1. Chromosomes and Gene Expression
  2. Structural Biology and Molecular Biophysics
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Folding of cohesin’s coiled coil is important for Scc2/4-induced association with chromosomes

  1. Naomi J Petela
  2. Andres Gonzalez Llamazares
  3. Sarah Dixon
  4. Bin Hu
  5. Byung-Gil Lee
  6. Jean Metson
  7. Heekyo Seo
  8. Antonio Ferrer-Harding
  9. Menelaos Voulgaris
  10. Thomas Gligoris
  11. James Collier
  12. Byung-Ha Oh
  13. Jan Löwe  Is a corresponding author
  14. Kim A Nasmyth  Is a corresponding author
  1. Department of Biochemistry, University of Oxford, United Kingdom
  2. MRC Laboratory of Molecular Biology, United Kingdom
  3. Institute of Medical Sciences, University of Aberdeen, United Kingdom
  4. Department of Biological Sciences, KAIST Institute for the Biocentury, Cancer Metastasis Control Center, Korea Advanced Institute of Science and Technology, Republic of Korea
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Cite this article as: eLife 2021;10:e67268 doi: 10.7554/eLife.67268

Abstract

Cohesin’s association with and translocation along chromosomal DNAs depend on an ATP hydrolysis cycle driving the association and subsequent release of DNA. This involves DNA being ‘clamped’ by Scc2 and ATP-dependent engagement of cohesin’s Smc1 and Smc3 head domains. Scc2’s replacement by Pds5 abrogates cohesin’s ATPase and has an important role in halting DNA loop extrusion. The ATPase domains of all SMC proteins are separated from their hinge dimerisation domains by 50-nm-long coiled coils, which have been observed to zip up along their entire length and fold around an elbow, thereby greatly shortening the distance between hinges and ATPase heads. Whether folding exists in vivo or has any physiological importance is not known. We present here a cryo-EM structure of the apo form of cohesin that reveals the structure of folded and zipped-up coils in unprecedented detail and shows that Scc2 can associate with Smc1’s ATPase head even when it is fully disengaged from that of Smc3. Using cysteine-specific crosslinking, we show that cohesin’s coiled coils are frequently folded in vivo, including when cohesin holds sister chromatids together. Moreover, we describe a mutation (SMC1D588Y) within Smc1’s hinge that alters how Scc2 and Pds5 interact with Smc1’s hinge and that enables Scc2 to support loading in the absence of its normal partner Scc4. The mutant phenotype of loading without Scc4 is only explicable if loading depends on an association between Scc2/4 and cohesin’s hinge, which in turn requires coiled coil folding.

Introduction

SMC complexes are highly conserved from prokaryotes to eukaryotes. Best characterised among this family are cohesin and condensin, both of which are DNA translocases (Ganji et al., 2018; Davidson et al., 2019; Kim et al., 2019; Golfier et al., 2020). Cohesin and condensin are thought to organise chromosomes in eukaryotes during interphase and mitosis respectively by producing long loops of DNA (Nasmyth, 1982), a process called loop extrusion (LE). Cohesin has an additional property, namely the ability to hold sister DNAs together from their genesis during S phase till their eventual disjunction to opposite poles of the cell during anaphase.

Cohesin is composed of two rod-shaped SMC proteins, Smc1 and Smc3, with a dimerisation interface at one end that is connected to an ABC-like ATPase domain via a 50-nm-long coiled coil (Figure 1A). Interaction via their dimerisation domains produces a V-shaped Smc1/3 heterodimer whose two arms are connected by a central ‘hinge’ domain. The two ATPase ‘head’ domains at the apices of this dimer are meanwhile inter-connected by a kleisin subunit, Scc1. Scc1’s N- and C-terminal domains bind respectively to the coiled coil emerging from Smc3’s head (its neck) and the base of Smc1’s ATPase, thereby creating a tripartite SMC-kleisin (S-K) ring (Figure 1A). Cohesin’s association with DNA as well as its abilities to hold sisters together and extrude DNA loops are facilitated by three large hook-shaped HAWK (HEAT repeat proteins associated with kleisins) proteins; Scc2, Scc3, and Pds5 (Figure 1A). Scc3 is thought to be permanently bound to the complex, whereas Scc2 and Pds5, which are mutually exclusive, are more dynamic. Of these, Scc2 has a crucial role in activating cohesin’s ATPase at least in vitro, whether in the presence or absence of DNA (Petela et al., 2018).

Figure 1 with 1 supplement see all
A mutation in the hinge domain of Smc1 restores viability in the absence of Scc4.

(A) Schematic representation of Saccharomycescerevisiae cohesin complex and its folding cycle. (B) Comparison of growth of wild-type (WT), scc4-4, and scc4-4 smc1D588Y strains at 35.5°C (K699, K8326, K19813). (C) Tetrad dissection of diploid strains containing SCC4/scc4Δ SMC1/smc1D588Y grown at 30°C. Spores expressing smc1D588Y are circled in red, and spores that lack Scc4 are indicated with blue hexagons. (D) Structure of the mouse Smc3-Smc1D574Y hinge domain (PDB: 7DG5). (E) Multiple sequence alignment indicating conservation of Smc1D588. (F) Structural superposition of the WT hinge and the D574Y mutant hinge. Tyr574 swings out relative to the position of D574 with a concomitant local conformational change of the mutated loop.

The discovery that anaphase is initiated through the opening of S-K rings due to cleavage of their kleisin moiety by the protease separase (Uhlmann, 2001) led to the suggestion that cohesion is mediated by the co-entrapment of sister DNAs within individual S-K rings (Haering et al., 2002). This hypothesis, known as the ring model, made the key prediction that site-specific chemical crosslinking of all three of the ring’s subunit interfaces would create a covalent topological linkage resistant to protein denaturation between small circular sister DNAs. Such catenated dimers (CDs) are indeed found in cells (Haering et al., 2008; Gligoris et al., 2014) and only under conditions in which cells form sister chromatid cohesion (Srinivasan et al., 2018).

The ring model envisages that once established during DNA replication, maintenance of sister chromatid cohesion during G2 and M phases would not require continued ATP hydrolysis. This notion, namely that cohesion is a passive process, explains why Scc2, though essential for loading and for maintaining cohesin’s association with unreplicated DNA in vivo, has no role in maintaining cohesion during G2/M phases (Ciosk et al., 2000; Srinivasan et al., 2019). Cohesin’s ATPase is strictly dependent on Scc2 in vitro and is presumably inactive in vivo upon Scc2’s departure. LE in contrast requires continuous ATP hydrolysis dependent on Scc2, at least in vitro (Davidson et al., 2019).

Yet another difference is that cohesion depends on passage of DNAs inside S-K rings while LE does not (Srinivasan et al., 2018; Davidson et al., 2019). Given that cohesion and LE involve at least some different mechanisms, it is perhaps not surprising that there is increasing evidence that the two processes are mutually exclusive in vivo (Srinivasan et al., 2018; Davidson et al., 2019). Complexes engaged in cohesion do not extrude loops and vice versa.

Though maintenance of cohesion may have little in common with LE, the process by which cohesion is created in the first place may utilise mechanisms common to LE. This is supported by the fact that Scc2 is required for entrapping DNA within S-K rings as well as for the DNA-dependent ATPase activity necessary for LE. DNA entrapment assays combined with cryo-EM structures suggest that a key intermediate common to both processes is the passage of DNA between disengaged ATPase heads followed by its ‘clamping’ by Scc2 on a surface on top of them created by ATP-dependent head engagement (Collier et al., 2020; Shi et al., 2020; Higashi et al., 2020). It is envisaged that DNA translocation during LE involves recurrent rounds of DNA clamping followed by its release upon ATP hydrolysis. If so, each round presumably involves clamping of DNA successively further along the chromatin fibre. Clamping in this manner may be an important feature of cohesin’s association with chromatin, at least during G1 when LE is possibly its main activity. Crucially, clamping in vitro does not require Scc3, which is necessary for cohesin’s stable association with chromatin in vivo and ensures, at least in vitro, that clamping is followed or accompanied by transient opening of the S-K ring and thereby entrapment of DNAs within (Collier et al., 2020). The key point is that clamping may be a feature not only of LE but also of the entrapment of DNAs within S-K rings necessary for cohesion.

Which interface of the S-K ring is opened through the action of Scc3 is uncertain as is the mechanism, either when individual DNAs are entrapped during G1 (or G2) or when sister DNAs are entrapped during the passage of replication forks. Complexes containing co-translational fusions, either between the C-terminus of Smc3 and the NTD of Scc1, or between Scc1’s C-terminus and the NTD of Smc1 are functional and capable of entrapping individual or sister DNAs within S-K rings. In contrast, the artificial connection of the Smc1 and Smc3 hinge domains using rapamycin blocks the establishment but not maintenance of sister chromatid cohesion (Gruber et al., 2006), leading to the suggestion that DNAs enter the S-K ring via a gate created by transient dissociation of the hinge. Whether this is really the case awaits more rigorous types of experiments.

Cohesin complexes defective in ATP hydrolysis, due to Smc1E1158Q and Smc3E1155Q (EQEQ) mutations, accumulate in the clamped state in vitro (Collier et al., 2020; Shi et al., 2020; Higashi et al., 2020). Along with Scc2, they also accumulate at Saccharomyces cerevisiae CEN sequences (Hu et al., 2011), which are sites at which cohesin loads onto chromosomes with especially high efficiency, due to an interaction between the kinetochore protein Ctf19 and Scc4 bound to Scc2’s largely unstructured N-terminal domain (Hinshaw et al., 2017). This suggests that in addition to being a recurrent feature of LE, formation of the clamped state may be an early step in cohesin’s de novo association with chromosomal DNA. Scc4 facilitates Scc2-mediated loading throughout chromosome arms as well as at CEN sequences, though how it does so is poorly understood.

When cohesin’s ATPase heads are disengaged, the coiled coils of Smc1 and Smc3 associate with each other along much of their length (Chapard et al., 2019). When this ‘zipping up’ includes the sections of coiled coils close to the ATPase heads, it forces them to adopt a configuration in which they are juxtaposed in a ‘J’ state that is distinct from, and incompatible with ATP-driven head engagement known as the ‘E’ state. Crucially, the zipping up of coiled coils in this manner is incompatible with the clamping of DNA by Scc2 on top of engaged heads and the latter is therefore accompanied by extensive unzipping, at least up to the elbow (Collier et al., 2020). Coiled coil zipping up is a feature of cohesin engaged in holding sister chromatids together, with sister DNAs entrapped within J-K compartments, namely between juxtaposed (J) heads and the kleisin associated with them (Chapard et al., 2019). Extensive zipping up may have an important role in preventing unregulated ATP hydrolysis or precocious head engagement.

Along with coiled coil zipping up, the generation of cohesive structures during S phase is accompanied by acetylation of Smc3’s K112 and K113 residues (Guacci et al., 2015; Beckouët et al., 2016). The double acetylation stabilises cohesin’s association with chromosomes and increases the residence time of Pds5, which unlike Scc2 is necessary for maintaining cohesion as well as preventing de-acetylation of K112 and K113 (Chan et al., 2012; Chan et al., 2013). Complexes occupied by Pds5 cannot hydrolyse ATP, and in addition to maintaining cohesive structures in post-replicative cells, replacement of Scc2 or its human orthologue Nipbl by Pds5 appears to block the DNA translocation necessary for LE throughout interphase (Petela et al., 2018; Wutz et al., 2017; Dauban et al., 2020).

An important property of complexes occupied by Pds5, but not those by Scc2, is their ability to dissociate from chromosomes (Chan et al., 2013). This releasing activity is blocked by acetylation of Smc3 K112 and K113 during S phase by Eco1, substitution of both residues by glutamine, fusion of Scc1’s NTD to Smc3’s C-terminus (Chan et al., 2012), or mutations that affect the interface between Smc1 and Smc3 ATPase heads when engaged in the presence of ATP (Elbatsh et al., 2016). These findings have led to the suggestion that dissociation of Scc1’s NTD from Smc3’s neck during head engagement in the presence of Pds5 has a key role in triggering release (Beckouët et al., 2016). This process normally requires binding of Wapl to Pds5 and Scc3, along with head engagement (Kueng et al., 2006; Muir et al., 2020). Crucially, neither Wapl nor Pds5 are intrinsic to the release process as neither protein is necessary when Scc2 is inactivated in G1 cells, suggesting that the dissociation of Scc1 from Smc3 necessary for release takes place when heads engage in the absence of Scc2 and that Pds5 and Wapl facilitate the process at least partly by occluding Scc2 (Srinivasan et al., 2019). How Smc3’s K112 and K113 residues contribute to release when unmodified is not understood. If release involved an intermediate similar to the clamped state, albeit with Pds5 replacing Scc2, then these residues could contribute to the binding of DNA to engaged heads.

As well as their tendency to zip up, a striking feature of cohesin’s SMC coiled coils is their folding around an elbow (Figure 1A) situated two thirds of the way between the heads and hinge (Bürmann et al., 2019). Folding around this discontinuity results in association of the hinge with a section of the coiled coil close to the so-called joint region, a break in the coiled-coils above the ATPase heads. Folding is a widely conserved feature of SMCs when observed using EM in vitro, both when heads are engaged (Collier et al., 2020) or disengaged (Bürmann et al., 2019), but whether folding occurs in vivo and has an important physiological function is not known. It has been postulated that the elbow could be involved in LE by coupling cycles of folding and unfolding with DNA translocation (Bürmann et al., 2019; Hassler et al., 2018). Further, it has been noted that a potential simultaneous interaction of a HAWK with the hinge and kleisin would require some sort of folding (Murayama and Uhlmann, 2015, Huis in 't Veld et al., 2014, Bürmann et al., 2019).

Despite the discovery that Scc2 facilitates binding of DNA to engaged ATPase heads in vitro (Shi et al., 2020; Higashi et al., 2020) and does so in the absence of Scc3 without entry inside the S-K ring (Collier et al., 2020), the mechanism by which Scc2 promotes cohesin’s association with and translocation along chromosomes in vivo remains poorly understood. Scc2’s unstructured NTD is bound by a superhelical array of 13 tetratricopeptide repeats (TPRs) belonging to its partner Scc4 (Hinshaw et al., 2015). Cells lacking either Scc4 or Scc2’s NTD are not viable and have greatly reduced levels of chromosomal cohesin. Nevertheless, a version of Scc2 lacking the NTD is fully capable of activating cohesin’s ATPase (Petela et al., 2018) and clamping DNA on top of engaged ATPase heads in vitro (Shi et al., 2020; Collier et al., 2020). To gain insight into the role of Scc4, we recently undertook a genetic screen, isolating mutations that suppress lethality caused by loss of Scc4 activity (Petela et al., 2018). This identified two different scc2 point mutations, E822K and L937F. scc2E822K lies in the interface between Scc2 and Smc3’s K112 and K113, and as such, all three residues are in the vicinity of DNA clamped by Scc2 on top of engaged ATPase heads. Because acetylation of Smc3 K112 K113 greatly reduces cohesin loading as well as release, Scc2E822K might bypass Scc4 by increasing the avidity with which DNA is clamped (Collier et al., 2020).

Here, we describe two other types of mutations that suppress scc4 lethality. One type includes mutations in histone H2A that loosen the association between nucleosomes and DNA, which conceivably act like Scc2E822K, by facilitating cohesin’s interaction with naked DNA. The other is an aspartic acid on the surface of Smc1’s hinge domain that is replaced by an aromatic residue: smc1D588Y. UV-induced crosslinking in cells whose Smc1 hinge contains p-benzoyl L-phenylalanine (BPA) at defined positions revealed that it contacts Scc2, Scc3, and Pds5. Inactivation of Scc4 reduced crosslinking with Scc2, but increased that with Pds5, while smc1D588Y had the opposite effect. These findings suggest that Smc1’s hinge contacts Scc2 and Pds5 directly, Scc4 facilitates association with Scc2 and hinders that with Pds5, and smc1D588Y does likewise and compensates for a lack of Scc4. To explain how Scc2 contacts Smc1’s hinge while also bound to Smc1’s ATPase, we suppose that cohesin’s coiled coil is folded around its elbow, thereby bringing the hinge into contact with HAWK regulatory subunits associated with cohesin’s ATPase heads, as recently observed in a cryo-EM structure of the ATP-bound clamped state (Collier et al., 2020).

Using cryo-EM, we have now determined the structures of the folded cohesin complex associated with either Scc2 or Pds5, both in the absence of ATP. The structures demonstrate that both Scc2 and Pds5, while attached to the ATPase domains of Smc1 and Smc3, respectively, reach up to the hinge, thus providing a clue regarding the effects of Smc1D588Y and an explanation for previously observed K620BPA crosslinks (Bürmann et al., 2019). The resolution of the folded coiled coils and hinge (5–6 Å) not only permitted the identification of the contacts involved in folding but also allowed identification of candidates for cysteine substitution for potential bismaleimidoethane (BMOE) crosslinking to assay folding. One such residue pair, Smc1R578C-Smc3V933C, gave rise to efficient BMOE-induced crosslinking in vivo even when Smc3 was acetylated. Therefore, folding takes place not only when Scc2 is bound but also when cohesin is engaged in holding sister chromatids together in post-replicative cells. Our findings demonstrate that folding of cohesin’s coiled coils is not an in vitro artefact. Folding occurs in living cells, it is a feature of cohesin engaged in holding sister chromatids together, and it is of physiological importance during Scc2-mediated cohesin loading.

Results

To understand better how Scc4 helps Scc2 to load cohesin onto chromosomes, we isolated mutations that enable temperature-sensitive scc4-4 cells to grow at the restrictive temperature (35.5°C). This yielded both intragenic and extragenic mutations. The scc4-4 allele was created by error-prone PCR (Ciosk et al., 2000) and contains several different mutations, including Y40N. Sequencing of intragenic revertants revealed that wild-type (WT) growth was restored either by restoring tyrosine at position 40 or by substituting it with histidine, implying that the mutation responsible for scc4-4’s thermosensitive proliferation is Y40N. When integrated at the LEU2 locus, this mutation alone conferred temperature sensitive (ts) growth (Figure 1—figure supplement 1A). Y40 is a highly conserved residue that is buried in Scc4’s superhelical array of TPR motifs (Figure 1—figure supplement 1B). It is unlikely that it contacts the Scc2 polypeptide directly, but despite this, Y40N disrupts co-immunoprecipitation of Scc4 and Scc2 (Figure 1—figure supplement 1C).

A mutation in the hinge domain of Smc1 restores viability in the absence of Scc4

All of the extragenic scc4-4 suppressors (Figure 1B) contained a mutation tightly linked to SMC1. Indeed, all 12 independently isolated mutations contained the same single base change causing substitution of aspartic acid by tyrosine at position 588 in Smc1 (Figure 1E). Tetrad dissection of SCC4/scc4Δ SMC1/smc1D588Y diploids revealed that smc1D588Y enabled cells to proliferate in the complete absence of Scc4 (Figure 1C). Smc1D588 is located in the hinge domain, at the C-terminal end of a β strand that interacts in an antiparallel fashion with a strand in Smc3 (Figure 1D). Despite its proximity to the Smc1-Smc3 interface, D588 does not appear to contact Smc3 residues. To address whether suppression arises due to the loss of a relatively conserved acidic residue (Figure 1E) or due to the substitution of a bulky aromatic, we tested the ability of a variety of other amino acid substitutions to rescue viability in the absence of Scc4. Mutant or WT alleles of SMC1 were introduced into the TRP1 locus of a SMC1/smc1Δ SCC4/scc4Δ diploid. Subsequent dissection revealed that mutation to phenylalanine or tryptophan was able to restore growth to a similar degree as tyrosine in the absence of Scc4 (Figure 1—figure supplement 1D) but not histidine, arginine, alanine, glutamic acid, or asparagine (Supplementary file 1). Suppression of scc4Δ lethality also occurred in the presence of WT SMC1 but was much less effective (Figure 1—figure supplement 1D). Thus, we conclude that suppression is due to the introduction of a bulky aromatic amino acid at this crucial position and not through loss of the conserved aspartic acid. It is notable that the DNA base change observed in all 12 suppressors is the only one capable of creating such a transition via a single change and that the equivalent position is never a bulky aromatic in SMC2, SMC3, and SMC4 as well as SMC1. smc1D588Y was able to rescue the proliferation defect of the temperature-sensitive scc2-4 allele at 30°C, but not scc2Δ (Figure 1—figure supplement 1E, F), implying that it acts by enhancing the activity of Scc2, not by replacing it.

To determine whether the smc1D588Y mutation alters the hinge structure, we introduced the equivalent mutation (D574Y) into an isolated mouse hinge. X-ray crystallography revealed that a clash between the tyrosine residue and neighbouring loop causes Y574 to instead swing out relative to the position of D574, causing a local conformational change of the mutated loop (Figure 1D, F, Figure 1—figure supplement 1G, Supplementary file 2). Importantly, the change had little or no impact on the overall structure of the hinge. Despite this, both D588Y and D588W reduced the amount of Smc1 hinge that co-precipitated with Smc3 hinges (Figure 1—figure supplement 1H). To address whether Smc1D588Y affects dissociation of pre-assembled Smc1/3 hinge complexes, we co-expressed either WT Smc1 or Smc1D588Y hinge domains with Smc3 hinges, purified Smc1/3 complexes, and compared their persistence in the presence of a fivefold excess of SNAP-tagged Smc1 hinge domains. This revealed that the amount of Smc1D588Y associated with Smc3 hinges declined more rapidly in the presence of a WT competitor than WT Smc1, indicating that Smc1D588Y at least increases the off rate (Figure 1—figure supplement 1I).

Our finding that Smc1D588Y increases dissociation of Smc1 from Smc3 hinge domains in vitro raises the possibility that suppression depends on, or indeed is caused by, the greater ease with which hinges can dissociate. We therefore tested whether other non-lethal mutations within the Smc1/Smc3 hinge interface are also capable of bypassing the need for Scc4. A highly conserved lysine residue within Smc3 (Smc3K652) that opposes Smc1D588 was mutated to tyrosine, alanine, or valine, with no effect. Similarly, previously published mutations in both hinges, designed to weaken their interaction, smc1L635K K639E; smc1I590K; smc1L564K; smc3E570K; smc3L672R (Mishra et al., 2010), were also unable to support growth in the absence of Scc4. The failure of these other mutations to suppress scc4Δ lethality, together with our finding that smc1D588Y was identified in 12 out of 12 spontaneous extragenic suppressors, suggests that decreased affinity of Smc1/Smc3 hinges is not the mechanism by which Smc1D588Y enables Scc2 to load cohesin without Scc4.

smc1D588Y restores cohesin occupancy on chromosome arms in the absence of Scc4

Calibrated ChIP-seq revealed that scc4-4 causes a substantial reduction in the level of chromosomal cohesin when G1 cells undergo S phase and enter G2/M at the restrictive temperature (37°C) (Figure 2A, Figure 2—figure supplement 1A). The reduction is more marked within pericentric sequences, where there is a 10-fold reduction, than along arms where is there merely a fourfold reduction. Average chromosome profiles centred around the centromeric CDEIII, plotted as a percentage of the reads obtained for WT, revealed that smc1D588Y restores cohesin occupancy to approximately WT levels on chromosome arms (>30 kb from the centromere), but not around centromeres (Figure 2A). The failure to restore loading around centromeres is perhaps not surprising as most pericentric cohesin is loaded at CENs in a process that involves binding of Scc4 to the kinetochore protein Ctf19, a requirement that is apparently not bypassed by smc1D588Y. Interestingly, smc1D588Y caused a substantial reduction of cohesin occupancy around centromeres even in the presence of WT SCC4 (Figure 2A), an effect that will also have contributed to the lack of suppression in this region of the chromosome.

Figure 2 with 1 supplement see all
smc1D588Y restores cohesin occupancy on chromosome arms in the absence of Scc4.

(A) Average calibrated ChIP-seq profiles of Scc1-PK6 in smc1D588Y, scc4-4, and smc1D588Y scc4-4 cells 60 kb either side of CDEIII plotted as a percentage of the average number of reads obtained for wild-type (WT) cells. Cells were pheromone arrested in G1 at 25°C before release at 37°C into medium containing nocodazole. Samples were taken 75 min after release (K22005, K22009, K21999, K22001). (B) Average calibrated ChIP-seq profiles of Scc1-PK6 in smc1D588Y, and smc1D588Y scc4Δ cells 60 kb either side of CDEIII plotted as a percentage of the average number of reads obtained for WT cells. Cells were pheromone arrested in G1 at 25°C before release at 25°C into medium containing nocodazole. Samples were taken 60 min after release (K22005, K22009, K19624). (C) Average calibrated ChIP-seq profiles of Scc2-PK6 2 kb either side of CDEIII in cycling WT, smc1D588Y, and smc1D588Y scc4Δ cells at 25°C (K21388, K24680, K24678). (D) Average calibrated ChIP-seq profiles of ectopically expressed Smc3E1155Q-PK6 2 kb either side of CDEIII in cycling WT, smc1D588Y, and smc1D588Y scc4Δ cells at 25°C (K24562, K24689, K24564). (E) ATPase activity of WT or mutant tetramers on addition of ATP and Scc2 in the presence and absence of DNA.

To investigate the effect of smc1D588Y at a more physiological temperature, we used calibrated ChIP-seq to compare cohesin’s occupancy of the genome in SCC4, SCC4 smc1D588Y, and scc4Δ smc1D588Y cells following their release from a pheromone-induced G1 arrest and subsequent arrest in G2/M phase at 25°C. Average chromosome profiles around centromeres plotted as a percentage of WT (SCC4) revealed that smc1D588Y increased cohesin occupancy on chromosome arms to 120–150% of WT levels, both in the presence and absence of SCC4 (Figure 2B). In other words, smc1D588Y enhances cohesin’s loading on chromosome arms via a mechanism that is completely independent of Scc4. In contrast, smc1D588Y reduced association around centromeres to approximately 60% of WT levels, which was further reduced to 20% by scc4Δ. In scc4Δ smc1D588Y cells, cohesin occupancy within pericentric chromatin resembles that along chromosome arms as if a single Smc1D588Y-driven mechanism is responsible for loading at both locations in these cells (Figure 2—figure supplement 1B). Importantly, smc1D588Y does not increase occupancy on chromosome arms merely because defective loading at CENs increases the amount of cohesin available to load onto chromosome arms because the scc4m35 mutation, which disrupts Scc4’s association with Ctf19 and also reduces loading at CENs, has no such effect (Petela et al., 2018; Hinshaw et al., 2015).

It is striking that in SCC4 cells smc1D588Y had far less effect on cohesin’s association at CEN loading sites themselves. For example, it was 110% of WT in cells growing at 25°C and 75% of WT at 37°C. Because association was greatly reduced in scc4-4 and scc4Δ cells (Figure 2A, B), it presumably arises as a consequence of Scc4’s association with Ctf19. If so, these complexes should be associated with Scc2, which was confirmed by calibrated ChIP-seq showing that Scc2’s association with CENs far from being reduced was in fact substantially increased by smc1D588Y and fully dependent on Scc4 (Figure 2C). Cohesin occupied by Scc2 at CENs could either be in the process of loading (Petela et al., 2018; Hu et al., 2011) or engaged in LE (Dauban et al., 2020; Paldi et al., 2020). In both cases, the complexes are likely to adopt at least transiently the clamped state, which is stabilised by smc1E1158Q and smc3E1155Q, at least in vitro (Collier et al., 2020). In cells, cohesin complexes containing these mutations accumulate to especially high levels at CENs, albeit with a short residence time (Hu et al., 2011), suggesting that they initiate an early step in the loading process, namely the clamped state, but in the absence of ATP hydrolysis fail to undergo a later step required for stable association and translocation into neighbouring pericentric sequences. Interestingly, smc1D588Y not only increased Scc2’s association with CENs but also caused a similar increase in Smc3E1155Q’s association (Figure 2D). This implies that the reduced loading around centromeres arises not from defective formation of the clamped state at CENs by Scc2/4 complexes associated with Ctf19 but from a defect in a subsequent step in the loading/translocation reaction that requires ATP hydrolysis. Because accumulation of Smc1D588Y complexes at CENs resembles that of complexes containing Smc3E1155Q, we tested the effect of Smc1D588Y on cohesin’s ATPase activity but found little or no effect either in the presence or absence of DNA (Figure 2E, Figure 2—figure supplement 1D).

Mutations in SCC2 and histone genes also suppress scc4Δ lethality

To address whether it is possible to identify extragenic scc4Δ suppressor mutations besides smc1D588Y, we isolated a second set in a smc1D588E yeast strain that cannot mutate residue 588 to an aromatic residue through a single base pair mutation (Petela et al., 2018). We identified using genetic crosses and genomic sequencing 12 mutations within SCC2 (described in Petela et al., 2018), 49 within HTA1 (one of two histone H2As), and a single mutation within HTB1 (one of two H2Bs). All permitted proliferation of scc4Δ cells, albeit to a greater or lesser extent (Figure 3A).

Figure 3 with 1 supplement see all
Mutations in SCC2 and histone genes also suppress scc4Δ lethality.

(A) Tetrad dissection of diploid strains containing SCC4/scc4Δ leu2/scc4-4 ΗTΑ1/hta1R31I. Spores in which scc4Δ is rescued by hta1R31I are circled in blue. (B) Structure of the yeast nucleosome (PDB: 1ID3; White et al., 2001). H2A is shown in blue and H2B in green. Suppressor mutations are shown in yellow. (C) Average calibrated ChIP-seq profiles of Scc1-PK6 in hta1R31I, scc4-4, and hta1R31I scc4-4 cells 60 kb either side of CDEIII plotted as a percentage of the average number of reads obtained for wild-type (W)T cells. Cells were pheromone arrested in G1 at 25°C before release at 35.5°C into medium containing nocodazole. Samples were taken 60 min after release (K22005, K24574, K24568, K22001).

Scc4 helps overcome inhibition of loading by nucleosomes

The H2A mutations affected three residues, namely G30, R31, and R34. These mutations (G30D, R31I/T/S/G, and R34I) are all located on a defined patch on the surface of the nucleosome that interacts with DNA and the single H2B mutation (Y44D) is located nearby (Figure 3B). Because substitution of two positively charged residues causes suppression, we surmise that the mutations act by weakening the association between histones and DNA. hta1R31I was made de novo and shown to suppress the lethality of scc4-4 cells (Figure 3A). Its effect on cohesin loading in SCC4 and scc4-4 cells was measured using calibrated ChIP-seq to measure Scc1’s association with the genome after cells had undergone DNA replication at 35.5°C following a pheromone-induced G1 arrest at 25°C. A lower restrictive temperature (35.5°C) was used in this instance because hta1R31I is itself lethal at 37°C. Consistent with its poor suppression of scc4Δ lethality (Figure 3A), hta1R31I increased loading along chromosome arms more modestly than smc1D588Y, raising loading in scc4-4 cells from 20% to 70% of WT (HTA1 SCC4) (Figure 3C). As in the case of both smc1D588Y and scc2E822K L937F (Petela et al., 2018), hta1R31I failed to suppress the loading defect of scc4-4 mutants in the vicinity of centromeres (Figure 3C). Interestingly, in the presence of WT SCC4, hta1R31I actually increased loading along chromosome arms over WT by 20%. This implies that the association between histones and DNA within the nucleosome restricts cohesin loading, at least along chromosome arms, not only in scc4 mutants but also in WT cells. Like scc2E822K L937F (Petela et al., 2018) but unlike smc1D588Y, hta1R31I does not per se reduce loading of cohesin around centromeres (Figure 3C), suggesting that hta1R31I and smc1D588Y affect different aspects of the loading process.

It has been suggested that the chromatin structure remodelling complex (RSC) has a key role in loading cohesin onto yeast chromosomes (Huang et al., 2004) and that an important function of Scc2/4 along chromosome arms is to facilitate nucleosome remodelling catalysed by RSC (Lopez-Serra et al., 2014). This raised the possibility that mutations like hta1R31I suppress the loading defects of scc4 mutants because they bypass the need for RSC and smc1D588Y might act likewise. To address this, we used calibrated ChIP-seq to reinvestigate the loading defects of sth1-3 cells, which contain a temperature sensitive mutation within RSC’s ATPase subunit. WT and sth1-3 cells were arrested in G1 by α-factor at 25°C and then released from the block at 37°C. sth1-3 delayed budding, DNA replication, and the onset of Smc3 acetylation (Figure 3—figure supplement 1A–C), complicating the comparison with WT. We therefore compared the calibrated ChIP-seq profiles sth1-3 cells 105 min after release, when most but not all cells had both budded and undergone DNA replication, with WT cells at 75 min, a time point at which their cell cycle progression was most similar. Western blotting confirmed that the levels of Scc1 in this pair of samples were also similar (Figure 3—figure supplement 1C). Surprisingly, their calibrated ChIP-seq profiles were also very similar not only in the vicinity of centromeres (Figure 1—figure supplement 1D, F) but also throughout an interval 60 kb either side of centromeres (Figure 3—figure supplement 1F). Crucially, sth1-3 caused only a modest reduction in the occupancy ratio (OR), which denotes the overall level of association throughout the genome (Figure 3—figure supplement 1D). These findings contradict the previous claim that RSC has a crucial role in cohesin loading, based on qPCR measurements at individual loci of the very same sth1-3 strain (Lopez-Serra et al., 2014). Because of this discrepancy, we compared the calibrated ChIP-seq profiles of WT (75 min) and sth1-3 (105 min) in the vicinity of three loci whose association was previously reported to be 30% of WT. There was little or no effect of the mutation at CEN3, a modest reduction at POA1, and more surprisingly an increase at MET10 (Figure 3—figure supplement 1E).

Given the pleiotropic consequences of sth1-3 on cell cycle progression, it is difficult to exclude the possibility that RSC has a modest effect on cohesin loading. However, if Scc4 promoted loading by helping chromatin remodelling by RSC, then smc1D588Y should suppress any apparent loading defect caused by RSC. The fact that the Scc1 calibrated ChIP-seq profile of smc1D588Y sth1-3 double mutants is indistinguishable to that of sth1-3 single mutants (Figure 3—figure supplement 1G) shows that insofar that there is any defect, it is clearly unaffected by smc1D588Y. In other words, a version of cohesin that no longer requires Scc4 does not alter sth1-3’s albeit modest defect. It may therefore be a pleiotropic consequence of the mutant’s retarded cell cycle progression and not due to an Scc4-dependent RSC activity that creates nucleosome-free regions necessary for cohesin loading.

Recent work has revealed that Scc2 has a key role in clamping DNA onto engaged heads and that Scc2E822K, which also suppresses scc4Δ, might function by enhancing DNA binding within the clamped state (Collier et al., 2020; Shi et al., 2020; Higashi et al., 2020). We therefore suggest that the reason why histone mutations suppress the lethality of scc4 mutants is because they increase the accessibility of DNA and thereby facilitate formation of the clamped state.

Scc4 regulates an interaction between the hinge domain and HAWKs

How might replacement of a specific surface residue on the Smc1 hinge by a bulky aromatic one help Scc2 function without Scc4? One possibility is that it strengthens a hydrophobic interaction with another cohesin subunit. We have previously described the UV-dependent crosslinking in living yeast cells between Pds5 and a version of Smc1 containing BPA at position K620, which is located in an alpha helix adjacent to the loop containing D588 (Figure 4ABürmann et al., 2019). Pds5 is not required for cohesin loading, and therefore strengthening its interaction with Smc1’s hinge cannot be responsible for suppression. We therefore tested whether UV induces crosslinking of Smc1K620BPA to other regulatory subunits. To do this, cells expressing FLAG-tagged versions of Scc2, Scc3, Scc4, or Pds5 in cells whose sole source of Smc1 was Myc-tagged Smc1K620BPA were exposed to UV, and subsequent western blotting was used to detect FLAG-tagged proteins in immunoprecipitates (IPs) of Scc1-containing complexes (Figure 4B).

Figure 4 with 1 supplement see all
Scc4 regulates an interaction between the hinge domain and HAWKs.

(A) Modelled structure of the yeast cohesin hinge domain based on bacterial SMC hinge from Thermotoga maritima (PDB: 1GXL; Haering et al., 2002). (B) Identification of proteins that crosslink to Smc1 hinge. Strains expressing various cohesin regulators tagged with either FLAG6 or HA6 in combination with Smc1K620BPA-myc were treated with UV prior to immunoprecipitation with PK-tagged Scc1 and the products analysed by western blotting (B1969, B1976, B1983, B2020, B2072, B2079). (C) Effect of Scc4 and Smc1D588Y on crosslinking between Pds5 and Smc1 hinge. Cells expressing Smc1K620BPA in the presence or absence of scc4-4 and Smc1D588Y were exponentially grown at 25°C and shifted to 35.5°C for 1 hr. Cells were irradiated with UV, and the cohesin complex was isolated by immunoprecipitation of PK-tagged Scc1. The Myc-tagged Smc1K620BPA was examined by western blot (B2072, B2212, B2214, B2215). (D) Quantification of the crosslinks in (C) as a percentage of the wild-type (WT) Smc1 crosslinking efficiency. (E) Effect of Scc4 and Smc1D588Y on crosslinking between Scc2 and Smc1 hinge. Strains were treated as described in (C) (B1969, B2213, B2216, B2217). (F) Quantification of the crosslinks in (E) as a percentage of the WT Smc1 crosslinking efficiency. The experiments shown in (CF) were performed twice with the same result. (G) In vivo cysteine crosslinking of Smc1 hinge with Scc2 protein. Yeast cells expressing Smc1K620C and Scc2N200C were incubated with bismaleimidoethane (BMOE) (B3082, B3107, B3114, and B3116). The crosslinked Smc1/Scc2 was isolated by immunoprecipitation of PK-tagged Scc1 and examined by western blot. * Unspecific crosslink band.

Western blotting for the Myc epitope confirmed that all samples contained a high molecular weight version of Smc1, consisting of proteins crosslinked to K620 (Figure 4B). As expected for a subunit that is stably associated with cohesin Smc-kleisin trimers, high levels of Scc3 were detected in IPs from Scc3-FLAG cells, most of which had an electrophoretic mobility expected of uncrosslinked protein, but a small fraction co-migrated with the high molecular weight version of Smc1, suggesting that UV also induces crosslinking of Smc1K620BPA to Scc3. Pds5 is less stably associated, explaining why only modest amounts of uncrosslinked Pds5 are detected in the IPs. Despite this, we observed much more Smc1-Pds5 than Smc1-Scc3 crosslinked protein, confirming that Smc1K620BPA crosslinks to Pds5 with high efficiency (Bürmann et al., 2019). Because co-precipitation of unstably associated proteins will be greatly enhanced by crosslinking, it is not possible to assess the actual fraction of crosslinked protein. Scc2’s residence time on chromosomal cohesin of approximately 2–4 s (Hu et al., 2011) is even less than that of Pds5 and the former is therefore difficult to detect in Smc1 IPs. Nevertheless, the level of Smc1-Scc2 crosslinked protein was comparable to that of Scc3, despite being overall threefold less abundant (Tóth et al., 1999). In contrast, we detected no Smc1-Scc4 crosslinked proteins in Scc4FLAG cells. Cryo-EM has revealed that the N-terminal HEAT repeats of Scc2 as well as those of its human ortholog Nipbl are found in close proximity to Smc1’s hinge within complexes that have clamped DNA on top of their engaged ATPase domains (Shi et al., 2020; Higashi et al., 2020; Collier et al., 2020) and the crosslinking between Smc1K620BPA and Scc2 may reflect this state. However, they could also reflect an alternative one in which Scc2 is bound to cohesin whose heads are disengaged as described in the next section.

As association of cohesin with Scc2 and Pds5 is mutually exclusive and the latter incapable of activating cohesin’s ATPase or association with chromatin (Petela et al., 2018), Scc4 and Smc1D588Y could facilitate loading either by enhancing association of the hinge with Scc2 or decreasing it with Pds5. To test this, we measured the effect of scc4-4 in the presence or absence of smc1D588Y on Smc1K620BPA-Pds5 and Smc1K620BPA-Scc2 crosslinking. This revealed that Scc4 inactivation (scc4-4) increased Pds5 crosslinking threefold while smc1D588Y had the opposite effect. Crucially, smc1D588Y was largely epistatic to scc4-4. In other words, the elevated crosslinking observed when Scc4 was inactivated dropped in the presence of smc1D588Y to the depressed level of SCC4 smc1D588Y cells (Figure 4C, D). The mutations had the opposite, albeit less dramatic, effects on Smc1K620BPA-Scc2 crosslinking. Scc4 inactivation halved it while smc1D588Y restored it 80% of WT levels (Figure 4E, F). These results are consistent with the notion that a key function of Scc4 is to facilitate interaction between Scc2 and the Smc1 hinge, either directly or indirectly by impeding the latter’s interaction with Scc2’s competitor Pds5 or conceivably via both mechanisms.

If the essential role of Scc4 were merely to hinder an interaction between the Smc1 hinge and Pds5, then Scc4 should be unnecessary for cohesin’s association with chromosome arms in cells lacking Pds5. We therefore used calibrated ChIP-seq to measure the effect of depleting Pds5 (using the auxin-dependent AID degron) on cohesin’s occupancy of chromosome arms after scc4-4 cells undergo S phase at 37°C, which revealed that Pds5 depletion had no effect (Figure 4—figure supplement 1A). In other words, Pds5 is not necessary for depressing cohesin’s association with chromosomes in scc4-4 mutants. Likewise, if by reducing Pds5’s interaction with the Smc1 hinge smc1D588Y reduced Pds5’s occupancy of chromosomal cohesin, then it should depress the fraction of chromosomal cohesin associated with Pds5. The fact that smc1D588Y has no such effect (Figure 1—figure supplement 1B, C) implies that though the mutation alters how Pds5 interacts with the Smc1 hinge, this does not in fact alter chromosomal cohesin’s occupancy by Pds5.

Our finding that Scc4 does not act solely by hindering Pds5 suggests that Scc4 and Smc1D588Y facilitate Scc2 activity by promoting its interaction with the hinge. To elucidate where the hinge contacts Scc2, we inserted TEV protease cleavage sites at various positions within Scc2 to determine whether Smc1K620BPA crosslinked to the N- or C-terminal fragments created by TEV cleavage (Figure 1—figure supplement 1D, E). Analysis of those TEV insertions that were functional in vivo revealed that crosslinking occurred within Scc2’s N-terminal sequences, between residues 150 and 215. This interval is between the N-terminal domain that binds Scc4 (Hinshaw et al., 2015) and the hook-shaped structure composed of HEAT repeats. This part of Scc2 is not sufficiently ordered to have been visualised in the cryo-EM structure of a complex containing DNA clamped between Scc2 and engaged Smc1/3 ATPases (Collier et al., 2020). To confirm the location, we measured BMOE-induced crosslinking in vivo between Smc1K620C and a variety of Scc2 cysteine substitutions between residues 153 and 212. Although Smc1K620C alone gave rise to a Smc1-Scc2 crosslinked species, the crosslinking was more efficient on the introduction of Scc2N200C, suggesting that Smc1 is likely also crosslinking to a natural cysteine in Scc2 (most likely Scc2C224, which sits on a small helix just below N200) (Figure 4G). Importantly, the region of Scc2 whose association with the Smc1 hinge is reduced by scc4-4 and restored by smc1D588Y is close to where Scc4 binds to Scc2 (Hinshaw et al., 2015). In other words, Scc4 would be close enough to directly influence Scc2’s interaction with the hinge.

Cryo-EM structures of cohesin trimers associated with Scc2 or Pds5 reveal folded coiled coils

The notion that smc1D588Y suppresses scc4Δ by altering the interaction between Smc1’s hinge domain and cohesin’s HAWK subunits Scc2 and Pds5 raises a conundrum: how can HAWK proteins, which are known to associate with cohesin’s kleisin subunit and its ATPase domains, interact with a hinge domain that is separated from the ATPase domains by a 50-nm-long coiled coil? One possibility is that the HAWK proteins interact with cohesin’s hinge and ATPase domains at different points in time. Alternatively, if in fact they interact with hinge and heads simultaneously, then the coiled coil cannot be fully extended. For example, folding at an elbow in the middle of the coiled coil (Bürmann et al., 2019) may bring the hinge into proximity of HAWKs associated with the ATPases. Folding has recently been observed at low resolution in a complex between DNA, Scc2, and hydrolysis-impaired EQEQ ATPases engaged in the presence of ATP (Collier et al., 2020; Shi et al., 2020; Higashi et al., 2020). To investigate this further, we used cryo-EM to determine the structures of the S. cerevisiae cohesin trimer (Smc1, Smc3, and Scc1 containing cysteines specifically crosslinking the three intermolecular interfaces; Smc1-Scc1, Smc3-Scc1, and the Smc1-Smc3 hinge [Collier et al., 2020] at an efficiency of 20% [data not shown]) bound to either Scc2 (Figure 5A) (EMD-12880) or Pds5 (Figure 6A) (EMD-12888) in the absence of nucleotide and DNA. The former revealed a coiled coil folded at its elbow (Figure 5B, C), causing the hinge to interact with sections of the coiled coil that are approximately 10 nm away from the point at which they emerge from the ATPase domains (Figure 5B). The cryo-EM reconstruction not only revealed the path of the coiled coils around the hinge-coiled coil interface (where the map is at 5–6 Å resolution; EMD-12887), but also enabled the production of a pseudo-atomic model of the folded form (PDB: 7OGT). Folding brings a pair of helices within Smc1’s hinge, namely the end of the coiled coil around A520-F526 and another short helix around L564-R578, into close proximity of a short stretch of Smc3’s coiled coil (Figure 5D). Very similar folding was observed when Pds5 was bound instead of Scc2 (Figure 6A, B). Though folding permits an association between the hinge and the N-terminal Scc2 sequences, which could in principle stabilise the folded conformation, we also observed similar, if not identical, folding in samples lacking all HAWK proteins (Figure 5—figure supplement 1E).

Figure 5 with 2 supplements see all
Folded cohesin allows interaction of hinge with Scc2 N-terminus.

(A) Views of cryo-EM reconstruction of Scc2-bound cohesin coloured by subunit. (B) Full pseudo-atomic model of folded cohesin trimer bound to Scc2. (C) Close-up of breaks in the coiled coils of Smc3 and Smc1 that constitute the elbow region of cohesin (PDB: 7OGT; EMD-12887). (D) Close-up of the interaction between the hinge and Smc3 that stabilises the folded state. (E) Close-up of Scc2 N-terminus in proximity of hinge residues K620 and D588Y. (FG) Comparison of cryo-EM densities between Scc2-bound and ATP-free cohesin seen in (F) (EMD-12880) and ATP-bound cohesin seen in (G) (EMD-12889), demonstrating that head engagement is not sufficient for coiled coil unzipping.

Figure 6 with 1 supplement see all
Pds5 binds to Smc3 head while contacting the hinge.

(A) Composite map of cryo-EM reconstructions of Pds5-bound cohesin (EMD-12888). (B) Full pseudo-atomic model of folded cohesin trimer bound to Pds5 coloured by subunit. (C) Close-up of interaction between hinge and Pds5 showing proximity of N-terminus of the HAWK to hinge residues D588 and K620. (D) 2D classes of Pds5-bound ATPase heads. (E) Close-up of Pds5 binding to K112- and K113-proximal region of the Smc3 head.

Coiled coil folding enables interaction of Scc2 or Pds5 with the Smc1 hinge

Initial 2D classes of Scc2-bound cohesin revealed floppiness not only within the HAWK, especially within its C-terminus, but also between the joint and the ATPase heads (Figure 5—figure supplement 1B). We therefore split the complex computationally into two regions, with a boundary at the joint, and processed their densities separately (Figure 5F). This yielded an overall resolution of 13 Å for the HAWK-bound part, which enabled fitting of a homologous Scc2 crystal structure (PDB: 5ME3; Chao et al., 2015) together with both head crystal structures (PDB: 1W1W; Haering et al., 2004; PDB: 4UX3; Gligoris et al., 2014) to produce a pseudo-atomic model. Analysis revealed that Scc2 binds rigidly to the Smc1 ATPase head in a manner resembling but distinct from its interaction in the clamped state (Figure 5—figure supplement 1BCollier et al., 2020). Scc2’s C-terminal HEAT repeats 18–24 (residues 1127–1493) dock onto Smc1’s F-loop (residues 1095–1118) as well as the emerging coiled coils, a mode of interaction analogous to that between condensin’s HAWK Ycs4 and Smc4 (Lee et al., 2020). This mode of interaction therefore takes place whether or not heads are engaged. Unlike the engaged and clamped head state, Scc2 makes no contact with Smc3 in the non-engaged, nucleotide-free structure. Contrary to its C-terminal part, Scc2’s N-terminal region adopts a range of conformations. Bending around the mid-region of Scc2 enables its N-terminus to contact the joint region of Smc3’s coiled coil in the clamped state. However, when heads are disengaged in our nucleotide-free structure, Scc2 is straightened and its N-terminal half adopts the conformation observed in crystals of Scc2 alone (Chao et al., 2015). Because cohesin’s elbow is further away from its hinge than is the case for condensin, folding of its coiled coils brings the hinge to within 12 nm of the ATPase heads. As a consequence, the N-terminal part of Scc2 molecules bound to Smc1 ATPase heads is in proximity to the hinge, thereby explaining not only its crosslinking to Smc1K620BPA in vivo but also how Smc1D588Y could circumvent the need for Scc4 (Figure 5E). We suggest that the addition of a bulky amino acid into Smc1 through D588Y may be sufficient to help bind an otherwise floppy Scc2 N-terminal domain, whose interaction with the hinge is normally stabilised by Scc4.

We processed data collected on Pds5-bound complexes in a similar manner, producing a 13 Å resolution structure, which revealed that Pds5 binds to Smc3 and not, like Scc2, to Smc1’s ATPase head domain (placed PDB: 5F0O; Lee et al., 2016Figure 6A, B). The contact takes place between the most C-terminal HEAT repeats of Pds5 and the top region of the N-terminal lobe of Smc3’s ATPase. Strikingly, this part of Smc3 contains the pair of highly conserved lysine residues K112 and K113 (Figure 6E), whose acetylation by Eco1 not only prevents releasing activity (Unal et al., 2008; Rolef Ben-Shahar et al., 2008) but also stabilises Pds5’s interaction with chromosomal cohesin complexes (Chan et al., 2012). Unlike Scc2, Pds5 does not rely on negatively charged amino acids for its interaction with the K112/K113 region and may therefore be better suited than Scc2 for binding the acetylated and less positively charged version of Smc3. Furthermore, binding in this manner shields both lysine residues when acetylated, hence explaining how Pds5 hinders de-acetylation during G2/M phases (Chan et al., 2013). Like Scc2, Pds5’s N-terminal HEAT repeats approach Smc1’s hinge domain, which explains the crosslinking to Smc1K620BPA in vivo (Figure 6C). The low resolution and flexibility apparent in our map mean that we cannot be sure whether Pds5’s C-terminal domain reaches beyond the hinge and contacts the coiled coils. Importantly, the modes of interaction of Scc2 and Pds5 with Smc subunits appear to be incompatible with each other, as has been postulated previously through in vivo and in vitro work (Petela et al., 2018).

Head engagement does not per se drive unzipping of cohesin’s coiled coil

A major difference between the apo state bound to Scc2 and the ATP-bound clamped state is the conformation of the Smc coiled coils. Though both folded, they are zipped up in the case of the former but splayed open up to the elbow in the case of the latter. Opening up could be driven by engagement per se. Alternatively, it might additionally require the binding of DNA to engaged heads in the presence of Scc2. In the course of our studies, we identified and solved with a resolution of 6 Å a form of cohesin lacking Scc2, Scc1, DNA, or crosslinker, whose ATPase heads were engaged in the presence of ATP (Figure 5G; EMD-12889). Contrary to previous studies with shortened constructs (Muir et al., 2020), which suggested that engagement per se might drive coiled coil unzipping, the coiled coils of our engaged heads are fully zipped up, at least from their joints to their hinge domains (Figure 5G). Thus, head engagement does not per se cause unzipping. Furthermore, 2D classes of heads-engaged cohesin bound to Scc2 demonstrate that addition of Scc2 to an ATP-bound state is insufficient to promote unzipping (Figure 5—figure supplement 1A). We therefore suggest that it is the binding of DNA to the surface on top of engaged heads that causes unzipping to make space for the DNA double helix, as well as the rearrangement of Scc2’s NTD necessary for its association with Smc3’s coiled coil.

Folding of cohesin’s coiled coils occurs in vivo and is a feature of sister chromatid cohesion

Though the interaction of Scc2 and Pds5 with Smc1’s hinge in vivo is fully consistent with coiled coil folding and vice versa, it does not prove that folding actually occurs in vivo. To address this, we identified regions of both Smc3 and Smc1 whose residues when substituted by cysteine should permit crosslinking by BMOE specifically if Smc1’s hinge interacted with Smc3’s coiled coil in the manner observed in our cryo-EM structure (Figure 7A). A pair of residues, Smc1R578C and Smc3V933C, were viable both as single and double mutants, and gave rise to efficient BMOE-dependent crosslinking between Smc1 and Smc3 in vivo only when combined (Figure 7B). Because the efficiency of crosslinking was 60% or even more, we conclude that a high fraction of cohesin complexes must be folded at the elbow in vivo.

Folding of cohesin’s coiled coils occurs in vivo and is a feature of sister chromatid cohesion.

(A) Sequence conservation analysis for the Smc3 coiled coil and Smc1 hinge helices shown in Figure 4D shows that the residues are highly conserved. (B) Whole-cell extract western blot analysis for the crosslink between Smc1R578C-HA6 and Smc3V933C-PK6 with single cysteine controls probing for hemagglutinin (HA) (top) and PK (bottom). A band shift is observed at the same molecular weight for both blots, confirming the identity of the crosslinked species. Crosslinking of the engaged heads (Chapard et al., 2019) was used as a positive control (K28401, K27359, K28585, K28546, K28583). (C) Western blot analysis of crosslinking measuring the folded state (Smc1R578C-HA6 and Smc3V933C-PK6) and Smc1-Smc3 hinge dimerisation (Haering et al., 2008) probing for acetylated Smc3 (top) and HA (bottom) in logarithmic or pheromone arrested cells (K26081, K28586).

If, as seems likely, Smc1D588Y bypasses the need for Scc4 by strengthening the interaction between the hinge and Scc2, then folding would appear to be a feature of cohesin complexes engaged in loading in vivo and the observation by cryo-EM that folding is a feature of the clamped state in vitro confirms this. To address whether folding is also a feature of cohesin complexes engaged in holding sister chromatids together, when the cohesin’s ATPase is thought to be inactive, we measured whether acetylated Smc3 molecules were also efficiently crosslinked to Smc1 using the cysteine pair that reports folding. Western blots using an antibody specific for acetylated Smc3 revealed that crosslinking between Smc3V933C and Smc1R578C was similar to that between a hinge cysteine pair (Haering et al., 2008). Because a large fraction of acetylated Smc3 was crosslinked to Smc1 in both cases (Figure 7C), we conclude that folding is a feature of many, if not most, cohesin complexes engaged in holding sister chromatids together. As expected, the Smc3Ac antibody failed to detect any protein in G1-arrested cells (Figure 7C), confirming its specificity.

Discussion

Folding occurs in vivo and is of functional importance

A major feature of all Smc-kleisin complexes, be they bacterial homodimers or eukaryotic heterodimers, are the 50-nm-long coiled coils connecting their hinge dimerisation domains to their ATPase heads. Recent structural and biochemical studies have revealed that the two coiled coils of Smc dimers have a strong tendency to self-associate or zip up throughout their length both in vitro and in vivo (Bürmann et al., 2019; Chapard et al., 2019; Diebold-Durand et al., 2017; Soh et al., 2015). In many cases, for example, MukBEF from Escherichia coli as well as the eukaryotic cohesin and condensin complexes, zipping up is accompanied by folding around an elbow, which leads to an association of hinges with sections of the coiled coil closer to the heads. Cryo-EM imaging suggests that complete zipping up may be an invariant property of apo-complexes. In the case of cohesin, clamping of DNA by Scc2 on top of engaged ATPase heads is accompanied by extensive unzipping (Collier et al., 2020). The finding reported here (Figure 5G), that a cohesin complex whose heads are engaged in the presence of ATP nevertheless possesses coiled coils that are extensively zipped up, suggests that unzipping is not caused by head engagement per se but instead by the binding of DNA to engaged heads. Indeed, crosslinking studies have confirmed that the coiled coils associated with engaged heads are at least sometimes zipped up even in vivo (Chapard et al., 2019). Whether DNA clamping causes unfolding as well as unzipping is a matter of considerable interest because it has been speculated that the folding and unfolding of Smc coiled coils might be a crucial aspect of ATP-driven mechanical cycle responsible for the translocation of Smc-kleisin complexes along DNA during LE (Hassler et al., 2018; Bürmann et al., 2019).

The observation that a large fraction of clamped complexes possess folded but extensively unzipped coiled coils suggests that unfolding is not a hard-wired response to clamping. This finding, along with the finding that some Smc coiled coils, for example, those in Bacillus subtilis, likely do not possess an elbow (Bürmann et al., 2019), the discovery that the hinges of cohesin and condensin occupy different positions, and the fact that folding has not hitherto been demonstrated in living cells all raised the possibility that the phenomenon might not in fact have an important physiological role. The work described here provides the first concrete evidence to the contrary. A 5.5 Å structure of cohesin’s coiled coils in a folded state enabled us to identify a pair of residues within the Smc1 hinge/Smc3 coiled coil interface, namely Smc1R578 and Smc3V933, whose substitution by cysteine led to efficient crosslinking by BMOE inside cells, demonstrating that folding takes place frequently in vivo. Because Smc1R578C/Smc3V933C crosslinking occurs when Smc3 is acetylated, folding would also appear to be a feature of complexes engaged in holding sister chromatids together. Unlike mammalian cells, yeast lack sororin and acetylation is strictly linked to replication and it is therefore a good marker for complexes associated with cohesion. Future crosslinking studies combining Smc1R578C/Smc3V933C with other cysteine pairs should make it possible to address whether folding is also a feature of other cohesin states in vivo, for example, while DNAs are clamped between Scc2 and engaged heads when cohesin loads onto and translocates along chromatin fibres.

That folding not only occurs but is of physiological importance stemmed from a very different approach; the isolation of extragenic mutations that suppress the loading defect and lethality caused by scc4 mutations. In addition to scc2 alleles (e.g. E822K) and mutations that affect the way histones H2A and H2B bind to nucleosomal DNA, substitution by tyrosine (or any other aromatic residue for that matter) of a conserved aspartate residue on the surface of Smc1’s hinge domain (Smc1D588Y) restored loading in scc4 mutants to levels that were in fact 30% higher than WT. Because smc1D588Y cannot suppress loss of Scc2 itself, the suppressor acts by enabling Scc2 to function without its auxiliary subunit and not by bypassing Scc2 entirely.

In vivo crosslinking using cohesin complexes in which a surface lysine residue nearby D588 within Smc1’s hinge was replaced by the non-canonical amino acid BPA (Smc1K620BPA) demonstrated that the hinge must be in proximity, at least some of the time, to Scc2, Scc3, and Pds5. Crucially, Scc4 facilitated crosslinking between Smc1K620BPA and Scc2 but inhibited that with Pds5, while smc1D588Y had a similar effect and compensated for loss of Scc4. To explain how Scc2 and Pds5, which bind to the Smc heads (Figure 5A and Figure 6A), are close enough to Smc1’s hinge domain for BPA-mediated crosslinking, we suggest that cohesin’s coiled coil must be folded not merely when cohesin holds sister chromatids together (Figure 7) but also during the loading reaction. This also explains how smc1D588Y exerts such a powerful effect on cohesin loading. Because Smc1K620BPA crosslinks to N-terminal sequences within Scc2 that are close to where Scc4 normally binds, we suggest that Scc4 helps Scc2 promote loading by facilitating the latter’s interaction with the Smc1 hinge, and that Scc4’s role can be substituted by insertion of an aromatic residue within the Smc1-Scc2 interface. How mechanistically Scc4 or Smc1D588Y facilitate interaction between the hinge and Scc2 while hindering that with Pds5 is presently unclear. The key point is that our genetic data confirm that the proximity between the N-terminal domain of Scc2 and Smc1 hinges observed in cryo-EM structures whether in the clamped (Collier et al., 2020; Shi et al., 2020) or ATP-free-state (Figure 5A) also occurs in vivo and more importantly has functional significance.

Though folding enables the hinge to interact with cohesin’s HAWK proteins, it is likely that the process has functions besides such interactions as folding appears to be more conserved than the HAWKS themselves. It has been suggested that an extension/folding cycle might have a role in cohesin’s translocation along DNA during LE (Bürmann et al., 2019). Another possibility is that by packing coiled coils on top of each other, folding helps to stabilise the zipping up of Smc1/3 coiled coils, which may have a role in ensuring that unzipping does not occur precociously, in other words, only when DNA is correctly clamped on top of engaged heads by Scc2. The notion that folding acts primarily to reinforce the zipped-up state helps explain why Smc proteins in organisms like B. subtilis do not appear to have an elbow around which their coiled coils are folded. Zipping up of B. subtilis Smc coiled coils might be strong enough that it does not need to be reinforced by folding. A third possibility is that, by bringing the hinge close to DNA clamped by Scc2 on top of engaged Smc1/3 ATPase heads, folding facilitates passage of DNA through a gate created by hinge opening, thereby mediating entrapment of DNA within S-K rings.

What is the function of Scc4?

In addition to recruiting the Scc2/4 complex to kinetochores and thereby promoting high rates of cohesin loading at CENs, Scc4 helps Scc2 promote cohesin’s efficient association with chromosome arms. It has previously been suggested that Scc4’s function involves the nucleosome remodelling complex RSC. Two rather different types of proposal have been made in separate papers by the Uhlmann group. According to the first, Scc2’s association with Scc4 enables RSC to create nucleosome-free regions necessary for cohesin loading; in other words, Scc2/4’s role is to positively regulate RSC’s cohesin loading activity (Lopez-Serra et al., 2014). This proposal is difficult to reconcile not only with our finding that RSC inactivation causes only modest, if any, defect in cohesin loading as measured by calibrated ChIP-seq but also with the now incontrovertible evidence that the Scc2/4 complex acts directly on cohesin, enabling it to clamp DNA on top of its Smc ATPase domains. Neither our ChIP-seq data nor recent structural work (Collier et al., 2020; Shi et al., 2020; Higashi et al., 2020) support the notion that that ‘Scc2/4 acts in sister chromatid cohesion by maintaining nucleosome-free regions’ (Lopez-Serra et al., 2014).

The second proposal shares with the first the notion that nucleosome-free regions created by RSC are necessary for cohesin loading but differs from the first in suggesting that by interacting with Scc4, RSC recruits Scc2 to regions that have been cleared of nucleosomes and permits Scc2 to catalyse their association with cohesin (Muñoz et al., 2019). Thus, instead of Scc2/4 activating RSC and thereby creating nucleosome-free regions needed for cohesin loading, RSC is envisioned to activate Scc2/4 by bringing it to nucleosome regions created by and associated with RSC. Deletion of Scc2’s NTD to which Scc4 binds is normally lethal, but fusion to RSC’s Sth1 subunit of a version of Scc2 (Scc2C) lacking its N-terminal Scc4-binding domain restores viability. Though this finding is consistent with the notion that Scc4’s function is merely to link Scc2 with RSC, the viability of Sth1-Scc2C fusions could equally well be explained if the artificially efficient recruitment of Scc2 to nucleosome-free regions (associated with RSC) enables Scc2 and cohesin to clamp DNA without the help of Scc4. Crucially, the notion that Scc4’s function is to connect Scc2 with RSC fails to explain why RSC inactivation causes only a modest, if any, defect in cohesin loading, and certainly not one comparable to that caused by Scc4 inactivation, nor how a hinge mutation (Smc1D588Y) that alters cohesin’s association with its HAWK regulatory subunits Scc2 and Pds5 is sufficient to bypass Scc4. One would have to suppose that Scc2/4’s recruitment to RSC involved not merely the latter’s association with Scc4 but also with cohesin, and Smc1D588Y acts by facilitating the interaction. It also provides no explanation for why mutations within Scc2 (E822K) that probably alter how the latter forms the clamped state also fully bypass Scc4.

Our finding that the major partners of the part of the hinge containing Smc1D588 are in fact cohesin’s HAWKs, principally Pds5 and Scc2, favours an alternative explanation, namely that Scc4 facilitates an interaction between the Scc2/4 complex and cohesin’s hinge that either stabilises Scc2’s association with cohesin and/or alters its conformation in a manner that enhances its ability to clamp DNA. It is nevertheless striking that the lethality of scc4 mutants can be bypassed, albeit less effectively than with smc1D588Y, by mutations in histones H2A and H2B that presumably reduce the affinity of their interaction with nucleosomal DNA and would therefore favour formation of nucleosome-free DNA. We suggest that nucleosome-free DNA is indeed important for loading because naked DNA is necessary for formation of the clamped state. However, creation of naked DNA is insufficient for efficient clamping in vivo. Association of Scc2 with cohesin’s hinge and/or a conformational change that is a consequence of this association is additionally required, a process normally facilitated by Scc4 but whose absence can be compensated by Smc1D588Y. In other words, association of Scc2/4 with cohesin’s hinge domain facilitates the binding of naked DNA to its engaged ATPase heads and it is ultimately the latter that promotes a productive association with chromatin. Our data do not exclude the possibility that RSC is in principle able to create the nucleosome-free DNA necessary for clamping, but whether it is normally necessary is at present unclear.

Why does smc1D588Y depress loading at CENs?

It is striking that while smc1D588Y facilitates cohesin’s association with chromosome arms, in the presence as well as the absence of Scc4, the mutation has the opposite effect on loading at CENs (Figure 2). To explain this paradox, we suggest that formation of the clamped state, which we propose is facilitated by smc1D588Y, is just the first step in cohesin’s productive association with and translocation along chromatin fibres and must be followed by a second step, likely involving ATP hydrolysis and head disengagement. Consistent with the notion that D588Y accelerates the first clamping step, we observed that despite lowering the overall level of pericentric cohesin arising from loading at CENs, smc1D588Y actually increased the amount of Scc2 associated with cohesin at CENs. The fact that it also increased the amount of cohesin containing Smc3E1155Q confirms that this population represents the clamped state. Because such complexes do not load productively (Hu et al., 2011), hydrolysis of ATP associated with clamped complexes must also be required for loading. In other words, a sequence of clamping, DNA loading, and unclamping while the DNA remains loaded, is necessary. If so, an important question is whether clamping or subsequent unclamping driven by ATP hydrolysis is rate-limiting during the loading process. We suggest that clamping is rate-limiting along chromosome arms but unclamping is rate-limiting at CENs and that this is the reason why smc1D588Y enhances arm loading while depressing loading at CENs.

In summary, we provide biochemical evidence that cohesin’s coiled coils are indeed folded in vivo and genetic evidence that folding is of physiological importance in loading cohesin onto chromosomes. Whether folding is regulated by cohesin’s ATPase cycle and cyclical unfolding has a role in DNA LE are important questions for the future.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Spodoptera frugiperda)Sf9 insect cellsThermo FisherCat# 11496015N/A
Genetic reagent (Saccharomyces cerevisiae)NCBITaxon:4932This paperYeast strainsSupplementary file 4
Biological sampleα-factor peptideCRUK Peptide Synthesis ServiceN/AN/A
AntibodyMouse monoclonal Anti-V5Bio-RadCat# MCA1360(1:1000)
AntibodyAnti-HA High Affinity (3F10) (Rat)RocheCat# 11867423001(1:1000)
AntibodyAnti-His (mouse)GenScriptCat# A00186(1:1000)
AntibodyAnti-c-Myc A-14 (9E10) (rabbit)Santa Cruz BiotechCat# sc-789(1:1000)
AntibodyAnti-Myc 4A6 (mouse)MilliporeCat# 05-724(1:1000)
AntibodyAnti-FLAG (rabbit)SigmaCat# F7425(1:1000)
Recombinant DNA reagentpACEbac1 2xStrepII-Scc2151-1493Collier et al., 2020N/AN/A
Recombinant DNA reagentpACEbac1 Smc1-8xHis-Smc3/pIDC Scc1-2xStrepII (trimer)Petela et al., 2018N/AN/A
Recombinant DNA reagentpIDS Pds5-FlagPetela et al., 2018N/AN/A
Commercial assay or kitTalon Superflow Metal Affinity ResinTakara Bio.Cat# 635669N/A
Commercial assay or kitNuPAGE 3–8% Tris-Acetate Protein gelsThermo FisherCat# EA0378BOXN/A
Commercial assay or kitTrans-Blot Turbo Midi0.2 µm Nitrocellulose Transfer PacksBio-RadCat# 1704159N/A
Commercial assay or kitProtein G DynabeadsThermo FisherCat# 300385N/A
Commercial assay or kitChIP DNA Clean and Concentrator kitZymo ResearchCat# D5205N/A
Commercial assay or kitNEBNext Fast DNA Library Prep Set for Ion TorrentNEBCat# Z648094N/A
Commercial assay or kitIon Xpress Barcode AdaptorsThermo FisherCat# 4471250N/A
Commercial assay or kitE-Gel SizeSelect II 2% Agarose gelsThermo FisherCat# G661012N/A
Commercial assay or kitKAPA Ion Torrent DNA standardsRocheCat# 07960395001N/A
Commercial assay or kitEnzChek phosphate assay kitThermo FisherCat# E6646N/A
Commercial assay or kitStrepTrap HPFisher ScientificCat# 11540654N/A
Commercial assay or kitSuperose 6 Increase10/300 GLVWRCat# 29-0915-96N/A
Chemical compoundNocodazoleSigmaCat# M1404N/A
Chemical compoundBismaleimidoethane (BMOE)Thermo FisherCat# 223235 mM
Chemical compoundComplete EDTA-free protease inhibitor cocktailRocheCat# 4693132001(1:50 mL)
Chemical compoundPMSFSigmaCat# 031158360011 mM
Chemical compoundImmobilon Western ECLMilliporeCat# WBLKS0500N/A
Chemical compoundRNase ARocheCat# 10109169001N/A
Chemical compoundProteinase KRocheCat# 03115836001N/A
Chemical compoundBPABachemCat# 4017646.0005N/A
Chemical compoundTCEPThermo FisherCat# 20490N/A
Chemical compoundDesthiobiotinFisher ScientificCat# 12753064N/A
Software, algorithmFastQCBabraham Bioinformaticshttps://www.bioinformatics.babraham.ac.uk/projects/fastqc/N/A
Software, algorithmFastx_trimmerHannon Labhttp://hannonlab.cshl.edu/fastx_toolkit/index.htmlN/A
Software, algorithmFilterFastq.pyPetela et al., 2018https://github.com/naomipetela/nasmythlab-ngsN/A
Software, algorithmBowtie2Langmead and Salzberg, 2012http://bowtie-bio.sourceforge.net/bowtie2/index.shtmlN/A
Software, algorithmSamtoolsSamtoolshttp://www.htslib.orgN/A
Software, algorithmIGB browserNicol et al., 2009https://www.bioviz.orgN/A
Software, algorithmchr_position.pyPetela et al., 2018https://github.com/naomipetela/nasmythlab-ngsN/A
Software, algorithmfilter.pyPetela et al., 2018https://github.com/naomipetela/nasmythlab-ngsN/A
Software, algorithmBcftools callSamtoolshttp://www.htslib.orgN/A
Software, algorithmMutationFinder.pyPetela et al., 2018https://github.com/naomipetela/nasmythlab-ngsN/A
Software, algorithmyeastmine.pyPetela et al., 2018https://github.com/naomipetela/nasmythlab-ngsN/A
Software, algorithmRELION 3.1doi:10.1016/j.jsb.2012.09.006N/AN/A
Software, algorithmCtfFind4doi:10.1016/j.jsb.2015.08.008N/AN/A
Software, algorithmCrYOLO 1.5doi:10.1038/s42003-019-0437N/AN/A
Software, algorithmChimerahttps://www.cgl.ucsf.edu/chimera/N/AN/A
Software, algorithmChimeraX 1.0https://www.cgl.ucsf.edu/chimera/N/AN/A
Software, algorithmCOOTdoi:10.1107/S0907444910007493N/AN/A
Software, algorithmMAINdoi:10.1107/S0907444913008408N/AN/A
Software, algorithmPhenix.real_ space_refinementdoi:10.1107/S2059798318006551N/AN/A
Software, algorithmPYMOL 2https://pymol.org/2/N/AN/A
Software, algorithmSWISS-MODELhttps://swissmodel.expasy.orgN/AN/A
OtherQuantifoil R 2/2 grid: Cu/Rh 200 cryoEM gridsQuantifoil GmbHN/AN/A
Table of structures
Map description and file name in ‘coordinates and maps’First appearance in figuresDatabase accession code
Elbow EM mapFigure 5FEMD-12887
Elbow coordinate mapFigure 5BPDB ID 7OGT
Scc2 bound to ATPase headsFigure 5FEMD-12880
Scc2 bound to ATPase heads masking hinge and N-terminusFig 5—figure supplement 1C
Pds5 bound to ATPase headsFigure 6AEMD-12888
Engaged ATPase headsFigure 5GEMD-12889
Mouse hinge D574YFigure 1DPDB ID 7DG5

Yeast strains and growth conditions

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All yeast strains were derived from W303 and grown in rich medium (YEP) supplemented with 2% glucose (YPD) at 25°C unless otherwise stated. Cultures were agitated at 200 rpm (Multitron Standard, Infors HT). Strain numbers and relevant genotypes of the strains used are listed in the Key resources table, Supplementary file 4. To arrest the cells in G1, α-factor was added to a final concentration of 2 mg/L/h, every 30 min for 2.5 hr. Release was achieved by filtration wherein cells were captured on 1.2 μm filtration paper (Whatman GE Healthcare), washed with 1 L YPD, and resuspended in the appropriate fresh media. To arrest the cells in G2, nocodazole (Sigma) was added to the fresh media to a final concentration of 10 μg/mL and cells were incubated until the synchronisation was achieved (>95% large-budded cells). To inactivate temperature-sensitive alleles, fresh media were pre-warmed prior to filtration (Aquatron, Infors HT). To produce cells deficient in Pds5 using the AID system, cells were arrested with α-factor as described above. 30 min prior to release, auxin was added to 5 mM final concentration. Cells were then filtered as described above and released into YPD medium containing 5 mM auxin.

Screening for suppressors of scc4-4

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Forty independent colonies of the parental strain (YCplac33::scc4-4::NATMX scc4Δ::HIS3 [K23967]) were picked and grown overnight at 25°C. Each was plated at 5 OD600 units per plate over three plates and incubated at 35.5°C until colonies appeared. Up to three colonies were picked from each plate and streaked for single colonies at 25°C before being retested for growth at 35.5°C. Those that grew at 35.5°C were checked by PCR from genomic DNA preparations for revertants of Scc4. Isolated suppressors that did not show revertant mutations were checked for 2:2 segregation and grouped into complementation groups prior to deep sequencing. To check for the ability to rescue the deletion of Scc4, suppressors were streaked onto 1 mg/mL 5-FOA plates and allowed to grow for 2 days.

Protein purification from E. coli

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BL21(DE3) strains containing plasmids encoding proteins for purification were grown at 37°C in 2XTY media supplemented with the appropriate antibiotic until an OD600 of 0.6 was reached. Expression was induced by addition of IPTG to a concentration of 1 mM for 16 hr at 20°C. Cells were harvested by centrifugation and mixed with five times the cell pellet volume of lysis buffer (250 mM NaCl, 50 mM Tris-HCl pH 7.5, 1 mM EDTA, 2 mM β-mercaptoethanol, 1 tablet/50 mL protease inhibitor cocktail [Roche]). Cells were lysed by passage through a cell disruptor (Constant Systems) at 20 kpsi. PMSF (Sigma) was added to the lysate to a final concentration of 1 mM. Samples were sonicated on ice for 1 min/50 mL in 30 s intervals with a Vibra-cell sonicator (VCX 130FSJ; Sonics and Materials) at 80% amplitude. Lysate was cleared by centrifugation at 50,000 g for 90 min in an Avanti J-26S XP centrifuge (Beckman Coulter).

Affinity purification was performed by incubating cleared lysates with pre-equilibrated Talon Superflow Metal Affinity Resin (500 μL/50 mL lysate; Takare Bio) for 1 hr at 4°C. Beads were sedimented by centrifugation at 700 g for 2 min and the supernatant decanted and discarded. The resin was washed five times with 50 mL lysis buffer containing 10 mM imidazole. Proteins wereeluted with a volume of elution buffer (lysis buffer +250 mM imidazole) equal to twice the volume of resin.

Size exclusion chromatography was performed by injecting up to 2 mL of eluent onto a HiLoad 16/60 Superdex 200 prep grade column (equilibrated with Buffer A: 95 mM NaCl, 20 mM HEPES pH 7.5, 2 mM β-mercaptoethanol) connected to an ÄKTApurifier 100 purification system controlled by UNICORN software (GE Healthcare). Peak fractions were concentrated using Vivaspin columns (Sartorius Stedim Biotech) with a molecular weight cutoff of 10 kDa. Concentration of purified protein was determined based on its absorbance at 280 nm, measured using a NanoDrop-1000 (Thermo Fisher Scientific).

In vitro hinge binding assay

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WT and mutant MBP-Smc1hinge-HIS6 and Smc3hinge-FLAG3-HIS6 proteins were expressed and purified as described above. Proteins were mixed at an equimolar concentration of 250 mM in Buffer A and incubated at 16°C with shaking at 1000 rpm for 15 min. 400 μL of protein mixture was added to 20 μL of pre-equilibrated ANTI-FLAG M2 affinity gel and incubated at 16°C with shaking at 1000 rpm for 15 min. Resin was sedimented by centrifugation for 1 min at 850 g and washed three times with Buffer A + 1% Triton X-100 (Sigma) before being boiled in 2× SDS sample buffer prior to immunoblotting.

Protein gel electrophoresis and western blotting

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The samples were mixed with 4× LDS sample buffer (NuPAGE Life Technologies), loaded onto 3–8% Tris-acetate gels (NuPAGE, Life Technologies) and the proteins separated using an appropriate current. The proteins were then transferred onto 0.2 μm nitrocellulose using Trans-blot Turbo transfer packs for the Trans-blot Turbo system (Bio-Rad). The following antibodies were used: anti-V5 (Bio-Rad), anti-HA (Roche), His-tag antibody (GenScript), and A-14 (Santa Cruz Biotech). For visualisation, the membrane was incubated with Immobilon Western Chemiluminescent HRP substrate (Millipore) before detection using an ODYSSEY Fc Imaging System (LI-COR).

Multiple sequence alignment

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Multiple sequence alignments were created using Clustal Omega (Sievers et al., 2011).

Calibrated ChIP-seq

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Cells were grown exponentially to 0.5 OD600 and the required cell cycle stage where necessary. 15 OD600 units of S. cerevisiae cells were then mixed with 5 OD600 units of Candida glabrata to a total volume of 45 mL and fixed with 4 mL of fixative (50 mM Tris-HCl, pH 8.0; 100 mM NaCl; 0.5 mM EGTA; 1 mM EDTA; 30% [v/v] formaldehyde) for 30 min at room temperature (RT) with rotation. Fixation was quenched with 2 mL of 2.5 M glycine incubated at RT for 5 min with rotation. The cells were then harvested by centrifugation at 3500 rpm for 3 min and washed with ice-cold 1× PBS. The cells were then resuspended in 300 μL of ChIP lysis buffer (50 mM HEPES-KOH, pH 8.0; 140 mM NaCl; 1 mM EDTA; 1% [v/v] Triton X-100; 0.1% [w/v] sodium deoxycholate; 1 mM PMSF; 1 tablet/25 mL protease inhibitor cocktail [Roche]) and an equal amount of acid-washed glass beads (425–600 μm, Sigma) added before cells were lysed using a FastPrep−24 benchtop homogeniser (M.P. Biomedicals) at 4°C (3 × 60 s at 6.5 m/s or until >90% of the cells were lysed as confirmed by microscopy).

The soluble fraction was isolated by centrifugation at 2000 rpm for 3 min, then sonicated using a Bioruptor (Diagenode) for 30 min in bursts of 30 s 'on' and 30 s 'off' at high level in a 4°C water bath to produce sheared chromatin with a size range of 200–1000 bp. After sonication, the samples were centrifuged at 13,200 rpm at 4°C for 20 min and the supernatant was transferred into 700 μL of ChIP lysis buffer. 30 μL of protein G Dynabeads (Thermo Fisher) were added, and the samples were pre-cleared for 1 hr at 4°C. 80 μL of the supernatant was taken as the whole-cell extract (WCE) and 5 μg of antibody (anti-PK; Bio-Rad) was added to the remaining supernatant which was then incubated overnight at 4°C. 50 μL of protein G Dynabeads were then added and incubated at 4°C for 2 hr before washing 2× with ChIP lysis buffer, 3× with high salt ChIP lysis buffer (50 mM HEPES-KOH, pH 8.0; 500 mM NaCl; 1 mM EDTA; 1% [v/v] Triton X-100; 0.1% [w/v] sodium deoxycholate; 1 mM PMSF), 2× with ChIP wash buffer (10 mM Tris-HCl, pH 8.0; 0.25 M LiCl; 0.5 % NP-40; 0.5% sodium deoxycholate; 1 mM EDTA; 1 mM PMSF), and 1× with TE pH 7.5. The immunoprecipitated chromatin was then eluted by incubation in 120 μL of TES buffer (50 mM Tris-HCl, pH 8.0; 10 mM EDTA; 1% SDS) for 15 min at 65°C and the collected supernatant termed the IP sample. The WCE extracts were mixed with 40 μL of TES3 buffer (50 mM Tris-HCl, pH 8.0; 10 mM EDTA; 3% SDS), and all samples were de-crosslinked by incubation at 65°C overnight. RNA was degraded by incubation with 2 μL RNase A (10 mg/mL; Roche) for 1 hr at 37°C, and protein was removed by incubation with 10 μL of proteinase K (18 mg/mL; Roche) for 2 hr at 65°C. DNA was purified using ChIP DNA Clean and Concentrator kit (Zymo Research).

Extraction of yeast DNA for deep sequencing

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Cultures were grown to exponential phase (OD600 = 0.5). 12.5 OD600 units were then collected and diluted to a final volume of 45 mL before fixation as described in the protocol for ChIP-seq. The samples were treated as specified in the ChIP-seq protocol up to the completion of the sonication step whereby 80 μL of the samples were carried forward and treated as WCE samples.

Preparation of sequencing libraries

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Sequencing libraries were prepared using NEBNext Fast DNA Library Prep Set for Ion Torrent Kit (New England Biolabs) according to the manufacturer’s instructions. To summarise, 10–100 ng of fragmented DNA was converted to blunt ends by end repair before ligation of the Ion Xpress Barcode Adaptors. Fragments of 300 bp were then selected using E-Gel SizeSelect2% Agarose gels (Life Technologies) and amplified with 6–8 PCR cycles. The DNA concentration was determined by qPCR using Ion Torrent DNA standards (Kapa Biosystems) as a reference. 12–16 libraries with different barcodes could then be pooled together to a final concentration of 350 pM and loaded onto the Ion PI V3 Chip (Life Technologies) using the Ion Chef (Life Technologies). Sequencing was performed on the Ion Torrent Proton (Life Technologies), typically producing 6–10 million reads per library with an average read length of 190 bp.

Data analysis, alignment, and production of BigWigs

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Quality of the reads was assessed using FastQC and trimmed as required using fastx_trimmer. Generally, this involved removing the first 10 bases and any bases after the 200th, but trimming more or fewer bases may be required to ensure the removal of kmers and that the per-base sequence content is equal across the reads. Reads shorter than 50 bp were removed using ‘FilterFastq.py’ and the remaining reads aligned to the necessary genome(s) using Bowtie2 with the default (--sensitive) parameters (Langmead and Salzberg, 2012).

To generate alignments of reads that uniquely align to the S. cerevisiae genome, the reads were first aligned to the C. glabrata (CBS138, Génolevures; Dujon et al., 2004) genome with the unaligned reads saved as a separate file. These reads that could not be aligned to the C. glabrata genome were then aligned to the S. cerevisiae (sacCer3, SGD) genome and the resulting BAM file converted to BigWigs for visualisation. Similarly, this process was done with the order of genomes reversed to produce alignments of reads that uniquely align to C. glabrata.

Visualisation of ChIP-seq profiles

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The resulting BigWigs were visualised using the IGB browser (Nicol et al., 2009). To normalise the data to show quantitative ChIP signal, the track was multiplied by the sample's OR and normalised to 1 million reads using the graph multiply function.

In order to calculate the average occupancy at each base pair up to 60 kb around all 16 centromeres, the BAM file that contains reads uniquely aligning to S. cerevisiae was separated into files for each chromosome and a pileup of each chromosome was then obtained using samtools mpileup. These files were then amended using our own script ‘chr_position.py’ to assign all unrepresented genome positions a value of 0. Each pileup was then filtered using another in-house script ‘filter.py’ to obtain the number of reads at each base pair within up to 60 kb intervals either side of the centromeric CDEIII elements of each chromosome. The number of reads covering each site as one successively moves away from these CDEIII elements could then be averaged across all 16 chromosomes and calibrated by multiplying by the samples OR and normalising to 1 million reads. All scripts written for this analysis method are available on request.

Identification of mutations from whole genome sequencing

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Pileups were created using samtools mpileup (-v --skip-indels –f sacCer3.fa –o sample name.vcf sample name.bam), then SNPs were called using bcftools call (-v –c –o sample name.bcf sample name.vcf). To find mutations unique to a suppressor strain, lists of SNPs from the parental strain or backcrossed clones of the suppressor strain were compared to the list of SNPs from the suppressor strain. In the case of parental strains, mutations that were present in both were removed, and in the case of backcrossed clones of the suppressor strain, mutations that were present in both were kept in order to identify the mutation that caused the suppression phenotype. This was done using ‘MutationFinder.py’ and the resulting lists further narrowed using ‘yeastmine.py’ which searches the Saccharomyces Genome Database (SGD) for genes that correspond to the position of each mutation so that those that lie outside of genes could be removed. From this it was possible to identify the mutation in each suppressor that gave rise to the suppressor phenotype.

ATPase assay

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ATPase activity was measured by using the EnzChek phosphate assay kit (Invitrogen) by following the provided protocol. Cohesin in various complexes were mixed to a final concentration of 50 nM in under 50 mM NaCl in the presence of 700 nM 40 bp dsDNA in those experiments testing the effect of duplex DNA. The reaction was started with addition of ATP to a final concentration of 1.3 mM, always in a final volume of 150 μL. ATPase activity was measured by recording absorption at 360 nm every 30 s for 1 hr 30 min using a PHERAstar FS. ΔΑU at 360 nm was translated to Pi release using an equation derived by a standard curve of KH2PO4 provided with the EnzChek kit and according to instructions. The reactions were assumed linear for at least the first 10 min of the experiment and rates calculated using this time period. On completion, a fraction of each reaction was analysed by SDS-PAGE and the gel stained with Coomassie Brilliant Blue in order to test that the complexes were intact throughout the experiment and that equal amounts were used when testing various mutants and conditions. At least two independent biological experiments were performed for each experiment.

Cohesin protein expression and purification

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WT or 6C cohesin trimer, Scc2, and Pds5 were expressed and purified as described in Collier et al., 2020 and Bürmann et al., 2019, respectively. In brief, vectors containing S. cerevisiae cohesin trimers were generated by combining pACEbac1 SMC1-His SMC3 containing the 6C cysteine mutations (Smc1K639C-Smc3E570C, Smc1G22C-Scc1A547C, and Smc3S1043C-Scc1C56) with pIDC SCC1-2xStrepII by a Cre recombinase reaction (New England Biolabs). Sequences of S. cerevisiae Scc2 and Pds5 were individually cloned as 2xStrepII-(151-1493)Scc2 and 2xStrepII-Pds5 into Multibac vectors, yielding 2xStrepII-∆N150-Scc2-pACEbac1 and 2xStrepII-Pds5-pACEbac1 with an HRV 3C protease site (LEVLFQ/GP) in the tag linker. Expression of the 6C trimer, Scc2, and Pds5 was done individually in Sf9 insect cells followed by the same previously described three-step purification protocol: proteins were purified via affinity pulldown of their StrepII and eluted with desthiobiotin, 3C protease was added to the eluents to cleave the affinity tags, the cleavage products were further purified by anion exchange columns, and finally buffer exchanged to Buffer 6C (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 1 mM TCEP, 10% glycerol). The purified trimer, Scc2, and Pds5 proteins were then frozen in liquid nitrogen and stored at −80°C until further use.

Cryo-EM grid preparation

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For imaging of cohesin with cryo-EM, the purified 6C trimer and Scc2 or Pds5 were mixed at a 1:1.5 molar ratio and injected onto a Superose 6 Increase 3.2/300 column (GE Healthcare) in buffer containing 25 mM HEPES-NaOH pH 7.5, 150 mM NaCl, 1 mM TCEP. The tetramer fraction was incubated with 2 mM BMOE for 3 min at room temperature, and then buffer exchanged into buffer 6C with Zeba spin buffer exchange columns (Sigma Aldrich). For ATP-containing sample 5 mM ATP, 2 mM MgCl2 was added to the buffers.

Grids were prepared by applying 3 μL of sample at a concentration of 0.2–0.3 mg/mL to freshly glow-discharged Cu/Rh 2/2 holey carbon 200 mesh grids (Quantifoil). The grids were blotted for 1.5–2 s at 4°C with humidity at 100% and were flash frozen using a Vitrobot (Thermo Fisher Scientific).

Cryo-EM data collection, processing, and modelling

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Images were recorded on a Titan Krios electron cryo-microscope (FEI) equipped with a K2 or K3 summit direct electron detector with the use of a Volta phase plate (VPP) and varying pixel sizes between 1.09 and 1.16 Å/pixel. Micrographs were collected with total doses of ~40 electrons per Å2, dose-fractionated into 40 movie frames, and at defocus ranges of 0.5–0.9 μm. All datasets containing the same sample were merged as described by Wilkinson et al., 2019, resulting in a final pixel size of 1.16 Å. Image processing was done in RELION 3.0 (Zivanov et al., 2018) and cryoSPARC (Punjani et al., 2017). Movies were aligned using 5 × 5 patches using MotionCor2 with dose-weighting (Zheng et al., 2017). CTF parameters were estimated with Gctf (Zhang, 2016). All refinements were performed using independent data half-sets (gold-standard refinement) and resolutions were determined based on the Fourier shell correlation (FSC = 0.143) criterion (Rosenthal and Henderson, 2003). Due to the elongated shape of cohesin, particle picking was done with the help of the machine learning-based crYOLO software (Wagner et al., 2019). Initial 2D classifications and the first initial model made with cryoSPARC revealed intrinsic flexibility between the upper part of the complex, containing the hinge and the coiled coils, and the lower part, containing the HAWK-bound heads. Therefore, after an initial round of 3D refinement, the two parts were extracted and re-centred separately for all downstream processing. Specific EM processing strategies are discussed in detail in Figure 5—figure supplement 2. All depictions of these structures within the paper were made with the use of UCSF ChimeraX (Goddard et al., 2018).

To produce the coordinate map of the folded elbow, a homology model of the yeast hinge dimer was obtained from SWISS-MODEL (Waterhouse et al., 2018) using a crystal structure of the hinge from Mus musculus (PDB: 2WD5) as the template (Kurze et al., 2011). MAIN (Turk, 2013), and COOT (Emsley et al., 2010) were used for manual rebuilding, followed by refinement using Phenix.real_space_refinement (Hu, 2018). Manual rebuilding and refinement were repeated for several cycles.

In vivo photo crosslinking

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Yeast stains bearing TAG-substituted Smc1-myc9 plasmid and pBH61 were grown in −Trp −Leu SD medium containing 1 mM BPA. Cells were collected and resuspended in 1 mL of ice-cold PBS buffer. The cell suspension was then placed in a Spectrolinker XL-1500a (Spectronics) and irradiated at 360 nm for 2 × 5 min. Extracts were prepared as described previously (Hu et al., 2011) and 5 mg of protein were incubated with 5 μL of Anti-PK antibody (Bio-Rad) for 2 hr at 4°C. Next, 50 μL of Protein G Dynabeads (Thermo Fisher) were added and incubated overnight at 4°C to immunoprecipitate Scc1. After washing five times with lysis buffer, the beads were boiled in 2× SDS-PAGE buffer. Samples were run on a 3–8% Tris-acetate gel (Life Technologies) for 3.5 hr at 150 V. For western blot analysis, anti-Myc (Millipore), anti-FLAG (Sigma), and anti-HA (Roche) antibodies were used to probe the indicated proteins.

In vivo cysteine crosslinking

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15 OD units of cells grown in exponential phase were washed with ice-cold PBS and kept on ice throughout the experiment. Cells were resuspended in 500 µL cold PBS and 300 µL was added to 2 × 2 mL bead beater tubes. 12.5 µL BMOE (125 mM in DMSO to a final concentration of 5 mM) or 12.5 µL DMSO was added before incubating on ice for 6 min. Cells were washed twice with 1 mL cold PBS containing 5 mM DTT.

Crosslinked cells were resuspended in 500 µL lysis buffer (150 mM NaCl; 5 mM EDTA; 0.5% [v/v] NP40; 500 mM Tris-HCl, pH 7.5; 1 mM PMSF; 1 tablet/50 mL protease inhibitor cocktail [Roche]; 1 mM DTT) and an equal volume of acid-washed glass beads (425–600 μm, Sigma) was added. The cells were lysed using a FastPrep−24 benchtop homogeniser (M.P. Biomedicals) at 4°C for 3 × 60 s at 6.5 m/s with a 5 min rest between cycles, until >90% of the cells were lysed as confirmed microscopically. The insoluble fraction was pelleted by centrifugation at 13,200 rpm for 10 min and the supernatant isolated and analysed by western blot.

Data and software availability

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All scripts written for this analysis method are available to download from https://github.com/naomipetela/nasmythlab-ngs (copy archived at https://archive.softwareheritage.org/swh:1:rev:d7509c6f3e0a0f34db71b485a9e332223084e7be). The accession number for the next-generation sequencing data (raw and analysed) reported in this paper is GSE167318.

Data availability

PDB validation reports of the crystal structures are included in the manuscript. All scripts written for this analysis method are available to download from https://github.com/naomipetela/nasmythlab-ngs (copy archived at https://archive.softwareheritage.org/swh:1:rev:d7509c6f3e0a0f34db71b485a9e332223084e7be).

References

    1. Hu B
    (2018)
    ATP hydrolysis is required for relocating cohesin from sites occupied by its Scc2/4 loading complex
    Current Biology : CB.

Decision letter

  1. Adèle L Marston
    Reviewing Editor; University of Edinburgh, United Kingdom
  2. Cynthia Wolberger
    Senior Editor; Johns Hopkins University School of Medicine, United States
  3. Adèle L Marston
    Reviewer; University of Edinburgh, United Kingdom
  4. Daniel Panne
    Reviewer; University of Leicester, United Kingdom

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This paper is of interest for biologists studying SMC proteins and their role in various aspects of genome biology. The paper demonstrates that folding of cohesin's coil-coil, which previously had been demonstrated to occur in vitro, actually occurs in cells. Through a series of biochemical and structural studies, the authors demonstrate that such folding likely enables DNA loading by enabling an association with SCC2, the cohesin loader. The underlying mechanism, and if such folding has a role in DNA loop extrusion, remains unclear.

Decision letter after peer review:

Thank you for submitting your article "Folding of cohesin's coiled coil is important for Scc2/4-induced association with chromosomes" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, including Adèle L Marston as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Cynthia Wolberger as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Daniel Panne (Reviewer #2).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

Specific points related to these 4 essential revisions and which should be addressed are given in the individual reviews below.

1) Please provide full details of how the structural analyses were done to allow a full review of the relevant parts of the paper. The authors should provide clarity on which maps and models the authors intend to submit to the databases (EMDB and PDB). A table should be provided indicating what the files are and where they appear in the updated manuscript. They should be aligned (PDB with corresponding maps, where applicable) so that their fitting can be evaluated. More details of specific information that should be included is provided in the comments from reviewers #2 and 3.

2) A key point of the paper is that the SMC1/3 hinge interacts directly with Scc2. The authors provide cross-linking data as evidence that this occurs in vivo. However some of the cross linking data are not always convincing as some controls are missing. FigS4F is lacking the WT control. Data shown in Fig4G is also not fully convincing: Cys crosslinks should only occur if both SMC1 and SCC2 carry the Cys mutation. The BPA crosslink shown in Fig4E is in principle sufficient (but the Figure legend needs to be revised as it it shows SCC2 not PDS5 cross linking). In addition, please provide a full description of how these experiments were done in the methods section. (Specific points from all three reviewers).

3) The paper covers a lot of ground and is therefore difficult to follow in places. In revising the paper, it would be helpful to focus on the main narrative and remove some of the discussion that is not directly relevant to the main conclusions. (Specific points from all three reviewers).

4) The section on the requirement for nucleosome free regions for cohesin loading should be revised. Some of the data presented is inconclusive and while the authors do not find evidence for a role of RSC in cohesin loading, in contrast with an earlier report, this message is a distraction from the more exciting main narrative. It could therefore be given less attention as suggested in specific points from reviewers #1 and 2.

Reviewer #1:

In this manuscript, the authors address the mechanism of how cohesin regulators regulate its loading onto chromosomes. Cohesin loading requires the Scc2 protein, which forms a complex with Scc4. Previous work had demonstrated that the role of Scc4 is two-fold. First, it specifically targets cohesin loading to centromeres. Second, it activates Scc2/cohesin in some unknown way to facilitate cohesin loading genome-wide. Here the authors address this second function of Scc4, starting by isolating suppressors of the scc4-4 temperature sensitive mutation. This leads them to a mutation in the SMC hinge (smc1D588Y). This was initially surprising since Scc2 binds cohesin close to the ATPase heads which are distant from the hinge when cohesin is in its extended, rod-like conformation. However, a beautiful cryo-electron microscopy structure of cohesin in complex with Scc2 revealed that cohesin bending at the elbow (as has been observed in other SMC complexes) brings the hinge into close contact with Scc2 and the Smc3 coiled-coil. Using engineered cysteines and a cross-linking approach, the authors also provide convincing evidence that cohesin folding occurs in vivo. Using the same approach, they show that smc1D588Y stabilizes the folded conformation in complex with Scc2, while impairing association with Pds5, a mutually exclusive cohesin maintenance factor. The authors surmise that smc1D588Y allows cohesin loading without Scc4 by favouring the interaction between Scc2 and the Smc1/Smc3 hinge.

In addition to this main narrative, the authors report several additional findings. These include isolation of additional suppressors of the scc4-4 mutation which map to histone residues that interact with DNA. Prompted by this discovery, the authors re-examine previous work which reported a role for the chromosome remodeller, RSC, in cohesin recruitment to chromosomes. While the authors find no strong evidence for a direct role of RSC in cohesin recruitment, the relationship between histones/remodellers and cohesin loading remains unclear and this part of the manuscript is not necessary for the main message of the manuscript.

The strength of this manuscript is the range of complementary approaches used to dissect a central part of the mechanism of a central player in chromosome biology. Experiments are rigorously designed and appropriately interpreted. The manuscript contains a lot of detailed information, some peripheral to the main message, which may challenge readers not immersed in the field. The manuscript may benefit from some streamlining/reorganisation to ensure the key messages are highlighted for the general reader.

Comments for the authors:

1. The authors should considering shortening the section on RSC describing Figure S3 as it detracts from the main message of the manuscript. In addition, while the ChIP-seq in Figure S3D argue against a major role for RSC in cohesin loading, the nucleosome mapping data in Figure S3S are not conclusive, as presented. Why do the upper two and lower two nucleosome profiles look different from each other? Potentially effects on nucleosome positioning will only be visible in metagene analysis: the sth1-3 mutant at least would be expected to alter nucleosome positioning but this is not apparent from the data presented. The authors should either remove this data or perform additional analysis.

2. Figure 7G: The conclusion that cohesin folding at the elbow is a feature of cohesin complexes that are holding sister chromatids together relies on the assumption that all acetylated cohesin is participating in cohesion. However, this may not be the case. In mammals, cohesin acetylation is also observed in G1, indicating that it may have a more general role. This potential caveat should be mentioned.

3. It is surprising that there is high Scc1 ChIP-seq signal at centromeres in scc4-4 cells if Scc4 is required for cohesin loading at centromeres (Figure S4A). How do the authors explain this?

4. Line 273: It looks like smc1D588Y CAN rescue the proliferation defect of scc2-4 in Figure S1E.

5. Figure 3A please show proliferation data for the different histone mutants.

6. Line 402: Smc3 acetylation

7. Supplementary Figure 4F: needs a negative control without cysteine substitutions.

8. Page 19: Line 547: Call to Figure S4D is incorrect.

Reviewer #2:

Patela et al. aim to understand how folding of 50nm long coiled-coils contributes to cohesin function. Using a combination of genetics and crosslinking experiments, the authors demonstrate that a mutation in the cohesin hinge enables cohesin loading in the absence of Scc4 and that the hinge directly interacts with Scc2 or Pds5. CryoEM structures confirm that the coiled-coils fold back into position in which the SMC hinges interact with the coiled-coil. Such a conformation is also detected in cross-linking experiments in cells, thus supporting the view that the hinges are frequently positioned in close proximity of Scc2 or PDS5. This study thus presents an important step forward in understanding how DNA loading is achieved by the cohesin complex. The described mechanism may be important for the control of loop extrusion or sister chromatid cohesion.

The conclusions are mostly well supported by the data, but some aspects of the biochemical and structural data require clarification.

1. Biochemical data: Figure 4B: The FLAG blot indicates that x-linked species migrate at different positions on SDS-PAGE while the top panel shows uniform mobility. How do we know that the band labeled SMC1-? corresponds to x-linked SMC1 and not some non-specific anti-myc background (e.g. there is an additional non-labeled band running lower)? Figure 4B: Different amounts of FLAG-HAWKs are recovered in the anti-Myc IP. Can the authors generate an 'input' blot showing expression levels to exclude the possibility that the variable crosslinking efficiency is not due to different expression levels?

2. Figure S4F: Show a WT negative control (no Cys mutation) to confirm that the bands observed do not arise due to non-specific background cross-linking.

3. Figure 4G: Top: Why is there a crosslinked band in the lane containing Smc1K620C alone? Would it not be expected to see cross-linking only when both Scc2N200C and Smc1K620C are present? Why is the non-specific crosslinking (labeled as *) present everywhere? Which cross-linked band they are referring to? While there is a small amount of a band visible above the non-specific band, this weak band is also present in Smc1K620C alone!

4. Structural data: Figure 5A: The authors need to clarify if in their cryoEM structure, the SMC ATPase heads are in the 'apo' state, as indicated in the text or in the engaged state as indicated in the Discussion (L. 641).

5. The authors need to explicitly state in their description of their structural data (Line 512 et seq) that they used a crosslinked version of the cohesin trimer containing 6 Cysteines, positioned at the SMC-SCC1 and SMC1-3 ATPase head heterodimerization interfaces.

6. Crosslinking would be expected to stabilize the SMC ATP heads in close proximity, potentially an engaged state. Would full SMC head disengagement be prevented by such cross-linking even in the absence of nucleotide?

7. From Figure 5 the state of the heads is not immediately apparent. Please include a Figure comparing the state of the 'disengaged' SMC heads obtained using their cross-linking method with that of the ATP engaged state published previously.

8. If indeed, crosslinking stabilizes an engaged but nucleotide-free form, the authors need explicitly discuss the potential implications. For example, crosslinking of the heads could prevent the heads from properly disengaging. This has potential implications for the conformation/interactions of Scc2 or PDS5.

9. Figure 5G: The authors need to clarify how they obtain the ATP-bound state. Did they also use a crosslinked cohesin trimer, hydrolysis-impaired mutant ATPase heads or non-hydrolyzable ATP variants?

10. What is the distance between Smc1D588 and SCC2? Figure 5B indicates ~25Å which would be too far away for a direct interaction. In contrast, Figure 5E and their text description suggest that 'folding of the coiled coil brings the hinge to within 12nm (they probably mean Å?) of the heads'. Figure 5E indicates a distance of 12Å but it is between Smc1K620BPA and Smc1D588Y and not SCC2.

11. If indeed no direct interaction is apparent, or if the low resolution and flexibility do not allow firm conclusions, they need to indicate this during their discussion of these results.

12. Figure 6: While biochemical data indicate that PDS5 inhibits ATP hydrolysis, the presented structural data does not reveal how this is accomplished. One caveat of using a cross-linked version is that the procedure may prevent access of PDS5 to fully disengaged SMC heads and thus prevent SMC head engagement (in analogy to the role of the condensin YCS4 subunit). The authors need to indicate potential pitfalls of their cross-linking procedure on PDS5 positioning and discuss if the conformation observed is potentially off pathway.

Comments for the authors:

1. Figure 4C-F: Please indicate if the experiment has been done once or repeated several times with consistency.

2. Can they indicate if their preparation used for CryoEM is fully or partially BMOE cross-linked (e.g. by SDS-PAGE analysis).

3. Figure 6: While at lower resolution, it would still be important to show how well the PDB models fit into the cryoEM density map shown in Figure 6A. This would give the reader a sense for how reliable positioning of the PDB models (shown in Figure 6B) are.

4. Can they indicate where the previously published (Rowland et al. 2009, Sutani et al. 2009) eco1-1 suppressor mutations of PDS5 are located? Are suppressor mutations located in observed interfaces? Would they be predicted to interfere with PDS5 interaction?

5. L.593 et seq: The authors mention that their data are in contrast to 'previous studies with shortened constructs'. Previous work needs to be cited.

6. L280 : This should read: 'relative to the position of D574'?

7. Figure S4E: Cartoon shows SMC3! Do they mean SMC1?

8. Details on CryoEM data processing and reconstruction information need to be included.

9. PDB validation reports of the modeled cryoEM structures are missing.

10. It is not clear how the PDB model for the cohesin elbow is derived. No details of CryoEM/Xray data collection are given and PDB validation reports are again missing.

11. L. 1029: Spelling error: Katsu Shirahiga

Reviewer #3:

Chemical crosslinking experiments have established that sister chromatid cohesion requires "topological embrace" of sister chromatids within a cohesin ring. Three contacts define the ring: Scc1-N:Smc3-head, Scc1-C:Smc1-head, and Smc1-hinge:Smc3-hinge. The strongest experimental evidence for a specific DNA entry point is that artificial dimerization of the Smc1-hinge:Smc3-hinge interface prevents cohesion establishment (Gruber et al. 2006; Srinivasan et al. 2018). In the current manuscript, Patela et al. present genetic, biochemical, and structural data that indicate that the cohesin hinge interacts with the cohesin loading complex (Scc2/4), that this interaction depends on Scc4, and that mutations in the hinge that appear to enhance the interaction positively affect cohesin's association with chromosomes. Considered together, these data provide new evidence that the cohesin hinge is connected to the loading reaction. The manuscript is really interesting, includes many very clever experiments, and extends previous models for how cohesin works.

There are major issues with the paper that need to be addressed before publication. First, the paper is confusing. It should be simplified so that the main points are clearer, and errors in the text and figures should be corrected. Second, the section about nucleosome remodeling must be rewritten. There is great value in pointing out differences between studies, but these points can be made more succinctly. Third, the descriptions of the structures are insufficient and do not permit comparison with published structures. The reader cannot currently judge the validity of the interpretations presented.

With the exception of some minor updates to the biochemical experiments, the majority of these changes can be made without adding new data. The rewriting and corrections I have requested will nevertheless require significant time and attention. In addition, I have also requested the EM maps and corresponding docked models be made available to reviewers before a preliminary decision is reached. After this is done, I would support publication of a substantially updated version of the current manuscript in eLife.

1. The manuscript contains a collection of interesting observations, all of which address the overall hypothesis that Scc2/4 stimulates cohesion establishment by interacting with the cohesin hinge.

– See below for comments on the structures. The Pds5 structure should be moved to the supplemental material or removed altogether. The new structure is good enough to discern the overall position of Pds5 relative to cohesin, but it does not provide much new reliable information beyond this. It is pertinent in that it confirms competition between Scc2 and Pds5. Finer details like the status of the head domains, the identity of specific contacts, and the relationship to cohesin acetylation/Eco1 binding will have to wait for higher-resolution structures that will ideally be trapped in defined identifiable biochemical states.

– Like the Smc1/3 E-to-Q mutants, smc1D588Y appears to slow or stall cohesin at an early/intermediate step in the loading process (Figure 2C-D). The cells are viable, so the stall is not terminal. How do the authors reconcile this observation with their finding that cohesin loading on chromosome arms is actually elevated relative to wild type?

– Related to the above question: Have the authors explored how the mutation affects engagement with chromosomal DNA analogous to the recent Chappard paper? Are S-K complexes depleted in favor of others? Can they observe so-called E or J heads, and is either enriched in the mutant background? One line 353, the authors stop short of suggesting the smc1D588Y mutant favors the clamped state but imply it may stop the loading reaction at a step after the one stalled by the clamped-favoring EQ mutations. The current data do not conclusively show that the step slowed by smc1D588Y is before or after the step stalled by the EQ mutations. The authors should remove speculation on this point from the Results section.

– The description of results that begins on line 336 is very confusing. Specifically, "This implies that the reduced loading arises…" but the authors say directly above that Smc1D588Y is at 110% (CENs) of WT. Also, the first two sentences of this paragraph use "it," but the referent is not clear. Last, a simple explanation of the phenotype shown in Figure 2 seems to be that smc1D588Y preserves CEN targeting but disrupts translocation to pericentromeres. Arm loading is not disrupted (slightly enhanced). A simpler description of the phenotype would make the paragraph much easier to read and focus the reader on the important question: what accounts for the subtle defects seen in the smc1D588Y mutant?

– Line 456 – "Nevertheless, the level of Smc1-Scc2 crosslinked protein was comparable to that of Scc3…" This is an inappropriate comparison. Crosslinking efficiency could be far more efficient if there is a fortuitous lysine on the target (Scc3 or Scc2 here).

– Line 562 – This is a concrete biochemical prediction that is totally testable. Since it is central to the proposed mechanism of suppression, it should be tested using pulldown assays with purified proteins (cohesin tetramer, Scc2/4 or Scc2-C).

2. The section describing the RSC-Scc4 interaction needs to be revised. The authors focus on and oversimplify a secondary conclusion from a previous paper (Lopez-Serra et al. 2014), which holds that Scc4 stimulates RSC. The majority of the work and discussion in this and a related paper from the same group (Lopez-Serra et al. 2014 and Munoz et al. 2020) indicate that Scc2/4 localization depends on RSC. The Munoz paper offers a biochemical explanation for this observation but does not follow up on the secondary observation from the first paper that Scc2/4 influences nucleosome positioning.

– The current findings regarding the genome-wide distribution of cohesin in sth1-3, smc1D588Y, or related mutants seem mostly consistent with the results presented in Lopez-Serra and in Munoz. The authors should revise their text to reflect this concordance. In fact, all three studies emphasize the importance of nucleosome-free DNA. This could be pointed out.

– Any differences in the cohesin distribution between the two studies (Patela and Lopez-Serra), especially in the sth1-3 mutant must be more carefully annotated. Direct comparisons of the original studied loci (Lopez-Serra et al. Figure 3B, for example) must be presented if the authors wish to question the previous findings.

– That artificial chromosome tethering of Scc2-C rescues scc4D (Munoz Figure 6B) suggests the partially Scc4-dependent hinge interaction described here is not an essential function of Scc4. The authors suggest that this Scc4 function is rendered irrelevant when the loader is brought near DNA. At this point, there is not enough information to discriminate between these very similar models, and it would be better not to set the current manuscript up in opposition to previous work when an experimental distinction has not been made.

– Was the MNase-seq calibrated? Also, it looks like there are differences in nucleosome occupancy around AST2, which was also highlighted in the Lopez-Serra work (Figure S3G here). How are the two WT panels different? They show very different results.

– The authors say that RSC is not uniquely required for Scc2/4 recruitment, but Figure 3A from Lopez-Serra et al. shows that sth1-3 produces a pronounced cohesion defect, and other chromatin modifying enzyme mutants do not.

– Line 413 -The double mutant experiment (smc1D588Y sth1-3) is not very informative. It's unclear to me that suppression (or not) of the RSC phenotype by any of the mutants described in the current work would prove the previous sth1-3 findings to be unreliable. In addition, it would be important to compare cohesion, which is not done here. The double mutant experiments can be included, but the conclusions drawn are very limited and need to be rewritten.

– Line 416 – "We were also unable to reproduce the reported effects of scc4-4 and sth1-3 on nucleosome positioning…" If the authors really wish to make this statement/comparison, they need to be much more deliberate about evaluating the reported effects and then comparing their own data. There are clearly differences in Figure S3G, but the authors do not look at them with sufficient granularity. In particular, the Lopez-Serra paper looks at Scc2-bound genes/promoters. The authors must also compare their data with a more recent report (Kubik…Shore, Mol Cell, 2018). This study shows effects specifically at the +1 nucleosome of transcribed genes and is much more in line with the Lopez-Serra observations.

3. The new structures presented in the paper are compelling. There are several issues that must be cleared up before the manuscript can be published.

– It is unclear how the structures were determined. The authors should present data processing flowcharts to show how the most important structures in the paper were generated. It would be very helpful to know whether the structures described are major or minor components of the samples that were analyzed (relative to other determine structures). Therefore, particle numbers should be included for 3D classification steps, and if any other medium-high resolution structures came from the same datasets, those should be included in the chart for the benefit of the reader. If the authors wish not to disclose unpublished structures, they may elect to exclude some of these, but the overall breakdown of the data needs to be clear. If other structures produced by the dataset are already published, this is fine.

– Data collection tables need to be included, as do tables describing any "pseudo-atomic models." This is especially true for structures that will be deposited in the PDB. It looks like 5F, 5G, and 6A show structures that should be treated this way. For parts the authors do not wish to model explicitly (low-resolution parts corresponding to Scc2/Pds5), they can exclude these from the PDB and carry out model-map correlation calculations using maps truncated to the modeled residues.

– 2D classification results can be unreliable, especially when there are flexible modules. The analysis shown in Figure S4A-C should be removed, and the text should be modified so that any conclusions made from these images are removed. In particular, the consequences of ATP binding, discussed in relation to Figure S4C is inappropriate given that (1) the authors cannot resolve nucleotide with these images (is it really bound? hydrolyzed? Were there other classes in which the opposite was true?); (2) designation of Scc2 density is reasonable but not probative; and (3) it is not clear what the relationship is between the cohesin head domains. Subtle changes in head engagement could make huge differences in the interpretation. In the displayed images, it is not entirely clear that the heads are even engaged (versus "juxtaposed").

– How confident are the authors in their positioning of the Pds5 module and cohesin head domains in Figure 6A? The density below the "joint" as drawn is very difficult to interpret and looks like it might be artifactual.

– The authors should compare the cohesin head domain conformations they observe, especially for the Scc2 complex, to those observed in the recent structures showing cohesin-nucleotide-Scc3-Scc2-DNA interactions. Are the current structures well enough resolved to make these comparisons? Can the authors see Scc1-N:Smc3-head domain contact?

– Do the authors truly know the cohesin structure displayed in 5G is ATP-bound? To make this statement, they would need to resolve nucleotide and a connected γ phosphate.

– Why did the authors leave out Scc3 from the cryo-EM samples? How do they relate their observations to the published structures, which show the hinge attached to Scc3/Psc3 and dislocated from the Smc1/3 head domains? Both conformations probably inform on mechanism, but the reader is not presented with a helpful way to think about this. Does Scc3 "grab" the hinge after head-Scc2 engagement, as proposed by Uhlmann's group recently (Higashi et al. 2020)? To be sure, Srinivasan et al. proposed an Scc3-dependent late step in loading already, so both of these could easily be referenced in a paragraph contextualizing the current structures with respect to Scc3.

4. There are disagreements between data and text and confusing inconsistencies in the figures:

o S1C: why is Y40A not an scc4-ts allele?

o S1E – It looks like smc1D588Y is an scc2-4 suppressor. The text says the opposite.

o Figure S3 is discussed below.

o Genotypes should be given in color for all tetrad dissections (as in Figure 1).

o Size markers are missing from the blots. These are important for the crosslinking experiments so that positions can be compared for different antibodies (Figure 4B, for example).

o In at least once case, the authors should show a myc blot for Smc1-WT (no Cys mutation) so that it is clear the higher band is indeed an Smc1 crosslinking product.

o For these assays, the text and legend disagree on which subunit was immobilized for the pulldown (Scc1 or Smc1). If it was different in different experiments, this should be stated.

o Figure S4F is not helpful. There is no control shown (either Scc2 or Smc1, and ideally both, lacking Cys mutation). The crosslinking efficiencies do not seem to correlate for the two subunits, and it is not clear why some Scc2 mutants show an even higher band.

In the end, the experiment would have only been valuable if a really great crosslink position were found. The experiment can be removed without seriously damaging the manuscript.

– The scatter plot in figure S3E does not contradict the Lopez-Serra finding and is not a good way to look at this data. The Lopez-Serra paper addressed a small subset of chromosomal locations. It is not clear how far away from the diagonal the distribution of dots would need to spread to constitute a truly different localization pattern (no control for this).

– If the authors are sure of their placements for Scc2 and Pds5, then why not directly compare the two structures by overlaying them anchored on the coiled-coiled/hinge domains? This would be a great way of showing that (i) Pds5 is indeed more closely connected to the hinge and (ii) both modules engage the same overall cohesin conformation/fold (or not, as the case may be).

– Figure 5F-G make it look like the two structures are different views of the same. Only G is required for the point that head engagement does not unzip the coiled coils, so there is no need to make it look like a comparison.

– Figure S5D does not give the reader enough information to evaluate what is being shown. What dataset does this map come from? Is the corresponding main figure 5A? Was Smc3 included in the fit? DNA? Was nucleotide included in the sample? Why is there a large part of Scc2 poking out (presumably related to mentioned "floppiness")?

– Neither the Murayama papers nor the Huis in 't Veld papers are discussed. The Huis in 't Veld paper reports contact between SA1 and the cohesin hinge. Murayama and Uhlmann (2015) reconstitute this interaction (Psm1/3-Psc3) for the fission yeast components and state that it means the ring must bend and that the loader is probably involved. While neither study proved hinge-Scc3 contact is essential for the loading reaction, they were important contributions and should be cited.

– Presentation issues for structures:

o At least once in each figure (but not necessarily each panel), colored text should be shown for each included protein as is done in Figure 1. This is missing in Figure 5 (for example).

o The overlay in Figure 1F is hard to understand. The mutated structure should be shaded differently so the reader can see the difference clearly (pale green vs green and same for red).

o The nucleosome structure in Figure 3B is impossible to understand. The entire particle should be shown so the reader can see the DNA wrap, which is relevant here (do the mutations enhance unwrapping?). Only the mutated histones and mutated residues (highlight color) need to be colored.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Folding of cohesin's coiled coil is important for Scc2/4-induced association with chromosomes" for further consideration by eLife. Your revised article has been evaluated by Cynthia Wolberger as the Senior Editor, and a Reviewing Editor.

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined in the points made by reviewers below. It is particularly important to clarify where the crosslinks reside in the cohesin variant used for structural analysis, as raised in Points 2 and 3 from Reviewer 2. If the crosslinks are in the heads, as indicated in the methods section, then this has important implications for the interpretation.

Reviewer #2:

The authors have addressed most comments. There are however a couple of issues that require further clarification:

1. Figure 4G: The author's response is only partially convincing: If Smc1K620C also crosslinks to Scc2C224, it is not fully clear that Scc2N220C specifically allows readout of contacts between the Smc1 hinge and Scc2. As they have tested a variety of Scc2 cysteine substitutions, it would have been important to test a Scc2 version that does not contain C224, in order to avoid such confounding non-specific crosslinking.

In any case, they need to further clarify how the different blots are developed. Presumably, the top panel shows an anti-FLAG western blot and the bottom panel an anti-myc blot (as indicated in the author's response)? The top panel is not labeled at all and the bottom panel is mislabeled as Anti-HA!

2. The authors state line 523 that they use a cohesin variant 'containing cysteines specifically crosslinking the three intermolecular interfaces'. They need to explicitly state which interfaces are crosslinked in the variant they use for structure determination. The author response indicates that 'the 6-cysteine version used interrogates the hinge dimerisation and not the head dimerisation'. Their methods section indicates however that the variant they use for structure determination does not contain Cys mutations in the hinges. Instead, they use Smc1G22C N1192C, Smc3S1043C R1222C, Scc1C56A547C.

These mutations are located (1) in the Smc3-NScc1 interface (Scc1C56 and Smc3S1043C), (2) Smc1-cScc1 (Scc1A547C and Smc1G22C) interface and (3), in the interface between the Smc1 and Smc3 ATPase heads (Smc1 N1192C and Smc3 R1222C). That is, the third pair of cysteines is clearly placed in the heads and not the hinges.

3. It is therefore not clear why the authors maintain (Reviewer #2 points 6, 7, 8, 12) that 'we did not cross-link the ATP head dimerisation interface'. Clearly, something is wrong! Maybe it is the methods section? If not, the authors need to clearly indicate that the version of cohesin they use is crosslinked at the SMC1/3 ATPase head domains. The implications of such head crosslinking need to be taken into account in the interpretation of their data and discussion of the results.

Reviewer #3:

The manuscript is greatly improved. The updated figures make it much clearer. The rewritten section describing Sth1, nucleosome positioning, and histone mutants is good. This is important work, and the breadth and strength of the experiments described are impressive.

Most of my comments have been adequately addressed. The requests below relate to the presentation and description of the data and should be addressed before publication but do not require extensive writing or any new experiments.

Line 593-597:

"In the presence of ATP" implies that ATP occupies both ATPase active sites. Neither γ phosphate can be resolved at this resolution. The structure could be a reaction intermediate resulting from slow π dissociation from one or both sites, for example. This is not to detract from the observations or statements made. Instead, it would be wise to state more clearly that ATP was included in the sample but avoid any suggestion that the structure provides detailed information about the SMC ATPase cycle, which will doubtless be a topic for future studies. This is rightly stated in the final paragraph of the Discussion.

Figure 5G needs to be modified. Figure 5S2 shows that 5G is a composite structure derived from two different samples. One contained ATP and the other did not. Overlaying them is inappropriate. Yes, the angle of the head-proximal coiled-coils is similar (Figure 5F vs. 5G), and it is likely that the remainder of the CC density beyond (hinge-proximal) the joint is similar/identical, but this is not proven by the data in the manuscript. In fact, it appears this section of the complex was excluded from analysis in the case of the sample containing nucleotide (Figure 5G and 5S2D). Without recalculating structures or even remaking complicated figures, it must be possible to present this data more clearly. One possibility: simply removing the "Arm" density from figure 5G. Another possibility is to compare the head domain density (including head-proximal cc) from Figure 5F, which is currently obscured by Scc2 density, with the head domain reconstruction from 5G.

Figure 5 – supplement 1A does not conclusively show complexes lacking Scc2. Compare with 5S1A, for example. One needs to consider variability in Scc2 occupancy, Scc2 position, and also the fraction and quality of the particle images contributing to these classes. The description "in samples lacking all HAWK proteins" (line 537) therefore overstates the certainty of the claim given the evidence. The statement and figure panel could be removed without damaging the overall message. Removing panel A has the added benefit that it eliminates the need to discuss a dataset not referenced in figure 5S2 (cohesin, Scc2, ATP).

Author response:

Reviewer #1:

Comments for the authors:

1. The authors should considering shortening the section on RSC describing Figure S3 as it detracts from the main message of the manuscript. In addition, while the ChIP-seq in Figure S3D argue against a major role for RSC in cohesin loading, the nucleosome mapping data in Figure S3S are not conclusive, as presented. Why do the upper two and lower two nucleosome profiles look different from each other? Potentially effects on nucleosome positioning will only be visible in metagene analysis: the sth1-3 mutant at least would be expected to alter nucleosome positioning but this is not apparent from the data presented. The authors should either remove this data or perform additional analysis.

We have removed the nucleosome positioning data in Figure S3G and amended the text accordingly.

2. Figure 7G: The conclusion that cohesin folding at the elbow is a feature of cohesin complexes that are holding sister chromatids together relies on the assumption that all acetylated cohesin is participating in cohesion. However, this may not be the case. In mammals, cohesin acetylation is also observed in G1, indicating that it may have a more general role. This potential caveat should be mentioned.

Our work is exclusively about what happens in yeast, where acetylation is linked to replication, where sororin is absent, and where acetylation (not sororin association) is the best (known) marker for cohesion. The fact that acetylation is clearly not a good marker for cohesion in mammalian cells does not therefore merit a “caveat”. As far as we know, acetylation is as good a marker in yeast as is sororin association in mammalian cells. The reviewer is correct that this should be pointed out and we have altered the text accordingly.

3. It is surprising that there is high Scc1 ChIP-seq signal at centromeres in scc4-4 cells if Scc4 is required for cohesin loading at centromeres (Figure S4A). How do the authors explain this?

It has been previously documented that there is an Scc4-independent mechanism that promotes centromeric loading in the presence of nocodazole (Petela et al. 2018). As this experiment was done in the presence of nocodazole, this mechanism is likely responsible for the high Scc1 ChIP-seq signal observed here.

4. Line 273: It looks like smc1D588Y CAN rescue the proliferation defect of scc2-4 in Figure S1E.

This is quite right, and the text has been amended accordingly.

5. Figure 3A please show proliferation data for the different histone mutants.

We show proliferation data for the mutant that we use for all subsequent experiments in Figure 3A. This mutant was made de novo and is a good representative for the other mutants. We think showing additional proliferation data for all the mutants is therefore unnecessary.

6. Line 402: Smc3 acetylation

Corrected, many thanks.

7. Supplementary Figure 4F: needs a negative control without cysteine substitutions.

In line with comments from the other reviewers, we agree this panel is not helpful and not needed and therefore have removed it and amended the text accordingly.

8. Page 19: Line 547: Call to Figure S4D is incorrect.

Corrected, many thanks again.

Reviewer #2:

The conclusions are mostly well supported by the data, but some aspects of the biochemical and structural data require clarification.

1. Biochemical data: Figure 4B: The FLAG blot indicates that x-linked species migrate at different positions on SDS-PAGE while the top panel shows uniform mobility. How do we know that the band labeled SMC1-? corresponds to x-linked SMC1 and not some non-specific anti-myc background (e.g. there is an additional non-labeled band running lower)?

The Myc blot shows multiple crosslinked species corresponding to Smc1 crosslinking to multiple proteins. As can be seen from the FLAG blot these species migrate at slightly different positions and have different efficiencies. The most efficient crosslink by far is to Pds5, corresponding to the intense band labelled “Smc1-?” in the Myc blot, which is present in all cases and does mask the other species. It should be noted that this band runs slightly higher in the Pds5-FLAG lane, due to the tag increasing the molecular weight. This crosslinking species has been documented previously, with additional controls, in Bürmann et al., 2019.

We do understand however that our labelling of the blot may have been misleading and so have tried to improve the clarity by amending the “Smc1-?” label to reflect the multiple species present.

Figure 4B: Different amounts of FLAG-HAWKs are recovered in the anti-Myc IP. Can the authors generate an 'input' blot showing expression levels to exclude the possibility that the variable crosslinking efficiency is not due to different expression levels?

We agree this would be useful and have modified the figure panel to include the expression levels of Scc2, Scc3 and Pds5 in whole cell extracts relative to a loading control. Indeed, the proteins are expressed to different levels as expected but this does not correlate to the crosslinking efficiency. Pds5 for example is not as abundant as Scc3 but crosslinks much more efficiently.

2. Figure S4F: Show a WT negative control (no Cys mutation) to confirm that the bands observed do not arise due to non-specific background cross-linking.

As detailed above, this figure panel has been removed and the text amended accordingly.

3. Figure 4G: Top: Why is there a crosslinked band in the lane containing Smc1K620C alone? Would it not be expected to see cross-linking only when both Scc2N200C and Smc1K620C are present? Why is the non-specific crosslinking (labeled as *) present everywhere? Which cross-linked band they are referring to? While there is a small amount of a band visible above the non-specific band, this weak band is also present in Smc1K620C alone!

The crosslinking band observed in the lane containing only Smc1K620C is most likely due to Smc1K620C crosslinking to a natural cysteine in Scc2, probably C224 which sits on a small helix just under that containing N200. This would explain the very similar molecular weight, the fact the band also appears in the Myc blot and the increased intensity of the band on addition of Scc2N200C. The text has been modified to reflect this. We do not know the identity of the non-specific crosslinking band present in the Myc blot but it is clearly independent of either cysteine mutations as it is observed in lane 1. To improve the clarity, we have amended the labelling.

4. Structural data: Figure 5A: The authors need to clarify if in their cryoEM structure, the SMC ATPase heads are in the 'apo' state, as indicated in the text or in the engaged state as indicated in the Discussion (L. 641).

Changed. Thank you, this was a typo that has now been corrected.

5. The authors need to explicitly state in their description of their structural data (Line 512 et seq) that they used a crosslinked version of the cohesin trimer containing 6 Cysteines, positioned at the SMC-SCC1 and SMC1-3 ATPase head heterodimerization interfaces.

Done, but please bear in mind, as below, that the 6-cysteine version used interrogates the hinge dimerisation and not the head dimerisation.

6. Crosslinking would be expected to stabilize the SMC ATP heads in close proximity, potentially an engaged state. Would full SMC head disengagement be prevented by such cross-linking even in the absence of nucleotide?

See last point: we did not cross-link the ATP head dimerisation interface and therefore would not expect head (dis)engagement to be affected in any way.

7. From Figure 5 the state of the heads is not immediately apparent. Please include a Figure comparing the state of the 'disengaged' SMC heads obtained using their cross-linking method with that of the ATP engaged state published previously.

Again, the cross-linking does not affect the head engagement of the molecule and at this resolution it would seem somewhat over-interpretative to analyse the precise placement of the head subunits.

8. If indeed, crosslinking stabilizes an engaged but nucleotide-free form, the authors need explicitly discuss the potential implications. For example, crosslinking of the heads could prevent the heads from properly disengaging. This has potential implications for the conformation/interactions of Scc2 or PDS5.

Again, see above. The head dimerisation domains were not cross-linked.

9. Figure 5G: The authors need to clarify how they obtain the ATP-bound state. Did they also use a crosslinked cohesin trimer, hydrolysis-impaired mutant ATPase heads or non-hydrolyzable ATP variants?

As described, the ATP-bound state was simply achieved by incubating the trimer with ATP. For clarification we have added that no cross-linker was used.

10. What is the distance between Smc1D588 and SCC2? Figure 5B indicates ~25Å which would be too far away for a direct interaction. In contrast, Figure 5E and their text description suggest that 'folding of the coiled coil brings the hinge to within 12nm (they probably mean Å?) of the heads'. Figure 5E indicates a distance of 12Å but it is between Smc1K620BPA and Smc1D588Y and not SCC2.

As described in Figure 5B, the distance between Scc2 and Smc1D588 is indeed 25 Å. This refers to the distance of the best cryo-EM map reconstruction (which by definition is an average) and therefore only represents a single snapshot of the entire range of Scc2’s movements above the neck as mentioned in the manuscript. The distance is supposed to highlight this flexibility when comparing it to the hinge-bound structures published previously (e.g. Shi et al. 2020).

We apologise if this was unclear and have reworded the following to aid with clarity:

“Initial 2D classes of Scc2-bound cohesin revealed (Figure S5B) floppiness not only within the HAWK, especially in its C-terminus, but also between the joint and the ATPase heads.”

We recognise the phrase 'folding of the coiled coil brings the hinge to within 12 nm of the heads’ is misleading and should have referred to the ATPase heads to be precise and so as to avoid confusion with the HAWK’s head. We have now changed this accordingly

11. If indeed no direct interaction is apparent, or if the low resolution and flexibility do not allow firm conclusions, they need to indicate this during their discussion of these results.

Our manuscript never claims to observe a rigid interaction between Scc2 and the hinge – and the 25 Å distance in Figure 5 reaffirms this – but rather that the folding of the elbow is fundamental in allowing for it and that the clear interaction observed in previously published structures must stem from an intrinsic floppiness in the head region of Scc2 that we also observe in our processing.

12. Figure 6: While biochemical data indicate that PDS5 inhibits ATP hydrolysis, the presented structural data does not reveal how this is accomplished. One caveat of using a cross-linked version is that the procedure may prevent access of PDS5 to fully disengaged SMC heads and thus prevent SMC head engagement (in analogy to the role of the condensin YCS4 subunit). The authors need to indicate potential pitfalls of their cross-linking procedure on PDS5 positioning and discuss if the conformation observed is potentially off pathway.

See above. Head engagement should not be affected in any way by the crosslinking as the third pair of cysteines are placed in the hinge and not the heads.

Comments for the authors:

1. Figure 4C-F: Please indicate if the experiment has been done once or repeated several times with consistency.

The experiments were performed twice with the same results. This is now stated in the figure legend.

2. Can they indicate if their preparation used for CryoEM is fully or partially BMOE cross-linked (e.g. by SDS-PAGE analysis).

The estimated efficiency of cross-linking is around 20% as previously mentioned in Collier et al., 2020 (also see SDS-PAGE gel in Author response image 1; the total intensity of the SMCs [including the smear] and Scc1 from lane 1 [trimer crosslinking no cysteines] compared to the 6C crosslinking from the top band from lane 8 [trimer crosslinking 6C]). We have now added that to the manuscript as suggested:

“…To investigate this further, we used cryo-EM to determine the structures of the S. cerevisiae cohesin trimer (Smc1, Smc3, Scc1, in their 6C version as reported in Collier et al., 2020; specifically cross-linked at the three intermolecular interfaces at an efficiency of 20% (data not shown)) bound to either Scc2 (Figure 5A)”

3. Figure 6: While at lower resolution, it would still be important to show how well the PDB models fit into the cryoEM density map shown in Figure 6A. This would give the reader a sense for how reliable positioning of the PDB models (shown in Figure 6B) are.

Agreed and done.

4. Can they indicate where the previously published (Rowland et al. 2009, Sutani et al. 2009) eco1-1 suppressor mutations of PDS5 are located? Are suppressor mutations located in observed interfaces? Would they be predicted to interfere with PDS5 interaction?

The suppressor mutation cluster around residues 81-89 in PDS5 reported by Rowland et al. would sit at the hinge:Pds5 interaction interface. The rest of the mutations do not reside in any of the other interfaces predicted by our model. However, as we have explained in the manuscript, the density around the N-terminus of Pds5 is not ideal and we would prefer to refrain from making any too detailed conclusions.

5. L.593 et seq: The authors mention that their data are in contrast to 'previous studies with shortened constructs'. Previous work needs to be cited.

We have added a reference to Muir et al., 2020.

6. L280: This should read: 'relative to the position of D574'?

Thank you, this has been corrected.

7. Figure S4E: Cartoon shows SMC3! Do they mean SMC1?

Yes it should read Smc1, this has also been corrected.

8. Details on CryoEM data processing and reconstruction information need to be included.

We have added a detailed overview of cryo-EM data processing and reconstruction workflows for all maps in Supplementary Figure 7.

9. PDB validation reports of the modeled cryoEM structures are missing.

PDB and EMDB validation reports for all the maps have been attached. The accession codes have been added both to the figure legends and the text. The Table of Structures” in Materials and methods details this for each map:

10. It is not clear how the PDB model for the cohesin elbow is derived. No details of CryoEM/Xray data collection are given and PDB validation reports are again missing.

Please be referred to point 8. We have also added more detail in the Methods section about the model building of the cohesin elbow PDB. In addition, we have added Supplementary Table 3 with information about data collection.

11. L. 1029: Spelling error: Katsu Shirahiga

Thank you, this has been corrected.

Reviewer #3:

Chemical crosslinking experiments have established that sister chromatid cohesion requires "topological embrace" of sister chromatids within a cohesin ring. Three contacts define the ring: Scc1-N:Smc3-head, Scc1-C:Smc1-head, and Smc1-hinge:Smc3-hinge. The strongest experimental evidence for a specific DNA entry point is that artificial dimerization of the Smc1-hinge:Smc3-hinge interface prevents cohesion establishment (Gruber et al. 2006; Srinivasan et al. 2018). In the current manuscript, Patela et al. present genetic, biochemical, and structural data that indicate that the cohesin hinge interacts with the cohesin loading complex (Scc2/4), that this interaction depends on Scc4, and that mutations in the hinge that appear to enhance the interaction positively affect cohesin's association with chromosomes. Considered together, these data provide new evidence that the cohesin hinge is connected to the loading reaction. The manuscript is really interesting, includes many very clever experiments, and extends previous models for how cohesin works.

There are major issues with the paper that need to be addressed before publication. First, the paper is confusing. It should be simplified so that the main points are clearer, and errors in the text and figures should be corrected. Second, the section about nucleosome remodeling must be rewritten. There is great value in pointing out differences between studies, but these points can be made more succinctly. Third, the descriptions of the structures are insufficient and do not permit comparison with published structures. The reader cannot currently judge the validity of the interpretations presented.

With the exception of some minor updates to the biochemical experiments, the majority of these changes can be made without adding new data. The rewriting and corrections I have requested will nevertheless require significant time and attention. In addition, I have also requested the EM maps and corresponding docked models be made available to reviewers before a preliminary decision is reached. After this is done, I would support publication of a substantially updated version of the current manuscript in eLife.

1. The manuscript contains a collection of interesting observations, all of which address the overall hypothesis that Scc2/4 stimulates cohesion establishment by interacting with the cohesin hinge.

– See below for comments on the structures. The Pds5 structure should be moved to the supplemental material or removed altogether. The new structure is good enough to discern the overall position of Pds5 relative to cohesin, but it does not provide much new reliable information beyond this. It is pertinent in that it confirms competition between Scc2 and Pds5. Finer details like the status of the head domains, the identity of specific contacts, and the relationship to cohesin acetylation/Eco1 binding will have to wait for higher-resolution structures that will ideally be trapped in defined identifiable biochemical states.

We respectfully disagree with the reviewer. The overall position of Pds5 relative to cohesin is of a clear interest to the wider cohesin community as it not only provides the first visualisation of the interaction but starts to hint at the answers to fundamental questions of this interaction. At the same time, it finally allows much needed further biochemical work to interrogate said interaction that would not be possible without it. It also serves as structural confirmation of the biochemical/genetic data reported in this paper and therefore plays an important role in the manuscript.

– Like the Smc1/3 E-to-Q mutants, smc1D588Y appears to slow or stall cohesin at an early/intermediate step in the loading process (Figure 2C-D). The cells are viable, so the stall is not terminal. How do the authors reconcile this observation with their finding that cohesin loading on chromosome arms is actually elevated relative to wild type?

We believe we addressed this point in the discussion “Why does smc1D588Y depress loading at CENs?”

In particular:

“We suggest that clamping is rate limiting along chromosome arms but unclamping is rate limiting at CENs and that this is the reason why smc1D588Y enhances arm loading while depressing loading at CENs.”

– Related to the above question: Have the authors explored how the mutation affects engagement with chromosomal DNA analogous to the recent Chappard paper? Are S-K complexes depleted in favor of others? Can they observe so-called E or J heads, and is either enriched in the mutant background? One line 353, the authors stop short of suggesting the smc1D588Y mutant favors the clamped state but imply it may stop the loading reaction at a step after the one stalled by the clamped-favoring EQ mutations. The current data do not conclusively show that the step slowed by smc1D588Y is before or after the step stalled by the EQ mutations. The authors should remove speculation on this point from the Results section.

Although these are valid questions that would be interesting to address, the experiments required to address them are not trivial and we do not think the answers are necessary for this manuscript, which is already quite long. Concerning the second point, we merely point out that the enhanced association of both Scc2 or EQ complexes at CENs caused by D588Y suggests that the mutation does not affect formation of the clamped state at CENs in vivo but some later step in the loading reaction. We do not agree that this is an unjustified speculation.

– The description of results that begins on line 336 is very confusing. Specifically, "This implies that the reduced loading arises…" but the authors say directly above that Smc1D588Y is at 110% (CENs) of WT. Also, the first two sentences of this paragraph use "it," but the referent is not clear. Last, a simple explanation of the phenotype shown in Figure 2 seems to be that smc1D588Y preserves CEN targeting but disrupts translocation to pericentromeres. Arm loading is not disrupted (slightly enhanced). A simpler description of the phenotype would make the paragraph much easier to read and focus the reader on the important question: what accounts for the subtle defects seen in the smc1D588Y mutant?

The sentence now reads: “This implies that the reduced loading around centromeres arises not from defective formation of the clamped state at CENs by Scc2/4 complexes associated with Ctf19 but from a defect in a subsequent step in the loading/translocation reaction that requires ATP hydrolysis.”

– Line 456 – "Nevertheless, the level of Smc1-Scc2 crosslinked protein was comparable to that of Scc3…" This is an inappropriate comparison. Crosslinking efficiency could be far more efficient if there is a fortuitous lysine on the target (Scc3 or Scc2 here).

The reviewer is of course correct. What we merely implied is that irrespective of such an effect, may or may not be the case, the amount of Smc1-Scc2 crosslinking was comparable to that of Smc1-Scc3 despite Scc2 being less abundant in the cell. This is surely a valid point to make.

– Line 562 – This is a concrete biochemical prediction that is totally testable. Since it is central to the proposed mechanism of suppression, it should be tested using pulldown assays with purified proteins (cohesin tetramer, Scc2/4 or Scc2-C).

This refers to the statement “We suggest that the addition of a bulky amino acid into Smc1 through D588Y may be sufficient to help bind an otherwise floppy Scc2 N-terminal domain, whose interaction with the hinge is normally stabilised by Scc4”. We disagree that this is a testable hypothesis using pulldown assays as we are referring to the stabilisation of a particular conformation of binding not binding at all, which itself was difficult to measure in this way due to the multiple contacts Scc2 makes with the tetramer (Petela et al. 2018). Although this would be good to address, doing so is not trivial.

2. The section describing the RSC-Scc4 interaction needs to be revised. The authors focus on and oversimplify a secondary conclusion from a previous paper (Lopez-Serra et al. 2014), which holds that Scc4 stimulates RSC. The majority of the work and discussion in this and a related paper from the same group (Lopez-Serra et al. 2014 and Munoz et al. 2020) indicate that Scc2/4 localization depends on RSC. The Munoz paper offers a biochemical explanation for this observation but does not follow up on the secondary observation from the first paper that Scc2/4 influences nucleosome positioning.

– The current findings regarding the genome-wide distribution of cohesin in sth1-3, smc1D588Y, or related mutants seem mostly consistent with the results presented in Lopez-Serra and in Munoz. The authors should revise their text to reflect this concordance. In fact, all three studies emphasize the importance of nucleosome-free DNA. This could be pointed out.

We are grateful to the reviewer for pointing out this deficiency in our discussion of the papers from the Uhlmann lab. We now point out in our revised manuscript that they have in fact proposed two, possibly contradictory, hypotheses for how Scc2/4 functions together with RSC. Crucially, both are inconsistent with our finding that sth1-3 causes only a modest if any defect in cohesin loading in vivo. We would also like to point out that the conclusion that Scc4 acts by stimulating RSC cannot be considered merely a minor secondary conclusion of the Lopez-Serra paper. The very title of their paper was “Scc2/4 acts in sister chromatid cohesion by …maintaining nucleosome regions”. We also respectfully disagree with the reviewer’s statement that the genome-wide distribution of cohesin in sth1-3, smc1D588Y, or related mutants seem mostly consistent with the results presented in Lopez-Serra and in Munoz. We do not believe in papering over the cracks in this manner. There is a clear inconsistency between our calibrated ChIP-seq data and their anecdotal qPCR measurements. We have nevertheless greatly revised our discussion of this issue and hope it that it more accurately describes the claims made by the Uhlmann lab while at the same time being easier to read.

– Any differences in the cohesin distribution between the two studies (Patela and Lopez-Serra), especially in the sth1-3 mutant must be more carefully annotated. Direct comparisons of the original studied loci (Lopez-Serra et al. Figure 3B, for example) must be presented if the authors wish to question the previous findings.

We have now included comparisons of the original studied loci.

– That artificial chromosome tethering of Scc2-C rescues scc4D (Munoz Figure 6B) suggests the partially Scc4-dependent hinge interaction described here is not an essential function of Scc4. The authors suggest that this Scc4 function is rendered irrelevant when the loader is brought near DNA. At this point, there is not enough information to discriminate between these very similar models, and it would be better not to set the current manuscript up in opposition to previous work when an experimental distinction has not been made.

We have now revised this aspect of the discussion.

– Was the MNase-seq calibrated? Also, it looks like there are differences in nucleosome occupancy around AST2, which was also highlighted in the Lopez-Serra work (Figure S3G here). How are the two WT panels different? They show very different results.

In line with comments from the other reviewers we have removed this data and amended the text accordingly.

– The authors say that RSC is not uniquely required for Scc2/4 recruitment, but Figure 3A from Lopez-Serra et al. shows that sth1-3 produces a pronounced cohesion defect, and other chromatin modifying enzyme mutants do not.

We do not discuss the involvement of nucleosome remodellers in sister chromatid cohesion. The observations we discuss relate only to the occupancy of cohesin on DNA and cannot distinguish cohesive and non-cohesive complexes. We note that there may be a greater defect in Smc3 acetylation in sth1-3 mutants than there is a defect in cohesin loading (Figure S3C), which might conceivably be accompanied by cohesion defects. We did not follow this up as we were concerned with cohesin loading and not with the process of cohesion establishment and besides which it was not our goal to re-investigate the role of RSC in this process. That said, this section has been rewritten.

– Line 413 -The double mutant experiment (smc1D588Y sth1-3) is not very informative. It's unclear to me that suppression (or not) of the RSC phenotype by any of the mutants described in the current work would prove the previous sth1-3 findings to be unreliable. In addition, it would be important to compare cohesion, which is not done here. The double mutant experiments can be included, but the conclusions drawn are very limited and need to be rewritten.

This has been re-written to make it clear that even if there is a modest defect in loading in sth1 mutants, it is not altered by smc1D588Y, which bypasses the requirement for Scc4. We hope our logic is now clearer.

– Line 416 – "We were also unable to reproduce the reported effects of scc4-4 and sth1-3 on nucleosome positioning…" If the authors really wish to make this statement/comparison, they need to be much more deliberate about evaluating the reported effects and then comparing their own data. There are clearly differences in Figure S3G, but the authors do not look at them with sufficient granularity. In particular, the Lopez-Serra paper looks at Scc2-bound genes/promoters. The authors must also compare their data with a more recent report (Kubik…Shore, Mol Cell, 2018). This study shows effects specifically at the +1 nucleosome of transcribed genes and is much more in line with the Lopez-Serra observations.

In line with the comments from the other reviewers, we have removed this data and amended the text accordingly.

3. The new structures presented in the paper are compelling. There are several issues that must be cleared up before the manuscript can be published.

– It is unclear how the structures were determined. The authors should present data processing flowcharts to show how the most important structures in the paper were generated. It would be very helpful to know whether the structures described are major or minor components of the samples that were analyzed (relative to other determine structures). Therefore, particle numbers should be included for 3D classification steps, and if any other medium-high resolution structures came from the same datasets, those should be included in the chart for the benefit of the reader. If the authors wish not to disclose unpublished structures, they may elect to exclude some of these, but the overall breakdown of the data needs to be clear. If other structures produced by the dataset are already published, this is fine.

We have addressed all of these points. Please see responses to points 8, 9, and 10 of reviewer #2.

– Data collection tables need to be included, as do tables describing any "pseudo-atomic models." This is especially true for structures that will be deposited in the PDB. It looks like 5F, 5G, and 6A show structures that should be treated this way. For parts the authors do not wish to model explicitly (low-resolution parts corresponding to Scc2/Pds5), they can exclude these from the PDB and carry out model-map correlation calculations using maps truncated to the modeled residues.

See above. We have also added a fit of the three fitted maps used in Figures 5 and 6 as requested in the form of Supplementary Figure 6.

– 2D classification results can be unreliable, especially when there are flexible modules. The analysis shown in Figure S4A-C should be removed, and the text should be modified so that any conclusions made from these images are removed. In particular, the consequences of ATP binding, discussed in relation to Figure S4C is inappropriate given that (1) the authors cannot resolve nucleotide with these images (is it really bound? hydrolyzed? Were there other classes in which the opposite was true?); (2) designation of Scc2 density is reasonable but not probative; and (3) it is not clear what the relationship is between the cohesin head domains. Subtle changes in head engagement could make huge differences in the interpretation. In the displayed images, it is not entirely clear that the heads are even engaged (versus "juxtaposed").

We assume the reviewer refers to Figure S5 here and not S4. In that case, we believe that the conclusions made from the Figures S5A-C are all within reason, i.e. the 2D classes clearly show that (1) cohesin folds without HAWKS, (2) that ATP heads are floppy relative to the folded coiled-coil, and (3) that head engagement and Scc2 binding are not enough for coiled-coil unzipping.

However, we have taken the reviewer’s points into consideration and agree that

comparison of (3) with a sample without ATP would make these points even clearer, which is why we have now added this to help clarify these points. They also demonstrate the consequences of ATP binding while lacking the atomic resolution to observe the ATP molecule. We also agree that S5A does not add anything to the manuscript that has not been shown in Figure 5 previously and so have removed it.

We have no reason to believe the heads in S5C are juxtaposed as the changes are a clear consequence of the presence of ATP in the sample, and that the removal of Scc2 allows us to produce maps that show clear engagement like that of Figure 5G.

– How confident are the authors in their positioning of the Pds5 module and cohesin head domains in Figure 6A? The density below the "joint" as drawn is very difficult to interpret and looks like it might be artifactual.

We are very confident in the positioning of the Pds5 model and refer the reviewer to the provided maps to have a look for themselves. The resolution, although not high enough for any detailed modelling, really does only allow a single pose of Pds5 and the heads.

There is no reason to believe this density is artefactual as none of the processing had included neither the atomic structure of Pds5 nor the heads as an initial model.

– The authors should compare the cohesin head domain conformations they observe, especially for the Scc2 complex, to those observed in the recent structures showing cohesin-nucleotide-Scc3-Scc2-DNA interactions. Are the current structures well enough resolved to make these comparisons? Can the authors see Scc1-N:Smc3-head domain contact?

Please see below. Figure S5D does this by comparing the binding pose of Scc2 of the ES/EK state with that of our manuscript.

We can indeed see both the Scc1-N:Smc1 and Scc1-C:Smc3 contacts at lower thresholds.

– Do the authors truly know the cohesin structure displayed in 5G is ATP-bound? To make this statement, they would need to resolve nucleotide and a connected γ phosphate.

We are confident that Figure 5G shows an ATP-bound map. The structure could only be found in the presence of ATP – a molecule that binds the ATPase heads and causes engagement – and it perfectly recapitulates the crystal structures of engaged heads bound to ATP. In light of this it would seem unreasonable to us to assume that the structure is in an apo state.

– Why did the authors leave out Scc3 from the cryo-EM samples? How do they relate their observations to the published structures, which show the hinge attached to Scc3/Psc3 and dislocated from the Smc1/3 head domains? Both conformations probably inform on mechanism, but the reader is not presented with a helpful way to think about this. Does Scc3 "grab" the hinge after head-Scc2 engagement, as proposed by Uhlmann's group recently (Higashi et al. 2020)? To be sure, Srinivasan et al. proposed an Scc3-dependent late step in loading already, so both of these could easily be referenced in a paragraph contextualizing the current structures with respect to Scc3.

We were looking at the putative simultaneous interaction of single HAWK proteins with both hinge and heads and therefore wanted to avoid the addition of another confounding variable in the form of Scc3. We do not present any structural data regarding Scc3 in our manuscript and therefore have abstained from making any conclusions about its potential “grabbing mechanism” as we cannot support it with evidence.

4. There are disagreements between data and text and confusing inconsistencies in the figures:

o S1C: why is Y40A not an scc4-ts allele?

Because it grows at the restrictive temperature of 37°C (Figure S1A). We don’t know why Y40A, like Y40N, disrupts co-immunoprecipitation of Scc4 and Scc2 but does not exhibit temperature sensitivity. Presumably, the temperature sensitivity of Y40N cannot solely be due to disruption of Scc2 binding.

o S1E – It looks like smc1D588Y is an scc2-4 suppressor. The text says the opposite.

As above, this has been amended.

o Figure S3 is discussed below.

See Below.

o Genotypes should be given in color for all tetrad dissections (as in Figure 1).

Done.

o Size markers are missing from the blots. These are important for the crosslinking experiments so that positions can be compared for different antibodies (Figure 4B, for example).

Size markers are now shown on crosslinking blots.

o In at least once case, the authors should show a myc blot for Smc1-WT (no Cys mutation) so that it is clear the higher band is indeed an Smc1 crosslinking product.

This is shown in Lane 1 of Figure 4G.

o For these assays, the text and legend disagree on which subunit was immobilized for the pulldown (Scc1 or Smc1). If it was different in different experiments, this should be stated.

This has been corrected, many thanks.

o Figure S4F is not helpful. There is no control shown (either Scc2 or Smc1, and ideally both, lacking Cys mutation). The crosslinking efficiencies do not seem to correlate for the two subunits, and it is not clear why some Scc2 mutants show an even higher band.

In the end, the experiment would have only been valuable if a really great crosslink position were found. The experiment can be removed without seriously damaging the manuscript.

As described previously, we agree and have removed the figure.

– The scatter plot in figure S3E does not contradict the Lopez-Serra finding and is not a good way to look at this data. The Lopez-Serra paper addressed a small subset of chromosomal locations. It is not clear how far away from the diagonal the distribution of dots would need to spread to constitute a truly different localization pattern (no control for this).

The scatterplots have been removed.

– If the authors are sure of their placements for Scc2 and Pds5, then why not directly compare the two structures by overlaying them anchored on the coiled-coiled/hinge domains? This would be a great way of showing that (i) Pds5 is indeed more closely connected to the hinge and (ii) both modules engage the same overall cohesin conformation/fold (or not, as the case may be).

We thank the reviewer for the suggestion and have added a comparison to the Supplemental Figure 5D to show the region of clashes.

– Figure 5F-G make it look like the two structures are different views of the same. Only G is required for the point that head engagement does not unzip the coiled coils, so there is no need to make it look like a comparison.

We believe it to be helpful to show the respective resolutions of each map discussed in the figure, but we agree it may cause some confusion, so we have removed the arrow signalling head engagement.

– Figure S5D does not give the reader enough information to evaluate what is being shown. What dataset does this map come from? Is the corresponding main figure 5A? Was Smc3 included in the fit? DNA? Was nucleotide included in the sample? Why is there a large part of Scc2 poking out (presumably related to mentioned "floppiness")?

Figure S5D intends to show a comparison between our map and the binding pose of Scc2 described for the ES/EK state in Collier et al., 2020. The map originates from the same data set as that of Figure 5A and has been processed to remove the floppy part to produce a map with a higher resolution of the binding of Scc2 to Smc1 to allow for a better comparison. We appreciate that this information was missing from the figure legend and have now included it.

It now reads: “(D) Fitting of atomic map from Collier et al. (6ZZ6) 2020 in cryo-EM map made by focused classification. The map originates from the same data as that of Figure 5A and has been processed to remove the floppy C-terminal head domain of Scc2”.

– Neither the Murayama papers nor the Huis in 't Veld papers are discussed. The Huis in 't Veld paper reports contact between SA1 and the cohesin hinge. Murayama and Uhlmann (2015) reconstitute this interaction (Psm1/3-Psc3) for the fission yeast components and state that it means the ring must bend and that the loader is probably involved. While neither study proved hinge-Scc3 contact is essential for the loading reaction, they were important contributions and should be cited.

Thank you. The papers have now been cited and the following added to the introduction:

“Further, it has been noted that a potential simultaneous interaction of a HAWK with the hinge and kleisin would require some sort of folding (Murayama and Uhlmann, 2015, Huis in 't Veld et al., 2014, Bürmann et al., 2019).”

– Presentation issues for structures:

o At least once in each figure (but not necessarily each panel), colored text should be shown for each included protein as is done in Figure 1. This is missing in Figure 5 (for example).

Done.

o The overlay in Figure 1F is hard to understand. The mutated structure should be shaded differently so the reader can see the difference clearly (pale green vs green and same for red).

Done.

o The nucleosome structure in Figure 3B is impossible to understand. The entire particle should be shown so the reader can see the DNA wrap, which is relevant here (do the mutations enhance unwrapping?). Only the mutated histones and mutated residues (highlight color) need to be colored.

Done.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined in the points made by reviewers below. It is particularly important to clarify where the crosslinks reside in the cohesin variant used for structural analysis, as raised in Points 2 and 3 from Reviewer 2. If the crosslinks are in the heads, as indicated in the methods section, then this has important implications for the interpretation.

We thank the reviewers for their keen eye, particularly with reference to the points on head crosslinking which was indeed a mistake on our part in the methods section. Please see our responses to each comment below for more details.

Reviewer #2:

The authors have addressed most comments. There are however a couple of issues that require further clarification:

1. Figure 4G: The author's response is only partially convincing: If Smc1K620C also crosslinks to Scc2C224, it is not fully clear that Scc2N220C specifically allows readout of contacts between the Smc1 hinge and Scc2. As they have tested a variety of Scc2 cysteine substitutions, it would have been important to test a Scc2 version that does not contain C224, in order to avoid such confounding non-specific crosslinking.

In any case, they need to further clarify how the different blots are developed. Presumably, the top panel shows an anti-FLAG western blot and the bottom panel an anti-myc blot (as indicated in the author's response)? The top panel is not labeled at all and the bottom panel is mislabeled as Anti-HA!

The reviewer is quite right, the top panel is anti-FLAG and the bottom is anti-myc. The anti-HA label corresponds the small panel in Figure 4B. We see how this is confusing and have modified the figure to make it clearer.

2. The authors state line 523 that they use a cohesin variant 'containing cysteines specifically crosslinking the three intermolecular interfaces'. They need to explicitly state which interfaces are crosslinked in the variant they use for structure determination. The author response indicates that 'the 6-cysteine version used interrogates the hinge dimerisation and not the head dimerisation'. Their methods section indicates however that the variant they use for structure determination does not contain Cys mutations in the hinges. Instead, they use Smc1G22C N1192C, Smc3S1043C R1222C, Scc1C56A547C.

These mutations are located (1) in the Smc3-NScc1 interface (Scc1C56 and Smc3S1043C), (2) Smc1-cScc1 (Scc1A547C and Smc1G22C) interface and (3), in the interface between the Smc1 and Smc3 ATPase heads (Smc1 N1192C and Smc3 R1222C). That is, the third pair of cysteines is clearly placed in the heads and not the hinges.

We profoundly apologise for the confusion caused by our mistake and thank the reviewer for pointing this out again. The correct cysteines are Smc1K639C-Smc3E570C, Smc1G22C-Scc1A547C, and Smc3S1043C-Scc1C56. We have amended the manuscript accordingly in the respective places. Therefore, our original interpretation stands, i.e. the ATPase heads have not been crosslinked to each other.

3. It is therefore not clear why the authors maintain (Reviewer #2 points 6, 7, 8, 12) that 'we did not cross-link the ATP head dimerisation interface'. Clearly, something is wrong! Maybe it is the methods section? If not, the authors need to clearly indicate that the version of cohesin they use is crosslinked at the SMC1/3 ATPase head domains. The implications of such head crosslinking need to be taken into account in the interpretation of their data and discussion of the results.

See above.

Reviewer #3:

The manuscript is greatly improved. The updated figures make it much clearer. The rewritten section describing Sth1, nucleosome positioning, and histone mutants is good. This is important work, and the breadth and strength of the experiments described are impressive.

Most of my comments have been adequately addressed. The requests below relate to the presentation and description of the data and should be addressed before publication but do not require extensive writing or any new experiments.

Line 593-597:

"In the presence of ATP" implies that ATP occupies both ATPase active sites. Neither γ phosphate can be resolved at this resolution. The structure could be a reaction intermediate resulting from slow π dissociation from one or both sites, for example. This is not to detract from the observations or statements made. Instead, it would be wise to state more clearly that ATP was included in the sample but avoid any suggestion that the structure provides detailed information about the SMC ATPase cycle, which will doubtless be a topic for future studies. This is rightly stated in the final paragraph of the Discussion.

We assume that the reviewer refers to one or both of these instances: “We identified and solved … a form of cohesin at 6Å … whose ATPase heads were engaged in the presence of ATP” and/or “A cohesin complex whose heads are engaged in the presence of ATP…”

If so, we respectfully disagree with the reviewer. What is precisely stated in our manuscript is that our engaged cohesin heads, which we explicitly mention are at no more than 6 Å resolution, were in the presence of ATP. We never allude to the state of the molecule or even hint at the possibility of seeing the gamma phosphate at this resolution. Adding that the heads could also be bound to ADP and Pi or even be in a transition would just serve to confuse the reader in an, in our opinion, very straightforward and factual statement of the sample conditions.

Figure 5G needs to be modified. Figure 5S2 shows that 5G is a composite structure derived from two different samples. One contained ATP and the other did not. Overlaying them is inappropriate. Yes, the angle of the head-proximal coiled-coils is similar (Figure 5F vs. 5G), and it is likely that the remainder of the CC density beyond (hinge-proximal) the joint is similar/identical, but this is not proven by the data in the manuscript. In fact, it appears this section of the complex was excluded from analysis in the case of the sample containing nucleotide (Figure 5G and 5S2D). Without recalculating structures or even remaking complicated figures, it must be possible to present this data more clearly. One possibility: simply removing the "Arm" density from figure 5G. Another possibility is to compare the head domain density (including head-proximal cc) from Figure 5F, which is currently obscured by Scc2 density, with the head domain reconstruction from 5G.

We thank the reviewer for the suggestion and we agree that the figure should be more transparent. We have removed the separating discontinuous line that suggested that these were two structures from different datasets and have added that the folded coiled-coil was derived from a dataset without ATP to the figure. In addition, we have referred the reader explicitly to figure 5S2D to see how the map was obtained. However, we believe that the composite structure does add valuable information that aids the reader’s understanding by putting the engaged ATPase heads into the structural context of a folded cohesion and have therefore decided to leave it with the mentioned remarks. Especially given the 2D classes in Figure 5 Supplement 1E.

Figure 5 – supplement 1A does not conclusively show complexes lacking Scc2. Compare with 5S1A, for example. One needs to consider variability in Scc2 occupancy, Scc2 position, and also the fraction and quality of the particle images contributing to these classes. The description "in samples lacking all HAWK proteins" (line 537) therefore overstates the certainty of the claim given the evidence. The statement and figure panel could be removed without damaging the overall message. Removing panel A has the added benefit that it eliminates the need to discuss a dataset not referenced in figure 5S2 (cohesin, Scc2, ATP).

The reviewer is absolutely right – this was due to a mistake while reorganising the supplementary figures 5 during the previous revision. We have added 2D classes showing exactly that, i.e, a folded cohesion complex in the absence of HAWK proteins to the figure (cf. Figure 5S1E) and have amended the text that referred to the wrong figure.

https://doi.org/10.7554/eLife.67268.sa1

Author response

Reviewer #1:

Comments for the authors:

1. The authors should considering shortening the section on RSC describing Figure S3 as it detracts from the main message of the manuscript. In addition, while the ChIP-seq in Figure S3D argue against a major role for RSC in cohesin loading, the nucleosome mapping data in Figure S3S are not conclusive, as presented. Why do the upper two and lower two nucleosome profiles look different from each other? Potentially effects on nucleosome positioning will only be visible in metagene analysis: the sth1-3 mutant at least would be expected to alter nucleosome positioning but this is not apparent from the data presented. The authors should either remove this data or perform additional analysis.

We have removed the nucleosome positioning data in Figure S3G and amended the text accordingly.

2. Figure 7G: The conclusion that cohesin folding at the elbow is a feature of cohesin complexes that are holding sister chromatids together relies on the assumption that all acetylated cohesin is participating in cohesion. However, this may not be the case. In mammals, cohesin acetylation is also observed in G1, indicating that it may have a more general role. This potential caveat should be mentioned.

Our work is exclusively about what happens in yeast, where acetylation is linked to replication, where sororin is absent, and where acetylation (not sororin association) is the best (known) marker for cohesion. The fact that acetylation is clearly not a good marker for cohesion in mammalian cells does not therefore merit a “caveat”. As far as we know, acetylation is as good a marker in yeast as is sororin association in mammalian cells. The reviewer is correct that this should be pointed out and we have altered the text accordingly.

3. It is surprising that there is high Scc1 ChIP-seq signal at centromeres in scc4-4 cells if Scc4 is required for cohesin loading at centromeres (Figure S4A). How do the authors explain this?

It has been previously documented that there is an Scc4-independent mechanism that promotes centromeric loading in the presence of nocodazole (Petela et al. 2018). As this experiment was done in the presence of nocodazole, this mechanism is likely responsible for the high Scc1 ChIP-seq signal observed here.

4. Line 273: It looks like smc1D588Y CAN rescue the proliferation defect of scc2-4 in Figure S1E.

This is quite right, and the text has been amended accordingly.

5. Figure 3A please show proliferation data for the different histone mutants.

We show proliferation data for the mutant that we use for all subsequent experiments in Figure 3A. This mutant was made de novo and is a good representative for the other mutants. We think showing additional proliferation data for all the mutants is therefore unnecessary.

6. Line 402: Smc3 acetylation

Corrected, many thanks.

7. Supplementary Figure 4F: needs a negative control without cysteine substitutions.

In line with comments from the other reviewers, we agree this panel is not helpful and not needed and therefore have removed it and amended the text accordingly.

8. Page 19: Line 547: Call to Figure S4D is incorrect.

Corrected, many thanks again.

Reviewer #2:

The conclusions are mostly well supported by the data, but some aspects of the biochemical and structural data require clarification.

1. Biochemical data: Figure 4B: The FLAG blot indicates that x-linked species migrate at different positions on SDS-PAGE while the top panel shows uniform mobility. How do we know that the band labeled SMC1-? corresponds to x-linked SMC1 and not some non-specific anti-myc background (e.g. there is an additional non-labeled band running lower)?

The Myc blot shows multiple crosslinked species corresponding to Smc1 crosslinking to multiple proteins. As can be seen from the FLAG blot these species migrate at slightly different positions and have different efficiencies. The most efficient crosslink by far is to Pds5, corresponding to the intense band labelled “Smc1-?” in the Myc blot, which is present in all cases and does mask the other species. It should be noted that this band runs slightly higher in the Pds5-FLAG lane, due to the tag increasing the molecular weight. This crosslinking species has been documented previously, with additional controls, in Bürmann et al., 2019.

We do understand however that our labelling of the blot may have been misleading and so have tried to improve the clarity by amending the “Smc1-?” label to reflect the multiple species present.

Figure 4B: Different amounts of FLAG-HAWKs are recovered in the anti-Myc IP. Can the authors generate an 'input' blot showing expression levels to exclude the possibility that the variable crosslinking efficiency is not due to different expression levels?

We agree this would be useful and have modified the figure panel to include the expression levels of Scc2, Scc3 and Pds5 in whole cell extracts relative to a loading control. Indeed, the proteins are expressed to different levels as expected but this does not correlate to the crosslinking efficiency. Pds5 for example is not as abundant as Scc3 but crosslinks much more efficiently.

2. Figure S4F: Show a WT negative control (no Cys mutation) to confirm that the bands observed do not arise due to non-specific background cross-linking.

As detailed above, this figure panel has been removed and the text amended accordingly.

3. Figure 4G: Top: Why is there a crosslinked band in the lane containing Smc1K620C alone? Would it not be expected to see cross-linking only when both Scc2N200C and Smc1K620C are present? Why is the non-specific crosslinking (labeled as *) present everywhere? Which cross-linked band they are referring to? While there is a small amount of a band visible above the non-specific band, this weak band is also present in Smc1K620C alone!

The crosslinking band observed in the lane containing only Smc1K620C is most likely due to Smc1K620C crosslinking to a natural cysteine in Scc2, probably C224 which sits on a small helix just under that containing N200. This would explain the very similar molecular weight, the fact the band also appears in the Myc blot and the increased intensity of the band on addition of Scc2N200C. The text has been modified to reflect this. We do not know the identity of the non-specific crosslinking band present in the Myc blot but it is clearly independent of either cysteine mutations as it is observed in lane 1. To improve the clarity, we have amended the labelling.

4. Structural data: Figure 5A: The authors need to clarify if in their cryoEM structure, the SMC ATPase heads are in the 'apo' state, as indicated in the text or in the engaged state as indicated in the Discussion (L. 641).

Changed. Thank you, this was a typo that has now been corrected.

5. The authors need to explicitly state in their description of their structural data (Line 512 et seq) that they used a crosslinked version of the cohesin trimer containing 6 Cysteines, positioned at the SMC-SCC1 and SMC1-3 ATPase head heterodimerization interfaces.

Done, but please bear in mind, as below, that the 6-cysteine version used interrogates the hinge dimerisation and not the head dimerisation.

6. Crosslinking would be expected to stabilize the SMC ATP heads in close proximity, potentially an engaged state. Would full SMC head disengagement be prevented by such cross-linking even in the absence of nucleotide?

See last point: we did not cross-link the ATP head dimerisation interface and therefore would not expect head (dis)engagement to be affected in any way.

7. From Figure 5 the state of the heads is not immediately apparent. Please include a Figure comparing the state of the 'disengaged' SMC heads obtained using their cross-linking method with that of the ATP engaged state published previously.

Again, the cross-linking does not affect the head engagement of the molecule and at this resolution it would seem somewhat over-interpretative to analyse the precise placement of the head subunits.

8. If indeed, crosslinking stabilizes an engaged but nucleotide-free form, the authors need explicitly discuss the potential implications. For example, crosslinking of the heads could prevent the heads from properly disengaging. This has potential implications for the conformation/interactions of Scc2 or PDS5.

Again, see above. The head dimerisation domains were not cross-linked.

9. Figure 5G: The authors need to clarify how they obtain the ATP-bound state. Did they also use a crosslinked cohesin trimer, hydrolysis-impaired mutant ATPase heads or non-hydrolyzable ATP variants?

As described, the ATP-bound state was simply achieved by incubating the trimer with ATP. For clarification we have added that no cross-linker was used.

10. What is the distance between Smc1D588 and SCC2? Figure 5B indicates ~25Å which would be too far away for a direct interaction. In contrast, Figure 5E and their text description suggest that 'folding of the coiled coil brings the hinge to within 12nm (they probably mean Å?) of the heads'. Figure 5E indicates a distance of 12Å but it is between Smc1K620BPA and Smc1D588Y and not SCC2.

As described in Figure 5B, the distance between Scc2 and Smc1D588 is indeed 25 Å. This refers to the distance of the best cryo-EM map reconstruction (which by definition is an average) and therefore only represents a single snapshot of the entire range of Scc2’s movements above the neck as mentioned in the manuscript. The distance is supposed to highlight this flexibility when comparing it to the hinge-bound structures published previously (e.g. Shi et al. 2020).

We apologise if this was unclear and have reworded the following to aid with clarity:

“Initial 2D classes of Scc2-bound cohesin revealed (Figure S5B) floppiness not only within the HAWK, especially in its C-terminus, but also between the joint and the ATPase heads.”

We recognise the phrase 'folding of the coiled coil brings the hinge to within 12 nm of the heads’ is misleading and should have referred to the ATPase heads to be precise and so as to avoid confusion with the HAWK’s head. We have now changed this accordingly

11. If indeed no direct interaction is apparent, or if the low resolution and flexibility do not allow firm conclusions, they need to indicate this during their discussion of these results.

Our manuscript never claims to observe a rigid interaction between Scc2 and the hinge – and the 25 Å distance in Figure 5 reaffirms this – but rather that the folding of the elbow is fundamental in allowing for it and that the clear interaction observed in previously published structures must stem from an intrinsic floppiness in the head region of Scc2 that we also observe in our processing.

12. Figure 6: While biochemical data indicate that PDS5 inhibits ATP hydrolysis, the presented structural data does not reveal how this is accomplished. One caveat of using a cross-linked version is that the procedure may prevent access of PDS5 to fully disengaged SMC heads and thus prevent SMC head engagement (in analogy to the role of the condensin YCS4 subunit). The authors need to indicate potential pitfalls of their cross-linking procedure on PDS5 positioning and discuss if the conformation observed is potentially off pathway.

See above. Head engagement should not be affected in any way by the crosslinking as the third pair of cysteines are placed in the hinge and not the heads.

Comments for the authors:

1. Figure 4C-F: Please indicate if the experiment has been done once or repeated several times with consistency.

The experiments were performed twice with the same results. This is now stated in the figure legend.

2. Can they indicate if their preparation used for CryoEM is fully or partially BMOE cross-linked (e.g. by SDS-PAGE analysis).

The estimated efficiency of cross-linking is around 20% as previously mentioned in Collier et al., 2020 (also see SDS-PAGE gel in Author response image 1; the total intensity of the SMCs [including the smear] and Scc1 from lane 1 [trimer crosslinking no cysteines] compared to the 6C crosslinking from the top band from lane 8 [trimer crosslinking 6C]). We have now added that to the manuscript as suggested:

“…To investigate this further, we used cryo-EM to determine the structures of the S. cerevisiae cohesin trimer (Smc1, Smc3, Scc1, in their 6C version as reported in Collier et al., 2020; specifically cross-linked at the three intermolecular interfaces at an efficiency of 20% (data not shown)) bound to either Scc2 (Figure 5A)”

Author response image 1

3. Figure 6: While at lower resolution, it would still be important to show how well the PDB models fit into the cryoEM density map shown in Figure 6A. This would give the reader a sense for how reliable positioning of the PDB models (shown in Figure 6B) are.

Agreed and done.

4. Can they indicate where the previously published (Rowland et al. 2009, Sutani et al. 2009) eco1-1 suppressor mutations of PDS5 are located? Are suppressor mutations located in observed interfaces? Would they be predicted to interfere with PDS5 interaction?

The suppressor mutation cluster around residues 81-89 in PDS5 reported by Rowland et al. would sit at the hinge:Pds5 interaction interface. The rest of the mutations do not reside in any of the other interfaces predicted by our model. However, as we have explained in the manuscript, the density around the N-terminus of Pds5 is not ideal and we would prefer to refrain from making any too detailed conclusions.

5. L.593 et seq: The authors mention that their data are in contrast to 'previous studies with shortened constructs'. Previous work needs to be cited.

We have added a reference to Muir et al., 2020.

6. L280: This should read: 'relative to the position of D574'?

Thank you, this has been corrected.

7. Figure S4E: Cartoon shows SMC3! Do they mean SMC1?

Yes it should read Smc1, this has also been corrected.

8. Details on CryoEM data processing and reconstruction information need to be included.

We have added a detailed overview of cryo-EM data processing and reconstruction workflows for all maps in Supplementary Figure 7.

9. PDB validation reports of the modeled cryoEM structures are missing.

PDB and EMDB validation reports for all the maps have been attached. The accession codes have been added both to the figure legends and the text. The Table of Structures” in Materials and methods details this for each map:

10. It is not clear how the PDB model for the cohesin elbow is derived. No details of CryoEM/Xray data collection are given and PDB validation reports are again missing.

Please be referred to point 8. We have also added more detail in the Methods section about the model building of the cohesin elbow PDB. In addition, we have added Supplementary Table 3 with information about data collection.

11. L. 1029: Spelling error: Katsu Shirahiga

Thank you, this has been corrected.

Reviewer #3:

Chemical crosslinking experiments have established that sister chromatid cohesion requires "topological embrace" of sister chromatids within a cohesin ring. Three contacts define the ring: Scc1-N:Smc3-head, Scc1-C:Smc1-head, and Smc1-hinge:Smc3-hinge. The strongest experimental evidence for a specific DNA entry point is that artificial dimerization of the Smc1-hinge:Smc3-hinge interface prevents cohesion establishment (Gruber et al. 2006; Srinivasan et al. 2018). In the current manuscript, Patela et al. present genetic, biochemical, and structural data that indicate that the cohesin hinge interacts with the cohesin loading complex (Scc2/4), that this interaction depends on Scc4, and that mutations in the hinge that appear to enhance the interaction positively affect cohesin's association with chromosomes. Considered together, these data provide new evidence that the cohesin hinge is connected to the loading reaction. The manuscript is really interesting, includes many very clever experiments, and extends previous models for how cohesin works.

There are major issues with the paper that need to be addressed before publication. First, the paper is confusing. It should be simplified so that the main points are clearer, and errors in the text and figures should be corrected. Second, the section about nucleosome remodeling must be rewritten. There is great value in pointing out differences between studies, but these points can be made more succinctly. Third, the descriptions of the structures are insufficient and do not permit comparison with published structures. The reader cannot currently judge the validity of the interpretations presented.

With the exception of some minor updates to the biochemical experiments, the majority of these changes can be made without adding new data. The rewriting and corrections I have requested will nevertheless require significant time and attention. In addition, I have also requested the EM maps and corresponding docked models be made available to reviewers before a preliminary decision is reached. After this is done, I would support publication of a substantially updated version of the current manuscript in eLife.

1. The manuscript contains a collection of interesting observations, all of which address the overall hypothesis that Scc2/4 stimulates cohesion establishment by interacting with the cohesin hinge.

– See below for comments on the structures. The Pds5 structure should be moved to the supplemental material or removed altogether. The new structure is good enough to discern the overall position of Pds5 relative to cohesin, but it does not provide much new reliable information beyond this. It is pertinent in that it confirms competition between Scc2 and Pds5. Finer details like the status of the head domains, the identity of specific contacts, and the relationship to cohesin acetylation/Eco1 binding will have to wait for higher-resolution structures that will ideally be trapped in defined identifiable biochemical states.

We respectfully disagree with the reviewer. The overall position of Pds5 relative to cohesin is of a clear interest to the wider cohesin community as it not only provides the first visualisation of the interaction but starts to hint at the answers to fundamental questions of this interaction. At the same time, it finally allows much needed further biochemical work to interrogate said interaction that would not be possible without it. It also serves as structural confirmation of the biochemical/genetic data reported in this paper and therefore plays an important role in the manuscript.

– Like the Smc1/3 E-to-Q mutants, smc1D588Y appears to slow or stall cohesin at an early/intermediate step in the loading process (Figure 2C-D). The cells are viable, so the stall is not terminal. How do the authors reconcile this observation with their finding that cohesin loading on chromosome arms is actually elevated relative to wild type?

We believe we addressed this point in the discussion “Why does smc1D588Y depress loading at CENs?”

In particular:

“We suggest that clamping is rate limiting along chromosome arms but unclamping is rate limiting at CENs and that this is the reason why smc1D588Y enhances arm loading while depressing loading at CENs.”

– Related to the above question: Have the authors explored how the mutation affects engagement with chromosomal DNA analogous to the recent Chappard paper? Are S-K complexes depleted in favor of others? Can they observe so-called E or J heads, and is either enriched in the mutant background? One line 353, the authors stop short of suggesting the smc1D588Y mutant favors the clamped state but imply it may stop the loading reaction at a step after the one stalled by the clamped-favoring EQ mutations. The current data do not conclusively show that the step slowed by smc1D588Y is before or after the step stalled by the EQ mutations. The authors should remove speculation on this point from the Results section.

Although these are valid questions that would be interesting to address, the experiments required to address them are not trivial and we do not think the answers are necessary for this manuscript, which is already quite long. Concerning the second point, we merely point out that the enhanced association of both Scc2 or EQ complexes at CENs caused by D588Y suggests that the mutation does not affect formation of the clamped state at CENs in vivo but some later step in the loading reaction. We do not agree that this is an unjustified speculation.

– The description of results that begins on line 336 is very confusing. Specifically, "This implies that the reduced loading arises…" but the authors say directly above that Smc1D588Y is at 110% (CENs) of WT. Also, the first two sentences of this paragraph use "it," but the referent is not clear. Last, a simple explanation of the phenotype shown in Figure 2 seems to be that smc1D588Y preserves CEN targeting but disrupts translocation to pericentromeres. Arm loading is not disrupted (slightly enhanced). A simpler description of the phenotype would make the paragraph much easier to read and focus the reader on the important question: what accounts for the subtle defects seen in the smc1D588Y mutant?

The sentence now reads: “This implies that the reduced loading around centromeres arises not from defective formation of the clamped state at CENs by Scc2/4 complexes associated with Ctf19 but from a defect in a subsequent step in the loading/translocation reaction that requires ATP hydrolysis.”

– Line 456 – "Nevertheless, the level of Smc1-Scc2 crosslinked protein was comparable to that of Scc3…" This is an inappropriate comparison. Crosslinking efficiency could be far more efficient if there is a fortuitous lysine on the target (Scc3 or Scc2 here).

The reviewer is of course correct. What we merely implied is that irrespective of such an effect, may or may not be the case, the amount of Smc1-Scc2 crosslinking was comparable to that of Smc1-Scc3 despite Scc2 being less abundant in the cell. This is surely a valid point to make.

– Line 562 – This is a concrete biochemical prediction that is totally testable. Since it is central to the proposed mechanism of suppression, it should be tested using pulldown assays with purified proteins (cohesin tetramer, Scc2/4 or Scc2-C).

This refers to the statement “We suggest that the addition of a bulky amino acid into Smc1 through D588Y may be sufficient to help bind an otherwise floppy Scc2 N-terminal domain, whose interaction with the hinge is normally stabilised by Scc4”. We disagree that this is a testable hypothesis using pulldown assays as we are referring to the stabilisation of a particular conformation of binding not binding at all, which itself was difficult to measure in this way due to the multiple contacts Scc2 makes with the tetramer (Petela et al. 2018). Although this would be good to address, doing so is not trivial.

2. The section describing the RSC-Scc4 interaction needs to be revised. The authors focus on and oversimplify a secondary conclusion from a previous paper (Lopez-Serra et al. 2014), which holds that Scc4 stimulates RSC. The majority of the work and discussion in this and a related paper from the same group (Lopez-Serra et al. 2014 and Munoz et al. 2020) indicate that Scc2/4 localization depends on RSC. The Munoz paper offers a biochemical explanation for this observation but does not follow up on the secondary observation from the first paper that Scc2/4 influences nucleosome positioning.

– The current findings regarding the genome-wide distribution of cohesin in sth1-3, smc1D588Y, or related mutants seem mostly consistent with the results presented in Lopez-Serra and in Munoz. The authors should revise their text to reflect this concordance. In fact, all three studies emphasize the importance of nucleosome-free DNA. This could be pointed out.

We are grateful to the reviewer for pointing out this deficiency in our discussion of the papers from the Uhlmann lab. We now point out in our revised manuscript that they have in fact proposed two, possibly contradictory, hypotheses for how Scc2/4 functions together with RSC. Crucially, both are inconsistent with our finding that sth1-3 causes only a modest if any defect in cohesin loading in vivo. We would also like to point out that the conclusion that Scc4 acts by stimulating RSC cannot be considered merely a minor secondary conclusion of the Lopez-Serra paper. The very title of their paper was “Scc2/4 acts in sister chromatid cohesion by …maintaining nucleosome regions”. We also respectfully disagree with the reviewer’s statement that the genome-wide distribution of cohesin in sth1-3, smc1D588Y, or related mutants seem mostly consistent with the results presented in Lopez-Serra and in Munoz. We do not believe in papering over the cracks in this manner. There is a clear inconsistency between our calibrated ChIP-seq data and their anecdotal qPCR measurements. We have nevertheless greatly revised our discussion of this issue and hope it that it more accurately describes the claims made by the Uhlmann lab while at the same time being easier to read.

– Any differences in the cohesin distribution between the two studies (Patela and Lopez-Serra), especially in the sth1-3 mutant must be more carefully annotated. Direct comparisons of the original studied loci (Lopez-Serra et al. Figure 3B, for example) must be presented if the authors wish to question the previous findings.

We have now included comparisons of the original studied loci.

– That artificial chromosome tethering of Scc2-C rescues scc4D (Munoz Figure 6B) suggests the partially Scc4-dependent hinge interaction described here is not an essential function of Scc4. The authors suggest that this Scc4 function is rendered irrelevant when the loader is brought near DNA. At this point, there is not enough information to discriminate between these very similar models, and it would be better not to set the current manuscript up in opposition to previous work when an experimental distinction has not been made.

We have now revised this aspect of the discussion.

– Was the MNase-seq calibrated? Also, it looks like there are differences in nucleosome occupancy around AST2, which was also highlighted in the Lopez-Serra work (Figure S3G here). How are the two WT panels different? They show very different results.

In line with comments from the other reviewers we have removed this data and amended the text accordingly.

– The authors say that RSC is not uniquely required for Scc2/4 recruitment, but Figure 3A from Lopez-Serra et al. shows that sth1-3 produces a pronounced cohesion defect, and other chromatin modifying enzyme mutants do not.

We do not discuss the involvement of nucleosome remodellers in sister chromatid cohesion. The observations we discuss relate only to the occupancy of cohesin on DNA and cannot distinguish cohesive and non-cohesive complexes. We note that there may be a greater defect in Smc3 acetylation in sth1-3 mutants than there is a defect in cohesin loading (Figure S3C), which might conceivably be accompanied by cohesion defects. We did not follow this up as we were concerned with cohesin loading and not with the process of cohesion establishment and besides which it was not our goal to re-investigate the role of RSC in this process. That said, this section has been rewritten.

– Line 413 -The double mutant experiment (smc1D588Y sth1-3) is not very informative. It's unclear to me that suppression (or not) of the RSC phenotype by any of the mutants described in the current work would prove the previous sth1-3 findings to be unreliable. In addition, it would be important to compare cohesion, which is not done here. The double mutant experiments can be included, but the conclusions drawn are very limited and need to be rewritten.

This has been re-written to make it clear that even if there is a modest defect in loading in sth1 mutants, it is not altered by smc1D588Y, which bypasses the requirement for Scc4. We hope our logic is now clearer.

– Line 416 – "We were also unable to reproduce the reported effects of scc4-4 and sth1-3 on nucleosome positioning…" If the authors really wish to make this statement/comparison, they need to be much more deliberate about evaluating the reported effects and then comparing their own data. There are clearly differences in Figure S3G, but the authors do not look at them with sufficient granularity. In particular, the Lopez-Serra paper looks at Scc2-bound genes/promoters. The authors must also compare their data with a more recent report (Kubik…Shore, Mol Cell, 2018). This study shows effects specifically at the +1 nucleosome of transcribed genes and is much more in line with the Lopez-Serra observations.

In line with the comments from the other reviewers, we have removed this data and amended the text accordingly.

3. The new structures presented in the paper are compelling. There are several issues that must be cleared up before the manuscript can be published.

– It is unclear how the structures were determined. The authors should present data processing flowcharts to show how the most important structures in the paper were generated. It would be very helpful to know whether the structures described are major or minor components of the samples that were analyzed (relative to other determine structures). Therefore, particle numbers should be included for 3D classification steps, and if any other medium-high resolution structures came from the same datasets, those should be included in the chart for the benefit of the reader. If the authors wish not to disclose unpublished structures, they may elect to exclude some of these, but the overall breakdown of the data needs to be clear. If other structures produced by the dataset are already published, this is fine.

We have addressed all of these points. Please see responses to points 8, 9, and 10 of reviewer #2.

– Data collection tables need to be included, as do tables describing any "pseudo-atomic models." This is especially true for structures that will be deposited in the PDB. It looks like 5F, 5G, and 6A show structures that should be treated this way. For parts the authors do not wish to model explicitly (low-resolution parts corresponding to Scc2/Pds5), they can exclude these from the PDB and carry out model-map correlation calculations using maps truncated to the modeled residues.

See above. We have also added a fit of the three fitted maps used in Figures 5 and 6 as requested in the form of Supplementary Figure 6.

– 2D classification results can be unreliable, especially when there are flexible modules. The analysis shown in Figure S4A-C should be removed, and the text should be modified so that any conclusions made from these images are removed. In particular, the consequences of ATP binding, discussed in relation to Figure S4C is inappropriate given that (1) the authors cannot resolve nucleotide with these images (is it really bound? hydrolyzed? Were there other classes in which the opposite was true?); (2) designation of Scc2 density is reasonable but not probative; and (3) it is not clear what the relationship is between the cohesin head domains. Subtle changes in head engagement could make huge differences in the interpretation. In the displayed images, it is not entirely clear that the heads are even engaged (versus "juxtaposed").

We assume the reviewer refers to Figure S5 here and not S4. In that case, we believe that the conclusions made from the Figures S5A-C are all within reason, i.e. the 2D classes clearly show that (1) cohesin folds without HAWKS, (2) that ATP heads are floppy relative to the folded coiled-coil, and (3) that head engagement and Scc2 binding are not enough for coiled-coil unzipping.

However, we have taken the reviewer’s points into consideration and agree that

comparison of (3) with a sample without ATP would make these points even clearer, which is why we have now added this to help clarify these points. They also demonstrate the consequences of ATP binding while lacking the atomic resolution to observe the ATP molecule. We also agree that S5A does not add anything to the manuscript that has not been shown in Figure 5 previously and so have removed it.

We have no reason to believe the heads in S5C are juxtaposed as the changes are a clear consequence of the presence of ATP in the sample, and that the removal of Scc2 allows us to produce maps that show clear engagement like that of Figure 5G.

– How confident are the authors in their positioning of the Pds5 module and cohesin head domains in Figure 6A? The density below the "joint" as drawn is very difficult to interpret and looks like it might be artifactual.

We are very confident in the positioning of the Pds5 model and refer the reviewer to the provided maps to have a look for themselves. The resolution, although not high enough for any detailed modelling, really does only allow a single pose of Pds5 and the heads.

There is no reason to believe this density is artefactual as none of the processing had included neither the atomic structure of Pds5 nor the heads as an initial model.

– The authors should compare the cohesin head domain conformations they observe, especially for the Scc2 complex, to those observed in the recent structures showing cohesin-nucleotide-Scc3-Scc2-DNA interactions. Are the current structures well enough resolved to make these comparisons? Can the authors see Scc1-N:Smc3-head domain contact?

Please see below. Figure S5D does this by comparing the binding pose of Scc2 of the ES/EK state with that of our manuscript.

We can indeed see both the Scc1-N:Smc1 and Scc1-C:Smc3 contacts at lower thresholds.

– Do the authors truly know the cohesin structure displayed in 5G is ATP-bound? To make this statement, they would need to resolve nucleotide and a connected γ phosphate.

We are confident that Figure 5G shows an ATP-bound map. The structure could only be found in the presence of ATP – a molecule that binds the ATPase heads and causes engagement – and it perfectly recapitulates the crystal structures of engaged heads bound to ATP. In light of this it would seem unreasonable to us to assume that the structure is in an apo state.

– Why did the authors leave out Scc3 from the cryo-EM samples? How do they relate their observations to the published structures, which show the hinge attached to Scc3/Psc3 and dislocated from the Smc1/3 head domains? Both conformations probably inform on mechanism, but the reader is not presented with a helpful way to think about this. Does Scc3 "grab" the hinge after head-Scc2 engagement, as proposed by Uhlmann's group recently (Higashi et al. 2020)? To be sure, Srinivasan et al. proposed an Scc3-dependent late step in loading already, so both of these could easily be referenced in a paragraph contextualizing the current structures with respect to Scc3.

We were looking at the putative simultaneous interaction of single HAWK proteins with both hinge and heads and therefore wanted to avoid the addition of another confounding variable in the form of Scc3. We do not present any structural data regarding Scc3 in our manuscript and therefore have abstained from making any conclusions about its potential “grabbing mechanism” as we cannot support it with evidence.

4. There are disagreements between data and text and confusing inconsistencies in the figures:

o S1C: why is Y40A not an scc4-ts allele?

Because it grows at the restrictive temperature of 37°C (Figure S1A). We don’t know why Y40A, like Y40N, disrupts co-immunoprecipitation of Scc4 and Scc2 but does not exhibit temperature sensitivity. Presumably, the temperature sensitivity of Y40N cannot solely be due to disruption of Scc2 binding.

o S1E – It looks like smc1D588Y is an scc2-4 suppressor. The text says the opposite.

As above, this has been amended.

o Figure S3 is discussed below.

See Below.

o Genotypes should be given in color for all tetrad dissections (as in Figure 1).

Done.

o Size markers are missing from the blots. These are important for the crosslinking experiments so that positions can be compared for different antibodies (Figure 4B, for example).

Size markers are now shown on crosslinking blots.

o In at least once case, the authors should show a myc blot for Smc1-WT (no Cys mutation) so that it is clear the higher band is indeed an Smc1 crosslinking product.

This is shown in Lane 1 of Figure 4G.

o For these assays, the text and legend disagree on which subunit was immobilized for the pulldown (Scc1 or Smc1). If it was different in different experiments, this should be stated.

This has been corrected, many thanks.

o Figure S4F is not helpful. There is no control shown (either Scc2 or Smc1, and ideally both, lacking Cys mutation). The crosslinking efficiencies do not seem to correlate for the two subunits, and it is not clear why some Scc2 mutants show an even higher band.

In the end, the experiment would have only been valuable if a really great crosslink position were found. The experiment can be removed without seriously damaging the manuscript.

As described previously, we agree and have removed the figure.

– The scatter plot in figure S3E does not contradict the Lopez-Serra finding and is not a good way to look at this data. The Lopez-Serra paper addressed a small subset of chromosomal locations. It is not clear how far away from the diagonal the distribution of dots would need to spread to constitute a truly different localization pattern (no control for this).

The scatterplots have been removed.

– If the authors are sure of their placements for Scc2 and Pds5, then why not directly compare the two structures by overlaying them anchored on the coiled-coiled/hinge domains? This would be a great way of showing that i) Pds5 is indeed more closely connected to the hinge and ii) both modules engage the same overall cohesin conformation/fold (or not, as the case may be).

We thank the reviewer for the suggestion and have added a comparison to the Supplemental Figure 5D to show the region of clashes.

– Figure 5F-G make it look like the two structures are different views of the same. Only G is required for the point that head engagement does not unzip the coiled coils, so there is no need to make it look like a comparison.

We believe it to be helpful to show the respective resolutions of each map discussed in the figure, but we agree it may cause some confusion, so we have removed the arrow signalling head engagement.

– Figure S5D does not give the reader enough information to evaluate what is being shown. What dataset does this map come from? Is the corresponding main figure 5A? Was Smc3 included in the fit? DNA? Was nucleotide included in the sample? Why is there a large part of Scc2 poking out (presumably related to mentioned "floppiness")?

Figure S5D intends to show a comparison between our map and the binding pose of Scc2 described for the ES/EK state in Collier et al., 2020. The map originates from the same data set as that of Figure 5A and has been processed to remove the floppy part to produce a map with a higher resolution of the binding of Scc2 to Smc1 to allow for a better comparison. We appreciate that this information was missing from the figure legend and have now included it.

It now reads: “(D) Fitting of atomic map from Collier et al. (6ZZ6) 2020 in cryo-EM map made by focused classification. The map originates from the same data as that of Figure 5A and has been processed to remove the floppy C-terminal head domain of Scc2”.

– Neither the Murayama papers nor the Huis in 't Veld papers are discussed. The Huis in 't Veld paper reports contact between SA1 and the cohesin hinge. Murayama and Uhlmann (2015) reconstitute this interaction (Psm1/3-Psc3) for the fission yeast components and state that it means the ring must bend and that the loader is probably involved. While neither study proved hinge-Scc3 contact is essential for the loading reaction, they were important contributions and should be cited.

Thank you. The papers have now been cited and the following added to the introduction:

“Further, it has been noted that a potential simultaneous interaction of a HAWK with the hinge and kleisin would require some sort of folding (Murayama and Uhlmann, 2015, Huis in 't Veld et al., 2014, Bürmann et al., 2019).”

– Presentation issues for structures:

o At least once in each figure (but not necessarily each panel), colored text should be shown for each included protein as is done in Figure 1. This is missing in Figure 5 (for example).

Done.

o The overlay in Figure 1F is hard to understand. The mutated structure should be shaded differently so the reader can see the difference clearly (pale green vs green and same for red).

Done.

o The nucleosome structure in Figure 3B is impossible to understand. The entire particle should be shown so the reader can see the DNA wrap, which is relevant here (do the mutations enhance unwrapping?). Only the mutated histones and mutated residues (highlight color) need to be colored.

Done.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined in the points made by reviewers below. It is particularly important to clarify where the crosslinks reside in the cohesin variant used for structural analysis, as raised in Points 2 and 3 from Reviewer 2. If the crosslinks are in the heads, as indicated in the methods section, then this has important implications for the interpretation.

We thank the reviewers for their keen eye, particularly with reference to the points on head crosslinking which was indeed a mistake on our part in the methods section. Please see our responses to each comment below for more details.

Reviewer #2:

The authors have addressed most comments. There are however a couple of issues that require further clarification:

1. Figure 4G: The author's response is only partially convincing: If Smc1K620C also crosslinks to Scc2C224, it is not fully clear that Scc2N220C specifically allows readout of contacts between the Smc1 hinge and Scc2. As they have tested a variety of Scc2 cysteine substitutions, it would have been important to test a Scc2 version that does not contain C224, in order to avoid such confounding non-specific crosslinking.

In any case, they need to further clarify how the different blots are developed. Presumably, the top panel shows an anti-FLAG western blot and the bottom panel an anti-myc blot (as indicated in the author's response)? The top panel is not labeled at all and the bottom panel is mislabeled as Anti-HA!

The reviewer is quite right, the top panel is anti-FLAG and the bottom is anti-myc. The anti-HA label corresponds the small panel in Figure 4B. We see how this is confusing and have modified the figure to make it clearer.

2. The authors state line 523 that they use a cohesin variant 'containing cysteines specifically crosslinking the three intermolecular interfaces'. They need to explicitly state which interfaces are crosslinked in the variant they use for structure determination. The author response indicates that 'the 6-cysteine version used interrogates the hinge dimerisation and not the head dimerisation'. Their methods section indicates however that the variant they use for structure determination does not contain Cys mutations in the hinges. Instead, they use Smc1G22C N1192C, Smc3S1043C R1222C, Scc1C56A547C.

These mutations are located (1) in the Smc3-NScc1 interface (Scc1C56 and Smc3S1043C), (2) Smc1-cScc1 (Scc1A547C and Smc1G22C) interface and (3), in the interface between the Smc1 and Smc3 ATPase heads (Smc1 N1192C and Smc3 R1222C). That is, the third pair of cysteines is clearly placed in the heads and not the hinges.

We profoundly apologise for the confusion caused by our mistake and thank the reviewer for pointing this out again. The correct cysteines are Smc1K639C-Smc3E570C, Smc1G22C-Scc1A547C, and Smc3S1043C-Scc1C56. We have amended the manuscript accordingly in the respective places. Therefore, our original interpretation stands, i.e. the ATPase heads have not been crosslinked to each other.

3. It is therefore not clear why the authors maintain (Reviewer #2 points 6, 7, 8, 12) that 'we did not cross-link the ATP head dimerisation interface'. Clearly, something is wrong! Maybe it is the methods section? If not, the authors need to clearly indicate that the version of cohesin they use is crosslinked at the SMC1/3 ATPase head domains. The implications of such head crosslinking need to be taken into account in the interpretation of their data and discussion of the results.

See above.

Reviewer #3:

The manuscript is greatly improved. The updated figures make it much clearer. The rewritten section describing Sth1, nucleosome positioning, and histone mutants is good. This is important work, and the breadth and strength of the experiments described are impressive.

Most of my comments have been adequately addressed. The requests below relate to the presentation and description of the data and should be addressed before publication but do not require extensive writing or any new experiments.

Line 593-597:

"In the presence of ATP" implies that ATP occupies both ATPase active sites. Neither γ phosphate can be resolved at this resolution. The structure could be a reaction intermediate resulting from slow π dissociation from one or both sites, for example. This is not to detract from the observations or statements made. Instead, it would be wise to state more clearly that ATP was included in the sample but avoid any suggestion that the structure provides detailed information about the SMC ATPase cycle, which will doubtless be a topic for future studies. This is rightly stated in the final paragraph of the Discussion.

We assume that the reviewer refers to one or both of these instances: “We identified and solved … a form of cohesin at 6Å … whose ATPase heads were engaged in the presence of ATP” and/or “A cohesin complex whose heads are engaged in the presence of ATP…”

If so, we respectfully disagree with the reviewer. What is precisely stated in our manuscript is that our engaged cohesin heads, which we explicitly mention are at no more than 6 Å resolution, were in the presence of ATP. We never allude to the state of the molecule or even hint at the possibility of seeing the gamma phosphate at this resolution. Adding that the heads could also be bound to ADP and Pi or even be in a transition would just serve to confuse the reader in an, in our opinion, very straightforward and factual statement of the sample conditions.

Figure 5G needs to be modified. Figure 5S2 shows that 5G is a composite structure derived from two different samples. One contained ATP and the other did not. Overlaying them is inappropriate. Yes, the angle of the head-proximal coiled-coils is similar (Figure 5F vs. 5G), and it is likely that the remainder of the CC density beyond (hinge-proximal) the joint is similar/identical, but this is not proven by the data in the manuscript. In fact, it appears this section of the complex was excluded from analysis in the case of the sample containing nucleotide (Figure 5G and 5S2D). Without recalculating structures or even remaking complicated figures, it must be possible to present this data more clearly. One possibility: simply removing the "Arm" density from figure 5G. Another possibility is to compare the head domain density (including head-proximal cc) from Figure 5F, which is currently obscured by Scc2 density, with the head domain reconstruction from 5G.

We thank the reviewer for the suggestion and we agree that the figure should be more transparent. We have removed the separating discontinuous line that suggested that these were two structures from different datasets and have added that the folded coiled-coil was derived from a dataset without ATP to the figure. In addition, we have referred the reader explicitly to figure 5S2D to see how the map was obtained. However, we believe that the composite structure does add valuable information that aids the reader’s understanding by putting the engaged ATPase heads into the structural context of a folded cohesion and have therefore decided to leave it with the mentioned remarks. Especially given the 2D classes in Figure 5 Supplement 1E.

Figure 5 – supplement 1A does not conclusively show complexes lacking Scc2. Compare with 5S1A, for example. One needs to consider variability in Scc2 occupancy, Scc2 position, and also the fraction and quality of the particle images contributing to these classes. The description "in samples lacking all HAWK proteins" (line 537) therefore overstates the certainty of the claim given the evidence. The statement and figure panel could be removed without damaging the overall message. Removing panel A has the added benefit that it eliminates the need to discuss a dataset not referenced in figure 5S2 (cohesin, Scc2, ATP).

The reviewer is absolutely right – this was due to a mistake while reorganising the supplementary figures 5 during the previous revision. We have added 2D classes showing exactly that, i.e, a folded cohesion complex in the absence of HAWK proteins to the figure (cf. Figure 5S1E) and have amended the text that referred to the wrong figure.

https://doi.org/10.7554/eLife.67268.sa2

Article and author information

Author details

  1. Naomi J Petela

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Conceptualization, Data curation, Software, Formal analysis, Validation, Investigation, Writing - original draft, Writing - review and editing
    Contributed equally with
    Andres Gonzalez Llamazares
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9607-0422
  2. Andres Gonzalez Llamazares

    MRC Laboratory of Molecular Biology, Cambridge, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Validation, Investigation, Writing - original draft, Writing - review and editing
    Contributed equally with
    Naomi J Petela
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5404-6360
  3. Sarah Dixon

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  4. Bin Hu

    Institute of Medical Sciences, University of Aberdeen, Aberdeen, United Kingdom
    Contribution
    Funding acquisition, Investigation
    Competing interests
    No competing interests declared
  5. Byung-Gil Lee

    MRC Laboratory of Molecular Biology, Cambridge, United Kingdom
    Contribution
    Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9565-6114
  6. Jean Metson

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  7. Heekyo Seo

    Department of Biological Sciences, KAIST Institute for the Biocentury, Cancer Metastasis Control Center, Korea Advanced Institute of Science and Technology, Daejeon, Republic of Korea
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  8. Antonio Ferrer-Harding

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  9. Menelaos Voulgaris

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  10. Thomas Gligoris

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  11. James Collier

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9904-9423
  12. Byung-Ha Oh

    Department of Biological Sciences, KAIST Institute for the Biocentury, Cancer Metastasis Control Center, Korea Advanced Institute of Science and Technology, Daejeon, Republic of Korea
    Contribution
    Supervision, Funding acquisition, Investigation
    Competing interests
    No competing interests declared
  13. Jan Löwe

    MRC Laboratory of Molecular Biology, Cambridge, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    jyl@mrc-lmb.cam.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5218-6615
  14. Kim A Nasmyth

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Validation, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    ashley.nasmyth@bioch.ox.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7030-4403

Funding

Wellcome Trust (107935/Z/15/Z)

  • Kim A Nasmyth

Wellcome Trust (202754/Z/16/Z)

  • Jan Löwe

Wellcome Trust (202062/Z/16/Z)

  • Bin Hu

Cancer Research UK (26747)

  • Kim A Nasmyth

H2020 European Research Council (294401)

  • Kim A Nasmyth

Medical Research Council (U105184326)

  • Jan Löwe

Biotechnology and Biological Sciences Research Council (BB/S002537/1)

  • Bin Hu

National Research Foundation of Korea (NRF2020R1A4A3079755)

  • Byung-Ha Oh

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We are grateful to Frank Uhlmann for sharing yeast strains, Katsu Shirahige for the anti-AcSmc3 antibody, and to Maria Demidova, Wentao Chen, and Christophe Chapard for invaluable technical assistance. We would like to thank all the members of the Nasmyth and Löwe groups for valuable discussions. This work was funded by the Wellcome Trust (107935/Z/15/Z to KN; 202754/Z/16/Z to JL; 202062/Z/16/Z to BH), Cancer Research UK (26747 to K N), the European Research Council (294401 to KN), the Medical Research Council (U105184326 to JL), the Biotechnology and Biological Sciences Research Council (BB/S002537/1 to B H), and the National Research Foundation of Korea (B-HO, NRF2020R1A4A3079755).

Senior Editor

  1. Cynthia Wolberger, Johns Hopkins University School of Medicine, United States

Reviewing Editor

  1. Adèle L Marston, University of Edinburgh, United Kingdom

Reviewers

  1. Adèle L Marston, University of Edinburgh, United Kingdom
  2. Daniel Panne, University of Leicester, United Kingdom

Publication history

  1. Received: February 5, 2021
  2. Accepted: June 21, 2021
  3. Version of Record published: July 14, 2021 (version 1)

Copyright

© 2021, Petela et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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