Chromatin-associated cohesin turns over extensively and forms new cohesive linkages in Drosophila oocytes during meiotic prophase

SUMMARY

In dividing cells, accurate chromosome segregation depends on sister chromatid cohesion, protein linkages that are established during DNA replication. Faithful chromosome segregation in oocytes requires that cohesion, first established in S phase, remain intact for days to decades, depending on the organism. Premature loss of meiotic cohesion in oocytes leads to the production of aneuploid gametes and contributes to the increased incidence of meiotic segregation errors as women age (maternal age effect). The prevailing model is that cohesive linkages do not turn over in mammalian oocytes. However, we have previously reported that cohesion-related defects arise in Drosophila oocytes when individual cohesin subunits or cohesin regulators are knocked down after meiotic S phase. Here we use two strategies to express a tagged cohesin subunit exclusively during mid-prophase in Drosophila oocytes and demonstrate that newly expressed cohesin is used to form de novo linkages after meiotic S phase. Cohesin along the arms of oocyte chromosomes appears to completely turn over within a two-day window during prophase, whereas replacement is less extensive at centromeres. Unlike S-phase cohesion establishment, the formation of new cohesive linkages during meiotic prophase does not require acetylation of conserved lysines within the Smc3 head. Our findings indicate that maintenance of cohesion between S phase and chromosome segregation in Drosophila oocytes requires an active cohesion rejuvenation program that generates new cohesive linkages during meiotic prophase.

eTOC Blurb

Findings of Haseeb et al. challenge the model that oocytes rely solely on the original cohesive linkages established during meiotic S phase. Using two different approaches, they demonstrate that generation of new cohesive linkages occurs during meiotic prophase and is required to maintain sister chromatid cohesion in Drosophila oocytes.

Graphical Abstract

INTRODUCTION

During cell division, sister chromatid cohesion is required for proper alignment of chromosomes on the spindle and for accurate chromosome segregation.^1–6 When sister chromatids are created during DNA replication, cohesin-mediated protein linkages are formed that keep the sisters physically associated. Within the cohesin complex, the head domains of the Smc1-Smc3 heterodimer interact with an α-kleisin subunit, forming a ring that encircles the DNA. The formation of stable cohesive linkages during S phase depends on Eco-mediated acetylation of conserved lysine residues located in the head domain of the cohesin subunit, Smc3.^7–11

In meiotic cells, sister chromatid cohesion is not only required for accurate segregation of sisters during meiosis II, but also for proper alignment and segregation of homologous chromosomes during the first meiotic division.^3,4,6 Within the bivalent, sister chromatid cohesion distal to the crossover keeps recombinant homologs physically associated until anaphase I. In mammalian oocytes, cohesion establishment and meiotic recombination are completed during fetal development and oocytes arrest in the dictyate stage of prophase I until ovulation. Several lines of evidence indicate that meiotic cohesion weakens as oocytes age and support the hypothesis that premature loss of cohesion in human oocytes contributes to the maternal age effect.^3,12–14 Because turnover of cohesive linkages has not been detected in mouse oocytes^15–17, the prominent model is that human oocytes rely solely on the original cohesive linkages established before birth.^3,12–14,18

We have previously demonstrated that knockdown of the cohesin loader (Nipped-B), the cohesion establishment factor (Eco), or individual cohesin subunits during meiotic prophase in Drosophila oocytes leads to phenotypes consistent with premature loss of cohesion.¹⁹ Based on these findings, we proposed that a cohesion rejuvenation program operates during meiotic prophase to establish new cohesive linkages after S phase and that this process is required to keep sister chromatids physically associated until chromosome segregation.

Using two independent approaches, we show here that a tagged cohesin subunit expressed exclusively during meiotic prophase loads onto oocyte chromosomes and is used to form new cohesive linkages after meiotic S phase. In contrast to cohesion establishment during DNA replication, our data indicate that acetylation of conserved lysines within the Smc3 head is not required for the formation of functional cohesive linkages during meiotic prophase. We observe extensive turnover of chromosome-associated cohesin on oocyte chromosomes and failure to load cohesin onto chromosomes during meiotic prophase significantly increases the incidence of cohesion defects. Together our findings demonstrate that loading of cohesin and formation of new cohesive linkages after S phase are required to maintain cohesion in Drosophila oocytes during meiotic prophase.

RESULTS

HA-tagged Smc3* expressed after meiotic DNA replication associates with oocyte chromosomes and colocalizes with the synaptonemal complex (SC)

To manipulate expression of cohesion proteins during Drosophila oogenesis, we utilized the Gal4/UAS system for both RNAi-mediated knockdown and Gal4-inducible protein expression. We designed a Gal4/UAS inducible cDNA construct to express Smc3 protein tagged with HA at its C-terminus. We engineered silent mutations in the Smc3 cDNA (denoted Smc3*) which rendered the mRNA expressed from this transgene to be insensitive to the SH00137.N hairpin used to knock down endogenous Smc3 protein (Figure 1B).

Figure 1. Smc3 expressed after S phase stably associates with meiotic chromosomes independent of K105–106 acetylation.

(A) Matα-Gal4 driver expression begins in germarial region 3 (R3) of the Drosophila ovariole and increases during subsequent stages.

(B) Conserved lysine residues (K105–106, blue) required for S-phase cohesion establishment are mutated in KK-Smc3*-HA.

(C) Left: UAS-Gal4 strategy uses matα-Gal4 driver to knock down endogenous Smc3 and simultaneously express HA-tagged Smc3* protein during meiotic prophase. Right: WT-Smc3*-HA and KK-Smc3*-HA colocalize with the synaptonemal complex protein, C(3)G in stage 2 oocytes. Scale bar, 2μm.

See also Figure S1.

The matα-Gal4-VP16 driver (hereafter named matα-Gal4) is germline specific and turns on approximately two days after exit from meiotic S phase¹⁹, specifically during mid-prophase (see Figures 1A & S1). Use of this driver allows us to modify protein levels after the original cohesive linkages have already been established during meiotic S phase. Using this strategy, we have previously demonstrated that matα-Gal4 induced knockdown of cohesion proteins during meiotic prophase in Drosophila oocytes causes phenotypes indicative of premature loss of cohesion.¹⁹ Here, we utilize “strong” and “weak” matα-Gal4 driver chromosomes, which we have previously described.²⁰ These driver chromosomes also contain an amorphic allele of the matrimony gene (mtrm^KG0805).²¹

Using the strong matα-Gal4 driver to knock down endogenous Smc3 and simultaneously express wild-type Smc3 tagged with HA (WT-Smc3*-HA), we asked whether HA-tagged Smc3* could associate with the meiotic chromosomes during prophase (Figure 1C). We confirmed that WT-Smc3*-HA signal is not detectable until germarial region 3 (R3, Figure S1), the stage at which we have previously documented matα-Gal4 to first drive expression.¹⁹ Cohesin-enriched chromosome axes form a scaffold for assembly of the synaptonemal complex (SC), a multi-protein structure that holds homologous chromosomes in close proximity during meiotic recombination.^22–25 WT-Smc3*-HA localization overlaps extensively with that of the SC protein C(3)G (Figure 1C) and mimics what we have previously described for endogenous cohesin subunits.²⁶ These data indicate that a cohesin subunit synthesized after meiotic S phase associates with the meiotic chromosomes during prophase.

SC defects caused by prophase knockdown of endogenous Smc3 protein are suppressed by simultaneous expression of HA-tagged-Smc3*

To assess whether WT-Smc3*-HA protein that loads onto chromosomes after meiotic S phase becomes part of functional cohesin complexes, we asked whether SC defects caused by prophase knockdown of endogenous Smc3 are suppressed by simultaneous expression of RNAi insensitive HA-tagged Smc3*. We have previously shown that matα-Gal4 driven knockdown of individual cohesion proteins causes premature disassembly of the SC as early as stage 2.¹⁹ In wild-type Drosophila oocytes, the SC fully assembles in germarial region 2A and usually remains intact until stages 5–6, although the timing of disassembly can be affected by genetic background.^27,28

To quantify SC defects, we visualized C(3)G immunostaining and assigned each oocyte to one of four categories (Figure 2A). Long continuous threads of C(3)G signal are indicative of normal, full-length SC. Broken threads, short threads, and spots correspond to SC defects with increasing severity. For control oocytes (Figure 2B), we utilized females that lacked a Gal4 driver but contained the UAS-Smc3 hairpin transgene as well as an “Empty” UASp vector insertion that does not code for protein. Inclusion of this “UAS-Empty” transgene allowed us to match the number of UAS binding sites in control and experimental genotypes and avoid confounding results due to Gal4 titration effects. We observed full-length SC in all control oocytes until stage 6, at which time a few oocytes had begun normal disassembly of the SC (Figure 2B).

Figure 2. Prophase expression of HA-tagged-Smc3* delays premature SC disassembly caused by knockdown of endogenous Smc3 protein.

(A) Synaptonemal complex (SC) integrity was assessed using C(3)G immunostaining and oocytes were assigned to one of the four categories shown. Scale bar, 2μm.

(B-E) SC defects were quantified from region 3 (R3) to stage 6 (S6) in the genotypes shown, all of which are also heterozygous for mtrm^KG0805.

See also Figures S2 and S3.

When the strong matα-Gal4 driver is combined with the UAS-Smc3 hairpin and UAS-Empty vector (Smc3 KD + Empty), SC defects are visible in stage 2 and increase in severity in subsequent stages (Figure 2C). These results are very similar to what we previously observed for matα-Gal4 driven expression of the same Smc3 hairpin in the absence of UAS-Empty.¹⁹ In contrast, when expression of WT-Smc3*-HA and knockdown of endogenous Smc3 are simultaneously driven by strong matα-Gal4 (Smc3 KD + WT-Smc3*-HA), SC defects do not arise until stage 3 and are less severe in stage 4 than for Smc3 KD oocytes lacking HA-tagged Smc3* (Figure 2C). This result supports the hypothesis that cohesin complexes that load onto meiotic chromosomes during prophase are functional. However, expression of WT-Smc3*-HA only marginally reduces the frequency and severity of SC defects in stage 5 oocytes (compared to Smc3 KD + Empty), and SC defects in stage 6 are equivalent in the two genotypes (Figure 2C). We reasoned that SC defects in later stages might arise if overexpression of WT-Smc3*-HA caused by the strong matα-Gal4 driver perturbs the normal stoichiometry of cohesin subunits and disrupts proper cellular localization of cohesin as described for cultured mammalian cells.²⁹ Consistent with this idea, WT-Smc3*-HA induced by strong matα-Gal4 is primarily cytosolic during stages 5 and 6, with little to no nuclear signal (Figure S2). In addition, the timing and magnitude of SC defects are similar when the strong matα-Gal4 driver is used to knock down endogenous Smc3 (Figure 2C) or express additional HA-tagged Smc3* protein in the absence of Smc3 knockdown (Figure 2E).

To express lower levels of RNAi insensitive HA-tagged Smc3* specifically during prophase, we utilized a previously described weak matα-Gal4-VP16 driver chromosome²⁰ for which UAS-inducible expression is substantially lower (Figure S3). As expected, expression of the Smc3 hairpin with the weak driver resulted in SC defects that were less severe and began later than those observed with the strong driver (Figure 2C–D). When we induced expression of both WT-Smc3*-HA and the Smc3 hairpin with the weak matα-Gal4 driver, premature disassembly of the SC started later than in Smc3 KD + Empty oocytes and the defects were mild (Figure 2D), again supporting the hypothesis that cohesin loaded onto meiotic chromosomes during prophase is functional.

Neither the strong nor weak matα-Gal4 driver chromosome results in Smc3*-HA expression that completely rescues the SC defects caused by knockdown of endogenous Smc3. The level of prophase-expressed HA-tagged Smc3* relative to endogenous cohesin subunits is likely critical for its functionality.²⁹ Although we have been unable to detect Smc3*-HA signal on oocyte chromatin when using the weak driver, SC defects in stages 5 and 6 are better suppressed when the weak driver is used. These data support the hypothesis that mislocalization of WT-Smc3*-HA caused by strong matα-Gal4 driver induced overexpression is responsible for the severe SC defects we observe in stages 5–6 in that genotype.

HA-tagged Smc3* expressed during meiotic prophase is used to establish new cohesive linkages

To directly assay the state of sister chromatid cohesion in Smc3 KD oocytes expressing low levels of RNAi insensitive WT-Smc3*-HA (weak matα-Gal4 driver), we utilized Fluorescence In Situ Hybridization (FISH). Using the 359-bp satellite repeat probe that binds to X chromosome pericentric heterochromatin and an Oligopaint arm probe that hybridizes to a distal 100kb region on the X chromosome (Figure 3A), we quantified cohesion defects at these two locations in mature Drosophila oocytes (stages 13 and 14). Sister chromatid cohesion distal to the crossover keeps recombinant homologs physically associated (Figure 3A), and premature loss of arm cohesion results in segregation errors during the first meiotic division. Using a different arm probe, we have previously reported an increase in arm cohesion defects, but not pericentric cohesion defects, when the original matα-Gal4 driver chromosome (mtrm⁺) was used to induce expression of the SH00137.N Smc3 hairpin.³⁰

Figure 3. New cohesive linkages are established during meiotic prophase and their formation does not require acetylation of Smc3 at conserved lysines K105–106.

(A) Recombinant X chromosome bivalent in Drosophila oocytes with gray centromeres and cohesion pictured as black lines. Sites of hybridization for pericentric (magenta) and Oligopaint arm (yellow) FISH probes.

(B) Examples of intact arm cohesion (1 or 2 yellow spots) and premature loss of arm cohesion (3 or 4 yellow spots). Scale bar, 2μm.

(C) Arm cohesion defects in the genotypes shown, which are also heterozygous for the mtrm^KG0805 allele. *** corresponds to P<0.001 and ** corresponds to P<0.002.

We first quantified cohesion defects in oocytes that contained both the UAS-Smc3 hairpin and the UAS-Empty transgene in the absence or presence of the weak matα-Gal4 driver. Expression of the Smc3 hairpin caused a significant increase in the percentage of oocytes exhibiting arm cohesion defects (Figure 3C, P < 0.001). Pericentric cohesion defects were negligible in Smc3 KD oocytes (Figure 3C), consistent with our previous studies.³⁰ We attribute the high incidence of arm cohesion defects in control (no driver) oocytes to heterozygosity of the mutant mtrm allele on the 3^rd chromosome of all four genotypes in Figure 3C (See Discussion). Our results demonstrate that even the weak matα-Gal4 driver results in Smc3 knockdown during meiotic prophase that is sufficient to cause a significant elevation of arm cohesion defects in prometaphase and metaphase I arrested Drosophila oocytes.

We then asked whether expression of RNAi insensitive WT-Smc3*-HA after meiotic S phase can suppress arm cohesion defects caused by knockdown of endogenous Smc3. When the weak matα-Gal4 driver was used to express both the Smc3 hairpin and the WT-Smc3*-HA transgene during meiotic prophase, the number of oocytes with premature loss of arm cohesion was significantly lower than for Smc3 KD + UAS-Empty oocytes (Figure 3C, P < 0.002). Moreover, the incidence of defects in Smc3 KD oocytes expressing WT-Smc3*-HA was similar (P = 0.79) to that in control oocytes that lack a Gal4 driver, consistent with complete rescue. These data demonstrate that new cohesive linkages are established after meiotic S phase in Drosophila oocytes.

Formation of new cohesive linkages during meiotic prophase does not require acetylation of Smc3 at conserved lysines 105–106

During DNA replication, the formation of stable cohesive linkages depends on Eco-mediated acetylation of conserved lysines within the Smc3 head domain.^7–11 These lysines correspond to K105–106 in Drosophila Smc3 (Figure 1B). We have previously demonstrated that matα-Gal4 induced knockdown of the acetyltransferase Eco in Drosophila oocytes causes premature disassembly of the SC and chromosome segregation errors consistent with premature loss of arm cohesion.¹⁹ These results support the hypothesis that acetylation of one or more targets by Eco is required for cohesion rejuvenation in Drosophila oocytes during meiotic prophase.

To determine whether acetylation of lysines 105–106 in Drosophila Smc3 is required for the formation of functional cohesive linkages during meiotic prophase, we utilized a second RNAi insensitive HA-tagged Smc3* cDNA construct in which these lysines are mutated to non-acetylatable arginine residues (KK-Smc3*-HA, Figure 1B). When expression of KK-Smc3*-HA was induced by the strong matα-Gal4 driver, we observed threads of HA signal colocalizing with the SC protein C(3)G (Figure 1C), indicating that acetylation of K105–106 is not required for association of Smc3 with oocyte chromosomes during meiotic prophase. Furthermore, KK-Smc3*-HA expressed during meiotic prophase was able to function as well as WT-Smc3*-HA in its ability to suppress SC defects caused by knockdown of endogenous Smc3 (Figure 2C–D). This was true whether we utilized the strong or weak matα-Gal4 driver chromosome. Finally, like WT-Smc3*-HA, expression of KK-Smc3*-HA during meiotic prophase rescued arm cohesion defects caused by Smc3 knockdown (Figure 3C, P < 0.002). These data indicate that formation of new cohesive linkages in Drosophila oocytes during meiotic prophase does not depend on Eco-dependent acetylation of Smc3 at K105–106.

Chromatin-associated cohesin turns over extensively during meiotic prophase

One drawback of matα-Gal4 induction of UAS-Smc3*-HA during meiotic prophase is that we have been unable to express HA-tagged Smc3* at a level comparable to endogenous Smc3. In addition, we wanted to rule out the possibility that a cohesin subunit expressed during prophase associates with chromatin and contributes to formation of new cohesive linkages only if we knock down the endogenous subunit.

To address the above concerns, we designed a construct that allowed us to visualize tagged-Smc1 protein expressed at physiological levels specifically during meiotic prophase (Figure 4A). This Smc1 tag-switch transgene, integrated at the attP40 site on chromosome 2, is composed of Smc1 genomic DNA, including upstream regulatory sequences that control its expression. In addition, the construct encodes a switchable fluorescent protein tag fused to the C-terminus of the encoded Smc1 protein (Figure 4A). In the absence of Flippase (FLP) expression, Smc1 encoded by this transgene is tagged with mCherry. However, because the mCherry coding sequence (including a stop codon) is flanked by FRT sites, induction of FLP expression causes excision of mCherry DNA and results in expression of Smc1 protein fused to superfolder-GFP (sGFP).³¹

Figure 4. Chromosome-associated cohesin turns over extensively during meiotic prophase.

(A) Smc1 tag-switch transgene and expression strategy.

(B-C) Smc1-mCherry and Smc1-sGFP on meiotic chromosomes from region 3 (R3) through stage 6 (S6) in Smc3 control and knockdown oocytes. Arrowheads mark centromeres (based on CID). Scale bars, 2μm.

(D-E) Quantification of Smc1-mCherry and Smc1-sGFP on chromosome arms and centromeres.

See also Figures S4 and S5.

Using Smc1 immunoblotting, we verified that the level of tagged Smc1 protein produced from the tag-switch transgene is similar to that encoded by a single copy of the endogenous smc1 gene (Figure S4A). In addition, we confirmed that the transgene rescues the lethality of smc1Δ homozygotes (Figure S4B), indicating that the tagged Smc1 protein is functional.

We utilized the original matα-Gal4 driver chromosome (mtrm⁺) to induce expression of a UAS-FLP transgene in the germline of females also containing the Smc1 tag-switch insertion but which were otherwise wild type (Figure 4B). In this genotype, the onset of FLP expression in germarial region 3 (see Figure 1A) should elicit a switch from Smc1-mCherry to Smc1-sGFP expression, allowing us to visualize sGFP-tagged Smc1 produced exclusively during meiotic prophase.

We monitored the localization and signal intensity of chromosome-associated Smc1-mCherry and Smc1-sGFP in oocytes from germarial region 3 (R3) to stage 6 (S6). The expected pattern of long continuous Smc1-mCherry threads on oocyte DNA was evident in R3 (Figure 4B) as well as earlier stages within the germarium (Figure S5). In contrast, we did not observe distinct Smc1-sGFP signal on oocyte chromosomes until stage 2 (Figures 4B & S5), and its intensity was relatively weak (Figure 4B & 4D). During subsequent stages, the signal for sGFP-tagged Smc1 became more prominent on both the chromosome arms and the centromeres (Figure 4B & 4D). In stages 2 and 3, we observed extensive colocalization of mCherry and sGFP on oocyte DNA. However, as the signal intensity for chromosomal sGFP-tagged Smc1 increased, we observed a concomitant decrease in Smc1-mCherry associated with the meiotic chromosomes (Figure 4B & 4D). In stages 5 and 6, we observed both Smc1-sGFP and Smc1-mCherry at oocyte centromeres, but distinct threadlike signal on the arms was only apparent for sGFP-tagged Smc1 (Figure 4B). These data indicate that chromosome-associated cohesin turns over extensively during meiotic prophase. Importantly, association of prophase-specific Smc1-sGFP with meiotic chromosomes occurs even when the level of endogenous Smc1 is not perturbed by knockdown.

To further characterize Smc1 tag-switch dynamics during meiotic prophase, we examined Smc1-mCherry and Smc1-sGFP localization in oocytes in which expression of the UAS-Smc3 hairpin was induced by the original matα-Gal4 driver chromosome (Figure 4C). When Smc3 was knocked down, loading of newly synthesized sGFP-tagged-Smc1 onto oocyte chromosomes was almost completely abolished (Figure 4C & 4E), although at later stages we detected faint Smc1-sGFP signal at some centromeres (Figure 4C, arrowheads). As we observed for Smc3 control oocytes, chromosome-associated Smc1-mCherry decreased during prophase progression in Smc3 KD oocytes, but the decline was more precipitous than in control oocytes (compare Figure 4D & 4E). Similar to Smc3 control oocytes (Figure 4B), mCherry-tagged Smc1 was still visible at oocyte centromeres at later stages in Smc3 KD oocytes (Figure 4C, arrowheads in S4-S6). In contrast to Smc3 control oocytes (Figure 4B), Smc1-sGFP and Smc1-mCherry signals were often most predominant in the cytoplasm, not the nuclei, of Smc3 KD oocytes, especially at later stages (Figure 4C). Based on these data, we conclude that loading of newly synthesized sGFP-tagged-Smc1 onto chromosomes during meiotic prophase depends on the other cohesin subunits and that the Smc1-sGFP dynamics that we observe reflect the behavior of the intact cohesin ring.

The data presented in Figure 4 indicate that cohesin association with meiotic chromosomes is highly dynamic during pachytene. We also wanted to determine whether cohesin turnover continues during diplotene, the stage at which human oocytes remain arrested for decades. Unfortunately, our ability to induce a switch from Smc1-mCherry to Smc1-sGFP during late pachytene or early diplotene was precluded by lack of a suitable Gal4-VP16 driver. However, quantitative analysis of matα-Gal4 → UAS-FLP → Smc1 tag-switch oocytes at later stages of oogenesis indicated that chromatin-associated Smc1-sGFP levels continue to increase in oocytes after they exit pachytene (Figure 5). These data support the hypothesis that cohesion rejuvenation continues throughout late prophase in Drosophila oocytes.

Figure 5. Cohesin loading onto oocyte chromosomes continues during diplotene.

(A) Smc1-sGFP and C(3)G immunostaining on oocyte chromosomes in control ovaries. Disassembly of the SC marks the transition to diplotene. Scale bar, 2μm.

(B) Quantification of chromatin-associated Smc1-sGFP in oocytes at indicated stages.

Failure to load cohesin onto chromosomes during meiotic prophase leads to premature loss of arm cohesion

Newly synthesized Smc1-sGFP is unable to associate with oocyte chromosomes during pachytene when its heterodimeric partner, Smc3, is knocked down (Figure 4C & 4E). Therefore, the Smc3 KD genotype provides an opportunity to ask whether failure to load “rejuvenation specific” cohesin onto meiotic chromosomes causes premature loss of cohesion.

We quantified Smc1 tag-switch localization in stage 10 oocytes for both Smc3 control and knockdown genotypes. This late prophase stage occurs approximately three days after the onset of matα-Gal4 expression.^19,32 We have been unable to detect endogenous or tagged-cohesin subunits on the condensed chromosomes of metaphase I arrested oocytes (stage 14), most likely due to technical limitations. However, in well-fed females undergoing normal egg-laying, it takes only a short time (~three hours) for a stage 10 oocyte to mature to stage 14 and arrest at metaphase I.³²

In the presence of normal levels of Smc3, sGFP-tagged Smc1 was enriched in the large nucleus of the stage 10 oocyte (Figure S6A–B) and associated with the meiotic chromosomes (Figure 6A). In contrast, when Smc3 was knocked down, Smc1-sGFP was largely excluded from the nuclei of stage 10 oocytes (Figure S6C–D) and failed to localize to the meiotic chromosomes (Figure 6A–B). Notably, even in stage 10 oocytes, mCherry-tagged Smc1 was still faintly visible at the centromeres in both genotypes (Figure 6A, arrowheads).

Figure 6. Failure to load newly synthesized cohesin onto oocyte chromosomes during meiotic prophase leads to premature loss of arm cohesion.

(A) Chromatin localization of Smc1-mCherry and Smc1-sGFP in stage 10 oocytes. Arrowheads mark the location of centromeres (based on CID). Scale bar, 2μm.

(B) Chromatin-associated Smc1-sGFP signal intensity in stage 10 oocytes in Smc3 control and knockdown.

(C) Quantification of arm cohesion defects in Smc3 control and knockdown oocytes expressing the Smc1 tag-switch transgene.

See also Figure S6.

To ask whether Smc1-sGFP containing cohesin complexes participate in the formation of new cohesive linkages during meiotic prophase, we performed FISH to quantify cohesion defects in Smc3 control and knockdown oocytes expressing the Smc1 tag-switch transgene (Figure 6C). In Smc3 knockdown oocytes, in which Smc1-sGFP fails to load onto the oocyte DNA during meiotic prophase, cohesion defects were significantly higher than in Smc3 control oocytes (Figure 6C, P < 0.03). These data indicate that normal turnover and loading of cohesin complexes during meiotic prophase leads to the formation of new cohesive linkages that are required to maintain arm cohesion in Drosophila oocytes.

DISCUSSION

Using two different approaches, we have observed that cohesin synthesized after meiotic S phase can load onto Drosophila oocyte chromosomes and facilitate the formation of new cohesive linkages during prophase. When we use the prophase-specific matα-Gal4 driver to knock down endogenous Smc3, simultaneous expression of a UAS-Smc3*-HA transgene can partially suppress knockdown-induced SC defects (Figure 2) and fully rescue arm cohesion defects (Figure 3). However, one challenge with this approach is achieving physiological levels of Smc3*-HA expression. Therefore, we used an alternative strategy in which genomic sequences within an Smc1 transgene resulted in expression of tagged Smc1 protein at levels that were comparable to the endogenous protein. Matα-Gal4 induced expression of FLP during meiotic prophase causes a switch from Smc1-mCherry to Smc1-sGFP expression from this transgene, allowing us to follow the behavior of a cohesin subunit synthesized exclusively during prophase. Using this Smc1 tag-switch construct, we observed nearly complete turnover of cohesin on oocyte chromosomes within a two-day timeframe (R3 to S6, 4B & 4D). In addition, an increase in chromatin-associated Smc1-sGFP after SC disassembly argues that cohesion rejuvenation continues during diplotene (Figure 5), the stage at which human oocytes remain arrested for decades. Given that meiotic prophase lasts approximately six days in Drosophila oocytes³², our data suggest that chromatin-bound cohesin complexes could be replaced three times during the course of meiosis. Importantly, when Smc3 is knocked down, chromosome association of Smc1-sGFP during meiotic prophase is nearly abolished (Figure 4C & 4E). Moreover, failure of sGFP-tagged Smc1/cohesin to associate with the meiotic chromosomes during meiotic prophase results in premature loss of arm cohesion (Figure 6). These data provide compelling evidence that turnover of cohesin on meiotic chromosomes is normal in Drosophila oocytes and leads to the formation of new cohesive linkages after DNA replication.

Our previous work indicated that cohesion rejuvenation during meiotic prophase requires the acetyltransferase Eco.¹⁹ During DNA replication, cohesion establishment depends on Eco-mediated acetylation of two lysines within the Smc3 head domain.^7–11 In contrast, we show here that formation of new linkages during meiotic prophase does not require acetylation of these conserved lysines (K105–6) within Drosophila Smc3; when endogenous Smc3 is knocked down, expression of KK-Smc3*-HA rescues arm cohesion defects as well as WT-Smc3*-HA (Figure 3, P = 0.99). Our data suggest that formation of stable cohesive linkages during meiotic prophase depends on Eco-dependent acetylation of other lysines within Smc3 or a completely different protein target. In mitotically dividing yeast cells, Eco acetylation of the mitotic α-kleisin (not Smc3) is required for genome-wide cohesion establishment during G2 in response to DNA damage.^33,34 However, when meiotic double strand breaks (DSBs) are prevented (Spo11 null oocytes), Eco is still required during meiotic prophase to prevent cohesion-related defects in Drosophila oocytes, indicating that the cohesion rejuvenation pathway does not depend on induction of DSBs.¹⁹ Therefore, formation of new cohesive linkages during meiotic prophase in Drosophila oocytes is mechanistically distinct from cohesion establishment during S phase and DNA-damage induced cohesion establishment during G2 in yeast cells. Future experiments will be required to identify the Eco target required for cohesion rejuvenation during meiotic prophase.

We show here that association of cohesin with chromosomes is highly dynamic during meiotic prophase in Drosophila oocytes. Interestingly, within a 2-day period, replacement at centromeres appears less complete than that on arms. However, the rate at which tag-switch Smc1 turns over at oocyte centromeres and arms appears to be similar (Figure 4D). One possibility is that clustering of centromeres during prophase in Drosophila oocytes³⁵ allows us to detect residual Smc1-mCherry more readily at centromeres than on arms. Additionally, a higher density of cohesin complexes in pericentric heterochromatin might take longer to completely replace. However, at this time, we cannot exclude the possibility that cohesive linkages within pericentric heterochromatin are more stable than those on chromosome arms. This could explain why our FISH analyses uncovered very few pericentric cohesion defects when we knock down Smc3 (³⁰ and this paper). Although matα-induced knockdown of Smc1 significantly increases the missegregation of recombinant homologs¹⁹, consistent with decreased rejuvenation of arm cohesion, segregation errors indicative of centromeric cohesion loss were not completely absent in Smc1 KD oocytes.¹⁹ Moreover, the large satellite DNA target (~11Mb) of the X-chromosome pericentric probe likely impedes, at least to some degree, our ability to detect cohesion defects within the pericentric heterochromatin of the X chromosome. Given these caveats, we think it premature to conclude that turnover of cohesive linkages is restricted to the chromosome arms.

Our experiments expressing UAS-Smc3*-HA utilized matα-Gal4 driver chromosomes (strong and weak) that also carried an amorphic allele of the matrimony gene (mtrm^KG08051, abbreviated mtrm^KG). In Drosophila oocytes, accurate segregation of achiasmate homologs relies on homology-dependent interactions within their pericentric heterochromatin^36,37, and heterozygosity for mtrm^KG causes missegregation of achiasmate homologs.²¹ Because the achiasmate segregation system also promotes accurate segregation of recombinant homologs that have lost chiasmata due to premature loss of arm cohesion, we often utilize a mtrm^KG/+ background for chromosome segregation assays to disable this mechanism.^20,30,38 Our previous creation of recombinant chromosomes that contain both the matα-Gal4-VP16 transgene and the mtrm^KG allele resulted in driver chromosomes of different strengths²⁰, which proved useful for this study.

In our FISH analyses, the baseline for arm cohesion defects in control oocytes was considerably higher in mtrm^KG/+ oocytes (Figure 3C) than in oocytes with two wild-type copies of mtrm (Figure 6C). Mtrm protein binds and inhibits the activity of Drosophila Polo-like kinase (PLK) in a dose dependent manner.³⁹ Furthermore, in both mitotic and meiotic cells, PLK phosphorylation of cohesin has been shown to cause its dissociation from chromosome arms during prophase in a cleavage-independent manner.^40–42 mtrm null oocytes exhibit a high incidence of completely separated chromatids⁴³, consistent with elevated PLK activity in late prophase inducing premature dissociation of cohesin from the meiotic chromosomes. Therefore, partial dissociation of cohesin from meiotic chromosomes caused by increased PLK activity in mtrm^KG/+ oocytes during late prophase is likely responsible for the increased frequency of arm cohesion defects that we observe in mtrm^KG/+ oocytes compared to mtrm⁺. This basal level of cohesion defects in mtrm^KG/+ control oocytes is further increased by knockdown of the cohesin subunit Smc3 during meiotic prophase (Figure 3C). Our FISH data argue that mtrm^KG/+ oocytes provide a useful sensitized background for measuring segregation errors caused by premature loss of arm cohesion not only because the achiasmate system is disabled but also because this genotype reduces (but does not eliminate) the number of functional cohesive linkages on meiotic chromosomes during late prophase.

Here we demonstrate that cohesin turnover and formation of new cohesive linkages during meiotic prophase are required to prevent premature loss of arm cohesion in Drosophila oocytes during normal oogenesis. In contrast, analysis of mouse oocytes has not uncovered evidence that new cohesive linkages are established during meiotic prophase.^15–17 However, orthologs of two proteins required for cohesion rejuvenation in Drosophila oocytes, the mammalian cohesin loader (NIPBL) and the cohesion establishment factor (ESCO2), both localize to mouse oocyte chromosomes during multiple prophase stages, including diplotene.^44–46 Because Drosophila lacks Smc1 or Smc3 variants, one strength of our approach is that it encompasses all cohesin complexes present in Drosophila oocytes. Although we acknowledge that cohesion rejuvenation may be unique to Drosophila oocytes, we question why this process would be necessary to maintain meiotic cohesion during a six-day timeframe (flies) but not during an extended prophase arrest that lasts months (mice) or years (humans).

In Drosophila oocytes, chromosome association of cohesin is highly dynamic during meiotic prophase and cohesin loading and establishment of new cohesive linkages after oocyte DNA replication are required to maintain arm cohesion. However, even with a cohesion rejuvenation program that operates during meiotic prophase, when diplotene Drosophila oocytes undergo aging, they exhibit a significant increase in chromosome missegregation.^20,38 Furthermore, our previous work revealed that oxidative damage contributes to age-induced meiotic segregation errors.^20,30 Our findings raise the possibility that oxidative damage compromises the rejuvenation program in aging Drosophila oocytes. If a meiotic cohesion rejuvenation pathway also operates in human oocytes, premature loss of cohesion may arise in the oocytes of older women, at least in part, because rejuvenation declines with age. Further delineation of the mechanism(s) underlying rejuvenation and its possible age-induced decline in Drosophila oocytes could potentially inform therapeutic strategies to bolster rejuvenation and slow cohesion loss in aging human oocytes.

STAR METHODS

RESOURCE AVAILABILITY

Lead Contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Sharon E. Bickel (sharon.e.bickel@dartmouth.edu).

Materials Availability

All Drosophila stocks and reagents used in this study are available upon request from the lead contact without restriction.

Data and Code Availability

• All data reported in this paper will be shared by the lead contact upon request.

• This paper does not report original code.

• Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

Fly Stocks and Crosses

All fly stocks and crosses were reared on standard cornmeal molasses medium in a humidified chamber at 25° C. Table S1 provides full genotypes for stocks used in this study, and Bickel stock numbers from this table are noted below for cross descriptions.

Creation of WT-Smc3*-HA and KK-Smc3*-HA transgenic flies

All site-directed mutagenesis was performed using the QuikChange II XL Site-Directed Mutagenesis Kit (Agilent 200521) and PAGE purified primers from Integrated DNA Technologies. All mutations were verified by sequencing. Primer sequences are provided in Table S1. Using a Bluescript KS vector containing 12 tandem HA tags (12X HA) (gift from Christian Lehner⁵⁰), we engineered a stop codon at the end of the HA open reading frame (ORF). The Smc3 ORF (including the consensus translational start sequence⁵¹) was PCR amplified using clone RE14758 (DGRC Stock 8502; https://dgrc.bio.indiana.edu//stock/8502; RRID:DGRC_8502) obtained from the Drosophila Genomics Resource Center. PCR primers introduced Kpn1 and Not1 sites at the 5’ end and an Xho1 site at the 3’ end to facilitate subcloning. Following Kpn1 and Xho1 digestion, the amplified Smc3 fragment was cloned into the above KS vector, immediately upstream of the 12X HA sequence. The entire ORF was sequenced to verify that no mutations occurred during PCR amplification.

Silent mutations were introduced into the Smc3 sequence so that the encoded mRNA (Smc3*) was insensitive to the Smc3 short hairpin SH00137.N used to knock down endogenous Smc3. The original Smc3 sequence: GAGTATATACGCTACGAA was mutated to GAATACATTCGGTATGAG (mutated nucleotides underlined), resulting in WT-Smc3*-HA.

Eco-dependent acetylation of two conserved lysines is required for cohesion establishment during S phase.^7–11 For the KK-Smc3*-HA construct, site-directed mutagenesis was used to convert these lysines (K105, K106 in Drosophila Smc3) to arginines, which cannot be acetylated. WT-Smc3*-HA and KK-Smc3*-HA were each subcloned into pUASP-attB⁴⁹ using Not1 and Spe1 sites and confirmed by sequencing. Injections were performed by Genetic Services, Inc (Sudbury, MA) to generate PhiC31 integrase mediated attP40 insertions on chromosome 2.⁵²

Generation of Smc1 tag-switch construct and transgenic flies

The Smc1 tag-switch construct (see Figure 4) was synthesized by Genscript Biotech Corp (Piscataway, NJ) using the Drosophila smc1 genomic sequence (FlyBase ID FBgn0040283). In addition to all exons and introns, 976 bp of upstream regulatory DNA (including the promoter) as well as the 5’ and 3’UTRs were included. At the end of the Smc1 ORF, a 48bp FRT site replaces the Smc1 stop codon and is followed by the mCherry coding sequence, a 3X HA tag (with stop codon), the Smc1 3’ UTR, and an additional 100bp of smc1 genomic DNA. Immediately downstream of the mCherry cassette lies an additional 48 bp FRT site, followed by DNA encoding superfolder GFP (sGFP), an EPEA epitope tag (with stop codon), the smc1 3’ UTR, and an additional 100bp of smc1 genomic DNA. Following synthesis, GenScript cloned the construct into the pUC57 vector, verified the sequence, and subcloned the insertion into the w+attB vector (Addgene plasmid # 30326). Both mCherry and sGFP sequences were codon optimized for Drosophila. Sequence of the Smc1 tag-switch construct in the w+attB vector will be provided upon request. BestGene Inc (Chino Hills, CA) performed injections to insert the Smc1 tag-switch transgene from w+attB into the attP40 landing site on chromosome 2.⁵²

Crosses to induce knockdown and/or protein expression

To induce expression of UASp-Smc3*-HA, we utilized two previously described²⁰ matα-Gal4-VP16 driver chromosomes (“strong” and “weak”), each of which also contains the mtrm^KG08051 allele (W-110 and W-107). Expression induced by our strong matα driver is similar to that of the original matα chromosome (Bloomington #7063, T-273). Gal4-inducible expression is considerably lower when we utilize our weak matα driver chromosome (²⁰ and Figure S3).

To knock down endogenous Smc3 during meiotic prophase and simultaneously express wild-type Smc3*-HA (WT), mutant Smc3*-HA (KK) or no additional Smc3 protein (Empty), strong matα, mtrm^KG /TM3 (W-110), weak matα, mtrm^KG /TM3 (W-107) or no driver, mtrm^KG /TM3 (W-109) males were crossed to UAS-WT-Smc3*-HA; Smc3 hairpin (T-622), UAS-KK-Smc3*-HA; Smc3 hairpin (T-625) or UAS-Empty; Smc3 hairpin (T-800) virgins. Ovaries from non-balancer female progeny were used for immunostaining and FISH. The strong matα driver was used for immunolocalization of HA-tagged WT- and KK-Smc3* (Figures 1C, S1, and S2) as well as quantification of SC defects (Figure 2C). Ovaries from the no driver and weak matα driver crosses were also used to assay SC defects (Figure 2B and 2D) and to quantify cohesion defects using FISH (Figure 3). To quantify SC defects when WT-Smc3*-HA was overexpressed in the absence of Smc3 knockdown (Figure 2E), strong matα, mtrm^KG /TM3 (W-110) males were crossed to UAS-WT-Smc3*-HA (T-615) virgins and ovaries dissected from non-balancer female progeny. For the immunoblots in Figure S3, no driver, mtrm^KG /TM3 (W-109), weak matα, mtrm^KG /TM3 (W-107), strong matα, mtrm^KG /TM3 (W-110) or nanos-Gal4-VP16 driver; + ; mtrm^KG /TM3 (T-764) males were crossed to UAS-WT-Smc3*-HA (T-615) virgins and ovaries from non-balancer female progeny were used to prepare protein extracts.

For Smc1 tag-switch experiments, we used the original matα-Gal4-VP16 driver chromosome (T-273, mtrm⁺) to induce expression of Flippase (FLP) after meiotic S phase. Excision of FRT-flanked mCherry sequences in the Smc1 tag-switch transgene induces a switch from Smc1-mCherry to Smc1-GFP expression during meiotic prophase. UASp-FLP; matα (T-778) males were crossed to Smc1 tag-switch (T-772) or Smc1 tag-switch; Smc3 hairpin (T-779) virgins. Ovaries of female progeny were used for immunolocalization of Smc1-mCherry, Smc1-sGFP, and CID on meiotic chromosomes (Figures 4, 5, 6, S5, and S6), for FISH analysis of cohesion defects (Figure 6), and for the immunoblots (Figure S4A).

To confirm that Smc1-mCherry and Smc1-sGFP were each functional, we performed a series of crosses to determine whether the lethality of flies transheterozygous for two different smc1 deletion alleles (smc1Δ^ex46/smc1Δ^ex89) was rescued when the Smc1 tag-switch transgene was present and solely expressing either Smc1-mCherry or Smc1-sGFP. In the absence of FLP, the intact Smc1 tag-switch transgene expresses only mCherry-tagged Smc1. Following a series of crosses (using T-815, OL-124 and M-743), we recovered Smc1-mCherry/+; smc1Δ^ex46/smc1Δ^ex89 males and females at the expected ratio and verified that they were fully fertile (Figure S4B). To generate flies solely expressing Smc1-sGFP, we generated a stock (T-816, Smc1-sGFP) starting with progeny from UASp-FLP/+; Smc1 tag-switch/+; matα/+ females. Progeny that inherit the FRT-excised transgene (Smc1-sGFP) can no longer express Smc1-mCherry. We confirmed the absence of Smc1-mCherry expression in this stock using immunostaining. Through a series of crosses (using T-817, OL-124 and M-743), we verified that Smc1-sGFP/+ ; smc1Δ^ex46/smc1Δ^ex89 males and females eclosed at the expected ratio and were fully fertile (Figure S4B).

METHOD DETAILS

Immunostaining

Newly eclosed females were fattened for 2–3 days in vials containing food, yeast, and males. Following dissection in 1X PBS, ovarioles were partially separated with a fine tungsten needle and fixed for 20 minutes in a mixture of 600μl heptane and 200μl of 2% formaldehyde (Ted Pella, 18505) containing 0.5% NP-40 (Thermo Fisher 28324). All incubations and washes were performed in glass dishes at room temperature (RT) with gentle agitation on a rotating platform unless otherwise stated. Fixed ovaries were rinsed three times in 1X PBST (1X PBS + 0.2% Tween-20, Thermo Fisher 28320), blocked in 1X PBST + 1% BSA (Fisher BP1605–100) for 1 hour, and incubated at 4°C overnight in antibody incubation buffer (1X PBS + 0.01% Tween-20 + 1% BSA) containing the appropriate primary antibodies. After rinsing three times in 1X PBST, ovaries were washed three times for 20 minutes in 1X PBST and placed in antibody incubation buffer containing the appropriate secondary antibodies for 1 hour. Following three rinses in 1X PBST, one 20-minute wash in 1X PBST, a 20-minute incubation in 1μg/ml DAPI (Invitrogen D1306) in 1X PBS, and one 20-minute wash in 1XPBS + 0.01% Tween-20, ovarioles were fully separated with tungsten needles. Ovarioles were transferred onto to 18mm poly-L-lysine coated #1.5 coverslips, and excess liquid removed before adding mounting media.

For HA-tagged-Smc3* and C(3)G immunolocalization (Figures 1C, 2A, S1 and S2) and quantification of SC defects (Figure 2), high-affinity rat anti-HA (Roche 3F10, 1:2000) and mouse anti-C3G (1A8–1G2, 1:1000, gift from Hawley lab⁴⁷ were used. Cy3 anti-rat (min-X mouse, 712–165-153) and Alexa 488 anti-mouse (min-X rat, 715–545-151) secondary antibodies from Jackson ImmunoResearch were used at 1:400. Slides were mounted in SlowFade Diamond Antifade (Molecular Probes S36967, Figures 1, 2 and S1) or Prolong Gold Antifade (Molecular Probes P36930, Figures 2 and S2).

For Smc1-mCherry and Smc1-sGFP detection (Figures 4, 5, 6, S5 and S6), GFP-Booster Alexa Fluor 488 (Alpaca anti-GFP, ChromTek gb2AF488) and RFP-Booster Alexa Fluor 568 (Alpaca anti-mCherry, ChromTek rb2AF568) primary antibodies were each used at 1:500 (with no secondary antibody). To visualize centromeres, rabbit anti-CID (1:10,000, gift from Rogers lab,⁴⁸) was used followed by Cy5 anti-rabbit (Jackson ImmunoResearch 711–175-152, 1:400). Samples were incubated overnight at 4°C with primary antibodies (no rotation) and 1 hr at RT with the secondary antibody. Slides were mounted in Vectashield Vibrance (Vector Laboratories, H-1700, Figure 4) or SlowFade Diamond (Figures 5, 6, S5 and S6).

FISH

For detecting arm cohesion defects in mature Drosophila oocytes (stage 13–14), we utilized an Alexa 647-labeled Oligopaint probe (OPP122) generated by the Joyce Lab, University of Pennsylvania. This mixture of oligonucleotides (each with 80 bases of homology) contains 937 unique oligos targeting a 100kb distal region of the X chromosome (dm6, nucleotides 1,400,000–1,500,000) and was used at a final concentration of 0.50pmol/μl. A Cy3-conjugated probe (5′-Cy3-AGGGATCGTTAGCACTCGTAAT; Integrated DNA Technologies) that hybridizes to the 359-bp repeat in pericentric heterochromatin of the X chromosome was used at a concentration of 1ng/μl. Fixation, hybridization, and washes were performed as previously described^30,53 except that oocytes were subjected to a pre-denaturation step (5min @ 37°C, 3 min @ 92°C, 20 min @ 60°C, hold @ 37°C) prior to denaturation and hybridization (5min @ 37°C, 3 min @ 92°C, hold @ 37°C overnight). Oocytes were mounted on 18 mm #1.5 coverslips in Prolong Gold mounting medium and slides were allowed to cure in a box containing desiccant for at least 14 days in the dark.

Image acquisition and processing

All images were acquired using an Andor Spinning Disk confocal on a Nikon Eclipse Ti inverted microscope equipped with an ASI MS-2000 motorized piezo stage and Zyla 4.2-megapixel sCMOS camera. Image acquisition utilized Nikon Elements software (5.11.02 Build 1369), a 50 μm pinhole disk, and up to four laser lines (405, 488, 561 and 637 nm). A Nikon CFI 60X Plan Apo oil objective (NA 1.4) was utilized for immunostaining experiments, and a Nikon CFI 100X oil Plan Apo DIC objective (NA 1.45) was used for FISH experiments. All image acquisition utilized 4X frame averaging. Z-series acquisition was as follows: Smc3*-HA (0.3 μm steps, 3 μm total), C(G)3 in Figure 2 (0.1 μm steps, 4 μm total), Smc1 tag-switch (0.5 μm steps, 2 μm total), FISH (0.1 μm steps, 4 μm total). For multi-channel Z-series, an entire Z-stack was captured for one fluor before switching to the next laser. When imaging oocyte chromatin in intact ovarioles, only a specified ROI was captured, but we also collected a full-frame single-plane DAPI image of the ovariole using a Nikon CFI 20X Plan Apo oil objective (NA 0.75) which allowed us to stage individual egg chambers using size and morphological criteria.^54–56

The images in Figures 2, 3, 4 and S6 are maximum intensity projections of deconvolved confocal Z series. The images presented in Figure S2 are single optical sections of deconvolved confocal Z series. The images in Figures 1, 5, 6, S1 and S5 are maximum intensity projections of confocal Z series (no deconvolution). All images were chosen as representative examples. For immunostaining comparisons presented in each figure, identical acquisition, deconvolution and processing were used for different genotypes, including the number of optical sections included in the projection shown in the figure.

Protein extracts and immunoblotting

Young females were fattened for 3 days in food vials with yeast and males. 20 sets of ovaries were dissected in 1X PBS, flash frozen in a microtube, and stored at −80°C. Frozen ovaries were homogenized in 200μl RIPA buffer (Sigma-Aldrich R0278) with 1X HALT protease inhibitor (Thermo Fisher 87786) using a motorized disposable pestle for 30 seconds. Benzonase digestion (Millipore Sigma E1014, 250 units, 15 min) on a nutator at RT was stopped by addition of EDTA to 5mM (Thermo Fisher 1861274). The sample was filtered through a 0.22μm centrifugal filter (Millipore UFC30GV00) at 12,000 × g for 4 minutes at RT and stored in aliquots at −80°C.

Total ovary extract was separated by SDS/PAGE on a 7.5% protein gel (BioRad Stain-Free), transferred to Immobilon-P membrane (Millipore IPVH00010) and the membrane was cut using pre-stained molecular weight standards (BioRad 1610374) as a guide. After blocking overnight in 1X TBS containing 2% BSA and 5% non-fat dry milk, blots were incubated for one hour at RT in antibody incubation buffer (1X TBS + 0.1% Tween-20 [BioRad 1610781] + 5% non-fat dry milk) containing primary antibody: high-affinity rat monoclonal anti-HA (Roche 3F10, 1:10,000) or mouse monoclonal anti-alpha-tubulin (12G10 DSHB, 1:10,000) for Figure S3; affinity-purified guinea pig anti-Smc1 (²⁶, 1:1000) or rabbit anti-GFP (Invitrogen A11122, 1:1000) for Figure S4. Following washes, blots were incubated for 30 min in alkaline phosphatase conjugated secondary antibody at the following dilutions: 1:5000 AP goat anti-rat (Sigma A8438), 1:5000 AP goat anti-mouse (Invitrogen A16069), 1:3000 AP goat anti-guinea pig (Southern Biotech 6090–04), or 1:7500 goat anti rabbit (Promega S373B) and washed before applying substrate (BioRad 1705018) for five minutes. Blots were imaged using a ChemiDoc Touch system (BioRad), and bands were quantified using BioRad Image Lab software (Version 6.1).

QUANTIFICATION AND STATISTICAL ANALYSIS

Scoring for SC defects

To quantify SC defects, ovarioles stained with anti-C(3)G antibody were viewed using a 63X Plan-Apochromat (NA 1.4) objective on a Zeiss AxioImager M1 microscope equipped with a Hamamatsu ORCA^R2 digital camera controlled by Volocity Acquisition software (v6.5.1). Slides were blinded, and scoring was performed by viewing an enlarged C(3)G image on the computer monitor while slowly focusing up and down through the oocyte nucleus. Oocytes were analyzed from germarial region 3 through stage 6. The integrity of the SC was assigned to one of four categories, for which representative images are provided in Figure 2A. Long continuous threads of C(3)G signal are indicative of normal, full-length SC. Broken threads, short threads, and spots correspond to SC defects with increasing severity.

For each genotype in Figure 2B–E, a stacked bar graph (Microsoft Excel) was created to present the percentage of each SC category observed for each stage. At least 23 oocytes were scored for each stage of each genotype.

Quantification of Smc1-mCherry and Smc1-sGFP on oocyte chromosomes

Volocity Quantification (v6.5.0) was used to quantify Smc1-sGFP and Smc1-mCherry signal intensities on non-deconvolved confocal Z series. For germarial region 3 to stage 6 in Figure 4, the volume occupied by oocyte DNA (405 channel) was cropped using a free-hand tool while viewing a maximum intensity projection (5 steps, 2 μm total). Smc1 signal in the cropped volume was quantified separately for mCherry and sGFP using a threshold corresponding to 0.5 standard deviation (SD). For each volume, centromeres were identified using CID intensity thresholding (637 nm channel, 3.0 SD). For each oocyte, total intensity was calculated for the centromere volume and this value was subtracted from the total Smc1-mCherry or Smc1-sGFP signal on the DNA volume to determine the total signal intensity on chromosome arms. For oocytes in Figures 5 and 6, voxels containing oocyte DNA were identified by thresholding the DAPI signal (0.5 SD) and the Smc1-sGFP signal on DNA was quantified.

A box and whiskers plot (Microscoft Excel) is used to graph the signal intensity per voxel for each stage in Figures 4D–E, 5B and 6B. The average intensity is indicated by an “X” and median and quartiles are depicted with horizontal lines. AU = arbitrary units. N values, which correspond to the number of individual oocytes scored, are provided at the bottom of each graph. An unpaired t test (Microsoft Excel) was used to calculate P values in Figure 5B and Figure 6B. P < 0.05 was considered significant.

Scoring cohesion defects

All FISH scoring was performed blind to genotype. Following image acquisition, a MATLAB (R2022a) script was generated and used to randomly mix and rename Nikon Elements .nd2 image stacks from two or more genotypes and output a spreadsheet with the key. Blinded Z-stacks were deconvolved (Volocity Restoration v6.5.0) and visualized in three dimensions (Volocity Visualization v6.5.0) to determine the number of X chromosome arm (Alexa 647) and pericentric (Cy3) spots present on the oocyte DNA. Probe signals were considered separate if they were separated in all three dimensions by at least half the diameter of the smallest spot. 3D scoring was essential to rule out connections in the Z dimension. Two spots connected by a thread-like signal were common and were not scored as separate foci. Following scoring, the MATLAB generated key was used to determine the genotype associated with each image stack, and the percentage of oocytes with cohesion defects was calculated.

For Figures 3B and 6C, a histogram (Microsoft Excel) provides a comparison of cohesion defects in different genotypes. The total number of oocytes scored (N) for each genotype is shown within each bar. The fraction of oocytes exhibiting pericentric defects is indicated below each bar. A two-tailed Fisher’s exact test (Graphpad) was used to determine whether the incidence of arm defects in two genotypes was significantly different (P < 0.05).

KEY RESOURCES TABLE

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
High-affinity rat monoclonal anti-HA (clone 3F10)	Roche	Cat#11863423001 Lot no. 14223800 RRID:AB_390918
Mouse monoclonal anti-C3G (clone 1A8–1G2)	Hawley Lab47	N/A
Rabbit anti-CID	Rogers Lab48	N/A
GFP-Booster Alexa Fluor 488 (Alpaca anti-GFP)	ChromTek	Cat#gb2AF488 RRID:AB_2827573
RFP-Booster Alexa Fluor 568 (Alpaca anti-mCherry)	ChromTek	Cat#rb2AF568 RRID:AB_2827576
Affinity-purified guinea pig anti-Smc1	Bickel Lab26	N/A
Rabbit anti-GFP	Invitrogen	Cat#A11122 RRID:AB_221569
Monoclonal mouse anti-alpha-tubulin (12G10)	Developmental Studies Hybridoma Bank (deposited by J. Frankel and E.M. Nelson)	Cat#AB_1157911 RRID:N/A
Cy3 anti-rat (min-X mouse)	Jackson ImmunoResearch Labs	Cat#712–165-153 RRID:AB_2340667
Cy5 anti-rabbit	Jackson ImmunoResearch Labs	Cat#711–175-152 RRID:AB_2340607
Alexa 488 anti-mouse (min-X rat)	Jackson ImmunoResearch Labs	Cat#715–545-151 RRID:AB_2341099
Alkaline phosphatase goat anti-rat	Sigma-Aldrich	Cat#A8438 RRID:AB_258391
Alkaline phosphatase goat anti-mouse	Invitrogen	Cat#A16069 RRID:AB_2534742
Alkaline phosphatase goat anti-guinea pig	Southern Biotech	Cat#6090–04 RRID:AB_2796156
Alkaline phosphatase goat anti-rabbit	Promega	Cat#S373B RRID:AB_430872
Chemicals, peptides, and recombinant proteins
Formaldehyde	Ted Pella	Cat#18505
NP-40	Thermo Fisher	Cat#28324
Tween-20 (for cytology)	Thermo Fisher	Cat#28320
BSA	Fisher Scientific	Cat#BP1605–100
DAPI	Invitrogen	Cat#D1306
Poly-L-lysine	Sigma-Aldrich	Cat#P8920
SlowFade Diamond Antifade	Molecular Probes	Cat#S36967
Prolong Gold Antifade	Molecular Probes	Cat#P36930
Vectashield Vibrance	Vector Laboratories	Cat#H-1700
RIPA buffer	Sigma-Aldrich	Cat#R0278
HALT protease inhibitor	Thermo Fisher	Cat#87786
Benzonase	Millipore	Cat#E1014
EDTA	Thermo Fisher	Cat#1861274
Tween-20 (for immunoblots)	BioRad	Cat#1610781
Centrifugal filter	Millipore	Cat#UFC30GV00
7.5% protein gel	BioRad	Cat#4568023
Precision Plus Protein Dual Color Standards	BioRad	Cat#1610374
Immobilon-P membrane	Millipore	Cat#IPVH00010
AP immune star substrate	BioRad	Cat#1705018
Experimental models: Organisms/strains
Drosophila stocks used in this study, see Table S1	This study	N/A
Oligonucleotides
Alexa 647-labeled Oligopaint probe (OPP122), Mixture of 80-base oligos targeting 100kb distal region of the X chromosome (dm6, nucleotides 1,400,000–1,500,000)	Joyce Lab, University of Pennsylvania	N/A
Cy3-conjugated probe (5′-Cy3-AGGGATCGTTAGCACTCGTAAT) hybridizes to 359-bp repeat in pericentric heterochromatin of the X chromosome	Integrated DNA Technologies	N/A
Smc3* Fwd: 5’GACAAGACGCGCCGCACGCTAGAATACATTCGGTATGAGACCGAGCTGAAGGACA	Integrated DNA Technologies	N/A
Smc3* Rev: 5’TGTCCTTCAGCTCGGTCTCATACCGAATGTATTCTAGCGTGCGGCGCGTCTTGTC	Integrated DNA Technologies	N/A
HA stop Fwd: 5’GCTTCTAGCGAAAAAGTGTAGACGGTATCGATAAGCTTG	Integrated DNA Technologies	N/A
HA stop Rev: 5’CAAGCTTATCGATACCGTCTACACTTTTTCGCTAGAAGC	Integrated DNA Technologies	N/A
Smc3-WT Fwd: 5’AGCACTGGTACCGCGGCCGCAATAAGATGCACATCAAGCAG	Integrated DNA Technologies	N/A
Smc3-WT Rev: 5’ACTGATCTCGAGGCGGCGTGGGTGCTGTCG	Integrated DNA Technologies	N/A
Smc3-KK Fwd: 5’CGCGTGATTGGCGCCAGAAGGGACCAGTACTTCC	Integrated DNA Technologies	N/A
Smc3-KK Rev: 5’GGAAGTACTGGTCCCTTCTGGCGCCAATCACGCG	Integrated DNA Technologies	N/A
Recombinant DNA
Clone RE14758 (Smc3 cDNA)	DGRC	Stock 8520 RRID:DGRC_8502
pUASP-attB	Gift from Hawley lab49	RRID: N/A
w+ attB vector	Addgene (donated by Jeff Sekelsky)	Plasmid# 30326 RRID: Addgene_30326
Software and algorithms
Volocity Visualization, Restoration, and Quantitation	Quorum Technologies, Canada	Version 6.5.0 https://www.volocity4d.com/download
Volocity Acquisition (for epifluorescence imaging)	Quorum Technologies, Canada	Version 6.5.1 https://www.volocity4d.com/volocity-acquisition
Nikon Elements (for spinning disc confocal imaging)	Nikon	Version 5.11.02 Build 1369
MATLAB	Mathworks	Version R2022a https://www.mathworks.com/products/new_products/release2022a.html
BioRad Image Lab	BioRad	Version 6.1 https://www.bio-rad.com/en-us/product/image-lab-software?ID=KRE6P5E8Z
Microsoft Office	Microsoft	Version 16.84
Affinity Designer	Affinity	Version 1.10.6.1665 https://affinity.serif.com/en-us/designer/
Other
Chemi Doc Touch Imaging system	BioRad	Serial no. 732BR0120

Highlights.

• Chromatin-associated cohesin turns over extensively in prophase oocytes.

• A rejuvenation program establishes new cohesive linkages after meiotic S phase.

• New linkages do not require Smc3 acetylation at canonical lysines.