Eco1-dependent cohesin acetylation anchors chromatin loops and cohesion to define functional meiotic chromosome domains

Cohesin organizes the genome by forming intra-chromosomal loops and inter-sister chromatid linkages. During gamete formation by meiosis, chromosomes are reshaped to support crossover recombination and two consecutive rounds of chromosome segregation. Here, we show that meiotic chromosomes are organized into functional domains by Eco1 acetyltransferase-dependent positioning of both chromatin loops and sister chromatid cohesion in budding yeast. Eco1 acetylates the Smc3 cohesin subunit in meiotic S phase to establish chromatin boundaries, independently of DNA replication. Boundary formation by Eco1 is critical for prophase exit and for the maintenance of cohesion until meiosis II, but is independent of the ability of Eco1 to antagonize the cohesin release factor, Wpl1. Conversely, prevention of cohesin release by Wpl1 is essential for centromeric cohesion, kinetochore mono-orientation, and co-segregation of sister chromatids in meiosis I. Our findings establish Eco1 as a key determinant of chromatin boundaries and cohesion positioning, revealing how local chromosome structuring directs genome transmission into gametes.


Introduction
The cohesin complex defines genome architecture to support DNA repair, gene expression and chromosome segregation (Davidson and Peters, 2021). Core cohesin is a DNA translocase comprising a V-shaped heterodimer of two structural maintenance of chromosomes proteins, Smc1 and Smc3, whose two ATPase heads are connected by a kleisin subunit. Cohesin folds the genome through ATPdependent extrusion of intra-molecular DNA loops. Cohesin also entraps newly replicated sister chromatids within its tripartite structure, to establish the cohesion needed for chromosome segregation. Loop extrusion and cohesion are biochemically distinct and dependent on cohesin accessory proteins, although the mechanisms are not completely understood (Srinivasan et al., 2018). Loading of cohesin onto DNA requires the Scc2-Scc4 (NIPBL-MAU2 in mammals) complex, which also drives loop extrusion (Ciosk et al., 2000;Davidson et al., 2019;Petela et al., 2018;Srinivasan et al., 2019). Chromosomal cohesin is destabilized by the cohesin release factor, Wpl1/Rad61 (WAPL in mammals), an activity that is counteracted by acetylation of cohesin's Smc3 subunit by the Eco1 acetyltransferase (Ben-Shahar et al., 2008;Unal et al., 2008).
Cohesin-mediated loops are positioned by boundary elements. In mammalian interphase cells, the insulation protein CTCF anchors cohesin at the sites of longrange interactions (Haarhuis et al., 2017;Rao et al., 2017;Schwarzer et al., 2017;Wutz et al., 2017). In yeast, CTCF is absent but chromosomes are nevertheless organized into cohesin-dependent loops in mitosis and convergent genes, which are known to accumulate cohesin, are found at loop boundaries (Costantino et al., 2020;Lazar-Stefanita et al., 2017;Lengronne et al., 2004;Paldi et al., 2020;Schalbetter et al., 2017). Budding yeast pericentromeres provide an exemplary model of how a functional chromosome domain is folded. Cohesin loaded at centromeres extrudes loops on both sides until it is stalled by convergent genes at flanking pericentromere boundaries, thereby establishing a structure that facilitates chromosome segregation in mitosis . Loop size and position are also controlled by cohesin regulators. Wpl1 restricts loop size, but does not affect their positioning (Costantino et al., 2020;Dauban et al., 2020;Haarhuis et al., 2017). Eco1 also limits long-range interactions, but additionally affects loop positioning (Dauban et al., 2020). Whether this is a direct effect, through Eco1-mediated acetylation and inhibition of loop-extruding cohesin, or indirect, as a result of acetylated cohesive cohesin forming a barrier to translocating cohesin, is unclear. Cells lacking Eco1 are inviable due to cohesion defects, which can be rescued by removal of Wpl1 to restore viability (Ben-Shahar et al., 2008;Rowland et al., 2009;Sutani et al., 2009;Unal et al., 2008).
However, cells lacking both Wpl1 and Eco1 show an additive increase in long-range interactions (Dauban et al., 2020). Together, these observations suggested that loop positioning is not essential for viability. Consistent with its essential function in establishing cohesion, in mitotically growing cells acetylation of Smc3 by Eco1 is coupled with DNA replication (Beckouët et al., 2010;Ben-Shahar et al., 2008). In mammals, ESCO2 similarly acetylates SMC3 during S phase to establish cohesion (Alomer et al., 2017) while, during interphase, an additional family member, ESCO1, is active and contributes to boundary formation in chromatin looping (Wutz et al., 2020). Together, these observations indicate that Eco1-dependent cohesin acetylation can both position loops and maintain cohesion.
During gamete formation by meiosis, chromosomes undergo extensive restructuring, underpinned by cohesin-dependent chromosome looping and cohesion.
In many organisms a meiosis-specific kleisin, Rec8, enables functions that cannot be carried out by canonical kleisin, Scc1/Rad21 (Severson et al., 2009;Tachibana-Konwalski et al., 2010;Toth et al., 2000;Yokobayashi et al., 2003). During meiotic prophase, chromosomes comprise a dense array of chromatin loops emanating from cohesin-rich axes that are zipped together in homologous pairs by a central corethe synaptonemal complex (Cahoon and Hawley, 2016). Budding yeast cohesin anchors loops at their base (Muller et al., 2018;Schalbetter et al., 2019) and supports crossover recombination and DNA repair to allow prophase exit (Klein et al., 1999). Between prophase I and metaphase I, Wpl1 removes a fraction of chromosomal Rec8 (Challa et al., 2016, 2019. WAPL also promotes meiotic release of cohesin in Arabidopsis thaliana and Caenorhabditis elegans, and reduces loop number in mouse oocytes, although the kleisin target differs between organisms. (Crawley et al., 2016;De et al., 2014;Silva et al., 2020). Following prophase exit, two distinct meiotic divisions ensue (reviewed in (Duro and Marston, 2015)). During meiosis I, sister kinetochores are monooriented to ensure sister chromatid cosegregation. In budding yeast, the monopolin complex fuses sister kinetochores, while in fission yeast and mammals centromeric Rec8-cohesin directs sister chromatid co-segregation (Chelysheva et al., 2005;Monje-Casas et al., 2007;Parra et al., 2004;Sakuno et al., 2009;Severson et al., 2009). Homolog segregation at meiosis I is triggered by separase-dependent cleavage of Rec8 on chromosome arms, while pericentromeric cohesin is retained and cleaved only at meiosis II to allow sister chromatid segregation. How cohesin-dependent loop formation and cohesion are spatially and temporally regulated to establish distinct functional chromosome domains in meiosis remains unclear.
Here we identify Eco1 acetyltransferase as a key determinant of localized meiotic chromosome structure in budding yeast. In meiosis, Eco1 acetylates Smc3 independently of DNA replication and is critical for viability, even in the absence of Wpl1. Eco1 counteracts Wpl1 to allow centromeric cohesion establishment and thereby kinetochore monoorientation. In contrast, arm cohesion and prophase exit all require an Eco1 function other than Wpl1 antagonism. While Eco1 and Wpl1 independently restrict chromatin loop size in prophase I, only Eco1 is critical for boundary formation, notably at pericentromere borders. We propose that cohesin acetylation by Eco1 traps both loop extruding and cohesive cohesin complexes at boundaries to define a chromosome architecture that is essential for meiotic recombination and chromosome segregation.

Eco1 acetylates cohesin during meiotic S phase, independently of DNA replication
To determine the timing of Eco1-dependent Smc3 acetylation during meiosis, wild type cells carrying functional ECO1-6HIS-3FLAG (Figure S1A), were released from a pre-meiotic S phase block (Berchowitz et al., 2013) allowing synchronous DNA replication and nuclear divisions ( Figure 1A and B). Eco1 levels increased after meiotic entry, were maximal during meiotic S phase and declined around the time of prophase exit, while Smc3-K112,K113 acetylation (henceforth Smc3-Ac), detected using a verified antibody ( Figure S1B and C), accumulated after Eco1 appearance, persisted throughout the meiotic divisions and declined during meiosis II ( Figure 1C).
To test whether DNA replication is required for Smc3 acetylation in meiosis, we analyzed pSCC1-CDC6 and clb5D clb6D cells that fail to assemble or fire, prereplicative complexes, respectively, resulting in little or no replication in pre-meiotic S phase ( Figure 1D; (Brar et al., 2009;Stuart and Wittenberg, 1998)). Surprisingly, Smc3-Ac appeared in pSCC1-CDC6 and clb5D clb6D meiotic cells with comparable timing to wild type and was only modestly reduced ( Figure 1E). In contrast, Smc3-Ac was greatly diminished in cells lacking the meiotic cohesin kleisin subunit (rec8D) ( Figure 1E). Because DNA double-strand breaks trigger Eco1-dependent cohesion establishment in mitotic cells (Strom et al., 2007;Unal et al., 2007), we tested whether Spo11 endonuclease-induced meiotic double strand breaks were required for Smc3-Ac. However, preventing double strand break formation (spo11D) did not reduce Smc3-Ac levels whether chromosomes were replicated or not ( Figure S1D and E). We conclude that cohesin acetylation on its Smc3 subunit during meiotic S phase occurs independently of DNA replication and programmed double strand break formation.
Eco1 is essential for meiosis, even in the absence of WPL1 Eco1 is essential for vegetative growth and eco1D viability is restored by deletion of WPL1 (eco1D wpl1D cells) (Ben-Shahar et al., 2008). To examine Eco1 function specifically in meiosis we fused it to the FKBP-rapamycin binding domain (FRB) to allow rapamycin-induced anchoring out of the nucleus (Haruki et al., 2008) ( Figure   1F; see also materials and methods). Growth of ECO1-FRB-GFP cells was inhibited in the presence of rapamycin, but restored by deletion of WPL1 ( Figure 1G), consistent with successful anchor-away (Ben-Shahar et al., 2008;Sutani et al., 2009;Unal et al., 2008). Upon induction of meiosis in the presence of rapamycin, Smc3-Ac was undetectable in ECO1-FRB-GFP cells and full-length Rec8 persisted long after its expected time of degradation ( Figure 1H). Furthermore, only a minor fraction of ECO1-FRB-GFP cells completed the first meiotic division ( Figure 1I). Even in the absence of rapamycin, ECO1-FRB-GFP cells showed defects in meiotic progression, Rec8 degradation and Smc3-Ac ( Figure S1F and G), despite supporting vegetative growth ( Figure 1G). Therefore, the FRB-GFP tag on Eco1 specifically affects meiosis. Henceforth, ECO1-FRB-GFP cells were induced to sporulate in the presence of rapamycin and are denoted eco1-aa. We conclude that Eco1 acetylates Smc3 on residues K112,113 during S phase of meiosis, and that Eco1 is required for efficient meiotic division.
We reasoned that Eco1 activity early in meiosis may be required to counter Wpl1-dependent cohesin destabilization during later stages of meiosis. If this is the case, deletion of Wpl1 may overcome the meiotic arrest of eco1-aa cells. Wpl1 promotes the non-proteolytic removal of cohesin between meiotic prophase and metaphase I (Challa et al., 2016(Challa et al., , 2019. Prior to meiotic S phase, functional Wpl1-6HA gains an activating phosphorylation (Challa et al., 2019) and its overall levels increase (Figure 2A-C; Figure S2A-C). Subsequently, and reminiscent of the sequential loss of cohesin from chromosome arms and pericentromeres, Wpl1 undergoes step-wise degradation during meiosis I and II ( Figure S2B and C). To test the idea that Eco1 allows meiotic progression by countering Wpl1, we sought to measure spore formation and viability ( Figure 2D; Figure S2D-F). However, deletion of WPL1 in the eco1-aa background only slightly increased the formation, but not the viability, of spores, while wpl1D cells showed a small decrease in spore viability, as reported ( (Challa et al., 2016); Figure 2D; Figure S2D,). Consistently, WPL1 deletion did not restore nuclei division to eco1-aa cells ( Figure S2E). Sporulating ECO1-FRB-GFP cells in the absence of rapamycin increased sporulation efficiency, but only slightly improved viability ( Figure S2D-F). Therefore, in contrast to vegetative cells, Eco1 is essential for meiosis, even in the absence of Wpl1.

Distinct requirements for Eco1 in cohesion establishment at centromeres and on chromosome arms
To determine whether eco1-aa cells establish functional cohesion during S phase, we labelled one homolog either at a centromere (CEN5-GFP) or at a chromosomal arm site (LYS2-GFP) and scored the percentage of cells with two GFP foci (indicating defective cohesion) as cells progressed through meiotic S phase into a prophase I arrest. In wild type prophase cells, sister chromatids are tightly cohered and a single focus is visible. In contrast, eco1-aa cells showed a profound cohesion defect ( Figure 2E-H). Remarkably, deletion of WPL1 restored cohesion at the centromere (CEN5-GFP), but not at the chromosomal arm site (LYS2-GFP) ( Figure   2F and H). Deletion of WPL1 alone also caused a modest cohesion defect at LYS2, but not at CEN5 ( Figure 2F and H). Therefore, the critical role of Eco1 in cohesion establishment at centromeres is to counteract Wpl1, while on chromosome arms Eco1 plays an additional, essential function.

Eco1 counteracts Wpl1-dependent cohesin destabilization during meiotic prophase
While Wpl1 promotes cohesin removal from chromosomes between prophase exit and metaphase I (Challa et al., 2019), paradoxically, Wpl1 levels and the slower migrating, presumed active, form decline at prophase exit and are instead highest during meiotic S and prophase ( Figure 2A). Therefore, Wpl1 may need to be counteracted by Eco1 prior to or during meiotic prophase. Calibrated ChIP-Seq revealed a global increase in the levels of the cohesin subunit Smc3 on chromosomes of wpl1D prophase I cells ( Figure 3A and B). Meiotic stage and Smc3 total protein levels were comparable in all conditions ( Figure 3B and C). In contrast, chromosomal Smc3 was reduced in eco1-aa genome-wide, with the notable exception of core centromeres ( Figure 3D), Interestingly, inactivation of Wpl1 increased the chromosomal levels of Smc3 in eco1-aa, particularly at centromeres, where Smc3 levels in eco1-aa wpl1D were comparable to the wpl1D mutant alone ( Figure 3D). Elsewhere, including at pericentromere borders and known chromosomal arm cohesin sites , Smc3 levels in eco1-aa wpl1D were similar to wild type ( Figure 3D). ChIP-Seq of the meiosis-specific kleisin Rec8 in prophase I revealed a similar pattern to that of Smc3 ( Figure S3). Although the different strains cannot be directly compared due lack of a suitable antigen for calibration (see materials and methods), inspection of Rec8 levels in individual strains confirmed that Eco1 is more important for Rec8 association with borders and arm sites than centromeres ( Figure S3). Taken together, our Rec8 and Smc3 ChIP-Seq show that the function of Eco1 at pericentromere borders and chromosome arms in meiotic prophase is two-fold. First, Eco1 protects border and arm cohesin from Wpl1-dependent removal, since cohesin levels at these sites are higher in eco1-aa wpl1D compared to eco1-aa cells. Second, Eco1 has an additional, Wpl1independent function in cohesin retention at border and arm sites, because cohesin levels are higher in wpl1D compared to eco1-aa wpl1D cells. In contrast, at centromeres, Wpl1 removes cohesin, but Eco1 has little influence on cohesin levels.
This implies that specialized cohesin loading and/or anchoring mechanisms at kinetochores (Hinshaw et al., 2017;Paldi et al., 2020) are the main antagonists of the cohesin removal activity of Wpl1 at centromeres, at least in prophase.
Nevertheless, the ability of Wpl1 deletion to rescue the centromeric cohesion detects of eco1-aa cells indicates that Eco1 must counteract Wpl1 at centromeres to allow cohesion establishment during S phase ( Figure 2F).

Eco1 establishes loop boundaries in meiotic prophase chromosomes and restricts long-range chromatin interactions independently from Wpl1
Meiotic prophase chromosomes are highly structured, with ordered chromatin loops emanating from linear protein axes which are zipped together by the synaptonemal complex ( Figure 4A). To understand how Eco1 and Wpl1 define chromosome structure in meiotic prophase, we performed Hi-C 6h after inducing ndt80D cells to sporulate and confirmed consistent DNA content for all conditions ( Figure 4B).
Analysis of contact probability on chromosome arms as a function of genomic distance revealed an increase in long-range interactions in both eco1-aa and wpl1D mutants, which was further exacerbated in eco1-aa wpl1D cells ( Figure 4C). Plotting the slope resulted in a right-ward shift of the curve maxima, which estimates the average size of the loops (Dauban et al., 2020; Figure 4C). This indicated an increase from ~10kb in wild type to ~20kb in eco1-aa or wpl1D cells and up to ~40 kb in eco1-aa wpl1D ( Figure 4C). Heat maps of individual chromosomes corroborated the additive increase in long range interactions in wpl1D eco1-aa double mutants ( Figure 4D). Further inspection revealed that spots and stripes on the Hi-C contact maps, indicative of positioned loops anchored on two or one sides, respectively, were stronger and increased in number in wpl1D, but were more diffuse in eco1-aa, even after Wpl1 deletion ( Figure 4D). This suggests loop boundaries/anchors are strengthened by wpl1D but lost in eco1-aa. Mirrored pile-ups of all 16 wild type pericentromeres centered on the centromeres confirmed the organization of the flanking chromatin into two separate domains, indicating that centromeres act as insulators. In contrast, the absence of active Eco1 reduced insulation across centromeres ( Figure 4E see also top right and bottom left quadrants on the ratio difference maps in Figure S4E). Interestingly, the length and intensity of the Hi-C contact stripe protruding from centromeres progressively increased in eco1-aa and wpl1D eco1-aa cells, compared to wild type ( Figure 4E, arrows; Figure S4E), suggesting that Eco1 limits the extent of loop-extrusion by cohesin complexes anchored at centromeres. Pile-ups centered on all 32 pericentromere borders  revealed strong boundaries in wild type, which increased in intensity in wpl1D cells ( Figure 4F; Figure S4F). In contrast, boundaries at pericentromere borders were barely detectable in eco1-aa and were only partially rescued by deletion of WPL1 ( Figure 4F), suggesting that Eco1 is critical to halt loop extrusion at borders, even in the absence of Wpl1. Note that border pile-ups also display a second centromere-proximal stripe, corresponding to loop extrusion from the centromeres that is increased in intensity in wpl1D, consistent with the centromere pile-ups. Both eco1-aa and eco1-aa wpl1D cells also exhibited reduced insulation at borders, indicating that Eco1 is also important to prevent loop extrusion across borders ( Figure 4F, see also difference maps in Figure S4F). Consistent with the pile-up analysis, examination of individual wild-type pericentromeres revealed the presence of Hi-C spots, indicative of positioned loops, between the centromere and each of the two borders, as marked by the characteristic tripartite Smc3 ChIP-seq signal ( Figure 4G). While stronger Hi-C spots were localized with tripartite Smc3 in wpl1D cells, both features were absent in eco1-aa and weaker in eco1-aa wpl1D cells ( Figure 4G). These data indicate that Wpl1 and Eco1 limit loop expansion through distinct mechanisms in meiotic prophase. While Wpl1 destabilizes loopextruding cohesin to reduce its lifetime, Eco1 anchors cohesin at boundary sites to stabilize and position loops. We further note that Eco1-dependent loop stabilization critically defines the boundaries that demarcate the pericentromeric domain. This is consistent with the requirement of Eco1 to maintain cohesin association with borders and chromosomal arm sites ( Figure 3).

Replication-independent Smc3 acetylation defines meiotic chromosome loops
Our findings show that Eco1 is a key determinant of loop anchors at pericentromere borders and other chromosomal boundaries in meiotic prophase cells. Interestingly, Eco1-dependent Smc3 acetylation also occurs in unreplicated cells, suggesting that loop anchors may form independently of DNA replication and the presence of a sister chromatid ( Figure 1E, Figure S1D). As such, Eco1 could form boundaries by directly acetylating loop extruding cohesin complexes, rather than complexes engaged in cohesion. To test this idea, we generated Hi-C maps of prophase I wildtype and clb5D clb6D cells. Flow cytometry confirmed that, in contrast to wild-type, clb5D clb6D cells failed to undergo bulk pre-meiotic DNA replication ( Figure 5A).
Although chromosome axis proteins assemble apparently normally in clb5D clb6D cells, double strand break formation and, consequently, homolog pairing are defective (Blitzblau et al., 2007;Brar et al., 2009;Smith et al., 2001). Therefore, both inter-sister and inter-homolog interactions are expected to be absent in clb5D clb6D cells, so that any stripes and dots observed on Hi-C maps can be attributed to cislooping along a single chromatid. Consistently, in clb5D clb6D cells, mid-to longrange contacts on individual chromosomes were strongly reduced, though the characteristic stripe and dot pattern indicating the presence of loops was still visible ( Figure 5B). Analysis of contact probability on chromosome arms found that contacts in the 5-100kb range are reduced in clb5D clb6D cells ( Figure 5C). Importantly, however, plotting the derivative revealed that the average loop size in clb5D clb6D was only slightly reduced (~8kb in clb5D clb6D vs 10kb in wild type; Figure 5C). This suggests that the decrease in mid-/long-range interactions might result from the absence of inter-sister and inter-homolog contacts. Together with the observation that clb5D clb6D cells do not display an increase in long range interactions, this indicates that replication and the presence of cohesion do not play a fundamental role in restricting loop extrusion. To confirm the notion that loop boundaries form in unreplicated cells, we examined individual pericentromeres of clb5D clb6D cells. As an example, Figure 5D shows that Hi-C spots and stripes are anchored at the pericentromere borders of chromosome IX in wild type and clb5D clb6D cells, respectively. The presence of Hi-C stripes, rather than spots, in clb5D clb6D cells suggests that cis-looping may be more dynamic than in replicated cells, perhaps due to the absence of sister chromatid cohesion. Centromere and border Hi-C pile-ups for the wild type and clb5D clb6D Hi-C datasets further corroborated these findings ( Figure 5E-H). Interestingly, centromere pile-ups and ratio maps revealed that the contact stripe originating from centromeres was greatly diminished in clb5D clb6D and accompanied by a loss of centromeric insulation ( Figure 5E and F). Although the underlying reasons are unclear, this suggests that Clb5 and Clb6 promote centromere-directed loop extrusion. In contrast, border pile-ups confirmed the presence of boundaries in clb5D clb6D cells, though they were not as strong as in wild type ( Figure 5G and H). Overall, these results indicate that although replication is not required for boundary formation, cohesion and/or the presence of a sister chromatid strengthen these boundaries. Importantly, boundary formation and loop anchoring by pericentromere borders in clb5D clb6D contrasts with eco1-aa cells which show greatly diminished border-anchored loops (compare Figure 5D with Figure 4G and Figure 5H with Figure S4B) and border/centromere contacts (compare Figure 5G with Figure 4F) both on individual pericentromeres or on border pile-ups and pile-up ratio plots. Since robust boundary formation in replicated cells relies on Eco1 (Figure 4), and because Eco1 is proficient in acetylating cohesin also in unreplicated cells ( Figure 1E and Figure S1D), it follows that Eco1 is capable of anchoring loops independently of DNA replication and sister chromatid cohesion.
Moreover, it implies that loop extruding cohesin can be acetylated by Eco1.

Recombination prevents prophase exit in eco1-aa cells
We next sought to understand how boundary formation and cohesion establishment by Eco1 impact meiotic chromosome segregation. Though eco1-aa cells complete bulk DNA replication in meiotic S phase with similar timing to wild type ( Figure 2E and G), only a small fraction of cells undergo nuclear divisions ( Figure 1I). Cohesin is required for meiotic recombination and rec8D cells undergo a recombinationdependent checkpoint arrest in meiotic prophase, due to the persistence of unrepaired double strand breaks (Klein et al., 1999). We found that eco1-aa cells similarly arrest in prophase as judged by a failure to separate spindle pole bodies (SPBs, marked by Spc42-tdTomato) and prophase exit was only modestly advanced by deletion of WPL1 (compare eco1-aa to wpl1D eco1-aa cells; Figure 6A). This indicates that although Eco1 facilitates prophase exit by counteracting Wpl1, other Eco1 functions are also important. To determine whether activation of the recombination checkpoint prevents timely prophase exit in eco1-aa and eco1-aa wpl1D cells, we abolished meiotic double strand break formation (by deletion of SPO11). Figure 6A shows that SPO11 deletion abolished the prophase exit delay of both eco1-aa and eco1-aa wpl1D cells, confirming that Eco1 is required for satisfaction of the recombination checkpoint to allow prophase exit.

Co-segregation of sister chromatids during meiosis I requires Wpl1 antagonism by Eco1
To understand how Eco1/Wpl1-dependent chromosome organization impacts meiotic chromosome segregation, we exploited the ability of spo11D to bypass of the prophase block of eco1-aa cells. Note that Eco1-dependent cohesin acetylation does not require recombination ( Figure S1E). Imaging of live cells carrying a heterozygous centromere label (CEN5-GFP) and Spc42-tdTomato confirmed that deletion of SPO11 permitted meiotic divisions in eco1-aa and eco1-aa wpl1D ( Figure 6B). In wild type cells, sister chromatids co-segregate in meiosis I ("reductional segregation") so that CEN5-GFP foci are inherited by just one of the two nuclei and this was also the case in spo11D, wpl1D and spo11D wpl1D cells, (2 separated SPBs; Figure 6C and D; Figure S5A). However, surprisingly, in ~40% of spo11D eco1-aa cells, CEN5-GFP foci segregated to opposite poles in meiosis I ("equational segregation") indicating loss of sister kinetochore monoorientation ( Figure 6C and D). Remarkably, deletion of WPL1 largely rescued the sister kinetochore monoorientation defect of spo11D eco1-aa cells in meiosis I ( Figure 6C and D), consistent with the restoration of centromere cohesion at prophase in eco1-aa wpl1D compared to eco1-aa cells ( Figure 2F). This supports the idea that Eco1 may enable sister kinetochore monoorientation through effects on cohesion establishment. Therefore, Eco1 antagonism of Wpl1 allows the establishment of centromeric cohesion and enables sister kinetochore monoorientation.

Wpl1 antagonism
During wild type meiosis II, segregation of sister chromatids to opposite poles is ensured by the pool of cohesin that persists on pericentromeres after bulk cohesin cleavage in anaphase I and which likely resides at borders. In wild type cells, this can be visualized by sister CEN5-GFP foci segregation in anaphase II, resulting in their association with two of the four Spc42-tdTomato foci ( Figure S5A). Similarly, spo11D, wpl1D, and spo11D wpl1D cells segregated sister CEN5-GFP foci to opposite poles in meiosis II ( Figure 6C and E; Figure S5A). In spo11D eco1-aa cells, among the cells that enter meiosis II ~30% had already segregated sister CEN5-GFP foci during meiosis I (green bar in Figure 6E), while in a further ~50% of cells, CEN5-GFP were found next to a single Spc42-tdTomato focus (blue bar in Figure   6E), indicating defective meiosis II segregation. Moreover, defective meiosis II segregation in eco1-aa cells is not rescued by deletion of WPL1, whether or not Spo11 is present ( Figure 6E). Therefore, Eco1 is essential for chromosome segregation during meiosis II, even in the absence of Wpl1. One potential explanation for these findings is that localization of the cohesin protector protein, shugoshin (Sgo1) or the meiotic protein Spo13, which is also required for cohesin protection during meiosis I (Galander et al., 2019b(Galander et al., , 2019aKatis et al., 2004;Lee et al., 2004), may require Smc3 acetylation. However, ChIP-qPCR revealed that both Sgo1 and Spo13 localize to chromosomes in eco1-aa cells ( Figure S5B-F). Both proteins follow a similar pattern to Rec8, showing reduced chromosomal association in eco1-aa cells that is rescued by WPL1 deletion ( Figure S5B-F), consistent with a requirement for cohesin for the chromosomal association of Sgo1 and Spo13 (Galander et al., 2019b;Kiburz et al., 2005). Therefore, the failure to build, rather than protect, pericentromeric cohesion is the cause of meiosis II mis-segregation in eco1-aa cells. Since Wpl1 rescues the loss of cohesin on pericentromeric borders in eco1-aa cells, but not the anchoring of loops, it is likely that cohesin anchoring at chromatin boundaries is the critical function of Eco1 in cohesion establishment and meiosis II segregation.

Mutation of Smc3-K112,113 results in meiotic lethality
In mitotically-growing cells, Eco1 protects cohesin from Wpl1 by acetylation of Smc3 residues K112 and K113 (Ben-Shahar et al., 2008;Rowland et al., 2009;Unal et al., 2008). To determine whether Smc3-Ac similarly allows cohesion establishment by protecting from Wpl1 and/or confers chromatin boundary function in meiosis, we analyzed a non-acetylatable smc3-K112,113R mutant. To support mitotic growth, cells carried wild type SMC3 under the CLB2 promoter, which is repressed in meiosis (homozygous pCLB2-3HA-SMC3), and heterozygous smc3-K112,113R or, as a control SMC3, at an ectopic locus expressed from the endogenous promoter ( Figure S6A). Smc3 levels in pCLB2-3HA-SMC3 cells without ectopic expression were largely repressed, though low levels of residual Smc3 were detectable ( Figure   S6B and C). Levels of heterozygously produced Smc3 and Smc3-K112,113R were approximately half that of Smc3 in wild type cells ( Figure S6B and C). Ectopic SMC3 expression rescued the unviability of pCLB2-3HA-SMC3 spores while smc3-K112,113R expression did not, indicating that Smc3 acetylation is essential for meiosis ( Figure S6D). To determine whether Smc3-K112,113R localizes normally to chromosomes and whether it is susceptible to destabilization by Wpl1, we performed calibrated Smc3 ChIP-Seq in prophase I cells where either Smc3 or Smc3-K112,113R were heterozgously produced and in the presence and absence of Wpl1 ( Figure 7A-D). Flow cytometry confirmed similar meiotic progression ( Figure 7A) and western blotting showed that total cellular Smc3 levels were comparable ( Figure 7B).
Therefore, like Eco1, Smc3-Ac is more important for retention of cohesin at pericentromere borders and arm sites than at centromeres, suggesting a role in boundary formation.
Next we asked whether Smc3-Ac underlies the functional effects of Eco1 in meiosis by conferring both cohesion establishment and boundary function. First, we scored sister CEN5-GFP or LYS2-GFP foci separation as cells progressed into a prophase arrest and found that smc3-K112,113R cells exhibited similar cohesion defects to cells depleted of Smc3 ( Figures S6E-J). In both smc3-K112,113R and pCLB2-SMC3 cells, the cohesion defect at centromeres was more modest than that of eco1-aa cells ( Figure 2F), potentially reflecting incomplete meiotic depletion of Smc3 when placed under the CLB2 promoter ( Figure S6B and C). Next, we used live-cell imaging to assay the requirement for Smc3-Ac in prophase exit and meiosis I and II chromosome segregation. Exit from prophase was impaired in pCLB2-3HA-SMC3 and, to a lesser extent, smc3-K112,113R cells, but in both cases was overcome by deletion of SPO11 ( Figure S7A; Figure 7E), indicating that Smc3-Ac is required for satisfaction of the recombination checkpoint, like Eco1. Analysis of sister chromatid segregation in meiosis I in the spo11D background ( Figure 7F; Figure   S7B) revealed that ~10-15% of pCLB2-SMC3 and smc3-K112, 113R cells aberrantly segregated sister chromatids to opposite poles during meiosis I ( Figure 7F, Figure   S7B). In the case of smc3-K112,113R, but not pCLB2-SMC3, this equational meiosis I segregation was rescued by deletion of WPL1 ( Figure 7F). Sister chromatid segregation during meiosis II was also greatly impaired in both pCLB2-SMC3 and smc3-K112,113R cells, but in neither case was it rescued by WPL1 deletion ( Figure   7G and Figure S7C). We note that smc3-K112,113R and pCLB2-SMC3 have a lesser effect on meiosis I sister chromatid segregation and centromeric cohesion as compared to eco1-aa (compare Figure 2F with Figure S6E). In contrast, meiosis II segregation was similarly defective in eco1-aa, pCLB2-SMC3 and smc3-K112,113R cells ( Figure 6E and Figure 7G). The most likely explanation for this difference is that pre-meiotic expression of pCLB2-SMC3 leads to persistence of functional Smc3 ( Figure S6C) which is preferentially loaded at centromeres to partially establish cohesion at this site. However, in both meiosis I and II segregation WPL1 deletion had the same effect on eco1-aa and smc3-K112,113R cells. We conclude that Eco1dependent Smc3-Ac functionally organizes meiotic chromosomes for their segregation. At centromeres, Eco1-dependent Smc3 acetylation counteracts Wpl1 to direct sister kinetochore monoorientation and thereby enforce sister chromatid cosegregation in meiosis I. Eco1 acetylation of Smc3 is also essential for prophase exit and sister chromatid segregation in meiosis II, even in the absence of Wpl1, likely as a result of cohesin anchoring to establish chromatin boundaries.

Cohesin regulators organize functionally distinct chromosomal domains
We have shown how cohesin regulators remodel chromosomes into functionally distinct domains to allow for the unique events of meiosis. We find that the control of loop formation and cohesion establishment by Eco1 and Wpl1 allows centromeres, pericentromeres and chromosome arms to adopt region-specific functions in sister kinetochore co-orientation, cohesion maintenance and recombination. These functions of Eco1 and Wpl1 that we uncovered in meiosis may also explain how other chromosome domains are established to support other genomic functions, including localized DNA repair and control of gene expression.

Loop anchoring allows the formation of specific chromosomal boundaries
Eco1 associates with replication factors and is proposed to couple cohesion establishment to DNA replication by travelling with replication forks (Ivanov et al., 2018;Ladurner et al., 2016;Lengronne et al., 2006;Song et al., 2012). In mitotically dividing yeast cells, Smc3 acetylation is largely dependent on DNA replication (Ben-Shahar et al., 2008), however during meiotic S phase substantial Smc3 acetylation, and anchoring of chromatin boundaries, occurs even in the absence of DNA replication. Cohesin acetylation also occurs without DNA replication in mammalian G1 cells, with a preference for STAG1-containing cohesin complexes (Alomer et al., 2017;Wutz et al., 2020). Therefore, Smc3-Ac can exist independently of cohesion establishment. Indeed, Smc3-Ac per se is not critical for cohesion since mitotic eco1D wpl1D yeast cells build sufficient cohesion to support viability. How Eco1 gains access to cohesin that is not associated with replication forks remains to be understood but it is interesting to speculate that specialized cohesin subunits, for example STAG1 in mammals or Rec8 in yeast, may allow cohesin targeting by Eco1 independent of the replication machinery.
In addition to Wpl1 antagonism, Eco1 and Smc3-Ac confer a more fundamental role in chromosomal loop positioning that we find to be indispensable for meiosis. We envisage that early in meiosis, prior to DNA replication, Scc2-Scc4-cohesin complexes load onto chromosomes and begin to extrude loops ( Figure 7H). Loaded cohesin, and loops, have a limited lifetime due to the removal of unacetylated cohesin by Wpl1.
However, a fraction of cohesin engaged in cis-looping is acetylated by Eco1, with two consequences. First, Smc3-Ac blocks cohesin's Wpl1-dependent release. Second, Smc3-Ac anchors cohesin at the base of loops, preventing loop migration and restricting extrusion, resulting in the stable positioning of moderately-sized loops. The absence of Wpl1 increases the lifetime of loop-extruding cohesin complexes resulting in loop extension (Figure 4; (Costantino et al., 2020;Dauban et al., 2020;Haarhuis et al., 2017)). The combined absence of both Eco1 and Wpl1 results in mobile cohesin with extended lifetimes on chromosomes and untempered loop extruding activity, leading to long, poorly positioned loops. The positioning activity of Eco1 also facilitates cohesion establishment along chromosome arms by anchoring cohesin complexes that are engaged in linking the two sister chromatids ("cohesive cohesin'). Whether it does this directly by acetylating cohesive cohesin to prevent its migration, or indirectly by generating cis-loops that form a barrier to translocation of cohesive cohesin is unclear. At centromeres, Smc3-Ac, which blocks the destabilizing activity of Wpl1, is sufficient for cohesion establishment, likely due to enhanced cohesin loading or specialized anchoring mechanisms at this site. A recent report indicates that Eco1-dependent Smc3 acetylation also establishes loop positioning in S phase of yeast mitotic growth (Bastié et al., 2021). Exactly how acetylation affects cohesin enzymology awaits detailed biochemical analysis.

Establishment of functional chromosomal units for meiotic recombination
Our studies on meiosis, where chromosomal domains must be defined to lose cohesin at chromosome arms in meiosis I and pericentromeres in meiosis II, provide a unique opportunity to dissect the functional importance of loop positioning. We

Centromeric cohesion directs sister chromatid co-segregation during meiosis I
Centromeres are unique in retaining higher cohesin levels in the absence of Eco1 function, but this is insufficient for centromeric cohesion. Moreover, Wpl1 inactivation in eco1-aa cells leads to an increase in chromosomal cohesin genome-wide, but cohesion is rescued specifically at centromeres. Our live-cell imaging revealed that the loss of centromeric cohesion in eco1-aa cells is accompanied by the aberrant segregation of sister chromatids to opposite poles in meiosis I, which was also rescued by Wpl1 inactivation. In Smc3-depleted and smc3-K112,113R cells, likely due to residual SMC3 expression from the CLB2 promoter, the centromeric cohesion defect was more modest and segregation of sister chromatids to opposite poles was less frequent, but was nevertheless rescued by Wpl1 inactivation. Although we cannot currently rule out the possibility that Eco1 has an additional substrate that confers its function in sister kinetochore co-orientation as has been suggested for fission yeast (Kagami et al., 2017), the simplest explanation is that acetylation of Smc3 by Eco1 counteracts Wpl1 at centromeres allowing the establishment of centromeric cohesion, which in turn facilitates kinetochore fusion via the monopolin complex (reviewed in (Duro and Marston, 2015)). Interestingly, budding yeast lacking Rec8 cohesin do not show co-orientation defects (Monje-Casas et al., 2007;Toth et al., 2000), raising the possibility that mitosis-like, Scc1-containing cohesin complexes confer the co-orientation function of cohesin at centromeres. It will be of great interest to understand how monopolin and cohesin-mediated co-orientation pathways intersect to ensure proper meiosis I chromosome segregation.

Pericentromere boundaries define persistent cohesion at meiosis II
During meiosis, cohesion on chromosome arms requires Eco1 and Smc3-Ac even in the absence of Wpl1. This argues against the idea that the key function of Smc3-Ac in cohesion is to antagonize Wpl1. Instead, we propose that the ability of Eco1 to reinforce chromatin boundaries is critical for cohesion during meiosis. This phenomenon is most apparent at pericentromere borders, where centromere-loaded cohesin is trapped by convergent genes in mitosis . In meiosis II, cells rely entirely on cohesin at pericentromere borders to hold sister chromatids together. We now show that Eco1 is required to retain cohesin at these sites and, consequently, to establish the boundaries that structure pericentromeres during meiosis. As a result, sister chromatids undergo random meiosis II segregation in cells lacking Eco1, whether or not Wpl1 is present. This can explain why cohesin anchoring is essential in meiosis but not in mitosis, where cohesin is present also along chromosome arms. Understanding how Eco1-dependent Smc3 acetylation and convergent genes together trap cohesin to elicit loop anchoring and cohesion at pericentromere borders is an important question for the future.

Importance of cohesin anchoring
A universal feature of cohesin-dependent chromosome organization is the existence of boundary elements which position both loops and cohesion. However, how these boundaries are established and the functional importance of cohesin anchoring were unclear. We found that Eco1-dependent cohesin acetylation confers boundary

Declaration of interests
The authors declare no competing interests. Xu, L., Ajimura, M., Padmore, R., Klein, C., and Kleckner, N. (1995).  -Ac is deposited in S phase, following Eco1 production. Wild type (strain AM21574) carrying ECO1-6HIS-3FLAG and pCUP1-IME1 pCUP1-IME4 was released from a pre-meiotic S phase block 120 min after sporulation induction by addition of 25 μM CuSO4. (A) S phase completion (4N) was monitored by flow cytometry. (B) The percentages of bi-and tetra-nucleate cells were scored at the indicated time points to monitor meiosis I and II nuclear division, respectively (n=200).

Figure 2. Counteracting Wpl1 is not the only essential role of Eco1 in meiosis
(A -C) Wpl1 is most abundant during meiotic S phase and prophase. Wild type (AM20916) carrying WPL1-6HA and pCUP1-IME1 pCUP1-IME4 was induced to undergo synchronous meiotic S phase as described in Figure 1A. (A) Western immunoblot shows total protein levels of Wpl1-6HA (a-HA) with Pgk1 loading control (α-Pgk1). (B) Flow cytometry profiles and (C) nuclear division (n=200) show the timing of bulk DNA replication and chromosome segregation, respectively. (D) Eco1 is essential for meiosis, even in the absence of Wpl1. Spore viability of wild type (AM24170), wpl1Δ (AM24265), eco1-aa (AM24171), and eco1-aa wpl1Δ (AM24289) strains, sporulated in the presence of 1 μM rapamycin. (E and F) Establishment of centromeric cohesion requires Eco1-dependent antagonism of Wpl1. Wild type (AM27183), wpl1Δ (AM27186), eco1-aa (AM27185), and eco1-aa wpl1Δ (AM27184) anchor away strains carrying heterozygous CEN5-GFP and ndt80D were induced to sporulate in 1 μM rapamycin and the percentage of cells with two visible GFP foci was scored at the indicated timepoints (F). Meiotic progression was monitored as DNA content (E). (G and H) Chromosomal arm cohesion requires Eco1, even in the absence of Wpl1. Wild type (AM27253), wpl1Δ (AM27256), eco1aa (AM27255), and eco1-aa wpl1Δ (AM27254) anchor away strains carrying heterozygous LYS2-GFP and ndt80D were treated and analyzed as described in (E and F). In (E-H) an average of 3 biological replicates is shown; 100 cells were scored for each timepoint in each experiment. Error bars show standard error; * p<0.05, **p<0.01, paired student T test when compared to wild type.     (E-G) Smc3 acetylation is required to ensure co-segregation of sister chromatids in meiosis I and accurate meiosis II chromosome segregation. Meiotic progression (E) and meiosis I (F) and II (G) chromosome segregation were scored after live cell imaging as in Figure 6 (B-E). Strains used were spo11D (AM30238), spo11D pCLB2-3HA-SMC3 (AM30240), spo11D SMC3 (AM30242), spo11 smc3-K112,113R (AM30244), spo11D wpl1D (AM30234), spo11D wpl1D pCLB2-3HA-SMC3 (AM30235), spo11D wpl1D SMC3 (AM30655) and spo11D wpl1D smc3-K112,113R (AM30237). (H) Model for Eco1 and Wpl1 roles in meiosis. In wild type cells (top panel), Eco1 cohesin acetylation is essential in three meiotic processes: it protects centromeric cohesin from Wpl1-mediated release, allowing sufficient cohesion to be built to establish monoorientation; it positions DNA loops along chromosome arms to promote recombination and prophase exit; and it positions loops and cohesion at pericentromeric borders to guide correct sister chromatid segregation in meiosis II. In the absence of Eco1 or Smc3-Ac (middle panel), boundaries are not respected and more cohesin complexes are released from DNA due to the action of Wpl1, with detrimental effects on recombination and meiosis I and II segregation. In the absence of both Eco1 or Smc3-Ac and Wpl1, cohesion is partially restored, specifically at the centromere, but loop boundaries are not, leading to the formation of long unpositioned loops that are not able to support prophase recombination and meiosis II segregation.

Yeast strains and plasmids
Yeast strains used in this study were either Saccharomyces cerevisiae SK1 or W303 derivatives or Schizosaccharomyces pombe, all of which are listed in Table S1.
Plasmids used in this study are listed in Table S2. Gene tags, gene deletions, and promoter replacements were introduced using standard PCR-based methods.

Growth conditions
For asynchronous mitotic culture, cells were inoculated into YPDA (1% Bacto yeast extract, 2% Bacto peptone, 2% Glucose, 0.3 mM adenine) and grown at 30 o C with shaking at 250 rpm for ~15 h. Cells were diluted to OD600=0.2 in YPDA and grown at room temperature for approximately 5 h until OD600 = 0.7-1.0. The OD600 was measured, and the cells were harvested for the experiment. For testing the viability of anchor-away strains YPDA agar plates contained 0.1 μM rapamycin.

Spore viability
Diploid strains were sporulated on agar (>48h) or liquid (24h) SPO media (with addition of appropriate drugs) at 30 o C. Cells were treated with 20 μl 1 mg/ml zymolyase (AMS Biotechnology) in 2 M sorbitol for 8 min, before diluting with 1 ml dH2O. A minimum of 50 tetrads were dissected into individual spores on YPDA agar using a micromanipulator on a Nikon Eclipse 50i light microscope and viable colonies were scored ~48h later. polylysine for 5 min, washing in dH2O and air-drying) for 10 min. The supernatant was aspirated, cell density confirmed by light microscopy, then the slide was incubated in 100% MeOH for 3 min followed by 10 sec in 100% acetone before air drying. To each well, 5 μl primary rat anti-tubulin antibody (Bio-Rad AbD Serotec) at a 1:50 dilution in PBS/1%BSA was added and the slide incubated in a dark wet chamber at room temperature for 2 h. Wells were washed individually five times in 5 μl PBS/BSA with aspiration between each wash. Secondary donkey anti-rat FITC antibody (Jackson ImmunoResearch) was added at a 1:100 dilution in PBS/BSA (5 µl), and incubated for a further 2 h before washing 5 times as above. DAPI-Mount (1 mg/ml pphenylenediamine, 0.05 μg/ml DAPI, 40 mM K2HPO4, 10 mM KH2PO4, 150 mM NaCl, 0.1% NaN3, 90% glycerol) was added to each well (3 µl) and the slide was covered with a 24x60 mm cover-slip and sealed with nail varnish. Slides were then stored at -20 o C before visualizing on a Zeiss Axioplan Imager Z2 fluorescence microscope with a 100 x Plan ApoChromat NA 1.45 oil lens. Spindle morphology was scored in 200 cells per timepoint.

Flow cytometry
For flow cytometry, 500 μl of cycling or 150 μl of meiotic culture was fixed in 70% EtOH and stored at 4 o C. Pelleted cells were resuspended in 1 ml 50 mM Tris pH 7.5 and sonicated in a Bioruptor Twin (Diagenode) on LOW 30 sec ON. Pellets were collected, resuspended in 475 μl of 50 mM Tris-HCl pH 7.5 with 25 μl 20 mg/ml RNase A (Amresco) and incubated at 37 o C overnight. Cell pellets were washed in 1 ml 50 mM Tris pH 7.5, resuspended in 500 μl 50 mM Tris pH 7.5 with 10 μl 20 mg/ml Proteinase K (Invitrogen) and placed at 50 o C for 2 h. Collected cell pellets were washed in 1 ml 50 mM NaCitrate, resuspended in 500 μl 50 mM NaCitrate with 9.17 μl 1 mg/ml propridium iodide (Sigma Aldrich), and sonicated on LOW 30 sec ON, 30 sec OFF for 10 times. Samples were measured on a Becton Dickinson FACSCalibur with CellQuest Pro programme or an Attune NxT flow cytometer (n = 20,000 cells per sample) and analyzed using FlowJo V10. Signal intensity is shown on a linear scale.

Visualization of GFP-labelled chromosomes
100 μl of culture was added to 10 μl of 37% formaldehyde and incubated for 8-9 min at room temperature before pelleting. The supernatant was aspirated, 1 ml of 80% EtOH added and tubes briefly vortexed. Cells were collected by short (~15 sec) centrifugation, all traces of 80% EtOH removed by pipetting, the pellet was resuspended in 20 μl of 1 μg/ml DAPI in PBS and stored at 4 o C. For visualization, 3ul of sample was transferred to a glass slide, a coverslip added, pressure applied to flatten the cells before viewing under a Zeiss Axioplan Imager Z2 fluorescence microscope with a 100 x Plan ApoChromat NA 1.45 oil lens. All unbudded cells were scored for the presence of either one or two GFP dots (n=100). All experiments were done in triplicate.

Western blotting
For mitotic protein extracts 10 ml of YPDA cell culture OD600=0.6-1 was collected, and for meiotic protein extracts 5 ml of SPO cell culture OD600=1.8 was collected. Cells were pelleted by centrifugation, resuspended in 5 ml 5% trichloroacetic acid and incubated on ice for a minimum of 10 min, before pelleting and transferring to a 2 ml fast-prep tube (MP Biomedicals). The fast-prep tube was centrifuged, the supernatant removed and snap-frozen in liquid nitrogen before storing at -80 o C. Thawed samples were resuspended in 1 ml acetone by vortexing, cells pelleted by centrifugation and the acetone removed. Samples were air-dried until the pellet was dry (>4h) and resuspended in 100 μl of protein breakage buffer (10 mM Tris-HCl pH 7.5, 1 mM EDTA pH 7.5, 2.75 mM DTT, 1 x Roche EDTA-free protease inhibitors) and one volume of glass beads (0.5mM zirconia/silica glass beads, Biospec Products) added. The cells were disrupted in a Fastprep Bio-Pulveriser FP120 at 6.5 speed for 3 cycles of 45 sec, placed on ice and 50 μl of 3xSDS sample buffer (187 mM Tris-HCl pH 6.8, 6% β-mercaptoethanol, 30% glycerol, 9% SDS, 0.05% bromophenol blue) was added to the lysate before immediately heating at 95 o C for 5 min, cooled and centrifuged before loading onto SDS-PAGE gels of the appropriate concentration (8-10%). PAGE was carried out using a Biorad Mini Trans-Blot System (Biorad) or a Biometra V17.15 lysate and debris were collected by centrifugation, transferred to a new tube and then pelleted by centrifugation at 13,000 rpm for 15 min at 4 o C. The cell pellet was resuspended in 1 ml 1xFA lysis buffer with 0.1% SDS, 1x Roche EDTA-free protease inhibitors and 1 mM PMSF, and re-centrifuged. The supernatant was removed, the pellet resuspended in 300 μl 1xFA lysis buffer with 0.1% SDS, 1x Roche EDTA-free Approximately 120 μl of supernatant was transferred into a new tube, and frozen at -20 o C. For qPCR either SYBR GreenER (Invitrogen; Figure S5B-C) or Luna (NEB; Figure S5E) mastermix was used. Input DNA was diluted 1:500 or 1:300 and the ChIP DNA was diluted 1:10 or 1: 6 in Hyclone Water for SYBR GreenER or Luna, respectively. For each biological replicate, qPCR reactions were carried out in technical triplicate on a Lightcycler 480 Roche machine with 40 cycles for SYBR GreenER and 45 cycles for NEB Luna. The threshold cycle (Ct) values were computed by the Lightcycler 480 Roche software using the 2 nd derivative maximum algorithm.
The geometric mean of technical replicate Ct values was determined and delta Ct was calculated according to the following formula: ΔCt = Ct(ChIP) (Ct(Input) log(primer efficiency)(Input dilution factor)). Enrichment values are given as ChIP/Input = (primer efficiency)^( -ΔCt) . Primers used are given in Table S3.

ChIP -sequencing
For calibrated ChIP-seq (Smc3) we used the method of , as modified by (Galander et al., 2019b). For each condition 200 ml of S. cerevisiae meiotic culture was grown and fixed as for ChIP-qPCR and frozen in four pellets (each from 50 ml culture). Each pellet was mixed with a fixed pellet from 100 ml S. pombe culture prepared as follows. Wild type S. pombe (strain spAM29) was inoculated into YES media (0.5% yeast extract, 3% glucose, 2% agar 225mg/l each of adenine, histidine, leucin, uracil and lysine hydrochloride), grown for 16 h at 30 o C before diluting to OD600=0.07-0.1 and growing for approximately 4-6 h at 30 o C until the OD600=0.4. Cell culture (1l) was mixed with 100 ml of fixing solution (33.3 ml 37% formaldehyde in 66.6 ml diluent) and incubated at room temperature at 90 rpm for 2h. Cells were harvested, washed twice with TBS and completely resuspended in 1xFA lysis/0.1% SDS. An equal amount of cells, equivalent to 100 ml of culture, was evenly distributed into  Table S5.
For Hi-C data analysis, Fastq reads were aligned to S. cerevisiae SK1 reference genome using HiC-Pro v2.11.4 bowtie2 v2.3.4.1 (--very-sensitive -L 30 --score-min L,-0.6,-0.2 --end-to-end --reorder), removing singleton, multi hit, duplicated and MAPQ<10 reads. Read pairs were assigned to restriction fragment (DpnII) and invalid pairs filtered out. Valid interaction pairs were converted into the .cool contact matrix format using the cooler library, and matrixes balanced using Iterative correction down to one kilobase resolution. Multi-resolution cool files were uploaded onto a local HiGlass server. To generate pile-ups at centromeres/pericentromeric borders, the cooltools library was used and cool matrices were binned at one kilobase resolutions. Plots were created around the midpoint of centromeres/pericentromeric borders, at the same coordinates used for ChIP-seq plots (listed in Table S4), with 25, 50, 100 kb flanks on each side, showing the log10 mean interaction frequency using a colour map similar to HiGlass 'fall'. All centromere annotations were duplicated in both the forward/reverse strand orientations to create an average image which is mirror symmetrical. Pile-ups at pericentromeric borders were oriented where positions are identical in relation to the centromere. The ratio pile-ups between samples were created in a similar fashion plotting the log2 difference between samples in the 'coolwarm' colour map, e.g. A/B; red signifying increased contacts in A relative to B and blue decreased contacts in B relative to A. Scripts for visualisation will be available at (https://github.com/danrobertson87/Barton_2021). Cooler 'show' was also used to generate individual plots for each chromosome. Contact probability P(s) and its derivative slope plots were generated using the cooltools library. White stripes on plots represent regions where data was lost stochastically during mapping due to stringency settings filtering out reads. Hi-C matrices were aligned to ChIP-seq tracks on HiGlass.