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bioRxiv
BIORXIV
bioRxiv
2692-8205
Cold Spring Harbor Laboratory

39229242
10.1101/2024.08.21.608908
preprint
1
Article
Distinct checkpoint and homolog biorientation pathways regulate meiosis I in Drosophila oocytes
Shapiro Joanatta G.
Changela Neha
Jang Janet K.
Joshi Jay N.
http://orcid.org/0000-0002-8990-1168
McKim Kim S.
Waksman Institute and Department of Genetics, Rutgers, the State University of New Jersey, Piscataway, New Jersey, United States of America.
Corresponding author: Kim S. McKim, 190 Frelinghuysen Road, Piscataway, NJ-08854, mckim@waksman.rutgers.edu
21 8 2024
2024.08.21.608908https://creativecommons.org/licenses/by/4.0/ This work is licensed under a Creative Commons Attribution 4.0 International License, which allows reusers to distribute, remix, adapt, and build upon the material in any medium or format, so long as attribution is given to the creator. The license allows for commercial use.
nihpp-2024.08.21.608908.pdf
Mitosis and meiosis have two mechanisms for regulating the accuracy of chromosome segregation: error correction and the spindle assembly checkpoint (SAC). We have investigated the function of several checkpoint proteins in meiosis I of Drosophila oocytes. Evidence of a SAC response by several of these proteins is found upon depolymerization of microtubules by colchicine. However, unattached kinetochores or errors in biorientation of homologous chromosomes does not induce a SAC response. Furthermore, the metaphase I arrest does not depend on SAC genes, suggesting the APC is inhibited even if the SAC is silenced. Two SAC proteins, ROD of the ROD-ZW10-Zwilch (RZZ) complex and MPS1, are also required for the biorientation of homologous chromosomes during meiosis I, suggesting an error correction function. Both proteins aid in preventing or correcting erroneous attachments and depend on SPC105R for localization to the kinetochore. We have defined a region of SPC105R, amino acids 123-473, that is required for ROD localization and biorientation of homologous chromosomes at meiosis I. Surprisingly, ROD removal, or “streaming”, is independent of the dynein adaptor Spindly and is not linked to the stabilization of end-on attachments. Instead, meiotic RZZ streaming appears to depend on cell cycle stage and may be regulated independently of kinetochore attachment or biorientation status. We also show that dynein adaptor Spindly is also required for biorientation at meiosis I, and surprisingly, the direction of RZZ streaming.

Author Summary

The Spindle Assembly Checkpoint (SAC) is known to delay cell cycle progression until chromosomes are properly attached to microtubules. Meiotic cells often have modified cell cycle phases, and natural arrest points such as metaphase I in Drosophila. We show that in Drosophila oocytes, the SAC is sensitive to loss of microtubules, but not sensitive to a variety of kinetochore attachment errors. Thus, the function of the SAC appears to be limited to monitoring oocyte spindle assembly, and not required for accurate chromosome segregation. However, two of the SAC genes, rod and Mps1, are required for the biorientation of homologous chromosomes during meiosis I, suggesting an error correction function. Rod is part of the RZZ complex and is notable for its property of streaming off the kinetochores. However, our results show that streaming off the kinetochore may not contribute to RZZ regulation of microtubule attachments, and only be associated with SAC function. Instead, the establishment of stable end-on attachments may occur while RZZ is still present at kinetochore. We suggest that RZZ interacts with multiple motors to promote bidirectional movement of kinetochores along microtubules, which allows chromosomes to find and attach to the correct pole.
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pmcIntroduction

Errors during meiotic chromosome segregation can result in inviable gametes leading to spontaneous abortions and infertility, or viable offspring with chromosomal abnormalities. Several mechanisms exist in dividing cells to prevent aneuploidy resulting from improper kinetochore-microtubule (KT-MT) attachments. The spindle assembly checkpoint (SAC) delays anaphase onset until attachments form between kinetochores and microtubules [1, 2]. Unattached kinetochores generate a signal involving SAC proteins such as MAD1, MAD2, BUB1, and BUB3 that have a global effect on all chromosomes and cell cycle progression. In contrast, error correction (EC) is specific to individual kinetochores whereupon interactions between microtubules and properly bi-oriented kinetochores are stabilized while those with improperly oriented kinetochores are weakened [3]. Multiple models have been proposed to explain how improperly oriented kinetochores are identified, involving both tension-dependent and - independent mechanisms, and how they are corrected, including phosphorylation of kinetochore proteins to destabilize attachments [4, 5].

Error rates are higher in human oocytes than in spermatocytes, which has been attributed to several causes [6-8]. One hypothesis is that the SAC is weak or inefficient [7, 9-11]. Indeed, Xenopus oocytes may not have a functional SAC [12, 13]. The observation that mammalian oocytes progress through anaphase with segregation errors may be a result of a weak checkpoint [14, 15]. The reason for a weak SAC in oocytes is not known, although possible contributing factors include the large size of oocytes or the lack of centrosomes [9, 16-19]. The SAC may also have other functions in oocytes, such as regulating the development of the oocytes and the timing of meiotic progression [10].[1, 2].

Drosophila oocytes are an excellent model for studying meiosis in females, but the SAC has not been examined. Drosophila oocytes have kinetochores that include the KMN complex (KNL1-MIS12-NDC80) [20]. Biorientation of homologous chromosomes involves types of kinetochore-microtubule attachments, lateral attachments that depend on KNL1/SPC105R and end-on attachments that depend on NDC80 [21, 22]. To understand the contributions of SAC and EC to biorientation in Drosophila oocytes, we characterized the SAC response during the first meiotic division, investigated how the SAC proteins are recruited to the kinetochores, and tested several SAC proteins for functions in meiotic EC. Our data suggest that the SAC is not sensitive to kinetochore attachment status and might be silenced prior to the metaphase I arrest. Furthermore, among seven SAC proteins, only two, ROD of the ROD-ZW10-Zwilch (RZZ) complex and MPS1, are required for biorientation of meiotic homologous chromosomes at metaphase I.

We examined the role of RZZ in Drosophila oocytes because it has not been studied before in meiosis. Previous studies in C. elegans embryos and Drosophila neuroblasts suggest that RZZ prevents errors by inhibiting premature stabilization of interactions between the ends of microtubules and NDC80 [23, 24]. RZZ also has the property of streaming off the kinetochore, which could alleviate the inhibition of end-on attachments [24-26]. We have shown that the RZZ complex is required for accurate homolog biorientation in meiosis I, which could indicate a role for the RZZ complex in delaying the formation of end-on attachments. We also show that Spindly, a dynein adaptor for RZZ, is also required for homolog biorientation in meiosis I. However, RZZ streaming in oocytes is not required for, nor does it depend on, the formation of end-on attachments. We propose that RZZ streams in response to the silencing of the SAC prior to the metaphase I arrest. Biorientation, however, depends on an interaction between RZZ, Spindly, and possibly other motor activities, that regulate chromosome movement and attachment status.

Results

Analysis of the SAC protein localization in oocytes

The localization of SAC proteins was examined in Drosophila oocytes. Mature Drosophila oocytes, known as stage 14, arrest at metaphase I with the chromosomes clustered together in a single mass, or karyosome, and the centromeres of homologous chromosomes oriented towards opposite poles. Several SAC proteins were detected at kinetochores in stage 14 oocytes, including BUBR1, BUB3, MPS1 and ROD (Figure 1A). In addition to its kinetochore localization, ROD localized to the spindle region between the pole and the centromeres (Figure 1). This redistribution of ROD from the kinetochores to the spindle is termed “streaming” and has been previously observed in mitotic cells [25, 27, 28]. Another RZZ subunit, ZW10 tagged with HA, overlapped with RODGFP staining, validating that RODGFP provides an accurate representation of the RZZ complex’s localization (Figure S 1A).

In contrast to the above proteins, we did not detect MAD1 in stage 14 oocytes (Figure 1A). Therefore, we attempted to increase SAC signaling by incubating stage 14 oocytes in colchicine for 60 minutes to remove nearly all microtubules. Following this treatment, robust kinetochore localization of MAD1 was observed in all oocytes (Figure 1B). Because colchicine induced a change in MAD1 localization, we tested if the localization of other checkpoint protein was increased by loss of microtubules. BUBR1 and MPS1 intensity increased in colchicine, suggesting their levels are sensitive to the SAC (Figure 1C). We did not, however, observe a SAC response with BUB3 localization.

A more dramatic response was observed with ROD. In colchicine-treated oocytes, ROD expanded around the kinetochores (Figure 1B). In some cases, ROD expanded to surround the karyosome and individual kinetochore foci could no longer be observed. This behavior of the RZZ complex likely corresponds to expansion of the fibrous corona, observed in miotic cells that lack kinetochore-microtubule (KT-MT) attachments [25, 29]. This expansion is thought to facilitate microtubule capture by increasing the surface area of the kinetochore which can contact the microtubules. Thus, in Drosophila meiotic oocytes, the fibrous corona expands when microtubules are absent.

Activation of the SAC is also associated with elevated Aurora B activity [30]. In Drosophila oocytes, the chromosomal passenger complex (CPC) is recruited to the chromosomes but then moves to the central spindle in prometaphase I oocytes [31]. CPC proteins are not detected at the meiotic centromeres in fixed samples [31, 32]. In colchicine-treated oocytes, however, the CPC component INCENP localized to the chromosomes and the centromeres (Figure 1B, C). These results suggest that, like MAD1, colchicine treatment causes increased localization of the CPC to the centromeres.

Effect of kinetochore-microtubule attachments on the SAC response

In all colchicine-treated oocytes, MAD1 localization was consistent and observed on all kinetochores, suggesting a global SAC response to the absence of microtubules. To investigate the mechanism of SAC activation, we examined MAD1 localization in oocytes where end-on attachments or tension was disrupted. NDC80 is required for end-on attachments, so nearly all attachments in Ndc80RNAi oocytes are lateral [22]. MAD1 was not detected in Ndc80RNAi oocytes unless they were treated with colchicine, suggesting that the SAC is insensitive to the loss of end-on attachments (Figure 2A-C). We also examined MAD1 in a mei-218 mutant (mei-218null), which has a high error rate and lacks tension on the kinetochores due to the absence of chiasmata [33]. MAD1 was not detected in mei-218null oocytes unless they were treated with colchicine, suggesting that loss of tenson does not induce a SAC response (Figure 2A, B, D). The absence of MAD1 in the untreated mei-218null oocytes is consistent with the prior observation that oocytes lacking chiasma bypass the metaphase I arrest and precociously enter anaphase in stage 14 oocytes [33]. These results show that the SAC is only activated in the absence of microtubules and does not respond to the lack of stable attachments, biorientation errors, or tension.

Localization of checkpoint proteins in Drosophila oocytes depends on SPC105R

The outer kinetochore protein KNL1/SPC105R is involved in the recruitment of several checkpoint proteins in mitotic cells [34, 35]. Therefore, we investigated the role of SPC105R, the Drosophila homolog of KNL1, in the recruitment of MAD1, ROD, MPS1, and the CPC to kinetochores in Drosophila oocytes. To test if SPC105R is required for the recruitment of INCENP and MAD1, we treated Spc105RRNAi oocytes with colchicine. Colchicine-induced kinetochore localization of both INCENP and MAD1 depended on SPC105R (Figure S 2A-D). However, both still localized to the chromosomes. In the absence of SPC105R, MAD1 moved to the pericentric regions adjacent to the centromeres, while INCENP localized to all chromatin and occasionally to regions adjacent to the chromatin. The observation that MAD1 localized in colchicine-treated oocytes depleted of SPC105R suggests that the oocyte can respond to the loss of microtubules even in the absence of the outer kinetochore.

We also examined localization of INCENP and MAD1 in Spc105R deletion mutants to investigate the domains of SPC105R required for the SAC response (Figure S 3). Because Spc105R is an essential gene, we used a combination of Spc105RRNAi and RNAi-resistant transgenes to examine mutants in oocytes. The control for each mutant was an RNAi-resistant wild-type transgene, Spc105RB (Figure S 3). MAD1 kinetochore localization was reduced in mutants such as Spc105RC or Spc105RΔM that deleted motifs (MELT, KI, and ExxEED) in the central region of SPC105R (Figure S 2A, B). INCENP kinetochore localization was also lost from the kinetochore in the majority of Spc105RC or Spc105RΔM oocytes, but due to the diffuse chromatin localization, this was not reflected in significant differences in intensity (Figure S 2C, D). These results suggest that the MELT-like, KI-like, and ExxEED repeats are involved in recruiting MAD1 and the CPC to the kinetochore as part of the SAC response.

MPS1 localization depends on SPC105R, as shown by its drastic reduction in Spc105RRNAi oocytes (Figure S 4A, B). Ndc80RNAi oocytes also had a decrease in MPS1 localization, but this was less severe than Spc105RRNAi oocytes, indicating that the majority of MPS1 is recruited by SPC105R (Figure S 4A, C). Similarly, the Spc105RC, Spc105RΔM, and Spc105RΔExxEED mutants had significant decreases in MPS1 localization, but not to the same degree as the Spc105RRNAi oocytes (Figure S 4B, D). Each of these mutants retains the C-terminal domain of SPC105R that recruits NDC80 [22]. Thus, these mutants may recruit more MPS1 than Spc105RRNAi oocytes because NDC80 recruits some MPS1. Therefore, these results suggest that MPS1 is recruited by domains within both SPC105R and NDC80.

ROD localization also depends on SPC105R, as shown by its reduction in Spc105RRNAi oocytes (Figure 3A, B). In comparison to the other SAC proteins, recruitment of RZZ depends on a smaller and specific domain of SPC105R (Figure S 3, Figure 3, D). Oocytes with a deletion of everything except the KT binding domain (Spc105RC) or a deletion of all the sequences between the N-terminal and C-terminal domains (MELT-like, KI-like, and ExxEED, termed Spc105RΔM) had almost no RODGFP localization, similar to Spc105RRNAi oocytes. Furthermore, a deletion of the MELT-KI (Spc105RΔMELT-KI), but not the ExxEED domain, had reduced ROD localization. These results suggest that ROD is recruited by the MELT-KI region. Smaller deletions of only the KI and MELT domains (Spc105RΔKI and Spc105RΔMELT) resulted in significantly reduced RODGFP localization at the kinetochores, although these intensities were higher than in the Spc105RΔMELT-KI deletion. This suggests that ROD is recruited by both the MELT and KI regions.

We also tested if the localization of ROD and MPS1 depended on other SAC proteins. ROD (Figure 4A, B) and MPS1 (Figure 4C, D) localization was significantly reduced in both Bub3RNAi and BubR1RNAi oocytes. The more significant reductions that were observed with Bub3 knockdown could be due to less efficient BubR1RNAi (see Methods), or redundancy between BUBR1 and BUB1. We also tested the effect of Cyclin A knockdown because it may regulate MPS1 localization [36]. CycARNAi oocytes had a significant reduction in MPS1 localization, although not as strong as Bub3RNAi, and a less significant reduction in ROD localization. Furthermore, MPS1 and ROD localization was inter-dependent, although there was a milder reduction of ROD in Mps1RNAi. These results suggest ROD and MPS1 localization depends on both SAC proteins like BUB3 and Cyclin A.

RZZ and MPS1, but not other SAC proteins, are required for meiosis I biorientation

To determine which SAC genes are required for biorientation during meiosis I, we used fluorescent in-situ hybridization (FISH) probes against the pericentromeric regions of the 2nd, 3rd, and X chromosomes. Biorientation was defined as two homologous centromeres at opposite ends of the karyosome (Figure 5A). A mono-orientation defect was defined as the two homologous centromeres being in the same half of the karyosome. Oocytes from females expressing rodRNAi or Mps1RNAi had increased chromosome mono-orientation (Figure 5B). The results with Mps1 are consistent with previous studies showing nondisjunction using fertile allele combinations of Mps1 [37, 38]. We also tested CycARNAi oocytes, and consistent with a previous report [39], found an elevated level of biorientation defects (Figure 5B).

In contrast, BubR1RNAi, Bub1RNAi, and Bub3RNAi oocytes had insignificant frequencies of mono-orientation (Figure 5B). Similarly, we have previously shown that flies with knock down of Bub1 by RNAi are fertile and do not have increased nondisjunction [31]. BUBR1 is required for sister chromatid cohesion protection in Drosophila, however, this may have not been detected by our FISH assay at metaphase I [22, 40]. Mutants of mad1 and mad2 are viable and fertile [41, 42], so we used genetic crosses to measure the frequency of nondisjunction (see Methods). Neither mutant had a significant increase in nondisjunction (Table 1). We also used a balancer chromosome to suppress crossing over on the X chromosome (Bwinscy), thus testing the role of mad1 and mad2 in an achiasmate system [43]. Only a mild increase in nondisjunction was observed. These results suggest that most of the SAC proteins are not required for biorientation of homologous chromosomes at metaphase I, while the RZZ complex and MPS1 may be involved in error correction in addition to their roles in the meiotic SAC.

Stage 14 oocytes arrest in metaphase I [33, 44]. If the metaphase I arrest depends on the SAC, we would predict that SAC mutants would precociously bypass this arrest and have increased oocytes with split karyosomes. However, this was not observed in most oocytes, suggesting the metaphase I arrest does not depend on SAC activity (Figure 5C, D). The most striking exceptions were rodRNAi and the mad2 mutant. The split karyosome phenotype is less than observed in mutants lacking crossing over [22, 33], suggesting these genes could have a minor role in the metaphase I arrest, or in the case of Rod, the precocious anaphase could result from the bi-orientation defects (see below).

ROD streaming is independent of end-on attachments

MSP1 and ROD may have two distinct functions, one in the SAC and another for biorientation. A striking feature of RZZ is its streaming off the kinetochores, but how streaming contributes to kinetochore attachments and biorientation in meiosis is not known. A small fraction of wild-type oocytes have no ROD streaming, which are likely oocytes in early prometaphase [45]. If streaming is in response to proper attachment of kinetochores to microtubules, we predicted that RZZ would not stream if errors were present, or if end-on attachments could not form. To determine if biorientation errors blocked streaming, we examined ROD in mei-218null oocytes, in which 90% of crossovers are absent, resulting in frequent chromosome segregation errors [46]. In the absence of crossovers, we still observed precocious anaphase and ROD streaming (Figure 6A,B). Thus, streaming occurs even in the absence of tension and in the presence of attachment errors.

To determine if attachment status regulates streaming, we examined ROD in Ndc80RNAi oocytes, which lack end-on attachments [21]. Despite the lack of end-on attachments, ROD streamed in all Ndc80RNAi oocytes, and there was no effect on ROD accumulation at the kinetochores (Figure 6A-C). Furthermore, oocytes lacking protein phosphatase 2A-B56 (PP2A-B56), which is also required for the creation of end-on attachments [47], still exhibited ROD streaming (Figure 6A-C). Indeed, there may even be a reduction of ROD at the kinetochore in PP2A-B56RNAi oocytes. These results demonstrate that end-on attachments are not required for the initiation of RZZ streaming.

ROD is not required for lateral or end-on attachments

Microtubule nucleation from the kinetochore could depend on RZZ [48]. Thus, the biorientation defects observed in the rodRNAi oocytes could be caused by the inability to create lateral or end-on attachments, or the creation of inaccurate attachments. If ROD is necessary for making end-on attachments, we would expect that oocytes depleted of rod alone would have lateral attachments. However, most attachments in rodRNAi oocytes were end-on (Figure 7A, B). Interestingly, there was an increase of lateral attachments in rodRNAi compared to wild-type oocytes. This could be explained if rodRNAi oocytes are delayed in making end-on attachments, possibly due to frequent biorientation errors. To test whether ROD was necessary for forming lateral attachments, we knocked down both rod and Ndc80 (rodRNAi, Ndc80RNAi). Similar to Ndc80RNAi, we observed that rodRNAi, Ndc80RNAi oocytes had primarily lateral attachments, suggesting that ROD is not required for making lateral attachments (Figure 7A, B).

As noted above, rodRNAi oocytes have an elevated frequency of separated karyosomes. This phenotype was not observed in rodRNAi, Ndc80RNAi oocytes, suggesting that separated karyosome phenotype is caused by the formation of end-on attachments is required for the (Figure 7A, C).

ROD localization to the kinetochore may delay, but does not prevent, end-on attachments

Because ROD is not required for making lateral or end-on attachments, we reasoned that it could be required for regulating the transition from lateral to end-on attachments. This would align with a model proposed in mitotic cells in which high levels of RZZ at the kinetochores inhibit NDC80 from prematurely establishing end-on attachments [23, 24]. In this model, RZZ streaming allows for the establishment of end-on attachments. If this was true in meiosis, we would expect oocytes in which ROD was exclusively on kinetochores and did not stream to have lateral attachments. This ROD behavior was observed in two Spc105R mutants. A deletion of the ExxEED repeats (Spc105RΔExxEED) or the N-terminal region (Spc105RΔN) had higher ROD intensities at the kinetochores (Figure 3) and less streaming than in Spc105RB oocytes (Figure 7D, E). However, oocytes with ROD only on kinetochores still exhibited end-on attachments (Figure 7D). These results suggest that the N-terminal region and ExxEED repeats of SPC105R may regulate streaming, and that RZZ streaming is not required for the creation of end-on attachments in Drosophila oocytes.

To further test whether end-on attachments are inhibited when the RZZ complex is localized to the kinetochore, we stably targeted RZZ to the kinetochore. The rationale for this experiment is that the recruitment of the RZZ components are interdependent [49]. We fused Zw10 to the KT-binding C-terminal domain of Spc105R (Zw10-Spc105RC), along with three copies of the HA epitope at the C-terminal end (Figure S 3). The ZW10-SPC105RC fusion protein was targeted to the kinetochores (Figure 7F). When Zw10-Spc105RC was combined with an shRNA targeting the endogenous Spc105R (Spc105RRNAi), loss of endogenous SPC105R was confirmed cytologically using an antibody against the first 400 amino acids of SPC105R, which is absent in the fusion construct (Figure S 5A). In addition, ROD localized to the kinetochores and did not stream in Zw10-Spc105RC oocytes, indicating that the RZZ complex was restricted to the kinetochores (Figure S 5B). RODGFP did not stream even when there was endogenous SPC105R present at the kinetochores. Thus, the ZW10-SPC105RC fusion protein prevented endogenous RZZ streaming from kinetochores. Zw10-Spc105RC oocytes exhibited an increase in lateral attachments compared to Spc105RC oocytes (Figure 7G). However, the majority of attachments were still end-on (~60%), indicating that RZZ may delay, but does not prevent, end-on attachments. Therefore, in agreement with the two Spc105R mutants (Figure 7D), streaming of RZZ is not required to establish end-on attachments in Drosophila oocytes. Indeed, end-on attachments occur while RZZ is localized to the kinetochore.

Spindly regulates ROD streaming

Streaming is proposed to be dependent on the microtubule motor protein Dynein [50]. To further examine the role of RZZ streaming and the interaction with Dynein in meiosis, we investigated two dynein-associated proteins previously implicated in RZZ removal from the Drosophila kinetochores: Spindly [51], and NudE [52]. We performed FISH on stage 14 oocytes to determine whether these two genes are required for chromosome biorientation (Figure 8A, B). There was only a slight increase in chromosome mono-orientation in nudERNAi oocytes compared to wild-type (9% and 0%, respectively). However, 35% of chromosomes were mono-oriented in SpindlyRNAi oocytes, suggesting that Spindly is necessary for accurate meiotic chromosome segregation.

To determine whether these proteins regulate streaming, we characterized ROD behavior in oocytes depleted of Spindly or nudE (Figure 8C-E). Both SpindlyRNAi and nudERNAi oocytes exhibited ROD streaming (Figure 8D). This was surprising for Spindly, which has been shown to be required for streaming in Drosophila S2 cells [51]. However, SpindlyRNAi oocytes also exhibited unusual behavior, notably ROD streaming in the central spindle region between homologous centromeres in approximately 50% of oocytes (Figure 8E, Figure S 6A). Thus, Spindly may be required for proper streaming towards the spindle pole. To confirm that ROD was being efficiently removed and not accumulating at the kinetochores, we measured the intensity of ROD in SpindlyRNAi and nudERNAi oocytes (Figure 8F). Both RNAi oocytes had ROD levels similar to those in wild-type oocytes, indicating that ROD can still stream from kinetochores without NudE or Spindly. Therefore, while neither protein is required for RZZ streaming, Spindly may be required for RZZ to stream towards the spindle poles.

RZZ is proposed to recruit Spindly to the kinetochore, which is followed by streaming of both Spindly and RZZ [51, 53]. To examine the localization of Spindly on the meiotic spindle, we used previously described Spindly-GFP fusion transgenes [54]. Oocytes expressing wild-type Spindly had weak localization throughout the spindle (Figure S 6B). We also examined Spindly mutants with deletions of domains implicated in streaming: the N-terminus (a deletion of amino acids 1-228) and the Spindly Box (a deletion of amino acids 229-250). Both mutants had a dominant sterility phenotype when expressed in oocytes using matα, suggesting that these mutants interfere with the activity of the wild-type Spindly protein. In contrast to the wild-type oocytes, which had weak localization throughout the spindle, both mutants had pronounced localization near the centromeres. SpindlyΔN localized to the kinetochores and spindle, while SpindlyΔSB primarily localized to the kinetochores (Figure S 6B, C). These results suggest that the Spindly Box domain is important for removal of Spindly from the kinetochores. The robust localization of these mutants was then used to test if ROD is required for Spindly localization, as shown in mitotic cells [51]. Indeed, Spindly was absent in rodRNAi oocytes, indicating ROD is required for Spindly localization in oocyte meiosis (Figure S 6C, D).

Discussion

The function of the SAC in oocytes

We analyzed the localization and meiotic phenotypes of several SAC proteins in Drosophila oocytes. Increased localization of MAD1, BUBR1, INCENP, and ROD at the kinetochores following loss of microtubules indicated Drosophila oocytes have a functioning SAC. This SAC response was observed even in the absence of a kinetochore. However, a SAC response was not observed following loss of microtubule attachments to the kinetochores, the lack of tension, or the presence of multiple biorientation errors. Thus, the function of the SAC in oocytes may be to promote spindle assembly, rather than proper microtubules attachments to kinetochores [28].

Only two of the SAC proteins we tested were required for biorientation of homologous chromosomes at meiosis I, MPS1 [37, 38] and ROD, suggesting that the SAC is not required for error correction. Similarly, C. elegans may correct attachment errors using a mechanism that does not rely on the SAC [55]. The SAC may have a more prominent role in chromosome segregation in mouse oocytes that arrest in meiosis II, where increased aneuploidy has been observed with loss of SAC proteins [56, 57]. However, the evidence comes from a variety of assays that may not be comparable to our FISH experiments, and one study concluded that Mad2 is dispensable for accurate meiotic chromosome segregation except in response to stress [58].

In most cases, the loss of SAC proteins in Drosophila oocytes does not result in progression into anaphase (Figure 5) or a rise in APC activity [59]. These observations suggest the SAC may not regulate the APC in metaphase I-arrested oocytes. Instead, the metaphase I arrest in Drosophila oocytes may be maintained by a SAC-independent mechanism relying on chiasmata and cohesion, with progression into anaphase I depending on cohesion release. This would explain why Drosophila oocytes lacking chiasmata and tension bypass metaphase I arrest and progress into anaphase I [33, 60].

Furthermore, several observations suggest that the SAC is silenced in oocytes, possibly to allow for developmental regulation of the metaphase I arrest, with progression into anaphase occurring only after oocytes enter the oviduct. Developmentally regulated SAC silencing would explain why ROD streams in wild-type metaphase I-arrested oocytes, even in the absence of end-on attachments or the presence of multiple biorientation errors in Ndc80RNAi and PP2A-B56RNAi oocytes [61]. Similarly, MAD2 localizes to centromeres in early (stage 13) but not late (stage 14) stage oocytes [45], and the metaphase I spindle has some anaphase-like characteristics such as high concentrations of the CPC on the central spindle [62]. Yeast meiotic cells silence their SAC to prevent prolonged arrest, indicating this may be a conserved feature of meiotic cells [63].

Recruitment of checkpoint proteins during meiosis

Our results are consistent with the known role of KNL1/SPC105R in the recruitment of checkpoint proteins to the kinetochore [1]. We previously showed that BUBR1 is recruited by both the MELT-KI and C-terminal regions of SPC105R [22]. MAD1, INCENP, and probably BUB3 depend on the MELT-KI and ExxEED repeat regions of SPC105R, which correspond to the repeated MELT motifs found in vertebrate KNL1. In the absence of SPC105R, recruitment of MAD1 shifts to the pericentromeric regions. Thus, some components of the SAC do not depend on SPC105R, and there is a SPC105R-independnet SAC response. Human cells may also have two SAC pathways, and one is active in the absence of KNL1 [64]. Thus, centromere components may recruit SAC proteins. In Drosophila, this could be facilitated by ZW10, which interacts with and is recruited by centromere protein CAL1 [65].

Multiple functions and recruitment factors for MPS1 and RZZ

MPS1 was originally discovered with the mutation ald as being required for meiotic chromosome segregation [66], and subsequent studies have characterized its role in Drosophila meiosis [37, 38] and mouse oocytes [67]. While MPS1 has been shown to be recruited by NDC80 [68-71], two observations show that a significant amount of MPS1 is also recruited by SPC105R. First, we observed a more severe decrease in MPS1 localization upon Spc105R depletion than Ndc80 depletion. Second, we observed a decrease in MPS1 localization in Spc105R mutants that still recruit NDC80. These data suggest that MPS1 is recruited by both NDC80 and SPC105R in Drosophila oocytes. The SPC105R-dependent MPS1 could function in the SAC while NDC80-dependent MPS1 could regulate attachments [68, 72].

Like MPS1, the RZZ complex has roles in error correction and in the SAC. Like MPS1, our data suggest there are two separate RZZ populations: one that is recruited by BUB3 and BUBR1 and is involved in the SAC and a second that is recruited by the MELT-KI domains of SPC105R and is important for EC. The SAC population of ROD that depends on BUB3 could recruit other checkpoint proteins like MAD1 and MAD2 [73, 74]. These conclusions are based on the observation that unlike rodRNAi, Bub3RNAi and BubR1RNAi oocytes do not have biorientation defects, suggesting that BUB3 and BUBR1 do not recruit the population of RZZ which is involved in error correction. Furthermore, there is a substantial amount of RODGFP remaining after Bub3 and BubR1 are knocked down, arguing that they do not recruit the total population of RZZ at kinetochores. However, the Spc105RΔMELT-KI mutant (deleting amino acids 154-840) abolishes nearly all RODGFP localization and does have biorientation defects [22]. Therefore, the MELT-KI domains of SPC105R are likely involved in recruiting the populations of RZZ involved in EC, independently of BUB3.

The MELT-KI region of SPC105R is highly conserved [75]. Indeed, our results refine observations made in HeLa cells showing that the N-terminal domain of KNL1 recruits RZZ [76, 77]. In Drosophila cell lines, a region within the KI-like domain (amino acids 266-384) and BUBR1 are required for RZZ recruitment to kinetochores [78]. Interestingly, this study also showed that the recruitment of RZZ by SPC105R could promote chromosome movement towards the spindle poles. Like this study, we found the KI domain recruits the majority of RZZ. It remains to be determined, however, why both the MELT and KI domains are required to recruit RZZ and for biorientation in meiosis I [22].

ROD and MPS1 localization was reduced in CycARNAi oocytes, and CycARNAi oocytes have biorientation defects in Drosophila oocytes [39]. Indeed, Cyclin A has been shown to regulate KT-MT attachments in mammalian cells [79, 80] and in Drosophila in mitotic cells, Cyclin A/CDK1 activates Polo kinase, which has recently been shown to regulate Spindly and prevent it from inhibiting RZZ [24]. One explanation for these results is that Cyclin A is required for the functions of ROD and MPS1 associated with ensuring the accuracy of biorientation.

The role of RZZ localization in attachment stabilization

Previous studies in C. elegans, Drosophila, and human mitotic cells have suggested a two-part model for how RZZ regulates KT-MT attachments [23, 24, 81, 82]. First, RZZ interacts with NDC80 to inhibit end-on attachments. Second, Spindly-dependent RZZ streaming alleviates RZZ inhibition to allow for the formation of end-on attachments. Three of our observations are consistent with this hypothesis. First, knockdown of Spindly resulted in a decrease in end-on and an increase in lateral attachments. This would be expected if Spindly negatively regulates RZZ. Second, Zw10 fused to the C terminus of Spc105R led to complete RZZ retention on the kinetochores and increased lateral attachments. Third, we observed an increase in karyosome separation in rodRNAi oocytes, which could be due to precocious stabilization of end-on attachments [22, 83]. These results suggest that the RZZ complex at the kinetochore has a role in preventing or delaying end-on attachments.

However, some of our results are not consistent with this model. Oocytes expressing Zw10-Spc105RC primarily had end-on attachments. A similar observation was made with two mutants (Spc105RΔExxEED and Spc105RΔN) that had reduced ROD streaming but still formed end-on attachments. Therefore, if KT-localized RZZ inhibits end-on attachments, this can be overridden in oocytes. Finally, ROD streaming was observed in the absence of end-on attachments. Thus, physical removal of RZZ (or streaming) may not be required for the formation of end-on microtubule attachments, and alternative mechanisms for regulating this population of RZZ may exist. For example, a conformational change in RZZ may affect its regulation of attachments [23]. Understanding the relationship between the localization of the RZZ complex and its effects on microtubule attachments is crucial for analyzing its role in promoting accurate chromosome segregation during female meiosis.

The role of Spindly in meiotic RZZ streaming and homolog biorientation

Spindly has a conserved role as a Dynein adaptor [84], and we have shown that RZZ recruits Spindly to kinetochores in Drosophila oocytes. Spindly has been shown in multiple organisms to recruit Dynein, resulting in removal of the RZZ complex from the kinetochores to stabilize microtubule attachments [24, 53, 81, 85]. This activity has been used to explain why depletion of RZZ can suppress defects in mitotic cells depleted of Spindly [23, 24, 81]. However, the role of Spindly in meiosis has not been reported.

Spindly is required for biorientation of homologous chromosomes during oocyte meiosis. The failure to remove RZZ from the kinetochores could be the reason for the meiotic biorientation defect. Contrary to the expectations from data in mitotic cells described above, however, knocking down Spindly did not abolish ROD streaming. Instead, we found that ROD displayed abnormal behaviors in oocytes depleted of Spindly, such as streaming in the opposite direction and localizing in the central spindle region between kinetochores. Therefore, it is likely that other proteins are involved in the removal of RZZ from the kinetochores in oocytes.

In the absence of Spindly in HeLa cells, alternative mechanisms are activated to remove kinetochore proteins [86]. One candidate is CENP-E, which is a plus-end directed motor required for the fibrous corona [87], and is required for homolog biorientation in Drosophila oocytes [21]. Given that RZZ can move SPC105R oligomers [88], and our observation that streaming off the kinetochore may not affect attachment status, an important role of RZZ may be to move kinetochores. Rather than being necessary for the direct removal of RZZ from the kinetochores, Spindly, CENP-E, and RZZ might form a complex capable of bidirectional movement of the kinetochores [89]. Understanding how RZZ regulates biorientation may depend on reconciling the relative importance of two activities, RZZ moving kinetochores with motors like Dynein and CENP-E, and RZZ blocking or delaying end-on attachments.

Materials and Methods

Drosophila Genotypes and Expression of Transgenes and shRNAs

The transgenes and mutants used in this study are listed in Table 2. The localization of SAC proteins was examined in Drosophila oocytes using GFP fusion transgenes, most under the control of their own promoter. To examine MPS1, we used a construct used extensively in cell culture but modified for making transgenics (P{EGFP-Mps1.F} = Mps1GFP) [90, 91]. To visualize the RZZ complex in oocytes, we used a construct where ROD was tagged with Green Fluorescent Protein (GFP) at the N-terminus and under the control of its own promoter (P{gEGFP-rod} = rodGFP) [25] (Figure S 1A). Expression of either transgene resulted in very low levels (~0.25%) of X-chromosome nondisjunction (Table S1), which was comparable to wild-type strains [92]. This is consistent with previous studies showing that this transgene can rescue the lethality of rod mutants [25, 93].

The UAS/GAL4 system was utilized to drive expression of many of the transgenes in oocytes. Matα was used to induce expression following DNA replication and early pachytene and throughout meiotic prophase.

A fusion between the C-terminal domain of Spc105R and the entire coding region of Zw10 was generated by GenScript in pENTR4. The C-terminal fragment of SPC105R has been characterized previously and is sufficient for kinetochore localization [22]. The fusion gene was transferred into pPWH, which fused the coding region to a 3XHA tag at the C-terminal end. This was injected into embryos for germline transformation by Model Systems Injections.

RNA Extraction and Quantification of RNAi Knockdown

Among three shRNA lines targeting rod, one (P{Val22:rod-153}1attP40, = rodRNAi)) was chosen because it had the strongest phenotype and most severe mRNA knockdown (4% of the wild-type). Expression of rodRNAi using matα produced sterile females and ubiquitous expression caused lethality. To confirm the efficacy of the rodRNAi in oocytes, we examined RODGFP in oocytes expressing the rod shRNA. The intensity of RODGFP at the kinetochores in rodRNAi oocytes was reduced to nearly background levels (Figure S 1B, C), indicating that the rod shRNA provides a strong knockdown of rod. Similarly, the intensity of MPS1GFP at the kinetochores in Mps1RNAi oocytes was significantly reduced (Figure S 1D, E).

RT-qPCR was performed to determine knockdown of mRNAs for each shRNA. After the parental flies were crossed, mature female offspring with the correct genotype were collected into yeasted vials for three days. These flies were then ground in 1X phosphate-buffered saline (PBS) and filtered through meshes to isolate the stage 14 oocytes. 1 mL of TRIzol reagent was added and RNA was extracted per the manufacturer’s instructions (Thermo Fisher Scientific). Taqman RT-qPCR was performed to quantify RNAi knockdowns. 2 μg of isolated RNA was reverse transcribed into cDNA using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems). Afterwards, qPCR was performed using a TaqMan Assay (Thermo Scientific) with four replicates per reaction. The efficacy of RNAi knockdowns has been published for Ndc80 GL00625 = 6% remaining, Spc105R GL00392 = 13% remaining [21], Bub1 GL00151 = 2% remaining [31], BubR1 GLV21065 = 22% remaining, BubR1 GL00236= 27% remaining, and Bub3 HMS00789 = 0.7% remaining.

Measuring X-Chromosome Nondisjunction

To measure nondisjunction of the X chromosome, virgin females with the desired genotype were crossed to males with a dominant Bar mutation on the Y chromosome (yw/BSY). This mutation led to a visible bar-eyed phenotype where the eye was thinner than wild-type eyes. The normal segregation of chromosomes would lead to female offspring (XX) with wild-type eyes and males (XY) with bar eyes. If a nondisjunction event occurred in the X chromosome, the following sex chromosome combinations were possible: XO, XXY, XXX, and YO. XO and XXY were viable and resulted in normal-eyed males and bar-eyed females, respectively. XXX and YO were lethal. To calculate nondisjunction rates while taking into account the lethal genotypes, the following equation was used: #flieswithnondisjunction×2(#flieswithnondisjunction×2)+(#ofnormalflies). This rate determined whether a particular genotype resulted in increased meiotic nondisjunction.

Cytology of Stage 14 Oocytes

Cytology of stage 14 oocytes was performed as previously described [94]. After the parental flies were crossed, mature female offspring with the correct genotypes were collected into yeasted vials for one to two days. These flies were then broken open in a blender while in 1x modified Robb’s buffer and filtered through meshes to isolate the stage 14 oocytes. If a colchicine treatment was needed, oocytes were incubated with 150 μM colchicine or 0.5% EtOH in modified Robb’s buffer for one hour to remove all microtubules. The oocytes were fixed with 5% formaldehyde and heptane before being rinsed in 1X PBS. The oocytes were then rolled between the frosted surface of a glass slide and a cover slip to remove the membranes and nutated for two hours in PBS/1% Triton X-100 to make them permeable to antibody staining. Afterwards, oocytes were washed in PBS/0.05% Triton X-100 and blocked for one hour in blocking buffer (0.5% BSA and 0.1% Tween-20 in 1X PBS). Subsequently, primary antibodies were added to the oocytes while the secondary antibodies were added to Drosophila embryos to remove any nonspecific binding. After nutating overnight at 4°C, the oocytes were washed in blocking buffer four times at room temperature for 15 minutes. The secondary antibodies were added and the oocytes were nutated for 3-4 hours at room temperature. The oocytes were then washed in blocking buffer with Hoechst33342 (10 μg/mL) to stain the DNA. The oocytes were washed twice more in blocking buffer and then were ready for mounting and imaging.

The primary antibodies used in this paper were tubulin monoclonal antibody DM1A at 1:50 conjugated to FITC (Sigma-Aldrich), tubulin E7 monoclonal antibody at 1:150 [95], rat anti-INCENP at 1:600 [96], guinea pig anti-CENP-C at 1:1000 [97], rat anti-tubulin at 1:300 (Millipore), rabbit anti-Spc105R (1:4000) [98], rabbit anti-GFP (1:200) (Thermo Fisher), rat anti-HA (Roche, 1:50). Multiply preabsorbed secondary antibodies were from Jackson ImmunoResearch, and were conjugated to Alexa 647, Cy3, Alexa 594 or Alexa 488.

To visualize pairs of homologous chromosomes, fluorescent in situ hybridization (FISH) was performed. After oocytes fixation, 2X SSC was added and the membranes were removed by rolling the oocytes between glass slides. The oocytes were then nutated in increasing concentrations of formamide solution (20%, 40%, and 50%) before being added to the hybridization solution and probes. The probes were synthesized by IDT, and recognize the 359 bp repeats on the X chromosome (labeled with AlexaFluor 594), the AACAC repeats on the second chromosome (labeled with Cy3), or the dodeca repeats on the third chromosome (labeled with Alexa 647). This was followed by incubation in 80°C for 20 minutes and then 37°C overnight. The next day, the oocytes were washed in 50% formamide solution twice at 37°C and 20% formamide once at room temperature. The oocytes were then rinsed three times in 2X SSCT, once in 2X PBST, and nutated in blocking buffer for 4 hours at room temperature. Afterwards, mouse anti-tubulin-FITC antibody (Sigma) was added (1:50) and the oocytes were nutated overnight at room temperature. The next day, the oocytes were washed in blocking buffer with Hoechst33342 (10 μg/mL). After washing twice more with blocking buffer, the oocytes were mounted and imaged.

Imaging Oocytes, Image Analysis, and Statistical Analysis

Oocytes were mounted on a glass slide in SlowFade Gold (Invitrogen) and images were taken with a 63x lens on a Leica SP8 or Stellaris microscope. Protein intensities were quantified using Imaris image analysis software (Bitplane). To measure the intensity of proteins at kinetochores, spots marking the kinetochores and random areas of the background were created. The average intensity of the protein of interest at the background was subtracted from the intensity at each kinetochore to generate the normalized intensity value. GraphPad Prism software was used to graph data and perform statistical analysis. All statistical tests and sample sizes are reported in the figure legends.

Supplementary Material

Supplement 1

Acknowledgements

We thank Marina Druzhinina for technical assistance, Christian Lehner, and Rager Karess for providing Drosophila stocks and antibodies. Stocks obtained from the Bloomington Drosophila Stock Center (NIH P40OD018537) were used in this study. The E7 tubulin monoclonal antibody was obtained from the Developmental Studies Hybridoma Bank, created by the NICHD of the NIH and maintained at The University of Iowa, Department of Biology, Iowa City, IA 52242. Image acquisition and analysis was made possible by the Waksman Institute Shared Imaging Core Facility and The Human Genetics Institute Imaging Core Facility at Rutgers. This work was supported by NIH grant GM101955 to K.S.M.

Data availability statement

All the data underlying this study are available in the published article and its online supplementary material.

Figure 1. Localization of SAC proteins in oocytes.

(A, B) Oocytes were incubated for one hour in Robb’s buffer with 0.5% EtOH (A) or 250 μM colchicine (B). BUBR1, BUB3, MPS1, ROD, and MAD1 were detected using GFP-tagged transgenes, while INCENP was detected using an anti-INCENP antibody (all proteins shown in green), DNA is in blue, CENP-C in white, and tubulin in red. Single channel images show SAC protein localization. All images are maximum intensity projections of z stacks. Scale bars represent 5 μm. (C) Quantification of SAC protein intensity at kinetochores, normalized to background signal (n for EtOH and colchicine = 313 and 174 for BUBR1, 67 and 83 for BUB3, 219 and 356 for MPS1, 139 and 184 for ROD, 138 and 185 for MAD1, and 74 and 200 for INCENP). Error bars show mean ± s.d.; ****P<0.0001, ns=no significance (unpaired two-tailed t test).

Figure 2. Induction of MAD1 localization is independent of tension and attachment status.

(A, B) MAD1GFP localization in wild-type, Ndc80RNAi, or mei-218null oocytes that were either untreated (A) or treated with 250 μM colchicine (B). MAD1GFP is shown in green, DNA in blue, CENP-C in white, and tubulin in red. Single channel images (bottom) show MAD1GFP with karyosome outlines in dashed blue in (A). All images are maximum intensity projections of z stacks. Scale bars represent 5 μm. (C, D) Quantification of MAD1GFP intensity at kinetochores, normalized to background GFP signal (n =106 and 127 kinetochores for (C) and 158 and 311 kinetochores for (D)). Error bars show mean ± s.d.; ****P<0.0001 (unpaired two-tailed t test).

Figure 3. ROD localization depends on the MELT-KI domain of SPC105R.

(A) RODGFP localization in control and an Spc105RRNAi oocytes, with RODGFP in green, DNA in blue, CENP-C in white, and tubulin in red. Single channel images (bottom) show RODGFP. (B) Quantification of RODGFP intensity at kinetochores normalized to background GFP signal in indicated oocytes (from left to right, n = 142, 169, 144, 144, 167, 138, 206, 159, 173, and 146 kinetochores). Error bars show mean ± s.d.; ****P<0.0001, ***P=0.0006, ns=no significance (unpaired two-tailed t test). All Spc105R mutants are in an Spc105RRNAi background targeting the endogenous Spc105R. (C) Schematic of SPC105RB domains in Drosophila melanogaster. (D) RODGFP localization in the indicated Spc105R mutant oocytes, with RODGFP in green, DNA in blue, CENP-C in white, and tubulin in red. Single channel images (bottom) show RODGFP. All images are maximum intensity projections of z stacks. Scale bars represent 5 μm.

Figure 4. ROD and MPS1 localization depend on SAC genes.

(A) RODGFP localization in wild-type, Bub3RNAi, BubR1RNAi, Mps1RNAi, and CycARNAi oocytes with RODGFP in green, DNA in blue, CENP-C in white, and tubulin in red. Single channel images (bottom) show RODGFP. (B) Quantification of RODGFP intensity at kinetochores, normalized to background GFP signal in indicated oocytes (from left to right, n = 142, 170, 66, 76, and 155 kinetochores). Error bars show mean ± s.d.; ****P<0.0001, *P≤0.0123 (unpaired two-tailed t test). (C) MPS1GFP localization in wild-type, Bub3RNAi, BubR1RNAi, rodRNAi, and CycARNAi oocytes with MPS1GFP in green, DNA in blue, CENP-C in white, and tubulin in red. Single channel images (bottom) show MPS1GFP. (D) Quantification of MPS1GFP intensity at kinetochores, normalized to background GFP signal in indicated oocytes (from left to right, n = 455, 97, 149, 180, and 152 kinetochores). Error bars show mean ± s.d.; ****P<0.0001 (unpaired two-tailed t test). All images are maximum intensity projections of z stacks. Scale bars represent 5 μm.

Figure 5. Meiotic chromosome alignment in SAC RNAi oocytes.

(A) Representative images depicting chromosome orientation in wild-type, rodRNAi, and Mps1RNAi oocytes. Images show DNA (blue), tubulin (green), the X chromosome (magenta), the second chromosome (red), and the third chromosome (white). Tubulin and FISH probes are also shown in a separate channel, with examples of mono-oriented chromosomes circled. All images are maximum intensity projections of z stacks. Scale bars represent 5 μm. (B) Percent of chromosomes per oocyte with the indicated orientation (from left to right, n = 132, 65, 114, 94, 65, 66, 87, 39, and 30 pairs of chromosomes). ****P<0.0001 (Fisher’s exact test). (C) Examples of wild-type, mad11, and mad2EY21687 mutant oocytes, with tubulin in green, DNA in blue, CENP-C in red, and INCENP in white. Single channel images show DNA. (D) Percent of oocytes with the specified karyosome morphology (from left to right, n=114, 102, 103, 41, 31, 32, and 31 oocytes). **P=0.0020, *P=0.0375 (Fisher’s exact test).

Figure 6. ROD streams in the absence of tension and end-on attachments.

(A) RODGFP localization in wild-type, mei-218null, Ndc80RNAi, and PP2A-B56RNAi oocytes with RODGFP in green, DNA in blue, CENP-C in white, and tubulin in red. Single channel images (bottom) show RODGFP. All images are maximum intensity projections of z stacks. Scale bars represent 5 μm. (B) Percent of oocytes with RODGFP streaming in indicated genotypes (from left to right, n = 25, 15, 22, and 16 oocytes), ns=no significance (Fisher’s exact test). (C) Quantification of RODGFP intensity at kinetochores, normalized to background GFP signal in indicated oocytes (from left to right, n = 142, 139, and 92 kinetochores). Error bars show mean ± s.d.; *P=0.0202 (unpaired two-tailed t test).

Figure 7. ROD is not required for microtubule attachments.

(A) Wild-type; rodRNAi; Ndc80RNAi; and rodRNAi, Ndc80RNAi oocytes with tubulin in green, DNA in blue, CENP-C in white, and INCENP in red. RodRNAi oocyte on the left has an intact karyosome and on the right a split karyosome. (B) Percent of kinetochores per oocyte with lateral kinetochore-microtubule attachments in indicated genotypes (from left to right, n = 40, 46, 30, 20, and 31 oocytes). Error bars show s.d.; ****P<0.0001, ***P=0.0007, ns=no significance (unpaired two-tailed t test). (C) Percent of oocytes with the specified karyosome morphology in the indicated genotypes (from left to right, n = 41, 46, 22, 20, and 31 oocytes). ***P=0.0002, *P=0.0260 (Fisher’s exact test). (D) RODGFP localization in Spc105RB, Spc105RΔN, and Spc105RΔExxEED oocytes with RODGFP in green, DNA in blue, CENP-C in white, and tubulin in red. Single channel images (bottom) show RODGFP. All Spc105R mutants are in an Spc105RRNAi background targeting the endogenous Spc105R. (E) Percent of oocytes with RODGFP streaming in indicated genotypes (from left to right, n = 25, 20, 20, and 28 oocytes); **P=0.0012 (Fisher’s exact test). (F) Spc105RC and Zw10-Spc105RC oocytes (both in an Spc105RRNAi background) with DNA in blue, CENP-C in white, tubulin in red, and ZW10-SPC105RC in red (right). Single channel images (bottom) show CENP-C (left) or ZW10-SPC105RC (right). (G) Percent of kinetochores per oocyte with lateral kinetochore-microtubule attachments in indicated genotypes (n = 42 and 51 oocytes). Error bars show s.d.; ****P<0.0001 (unpaired two-tailed t test). All Spc105RC and Zw10-Spc105RC oocytes are in an Spc105RRNAi background targeting the endogenous Spc105R. All images are maximum intensity projections of z stacks. Scale bars represent 5 μm.

Figure 8. Requirement for Spindly in meiosis and interactions with ROD.

(A) Representative images depicting chromosome orientation in wild-type, SpindlyRNAi, and nudERNAi oocytes. Images show DNA (blue), tubulin (green), the X chromosome (magenta), the second chromosome (red), and the third chromosome (white). Tubulin and FISH probes are also shown in a separate channel, with examples of mono-oriented chromosomes circled. (B) Percent of chromosomes per oocyte with the indicated orientation (from left to right, n = 86, 66, and 54 pairs of chromosomes); ****P<0.0001, **P=0.0076 (Fisher’s exact test). (C) RODGFP localization in wild-type, SpindlyRNAi, and nudERNAi oocytes with RODGFP in green, DNA in blue, CENP-C in white, and tubulin in red. Single channel images (bottom) show RODGFP. Arrow points to abnormal streaming in the SpindlyRNAi oocyte. See also Figure S 6A. (D) Percent of oocytes with RODGFP streaming in indicated genotypes (from left to right, n = 25, 41, and 10 oocytes); ns=no significance (Fisher’s exact test). (E) Percent of oocytes that abnormal streaming, defined as RODGFP in the central spindle, in wild-type or SpindlyRNAi oocytes (n=24 and 43 oocytes); ***P=0.0002 (Fisher’s exact test). Abnormal streaming is defined as having RODGFP in the central region between the centromeres. (F) Quantification of RODGFP intensity at kinetochores, normalized to background GFP signal in indicated oocytes (from left to right, n = 142, 256, and 72 kinetochores. Error bars show mean ± s.d.; ns=no significance (unpaired two-tailed t test). All images are maximum intensity projections of z stacks. Scale bars represent 5 μm.

Table 1: Nondisjunction in mad mutants

Genotype	XX	XY	XXY	XO	% NDJ	
mad2EY21687/Df	3857	3558	6	17	0.6	
mad11/Df	805	769	1	6	1.0	
Bwinscy/+; mad2EY21687/Df	357	263	4	12	4.9	
Bwinscy/+; mad11/mad11	427	321	9	4	3.4	
Bwinscy/+; mad11/+	588	375	2	4	1.2	
Bwinscy/+; mad2EY21687/+	509	301	2	2	1.0	
mad2 Df = Df(3L)Exel6107 and mad1 Df = Df(2R)ED1791

Table 2. Drosophila genotypes used in this work.

Genotype	Abbreviation	Reference	
P{w+mC=matalpha-GAL-VP16}V37	matα	[99]	
P{Val22:rod-153}attP40	rodRNAi	This paper	
P{gEGFP-rod}	rodGFP	[25]	
P{EGFP-Mps1.F}	Mps1GFP	[90]	
P{Mad1-GFP}	Mad1GFP	[42]	
P{BubR1.GFP}	BubR1GFP	[73]	
P{gEGFP-Bub3}III.3	Bub3GFP	[100]	
P{w+mC=UASp-Spc105RB}	Spc105RB	[22]	
P{UASp-Spindly.RR.GFP}	SpindlyGFP	[54]	
P{w+mC=UASp-Zw10-Spc105RC}	Zw10-Spc105RC	This paper	
P{TRiP.GL00392}attP2	Spc105RRNAi	[101]	
P{TRiP.GL00625}attP40	Ndc80RNAi	[101]	
P{TRiP.HMS01283}attP2	SpindlyRNAi	[101]	
P{TRiP.HMS01868}attP40	nudERNAi	[101]	
P{TRiP.GL00151}attP2	Bub1RNAi	[101]	
P{wrdGL00671wdbHMS01864}/TM3	PP2A-B56RNAi	[101]	
P{TRiP.GL00236}attP2	BubR1RNAi	[101]	
P{TRiP.HMS00789}attP2	Bub3RNAi	[101]	
Mad11	Mad1	[42]	
Mad2EY21687	mad2	[41]	
mei-2186	mei-218null	[102]	
Mps1ald	Mps1ald	[66]	
Mps1C3	Mps1C3	[37]
==== Refs
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