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Structural and functional characterization of the Spo11 core complex

Spo11, which makes DNA double-strand breaks (DSBs) essential for meiotic recombination, has long been recalcitrant to biochemical study. We provide molecular analysis of S. cerevisiae Spo11 purified with partners Rec102, Rec104 and Ski8. Rec102 and Rec104 jointly resemble the B subunit of archaeal Topoisomerase VI, with Rec104 occupying a position similar to the Top6B GHKL-type ATPase domain. Unexpectedly, the Spo11 complex is monomeric (1:1:1:1 stoichiometry), consistent with dimerization controlling DSB formation. Reconstitution of DNA binding reveals topoisomerase-like preferences for duplex-duplex junctions and bent DNA. Spo11 also binds noncovalently but with high affinity to DNA ends mimicking cleavage products, suggesting a mechanism to cap DSB ends. Mutations that reduce DNA binding in vitro attenuate DSB formation, alter DSB processing, and reshape the DSB landscape in vivo. Our data reveal structural and functional similarities between the Spo11 core complex and Topo VI, but also highlight differences reflecting their distinct biological roles.


INTRODUCTION
Spo11 evolved from the DNA cleaving subunit of archaeal topoisomerase VI 1,2 , which is a tetramer of two A subunits (DNA cleaving) and two B subunits (GHKL-type ATPase) 3,4 . Concerted attack of the A subunits on opposite strands of a duplex using active-site tyrosines breaks the DNA and produces covalent 5'-phosphotyrosyl links 5 . Topo VI passes an intact duplex through the transient DSB to modulate DNA topology.
In contrast, Spo11 remains in the covalent complex 6 . Endonucleolytic cleavage of Spo11bound strands then releases Spo11 with a short oligonucleotide still covalently attached (Spo11-oligo complexes) 7 . This frees DSB ends for further exonucleolytic resection followed by homology search and strand exchange 8 .
Spo11 requires a cohort of partners 6 that are conserved functionally 9-14 despite having highly diverged sequences 15 . One critical partner (Top6BL) is homologous to the Topo VI B subunit 10,13 . Discovery of Top6BL in plants and mouse led to identification of putative B subunits in yeasts and flies. This protein in S. cerevisiae, Rec102, was long known to be important for Spo11 activity 16,17 , but its relationship with the B subunit had remained unnoticed. Structural modeling suggested that the yeast and fly B-type subunits lack the GHKL domain that mediates ATP-dependent dimerization in Topo VI, whereas the mouse and plant counterparts contain a degenerate version of this domain 10,13 .
Biochemical studies of the meiotic DSB machinery have been lacking, largely due to difficulties in purifying components. Consequently, how Spo11 engages DNA, relationships with Topo VI, and roles of Spo11 partners have remained a matter of conjecture. We present in vitro characterization of a stoichiometric Spo11 core complex from yeast. We elucidate similarities with the ancestral Topo VI but also uncover unique features related to its exaptation as the meiotic DSB generator.
Rec102 and Rec104-argued to behave as a functional unit 20 -were efficiently crosslinked (orange lines in Fig. 2a), with prominent links mapping to the top of the putative transducer domain of Rec102, at the edge of a five-stranded β-sheet (Fig. 2c). This is where the GHKL domain is seated on the transducer domain in Topo VI 3,4 (Fig. 1e).
Nearly all tested alanine substitutions in this region of Rec102 (Fig. 2c) abolished or greatly reduced interaction with Rec104 and compromised meiotic recombination without reducing protein levels ( Fig. 2e; e.g., F90A/W91A double substitution). Exceptions were the triple H134A/T135A/H136A substitution that showed reduced meiotic recombination but normal two-hybrid interaction, L149A that showed the opposite effect, and Y145A that behaved like wild type. The XL-MS and mutational data strongly indicate that Rec104 occupies the position expected for the GHKL domain.

DNA bending and binding to duplex junctions
AFM analysis of the core complex binding to linear, relaxed circle, and supercoiled plasmid DNA revealed several unanticipated DNA-binding modes (Fig. 3a). With all three DNA forms, 40-65% of the particles were bound internally to the DNA duplex (Fig. 3b, c), the presumed pre-DSB substrate of Spo11. Particles were frequently associated with DNA bends (Fig. 3d, e), suggesting that the core complex either bends its substrate or prefers prebent DNA. Two apparent subpopulations with modal bend angles of ~60° and ~120° (Fig.  3e) may represent structural isoforms of the nucleoprotein complexes.
With circular substrates, 35% and 60% of the particles (relaxed and supercoiled, respectively) fell where DNA duplexes crossed one another (Fig. 3b, c). Internal duplex binding sites are in molar excess over sites where duplexes happen to cross, so we infer that the core complex prefers DNA crossings, similar to type II topoisomerases including Topo VI [27][28][29][30] .

Tight non-covalent interactions with DSB ends
One third of particles on linear DNA were at an end and ~7% were at locations from which three DNA arms emanated, presumably simultaneous binding to a DNA end and internally to a duplex (Fig. 3b, c). We therefore surmised that Spo11 has intrinsic affinity for the cleavage product, even without a covalent link. Two-base 5′ overhangs outcompeted other lengths and end-binding did not require a 5′ phosphate (Extended Data Fig. 5a,b).
To compare end binding with internal binding to duplexes, we used 100 and 400 bp minicircles. Compared to hairpins (apparent K D ≈ 0.1 nM), mini-circles were bound with lower affinity (K D = 0.5 ± 0.04 nM for 100 bp; 3.8 ± 0.07 nM for 400 bp) (Fig. 4c). We interpret the difference between the circles as reflecting a preference for pre-bent substrates, because the 100 bp circle is more strained 34 . These findings agree with AFM of plasmid substrates.
Mg 2+ enhanced DNA binding (Extended Data Fig. 5c). Mutating the Spo11 catalytic tyrosine (Y135F) had no effect on DNA binding but mutating a presumptive metal-ioncoordinating glutamate (E233A) decreased binding ~10-fold (Fig. 4d, e). These results validate use of the Y135F mutant to compromise catalytic activity while maintaining DNA binding in vivo (e.g. refs 1,21 ) and may explain why Y135F but not E233A can be semidominant 35 .

Mapping DNA-binding sites on the core complex
To map DNA-binding surfaces, we used FeBABE footprinting, in which chelated Fe 3+ ions covalently attached to DNA generate hydroxyl radicals that cleave nearby proteins. Estimating sizes of cleavage fragments by western blotting of terminally-tagged proteins identifies protein residues that are close to DNA 36,37 .
We first used two probes based on a 25-bp duplex with a two-base 5′ overhang on one end and a biotin-streptavidin block on the other, each conjugated to five FeBABE moieties that were either proximal or distal to the overhang (Fig. 5a). Each core complex subunit was tagged in turn, all with similar DNA-binding activities (Extended Data Fig. 6). Hydroxyl radical cleavage yielded specific banding patterns for Spo11, Ski8, and Rec102, and a weak signal for Rec104 ( Fig. 5b-e, lanes 2 and 3). Cleavage required presence of the FeBABE probe, was strongly stimulated by hydrogen peroxide (compare lanes 2 and 3 with lanes 1, 4-6), and was more efficient with the proximal probe than the distal one (compare lanes 2 and 3), confirming binding to the overhang-containing end.
Cleavage positions and candidate DNA-binding (basic) residues are summarized in Fig. 5f, g, and mapped onto the model in Fig. 5h. Cleavage sites in Spo11 mapped to positions equivalent to the groove that is thought to accommodate DNA in Topo VI 3,4 . This predicted groove is positively charged in Spo11 (Fig. 5i). The main cleavage position in Rec102 (residue ~207) corresponds to the base of the transducer domain, close to the Spo11 DNAbinding groove (Fig. 5h, top right). Ski8 was cleaved along the surface predicted to point toward the Spo11 groove (Fig. 5h, top left). Ski8 may directly contact DNA, in which case the end-bound complex must be more closed upon itself than the model based on the Topo VI apoenzyme suggests. Additional support for conformational differences of the end-bound complex is presented below.
bound to a DNA end, highlighting specific residues predicted to lie on the surface of the DNA-binding groove (Fig. 5j, k, bottom panels). The spatial correlation between FeBABE positions and the predicted positions of DNA-binding residues provides strong experimental support for our model of Spo11 binding to the DSB end.
Based on these results, we mutated candidate DNA-interacting residues in Spo11. K173A and R344A mutations reduced binding, both to DNA ends and internally to duplexes (Fig.  6a). In vivo, these mutations caused reduced and delayed DSB formation as assessed by labeling of Spo11-oligo complexes (Fig. 6b) and direct DSB measurements at the CCT6 hotspot (Extended Data Fig. 7a, b). Meiotic divisions were also delayed (Extended Data Fig.  7c), possibly in part because frequently achiasmate chromosomes trigger the spindle checkpoint 38 .

Conformational changes upon DNA binding
When crosslinks within Spo11 were mapped on our model, at least six lysine pairs were further apart than the crosslinker's 27.4 Å limit (Fig. 7a, Supplementary Table 2), suggesting that Spo11 might be structurally flexible. Variability in 2D class averages from nsEM supports this interpretation (Extended Data Fig. 2). The polypeptide linker between Toprim and 5Y-CAP domains is a likely flex point 39 .
Model-clashing crosslinks in Spo11 were not observed in the presence of DNA (Fig. 7a, b, Supplementary Table 2). We interpret that Spo11 is flexible but adopts a configuration more consistent with the model when it binds to DNA. Moreover, DNA substantially changed the crosslinking spectrum across the entire core complex (Fig. 7b). Noteworthy changes included (i) loss of prominent intramolecular and intermolecular crosslinks involving the Cterminus of Rec102, suggesting that this domain is more constrained in the presence of DNA; (ii) loss of prominent crosslinks between Ski8 and Spo11 (highlighted in Fig. 2b) presumably because occupancy of the DNA-binding groove of Spo11 obstructs crosslinker access; and (iii) appearance of crosslinks involving the C-terminus of Ski8 with Rec104 and internal parts of Spo11, consistent with end-bound complexes adopting a more closed configuration. In nsEM, the core complex tended to show an elongated shape in the absence of DNA but a more compact, closed configuration with DNA (Fig. 7c).

A DNA-binding mutant of Spo11 alters DSB processing in vivo
To elucidate how DNA binding contributes to Spo11 function in vivo, we analyzed the F260A mutant, which was originally constructed because the residue was predicted to be in the DNA-binding groove 35 (Fig. 8a). Mutant core complexes bound DNA with reduced affinity, both at ends (12-fold) and internally (3.2-fold) (Fig. 8b). This mutant has a moderate DSB defect at specific hotspots 35 . Quantification of Spo11-oligo complexes indicated that global DSB frequency is reduced to about 40% of wild type (Fig. 8c).
We also observed an unexpected shift in the size distribution of Spo11 oligos. In wild type, radiolabeled Spo11-oligo complexes migrate on SDS-PAGE as two predominant species that differ by oligo size and are about equally abundant 7 , but the F260A mutant gave a strong bias toward shorter oligos ( Fig. 8c and Extended Data Fig. 8a). A similar bias was seen with the other DNA-binding mutants above (K173A and R344A; Fig. 6b). These findings suggest that nucleolytic processing of DSBs is influenced by direct DNA binding by Spo11 (see Discussion).

Spo11 DNA binding shapes the DSB landscape
S. cerevisiae DSB distributions are highly nonrandom 33,[40][41][42][43] . Most DSBs form in hotspots that mostly coincide with nucleosome-depleted regions at gene promoters 6,33,40,44 . At broader scales, chromosomes show large domains (tens of kb) that are relatively DSB-hot or DSB-cold, and at finer scale DSBs are distributed nonrandomly within hotspots. The factors shaping these distributions over different size scales remain poorly understood 6,45 .
We hypothesized that DNA binding by Spo11 influences the finest scale of the DSB landscape by selecting cleavage sites within hotspots, supported by studies of a few hotspots 32,35 . To test this idea genome-wide, we sequenced Spo11 oligos. Because a larger fraction of oligos in the F260A mutant were of the unmapped shorter class, we also mapped unresected DSBs in a sae2 background using S1-seq, which uses S1 nuclease to remove 3′-ssDNA tails of resected DSBs and 5′ overhangs of unresected DSBs 46,47 . Biological replicate maps were highly reproducible (Extended Data Fig. 8b, c). For results described below, Spo11-oligo mapping and S1-seq were largely indistinguishable. Hotspots in the mutant had similar widths as in wild type (Fig. 8d), reflecting the constraining influence of the surrounding nucleosomes 33 . However, as predicted, there was a dramatically different DSB distribution within hotspots (Fig. 8e), irrespective of how strong the hotspots were in wild type (Extended Data Fig. 8d). For example, whereas DSBs occurred across ~880 bp of the CCT6 hotspot in wild type, most of the F260A signal collapsed to a single site (Fig. 8f). Similarly at the GAT1 hotspot, DSBs formed at a smaller number of prominent sites in the mutant than in wild type (Fig. 8g). Some of the newly prominent sites were also targeted by wild-type Spo11, but often they were either relatively minor sites in wild type or not cleaved at all (e.g., the uppermost band at GAT1 for the mutant, Fig. 8g). Thus, Spo11-F260A has substantially altered site selectivity, possibly attributable to its altered affinity for DNA.
Map positions of Spo11 oligos tend to occur in the context of a rotationally symmetric base composition bias that implies preferences of Spo11 for DNA binding and catalysis 33 . We speculated that the altered site selectivity of F260A would be accompanied by changes in these preferences. To test this, we used the S1-seq maps because these are not subject to a mapping ambiguity caused by rGMP-tailing of Spo11 oligos during library preparation 33 . In wild-type, C residues were favored and G residues were disfavored at the position 5′ of the scissile phosphate, and dinucleotides encompassing the scissile phosphate were enriched for 5′-C[A/C/T], TA, AT and AC, but were depleted for GG and GA (Extended Data Fig. 8e, f). Moreover, there was a ~20-bp zone of strong sequence bias centered on the dyad axis ("a" region in Fig. 8h, which corresponds to the footprint deduced by docking Topo VI onto DNA) flanked by sites of weaker bias 11-16 bp on either side of the dyad axis ("b" regions in Fig. 8h, outside the predicted footprint). These patterns agree with prior findings 33 .
By contrast, F260A reduced or eliminated most of these base composition signatures. There was less enrichment for C and depletion of G 5′ of the scissile phosphate (Extended Data Other changes within the predicted Spo11 footprint included substantial weakening of the normally strong preference for A at positions -2 to -5 and for T at +2 to +5 (Extended Data Unexpectedly, we also observed that relative strengths of hotspots changed substantially (Fig. 8i). To examine these changes, we scaled the F260A map based on the global 2.5-fold decrease in Spo11-oligo complexes (Fig. 8c) and plotted the absolute log-fold change in the mutant as a function of hotspot strength in wild type (Fig. 8j). Compared with other mutants that change the DSB landscape (tel1Δ and zip3Δ), Spo11-F260A caused a wider dispersion of fold changes, and the fold changes showed a modest negative correlation with hotspot strength in wild type. In other words, hotspots vary markedly from one another in their response to F260A, with a tendency for weaker hotspots to decrease more. In tel1Δ and zip3Δ mutants, large-scale changes in relative hotspot strength occurred in a domain-wise fashion in which the behavior of individual hotspots was correlated with other hotspots within 50 kb or more 48,49 . In contrast, the F260A mutant showed no evidence of such local correlation ( Fig. 8k), indicating that the mutation affects DSB frequency in a "hotspotautonomous" rather than domain-wise fashion. We conclude that Spo11 DNA-binding activity contributes to a surprising degree to the selection of which hotspot will be cleaved, not just to the selection of cleavage site within a hotspot.

DISCUSSION
Biochemical characterization of the meiotic DSB core complex from S. cerevisiae shows that it retains molecular architecture and DNA-binding properties similar to ancestral Topo VI. Nevertheless, key differences stand out, presumably reflecting distinct biological requirements in recombination initiation vs. control of DNA topology.

Architecture of the core complex
The 1:1:1:1 stoichiometry was unanticipated because Topo VI forms a stable A 2 B 2 dimer of dimers 3,4 and because DSB formation requires two Spo11 subunits. Mouse SPO11 and TOP6BL were suggested to form 2:2 complexes when expressed in E. coli 13 . However, this conclusion was based on low-resolution gel filtration and glycerol gradient sedimentation analysis and the proteins were not shown to retain DNA-binding activity, so misfolding or nonspecific aggregation cannot be excluded. Thus, while it is possible that the purified yeast and mouse proteins behave differently, available data do not definitively establish the stoichiometry of the mammalian complex. We propose that controlled dimerization of yeast Spo11 controls DSB formation in vivo. More specifically, we suggest that dimerization of the core complex occurs in the context of recruitment to the chromosome axis by Rec114. Indeed, coimmunoprecipitation experiments indicated that Spo11 self-association depends on Rec102, Rec104, and Rec114 22 .
The core complex retains an overall topology similar to Topo VI, in which ATP-dependent dimerization of the GHKL fold is communicated by the transducer domain to the A subunit to coordinate DSB formation with passage of an intact duplex through the DNA gate 3 . The mouse and plant TOP6BL subunits have both GHKL and transducer domains 10,13 . In yeast, the B subunit appears to be split into two polypeptides with Rec102 forming the transducer domain and Rec104 occupying the position where the GHKL domain should be. Perhaps it is similar in S. pombe and Drosophila, whose TOP6BL homologs also lack GHKL domains 13,50 .
One possibility is that Rec104 evolved from part of the ancestral Top6BL subunit and now remains as a highly divergent GHKL-like fold. In potential support of this idea, although secondary structure prediction was uninformative about a potential relationship with GHKL (Extended Data Fig. 9a), iTasser 51 generated a three-dimensional model for Rec104 with similarity to the GHKL fold when Topo VIB was the template (Extended Data Fig. 9b). This model also included partial overlap with the helix-two-turns-helix (H2TH) domain of Top6B that was not identified in the mouse homolog but was proposed to remain in vestigial form in the plant homolog 10,13 . Another possibility is that Rec104 is a wholly unrelated structure occupying the GHKL position. In either case, Rec104 lacks recognizable ATPase motifs. The mouse and plant TOP6BL proteins also lack important ATPase residues 10,13 . Topoisomerases are proposed to use ATP to avoid entrapment in the covalent complex 52 . Perhaps loss of ATPase activity was a route to yield the meiotic DSB enzyme from the ancestral topoisomerase.
The S. cerevisiae core complex also includes Ski8, which is separately part of the Ski complex that provides mRNA to the exosome for degradation 19,25 . Spo11-Ski8 interaction is essential for DSB activity 19 and integrity of the core complex, but its specific function is unclear. Ski8 has a WD40 structure, something often involved in protein-protein interactions 23 . Ski8 may also directly participate in DNA binding, as suggested by FeBABE footprinting. However, FeBABE assays for proximity, not interaction, so whether Ski8 directly contacts DNA is unclear. The role of Ski8 in DSB formation is conserved in S. pombe and S. macrospora, but not Arabidopsis 53-55 .

Substrate specificities and reaction dynamics
We propose that the different modes of interaction between the core complex and DNA revealed by AFM correspond to intermediates in a multi-step reaction pathway. Spo11 may introduce a bend as it engages DNA, then may trap a second duplex within a closed dimer, similar to Topo VI (Extended Data Fig. 10a) 3,30 . In this model, catalysis occurs on a trapped junction and the core complex then remains tightly associated to the DSB end through both non-covalent protein-DNA interactions and the covalent phosphotyrosyl link.
An important caveat is that our data do not establish a temporal relationship between the binding modes. Moreover, although the end-bound complex is clearly monomeric, the stoichiometry of the other DNA-bound complexes is unclear. It is possible that binding to two crossed DNA duplexes and/or the three-armed complexes containing an internally bound duplex and a DNA end reflect assembly of dimers of the core complex (heterooctamers).
The similarities in DNA-binding properties between the core complex and type II topoisomerases are intriguing. The biological function of Spo11 does not require strand passage, so it is unclear why Spo11 retained affinity for DNA junctions [27][28][29][30] . One possibility is that Spo11 captures both sister chromatids to restrict cleavage to just one. An alternative invokes tethering of the core complex to the chromosome axis: perhaps the uncleaved duplex corresponds to axis DNA while the duplex that experiences the DSB corresponds to the loop.

DNA binding residues and substrate selection by Spo11
FeBABE footprinting provided direct experimental support for the predicted DNA-binding surface on Spo11 and identified residues K173 and R344 as likely DNA-interacting residues that affect cleavage in vivo. Genome-wide DSB mapping with the DNA-binding mutant F260A revealed profound effects on the DSB landscape, showing that Spo11 directly influences both fine-scale and hotspot-scale selection of cleavage sites.
The attenuated binding affinity of F260A means that Spo11 preferences contribute less to site selection, and other factors now contribute more, such as chromatin structure (including nucleosome occupancy, H3K4me3 binding by Spp1), competition with other proteins for DNA access, and DNA binding by Spo11-associated proteins. The remaining base composition bias in the mutant is strongly concentrated around the dyad axis, suggesting that it reflects direct constraints on catalysis and/or binding.

The effect of Spo11 DNA binding on DSB processing
The DNA-binding mutants K173A, F260A and R344A yielded fewer of the longer Spo11 oligos. The provenance of the long and short oligo classes was previously unclear 33,56 , but was recently explained through studies of DSB patterns 57 . Spo11 can often cut the same DNA molecule twice in close proximity; when it does, the cuts are spaced at intervals with a 10-bp periodicity that is interpreted to mean that adjacent Spo11 complexes engage the same side of a DNA duplex. This suggests that Spo11 complexes are tethered on a surface 57 . Rec114-Mei4 and Mer2 complexes undergo DNA-driven macromolecular condensation 58 , so we propose that Rec114-Mei4-Mer2 nucleoprotein assemblies attached to the chromosome axis provides a surface that is occupied by co-oriented Spo11 core complexes that can engage DNA simultaneously (Extended Data Fig. 10b).
Short oligos are enriched at the edges of DSB hotspots, suggesting that oligo size is dictated in part by the access of MRX/Sae2, which is proposed to be inhibited at the center of the hotspot by arrays of Spo11 complexes 57 . The depletion of the long-oligo class we observed with Spo11 DNA-binding mutants is consistent with this model, because reduced occupancy of hotspot DNA by mutant complexes should improve the access of Mre11 (Extended Data Fig. 10c).

Spo11 end binding and DSB repair
Usual models envision that Spo11-oligo complexes dissociate from the complementary 3'end after MRX/Sae2-mediated clipping, although it has been argued that longer oligos might require active displacement 7,15 . In fact, the remarkably tight end binding by the core complex suggests that Spo11-oligo complexes may cap most DSB ends, potentially affecting subsequent repair. In support, recent mapping of ssDNA-dsDNA junctions in mouse spermatocytes revealed patterns of recombination intermediates consistent with the 3' end remaining annealed to the Spo11 oligo 59,60 . Because the core complex is tethered to the chromosome axis, probably embedded in Rec114-Mei4-Mer2 condensates, end-capping might maintain a physical connection between the two broken ends during recombination, facilitating repair and reducing the risks of gross chromosomal rearrangements.

Preparation of expression plasmids and baculoviruses
Oligonucleotides (Integrated DNA Technologies) are in Supplementary Table 3. Plasmids are in Supplementary Table 4. Expression vectors for untagged Spo11, Ski8, Rec102 and Rec104 were generated by cloning sequences from SK1 in pFastBac1 (Invitrogen). Cterminal 10xHis-3xFlag tags on Spo11 or Ski8 were introduced by inverted PCR and self ligation; N-terminal tagging of Rec102 was achieved by cloning in pFastBac-HTb-Flag, which codes for 6xHis and 2xFlag tags. N-terminal MBP-tagged vectors were generated by amplifying MBP from pMAL-c2x and cloning into the untagged expression construct using appropriate restriction enzymes. Point mutants were generated by QuikChange mutagenesis. The viruses were produced by a Bac-to-Bac Baculovirus Expression System (Invitrogen) following the manufacturer's instructions. The C-terminal tag on Spo11 is known to be functional in vivo 61 .

Expression and purification of recombinant proteins
Spo11 core complexes were expressed by co-infecting Spodoptera frugiperda Sf9 cells with a combination of viruses at a multiplicity of infection (MOI) of 2.5 each. Spo11 HisFlag :Ski8:Rec102:Rec104 complex used viruses produced from vectors pCCB592, pCCB587, pCCB588 and pCCB589. Spo11 HisFlag :Ski8: MBP Rec102:Rec104 complex used pCCB592, pCCB587, pCCB617 and pCCB589. Other tagged or mutant complexes were prepared using the appropriate combination of viruses (vector list in Supplementary Table  4).
Cells were harvested 62 h after infection, washed with phosphate buffered saline (PBS), frozen in dry ice and kept at −80 °C until use. A typical purification was performed with cell pellets from 0.5-2 l culture. All purification steps were carried out at 0-4 °C. Cell pellets were resuspended in 4 volumes of lysis buffer (25 mM HEPES-NaOH pH 7.5, 500 mM NaCl, 0.1 mM DTT, 20 mM imidazole, 1× Complete protease inhibitor tablet (Roche) and 0.1 mM phenylmethanesulfonyl fluoride (PMSF)). Cells were lysed by sonication and centrifuged at 43,000 g for 30 min. Cleared extract was loaded onto 1 ml pre-equilibrated NiNTA resin (Qiagen). The column was washed extensively with Nickel buffer (25 mM HEPES-NaOH pH 7.5, 500 mM NaCl, 10% glycerol, 0.1 mM DTT, 20 mM imidazole, 0.1 mM PMSF). The tagged complexes were then eluted in Nickel buffer containing 250 mM imidazole. MBP-tagged complexes were further purified on amylose resin (NEB). The core complex without MBP was instead further purified on Anti-Flag M2 affinity resin (Sigma). Fractions containing protein were pooled and diluted in 3 volumes of Amylose/Flag buffer (25 mM HEPES-NaOH pH 7.5, 500 mM NaCl, 10% glycerol, 2 mM DTT, 5 mM EDTA). Next, the complexes were bound to 1 ml of the appropriate resin in a poly-prep chromatography column (Bio-Rad) and the resin was washed extensively with buffer. Complexes were eluted from amylose resin with buffer containing 10 mM maltose, or from anti-Flag resin with buffer containing 250 μg/ml 3xFlag peptide (Sigma). Fractions containing protein were pooled and loaded on a Superdex 200 column preequilibrated in Amylose/Flag buffer. Fractions containing protein were concentrated in 50-kDa-cutoff Amicon centrifugal filters (Millipore). Aliquots were frozen in dry ice and stored at −80 °C. Mutant core complexes were prepared using the same procedure as the wild-type proteins.

Electrophoretic mobility shift assays
Binding reactions (20 μl) were carried out in 25 mM Tris-HCl pH 7.5, 7.5% glycerol, 100 mM NaCl, 2 mM DTT, 5 mM MgCl 2 and 1 mg/ml BSA with 0.1 nM DNA (for hairpin substrates) or 1 nM DNA (for mini-circles) and the indicated amounts of protein complexes. Complexes were assembled for 30 minutes at 30 °C and separated on a 5% Tris-acetatepolyacrylamide/bis (80:1) gel containing 0.5 mM MgCl 2 at 200 V for two hours. Gels were dried then analyzed by phosphorimaging (Fuji). Apparent dissociation constants (K D ) were estimated from protein titration EMSA experiments as the concentration of protein at which 50% of the substrate was bound. Because binding to 5′ overhang substrates was so tight (apparent K D close to the DNA substrate concentration in the assay), these measurements are approximations of the true K D .

Mapping of the protein-DNA interface
The sizes of Flag-tagged fragments of Spo11, Ski8, Rec102 and Rec104 generated by hydroxyl radical cleavage were determined by comparing their migration with MagicMark™ XP Western protein standards (Invitrogen). Lane profiles were quantified using ImageGauge. Peaks identified for molecular weight standards were fitted to a fourth order polynomial equation of migration distance as a function of molecular weight. The accuracy with which fragment sizes are estimated was assessed by comparing deduced trypsin and chymotrypsin cleavage sites with expected cleavage sites. Based on multiple experiments, we estimate that cleavage sites are mapped within 5-10 residues.

Structural modeling
The Ski complex is from PDB accession number 4BUJ 25 . Ski8 intramolecular crosslinks were mapped on 1SQ9 23 . The homology model for Spo11 and Rec102 was reported previously 13 . Secondary structure predictions were generated by PsiPred 63 . The 3D model of Rec104 was generated by iTasser 51 with Topo VIB (PDB: 2Q2E) as a template (C-score = −2.15, estimated TM-score = 0.46 ± 0.15, estimated RMSD = 10.0 ± 4.6 Å). The model of the core complex was generated using Pymol. The pdb files are available upon request.

AFM imaging
Relaxed and linear plasmids for AFM were prepared by treatment of pUC19 with topoisomerase I and NdeI, respectively. Core complexes were diluted to a final concentration of 4 nM in the presence of 1 nM DNA in 25 mM HEPES-NaOH pH 6.8, 5 mM MgCl 2 , 50 mM NaCl, 10% glycerol. Complexes were assembled at 30 °C for 30 minutes. A volume of 40 μl of the protein-DNA binding reaction was deposited onto freshly cleaved mica (SP1) for 2 minutes. The sample was rinsed with 10 ml ultrapure deionized water and the surface was dried using a stream of nitrogen. AFM images were captured using an Asylum Research MFP-3D-BIO (Oxford Instruments) microscope in tapping mode at room temperature. An Olympus AC240TS-R3 AFM probe with resonance frequencies of approximately 70 kHz and spring constant of approximately 1.7 N/m was used for imaging. Images were collected at a speed of 0.5-1 Hz with an image size of 2 μm at 2048 × 2048 pixel resolution.
For AFM imaging of proteins alone, samples were diluted in 25 mM HEPES-NaOH pH 7.5, 500 mM NaCl, 10% glycerol, 2 mM DTT, 5 mM EDTA. Immobilized proteins were imaged with a Bruker FMV-A AFM probe with resonance frequencies of approximately 75 kHz and spring constant of approximately 2.5 N/m. Images were collected at a speed of 0.8 Hz with an image size of 1 μm at 512 × 512 pixel resolution. For volume analyses, raw data were exported into 8-bit grayscale tiff images using the Asylum Research's Igor Pro software. Tiff images were then imported into FIJI/ImageJ (NIH) for quantification of volume using a custom written FIJI code. A height threshold of 0.25 nm was set for each protein sample, as well as a pixel area threshold, to exclude noise from the image. The volume of each structure was calculated using the formula V = I avg · Z conversion · A p · XY conversion 2 where I avg is the average intensity, Z conversion is the conversion of one gray scale unit of intensity into height in nanometers, XY conversion is the pixel to nanometer conversion for the image in xy, and A p is the area of particles in pixels. The predicted volume of protein complexes can be calculated according to the equation: V c = V 1 × M 0 / N 0 , where M 0 is the molecular weight, N 0 is Avogadro's number and V 1 is the specific volume for proteins (0.73 cm 3 g −1 ) 64 . Based on this equation, the predicted volume of a 1:1:1:1 core complex with MBP tag (187.8 kDa) or without MBP tag (145.3 kDa) is 226 nm 3 and 176 nm 3 , respectively.
Additional FIJI analysis was done for the quantification of DNA bending angles at protein-DNA interaction sites. A FIJI macro was written to allow for two lines to be manually drawn on DNA strands on either side of a DNA-protein binding event to allow for automated calculation of the angles created by the DNA strand.

Yeast strains and targeting vectors
Yeast strains were from the SK1 background (Supplementary Table 5). Flag-tagged Spo11 strains were constructed by transformation of Sph1 fragments of plasmids pSK806 (WT), pSK809 (F260A), pCCB822 (K173A) and pCCB823 (R344A), replacing the endogenous locus with the tagged construct together with a hygromycin resistance cassette.

XL-MS
For protein-alone samples, ~25 μg protein complexes were cross linked in 160 μl reactions in the presence of 2 mM disuccinimidyl suberate (DSS) in buffer containing 25 mM HEPES-NaOH pH 7.5, 500 mM NaCl, 10% glycerol, 2 mM DTT, 5 mM EDTA. After 10-20 minutes at 30 °C, reactions were quenched with 100 mM Tris-HCl pH 7.5. For crosslinking in the presence of DNA, 20 μg core complexes was bound to equimolar concentrations (100 nM) of a 25 bp hairpin substrate with 2 nt 5′-overhang in a 6-ml reaction volume. Binding reactions and crosslinking were performed as above, but with 100 mM NaCl and 5 mM MgCl 2 (instead of EDTA). After crosslinking, proteins were concentrated by acetone precipitation and resuspended in 1× Laemmli buffer. Samples were boiled, separated by SDS-polyacrylamide gel electrophoresis and stained with SimplyBlue SafeStain (Invitrogen). Slower-migrating bands representing protein-protein crosslinks were excised and digested in situ with trypsin digestion as described 65 . The tryptic peptides were purified using a 2-μl bed volume of Poros 50 R2 (Applied Biosystems) reversed-phase beads packed in Eppendorf gel-loading tips 66 . The digested peptides were diluted to 0.1% formic acid, and each sample was analyzed separately by microcapillary LC with tandem MS by using the NanoAcquity system (Waters) with a 100 μm inner diameter × 10 cm length C18 column (1.7 μm BEH130; Waters) configured with a 180 μm × 2 cm trap column coupled to a Q-Exactive Plus mass spectrometer (Thermo Fisher Scientific). A proxeon nanoelectrospray source set at 1800 V and a 75 μm (with 10 μm orifice) fused silica nanoelectrospray needle (New Objective, Woburn, MA) was used to complete the interface. 1 μl of sample was loaded onto the trap column, washed with 3× loop volume of buffer A (0.1% formic acid) and the flow was reversed through the trap column and the peptides eluted with a 1-50% acetonitrile (with 0.1% formic acid) gradient over 50 min at a flow rate of 300 nl/min over the analytical column. The QE Plus was operated in automatic, data-dependent MS/MS acquisition mode with one MS full scan (370-1700 m/z) at 70,000 mass resolution and up to ten concurrent MS/MS scans for the ten most intense peaks selected from each survey scan. Survey scans were acquired in profile mode and MS/MS scans were acquired in centroid mode at 17500 resolution and isolation window of 1.5 amu. AGC was set to 1 × 10 6 for MS1 and 5 × 10 5 and 100 ms maximum IT for MS2. Charge exclusion of 1, 2 and greater than 8 enabled with dynamic exclusion of 15 s. To analyze the cross-linked peptides we used pLink 67 . The raw MS data was analyzed using pLink search with the following parameters: precursor mass tolerance 50 p.p.m., fragment mass tolerance 10 p.p.m., crosslinker DSS (cross-linking sites K and protein N terminus, xlink mass-shift 138.068, monolink mass-shift 156.079), fixed modification C 57.02146, variable modification oxidized methionine, deamidation N,Q, protein N-acetyl, peptide length minimum 4 amino acids and maximum 100 amino acids per chain, peptide mass minimum 400 and maximum 10,000 Da per chain, enzyme trypsin, two missed cleavage sites per chain (four per crosslink). The data were imported on the xiNET online tool to generate crosslinking maps 68 . All identified crosslinks can be found in Supplementary Table 1.

SEC-MALS
The light scattering data were collected using a Superdex 200, 10/300, HR Size Exclusion Chromatography (SEC) column (GE Healthcare, Piscataway, NJ), connected to High Performance Liquid Chromatography System (HPLC), Agilent 1200, (Agilent Technologies, Wilmington, DE) equipped with an autosampler. The elution from SEC was monitored by a photodiode array (PDA) UV/VIS detector (Agilent Technologies, Wilmington, DE), differential refractometer (OPTI-Lab rEx Wyatt Corp., Santa Barbara, CA), static and dynamic, multiangle laser light scattering (LS) detector (HELEOS II with QELS capability, Wyatt Corp., Santa Barbara, CA). The SEC-UV/LS/RI system was equilibrated in buffer 25 mM Hepes pH 7.5, 500 mM NaCl, 10% glycerol, 2 mM EDTA at the flow rate of 0.5 ml/min or 1.0 ml/min. Two software packages were used for data collection and analysis: the Chemstation software (Agilent Technologies, Wilmington, DE) controlled the HPLC operation and data collection from the multi-wavelength UV/VIS detector, while the ASTRA software (Wyatt Corp., Santa Barbara, CA) collected data from the refractive index detector, the light scattering detectors, and recorded the UV trace at 280 nm sent from the PDA detector. The weight average molecular masses were determined across the entire elution profile in intervals of 1 sec from static LS measurement using ASTRA software as previously described 69 .

Negative-stain EM
The sample (4 μl of protein at 100-150 nM in buffer containing 25 mM HEPES-NaOH pH 7.5, 500 mM NaCl, 10% glycerol, 2 mM DTT, 5 mM EDTA) was applied to fresh, glowdischarged homemade continuous carbon grids. For the DNA-bound sample, the complex was mixed with a 25-bp hairpin substrate with 5′-TA overhang in 25 mM HEPES-NaOH pH 7.5, 100 mM NaCl, 10% glycerol, 2 mM DTT, 2 mM MgCl 2 . After 1-minute incubation, the grids were stained in five consecutive 40-μl drops of 2% uranyl acetate during a total of 45 seconds prior to blotting completely dry. Negatively-stained specimens were examined under an FEI Tecnai-12 TWIN electron microscope equipped with a LaB 6 filament and operated at 100 kV acceleration voltage. Data were acquired using a dose of ~25 e − /Å 2 at a nominal magnification of 49,000 × and binned to 4.14 Å per pixel. All images were recorded on a FEI Eagle 4k × 4k pixel camera (Thermo) utilizing LEGINON data collection software 70 . Particle picking, contrast transfer function estimation and phase-flipping correction were performed using EMAN 71,72 . Reference-free 2D methods implemented in EMAN 71 , and 2D maximum-likelihood classification as implemented in RELION 73 , were used to analyze an average of 10,000 particles for each condition. The particles were too conformationally heterogeneous to permit 3D volume reconstruction.

Spo11-oligo labeling
The procedure has been described 56 . Briefly, yeast cultures were harvested at the indicated time in meiosis and denatured extracts were prepared by trichloroacetic acid precipitation.
Proteins were solubilized in 2% SDS, 500 mM Tris-HCl pH 8.1, 10 mM EDTA. Extracts were diluted in an equal volume of 2× IP Buffer (2% Triton X100, 30 mM Tris-HCl pH 8.1, 300 mM NaCl, 2 mM EDTA, 0.02% SDS) and Flag-tagged Spo11-oligo complexes were immunoprecipitated on agarose beads conjugated with mouse monoclonal M2 anti-Flag antibody. DNA was labeled on the beads with terminal deoxynucleotidyl transferase and [α − 32 P]-dCTP. After washing the beads in 1× IP buffer, proteins were eluted with LDS sample buffer and separated by SDS-PAGE. The gel was dried and developed by autoradiography. Total DSB levels were estimated from relative areas under the curves for time courses. For analysis of Spo11-oligo length, radiolabeled Spo11-oligo complexes were digested with proteinase K and deproteinized oligos were separated on a 15% TBE-UREA polyacrylamide gel.

Southern blot analysis of meiotic DSB formation
Meiotic DSB analysis by Southern blotting was performed as described 74 . Briefly, synchronized cultures undergoing meiosis were harvested at the indicated times. After DNA purification, 800 ng of genomic DNA was digested with PstI and separated on a 1% TBEagarose gel. For direct comparison of break formation at GAT1 and CCT6 with genomewide DSB maps, genomic DNA from sae2 strains were prepared in agarose plugs as described 74 . Plugs were equilibrated in NEB CutSmart buffer, digested with PstI-HF, and digested DNA was separated on a 0.65% TBE-agarose gel. DNA was transferred to Hybond-XL nylon membranes by vacuum transfer, hybridized with GAT1 probe (amplified with primers 5′-CGCGCTTCACATAATGCTTCTGG and 5′-TTCAGATTCAACCAATCCAGGCTC) or CCT6 probe (amplified with primers 5′-GCGTCCCGCAAGGACATTAG and 5′-TTGTGGCTAATGGTTTTGCGGTG), and developed by autoradiography.

Heteroallele recombination assays
Heteroallele recombination assays were done in diploid strains carrying the arg4-Bgl and arg4-Nsp alleles 75 . Three independent colonies were cultured overnight in selective dropout medium, then transferred to YPA for 13.5 hours prior to meiotic induction in 2% KOAc. After at least 6 hours in meiosis, appropriate dilutions of cultures were plated on arginine dropout medium to measure frequency of Arg+ recombinants and on YPD to measure colony forming units.

Yeast extracts and western blotting
Denaturing whole-cell extracts were prepared in10% trichloroacetic acid by agitation with glass beads.

Yeast two-hybrid assays
Yeast two-hybrid vectors were transformed separately into haploid strains SKY661 and SKY662 and selected on appropriate synthetic dropout medium. Strains were mated and streaked for single diploid colonies on medium lacking tryptophan and leucine. Single colonies were grown overnight in selective medium containing 2% glucose. Cultures were diluted in fresh medium containing 2% galactose and 1% raffinose and grown until log phase (4 h). Cells were lysed and quantitative β-galactosidase assay was performed using ONPG substrate following standard protocols (Clontech Laboratories). For two-hybrid experiments in meiotic conditions, after overnight culture in selective medium with glucose, cultures were washed twice then incubated with 2% KOAc for 20 hours to induce meiosis.

Spo11-oligo sequencing and S1-seq
Processing of cell lysates for Spo11 oligo purification and sequencing library preparation were essentially as described 76 . Paired-end Illumina sequencing was performed by the Integrated Genomics Operation at MSKCC. Adapter clipping and mapping of reads was performed by the Bioinformatics Core Facility (MSKCC) as described 33 . Statistical analyses were performed using R (http://www.r-project.org/). Sequence read totals and mapping statistics are in Supplementary Table 6. Only uniquely mapping reads were analyzed. Each map was normalized to the total number of uniquely mapped reads, excluding mitochondrial DNA or the 2μ plasmid, then biological replicates were averaged. In analyses evaluating the fold change in DSB activity, the ratio was scaled according to the decrease in DSBs as measured through quantification of Spo11-oligo complexes (0.4 for F260A).
For S1-seq, yeast cultures in the sae2 background were harvested 4 hours after meiotic induction, washed with 50 mM EDTA pH 8.0 and stored at −80°C. DNA preparation in plugs and generation of sequencing libraries were done as described 47 . The rDNA locus was masked. In addition, to reduce background signal, a thresholding was applied, calculated from the frequency of reads in the coldest 10% of coding regions. A detailed description of the bioinformatic processing of S1-seq reads was previously provided 47 .

Reporting summary
Further information on experimental design is available in the Nature Research Reporting Summary linked to this article.

Code availability
Code for processing Spo11-oligo reads 48,49 is available at http://cbio.mskcc.org/public/ Thacker_ZMM_feedback/. Code for processing S1-seq reads 47 is available at https:// github.com/soccin/S1seq.      a. Binding of wild-type or mutant Spo11-containing core complexes to a 25-bp hairpin substrate with 5′-TA overhang (DSB) or a DNA mini-circle. The affinity of R344A was decreased 3-fold for DNA duplex and 4-fold for DNA ends. For K178A, the affinity was decreased 27-fold for the ends and could not be measured on duplex DNA because binding did not reach saturation. b. Labeling of Spo11-oligo complexes. Representative gels of n = 2 assays are shown above; quantification is plotted below (mean and range from two experiments). Uncropped blot and gel images and data for graphs are provided in the Source Data.   e. Altered DSB distribution within hotspots. For each hotspot (n=3908), we calculated Pearson's r between the consensus Spo11-oligo pattern in wild type (average of independent maps 49 ) and either F260A or wild type from this study, or tel1Δ and zip3Δ. f, g. DSBs in sae2Δ genomic DNA detected by Southern blotting at CCT6 (f) and GAT1 (g).
h. Nonrandom base composition (summed deviation from local average of dinucleotide frequencies) at DSB sites. Schematic illustrates where Spo11 cuts (arrows) relative to S1-seq reads. I,j. Altered hotspot strengths. (i) Unscaled RPM (reads per million mapped) compares relative strengths between example hotspots, not absolute changes.