Abstract
DNA double-strand breaks that initiate meiotic recombination are formed by the topoisomerase-relative enzyme Spo11, supported by conserved auxiliary factors. Because high-resolution structural data have not been available, many questions remain about the architecture of Spo11 and its partners and how they engage with DNA. We report cryo-electron microscopy structures at up to 3.3-Å resolution of DNA-bound core complexes of Saccharomyces cerevisiae Spo11 with Rec102, Rec104 and Ski8. In these structures, monomeric core complexes make extensive contacts with the DNA backbone and with the recessed 3′-OH and first 5′ overhanging nucleotide, establishing the molecular determinants of DNA end-binding specificity and providing insight into DNA cleavage preferences in vivo. The structures of individual subunits and their interfaces, supported by functional data in yeast, provide insight into the role of metal ions in DNA binding and uncover unexpected structural variation in homologs of the Top6BL component of the core complex.
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Main
Spo11 is related to the DNA-cleaving A subunit (Top6A) of archaeal topoisomerase VI (Topo VI)1. Both enzymes cut DNA through a transesterification reaction in which a tyrosine side chain severs the DNA backbone and attaches covalently to the 5′ terminus by a tyrosyl phosphodiester linkage. Two Top6A or Spo11 proteins work together to make a double-strand break (DSB) with two-nucleotide 5′ overhangs2,3.
Topo VI is a heterotetramer of two A and two B subunits. Top6A has a winged-helix (WH) domain that carries the DNA-cleaving tyrosine and a metal-ion-binding Rossmann fold known as the Toprim domain4,5,6 (Fig. 1a). Scission of each DNA strand involves the tyrosine of one Top6A monomer interacting with the Mg2+-binding pocket of the second Top6A monomer to form a hybrid active site, thus requiring dimer formation. Sequence alignments and computational modeling predict equivalent domains in Spo11 (refs. 7,8,9,10) (Fig. 1a,b).
Top6B has an N-terminal GHKL-family adenosine triphosphate (ATP) hydrolase (ATPase) domain that dimerizes upon binding to ATP and a ‘transducer’ domain consisting of a β-sheet backed by α-helices, one of which provides a long lever arm connecting the GHKL domain to the Top6A interface (Fig. 1a)4,5,11. Mouse and plant Top6B homologs (Top6BL) have the transducer and GHKL folds but the homologs from some species, including yeasts, resemble only the transducer1,12,13. Top6B also has a helix-2–turn–helix (H2TH) domain and sometimes a C-terminal domain (CTD), of unknown function. The eukaryotic homologs appear to lack these domains.
In Saccharomyces cerevisiae, Spo11 forms a ‘core complex’ with Rec102, Rec104 and Ski8 (ref. 14). Core complexes are found in most eukaryotic taxa but with substantial diversity15,16,17,18. Rec102 is homologous to the Top6B transducer but lacks a clear GHKL equivalent1,16 (Fig. 1b). GHKL loss has occurred in multiple eukaryotic lineages16. Moreover, Rec104 is enigmatic; its narrow phylogenetic distribution15 and consequent shallow multiple-sequence alignment preclude a high-confidence prediction by AlphaFold2. It has been hypothesized that Rec104 may have either evolved from or replaced the GHKL domain1,13,14,16.
Purified recombinant core complexes have a 1:1:1:1 stoichiometry, which was unexpected because Top6A forms a stable dimer4,5,6 and because DNA cleavage requires hybrid active sites. The monomeric core complexes bind noncovalently with high affinity (subnanomolar dissociation constant (Kd)) to DNA ends that mimic the cleavage product in having a two-nucleotide 5′ overhang14. This tight binding may allow Spo11 to cap DSB ends, potentially affecting subsequent repair14,19,20.
Crystal structures have been reported for Topo VI (refs. 4,5,6,11) but not for eukaryotic Spo11. Homology models for yeast core complexes templated on Topo VI (refs. 12,13,14) or predicted by RosettaFold and AlphaFold2 (ref. 9) differ from one another in terms of subunit tertiary structure and protein–protein contacts9. Moreover, no structure determination has been reported for any Spo11 or Topo VI relative bound to DNA. We address these issues with cryo-electron microscopy (cryo-EM) structures of DNA-bound yeast Spo11 core complexes.
Results and discussion
Cryo-EM structure of the core complex bound to hairpin DNA
S. cerevisiae core complexes were conformationally variable in negative-stain EM but adopted a more uniform structure when bound to DNA14. We leveraged the tight binding to DNA ends to determine a cryo-EM structure of core complexes bound to a 23-bp hairpin DNA with a two-nucleotide 5′ overhang. Spo11 binds tightly to the free end of this substrate and with much lower affinity to the hairpin end14.
Core complexes were purified after expression in insect cells (Extended Data Fig. 1a) and incubated with Mg2+ and hairpin DNA (Fig. 1c and Extended Data Fig. 1b). The structure of the protein–DNA complexes was solved at 3.7-Å resolution. The cryo-EM reconstruction is shown in Extended Data Fig. 2a–d, cryo-EM statistics are provided in Table 1 and the density representation is presented in Extended Data Fig. 2e. The model was built using subunit structures predicted by AlphaFold2 (ref. 10) and manually refined by positioning of bulky amino acid side chains.
The core complex is monomeric (1:1:1:1 stoichiometry) and adopts a V shape with Spo11 at the base contacting the three other subunits (Fig. 1d,e). The hairpin’s overhang end is anchored in a deep cleft between the Spo11 WH and Toprim domains (Fig. 1d). The remainder of the duplex makes non-sequence-specific contacts primarily with Spo11 and fewer with Rec102 and Rec104 but not with Ski8 (Fig. 1e). The proteins form a left-handed wrap around the DNA. Protein–DNA contacts in the complex are summarized in Fig. 1c.
We could trace the base, sugar and 5′-phosphate of the first nucleotide (A2) in the overhang adjacent to the terminal base pair. By contrast, the next overhang base (T1) could not be traced because of weak density. Residue Y135 involved in DNA covalent bond formation is on the opposite side of the DNA end from the Mg2+-coordinating E233, D288 and D290 residues (Fig. 1f). This separation of active site components is expected given that, with only one copy of Spo11, the structure shows the configuration of two catalytic half-sites (Fig. 1a). It is likely that the observed structure mimics the post-DSB product complex, except that Y135 would be covalently bound to the 5′ end in a true product complex.
Cryo-EM structure of the core complex bound to gapped DNA
We also solved the structure of the core complex bound to a double-hairpin substrate with a single-stranded DNA (ssDNA) gap (Fig. 2a and Extended Data Fig. 1c) at 3.3-Å resolution. The cryo-EM reconstruction is shown in Extended Data Fig. 3, cryo-EM statistics are provided in Table 1 and the density maps are presented in Extended Data Fig. 4a–e). Two views are shown in Fig. 2b,c. The preference for binding to bent DNA14 motivated these experiments, on the premise that the ssDNA region would provide the flexibility needed to assemble dimeric complexes that resemble the pre-DSB state. However, the majority of particles again had just a single core complex (Extended Data Fig. 3a).
We can trace the duplex of one hairpin plus the first nucleotide of the ssDNA (A39) including its 5′-phosphate. The bound hairpin is the one with a free 3′-OH and ssDNA extending from the 5′ end (Fig. 2a), which is structurally analogous to the single hairpin initially used. Weak density in the unsharpened map spanned the single-stranded nucleotides T38-T37-A36 and a portion of the second hairpin, with the two duplex segments aligned at a relative angle of 130° (Fig. 2d). Protein–DNA contacts are summarized in Fig. 2a.
The α-helical and β-strand segments for the four subunits are shown in Extended Data Fig. 5. Spo11 is a central hub making extensive protein–protein and protein–DNA contacts. The two arms of the V-shaped protein scaffold comprise the Spo11 WH domain interacting with Rec102 and Rec104 and the Toprim domain interacting with Ski8 (Fig. 2b,c). The structures with hairpin DNA and gapped DNA superpose well (root-mean-square deviation (r.m.s.d.) = 1.23 Å); hence, the following sections focus on the higher-resolution gapped-DNA structure.
Validation of the Spo11–Ski8 interface
The Spo11–Ski8 interface was previously modeled using a crystal structure of the Ski complex, in which two Ski8 copies contact two copies of a QRxxΦ motif in Ski3 that is also present in fungal Spo11 proteins14,21. This motif is critical for Spo11–Ski8 interaction in S. cerevisiae both in vivo and in vitro14,22. This same interface was also predicted computationally9. The cryo-EM structure matched these predictions, showing that Ski8 recognizes Q376REIFF in the Spo11 Toprim domain (Extended Data Fig. 6a). The interface involves both hydrophobic and hydrogen-bonding interactions (Extended Data Fig. 6b,c), including a computationally predicted extended interaction surface (Extended Data Fig. 6d).
Rec102 and its interaction with Spo11
The overall core complex architecture resembles Top6A–Top6B, as expected (Fig. 3a). Rec102 comprises a six-stranded β-sheet extended by multiple α-helices (Figs. 2b and 3b and Extended Data Fig. 5c). It has structural elements in common with the Top6B transducer domain, as anticipated, but with substantial differences. We consider here five main features from Top6B: a central β-sheet, a WKxY motif, a ‘switch loop’ that interacts with the ATP-binding site in the GHKL domain, the α-helical lever arm (‘stalk’) and the interface with Top6A (Fig. 3b, top). Conserved β-strands are indicated by uppercase letters ordered by tertiary-structure position, while numerical designations for β-strands correspond to primary-structure position (for example, Extended Data Fig. 5c); thus, these differ between species. Our structure agrees with crosslinking data14 where K60 and K64 in β-strand D crosslinked with K79 in nearby α-helix 1.
In Top6B, the central β-sheet is a scaffold supporting the stalk helix and the GHKL and H2TH domains4,5,11. Rec102 has a similar β-sheet, with the first four strands corresponding to segments of homology with Top6B (ref. 13) (Fig. 3b, strands labeled A–D, and Extended Data Fig. 7a,b). These strands are mostly encoded in REC102 exon 1, which is essential in vivo23. The sheet is extended by one (Top6B) or two (Rec102) additional β-strands but these arise differently in the sequence; the short strand in Top6B is immediately after strand D but the two longer strands in Rec102 instead occur after a conserved helix (Fig. 3b).
Another conserved element is a tryptophan within a motif that has the sequence WKxY in archaea13 and that starts at the end of a set of short α-helices and continues into the subsequent turn (Fig. 3b and Extended Data Fig. 7b). The lysine in this motif is part of a putative DNA-binding surface in Topo VI that is critical for activity24. The tryptophan is in the hydrophobic core of Saccharolobus Top6B, packing against the stalk and hydrophobic residues at the beginning of strand C (Extended Data Fig. 7c). The tryptophan is nearly invariant in eukaryotes but the rest of the motif is highly variable (W91EEQ in Rec102) (Extended Data Fig. 7b). In Rec102, W91 packs against the equivalent of the stalk helix, analogous to Top6B, but contacts different strands in the β-sheet: I59 in strand D (β4) and L113 in strand E (β6) (Extended Data Fig. 7c). This motif is ≥16 Å away from the DNA; thus, it does not appear to contribute to DNA binding in the end-bound complex.
The switch loop is a 16-residue sequence in Saccharolobus Top6B that connects strand C with the stalk helix (Fig. 3b and Extended Data Fig. 7b). It contains a lysine that contacts ATP bound by the GHKL domain’s active site and that is essential for ATPase activity in Topo VI and other topoisomerases25,26,27. Our structure confirms the earlier deduction of the position in the Rec102 sequence equivalent to the switch loop13 but the loop is smaller in Rec102 than previously appreciated (just eight residues, S157KEGNYVE). We tested the importance of this element by assessing the ability of rec102 mutant alleles to complement a rec102-null strain in a heteroallele recombination assay, in which arginine-positive prototrophs can be generated by recombination between two different arg4 mutant alleles (Extended Data Fig. 7d). Where tested, alanine substitutions across the switch loop had no effect on Rec102 activity in vivo and completely replacing the loop sequence with a flexible glycine or serine linker only reduced recombination about fourfold (Extended Data Fig. 7e). These results suggest that the specific amino acid sequence of the switch loop is not essential for Rec102 function, in keeping with the absence of a GHKL domain.
The stalk of Rec102 begins at residue I165, after the switch loop. As in Top6B, the first part of the stalk is an amphipathic helix whose hydrophobic face runs diagonally across strands D through A of the central β-sheet (Fig. 3b and Extended Data Fig. 7f). However, unlike the long, continuous α-helix that makes up the stalk of Top6B, the equivalent region of Rec102 is distorted into four helical segments to make a kinked path that is more compact and engages in more extensive intramolecular and intermolecular contacts within the core complex (Fig. 3b)9. We examined substitutions of four stalk residues predicted to contact Spo11 (I195A, L198A, R199A and W202A). Of these, R199A completely eliminated both meiotic recombination in vivo and yeast two-hybrid (Y2H) interaction with Spo11, without diminishing protein levels (green shading in Fig. 3c,d). R199 forms hydrogen bonds with multiple residues in Spo11 (Extended Data Fig. 8a). The other three substitutions reduced the Y2H interaction but did not disrupt recombination in vivo (Fig. 3c,d). Perhaps other interactions within the DSB-forming machinery can compensate when this interface with Spo11 is only partially weakened.
The Rec102 stalk winds its way to interact with the Spo11 WH domain. A helical hairpin at the end of the stalk aligns with a pair of α helices from Spo11 (Fig. 3b,e,f). The structure of this part of the Spo11–Rec102 interface resembles the computational Spo11–Rec102 model, which was earlier noted to resemble the Top6A–Top6B interface9. However, Spo11 and Rec102 have a much more extensive interface than Top6A and Top6B (1,681 Å2 for Spo11–Rec102 versus 958 Å2 for Topo VI; Extended Data Fig. 8b), mediated by both hydrophobic and hydrogen-bonding interactions (Fig. 3f and Extended Data Fig. 8a). Substitutions across the Rec102 helical hairpin (L214 and residues 220–236) were previously shown to disrupt the Y2H interaction with Spo11 and to eliminate meiotic recombination initiation in vivo14. Meiotic recombination was also compromised by single-alanine substitutions for interfacial residues L207 of Rec102 (yellow shading in Fig. 3c, left) and L60 and L112 in Spo11 (peach shading in Fig. 3c, right). Of these, Spo11-L112A was also deficient for the Y2H interaction with Rec102 despite having normal protein levels (Fig. 3d) but Rec102-L207A and Spo11-L60A retained the Y2H interaction, suggesting that their recombination defects are attributable to another biochemical property of the interface separate from stabilizing the interaction per se. Other tested residues in this interface were dispensable for meiotic recombination in vivo (Spo11 F103, which packs against Rec102 W202, and Spo11 L105) (Fig. 3c,d).
In addition to these elements that are conserved with Top6B, Rec102 sports a 34-residue C-terminal extension beginning at D231. This segment wraps around the stalk, contributes to the extended interface with Spo11 and also makes contacts with the DNA and Rec104 (Fig. 3e,g), as discussed below. Double-alanine substitutions for Rec102 D231;K232 or T235;T236 compromise both meiotic recombination and the Spo11 Y2H interaction14. Rec102 S233 and Q239 form hydrogen bonds with the Spo11 peptide backbone at G95 and L98, respectively (Extended Data Fig. 8a). Rec102-S233A, Rec102-Q239A and Spo11-G95A severely compromise meiotic recombination in vivo and diminish the Spo11–Rec102 Y2H interaction without affecting protein levels (purple shading in Fig. 3c, left, salmon shading in Fig. 3c, right and Fig. 3d).
There is also an extension (25 residues) of the N terminus of Spo11 before the WH domain, of unknown function16. This α-helical segment contacts Rec102’s C-terminal extension and stalk and the N terminus of Rec104 through hydrophobic interactions and hydrogen bonds (Fig. 3e,h). The Spo11 extension places the N termini of Spo11, Rec102 and Rec104 in close proximity, which agrees with crosslinking data and explains why an N-terminal maltose-binding protein tag on any of these proteins resulted in extra density in nearly the same position in negative-stain EM14. Several substitutions in the Spo11 extension (L3A, R7D and L20A) severely compromised meiotic recombination in vivo and diminished Y2H interactions with Rec102 or Rec104 (blue shading in Fig. 3c, right and Extended Data Fig. 8c). Most of the other tested mutants in this region also decreased the Y2H interactions without affecting recombination (Fig. 3c and Extended Data Fig. 8c).
Rec104 structure and protein interactions
Only residues 9–58 (of 182) of Rec104 are visible in the structure, forming three α-helices that interact with both Rec102 and Spo11 (Fig. 4a–c). The structure agrees with previously observed14 crosslinks between Rec104 K43 and Rec104 K10, K36 and K39 (α-carbon-to-α-carbon distances of 12–20 Å). However, the resolution is lower for this part of the structure (Extended Data Fig. 2d and Extended Data Fig. 3d); thus, the details of interfacial contacts should be viewed cautiously.
Previous efforts to identify Rec104 orthologs failed to find them outside of Saccharomyces species28. The structure of the core complex allowed us to revisit this because the visible parts of Rec104 correspond to a previously unrecognized conserved domain (Fig. 4d). We expanded the collection of Rec104 homologs by focusing on this domain but we were still only able to detect them in family Saccharomycetaceae (Fig. 4d and Methods). Particularly well conserved residues (Fig. 4d) line the interfaces with Rec102 and Spo11, including Rec104 Y21 (Fig. 4a), Rec104 F17, L18, V23, V27, F31 and L33 (Fig. 4b) and Rec104 I41 and F46 (Fig. 4c). The C-terminal portion of Rec104 that is not visible in the cryo-EM structure is poorly conserved (Fig. 4d).
We tested the functional importance of interfacial residues. A Rec104-Y21A mutant was strongly defective for meiotic recombination (Extended Data Fig. 8d) and the Y2H interaction with Spo11 (Extended Data Fig. 8c) but was expressed at normal levels and retained the Y2H interaction with Rec102 (Extended Data Fig. 8e). There was little if any effect for the other Rec104 mutants tested (Extended Data Fig. 8d,e) or for Rec102 β-sheet substitutions L8A, V10A, F11A and F156A (pink shading in Fig. 3c, left and Extended Data Fig. 8e). Interestingly, Rec102 stalk substitutions L179A and Q183A disrupted the Y2H interaction with Rec104 but had only a modest effect (L179A) or no effect (Q183A) in recombination (green shading in Fig. 3c and Extended Data Fig. 8e). Because this Y2H assay is performed in vegetative cells, a plausible interpretation is that these substitutions weaken the Rec102–Rec104 interaction but are compensated for in the context of the full DSB apparatus.
Prior crosslinking experiments suggested that Rec104 lies near where the GHKL domain abuts the transducer domain in Topo VI, supporting the proposal that Rec104 replaces the GHKL domain1,14. We explored this idea using our structure.
The distances between Rec104 K43 and either Rec102 K60 or K64 agree well with crosslinking data (Extended Data Fig. 8f). Rec102 K60 and K64 are in strand D of the central β-sheet, with their side chains on the opposite face of the sheet from the stalk (Extended Data Fig. 7b and Extended Data Fig. 8f). The equivalent region in Top6B is at the interface with the GHKL domain, as previously noted14. However, when we superimposed the core complex on a crystal structure of Top6B (ref. 4), the structured elements of Rec104 were strikingly distinct from the GHKL or HT2H domains of Top6B (Fig. 4e). Although we do not know the disposition of the C-terminal two thirds of Rec104, the end of helix 3 points toward the location of the Top6B GHKL domain (Fig. 4e). Moreover, Rec104 K61, K65, K78 and K79 (which lie just outside the structured segment) crosslink to Rec102 K60, K64 and K79 (ref. 14), all three of which are near the position occupied by the GHKL domain in Top6B.
We conclude that the structured part of Rec104 is not equivalent to any of the domains in Top6B but that the C-terminal part of Rec104 is near or in the GHKL domain position, consistent with the previous proposal about Rec104’s location14. However, it remains unclear whether the unstructured portion of Rec104 evolved from the GHKL domain or is a wholly unrelated fold that replaced it.
Protein–DNA contacts and the specificity of DNA binding
An electrostatic surface representation of the core complex identified three positively charged patches that line the DNA-binding channel and mediate non-sequence-specific protein–DNA contacts (Fig. 5a). Patch 1 anchors overhang nucleotide A39 and its adjacent duplex in the pocket between the Spo11 WH and Toprim domains. R131 on the second α-helix of the WH domain forms a hydrogen bond with the 5′-phosphate of G40, which is the bridging phosphate between the first base of the duplex (G40) and the beginning of the 5′ overhang (A39) (Fig. 5b). This interaction stabilizes the orientation relative to the DNA of the catalytic Y135, which is on the same α-helix (Fig. 5b). In addition, the third α-helix of the WH domain inserts into the major groove, with Q144 interacting with the same phosphate as R131 (Fig. 5b) and R143 interacting with the phosphate backbone of the opposite strand (Fig. 5c). Furthermore, residue K173 at the beginning of the flexible linker connecting the WH and Toprim domains makes one of the rare contacts of Spo11 with a base, forming a hydrogen bond with the N-3 position of G40 (Fig. 5b). This contact is interesting because G is favored at this position in vivo29. Toprim domain residues in patch 1 include R266, which forms a hydrogen bond with the phosphate of A42, and E233, which forms a hydrogen bond with the 3′-terminal hydroxyl group of C71 (Fig. 5b).
These DNA interactions explain protein sequence conservation and prior experimental findings. Specifically, R131, Q144, R266 and E233 are invariant in Spo11 and Top6A and K173 is a basic residue in nearly all eukaryotes (Extended Data Fig. 9a). Moreover, the contacts of R131 and Q144 with the 5′-phosphate at the beginning of the duplex and of E233 with the 3′-hydroxyl explain the strong selectivity for a free 3′-OH end and 5′ ssDNA overhang14. R131 and E233 are essential for Spo11 function in vivo7; substituting K173 to alanine reduced DNA binding in vitro and delayed and reduced DSB formation in vivo14, while substituting E233 to alanine reduced DNA binding in vitro ~10-fold14.
Patch 2 is in the WH domain (Figs. 2a and 5a). The basic K74KKK loop surrounds the T63-T64-A65-containing strand, while K104 hydrogen bonds with the phosphate of C66 (Figs. 2a and 5c). The density is weak for the K74KKK loop, with K74 and K76 directed toward the DNA backbone. This loop is part of a β-hairpin (the ‘wing’ in the WH domain) and is cleaved by hydroxyl radicals produced by iron-chelating moieties placed nearby on the DNA14 but is highly variable in length and sequence in Spo11 proteins (Extended Data Fig. 9a).
Patch 3 is on the outer segment of the DNA-binding channel (Figs. 2a and 5a). It includes Rec102 R245 and Rec104 R26, which form hydrogen bonds with the phosphates of C50 and G58, respectively (Fig. 5d).
We also identified two sets of residues (‘fingers’) projecting from Spo11 into the minor groove. Finger 1 (F260, Y292 and R344 from the Toprim domain) tracks along the first five base pairs and finger 2 (H101 from the WH domain) forms a main chain amide hydrogen bond with the phosphate of T49 and inserts its imidazole ring into the minor groove (Fig. 5e). F260 is a bulky hydrophobic residue in Spo11 orthologs but glutamine in most Top6A proteins (Extended Data Fig. 9a). The minor groove binding by finger 1 is particularly striking because the base composition bias around Spo11 cleavage sites in vivo suggested a tendency toward a relatively wide and shallow minor groove across precisely this region29. An F260A substitution dramatically changes DSB site preference in vivo and reduces DNA-binding affinity in vitro14, while substituting Y292 to arginine disrupts DSB formation in vivo7.
Metal-ion binding
Type IIA topoisomerases (eukaryotic Topo II and bacterial gyrase and Topo IV) are thought to use a two-metal mechanism involving divalent cations bound by acidic residues of the Toprim domain30,31,32. In this model, metal ion A interacts with both bridging and nonbridging oxygens of the scissile phosphate and has a direct role in catalysis, while metal ion B interacts with an adjacent (nonscissile) phosphate and has a structural role in stabilizing protein–DNA interactions (Fig. 6a). Comparatively little is known in type IIB enzymes3,33.
We observed density in both cryo-EM structures consistent with a single metal ion, most likely Mg2+, bound by D288 and D290 of Spo11 (Fig. 6b). When the Spo11 metal-binding pocket was compared with that of Methanocaldococcus jannaschii Top6A (ref. 6), the trio of conserved acidic residues was highly congruent but the single Mg2+ bound by each protein was in a different position, bound by E197 and D249 in Top6A (equivalent to Spo11 E233 and D288) (Fig. 6b). By comparison to yeast Top2 (ref. 30), we infer that site A is occupied in the Top6A structure, whereas site B is occupied in our Spo11 structure (Fig. 6a,b). In type IIA enzymes, site A has a higher affinity for metal than site B but is thought to rely on the scissile phosphate for two of the coordination contacts32. If the same is true for Spo11, the absence of an equivalent of the scissile phosphate at the 3′ end in our structure may explain why Mg2+ is not stably bound at site A. The observed binding in site B is consistent with the presumed structural role for the metal in this position. Occupancy of site B is consistently observed in postcleavage structures of eukaryotic and bacterial topoisomerases30,34, supporting the conclusion that our structure mimics the postcleavage state.
These findings provide a framework for understanding the conservation and functional importance of the metal-binding acidic residues in Spo11. E233 and D288 are invariant in Spo11 and Top6A proteins (Extended Data Fig. 9a) and are essential for DSB formation in vivo in yeast7. Extrapolating from type IIA enzymes, we propose that the essentiality of both residues traces to them directly coordinating the catalysis-critical Mg2+ in site A (Fig. 6a,b). By contrast, D290 is mostly but not strictly conserved (Extended Data Fig. 9a) and it is dispensable for DSB formation in vivo, whereby the substitution with asparagine has little effect on recombination activity in S. cerevisiae while the alanine substitution partially decreases DSB formation7. It may be that this residue is not essential because it contributes to metal binding only in the noncatalytic site B (Fig. 6a,b).
Modeling higher-order complexes
Spacing between adjacent DSBs
Our cryo-EM structure sheds light on spatial patterns of Spo11–DNA interactions both in vivo and in vitro. Multiple Spo11 complexes can sometimes introduce two or more DSBs close together on the same DNA molecule35,36,37. When such double-cutting occurs, it has a preferred spacing between DSBs with a minimum of ~33 bp (measured from the center of each DSB’s 5′ overhang, corresponding to 31 bp of duplex DNA plus the overhangs) and increasing in steps of ~10 bp. The 10-bp periodicity has been proposed to reflect a geometric constraint, in which adjacent Spo11 dimers are co-oriented with their active sites facing in the same direction35 (Extended Data Fig. 9b). In this model, the observed minimum distance reflects steric constraints that prevent adjacent co-oriented Spo11 complexes from being fewer than three helical turns apart.
Electrophoretic mobility shift assay (EMSA) experiments with yeast Spo11 core complexes support this interpretation; both 5′ overhang ends of a DNA substrate are readily bound if the duplex DNA segment between the overhangs is ≥28 bp but duplex lengths of 24–27 bp can only be bound at a single end35 (Fig. 6c). Double-end binding by monomeric core complexes in vitro likely mimics how close two adjacent Spo11 dimers can be on an intact DNA molecule before cleavage (Fig. 6d, top). Interestingly, reducing the duplex length even further (to 22–23 bp) allows core complexes to again bind at both ends35 (Fig. 6c). These results suggest that steric clashes that preclude double-end binding at 24–27 bp are relieved if the two DNA ends are rotated relative to one another. Different orientations of adjacent Spo11 complexes are possible with free proteins in vitro but not with the constrained complexes presumed to be present in vivo (Extended Data Fig. 9b).
These patterns of DNA cleavage in vivo and DNA binding in vitro are well explained by the extensive left-handed wrap of the core complex around the DNA in the cryo-EM structure. Two core complexes could be modeled on each end of a 28-bp DNA duplex with essentially no steric clashes but modeling on a shorter DNA segment (24 bp) resulted in substantial overlap between the Rec102 and Rec104 moieties of the two core complexes (Fig. 6c,d). Consistent with EMSA data, the clashes could be almost entirely resolved by shortening the DNA still further to 22 bp, which rotates the core complexes relative to one another and allows them to interdigitate (Fig. 6c,d). Moreover, a 31-bp duplex is the shortest distance that can accommodate a pair of core complexes that is both co-oriented and lacking in steric clashes (Fig. 6d). This agrees well with in vivo spacing because a 31-bp duplex corresponds to a center-to-center DSB distance of 33 bp (that is, the minimum preferred distance for double cuts)35,37.
Model of a pre-DSB Spo11 dimer
We attempted to model a catalytically competent Spo11 dimer by docking two copies of the monomeric DNA-bound core complex together. If the two DNA copies are aligned to make a roughly B-form duplex, the WH domain from each Spo11 clashes sterically with the Toprim domain from the other (Fig. 7a). In addition, the nonoverhang strand from one complex clashes with the C-terminal part of α-helix 8 of Spo11 from the other complex and the overhang strand collides with a loop near finger 1 (Fig. 7b). Thus, the configurations of both protein and DNA in the cryo-EM structure are incompatible with a plausible structure of a pre-DSB complex on B-form DNA.
Because Spo11 core complexes are flexible in solution14, we asked whether simple rigid-body motions of their domains could resolve the clashes. To answer this, we separately aligned the Spo11 WH domain (together with DNA, Rec102 and Rec104) and the Toprim domain (together with Ski8) to the cognate domains of Top6A in the Saccharolobus shibatae Topo VI dimer structure4 (Extended Data Fig. 9c) because the segment between the domains is thought to be a flexible linker6,14. This alignment rotates the WH domain 45° relative to the Toprim domain and, notably, eliminates all of the steric clashes (Fig. 7c). The model also matches segments of Spo11 well with the two major Top6A–Top6A interfaces in Topo VI dimer structures4,5,6: a pseudocontinuous β-sheet on the underside of the Toprim domain away from the DNA (K206 to P211 from one Spo11 interacting with its match on the other Spo11) and an interface between the WH domain of one Spo11 near the catalytic tyrosine and a C-terminal region of the other Spo11 that includes an invariant residue E386 that is brought in close proximity to the catalytic tyrosine (Fig. 7d and Extended Data Fig. 9a). This model is, thus, a plausible representation of a dimeric pre-DSB complex. We further speculate that DSB formation is accompanied by a Spo11 conformation change into a postcleavage state resembling our cryo-EM structures and that the steric clashes described above may prevent DNA end religation, rendering DSB formation irreversible.
The model predicts an intriguing deformation of the DNA. We preserved the relationship of the DNA to the WH domain because this also preserves the large majority of the protein–DNA contacts from the cryo-EM structure. In doing so, the overhang ends of the two DNA segments come into close proximity (~4.8 Å separating the inferred positions of 5′ and 3′ ends of each strand) but make a V shape with a 150° angle and with the helical axes slightly offset by ~5 Å (Fig. 7e). Empirical evidence for DNA deformation by Spo11 comes from the 130° angle of the DNA in our gapped double-hairpin structure (Fig. 2d) and by previous DNA-binding experiments that revealed bent DNA by atomic force microscopy and apparent preferential binding to bent duplex DNA (inferred from higher affinity for binding 100-bp versus 400-bp minicircles)14. Topo VI and other type II topoisomerases bend DNA before cleavage6,14,38; hence, it is likely that Spo11 core complexes may do so as well. Our model, thus, provides a framework for understanding the structural determinants of DNA bending, for which no high-resolution structural information is currently available for any dimeric Topo VI or Spo11 complex.
Conclusions
Ever since the discovery of the DSB-forming activity of Spo11 more than a quarter-century ago39,40, there has been a lack of empirical structures of Spo11 and its accessory proteins. Our structures provide insight into the architectures of the individual subunits and of their interfaces with each other and DNA. We confirm the structure and position of Ski8, uncover unexpected differences between Rec102 and archaeal Top6B proteins and partially resolve outstanding questions about the structure and spatial disposition of Rec104. Our findings further explain the high affinity and exquisite selectivity of core complex binding to DNA ends that have a recessed 3′-OH. Lastly, the structures provide a molecular framework to understand nonrandom DSB patterns in vivo, including the biased base composition around cleavage sites and the spacing of double cuts.
It remains unclear why yeast core complexes are thus far not competent to cleave DNA in vitro. Perhaps the high affinity for DNA ends precludes dimer assembly on DNA when ends are available. A related issue is that the high-affinity DNA-binding state we observe has the proteins in a conformation that is not compatible with dimer formation. The ability of Spo11 accessory proteins Rec114, Mei4 and Mer2 to recruit Spo11 core complexes and present them in a coherent orientation (Extended Data Fig. 9b) may facilitate formation of DSB-competent dimers41,42,43,44. Inclusion of these other proteins may, thus, be needed for DNA cleavage in vitro. Alternatively, it may be possible to overcome the barrier to dimer formation by using DNA substrates that disfavor the monomer configuration. In any case, our results point to the Spo11 monomer–dimer dynamic as being key to controlling the DSB-forming activity.
Methods
Purification of S. cerevisiae Spo11 core complexes
The expression plasmids and baculoviruses were prepared as described previously14. The Spo11 core complex was expressed by coinfecting Gibco Spodoptera frugiperda Sf9 cells (Thermo Fisher 11496015) with a combination of viruses at a multiplicity of infection of 2.5 each as described previously14. Typically, 500 ml of suspension culture was collected 62 h after infection. The cells were lysed by sonication in 25 mM HEPES-NaOH pH 7.4, 500 mM NaCl, 0.1 mM DTT, 20 mM imidazole, 1× cOmplete protease inhibitor tablet (Roche) and 0.1 mM PMSF and centrifuged at 43,000g for 30 min. Cleared extract was loaded onto 1 ml of Ni-NTA resin (Qiagen) pre-equilibrated with nickel buffer (25 mM HEPES-NaOH pH 7.4, 500 mM NaCl, 10% glycerol, 0.1 mM DTT, 20 mM imidazole and 0.1 mM PMSF) and then washed extensively with nickel buffer. Core complexes were eluted in nickel buffer containing 250 mM imidazole and then further purified on anti-Flag M2 affinity resin (Sigma). To do so, fractions from the Ni-NTA elution containing protein were pooled and diluted in three volumes of Flag buffer (25 mM HEPES-NaOH pH 7.4, 500 mM NaCl, 10% glycerol and 1 mM EDTA) before binding to the anti-Flag resin. Core complexes were eluted from anti-Flag resin with Flag buffer containing 250 μg ml−1 3xFlag peptide (Sigma). Fractions containing protein were pooled and loaded on a Superdex 200 Increase 10/300 GL column (Cytiva) pre-equilibrated in 25 mM HEPES-NaOH pH 7.4, 300 mM NaCl, 5 mM EDTA and 2 mM DTT. Fractions containing protein were concentrated in 50-kDa-cutoff Amicon centrifugal filters (Millipore). Aliquots were frozen in liquid nitrogen and stored at −80 °C.
Cryo-EM structure determination
Purified core complexes were diluted to a final concentration of ~0.2 mg ml−1 and incubated with the hairpin (5′-TAGCAATGTAATCGTCTATGACGTTAACGTCATAGACGATTACATTGC) or gapped DNA (5′-TAGGCCGTCGGCTACTAAAAGTAGCCGACGGCCGGATTAGCAATGTAATCGTCTTAAGACGATTACATTGC) substrates at a molar ratio of 1:2 (protein:DNA) in binding buffer (25 mM HEPES-NaOH pH 7.5, 150 mM NaCl, 5 mM MgCl2, 2 mM DTT and 2% glycerol) for 1 h on ice. An aliquot of the incubated mixture (3.5 μl) was applied onto glow-discharged UltrAuFoil 300 mesh R 1.2/1.3 grids (Quantifoil) at ~4 °C. Grids were blotted for 2 s at 100% humidity at 4 °C and flash-frozen in liquid ethane using a Vitrobot Mark IV (Thermo Fisher Scientific). The twofold molar excess of DNA was expected to favor singly-bound DNA molecules but would allow recovery of doubly-bound molecules if the affinity for binding by the second core complex were comparable to binding by the first. Because binding to the hairpin end is much lower affinity than binding to the end with the 5′ overhang, only singly-bound complexes were expected for the hairpin substrate. However, we did not know a priori whether a dimer of core complexes might be able to bind to the gapped substrate.
Images were collected on a Titan Krios G2 (FEI) transmission EM instrument operating at 300 kV with a K3 direct detector (Gatan) using a 1.064-Å pixel size at the Memorial Sloan Kettering (MSK) Cancer Center. The defocus range was set from −1.0 to −2.5 μm. Videos were recorded in super-resolution mode at an electron dose rate of 20 e− per pixel per s with a total exposure time of 3 s and intermediate frames were recorded every 0.075 s for an accumulated electron dose of 53.00 e− per Å2.
The protocol for cryo-EM reconstruction is presented in Extended Data Figs. 2a–d and 3a–d. The super-resolution movies (0.532 Å per pixel) were motion-corrected and Fourier-cropped twice using MotionCor2 (ref. 47). Contrast transfer function parameters were estimated by CTFFIND-4 (ref. 48). All other steps of image processing were performed by RELION 3.0 (ref. 49) and Cryosparc version 3.3.0 (ref. 50).
For hairpin-bound core complexes, after blob picking from 2,338 images without reference, a total of 1,829,931 particles extracted after three-pixel binning were applied to two rounds of two-dimensional (2D) classification. All particles were applied to multiple rounds of heterogeneous refinement by using the 4.8-Å-resolution model generated from RELION 3.0. The final 128,787 particles were polished and yielded a reconstruction EM map with an average resolution of 3.66 Å. All reported map resolutions are from gold-standard refinement procedures with the Fourier shell correlation (FSC) cutoff being 0.143 (Extended Data Fig. 2).
For gapped-DNA-bound core complexes, after blob picking from 5,521 images without reference, a total of 7,804,373 particles extracted after four-pixel binning were applied to the first round of 2D classification. A total of 6,930,069 particles selected from 2D classification were applied to multiple rounds of three-dimensional (3D) classification. One of the 3D classes with good secondary-structure features and the corresponding 1,314,980 particles were re-extracted without binning. The selected particles were imported to new initial models and hetero refinement and 548,674 particles were selected from the best class for homogeneous refinement. After nonuniform refinement and local refinement, we obtained an EM map with an average resolution of 3.29 Å. All reported map resolutions are from gold-standard refinement procedures with the FSC cutoff being 0.143 (Extended Data Fig. 3).
Spo11, Ski8 and Rec102 models were downloaded from the AlphaFold2 database10 and fitted into the cryo-EM density map as rigid bodies. The DNA model was generated using Coot51. We also generated a model of the Rec102–Rec104 subcomplex using ColabFold52 and fitted this model into the cryo-EM density map as well. The side chains of the visible N-terminal portion of Rec104 could be traced in the density map of the hairpin-bound core complex. Atomic coordinates were refined against the map by real-space refinement in PHENIX53 by applying geometric and secondary-structure restraints. Details of data collection, image processing and model building are listed in Table 1 and shown in Extended Data Figs. 2 and 3. All figures were prepared using PyMol (https://pymol.org/2/), UCSF Chimera54 or UCSF ChimeraX55.
DNA substrates and EMSAs
EMSAs used the same DNA substrates as in the cryo-EM structures. Both the hairpin and the gapped substrates were assembled by self-annealing in annealing buffer (100 mM NaCl, 10 mM Tris-HCl pH 8 and 1 mM EDTA). Substrates were 5′-end-labeled with [γ-32P]ATP (Perkin Elmer) and T4 polynucleotide kinase (New England Biolabs) and purified by native PAGE. Binding reactions (10 µl) were carried out in 25 mM Tris-HCl pH 7.5, 7.5% glycerol, 100 mM NaCl, 2 mM DTT, 5 mM MgCl2 and 1 mg ml−1 BSA with 0.1 nM DNA. Complexes were assembled for 30 min at 30 °C and separated on 6% polyacrylamide DNA retardation gels (Invitrogen). Gels were dried and analyzed by phosphor imaging (Fuji).
Modeling higher-order assemblies of Spo11 core complexes
All modeling was performed in ChimeraX56. The Spo11 double-end binding complex was modeled by aligning the DNA from a Spo11 core complex structure to each end of a 34-bp B-form DNA. To generate models with different lengths of DNA, the core complex at one end was fixed, while the other Spo11 core complex was moved along the B-form DNA path by the specified number of base pairs.
To model a potential pre-DSB assembly containing two core complexes bound to B-form DNA, the individual DNA sequences in two copies of the cryo-EM structure were each aligned to a B-form DNA segment such that the first nucleotide in the 5′ overhang from one DNA sequence was one nucleotide away from the 3′ end of the other DNA sequence.
To model a pre-DSB dimer based on a Topo VI dimer, the cryo-EM structure was separated into two parts; one included residues 1–172 of Spo11 plus Rec102 and Rec104, while the other included the remaining C-terminal part of Spo11 plus Ski8. The two parts were aligned separately to the cognate domains of the S. shibatae Top6A subunit from a crystal structure of the Topo VI holoenzyme (Protein Data Bank (PDB) 2ZBK)4. The alignment of N-terminal part of Spo11 resulted in an r.m.s.d. of ~1.2 Å over 32 atom pairs; alignment of the C-terminal part of Spo11 resulted in an r.m.s.d. of ~1.1 Å over 70 atom pairs. The session files are available from Mendeley (https://doi.org/10.17632/dx3hx827fp.1).
Yeast strains and targeting vectors
All yeast strains were from the SK1 background (Supplementary Table 1). All vectors were generated on the basis of pSK275, pSK276, pSK282, pSK293, pSK305 and pSK310 for both yeast two-hybrid assays and heteroallele recombination assays (Supplementary Table 2).
Yeast two-hybrid assays
Y2H 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. Cells were lysed and a quantitative β-galactosidase assay was performed using the yeast β-galactosidase assay kit following the manufacturer’s protocols (Thermo Scientific). For Y2H experiments in meiotic conditions, after overnight culture in selective medium with glucose, cultures were washed twice then incubated with 2% potassium acetate for 20 h to induce meiosis.
Y2H assays do not necessarily reflect only those interactions that are direct, as the interactions may be indirect or supported by other proteins. Interactions of Spo11 with either Rec102 (Fig. 3d) or Rec104 (Extended Data Fig. 8c) were measured in meiotic conditions because these interactions require the presence of other members of the core complex (for example, interaction of Spo11 with Rec102 requires Rec104 to also be present)23. Interaction of Rec102 with Rec104 (Extended Data Fig. 8e) was measured in vegetatively growing reporter cells.
For immunoblotting, yeast cells expressing the Y2H protein variants were cultured overnight in selective medium. Equivalent cell amounts based on optical density measurements at 600 nm were harvested and treated with 0.2 M NaOH and then proteins were solubilized in 100 µl of 2× Laemmli sample buffer (Bio-Rad). A 10-µl aliquot of each sample was run on a 4–12% Bis-Tris PAGE gel (Invitrogen) and transferred to a 0.2-µm PVDF membrane (Thermo Scientific). The membranes were blocked using StartingBlock Blocking Buffer (Thermo Scientific) for 20 min at room temperature and probed with primary antibodies to LexA (rabbit, 1:2,000, Sigma-Aldrich), hemagluttinin (HA; rat, horseradish peroxidase (HRP)-conjugated, 1:1,000, Roche) and α-tubulin (rat, 1:2,000, Invitrogen, for Rec104 blots; rat, HRP-conjugated, 1:2,000, Santa Cruz, for Spo11 and Rec102 blots). Following three 10-min washes with PBS-T at room temperature, the membranes were developed using an enhanced chemiluminescence western blotting substrate (Thermo Scientific) and imaged with a ChemiDoc imaging system for HRP-conjugated primary antibodies. Alternatively, membranes were incubated for 1 h at room temperature with fluorophore-conjugated species-specific secondary antibodies (donkey antirabbit IgG (Alexa Fluor 488, 1:5,000, Invitrogen) or donkey antirat IgG (Alexa Fluor 594, 1:5,000, Invitrogen)); then, the membranes were washed three times with PBS-T for 10 min each and imaged using a ChemiDoc imaging system.
Heteroallele recombination assays
Heteroallele recombination assays were performed in diploid strains heterozygous for the arg4-Bgl and arg4-Nsp alleles57 along with homozygous deletion mutations for spo11, rec102 or rec104. Complementation test plasmids expressing wild-type or mutant versions of Spo11, Rec102 or Rec104 (Supplementary Table 2) were introduced by lithium acetate transformation. The Y2H fusion constructs were used for this purpose. Three independent colonies were cultured overnight on synthetic complete medium lacking tryptophan and leucine to select for the test plasmid and then transferred to YPA (1% yeast extract, 2% peptone and 1% potassium acetate) for 13.5 h before meiotic induction in 2% potassium acetate. After at least 3 h in meiosis, appropriate dilutions of cultures were plated on synthetic complete medium lacking arginine to measure the frequency of arginine-positive recombinants and on YPD (1% yeast extract, 2% peptone and 2% dextrose) to measure colony-forming units.
Phylogenetic analyses
Protein-coding sequences of Rec104 from different fungal species were identified using the National Center for Biotechnology Information (NCBI) basic local alignment search tool (BLAST). Conserved regions were identified using a subset of 12 species and visualized using COBALT45. For the analysis of structural conservation in Rec102 and Top6BL homologs, AlphaFold2 structure predictions were downloaded from the AlphaFold2 database58. A BLAST search for a Rec102 homolog in Sordaria macrospora identified a predicted structure (AF-F7VPQ4-F1, NCBI XP_003350816.1) that lacks one of the conserved β-strands and the characteristic WKxY motif. Inspection of the genomic locus suggests that a splice site is misannotated; correcting this results in a longer second exon, adding the missing sequence. We used the new sequence to model the structure using ColabFold52.
Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.
Data availability
The atomic coordinates and cryo-EM density maps for the core complex bound to hairpin DNA (PDB 8URU and EMD-42501) or gapped DNA (PDB 8URQ and EMD-42497) were deposited to the Research Collaboratory for Structural Bioinformatics PDB and EM Data Bank, respectively. We used the following published atomic coordinate accessions: PDB 2ZBK, PDB 2Q2E, PDB 1D3Y, AF-Q02721-F1, AF-P33323-F1 and AF-P23179-F1. Source data are provided with this paper.
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Acknowledgements
This article is subject to the Open Access to Publications policy of the Howard Hughes Medical Institute (HHMI). HHMI lab heads previously granted a nonexclusive CC BY 4.0 license to the public and a sublicensable license to HHMI in their research articles. Pursuant to those licenses, the author-accepted manuscript of this article can be made freely available under a CC BY 4.0 license immediately upon publication. We thank members of the Keeney and Patel laboratories for discussions and experimental advice and J. De La Cruz (MSK) for support in cryo-EM data collection. MSK core facilities are supported by National Cancer Institute Cancer Center support grant P30 CA08748. K.L. was supported in part by the Damon Runyon Cancer Research Foundation (DRG-[2389-20]). M.A. was supported in part by an EMBO long-term fellowship (ALTF 905-2019). This work was supported by National Institutes of Health (NIH) grant R01 HD110120 (to S.K. and D.J.P.), an MSK Basic Research Innovation Award (BRIA; to S.K. and D.J.P.), a postdoctoral BRIA (to M.A.), a Leukemia and Lymphoma SCOR 7021020 grant (to D.J.P.), the Maloris Foundation (to D.J.P.) and NIH grant R35 GM118092 (to S.K.). S.K. is an HHMI investigator. The funders had no role in study design, data collection and analysis, decision to publish or preparation of the manuscript.
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C.C.B., S.K. and D.J.P. conceptualized the project. Y.Y., J.W., K.L., Z.Z., M.A., C.C.B., S.K. and D.J.P. designed the experiments. Y.Y., J.W., K.L., Z.Z., M.A., C.C.B. and S.P. performed the experiments. Y.Y., J.W., K.L., Z.Z., M.A., C.C.B., S.K. and D.J.P. analyzed the data. S.K. and D.J.P. supervised the research. K.L., M.A., S.K. and D.J.P. secured funding. Y.Y., J.W., K.L., S.K. and D.J.P. wrote the paper with input from Z.Z. and M.A. All authors edited the manuscript.
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Extended data
Extended Data Fig. 1 Spo11 core complex purification and EMSA assays.
a, SDS-PAGE gel of purified core complex ( ~ 1 µg), stained with Coomassie. The image is representative of results from > 6 independent preparations. b,c, EMSA assays and quantification of core complex binding to the hairpin DNA substrate (b) and the gapped DNA substrate (c). In panel c, the position of the second core complex that gives the slowest migrating band is unknown; it could be at the nick location as shown or it could associate with one of the hairpin ends. Error bars indicate mean ± SD of three replicates.
Extended Data Fig. 2 Cryo-EM reconstruction of the core complex bound to hairpin DNA.
a, Flow chart of cryo-EM image processing. b, Global Fourier shell correlation (FSC) curves. The overall cryo-EM map resolution is 3.7 Å with FSC set at 0.143. c,d, Euler angle distribution (panel c) and final 3D reconstructed map colored according to local resolution (panel d). e, Density map.
Extended Data Fig. 3 Cryo-EM reconstruction of the core complex bound to gapped DNA.
a, Flow chart of cryo-EM image processing. b, Global FSC curves. The overall cryo-EM map resolution is 3.3 Å with FSC set at 0.143. c,d Euler angle distribution (panel c) and final 3D reconstructed map colored according to local resolution (panel d).
Extended Data Fig. 4 Examples of protein side chain and DNA base-sugar-phosphate identification in the cryo-EM structure of the core complex bound to gapped DNA.
a–c, Examples of fitting of protein amino acid side chains into the density map for interacting helices of Rec102 and Rec104 (panel a), a segment of β sheet from Ski8 (panel b), and a segment of α helix from Spo11 (panel c). d, Example of fitting DNA base-sugar-phosphate and interacting protein side chains into the density map for Spo11 interaction with the DNA. e, Example of fitting DNA base-sugar-phosphate into the density map, highlighting the 3′-OH and first nucleotide of the 5′ overhang.
Extended Data Fig. 5 Protein secondary structures.
a–d, The secondary structures for Spo11 (panel a), Ski8 (panel b), Rec102 (panel c), and Rec104 (panel d) were determined from the cryo-EM structure of the core complex bound to gapped DNA. Gray-colored regions are not visible in the cryo-EM structure. Red letters in parentheses in panel c indicate the conserved strand designations shown in Fig. 3b.
Extended Data Fig. 6 Interactions between Ski8 and Spo11.
All images are from the cryo-EM structure of the core complex bound to gapped DNA. a, QREIFF motif from Spo11 (magenta) and the WD40 repeats from Ski8 (cyan). Ski8 in the cryo-EM structures adopts a propeller folding topology formed by seven WD40 repeats and recognizes the sequence Q376REIFF in the Spo11 Toprim domain. b, Hydrophobic interactions between Ski8 WD40 repeats and Spo11. The hydrophobic core of the interface is formed by Spo11 residues F380 and F381 and surrounding Ski8 aromatic residues involved in CH-π and π-π stacking interactions. c, Hydrogen bond interactions between Ski8 and Spo11. A hydrogen bonding network surrounds the hydrophobic core illustrated in panel b, including bonds between Q376 and R377 of Spo11 and Ski8 residues S63 and D18, respectively. d, Additional hydrogen bonding interactions between Spo11 and the extended loop in Ski8. Computational prediction had suggested that an extended loop in Ski8 expands the interface with Spo119. In agreement, we observed additional hydrogen bonding interactions that further stabilize the interface.
Extended Data Fig. 7 Sequence and structural conservation of Rec102 and Top6B.
a, Top6B and Rec102/Top6BL architectures. Strands, helices, and switch loops are colored as in Fig. 3b. All structures show truncated proteins, starting at the first β strand (strand A) and ending with the switch loop before the start of the stalk. The switch loop is truncated for the N. crassa and S. macrospora proteins for clarity. b, Secondary structure and sequence information for models shown in a. The conserved WKxY motif is highlighted in red and β strands are labeled as in a. c, Packing for S. shibatae Top6B (PDB: 2zbk) in comparison to Rec102 for the W in the WKxY motif. Ribbons and sticks are colored as in Fig. 3b. d, Cartoon illustrating the heteroallele recombination assay. In a strain heterozygous for two different mutant versions of arg4 (gray arrows with mutations marked with red x’s), meiotic recombination can generate a functional copy of ARG4 (green arrow) by gene conversion and/or crossing over. Below are representative images of Arg+ prototrophic colonies from a rec102∆ strain carrying complementing plasmids expressing the indicated version of Rec102. e, Effects of switch-loop mutations on Rec102 activity in vivo. LexA-Rec102 fusion proteins with the indicated switch-loop sequence were used to complement a rec102∆ mutation. The wild-type switch-loop sequence is shown in black and the amino acid substitutions are in red. The graph shows the frequency of Arg+ prototrophs generated by heteroallelic recombination. Bars show mean ± SD of six replicates; points show individual measurements. Below, Rec102 levels were assessed by anti-LexA immunoblots; anti-tubulin immunoblots served as loading controls. The shown lanes were from a single exposure of the same blot; the vertical dashed line indicates where the image was spliced to remove irrelevant lanes. The full immunoblot is provided in Source Data. f, Detail view of the hydrophobic interface between the amphipathic stalk and the β strands; related residues in Top6B and Rec102 are indicated.
Extended Data Fig. 8 Functional analyses of protein-protein interfaces within the core complex.
a, Hydrogen bonding interactions between Spo11 and Rec102. b, Comparing interfacial areas for Spo11–Rec102 (1,680 Å2, complex with gapped DNA) vs. Top6A–Top6B (M. mazei, 958 Å2, PDB: 2q2e). Interfacial residues are shown as spheres. c, Y2H interactions of mutants of Gal4AD-Spo11 and Rec104-LexA. Bars show mean ± SD of three biological replicates; points show individual measurements. Immunoblots below are presented as in Fig. 3d. Several mutants yielded notable differences in recombination phenotype compared with the Y2H assay. There were two main types of such discrepancy. In one type, the mutation strongly compromised recombination without diminishing the Y2H interaction (for example, rec102-L207A, highlighted in yellow in Fig. 3c,d). Because the core complex presumably must do many things besides just coming together as a single multiprotein entity, it is not surprising that some mutations disrupt biological function without apparently eliminating core complex assembly. The second discrepant type is those that disrupt the Y2H interaction without disrupting recombination (for example, the spo11-L105A, highlighted in peach color in Fig. 3c,d). Such mutants are more surprising than the first type since we assume that integrity of the core complex is critical for DSB formation. We do not know the explanation for these mutants, other than to consider that the artificial Y2H system may fail to recapitulate protein interactions in the more complex environment of the DSB formation machinery that can stabilize protein-protein interfaces that are weakened by some mutations. d, Heteroallele recombination assays with different Rec104 variants introduced into a rec104∆ reporter strain. Bars show mean ± SD of three biological replicates; points show individual measurements. e, Y2H interactions of mutants of LexA-Rec102 and Gal4AD-Rec104. Bars show mean ± SD of three biological replicates; points show individual measurements. Immunoblots below are presented as in Fig. 3d. f, Positions of lysines in Rec102 and Rec104 that are readily crosslinked. Red dashed lines connect the α carbons of lysines that were crosslinked in a prior study14. The distances between their Cα atoms are less than the 27.4 Å maximum for the crosslinker used (19.1 Å for K43 to K64; 24.3 Å for K43 to K60).
Extended Data Fig. 9 Spo11 sequence conservation and structural constraints on Spo11 double cuts and pre-DSB dimers.
a, Multiple sequence alignment of Spo11 and Top6A orthologs at protein-protein and protein-DNA contact sites. b, Cartoons illustrating how physically constraining the orientation of adjacent Spo11 core complexes can explain the minimum length and 10-bp periodicity in the size of double-cut DSB fragments14,35. At right, Spo11 core complexes are proposed to be associated with the surface of chromosome-associated biomolecular condensates containing Rec114, Mei4, and Mer2 proteins14. Dimeric core complexes can capture a DNA molecule and break it. At left, if two adjacent dimeric complexes cut the same DNA molecule, the double cuts can be three or more helical turns apart, but not closer than that. The constrained orientation of the Spo11 core complexes further imposes a 10-bp periodicity on the allowed spacing of double cuts. c, Structural alignment of the hypothetical model of a Spo11 core complex pre-DSB dimer and the crystal structure of the dimeric S. shibatae Topo VI holoenzyme (yellow). One copy of the Spo11 core complex is colored as Fig. 1b, and the other copy is colored in gray. Ski8 is not shown for simplicity.
Supplementary information
Source data
Source Data Fig. 3
Uncropped immunoblot images.
Source Data Fig. 3
Statistical source data.
Source Data Extended Data Fig. 1
Uncropped gels.
Source Data Extended Data Fig. 1
Statistical source data.
Source Data Extended Data Fig. 7
Unprocessed immunoblots.
Source Data Extended Data Fig. 7
Statistical source data.
Source Data Extended Data Fig. 8
Unprocessed immunoblots.
Source Data Extended Data Fig. 8
Statistical source data.
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Yu, Y., Wang, J., Liu, K. et al. Cryo-EM structures of the Spo11 core complex bound to DNA. Nat Struct Mol Biol (2024). https://doi.org/10.1038/s41594-024-01382-8
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DOI: https://doi.org/10.1038/s41594-024-01382-8
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