1. Chromosomes and Gene Expression
  2. Structural Biology and Molecular Biophysics
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Structural basis for Scc3-dependent cohesin recruitment to chromatin

  1. Yan Li
  2. Kyle W Muir  Is a corresponding author
  3. Matthew W Bowler
  4. Jutta Metz
  5. Christian H Haering
  6. Daniel Panne  Is a corresponding author
  1. European Molecular Biology Laboratory, France
  2. European Molecular Biology Laboratory, Germany
  3. University of Leicester, United Kingdom
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Cite as: eLife 2018;7:e38356 doi: 10.7554/eLife.38356

Abstract

The cohesin ring complex is required for numerous chromosomal transactions including sister chromatid cohesion, DNA damage repair and transcriptional regulation. How cohesin engages its chromatin substrate has remained an unresolved question. We show here, by determining a crystal structure of the budding yeast cohesin HEAT-repeat subunit Scc3 bound to a fragment of the Scc1 kleisin subunit and DNA, that Scc3 and Scc1 form a composite DNA interaction module. The Scc3-Scc1 subcomplex engages double-stranded DNA through a conserved, positively charged surface. We demonstrate that this conserved domain is required for DNA binding by Scc3-Scc1 in vitro, as well as for the enrichment of cohesin on chromosomes and for cell viability. These findings suggest that the Scc3-Scc1 DNA-binding interface plays a central role in the recruitment of cohesin complexes to chromosomes and therefore for cohesin to faithfully execute its functions during cell division.

https://doi.org/10.7554/eLife.38356.001

Introduction

To ensure that each daughter cell receives an equal complement of genetic information, cognate chromatids are paired through replication-coupled sister chromatid cohesion. Cohesion is then actively maintained and eventually enables attachment of kinetochores to the mitotic spindle microtubules emanating from opposite poles to ensure chromosome bi-orientation, prior to subsequent segregation of sister chromatids into daughter cells (Nasmyth and Haering, 2009; Peters and Nishiyama, 2012).

Cohesion is facilitated by cohesin, a member of the Structural Maintenance of Chromosomes (SMC) family of protein complexes, which is responsible for genome organisation across all domains of life (Palecek and Gruber, 2015; Wells et al., 2017). Cohesin complexes form tripartite rings, comprising Smc1-Smc3 and the kleisin subunit Scc1, that are proposed to topologically entrap sister DNA molecules (Gligoris et al., 2014; Nasmyth and Haering, 2009).

The chromosomal addresses of cohesin loading are determined by the Scc2-Scc4 complex, which is enriched at centromeres via direct contacts with kinetochore proteins and promotes DNA-stimulated ATP hydrolysis by the Smc1-Smc3 ATPase heads to drive chromatin entrapment (Ciosk et al., 2000; Hinshaw et al., 2017; Murayama and Uhlmann, 2014). Conversely, dynamic release of DNA from the ring is achieved either by the proteolytic cleavage of the Scc1 kleisin subunit by separase protease (Uhlmann et al., 2000), or by the opening of an ‘exit gate’ formed at the Scc1 and Smc3 interface. Release of the latter is inhibited by Smc3 acetylation and is controlled by the accessory factors Scc3, Wapl and Pds5 (Beckouët et al., 2016; Rolef Ben-Shahar et al., 2008; Rowland et al., 2009; Unal et al., 2008).

Cohesin can associate dynamically with chromatin through alternating cycles of DNA entrapment and release until ring opening is inhibited by acetylation of the Smc3 ATPase head. At this point, DNA is thought to remain topologically entrapped within the SMC-kleisin ring (Gligoris et al., 2014Beckouët et al., 2016). A prevailing model of genome organisation posits that chromatin loops entrapped within the lumen of the SMC ring are then processively enlarged by extrusion, which presumably forms the basis for higher-order chromatin structure (Alipour and Marko, 2012).

Whereas the essential role of cohesin in modulating genome architecture, gene expression and DNA damage repair is increasingly well established, comparatively little information is available concerning the molecular determinants of its association with chromatin (Busslinger et al., 2017; Haarhuis et al., 2017; Schwarzer et al., 2017; Wu and Yu, 2012). Considerable advances have been made in dissecting the compositional and biochemical prerequisites for topological entrapment of DNA by the cohesin complex (Davidson et al., 2016; Kanke et al., 2016; Stigler et al., 2016; Murayama and Uhlmann, 2014; 2015). However, how cohesin engages its DNA substrate and how such interactions might be regulated is yet to be fully elucidated. Recent work identified a direct DNA-binding site in the paralogous condensin complex, in which the HEAT repeat subunit Ycg1, in complex with the kleisin subunit Brn1, contacts the DNA double helix backbone and stabilizes its association through DNA entrapment within the Brn1 peptide loop (Kschonsak et al., 2017).

To investigate whether cohesin can similarly establish direct contacts with DNA, we produced a series of structurally well-defined globular cohesin subcomplexes and systematically interrogated their ability to bind to DNA. We found that the essential HEAT-repeat subunit Scc3, when in complex with Scc1, binds double-stranded DNA (dsDNA). We further determined the crystal structure of the yeast Scc3-Scc1 complex bound to DNA, which revealed that DNA binding is mediated by a composite surface comprising positively charged amino acid residues from Scc3 and Scc1. Charge-reversal mutagenesis of this interface demonstrates that DNA substrate engagement by the Scc3-Scc1 complex is essential for cohesin to bind to DNA in vitro or chromosomes in vivo and is therefore indispensable for cell viability. Thus, direct DNA substrate engagement through the newly discovered DNA-binding interface of the Scc3-Scc1 subcomplex is a key element for the cohesin cycle.

Results

To investigate the DNA-binding properties of cohesin, we co-expressed and purified defined globular domains and subcomplexes of the cohesin complex from the budding yeast Saccharomyces cerevisiae (Figure 1A, Figure 1—figure supplement 1, Figure 1—figure supplement 2). These encompassed the Smc3 ATPase head domain bound to an N-terminal fragment of Scc1 (Smc3hd-NScc1), the Smc1 ATPase head domain bound to a C-terminal fragment of Scc1 (Smc1hd-CScc1), as well as an Scc3-Scc1 subcomplex (Scc3T-Scc1K) (Figure 1B). In addition, we produced an Smc1-Smc3 hinge heterodimer, Pds5 bound to a Scc1 fragment (Muir et al., 2016) as a full-length (Pds5fl) or truncated variant (Pds5T), as well as Wapl as full-length (Waplfl) or truncated variants (WaplC; Figure 1—figure supplement 1). Consistent with prior studies (Murayama and Uhlmann, 2014), we found that the Scc3T-Scc1K subcomplex and the Smc1-Smc3 hinge heterodimer bound DNA, as seen by the appearance of slower-migrating species in electrophoretic mobility shift assays (EMSAs), as did the Smc3hd-NScc1 module, which has previously been implicated as a DNA sensor in cohesin (Murayama and Uhlmann, 2014) (Figure 1C, Figure 1—figure supplement 2). As expected for non-sequence specific DNA-binding factors, longer DNA fragments (>21 base pairs (bp)) bound more efficiently than shorter DNA duplexes (15 bp) (Figure 1C, Figure 1—figure supplement 2). The ability of the Scc3T-Scc1K subcomplex to bind DNA depended on the presence of Scc1K. Conversely, the other HEAT-repeat-kleisin subcomplex of cohesin, Pds5-Scc1, the Smc1hd-CScc1 subcomplex and the Wapl subunit did not interact with DNA in this assay. Thus, as in the paralogous condensin complex, the HEAT-repeat protein bound to the C-terminal region of the kleisin subunit directly engages DNA (Kschonsak et al., 2017).

Figure 1 with 2 supplements see all
DNA binding by the Scc3-Scc1 subcomplex.

(A) Cartoon of the cohesin complex. (B) Domain structure of Scc3 and Scc1. Construct boundaries used and their acronyms are shown below. (C) SDS-PAGE analysis of purified Scc3T-Scc1K and DNA binding analysis by EMSA. Scc3 binds to longer DNA more efficiently compared to shorter DNA. The DNA binding capacity of Scc3T is enhanced by Scc1K.

https://doi.org/10.7554/eLife.38356.002

To identify the molecular basis of this interaction, we crystallized the Scc3T-Scc1K complex from budding yeast bound to 19 bp of dsDNA. Optimized crystals diffracted anisotropically to a minimum Bragg spacing of 3.6 Å in the best, and ~5.7 Å in the worst direction (Table 1). We determined the structure by molecular replacement, using the structures of an Scc3 ortholog from Zygosaccharomyces rouxii and a C-terminal fragment of Saccharomyces cerevisiae Scc3 as search models (Roig et al., 2014). The resulting electron-density map provided a continuous trace of the polypeptide main chain, but with a limited level of detail owing to the anisotropy of the data. Despite these drawbacks, we successfully traced the structure using a selenomethionine derivative and refined a model encompassing amino acid residues 134 to 1064 of Scc3 in complex with residues 309 to 400 of Scc1, bound to a 19 bp DNA molecule (Table 1).

Table 1
Data collection and refinement statistics.
https://doi.org/10.7554/eLife.38356.005
Scc3T/Scc1K nativeScc3T/Scc1K
SeMet
Data collection
Space groupP21212P21212
Cell dimensions
 a, b, c (Å)109.9, 115.4, 295.6109.9, 115.6, 296.2
Wavelength (Å)1.2821.282
Resolution (Å)50–3.6049.9–4.79
No. reflections2096310279
Rmerge5.8 (122.6)*4.6 (112.3)*
I / σI11.9 (1.6)*10.6 2(.1)
CC 1/20.99 (0.56)0.99 (0.52)
Completeness (%)91.4 (63.5)*93.6 (71.2)*
Redundancy4.4 (6.0)*1.8 (1.8)
Refinement
Resolution (Å)50–3.60
Rwork/Rfree0.28/0.31
No. atoms16465
 Protein14909
 DNA1556
B-factors (mean)
 Protein254.5
 DNA266.4
R.m.s deviations
 Bond lengths (Å)0.002
 Bond angles (°)0.5
  1. *Values in parentheses are for highest-resolution shell.

As seen in other structures of Scc3, the protein is hook-shaped in the C-terminal section and contains an N-terminal ‘nose’ formed by a pair of extended antiparallel α-helices (Figure 2A). Similarly to the interaction of human Scc1 with Scc3, the yeast Scc1 in our structure binds along the convex surface of the hook-shaped HEAT-repeat subunit. We detected additional electron density corresponding to dsDNA within the cradle of this hook (Figure 2—figure supplement 1A,B). Whereas the DNA duplexes aligned to form pseudocontinuous double helices throughout the crystal, the DNA duplex was slightly too short for tight end-to-end stacking (Figure 2—figure supplement 1A,B). As a result, the DNA density was only partially resolved, apparently due to rotational and translational disorder of the DNA in the binding cavity.

Figure 2 with 1 supplement see all
Structure of the Scc3-Scc1 subcomplex bound to DNA.

(A) Cartoon representation of the Scc3-Scc1 complex bound to a 19 bp dsDNA substrate. The N- and C- termini of Scc3 (violet) and Scc1 subunits (green) are shown. The inset shows a close-up view of the Scc1 amino acid K363. (B) Electrostatic surface potential representation of the Scc3-Scc1 subcomplex with bound dsDNA (calculated with APBS and displayed with Pymol).

https://doi.org/10.7554/eLife.38356.006

To identify amino acid residues potentially involved in DNA binding, we mapped the electrostatic surface potential onto the Scc3-Scc1 structure (Figure 2B). This revealed that DNA is nested within an extended cradle spanning the majority of Scc3-Scc1, lined by a set of positively charged residues that directly contact the DNA phosphate backbone. The DNA is aligned almost parallel to the N-terminal ‘nose’ of Scc3, which interacts through a set of positively charged amino acid residues with the DNA of a neighbouring complex related by crystallographic symmetry (Figure 2—figure supplement 1C). We observed no direct nucleotide base–amino acid interactions, which explains the apparent lack of DNA sequence specificity.

To ascertain the amino-acid register of Scc1, we used an anomalous difference map peak for M373 in the selenomethionine-derivative data (Figure 2—figure supplement 1D). The deduced register places Scc1 residue K363 in close proximity to the DNA (Figure 2A, Figure 2—figure supplement 1D). Direct interactions between Scc1 and the DNA phosphate backbone potentially explain why the Scc3T-Scc1K subcomplex has greater DNA binding affinity than does isolated Scc3T (Figure 1C).

Mapping of sequence conservation onto the structure revealed that amino acid residues in the DNA binding groove are generally well conserved among yeast Scc3 orthologs (Figure 3A). In particular, amino acid residues that are located proximal to the DNA phosphate backbone showed strong conservation. To further evaluate the contributions made by individual segments of the DNA-binding surface, we subdivided participating residues into a series of three patches, based on their physical proximity to the DNA (Figure 3B), and subjected these patches to site-directed mutagenesis.

Figure 3 with 2 supplements see all
A conserved DNA binding domain in the Scc3-Scc1 subcomplex is required for cohesin association with chromatin.

(A) Surface amino acid conservation of yeast Scc3. Residues in the DNA binding domain are well conserved. (B) DNA binding residues are located in three surface patches of Scc3. (C) DNA binding fluorescence polarization of 6-FAM labelled 32 bp dsDNA by variants of the Scc3-Scc1 subcomplex. Data points corresponding to the average of three independent experiments were fitted to a standard binding equation assuming a single binding site using Kaleidagraph. Standard deviations are depicted as vertical error bars. Apparent dissociation constants (KD) are noted below. (D) Tetrad analysis of diploid budding yeast strains expressing ectopic wild-type or mutant versions of Scc3 under control of the endogenous promoter in an SCC3/scc3Δ background (strains C5013, C5014, C5015, C5043, C5033). Images were recorded after three days at 30°C on rich media. Genetic marker analysis identified Scc3(mutant), scc3Δ cells (circles). (E) ChIP-qPCR analysis of binding to centromeric (cen), pericentromeric (pericen) or chromosome arm (arm) regions (chromosomes IV, V, and VI as indicated) of untagged (strain C3) or PK6-tagged wild-type or mutant versions of Scc3 expressed from an ectopic locus under its endogenous promoter (strains C5013, C5043, C5033). The fractions of immunoprecipitated DNA relative to input DNA are plotted as circles for two biological repeats with two technical repeats each (same colour pairs). Mean values of all four data points are shown as lines.

https://doi.org/10.7554/eLife.38356.008

We measured the DNA equilibrium dissociation constant by fluorescence polarization, using a 32-bp 6-FAM labelled DNA substrate, which is sufficient to bridge the entire DNA binding surface present in the Scc3-Scc1 crystal structure (Figure 3C). Whereas the wild-type Scc3T-Scc1K complex bound this substrate with an equilibrium dissociation constant of 2.2 μM, charge-inversion mutations in patches 1 and 3, located at the crest of and within the Scc3 cradle, had only modest effects on affinity (equilibrium dissociation constants of 9.5 μM and 7.3 μM, respectively). Patch 2 mutants, residing in the Scc3 'nose', exhibited essentially unaltered DNA binding affinity. In contrast, the simultaneous mutation of all three patches (a heptamutant) reduced the binding affinity of the patch 1 or 3 mutants even further (to 29.3 μM, Figure 3C). The defect in binding DNA was not due to any impact of the mutations on the structural integrity of Scc3-Scc1, as all mutant complexes eluted indistinguishably from wild-type Scc3-Scc1 during size-exclusion chromatography (Figure 3—figure supplement 1A) and efficiently formed a complex with Scc1 in vitro (Figure 3—figure supplement 1B) or in vivo (see below). We conclude that the positively charged Scc3 cradle comprises the major DNA-binding surface of Scc3 and that the positively charged amino acid residues located in patch 1 and 3 constitute a composite DNA binding surface. The distribution of these residues across an extended surface of Scc3 might explain why their significance has thus far eluded cell-biological and genetic characterization. To test whether the direct interactions between Scc1 and the DNA phosphate backbone contribute to DNA binding we introduced the double mutation K363E/R364E into Scc1K. This mutant showed an essentially indistinguishable equilibrium dissociation constant as compared to that of the wild-type complex, thus indicating that this positively charged patch of Scc1 does not contribute to DNA binding (Figure 3C).

We then assayed whether DNA binding by Scc3 is important for cohesin function in vivo. We integrated wild-type or mutant versions of Scc3 into a diploid yeast strain in which one of the two endogenous SCC3 genes had been deleted (Supplementary file 2) and analysed the competence of these Scc3 variants to complement the SCC3 deletion by tetrad dissection (Figure 3D). Whereas all three individual patch mutants could support cell proliferation, cells expressing the heptamutant mutant version as their only source of Scc3 failed to divide, despite expressing the mutant Scc3 protein at wild-type levels (Figure 3—figure supplement 2A).

To test whether the inability of the heptamutant version of Scc3 to support cohesin function was the result of a defect in the association of the mutant cohesin complex with chromosomes, we measured the levels of wild-type and mutant cohesin complexes at five independent binding sites in the budding yeast genome by chromatin immunoprecipitation followed by quantitative PCR (ChIP-qPCR). The amounts of chromosomal DNA that co-immunoprecipitated with the Scc3 patch 3 mutant were on average 40% lower than those that co-immunoprecipitated with wild-type Scc3. The Scc3 heptamutant failed to bind chromatin entirely (Figure 3E), although it was incorporated normally into cohesin complexes (Figure 3—figure supplement 2B). We conclude that DNA binding by Scc3-Scc1 is essential for the stable association of cohesin complexes with chromosomes in vivo and hence an important determinant of cohesin function.

Discussion

Targeting of cohesin to the genome is essential for numerous aspects of chromosome biology, including sister chromatid cohesion, DNA damage repair, and transcriptional regulation (Uhlmann, 2016). In this study, we identified a direct DNA-binding site in the Scc3 subunit of the cohesin complex and determined its interaction with DNA at near-atomic resolution. These findings provide evidence for a site of direct DNA contact in cohesin complexes, which presumably contributes to the initial step of chromosome entrapment and/or DNA translocation (Figure 4C).

A conserved DNA binding interface in cohesin and condensin.

(A) Structure of the DNA-bound Ycg1–Brn1 subcomplex from condensin. (B) Structure of the Scc3-Scc1 complex. In the condensin structure, a peptide loop (green dashed line) of the Brn1 kleisin subunit encircles the bound DNA and prevents its dissociation. Alignments were generated by secondary structure matching using Cα atoms from the Scc3 HEAT-repeats and the structurally equivalent region of the condensin Ycg1 HEAT-repeat subunit. (C) Model for Scc3-mediated DNA binding by cohesin complexes. Scc3-Scc1 enables direct chromatin binding. Cohesin is loaded by Scc2-Scc4 in an ATP dependent fashion thus resulting in topological DNA entrapment.

https://doi.org/10.7554/eLife.38356.011

We show here that cohesin interacts directly with DNA through a complex formed by the HEAT-repeat subunit Scc3 and the kleisin subunit Scc1. We have determined the DNA-bound structure of Scc3 in complex with a minimal fragment of Scc1 and demonstrated that DNA binding depends on conserved positively charged residues of a composite Scc3-Scc1 interface. We have used this structure to derive DNA-binding deficient Scc3 variants, which fail to support cohesin recruitment to chromatin and consequently cell division. In addition to providing a scaffold for the assembly of cohesin regulators (Hara et al., 2014; Murayama and Uhlmann, 2014; Roig et al., 2014) and thereby participating in the maintenance of cohesion, the Scc3-Scc1 subcomplex also plays a key role in DNA substrate recognition and hence the efficient association of the cohesin holocomplex with chromatin.

The cohesin ring has been proposed to topologically embrace chromosomal DNA, which requires that DNA is loaded into and also released from the ring complex during cohesin’s reaction cycle (Nasmyth, 2011). As mutations in Scc3 that prevent DNA engagement by the Scc3-Scc1 subcomplex in vitro fail to stably bind to chromatin in vivo, it is likely that direct DNA-cohesin interactions contribute to such DNA entry and exit reactions (Figure 4C).

We propose that the Scc3-Scc1 subcomplex provides a dynamic DNA anchoring point that is required for the efficient loading and/or maintenance of cohesin on chromatin. Such a model for Scc3 is supported by previous data, which indicate that Scc3 contributes to cohesin loading (Hu et al., 2011; Murayama and Uhlmann, 2014; Orgil et al., 2015; Roig et al., 2014). In agreement with this model, Scc1 deletion mutants that lack the sequence responsible for binding Scc3 fail to load onto yeast chromosomes (Hu et al., 2011). Furthermore, Scc3 enhances topological DNA entrapment by cohesin in in vitro loading assays (Murayama and Uhlmann, 2014). Thus, DNA binding by the Scc3-Scc1 subcomplex might be the first step in moving chromosomal DNA into the cohesin ring. The subsequent entrapment reaction is then presumably catalysed by the Scc2-Scc4 complex (Davidson et al., 2016; Murayama and Uhlmann, 2014; Stigler et al., 2016).

Surface patch 3 mutations in Scc3, which partially ablate DNA binding by the Scc3-Scc1 subcomplex and reduce cohesin levels on chromatin, do not exhibit any obvious growth defects and are therefore presumably competent to establish sister chromatid cohesion. Indeed, partial depletion of cohesin does not seem to impact some of its core functions, including sister-chromatid cohesion and chromosome segregation (Elbatsh et al., 2016; Heidinger-Pauli et al., 2010). In contrast, even mutations that only slightly impair DNA binding of the equivalent HEAT-repeat/kleisin module of the condensin complex are sufficient to abolish stable chromatin association and to interfere with cell division (Kschonsak et al., 2017). Such discrepancies might be due to alternate loading and/or maintenance mechanisms of cohesin and condensin. Whilst cohesin is loaded by the Scc2-Scc4 complex, no such independent loading factor has been identified for condensin thus far, which could explain why the latter depends more strongly on the direct DNA binding site formed by its HEAT-repeat and kleisin subunits.

Binding of condensin to DNA is further stabilized by the entrapment of the bound DNA helix within a kleisin peptide loop (Figure 4A-) (Kschonsak et al., 2017). The relevant section of Scc1 that would contribute to such topological DNA entrapment is disordered in our structure(Figure 4B). As Scc1 is clearly required for DNA binding of the Scc3-Scc1 subcomplex, but apparently not through direct DNA interactions (Figure 3C), it is possible that cohesin uses a similar mode of chromatin engagement. These findings thus point towards a conserved molecular mechanism that enables chromatin substrate engagement by condensin and cohesin. This mechanism potentially facilitates topological loading, chromatin looping and tracking along chromatin fibres by these SMC complexes (Ganji et al., 2018).

Materials and methods

Constructs, expression and purification

Scc3, Scc1 and other cohesin subunits were amplified from yeast Genomic DNA (Millipore). Scc3 and mutant variants thereof were cloned into pETM-30 vector using NcoI and NotI restriction enzyme cleavage sites. Scc1 constructs were cloned using the NcoI and NotI sites of a pACYC-DUET vector for co-expression with Scc3, or into vectors containing the cognate Smc head domain in their second ORF for Smc/kleisin complexes (see below). Codon optimised genes comprising the Smc1 and Smc3 ATPase domains were produced by gene synthesis (Thermofisher), to include C-terminal 6xHistidine tags, and ligated into the NdeI-XhoI sites of pRsf-DUET1 and pACYC-DUET1 respectively via Gibson Assembly (NEB). Pds5 and Wapl were cloned and expressed as described previously (Muir et al., 2016). For the expression and purification of Pds5, media and buffers were supplemented with 20 μM and 5 μM of inositol hexa-kis-phosphate, respectively (Ouyang et al., 2016).

Proteins were expressed in Escherichia coli BL21(DE3) by auto-induction (Studier, 2005). Cells were grown at 37°C until OD600nm = 0.6 and then shifted to 18°C for 16 hr. Cells were harvested and washed once with ice-cold PBS buffer. Pellets were resuspended in buffer 1 (40 mM TRIS, pH 7.5, 500 mM NaCl, 20 mM imidazole, 0.5 mM TCEP) containing one tablet of complete, Mini, EDTA-free protease inhibitors (Roche). Cells were lysed using a microfluidiser (Microfluidics) and centrifuged at 15000 rpm for 1 hr using a JA-20 rotor (Beckman). The supernatant was loaded onto 5 ml Co2+–conjugated IMAC beads (GE healthcare) by using a peristaltic pump (GILSON). The column was washed with 10 column volumes of buffer 1 and the protein eluted with buffer 2 (40 mM TRIS, pH 7.5, 300 mM NaCl, 300 mM imidazole, 0.5 mM TCEP). The His-GST tag was cleaved by addition of His-tagged TEV protease during overnight dialysis against 40 mM TRIS, pH 7.5, 300 mM NaCl, 0.5 mM TCEP at 4°C. For Smc3-NScc1, Smc1-CScc1, and the Smc3-Smc1 hinge, this cleavage step was bypassed.

The HIS-GST tag, protease and uncleaved protein were removed by passing this mixture over Co2+ IMAC resin. The flow through was concentrated using an Amicon Ultra −15 concentrator (Millipore) and applied onto a MonoQ 5/50 GL column (GE healthcare) in buffer 3 (150 mM NaCl, 40 mM TRIS, pH7.5, 0.5 mM TCEP). Proteins were eluted using a linear gradient in buffer 4 (1 M NaCl, 40 mM TRIS, pH 7.5, 0.5 mM TCEP). The final purification step was performed by using a HiLoad 16/60 Superdex 200 prep–grade column in buffer 5 (150 mM NaCl, 20 mM TRIS, pH7.5, 0.5 mM TCEP).

DNA binding assays

For analysis of DNA-binding by EMSA and for co-crystallization, DNA substrates were generated by annealing complementary oligonucleotides (MWG Eurofins) at a final concentration of 1 mM in 20 mM TRIS, pH 7.5, 150 mM NaCl (Supplementary file 1). Successful annealing and purity of the oligonucleotides were confirmed by native PAGE on a 6% gel.

For EMSA experiments, varying concentrations of protein samples were incubated at the indicated molar ratios with 1 μM the 32mer DNA in 150 mM NaCl, 20 mM TRIS, pH 7.5, 0.5 mM TCEP. Samples were incubated on ice for 30 min. 5% Glycerol was added and the samples were analysed on a 6% native 1x TRIS-Glycine (25 mM TRIS, 250 mM glycine, pH 8.3, 5% Glycerol) polyacrylamide gel using 1x TRIS-Glycine running buffer. Gels were stained with SYBR Safe (Thermo Fisher Scientific) to visualize DNA-bound complexes or Coomassie Blue for protein staining.

Fluorescence Polarization (FP)

32 bp 6-FAM labelled DNA was prepared by annealing two complementary DNA strands, essentially as described for the crystallisation oligonucleotides, albeit under low-light conditions (Supplementary file 1). Fluoresence polarisation assays were conducted in a buffer containing 50 mM TRIS pH 7.5, 150 mM NaCl, 0.1% tween and 0.5 mM TCEP. A series of protein concentrations, ranging from 0.5 μM to 25 μM, were incubated in the presence of 50 nM DNA for 30 min at room temperature in order to attain equilibrium. Immediately thereafter, fluorescence polarization was recorded using 485 nm and 520 nm excitation and emission filters, respectively (CLARIOstar, BMG Labtech, Germany). The change in fluorescence polarization was then plotted as mean values of three independent replicates and the dissociation constant for each complex determined.

Crystallization and data collection

Crystals of the Scc3-Scc1-DNA complex were obtained by mixing 8 mg ml−1 protein with DNA at a 1:1.1 ratio. 1 μl of the protein:DNA complex were mixed with 1 μl 10% PEG 8000, 0.1M Bis-TRIS, pH 6.5 crystallization buffer and equilibrated against the crystallization buffer at 4°C. Initial crystals with a 17 mer DNA were obtained after 5 days. These crystals were used as seeds for crystallization of Scc3-Scc1 bound to a 19mer DNA using the same crystallization condition. Crystals were cryo protected by adding 20% Glycerol to the crystallization buffer and flash frozen in liquid nitrogen. Diffraction data for native and selenomethionine-derivatised Scc3-Scc1-DNA crystals were collected at 100 K at an X-ray wavelength of 0.966 Å at beamline ID30A-1/MASSIF-1 (Bowler et al., 2015) of the European Synchrotron Radiation Facility, with a Pilatus3 2M detector using automatic protocols for the location and optimal centring of crystals (Svensson et al., 2015). The beam diameter was selected automatically to match the crystal volume of highest homogeneous quality (Svensson et al., 2018). Data were processed with XDS (Kabsch, 2010) and imported into CCP4 format using AIMLESS (Winn et al., 2011).

Structure determination refinement and analysis

The structure of the Scc3-Scc1-DNA complex was determined by molecular replacement by Phaser (McCoy et al., 2007) using a Scc3 structure from Zygosaccharomyces rouxii (PDB code 4UVK) and a structure of a C-terminal fragment of Scc3 from Saccharomyces cerevisiae (PDB code 4UVL) (Roig et al., 2014) in spacegroup P21212 at a resolution of 3.6 Å (Table 1). The model was build by iterative rounds of manual adjustments with Coot and of restrained refinements with Phenix (Afonine et al., 2012; Emsley et al., 2010). Sequence register was confirmed using a selenomethionine-derivatized crystal. Analysis of the refined structure in MolProbity showed that there were no residues in the disallowed and 94% in the favoured region of the Ramachandran plot. The MolProbity all atom clash score was 4.1 (Chen et al., 2010). Structures were visualized with PyMOL (Schrödinger, LLC). Surface conservation graphics were created using the ConSurf server (Ashkenazy et al., 2016) using a multi-sequence alignment containing Scc3 orthologs from Saccharomyces_cerevisiae (P40541), Saccharomyces_kudriavzevii (J5PHP0), Candida glabrata (A0A0W0DN34), Naumovozyma castellii (G0VFI2), Vanderwaltozyma polyspora (A7TNN6), Tetrapisispora phaffii (G8BRB9), Saccharomyces kudriavzevii (J5PHP0), Zygosaccharomyces rouxii (A0A1Q3A0R6), Torulaspora delbrueckii (G8ZZR1), Kluyveromyces lactis (Q6CIC3), Lachancea quebecensis (A0A0P1KU08), Eremothecium cymbalariae (G8JTR2), Ashbya gossypii (Q75AL6), Pichia sorbitophila (G8YM42) and Yarrowia lipolytica(Q6C144). The electrostatic surface potential graph was created with APBS (Baker et al., 2001).

Chromatin immunoprecipitation and ChIP-qPCR

Chromatin immunoprecipitation followed by quantitative PCR ChIP-qPCR was performed from asynchronous yeast cell cultures as described (Cuylen et al., 2011), with the exception that sonication was performed with a Bioruptor Plus (Diagenode) at 4°C using 6 cycles of 30 s on, 60 s off at ‘high’ level. Quantitative PCR was performed with primers listed Supplementary file 3.

Tetrad analysis

Wild-type or mutant alleles of Scc3 fused to a C-terminal PK6 epitope tag were integrated into the ura3 locus of a SCC3/scc3::HIS3 heterozygous diploid yeast strain (C1073). Correct integration was confirmed by PCR (Supplementary file 2). Following sporulation, strains were tetrad dissected and cultured on YPAD media for 3 days at 30°C before genotyping by replica plating.

Data availability

Coordinates for the Scc3-Scc1-DNA complex are available from the Protein Data Bank under accession number 6H8Q.

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Decision letter

  1. Andrea Musacchio
    Senior and Reviewing Editor; Max Planck Institute of Molecular Physiology, Germany

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Structural basis for Scc3-dependent cohesin recruitment to chromatin" for consideration by eLife. Your article has been reviewed by Andrea Musacchio as the Senior Editor, a Reviewing Editor, and three reviewers.. The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission. We hope you will be able to submit the revised version within two months.

Summary:

Li et al., describe a crystal structure of Scc3 (slightly truncated at both ends and designated Scc3T) with a fragment of Scc1 ("Scc1K") bound with a 19bp fragment of dsDNA. The structure, which closely resembles a corresponding condensin DNA complex published last year by Haering's group, shows that DNA binds along a groove created by the curvature of the Scc3 heat repeats. Mutations of residues at points of contact with DNA backbone diminish affinity in vitro and (in appropriate combination) prevent loading in vivo, as shown by ChIP-qPCR and by tetrad analysis of strains ectopically expressing wt or mutant versions of Scc3 in an SCC3/scc3Δ background. The biochemical and structural work clearly identify the molecular basis of an interaction between Scc3 and DNA and demonstrate that a fragment of Scc1 enhances this interaction. While not entirely unsurprising, the results add to our understanding of how cohesin might function mechanistically. With appropriate revision, the MS can be made suitable for publication in eLife.

Essential revisions:

1) In the Discussion section (first paragraph), the authors set up a straw man. The only alternative to direct protein-DNA contacts (either by a cohesin subunit or by some adaptor protein) is that cohesin is snapping open and closed all the time and should it happen to entrap DNA, it somehow sticks. That silly alternative is obviously hugely improbable. Thus, the real conclusion is that "these findings provide evidence for a site of DNA contact, probably during the initial step in chromosome entrapment" – or something like that. See also point 3, below. Incidentally, "topological principles" don't "drive" anything in biology – they explain various mechanisms or activities and suggest why they may have evolved, but biology is never "driven" by "principles" other than natural selection (except in the minds of the unreconstructed Cartesians still running around in France).

2) The anisotropy of the crystals, presumably due to a less than optimal DNA length, limits the accuracy and the information content of the structure. Did the authors, seeing the result, take the obvious next step of trying to get better crystals with other DNA lengths (e.g., 20 or 21 bp)? (That is, were all DNAs in Supplementary file 2 used in crystallization trials, or just for FP measurements? If the former, why not also 20 bp?) In this reviewer's view, that would have been an easier and better path than the one they took by validating the Scc1 trace with SeMet. In any case, Figure 3—figure supplement 1 should show either the DNA density in the initial MR map or (if the MR is good enough – it might not be) an Fo-Fc map after that initial step (i.e., phases from MR but 2Fo-Fc, showing in principle what's missing). In any case, a map with the final 2Fo-Fc phases, which included this DNA contributions, is not helpful. Incidentally, in Supplementary file 1, the Rmerge in the last bin is truly miserable. Was there an "elliptical" (i.e., anisotropic) cutoff, or did the meaningless reflections in the "bad" directions contribute? If the latter, then please recalculate with the correct anisotropic cutoff for each frame or set of frames, so that pure noise doesn't contribute to the data used. Also, "3.99" is 4.0 in my book, not "3.9".

3) The text overstates some of the biological and mechanistic conclusions. Although the correlation of affinity in vitro with function in vivo does permit the inference that the observed interaction is part of the DNA docking mechanism, the results do not rule out the participation of other contacts. Indeed, were those contacts strengthened by compensating mutations, it is possible that this contact would not be "indispensible", as the Abstract states.

4) The most useful conclusion is the similarity with condensin. For understandable "psychological" reasons, the authors do not mention the Kschonsak et al. (2017) paper in the Introduction. They should do so, as it surely guided their strategy at some point, either consciously or otherwise. Is the peptide loop definitely absent here, or could its absence be a consequence of truncating Scc1?

5) At the end of the Discussion section, the authors write that the mechanism they describe would enable cohesin to entrap a second DNA helix without releasing the first, etc. Not obvious to this reader why or how, perhaps because Figure 4C is so vague and incomprehensible.

6a) Is K363 on Scc1 important for the DNA binding ability of the complex? It would also be useful to highlight this residue on Figure 2. This experiment is required to confirm the in vivo relevance of the enhancement of DNA association by the Scc1 fragment in the in vitro experiments.

6b) Scc1 is clearly required for the DNA-binding activity of Scc3-Scc1. The structure suggests that Scc1 K363 might contact DNA. Does Scc1 K363E reduce the binding of Scc3-Scc1? Related to this, even though the authors cannot see any density of the N-terminal region of Scc1K, this region might contribute to DNA binding. This should be experimentally tested. In the human SA2-Scc1K structure, the corresponding region in Scc1K forms a helix that is located at the base of the "nose" of SA2. Can the authors build a model of the SA2-Scc1-DNA complex and see if this N-terminal region of Scc1K might be close to DNA?

NOTE: Concern on the function of K363 was raised by two reviewers and is reported here in its original wording as points 6a and 6b.

7) The ChIP-qPCR is essential for the conclusions of the paper but there are some issues with the presented experiment in Figure 3D. A minimum of 3 biological repeats are required to compute standard deviation, so the error bars here are not appropriate and should be removed. The authors could show the data for the two biological replicates side by side without error bars as an indication of reproducibility or, better, repeat the experiment a third time and calculate standard deviation. What do the percentages mean above the bars? How did the authors analyse the heptamutant given that it is not viable? Do the cells also carry endogenous Scc3? In this case, do all the strains in this experiment carry untagged Scc3 in addition to the tagged wild type or mutant protein? The fact that patch 2 mutants still bind DNA in vitro predicts that the patch 2 mutant protein should also associate with the chromosome in vivo, but this was not tested.

8) The authors are proposing that the reported interactions are required for cohesin loading. However, an alternative possibility is that "core" cohesion can load but that Scc3 fails to associate with it. This should be tested by assessment of the association of other "core" cohesin subunits (Smc1/Smc3/Scc1) with chromosomes in the patch 3 mutant cells.

9) What is the effect of the observed mutations on sister chromatid cohesion? The authors should test this using the TetR-GFP or LacI-GFP system.

10) Hara et al. (2014) showed that mutating conserved basic residues in the N-terminal and middle regions of Scc1K did not affect Scc1K binding to SA2. These regions may transiently dissociate from SA2/Scc3 while the C-terminal region of Scc1K is still anchored to SA2/Scc3. It is thus possible that Scc3-Scc1K form a topological embrace of DNA, similar to the condensin sub-complex. This possibility needs to be discussed, especially if the N-terminal region of Scc1K is required for DNA binding (see point 6b).

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

Thank you for submitting your article "Structural basis for Scc3-dependent cohesin recruitment to chromatin" for consideration by eLife.

I have now examined your resubmission and I am happy to inform you that I consider it essentially ready for acceptance. However, before formal acceptance, I would like to note the following three points:

1) You seem to be using two somewhat different color schemes for Scc3 in the different figures, more bluish in Figure 2 and Figure 4, and more violet in Figure 1. May I suggest that you make the colours more uniform?

2) In Figure 2—figure supplement 1E, the right hand panel appears to be a composite of pasted lanes. If this is the case, could you please clearly mark this on the figure with a black vertical line and add a short reference to lane pasting in the legend?

https://doi.org/10.7554/eLife.38356.021

Author response

Essential revisions:

1) In the Discussion section (first paragraph), the authors set up a straw man. The only alternative to direct protein-DNA contacts (either by a cohesin subunit or by some adaptor protein) is that cohesin is snapping open and closed all the time and should it happen to entrap DNA, it somehow sticks. That silly alternative is obviously hugely improbable. Thus, the real conclusion is that "these findings provide evidence for a site of DNA contact, probably during the initial step in chromosome entrapment" – or something like that. See also point 3, below. Incidentally, "topological principles" don't "drive" anything in biology – they explain various mechanisms or activities and suggest why they may have evolved, but biology is never "driven" by "principles" other than natural selection (except in the minds of the unreconstructed Cartesians still running around in France).

We agree with the reviewers’ point and revised our statement accordingly (Discussion section et seq.): ‘In this study, we identified a DNA-binding site in the Scc3 subunit of the cohesin complex and determined its interaction with DNA at near-atomic resolution. These findings provide evidence for direct DNA-protein contacts in the cohesin complex, which presumably play a major role in the initial step of chromosome entrapment and/or possibly DNA translocation.’

2) The anisotropy of the crystals, presumably due to a less than optimal DNA length, limits the accuracy and the information content of the structure. Did the authors, seeing the result, take the obvious next step of trying to get better crystals with other DNA lengths (e.g., 20 or 21 bp)? (That is, were all DNAs in Supplementary file 2 used in crystallization trials, or just for FP measurements? If the former, why not also 20 bp?) In this reviewer's view, that would have been an easier and better path than the one they took by validating the Scc1 trace with SeMet.

Improving crystal quality by testing different DNA lengths was indeed our strategy. As indicated in the Materials and methods section, the initial DNA duplex length for spontaneous crystallization was 17 bp. We have tried extensively to improve these crystals using DNA of various lengths, including those shown in Supplementary file 1 (previously Supplementary file 2). We obtained the best-diffracting crystals by using the 17-bp DNA crystals as ‘seeds’ for crystallization of a complex containing a 19mer DNA. Our expectation was that a DNA substrate of 20/21bp would be ideal, but none of the longer DNA duplexes allowed crystallization.

In any case, Figure 3—figure supplement 3 should show either the DNA density in the initial MR map or (if the MR is good enough – it might not be) an Fo-Fc map after that initial step (i.e., phases from MR but 2Fo-Fc, showing in principle what's missing). In any case, a map with the final 2Fo-Fc phases, which included this DNA contributions, is not helpful. Incidentally, in Supplementary file 1, the Rmerge in the last bin is truly miserable. Was there an "elliptical" (i.e., anisotropic) cutoff, or did the meaningless reflections in the "bad" directions contribute? If the latter, then please recalculate with the correct anisotropic cutoff for each frame or set of frames, so that pure noise doesn't contribute to the data used. Also, "3.99" is 4.0 in my book, not "3.9".

We now show a 2Fo-Fc map after the initial MR step, in which the DNA was omitted from the model (Figure 3—figure supplement 1A).

We thank the reviewer for pointing out the high Rmerge in the highest resolution bin. Indeed, we had not applied an elliptical cutoff and noisy reflections contributed to the data used. We have reprocessed the data using an elliptical cutoff, as suggested. Overall statistics improved and we were now able to include reflections up to 3.6Å. We have refined the model (maintaining and extending the previous set of Rfree reflections) against this reprocessed data and have updated Table 1 accordingly.

To ensure that potential users have access to the data prior to additional processing, we have deposited unprocessed structure factors. Table 1 now shows data statistics after additional anisotropic data processing.

3) The text overstates some of the biological and mechanistic conclusions. Although the correlation of affinity in vitro with function in vivo does permit the inference that the observed interaction is part of the DNA docking mechanism, the results do not rule out the participation of other contacts. Indeed, were those contacts strengthened by compensating mutations, it is possible that this contact would not be "indispensible", as the Abstract states.

We have revised our Abstract to avoid any overstatement: ‘These findings suggest that the Scc3-Scc1 DNA-binding interface plays a central role in the recruitment of cohesin complexes to chromosomes and therefore for cohesin to faithfully execute its functions during cell division.’

4) The most useful conclusion is the similarity with condensin. For understandable "psychological" reasons, the authors do not mention the Kschonsak et al. (2017) paper in the Introduction. They should do so, as it surely guided their strategy at some point, either consciously or otherwise. Is the peptide loop definitely absent here, or could its absence be a consequence of truncating Scc1?

We now refer to the paper by Kschonsak et al. (2017) in the Introduction: ‘Recent work identified a direct DNA-binding site in the paraloguous condensin complex, where the HEAT repeat subunit Ycg1 in complex with the kleisin subunit Brn1 contact the DNA double helix backbone and stabilize its association through DNA entrapment within a Brn1 peptide loop (Kschonsak et al., 2017).’

We cannot exclude for certain that the peptide loop is lacking in Scc1, as it might be absent due the truncation construct used in our experiments. However, as mentioned previously, data from the Nasymth lab (Chan et al., 2012; Roig et al., 2014; Petela et al., 2018) indicate that Scc1 constructs with mutations/truncations in the relevant area are viable, thus indicating that such a peptide loop, if present at all, may not be essential for cohesin function in budding yeast.

5) At the end of the Discussion section, the authors write that the mechanism they describe would enable cohesin to entrap a second DNA helix without releasing the first, etc. Not obvious to this reader why or how, perhaps because Figure 4C is so vague and incomprehensible.

DNA translocation during loop extrusion. We have modified the Discussion section to clarify this point.

6a) Is K363 on Scc1 important for the DNA binding ability of the complex? It would also be useful to highlight this residue on Figure 2. This experiment is required to confirm the in vivo relevance of the enhancement of DNA association by the Scc1 fragment in the in vitro experiments.

6b) Scc1 is clearly required for the DNA-binding activity of Scc3-Scc1. The structure suggests that Scc1 K363 might contact DNA. Does Scc1 K363E reduce the binding of Scc3-Scc1? Related to this, even though the authors cannot see any density of the N-terminal region of Scc1K, this region might contribute to DNA binding. This should be experimentally tested. In the human SA2-Scc1K structure, the corresponding region in Scc1K forms a helix that is located at the base of the "nose" of SA2. Can the authors build a model of the SA2-Scc1-DNA complex and see if this N-terminal region of Scc1K might be close to DNA?

NOTE: Concern on the function of K363 was raised by two reviewers and is reported here in its original wording as points 6a and 6b.

To address questions 6a and 6b, we have mutated Scc1 residue K363, as well as the neighboring R364, to arginine. We have also updated Figure 2 to highlight residue K363, as requested.

We found that Scc3-Scc1 complexes with the K363E/R364R double mutation display a DNA equilibrium dissociation constant that is essentially indistinguishable from that of the wild-type complex. We have added a description of these data in the Results section et seq. and updated Figure 3C accordingly. As we observed no phenotype of this mutation in vitro, we anticipate that yeast cells that express this mutant version of Scc1 would most likely show no advert phenotype, taking into account that even cells that express individual Scc3 patch mutants are perfectly viable (Figure 3E). We therefore did not further test this mutant Scc1 in vivo.

To further test the possibility that the disordered N-terminus might contribute to DNA binding, we removed the disordered region of Scc1K spanning amino acids 301–354. Unfortunately, the truncated Scc1K fragment (amino acids 355–400) failed to co-purify with Scc3. As shown in the SDS-PAGE gel (Author response image 1), we did not detect a band for the truncated Scc1K at the expected position, even when we highly overloaded the gel. Our interpretation is that the disordered region of Scc1K might contribute to the solubility of this fragment under overexpression conditions. This prevented us from testing DNA binding of a minimal Scc3T/Scc1 complex to investigate whether the disordered segment of Scc1K contributes to DNA binding.

In an alternative attempt to address whether the missing region of Scc1 might potentially interact with DNA, we built a composite model of the SA2-Scc1-DNA complex, as suggested by the reviewers. As shown in Author response image 2, the DNA is principally accommodated in the DNA binding ‘cradle’ of SA2 in this model. The N-terminal region of Scc1, including the mentioned a helix334-342, is clearly located too far away to engage in direct DNA contacts. Thus, while we cannot exclude the possibility that Scc1 topologically embraces DNA, similar to the situation in the condensin Ycg1-Brn1 subcomplex, there is no evidence that it directly contributes to the DNA-binding activity of Scc3-Scc1.

7) The ChIP-qPCR is essential for the conclusions of the paper but there are some issues with the presented experiment in Figure 3D. A minimum of 3 biological repeats are required to compute standard deviation, so the error bars here are not appropriate and should be removed. The authors could show the data for the two biological replicates side by side without error bars as an indication of reproducibility or, better, repeat the experiment a third time and calculate standard deviation. What do the percentages mean above the bars? How did the authors analyse the heptamutant given that it is not viable? Do the cells also carry endogenous Scc3? In this case, do all the strains in this experiment carry untagged Scc3 in addition to the tagged wild type or mutant protein? The fact that patch 2 mutants still bind DNA in vitro predicts that the patch 2 mutant protein should also associate with the chromosome in vivo, but this was not tested.

As requested, we are now showing the original data points in a revised version of the graph. The effects of the patch 3 mutation and heptamutant on condensin binding to chromosomes are very clear and consistent over all loci tested, and we are convinced that this wouldn’t change in a third biological repeat.

To be able to measure also nonviable mutants, we performed the ChIP experiments in diploid yeast strains that express an ectopic copy of Scc3-PK6 (including the wild-type control) and an untagged wild-type copy of Scc3 from one of its endogenous alleles (see detailed genotypes in Supplementary file 3).

To test the relevance of the DNA binding site revealed in our crystal structure, we decided to test one mutant that reduces (patch 3) and one mutant that largely abolishes (heptamutant) the positive charge in the DNA binding groove for ChIP experiments. Since the in vivo chromosome-binding data excellently correlates with the in vitro DNA-binding data (compare Figure 3C and 3D), we are convinced that the data presented in this figure are reasonably representative. The central conclusion of these data is that integrity of the DNA-binding surface of Scc3 is important for the association of cohesin with chromosomes. In our view, testing patch 1 or 2 mutant data in vivo would not provide any major additional insights.

8) The authors are proposing that the reported interactions are required for cohesin loading. However, an alternative possibility is that "core" cohesion can load but that Scc3 fails to associate with it. This should be tested by assessment of the association of other "core" cohesin subunits (Smc1/Smc3/Scc1) with chromosomes in the patch 3 mutant cells.

Previous work has shown beyond doubt that chromosome association of the other three core cohesin subunits strictly depends on the Scc3 subunit (Toth et al., 1999). Furthermore, we show that heptamutant and wild-type Scc3 bind Scc1 to a similar degree (Figure 3C). By extension, they must be incorporated into cohesin complexes in yeast cells with equal efficiency, indicating that mutagenesis of the DNA-binding interface does not perturb recruitment of Scc3 to cohesin.

9) What is the effect of the observed mutations on sister chromatid cohesion? The authors should test this using the TetR-GFP or LacI-GFP system.

The ability of cells to divide (Figure 3E) is an excellent read-out for the capacity of cohesin mutants to generate sister chromatid cohesion. As none of the three mutant patches alone has any obvious impact on cell division, these cells must be able to generate sister chromatid cohesion to a degree that enables efficient chromosome segregation. Hence, the combination of viability assays and ChIP-qPCR experiments provide adequate biological context to the structural biochemistry presented in this manuscript.

We would also like to draw the reviewers’ attention to another recent study (Elbatsh et al., 2016), which demonstrated that cohesin levels can be reduced quite significantly (to an extent mirrored by the patch 3 mutant) in human cells without any obvious impact on sister chromatid cohesion, consistent with previous work in budding yeast (Heidinger-Pauli et al., 2010).

Since complexes harbouring the heptamutant version of Scc3 fail to load onto chromosomes (Figure 3D), they by definition cannot generate sister chromatid cohesion and consequently cells fail to divide.

10) Hara et al. (2014) showed that mutating conserved basic residues in the N-terminal and middle regions of Scc1K did not affect Scc1K binding to SA2. These regions may transiently dissociate from SA2/Scc3 while the C-terminal region of Scc1K is still anchored to SA2/Scc3. It is thus possible that Scc3-Scc1K form a topological embrace of DNA, similar to the condensin sub-complex. This possibility needs to be discussed, especially if the N-terminal region of Scc1K is required for DNA binding (see point 6b).

Considering that Scc1 is clearly required for DNA binding, such a scenario is of course plausible. We now explain in reference to condensin (Discussion section et seq.): ‘The relevant section of Scc1 that would contribute to such topological DNA entrapment is disordered in our structure. As Scc1 is clearly required for DNA binding of the Scc3-Scc1 subcomplex but not through direct DNA interactions (Figure 3C), it is possible that cohesin uses a similar mode of chromatin engagement.’

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

I have now examined your resubmission and I am happy to inform you that I consider it essentially ready for acceptance. However, before formal acceptance, I would like to note the following three points:

1) You seem to be using two somewhat different color schemes for Scc3 in the different figures, more bluish in Figure 2 and Figure 4, and more violet in Figure 1. May I suggest that you make the colours more uniform?

I have revised Figure 1 accordingly. Colors are now matching.

2) In Figure 2—figure supplement 1E, the right hand panel appears to be a composite of pasted lanes. If this is the case, could you please clearly mark this on the figure with a black vertical line and add a short reference to lane pasting in the legend?

Indeed, Figure 2—figure supplement 1E is a composite figure as non-relevant lanes have been cropped out. I have added a black vertical line and mentioned the following in the figure legend: “For the gel showing WaplFL, the black vertical line indicates the position where the gel has been cropped.”

https://doi.org/10.7554/eLife.38356.022

Article and author information

Author details

  1. Yan Li

    European Molecular Biology Laboratory, Grenoble, France
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon 0000-0001-8190-1633
  2. Kyle W Muir

    European Molecular Biology Laboratory, Grenoble, France
    Present address
    MRC Laboratory of Molecular Biology, Cambridge, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Supervision, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing
    For correspondence
    kmuir@mrc-lmb.cam.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon 0000-0002-0294-5679
  3. Matthew W Bowler

    European Molecular Biology Laboratory, Grenoble, France
    Contribution
    Methodology
    Competing interests
    No competing interests declared
    ORCID icon 0000-0003-0465-3351
  4. Jutta Metz

    1. Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
    2. Structural and Computational Biology Unit, European Molecular Biology Laboratory, Heidelberg, Germany
    Contribution
    Data curation, Methodology
    Competing interests
    No competing interests declared
  5. Christian H Haering

    1. Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
    2. Structural and Computational Biology Unit, European Molecular Biology Laboratory, Heidelberg, Germany
    Contribution
    Formal analysis, Supervision, Visualization, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon 0000-0001-8301-1722
  6. Daniel Panne

    Leicester Institute of Structural and Chemical Biology, Department of Molecular and Cell Biology, University of Leicester, Leicester, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing
    For correspondence
    panne@embl.fr
    Competing interests
    No competing interests declared
    ORCID icon 0000-0001-9158-5507

Funding

European Molecular Biology Laboratory

  • Yan Li
  • Kyle W Muir
  • Matthew W Bowler
  • Jutta Metz
  • Christian H Haering
  • Daniel Panne

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

Acknowledgements

This work was funded by EMBL. We thank Stefan Reich for assistance with FP experiments.

Senior and Reviewing Editor

  1. Andrea Musacchio, Max Planck Institute of Molecular Physiology, Germany

Publication history

  1. Received: May 14, 2018
  2. Accepted: August 13, 2018
  3. Accepted Manuscript published: August 15, 2018 (version 1)
  4. Version of Record published: September 3, 2018 (version 2)

Copyright

© 2018, Li 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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