R-loops acted on by RNase H1 are a determinant of chromosome length-associated DNA replication timing and genome stability inLeishmania

R-loops acted on by RNase H1 are a determinant of chromosome length-associated DNA replication timing and genome stability in Leishmania | 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 R-loops acted on by RNase H1 are a determinant of chromosome length-associated DNA replication timing and genome stability in Leishmania View ORCID Profile Jeziel D. Damasceno , Emma M. Briggs , Marija Krasilnikova , Catarina A. Marques , Craig Lapsley , Richard McCulloch doi: https://doi.org/10.1101/2024.04.29.591643 Jeziel D. Damasceno 1 The Wellcome Centre for Integrative Parasitology, University of Glasgow, School of Infection and Immunity, Sir Graeme Davies Building, 120 University Place , Glasgow, G12 8TA, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jeziel D. Damasceno For correspondence: jeziel.damasceno{at}glasgow.ac.uk richard.mcculloch{at}glasgow.ac.uk Emma M. Briggs 2 University of Edinburgh, Institute for Immunology and Infection Research, School of Biological Sciences , Edinburgh, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site Marija Krasilnikova 1 The Wellcome Centre for Integrative Parasitology, University of Glasgow, School of Infection and Immunity, Sir Graeme Davies Building, 120 University Place , Glasgow, G12 8TA, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site Catarina A. Marques 1 The Wellcome Centre for Integrative Parasitology, University of Glasgow, School of Infection and Immunity, Sir Graeme Davies Building, 120 University Place , Glasgow, G12 8TA, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site Craig Lapsley 1 The Wellcome Centre for Integrative Parasitology, University of Glasgow, School of Infection and Immunity, Sir Graeme Davies Building, 120 University Place , Glasgow, G12 8TA, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site Richard McCulloch 1 The Wellcome Centre for Integrative Parasitology, University of Glasgow, School of Infection and Immunity, Sir Graeme Davies Building, 120 University Place , Glasgow, G12 8TA, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: jeziel.damasceno{at}glasgow.ac.uk richard.mcculloch{at}glasgow.ac.uk Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract Genomes in eukaryotes normally undergo DNA replication in a choreographed temporal order, resulting in early and late replicating chromosome compartments. Leishmania , a human protozoan parasite, displays an unconventional DNA replication program in which the timing of DNA replication completion is chromosome size-dependent: larger chromosomes complete replication later then smaller ones. Here we show that both R-loops and RNase H1, a ribonuclease that resolves RNA-DNA hybrids, accumulate in Leishmania major chromosomes in a pattern that reflects their replication timing. Furthermore, we demonstrate that such differential organisation of R-loops, RNase H1 and DNA replication timing across the parasite’s chromosomes correlates with size-dependent differences in chromatin accessibility, G quadruplex distribution and sequence content. Using conditional gene excision, we show that loss of RNase H1 leads to transient growth perturbation and permanently abrogates the differences in DNA replication timing across chromosomes, as well as altering levels of aneuploidy and increasing chromosome instability in a size-dependent manner. This work provides a link between R-loop homeostasis and DNA replication timing in a eukaryotic parasite and demonstrates that orchestration of DNA replication dictates levels of genome plasticity in Leishmania . Introduction In all organisms, DNA replication normally initiates from genomic locations known as origins 1 , which are specified in eukaryotes by the binding of the Origin Recognition Complex (ORC) 2 . Eukaryotic cells normally complete replication of their genome during S phase of the cell cycle, before segregating the resulting chromosome copies into offspring cells. To ensure completion of genome replication in this window of the cell cycle, numerous origins are normally specified and then activated on each eukaryotic chromosome. However, not all origins activate simultaneously at the onset of S phase. Instead, spatial and temporal regulation leads to differential origin activation across the genome as S phase progresses. As a result, the completion of genome duplication occurs in a staggered manner across different chromosome and genome compartments, a phenomenon referred to as ‘replication timing’ 3 – 5 . Most of our knowledge about eukaryotic DNA replication timing comes from studies using model organisms, and mainly mammalian cells and yeast, revealing correlations between the orchestration of origin activation and primary and tertiary genomic features. In Saccharomyces cerevisiae and related yeasts, origins are conserved DNA elements that bind ORC 6 , but replication mapping shows that not all are activated equally 7 . In all yeasts examined, centromeric origins are activated early 8 , 9 , while subtelomeric origins are activated later 10 . Further domains containing groups of late- or early-activated origins can also be detected 7 , 11 , 12 , but the basis for such co-ordination is not fully understood, though origin sequence, chromatin and transcription have all been shown to relate to origin activity 13 – 15 . In the larger genomes of mammals, identification of origins has proved more challenging due to a lack of consistency between datasets that map DNA replication initiation, or localisation of ORC 1 , 16 . In part, such inconsistency may be due to mammalian origins not being sequence-conserved genome features, but could also reflect greater use of stochastic, potentially ORC-independent initiation than is seen in yeast 17 – 21 . Nonetheless, mammalian replication timing appears to be organised over several scales. The smallest scale is clusters of origins within initiation zones of tens of kb, the earliest activating of which are bounded by efficient origins 22 . Replication timing is not fixed between cell types during mammalian development 23 and cell type-specific ‘constant timing regions’ have been described, which are large, near megabase-sized regions that comprise smaller (hundreds of kb) replication domains in which origins show coordinated changes in replication timing during development 23 , 24 . Replication domains display considerable correlation with topologically associated domains revealed by chromatin capture analyses 25 , 26 . On the largest scale, replication timing is related to nuclear substructuring 27 . Lowly expressed genes are often found towards the periphery of the nucleus in a heterochromatic, lamina-associated domain and are typically late replicating 28 ; indeed, increasing the expression of a gene can result in disassociation from the lamina and can advance replication in S-phase for the gene and surrounding sequence 29 . At least some of the genome is associated with the nucleolus and is also characterised by heterochromatin, low levels of gene expression and late replication 30 , 31 . In contrast, greater gene expression and gene density, as well as higher GC content, is associated with euchromatin and early replication 24 . Indeed, proximity to nuclear speckles (mRNA splicing factor-enriched nuclear bodies) correlates strongly with expression levels, gene density and early replication 32 . DNA replication timing has important implications for genome stability, composition and, ultimately, evolution 33 – 35 . In both mammals 36 , 37 and yeast 38 , 39 , levels of single nucleotide polymorphisms have been shown to increase in later replicating parts of the genome. Conversely, in mammals, amplification events through copy number variation 40 and translocations 41 are more abundant in early replicating genomic regions; indeed, early replicating regions are associated with cancerous translocations and altering the timing of DNA replication changes the frequency of such translocations 42 , 43 . Finally, replication timing correlates with the density of transposable elements in the human genome 23 , 44 , 45 . Despite such evidence of its importance, few cis and trans determinants of DNA replication timing have been described. Recent work identified DNA sequence elements that dictate early replication timing, subnuclear localisation and transcription in mouse embryonic stem cells 46 , but whether they are common to other mouse cells or wider eukaryotes is unclear. RIF1 (Rap1 interacting factor 1) defines late replicating domains 10 , 47 , 48 , both through controlling origin activation 49 , 50 and defining chromatin architecture 51 , 52 . In contrast, Fkh1/2 (forkhead transcription factor 1/2) defines early replicating domains in yeast 14 , 53 , 54 . Finally, in mammals, very long non-coding RNAs, termed asynchronous replication and autosomal RNAs (ASARs), are encoded across the genome and remain associated with the chromosome region from which they are generated 55 , 56 . ASARs can be expressed from just one allele (termed monoallelic expression) 57 , 58 or can show different levels of expression from the two alleles 55 , but in both cases are correlated with differential replication timing of the chromosomal locus. Disruption of all ASARs so far tested results in delayed replication and instability of the entire chromosome 55 , 58 , 59 . How ASARs act in DNA replication timing determination is unclear, but their role can be mediated via features of LINE retrotransposons, which are highly abundant in mammalian genomes 60 . Whether ASARs form large or punctuated RNA-DNA hybrids (see below) has not been tested. In contrast to the advanced understanding in model eukaryotes, we are only beginning to unveil how genome duplication is orchestrated in protozoans, which represent much of eukaryotic diversity 61 , 62 . Giardia and Tetrahymena cells are unusual amongst eukaryotes in possessing two nuclei, and DNA replication timing of the two genomes appears uncoordinated 63 or constitutively asynchronous 64 , respectively. Replication timing within these genomes has not been assessed. In Plasmodium , ChIP-seq reveals abundant localisation of ORC across the genome that shows limited correlation with mapped origins during schyzogony 65 , 66 , and so it is unclear how an observed increase in number of origins activated as S-phase proceeds occurs 66 . In contrast, in the highly transcribed core of the 11 megabase chromosomes of Trypanosoma brucei , mapped origins show clear correspondence with ORC binding 67 . In common with all kinetoplastids, virtually all genes in T. brucei are expressed from polycistrons 68 , 69 and ORC and origins localise to the start and ends of these transcription units, meaning they are unusually widely separated and limited in number in the genome 70 . Nonetheless, origin mapping by Marker Frequency sequencing (MFA-seq, equivalent to sort-seq in yeast) 71 suggests variation in origin activation that is conserved across the life cycle of T. brucei and between strains 72 . T. brucei centromeres appear to be the earliest replicating origins but what dictates variable timing at other origins is unclear, though the wide spacing of origins in T. brucei appears to preclude coordinated activation of proximal origins. In addition, replication timing of the large, variable subtelomeres 73 remains uncertain. Leishmania spp are also parasites belonging to the kinetoplastid grouping but show striking differences in genome stability and DNA replication programming relative to T. brucei . Leishmania parasites display genome-wide intra- and extra-chromosomal gene copy number variation 74 , 75 and mosaic aneuploidy 76 – 78 , each providing a means of gene expression control and adaptation. Such genome instability is more pervasive than is seen in T. brucei , where aneuploidy appears rare 79 and few episomes have been described 80 . An explanation for Leishmania ’s remarkable genome plasticity could reside in novelties in DNA replication timing. MFA-seq mapped just a single origin in each Leishmania chromosome 81 , indicating a difference in replication programming compared to the multiple origins/chromosome see in T. brucei . Later refinement of MFA-seq in Leishmania major indicated persistent, subtelomere-localised DNA synthesis throughout the cell cycle and revealed chromosome length-related replication timing, with larger chromosomes replicated later than smaller ones 82 . Whether such chromosome size-dependent replication timing results from the use of a single predominant origin in each chromosome, with the result that larger chromosomes take longer to be copied 81 , or may relate to use of abundant replication initiation activity not detected by MFA-seq 83 , is unclear. Here, we sought to determine how this potentially novel chromosome size-dependent replication timing might arise. We show that the distribution of R-loops acted upon by RNase H1 has a striking correlation with L. major chromosome length, and that loss of RNase H1 alters both DNA replication timing and genome stability. Thus, our works reveals that RNA-DNA hybrids, which are ubiquitous features of all genomes 84 , 85 , are integral to the programming of DNA replication in Leishmania and provide a mechanistic link to adaptive genome plasticity. Results Genome-wide detection of R-loops in Leishmania major R-loops have been characterised from bacteria to mammals, where they play a variety of roles under both physiological and pathological conditions 86 . For instance, R-loops are required for targeted genome rearrangements to mediate antibody class-switch recombination in mammals 87 and can contribute to genome stability by controlling various steps of DNA double strand break repair 88 – 91 . Moreover, R-loop accumulation is associated with increased local chromatin accessibility 92 , activation of gene expression 93 , 94 , and modulation of transcription termination 95 , 96 . In T. brucei R-loops have been implicated in transcription initiation and mRNA maturation 97 , 98 , telomere function 99 , and host immune evasion by antigenic variation 100 – 102 . Reflecting the myriad functions of R-loops, a wide range of proteins have been shown to interact with RNA-DNA hybrids in mammals and T. brucei 103 , 104 . Amongst these are factors that act to remove the RNA-DNA hybrids, including helicases to unwind the structures 105 , 106 and ribonucleases RNase H1 and RNase H2, which hydrolyse the RNA moiety in R-loops 107 . Unlike in T. brucei 97 , the distribution of RNA-DNA hybrids has not been examined in Leishmania . To assess this, we first examined the subcellular localization of R-loops in L. major by performing immunofluorescence using the S9.6 antibody 108 in wild type (WT) cells. Prior to detection with the S9.6 antibody, fixed cells were either left untreated or were treated with recombinant Escherichia coli RNase HI. Signal was readily detected in untreated samples but was drastically reduced in all cells examined upon pre-treatment with E. coli RNase HI, demonstrating the specificity of the S9.6 antibody for RNA-DNA hybrids in Leishmania , unlike in mammalian cells 109 . S9.6 signal localized only to the nucleus, where it was detected in a punctuated pattern ( Fig. 1A ). Download figure Open in new tab Figure 1. Subcellular localization and genome-wide mapping of R-loops in L. major . A) Immunofluorescence analysis to detect R-loops in wild type cells using S9.6 antibody; -RNase H and +RNase H indicate mock or treatment with recombinant RNase HI prior to incubation with antibody, respectively; n and k , nuclear and kinetoplast DNA, respectively. B) Snapshot showing DRIP-seq signal relative to the indicated features in a representative genomic region; from top to bottom: track 1 and 2 (green), R-loop enriched regions relative to input material; R-loop peaks are indicated as purple horizontal bars below track 1; track 3 (dark red), chromatin accessibility as determined by MNase-seq; track 4 (blue), G quadruplex structures (G4s) as determined by G4-seq; track 5 (purple), splice leader (SL) acceptor sites as determined by RNA-seq; track 6 (yellow), polyadenylation (Poly A) acceptor sites as determined by RNA-seq; grey arrows at the bottom indicate annotated coding sequences (CDSs); further genomic regions are shown in Supplementary Figure S1 and Figure S2. C) Metaplots showing global DRIP-seq signal around CDSs relative to chromatin accessibility, G4 localisation, and SL and Poly A sites; shaded areas represent 95% confidence interval. D) R-loops peak length distribution. E) Top enriched DNA sequences motifs found in R-loop peaks, as identified by MEME analysis; e -values for each motif are shown on top of each panel. To provide a genome-wide map of R-loops in L. major , we used D NA- R NA hybrid i mmuno p recipitation followed by deep seq uencing (DRIP-seq) with the S9.6 antibody. Similar to the immunofluorescence analysis, prior to DRIP, genomic DNA was either left untreated or was treated with recombinant E. coli RNase HI. Then, DRIP material was subjected to Illumina sequencing (DRIP-seq) and reads mapped to the L. major reference genome. While DRIP-seq signal was widely detected in untreated samples, it was drastically reduced at all sites analysed upon RNase HI pre-treatment, indicating DRIP-seq detects locations of RNA-DNA hybrids ( Fig. 1B, C ; see also Fig. 2A and Fig. S1, S2 and S3). To understand the localisation of RNA-DNA hybrids, we compared the distribution of DRIP-seq signal to a number of genome features, revealing that R-loops display pronounced accumulation at intergenic regions between coding sequences (CDSs) in polycistronic transcription units, where chromatin occupancy is lower (as determined by MNase-seq) 83 and G-quadruplex (G4) 110 occurrence is higher ( Fig. 1B and 1C ). At a finer level, R-loop accumulation in intergenic regions overlapped with splice leader (SL) and polyadenylation (Poly A) acceptor sites ( Fig. 1B and 1C ). To examine this association further, we identified 12,219 DRIP-seq peaks (size range ∼100-500 bp) across the L. major genome ( Fig. 1D ) and used MEME 111 to identify DNA sequences that were enriched, which revealed two motifs composed of polypyrimidines (TC n and CCT n ) and two other motifs composed of either TA or TG repeats ( Fig. 1E ). Since polypyrimidine tracts and their binding proteins have been shown to act in trans- splicing and polyadenylation across kinetoplastids 112 – 114 , these data indicate an association between R-loops and RNA processing events to generate mature mRNAs during multigene transcription in L. major that is consistent with previous observations in T. brucei 97 . Download figure Open in new tab Figure 2. Chromosome-size dependent distribution of R-loops is reflected in a range of further genetic features. A) Colourmap showing distribution of DRIP-seq signal in all 36 L .major chromosomes; chromosomes are ordered by size; -RNase H and +RNase H indicate mock or treatment with recombinant RNase HI prior to immunoprecipitation, respectively; random, indicatesDRIP-seq signal plotted after R-loops peaks positions were randomly distributed throughout the genome; an independent experiment is shown in Fig. S3. B) , C), D), E), F) and G) Colourmaps showing distribution patterns of DNA replication timing, putative origins of DNA replication (ORIs) predicted by SNS-seq, chromatin accessibility determined by MNase-seq, G4s and s hort i nterspersed de generate r etroposons 1 (SIDER1), respectively; chromosome 31, which does not follow the pattern of all other chromsomes for A, C and D, is indicated. G), H), I), J), K), L) and M) Linear regression analysis showing correlation between chromosome size and chromosome-averaged signals of DRIP-seq, DNA replication timing, ORIs, chromatin accessibility, G4s and SIDER1 sequences, respectively; R and P values are indicated at the top of each panel. N), O), P), Q) and R) Linear regression analysis showing correlation between averaged DRIP-seq signal at each chromosome and averaged signals of DNA replication timing, ORIs, chromatin accessibility, G4s and SIDER sequences, respectively; R and P values are indicated at the top of each panel; shaded areas around in panels G-S represent 95% confidence interval. The global distribution of R-loops correlates with Leishmania chromosome size-related DNA replication timing In analysing R-loop distribution we noted that, remarkably, DRIP-seq density displayed a significant correlation with L. major chromosome length ( Fig. 2A,H ; independent replicate shown in Fig. S3A), with R-loops becoming more abundant as chromosome size increases. Moreover, DRIP-seq signal increased towards the centre of each chromosome, with concomitant depletion from the sub-telomeres, indicating a gradient of R-loop levels from the chromosome cores to their ends (Fig. S3B). Importantly, these patterns were lost in samples pre-treated with E. coli RNase HI and upon randomization of DRIP-seq signal, ruling out intrinsic bias from the immunoprecipitation or mapping strategy ( Fig. 2A and Fig. S3). It could be argued that chromosome length is unlikely to be the determinant of such an unusual R-loop distribution, but that it reflects asymmetric activities across the genome. Therefore, we tested how R-loop distribution correlates with known features of the L. major DNA replication programme. Our previous work 82 using MFA-seq showed larger L. major chromosomes are, on average, replicated later when compared to smaller chromosomes ( Fig. 2B and 2I ), with R-loop density and chromosome replication timing therefore showing a significant anti-correlation ( Fig. 2O ). Lombrana et al 83 used SNS-seq to map thousands of predicted DNA replication origins in L. major . In light of our R-loop mapping, we reanalysed the SNS-seq data and again found a significant chromosome size-dependence, with larger chromosomes presenting a lower density of predicted origins than smaller ones ( Fig. 1C and 1J ). As a result, SNS-seq origin density significantly negatively correlates with the average R-loops level of each chromosome ( Fig. 2P ). This analysis indicates that chromosome size-dependent replication timing is reflected in a similarly skewed distribution of R-loops and SNS-seq predicted DNA replication initiation events. Global R-loop distribution correlates with chromosome size-related chromatin accessibility, G-quadruplex levels and DNA sequence content We next asked if the global DRIP-seq pattern is reflected in wider features of L. major chromosome activity and sequence composition. First we looked at chromatin occupancy by remapping available MNase-seq 83 data across the genome, which revealed higher occupancy as chromosomes reduce in size ( Fig. 2D, 2K ), meaning there is a significant anticorrelation with DRIP-seq signal ( Fig. 2Q ). This finding indicates that larger chromosomes with higher R-loop levels present relatively more relaxed chromatin when compared to smaller ones, suggesting that chromatin accessibility is not just a determinant of local R-loop accumulation ( Fig.1B, C ), but of global distribution as well. Re-analysis of G4-seq data revealed a similar and significant anticorrelation with DRIP-seq, with higher G4s level in the smaller chromosomes than the larger ( Figs. 2E, L, R ). Thus, the presence of G4s is a good predictor of R-loop accumulation at the local level, but not at a genome-wide level. Remarkably, we next observed that R-loop, chromatin and G4 levels are reflected in size-dependent variation in L. major chromosome sequence content. First, we examined the distribution of direct and inverted repeat DNA sequences, including s hort i nterspersed de generate r etroposons 1 and 2 (SIDER1 and SIDER2, respectively), which are related to gene copy number variation (CNV) 115 and regulation of gene expression 116 , 117 in Leishmania . Both non-SIDER and SIDER2 repeats appear to be present in homogenous levels across chromosomes (Fig. S4A, S4B, S4E and S4F). Also, we found no evidence that GC or AT skew displays any correlation with chromosome length (Fig. S4C, S4D, S4G and S4H). However, SIDER1 repeats were found to be enriched in larger chromosomes when compared to smaller ones ( Fig. 2F and 2M ), significantly correlating with DRIP-seq levels ( Fig. 2S ), and GC content decreased as chromosome size increased ( Fig. 2G and 2N ), displaying a significant anticorrelation with DRIP-seq levels ( Fig. 2T ). Altogether, these observations indicate a previously undetected asymmetry in many aspects of chromosome content and function in L. major : patterns of R-loop accumulation among chromosomes can be correlated not only with chromatin accessibility and the unconventional DNA replication timing programme of the parasite, but also with the evolution of genome architecture, as reflected in sequence content biases among chromosomes. L. major RNase H1 localizes to the nucleus and associates with strand switch regions To test for potential functional consequences of R-loop distribution in L. major , we next focused on the factors involved in RNA-DNA hybrid resolution. Two ribonuclease H enzymes are known to participate in the resolution of R-loops in eukaryotes 107 , including T. brucei 98 , 100 . In L. major one predicted subunit of trimeric RNase H2 has been reported to act in the mitochondrion 118 , and so we focused on RNase H1, which has not been studied in Leishmania but provides nuclear functions in T. brucei 100 . We used CRISPR/Cas9 to flank the endogenous RNase H1 ORF with loxP sites (flox), allowing the gene to be deleted by rapamycin-mediated induction of DiCre activity (Fig. S5A). In addition, the RNase H1 ORF was translationally fused with 6 copies of the HA epitope at the C-terminus to allow us to monitor protein levels and location before and after gene excision. PCR showed that all copies of RNase H1 were floxed and HA-tagged after a single round of transformation (Fig.S5B); the resulting cell line is hereafter referred to as RNase H1-HA Flox . Simultaneous addition of loxP sites and the HA tag did not result in any significant growth defect in RNAse H1-HA Flox cells compared with WT (Fig.S5C). To investigate the subcellular localization of RNase H1 in L. major and its potential relationship with DNA replication, we employed immunofluorescence analysis with RNAse H1-HA Flox cells pulsed with EdU to label cells in S phase. These experiments indicated that RNase H1-HA primarily localized to the nucleus and showed a higher abundance in non-replicating cells ( Fig. 3A ). Based on this, we conclude that RNase H1 provides nuclear functions in L. major and does not appear to be temporally or spatially linked to replicative DNA synthesis. Download figure Open in new tab Figure 3. Subcellular localization and genome wide mapping of RNase H1 in L. major A) Immunofluorescence analysis to detect RNase H1-HA using anti-HA antibody; cells undergoing DNA replication are shown by EdU signal; n and k indicate nuclear and kinetoplast DNA, respectively. B) Snapshot of RNase H1-HA ChIP-seq in two representative chromosomes; from top to bottom: track 1 (green), enriched regions of acetylated Histone H3 (AcH3); track 2 (dark red), β-D-glucosyl-hydroxymethyluracil (Base J) enriched regions; track 3 (dark grey), RNase H1-HA enriched regions relative to input material; grey arrows at the bottom indicate the position and orientation of polycistronic transcription units (PTUs). C) Metaplots (top) and colourmap (bottom) showing RNase H1-HA ChIP-seq signal around convergent, divergent and head-to-tail strand switch regions (SSR-Conv, SSR-Div and HT, respectively); metaplots for AcH3 and Base J are also shown. D) Metaplot showing global RNase H1-HA ChIP-seq signal around CDSs. E) Linear regression analysis showing correlation between chromosomes length and averaged signals of RNase H1-HA ChIP-seq at each chromosome in unperturbed exponentially growing cells (NT) and after release from synchronization with hydroxyurea (HU); cell cycle progression analysis upon HU synchronization is shown in Supplementary Figure S9; R and P values are indicated at the top of each panel; shaded areas in panels D and E represent 95% confidence interval. Next, we performed chromatin immunoprecipitation followed by deep sequencing (ChIP-seq) with anti-HA antiserum to characterise the genome-wide binding profile of RNase H1-HA. This analysis showed that RNase H1-HA is most strongly enriched at the boundaries of the polycistronic transcription units (PTUs), termed strand switch regions (SSRs), co-localizing with acetylated histone H3 (AcH3) 119 and β-D-glucosyl-hydroxymethyluracil (Base J) 120 , markers of transcription initiation and termination sites, respectively ( Fig. 3B and 3C ). We also observed RNase H1-HA enrichment at intergenic regions ( Fig. 3D ), where R-loops are found to accumulate ( Fig. 1B,C ). Given the chromosome-size dependent distribution of R-loops ( Fig.2 ), we next asked if RNase H1 might accumulate across the genome in a similar way. Quantification of the ChIP-seq signal indicated that RNase H1-HA does not associate equally to all chromosomes, since we found a significant correlation between averaged RNase H1-HA ChIP-seq signal for each chromosome and its length in asynchronous cultures ( Fig. 3E , first panel). Moreover, this correlation was lost when L. major cells were arrested in G1 by hydroxyurea treatment, but increased as cells were released from G1 arrest and synchronously navigated through S-phase (3 hrs) and G2/M (6 hrs; Fig. 3E , second to fourth panels; cell cycle progression upon synchronisation with hydroxyurea is shown in Fig. S9A). These analyses suggest two things. First, the global distribution of L. major RNase H1 mirrors that of the R-loops that it acts to dissolve, with greater accumulation in larger chromosomes. Second, despite no evidence of overlap between RNase H1-HA signal and DNA synthesis detected by EdU, chromosome-size dependent accumulation of RNase H1 is cell cycle-dependent and is more marked during S-phase to G2/M transition, perhaps reflecting Leishmania chromosome size-dependent DNA replication timing. Loss of L. major RNase H1 results in a transient growth defect and R-loop accumulation under DNA replication stress To evaluate the consequences of RNase H1 loss, conditional knockout (KO) of the floxed RNase H1-HA gene was induced by rapamycin-mediated DiCre activation in logarithmically growing cultures of L. major ( Fig. 4A ). DiCre-mediated loss of the protein was confirmed by western blotting, where signal for RNase H1-HA was no longer detectable after 48 h of the second round of KO induction ( Fig. 4B , Fig.S6). Genome-wide DNA and RNA sequencing showed loss of the RNase H1 gene and its RNA upon induction of DiCre activity, confirming precise DiCre excision (Fig. S7). The timing of RNase H1-HA loss coincided with marked slowing in growth of the induced cells compared with uninduced, an effect that continued for around 20 days (until passage 5, approximately 40 population doublings for WT cells) but was followed by recovery, with the DiCre-induced cells’ growth becoming indistinguishable from uninduced cells by passage 6 ( Fig.4C ). Importantly, PCR analysis showed that unexcised copies of RNase H1-HA were undetectable in induced cells throughout the course of the experiment, ruling out the possibility that reversion of the growth defect was due to restoration of the RNase H1 gene or outgrowth of cells with unexcised floxed gene ( Fig. 4D ). These data suggest that loss of RNase H1 leads to deleterious effects in L. major promastigotes in the short-term, from which cells recover in the long-term. The data also suggest that loss of RNase H1 is not lethal. To test this prediction, we subcloned the DiCre induced cells after passage 2 ( Fig.4C , Fig.S6). A clonal cell line (Fig. S5D and S7), hereafter referred as KO, was recovered that showed no evidence for the presence of RNase H1 gene copies or mRNA (Fig.S7) and did not present any detectable growth defect compared with WT cells ( Fig. 4E ). Altogether, these data indicate that loss of RNase H1 is transiently detrimental to L. major proliferation, but the parasites can adapt and recover fecundity. In addition, the KO cells provides a means of comparing short- and long-term effects resulting from RNase H1 loss. Download figure Open in new tab Figure 4. Effects of RNase H1 loss on growth, R-loop accumulation and DNA replication timing. A) Schematic representation of the DiCre-mediated RNase H1 gene deletion strategy; CRISPR-Cas9 was used to flank the RNase H1 ORF with LoxP sites and fuse it with an HA tag ( RNase H1-HA Flox ); rapamycin-mediated activation of DiCre was used to catalyse excision of RNase H1-HA Flox ; refer to Supplementary Figure S6 for the rapamycin induction strategy; a and b , annealing position of primers used in D. B) Western blotting analysis of whole cell extracts from RNase H1-HA Flox cells ∼48 h after growth in the absence (-RAP) or in the presence (+RAP) of rapamycin in two passage stages of growth at passages 1 and 2 (P1 and P2, as shown in C); extracts were probed with anti-HA antibody and anti-EF1α was used as loading control. C) Growth profile of the RNase H1-HA Flox cell line cultivated in the absence (- RAP, black) or presence (+ RAP, grey) of rapamycin; cells were seeded at ∼10 5 cells.mL -1 at day 0 and diluted back to that density every 4 - 5 days for seven passages (P1 to P7); cell density was assessed every 24 h and error bars depict standard error of the mean (SEM) from two independent experiments. D) PCR analysis of genomic DNA xtracted from RNase H1-HA Flox cells ∼48 h in the indicated passages, after growth in the absence (-RAP) or in the presence (+RAP) of rapamycin; annealing positions for primers a and b are shown in A. E) Growth profile of a clonal RNase H1 KO cell line, selected after DiCre-mediated RNase H1 gene deletion compared to wild type (WT) cells . F) Immunofluorescence analysis using S9.6 antibody to detect R-loops with (+HU) or without (-HU) 5 mM hydroxyurea treatment for 6 hours. G) Quantification of R-loops levels detected via immunofluorescence using S6.9 antibody in the indicated conditions; -RNase H and +RNase H indicate mock or treatment with recombinant RNAse HI prior to incubation with antibody, respectively; (****),(**) and ns: p<0.0001, p=0.0089 and not significant, respectively, as determined by Kruskal-Wallis test. H) Representative snapshot showing DNA replication timing as determined by MFA-seq using exponentially growing cells normalised with stationary cells; positive and negative values indicate early and late replicating regions, respectively; grey arrows at the bottom indicate the position and orientation of polycistronic transcription units (PTUs). I) and J) Metaplots showing global MFA-seq signal in early and late replicating regions, respectively; (****),(**), (*) and ns: p<0.0001, p=0.0089, p=0.0188 and not significant, respectively, as determined by Kruskal-Wallis test. K) Linear regression analysis showing correlation between chromosome size and averaged MFA-seq signal; R and P values are indicated at the top of each panel; shaded areas represent 95% confidence interval. Next, we asked if RNase H1 loss leads to accumulation of R-loops. For this, we performed immunofluorescence analysis using the S9.6 antibody, comparing the levels of nuclear R-loops signal in uninduced cells relative to when RNase H1 excision caused growth impairment and also in the KO cells. Surprisingly, we did not observe any significant difference in S9.6 signal in any of these conditions ( Fig. 4F and 4G ). However, when cell cycle progression at G1/S was blocked via hydroxyurea treatment, we observed a significant accumulation of R-loops upon both short- and long-term RNase H1 loss ( Fig. 4F,G ). Importantly, this effect appears specific to DNA replication stress resulting from hydroxyurea exposure, since treatment with camptothecin (blocks DNA replication by inhibiting the activity of topoisomerases) 121 , actinomycin D (blocks transcription elongation) 122 , flavopiridol (blocks cells cycle at G2/M by inhibiting CRK3 Cyclin-Dependent Kinase) 123 or AB1 (blocks cells cycle at G2/M by inhibiting kinetochore assembly) 124 did not result in significant change in nuclear R-loop signal (Fig. S8). These data suggest RNase H1 is dispensable for R-loop resolution either because R-loops do not accumulate under normal growth conditions, or because of functional compensation by other activities, such as RNase H2. Alternatively, it is possible that R-loops do accumulate upon loss of RNase H1 under unstressed conditions, but only in discrete genomic locations, thus escaping detection via immunofluorescence. Nonetheless, the pronounced change in R-loop levels in the absence of RNase H1 after hydroxyurea treatment lends further weight to a cell cycle-dependent role for RNase H1. Loss of RNase H1 abrogates Leishmania chromosome size-dependent DNA replication timing Since the distribution of R-loops and RNase H1 has parallels with chromosome size-dependent timing of L. major DNA replication 125 ( Fig. 2 and Fig. 3 ), we reasoned that the ribonuclease could be a hitherto undetected player that directs the DNA replication programme of the parasite. To test this prediction, we performed MFA-seq analysis in uninduced and induced RNase H1-HA Flox cells at passage 2, as well as in the KO cells, allowing us to compare the short- and long-term effects of RNase H1 loss. The MFA-seq profile in uninduced RNase H1-HA Flox cells corresponded with previous reports 81 , 125 , with a single pronounced peak in each chromosome that overlapped with an SSR ( Fig. 4H ). However, when comparing the MFA-seq profile with both the induced RNase H1-HA Flox cells and the KO cells a striking change emerged: MFA-seq signal around every early-replicating SSR was decreased ( Fig. 4H and 4I ). Concomitantly, increased MFA-seq signal was observed across late replicating PTUs in both the induced RNase H1-HA Flox cells and the KO cells ( Fig. 4H and 4J ). This result suggests that RNase H1 activity is required for maintenance of the DNA replication programme in L. major . To further test this, we performed linear regression analysis of averaged MFA-seq signal for each chromosome against its length and compared the resulting profile from uninduced cells with those from induced and from KO cells. Strikingly, the correlation between replication timing and chromosome length was dramatically weaker in induced cells and absent in the KO cells ( Fig. 4K ). Interestingly, this change in DNA replication timing upon loss of RNase H1 was not accompanied by pronounced cell cycle progression defects (Fig. S9A), or by alterations in the proportion of cells in S phase (Fig. S9B). Altogether, these data suggest that RNase H1 has a pivotal role in the DNA replication programme of L. major and that reversion of a growth defect seen shortly after RNase H1 loss correlates with abrogation of the major DNA replication timing dynamic, which is chromosome length-dependent. Loss of RNase H1 leads to genome instability Because DNA replication is key for genome maintenance and transmission, we set out to investigate if the shift in the replication program upon loss of RNase H1 might affect genome stability by performing short-read Illumina whole genome sequencing to test for levels of CNV events during growth. We did this by calculating fold change in normalized read numbers from uninduced and induced RNase H1-HA Flox cells upon short-term cultivation (three passages), as well as from long-term cultivation (KO cell line) across the entirety of each chromosome ( Fig. 5A ). This genome-wide analysis revealed that short term cultivation of induced RNase H1-HA Flox cells displayed a modest increase in CNV relative to uninduced cells, and that this change was more prominent in the cores of chromosomes (where R-loops are more abundant; Fig. 2 ) and increased to a greater extent in the larger chromosomes compared with the small ones (again reflecting R-loop abundance; Fig. 5B ). This chromosome size- and location-dependent change in CNV was more clearly seen when comparing the KO cells to uninduced RNase H1-HA Flox cells ( Fig. 5B ), indicating that the CNV change was greater after prolonged cultivation of RNase H1 KO cells. In addition, this analysis revealed apparent loss of DNA sequences in the KO cells at subtelomeres and at the repetitive ribosomal- and SL-RNA-encoding loci ( Fig. 5C ), which in L. major (Fig. S2) and other organisms are a pronounced region of R-loop accumulation and where loss of RNase H affects their stability 97 , 126 – 128 . Download figure Open in new tab Figure 5. Analysis of genome alterations upon DiCre-mediated RNase H1 gene deletion. A) Schematic representation showing time points from which cells were collected for whole genome sequencing (WGS). B) Colourmap showing genome-wide relative copy number variation (CNV) analysis; chromosomes are ordered by size; CNV in RNase H1-HA Flox cells is expressed as log 2 [ratio(normalised reads from P7/normalised reads from P4)] for either –RAP or +RAP conditions; CNV in KO cell line is expressed as log 2 [ratio(normalised reads from KO/ normalised reads from P4 - RAP)]; chromosomes 05 and 12, showing decreased copy number, are indicated. C) Relative CNV analysis at the indicated loci ; CNV is expressed as in B. D) Absolute chromosome CNV analysis for the indicated chromosomes in the indicated conditions and passages; violin plots represent the distribution of normalized read counts relative to the haploid genome content; (****),(***) and (*): p<0.0001, p=0.0005 and p=0.0188, respectively, as determined by Kruskal-Wallis test. E) and F) Metaplots (top) and colourmaps (bottom) showing normalised density of new SNPs and InDels, respectively, around annotated coding sequences (CDS). G) and H) SNP and InDel densities in chromosomes grouped by length; smaller: 0.268 – 0.622 Mb, medium: 0.629 – 0.840 Mb, larger: 0.913 – 2.68 Mb; (**), (*) and ns: p=0.0068, p=0.0155 and not significant, respectively, as determined by Kruskal-Wallis test. Genome sequencing also revealed both dramatic (chromosomes 05 and 12) and milder (chromosomes 04, 14 and 31) whole-chromosome CNV in the KO cells, as well as modest but significant whole-chromosome CNV during the limited growth period after DiCre induction (chromosomes 04, 05, 14, 20 and 31; Fig. 5B and 5D ). The median of normalized coverage for chromosomes 04, 12, 14 and 20 from the starting, uninduced cultures indicate they were nearly euploid, with 2.1, 3.1, 2.1 and 3.0 copies, respectively. Upon either short- or long-term growth after RNase H1 loss their copy number dropped to 1.84, 2.16, 2.95 and 2.56, respectively. On the other hand, chromosomes 05 and 31, which were aneuploidy in the starting uninduced cells with 2.7 and 5.2 median copies, respectively, showed a drop to 2.24 and 4.86 during short-term cultivation, respectively. Altogether, these data suggest that reprograming of the DNA replication landscape upon RNase H1 loss leads to genome-wide CNV and alterations in ploidy control, with such effects being more severe after long-term growth. The loss of subtelomere, ribosomal and SL sequences suggests that RNase H1 is required for the maintenance of repetitive DNA, possibly by preventing R-loops accumulation at DNA replication stress-prone regions. It is also possible that at least some of the stable changes in karyotype may explain reversion of the short-term growth defect after RNase H1 loss, consistent with the idea that aneuploidy is an adaptative strategy relying on polyclonal selection of pre-existing karyotypes 129 , 130 . Loss of RNase H1 leads to chromosome size-dependent mutagenesis To ask if genome instability arising during short- and long-term growth after RNase H1 loss is limited to CNV and karyotype changes, we tested for the appearance of single nucleotide polymorphisms (SNPs) and small insertions and deletions (InDels). For this, we identified new SNPs and InDels that arose during growth between passage four and passage seven for induced and uninduced RNase H1 Flox cells, as well as in KO cells relative to uninduced RNase H1 Flox cells at passage four. First, we examined the density of these mutations at intergenic regions, where R-loops are enriched ( Fig. 1C ) and observed the mutation patterns differed when comparing short- and long-term growth in the absence of RNase H1: a more pronounced accumulation of SNPs in these regions was seen in the induced RNase H1 Flox cells ( Fig. 5E ), while an increased accumulation of InDels was seen in the KO cells ( Fig. 5F ). In both cases, SNPs and InDels accumulated asymmetrically around many CDSs, which we speculate may be due to differing orientations of collisions between the transcription and DNA replication machineries, leading to uneven accumulation of R-loops 131 . We also observed a slight increase in transitions (Ts) relative to transversions (Tv) in the KO cells compared to the uninduced and induced RNase H1 Flox cells (Fig. S10A). pLogo enrichment analysis 132 confirmed this by showing enrichment of mutations in T and A residues, and further revealing that these mutations were not randomly distributed, but were preferentially flanked by G or C residues in the KO cells (Fig. S10B). We conclude this demonstrates that RNase H1 is required for genome stability and the distinct mutagenic profiles upon short- and long-term cultivation is consistent with reversion of the short-term growth defect after RNase H1 loss. We have previously observed that SNP occurrence among chromosomes correlates with their length 133 , and therefore possibly with differential DNA replication timing. To ask how the loss of size-dependent chromosome DNA replication timing seen in the absence of RNase H1 ( Fig. 4K ) would affect SNPs and InDels accumulation, we examined the density of these mutations, grouping chromosomes by length. This analysis confirmed the chromosome length-related accumulation of these mutations in uninduced cells and revealed that this effect is more pronounced in both short- and long-term growth in the absence of RNase H1 ( Fig. 5G and 5H ). Altogether, these analyses, when considered alongside the CNV data, illustrate that R-loops acted upon by RNase H1 have widespread effects on L. major genome instability, which vary in pattern across the genome and during growth in absence of the RNase H1, reflecting the complexity of global and localised R-loop association with sequence features and chromosome size. Thus, DNA replication timing is intimately linked with Leishmania genome plasticity through R-loops. Discussion Here, we have used DRIP-seq and genetic analysis to map the localization and function of R-loops and RNase H1 in the nuclear genome of the eukaryotic parasite L. major . R-loops are remarkably abundant in the parasite’s genome and display a distribution that, both locally and globally, correlates with features such as chromatin accessibility, G4 formation and sequence composition, all of which show a remarkable parallel with Leishmania ’s unconventional chromosome length-related DNA replication timing programme. Our work indicates that these correlations stem from a functional relationship between R-loops and DNA replication, since we show that loss of RNase H1 abrogates the differences in DNA replication timing between large and small chromosomes and results in genome-wide, chromosome size-dependent increases in genome instability. Since R-loops are ubiquitous epigenetic features of all genomes, we suggest that RNA-DNA hybrids may be widespread, hitherto unappreciated determinants of DNA replication programming and resulting patterns of genome variation in many eukaryotes. The local patterns of R-loop and RNase H1 enrichment in L. major reveal considerable intersection with transcription. R-loop enrichment at intergenic regions in the L. major genome is remarkably similar to R-loops mapped in T. brucei 97 , 98 . Hence, this work substantiates a novel association between R-loops and pre-mRNA processing across kinetoplastids, which appears distinct from roles in transcription initiation and termination seen in other eukaryotes and reflects the ubiquity of multigenic RNA Pol II transcription in kinetoplastids 105 , 134 . Nonetheless, it remains unclear if kinetoplastid R-loops in inter-CDSs locations simply form as a by-product of pre-mRNA processing, perhaps due to RNA Pol slowing, or if they actively participate to concentrate or organise RNA processing factors and maximize the efficiency of co-ordinated mRNA maturation during multigenic transcription. It is conceivable that G4s, together with R-loops, are a hitherto unrecognised part of mRNA processing orchestration or act to modulate RNA Pol movement. To date, no work has localised any RNase H enzyme in a kinetoplastid genome. Here, we describe mapping of RNase H1 by ChIP-seq and show that its enrichment across the PTUs largely mirrors DRIP-seq, indicating the enzyme acts on intergenic R-loops that arise during transcription elongation. This may suggest these R-loops are the equivalent of class II RNA-DNA hybrids that form during transcription and have been described in other eukaryotes 135 , though as noted above, they may be unique to kinetoplastids due to the ubiquitous need for pre-mRNA processing by trans-splicing and polyadenylation. More pronounced RNase H1 enrichment is seen at transcription start and stop sites in L. major , with the former localisation consistent with the formation of class I R-loops during RNA Pol II pausing as transcription initiates 135 . These data are also consistent with R-loop 97 and RNA Pol II enrichment 136 , 137 at transcription start sites in T. brucei , indicating widely conserved RNA Pol II pausing during transcription initiation. Loss of RNase H2A in T. brucei leads to pronounced damage accumulation at transcription start sites 98 , which is not seen in RNase H1 mutants 100 . As we have not mapped DNA damage in L. major RNase H1 mutants, and no analysis of Leishmania RNase H2 function has been described, it is too early to say if the two kinetoplast parasites differ in how RNA Pol II transcription initiates. On a global level, the unanticipated chromosome size-dependent distribution of R-loops in L. major has not to our knowledge been described to date in any other eukaryote, suggesting it may be a novel feature of genome biology in Leishmania . Moreover, this global distribution of R-loops is reflected in several other chromosome-size related genome features ( Fig.2 ): chromatin compaction (based on MNase-seq), G4 density (G4s-seq), putative origin density (SNS-seq) and GC content are all greater on the smaller chromosomes than the larger, while SIDER1 is more concentrated in larger chromosomes. These observations suggest that global R-loop distribution reflects activities that have resulted in a chromosome size-dependent patterning of many aspects of genome content and activity in L. major . One explanation that might be considered as the basis for the connection between these aspects of the genome is reduced nucleosome density in the larger chromosomes, as this may allow greater levels of R-loops to accumulate, meaning the global correlation between chromatin status and R-loops reflects the localised coordination of these features at inter-CDS regions. Such decreased chromatin compaction may also allow for better resolution of G4s structures in the larger chromosomes, leading to reduction in the replication initiation activity detected by SNS-seq (which is associated with the presence of G4s) 138 . However, what aspect of Leishmania biology might necessitate a gradient of nucleosome density across its chromosomes is unclear. For instance, no work has described greater levels of gene expression as chromosome size increases. A more compelling activity to explain all the above observations is Leishmania DNA replication programming and, as a result, timing. We have previously documented, through MFA-seq, that coordinated initiation of DNA replication in early S phase is localised to a single locus in each chromosome of L. major promastigotes 81 , 82 . Whether each locus is just a single origin is unclear 139 , as is whether or not they are the sole site of DNA replication initiation in each chromosome 83 , 140 . Nonetheless, programming of DNA replication to predominantly initiate from a single origin or locus per chromosome would explain the chromosome size-dependent timing of L. major DNA replication we describe here and previously 82 . As we have argued 70 , 81 , 141 , 142 , it is unlikely that whole genome duplication in Leishmania can be accomplished during S-phase using a single origin per chromosome, and so DNA replication may be supported by further, less efficient initiation activities. Such organisation could then explain Leishmania DNA replication timing, feeding into the other genome features we describe. An unknown feature of the above model is the nature of any DNA replication initiation events beyond the predicted single MFA-seq-detectable centromeric 143 origin on each Leishmania chromosome. Any explanation for this putative ‘supplementary’ replication activity must account for chromosome size-dependent replication timing. In this light, two possibilities may be considered. First, the MNase-seq gradient we describe may not reflect decreasing chromatin compaction as chromosome size increases, but instead that the larger chromosomes are closer to the nuclear periphery and therefore more susceptible to MNase digestion, given that the dataset was generated using isolated and permeabilised nuclei 83 . If so, chromosome positioning within the L. major nucleus may be a determinant of replication timing in a similar way to other eukaryotes, where less efficient, late replicating loci are at the lamina-rich nuclear periphery 28 , 29 . If correct, chromosome subnuclear positioning does not influence replication activation at the single, main locus in each chromosome, most likely because it corresponds to the centromere 143 and these early-acting origins may overcome spatial controls 1 , 9 . Instead, the use of ‘supplementary’ origins may be more influenced by nuclear position, explaining why SNS-seq predicts greater numbers of initiation events in the smaller chromosomes ( Fig.2 ). R-loops can be an impediment to DNA replication that is exacerbated upon RNase H1 loss 144 . Hence, the greater abundance of R-loops as L. major chromosome size increases may reflect less efficient DNA replication as initiation events detected by SNS-seq become sparser. A problem with this explanation is our demonstration that loss of L. major RNase H1 leads to earlier replication of the larger chromosomes, since the likely increase in R-loops would be predicted to further impede DNA replication. Hence, a second explanation for DNA replication timing in L. major could be that R-loops mediate replication initiation, as has been suggested elsewhere 145 – 148 . If correct, the chromosome-size dependent enrichment of R-loops may be explained: greater numbers of R-loops are needed to support replication of the larger chromosomes, where replicative DNA synthesis from a single constitutive origin duplicates less of the molecule during S-phase compared with a smaller chromosome. In addition, this explanation would explain MFA-seq mapping after RNase H1 loss: increased levels of R-loops would allow earlier replication of the larger chromosomes, resulting in loss of size-dependent DNA replication timing. In yeast 145 and bacteria 149 priming of DNA replication initiation upon aberrant accumulation of R-loops has been seen previously. However, the analysis we conducted here did not reveal global R-loop accumulation upon RNase H1 loss under normal growth conditions, but only under DNA replication stress, which suggests two things (that are not mutually exclusive): accumulation of R-loops after RNase H1 loss may be concentrated in a few regions of the genome (possibly in those prone to DNA replication stress), but is sufficient to mediate changes in the DNA replication programme; a hitherto undetected R-loop resolution-independent function of L. major RNase H1 may mediate the temporal order of DNA replication among chromosomes. Further work is needed to define where and how DNA replication initiation occurs across the Leishmania genome, including asking why changes in chromosome replication timing upon loss of RNase H1 are not associated with detectable change in cell cycle progression. Nonetheless, our demonstration that loss of RNase H1 results in increased L. major genome variability in patterns reflecting local and global accumulation of R-loops, as well as chromosome ploidy change, provides a mechanistic link between RNA-DNA hybrids, differential chromosome replication timing and homeostasis of the plasticity of the parasite’s genome. The accumulation of SNPs and InDels upon RNase H1 loss is locally most prominent in intergenic regions and may be explained by R-loop-mediated DNA synthesis reactions that arise from unscheduled (re-)initiation, or are mediated by error prone 150 or repair-associated DNA polymerases 151 . In this regard, in Saccharomyces cerevisiae , loss of both RNase H enzymes leads to unscheduled DNA synthesis, which is associated with increased nuclear DNA damage 152 . R-loops can also lead to genome instability by impeding DNA replication 145 and through replication-transcription clashes 131 , 153 . Such processes may explain the global change in mutation patterns that are most prominent after upon RNase H1 KO. Alternatively, or in addition, earlier traversal of DNA replication across R-loops in the smaller chromosomes relative to the larger could explain the increasing burden of SNPs and InDels as chromosome size decreases. It will be interesting to determine how RNase H1 mutation affects the ability of Leishmania to adapt to changing environments, which has been associated with genome instability 129 , 154 , 155 . Though this study details a factor that directs Leishmania DNA replication programming and links it to genome stability, the biological rationale for the parasite organising DNA replication timing based on chromosome size is unclear. In this regard, we cannot currently say in Leishmania if R-loops might be allele-specific and provide a form of potentially non-coding RNA-associated ASARs that contribute to replication timing 55 . However, it is notable that L. major chromosome 31, which is always greater than diploid 156 , does not follow the global pattern of R-loop or SNS-seq density, or of chromatin accessibility. In T. brucei , monoallelic transcription of one the ∼15 telomeric VSG expression sites 157 , 158 is intimately tied with DNA replication timing 72 , and mutation of RNase H1 or RNaseH2A impairs this expression control, an effect that is associated with increased R-loops across all VSG expression sites 98 , 100 . Such connections between replication asynchrony, monoallelic expression and chromatin accessibility appear to mirror activities described in mammals 159 and, thus, R-loops may have wider and so far unexplored impacts on how gene expression and DNA replication intersect in kinetoplastids. Methods Parasite culture and generation of an RNase H1-HA Flox cell line Promastigotes derived from Leishmania major strain LT252 (MHOM/IR/1983/IR) were cultured at 26 °C in HOMEM medium supplemented with 10% heat-inactivated foetal bovine serum. For transfections, exponentially growing cells were electroporated using Amaxa Nucleofactor™ II (pre-set program X-001). The RNase H1-HA Flox cell line was generated in three sequential transfection and selection rounds. First, a cell line expressing DiCre from the rRNA encoding locus was established. For this, a wild type strain was transfected with pGL2339 plasmid 160 , previously digested with Pac I and Pme I. DiCre-expressing clones were selected with 10 μg.mL -1 blasticidin. Second, this cell line was further modified to concomitantly express Cas9 and T7 RNA Pol from the β-tubulin array. For this, the DiCre-expressing cell line was transfected with plasmid pTB007 161 , previously digested with Pac I. DiCre, Cas9 and T7-expressing cells were selected in with 10 μg.mL -1 blasticidin and 20 μg.mL -1 hygromycin. Finally, by taking advantage of the Cas9/T7 system, as previously described 161 , we fused all copies of RNase H1 with six copies of HA while also flanking RNase H1-6xHA with LoxP sites, in a single round of transfection. For this, ORF LmjF.06.0290 encoding RNase H1 was PCR-amplified and cloned between the Nde I and Spe I restriction sites in the vector pGL2314 162 . This construct was used as a template in a PCR reaction to generate the donor fragment containing homology flanking arms. Following ethanol precipitation, the donor fragment was transfected together with sgRNA templates into the DiCre, Cas9 and T7-expressing cell line. RNase H1-HA Flox cells were selected with 10 μg.mL -1 blasticidin, 20 μg.mL -1 hygromycin and 10 μg.mL -1 puromycin. Generation of sgRNAs templates and homology arms was performed as previously described 161 . In each transfection, selection was carried out by limiting dilution in 96-well plates in the presence of the appropriate antibiotics. Integration into the expected locus was confirmed by PCR analysis. Induction of DiCre for RNase H1 KO was performed with rapamycin, as previously reported 162 , 163 . Antibodies Mouse anti-HA (1: 5000, Sigma), mouse anti-EF1α (1: 40 000, Merck Millipore), anti-BrdU clone B44 (1: 500, BD Bioscience), and anti-DNA-RNA hybrid clone S9.6 (1:500, Sigma) primary antibodies were used here. Goat anti-Mouse IgG HRP-conjugated (ThermoFisher), goat anti-Mouse IgG Alexa Fluor 488-conjugated (ThermoFisher) and goat anti-Mouse IgG Alexa Fluor 594-conjugated (ThermoFisher) secondary antibodies were also used. Western blotting Whole cell extracts were prepared by collecting cells by centrifugation, washing with 1xPBS, resuspending in NuPAGE™ LDS Sample Buffer (ThermoFisher) supplemented with 5% β- mercaptoethanol, and heating to 95°C for 10 minutes. Whole cell extracts were resolved on 4-12% gradient Bis-Tris Protein Gels (ThermoFisher) and transferred to Polyvinylidene difluoride (PVDF) membranes (GE Life Sciences). Membranes were first blocked with 10% non-fat dry milk dissolved in 1xPBS supplemented with 0.05% Tween-20 (PBS-T) for 1 h at room temperature. Next, membranes were probed with primary antibody for 2 h at room temperature, diluted in PBS-T supplemented with 5% non-fat dry milk. After extensive washing with PBS-T, membranes were incubated with HRP-conjugated secondary antibodies in the same conditions as the primary antibodies. Finally, membranes were extensively washed with PBS-T. To detect and visualize bands, membranes were incubated with ECL Prime Western Blotting Detection Reagent (GE Life Sciences) and exposed to Hyperfilm ECL (GE Life Sciences). Detection of cells in S phase using flow cytometry Exponentially growing cells were incubated with 150 μM BrdU for 30 mins, collected by centrifugation and washed with 1xPBS. Washed cells were resuspended in ethanol:1xPBS (7:3) and then incubated at -20 °C for at least 16 h. Fixed cells were collected by centrifugation, washed with washing buffer (1xPBS supplemented with 1% BSA) and subjected to DNA denaturation with 2N HCL for 30 mins at room temperature. The reaction was neutralised with phosphate buffer (0.2 M Na 2 HPO 4 , 0.2 M KH 2 PO 4 , pH 7.4), cells were collected by centrifugation, and further incubated with phosphate buffer at room temperature for 30 mins. Next, phosphate buffer was removed and cells were incubated with anti-BrdU antibody diluted in washing buffer supplemented with 0.2% Tween20 for 1h at room temperature. Cells were washed with washing buffer, then incubated with anti-mouse secondary antibody conjugated with Alexa Fluor 488 diluted in washing buffer supplemented with 0.2% Tween20 for 1 h at room temperature, followed by multiple washes with washing buffer. Finally, cells were incubated for 30 mins at room temperature with 10 μg.mL -1 Propidium Iodide and 10 μg.mL -1 RNase A diluted in 1xBPS, passed through a 35 μm nylon mesh and examined with FACS Celesta (BD Biosciences). Further data analysis and processing was performed with FlowJo software. Negative controls, in which anti-BrdU antibody was omitted, were used to discriminate BrdU-positive and BrdU-negative events. Detection of cells in S phase and RNase H1-HA using microscopy Exponentially growing cells were incubated with 150 μM of EdU (Click-iT; Thermo Scientific) for 30 mins, collected by centrifugation and washed with 1xPBS. Then, cells were fixed with 3.7% paraformaldehyde at room temperature for 15 min, collected by centrifugation and washed with 1xPBS. Fixed cells were adhered to poly-L-lysine coated slides followed by permeabilization with 0.5% TritonX-100 at room temperature for 20 mins. Cells were washed with 1xPBS supplemented with 3% BSA and subjected to Click-iT reaction, according to the manufacturer’s instructions. After completion of the Click-iT reaction, cells were blocked with 1xPBS supplemented with 3% BSA for 1h at room temperature followed by washing with 1xPBS. Cells were incubated with anti-HA antibody diluted in 1xPBS supplemented with 1% BSA and 0.01% Tween20 for 2 h at room temperature. Cells were washed with 1xPBS and further incubated with anti-mouse secondary antibody conjugated with Alexa Fluor 594 diluted in 1xPBS supplemented with 1% BSA and 0.01% Tween20 for 1 hr at room temperature. After washing cells with 1xPBS, DNA was stained with DAPI. Images were acquired with a Zeiss Elyra Super-resolution microscope and further processed with ImageJ software. Detection of R-loops using microscopy 2x10 7 exponentially growing cells were collected by centrifugation and resuspended in 1.5 ml of 1xPBS supplemented 5 mM EDTA. Next, 3.5 mL methanol was added slowly and the cell suspension was incubated for 2 h at 4 °C under constant agitation. Cells were collected by centrifugation and washed with 1xPBS supplemented with 5 mM EDTA. Next, cells were resuspended in 1xPBS supplemented with 0.5% TritonX-100 and incubated on ice for 10 min. After centrifugation, cells were resuspended in staining buffer (1xPBS supplemented with 0.01% Tween20 and 0.1% BSA) and split into two aliquots. To one of the aliquots, 5 mM MgCl 2 and 2 U of recombinant Escherichia coli RNase HI (NEB) were added. Both aliquots were incubated at 37 °C for 1 h under constant agitation. Cells were collected by centrifugation and incubated with S9.6 antibody diluted in staining buffer at 37 °C for 1 h under constant agitation. Cells were washed with staining buffer, resuspended in 1xPBS and adhered to poly-L-lysine coated slides at room temperature for 30 min. DNA was stained with DAPI and images acquired with Zeiss Elyra Super-resolution microscope or Leica DMi8 microscope. Further image processing was performed with ImageJ software. DNA-RNA hybrid immunoprecipitation and sequencing (DRIP-seq) 2x10 9 exponentially growing cells were collected and washed with 1xPBS. Cells were resuspended in lysis buffer (10 mM TrisHCl pH8.0; 100 mM NaCl; 25 mM EDTA) at the concentration of 1x10 8 cells.mL -1 . The cell suspension was supplement with 0.1 mg.mL -1 proteinase K and 0.5% SDS and incubated overnight at 37 °C. Nucleic acids were extracted with equal volume of ultrapure phenol:chloroform:isoamylic alcohol (25:24:1) (ThermoFisher) followed by extraction with chloroform alone. Then, nucleic acids were precipitated by adding 0.1 V of 3 M sodium acetate pH5.2 and 2 V of ice-cold absolute ethanol, followed by centrifugation at 16,000 g under refrigeration for 10 min. The pellet was resuspended in 1.5 mL ice-cold 75% ethanol and centrifuged for 10 min under refrigeration. Next, the pellet was dissolved in 0.3 mL of ultrapure TE pH 8.0 (ThermoFisher) and incubated at 4 °C for 12 h. The final nucleic acid suspension was carefully homogenized and divided into two aliquots. One aliquot was left untreated, and the other was treated with 20 U of recombinant Escherichia coli RNase HI (Invitrogen) overnight at 37 °C. Nucleic acids were extracted once with an equal volume of ultrapure phenol:chloroform:isoamylic alcohol (25:24:1) and precipitated with 0.1 V 3 M sodium acetate pH5.2 and 2 V of ice-cold 100% ethanol. Pellets were resuspended and subjected to digestion at 37 °C for 16 h with one of the following digestion cocktails: Cocktail 1 ( Hind III, Eco RI, Bsr GI, Xba I and Ssp I) was used for DRIP-seq data shown in main figures, and Cocktail 2 ( Hind III, Eco RI, Bsr GI, Xba I and Ssp I Bam H1, Nco I, Apa LI and Pvu II) was used for DRIP-seq data shown in supplementary figures. The digestion cocktail for RNase HI-negative samples was further supplemented with 120 U RNaseOUT (Invitrogen), while the digestion cocktail for RNase HI-positive samples were supplemented with 10 U recombinant Escherichia coli RNase HI (Invitrogen). After digestion, nucleic acids were extracted with an equal volume of ultrapure phenol:chloroform:isoamylic alcohol (25:24:1), precipitated with 0.1 V 3 M sodium acetate pH5.2 and 2 V of ice-cold 100% ethanol. Pellets were resuspended in 0.1 mL of ultrapure TE pH 8.0 and 5% of the final solution was saved as input. DNA concentration was determined using Qubit (Invitrogen) and 40 ug DNA was diluted in 0.9 mL of IP buffer (10 mM Sodium Phosphate pH 6.8; 140 mM NaCl; 0.5% Triton X-100). This solution was mixed with 0.4 mL of Dynabeads M-280 Sheep Anti-Mouse IgG (Invitrogen) previously conjugated with 25 ug of S9.6 antibody (Millipore) and incubated overnight at 4 °C under constant agitation. Beads were collected using a magnetic rack and washed four times with IP buffer for 10 minutes each at 4 °C under constant agitation. Then, beads were resuspended in 0.2 mL of 1xPBS supplemented with Proteinase K from the DNeasy Blood and Tissue Kit (QIAGEN) and incubated at 37 °C in a thermomixer for 4 h. Samples were centrifuged and supernatant was collected, then DNA was extracted following DNeasy Blood and Tissue Kit standard protocols. Library preparation and sequencing is described below. Enrichment of immunoprecipitated material over input is expressed as ratios and was determined using bamCompare (DeepTools) over a 60 bp rolling window. For this, only reads with mapping quality >10 were considered. Libraries sizes were normalized using the SES method, pair-ended extension was employed, PCR duplicates were ignored and regions were centred with respect to the fragment length. All snapshot representations were performed using Gviz 164 . In the DRIP-seq analysis, peaks were identified based on regions exhibiting an enrichment greater than 2.5-fold in the DRIP material compared to the input control, as well as a more than 2.5-fold enrichment in samples untreated with RNase H compared to those treated with RNase H. Regions smaller than 60 bp were excluded from the analysis, and adjacent regions within 60 bp of each other were merged. Chromatin immunoprecipitation (ChIP-seq) Exponentially growing RNase H1-HA Flox cells were collected by centrifugation, resuspended in 1xPBS supplemented with 1% paraformaldehyde and incubated at room temperature for 15 min under constant agitation. Cells were collected and resuspended in lysis buffer (100 mM Tris pH 8.8; 200 mM NaCl; 1% NP40; 10% glycerol; 10 mM EDTA; 5 mM PMSF; 5 mM 1,10-phenantroline; 2x EDTA-free Pierce™ Protease Inhibitor) and subjected to 10 rounds of sonication of 20 sec each. Lysates were clarified by centrifugation and 5% was saved as input. The remainder of the lysate was incubated with 0.1 mL of Dynabeads M-280 Sheep Anti-Mouse IgG (Invitrogen) previously conjugated with 10 ug of anti-HA antibody (Millipore) and incubated at 4 °C for 16 h under constant agitation. Beads were collected using a magnetic rack and washed three times with lysis buffer. Next, beads were resuspended in elution buffer (50 mM Tris pH 7.6; 1% SDS) and incubated at 55 °C for 10 min under constant agitation. Crosslink reversal of eluted and input materials was performed by incubating samples at 65 °C for 12 h. DNA was further purified using DNeasy Blood and Tissue Kit (QIAGEN) following the manufacturer’s instructions. Library preparation and sequencing is described below. Replication timing profiling using Marker Frequency Analysis coupled with Illumina sequencing (MFA-seq) Genomic DNA was extracted from exponentially growing and stationary cells using DNeasy Blood & Tissue Kit (QIAGEN). After library preparation and Illumina sequencing (see details below) BamCompare (DeepTools) was used to determine reads abundance from exponentially growing cells relative to the reads from stationary culture. Ratios was first calculated in 1 kb consecutive windows using the reads counts method for normalisation. Raw ratios were further transformed into Z scores relative to the whole genome ratios average as calculated in 15 kb sliding windows. MFAseq snapshots were represented in a graphical form using Gviz 164 . Library preparation, sequencing and analysis All libraries were prepared using using QIAseq FX DNA Library Kit (QIAGEN) and were sequenced as 75 nucleotide paired-end reads. Sequencing was performed at Glasgow Polyomics ( www.polyomics.gla.ac.uk/index.html ) using a NextSeq™ 500 Illumina platform. The Galaxy web platform ( usegalaxy.org ) 165 was used for most of the downstream data processing. For quality control and removal of adaptors, FastQC ( http://www.bioinformatics.babraham.ac.uk/projects/fastqc/ ) and trimomatic 166 were used, respectively. Trimmed reads were mapped to the reference genome ( Leishmania major Friedlin v39, available at Tritrypdb - http://tritrypdb.org/tritrypdb/ ) using BWA-mem 167 . Cell cycle synchronization and DNA content analysis using fluorescent activated cell sorting (FACS) Exponentially growing cells were incubated with 5 mM hydroxyurea for 8 h. Then, cells were washed and re-seeded into hydroxyurea-free medium and collected by centrifugation every three hours. Cells were washed in 1×PBS supplemented with 5 mM EDTA, collected by centrifugation, resuspended in methanol:1xPBS (7:3) and then incubated at 4 °C for at least 2 h. Fixed cells were collected by centrifugation, washed with 1×PBS supplemented with 5 mM EDTA and incubated for 30 min at room temperature with 10 μg.mL -1 Propidium Iodide and 10 μg.mL -1 RNase A diluted in 1×PBS containing 5 mM EDTA. Cells were then passed through a 35 μm nylon mesh and examined with FACS Celesta (BD Biosciences). Further data analysis and data processing was performed with FlowJo software. SNPs, InDels and CNV analysis SNPs and InDels relative to the reference genome were detected in P4, P7 and in KO cells using freeBayes 168 . Only those SNPs and InDels in regions with read depth of at least 5, with at least 2 supporting reads, and a map quality of 30 were considered. To better capture the genomic variability in the time frame of the experiments, variants present simultaneously in P4 and P7 or P4 and KO cells were excluded from the analysis using VCF-VFC intersect function from VCFtools package 169 . The SNP density function from VCFtools was used to calculate SNPs and InDels density in consecutive 1 Kb windows. Genome-wide CNVs were calculated with bamCompare (DeepTools), using the RPKM normalisation method. Pair-ended extension was employed, PCR duplicates were ignored, and regions were centered with respect to the fragment length. Fold-change was expressed as log2. Data availability Sequences used in this study will be deposited in the European Nucleotide Archive. Competing interests The authors declare no competing interests. Acknowledgements We thank all current and previous members of the McCulloch lab for input. This work was supported by the Wellcome Trust [224501/Z/21/Z], the BBSRC [BB/N016165/1, BB/R017166/1, BB/W001101/1], the MRC [MR/S019472/1], and the European Union’s Horizon 2020 research and innovation programme under the Marie Sklodowska-Curie grant agreement No 750259 [Individual Fellowship, RECREPEMLE]. The Wellcome Centre for Integrative Parasitology was supported by core funding from the Wellcome Trust [104111]. References 1. ↵ Hu , Y. & Stillman , B . Origins of DNA replication in eukaryotes . Molecular cell 83 , 352 – 372 ( 2023 ). doi: 10.1016/j.molcel.2022.12.024 OpenUrl CrossRef 2. ↵ Costa , A. & Diffley , J. F. X . The Initiation of Eukaryotic DNA Replication . Annual review of biochemistry 91 , 107 – 131 ( 2022 ). doi: 10.1146/annurev-biochem-072321-110228 OpenUrl CrossRef 3. ↵ Lee , C. S. K. , Weibeta , M. & Hamperl , S . Where and when to start: Regulating DNA replication origin activity in eukaryotic genomes . Nucleus 14 , 2229642 ( 2023 ). doi: 10.1080/19491034.2023.2229642 OpenUrl CrossRef 4. Chen , N. & Buonomo , S. C. B . Three-dimensional nuclear organisation and the DNA replication timing program . Current opinion in structural biology 83 , 102704 ( 2023 ). doi: 10.1016/j.sbi.2023.102704 OpenUrl CrossRef 5. ↵ Vouzas , A. E. & Gilbert , D. M . Mammalian DNA Replication Timing . Cold Spring Harbor perspectives in biology 13 ( 2021 ). doi: 10.1101/cshperspect.a040162 OpenUrl Abstract / FREE Full Text 6. ↵ Nieduszynski , C. A. , Knox , Y. & Donaldson , A. D . Genome-wide identification of replication origins in yeast by comparative genomics . Genes & development 20 , 1874 – 1879 ( 2006 ). doi: 10.1101/gad.385306 OpenUrl Abstract / FREE Full Text 7. ↵ Muller , C. A. & Nieduszynski , C. A . Conservation of replication timing reveals global and local regulation of replication origin activity . Genome research 22 , 1953 – 1962 ( 2012 ). doi: 10.1101/gr.139477.112 OpenUrl Abstract / FREE Full Text 8. ↵ Hayashi , M. T. , Takahashi , T. S. , Nakagawa , T. , Nakayama , J. & Masukata , H . The heterochromatin protein Swi6/HP1 activates replication origins at the pericentromeric region and silent mating-type locus . Nat.Cell Biol . 11 , 357 – 362 ( 2009 ). OpenUrl CrossRef PubMed Web of Science 9. ↵ Natsume , T. et al. Kinetochores coordinate pericentromeric cohesion and early DNA replication by Cdc7-Dbf4 kinase recruitment . Molecular cell 50 , 661 – 674 ( 2013 ). doi: 10.1016/j.molcel.2013.05.011 OpenUrl CrossRef PubMed Web of Science 10. ↵ Hayano , M. et al. Rif1 is a global regulator of timing of replication origin firing in fission yeast . Genes & development 26 , 137 – 150 ( 2012 ). doi: 10.1101/gad.178491.111 OpenUrl Abstract / FREE Full Text 11. ↵ Kaykov , A. & Nurse , P . The spatial and temporal organization of origin firing during the S-phase of fission yeast . Genome research 25 , 391 – 401 ( 2015 ). doi: 10.1101/gr.180372.114 OpenUrl Abstract / FREE Full Text 12. ↵ Agier , N. , Romano , O. M. , Touzain , F. , Cosentino Lagomarsino , M. & Fischer , G . The spatiotemporal program of replication in the genome of Lachancea kluyveri . Genome biology and evolution 5 , 370 – 388 ( 2013 ). doi: 10.1093/gbe/evt014 OpenUrl CrossRef PubMed 13. ↵ Kelly , T. & Callegari , A. J . Dynamics of DNA replication in a eukaryotic cell . Proceedings of the National Academy of Sciences of the United States of America 116 , 4973 – 4982 ( 2019 ). doi: 10.1073/pnas.1818680116 OpenUrl Abstract / FREE Full Text 14. ↵ Hoggard , T. , Shor , E. , Muller , C. A. , Nieduszynski , C. A. & Fox , C. A . A Link between ORC-origin binding mechanisms and origin activation time revealed in budding yeast . PLoS genetics 9 , e1003798 ( 2013 ). doi: 10.1371/journal.pgen.1003798 OpenUrl CrossRef PubMed 15. ↵ Muller , C. A. & Nieduszynski , C. A . DNA replication timing influences gene expression level . The Journal of cell biology 216 , 1907 – 1914 ( 2017 ). doi: 10.1083/jcb.201701061 OpenUrl Abstract / FREE Full Text 16. ↵ Hulke , M. L. , Massey , D. J. & Koren , A . Genomic methods for measuring DNA replication dynamics . Chromosome research : an international journal on the molecular, supramolecular and evolutionary aspects of chromosome biology 28 , 49 – 67 ( 2020 ). doi: 10.1007/s10577-019-09624-y OpenUrl CrossRef 17. ↵ Hennion , M. et al. FORK-seq: replication landscape of the Saccharomyces cerevisiae genome by nanopore sequencing . Genome biology 21 , 125 ( 2020 ). doi: 10.1186/s13059-020-02013-3 OpenUrl CrossRef 18. Muller , C. A. et al. Capturing the dynamics of genome replication on individual ultra-long nanopore sequence reads . Nature methods 16 , 429 – 436 ( 2019 ). doi: 10.1038/s41592-019-0394-y OpenUrl CrossRef 19. DePamphilis , M. L . Origins of DNA replication in metazoan chromosomes . The Journal of biological chemistry 268 , 1 – 4 ( 1993 ). OpenUrl Abstract / FREE Full Text 20. Wang , W. et al. Genome-wide mapping of human DNA replication by optical replication mapping supports a stochastic model of eukaryotic replication . Molecular cell 81 , 2975 – 2988 e2976 ( 2021 ). doi: 10.1016/j.molcel.2021.05.024 OpenUrl CrossRef 21. ↵ Tian , M. et al. Integrative analysis of DNA replication origins and ORC/MCM binding sites in human cells reveals a lack of overlap . bioRxiv ( 2023 ). doi: 10.1101/2023.07.25.550556 OpenUrl Abstract / FREE Full Text 22. ↵ Guilbaud , G. et al. Determination of human DNA replication origin position and efficiency reveals principles of initiation zone organisation . Nucleic Acids Res 50 , 7436 – 7450 ( 2022 ). doi: 10.1093/nar/gkac555 OpenUrl CrossRef 23. ↵ Rivera-Mulia , J. C. et al. Dynamic changes in replication timing and gene expression during lineage specification of human pluripotent stem cells . Genome research 25 , 1091 – 1103 ( 2015 ). doi: 10.1101/gr.187989.114 OpenUrl Abstract / FREE Full Text 24. ↵ Hiratani , I. et al. Global reorganization of replication domains during embryonic stem cell differentiation . PLoS biology 6 , e245 ( 2008 ). doi: 10.1371/journal.pbio.0060245 OpenUrl CrossRef PubMed 25. ↵ Pope , B. D. et al. Topologically associating domains are stable units of replication-timing regulation . Nature 515 , 402 – 405 ( 2014 ). doi: 10.1038/nature13986 OpenUrl CrossRef PubMed Web of Science 26. ↵ Moindrot , B. et al. 3D chromatin conformation correlates with replication timing and is conserved in resting cells . Nucleic Acids Res 40 , 9470 – 9481 ( 2012 ). doi: 10.1093/nar/gks736 OpenUrl CrossRef PubMed Web of Science 27. ↵ Wang , Y. et al. SPIN reveals genome-wide landscape of nuclear compartmentalization . Genome biology 22 , 36 ( 2021 ). doi: 10.1186/s13059-020-02253-3 OpenUrl CrossRef 28. ↵ Peric-Hupkes , D. et al. Molecular maps of the reorganization of genome-nuclear lamina interactions during differentiation . Molecular cell 38 , 603 – 613 ( 2010 ). doi: 10.1016/j.molcel.2010.03.016 OpenUrl CrossRef PubMed Web of Science 29. ↵ Brueckner , L. et al. Local rewiring of genome-nuclear lamina interactions by transcription . The EMBO journal 39 , e103159 ( 2020 ). doi: 10.15252/embj.2019103159 OpenUrl CrossRef 30. ↵ Ragoczy , T. , Telling , A. , Scalzo , D. , Kooperberg , C. & Groudine , M . Functional redundancy in the nuclear compartmentalization of the late-replicating genome . Nucleus 5 , 626 – 635 ( 2014 ). doi: 10.4161/19491034.2014.990863 OpenUrl CrossRef PubMed 31. ↵ Vertii , A. et al. Two contrasting classes of nucleolus-associated domains in mouse fibroblast heterochromatin . Genome research 29 , 1235 – 1249 ( 2019 ). doi: 10.1101/gr.247072.118 OpenUrl Abstract / FREE Full Text 32. ↵ Chen , Y. et al. Mapping 3D genome organization relative to nuclear compartments using TSA-Seq as a cytological ruler . The Journal of cell biology 217 , 4025 – 4048 ( 2018 ). doi: 10.1083/jcb.201807108 OpenUrl Abstract / FREE Full Text 33. ↵ Bracci , A. N. et al. The evolution of the human DNA replication timing program . Proceedings of the National Academy of Sciences of the United States of America 120 , e2213896120 ( 2023 ). doi: 10.1073/pnas.2213896120 OpenUrl CrossRef 34. Briu , L. M. , Maric , C. & Cadoret , J. C. Replication Stress, Genomic Instability, and Replication Timing: A Complex Relationship . International journal of molecular sciences 22 ( 2021 ). doi: 10.3390/ijms22094764 OpenUrl CrossRef 35. ↵ Sima , J. & Gilbert , D. M . Complex correlations: replication timing and mutational landscapes during cancer and genome evolution . Curr Opin Genet Dev 25 , 93 – 100 ( 2014 ). doi: 10.1016/j.gde.2013.11.022 OpenUrl CrossRef PubMed 36. ↵ Stamatoyannopoulos , J. A. et al. Human mutation rate associated with DNA replication timing . Nature genetics 41 , 393 – 395 ( 2009 ). doi: 10.1038/ng.363 OpenUrl CrossRef PubMed Web of Science 37. ↵ Chen , C. L. et al. Impact of replication timing on non-CpG and CpG substitution rates in mammalian genomes . Genome research 20 , 447 – 457 ( 2010 ). doi: 10.1101/gr.098947.109 OpenUrl Abstract / FREE Full Text 38. ↵ Lang , G. I. & Murray , A. W . Mutation rates across budding yeast chromosome VI are correlated with replication timing . Genome biology and evolution 3 , 799 – 811 ( 2011 ). doi: 10.1093/gbe/evr054 OpenUrl CrossRef PubMed 39. ↵ Agier , N. & Fischer , G . The mutational profile of the yeast genome is shaped by replication . Molecular biology and evolution 29 , 905 – 913 ( 2012 ). doi: 10.1093/molbev/msr280 OpenUrl CrossRef PubMed Web of Science 40. ↵ De , S. & Michor , F . DNA replication timing and long-range DNA interactions predict mutational landscapes of cancer genomes . Nat Biotechnol 29 , 1103 – 1108 ( 2011 ). doi: 10.1038/nbt.2030 OpenUrl CrossRef PubMed 41. ↵ Yaffe , E. et al. Comparative analysis of DNA replication timing reveals conserved large-scale chromosomal architecture . PLoS genetics 6 , e1001011 ( 2010 ). doi: 10.1371/journal.pgen.1001011 OpenUrl CrossRef PubMed 42. ↵ Janoueix-Lerosey , I. et al. Preferential occurrence of chromosome breakpoints within early replicating regions in neuroblastoma . Cell Cycle 4 , 1842 – 1846 ( 2005 ). doi: 10.4161/cc.4.12.2257 OpenUrl CrossRef PubMed Web of Science 43. ↵ Peycheva , M. et al. DNA replication timing directly regulates the frequency of oncogenic chromosomal translocations . Science 377 , eabj5502 ( 2022 ). doi: 10.1126/science.abj5502 OpenUrl CrossRef 44. ↵ Cohen , S. M. , Furey , T. S. , Doggett , N. A. & Kaufman , D. G . Genome-wide sequence and functional analysis of early replicating DNA in normal human fibroblasts . BMC Genomics 7 , 301 ( 2006 ). doi: 10.1186/1471-2164-7-301 OpenUrl CrossRef PubMed 45. ↵ Sultana , T. et al. The Landscape of L1 Retrotransposons in the Human Genome Is Shaped by Pre-insertion Sequence Biases and Post-insertion Selection . Molecular cell 74 , 555 – 570 e557 ( 2019 ). doi: 10.1016/j.molcel.2019.02.036 OpenUrl CrossRef 46. ↵ Sima , J. et al. Identifying cis Elements for Spatiotemporal Control of Mammalian DNA Replication . Cell 176 , 816 – 830 e818 ( 2019 ). doi: 10.1016/j.cell.2018.11.036 OpenUrl CrossRef PubMed 47. ↵ Richards , L. , Das , S. & Nordman , J. T . Rif1-Dependent Control of Replication Timing . Genes (Basel ) 13 ( 2022 ). doi: 10.3390/genes13030550 OpenUrl CrossRef 48. ↵ Yamazaki , S. et al. Rif1 regulates the replication timing domains on the human genome . The EMBO journal 31 , 3667 – 3677 ( 2012 ). doi: 10.1038/emboj.2012.180 OpenUrl Abstract / FREE Full Text 49. ↵ Hiraga , S. et al. Rif1 controls DNA replication by directing Protein Phosphatase 1 to reverse Cdc7-mediated phosphorylation of the MCM complex . Genes & development 28 , 372 – 383 ( 2014 ). doi: 10.1101/gad.231258.113 OpenUrl Abstract / FREE Full Text 50. ↵ Hiraga , S. I. et al. Human RIF1 and protein phosphatase 1 stimulate DNA replication origin licensing but suppress origin activation . EMBO reports 18 , 403 – 419 ( 2017 ). doi: 10.15252/embr.201641983 OpenUrl Abstract / FREE Full Text 51. ↵ Foti , R. et al. Nuclear Architecture Organized by Rif1 Underpins the Replication-Timing Program . Molecular cell 61 , 260 – 273 ( 2016 ). doi: 10.1016/j.molcel.2015.12.001 OpenUrl CrossRef PubMed 52. ↵ Gnan , S. et al. Nuclear organisation and replication timing are coupled through RIF1-PP1 interaction . Nature communications 12 , 2910 ( 2021 ). doi: 10.1038/s41467-021-22899-2 OpenUrl CrossRef 53. ↵ Knott , S. R. et al. Forkhead transcription factors establish origin timing and long-range clustering in S. cerevisiae . Cell 148 , 99 – 111 ( 2012 ). doi: 10.1016/j.cell.2011.12.012 OpenUrl CrossRef PubMed Web of Science 54. ↵ Reinapae , A. et al. Interactions between Fkh1 monomers stabilize its binding to DNA replication origins . The Journal of biological chemistry 299 , 105026 ( 2023 ). doi: 10.1016/j.jbc.2023.105026 OpenUrl CrossRef 55. ↵ Heskett , M. B. et al. Epigenetic control of chromosome-associated lncRNA genes essential for replication and stability . Nature communications 13 , 6301 ( 2022 ). doi: 10.1038/s41467-022-34099-7 OpenUrl CrossRef 56. ↵ Stoffregen , E. P. , Donley , N. , Stauffer , D. , Smith , L. & Thayer , M. J . An autosomal locus that controls chromosome-wide replication timing and mono-allelic expression . Hum Mol Genet 20 , 2366 – 2378 ( 2011 ). doi: 10.1093/hmg/ddr138 OpenUrl CrossRef PubMed Web of Science 57. ↵ Donley , N. , Stoffregen , E. P. , Smith , L. , Montagna , C. & Thayer , M. J . Asynchronous replication, mono-allelic expression, and long range Cis-effects of ASAR6 . PLoS genetics 9 , e1003423 ( 2013 ). doi: 10.1371/journal.pgen.1003423 OpenUrl CrossRef PubMed 58. ↵ Heskett , M. B. , Smith , L. G. , Spellman , P. & Thayer , M. J . Reciprocal monoallelic expression of ASAR lncRNA genes controls replication timing of human chromosome 6 . Rna 26 , 724 – 738 ( 2020 ). doi: 10.1261/rna.073114.119 OpenUrl Abstract / FREE Full Text 59. ↵ Donley , N. , Smith , L. & Thayer , M. J . ASAR15, A cis-acting locus that controls chromosome-wide replication timing and stability of human chromosome 15 . PLoS genetics 11 , e1004923 ( 2015 ). doi: 10.1371/journal.pgen.1004923 OpenUrl CrossRef PubMed 60. ↵ Platt , E. J. , Smith , L. & Thayer , M. J . L1 retrotransposon antisense RNA within ASAR lncRNAs controls chromosome-wide replication timing . The Journal of cell biology 217 , 541 – 553 ( 2018 ). doi: 10.1083/jcb.201707082 OpenUrl Abstract / FREE Full Text 61. ↵ Adl , S. M. et al. Revisions to the Classification, Nomenclature, and Diversity of Eukaryotes . The Journal of eukaryotic microbiology 66 , 4 – 119 ( 2019 ). doi: 10.1111/jeu.12691 OpenUrl CrossRef PubMed 62. ↵ Burki , F. , Roger , A. J. , Brown , M. W. & Simpson , A. G. B . The New Tree of Eukaryotes . Trends Ecol Evol 35 , 43 – 55 ( 2020 ). doi: 10.1016/j.tree.2019.08.008 OpenUrl CrossRef PubMed 63. ↵ da Silva , M. S. et al. Clues on the dynamics of DNA replication in Giardia lamblia . Journal of cell science 136 ( 2023 ). doi: 10.1242/jcs.260828 OpenUrl CrossRef 64. ↵ Zhang , L. et al. Transcriptome analysis of the binucleate ciliate Tetrahymena thermophila with asynchronous nuclear cell cycles . Molecular biology of the cell 34 , rs1 ( 2023 ). doi: 10.1091/mbc.E22-08-0326 OpenUrl CrossRef 65. ↵ Castellano , C. M. et al. The genetic landscape of origins of replication in P. falciparum . Nucleic Acids Res ( 2023 ). doi: 10.1093/nar/gkad1103 OpenUrl CrossRef 66. ↵ Totanes , F. I. G. et al. A genome-wide map of DNA replication at single-molecule resolution in the malaria parasite Plasmodium falciparum . Nucleic Acids Res 51 , 2709 – 2724 ( 2023 ). doi: 10.1093/nar/gkad093 OpenUrl CrossRef 67. ↵ Tiengwe , C. et al. Genome-wide analysis reveals extensive functional interaction between DNA replication initiation and transcription in the genome of Trypanosoma brucei . Cell reports 2 , 185 – 197 ( 2012 ). doi: 10.1016/j.celrep.2012.06.007 OpenUrl CrossRef PubMed Web of Science 68. ↵ Clayton , C . Regulation of gene expression in trypanosomatids: living with polycistronic transcription . Open biology 9 , 190072 ( 2019 ). doi: 10.1098/rsob.190072 OpenUrl CrossRef PubMed 69. ↵ El-Sayed , N. M. et al. Comparative genomics of trypanosomatid parasitic protozoa . Science 309 , 404 – 409 ( 2005 ). doi: 10.1126/science.1112181 OpenUrl Abstract / FREE Full Text 70. ↵ Damasceno , J. D. , Marques , C. A. , Black , J. , Briggs , E. & McCulloch , R. Read, Write, Adapt: Challenges and Opportunities during Kinetoplastid Genome Replication . Trends in genetics : TIG 37 , 21 – 34 ( 2021 ). doi: 10.1016/j.tig.2020.09.002 OpenUrl CrossRef 71. ↵ Batrakou , D. G. , Muller , C. A. , Wilson , R. H. C. & Nieduszynski , C. A . DNA copy-number measurement of genome replication dynamics by high-throughput sequencing: the sort-seq, sync-seq and MFA-seq family . Nature protocols 15 , 1255 – 1284 ( 2020 ). doi: 10.1038/s41596-019-0287-7 OpenUrl CrossRef PubMed 72. ↵ Devlin , R. et al. Mapping replication dynamics in Trypanosoma brucei reveals a link with telomere transcription and antigenic variation . eLife 5 , e12765 ( 2016 ). doi: 10.7554/eLife.12765 OpenUrl CrossRef 73. ↵ Muller , L. S. M. et al. Genome organization and DNA accessibility control antigenic variation in trypanosomes . Nature 563 , 121 – 125 ( 2018 ). doi: 10.1038/s41586-018-0619-8 OpenUrl CrossRef PubMed 74. ↵ Ubeda , J. M. et al. Modulation of gene expression in drug resistant Leishmania is associated with gene amplification, gene deletion and chromosome aneuploidy . Genome biology 9 , R115 ( 2008 ). doi: 10.1186/gb-2008-9-7-r115 OpenUrl CrossRef PubMed 75. ↵ Rogers , M. B. et al. Chromosome and gene copy number variation allow major structural change between species and strains of Leishmania . Genome research 21 , 2129 – 2142 ( 2011 ). doi: 10.1101/gr.122945.111 OpenUrl Abstract / FREE Full Text 76. ↵ Prieto Barja , P., et al. Haplotype selection as an adaptive mechanism in the protozoan pathogen Leishmania donovani . Nat Ecol Evol 1 , 1961 – 1969 ( 2017 ). doi: 10.1038/s41559-017-0361-x OpenUrl CrossRef PubMed 77. Sterkers , Y. , Lachaud , L. , Crobu , L. , Bastien , P. & Pages , M . FISH analysis reveals aneuploidy and continual generation of chromosomal mosaicism in Leishmania major . Cellular microbiology 13 , 274 – 283 ( 2011 ). doi: 10.1111/j.1462-5822.2010.01534.x OpenUrl CrossRef PubMed 78. ↵ Bussotti , G. et al. Leishmania Genome Dynamics during Environmental Adaptation Reveal Strain-Specific Differences in Gene Copy Number Variation, Karyotype Instability, and Telomeric Amplification . mBio 9 ( 2018 ). doi: 10.1128/mBio.01399-18 OpenUrl Abstract / FREE Full Text 79. ↵ Almeida , L. V. et al. Chromosomal copy number variation analysis by next generation sequencing confirms ploidy stability in Trypanosoma brucei subspecies . Microb Genom 4 ( 2018 ). doi: 10.1099/mgen.0.000223 OpenUrl CrossRef PubMed 80. ↵ Alsford , N. S. et al. The identification of circular extrachromosomal DNA in the nuclear genome of Trypanosoma brucei . Mol.Microbiol . 47 , 277 – 289 ( 2003 ). OpenUrl CrossRef PubMed Web of Science 81. ↵ Marques , C. A. , Dickens , N. J. , Paape , D. , Campbell , S. J. & McCulloch , R . Genome-wide mapping reveals single-origin chromosome replication in Leishmania, a eukaryotic microbe . Genome biology 16 , 230 ( 2015 ). doi: 10.1186/s13059-015-0788-9 OpenUrl CrossRef 82. ↵ Damasceno , J. D. et al. Genome duplication in Leishmania major relies on persistent subtelomeric DNA replication . eLife 9 ( 2020 ). doi: 10.7554/eLife.58030 OpenUrl CrossRef 83. ↵ Lombrana , R. et al. Transcriptionally Driven DNA Replication Program of the Human Parasite Leishmania major . Cell reports 16 , 1774 – 1786 ( 2016 ). doi: 10.1016/j.celrep.2016.07.007 OpenUrl CrossRef 84. ↵ Brickner , J. R. , Garzon , J. L. & Cimprich , K. A . Walking a tightrope: The complex balancing act of R-loops in genome stability . Molecular cell 82 , 2267 – 2297 ( 2022 ). doi: 10.1016/j.molcel.2022.04.014 OpenUrl CrossRef 85. ↵ Petermann , E. , Lan , L. & Zou , L . Sources, resolution and physiological relevance of R-loops and RNA-DNA hybrids . Nature reviews. Molecular cell biology 23 , 521 – 540 ( 2022 ). doi: 10.1038/s41580-022-00474-x OpenUrl CrossRef 86. ↵ Skourti-Stathaki , K. & Proudfoot , N. J . A double-edged sword: R loops as threats to genome integrity and powerful regulators of gene expression . Genes & development 28 , 1384 – 1396 ( 2014 ). doi: 10.1101/gad.242990.114 OpenUrl Abstract / FREE Full Text 87. ↵ Yu , K. , Chedin , F. , Hsieh , C. L. , Wilson , T. E. & Lieber , M. R . R-loops at immunoglobulin class switch regions in the chromosomes of stimulated B cells . Nat Immunol 4 , 442 – 451 ( 2003 ). doi: 10.1038/ni919 OpenUrl CrossRef PubMed Web of Science 88. ↵ Ohle , C. et al. Transient RNA-DNA Hybrids Are Required for Efficient Double-Strand Break Repair . Cell 167 , 1001 – 1013 e1007 ( 2016 ). doi: 10.1016/j.cell.2016.10.001 OpenUrl CrossRef PubMed 89. Yasuhara , T. et al. Human Rad52 Promotes XPG-Mediated R-loop Processing to Initiate Transcription-Associated Homologous Recombination Repair . Cell 175 , 558 – 570 e511 ( 2018 ). doi: 10.1016/j.cell.2018.08.056 OpenUrl CrossRef PubMed 90. Lu , W. T. et al. Drosha drives the formation of DNA:RNA hybrids around DNA break sites to facilitate DNA repair . Nature communications 9 , 532 ( 2018 ). doi: 10.1038/s41467-018-02893-x OpenUrl CrossRef PubMed 91. ↵ Cohen , S. et al. Senataxin resolves RNA:DNA hybrids forming at DNA double-strand breaks to prevent translocations . Nat Commun 9 , 533 ( 2018 ). doi: 10.1038/s41467-018-02894-w OpenUrl CrossRef PubMed 92. ↵ Powell , W. T. et al. R-loop formation at Snord116 mediates topotecan inhibition of Ube3a-antisense and allele-specific chromatin decondensation . Proceedings of the National Academy of Sciences of the United States of America 110 , 13938 – 13943 ( 2013 ). doi: 10.1073/pnas.1305426110 OpenUrl Abstract / FREE Full Text 93. ↵ Chen , P. B. , Chen , H. V. , Acharya , D. , Rando , O. J. & Fazzio , T. G . R loops regulate promoter-proximal chromatin architecture and cellular differentiation . Nat Struct Mol Biol 22 , 999 – 1007 ( 2015 ). doi: 10.1038/nsmb.3122 OpenUrl CrossRef PubMed 94. ↵ Chen , L. et al. R-ChIP Using Inactive RNase H Reveals Dynamic Coupling of R-loops with Transcriptional Pausing at Gene Promoters . Molecular cell 68 , 745 – 757 e745 ( 2017 ). doi: 10.1016/j.molcel.2017.10.008 OpenUrl CrossRef PubMed 95. ↵ Skourti-Stathaki , K. , Proudfoot , N. J. & Gromak , N . Human senataxin resolves RNA/DNA hybrids formed at transcriptional pause sites to promote Xrn2-dependent termination . Molecular cell 42 , 794 – 805 ( 2011 ). doi: 10.1016/j.molcel.2011.04.026 OpenUrl CrossRef PubMed Web of Science 96. ↵ Skourti-Stathaki , K. , Kamieniarz-Gdula , K. & Proudfoot , N. J . R-loops induce repressive chromatin marks over mammalian gene terminators . Nature 516 , 436 – 439 ( 2014 ). doi: 10.1038/nature13787 OpenUrl CrossRef PubMed 97. ↵ Briggs , E. , Hamilton , G. , Crouch , K. , Lapsley , C. & McCulloch , R . Genome-wide mapping reveals conserved and diverged R-loop activities in the unusual genetic landscape of the African trypanosome genome . Nucleic Acids Res 46 , 11789 – 11805 ( 2018 ). doi: 10.1093/nar/gky928 OpenUrl CrossRef 98. ↵ Briggs , E. et al. Trypanosoma brucei ribonuclease H2A is an essential R-loop processing enzyme whose loss causes DNA damage during transcription initiation and antigenic variation . Nucleic Acids Res 47 , 9180 – 9197 ( 2019 ). doi: 10.1093/nar/gkz644 OpenUrl CrossRef 99. ↵ Nanavaty , V. , Sandhu , R. , Jehi , S. E. , Pandya , U. M. & Li , B . Trypanosoma brucei RAP1 maintains telomere and subtelomere integrity by suppressing TERRA and telomeric RNA:DNA hybrids . Nucleic Acids Res 45 , 5785 – 5796 ( 2017 ). doi: 10.1093/nar/gkx184 OpenUrl CrossRef 100. ↵ Briggs , E. , Crouch , K. , Lemgruber , L. , Lapsley , C. & McCulloch , R . Ribonuclease H1-targeted R-loops in surface antigen gene expression sites can direct trypanosome immune evasion . PLoS genetics 14 , e1007729 ( 2018 ). doi: 10.1371/journal.pgen.1007729 OpenUrl CrossRef 101. Girasol , M. J. et al. RAD51-mediated R-loop formation acts to repair transcription-associated DNA breaks driving antigenic variation in Trypanosoma brucei . Proceedings of the National Academy of Sciences of the United States of America 120 , e2309306120 ( 2023 ). doi: 10.1073/pnas.2309306120 OpenUrl CrossRef 102. ↵ Eisenhuth , N. , Vellmer , T. , Rauh , E. T. , Butter , F. & Janzen , C. J . A DOT1B/Ribonuclease H2 Protein Complex Is Involved in R-Loop Processing, Genomic Integrity, and Antigenic Variation in Trypanosoma brucei . mBio 12 , e0135221 ( 2021 ). doi: 10.1128/mBio.01352-21 OpenUrl CrossRef 103. ↵ Cristini , A. , Groh , M. , Kristiansen , M. S. & Gromak , N . RNA/DNA Hybrid Interactome Identifies DXH9 as a Molecular Player in Transcriptional Termination and R-Loop-Associated DNA Damage . Cell reports 23 , 1891 – 1905 ( 2018 ). doi: 10.1016/j.celrep.2018.04.025 OpenUrl CrossRef 104. ↵ Girasol , M. J. et al. Immunoprecipitation of RNA-DNA hybrid interacting proteins in Trypanosoma brucei reveals conserved and novel activities, including in the control of surface antigen expression needed for immune evasion by antigenic variation . Nucleic Acids Res 51 , 11123 – 11141 ( 2023 ). doi: 10.1093/nar/gkad836 OpenUrl CrossRef 105. ↵ Santos-Pereira , J. M. & Aguilera , A . R loops: new modulators of genome dynamics and function . Nature reviews. Genetics 16 , 583 – 597 ( 2015 ). doi: 10.1038/nrg3961 OpenUrl CrossRef PubMed 106. ↵ Hamperl , S. & Cimprich , K. A . The contribution of co-transcriptional RNA:DNA hybrid structures to DNA damage and genome instability . DNA Repair (Amst ) 19 , 84 – 94 ( 2014 ). doi: 10.1016/j.dnarep.2014.03.023 OpenUrl CrossRef PubMed 107. ↵ Cerritelli , S. M. & Crouch , R. J . Ribonuclease H: the enzymes in eukaryotes . The FEBS journal 276 , 1494 – 1505 ( 2009 ). doi: 10.1111/j.1742-4658.2009.06908.x OpenUrl CrossRef PubMed 108. ↵ Hu , Z. , Zhang , A. , Storz , G. , Gottesman , S. & Leppla , S. H . An antibody-based microarray assay for small RNA detection . Nucleic Acids Res 34 , e52 ( 2006 ). doi: 10.1093/nar/gkl142 OpenUrl CrossRef PubMed 109. ↵ Smolka , J. A. , Sanz , L. A. , Hartono , S. R. & Chedin , F . Recognition of RNA by the S9.6 antibody creates pervasive artifacts when imaging RNA:DNA hybrids . The Journal of cell biology 220 ( 2021 ). doi: 10.1083/jcb.202004079 OpenUrl CrossRef 110. ↵ Marsico , G. et al. Whole genome experimental maps of DNA G-quadruplexes in multiple species . Nucleic Acids Res 47 , 3862 – 3874 ( 2019 ). doi: 10.1093/nar/gkz179 OpenUrl CrossRef PubMed 111. ↵ Bailey , T. L. , Johnson , J. , Grant , C. E. & Noble , W. S. The MEME Suite . Nucleic Acids Res 43 , W39 – 49 ( 2015 ). doi: 10.1093/nar/gkv416 OpenUrl CrossRef PubMed 112. ↵ Curotto de Lafaille , M. A. , Laban , A. & Wirth , D. F. Gene expression in Leishmania: analysis of essential 5’ DNA sequences . Proceedings of the National Academy of Sciences of the United States of America 89 , 2703 – 2707 ( 1992 ). doi: 10.1073/pnas.89.7.2703 OpenUrl Abstract / FREE Full Text 113. Vassella , E. , Braun , R. & Roditi , I . Control of polyadenylation and alternative splicing of transcripts from adjacent genes in a procyclin expression site: a dual role for polypyrimidine tracts in trypanosomes? Nucleic Acids Res 22 , 1359 – 1364 ( 1994 ). doi: 10.1093/nar/22.8.1359 OpenUrl CrossRef PubMed Web of Science 114. ↵ Stern , M. Z. et al. Multiple roles for polypyrimidine tract binding (PTB) proteins in trypanosome RNA metabolism . Rna 15 , 648 – 665 ( 2009 ). doi: 10.1261/rna.1230209 OpenUrl Abstract / FREE Full Text 115. ↵ Ubeda , J. M. et al. Genome-wide stochastic adaptive DNA amplification at direct and inverted DNA repeats in the parasite Leishmania . PLoS biology 12 , e1001868 ( 2014 ). doi: 10.1371/journal.pbio.1001868 OpenUrl CrossRef PubMed 116. ↵ Bringaud , F. et al. Members of a large retroposon family are determinants of post-transcriptional gene expression in Leishmania . PLoS pathogens 3 , 1291 – 1307 ( 2007 ). doi: 10.1371/journal.ppat.0030136 OpenUrl CrossRef PubMed Web of Science 117. ↵ Boucher , N. et al. A common mechanism of stage-regulated gene expression in Leishmania mediated by a conserved 3’-untranslated region element . The Journal of biological chemistry 277 , 19511 – 19520 ( 2002 ). doi: 10.1074/jbc.M200500200 OpenUrl Abstract / FREE Full Text 118. ↵ Misra , S. et al. A type II ribonuclease H from Leishmania mitochondria: an enzyme essential for the growth of the parasite . Molecular and biochemical parasitology 143 , 135 – 145 ( 2005 ). doi: 10.1016/j.molbiopara.2005.05.009 OpenUrl CrossRef PubMed 119. ↵ Thomas , S. , Green , A. , Sturm , N. R. , Campbell , D. A. & Myler , P. J . Histone acetylations mark origins of polycistronic transcription in Leishmania major . BMC Genomics 10 , 152 ( 2009 ). 120. ↵ van Luenen , H. G. et al. Glucosylated hydroxymethyluracil, DNA base J, prevents transcriptional readthrough in Leishmania . Cell 150 , 909 – 921 ( 2012 ). OpenUrl CrossRef PubMed Web of Science 121. ↵ Hsiang , Y. H. , Hertzberg , R. , Hecht , S. & Liu , L. F . Camptothecin induces protein-linked DNA breaks via mammalian DNA topoisomerase I . The Journal of biological chemistry 260 , 14873 – 14878 ( 1985 ). OpenUrl Abstract / FREE Full Text 122. ↵ Goldberg , I. H. , Rabinowitz , M. & Reich , E . Basis of actinomycin action. I. DNA binding and inhibition of RNA-polymerase synthetic reactions by actinomycin . Proceedings of the National Academy of Sciences of the United States of America 48 , 2094 – 2101 ( 1962 ). doi: 10.1073/pnas.48.12.2094 OpenUrl FREE Full Text 123. ↵ Hassan , P. , Fergusson , D. , Grant , K. M. & Mottram , J. C . The CRK3 protein kinase is essential for cell cycle progression of Leishmania mexicana . Molecular and biochemical parasitology 113 , 189 – 198 ( 2001 ). doi: 10.1016/s0166-6851(01)00220-1 OpenUrl CrossRef PubMed 124. ↵ Saldivia , M. et al. Targeting the trypanosome kinetochore with CLK1 protein kinase inhibitors . Nat Microbiol 5 , 1207 – 1216 ( 2020 ). doi: 10.1038/s41564-020-0745-6 OpenUrl CrossRef 125. ↵ Damasceno , J. D. et al. Genome duplication in Leishmania major relies on DNA replication outside S phase . bioRxiv , 799429 ( 2019 ). doi: 10.1101/799429 OpenUrl Abstract / FREE Full Text 126. ↵ Amon , J. D. & Koshland , D . RNase H enables efficient repair of R-loop induced DNA damage . eLife 5 ( 2016 ). doi: 10.7554/eLife.20533 OpenUrl CrossRef PubMed 127. Wahba , L. , Amon , J. D. , Koshland , D. & Vuica-Ross , M . RNase H and multiple RNA biogenesis factors cooperate to prevent RNA:DNA hybrids from generating genome instability . Molecular cell 44 , 978 – 988 ( 2011 ). doi: 10.1016/j.molcel.2011.10.017 OpenUrl CrossRef PubMed Web of Science 128. ↵ Stuckey , R. , Garcia-Rodriguez , N. , Aguilera , A. & Wellinger , R. E . Role for RNA:DNA hybrids in origin-independent replication priming in a eukaryotic system . Proceedings of the National Academy of Sciences of the United States of America 112 , 5779 – 5784 ( 2015 ). doi: 10.1073/pnas.1501769112 OpenUrl Abstract / FREE Full Text 129. ↵ Negreira , G. H. et al. The adaptive roles of aneuploidy and polyclonality in Leishmania in response to environmental stress . EMBO reports 24 , e57413 ( 2023 ). doi: 10.15252/embr.202357413 OpenUrl CrossRef 130. ↵ Dumetz , F. et al. Modulation of Aneuploidy in Leishmania donovani during Adaptation to Different In Vitro and In Vivo Environments and Its Impact on Gene Expression . mBio 8 ( 2017 ). doi: 10.1128/mBio.00599-17 OpenUrl Abstract / FREE Full Text 131. ↵ Hamperl , S. , Bocek , M. J. , Saldivar , J. C. , Swigut , T. & Cimprich , K. A . Transcription-Replication Conflict Orientation Modulates R-Loop Levels and Activates Distinct DNA Damage Responses . Cell 170 , 774 – 786 e719 ( 2017 ). doi: 10.1016/j.cell.2017.07.043 OpenUrl CrossRef PubMed 132. ↵ O’Shea , J. P. et al. pLogo: a probabilistic approach to visualizing sequence motifs . Nat Methods 10 , 1211 – 1212 ( 2013 ). doi: 10.1038/nmeth.2646 OpenUrl CrossRef PubMed Web of Science 133. ↵ Damasceno , J. D. et al. Conditional knockout of RAD51-related genes in Leishmania major reveals a critical role for homologous recombination during genome replication . PLoS genetics 16 , e1008828 ( 2020 ). doi: 10.1371/journal.pgen.1008828 OpenUrl CrossRef 134. ↵ Hegazy , Y. A. , Fernando , C. M. & Tran , E. J . The Balancing Act of R-loop Biology: The Good, the Bad, and the Ugly . The Journal of biological chemistry ( 2019 ). doi: 10.1074/jbc.REV119.011353 OpenUrl Abstract / FREE Full Text 135. ↵ Castillo-Guzman , D. & Chedin , F . Defining R-loop classes and their contributions to genome instability . DNA Repair (Amst ) 106 , 103182 ( 2021 ). doi: 10.1016/j.dnarep.2021.103182 OpenUrl CrossRef PubMed 136. ↵ Wedel , C. , Forstner , K. U. , Derr , R. & Siegel , T. N . GT-rich promoters can drive RNA pol II transcription and deposition of H2A.Z in African trypanosomes . The EMBO journal 36 , 2581 – 2594 ( 2017 ). doi: 10.15252/embj.201695323 OpenUrl Abstract / FREE Full Text 137. ↵ Cordon-Obras , C. et al. Identification of sequence-specific promoters driving polycistronic transcription initiation by RNA polymerase II in trypanosomes . Cell reports 38 , 110221 ( 2022 ). doi: 10.1016/j.celrep.2021.110221 OpenUrl CrossRef 138. ↵ Lombrana , R. et al. Transcriptionally Driven DNA Replication Program of the Human Parasite Leishmania major . Cell Rep 16 , 1774 – 1786 ( 2016 ). doi: 10.1016/j.celrep.2016.07.007 OpenUrl CrossRef 139. ↵ Rocha-Granados , M. C. & Klingbeil , M. M . Leishmania DNA Replication Timing: A Stochastic Event? Trends in parasitology 32 , 755 – 757 ( 2016 ). doi: 10.1016/j.pt.2016.05.011 OpenUrl CrossRef 140. ↵ Stanojcic , S. et al. Single-molecule analysis of DNA replication reveals novel features in the divergent eukaryotes Leishmania and Trypanosoma brucei versus mammalian cells . Scientific reports 6 , 23142 ( 2016 ). doi: 10.1038/srep23142 OpenUrl CrossRef 141. ↵ Briggs , E. M. et al. Profiling the bloodstream form and procyclic form Trypanosoma brucei cell cycle using single-cell transcriptomics . eLife 12 ( 2023 ). doi: 10.7554/eLife.86325 OpenUrl CrossRef 142. ↵ Marques , C. A. & McCulloch , R . Conservation and Variation in Strategies for DNA Replication of Kinetoplastid Nuclear Genomes . Current genomics 19 , 98 – 109 ( 2018 ). doi: 10.2174/1389202918666170815144627 OpenUrl CrossRef 143. ↵ Garcia-Silva , M. R. et al. Identification of the centromeres of Leishmania major: revealing the hidden pieces . EMBO reports 18 , 1968 – 1977 ( 2017 ). doi: 10.15252/embr.201744216 OpenUrl Abstract / FREE Full Text 144. ↵ Parajuli , S. et al. Human ribonuclease H1 resolves R-loops and thereby enables progression of the DNA replication fork . The Journal of biological chemistry 292 , 15216 – 15224 ( 2017 ). doi: 10.1074/jbc.M117.787473 OpenUrl Abstract / FREE Full Text 145. ↵ Costantino , L. & Koshland , D . Genome-wide Map of R-Loop-Induced Damage Reveals How a Subset of R-Loops Contributes to Genomic Instability . Molecular cell 71 , 487 – 497 e483 ( 2018 ). doi: 10.1016/j.molcel.2018.06.037 OpenUrl CrossRef PubMed 146. Lombrana , R. , Almeida , R. , Alvarez , A. & Gomez , M . R-loops and initiation of DNA replication in human cells: a missing link? Frontiers in genetics 6 , 158 ( 2015 ). doi: 10.3389/fgene.2015.00158 OpenUrl CrossRef 147. Veetil , R. T. , Malhotra , N. , Dubey , A. & Seshasayee , A. S. N . Laboratory Evolution Experiments Help Identify a Predominant Region of Constitutive Stable DNA Replication Initiation . mSphere 5 ( 2020 ). doi: 10.1128/mSphere.00939-19 OpenUrl Abstract / FREE Full Text 148. ↵ Kogoma , T . Stable DNA replication: interplay between DNA replication, homologous recombination, and transcription . Microbiology and molecular biology reviews : MMBR 61 , 212 – 238 ( 1997 ). OpenUrl 149. ↵ Maduike , N. Z. , Tehranchi , A. K. , Wang , J. D. & Kreuzer , K. N . Replication of the Escherichia coli chromosome in RNase HI-deficient cells: multiple initiation regions and fork dynamics . Molecular microbiology 91 , 39 – 56 ( 2014 ). doi: 10.1111/mmi.12440 OpenUrl CrossRef PubMed 150. ↵ Rondon , A. G. & Aguilera , A . What causes an RNA-DNA hybrid to compromise genome integrity? DNA Repair (Amst ) 81 , 102660 ( 2019 ). doi: 10.1016/j.dnarep.2019.102660 OpenUrl CrossRef 151. ↵ Laughery , M. F. , Mayes , H. C. , Pedroza , I. K. & Wyrick , J. J . R-loop formation by dCas9 is mutagenic in Saccharomyces cerevisiae . Nucleic Acids Res 47 , 2389 – 2401 ( 2019 ). doi: 10.1093/nar/gky1278 OpenUrl CrossRef 152. ↵ (!!! INVALID CITATION !!! (20)). 153. ↵ Matos , D. A. et al. ATR Protects the Genome against R Loops through a MUS81-Triggered Feedback Loop . Molecular cell 77 , 514 – 527 e514 ( 2020 ). doi: 10.1016/j.molcel.2019.10.010 OpenUrl CrossRef 154. ↵ Bussotti , G. et al. Genome instability drives epistatic adaptation in the human pathogen Leishmania . Proceedings of the National Academy of Sciences of the United States of America 118 ( 2021 ). doi: 10.1073/pnas.2113744118 OpenUrl Abstract / FREE Full Text 155. ↵ Piel , L. et al. Experimental evolution links post-transcriptional regulation to Leishmania fitness gain . PLoS pathogens 18 , e1010375 ( 2022 ). doi: 10.1371/journal.ppat.1010375 OpenUrl CrossRef 156. ↵ Reis-Cunha , J. L. et al. Ancestral aneuploidy and stable chromosomal duplication resulting in differential genome structure and gene expression control in trypanosomatid parasites . Genome research ( 2024 ). doi: 10.1101/gr.278550.123 OpenUrl Abstract / FREE Full Text 157. ↵ Faria , J. , Briggs , E. M. , Black , J. A. & McCulloch , R . Emergence and adaptation of the cellular machinery directing antigenic variation in the African trypanosome . Current opinion in microbiology 70 , 102209 ( 2022 ). doi: 10.1016/j.mib.2022.102209 OpenUrl CrossRef 158. ↵ Faria , J. et al. Spatial integration of transcription and splicing in a dedicated compartment sustains monogenic antigen expression in African trypanosomes . Nat Microbiol 6 , 289 – 300 ( 2021 ). doi: 10.1038/s41564-020-00833-4 OpenUrl CrossRef 159. ↵ Blumenfeld , B. et al. Chromosomal coordination and differential structure of asynchronous replicating regions . Nature communications 12 , 1035 ( 2021 ). doi: 10.1038/s41467-021-21348-4 OpenUrl CrossRef 160. ↵ Santos , R. et al. A DiCre recombinase-based system for inducible expression in Leishmania major . Molecular and biochemical parasitology 216 , 45 – 48 ( 2017 ). doi: 10.1016/j.molbiopara.2017.06.006 OpenUrl CrossRef 161. ↵ Beneke , T. et al. A CRISPR Cas9 high-throughput genome editing toolkit for kinetoplastids . R Soc Open Sci 4 , 170095 ( 2017 ). doi: 10.1098/rsos.170095 OpenUrl CrossRef PubMed 162. ↵ Duncan , S. M. et al. Conditional gene deletion with DiCre demonstrates an essential role for CRK3 in Leishmania mexicana cell cycle regulation . Molecular microbiology 100 , 931 – 944 ( 2016 ). doi: 10.1111/mmi.13375 OpenUrl CrossRef 163. ↵ Damasceno , J. D. et al. Conditional genome engineering reveals canonical and divergent roles for the Hus1 component of the 9-1-1 complex in the maintenance of the plastic genome of Leishmania . Nucleic Acids Res 46 , 11835 – 11846 ( 2018 ). doi: 10.1093/nar/gky1017 OpenUrl CrossRef 164. ↵ Hahne , F. & Ivanek , R . Visualizing Genomic Data Using Gviz and Bioconductor . Methods Mol Biol 1418 , 335 – 351 ( 2016 ). doi: 10.1007/978-1-4939-3578-9_16 OpenUrl CrossRef PubMed 165. ↵ Afgan , E. et al. The Galaxy platform for accessible, reproducible and collaborative biomedical analyses: 2018 update . Nucleic Acids Res 46 , W537 – W544 ( 2018 ). doi: 10.1093/nar/gky379 OpenUrl CrossRef PubMed 166. ↵ Bolger , A. M. , Lohse , M. & Usadel , B . Trimmomatic: a flexible trimmer for Illumina sequence data . Bioinformatics 30 , 2114 – 2120 ( 2014 ). doi: 10.1093/bioinformatics/btu170 OpenUrl CrossRef PubMed Web of Science 167. ↵ Li , H. & Durbin , R . Fast and accurate short read alignment with Burrows-Wheeler transform . Bioinformatics 25 , 1754 – 1760 ( 2009 ). doi: 10.1093/bioinformatics/btp324 OpenUrl CrossRef PubMed Web of Science 168. ↵ Garrison , E. & Marth , G . Haplotype-based variant detection from short-read sequencing . arXiv e-prints ( 2012 ). . 169. ↵ Danecek , P. et al. The variant call format and VCFtools . Bioinformatics 27 , 2156 – 2158 ( 2011 ). doi: 10.1093/bioinformatics/btr330 OpenUrl CrossRef PubMed Web of Science View the discussion thread. Back to top Previous Next Posted May 01, 2024. Download PDF Supplementary Material Email Thank you for your interest in spreading the word about bioRxiv. NOTE: Your email address is requested solely to identify you as the sender of this article. Your Email * Your Name * Send To * Enter multiple addresses on separate lines or separate them with commas. You are going to email the following R-loops acted on by RNase H1 are a determinant of chromosome length-associated DNA replication timing and genome stability in Leishmania Message Subject (Your Name) has forwarded a page to you from bioRxiv Message Body (Your Name) thought you would like to see this page from the bioRxiv website. Your Personal Message CAPTCHA This question is for testing whether or not you are a human visitor and to prevent automated spam submissions. Share R-loops acted on by RNase H1 are a determinant of chromosome length-associated DNA replication timing and genome stability in Leishmania Jeziel D. Damasceno , Emma M. Briggs , Marija Krasilnikova , Catarina A. Marques , Craig Lapsley , Richard McCulloch bioRxiv 2024.04.29.591643; doi: https://doi.org/10.1101/2024.04.29.591643 Share This Article: Copy Citation Tools R-loops acted on by RNase H1 are a determinant of chromosome length-associated DNA replication timing and genome stability in Leishmania Jeziel D. Damasceno , Emma M. Briggs , Marija Krasilnikova , Catarina A. Marques , Craig Lapsley , Richard McCulloch bioRxiv 2024.04.29.591643; doi: https://doi.org/10.1101/2024.04.29.591643 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Microbiology Subject Areas All Articles Animal Behavior and Cognition (7644) Biochemistry (17725) Bioengineering (13915) Bioinformatics (42031) Biophysics (21486) Cancer Biology (18635) Cell Biology (25548) Clinical Trials (138) Developmental Biology (13397) Ecology (19937) Epidemiology (2067) Evolutionary Biology (24361) Genetics (15619) Genomics (22538) Immunology (17763) Microbiology (40468) Molecular Biology (17206) Neuroscience (88733) Paleontology (667) Pathology (2842) Pharmacology and Toxicology (4834) Physiology (7657) Plant Biology (15172) Scientific Communication and Education (2046) Synthetic Biology (4304) Systems Biology (9831) Zoology (2272)