Structural evidence for Scc4-dependent localization of cohesin loading
Tight association between sister chromatids is crucial for successful chromosome segregation in eukaryotic cell division . Cohesin , a ring-shaped protein complex that wraps around sister chromatids ( Gruber et al . , 2003; Haering et al . , 2008 ) is the molecular agent of sister chromatid cohesion , which persists from the time of DNA replication until anaphase . A distinct protein complex containing Scc2NIPBL and Scc4Mau2 ( Scc2/4 ) initiates linkage of cohesin with DNA ( Ciosk et al . , 2000 ) , and an in vitro reconstitution of cohesin loading by Scc2/4 suggests that the product of this reaction is a topological protein–DNA linkage ( Murayama and Uhlmann , 2013 ) . In metazoans , cohesin deposition by Scc2/4 is required for normal development ( Dorsett et al . , 2005; Kawauchi et al . , 2009 ) , and mutations in NIPBL , the human homolog of Scc2 , are dominant and causally linked to a severe developmental disorder , Cornelia de Lange Syndrome ( CDLS ) ( Krantz et al . , 2004 ) . In addition to initiating a connection between cohesin and chromatin , Scc2/4 determines the timing and location of cohesin loading ( Ciosk et al . , 2000; Kogut et al . , 2009 ) . Cohesin enrichment at mitotic centromeres and pericentromeres results in tension across sister chromatids when paired kinetochores attach to opposite spindle microtubules ( Tanaka , 2000 ) . Mitotic spindle checkpoint signaling senses the tension between sister centromeres to ensure correct kinetochore–microtubule attachments ( Stern and Murray , 2001 ) . Defective centromeric cohesion therefore leads to elevated rates of chromosome missegregation ( Eckert et al . , 2007; Fernius and Marston , 2009 ) . Centromeric cohesion depends on recruitment of Scc2/4 to centromeres in late G1/early S phase ( Hu et al . , 2011; Fernius et al . , 2013 ) . A group of conserved kinetochore proteins—the Ctf19 complex in yeast ( homologous to the human CCAN ) —participates in this recruitment pathway , along with the S phase kinase complex , DDK ( Fernius and Marston , 2009; Hu et al . , 2011; Natsume et al . , 2013 ) . Deletion of any of several Ctf19 complex members leads to impaired centromeric cohesion and chromosome missegregation ( Eckert et al . , 2007; Fernius and Marston , 2009; Ng et al . , 2009; Hu et al . , 2011 ) , but whether individual components make direct contact with Scc2/4 is not yet known . The cohesin loading activity of Scc2/4 in vitro requires only Scc2 ( Murayama and Uhlmann , 2013 ) . Scc4 is essential in yeast , however , and in humans , de novo Mau2Scc4 missense and nonsense mutations are significantly underrepresented in exome sequences , indicating that disruption of Mau2Scc4 function is probably dominant and lethal ( Table 1 ) . Thus , Scc2 activity in vivo must depend on Scc4 in ways not recapitulated by the in vitro loading reaction . We report here the structure of yeast Scc4 in complex with an N-terminal fragment of Scc2 and demonstrate that Scc4 determines cohesin localization through a conserved patch on its surface . These findings show that Scc4 targets cohesin loading to a specific genomic locus and that this function is separable from its essential role in establishing sister chromatid cohesion across the genome . 10 . 7554/eLife . 06057 . 019Table 1 . De novo mutation profiles for human NIPBL and Mau2DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 019ObservedExpectedNIPBL Synonymous5858 . 4 Missense88160 . 9 Loss of function018 . 4Mau2 Synonymous2526 . 6 Missense1352 . 7 Loss of function03 . 8
We prepared full-length Scc2/4 by co-expressing both proteins in baculovirus-infected insect cells . Scc2 is a 1493 amino acid residue protein predicted to have an unstructured N-terminal segment and a C-terminal HEAT ( Huntington , EF3 , PP2A , TOR1 ) /ARM ( Armadillo ) repeat domain ( Figure 1A ) . Scc4 , also conserved among nearly all eukaryotes , is predicted to have an extensive TPR ( TetratricoPeptide Repeat ) architecture . Negative stain electron microscopy of the full-length complex showed two large globular structures , variably positioned relative to each other , which we interpret as corresponding to the Scc2 HEAT repeat module and Scc2N-Scc4 ( Figure 1B ) . We used limited proteolysis and mass spectrometry to identify an Scc4-containing subcomplex ( Figure 1—figure supplement 1A ) . An N-terminal fragment of Scc2 ( residues 1–181 or 1–205 ) is sufficient for stable association with full-length Scc4 ( Figure 1C ) . Both the truncated complex of Scc21–181/Scc4 and full-length Scc2/4 are heterodimers in solution ( Figure 1—figure supplement 1B ) . 10 . 7554/eLife . 06057 . 003Figure 1 . Purification of the cohesin-loading complex . ( A ) Domain organization of Scc2/4 . Dotted lines show the Scc2–Scc4 interaction . An arrow indicates the position of a regulated cleavage site ( Woodman et al . , 2014 ) . ( B ) Negatively stained Scc2/4 visualized by electron microscopy . Individual particles are shown . ( C ) Gel filtration chromatograms and SDS-PAGE show that Scc2FL/Scc4 ( magenta , left inset ) and Scc21–181/Scc4 ( purple , right inset ) form stable complexes ( * marks an Scc2 cleavage product ) . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 00310 . 7554/eLife . 06057 . 004Figure 1—figure supplement 1 . Purification and characterization of an Scc2N–Scc4 complex . ( A ) Full-length recombinant Scc2/4 was digested with trypsin and analyzed by gel filtration . Fractions from peak B contain full-length Scc4 and the N-terminal 181 or 205 residues of Scc2 . A scaled absorbance ( A280 ) trace is shown above for reference . ( B ) SEC-MALS to determine the molecular weight of Scc2FL-Scc4 ( top ) and Scc21–181-Scc4 ( bottom ) . Dotted lines show calculated molecular weights for intact complexes . The measured molecular weights were as follows: 238 . 3 kDa for Scc2FL-Scc4; 104 . 1 kDa for Scc21–-181-Scc4WT; 100 . 3 kDa for Scc21–181-Scc4F324A; K327A; K331A; 103 . 0 kDa for Scc21–181-Scc4K541A; K542A; and 98 . 8 kDa for Scc21–181-Scc4L256A; Y298A; K299A; Y313A; F324A; K327A; K331A . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 004 We obtained crystals of full-length Scc4 in complex with an N-terminal , 181-residue fragment of Scc2 . Diffraction data collected from selenomethionine-substituted derivatives of these crystals allowed us to determine initial phases by single wavelength anomalous dispersion ( SAD ) . We used an incomplete model , built into a 2 . 8 Å resolution map , to obtain phase information by molecular replacement for diffraction data from crystals of a native protein complex , extending to a minimum Bragg spacing of 2 . 1 Å . Our final model includes residues 5–383 , 390–527 , and 536–622 of Scc4 and residues 1–64 , 73–95 , and 106–132 of Scc2 ( Figure 2A ) . 10 . 7554/eLife . 06057 . 005Figure 2 . Crystal structure of the Scc21–181/Scc4 complex . ( A ) Rotated views of the Scc21–181/Scc4 complex . Scc2 is shown in gray as a cartoon and transparent surface . Individual Scc4 repeats ( R1-13 ) are colored as indicated in ( B ) . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 00510 . 7554/eLife . 06057 . 006Figure 2—figure supplement 1 . Comparison of Scc2N-Scc4 with structural homologs . ( A ) Scc2N–Scc4 crystal structure superimposed onto the crystal structure of the TPR domain of kinesin light chain 2 ( teal ) in complex with a cargo peptide ( green; PDB 3ZFW ) . ( B ) Scc2N–Scc4 crystal structure superimposed onto the crystal structure of a complex between LGN ( teal ) and NuMA ( green; PDB 3RO2 ) . In both cases , Scc4 TPRN is colored dark gray , and Scc4 TPRC is colored light gray . Only Scc2 residues in contact with the internal surface of Scc4 TPRN ( Scc2G31-V64 ) are shown in magenta . For clarity , the remainder of Scc2 is either colored light gray ( Scc21–30 ) or it is not shown ( Scc265–132 ) . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 00610 . 7554/eLife . 06057 . 007Figure 2—figure supplement 2 . Structure and conservation of Scc2N . ( A ) Scc2 amino acid residues 5–40 are shown and colored according to conservation across diverse eukaryotes . 2Fo − Fc map contoured at 1 . 8σ is shown . Boxes indicate a subset of residues mutated in ( C ) . ( B ) Alignment of Scc2 amino acid sequences from diverse eukaryotes . The N-terminal fragments from all included sequences were selected , and the alignment was recalculated for this region ( shown ) . Redundant sequences and sequences lacking this fragment of Scc2 have been removed . ( C ) SCC2 alleles under the control of their native promoter were tested for the ability to restore viability to an SCC2-AID strain in depletion conditions ( 500 µM auxin ) . Δ1-138 indicates removal of Scc2 residues 1–138 . 192STOP indicates removal of all residues following Scc2Leu192 . Scc2V1361R mimics a CDLS-associated NIBPL allele found in a human patient ( Tonkin et al . , 2004 ) . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 00710 . 7554/eLife . 06057 . 008Figure 2—figure supplement 3 . Complementation of Scc4 repression by Scc2 . SCC2 or SCC4 under the control of its native promoter was tested for the ability to restore viability to a pGAL1-SCC4 strain in depletion conditions ( 2% glucose [wt:vol] ) . scc2Δ1–138 indicates removal of Scc2 residues 1–138 . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 008 Scc4 is a superhelical array of 13 TPR modules with a beta ribbon insertion between repeats 6 and 7 ( Figure 2B ) . This tightly wrapped solenoid brings successive turns in contact with each other , and the concave surface becomes an axial groove . Repeat 8 , which has a particularly long first helix , lacks a second helix and has instead an extended segment with a disordered surface loop . This irregularity divides the solenoid into two subdomains , TPRN ( Scc41–384 ) and TPRC ( Scc4391–624 ) ( Figure 2A ) . Residues 10–50 of Scc2 snake along the continuous inner cavity of Scc4 and emerge at both ends , with the N-terminus of Scc2 close to the C-terminus of Scc4 . Examples of similar TPR-peptide interfaces include the interaction of the kinesin light chain with cargo peptides ( Figure 2—figure supplement 1A ) ( Pernigo et al . , 2013 ) and the interaction of the cell polarity-determining LGN protein with its binding partners nuclear mitotic apparatus protein 1 ( NuMA ) and mInscutable ( Figure 2—figure supplement 1B ) ( Zhu et al . , 2011 ) . The extended conformation of Scc21–181 clearly depends on its contacts with the surrounding Scc4; it is likely to be unstructured in the absence of its partner . The Scc2–Scc4 interaction has two unusual features . First , the concave surface of Scc4 is entirely enclosed , and Scc2 dissociation would therefore require either unfolding or proteolysis of Scc2 or Scc4 . In fact , regulated proteolysis of Scc2 has been reported recently ( Woodman et al . , 2014 ) . Second , residues 58–132 of Scc2 extend beyond the axial groove and make extensive contact with the external surface of Scc4 . Human Mau2Scc4 binds the N-terminus of human NIPBLScc2 ( Braunholz et al . , 2012 ) , but the primary sequence of NIPBL bears little resemblance to that of Scc21–181 . We compiled sequence alignments for the N-terminus of Scc2 from divergent eukaryotes , including yeasts and humans , and mapped amino acid conservation onto its structure ( Figure 2—figure Supplement 2A , B ) . Despite their low level of overall sequence conservation , Scc2/NIPBL proteins have conserved amino acid residues at buried positions that contact Scc4 . Several Scc2–Scc4 contacts on the external face of Scc4 are also conserved , including Scc2F86 , which stacks onto Scc4Y40-Y41 , and Scc2112–120 , which forms a helix that fits over a hydrophobic surface on Scc4 TPRN . We found that Scc2 variants mutated at conserved residues contacting either the concave or the external surface of Scc4 were less effective than wild-type Scc2 in restoring viability to an Scc2-degron ( SCC2-AID ) strain under depletion conditions ( Figure 2—figure supplement 2C ) . The phenotype of these mutants was comparable to the phenotype we observed when we removed the entire Scc2 fragment visible in the crystal structure ( Δ1–138 ) . If cohesin loading can occur in the absence of Scc4 in vitro ( Murayama and Uhlmann , 2013 ) and to some extent in the absence of the Scc4-binding region of Scc2 in vivo , why is SCC4 an essential gene ? We found that supplementing the chromosomal copy of SCC2 with a plasmid coding for Scc2 or Scc2Δ1–138 complemented transcriptional repression of SCC4 ( pGAL1-SCC4 ) . That is , increasing the SCC2 gene copy number bypassed the effect on viability of transcriptional repression of SCC4 , and the bypass did not require the Scc4-binding region of Scc2 ( Figure 2—figure supplement 3 ) . Because this region of Scc2 is probably unstructured when not bound by Scc4 , it is a plausible cause of instability or aggregation when expressed unprotected . One reason why Scc4 is essential may therefore be that Scc2 is unstable in its absence . Mapping Scc4 sequence conservation onto our structure revealed a cluster of extremely conserved , solvent-facing residues , some of them invariant among all eukaryotes we examined ( Figure 3A , B ) . We found unaccounted-for electron density in this patch , into which we modeled a sulfate group . Because bound sulfates often mark phosphate binding sites in crystal structures ( Bax et al . , 2001 ) , we tested the effects of mutating conserved residues that contribute to this patch . Strains in which the endogenous copy of SCC4 had been replaced by SCC4 coding for mutations at these residues were viable but displayed a plasmid missegregation phenotype ( Figure 3—figure supplement 1A ) . Combined mutation of seven of the most conserved residues ( scc4L256L; Y298A; K299D; Y313A; F324A; K327D; K331D; scc4m7 ) did not inactivate Scc4 , as strains bearing these substitutions were fully viable and had a plasmid segregation phenotype comparable in strength to that of strains bearing mutations only at positions 324 , 327 , and 331 ( scc4F324A; K327D; K331D; scc4m3 ) ( Figure 3—figure supplement 1B , C ) . Recombinant Scc21–181-Scc4 complexes bearing these mutations behaved identically to wild-type preparations ( Figure 1—figure supplement 1B ) , indicating that perturbation of the conserved Scc4 patch does not impair the stability or folding of the rest of the protein . 10 . 7554/eLife . 06057 . 009Figure 3 . A conserved patch on the surface of Scc4 . ( A ) Surface view of Scc4 colored according to primary sequence conservation across eukaryotes . Inset shows the Scc4-conserved patch with mutated residues labeled and colored according to their effect on plasmid segregation fidelity ( Figure 3—figure supplement 1 ) . ( B ) Multiple sequence alignment of Scc4 and homologs from fungi and metazoans . Alignment is colored by conservation according to the color scheme in ( A ) . ( C ) Plasmid missegregation was measured for the indicated strains ( scc4m3—scc4F324A; K327D; K331D; error bars indicate SD; * p < 0 . 05 , Student's t-test vs WT , two tails; n . s . indicates p > 0 . 05 . ) . The dotted line shows the rate of plasmid segregation in a WT background . ( D ) Spindle length measurements for each indicated strain arrested in S phase with hydroxyurea . The dotted line shows the WT mean spindle length ( error bars indicate SD; * p < 0 . 05 , Student's t-test vs WT , two tails ) . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 00910 . 7554/eLife . 06057 . 010Figure 3—figure supplement 1 . ( A ) Plasmid segregation defects in Scc4-conserved patch mutants . Plasmid segregation in the absence of selection was measured in a chl4Δ strain and in strains with the listed Scc4 amino acid substitutions ( error bars indicate SD; *p < 0 . 05 , Student's t-test vs WT , two tails ) . The dotted line shows the rate of plasmid segregation in a WT background . ( B ) Plasmid missegregation was measured for the indicated strains ( scc4m3—scc4F324A; K327D; K331D; n . s . indicates p > 0 . 05 ) . ( C ) DNA content was measured by flow cytometry in log-phase cultures of homozygous diploids with the indicated genotypes . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 010 Strains lacking the Ctf19 complex subunit Chl4 ( CENP-N in humans ) are defective in centromeric cohesin loading because they cannot preferentially localize Scc2/4 to centromeres , although cohesin loads normally elsewhere in the genome ( Fernius et al . , 2013 ) . To determine whether the Scc4-conserved patch functions in the same pathway as Chl4 , we introduced the scc4m3 mutation into a chl4Δ strain and tested for its effect on plasmid segregation ( Figure 3C ) . Inclusion of scc4m3 does not augment the severe plasmid loss phenotype caused by CHL4 deletion . This relationship also holds true for scc4m7 ( Figure 3—figure supplement 1B ) . Strains lacking Chl4 have extended metaphase spindles , and this phenotype corresponds to weakened centromeric cohesion ( Fernius and Marston , 2009; Laha et al . , 2011 ) . We found that an scc4m3-bearing strain exhibited increased inter-spindle pole distances and that the scc4m3 mutation did not exacerbate the spindle extension phenotype of a chl4Δ strain ( Figure 3D ) . Moreover , GFP-labeled sister centromeres ( Figure 4—figure supplement 1A ) , but not chromosome arms ( Figure 4—figure supplement 1D–G ) , were separated more frequently and to greater distances in strains mutated at 5 positions in the conserved patch on Scc4 ( scc4F324A; K327A; K331A; K541A; K542A; scc4m35 ) . We conclude that the conserved Scc4 patch promotes centromeric cohesion and that it does so in a manner that may also depend on CHL4 . These results suggest that interactions at the Scc4-conserved patch target Scc2/4 specifically to centromeres . To test this hypothesis , we measured Scc2 and cohesin localization by chromatin immunoprecipitation ( ChIP ) . Perturbation of the conserved patch ( either scc4m35 or scc4m7 ) eliminates centromeric localization of Scc2 in mitotic cells and reduces association of the cohesin subunit , Scc1 , with the centromere and pericentromere , but not with chromosome arms ( Figure 4B , C , Figure 4—figure supplement 2A–D ) . We also observed this pattern of Scc2 localization in cells progressing through S phase ( Figure 4—figure supplement 2E , F ) , the stage at which cohesin loading is initiated ( Kogut et al . , 2009; Natsume et al . , 2013 ) . 10 . 7554/eLife . 06057 . 011Figure 4 . Defective centromeric cohesin loading in Scc4 conserved patch mutants . ( A ) Sister centromeres ( +2 . 4CEN4-GFP ) are separated earlier and more frequently in an scc4m35 strain entering the cell cycle after a G1 arrest . pMET-CDC20 strains of the indicated genotypes ( gray—wild type; magenta—SCC4 integrated; blue—scc4m35 integrated ) were arrested in G1 with alpha factor and released into the cell cycle in the presence of methionine to repress CDC20 expression . Solid lines show the percent of cells with separated CEN4 dots , and dotted lines show the percent of cells with separated spindle pole bodies ( Spc42-tdTomato ) . ( B ) Strains of the indicated genotypes and either Scc2-His6-3FLAG ( top ) or Scc1-6HA ( bottom ) were arrested in metaphase of mitosis following treatment with nocodazole and benomyl ( to depolymerize microtubules ) . Cells were harvested after 2 hr . Anti-FLAG or anti-HA antibodies were used for ChIP , and pulldown samples were analyzed by qPCR . Mean values of four independent experiments are shown ( error bars indicate ±SD; * p < 0 . 05; **p < 0 . 01 paired two-tailed t-test ) . ( C ) Schematic of a fragment of chromosome IV showing the location of qPCR amplicons used in ( B ) . ( D ) Scc1 enrichment in a 50-kb domain surrounding all 16 budding yeast centromeres is shown for wild-type and scc4m35 cells . For both wild type and scc4m35 , the ratio of reads ( normalized to RPM ) over input in a 100-bp window was calculated separately for each chromosome at the indicated position . The median count value for each window was then plotted to give a composite view of all 16 pericentromeres . ( E ) Scc1 enrichment along chromosome V together with a magnification of a 50-kb region including the centromere is shown . The number of reads at each position was normalized to the total number of reads for each sample ( RPM: reads per million ) and shown in the Integrated Genome Viewer from the Broad Institute ( Robinson et al . , 2011 ) . ( F ) Live cell imaging of homozygous diploid cells expressing Scc2-GFP , Mtw1-tdTomato to mark centromeres , and the indicated version of Scc4 ( left—wild type; right scc4m7 ) . GFP dots observed in wild-type cells are marked with white arrows in the first frame in which they are visible , and these foci were not observed in scc4m7 cells . Time is given relative to the start of the imaging session ( hr:mm ) . ( G ) Quantification of live cell imaging . At least 10 budding cells per field ( three fields of view each for each strain for three separate experiments ) were scored for the presence of an Scc2-GFP focus ( error bars indicate ±SD; *p < 0 . 005 , two-tailed t-test ) . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 01110 . 7554/eLife . 06057 . 012Figure 4—figure supplement 1 . Spindle pole and CEN separation but not chromosome arm separation in an Scc4-conserved patch mutant . ( A–B ) Quantification of CEN4 ( A ) and SPB ( B ) spreading observed in Figure 4A . Representative live cell images are shown at right ( green—centromeres; red—Spc42—spindle poles ) . ( C ) Budding index corresponding to measurements shown in Figure 4A . ( D–G ) Cells of the indicated genotypes carrying pMET-CDC20 , SPC42-tdTomato , and GFP at various loci were arrested in G1 with α-factor and released into the medium containing methionine . Percentages of separated GFP loci were scored at the indicated time points with tet operators integrated at: ( A ) +4 . 5CEN6-GFP in wild type ( AM5329 ) and scc4m35 ( AM15971 ) ; ( B ) −12 . 6CEN5-GFP in wild type ( AM5545 ) and scc4m35 ( AM16203 ) ; ( C ) −17 . 8CEN5-GFP in wild type ( AM5533 ) and scc4m35 ( AM15973 ) ; and ( D ) URA3-GFP in wild type ( AM1081 ) and scc4m35 ( AM16084 ) . Distances are given from the start of the tetO array from the centromere . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 01210 . 7554/eLife . 06057 . 013Figure 4—figure supplement 2 . Scc2 and Scc1 association with centromeres and chromosome arms . ( A ) Strains from Figure 4B were arrested in metaphase of mitosis following treatment for 2 hr with nocodazole and benomyl ( to depolymerize microtubules ) . Anti-FLAG ( top ) or anti-HA ( bottom ) antibodies were used for ChIP , and samples were analyzed by qPCR . Mean values of four independent experiments are shown ( error bars indicate ±SD; * p < 0 . 05; **p < 0 . 01 paired two-tailed t-test ) . ( B–C ) Whole cell extracts from strains used in ( A ) were analyzed by Western blot for expression of Scc2-6HIS-3FLAG ( B ) , Scc1-6HA ( C ) , and Pgk1 . ( D ) Strains AM1176 ( no tag ) , AM6006 ( SCC2-6HIS-3FLAG ) , AM15307 ( SCC2-6HIS-3FLAG SCC4 ) and AM17882 ( SCC2-6HIS-3FLAG scc4m7 ) or Strains AM1176 ( no tag ) , AM1145 ( SCC1-6HA ) , AM15540 ( SCC1-6HA SCC4 ) and AM17885 ( SCC1-6HA scc4m7 ) were arrested in metaphase of mitosis following treatment with nocodazole and benomyl ( to depolymerize microtubules ) . Cells were harvested after 2 hr . Anti-FLAG or anti-HA antibodies were used for ChIP , and samples were analysed by qPCR . Mean values of four independent experiments are shown ( error bars indicate ±SD; *p < 0 . 05; **p < 0 . 01 paired two-tailed t-test ) . ( E ) Strains shown are AM1176 ( no tag ) , AM6006 ( SCC2-6HIS-3FLAG ) , AM15307 ( SCC2-6HIS-3FLAG SCC4 ) , and AM15311 ( SCC2-6HIS-3FLAG scc4-m35 ) . Cells were first arrested in G1 using alpha factor and harvested 15 min after release from G1 . An anti-FLAG antibody was used for ChIP , and isolated chromatin was analyzed by qPCR . Mean values of four independent experiments are shown ( error bars indicate ±SD; *p < 0 . 05; paired two-tailed t-test ) . Cells were stained with propidium iodide and analyzed by flow cytometry ( FACS ) to confirm entry into S phase . Samples were taken prior to release from G1 ( 0 min ) and 15 min after release . ( F ) Cells were stained with propidium iodide and analyzed by flow cytometry ( FACS ) to confirm entry into S phase . Samples were taken prior to release from G1 ( 0 min ) and 15 min after release . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 01310 . 7554/eLife . 06057 . 014Figure 4—figure supplement 3 . Scc1 is reduced around all 16 individual centromeres in scc4m35 cells . ChIP-seq data from Figure 4D , E is shown for each individual centromere . Transformed sample-to-input ratios are shown for a 20-kb region surrounding each centromere ( blue—wild type; red; scc4m35 ) . DOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 014 We further analyzed Scc1 localization by ChIP followed by high-throughput sequencing ( ChIP-seq ) and found that all 16 centromeres were specifically depleted of Scc1 in the scc4m35 background ( Figure 4—figure supplement 3 ) . Scc1 depletion extended roughly 10 kilobases to either side of the core centromere but not to chromosome arms ( Figure 4D , E ) . Moreover , centromeric Scc2-GFP signal , normally visible in wild-type cells and eliminated by CHL4 deletion ( Fernius et al . , 2013 ) , was lost upon mutation of the Scc4-conserved patch ( Figure 4F , G ) . These results indicate that , like Chl4 , a specific interaction surface on Scc4 is required to target cohesin loading to centromeres .
Cohesin targeting to specific genomic locations is required in all species for robust centromeric cohesion and for tissue-specific transcriptional programs in multicellular organisms ( Kawauchi et al . , 2009; Fay et al . , 2011 ) . DNA sequences per se do not drive cohesin loading ( Onn and Koshland , 2011; Murayama and Uhlmann , 2013 ) . Instead , targeting likely depends on interactions between the loading complex and chromatin-associated factors . We have determined the structure of Scc4 bound to a minimal fragment of Scc2 , and we have used this structure to derive separation of function alleles that uncouple cohesin loading from cohesin targeting to centromeres . These experiments demonstrate that cohesin targeting in yeast depends on Scc4 . This finding is consistent with unpublished genetic evidence for an Scc4-dependent cohesin localization pathway ( Nasmyth , personal communication ) . Because the pathway that targets cohesin to centromeres requires Scc4 residues that are , in some cases , invariant across diverse eukaryotes , we suggest that the Scc4-dependent cohesin targeting we describe is a general feature of the control of cohesin loading . Consistent with previous reports ( Fernius et al . , 2013; Natsume et al . , 2013 ) , we find that the centromeric enrichment of cohesin loading is not essential for viability and that arm cohesion is unaffected in strains in which this pathway is compromised . These results probably reflect two modes for cohesin loading: one that depends on a conserved patch of Scc4 and happens at centromeres , and a second opportunistic mode that happens everywhere on the chromosome and is sufficient to support viability in the absence of the first mode . If so , Scc4-conserved patch mutations would not eliminate cohesin loading but would redistribute loading events , resulting in similar cohesin levels at centromeres and on chromosome arms . This prediction is borne out both by strains bearing Scc4-conserved patch mutations and by strains lacking CHL4 ( Fernius and Marston , 2009 ) . Recent reports have shown that cohesin targeting to specific locations along the chromosome arms of fission yeast ( Mizuguchi et al . , 2014 ) and flies ( Oliveira et al . , 2014 ) is critical for normal three-dimensional chromosome structure . In addition to findings from studies conducted in mammalian cells ( Dowen et al . , 2014 ) , one of these reports ( Mizuguchi et al . , 2014 ) suggests that cohesin complexes residing at specified sites in the genome function in gene looping and in defining chromosome territories . Because our ChIP-seq experiments show that Scc4 directs cohesin localization to broad centromeric domains but does not affect the local distribution of cohesin peaks , we suggest that the Scc4-dependent localization pathway we discuss here is overlaid upon local determinants of cohesin positioning on chromosomes . These local determinants would be a second level of cohesin regulation , operating at roughly the resolution of the transcriptional units in yeast . Integration of these spatial cues with temporal cues , including the kinase activity of DDK , could then generate the final cohesin distribution observed in metaphase cells . The Scc4-conserved patch presents a possible explanation for how broad cohesin-dense domains may be specified by the cohesion-loading complex , and it provides a molecular foundation for the study of chromatin-associated factors , both known and unknown , that localize cohesin loading .
Analysis of exome data sets was performed as described ( Samocha et al . , 2014 ) . Coding sequences for Scc2 and Scc4 were amplified from yeast genomic DNA and inserted into a modified version of pFastbac ( Life Technologies , Carlsbad , CA ) suitable for ligation-independent cloning . The expression vector contains an N-terminal 6-His tag followed by a TEV protease sequence . Full-length Scc4 and fragments of the Scc2 coding sequence were cloned by the same procedure into a bacterial expression vector for expression from a single mRNA . The coding sequences for both genes were augmented such that each contains an N-terminal 6-His tag followed by a TEV protease cleavage site . For complementation experiments , the Scc2 or Scc4 locus was amplified from Saccharomyces cerevisiae genomic DNA by PCR and cloned by restriction digest into a plasmid containing a CEN-ARS cassette and a selectable auxotrophic marker ( LEU2 ) . Genomic regions included were as follows: Scc2—chrIV:820797–825978; Scc4—chrV:465380–462755 . We used PCR stitching and isothermal assembly to generate mutated versions of these constructs . Yeast strains bearing Scc4 point mutations or gene deletions were constructed using PCR methods as previously described ( Longtine et al . , 1998 ) . All Scc4-conserved patch mutations described in this text were achieved by replacement of the native SCC4 locus . Viability of s288c strains expressing SCC4 mutants was first confirmed by complementation of pGAL1-SCC4 repression as shown for Figure 2—figure supplement 3 using plasmids bearing the SCC4 chromosomal locus . Viability of the scc4m7 strain was determined by FACS analysis of homozygous diploid cells and during strain construction by sequential sporulation of the diploid imaging strains . To generate w303 strains carrying scc4 alleles , diploid strain ( AM14499 ) carrying a heterozygous deletion ( scc4Δ::KanMX6 ) was transformed with a PCR product corresponding to full-length SCC4 ( or its mutant derivatives ) and a downstream marker ( HIS3 ) . G418-sensitive , histidine prototrophs were sporulated , and SCC4 mutations were confirmed in the haploids by sequencing . All Scc2-GFP strains are derivatives of Scc2-GFP from the Yeast GFP Clone Collection ( Huh et al . , 2003 ) . Strains for live cell imaging were constructed by integration of an Mtw1-tdTomato PCR into the Scc2-GFP strain followed by mating to achieve the final diploid strains . Plasmid segregation experiments were performed essentially as described previously ( Hinshaw and Harrison , 2013 ) . Auxin-inducible degron-tagged Scc2 ( SCC2-AID ) was generated using PCR methods ( Nishimura et al . , 2009 ) . For complementation assays , SCC2-AID cells bearing a CEN-ARS ( Chr VI ) plasmid encoding Scc2 flanked by its native control elements were grown to mid-log phase in synthetic complete ( SC ) medium lacking leucine ( to select for the plasmid ) . Cells were plated in a fivefold dilution series on a solid SC medium lacking leucine and supplemented with the indicated amount of 1-Naphthaleneacetic acid ( Auxin; Sigma-Aldrich , St . Louis , MO ) . Benomyl and nocodazole were used at 30 μg/ml and 15 μg/ml , respectively . Recombinant baculoviruses for His6-Scc2 and His6-Scc4 were amplified separately in Sf21 cells ( Life Technologies ) for three passages . For protein expression , Trichoplusia ni cells grown in suspension in Ex-Cell405 medium ( Sigma–Aldrich ) were infected with equal amounts of both viruses , and cells were pelleted and resuspended for freezing in lysis buffer ( 40 mM HEPES pH 7 . 5 , 20 mM imidazole , 50 mM NaCl , 10% glycerol , and 2 mM β-mercaptoethanol ) after 72 hr . Upon thawing and addition of protease inhibitors ( 4 μM aprotinin , 1 μM leupeptin , 1 . 4 μM pepstatin , and 1 mM PMSF ) , NaCl was added to a final concentration of 800 mM , and cells were broken by Dounce homogenization and sonication . Insoluble material was pelleted by centrifugation for 30 min at 18 , 000 rpm in a JA-20 rotor ( Beckman-Coulter , Pasadena , CA ) . 6-His-tagged Scc2/4 complexes were isolated from the supernatant by Co2+ affinity chromatography followed by ion exchange chromatography ( HiTrap SP HP , GE Healthcare , UK ) and size exclusion chromatography ( Superdex 200 20/16 , GE ) in gel filtration buffer ( 20 mM Tris–HCl pH 8 . 5 , 200 mM NaCl , 1 mM TCEP ) . To isolate Scc2N–Scc4 complexes , polycistronic expression vectors ( described above ) encoding these proteins under the control of a single T7 promoter were transformed into the Escherichia coli strain Rosetta 2 ( DE3 ) pLysS ( Millipore , Billerica , MA ) , and protein expression was induced with 400 μM IPTG at an OD600 of approximately 0 . 5 . Bacterial cultures were further incubated overnight at 18°C . Cells were resuspended and frozen in lysis buffer containing 800 mM NaCl . Upon thawing and addition of protease inhibitors , cells were lysed by sonication , and 6-His-tagged proteins were purified as described for full-length Scc2/4 . Selenomethionine-derivatized ( SeMet ) Scc2181-Scc4 samples were prepared as described for native protein samples with the exception that growth medium was prepared as described previously ( Hinshaw and Harrison , 2013 ) . Purified Scc2/4 was diluted in gel filtration buffer and adsorbed to glow discharged carbon-coated copper grids . After staining with 0 . 75% ( wt/vol ) uranyl formate , grids were imaged using a CM10 electron microscope ( Philips , Amsterdam ) . For size exclusion chromatography coupled to multiple angle light scattering ( SEC-MALS ) , experiments were performed essentially as described previously ( Hinshaw and Harrison , 2013 ) with the exception that a 3-ml size exclusion column was used for analysis of truncated Scc2N–Scc4 complexes ( Superdex 200 5/150 GL; GE Healthcare ) . Crystals of Scc21–181-Scc4 formed overnight at 18°C . For native crystals , the protein was concentrated in gel filtration buffer to 18 mg/ml and mixed in a 1-to-1 ratio ( vol:vol ) with crystallization buffer ( 0 . 2M ammonium sulfate , 16% [wt:vol] PEG 3350 ) . Crystals were washed first in wash buffer ( 160 mM NaCl , 16 mM Tris–HCl pH 8 . 5 , 1 mM TCEP , 14 . 4% PEG 3350 , and 0 . 16M ammonium sulfate ) and then in wash buffer supplemented with 30% ( vol:vol ) glycerol before flash freezing in liquid nitrogen . SeMet-derivative crystals were concentrated to 18 mg/ml , and diffracting crystals formed in crystallization buffer with 20% ( wt:vol ) PEG 3350 . These crystals were frozen as described for native versions . All diffraction data were collected on NE-CAT beamline 24ID-E . Data were indexed and scaled with XDS ( SeMet ) ( Kabsch , 2010 ) or HKL2000 ( native data ) ( Otwinowski and Minor , 1997 ) . The structure of Scc21–181-Scc4 was initially determined by SAD . To locate selenium atoms , we used SHELXD as implemented by HKL2MAP ( Pape and Schneider , 2004 ) . A search for 20 heavy atom sites with a resolution cutoff of 4 Å yielded a solution with 28 heavy atom positions . A truncated list of coordinates for 18 heavy atoms was used to generate an initial map at 3 Å resolution using Phenix Autosol ( Adams et al . , 2010 ) . After density modification using Resolve ( as implemented by Phenix ) , the map displayed extensive density corresponding to alpha helices . Placement of ideal helices and refinement using Phenix Refine yielded a partial structure , which was then used as a search model for phase determination by molecular replacement using a high-resolution native data set and Phaser-MR . During later stages of refinement , riding hydrogens were included , and TLS groups were invoked for Scc4 ( Painter and Merritt , 2006 ) . Crystallography statistics are shown in Table 2 , and the coordinates have been deposited in the Protein Data Bank , accession number 4XDN . 10 . 7554/eLife . 06057 . 020Table 2 . Crystallographic data collection and refinement statisticsDOI: http://dx . doi . org/10 . 7554/eLife . 06057 . 020Scc21–181; Scc4 ( SeMet ) Scc21–181; Scc4 ( Native ) Data collection Resolution ( Å ) 30 . 0–2 . 8178–2 . 0 Wavelength ( Å ) 0 . 9792100 . 979240 Space groupP212121P21 Unit cell dimensions ( a , b , c ) ( Å ) 58 . 6 , 89 . 0 , 178 . 051 . 9 , 178 . 1 , 52 . 7 Unit cell angles ( α , β , γ ) ( ° ) 90 , 90 , 9090 , 111 . 7 , 90 I/σ ( last shell ) 11 . 6 ( 1 . 9 ) 6 . 0 ( 1 . 3 ) Rsym ( last shell ) ( % ) 14 . 1 ( 92 . 3 ) 11 . 0 ( 72 . 9 ) Completeness ( last shell ) ( % ) 99 . 7 ( 90 . 0 ) 93 . 0 ( 89 . 7 ) Number of reflections168241154940 unique2346050878 Number of Se sites18–Refinement Resolution ( Å ) –28 . 7–2 . 1 Number of reflections–47188 working–45322 free–1866 Rwork ( last shell ) ( % ) –18 . 5 ( 28 . 7 ) Rfree ( last shell ) ( % ) –21 . 0 ( 28 . 0 ) Structure Statistics Number of atoms ( protein ) –5845 sulfate–24 solvent–301 r . m . s . d . bond lengths–0 . 004 r . m . s . d . bond angles–0 . 661 Log-phase cultures grown in SC medium were arrested in S phase with 10 mg/ml hydroxyurea for 90 min . Cells were fixed at room temperature for 10 min with 3 . 7% formaldehyde , washed twice with phosphate buffered saline , pH 8 . 5 , and resuspended in wash buffer containing 1 . 2 M sorbitol . Cells were immobilized on concanavalin-A-coated cover slips and imaged using a Nikon Ti motorized inverted microscope with a 60× objective lens ( NA 1 . 4 ) and a Hamamatsu ORCA-R2 cooled digital camera . Z-stacks ( 11 × 0 . 3 µm ) were acquired with MetaMorph image acquisition software , and maximum z-projections were generated with ImageJ . To calculate spindle pole distances , we wrote a Matlab script that identifies Spc110-mCherry foci and calculates a distance to the nearest neighbor for each instance . The list of distances was filtered to remove redundant measurements and to remove measurements arising from S phase spindles that straddled the edge of the image ( distance measurements surpassing 32 . 9 μm ) . Cell growth and measurements were carried out as described previously ( Fernius et al . , 2013 ) . Strain genotypes are listed in the strain table ( Supplementary file 1 ) . Cells were grown in an SC medium overnight and diluted 1:20 ( vol:vol ) the next morning . After 6 hr , cells were immobilized on concanavilin A-coated cover slips and a fresh SC medium was applied . Experimental and control strains were loaded in adjacent imaging chambers , and cells were maintained at 30°C with high humidity using a Tokai Hit stage top incubator . Live cell images were captured using the imaging setup described above with the exception that Z-stacks ( 8 × 0 . 3 µm ) were acquired every 8 min . We used exposure times of 10 ms ( tdTomato ) and 200 ms ( GFP ) for each image . Maximum intensity projections were generated with ImageJ for each timepoint , and figures were created with Nikon Elements software using identical processing steps and settings for each image . ChIP-qPCR and sequencing experiments were carried out as described previously ( Fernius et al . , 2013; Verzijlbergen et al . , 2014 ) . Scripts , data files , and workflows used to create the ChIP-Seq figures can be found on the github repository at https://github . com/AlastairKerr/Hinshaw2015 . ChIP-Seq data sets have been deposited with the NCBI Gene Expression Omnibus under the accession number GSE68573 . Flow cytometry was performed as previously described ( Fernius et al . , 2013 ) . 5 , 000 cells were analyzed for each sample .
The cohesin ring holds newly replicated sister chromatids together until their separation at anaphase . Initiation of sister chromatid cohesion depends on a separate complex , Scc2NIPBL/Scc4Mau2 ( Scc2/4 ) , which loads cohesin onto DNA and determines its localization across the genome . Proper cohesin loading is essential for cell division , and partial defects cause chromosome missegregation and aberrant transcriptional regulation , leading to severe developmental defects in multicellular organisms . We present here a crystal structure showing the interaction between Scc2 and Scc4 . Scc4 is a TPR array that envelops an extended Scc2 peptide . Using budding yeast , we demonstrate that a conserved patch on the surface of Scc4 is required to recruit Scc2/4 to centromeres and to build pericentromeric cohesion . These findings reveal the role of Scc4 in determining the localization of cohesin loading and establish a molecular basis for Scc2/4 recruitment to centromeres .
DNA replication copies the genetic information contained in a cell's chromosomes . A ring-like protein complex , cohesin , holds together each pair of newly-replicated chromosomes , known as ‘sister chromatids’ . When the cell divides , cohesin is cleaved; this allows sister chromatids to separate , so that each daughter cell receives one member of each sister chromatid pair and thereby inherits a full complement of genes . Defects in this process result in severe developmental abnormalities . Moreover , the genes that underlie this process are among the most frequently mutated in cancer . Cohesin is enriched at centromeres: the chromosomal points of attachment to the apparatus ( called the ‘mitotic spindle’ ) that segregates the sister chromatids into the daughter cells . The protein complex that loads cohesin onto chromosomes determines this preferential localization . The two components of the loading complex , Scc2 and Scc4 , associate as a 1:1 pair . Hinshaw et al . used a technique called X-ray crystallography to determine the structure of Scc4 bound with a large fragment of Scc2 . The result showed that the elongated Scc4 twists around the fragment of Scc2 , forming an extended groove . Scc2 snakes through the Scc4 groove and emerges at both ends . Hinshaw et al . then performed a series of experiments in yeast cells to probe how Scc4 determines the location at which cohesin loads onto chromosomes . These experiments revealed that a region on the surface of Scc4 targets both Scc4 and Scc2 to centromeres . The amino-acid sequence of this centromere-targeting patch on the surface of Scc4 is conserved across species . Thus , the mechanism by which Scc4 localizes cohesin to centromeres may be similar in all eukaryotic organisms .