Precision-edited histone tails disrupt polycistronic gene expression controls in trypanosomes

Precision-edited histone tails disrupt polycistronic gene expression controls in trypanosomes | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Precision-edited histone tails disrupt polycistronic gene expression controls in trypanosomes Markéta Novotná , View ORCID Profile Michele Tinti , View ORCID Profile Joana R. C. Faria , View ORCID Profile David Horn doi: https://doi.org/10.1101/2025.03.21.644654 Markéta Novotná 1 Wellcome Centre for Anti-Infectives Research, School of Life Sciences, University of Dundee , Dow Street, Dundee DD1 5EH, UK Find this author on Google Scholar Find this author on PubMed Search for this author on this site Michele Tinti 1 Wellcome Centre for Anti-Infectives Research, School of Life Sciences, University of Dundee , Dow Street, Dundee DD1 5EH, UK Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Michele Tinti Joana R. C. Faria 1 Wellcome Centre for Anti-Infectives Research, School of Life Sciences, University of Dundee , Dow Street, Dundee DD1 5EH, UK Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Joana R. C. Faria David Horn 1 Wellcome Centre for Anti-Infectives Research, School of Life Sciences, University of Dundee , Dow Street, Dundee DD1 5EH, UK Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for David Horn For correspondence: d.horn{at}dundee.ac.uk Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract Transcription of protein coding genes in trypanosomatids is atypical and almost exclusively polycistronic. In Trypanosoma brucei , approximately 150 polycistrons, and 8000 genes, are constitutively transcribed by RNA polymerase II. RNA polymerase II promoters are unconventional and characterised by regions of chromatin enriched for histones with specific patterns of post-translational modification on their highly divergent N-terminal tails. To investigate the roles of histone tail-residues in gene expression control in T. brucei , we engineered strains exclusively expressing novel mutant histones. We used an inducible CRISPR-Cas9 system to delete >40 native copies of histone H4 , complementing the tandem arrays with a single ectopic H4 gene. The resulting ‘hist one H4’ strains were validated using whole-genome sequencing and transcriptome analysis. We then performed saturation mutagenesis of six histone H4 N-terminal tail lysine (K) residues and used multiplex amplicon-seq to profile the relative fitness of 384 distinct precision edited mutants. H4 K10 mutations were not tolerated, but we could derive a panel of nineteen strains exclusively expressing novel H4 K4 or H4 K14 mutants. Both proteomic and transcriptomic analysis of H4 K4Q mutants revealed significantly reduced expression of genes adjacent to RNA polymerase II promoters, where the glutamine (Q) mutation mimics an abnormally high level of acetylation. Thus, we present direct evidence for polycistronic expression control by histone H4 N-terminal tails in trypanosomes. Introduction Trypanosomatids are excavate protozoa that include several flagellated and vector-transmitted parasites 1 . The African trypanosome, Trypanosoma brucei , for example, is transmitted by tsetse flies and causes lethal human and animal diseases. Trypanosomatids display several unusual features for eukaryotes, one of which is a genome organised into very long, constitutively active, polycistronic, RNA polymerase II transcription units, comprising many genes with unrelated functions 2 . T. brucei also employs RNA polymerase I to transcribe both rRNA and some protein-coding genes, including sub-telomeric variant surface glycoprotein (VSG) genes, which are expressed in a monoallelic fashion in bloodstream-form cells 3 , 4 . Trypanosomatid histones display substantial differences relative to other model eukaryotes, and impacts on the T. brucei octameric nucleosome core particle, comprising approximately 146 bp of DNA wrapped around two molecules each of histones H2A, H2B, H3, and H4, have recently been elucidated using cryo-EM structural analysis 5 . Histones, histone tails that extend beyond the core nucleosome, histone post-translational modifications, and histone variants can impact gene expression. Histone N-terminal tails in particular display disordered and exposed surfaces that may recruit regulatory factors. Indeed, modifications, added by enzymes called writers, removed by erasers, and interpreted by proteins with reader domains, have been investigated in T. brucei . For example, histone modifications have been mapped using mass spectrometry 6 , 7 , while the roles of histone writers, erasers and readers have been investigated following knockdown 6 , by chromatin immunoprecipitation, and by subcellular localisation 8 . In a few cases, T. brucei writers have been linked to particular histone substrates and post-translational modifications. The histone acetyltransferase, HAT2 acetylates histone H4 at the lysine-10 position, for example 6 , 9 , while HAT3 acetylates histone H4 at the lysine-4 position 10 . Although the long polycistrons in trypanosomatids are typically constitutively transcribed by RNA polymerase II, locus-specific regulation has been reported in both Leishmania 11 and T. brucei 12 . In T. brucei , the regions that recruit RNA polymerase II to initiate transcription contain GT-rich tracts and display characteristics of highly dispersed or distributed promoters, in that they lack conventional promoter-associated sequences 13 , 14 . These unconventional promoters are characterised by regions of 5-10 kbp that are enriched for a large number of putative chromatin regulatory factors 8 , variant histones H2A.Z and H2B.V, and modified histones; histone H4 acetylated at the lysine-2 and -10 positions, for example 6 , 15 . Transcription termination sites, on the other hand, are enriched for histone variants H3.V and H4.V, and for a DNA base J modification 12 , 15 . In terms of the three-dimensional organisation of the nucleus, RNA polymerase II transcription initiation sites cluster in inter-chromosomal transcription hubs 16 , while a single active VSG is associated with an inter-chromosomal RNA polymerase I transcription and trans -splicing compartment 17 . Since histone writers, readers and erasers often operate as components of multi-subunit complexes, and since they often have multiple, and combinatorial histone and non-histone substrates, it can be challenging to establish the precise role of a particular histone post-translational modification. Homogeneous mutant genotypes can be generated to either prevent or mimic specific histone post-translational modifications, and this was first achieved in Saccharomyces cerevisiae , which has only two copies of the histone H4 gene; early studies in this yeast revealed surprising and substantial redundancy among histone N-terminal tails 18 , 19 . The generation of homogeneous mutant histone genotypes is typically challenging in other species, however. This required the expression of twelve copies of mutant histone H4 in Drosophila , for example 20 . Analysis of these strains clearly illustrated the benefits of assessing mutant histones directly, since histone H4 lysine 20 mutants failed to display the range of expected gene expression and developmental phenotypes, counter to prior interpretations based on studies in both humans and flies 21 . Interpretations from studies involving writer, reader or eraser perturbation are similarly challenging in trypanosomatids, which have more than 150 distinct histone modifications 6 . Transcription was perturbed following depletion of T. brucei histone acetyltransferase 2 (HAT2), for example, but acetylation was significantly reduced at histone H4 lysines 2, 5 and 10, as well as acetylation at multiple sites on the histone variants, H2A.Z and H2B.V 6 . Similarly, histone H4 lysine 4 acetylation is reduced but is not eliminated in hat3 null cells, indicating acetylation by another acetyltransferase 22 . In T. brucei , all four core histone genes are present in tandem arrays with approximately 43 copies of the histone H4 gene, for example. It was possible, using a CRISPR-Cas9 system over five months to edit many of these H4 genes, but 10 % remained intact and 40 % were deleted 23 . Using an alternative approach, mutant histone H4 genes were inducibly expressed in Trypanosoma cruzi , but these histones contributed only 1 % of the total histone H4 pool that was incorporated into chromatin 24 . We have engineered T. brucei strains that express a single ectopic H4 gene and that lack the histone H4 tandem arrays. These hist one H4 strains provide a platform for generating and analysing homogeneous mutant histone H4 genotypes. Accordingly, precision site-saturation mutagenesis was used to generate and assess hundreds of histone H4 N-terminal tail lysine mutants. Analysis of strains exclusively expressing mutant histones provided direct evidence for polycistronic gene expression control by H4 N-terminal tails. Results Replacement of >40 native T. brucei histone H4 genes Replication-dependent core histones are encoded by tandem gene arrays in T. brucei. Specifically, in the case of the histone H4 genes, there are estimated to be 43 copies in the diploid genome in two arrays each of approximately 15 kbp in length 23 . To facilitate H4 gene editing, we generated T. brucei strains with a single copy of the histone H4 gene. We first integrated a cassette that incorporated an ectopic H4 ECT gene and that encoded alternative H4 NAT A or H4 NAT B single guide RNAs (sgRNAs) in the 2T1 T7-Cas9 strain 25 , which inducibly expresses Cas9 ( Fig. 1a ). Both sgRNAs targeted the native H4 arrays, but not the H4 ECT gene, which encoded the same 100 amino acid protein but was recoded with 53 synonymous changes ( Fig. 1a ) such that every third codon position is a G or a C, which favours increased expression 26 , 27 . The synonymous changes additionally eliminated protospacer adjacent motifs associated with the sgRNAs, and altered 2-4 bases within each sgRNA target sequence, protecting the H4 ECT gene from Cas9-targeting. The H4 ECT gene was flanked by native H4 untranslated and cis -regulatory regions thought to support S phase specific expression 28 , and each expression cassette also incorporated a T7 phage promoter to drive transcription by the T7 polymerase, also expressed in the 2T1 T7-Cas9 strain 25 ; T7 polymerase achieves approximately 40-fold higher expression relative to RNA polymerase II in T. brucei 29 , and can drive expression of protein-coding genes in trypanosomatids because mRNA capping is achieved via trans -splicing. The system was, therefore, designed to replicate native H4 expression, and to complement for the loss of >40 native H4 genes with a single recoded H4 ECT gene. Download figure Open in new tab Fig. 1: Complementation of >40 T. brucei histone H4 genes with a recoded H4 gene a The schematic shows hist one H4 strain construction. The 2T1 T7-Cas9 strain was transfected with cassette containing a recoded, and ectopic histone H4 gene ( H4 ECT , synonymous changes highlighted) and a single guide RNA (sgRNA) targeting H4 NAT genes, both under the control of a T7 RNA polymerase promoter (T7). Cas9 was induced for 24 h in the resulting cell line, which was then transfected with an NPT (neomycin phosphotransferase) cassette to delete the native histone H4 ( H4 NAT ) tandem arrays. b The Southern blots were probed for the 5’ end of the native H4 genes (left), which detected the expected 736 bp band (approx. 43 copies), and 1505 bp band (1 copy from each of 2 alleles) only in the parental cell line. The blot was stripped and re-probed for the ectopic H4 gene (right) revealing the expected 4664 bp and 13803 bp bands in the hist one H4 strains, but not in the parent. Some residual signal can be seen from the 736 bp band. M, digoxigenin (DIG)-labelled DNA ladder. c The circular plot shows whole-genome sequencing data for the hist one H4 strains compared to the parental strain, and the parental strain plus the H4 ECT gene. The zoom at top shows precise deletion of the native H4 arrays; clone 1, blue; clone 2, magenta. d RNA-seq analysis. A strain expressing the H4 ECT gene was compared to the parent strain in the upper panels. A hist one H4 strain was compared to the strain expressing the H4 ECT gene in the lower panels. The H4 NAT and H4 ECT transcripts (orange) and the super-abundant VSG transcript are highlighted in the left-hand panels, while five genes immediately downstream of the H4 array are highlighted (blue) in the right-hand panels. n = 8934. Taking strains expressing H4 ECT and either the H4 NAT A or H4 NAT B sgRNA, we induced Cas9 expression, and delivered a Neomycin PhosphoTransferase (NPT) cassette, designed to replace the native H4 arrays, 24 h later ( Fig. 1a ). Screening of transformed clones using a PCR-assay revealed correct integration of the NPT cassette, driven by the H4 NAT B sgRNA, but not by the H4 NAT A sgRNA ( Supplementary Fig. 1a ). A second PCR assay suggested that both histone H4 arrays had been replaced in one of three ‘H4 NAT B’ clones ( Supplementary Fig. 1b ). We repeated the native H4 deletion process using the H4 NAT B strain, screened a second panel of clones, and isolated a second independent hist one H4 clone. Both hist one H4 strains were further validated using Southern blotting ( Fig. 1b ) and whole genome sequencing ( Fig. 1c ). Both approaches confirmed deletion of all native H4 genes in the hist one H4 strains. Genome sequencing also revealed that removal of the native H4 gene arrays on chr. 5 in the hist one H4 strains was both precise and specific ( Fig. 1c ). Download figure Open in new tab Supplementary Fig. 1: Generation and initial validation of hist one H4 strains. a PCR assay to test incorporation of the NPT cassette and H4 array knockout in T. brucei strains expressing the H4 NAT A (upper panel) or H4 NAT B sgRNA (lower panel). Only the H4 NAT B sgRNA yielded correct incorporation, indicated by 2020 bp amplicon in clones 1, 3 and 6. b Native and ectopic H4 genes were PCR-amplified from clones 1, 3 and 6, and digested with SacII. The gel shows that only clone 1 lacked native H4 genes. The process was repeated to generate a second independent hist one H4 strain. c The cumulative growth curves show that growth of the hist one H4 strains is comparable to the parental 2T1 T7-Cas9 strain. Two technical replicates. Some error bars are obscured by the datapoints. d Native and ectopic H4 transcripts were reverse-transcribed and PCR-amplified from both hist one H4 strains and controls and digested with SacII. The gel indicates similar abundance transcripts from native and ectopic H4 genes and only transcripts from the ectopic H4 gene in the hist one H4 strains, as expected. A recoded H4 gene complements for the loss of native H4 genes We next characterised the hist one H4 strains to assess complementation for the loss of >40 native H4 genes by the single ectopic H4 ECT gene. We first demonstrated similar growth rates for the 2T1 T7-Cas9 and hist one H4 strains ( Supplementary Fig. 1c ). A Reverse-Transcription PCR assay was then used to detect transcripts from both H4 ECT and native H4 genes, suggesting exclusive expression of the ectopic H4 ECT gene in both hist one H4 strains, as expected ( Supplementary Fig. 1d ). We next progressed to RNA-seq analysis of parental 2T1 T7-Cas9 cells, cells additionally expressing the H4 ECT gene, and hist one H4 cells lacking the native H4 genes. H4 ECT gene recoding meant that we could readily distinguish between native H4 and H4 ECT expression and indeed, the transcriptomes revealed highly significant changes in the expression of these two genes. Specifically, the H4 ECT gene registered robust expression in the presence and absence of the native H4 genes ( Fig. 1d , left-hand panels; log 2 CPM = 8.9, 97 th percentile). Additionally, the native H4 signal was depleted >1000-fold following H4 array deletion, as expected ( Fig. 1d , left-hand panels). A closer inspection of genes that registered significantly different expression highlighted 20-42 % increased expression of five adjacent genes immediately downstream of the H4 arrays, specifically following deletion of the H4 arrays ( Fig. 1d , right-hand panels). We speculate that this may be due to a post-transcriptional increase in mRNA maturation, since histone gene arrays have been shown to associate with trans -splicing loci in chromosome conformation capture assays 17 . We concluded that the single ectopic H4 ECT gene complemented for the precise deletion of >40 native H4 genes in two independent hist one H4 strains. H4 tail lysine saturation mutagenesis and multiplex fitness-profiling While human and S. cerevisiae histone H4 N-terminal tail sequences differ at only V 21 I, there are substantial differences in T. brucei ( Fig. 2a ). The T. brucei sequence, like the sequences in human and S. cerevisiae , is lysine (K)-rich, however; seven of the first eighteen residues are K in T. brucei . To enable H4 ECT gene editing, we replaced the H4 NAT B sgRNA in the hist one H4 strains with an sgRNA that reprogramed Cas9 to target the H4 ECT gene ( Fig. 2b ). To facilitate editing of K residues in the H4 ECT N-terminal tail, we used an sgRNA that guided the introduction of a break 41 bp from the ATG start-codon. Hist one H4 strains, now also incorporating the H4 ECT sgRNA, were validated by PCR and Sanger sequencing and independent clones were selected for editing. Download figure Open in new tab Fig. 2: H4 tail lysine saturation mutagenesis and multiplex fitness-profiling a The alignment shows the N-terminal tail sequence of T. brucei histone H4, compared to the equivalent sequences from yeast and human. Lysine residues are highlighted in the T. brucei sequence. b Schematic of the saturation mutagenesis and amplicon-sequencing strategy. An sgRNA cassette targeting the 5’ end of the H4 ECT gene was introduced into an hist one H4 strain. Cas9 was induced, and 24 h later the cells were transfected with single-stranded oligodeoxynucleotide (ssODNs) editing templates. DNA was extracted at different timepoints, the edited region of the H4 ECT gene was PCR-amplified, and the amplicons were deep-sequenced. c The radial plots show the abundance of sequence-reads representing each codon at each edited position during the time course. Data for six edited N-terminal tail lysine’s are shown. Values represent the averages of duplicate samples and are relative to codon scores obtained at the 12 h timepoint. D2, day-2; D4, day-4; D6, day-6. d The logo shows relative overrepresentation or underrepresentation at day-6 for those cognate amino acids that were either tolerated or not tolerated following editing at each of the six lysine positions targeted. We designed editing templates for marker-free site saturation mutagenesis at H4 K4 , H4 K10 , and H4 K14 ; lysine residues that are known to be acetylated and/or methylated 6 , 7 , that have been characterised in trypanosomes 6 , 9 , 10 , 24 , and that are also conserved in Leishmania histone H4. Each single-stranded DNA template incorporated a randomised codon and several synonymous bases, flanked by 25 b homology arms ( Supplementary Fig. 2a , Supplementary Data 1). To protect edited H4 ECT genes from further Cas9-targeting, the synonymous bases both eliminated the targeted protospacer adjacent motif and altered bases within the sgRNA sequence. The introduction of synonymous bases also served a second purpose here, by generating novel PCR primer binding-sites to facilitate the amplification of edited sequences. Taking the hist one H4 strains expressing the H4 ECT sgRNA, we induced Cas9 expression and delivered each editing template 24 h later; Cas9 induction was maintained throughout, and samples were collected for DNA extraction before template delivery, and 0.5, 2, 4, and 6 days after template delivery ( Fig. 2b ). Download figure Open in new tab Supplementary Fig. 2: Ectopic H4 N-tail lysine mutagenesis. a The sequence traces show a r epresentative example of the unedited ectopic H4 gene and an edited gene with a repair template illustrated above. Repair-templates contained 5’ and 3’ homology arms (5’, 3’ hom.), a series of synonymous edits designed to generate a novel primer binding site (b.s.), a degenerate codon (NNN, in this case at lysine 14), and a synonymous base edit distal to the primer b.s. (*). Asterisks above the edited sequence-trace highlight introduction of the desired base-edits. b The PCR assays show the presence of the ectopic H4 gene in all samples across all six 6-day time-courses (left-hand panels) and, exploiting the novel primer binding-site, the presence of edited sequences following delivery of editing templates (right-hand panels). The amplicons shown on the right were deep-sequenced. To profile the edits, we selectively amplified edited sequences and also amplified unedited sequences in parallel ( Supplementary Fig. 2b ). PCR products for unedited sequences were obtained from all thirty samples. In contrast, and as expected, edited products were only detected in samples taken after delivery of an editing template, both confirming editing and demonstrating specific amplification of edited H4 ECT sequences. We deep-sequenced the multiplexed and edited amplicons to profile the relative fitness of cells harbouring each of the 192 possible edits; 64 codons at either K 4 , K 10 , or K 14 . We then scored sequence-reads for all possible codons at each position in the duplicate experiments and visualised the outputs on radial plots ( Fig. 2c , upper panels). By comparing the 2-day samples to the 0.5-day baseline, we were able to confirm the introduction of all 192 possible edits ( Fig. 2c , upper panels, blue inner datasets). Notably, the three stop codons were the most rapidly depleted edits, regardless of the K-residue edited, consistent with the expected loss-of-function phenotypes in strains expressing a single H4 gene with a nonsense mutation. We next turned our attention to assessing the relative fitness of cells with mis-sense mutations. Samples across the 6-day time-course revealed broadly consistent trends for synonymous edits that encode common amino acids ( Fig. 2c , upper panels). The analysis revealed that H4 K10 mutations were not tolerated; although all mis-sense edits at the K 10 position were similarly represented in the day-2 samples, all but the pair of lysine codons were severely diminished in the day-4 and day-6 samples ( Fig. 2c , upper middle radial plot). These results suggested a severe growth defect regardless of the alternative amino acid at this position, and an important role for K 10 . K 4 , in contrast, can be replaced by positively charged (histidine, H; or arginine, R) or aromatic (phenylalanine, F; or tyrosine, Y) amino acids, while codons representing negatively charged amino acids (aspartic acid, D; or glutamic acid, E) were rapidly depleted ( Fig. 2c , upper left-hand radial plot). K 14 can also be replaced, but primarily with non-polar residues in this case (isoleucine, I; methionine, M; valine, V; alanine, A; or glycine, G), while codons representing the aromatic amino acids (phenylalanine, F; tryptophan, W; or tyrosine, Y) were rapidly depleted ( Fig. 2c , upper right-hand radial plot). To extend and further test the utility of the hist one H4 strains and the platform we developed for assessing fitness associated with homogeneous histone H4 mutant genotypes ( Fig. 2b ), we designed three further editing templates and implemented site saturation mutagenesis at H4 K2 , H4 K17 , and H4 K18 . These lysine residues are also known to be acetylated and/or methylated 6 , 7 , and are conserved in Leishmania . By comparing the 2-day samples to the 0.5-day baseline, we were once again able to confirm the introduction of all 192 possible edits and rapid depletion of all three nonsense mutations, regardless of the K-residue edited ( Fig. 2c , lower panels, blue inner datasets). Samples across the 6-day time-course, once again revealed broadly consistent trends for individual amino acids ( Fig. 2c , lower panels). Nonpolar residues were tolerated at the K 2 position, as also seen for K 14 above. The K 17 position appeared to show some tolerance for polar residues, while mis-sense edits were typically not well-tolerated at the K 18 position. The results for each of the six edited histone H4 tail lysines are summarised as sequence logos, ranking those edits that were tolerated and those that were not tolerated ( Fig. 2d ). Only lysine registered as tolerated at all six sites, while only nonsense codons (X), cysteine (C) and aspartic acid (D) codons registered as not tolerated at any of the six sites. A panel of strains exclusively expressing histone H4 mutants The analyses above revealed those histone N-terminal tail lysine edits that were tolerated at the K 4 or K 14 positions but did not directly demonstrate the viability of strains homogeneously expressing these edited histones. To isolate individual histone H4 mutants, saturation mutated H4 K4 , H4 K10 or H4 K14 cultures were subcloned two to ten days after mutagenesis. Clones were assessed using the PCR-assays described above to amplify unedited and edited sequences, and the latter amplicons from those clones that registered as negative and positive, respectively, were sequenced. More than 200 clones were assessed in total, 149 following K4 editing, and 53 following K14 editing, yielding strains exclusively expressing H4 K4K , H4 K10K or H4 K14K and a range of novel H4 K4 or H4 K14 mutants ( Fig. 3a ). The panel of mutants we isolated was highly predictable, based on the fitness profiles described above ( Fig. 2c-d ); A, F, I, L, R or Y replaced K 4 K, only K ‘replaced’ K10K, and A, G, I, M, P, R, T or V replaced K 14 K. Download figure Open in new tab Fig. 3: A panel of strains exclusively expressing histone H4 mutants a The plots show the fitness of a panel of K4 and K14 mutants relative to H4 K4K or H4 K14K strains, respectively. Cell density was recorded every 24 h for 4-5 days. Sanger sequencing traces show the edited codon for each mutant. b Protein blotting analysis of selected K4 mutants, showing H4 K4 acetylation, non-acetylated H4 K4 , and H4 K10 acetylation. EF-1α (elongation factor 1 α; red) served as a loading control. Although K 4 Q (glutamine), K 14 Q, and K 4 H mutants were not obtained using the approach above, the fitness profiles ( Fig. 2c-d ) suggested that these mutants would be viable. We also wondered whether cells lacking the K4 or K14 residues would be viable. To obtain K 4 Q, K 4 Δ, K 14 Q, K 14 Δ, and K 4 H mutants, we performed mutagenesis of these residues as above ( Fig. 2b ), but in this case using templates containing specific codons or codon deletions rather than randomised codons. Editing, subcloning and assessment as above, yielded all five of these mutants. The full panel of nineteen mutants, plus two additional and independent K 4 Q mutants, since we were particularly interested in this edit (see below), were assessed for relative fitness, and compared to H4 K4K or H4 K14K cells, as appropriate ( Fig. 3a ). Relative fitness of the K4 and K14 mutants were broadly predictable, again consistent with the fitness profiles described above ( Fig. 2c-d ). Most mutations were well-tolerated, resulting in <10 % reduced growth over 4-5 days ( Fig. 3a ). The greatest loss of fitness among the K4 mutants was for the K 4 Q mutants. K4 deletion was also tolerated, although this mutant is in fact equivalent to a K 5 Δ mutant, in that it retains a K residue at the fourth position (see Fig. 2a ). Substitution of lysine with arginine (R) or glutamine (Q) serves to mimic non-acetylated or constitutively acetylated lysine, respectively. We observed a greater fitness cost associated with both the H4 K4Q or H4 K14Q mutants relative to the H4 K4R or H4 K14R mutants ( Fig. 3a ), suggesting that mimicking constitutive acetylation at these sites is more deleterious than blocking acetylation. To further assess and validate the H4 K4 mutant strains, we used protein blotting with a series of H4-tail modification-specific antibodies, that identify acetylated or non-acetylated H4 K4 10 or acetylated H4 K10 9 . Acetylated and non-acetylated H4 K4 were detected in wild-type cells, in hist one H4 cells and in edited H4 K4K cells, as expected ( Fig. 3b ; top and middle panels). In contrast, and also as expected, neither antibody recognised edited H4 K4R or H4 K4Q histones; although Q mimics acetylated lysine, this modification is not recognised by anti-H4 K4 ac. Notably, anti-H4 K4 ac did recognise edited K 4 Δ histones, consistent with the view that this mutant does in fact retain a K residue at the fourth position. Finally, anti-H4 K10 ac recognised H4 in all of these strains, indicating that these mutations do not substantially disrupt acetylation at this site, shown above to play an important role in terms of maintaining viability ( Fig. 2c-d ). These results confirm homogeneous expression of mutant histones in the H4 K4Q and H4 K4R strains. Proteomic analysis of histone H4 K4Q and H4 K14Q mutants The finding that both H4 K4Q and H4 K4R mutants were viable suggested that dynamic H4 K4 acetylation is not required for viability and presented an opportunity to explore how histone H4 tail lysine residues impact gene expression. We were particularly interested in the H4 K4Q mutants because H4 K4 acetylation is the only histone acetylation mark that is not enriched in regions where histone H4 K10 acetylation demarcates RNA polymerase II promoters; indeed this mark is diminished at these sites 6 . We initially used data-independent acquisition mass spectrometry to examine the proteomes of H4 K4K , H4 K4Q , H4 K14K , and H4 K14Q mutants. We analysed three independent H4 K4Q mutants, the H4 K4K mutant served as an otherwise isogenic control, and the H4 K14K and H4 K14Q mutants allowed us to assess the impact of a K-Q mutation at a different site. We first assessed the abundance of the replication-dependent histones and the variant histones, H2A.Z, H2B.V, H3.V and H4.V, none of which were significantly different in abundance in the H4 K4Q or H4 K14Q mutants, relative to their H4 K4K or H4 K14K counterparts ( Fig. 4a ; all with FDR >0.1). This analysis also revealed that the variant histones were approximately 30-fold less abundant on average that the replication-dependent histones. Further comparison of the proteomes of H4 K4K and H4 K4Q strains ( Fig. 4a , top panels) and H4 K14K and H4 K14Q strains ( Fig. 4a , bottom panels) revealed more substantial differences in the H4 K4Q cells, relative to their H4 K4K counterparts. Considering an FDR threshold of <0.01, 289 and 795 proteins were significantly increased or decreased in abundance respectively in these mutants, while 144 and 166 proteins were significantly increased or decreased in abundance in the H4 K14Q mutants. Thus, the abundance of 3.5 times more proteins was significantly different in the H4 K4Q mutants and 73 % of these were significantly decreased in abundance. Download figure Open in new tab Fig. 4: Proteomic analysis of histone H4 K4Q and H4 K14Q mutants a Proteomics analysis. Three strains expressing the H4 K4Q mutant were compared to an H4 K4K control in the upper panels. A strain expressing the H4 K41Q mutant was compared to an H4 K14K control in the lower panels. The core and variant histones are highlighted. n = 5776. b The boxplot shows log 2 fold-change for the two comparisons in a, and for either genes within 10 kbp of transcription start-sites (n = 468) or >10 kbp distal from those sites (n = 5308). Boxes indicate the interquartile range (IQR), the whiskers show the range of values within 1.5*IQR and a horizontal line indicates the median. The notches represent the 95% confidence interval for each median. p- values were calculated using t -tests. To ask whether changes in expression were associated with gene location within polycistronic transcription units, we assessed the relative abundance of proteins encoded by those genes within 10 kbp of a transcription start-site. This cohort of proteins displayed significantly reduced abundance, specifically in the H4 K4Q mutants ( Fig. 4b ). We concluded that proteomics analysis of trypanosomes homogeneously expressing a histone H4 K4Q mutant, suggested reduced expression of promoter-adjacent genes, without significantly changes in the abundance of histones or histone variants. Expression of promoter-adjacent genes is disrupted in H4 K4Q mutants To investigate gene expression in the context of polycistronic transcription units in histone H4 K4Q mutants in more detail, we turned to transcriptome analysis. We identified transcription start-sites and transcription termination-sites (see Materials and Methods), and find that T. brucei polycistrons contain approximately fifty genes on average, with the first and last genes located 2.7 kbp and 120 kbp on average from transcription start-sites ( Fig. 5a ). Consistent with the proteomics analysis above, transcripts from genes within 10 kbp of a transcription start-site displayed significantly ( p = 1.5e -26 ) reduced abundance in the H4 K4Q mutants ( Fig. 5a ), as did transcripts from genes within 10 kbp of a transcription termination-site ( p = 1e -6 ). Many start-sites are situated where two polycistrons diverge and many termination-sites are situated where two polycistrons converge, while other termination-sites are situated adjacent to where another polycistron starts (see Fig. 5b ). To distinguish between these regions, we first considered cohorts of genes within 10 kbp of divergent start-sites, and within 10 kbp of convergent termination-sites. Genes adjacent to divergent start-sites were significantly ( p = 2.9e -19 ) enriched among transcripts that were reduced in abundance in the H4 K4Q mutants ( Fig. 5b , top left panel). In contrast, the representation of genes adjacent to convergent termination-sites that were reduced or increased in transcript abundance, was not significantly different ( Fig. 5b , p = 0.6, top right panel). We concluded that the expression of genes adjacent to RNA polymerase II promoters was specifically reduced in H4 K4Q mutants. Download figure Open in new tab Fig. 5: Expression of promoter-adjacent genes is disrupted in H4 K4Q mutants a The schematic illustrates a canonical RNA polymerase II transcribed polycistronic transcription unit in T. brucei , showing the distribution of genes closest to the start-site (orange), genes at the end (blue) and all other genes (grey). The boxplot shows log 2 fold-change in RNA-seq data when we compared three strains expressing the H4 K4Q mutant to an H4 K4K control. Genes within 10 kbp of transcription start-sites (orange, n = 736), genes within 10 kbp of a termination site (blue, n = 576), all other genes (grey, n = 8755). Boxes indicate the interquartile range (IQR), the whiskers show the range of values within 1.5*IQR and a horizontal line indicates the median. The notches represent the 95% confidence interval for each median. p- values calculated using t -tests. b RNA-seq analysis focussing on genes adjacent to divergent start-sites (top left), genes adjacent to convergent termination-sites (top right), genes adjacent to start-sites where another polycistron ends (bottom left), and genes adjacent to termination-sites where another polycistron starts (bottom right). Distances from the start or end are shown on the x-axis. Numbers of transcripts within 10 kbp of a start- or end-site that were significantly (FDR <0.01) increased or reduced in abundance in the H4 K4Q mutant relative to the H4 K4K control are indicated. These numbers were used to calculate p- values using χ 2 tests. FDR, False Discovery Rate. c The circular plot shows the full RNA-seq dataset mapped to the T. brucei chromosome cores (data in grey). Transcripts from genes that are within 10 kbp of a promoter (black bars, as defined by H4 K10 acetylation footprints), and that are significantly (FDR <0.01) increased or reduced in abundance in the H4 K4Q mutant relative to the H4 K4K control, are highlighted. To determine whether reduced expression was associated with transcription start-sites per se , or with H4 K10 acetylation footprints, we next considered termination-sites where another polycistron starts. At these sites, both cohorts of genes at the ends and at the beginning of polycistronic transcription units were over-represented among transcripts that were reduced in abundance in the H4 K4Q mutants ( Fig. 5b , bottom panels). We concluded that the expression of genes adjacent to promoters (H4 K10 acetylation footprints) was disrupted in H4 K4Q mutants, regardless of the direction of transcription in relation to the promoter. Transcriptome-wide visualisation of these changes revealed significantly reduced expression adjacent to most promoters, with few examples of significantly increased expression adjacent to these sites ( Fig. 5c ). T. brucei employs RNA polymerase I to transcribe both rRNA and some protein-coding genes, including sub-telomeric variant surface glycoprotein (VSG) genes, which are expressed in a monoallelic fashion in bloodstream-form cells. We assessed the abundance of expression site associated gene ( ESAG s) transcripts derived from these polycistronic VSG transcription units 30 and found that expression of genes within 10 kbp of an RNA polymerase I promoter was, in contrast to genes adjacent to RNA polymerase II promoters, increased in H4 K4Q mutants ( Supplementary Fig. 3 ). Download figure Open in new tab Supplementary Fig. 3: Expression of RNA polymerase I promoter-adjacent genes is disrupted in H4 K4Q mutants a The schematic illustrates a canonical RNA polymerase I transcribed polycistronic VSG expression site in T. brucei . b RNA-seq analysis highlighting expression site associated genes. Distances from the promoter are shown on the x-axis (left-hand panel). FDR, False Discovery Rate. Discussion It remains unclear to what extent, and by which mechanisms, polycistronic gene expression control relies upon chromatin in trypanosomatids. N-terminal histone tails, and tail modifications, such as lysine acetylation, play key roles in transcription control in other eukaryotes. However, trypanosomatid histone N-terminal tails, and polycistronic transcription, are highly divergent relative to the usual model eukaryotes, suggesting novel mechanisms. Many putative chromatin regulatory factors, including acetyltransferases, methyltransferases, and histone variants, are enriched in association with the unconventional promoters in T. brucei 8 , 15 , 31 , 32 , but it remains unclear whether accumulation of these factor is a cause or a consequence of transcription. In addition, and as in all eukaryotes, interpretation of histone writer, reader and eraser-defective phenotypes is complicated by potential impacts on diverse histone and non-histone substrates and binding sites. We describe establishment of a system to directly assess the impacts of specific histone H4 residues in T. brucei . This required the replacement of tandem arrays of native histone H4 genes with a single-copy ectopic and recoded H4 gene, which was then amenable to editing. Since post-translational modifications change the properties of histones, by altering DNA-histone interactions, or by creating binding sites that recruit chromatin-interacting proteins, we focussed on lysine residues in the N-terminal tail that are known to be either acetylated or methylated. We used site-saturation mutagenesis to profile 384 distinct mutants and isolated a panel of strains exclusively expressing mutant histones. We found that H4 K10 , consistent with the role of acetylation at this site in demarcating transcriptional start sites 6 , was essential for viability. In contrast, H4 K4 or H4 K14 could be replaced by a number of different amino acids. Indeed, we observed relatively few changes in gene expression in an H4 K14Q mutant. Since acetylation on H4 K4 is depleted at promoters, in contrast to increased acetylation on other histone residues in these regions, we assessed the impact of an H4 K4Q mutation, which mimics the acetylated state, on gene expression profiles. Both proteomic and transcriptomic analysis showed that the H4 K4Q mutation specifically reduced the expression of genes adjacent to promoters, consistent with the view that histones and their modifications serve to focus the recruitment and action of RNA polymerase II at these sites. Destabilised nucleosome appear to be a conserved feature at promoters and transcription initiation sites in eukaryotes 33 . Transcription must be further coordinated at such dispersed promoters, however, and this requires a mechanism to both delimit the boundaries of initiation and also to ensure that transcription is correctly oriented. The phenotype we observe in histone H4 K4Q mutants may reflect defects in boundary control and/or in orientation control. Defective boundary control could allow for transcription initiation over a wider region, thereby producing incomplete and poorly processed transcripts, while defective orientation control could create conflicts between RNA polymerase complexes travelling in opposite directions. Our results are consistent with the view that histone tails contribute to boundary control and/or orientation control at highly dispersed promoters in trypanosomes. Indeed, these histone tails likely contribute to the recruitment of regulatory factors and to the assembly of transcription factor hubs 16 . Histone H4 N-terminal tails are required for repressing silent mating loci in yeast 18 and we find that the expression of genes close to RNA polymerase I promoters in silent VSG expression sites is increased in T. brucei histone H4 K4Q mutants. Although RNA polymerase I transcription is thought to initiate at all of these sites in bloodstream-form cells, transcription attenuation effectively silences all but one of these sites 34 , and these silent subtelomeric polycistrons are folded into highly compact nuclear compartments 35 . The phenotype we observe could reflect defective attenuation of RNA polymerase I transcription, or inappropriate read-through of RNA polymerase II transcription 36 . Notably, inhibition of acetyl-lysine binding bromodomain factors also disrupts silencing at VSG expression sites 31 . Thus, histone tails are involved in controlling RNA polymerase I and RNA polymerase II polycistrons in T. brucei . Indeed, RNA polymerase III transcribed genes are also likely affected 37 . In summary, we show that a histone H4 K4Q mutation impacts RNA polymerase II polycistronic transcription units in T. brucei , specifically reducing the expression of genes adjacent to promoters. The system and approach we describe could also be used to explore how histones impact gene silencing 38 , DNA replication 39 , DNA recombination and repair 40 , or chromosome segregation 41 . Our findings also suggest that similar approaches could be used to edit and interrogate the roles of specific residues in other genes found in tandem arrays in T. brucei . Our findings support the view that histone acetylation contributes to delimiting the regions where RNA polymerase II can initiate transcription in trypanosomes. We conclude that histone H4 mutagenesis provided direct evidence that histone H4 tails impact polycistronic RNA polymerase II gene expression controls, and also RNA polymerase I mediated expression controls in T. brucei . Materials and Methods Trypanosoma brucei growth and manipulation Bloodstream form T. brucei Lister strain 427 (MITat 1.2), clone 221a cells, 2T1 T7-Cas9 cells 25 , and derivatives were grown in HMI-11 medium at 37 °C with 5 % CO 2 . Unless otherwise stated, strains were maintained in antibiotics at the following concentrations: 1 µg/mL of hygromycin (Sigma), phleomycin, G418 or puromycin and 2 µg/mL of blasticidin (all Invivogen); selection for new transformed strains was applied at 2.5, 2, 2, 10 and 10 µg/mL, respectively. Transfections were performed using the Human T Cell Nucleofector™ Kit and Amaxa 2b device (Lonza), and programme Z-001. Ten μg of linearised plasmid DNA or PCR-product, or 40 μg of each mutagenic oligonucleotide, were typically used for transfection. After transfection, cells were typically grown in 50 mL of HMI-11 medium without antibiotics for 6 h and then transferred to 48-well plates with the appropriate antibiotic selection, where applicable. The pT7 sgRNA H4 constructs were linearised using NotI prior to transfecting 2T1 T7- Cas9 cells. When replacing the native H4 arrays (H4 NAT sgRNA), or when editing the ectopic H4 gene in hist one H4 strains (H4 ECT sgRNA), Cas9 expression was induced by adding tetracycline (Sigma) at 1 μg/mL for 24 h prior to transfection, and tetracycline was maintained until clone or sample collection, respectively. When replacing the native H4 arrays, transfected cultures were selected with G418 for 3-6 days, diluted and transferred to 96-well plates to obtain clonal populations. The replace.sgRNA construct was digested with BamHI and XbaI prior to transfection with hist one H4 cells. Puromycin resistant clones were checked for phleomycin sensitivity, to confirm correct integration. When editing the ectopic H4 gene, transfected cultures were diluted and transferred to 96-well plates to obtain clonal populations 2-10 days after transfection. To determine growth rates, T. brucei cell density was measured using a haemocytometer every 24 h, and cultures were diluted to 10 5 cells/mL in the absence of antibiotics. Plasmids and editing templates Two single guide RNA (sgRNA) sequences targeting native histone H4 genes were assembled in the pT7 sgRNA plasmid as described 25 . These were H4 NAT A (TTCCGCGCACATTCTCACGG) and H4 NAT B (GAGGCGGCGGATGGAACCGC). A recoded ectopic histone H4 gene, H4 ECT (GenScript) was then, as a 593 bp XbaI / PstI fragment, ligated to similarly digested pT7 sgRNA , containing either the H4 NAT A or H4 NAT B sequence. For Cas9-driven deletion of the native H4 arrays, a repair cassette was amplified by PCR, using the NPT-H4.F and NPT-H4.R primers (Supplementary Data 1). The resulting template incorporated a neomycin phosphotransferase ( NPT ) expression cassette flanked by 50-bp sequences that targeted regions on either side of each array for homologous recombination. To exchange the sgRNA cassette targeting native histone H4 genes for an sgRNA cassette targeting the ectopic histone H4 gene, we assembled the replace.sgRNA construct. First, a PAC cassette was amplified using the TUBrFse and PACfBst primers (Supplementary Data 1). The 1023 bp amplicon and the pT7 sgRNA plasmid were then digested with FseI and Bst17I and ligated. The resulting construct and a fragment containing a VSG promoter and a PFR2 5’-UTR (GenScript) were then digested with NheI and BstZ17I and ligated. Finally, an H4 ECT sgRNA sequence (GCACCTTCTTCTGCCGCTTC) targeting the ectopic histone H4 gene was assembled in the modified pT7 sgRNA plasmid as described 25 . All constructs derived from pT7 sgRNA were validated by Sanger sequencing performed using an Applied Biosystems 3730 DNA analyser. Distinct oligonucleotide repair templates (ThermoFisher) were used for site-saturation mutagenesis or to generate specific mutants, H4 K4H , H4 K4Q , H4 K4Δ , H4 K14Q , and H4 K14Δ (Supplementary Data 1). T. brucei genomic DNA analysis T. brucei genomic DNA was extracted using DNazol (ThermoFisher), following the manufacturer’s instructions. A PCR assay was used to assess replacement of H4 arrays with the NPT cassette. Clones were screened using the H4.out.tandem.R and H4.out.tandem.F primers (Supplementary Data 1) that annealed to regions flanking the NPT -editing template detailed above. Clones with correctly integrated NPT cassettes were then assessed using a second PCR assay with the H4.F.new and H4.R.new primers to amplify both the ectopic and any remaining native H4 genes. The resulting PCR products were digested with SacII and separated on 2% agarose gels. To assess exchange of the H4 NAT B sgRNA with the H4 ECT sgRNA using the replace.sgRNA construct, clones were screened using a PCR assay with the sgRNA.PAC.inc.F and sgRNA.inc.R primers (Supplementary Data 1). The expected PCR products were also checked by Sanger sequencing. All PCR reactions were performed using Q5 High-Fidelity DNA Polymerase and Q5 polymerase buffer (New England BioLabs) and a ProFlex PCR System (ThermoFisher). For Southern blotting, T. brucei genomic DNA (10 μg) was digested with SacII and separated on 0.8% agarose gels. Digoxigenin labelled DNA ladder (Roche) and GeneRuler 1 kb plus DNA ladder (ThermoFisher) were run in parallel. Gels were then processed using standard protocols, DNA was transferred to nylon membranes (Amersham), and UV crosslinked. A native H4 probe was generated by PCR using the H4.F.new and H4.R.new primers (Supplementary Data 1) with wild type T. brucei genomic DNA as template. The product was digested with SacII, separated on an agarose gel, and the 124 bp fragment was isolated; this fragment corresponded to the 5’-end of the native H4 gene. An ectopic H4 probe was generated by digesting the pT7sgRNA.H4 plasmid with AflIII, separating the products on an agarose gel, and isolating the 359 bp fragment. A digoxigenin High Prime DNA Labelling and Detection Starter Kit II (Roche) was then used to label the probes, which were applied at 25 ng/mL at 45 °C ( H4 NAT ) or 53 °C ( H4 ECT ) overnight. Imaging was performed using a ChemiDoc XRS+ (BioRad) or iBright™ CL750 Imaging System. Genome sequencing for hist one H4 and control strains was performed on a DNBseq platform (BGI); 3.5 Gbp/sample, 150 bp read length. Initial quality control of fastq files was performed using FastQC ( https://www.bioinformatics.babraham.ac.uk/projects/fastqc/ ), followed by adapter trimming and quality filtering with Fastp (0.20.0) 42 . Processed reads were aligned to the reference genomes 427_2018 (TriTrypDB v68) 43 using Bowtie2 (2.3.5) 44 with ‘--very-sensitive-local’ parameters. The resulting alignments were processed with SAMtools (1.9) 45 for sorting and indexing. Bam files where were transformed to bigWig track file using bamCoverage from DeepTools (3.5) 46 with --binSize 5 and --normalizeUsing RPKM. The linear coverage visualization was performed with the pyGenomeTracks 47 python package. Bam files from Parental and derived clones were analysed using bamcompare from DeepTools (3.5) 46 with --bin 200, smooth 500, --operation log2, --normalizeUsing RPKM –extendReads, --scaleFactorsMethod None, and --outFileFormat bedgraph. The circular fold change visualization of the bedgraph files was performed with the pyCirclize (1.6) Python package ( https://github.com/moshi4/pyCirclize ). Specific T. brucei H4 mutants were screened using H4 PCR assays, as described above. Clones that were positive for the edited sequence and negative for the unedited sequence were confirmed by Sanger sequencing. RNA extraction and RT-PCR T. brucei RNA was extracted using the RNeasy Mini Kit (Qiagen), following the manufacturer’s instructions. RNA was reverse transcribed using a high-capacity RNA-to-cDNA kit (ThermoFisher), followed by PCR with the SL22 and MN.H4.3 primers (Supplementary Data 1); the SL22 primer is complementary to the spliced leader sequence found at the 5’ end of every T. brucei mRNA while the MN.H4.3 primer is complementary to the 3’ end of both the native and ectopic H4 genes. The resulting products were digested with SacII and separated on 2 % agarose gels. Codon-scoring following saturation mutagenesis of H4 N-tail residues Samples were collected 12 h, 2, 4, and 6 days after transfection with oligonucleotide editing templates. DNA was isolated and PCR assays were performed to amplify unedited H4 (primers H4K4.cont.R and H4K4.F for K2 and K4, H4K10.cont.F and H4K10/14.R for K10, H4K14.cont.F and H4K10/14.R for K14, H4K17.18.F.cont and H4K17.18.R for K17 and K18), or edited H4 (primers: H4K4.mut.R and H4K4.F for K2 and K4, H4K10.mut.F and H4K10/14.R for K10, H4K14.mut.F and H4K10/14.R for K14, H4K17.18.F.mut and H4K17.18.R for K17 and K18). Sequencing of edited amplicons (K2 and K4: 217 bp, K10: 220 bp, K14: 208 bp, K17 and K18: 200 bp) was performed on a BGISEQ-500 platform at BGI; 100 b paired-end reads 3.5 Gbp per sample. Oligo counting was performed with the OligoSeeker (0.0.5) ( https://doi.org/10.5281/zenodo.15011916 ) Python package designed to process paired FASTQ files and count occurrences of specific codons as described in 48 . Raw codon counts were normalized to account for differences in sequencing depth between samples by dividing each sample’s counts by a correction factor (sample total/mean total across samples). For time course experiments, biological replicates were averaged at each. To visualize relative changes in codon frequency over time, counts were further normalized to the 12-hour timepoint. The circular visualization of the codon data was performed with a custom Python script (3.7) using the matplolib library. Protein blotting We used antibodies specific for acetylated H4 K4 , non-acetylated H4 K4 10 or acetylated H4 K10 9 . Two million T. brucei cells were resuspended in LiCOR lysis buffer (137 mM Tris-HCl, 140 mM Tris base, 1% SDS, 513 μM EDTA, 7.5% glycerol, 1.2 Orange G (Invitrogen)). Samples were then treated with Pierce universal nuclease (ThermoFisher) to reduce viscosity and incubated at 70 °C for 10 min prior to loading on 12 % BisTris polyacrylamide gels alongside All Blue MW standard (BioRad). MES buffer was used for electrophoresis. Proteins were transferred to nitrocellulose membranes using an iBlot 2NC stack, and iBlot 2 device (Invitrogen) set at 25 V for 7 min. Membranes were stained with Ponceau to confirm correct protein transfer and washed with PBS. The membranes were then blocked using LiCOR blocking buffer (50 mM Tris-HCl pH 7.4, 0.15 M NaCl, 0.25% bovine serum albumin, 0.05% (w/V) Tween-20, 0.05% NaN 3 , 2% (w/V) fish scale gelatine). Membranes were incubated with primary antibodies: α-H4 K4 ac (rabbit, 1:500) and mouse α-EF-1α (Millipore, 1:10 000) for 2 h at RT or with α-H4 K4 non-ac (rabbit, 1:500), or α-H4 K10 ac (rabbit, 1:500), and mouse α-EF-1α (1:10 000) at 4 °C overnight. Membranes were washed three times with 0.1 % w/V Tween-20 in PBS and incubated with secondary antibodies: α-rabbit IRDye800 and α-mouse IRDye680 (1:15 000 and 1:10 000, respectively), for 1 h 30 min at RT; all antibodies were diluted in LiCOR blocking buffer. Membranes were then washed three times with 0.01% w/V Tween-20 in PBS, once with PBS. Blocking and washing steps were performed using a SNAPid vacuum platform, after inserting membranes in a SNAPid 2.0 miniblot holder and plastic mainframe (all Invitrogen). Blots were finally imaged using a LiCOR Odyssey CLx scanner. Images were adjusted and analysed in Image Studio ver 5.2. RNA-seq RNAseq was carried out using a DNBSEQ™ platform at the Beijing Genomics Institute (BGI), 100 b paired-end reads, 30 Mbp per sample. We assembled a comprehensive set of Variant Surface Glycoprotein (VSG) coding sequences from Trypanosoma brucei strain 427. To ensure our analysis focused exclusively on the unique regions of each VSG and to minimize the confounding effects of highly conserved C-terminal domains, all sequences were truncated at 1,200 base pairs from their ATG site. The dataset comprised 299 unique VSG sequences distributed across four categories: 14 from Bloodstream Expression Sites (BES), 5 from Metacyclic Expression Sites (MES), 63 classified as Metacyclic (MC), and 217 from chromosomal arrays. Initial quality control of fastq files was performed using FastQC ( https://www.bioinformatics.babraham.ac.uk/projects/fastqc/ ), followed by adapter trimming and quality filtering with Fastp (0.20.0) 42 . Processed reads were aligned to the reference genomes 427_2018 (TriTrypDB v68) 43 supplemented with the truncated set of VSG sequences and the pT7sgRNA_H4B sequence using Bowtie2 (2.3.5) 44 with ‘--very-sensitive-local’ parameters. The resulting alignments were processed with SAMtools (1.9) 45 for sorting and indexing, and PCR duplicates were marked using Picard MarkDuplicates (2.22.3) 49 . Read counts per coding sequence were quantified using featureCounts (1.6.4) 50 with parameters accounting for multi-mapping reads (-M) and overlapping features (-O) and configured to count only reads where both ends were mapped (-B) and to exclude chimeric reads that mapped to different chromosomes (-C). A minimum overlap fraction of 1.0 was required (--fracOverlap 1.0), ensuring that only reads completely contained within CDS features were counted. Gene features were identified using the ‘gene_id’ attribute (-g gene_id) from the reference genome annotation file (GTF) download from TriTrypDB. For the analysis of bloodstream expression site genes, we downloaded 14 sequence and annotation files from NCBI GeneBank corresponding to the BES/TAR clones 30 . The GFF files obtained from NCBI were converted to GTF format using GFF utilities to ensure compatibility with our analytical pipeline. These expression site regions were aligned following the same strategy described previously and mapped alignments were filtered for MAPQ > 1 before counting using featureCounts. Following the alignment processes, read counts from the expression site regions, were merged with counts from the main genome and VSG dataset. The combined dataset was then used as input for differential expression analysis using the edgeR package (3.28) 51 in R (3.6.1). We retained only genes that had a minimum count of 10 reads in at least one sample and a minimum total count of 30 reads across all samples. The cqn package (1.32) was used to compute gene length and GC content bias. The resulting offsets were incorporated into the DGEList object before computing dispersions. We fitted our data to a generalized linear model (GLM) using the quasi-likelihood (QL) method via glmQLFit and we performed differential expression testing using the quasi-likelihood F-test through the glmQLFTest function. The output from this statistical test was then processed using the topTags function, which extracted the complete set of test results. We configured topTags to return all tested genes (n=Inf) without applying any sorting (sort.by="none"), ensuring that the original gene order was preserved in the output. For multiple testing correction, we employed the Benjamini-Hochberg (BH) procedure (adjust.method="BH") to control the false discovery rate. The final result_table contained the complete set of statistical results for all tested genes, including log-fold changes, p-values, and adjusted p-values (FDR). RNA-seq visualization To accurately define the boundaries of polycistronic transcription units in the T. brucei 427 genome, we performed manual annotation of transcription start sites (TSS) and transcription termination sites (TTS). This manual curation process was guided by ChIP-seq footprint data for histones available at TriTrypDB in the 427_2018 genome 43 . The genomic distance for each gene annotated in the BES/TAR clones (ESAG, Expression Site Associated Genes) to its respective promoter was calculated using the annotated promoter sites identified in the GFF files downloaded from GenBank. A custom Python (3.7) script was used to visualize differential expression values (log2 fold changes) respect to the distance from TSS or TTS using the Matplotlib (3.6) and Pandas (1.4.2) python libraries. The circular visualization was performed with the pyCirclize (1.6) Python package ( https://github.com/moshi4/pyCirclize ). Proteomics For each T. brucei strain, three 50 ml cultures at 10 6 cells/ml (5 x 10 7 cells per each technical replicate) were harvested and resuspended in 5 % SDS, 100 mM triethylammonium bicarbonate in water. All samples were submitted for direct data-independent acquisition mass spectrometry. Peptides (equivalent of 1.5 µg) were injected onto a nanoscale C18 reverse-phase chromatography system (UltiMate 3000 RSLC nano, Thermo Scientific) and electrosprayed into an Orbitrap Exploris 480 Mass Spectrometer (Thermo Fisher) using Orbitrap Exploris 480 Mass Spectrometer. For proteomics analysis, we utilized a protein dataset derived from the T. brucei strain 42_2018 that matched directly to the transcript sequences used in our RNA-seq analysis. Raw files were analysed with DIA-NN (1.8.1) with C-carbamidomethilation as fixed modification, MBR option activated, the FASTA digest library-free search option activated and selection of unrelated runs activated. We extracted protein groups from the report.tsv file using the diann (1.0.1) library in R (4.2.3) with Q.Value <= 0.01 and PG.Q.Value <= 0.01. The protein groups identified as single peptide hit were considered missing values. One proteomic sample (hist one H4 c1, technical replicate 2) exhibited an anomalously high number of missing values, likely due to technical issues during protein injection at the mass spectrometry stage. This sample was excluded from subsequent analyses. Before batch effect correction using limma removeBatchEffect function (3.54), the proteomic data was normalized by equalizing the median intensities. Missing values were imputed using the missForest (1.5) algorithm in R, which was applied separately to each set of replicated experimental conditions. The differential expression analysis was performed with the limma package using the eBayes fitting function. FDR values were computed with the toptable function in limma. FUNDING This work was funded by a Wellcome Trust Investigator Award to D.H. (217105/Z/19/Z) and a Wellcome Trust PhD four-year Studentship award to M.N (222326/Z/21/Z). Conflict of interest statement. None declared. AUTHOR CONTRIBUTIONS Conceptualization; MN, DH. Investigation; MN. Data curation and analysis; MN, MT, DH. Supervision; JRCF, DH. Original draft; MN, MT, DH. Review & editing; all authors. Data and materials availability Genomic, transcriptomic, and amplicon-sequencing data have been deposited in the Sequence Read Archive, https://www.ncbi.nlm.nih.gov/sra (BioProject: PRJNA1234166). The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD061709. The OligoSeeker Python package for counting the occurrence of specific codons has been deposited in Zenodo ( https://doi.org/10.5281/zenodo.15011916 ). ACKNOWLEDGMENTS We thank A. Score for assistance with proteomics, G. 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