Distinct chromatin regulators downmodulate meiotic axis protein deposition and DNA break induction at chromosome ends

13 In many organisms, meiotic crossover recombination is suppressed near the extreme ends of 14 chromosomes. Here, we identified two chromatin modifiers, the histone methyltransferase Dot1 15 and the Sir silencing complex, as regulators of this process in Saccharomyces cerevisiae. We 16 show that the recombination-promoting axis proteins Red1 and Hop1, but not the axis-associated 17 cohesin Rec8, are significantly reduced within 20 kb of telomeres compared to the chromosome 18 interior. Dot1, which preferentially methylates histones in the chromosome interior, is required for 19 this pattern by directing Red1 binding toward the chromosome interior. In parallel, the Sir complex 20 suppresses the induction of meiotic DNA double-strand breaks (DSBs) at chromosome ends. Sir-21 dependent DSB suppression is independent of axis deposition and occurs in a chromosome end-22 specific manner that mirrors the spreading and transcriptional silencing activity of the complex, 23 suggesting that the Sir complex suppresses DSB formation by limiting the openness of promoters, 24 the preferred sites of meiotic DSB formation. We conclude that multiple chromatin -based 25 mechanisms collaborate to achieve a robust reduction of meiotic recombination near 26 chromosome ends. 27

meiotic DSBs, subtelomeres, axis proteins, chromatin modifiers, Dot1, Sir2, Sir3 28 29 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 3

30 Meiosis is a specialized type of cell division that generates haploid gametes from diploid 31 progenitor cells and plays an essential role in promoting genetic diversity 1. Meiosis initiates with 32 a single round of DNA replication followed by two rounds of chromosome segregation: the first 33 segregates homologous chromosomes, and the second segregates sister chromatids2,3. Accurate 34 segregation during meiosis I relies on meiotic crossover recombination, which exchanges DNA 35 between homologous chromosome pairs and, together with sister chromatid cohesion, forms a 36 physical connection between them 4. Meiotic recombination is initiated by programmed double -37 strand breaks (DSBs), which are catalyzed by Spo11, a highly conserved topoisomerase -like 38 enzyme5. 39 Although DSBs can occur throughout the genome, their frequency varies widely. Chromosomal 40 regions classified as “hot” experience frequent DSBs, whereas “cold” regions rarely undergo 41 breakage6. This spatial regulation results from a complex interplay of factors, including local base 42 composition, DNA accessibility, chromatin modifications, and meiotic chromosome 43 architecture5,7,8. Meiosis-specific axis proteins, which localize at the base of the loop-axis structure 44 of meiotic chromosomes, are also linked to DSB activity and the subsequent homolog-directed 45 repair6,9-11. In S. cerevisiae, the axis proteins Red1 and Hop1 are required for both processes11-46 14, and elevated levels of Red1 and Hop1 correlate with increased DSB formation and markers of 47 crossover repair15,16. Red1 and Hop1 are recruited to the base of chromatin loops by the meiotic 48 Rec8 cohesin complex but can also bind directly to chromatin through the nucleosome -binding 49 activity of Hop114,15,17, leading to two independent modes of axis-dependent patterning of meiotic 50 recombination. 51 In budding yeast, DSB activity is notably reduced within ~20 kb from telomeres18. This depletion 52 is likely important for two reasons. First, the ends of chromosomes are enriched for repetitive DNA 53 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 4 sequences that are vulnerable to non -allelic homologous recombination (NAHR) and genome 54 rearrangements19. Second, crossovers near chromosome ends are less effective at forming 55 stable connections between homologous chromosomes and may lead to chromosome mis -56 segregation if they are the sole link between homolog pairs20. Indeed, recombination events near 57 telomeres are linked to a higher risk of Trisomy 21 (Down syndrome) in humans21,22. 58 S. cerevisiae chromosome ends consist of several distinct regions ( Fig. 1a)23,24. In addition to 59 telomerase-templated telomeric repeats that cap chromosome ends, “telomere -associated 60 sequences” contain repetitive Y’ and X elements, while the “subtelomeric domains” extend inward 61 for an average of about 20 kb from telomeres. Subtelomeric domains are relatively gene -poor 62 regions enriched in gene families 25,26. The telomeric repeats and X elements recruit the 63 Sir2/Sir3/Sir4 histone deacetylase complex, establishing transcriptional silencing. This silent 64 chromatin can spread over limited distances into the subtelomeric domains 27. The subtelomeric 65 domains, in turn, are defined by a unique chromatin signature, which includes a relative depletion 66 of active chromatin marks25,28. 67 Previous studies of meiotic recombination in S. cerevisiae have consistently shown a depletion of 68 DSBs in X and Y’ elements as well as in subtelomeric domains 10,18,29. While the mechanisms 69 governing DSB suppression in X and Y’ elements remain largely unexplored, the reduced DSB 70 levels in subtelomeric domains are accompanied by lower recruitment of DSB-promoting factors, 71 such as Rec11430, and diminished abundance of the axis protein Hop1 31. Hop1, which recruits 72 Rec11411, is currently the most upstream regulator of recombination known to be depleted about 73 20 kb from the chromosome ends. However, the specific mechanisms controlling this altered 74 distribution of Hop1 —and whether its depletion is the sole driver of suppressed recombination 75 near telomeres—are not understood. 76

77 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 5 The meiotic axis extends into the telomere-associated X and Y’ sequences 78 Given the reduced axis protein enrichment near chromosome ends and their critical role in meiotic 79 DSB formation and repair, we investigated how axis proteins interact with telomere -associated 80 sequences and subtelomeric domains. These regions are often excluded from sequence -based 81 analyses due to variations in telomere organization even among closely related yeast 82 strains24,32,33, and because the abundance of repetitive sequences and gene families creates 83 challenges for uniquely assigning sequencing reads. Therefore, we tailored our analysis pipeline 84 to account for these unique biological features. To reduce structural mapping artifacts, we 85 mapped reads to high -quality end -to-end chromosome scaffolds of our experimental strain 86 (SK1)32, rather than the commonly used S288c reference. Further, by exclusively considering 87 optimal matches, we maximized the number of confidently mapped reads, leveraging inherent 88 sequence polymorphisms within telomeric repeat sequences for unique mapping. This approach 89 resulted in high mapping success across all telomere-proximal regions (Supplementary Fig. 1a). 90 Even in the highly repetitive X and Y’ elements, our pipeline uniquely mapped 12.81% and 0.87% 91 of single -end reads and 15.18% and 12.80% of paired -end reads, respectively. In the 92 subtelomeric domains, unique mapping rates were higher, averaging 24.07% for single-end reads 93 and 32.21% for paired-end reads. 94 If a read had multiple equally good mapping results, our pipeline randomly selected one location 95 as the primary alignment for the read. This approach did not affect metagene analyses but 96 significantly improved signal clarity at individual chromosome ends by closing coverage gaps 97 caused by a lack of polymorphisms. Comparative analysis of profiles with and without multiple 98 mapping reads revealed no qualitative differences (Supplementary Fig . 1b), supporting the 99 robustness of our approach. 100 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 6 With this strategy, we analyzed the distribution of axis proteins at chromosome ends using 101 previously published ChIP-seq datasets14,31,34. Meta-analysis revealed that Red1 and Hop1 were 102 significantly less enriched than the genome average within 20 kb of chromosome ends (Fig. 1b-103 c), consistent with previous studies31. Axis protein depletion was observed across all chromosome 104 ends, and thus could not be attributed to individual telomere outliers (Supplementary Fig. 2). 105 Yeast chromosome ends are characterized by telomeric repeats, (TG 1–3)n, and telomere-106 associated sequences, which include the Y’ and X elements35. The long Y’ elements (4-7 kb) are 107 not found on all ends and can occur as one or multiple copies, whereas a short X element (~500 108 bp) is present at essentially all chromosome ends 32,36 (Fig. 1a). Based on these features, we 109 classified chromosome ends into two categories: X-only ends, which contain only the X element, 110 and XY’ ends, which include both X and Y’ elements. Meta -plots of Red1 and Hop1 profiles 111 revealed that axis protein enrichment differed between these two categories. X elements at X -112 only ends exhibited axis protein enrichment close to the genome average, indicating that isolated 113 X elements have a substantial propensity for axis protein recruitment (Fig. 1d). By contrast, X 114 elements at XY’ ends showed Red1 and Hop1 enrichment below the genome average (Fig. 1d), 115 suggesting that the presence of Y’ elements reduces axis protein binding on adjacent X elements 116 (Fig. 1f). This reduction may result from axis protein repositioning, as meta-analyses revealed an 117 axis protein binding site at the telomere-proximal side of the Y’ element (Fig. 1e). 118 Multiple cis-acting features contribute to axis protein depletion near telomeres 119 Altered axis protein binding on adjacent X elements suggests that Y’ sequences may partially 120 drive the telomere-proximal depletion of axis proteins observed in our meta-analysis (Fig. 1b-c). 121 In support of this, although both XY’ and X -only chromosome ends show reduced axis protein 122 binding compared to the genome average, the depletion is stronger at XY’ ends (Fig. 2a). These 123

indicate that Y’ elements contribute to the overall reduction of axis proteins at chromosome 124 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 7 ends. However, the observation that axis protein levels are also lower at X-only ends implies that 125 additional mechanisms further suppress axis protein recruitment near telomeres. 126 To determine whether this additional mechanism is due to telomere proximity or intrinsic 127 sequence features, we analyzed chromosome fusions, in which subtelomeric domains were 128 relocated to the chromosome interior 37. We observed that axis protein distribution at fused 129 chromosome ends mirrored that of native, unfused ends (Fig. 2b), suggesting that reduced axis 130 protein enrichment at chromosome ends is driven by specific sequences encoded at chromosome 131 ends rather than by telomere proximity. 132 Across the genome, gene -rich regions exhibit an overall higher axis protein enrichment 14,15. As 133 the subtelomeric domains are comparatively gene -poor25,26, we plotted coding density (the 134 fraction of DNA encoding open reading frames) as a function of distance from the telomeres. This 135 analysis revealed a strong correlation between the regions of reduced coding density and axis 136 protein depletion (Fig. 2c), suggesting that reduced coding density may partially underlie the 137 reduced axis protein recruitment within 20 kb of telomeres. 138 Rec8-dependent and independent pathways mediate axis protein localization at 139 chromosome ends 140 Chromosomal recruitment of Red1 and Hop1 is mediated by two parallel pathways. The meiotic 141 Rec8-cohesin complex preferentially recruits axis proteins to sites of convergent transcription14, 142 whereas the chromatin binding region (CBR) of Hop1 directs axis proteins to nucleosome-dense 143 regions—“islands” characterized by higher coding density 15,17. We sought to determine whether 144 the depletion of axis proteins near chromosome ends could be attributed to the inactivity of one 145 of these pathways. 146 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 8 Analysis of ChIP-seq data revealed that Rec8 is abundantly present at chromosome ends. Unlike 147 Red1 and Hop1, Rec8-cohesin levels in the subtelomeric regions were comparable to the genome 148 average (Fig. 1b, c). Moreover, the regions enriched for Red1 and Hop1 in the telomere -149 associated sequences were also enriched for Rec8: Rec8 was bound more strongly to the X 150 elements of X-only ends than to XY’ ends (Fig. 1d, f) and formed a strong peak at the telomere-151 proximal side of Y’ elements (Fig. 1e). This peak coincided with the 3’ end of the Y’-encoded open 152 reading frame (ORF), consistent with other chromosomal Rec8 peaks, which are typically 153 enriched downstream of ORFs14,38. Previous studies have demonstrated that active transcription 154 can induce the sliding of the cohesin ring and direct axis protein association to the end of 155 ORFs14,39,40. Therefore, transcription of the Y’ ORF may similarly influence axis protein deposition 156 within Y’ elements. 157 The coincident binding of axis proteins and Rec8 supports the hypothesis that Rec8 -cohesin is 158 an important contributor to axis protein deposition at chromosome ends. To directly test this 159 possibility, we examined Red1 binding in a rec8 mutant strain using spike -in normalized ChIP-160 seq datasets15,34. In the subtelomeric domains, Red1 levels were significantly reduced in the rec8 161 mutant compared to wild -type (Fig. 3a-b). However, Red1 binding was still detectable above 162 baseline levels. Axis protein binding in the absence of Rec8 depends on the CBR domain of 163 Hop115,17. Indeed, the persistent subtelomeric axis protein signal in rec8 mutants was abolished 164 upon introduction of a Hop1 mutant (hop1-phd) lacking the CBR15,17 (Fig. 3a-b). By contrast, the 165 hop1-phd mutation alone had comparatively minor effects in the subtelomeric domains, although 166 we consistently noted reduced Red1 binding in the neighboring end-adjacent regions (EARs, 20-167 120 kb)31 (Fig. 3a-b). These data indicate that both pathways of axis recruitment additively 168 contribute to Red1 binding at chromosome ends, with Rec8-cohesin responsible for recruiting the 169 majority of Red1, consistent with the low coding density in these regions (Fig. 2c). Similar additive 170 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 9 effects were observed in the X and Y’ sequences (Fig. 3c, d). Therefore, the selective inactivation 171 of one recruitment pathway cannot explain the depletion of axis proteins in subtelomeric domains. 172 The methyltransferase Dot1 is required for subtelomeric depletion of axis proteins 173 Subtelomeric regions share qualitative similarities with pericentromeric regions, particularly in the 174 differential enrichment of axis proteins and Rec8 -cohesin. Both regions show a relatively higher 175 abundance of Rec8-cohesin compared to Red1, with most data points falling below the genome-176 wide regression line (Fig. 4a). At pericentromeres, this differential enrichment of axis factors 177 persists even if the centromere itself is inactivated 37, indicating that local DNA or chromatin 178 environment influences axis protein binding. Given the distinct chromatin state of subtelomeric 179 domains, we investigated whether chromatin modifiers specific to these regions contribute to axis 180 protein depletion. 181 We focused our analysis on histone marks related to meiotic DSB formation 41,42 or those 182 specifically different in the subtelomeric domains28,42-44. ChIP-seq analysis identified two marks - 183 H3K4me3 and H3K79me3 - that closely matched the pattern of axis protein depletion (Fig. 4b). 184 Both marks are long -lasting indicators of active gene expression that are depleted from 185 subtelomeric domains in vegetative cells 25,28 and have been implicated in the control of meiotic 186 DSB formation41,45. To explore the role of these histone modifications in axis protein depletion, we 187 used spike-in normalized ChIP-seq to analyze Red1 in mutants lacking Set1 or Dot1, the enzymes 188 responsible for the trimethylation of H3K4 and H3K79, respectively 34. In set1Δ mutants, 189 subtelomeric Red1 binding patterns were indistinguishable from wild-type (Fig. 4c-e). In contrast, 190 Red1 levels in dot1Δ mutants were markedly elevated and no longer significantly different from 191 the genome average (Fig. 4c-e), suggesting that Dot1 suppresses axis protein binding in 192 subtelomeric domains. Red1 levels were also elevated across Y’ elements and moderately over 193 X elements (Fig. 4g, h). Notably, pericentromeric axis protein recruitment in dot1Δ mutants did 194 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 10 not show a statistically significant difference by bootstrapping analysis, although the average 195 binding values were slightly reduced compared to wild-type (Fig. 4f). In contrast, subtelomeric 196 regions exhibited the opposite effect, with average binding levels increasing in dot1Δ mutants 197 (Fig. 4e). 198 Subtelomeric regions in wild-type cells are undermethylated on H3K79 (Fig. 4b), suggesting that 199 the role of Dot1 in axis protein regulation may be indirect. In line with this interpretation, spike -in 200 normalized ChIP-seq data showed that the increased Red1 binding in subtelomeric regions of 201 dot1Δ mutants was accompanied by lower Red1 levels in the chromosome interior (Fig. 4c-e). 202 To probe which axis recruitment pathway is regulated by Dot1, we separated Red1 signals into 203 genic and intergenic regions, as Hop1 primarily recruits Red1 to gene bodies, whereas Rec8 -204 cohesin leads to Red1 enrichment in intergenic regions 14. The reduced binding of Red1 in the 205 chromosome interior in dot1Δ mutants primarily affected intergenic regions, implying that Dot1 206 promotes Rec8-dependent axis binding. Consistent with this interpretation, the increase in Red1 207 association near chromosome ends in dot1Δ mutants occurred predominantly on gene bodies 208 (Fig. 4i). These data indicate that the chromatin requirements for axis recruitment differ between 209 chromosome ends and the chromosome interior, and that Dot1 helps direct Red1 to chromosome 210 interiors by promoting Rec8-dependent axis recruitment. 211 Conservation of Sir-dependent telomeric heterochromatin during meiotic recombination 212 One consequence of DOT1 disruption is the spreading of the Sir complex beyond its usual 213 boundaries46,47. Indeed, the mislocalization of Sir2 and Sir3 in dot1 mutants was previously linked 214 to checkpoint defects during meiotic recombination48. To investigate this link further, we examined 215 the role of the Sir complex in suppressing axis protein deposition and DSB formation at 216 chromosome ends. 217 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 11 In vegetative cells, the Sir complex establishes silent chromatin domains at mating-type loci and 218 telomere-associated sequences49,50. At some chromosome ends, Sir chromatin also spreads from 219 the X element into the neighboring subtelomeric domains 27,51, although spreading across entire 220 subtelomeric domains is only observed under conditions of Sir overexpression28. We used ChIP-221 seq analysis to assess Sir3's chromatin association and spreading during meiotic prophase. 222 Our observations at the time of meiotic induction and during the peak of DSB formation (3 hours 223 post-induction) revealed consistent binding patterns (Fig. 5a). Although Sir3 was enriched at both 224 types of X elements, its binding on X elements at XY’ ends was lower than that observed at X -225 only ends (Fig. 5a, c). Sir3 was also enriched upstream of the Y’-encoded ORF, but binding was 226 relatively diminished across the Y’ ORF (Fig. 5b). Additionally, we observed heterogeneous 227 spreading of Sir3 from subtelomeric X elements into adjacent subtelomeric domains on certain 228 chromosome ends. For example, on chrVII-L, Sir3 spread approximately 6 kb from the X element 229 during early prophase I (3 hours post -meiotic induction) (Fig. 5d), while there was no apparent 230 spreading on chrVI -L (Fig. 5e). The extent of Sir3 spreading varied substantially and did not 231 correlate with the presence of Y’ elements but mirrored the distribution of Sir proteins in vegetative 232 cells27,28. These findings indicate that Sir -dependent telomeric heterochromatin remains largely 233 unchanged as cells initiate meiotic recombination. 234 Sir3 spreading into subtelomeric domains suppresses meiotic transcription but does not 235 affect axis protein deposition. 236 To determine whether subtelomeric domains with Sir3 spreading exhibit altered transcription of 237 underlying genes, we conducted mRNA-seq analysis on samples collected 3 hours post-meiotic 238 induction in the presence or absence of SIR3. Plotting relative fold changes in mRNA levels as a 239 function of the average Sir3 occupancy in the associated promoter region (250bp upstream of 240 each gene), revealed a significant correlation between Sir3 occupancy and increased mRNA 241 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 12 levels upon SIR3 deletion (Fig. 6a) in the 5 kb regions adjacent to the X element where Sir3 242 frequently spreads. These results indicate that the transcriptional effects of Sir3 are confined to 243 sequences where Sir3 spreading occurs, as detected by ChIP-seq. 244 Given the prominent Sir3 peak observed at the start of the Y’ element metaplot (Fig. 5b), we also 245 analyzed Y’ element expression at 3 hours post -meiotic induction. The average Y’ element 246 expression was significantly increased in the absence of SIR3 (Supplementary Fig. 3). These 247

suggest that the absence of SIR3 impacts Y’ element transcription. 248 We asked whether deletion of SIR3 also affects the deposition of axis proteins near chromosome 249 ends by determining the distribution of Red1 in sir3 mutants using spike-in normalized ChIP-seq 250 analysis. Meta-analysis revealed no significant difference in the average depletion of Red1 in the 251 last 20 kb between sir3 and wild-type strains (Supplementary Fig. 4a, b). Similarly, meta-plots 252 of average Red1 profiles on X and Y’ elements in sir3 mutants showed no new peaks compared 253 to wild-type (Supplementary Fig. 4c, d), although enrichment levels were lower on both types of 254 X element (Supplementary Fig. 4c). The overall genome -wide median binding levels of Red1 255 were higher in the sir3 mutants (Supplementary Fig. 4b), aligning with the previously reported 256 analyses of Hop1 31. These findings indicate that Sir3 binding, and the establishment of silent 257 chromatin domains do not result in large-scale alterations to the binding landscape of axis proteins 258 at chromosome ends. They also imply that the altered axis protein binding seen in dot1 mutants 259 is not a consequence of the increased Sir complex spreading in this mutant. 260 Sir proteins protect X elements and regions of Sir spreading from meiotic DSBs 261 We wondered how Sir-dependent heterochromatin and Dot1-dependent axis protein suppression 262 interface with the meiotic DSB machinery near chromosome ends. Genome-wide DSB levels had 263 previously been determined in sir2Δ strains using Spo11-oligo sequencing, which sequences the 264 DNA fragments that remain covalently attached to Spo11 after cleavage 31,52, but telomere -265 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 13 proximal DSB formation had not been investigated in that study. Analysis of this dataset revealed 266 a significant increase in DSB levels on X elements but not on Y’ elements in sir2Δ strains (Fig. 267 6b). This pattern mirrors the relative enrichment of Sir3 in these elements, indicating that Sir 268 chromatin suppresses DSB formation in the X elements. 269 We also observed increased DSB formation in the subtelomeric domains in sir2Δ mutants. 270 Increased DSB induction was correlated with the extent of Sir3 occupancy in wild-type cells (Fig. 271 6c). Accordingly, increased DSB formation also correlated with elevated gene expression in the 272 same regions in sir3 mutants but not in the rest of the genome (Fig. 6d). These findings suggest 273 that elevated promoter openness, which enables increased gene expression, also creates a 274 window for meiotic DSB formation. 275 To complement and expand this analysis, we analyzed sir3 and dot1Δ mutants by TrAEL-seq, 276 which sequences the exposed 3’ ends that result from Spo11 removal53. To avoid signal changes 277 due to DSB repair, this analysis was conducted in a repair-defective dmc1Δ background54. TrAEL-278 seq analysis at the 5-hour time point showed a distinct increase in DSB formation within 20 kb of 279 telomeres in sir3 mutants, with the number of significant DSB hotspots nearly doubling within 5 280 kb of X elements (Fig. 6e, f). On the other hand, DSB formation was unaffected in the same 281 regions in dot1Δ mutants (Fig. 6e, f). Accordingly, TrAEL-seq analysis of sir3 dot1Δ double 282 mutants revealed DSB patterns similar to sir3 single mutants (Fig. 6e, f). Moreover, a substantial 283 level of DSB suppression persisted even in the double mutant (Fig. 6e, Supplementary Fig. 5b), 284 indicating that additional layers of regulation contribute to DSB suppression at chromosome ends 285 (see Discussion). 286 Intriguingly, TrAEL-seq analysis indicated differing genetic interactions between DOT1 and SIR3 287 in other parts of the genome. DSB levels dropped in the subtelomere-adjacent EARs, specifically 288 in dot1Δ sir3 double mutants, suggesting redundant functions (Supplementary Fig. 5a), whereas 289 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 14 DSB formation around centromeres w as decreased to a similar extent in dot1Δ and sir3 single 290 mutants (Supplementary Fig. 5b). Finally, around the ribosomal DNA locus, DSB levels were 291 increased in a Sir3 -dependent manner in dot1Δ mutants (Supplementary Fig. 5c) , consistent 292 with previously observed Sir -dependent DSB induction in this region 55. These findings imply a 293 combinatorial code that allows these two chromatin regulators to adjust meiotic recombination in 294 a region-specific manner. 295

296 Here we show that meiotic recombination potential at chromosome ends is suppressed by at least 297 two mechanisms. First, the local depletion of axis proteins, which regulate the recruitment of key 298 DSB factors and help target DSBs toward recombination with the homologous chromosome. 299 Second, telomeric heterochromatin restricts access to gene promoters, the preferred DNA 300 substrates for DSB formation in yeast 10 (Fig. 7) . These mechanisms thus represent distinct 301 strategies for regulating meiotic recombination potential. 302 The range of these two mechanisms differs significantly. Dot1 -dependent axis protein depletion 303 spans the entire subtelomeric domain and exhibits consistent patterns across chromosome ends, 304 with variability largely driven by the distribution of axis binding sites14. In contrast, Sir-dependent 305 DSB suppression is highly variable among chromosome ends, driven by the extent of Sir-306 dependent chromatin spreading. This variability in heterochromatin spread is also observed in 307 vegetative cells, where boundaries are influenced by transcription factor binding sites, genic 308 elements, and nucleosome-disfavoring DNA sequences 25,56-60. 309 While Sir -dependent chromatin spreading directly suppresses DSB hotspots , Dot1 likely 310 influences axis deposition indirectly (Fig. 7), as H3K79 trimethylation occurs predominantly in the 311 chromosome interior25. Our findings demonstrate that the dot1Δ mutation causes a genome-wide 312 redistribution of Red1, with reduced binding in the chromosomal interior. Because Red1 levels 313 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 15 are limiting61, one potential explanation is that the absence of Dot1 releases Red1 from intergenic 314 regions, enabling it to bind gene bodies in the subtelomeric domains (Supplementary Fig. 6). 315 Intriguingly, the hop1-phd mutant, which should also release some Red1 because of its failure to 316 bind to gene bodies across the genome 15, does not exhibit similar increases in Red1 binding at 317 subtelomeric domains (Fig. 2a), raising the possibility that the increased axis recruitment to 318 subtelomeric domains in dot1Δ mutants requires the CBR domain. 319 Although subtelomeric regions in dot1Δ mutants exhibit increased DSB potential due to Red1 320 redistribution, this does not result in substantial increases in DSB formation, even in sir3 dot1Δ 321 double mutants in which the suppressive effect of the Sir complex is eliminated. We speculate 322 that the absence of a stronger effect may be the consequence of increased DSB turnover, which 323 would not be detected by our TrAEL-seq analyses. Dot1 is required for the meiotic arrest of dmc1Δ 324 mutants48. As a result, repair foci disappear in dot1Δ dmc1Δ mutants as cells progress into 325 meiosis I, with breaks repaired via Rad54 -dependent inter -sister recombination 48. Moreover, 326 additional regulatory layers of telomere -proximal DSB suppression must exist because a 327 significant level of DSB suppression persists even in sir3 dot1Δ double mutants. These layers 328 may include further chromatin-based mechanisms. Additionally, one likely contributor is the low 329 coding density of subtelomeric regions as the lower density of gene promoters inherently results 330 in a smaller number of potential DSB hotspots. Indeed, the regional drop in coding density 331 correlates well with reduced axis protein recruitment and DSB formation. Our analysis of axis 332 deposition in chromosome fusion strains also supports this notion. Thus, to some extent the low 333 DSB potential may be hardwired into the genome sequence of chromosome ends. The combined 334 action of multiple pathways suppressing recombination at chromosome ends likely ensures that 335 meiotic crossovers are strongly disfavored near telomeres. Telomere -proximal crossovers are 336 less effective at maintaining linkages between homologous chromosomes 62 and are associated 337 with increased risk of Down syndrome in humans 21,22,63,64. The downmodulation of axis proteins 338 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 16 in subtelomeric regions may play a critical role in this process, as axis proteins not only recruit 339 DSB factors but also help target DSB repair to homologous chromosomes by preventing sister 340 chromatid recombination65. Although a low level of DSBs occurs in telomere -associated regions 341 and subtelomeric domains 10, reduced axis protein abundance likely makes these regions less 342 prone to crossover formation. Favoring repair from sister chromatids may also reduce the risk of 343 non-allelic recombination events. Non -allelic recombination is especially risky at chromosome 344 ends due to their repetitive sequences and the close physical proximity of telomeres along the 345 nuclear envelope during meiotic prophase19,66,67. These multiple regulatory layers thus establish 346 a robust protective mechanism to shield chromosome ends from non-allelic and unproductive 347 recombination and ensure proper meiotic chromosome segregation. 348

and Materials: 349 Yeast strains and growth conditions: 350 All the strains utilized in this study belonged to the SK1 background (except for the analysis using 351 the published fusion chromosomes, which are SK1 and S288C hybrids) . The genotypes are 352 detailed in Supplementary Table 1. For experiments using vegetative cells, m id-logarithmic 353 phase cultures were grown in YPD medium overnight at room temperature until saturation. On 354 the following day, the cells were diluted to an OD600 of 0.2 and incubated at 30°C until they 355 reached an OD600 of 1. At this stage, 50 ml of culture was collected for ChIP-seq. 356 To induce synchronous meiotic cultures, strains were grown for 24 hours in YPD medium at room 357 temperature. Cells were enriched in the G1 phase by inoculating them at OD600 = 0.3 in pre -358 sporulation BYTA medium (1% yeast extract, 2% bactotryptone, 1% potassium acetate, 50 mM 359 potassium phthalate) for 16.5 hours at 30°C. Cells were washed twice with sterile water and 360 transferred into SPO medium at OD600 = 1.9 (SPO: 0.3% potassium acetate, 0.001% acetic 361 acid). Cultures were grown at 30°C on a shaker, and the time of inoculation into SPO was defined 362 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 17 as time = 0. At specific timepoints, 25 ml of culture was collected for ChIP -seq and 1.6 ml for 363 mRNA-seq. The synchrony of meiotic cultures was validated using Fluorescence -activated cell 364 sorting (FACS). 365 FACS: 366 Cells collected at various timepoints were fixed overnight at 4°C in 350 μl of absolute ethanol. 367 Cells were pelleted and incubated in 500 μl FACS buffer (50 mM sodium citrate) with 0.7 μl RNase 368 A (20–40 mg/ml stock; Sigma -Aldrich) and 5 μl proteinase K (20 mg/ml; VWR) for 24 hours at 369 50°C. Prior to FACS analysis, cells were stained with 0.1 μl SYTOX Green (5 mM solution in 370 DMSO; Invitrogen) and briefly sonicated for 5 seconds at 10% amplitude. DNA content was 371 analyzed using the BD FACSAria II system at the Genomics Core at New York University Center 372 for Genomics and Systems Biology. 373 Chromatin immunoprecipitation (ChIP) & Illumina sequencing: 374 The cells collected at the indicated timepoints were pelleted and immediately crosslinked for 30 375 minutes in a 1% formaldehyde solution at room temperature with gentle shaking. Subsequently, 376 the crosslinking reaction was quenched by incubating the cells for 5 minutes at room temperature 377 in a 125 mM glycine solution. Following this, the cells were processed according to the protocol 378 outlined in68 and immunoprecipitated with 2.5 μl of either anti-Sir3 (HM01065, a kind gift from the 379 Bell lab) or anti-Red1 (#16440, kind gift of N. Hollingsworth) or anti-H3K79me3 (abcam, ab2621). 380 Library preparation, quality, and quantity checks were then performed as described in 17. All the 381 prepared chromatin immunoprecipitation (ChIP) libraries were sequenced as 150 -bp paired-end 382 reads on Illumina NextSeq 500 instruments. The sequencing run was conducted by the Genomics 383 Core at New York University Center for Genomics and Systems Biology. 384 Processing ChIP -seq reads from Illumina sequencing: 385 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 18 Illumina sequencing reads were aligned to the SK1 genome 32 using Bowtie 2 (Version 2.4.2) 69. 386 To improve mapping of reads to repetitive subtelomeric regions, the Bowtie 2 read reporting mode 387 was configured to allow multiple alignments, reporting the best match69. MACS2 2.1.1 was used 388 to extend reads in the 5'-> 3' direction to a final length of 200 bp. SPMR (signal per million reads) 389 normalization was applied to both Input and ChIP pileups using MACS2. The resulting fold -390 enrichment files were used for downstream analysis in R. The ChIP -seq pipeline and analysis 391 scripts are available on the Hochwagen Lab GitHub page 392 https://github.com/hochwagenlab/ChromosomeEnds.git. 393 mRNA-sequencing & analysis: 394 mRNA extraction, first - and second -strand synthesis, and library preparation were conducted 395 following the procedures outlined in 70. The resulting libraries were subjected to sequencing as 396 150-bp paired-end reads on Illumina NextSeq 500 instruments. The sequencing run was executed 397 by the Genomics Core at New York University Center for Genomics and Systems Biology. The 398 GTF (General Transfer Format) file was modified to include all completely annotated Y’ elements. 399 Subsequently, reads obtained from Illumina sequencing were aligned to the SK1 genome32, and 400 the counts mapping to the modified SK1 gtf file were determined using the nf -core RNA-Seq 401 pipeline71,72. The relative abundances, measured as Transcripts per million (TPM) in the Salmon73 402 output file "salmon.merged.gene_counts_length_scaled.rds ", were utilized for downstream 403 analysis in R. 404 Spo11-oligos mapping: 405 In this study, previously published Spo11-oligos datasets from WT and sir2Δ strains31,52 were 406 utilized. Adaptors were removed from the reads using fastx_clipper from the FASTX -Toolkit 407 (version 0.0.14) and reads shorter than 15 bp were discarded. Subsequently, the clipped reads 408 were aligned to the SK1 genome 32 using Bowtie 2 (Version 2.4.2) 69 . The read alignment mode 409 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 19 was set to local, and the reporting mode allowed multiple read alignments while reporting the best 410 alignment69. Following alignment, MACS2 2.2 (https://github.com/taoliu/MACS) was employed to 411 extend reads in the 5' →3' direction to a final length of 37 bp 74. SPMR (signal per million reads) 412 normalization was performed on the pileups. 413 Plug preparation for TrAEL-seq: 414 Plugs were prepared from meiotic cultures harvested 5 hours into meiosis. For each strain, 20 ml 415 of culture was pelleted and washed twice with CHEF TE buffer (10 mM Tris-HCl, pH 7.5, 50 mM 416 EDTA, pH 8.0). The cell pellet was resuspended in 300 µl CHEF TE, followed by the addition of 417 4 µl zymolyase (10 mg/ml). The mixture was briefly vortexed and incubated at 42°C for 418 approximately 30 seconds. Low-melting-point agarose (1% SeaPlaque GTG in 125 mM EDTA, 419 pH 8.0), prewarmed to 42°C, was added and transferred into plug molds. Plugs were solidified on 420 ice for 10 minutes before being transferred into LET buffer (10 mM Tris-HCl, pH 7.5; 0.5 M EDTA, 421 pH 8.0). Plugs were incubated overnight at 37°C. 422 The following day, plugs were treated with proteinase K in NDS buffer (10 mM Tris -HCl, pH 7.5; 423 0.5 M EDTA ; 1% N -lauroylsarcosine) at 50°C overnight. After digestion, plugs were washed 424 sequentially: first with CHEF TE for 1 hour at room temperature, followed by two washes with 425 CHEF TE containing 10 µl freshly prepared PMSF (100 mM) in ethanol, with each wash incubated 426 for 1 hour in the cold room. A fourth wash was performed with RNase T1 in CHEF TE, incubated 427 at 37°C for 1 hour, followed by a final wash with CHEF TE at room temperature for 1 hour. Plugs 428 were stored at 4°C in CHEF TE until further use. 429 TrAEL-seq Library Preparation, Sequencing, and Data Analysis: 430 TrAEL-seq library preparation, sequencing and data analysis were conducted following the 431 procedures outlined in 53. TrAEL-seq signal pileups were generated using MACS2 (v2.2) and 432 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 20 normalized pileups were generated using SPMR normalization. Downstream analyses were done 433 in R. 434 Supporting information: Supplementary Figures and Tables are attached in a word document. 435 Data reporting: The datasets generated and analyzed in this paper, excluding published 436 datasets, have been deposited in the Gene Expression Omnibus (GEO). The datasets can be 437 accessed with the accession number for ChIP -seq: GSExxxxxx, for RNA-seq: GSExxxxxx, and 438 for TrAEL-seq: GSExxxxxx. Additionally, all ChIP-seq datasets used in this study are described 439 in Supplementary Table 2 , and all Spo11 -oligos, RNA -seq and TrAEL -seq datasets are 440 described in Supplementary Table 3. 441 Conflict of interest: The authors declare no conflicting interests. 442

443 We express our sincere gratitude to Stephen P. Bell for generously providing the Sir3 antibody 444 and to Nancy Hollingsworth for the Red1 antibody. We also acknowledge the Genomics Core at 445 the New York University Center for Genomics and Systems Biology for their valuable technical 446 assistance and expertise in data processing. This work was supported in part by the NYU IT High-447 Performance Computing resources, services, and staff expertise. We are grateful to the Zegar 448 Family Foundation for their generous support. TrAEL-seq library sequencing and processing were 449 performed by the Genomics (Geno06) and Bioinformatics (Bioinf01) teams at the Babraham 450 Institute, which receive financial support from the Institute Core Capability Grant (BBSRC CCG). 451 This research was financially supported as part of NIH grant R35 GM148223 to AH. ARR 452 acknowledges support from a Fleur Strand Graduate Fellowship from the Department of Biology, 453 as well as a Henry MacCracken Fellowship and a Dean’s Dissertation Fellowship from the NYU 454 Graduate School of Arts and Science. JH acknowledges funding from the BBSRC (BI Epigenetics 455 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 21 ISP; BBS/E/B/000C0523), and KM acknowledges funding from the BBSRC (BB/W509917/1). The 456 funders had no role in the preparation of this manuscript. 457 Author contributions: Conceptualization - A.R.R., V.V.S., H.G.B., and A.H; Investigation & 458 Formal analysis - A.R.R., K.M., V.V.S., H.G.B., N.J.P., J.H., A.H; Computational Pipeline 459 development - A.R.R.; Manuscript writing (initial draft) - A.R.R. and A.H.; Manuscript editing - all 460 authors. 461 Figures: 462 463 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 22 464 465 Fig. 1: Meiotic axis extends to the telomere -associated sequences. ( a) Schematic 466 representation of the distinct regions present at chromosome ends in Saccharomyces cerevisiae. 467 Black dot represents the centromere. (b) Mean fold enrichment of Rec8, Red1, and Hop1 relative 468 to their distance from telomeres, using published ChIP -seq datasets from WT cells at early 469 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 23 prophase I (3 hours) 14. (c) Bootstrapped distributions of fold enrichment within thirty -two 20 kb 470 bins spanning the genome. Median values and two-sided 95% confidence intervals are shown as 471 black horizontal lines, with orange/red circles highlighting the mean enrichment in the last 20 kb 472 of chromosome ends. (d, e) Meta-plots of average Rec8, Red1, and Hop1 enrichment at 473 chromosome end-associated X and Y’ elements. Vertical dotted lines mark the positions of the X 474 or Y’ element. For X elements, regions of equal size adjacent to the X element on both the 475 centromere-proximal (Cen) and telomere -proximal (Tel) sides are shown. For Y’ elements, 476 adjacent regions equal to half the size of the Y’ element are shown. Genome averages are 477 denoted by gray horizontal dashed lines, and colored solid lines with shaded areas represent the 478 mean and two-sided 95% confidence intervals for each protein. (f) Bar plots comparing Rec8, 479 Hop1, and Red1 levels on X elements in X-only and XY’ ends. Normality of the data was assessed 480 using the Shapiro-Wilk test. Significance was determined using Student’s two-sided t-test (*p ≤ 481 0.05) (raw p-values: 0.0186, 0.0058, 0.0079), with Benjamini-Hochberg p-value adjustment (BH 482 corrected p-values: 0.0186, 0.0118, 0.0118). Effect sizes were estimated using Cohen’s d: Rec8 483 (d = 1.62), Hop1 (d = 1.13), and Red1 (d = 0.89). All the data presented here are averages of two 484 independent biological replicates, and the results were reproducible between replicates. 485 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 24 486 Fig. 2: Multiple cis-acting features contribute to axis protein depletion near telomeres. (a) 487 Meta-plots of Rec8, Hop1, and Red1 ChIP enrichment (ChIP/Input) at XY’ and X -only 488 chromosome ends, aligned by their distance from telomeres. The genome -wide average 489 enrichment is indicated by a dashed gray line. Data are derived from published ChIP-seq datasets 490 of WT cells at early prophase I (3 hours)14,31,34. (b) Analysis of Red1 binding in SK1/S288C hybrids 491 containing unfused (WT) or homozygous fusion (chrIV -chrI) chromosomes, using published 492 datasets37. Dashed lines represent signal traces, and solid -colored lines represent loess -493 normalized regressions. The black dot marks the position of CEN1. (c) Coding density as a 494 function of distance from telomeres, averaged across chromosome ends per bin. Blue dots 495 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 25 indicate mean coding density per bin, and the black line connects these means. Coding density 496 is calculated as the fraction of DNA encoding open reading frames. All the data presented here 497 are averages of two independent biological replicates, and the results were reproducible between 498 replicates. 499 500 501 502 Fig. 3: Differential recruitment of Red1 at chromosome ends by Rec8 -dependent and 503 independent pathways. ( a) Meta-plots of average Red1 enrichment (spike -in normalized) at 504 various distances from telomeres in WT, rec8, hop1-phd, and hop1-phd rec8 strains during early 505 prophase I (3 hours). Data are derived from spike -in normalized SNP ChIP-seq datasets15. The 506 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 26 colored dashed lines represent the genome average for each strain after spike-in normalization. 507 (b) Bootstrapped distributions of Red1 enrichment within thirty-two 20 kb bins across the genome, 508 with medians and two-sided 95% confidence intervals indicated by black lines. Mean enrichment 509 in the last 20 kb of chromosome ends is highlighted with orange/red circles . (c, d) Meta-plots of 510 average Red1 enrichment on X and Y’ elements at chromosome ends. Vertical dotted lines mark 511 the positions of X and Y’ elements. Adjacent regions —of the same size for X elements or half -512 sized for Y’ elements —are included both telomere (Tel) and centromere (Cen) proximal to the 513 elements. Colored solid lines represent mean enrichment, with shaded areas depicting two-sided 514 95% confidence intervals for each strain. The blue arrow denotes the transcription direction of the 515 Y’ ORF. All the data presented here are averages of two independent biological replicates , and 516 the results were reproducible between replicates. 517 518 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 27 519 Fig. 4: Dot1 regulates axis protein distribution at chromosome ends and interior regions. 520 (a) Scatter plot comparing Red1 and Rec8 levels at individual Rec8 peaks across three genomic 521 regions: pericentromeric regions (±10 kb from centromeres), non-pericentromeric/non-telomeric 522 regions, and chromosome ends (last 20 kb). Genome -wide trend lines illustrate the overall 523 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 28 correlation between Red1 and Rec8 (slope of linear regression trendline = 1.34). Data are derived 524 from published WT ChIP -seq datasets14. (b) Mean histone acetylation and methylation marks 525 (H4K44ac, H3K56ac, H3K4me3, and H3K79me3), normalized to H4 or H3, plotted as a function 526 of distance from chromosome ends using published datasets as well as experimental 527 datasets42,44. (c) Spike-in normalized Red1 levels in WT, dot1Δ, and set1Δ mutants, plotted as a 528 function of distance from chromosome ends using published datasets 34. (d) Boxplots of mean 529 Red1 levels per chromosome end (20 kb) in WT, dot1Δ, and set1Δ mutants. Circles represent 530 mean signals for individual chromosome ends, with lines connecting matched chromosome ends 531 across strains. Normality of the data was assessed using the Shapiro -Wilk test. Student’s two-532 sided t-test **p ≤ 0.01, ns – non significant (raw p-values: 0.001891863, 0.181274230), Benjamini-533 Hochberg p -value adjusted (BH corrected p -values: 0.003783726, 0.181274230). Effect sizes 534 were estimated using Cohen’s d: WT vs. dot1Δ (d = -0.82); WT vs. set1Δ (d = 0.34). (e, f) 535 Bootstrapped distributions of Red1 enrichment within (e) thirty-two 20 kb bins and (f) sixteen 20 536 kb bins, spanning the genome. Median values and two-sided 95% confidence intervals are shown 537 as black horizontal lines. Orange/red circles in (e) indicate mean enrichment at the last 20 kb of 538 chromosome ends, and blue circles in (f) highlight mean enrichment at ±10 kb around 539 centromeres. (g, h) Meta-plots of Red1 enrichment on X and Y’ elements. Vertical dotted lines 540 indicate the positions of X and Y’ elements. Adjacent regions, of the same size for X elements or 541 half-sized for Y’ elements, are shown telomere (Tel) and centromere (Cen) proximal. Colored 542 solid lines represent mean enrichment, with shaded areas showing two-sided 95% confidence 543 intervals for each strain. The blue arrow denotes the transcription direction of the Y’ ORF. (i) Meta-544 analysis of Red1 distribution in WT and dot1Δ mutants, separated into intergenic (top panel) and 545 genic regions (bottom panel), plotted as a function of distance from chromosome ends using 546 spike-in normalized values. All the data presented here are averages of two independent 547 biological replicates, and the results were reproducible between replicates. 548 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 29 549 Fig. 5: Sir3 spreads heterogeneously from the X element during meiotic prophase I. (a, b) 550 Meta-plots of Sir3 ChIP enrichment in wild-type (WT) strains at X elements and Y’ elements during 551 mid-log phase, meiotic induction (time = 0 hours), and early prophase I (time = 3 hours). Vertical 552 dotted lines indicate the positions of X and Y’ elements. Adjacent regions —of the same size for 553 X elements or half-sized for Y’ elements—are shown both telomere (Tel) and centromere (Cen) 554 proximal. (c) Bar plots comparing Sir3 enrichment at X elements in X-only and XY’ chromosome 555 ends during mid -log phase, meiotic induction (time = 0 hours), and early prophase I (time = 3 556 hours). Normality of the data was assessed using the Shapiro -Wilk test. Student’s two-sided t-557 test **p ≤ 0.01 (raw p-values: 0.007263657, 0.007243783, 0.001837521), Benjamini-Hochberg p-558 value adjusted (BH corrected p-values: 0.007263657, 0.007263657, 0.005512564). Effect sizes 559 were estimated using Cohen’s d: Sir3 mid-log phase (d = 1.22); meiotic induction (d = 1.32); early 560 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 30 prophase I (d = 1.11). (d, e) Sir3 binding profiles on chrVII -L (d) and chrVI-L (e) during mid-log 561 phase and early meiotic prophase I (time = 3 hours). ChrVII-L shows detectable spreading of Sir3 562 from the X element into adjacent gene -poor regions, while no detectable spreading is observed 563 on chrVI -L. ORFs are annotated in blue. All the data presented here are averages of two 564 independent biological replicates, and the results were reproducible between replicates. 565 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 31 566 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 32 Fig. 6: Sir proteins protect X elements and regions of spread from meiotic DSBs. (a) Scatter 567 plot of gene expression fold change ( sir3/WT, measured in ndt80Δ background) versus Sir3 568 binding (log10 ChIP/Input) in promoter regions for 5 kb genes (yellow) and non-5 kb genes (blue) 569 at 3 hours post-meiotic induction. Pearson correlation coefficients (r) were calculated in R using 570 the cor() function (Pearson method) and serve as the measure of effect size. p -values were 571 computed using Fisher Z-transformation (two-sided test) to enable statistical testing of correlation 572 significance. Outliers (top and bottom 1% quantiles for 5 kb genes and three non -5 kb gene 573 outliers) were excluded. (b) Boxplots of log10-transformed Spo11-oligo signals (hits per million) 574 at X and Y’ elements in WT and s ir2Δ strains. Data are derived from published Spo11 -oligo 575 datasets29,50. Circles represent total scores for individual elements, with lines connecting matched 576 elements across strains. Normality of the data was assessed using the Shapiro -Wilk test. Since 577 the data did not meet normality assumptions, the Wilcoxon rank -sum test was used to compare 578 groups. Wilcoxon rank -sum test (two-sided) **p ≤ 0.01; ns - not significant (raw p -values: 579 0.00179054, 0.15670080), with Benjamini-Hochberg p-value adjustment (BH adjusted p-values: 580 0.003581081, 0.156700804). Effect sizes were estimated using Rank-Biserial Correlation (RBC) 581 (r = -0.707, -0.408). (c) Scatter plot of Spo11-oligo fold change (sir2Δ/WT) in promoter regions as 582 a function of average Sir3 binding (log10 ChIP/Input) in the same promoter regions. Yellow points 583 represent genes within 5 kb of X elements, and blue points represent non -5 kb genes. Pearson 584 correlation coefficients (r) serve as the measure of effect size, and p-values were computed as in 585 (a). Outliers (top and bottom 1% quantiles for 5 kb genes and three outliers from non-5 kb genes) 586 were excluded. (d) Scatter plot of gene expression fold change ( sir3/WT, measured in ndt80Δ 587 strains) as a function of Spo11-oligo fold change (sir2Δ/WT) in promoter regions for genes within 588 5 kb of X elements (yellow) and non-5 kb genes (blue). Pearson correlation coefficients (r) serve 589 as the measure of effect size, and p-values were computed as in (a). (e) TrAEL-seq signal tracks 590 for chromosome V -R subtelomeric regions in the indicated strains. Blue boxes represent 591 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 33 annotated ORFs. The highlighted region (gray) marks a peak present in sir3 dmc1Δ and sir3 592 dot1Δ dmc1Δ but absent in dmc1Δ and dot1Δ dmc1Δ. (f) Mean hotspot counts per 5 kb bin as a 593 function of distance from X elements for the indicated strains. Hotspots were identified using 594 peaks called as significant from TrAEL -seq data. Error bars represent the standard error of the 595 mean. Normality of the data was assessed using the Shapiro -Wilk test. Since the data did not 596 meet normality assumptions, the Wilcoxon rank-sum test was used to compare groups. Statistical 597 significance was assessed using a Kruskal -Wallis two-sided test to compare hotspot counts 598 across bins. *p ≤ 0.05; ns - not significant (Raw p-values: 0.04066, 0.8046, 0.9805, 0.9118). All 599 the data presented here are averages of two independent biological replicates , and the results 600 were reproducible between replicates. 601 602 603 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 34 Fig. 7: Model showing distinct mechanisms of DSB regulation at chromosome ends by Sir3 604 and Dot1. Sir proteins restrict promoter accessibility at chromosome ends, thereby influencing 605 DSB formation in these regions. In contrast, Dot1 prevents Red1 accumulation at chromosome 606 ends by directing it to chromosome interiors . Whether the decreased Red1 levels contribute to 607 lower DSB levels or increased sister repair at chromosome ends similar to the rest of the genome 608 remains unresolved (dashed line with a question mark). Orange shading represents 609 coding/hotspot density. 610 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 35

611 1. Mozzachiodi, S. & Liti, G. Evolution of yeast hybrids by aborted meiosis. Current Opinion 612 in Genetics and Development 77, 101980-101980 (2022). 613 2. Börner, G.V., Hochwagen, A. & MacQueen, A.J. Meiosis in budding yeast. Genetics 225, 614 1-33 (2023). 615 3. Duro, E. & Marston, A.L. From equator to pole: Splitting chromosomes in mitosis and 616 meiosis. Genes and Development 29, 109-122 (2015). 617 4. Hunter, N. Meiotic recombination: The essence of heredity. Cold Spring Harbor 618 Perspectives in Biology 7(2015). 619 5. Arter, M. & Keeney, S. Divergence and conservation of the meiotic recombination 620 machinery. Nature Reviews Genetics 25, 309-325 (2024). 621 6. Lichten, M. Meiotic chromatin: The substrate for recombination initiation. Genome 622 Dynamics and Stability 3, 165-193 (2008). 623 7. Raghavan, A.R. & Hochwagen, A. Keeping it safe: control of meiotic chromosome 624 breakage. Trends in Genetics (2024). 625 8. Ito, M. & Shinohara, A. Chromosome architecture and homologous recombination in 626 meiosis. Front Cell Dev Biol 10, 1097446 (2022). 627 9. Mohibullah, N. & Keeney, S. Numerical and spatial patterning of yeast meiotic DNA 628 breaks by Tel1. Genome Research 27(2017). 629 10. Pan, J. et al. A hierarchical combination of factors shapes the genome-wide topography 630 of yeast meiotic recombination initiation. Cell 144, 719-731 (2011). 631 11. Panizza, S. et al. Spo11-accessory proteins link double-strand break sites to the 632 chromosome axis in early meiotic recombination. Cell 146, 372-383 (2011). 633 12. Schwacha, A. & Kleckner, N. Interhomolog Bias during Meiotic Recombination: Meiotic 634 Functions Promote a Highly Differentiated Interhomolog-Only Pathway. in Cell Vol. 90 635 1123-1135 (Keeney and Kleckner, 1997). 636 13. Hollingsworth, P.L. Making it Through Parenting. Gifted Child Today Magazine 13, 2-7 637 (1990). 638 14. Sun, X. et al. Transcription dynamically patterns the meiotic chromosome-axis interface. 639 eLife 4, 1-23 (2015). 640 15. Heldrich, J. et al. Two pathways drive meiotic chromosome axis assembly in 641 Saccharomyces cerevisiae. Nucleic Acids Research 50, 4545-4556 (2022). 642 16. Joshi, N., Barot, A., Jamison, C. & Börner, G.V. Pch2 links chromosome axis remodeling 643 at future crossover sites and crossover distribution during yeast meiosis. PLoS Genet 5, 644 e1000557 (2009). 645 17. Milano, C.R. et al. Chromatin binding by HORMAD proteins regulates meiotic 646 recombination initiation. EMBO Journal 43, 836-867 (2024). 647 18. Blitzblau, H.G., Bell, G.W., Rodriguez, J., Bell, S.P. & Hochwagen, A. Mapping of Meiotic 648 Single-Stranded DNA Reveals Double-Strand-Break Hotspots near Centromeres and 649 Telomeres. Current Biology 17, 2003-2012 (2007). 650 19. Sasaki, M., Lange, J. & Keeney, S. Genome destabilization by homologous recombination 651 in the germ line. Nature Reviews Molecular Cell Biology 11, 182-195 (2010). 652 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 36 20. Ross, L.O., Maxfield, R. & Dawson, D. Exchanges are not equally able to enhance meiotic 653 chromosome segregation in yeast. Proceedings of the National Academy of Sciences of 654 the United States of America 93(1996). 655 21. Oliver, T.R. et al. New insights into human nondisjunction of chromosome 21 in oocytes. 656 PLoS Genet 4, e1000033 (2008). 657 22. Ghosh, S., Feingold, E. & Dey, S.K. Etiology of Down syndrome: Evidence for consistent 658 association among altered meiotic recombination, nondisjunction, and maternal age 659 across populations. Am J Med Genet A 149a, 1415-20 (2009). 660 23. Pryde, F.E., Gorham, H.C. & Louis, E.J. Chromosome ends: all the same under their caps. 661 Curr Opin Genet Dev 7, 822-8 (1997). 662 24. Louis, E.J. The chromosome ends of Saccharomyces cerevisiae. Yeast 11, 1553-1573 663 (1995). 664 25. Hocher, A. & Taddei, A. Subtelomeres as Specialized Chromatin Domains. BioEssays 665 42(2020). 666 26. Fabre, E. & Spichal, M. Subnuclear Architecture of Telomeres and Subtelomeres in Yeast. 667 in Subtelomeres (eds. Louis, E.J. & Becker, M.M.) 13-37 (Springer Berlin Heidelberg, 668 Berlin, Heidelberg, 2014). 669 27. Brothers, M. & Rine, J. Distinguishing between recruitment and spread of silent 670 chromatin structures in Saccharomyces cerevisiae. eLife 11, 1-23 (2022). 671 28. Hocher, A. et al. Expanding heterochromatin reveals discrete subtelomeric domains 672 delimited by chromatin landscape transitions. Genome Research 28, 1852-1866 (2018). 673 29. Buhler, C., Borde, V. & Lichten, M. Mapping meiotic single-strand DNA reveals a new 674 landscape of DNA double-strand breaks in Saccharomyces cerevisiae. PLoS Biology 5, 675 2797-2808 (2007). 676 30. Murakami, H. et al. Multilayered mechanisms ensure that short chromosomes 677 recombine in meiosis. Nature (2020). 678 31. Subramanian, V.V. et al. Persistent DNA-break potential near telomeres increases 679 initiation of meiotic recombination on short chromosomes. Nature Communications 10, 680 1-15 (2019). 681 32. Yue, J.X. et al. Contrasting evolutionary genome dynamics between domesticated and 682 wild yeasts. Nature Genetics 49, 913-924 (2017). 683 33. D'Angiolo, M. et al. Telomeres are shorter in wild Saccharomyces cerevisiae isolates than 684 in domesticated ones. Genetics 223(2023). 685 34. Vale-Silva, L.A., Markowitz, T.E. & Hochwagen, A. SNP-ChIP: A versatile and tag-free 686

to quantify changes in protein binding across the genome. BMC Genomics 20, 1-687 10 (2019). 688 35. Kwapisz, M. & Morillon, A. Subtelomeric Transcription and its Regulation. Journal of 689 molecular biology 432, 4199-4219 (2020). 690 36. Wellinger, R.J. & Zakian, V.A. Everything you ever wanted to know about Saccharomyces 691 cerevisiae telomeres: Beginning to end. Genetics 191, 1073-1105 (2012). 692 37. Luo, J. et al. Synthetic chromosome fusion: Effects on mitotic and meiotic genome 693 structure and function. Cell Genomics 3(2023). 694 38. Glynn, E.F. et al. Genome-wide mapping of the cohesin complex in the yeast 695 Saccharomyces cerevisiae. PLoS Biology 2(2004). 696 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 37 39. Ocampo-Hafalla, M.T. & Uhlmann, F. Cohesin loading and sliding. Journal of Cell Science 697 124, 685-691 (2011). 698 40. Bausch, C. et al. Transcription Alters Chromosomal Locations of Cohesin in 699 Saccharomyces cerevisiae. Molecular and Cellular Biology 27, 8522-8532 (2007). 700 41. Borde, V. et al. Histone H3 lysine 4 trimethylation marks meiotic recombination 701 initiation sites. EMBO Journal 28, 99-111 (2009). 702 42. Hu, J. et al. H4K44 Acetylation Facilitates Chromatin Accessibility during Meiosis. Cell 703 Reports 13, 1772-1780 (2015). 704 43. Gothwal, S.K. et al. The double-strand break landscape of meiotic chromosomes is 705 shaped by the Paf1 transcription elongation complex in Saccharomyces cerevisiae. 706 Genetics 202, 497-512 (2016). 707 44. Zhu, X. & Keeney, S. High-resolution global analysis of the influences of Bas1 and Ino4 708 transcription factors on meiotic DNA break distributions in saccharomyces cerevisiae. 709 Genetics 201, 525-542 (2015). 710 45. Bani Ismail, M., Shinohara, M. & Shinohara, A. Dot1-dependent histone H3K79 711 methylation promotes the formation of meiotic double-strand breaks in the absence of 712 histone H3K4 methylation in budding yeast. PLoS ONE 9(2014). 713 46. Takahashi, Y.-H. et al. Dot1 and Histone H3K79 Methylation in Natural Telomeric and 714 HM Silencing. Molecular Cell 42, 118-126 (2011). 715 47. Shilatifard, A. Chromatin modifications by methylation and ubiquitination: Implications 716 in the regulation of gene expression. Annual Review of Biochemistry 75, 243-269 (2006). 717 48. San-Segundo, P.A. & Roeder, G.S. Role for the Silencing Protein Dot1 in Meiotic 718 Checkpoint Control. in Molecular Biology of the Cell Vol. 11 3601-3615 (2000). 719 49. Hoppe, G.J. et al. Steps in Assembly of Silent Chromatin in Yeast: Sir3-Independent 720 Binding of a Sir2/Sir4 Complex to Silencers and Role for Sir2-Dependent Deacetylation. 721 Molecular and Cellular Biology 22, 4167-4180 (2002). 722 50. Rusche, L.N., Kirchmaier, A.L. & Rine, J. The Establishment, Inheritance, and Function of 723 Silenced Chromatin in Saccharomyces cerevisiae. Annual Review of Biochemistry 72, 724 481-516 (2003). 725 51. Pryde, F.E. & Louis, E.J. Limitations of silencing at native yeast telomeres. EMBO Journal 726 18, 2538-2550 (1999). 727 52. Thacker, D., Mohibullah, N., Zhu, X. & Keeney, S. Homologue engagement controls 728 meiotic DNA break number and distribution. Nature 510, 241-246 (2014). 729 53. Kara, N., Krueger, F., Rugg-Gunn, P. & Houseley, J. Genome-wide analysis of DNA 730 replication and DNA double-strand breaks using TrAEL-seq. PLoS Biology 19(2021). 731 54. Bishop, D.K., Park, D., Xu, L. & Kleckner, N. DMC1: A meiosis-specific yeast homolog of E. 732 coli recA required for recombination, synaptonemal complex formation, and cell cycle 733 progression. Cell 69, 439-456 (1992). 734 55. Vader, G. et al. Protection of repetitive DNA borders from self-induced meiotic 735 instability. Nature 477, 115-121 (2011). 736 56. Fourel, G., Revardel, E., Koering, C.E. & Gilson, É. Cohabitation of insulators and silencing 737 elements in yeast subtelomeric regions. EMBO Journal 18, 2522-2537 (1999). 738 57. Radman-Livaja, M. et al. Dynamics of Sir3 spreading in budding yeast: Secondary 739 recruitment sites and euchromatic localization. EMBO Journal 30, 1012-1026 (2011). 740 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint 38 58. Fourel, G. et al. An activation-independent role of transcription factors in insulator 741 function. in EMBO reports Vol. 2 124-132 (2001). 742 59. Saxton, D.S. & Rine, J. Nucleosome Positioning Regulates the Establishment, Stability, 743 and Inheritance of Heterochromatin in Saccharomyces cerevisiae. PNAS 117, 27493-744 27501 (2004). 745 60. Ichikawa, Y., Morohashi, N., Nishimura, Y., Kurumizaka, H. & Shimizu, M. Telomeric 746 repeats act as nucleosome-disfavouring sequences in vivo. Nucleic Acids Research 42, 747 1541-1552 (2014). 748 61. Markowitz, T.E. et al. Reduced dosage of the chromosome axis factor Red1 selectively 749 disrupts the meiotic recombination checkpoint in Saccharomyces cerevisiae. PLoS 750 Genetics 13(2017). 751 62. Su, Y., Barton, A.B. & Kaback, D.B. Decreased meiotic reciprocal recombination in 752 subtelomeric regions in Saccharomyces cerevisiae. Chromosoma 109, 467-475 (2000). 753 63. Lamb, N.E., Yu, K., Shaffer, J., Feingold, E. & Sherman, S.L. Association between maternal 754 age and meiotic recombination for trisomy 21. American Journal of Human Genetics 76, 755 91-99 (2005). 756 64. Sherman, S.L., Freeman, S.B., Allen, E.G. & Lamb, N.E. Risk factors for nondisjunction of 757 trisomy 21. Cytogenetic and Genome Research 111, 273-280 (2005). 758 65. Humphryes, N. & Hochwagen, A. A non-sister act: Recombination template choice 759 during meiosis. Experimental Cell Research 329, 53-60 (2014). 760 66. Medhi, D., Goldman, A.S. & Lichten, M. Local chromosome context is a major 761 determinant of crossover pathway biochemistry during budding yeast meiosis. (2016). 762 67. Scherthan, H. A bouquet makes ends meet. Nature Reviews Molecular Cell Biology 2, 763 621-627 (2001). 764 68. Blitzblau, H.G. & Hochwagen, A. ATR/Mec1 prevents lethal meiotic recombination 765 initiation on partially replicated chromosomes in budding yeast. eLife 2, 1-20 (2013). 766 69. Langmead, B. & Salzberg, S.L. Fast gapped-read alignment with Bowtie 2. Nature 767

9, 357-359 (2012). 768 70. Kar, F.M., Vogel, C. & Hochwagen, A. Meiotic DNA breaks activate a streamlined 769 phospho-signaling response that largely avoids protein-level changes. Life Science 770 Alliance 5, 1-15 (2022). 771 71. Ewels, P.A.P.A.F.S. & et al. The nf-core framework for community-curated bioinformatics 772 pipelines. Nature Biotechnology 38, 271-271 (2020). 773 72. Patel H, E.P.P.A.H.n.R.B.O.S.G.M.D.V.P.M.S.L. & et al. nf-core/rnaseq: Nf-core/ rnaseq 774 v3.1 - Lead Alligator (3.1). (2021). 775 73. Patro, R., Duggal, G., Love, M.I., Irizarry, R.A. & Kingsford, C. Salmon provides fast and 776 bias-aware quantification of transcript expression. Nature Methods 14, 417-419 (2017). 777 74. Heldrich, J., Sun, X., Vale-Silva, L.A., Markowitz, T.E. & Hochwagen, A. Topoisomerases 778 Modulate the Timing of Meiotic DNA Breakage and Chromosome Morphogenesis in 779 Saccharomyces cerevisiae. Genetics 215, 59-73 (2020). 780 781 .CC-BY-NC-ND 4.0 International licenseavailable under a (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprintthis version posted March 3, 2025. ; https://doi.org/10.1101/2025.02.27.640173doi: bioRxiv preprint