We first focussed on the described interaction between Mer2 and Spp1. In order to probe the various possible functions of Mer2, we made use of four principal expression constructs, the full-length protein (residues 1–314 from hereon abbreviated to ‘Mer2^FL’), Mer2 amino acids 1–256, lacking the C-terminal 58 residues (from hereon ‘Mer2^∆C’), Mer2 residues 140–314 (i.e. lacking the N-terminal 139 residues; from hereon ‘Mer2^∆N’) and Mer2 containing residues 140–256 (from hereon referred to as ‘Mer2^core’) and a putative coiled-coil domain (Figure 1B). Previous work has identified the Spp1 interaction region to be contained within Mer2^core (specifically Mer2 residues 165–232; Acquaviva et al., 2013). Using our in-house expression system, ‘InteBac’(Altmannova et al., 2021), we produced full-length Spp1 and all Mer2 proteins in Escherichia coli with N-terminal MBP tags to facilitate protein solubility. We could successfully remove the MBP tag using the 3C protease from both Spp1 and Mer2, though in the case of Mer2^∆N and Mer2^Core the 3C cleavable MBP tag could not be removed, presumably due to steric hindrance by the MBP tag that precludes efficient cleavage. In co-lysis experiments, we found that we could purify a complex of Mer2 and Spp1 to homogeneity, and free of nucleic acid contamination (Mer2^FL with Spp1 shown as an example in Figure 1C, note the apparent A²⁶⁰ to A²⁸⁰ ratio as evidence of a lack of nucleic acid contamination), thus indicating that the interaction between Mer2 and Spp1 does not require any PTMs or additional cofactors and that the interaction was robust enough to survive extensive co-purification in >300 mM NaCl.

Using microscale thermophoresis (MST) we measured the binding affinity of Spp1 to Mer2^FL (Figure 1D orange trace, squares) and Spp1 to Mer2^Core (Figure 1D green trace, diamonds). Spp1 bound Mer2^FL with a K_D of 24 nM (±2), and to Mer2^Core with a K_D of 137 nM (±4). Mer2 constructs lacking the ‘core’ showed comparatively weak binding (Figure 1—figure supplement 1). Thus, we confirm that the majority of the Spp1 binding interface is indeed within the core of Mer2, as reported earlier (Acquaviva et al., 2013; Adam et al., 2018), although there does appear to be some contribution to Spp1 binding provided by the N- and C-terminal regions of Mer2. We next tested the reciprocal interaction using a C-terminally 2xStrep-II-tagged Mer2^FL against full-length Spp1, and two additional Spp1 constructs, Spp1^∆C, containing amino acids 1–170, and Spp1^∆PHD containing amino acids 169–353. We found that in a Strep-Mer2^FL pulldown on Streptactin beads Spp1^FL and Spp1^∆PHD interacted with Mer2, but Spp1^∆C did not, consistent with previous studies (Figure 1—figure supplement 2).

We measured the molecular mass of Mer2 by size exclusion chromatography coupled to multi-angle light scattering (SEC-MALS) and concluded that Mer2^core is the tetramerisation region (Figure 1E, orange trace and Figure 1—figure supplement 3A-D), consistent with recent observations of Mer2 (Claeys Bouuaert et al., 2021). Interestingly, while Mer2^∆C∆core is monomeric (Figure 1—figure supplement 3D), Mer2^∆N∆core is dimeric (Figure 1—figure supplement 3C) indicating the presence of a dimerisation region in the C-terminal region of Mer2 between residues 255 and 314, which presumably aids in the stability of a full coiled-coil tetramer.

Given that the tetramerisation region of Mer2 is also the principal Spp1 binding region (Figure 1D), we determined the stoichiometry of the Mer2-Spp1 complex. First, we determined that full-length Spp1 alone is a monomer (Figure 1E yellow trace, Figure 1—figure supplement 3E-F). We next analysed the stoichiometry of Mer2:Spp1 complexes. We measured the size of a complex of MBP-Mer2^core with Spp1 (Figure 1E green trace) and determined its mass to be 290 kDa. The theoretical mass of a 4:2 (MBP-Mer2^core:Spp1) complex is 310 kDa, whereas a 4:1 (MBP-Mer2^core:Spp1) complex is 268 kDa. Given the possible ambiguity in determination of stoichiometry, we also measured complexes of MBP-Mer2^FL and untagged Mer2^FL together with Spp1 (Figure 1—figure supplement 3G and H) which gave complex sizes best fitting a 4:2 (Mer2:Spp1) stoichiometry. Taken together we conclude that the Mer2 tetramer binds two copies of Spp1, establishing a complex in a 4:2 stoichiometry. Thus we show a novel function for Mer2 in not simply binding Spp1, but importantly, in mediating the dimerisation of Spp1. In light of the inherent 2-fold symmetry of nucleosomes, we suggest that this 4:2 constellation might aid in the recognition of modified nucleosome tails by Spp1.

Next we probed the structural organisation of Mer2-Spp1 further using cross-linking coupled to mass spectrometry (XL-MS) using the 11 Å spacer crosslinker disuccinimidyl dibutyric urea (DSBU) (Figure 1F). XL-MS of Mer2-Spp1 revealed that, while the ‘core’ of Mer2 showed many cross-links with Spp1, these were, unexpectedly, not with the previously described C-terminal interaction domain ‘Mer2-ID’ of Spp1 (Acquaviva et al., 2013). Instead the Mer2 core showed numerous cross-links with a region of Spp1 immediately C-terminal to the PHD domain. Furthermore there were additional cross-links between Spp1 and the N- and C-terminal regions of Mer2, consistent with the residual binding affinity we observed in MST (Figure 1—figure supplement 1). The intramolecular cross-linked pattern of Mer2 alone (Figure 1—figure supplement 4A) was very similar in the presence and absence of Spp1. As such we can possibly exclude a significant structural rearrangement of Mer2 upon association with Spp1. We also compared the cross-linking pattern observed previously for Mer2 alone (Claeys Bouuaert et al., 2021; Figure 1—figure supplement 4B). This revealed that the pattern was broadly similar with a mixture of long- and short-distance cross-links. One striking difference is the extensive cross-links emanating from the N-term of our Mer2. The most likely explanation is that the overhang remaining on our Mer2 preparation after removal of the N-terminal fusion protein is four amino acids longer, and thus more flexible.

In order to study the role of the Mer2-Spp1 complex binding to H3K4^me3 nucleosomes, we created synthetic H3K4^me3 mononucleosomes. Briefly, we mutated K4 of histone H3 to cysteine whereas the single naturally occurring cysteine of the natural H3 sequence was mutated to alanine (C110A). H3C4 was converted to H3K4^me3 using a trimethylysine analogue as previously described (Simon et al., 2007). We then reconstituted H3K4^me3 into octamers, and subsequently into mononucleosomes using 167 bp Widom sequences (see Materials and methods for further details). Due to the dimeric nature of nucleosomes, and because our reconstitutions showed a 4:2 Mer2:Spp1 complex stoichiometry, we hypothesised that dimerisation of Spp1 might lead to more tight binding to H3K4^me3 nucleosomes, and set out to test this idea. We compared the apparent nucleosome binding affinity of Spp1, GST-tagged Spp1 (which mediates dimerisation), and the Mer2-Spp1 complex using both electrophoretic mobility shift assays (EMSAs) (Figure 2A) and pulldowns on biotinylated nucleosomes (Figure 2B). We observed that the Mer2-Spp1 assembly (in which two copies of Spp1 are present) bound more tightly to nucleosomes, as compared to monomeric Spp1 alone (which bound relatively weakly to nucleosomes, consistent with the reported ~1 µM affinity of the PHD domain with H3K4^me3 peptide; He et al., 2019). Interestingly, when we compared the observed binding of Spp1:Mer2 with GST-Spp1, we found that GST-Spp1 exhibited an intermediate apparent binding affinity. A potential corollary of this observation is that Mer2 in addition to triggering Spp1 ‘dimerisation’ might directly contribute to nucleosome binding.

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Raw gel data for electrophoretic mobility shift assays (EMSAs) (triplicate), pulldowns, and size exclusion chromatography (SEC) experiments.

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We established that the Mer2-Spp1 complex was capable of forming a stable complex with H3K4^me3 nucleosomes in solution using analytical SEC (Figure 2C). Note that we do not observe a complete shift of nucleosomes; we suspect that this is due to not having an optimal buffer condition (in this experiment we have tried to balance the buffer conditions required for Mer2 [high salt], Spp1 [Zn²⁺ ions for the PHD domain], and nucleosomes [EDTA]). As our complex is currently not suitable for high-resolution structural studies, we made use again of XL-MS to determine a topological architecture of the Mer2-Spp1-H3K4^me3 mononucleosome complex (Figure 2D). We detected many more internal Spp1 cross-links than observed in the Mer2-Spp1 complex alone (Figure 1F), suggesting that either the binding to nucleosomes brings the two Spp1 moieties (in the 2:4 complex) closer to one another, or that there is an internal rearrangement of domains of Spp1. Most strikingly however, the cross-linking revealed that most of the cross-links between the Mer2-Spp1 complex and the nucleosomes are via regions of Mer2, in the N-term, core, and C-term regions. This observation strengthened the idea that Mer2 might directly contribute to nucleosome binding. We modelled the location of the Mer2-Spp1 cross-links onto the previously determined structure of a mononucleosome (Davey et al., 2002; Figure 2E). We observe that the cross-links cluster around histone H3, but more generally around the DNA entry/exit site on the nucleosomes. These observations suggest a large Mer2:nucleosome interface, and a considerably smaller Spp1:nucleosome interface.

Our pulldown data, SEC experiments, and XL-MS data all suggested that Mer2 might bind to mononucleosomes directly, perhaps providing additional affinity to the Spp1:nucleosome interaction. If true, this would be a previously unreported function of Mer2. We tested whether Mer2^FL could bind to unmodified mononucleosomes in an SEC experiment, and surprisingly found that it formed a stable complex (Figure 3A). Given that Mer2 is a tetramer that binds two copies of Spp1, and given that Mer2 has been previously shown to form large assemblies on DNA (Claeys Bouuaert et al., 2021), we asked what the stoichiometry of a Mer2-mononucleosome complex was. To do this, we first used mass photometry (MP), a technique that determines molecular mass in solution at low concentrations based on the intensity of scattered light on a solid surface (Young et al., 2018). In MP, we observe a mix of three species, free Mer2 tetramer (measured at 127 kDa; theoretical mass 142 kDa), free mononucleosomes (measured at 187 kDa; theoretical mass 202 kDa), and a complex at 303 kDa, which most likely corresponds to a 4:1 complex of Mer2:nucleosomes (theoretical mass 344 kDa) (Figure 3B). The MP experiment was carried out at a protein concentration of 60 nM, which suggests that the dissociation constant (K_D) is somewhat less than 60 nM (at K_D under equilibrium one would observe 50% complex formation, we observe less than 50% in Figure 3B). We next asked whether Mer2 might form larger assemblies on nucleosomes as higher concentrations. Using SEC-MALS (Figure 3C), we observed a complex of 341.0 kDa which matches a theoretical complex consisting of one Mer2 tetramer plus one mononucleosome (340 kDa, summarised in Figure 3D). Additionally, we observe a small fraction of a very high molecular weight assembly (though not an aggregate) of 10.96 MDa. This could be an oligomer of Mer2, of Mer2 on nucleosomes, or Mer2 on free DNA (it has recently been reported that Mer2 binds directly to DNA; Claeys Bouuaert et al., 2021). We also observe a shoulder on the Mer2-nucleosome peak, and find that this is a mixture of molecular masses (Figure 3—figure supplement 1A). Given that we observe Mer2 cross-links at the nucleosome DNA entry/exit site, we therefore tested whether Mer2 might simply be binding the free DNA ends on mononucleosomes. Using analytical EMSAs we found that Mer2 binds with a 6-fold higher apparent affinity to nucleosomes (5 nM vs. 30 nM) than to the same 167 bp DNA used to reconstitute the nucleosomes (Figure 3E). The discrepancy between K_D determined by EMSA and the apparent K_D from MP is presumably because EMSAs are non-equilibrium experiments, carried out by necessity at very low salt (Fried and Bromberg, 1997).

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Next we asked what effect using a smaller length of DNA to reconstitute nucleosomes might have. We reconstituted histone octamers on 147 bp DNA from now on referred to as nucleosome core particles (NCP; Lohr and Van Holde, 1975; Sollner-Webb et al., 1976). We found that with no free DNA ends Mer2 did bind with a lower affinity to NCPs, but nonetheless still with an apparent K_D of ~40 nM (Figure 3—figure supplement 2A). We then asked whether mutating the common binding site, the ‘acidic patch’ on H2A (E56T-E61T-E64T-D90S-E91T-E92T) (Kalashnikova et al., 2013) might have an effect on Mer2 binding. In order to enhance potential differences in binding, this was also done on NCPs. Mer2 bound to NCP acidic patch mutants essentially as well as wildtype NCPs (Figure 3—figure supplement 2A). We then asked whether Mer2 might be recognising the histone tails. We therefore prepared ‘tailless’ NCPs (see Materials and methods). Surprisingly Mer2 bound very tightly to tailless NCPs with an apparent K_D of ~5 nM (Figure 3—figure supplement 2A). We suggest that this might indicate that under the conditions of an EMSA, the histone tails are shielding either the histone cores or the NCP DNA and interfering with binding by Mer2.

We next asked which region of Mer2 might be involved in binding nucleosomes (reconstituted with 167 bp DNA). Initially we carried out EMSAs with Mer2^core, Mer2^∆N, and Mer2^∆C. Mer2^∆C and Mer2^∆N appeared to bind slightly weaker than Mer2^FL (K_D ~12.5 and ~30 nM, respectively), whereas Mer2^core showed no binding at all (Figure 3—figure supplement 2B). Apparently there was equal contribution to nucleosome binding from both termini. In order to refine this further, we carried streptavidin pulldown using mononucleosomes reconstituted with biotinylated DNA against different Mer2 constructs. This approach had the advantage of being able to use a more physiological, and as such more stringent, buffer. We confirmed that the Mer2^core did not bind nucleosomes, but neither did Mer2^∆C, strongly suggesting that the main nucleosome interaction region of Mer2 lies in the C-terminal 58 amino acids (Figure 3F), with some additional contribution from the N-terminus of Mer2 (summarised in Figure 3G). We propose that Mer2, in addition to enabling the ‘dimerisation’ of Spp1 via its central tetramerisation domain, provides a direct binding interface with nucleosomes.

In addition to Spp1, several lines of evidence point to an association between Mer2 and meiotic HORMA domain-containing factors. In fission yeast and mouse, the functional homologs of Mer2 (named Rec15 and IHO1, respectively) have been shown to interact with Hop1/HORMAD1 (Kariyazono et al., 2019; Stanzione et al., 2016). Likewise, in budding yeast Mer2 exhibits a chromatin-association pattern which is very similar to Hop1 (Panizza et al., 2011). Meiotic HORMA proteins are integral members of the meiotic chromosome axis and are needed to recruit Mer2 to chromosomes (Panizza et al., 2011; Stanzione et al., 2016). Hop1 (like most other known HORMA domains) can exist in two topological states (‘open/unbuckled’ [O/U] or ‘closed’ [C]), in which the closed state can embrace a binding partner via a closed HORMA-closure motif ‘safety belt’ binding architecture (West et al., 2018). The closure motif (also referred to as CM) is a loosely conserved peptide sequence encoded in HORMA binding partners. The meiotic HORMA proteins are unique among HORMA proteins in the fact that they contain a CM at the end of their own C-terminus (Hop1 residues 585–605; West et al., 2018), endowing these factors with the ability to form an intramolecular (closed) HORMA-CM configuration. The association of Hop1 with chromosomes is mediated by an interaction with Red1 which depends upon a similar CM:HORMA-based interaction. The CM of Red1 (located 340–362; West et al., 2018) binds to Hop1 with a higher affinity than the CM of Hop1 (West et al., 2018). There is mounting evidence in budding yeast and Arabidopsis, that, in addition to a chromosomal pool, a significant pool of Hop1 is non-chromosomal (in the nucleoplasm or cytoplasm) (Herruzo et al., 2021; Raina and Vader, 2020; Yang et al., 2020). Once bound to its own CM, Hop1 should not be able to interact with its chromosomal axis binding partner Red1. Due to the high local concentration of the intramolecular CM, free Hop1 is expected to rapidly transition from the ‘open/unbucked’ into the intramolecular ‘closed’ state (West et al., 2018). The closed state (whether it is intramolecular or, for example, with the CM of Red1) can be reversed by the action of the AAA+ ATPase Pch2/TRIP13 (Rosenberg and Corbett, 2015; Vader, 2015). As such, within the nucleoplasm/cytoplasm, Pch2/TRIP13 activity serves to generate enough ‘open/unbuckled’ Hop1 that is proficient for incorporation into the chromosomal axis, via a CM-based interaction with Red1. On the other hand, when recruited to chromosomes, that same Pch2/TRIP13 activity is expected to dismantle Hop1-Red1 assemblies, as such leading to removal of Hop1 from chromosomes (Deshong et al., 2014; Subramanian et al., 2016).

We tested the ability of Mer2 to interact with the proteins of the meiotic axis (i.e. Hop1 and Red1), with a focus on Hop1. Initially, we purified Hop1 using an N-terminal 2x-Strep-II tag and used it to pull down Mer2. We observed a faint band corresponding to Mer2 in the Hop1 pulldown, indicative of a weak interaction (Figure 4—figure supplement 1, lane 1, control in lane 2). Note that based on the high relative concentration of the CM in Hop1, this Hop1 is expected to largely consist of (intramolecular) closed Hop1. We then co-expressed Red1-MBP containing a I743R (from hereon referred to as Red1^I743R-MBP) mutation with Hop1 in insect cells. The I743R mutation should prevent Red1 from forming filaments, but still allow Red1 to form tetramers (West et al., 2019). Given that we have an excess of Hop1 in our Hop1-Red1 purification, we carried out a pulldown on the MBP tag of Red1 (using amylose beads) with Mer2 as prey. In this case, we observed considerably more Mer2 binding, when measured relative to Hop1, its putative direct binding partner (Figure 4—figure supplement 1, lane 3, control in lane 4). We quantitated the Mer2 intensity relative to the Hop1 band in both pulldowns, and from three independent experiments, and observed an ~6-fold increase in Mer2 binding (Figure 4A). We reasoned that this difference could either be due to Red1 interacting directly with Mer2, or that Red1 induces a conformational change in Hop1 that facilitates Mer2 binding. To test the former idea we purified Red1^I743R-MBP in the presence or absence of Strep-Hop1. Using amylose affinity beads to capture the MBP moiety of Red1 we tested the capture of Mer2 both in the presence and in the absence of Hop1 (Figure 4B). We detected no Mer2 interaction when pulling on Red1^I743R-MBP in the absence of Hop1. As such, we conclude that Mer2 does not have significant affinity for Red1. These data argue that Hop1, when bound to Red1, is acting as an efficient recruiter of Mer2 to this complex. How could this increased affinity of Hop1 for Mer2 be influenced by association with Red1? Based on the known biochemical basis of the Hop1-Red1 interaction, we can imagine that the interaction of Red1 with Hop1 ‘releases’ the C-terminus of Hop1 which could create a ‘chain’ of Hop1 moieties (akin to what has been observed in Caenorhabditis elegans; Kim et al., 2014) or could simply liberate the C-terminus of Hop1 for binding to non-self partners. We propose that such configurations would create or expose Mer2-specific binding interfaces (which can conceivably be located on either the HORMA domain or within the C-terminal non-HORMA domain) that are otherwise shielded when Hop1 is bound intramolecularly to its own C-terminal CM. This predicts that impairing intramolecular CM binding to the HORMA domain should ‘unlock’ the binding ability of Hop1 with Mer2, regardless of Red1 presence. To test this prediction and to delineate the binding interface between Mer2-Hop1, we created two additional Hop1 constructs, one where the N-terminal HORMA domain is missing (Hop1^∆HORMA) and another where the conserved lysine in the CM has been mutated to alanine (Hop1^K593A) disrupting the ability of the CM of Hop1 to interact with its HORMA domain thus forcing Hop1^K593A into an ‘unlocked’ state (West et al., 2018). Both Hop1^∆HORMA and Hop1^K593A were purified with an N-terminal 2xStrep-II tag (as for Hop1^WT) and their ability to bind to Mer2 was tested. In order to prevent background binding, stringent binding conditions were used. We saw that both Hop1^WT and Hop1^K593A bind to Mer2^FL, whereas Hop1^∆HORMA does not (Figure 4C). Quantification of the pulldown analyses showed that Hop1^K593A has ~2-fold stronger binding to Mer2 than does Hop1^WT (Figure 4D). Based on these data, and taking into consideration the effect of Red1 on the Hop1-Mer2 interaction, we suggest the following model: Mer2 binding to Hop1 is driven largely by HORMA domain-mediated interactions, and promoted under conditions where the HORMA is not bound to its intramolecular CM. The most parsimonious molecular explanation of this behavior is that the C-terminal region of Hop1, when kept in close proximity of the HORMA domain (by virtue of CM-HORMA binding), can sterically interfere with the establishment of a Mer2-Hop1 association. Furthermore, the robust Mer2 binding we observe for the Hop1-Red1 complex (Figure 4B) suggests that there is additional contribution from Red1, likely through promoting the closed conformation of the HORMA domain through Red1-CM-HORMA domain binding.

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Raw gel data for pulldowns (triplicate and singular), anti-Mer2 Western blot raw blot data.

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We next asked which region of Mer2 is necessary for Hop1 interaction. Using N-terminally 2xStrepII tagged Hop1^K593A as bait, we queried a variety of Mer2 constructs using Streptactin beads (Figure 4E). Due to the weak staining of Mer2 under these conditions, we also carried out an anti-Mer2 Western blot (Figure 4E lower panel). We determined that Hop1 was capable of interacting, apparently equally well, with all Mer2 constructs, except for Mer2^∆C. Why would the Mer2^core be capable of binding to Hop1, but the ∆C not? The answer may lie in the complex arrangement of the Mer2 coiled-coils and thus N- and C-termini of Mer2 relative to one another and to the core of Mer2. As such one could imagine that part of the interaction region for Hop1 lies in the core of Mer2, which might be shielded by the N-terminus when the C-terminus of Mer2 is not present. These results hint at a complex regulation, on the Mer2 side, underlying Mer2-Hop1 interaction, and we believe that high-resolution structural studies of the Mer2-Hop1 interface should eventually be able to provide deeper insight into this interaction. Taken together we propose a new model for Mer2 recruitment to the meiotic axis in which a ‘locked’ Hop1 cannot bind to Mer2 but once incorporated into Red1, Hop1 recruits Mer2 to the meiotic axis.

We sought to further characterise the interaction between Mer2 and Hop1 by again making use of XL-MS and the DSBU cross-linker (Figure 4F). We observed the most cross-links between the core of Mer2 (consistent with the binding of Mer2^core to Hop1, Figure 4E) and both the HORMA domain of Hop1 and the C-terminal part of the protein (few cross-links were detected within the putative PHD/wHTH like region of Hop1, located in the non-HORMA domain C-terminal part of Hop1; Ur and Corbett, 2021). We also observed additional cross-links with the C-terminal half of Mer2, again consistent with the pulldown data (Figure 4E). Using a model generated using open-Mad2 as a template (Luo et al., 2000), we mapped Mer2-Hop1 cross-links onto the HORMA/SB domain of Hop1 (Figure 4G). This revealed that multiple Mer2 cross-links were concentrated on one ‘face’ of the HORMA domain, whereas the other ‘face’ of the HORMA domain is essentially free of Mer2-Hop1 cross-links. We observed several cross-links in close proximity to the safety belt of the HORMA domain of Hop1 (Figure 4G). While we assume that the Hop1^K593A used in the cross-linking is mostly ‘unlocked’, we acknowledge that the safety belt likely has a somewhat different orientation in ‘unlocked’ Hop1 vs. ‘open’ Mad2 (West et al., 2018). Taken together these data reveal additional molecular details of the Mer2-Hop1 binding mode. The concentration of Mer2 cross-links on one face of Hop1 hints at a potential role for Red1 in enforcing a particular Hop1 orientation that is compatible with efficient recruitment of Mer2.

Due to the potential difficulties in assigning defined interaction regions within Mer2 when using truncation constructs, as so far described, we instead aimed to obtain separation of function alleles by introducing selected point mutants. Making use of sequence alignments from (evolutionary closely and more distantly related) Mer2 orthologs, we identified a conserved region, in the N-terminal domain between amino acids 52 and 71 (Figure 5A, Figure 5—figure supplement 1), that has also been previously annotated as Mer2 SSM1 (Tessé et al., 2017 ). This particular stretch of amino acids stands out in the protein sequence of Mer2 (and homologs) since, in addition to the central coiled-coil region, this region is one of the few regions which shows sequence similarities across evolutionary distant species (such as yeast and human). To probe a potential function of this conserved region, we created two different alleles containing mutations in this region, which we here refer to as mer2-3a and mer2-4a. In mer2-3a, we mutated three conserved residues W58, K61, and L64 to alanine (Figure 5A, red stars). The significance here is both in the high level of conservation, and, in the case of W58, in the presence of a bulky hydrophobic residue that might be involved in protein-protein interactions. Finally, the periodicity of the conserved residues combined with the secondary structure prediction hinted that these residues might be on the same face of an alpha-helical element (see Figure 5—figure supplement 1). mer2-4a contains the following mutations within the same domain: D52A, E68A, R70A, and E71A (Figure 4A, yellow stars). We note that the residues in mer2-4a are less well conserved than those in the mer2-3a allele, but we wondered whether they might function together or in separate pathways. We integrated plasmids carrying C-terminally 3HA-tagged versions of wildtype MER2, mer2-3a, or mer2-4a (note that all these constructs are driven by pMER2) in mer2∆ strains. All three constructs lead to comparable expression levels of Mer2 during meiotic prophase, although we note different mobility of the Mer2^3A-3HA or Mer2^4A-3HA as compared to Mer2-3HA, which might indicate altered post-translational modifications, such as phosphorylation or SUMOylation, since Mer2 was recently shown to be heavily SUMOylated including sites in the region covered by the 3A and 4A mutations (Mer2 K53 and K61; Bhagwat et al., 2021; Figure 5B). Alternatively, it might reflect an inherent effect of the introduced mutations on electrophoretic behavior (compare for example also the migrating patterns of wildtype Mer2 with Mer2^3A on SDS-PAGE from our in vitro preparations, where meiosis-specific post-translational modifications are most likely absent; see Figure 6B).

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The role of Mer2 in meiotic DSB activity is key in enabling faithful meiotic chromosome segregation and viable spore formation. Consequently, mer2Δ strains exhibit very strong spore viability defects, as reported previously (Rockmill et al., 1995 Figure 5C). As a first test of functionality, we investigated the effect of our MER2 alleles on spore viability. Expression of Mer2-3HA in mer2Δ overcame the spore viability defect seen in cells lacking Mer2 (Figure 5C, blue bar) demonstrating functionality of our MER2 expressing constructs. Strikingly, expression of Mer2^3A-3HA or Mer2^4A-3HA failed to restore spore viability in mer2Δ; these strains essentially behaved indistinguishably from the mer2Δ strain. This suggests that the designed mutants disrupt a functionality of Mer2 that is key to its role during meiotic prophase. When we probed for the activation of Mek1 kinase, which is regulated by DSB-dependent Mec1/Tel1 activation (Chuang et al., 2012; Niu et al., 2005), we noticed that expression of the Mer2^3A/4A mutants (in contrast to wildtype Mer2) prevented the appearance of the phosphorylated version of histone H3-threonine 11, a bona-fide Mek1 substrate (Kniewel et al., 2017; Figure 5B). Together with the fact that expression of Mer2^3A/4A mutants did not support the phosphorylation of Hop1 (mediated by Mec1/Tel1 as a response to DSBs) (Carballo et al., 2008; Figure 5B; as seen by an apparent electrophoretic shift in Hop1), these data show that the expression of our Mer2 mutants leads to defective Mec1/Tel1-Mek1 signaling. This hints that DSB formation might be negatively affected by the mutations that we introduced in Mer2.

Since the investigated region of Mer2 is located within the N-terminal region of the protein, we expected that these mutants would not disrupt the interactions with Spp1, Hop1, and nucleosomes described above. In line with this idea, we observed that in vitro, recombinant Mer2^3A and Mer2^4A are able to interact with nucleosomes (Figure 3F) and Hop1 (Figure 4E), albeit with slightly lower affinity for nucleosomes as determined by EMSA (Figure 3—figure supplement 2C).

To attempt to trace the defect of these MER2 alleles, we first investigated the association of Mer2 on spread meiotic chromosomes during meiotic prophase. We found that the recruitment of Mer2 to defined chromosomal foci was not disrupted in Mer2^3A and Mer2^4A expressing situations (Figure 5D). In contrast, quantification of the number of Mer2 foci that were observed during meiotic prophase revealed an increase in cells expressing these mutant proteins (Figure 5E). We currently do not understand the reason underlying this apparent increase in chromosome-associated Mer2 foci, but it could conceivably be related to known feedback regulation that ensures sustained DSB activity in conditions where chromosome synapsis fails or is delayed (Thacker et al., 2014). In any case, recruitment of Mer2 to chromatin-associated foci was not disrupted, suggesting that Mer2^3A and Mer2^4A are proficient to interact with upstream factors that lead to chromatin recruitment.

In order to identify a potential protein-protein interaction that is disrupted in the Mer2 mutants, we carried out a quantitative mass spectrometry analysis of samples immunoprecipitated from meiotic mer2∆ SK1 yeast expressing Mer2^4A-3HA or Mer2^WT-3HA. Wildtype (carrying an untagged MER2 allele) SK1 cells were used as a control. In order to prevent possible confounding effects due to potential differences in DSB forming potential (based on the effect on spore viability and the decrease in phospho-histone H3-T11 signal drop we observed in our mutants, Figure 5B and C), we performed the analysis in a DSB-deficient condition (by using the catalytic-dead spo11-Y135F allele) in both Mer2^4A-3HA and Mer2^WT-3HA. The summary of the differences between Mer2^WT and Mer2^4A are shown in the volcano plot in Figure 5F. In summary, the biggest change we see in the Mer2^4A mutant vs. Mer2^WT is a reduced binding to Hht1 (histone H3). We also saw a reduced binding to the other histones Htb1 and Htb2 (histone H2B). Interaction with Hop1 was largely unaffected (Figure 5G) – in agreement with our in vitro analysis (Figure 4E), as was association with Spp1 and Red1; the latter presumably associated via association with Hop1 (Figure 5—figure supplement 2A and B). Taken together, it appears from these analyses that Mer2^4A exhibits reduced direct nucleosome association. Nonetheless, the effect is likely mild, since we see in vitro association is at most reduced 5-fold in EMSAs for Mer2^3A (Figure 3—figure supplement 2C), and both Mer2^3A and Mer2^4A robustly interact with nucleosomes in a pulldown (Figure 3F), and Mer2 still localises to chromatin (Figure 5D and E).

What might be the cause of the penetrant phenotype seen in Mer2^3A and Mer2^4A? Since HORMA proteins are suggested to drive recruitment of Mer2 to the chromosome axis (Kariyazono et al., 2019; Panizza et al., 2011; Stanzione et al., 2016; and see our earlier observations), these results argue that the defect that is triggered by mer2-3a and mer2-4a is independent of its interaction with Hop1. Furthermore, the observation that spore viability was severely impaired in cells expressing mer2-3a and mer2-4a is in contrast with the relatively mild phenotype caused by the specific disruption of the Mer2-Spp1 interaction (Adam et al., 2018). Finally, our Western blots analysis revealed a loss of Mek1-dependent histone H3-pT11 phosphorylation (a proxy for meiotic DSB formation) in the Mer2^3A/4A mutants (Figure 5B, and see above). Together, these observations, in combination with the strong evolutionary conservation of the affected region, argue that the observed defects are caused by the disruption of yet another key Mer2 interaction, potentially with another essential DSB factor, leading to possible defects in DSB formation.

Strikingly, we detected no Spo11-associated machinery in the Mer2^WT IP (Figure 5—figure supplement 2C and Supplementary mass spectrometry data), despite a large total number of enriched proteins. We therefore postulated that we might be missing key Mer2 interactions – which are potentially difficult to detect using in vivo biochemical purification (ostensibly due to transient/low affinity interactions). The disruption of those interactions, as a result of the Mer2 mutants, might explain observed in vivo phenotypes. As such, we carried out a directed Y2H analysis against previously identified Mer2 interactors involved in DSB formation (Xrs2, Mre11, Ski8, Spo11, Rec104) (Arora et al., 2004) and Spp1 (as a positive control). Analysis of some of these interactions was complicated by the presence of apparent self-activation in our Y2H system. Nonetheless, we observed a clear difference in binding between the Mer2 and Mre11 when comparing MER2 to the mer2-3a or mer2-4a mutants (Figure 6A, Figure 6—figure supplement 1). Importantly, these mutants were proficient to interact with Spp1, similar to what was indicated by our in vitro binding assays and in accordance with the mass spectrometry data (Figure 5—figure supplement 2A).

We aimed to confirm the interaction between Mer2 and Mre11 by other means, and sought to query whether the interaction with other DSB factors was not perturbed in Mer2^3A/4A mutants. We initially used in vivo co-immunoprecipitation (co-IP) to test for Mer2 interaction with Mre11 and Rec114 (Figure 6—figure supplement 2A and B). Using this setup, we were not able to detect Mre11 nor Rec114, although we reliably detected Hop1 in Mer2^WT, Mer2^3A, and Mer2^4A co-IPs, indicating efficient purification of a relevant assembly. The apparent discrepancy between our Y2H and co-IP experiments might reflect an inherently transient or infrequently occurring interaction between Mer2 and Mre11 (and Mer2 and Rec114), precluding detection using our current in vivo methodology. We note that the results obtained with our co-IP experiments (i.e. no detected interaction with Mer2 or Rec114, but robust interaction with Hop1) were similar to the results we obtained using our IP-MS approach comparing Mer2^WT and Mer2^4A (Figure 5F and G). Therefore, we sought to corroborate an interaction between Mer2 and Mre11 using purified components. To this end, we expressed and purified recombinant Mre11 carrying a C-terminal 2xStrep-II tag using baculovirus-based expression. We then used this as a bait to pull down Mer2^WT, Mer2^3A and Mer2^4A. We detected a robust interaction between Mer2^WT and Mre11, and in line with our earlier result, we observed a reduced interaction between Mre11 and Mer2 when Mer2^3A and Mer2^4A were used in the pulldown assay (Figure 6B). We quantified the amount of Mer2 in the pulldown as a factor of Mre11 (Figure 6C): Mer2^3A leads to a 2-fold reduction in binding, and Mer2^4A a reduction closer to 10-fold. So, despite the fact that the effects we observed in pulldown between recombinant Mre11 and Mer2^3A and Mer2^4A were not as striking the ones we detected in our Y2H analysis, we were able to confirm that Mer2 directly associates with Mre11, and that our mutant Mer2 proteins exhibit reduced Mre11 association.

DSB factors are recruited to chromosomes during meiotic prophase (Panizza et al., 2011), and we were interested in addressing whether the introduction of Mer2 mutants affected the chromosomal association of Mre11 and Rec114. We carried out immunofluorescence on meiotic chromosome spreads and analysed the signal corresponding to Rec114 or Mre11 (both carrying a COOH-terminal 13XMYC tag) in cells expressing the different MER2 alleles (Figure 6—figure supplement 3A and B). We did not detect any obvious differences in the levels of chromosomally associated Rec114 or Mre11 in cells expressing wildtype or mutant versions of Mer2. As such, while the interaction between Mer2 and Mre11 might be disrupted in the mutants, the ability of Mre11 (and Rec114) to localise to chromosomes during meiosis appeared unaffected.

We next were interested to shed more light on the interaction between Mer2 and Mre11, and to this end, we carried out XL-MS analysis on a complex of Mer2 and Mre11. These data revealed extensive cross-links across the length of Mre11 and Mer2, with the exception of the C-terminal region of Mer2 (Figure 6D). We paid particular attention to those cross-links that emanate from the residues that are in proximity of the Mer2^3A/4A mutant region (amino acids 52–72) (Figure 6D, pale orange) and noted that these cross-links are predominantly formed with a region at the very C-terminus of Mre11. This region is predicted to be unstructured, but we noted that this region harbors a high-confidence SUMO interaction motif (SIM; residues 633–637) based on the SUMO-GPS prediction algorithm (Zhao et al., 2014). Intriguingly, a recent study on the role SUMOylation in meiosis (Bhagwat et al., 2021) reported Mer2 as a SUMO-target, and found that Mer2^K61 (which is one of the residues mutated in Mer2^3A) is directly SUMOylated. By comparing the location of the SIM (Figure 6D, green bar) with the Mer2-Mre11 cross-links, we noted that the SIM sits immediately adjacent to the most cross-linked region of Mre11. Taken together, we consider it a distinct possibility that the interaction between Mer2 and Mre11 might be influenced (and if so, likely enhanced) by SUMOylation of Mer2 during meiotic prophase.

Our earlier analysis of the phenotypes exhibited by cells expressing our Mer2 mutants, together with the fact that the MRX complex (including Mre11 specifically; Johzuka and Ogawa, 1995) is generally required for meiotic DSB formation (reviewed in Borde, 2007), prompted us to investigate meiotic DSB formation in mer2-3a and mer2-4a mutants. We used Southern blotting to track meiotic DSB formation at YCR047C, a confirmed DSB hotspot. (Note that we utilised the sae2Δ resection and repair-deficient background in order to enable DSB detection.) We detected a strong impairment of meiotic DSB activity specifically in mer2-3a or mer2-4a, to an extent that was comparable to what was observed in mer2Δ strains (Figure 6E). Thus, Mer2^3A and Mer2^4A disrupt a key functionality of Mer2 that is required for meiotic DSB formation. Based on this and our earlier data, we propose that, in addition to its centrally positioned role in mediating chromosome axis and chromatin loop-tethering of the DSB machinery, a third key contribution that Mer2 (and its conserved N-terminal region) makes to enable DSB activity is establishing an interaction Mre11. While we cannot comprehensively rule out an alternative mechanism, our data suggest that not only the presence of Mre11 is required, but its specific interaction with Mer2 is essential for the initiation of DSB formation during meiotic prophase.

Sequences of S. cerevisiae SPP1, HOP1, RED1, and MRE11 were derived from SK1 strain genomic DNA. Due to the presence of an intron in MER2, this was amplified as two separate fragments and Gibson assembled together.

All Mer2 constructs were expressed as an 3C HRV cleavable N-terminal MBP fusion in chemically competent C41 E. coli cells. Protein expression was induced by addition of 250 µM IPTG and the expression continued at 18°C overnight. Cells were washed with 1× PBS and resuspended in lysis buffer (50 mM HEPES pH 7.5, 300 mM NaCl, 5% glycerol, 0.1% Triton-X 100, 1 mM MgCl₂, 5 mM β-mercaptoethanol). Resuspended cells were lysed using an EmulsiFlex C3 (Avestin) in the presence of DNAse (10 μg/mL) and AEBSF (25 μg/mL) before clearance at 20,000 g at 4°C for 30 min. Cleared lysate was applied on a 5 mL MBP-trap column (GE Healthcare) and extensively washed with lysis buffer. Mer2 constructs were eluted with a lysis buffer containing 1 mM maltose and passed through a 6 mL ResourceQ column (GE Healthcare) equilibrated in 50 mM HEPES pH 7.5, 100 mM NaCl, 5% glycerol, 5 mM β-mercaptoethanol. The proteins were eluted by increasing salt gradient to 1 M NaCl. Protein containing elution fractions were concentrated on Amicon concentrator (100 kDa MWCO) and loaded a Superose 6 16/600 (GE Healthcare) pre-equilibrated in SEC buffer (50 mM HEPES pH 7.5, 300 mM NaCl, 10% glycerol, 1 mM TCEP). Untagged Mer2^FL was prepared likewise until concentration of protein eluted from ResourceQ. The concentrated eluent was mixed with 3C HRV protease in a molar ratio of 50:1 and incubated at 4°C for 6 hr. Afterwards, the cleaved protein was loaded on a Superose 6 16/600 pre-equilibrated in SEC buffer for cleaved Mer2 (20 mM HEPES pH 7.5, 500 mM NaCl, 10% glycerol, 1 mM TCEP, 1 mM EDTA, AEBSF).

Spp1 constructs were produced as an 3C HRV cleavable N-terminal MBP or GST fusion in a similar manner as MBP-Mer2. To purify GST-Spp1, cleared lysate was applied on GST-Trap (GE Healthcare) before extensive washing with lysis buffer. The protein was eluted with a lysis buffer with 40 mM reduced glutathione and passed through ResourceQ. Both GST and MBP could be cleaved by adding 3C HRV protease to concentrated protein (using an Amicon concentrator with 30 kDa cutoff) in 1:50 molar ratio. After an ~6 hr incubation at 4°C, the cleaved protein was loaded on Superdex 200 16/600 pre-equilibrated in SEC buffer (50 mM HEPES pH 7.5, 300 mM NaCl, 10% glycerol, 1 mM TCEP).

Hop1 constructs were produced as 3C HRV cleavable N-terminal Twin-StrepII tag in BL21 STAR E. coli cells. The expression was induced by addition of 250 µM IPTG and the expression continued at 18°C for 16 hr. Cleared lysate was applied on Strep-Tactin Superflow Cartridge (Qiagen) before extensive washing in lysis buffer. The bound protein was eluted with a lysis buffer containing 2.5 mM desthiobiotin and loaded on HiTrap Heparin HP column (GE Healthcare) and subsequently eluted with increasing salt gradient to 1 M NaCl. Eluted Strep-Hop1 constructs were concentrated on a 30 kDa MWCO Amicon concentrator and loaded on Superdex 200 16/600 pre-equilibrated in SEC buffer.

Red1 was produced in insect cells as a C-terminal MBP-fusion either alone or co-expressed with Strep-Hop1. Bacmids were in both cases produced in EmBacY cells and subsequently used to transfect Sf9 cells to produce baculovirus. Amplified baculovirus was used to infect Sf9 cells in 1:100 dilution prior to 72 hr cultivation and harvest. Cells were extensively washed and resuspended in Red1 lysis buffer (50 mM HEPES pH 7.5, 300 mM NaCl, 10% glycerol, 1 mM MgCl₂, 5 mM β-mercaptoethanol, 0.1% Triton-100). Resuspended cells were lysed by sonication in the presence of Benzonase and a protease inhibitor cocktail (Serva) before clearance at 40,000 g at 4°C for 1 hr. Cleared lysate was loaded on Strep-Tactin Superflow Cartridge (in case of Red1-Hop1 complex) or MBP-trap column (in case of Red1 alone). Proteins were eluted using a lysis buffer containing 2.5 mM desthiobiotin and 1 mM maltose, respectively. Partially purified proteins were further passed through HiTrap Heparin HP column and eluted with increasing salt gradient to 1 M NaCl. Purified proteins were subsequently concentrated using Pierce concentrator with 30 kDa cutoff in 50 mM HEPES pH 7.5, 300 mM NaCl, 10% glycerol, 1 mM TCEP. Because of the small yield of the proteins, the SEC purification step was neglected and the purity of the proteins was checked using the Refeyn One mass photometer.

Mre11 was produced as a C-terminal Twin-StrepII tag in insect cells using the same expression conditions as for Red1 protein. The cell pellet was resuspended in Mre11 lysis buffer (50 mM HEPES pH 7.5, 300 mM NaCl, 5% glycerol, 0.01% NP40, 5 mM β-mercaptoethanol, AEBSF, Serva inhibitors). Resuspended cells were lysed by sonication before clearance at 40,000 g at 4°C for 1 hr. Cleared lysate was loaded on a 5 mL Strep-Tactin XT Superflow Cartridge (IBA) followed by first wash using 25 mL of high-salt wash buffer (20 mM HEPES pH 7.5, 500 mM NaCl, 5% glycerol, 0.01% NP40, 1 mM β-mercaptoethanol) and second wash step using 25 mL of low-salt wash buffer (20 mM HEPES pH 7.5, 150 mM NaCl, 5% glycerol, 0.01% NP40, 1 mM β-mercaptoethanol). The Mre11 protein was eluted with 50 mL of low-salt wash buffer containing 50 mM biotin. Partially purified protein was further loaded onto a 5 mL Heparin column (GE Healthcare) pre-equilibrated in a low-salt wash buffer and eluted with increasing salt gradient to 1 M NaCl. The fractions containing Mre11 protein were concentrated on a 50 kDa MWCO Amicon concentrator and applied onto a Superdex 200 10/300 column (GE Healthcare) pre-equilibrated in Mre11 SEC buffer (20 mM HEPES pH 7.5, 300 mM NaCl, 5% glycerol, 1 mM β-mercaptoethanol, 1 mM TCEP).

Fifty µL samples at 5–10 µM concentration were loaded onto a Superose 6 5/150 analytical size exclusion column (GE Healthcare) equilibrated in buffer containing 50 mM HEPES pH 7.5, 1 mM TCEP, 300 mM NaCl (for samples without nucleosomes) or 150 mM NaCl (for samples with nucleosomes) attached to an 1260 Infinity II LC System (Agilent). MALS was carried out using a Wyatt DAWN detector attached in line with the size exclusion column.

Triplicates of MST analysis were performed in 50 mM HEPES pH 7.5, 300 mM NaCl, 5% glycerol, 1 mM TCEP, 0.005% Tween-20 at 20°C. The final reaction included 20 μM RED-NHS labelled untagged Spp1 (labelling was performed as in manufacturer’s protocol – Nanotemper) and titration series of MBP-Mer2 constructs (concentrations calculated based on oligomerisation stage of Mer2). The final curves were automatically fitted in Nanotemper analysis software.

Recombinant Xenopus laevis histones were purchased from ‘The Histone Source’ (Colorado State) with the exception of H3-C110A_K4C cloned into pET3, which was kindly gifted by Francesca Matirolli. The trimethylated H3 in C110A background was prepared as previously described (Simon et al., 2007). X. laevis histone expression, purification, octamer refolding, and mononucleosome reconstitution were performed as described (Luger et al., 1999). Plasmids for the production of 601–147 (pUC19) and 601–167 (pUC18) DNA were kindly gifted by Francesca Matirolli (Hubrecht Institute, Utrecht) and Andrea Musacchio (MPI Dortmund), respectively. DNA production was performed as previously described (Luger et al., 1999). Reconstituted mononucleosomes were shifted to 20 mM Tris pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM TCEP with addition of 20% glycerol prior to freezing in –80°C.

Quadruplicate EMSAs were carried out as previously described (Weir et al., 2016), at a constant nucleosome/NCP/DNA concentration of 10 nM with the DNA being post-stained with SYBRGold (Invitrogen). Gels were imaged using a ChemiDocMP (Bio-Rad Inc). Nucleosome depletion in each lane was quantitated by ImageJ, using measurements of triplicate of the nucleosome alone for each individual gel as a baseline. Binding curves were fitted using Prism software and the following algorithm (Y = Bmax*X^h/(K_D^h + X^h)). It was necessary in each Mer2 case to add a Hill coefficient to obtain the best fit.

Streptactin pulldowns were performed using pre-blocked streptavidin magnetic beads (Pierce) in a pulldown buffer (20 mM HEPES pH 7.5, 300 mM NaCl, 5% glycerol, 1 mM TCEP). One µM Strep-Hop1 or Mre11-Strep as a bait was incubated with 3 µM Mer2 as a prey in 40 µL reaction for 2 hr on ice without beads and another 30 min after addition of 10 µL of beads pre-blocked with 1 mg/mL BSA for 2 hr. After incubation, the beads were washed twice with 200 µL of buffer before elution of the proteins with a 1× Laemmli buffer. Samples were loaded on 10% SDS-PAGE gel and afterwards stained with InstantBlue.

Amylose pulldowns were performed using pre-blocked Amylose beads (New England BioLabs) in a pulldown buffer (20 mM HEPES pH 7.5, 300 mM NaCl, 5% glycerol, 1 mM TCEP). One µM Red^I743R-MBP or Red^I743R-MBP/Strep-Hop1 as a bait was incubated with 3 µM Mer2 as a prey in 40 µL reaction for 2 hr on ice without beads and another 1 hr after addition of 10 µL of beads pre-blocked with 1 mg/mL BSA for 2 hr. After incubation, the beads were washed twice with 200 µL of buffer before elution of the proteins with a buffer containing 1 mM maltose. Samples were loaded on 10% SDS-PAGE gel and afterwards stained with InstantBlue.

Biotinylated nucleosomes (0.5 μM) or NCP (0.4 µM) were incubated with prey proteins (1.5 μM) for 30 min on ice in buffer containing 20 mM HEPES pH 7.5, 150 mM NaCl, 5% glycerol, 1 mM EDTA, 0.05% Triton-X100, 1 mM TCEP in a reaction volume of 40 µL. Ten µL of protein mix were taken as an input before adding 10 µL of pre-equilibrated magnetic Dynabeads M 270 streptavidin beads (Thermo Fisher Scientific) to the reaction. The samples with beads were incubated on ice for 2 min before applying magnet and removing the supernatant. The beads were washed twice with 200 µL of buffer. To release the streptavidin from the beads, Laemmli buffer (1×) was added to the beads and incubated for 10 min. Samples were analysed on 10–20% SDS-PAGE gel and stained by InstantBlue.

Analytical SEC was performed using Superose 6 5/150 GL column (GE Healthcare) in a buffer containing 20 mM HEPES pH 7.5, 150 mM NaCl, 5% glycerol, 1 mM TCEP, 1 mM EDTA. All samples were eluted under isocratic elution at a flow rate of 0.15 mL/min. Protein elution was monitored at 280 nm. Fractions were subsequently analysed by SDS-PAGE and InstantBlue staining. To detect complex formation, proteins were mixed at 5 µM concentration in 50 µL and incubated on ice for 1 hr prior to SEC analysis.

MP was performed in 20 mM HEPES pH 7.5, 150 mM NaCl, 5% glycerol, 1 mM TCEP, 1 mM EDTA. Mer2 and mononucleosomes (600 nM) were mixed and incubated for 1 hr on ice prior to analysis using the Refeyn One mass photometer. Immediately before analysis, the sample was diluted 1:10 with the aforementioned buffer. Molecular mass was determined in Analysis software provided by the manufacturer using a NativeMark- (Invitrogen) based standard curve created under the identical buffer composition.

For XL-MS analysis proteins were dissolved in 200 μL of 30 mM HEPES pH 7.5, 1 mM TCEP, 300 mM NaCl (for samples without nucleosomes) or 150 mM NaCl (for samples with nucleosomes) to final concentration of 3 μM, mixed with 3 μL of DSBU (200 mM) and incubated for 1 hr in 25°C. The reaction was quenched by addition of 20 μL of Tris pH 8.0 (1 M) and incubated for another 30 min in 25°C. The crosslinked sample was precipitated by addition of 4× volumes of 100% cold acetone ON in –20°C and subsequently analysed as previously described (Pan et al., 2018).

All strains, except those used for Y2H analysis, are of the SK1 background. See Supplementary material for a description of genotypes of strains used per experiment. For MER2 alleles, constructs containing pMER2(−1000–1), the coding sequence of MER2 lacking its intron (i.e. wildtype, or 3a/4a-containing sequences), C-terminal 3HA tag, and 500 base pairs of downstream sequence flanked by HindIII and NarI restriction enzymes were custom-synthesised by Genewiz Inc These constructs were recloned in a YIPlac128 plasmid carrying LEU2 using restriction cloning. These plasmids were integrated at pMER2 in front of mer2::HISMX6 following EcoRI linearisation. Correct single copy integration was confirmed by PCR.

Yeast strains were patched onto YP-Glycerol plates and transferred to YP-4%Dextrose plates. After this, cells were grown overnight in liquid YPD culture (room temperature) followed by inoculation in pre-sporulation media (BYTA; 50 mM Sodium Phthalate-buffered, 1% yeast extract, 2% tryptone, and 1% acetate) at OD₆₀₀ = 0.3. Cells were grown for 18 hr in BYTA at 30°C, washed twice in water and resuspended in sporulation media (0.3% potassium acetate) at OD₆₀₀ = 1.9 to induce meiosis at 30°C.

Cells were synchronously induced into meiosis and incubated for 24 hr. Of each strain, 200 cells were counted using a standard bright-field microscope, and monads, dyads, and tetrads were scored. For viability, the indicated number of tetrads was dissected using standard manipulation methods and grown on YPD plates. Spore viability was calculated as a percentage of the total number of viable spores.

Two mL of meiotic cells (t = 3 hr induction) were collected at indicated time points, killed by addition of 1% sodium azide and processed. Cells were treated with 200 mM Tris pH 7.5, 20 mM dithiothreitol for 2 min at room temperature followed by spheroplasting at 30°C in 1 M sorbitol, 2% potassium acetate, 0.13 µg/µL zymolyase (20 min). Spheroplasts were washed two times in 1 mL ice-cold MES-Sorbitol solution (1 M sorbitol, 0.1 M MES pH 6.4, 1 mM EDTA, 0.5 mM MgCl₂) and resuspended in 55 µL of MES-Sorbitol. Twenty µL of spheroplasts were placed on clean glass slides (that were dipped in ethanol overnight and air-dried) and 2× volumes of fixing solution (3% paraformaldehyde, 3.4% sucrose) were added. This was followed by addition of four volumes of 1% Lipsol, and mixing through gentle rotation. After 1 min, 4× volumes of fixing solution were added. A glass rod was used to mechanically spread chromosomes, after which samples were dried overnight at room temperature, and stored at −20°C. Slides were treated with 0.4% Photoflo (Kodak)/PBS for 3 min, after which slides were dipped in PBS with gentle shaking (5 min). Samples were blocked by incubation with 5% BSA in PBS for 15 min at room temperature. Overnight incubation with desired primary antibodies was performed in a humidified chamber at 4°C, after which slides were subjected to 2× washes of 10 min in PBS with gentle shaking followed by incubation with fluorescent-conjugated secondary antibody for 3 hr at room temperature. The slides were washed 2× and mounted using 20 µL of Vectashield mounting solution containing 4’,6-diamidine-2’-phenylindole dihydrochloride (DAPI) (Vector Laboratories). Chromosome surface spreads were immunostained with rat α-HA at 1:200 (Roche), mouse α-MYC (9E10) at 1:200, and rabbit α-Hop1 at 1:200 (home made). Hop1 antibody production was performed at the antibody facility of the Max-Planck-Institute of Molecular Cell Biology and Genetics (Dresden, Germany) using affinity purified full length 6xHis-tagged Hop1. Secondary antibodies were used at the following concentrations: Alexa 488-conjugated donkey α-rat at 1:500; Texas Red 594-conjugated donkey α-rabbit at 1:500, and Cy3-conjugated donkey α-mouse at 1:500. Image acquisition was done by obtaining serial z-stacks of 0.2 μm thickness at room temperature using 100 × 1.42 NA PlanApo-N objective (Olympus) on a DeltaVision imaging system (GE Healthcare) equipped with an sCMOS camera (PCO Edge 5.5). The z-stack images were deconvolved using SoftWoRx software. Quantifications for the number of foci of Mer2^WT-3HA, Mer2^3A-3HA Mer2^4A-HA were done using the ‘Spots’ function of the Imaris software (Bitplane). Prism 8 (GraphPad) was used to generate the Scatter plots. Statistical significance was assessed by performing Mann-Whitney U-test. For representative images, Fiji/ImageJ software was used to obtain maximum intensity projection images.

One-hundred mL of meiotic cultures (at 4 hr into a meiotic time course) were harvested by spinning down at 3000 rpm for 3 min. Samples were washed with cold H₂O and snap frozen. To the pelleted cells, the following was added: 300 μL of ice-cold IP buffer (consisting of 50 mM Tris–HCl pH 7.5, 150 mM NaCl, 1% Triton X-100 and 1 mM EDTA pH 8.0, and a cocktail of protease inhibitors which was freshly added) and acid-washed glass beads. A FastPrep-24 disruptor (MP Biomedicals) was used to break the cells (MP Biomedicals) (setting: two 50 s cycles (speed 6)). Lysates were spun 30 s at 500 rpm, after which the supernatant was transferred to a 15 mL falcon tube. The resulting lysate was sonicated for 25 cycles (30 s on/30 s off, high power range) using a Bioruptor-Plus sonicator (Diagenode), and subsequently spun down 30 min at maximum speed at 4°C. The supernatant was transferred into new microcentrifuge tubes, and 10% of the supernatant (~50 μL) was collected as input. One μL of antibody (⍺-HA; BioLegend) was added, and the lysate was rotated for 3 hr at 4°C; 30 μL of buffer-washed Dynabeads protein G (Invitrogen, Thermo Fisher Scientific) was subsequently added, and the lysate was rotated overnight at 4°C. The next day, Dynabeads were washed four times with 500 μL of ice-cold IP buffer. For the final wash, beads were transferred to a new microcentrifuge tube. After removal of the supernatant, the beads were resuspended in 40 μL IP buffer, and 20 μL of 3XSDS-loading buffer was added. Samples were incubated for 5 min at 95°C. For the inputs, the following protocol was used: Inputs were precipitated by the addition of 5 μL 100% trichloroacetic acid (TCA) (10% final concentration), and incubated on ice for 30 min. Precipitates were collected by centrifugation, for 1 min at maximum speed, and washed with ice-cold acetone. Precipitations were dried and resuspended in resuspension buffer (40 μL, 50 mM Tris-HCl 7.5, and 6 M urea) for 30 min (on ice). Dissolution was encouraged by pipetting and vortexing. Ten μL of 5XSDS-loading buffer was added, and samples were incubated for 5 min at 95°C.

For Western blot analysis, protein lysates from yeast meiotic cultures were prepared using TCA precipitation and run on 8% or 10% SDS gels, transferred for 90 min at 300 mA and blotted with the selected antibodies, as described (Kuhl et al., 2020). Primary antibodies with respective dilutions were used: rabbit α-Hop1 (made in-house; 1:10,000), rabbit α-Mer2^40-271 (made in-house; 1:10,000); mouse α-Pgk1 (Thermo Fisher, 1:5000); rabbit α-phospho-Histone-H3-Thr11 (Abcam, 1:1000), mouse α-HA (Biolegend, diluted 1:500), α-Myc-9E11 (Abcam, ab56, 1:1000).

For Southern blot assay, DNA from meiotic samples was prepared as described (Vader et al., 2011). DNA was digested with HindIII (to detect DSBs at the control YCR047C hotspot) followed by gel electrophoresis, blotting of the membranes and radioactive (32P) hybridisation using a probe for YCR047C (chromosome III; 209,361–201,030) (Raina and Vader, 2020). DSBs signals were monitored by exposure of an X-ray film which was analysed using a Typhoon Trio scanner (GE Healthcare) after 1 week.

MER2 variants and their potential interactors were cloned into pGAD-C1 or pGBDU-C1 vectors, respectively. The resulting plasmids were co-transformed into the S. cerevisiae reporter strain (yWL365) and plated onto the selective medium lacking leucine and uracil. For drop assay, 2.5 μL from 10-fold serial dilutions of cell cultures with the initial optical density (OD₆₀₀) of 0.5 were spotted onto -Leu/-Ura (control) and -Leu/-Ura/-His plates. Cells were grown at 30°C for up to 7 days and imaged at time points indicated in the figures.

Cells (450 mL) were harvested after 3 hr of synchronised meiosis. Three independent cultures were prepared for each strain. Pellets were lysed in cryo-mill and the resulting powder was resuspended in 25 mL of lysis buffer (50 mM HEPES pH 7.5, 300 mM NaCl, 5% glycerol, 0.01% NP40, 5 mM β-mercaptoethanol, AEBSF, Serva protease inhibitors, 1 mM NEM, 1× Complete Mini EDTA-Free protease inhibitors; Roche). The lysate was cleared by centrifugation at 10,000 rpm for 1 hr. The supernatant was incubated with 5 μL of α-HA antibody (Sigma-Aldrich, H6908) for 3 hr at 4°C followed by the addition of 25 μL magnetic Dynabeads Protein G (Invitrogen, 10,004D) pre-equilibrated with wash buffer (20 mM HEPES pH 7.5, 100 mM NaCl, 5% glycerol, 0.01% NP40, 1 mM β-mercaptoethanol). The samples were incubated overnight at 4°C. Next day, the samples were centrifuged at 2500 rpm for 1 min at 4°C and the beads were washed six times with 150 μL of 20 mM ammonium bicarbonate. Proteins were eluted with 75 μL of 2× SDS Laemmli buffer and boiled for 5 min at 95°C. The samples were analysed by Proteomics Core Facility (EMBL, Heidelberg).

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

We are extremely grateful to Francesca Mattiroli (Hubrecht Institute, Utrecht), for assistance with producing mononucleosomes. The histone H3 expression plasmid pET3 and the plasmid pUC19_601–147 DNA were a kind gift from Francesca Mattiroli. We are also grateful to Luke Berchowitz (Columbia University, New York), Andreas Hochwagen (NYU, New York), and Eric Greene (Columbia University, New York) for support, reagents, and expertise. The plasmid pUC18_601–167 DNA was a kind gift from Andrea Musacchio (MPI of Molecular Physiology, Dortmund). We thank Christopher Heim (MPI Developmental Biology) for technical help with SEC-MALS and MST measurements. We thank Andreas Blaha and Constanze Gremmelmaier for their technical support. Quantitative mass spectrometry and analysis was made possible by the EMBL Proteomics Core Facility (EMBL, Heidelberg). Work in the Weir lab is funded by the Max Planck Society and DFG grant WE 6513/2–1. Work in the Vader lab was funded by Max Planck Society and the European Research Council (ERC StG URDNA; agreement no. 638197), and an Amsterdam UMC Research Fellowship. SKF is funded by a studentship from the International Max Planck Research School (IMPRS) 'From Molecules to Organisms'.

1. Preprint posted: July 30, 2020 (view preprint)
2. Received: July 20, 2021
3. Accepted: December 23, 2021
4. Accepted Manuscript published: December 24, 2021 (version 1)
5. Version of Record published: February 14, 2022 (version 2)

© 2021, Rousová et al.

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