, are enriched with particular patterns of modifications. In a similar 78 manner, broader regions of the genome, such as permissive euchromatin and its 79 antithesis, repressive heterochromatin, display their own characteristic modification 80 profiles. On a global scale, both euchromatin and heterochromatin regions are 81 partitioned into 3D topologically associating domains (TADs) 10, 11. Recent work has 82 shown that genomic organization at various levels, for example, heterochromatin 83 domains and super-enhancers with their target promoters, possibly involves the 84 formation of biomolecular condensates via phase transition (Box 1). A better 85 understanding of how histone modifications are related to these processes and 86 structures is required and will be fundamental to understanding their true roles in the 87 cell. 88 In this Review, by citing examples across species, we discuss how histone 89 modifications are affected, regulated and interdependent on different DNA processes, 90 genome topology and other epigenetic events. Focusing on transcription, we critically 91 assess whether histone modifications can be viewed as mechanistically instructive cues 92 for different nuclear machineries, or whether they should be considered critical 93 components of a regulatory network, bolstering already active processes and/or as a 94 mechanism of recording genomic events. 95 96 [H1] Histone PTMs in transcription 97 Histone PTMs are frequently enriched at distinct genomic locations, particularly at 98 genes, where they have been correlated, either positively or negatively, with 99 transcription. Indeed, the association between histone modifications and gene 100 expression was first documented over 50 years ago, when seminal work demonstrated 101 that histone acetylation can greatly suppress the inhibitory effect of histone 102 incorporation into DNA in vitro12. This effect is mainly due to neutralization of the 103 positive charge of lysine residues, and thus the basic properties of histones, and can 104 lead to a less compact chromatin structure13-15. Consequently, histone acetylation 105 generally correlates with transcriptional activity and is enriched on active promoters and 106 enhancers and other accessible regions of chromatin16. Importantly, acetylation can 107 directly increase the rate of transcription in vitro17, 18. It is thought that histone 108 acetylations act in a cumulative and redundant manner, as removal of single histone tail 109 acetylations generally has limited effects on transcription19. In addition to acetylation, 110 numerous longer chain acylations also occur on histone lysines (Fig.1). These also 111 generally correlate with transcription, although their abundance is typically much lower 112 than acetylation20. One of the best studied examples is histone crotonylation21, which 113 was originally identified as a positive regulator of transcription22. Interestingly, however, 114 crotonylation has also more recently been implicated in the repression of gene 115 expression in yeast23. This apparent dichotomy could be explained by the recruitment of 116 different binding proteins. 117 Early studies of histone bulk methylation did not correlate it with transcription24, 25. 118 However, it is now known that the effect of histone methylation is more complex, 119 occurring dynamically in three distinct states on both arginine (Rme1, me2 asymmetric, 120 me2 symmetric) and lysine (Kme1, me2, me3) sidechains. Moreover, the effects are 121 site-specific and unlikely to directly affect nucleosome structure. 122 However, despite much research, a causal role for most histone modifications in 123 regulating transcription still lacks support26. Below, we consider the cause or 124 consequential nature of different histone PTMs in transcription through various 125 processes. 126 127 [H2] Histone tail PTMs in transcription 128 One of the most well-characterized histone modifications associated with transcription is 129 histone 3 lysine 4 trimethylation (H3K4me3). It is enriched in the promoters of most 130 active genes in eukaryotes, peaking around the transcription start site (TSS)27, where 131 both its peak strength and breadth correlate with transcription28, 29. The mark enables 132 recruitment of transcriptional machinery and thus potentially facilitates transcription30. In 133 fact, it has been proposed that H3K4me3-dependent TAF4 recruitment is important for 134 the expression of select p53 target genes31. However, evidence from functional 135 experiments in numerous systems suggests H3K4me3 is not required for the majority of 136 transcription26. By contrast, local targeting of H3K4me3 modestly activates gene 137 expression in a strictly context-dependent manner32. Thus, the precise role and a 138 causative function of the remarkably conserved H3K4me3 enrichment at most active 139 promoters remains unclear. It is likely though that deposition of this mark reinforces 140 transcription, as transcription-dependent recruitment of H3K4me3-methylating 141 complexes has been demonstrated in several systems27. Another possibility is that 142 H3K4me3 influences transcriptional consistency, as H3K4me3 anti-correlates with 143 noise, in contrast to H3K4me1/229, 33. 144 In mammals, H3K4me3 can be maintained during transcriptional quiescent 145 periods, such as in mature oocytes and sperm and in fertilized embryos prior to genome 146 activation34-36. This phenomenon is consistent with a longer-term function, downstream 147 of gene expression, such as in epigenetic memory, a function originally suggested for 148 the Trithorax (Trx) H3K4 methyltransferase complex in Drosophila37, 38. This hypothesis 149 has gained traction from reprogramming work in Xenopus showing H3K4me3 is 150 necessary for memory of active transcriptional states39. In mammals, H3K4me3 at 151 retained nucleosomes in sperm seems to be important for establishing gene expression 152 patterns and development in the resulting embryos40, 41. This seems unlikely to be a 153 feature unique to H3K4me, since other marks are also present on embryonic chromatin 154 prior to transcriptional activation, potentially suggesting a more widespread role for 155 histone PTMs in epigenetic inheritance (Fig. 3a). 156 Active enhancers are marked with H3K4me1 and H3K27ac in a cell-type specific 157 manner42, 43. Interestingly, however, the writers of H3K4me1, mammalian MLL3/4 or Trx 158 in Drosophila, seem to be more important than the mark itself for enhancer activity, as 159 loss of H3K4me1 results in only minor effects on gene activity and development44, 45. 160 Similarly, H3K27ac also seems to be dispensable for enhancer activity in mouse cells46. 161 Interestingly, persistence of H3K4me1 at enhancers has recently been shown to be 162 important to maintain germline competence in a primordial germ cell culture model, 163 suggesting a more general role for H3K4me in epigenetic inheritance47. 164 It may be that H3K4me is more related to low levels of DNA methylation at CpG-165 rich sequences than transcription. H3K4me is generally found in a mutually exclusive 166 manner with DNA methylation, irrespective of transcription, and is recruited to an 167 engineered hypomethylated CpG island without promoters or transcription48. Prior to 168 transcriptional activation in Xenopus and Zebrafish embryos and mouse oocytes, 169 H3K4me is enriched on hypomethylated regions of the genome34, 35, 49, 50. In particular, 170 H3K4me3 in mature mouse oocytes shows a non-canonical distribution (ncH3K4me3) 171 including unusually broad domains that cover around one fifth of the genome34, 35, 51, 52 172 and erasure of ncH3K4me3 after fertilization is required for zygotic genome activation34 173 (Fig. 3b). As methylation of H3K4 strongly impairs binding of the essential de novo DNA 174 methyltransferases to chromatin, H3K4me may protect genomic regions from 175 inappropriate DNA methylation, particularly during development50, 53. By contrast, DNA 176 methylation also impairs the binding of H3K4me3 writers to chromatin, and deletion of 177 DNA methyltransferases in mouse oocytes results in the acquisition of H3K4me3 in 178 previously DNA methylated gene bodies rich in CpG52. Therefore, a highly conserved 179 negative feedback relationship between H3K4me3 and DNA methylation is likely to be 180 important in defining the genomic distribution of these marks, although the functional 181 role of this feedback relationship requires further investigation. 182 H3K36me3 is tightly correlated to actively transcribed regions, due to recruitment 183 of the H3K36 methyltransferase Setd2 by elongating RNA polymerase II54-56. While 184 HK36me3 does not seem to be required for transcriptional elongation, it functions to 185 inhibit cryptic transcription via deacetylation of histones and DNA methylation57, 186 regulates splicing58 and guides co-transcriptional N6-methyladenosine (m6A) 187 methylation of mRNA59 across phyla (see also Section 3). However, flies bearing 188 H3K36R mutations display widespread changes in gene expression, likely due to post-189 transcriptional effects rather than alternative splicing or suppression of cryptic 190 transcription60. In humans, H3K36me2 is enriched on multi-megabase scale domains 191 and it also interplays with DNA methylation in shaping the intergenic DNA methylation 192 landscape61, 62. Thus, while H3K36me is strongly associated with the post-193 transcriptional control of gene expression and DNA methylation, more research is 194 required to dissect the precise contribution of the methylation itself in different 195 organisms. 196 Dynamic histone phosphorylation, like acetylation, reduces the basic charge of 197 histones and is similarly thought to facilitate transcription. For instance, H3Y41ph, which 198 is present at a subset of actively transcribed genes 63, directly increases DNA 199 accessibility in vitro by enhancing nucleosome unwrapping 64. In addition, histone 200 phosphorylation has been shown to promote histone acetylation65, 66. For example, 201 phosphorylation of Ser31, which is unique to H3.3 and was previously considered a 202 mitosis-specific mark67, enhances interphase stimulus-dependent transcription, at least 203 in part by stimulating the catalytic activity of both the histone acetyltransferase p300 and 204 methyltransferase SETD2, and is important for Xenopus development68-70. 205 Phosphorylation of H3S10 also promotes binding of specific factors whilst inhibiting 206 others. For example, the 14-3-3 protein is inducibly recruited, via direct binding of 207 H3S10ph, to the c-Jun promoter upon its induction71. In contrast, the same modification 208 prevents binding of the INHAT (inhibitor of acetyltransferases) repressor to H3, at least 209 in vitro72. Similarly H3S10ph and H3S28ph counteract binding of HP1 and PcG proteins 210 to the repressive histone modifications H3K9me3 and H3K27me3, respectively, through 211 ‘phospho–methyl switches’73, 74 (Fig. 2b). Although displacement of PcG proteins is 212 likely linked to gene activation 74, the role of H3S10ph-dependent displacement of HP1 213 in transcription, especially from constitutive heterochromatin regions, is less clear. 214 Nevertheless, when present within specific promoters, histone phosphorylation can act 215 as a facilitator of transcription, predominantly via the recruitment or exclusion of specific 216 effector proteins (Fig. 2a). 217 H3K27me3 and H2Aub are intimately connected features of facultative 218 heterochromatin, and are produced by distinct activities of the Polycomb complexes 219 PRC2 and PRC1, respectively75. Whilst it is clear that Polycomb complexes are crucial 220 to maintain repression of cell-type specific genes76, the contribution of these two histone 221 modifications to silencing is still not fully understood77. It seems that H3K27me3 is 222 crucial for silencing, as the point mutation of H3K27 phenocopies the PRC2 knockout in 223 Drosophila78. By contrast, the role of H2Aub in gene silencing is less clear79-81. PRC1 224 also induces chromatin compaction independently of H2Aub, which may be linked to its 225 recently described role in phase separation, although the evidence for both activities in 226 vivo remains fairly weak82-85. Furthermore, while H3K27me3 and H2Aub are strongly 227 associated in most cell types, an unusual decoupling of these marks has recently been 228 revealed in early mammalian development, in which H2Aub is enriched at Polycomb 229 targets before H3K27me3, while H3K27me3 plays a role in non-canonical genomic 230 imprinting86-88. By contrast, inheritance of H3K27me3 in Drosophila is necessary to 231 establish silencing of developmental genes, such as the Hox cluster89, 90, and in 232 mammalian embryonic stem cells, repressed chromatin marked by H3K27me3 can be 233 inherited across cell division91. H3K27me3 has also been shown to be downstream of 234 transcription, as for example, inhibition of transcription results in recruitment of 235 H3K27me3 to classic Polycomb targets92. This finding again highlights the non-linear 236 relationship between histone modifications and transcription, demonstrating that they 237 form an integral part of a complex regulatory network. 238 H3K9me3, the classical marker of constitutive heterochromatin, is enriched on 239 transcriptionally silent regions of the genome93. For example, H3K9me3 marks 240 centromeric, telomeric and other repetitive sequences in diverse organisms, but also 241 silenced genes94. It is bound by heterochromatin protein 1 (HP1)95, 96, which contributes 242 to the compaction and repression of H3K9me3-marked regions, via self-oligomerization, 243 possibly phase transition and recruitment of other heterochromatic proteins, such as 244 histone deacetylases and writers of downstream histone methylation marks including 245 H3K56me3, H3K64me3 and H4K20me397-100. H3K9me3-marked regions occlude 246 replication initiation factor TRESLIN, regulating its recruitment to origins of replication148. 433 Thus, histone acetylation has an impact on both origin establishment and activity. 434 Regulation of histone H4K20me levels has also been shown to play a key role in 435 replication initiation across metazoan genomes. The bromo-adjacent homology (BAH) 436 domain of ORC1 binds H4K20me2, and mutating this domain reduces origin recognition 437 complex (ORC) binding to origins149. Consistent with this finding, artificial tethering of 438 the H4K20me1 methyltransferase PR-Set7 to a specific locus induces ORC binding in a 439 manner dependent on Suv4-20h1/2, the enzymes responsible for H4K20me2/me3150. 440 Moreover, PR-Set7 is targeted for proteasome degradation during S-phase, and de-441 regulation of its degradation results in DNA re-replication151. Interestingly, the 442 counteracting demethylase for H4K20me1, PHF8, is also cell-cycle regulated152. 443 Nonetheless, almost 80% of the total H4 is dimethylated at K20, and thus it is difficult to 444 envision it as a specific determinant of replication origin selection at the genome-wide 445 scale. Instead, it seems plausible that H4K20 methylation may function to stabilize ORC 446 on chromatin at already defined origins. In agreement with this hypothesis, it has 447 recently been suggested that H2A.Z regulates the selection and activation of early-448 replication origins by recruiting SUV420H1 to establish H4K20me2 at specific locations 449 (Fig. 4e)153. 450 Other histone methylations, and combinations thereof, have also been shown to 451 influence DNA replication. H3K4me3 and H3K9me3 demethylation by KDM5C and 452 KDM4D demethylases, respectively, are important for efficient DNA replication initiation 453 in different chromatin contexts154, 155. In particular, these demethylases are specifically 454 required for origin activation, but not establishment. Remarkably, the tudor domain of 455 KDM4 family of demethylases binds to H3K4me3, indicating that combinations of these 456 histone PTMs can exert a crosstalk that facilitates site-specific replication initiation156. 457 Hence, in addition to activating pathways, repressive mechanisms also play an integral 458 part in the regulation of the DNA replication programme. In line with this, KDM4A, which 459 is predominantly expressed during the G1/S transition, binds components of the 460 replication machinery, and its overexpression induces site-specific DNA re-461 replication157. 462 In higher eukaryotes, large chromosomal domains replicate in a characteristic 463 temporal order, establishing a replication timing (RT) programme. Importantly, it has 464 recently been shown that RT alteration causes replication-dependent disruption of 465 histone PTMs, including H3K9me3, H3K27ac and H3K4me3, and genome 466 compartmentalization158. Remarkably, TAD boundaries frequently demarcate 467 mammalian RT domains boundaries159. Together, these findings suggest that the timing 468 of chromatin replication is important for maintaining the global histone modification 469 landscape, which may affect the 3D genome architecture (see below). All in all, it is 470 becoming increasingly apparent that dynamic regulation of histone PTMs can play an 471 instructive role in the regulation of DNA replication, while, vice versa, DNA replication 472 can influence the histone modification landscape. 473 474 [H1] Histone PTMs and genome topology 475 Euchromatic and heterochromatic regions of animal and plant genomes are organized 476 into structurally distinct A and B compartments, respectively10. These compartments are 477 subdivided into TADs, whose anchor points are often binding sites for the CTCF 478 ‘insulator’ protein, which, in concert with cohesin, regulates their formation by chromatin 479 loop extrusion11. Evidence suggests that the formation of TADs antagonizes 480 compartment formation, as disruption of TADs by acute depletion of cohesin in various 481 systems leads to a strengthening of compartments 160. Despite the long-observed 482 correlation, it remains unclear whether histone modifications can actively regulate these 483 topological structures. 484 Studies using mouse cells with inverted (rod photoreceptors) and conventional 485 nuclear organization suggest that A/B compartmentalization is driven predominantly by 486 interactions between heterochromatic regions, probably via HP1-dependent phase 487 separation161. In line with this finding, dCas9-directed H3K9me3 in human cells 488 promotes chromatin anchoring to HP1 condensates and induces an extensive re-489 arrangement of existing chromatin compartments162. Moreover, mutant Caenorhabditis 490 elegans embryos lacking H3K9 methylation substantially lose compartmentalization163. 491 Similarly, mouse embryonic fibroblasts mutant for all six H3K9 methyltransferases no 492 longer maintain heterochromatin organization164. H3K9me2/3 is enriched in chromatin 493 associated with the nuclear periphery that forms lamina-associated domains (LADs), 494 which highly correlate with the B compartment165. Importantly, H3K9me2/3 is also 495 implicated in LADs positioning and inheritance through cell division166-168. Together, 496 these recent findings provide compelling indications for an important function of H3K9 497 methylation in 3D genome compartmentalization. 498 Growing evidence suggests that H3K27me3 also plays an important role in the 499 spatial organization of the genome. Super-resolution imaging experiments have shown 500 that polycomb-repressed chromatin domains adopt unique folded states169. Specifically, 501 PRC1 binding generates chromatin domains that are distinct in size and boundary 502 characteristics from TADs170. These domains, called polycomb-associating domains 503 (PADs), are highly prevalent in late-stage mouse oocytes, where they can occupy up to 504 half of the genome171. Similarly to compartments, PADs are also strengthened by 505 depletion of cohesin 171, 172. H3K27me3 has been shown to be important for the 506 formation, via the recruitment of PRC1, but not the maintenance of these chromatin 507 domains171. Consistently, acute depletion of MLL2, which leads to enhanced levels of 508 H3K27me3 at bivalent promoters, increases PRC1 occupancy and induces a 509 compartment switch of proximal regions173. Furthermore, in Arabidopsis thaliana, where 510 TAD-like structures are absent, alteration of H3K27me3 levels induces a strong 511 reconfiguration of chromatin repressive loops174. Thus, H3K27me3-driven facultative 512 heterochromatin formation also seems to be important for defining 3D genome 513 organization. 514 Our knowledge about the roles of histone PTMs in TAD formation is mainly 515 restricted to C. elegans, where TAD-like structures are formed on the X chromosome by 516 the dosage compensation complex (DCC)175. This complex, which includes a 517 H4K20me2 demethylase, is thought to drive TAD formation through a loop extrusion 518 mechanism, as has been proposed in mammals176. Importantly, selective inactivation of 519 the demethylase activity disrupts X chromosome conformation by diminishing the 520 formation of TADs177. By contrast, mutant C. elegans embryos lacking H3K9 521 methylation also display significantly weakened DCC-dependent TAD boundaries on the 522 X chromosome163. These findings, together with reports indicating that TAD boundaries 523 are disrupted in mammals by DNA methylation178, suggest that TAD formation may be 524 subjected to an intricate regulation by the interplay between DNA and histone 525 modifications. 526 527 [H1] Conclusion and perspectives 528 The vast portfolio of histone PTMs can integrate (take) signalling information into 529 chromatin to regulate access to and expression of DNA. The modifications are present 530 in both the terminal ‘tails’ of histones and the globular core domains. Most tail 531 modifications have little to no effect on the structure or stability of a mono-nucleosome 532 and consequently do not act directly, but rather by providing binding platforms for 533 downstream effectors (Fig. 2). Core modifications, on the other hand, often directly 534 perturb mono-nucleosome structure, facilitating more direct action (give). However, in 535 vivo, the overwhelming majority of nucleosomes are close to each other, sometimes 536 even in direct contact. Still, even in this organization, most histone tail modifications, 537 with the notable exception of H4K16ac, have little direct effect on chromatin structure, at 538 least as measured on nucleosomal arrays in vitro. Thus, fundamental mechanistic 539 differences exist between tail modifications and those in the core in how they exert their 540 actions. The situation is further complicated by the requirement for a continual dynamic 541 turnover of specific modifications, such as H3S10ph and H3K9ac9 in the promoters of 542 certain active genes, meaning effectors must ‘sense’ the modification flux. 543 But are histone PTMs cause or consequence of an associated biological effect, 544 such as transcription? It seems likely that core modifications generally serve a more 545 direct and instructive role by modifying nucleosome dynamics. Tail modifications, 546 however, seem to operate, depending on the modification, both instructively and as a 547 maintenance mechanism, whereby combinations of modifications reinforce or record a 548 particular functional state, as we have discussed throughout this Review. So, histone 549 PTMs can be a cause and consequence of DNA processes, can give and take, 550 depending on the chromatin and functional contexts. In general, they should be 551 considered as key components of a complex non-linear regulatory network, which 552 evolved to provide robustness and regulatory potential. Most excitingly, new 553 technologies, including the targeted recruitment of histone modifiers to individual loci 554 (Box 2), are beginning to enable better characterization of the causal influence of 555 histone PTMs on genomic processes. 556 An open question remains concerning the number of modifications yet to be 557 identified. The very recent and important discovery of novel types of acylation and 558 hormone-based (e.g., serotonylation and dopamination107, 108) modifications regulating 559 specific genes, for example, in the brain, highlights that the repertoire is still 560 expanding. Many of these recently identified dynamic modifications can be 561 enzymatically catalysed or occur due to the chemical reactivity of, for example, the 562 corresponding cofactors used by the modifying enzymes179. Furthermore, the continuing 563 improvement in our understanding of how the environment and metabolome affect and 564 regulate histone modifications is opening new avenues of research. Although these 565 lines of investigation are often challenging, they hold great potential for fundamental and 566 important discoveries in the future. 567 568 Acknowledgements 569 570 The R.S. laboratory is supported by the DFG through SFB 1064 (Project-ID 213249687) 571 and SFB 1309 (Project-ID 325871075), as well as the Helmholtz Gesellschaft. A.J.B is 572 supported by grants from Cancer Research UK (RG96894 and C6946/A24843) and the 573 Wellcome Trust (WT203144). G. M-Z is supported by a Juan de la Cierva postdoctoral 574 contract (IJC2019-039988) from the Spanish Ministry of Science and Innovation and a 575 fellowship from ”la Caixa”Foundation (ID 100010434) and from the European Union’s 576 Horizon 2020 research and innovation programme under the Marie Skłodowska-Curie 577 grant agreement No 847648”. The fellowship code is “LCF/BQ/PR21/11840007”. A.B. is 578 supported by the DFG through SFB 1064 (Project-ID 213249687), as well as the 579 Helmholtz Gesellschaft. The authors apologize to colleagues whose important studies 580 they were unable to cite due to space constraints. 581 582 Competing interests 583 The authors declare no competing interests. 584 585 Table 1. Selected omics techniques used to study the genomic localisation of 586 histone modifications 587 Approach Principle Pros Cons Chromatin immunoprecipitation followed by sequencing (ChIP– seq)180, 181 Cross-linked (e.g. formaldehyde) (X- ChIP) and sonication or micrococcal digestion OR native (N-ChIP) and micrococcal digestion Single-nucleotide resolution, possible with low input material (ULI-NChIP; 182). For many years it was the goldstandard, countless reference data-sets available, many ChIP grade antibodies commercially available X-ChIP well suited for transient interactions High background noise, standard protocols require high cellular input and high sequencingdepth X-ChIP: cross-linking can mask epitopes recognized by antibodies, time-intensive protocol Cleavage under targets & release using nuclease (CUT&RUN)183 Recombinant MNase fused to protein-A and/or -G binding to specific antibody against PTM of interest. Avoids cross-linking and fragmentation of DNA, low background noise, possible with low input, fast protocol, only low sequencing depth required, used for single cells184, CUT&RUN.ChIP185 to assess histone PTM cooccupancy. MNase digestion needs careful optimization, in addition to cleaving DNA next to PTM, MNase can also cleave DNA that is far away but close in 3D, often antibodies are only validated for X-ChIP, transient interactions might be missed Cleavage under targets and tagmentation (CUT&TAG)186 Recombinant Tn5 transposase fused to protein-A and/or -G binding to specific antibody against PTM of interest. No cross-linking and library preparation step, sensitive, easy workflow, low sequencing depth required, can be performed at single cell leve186 and used for multiple chromatin targets/PTMs in the same assay (MulTI-Tag; 187, 188. Tn5 enzyme biases: Tn5 preferentially tags accessible chromatin, potential background from mitochondrial DNA Chromatin integration labelling followed by sequencing (ChIL- seq)189 Combines immunostaining, transposase tagging and linear amplification for low-input PTM profiling. Applicable to very low cell numbers due to RNA-mediated linear amplification, can be combined with cell biology assays. Tn5 enzyme biases, requires fixation, relatively long protocol, poor mapping efficiency, necessitates high number of sequencing reads. Directed Methylation with Long-read sequencing (DiMeLoseq)190 Directs m6A DNA methylation to antibody-protein A fusion (pA-Hia5), coupled to long-read sequencing. Ability to map highly repetitive regions, concurrent protein/PTM mapping with DNA cytosine methylation, multiple binding events on single molecules can be determined. Low sensitivity, not applicable to low cell numbers. 588 Figure legends 589 Figure 1. Sites of PTMs within the 4 core histones. The single letter amino acid code 590 (numbered below) is shown for each histone, H2A, H2B, H3 and H4. Gaps in the 591 sequence are indicated by dots. Amino acids within central globular domains are 592 coloured ‘purple’. The relatively common modifications of methylation (me) and 593 phosphorylation (ph) are indicated in blue and purple text, respectively. Modifications of 594 lysine are grouped as indicated in the key: Acylations are annotated in red, ubiquitin-like 595 in green and other PTMs in black text. Non-lysine modifications are also grouped in the 596 key. Yeast H3Q105 and mammalian H3Q104 are equivalent (shown in red text). A 597 cartoon representation of a nucleosome is shown bottom right, which comprises 2 each 598 of the core histones (coloured quarters) around which the DNA (thick blue line) is 599 wrapped. 600 601 Figure 2. Mechanisms underlying histone PTM action and modification enzyme 602 recruitment. A | Histone PTMs (red filled circles) can act directly, for example, by 603 affecting nucleosome–nucleosome interactions (top), or indirectly, via binding of reader 604 proteins (blue shape) or by repelling them (green shape) (middle and bottom panels, 605 respectively). B | Mechanisms of recruiting histone-modifying enzymes. B(i) | Histone 606 acetyltransferases (e.g. CBP) are recruited by transcription factors (TFs) or use their 607 intrinsic bromodomains to bind pre-existing acetyl-lysines to robustly acetylate histones 608 at active promoters. B(ii) | Heterochromatin protein 1 (HP1) is recruited (dashed arrow) 609 to chromatin via its chromodomain binding pre-existing H3K9me2/3. HP1 interacts with 610 the Suv3-9 histone methyltransferase that lays down further H3K9me (solid arrow) to 611 create a positive feedback loop. H3S10ph adjacent to H3K9me (e.g. as occurs at 612 mitosis) displaces (red dashed arrow) HP1 and prevents its rebinding (indicated by X) 613 via a phospho-switch mechanism. B(iii) | Chromatin-modifying enzymes interact with 614 specific forms of the RNA polymerase II complex (examples shown, yeast Set 1 and 615 Set2, interact with the Ser-5ph and Ser-2ph forms of RNAPII CTD, respectively, to 616 establish H3K4me3 at transcriptional start sites (TSSs: black arrow) and H3K36me3 617 within transcribed regions, respectively. B(iv) | Non-coding RNA (ncRNA) helps to 618 target PRC2 to specific sites and/or effects the activity of histone modifying enzymes. 619 Example shown is HOTAIR ncRNA which is required for recruitment of PRC2 (to 620 methylate H3K27) and LSD1 (to demethylate H3K4) in polycomb repressed gene 621 promoters. Feedback exists as H3K7me3 recruits further PRC2. C Selected PTMs in 622 the globular domain (model based on PDB 1KX5), on the lateral surface of the octamer 623 and close to the DNA entry and exit site (black), the dyad axis of the nucleosome (blue) 624 or at the interface between histones (grey). D Left: PTMs on histone tails (yellow filled 625 circles) are readily accessible and can be recognized and bound by reader or effector 626 proteins (blue shape). In most cases, the recruited proteins, rather than the tail 627 modifications themselves, then regulate chromatin function. Right: Modifications in the 628 globular domains of histones (red filled circle) can, depending on their location, either 629 affect histone–histone interactions (for modifications on the histone interactions surface, 630 top right) and thus destabilize nucleosomes, or affect histone–DNA interactions (for 631 modifications on the lateral surface of the histone octamer, bottom right) and thereby 632 impact nucleosome dynamics and chromatin function, often without effector proteins. 633 634 Figure 3. Patterns of histone PTMs in relation to zygotic genome activation (ZGA) 635 during embryonic development. (A) Global pattern of histone modifications before 636 and during zygotic genome activation in human191, 192, mouse34, 35, 88, 193-195, 637 Zebrafish:196-199, Xenopus:49, 200, 201, Drosophila:89, 202-204, C. elegans:205-208). The 638 intensity of the shading indicates the global relative prevalence of the respective 639 modification. Canonical (c) or non-canonical (nc) distribution is indicated for H3K4me3. 640 (B) Non-canonical H3K4me3 in mouse oocytes, catalysed by KMT2B (MLL2) in a 641 transcription-independent manner, encompasses broad domains enriched in DNA 642 hypomethylated regions. This distribution is extensively remodelled by the late 2-cell 643 stage at ZGA by the demethylases KDM5A/B, which is essential for proper ZGA and 644 early embryo development34, 35, 52. (C) H3K9me3 occupies lineage-specific genes during 645 development in mammals and C. elegans. Loss of H3K9me3 does not lead to 646 precocious depression of such genes; activation of gene expression requires the 647 additional presence of lineage-specific transcription factors (TFs)103, 104, 209. 648 649 Figure 4. Histone PTMs in the coordination of different genomic processes. (A) 650 Meiotic recombination at LAPs. Schematic model depicting initiation of DSBs at the 651 onset of MR at LAPs. PRDM9 is recruited to LAPs via association with CTCF, direct 652 binding to PRDM9 DNA binding sites, or association with the cohesin complex 653 containing STAG3 (blue circle). PRDM9 catalyses H3K4me3 and H3K36me3 in 654 adjacent nucleosomes to complement pre-existing H3K4me3. This creates a favourable 655 environment for recruitment of the HORMAD1-IHO1 and MEI4 complexes, and 656 ultimately SPO11 which initiates a DSB)210. (B) γH2AX domain formation at DSBs. A 657 model for megabase scale γH2AX domain formation133, in which H2AX in nucleosomes 658 is phosphorylated as it passes through a CTCF–cohesin mediated chromatin loop. (C) 659 H4K20me-dependent DNA repair pathway choices. H4K20me-dependent coordination 660 of DNA repair pathway choice with DNA replication status. Both 53BP1 and BARD1-661 BRCA1 bind to H2AK15ub, which is specifically accumulated at sites of DNA damage. 662 53BP1 dual binding to H4K20me1/2 and H2AK15ub restricts DNA end resection, 663 thereby favouring NHEJ during G1. H4K20me dilution during DNA replication triggers 664 preferential recruitment of BARD1-BRCA1 via its newly described ankyrin repeat 665 domain (ARD), channeling DSB repair towards HR (D) H3K36me3-dependent DNA 666 repair pathway choices. H3K36me3-dependent coordination of DNA repair pathway 667 choice with transcriptional status. 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