Mcp5, a meiotic cell cortex protein, is required for nuclear movement mediated by dynein and microtubules in fission yeast

The formation of heritable haploid gametes from diploid parental cells requires meiosis, a special type of cell division. After premeiotic DNA replication, homologous chromosome pairing and genetic recombination occur during prophase I with formation of bouquet arrangement conserved among species (for review see Harper et al., 2004). During prophase I of fission yeast, horsetail nuclear movement occurs. It starts when all the telomeres become bundled at the spindle pole body (SPB; Hiraoka, 1998). Subsequently, the nucleus undergoes a dynamic oscillation, resulting in elongated nuclear morphology (Chikashige et al., 1994). This event has been proposed to facilitate the pairing of homologous chromosomes because it aligns the chromosomes in the same direction (through the bundling of their telomeric ends) and then shuffles them around each other (Chikashige et al., 1994; Kohli, 1994; Niwa et al., 2000; Ding et al., 2004).

It has been proposed that horsetail nuclear movement is predominantly due to the pulling of astral microtubules that link the SPB to cortical microtubule-attachment sites at the opposite end of the cell; the pulling force is believed to be provided by cytoplasmic dynein (Ding et al., 1998; Yamamoto et al., 1999; Yamamoto and Hiraoka, 2003) and the dynactin (Yamashita et al., 1997; Niccoli et al., 2004) complex. However, the details of these mechanisms are still almost completely unknown. A key to understanding nuclear oscillation is the identification and characterization of the anchor protein at the cell cortex that binds to dynein. We report that a novel meiosis-specific gene named mcp5⁺ (from meiotic coiled-coil protein) encodes a protein, Mcp5 (Saito et al., 2005), that functions as this anchor during the meiosis of Schizosaccharomyces pombe. Recently, we found that it is also registered as Mug21 (Martin-Castellanos et al., 2005). Mcp5/Mug21 is homologous to Num1 (nuclear migration) of Saccharomyces cerevisiae, ApsA (anucleate primary sterigmata) of Aspergillus nidurans, and Ami1 (anucleate microconnidia) of Podospra anserina, but these Num1-like proteins function during vegetative growth rather than in meiosis (Kormanec et al., 1991; Fischer and Timberlake, 1995; Graïa et al., 2000; Heil-Chapdelaine et al., 2000), unlike Mcp5. We demonstrate a contribution for such an anchoring mechanism in the process of homologue pairing during meiosis and establish a link between meiotic and mitotic nuclear behavior, albeit in different organisms.

We first found that Mcp5 shows meiosis-specific expression and is essential for sporulation as shown in Fig. S1 . Because S. cerevisiae Num1 is involved in nuclear migration during the budding process, we investigated the function of Mcp5 during the horsetail nuclear movement of S. pombe, which is the nuclear migration event in this species and which is specifically observed in meiotic prophase. We subjected h⁹⁰ cells expressing DNA polymerase α-GFP to time-lapse observation under a microscope. In wild-type (WT) cells, sequential oscillatory nuclear movement is observed from karyogamy to slightly before meiosis I (Fig. 1 A and Video 1). In contrast, such oscillatory nuclear movement was not observed throughout prophase I in mcp5Δ cells (Fig. 1 B and Video 2). This abnormal oscillation was also observed in the azygotic meiosis of mcp5Δ cells (Videos 3 and 4). These data suggest that Mcp5 plays a significant role in nuclear oscillation.

In mcp5Δ cells, which show impaired horsetail movement (Fig. 1), the rates of pairing (Fig. S2, A and B; and Videos 5–8; available) and recombination (Fig. S2, C and D) of homologous chromosomes are also reduced. This is similar to what has been observed for the mcp6/hrs1Δ (Saito et al., 2005; Tanaka et al., 2005), kms1Δ (Shimanuki et al., 1997), dhc1 mutant (Yamamoto et al., 1999), dlc1Δ (Miki et al., 2002), and ssm4Δ (Niccoli et al., 2004) cells, in which horsetail movement is also inhibited. Ectopic recombination rates are also increased in mcp5Δ cells (Fig. S2 E), as has also been observed for the mcp6/hrs1Δ (Saito et al., 2005) and kms1Δ (Niwa et al., 2000) cells.

We also found that the period from karyogamy to meiosis I in zygotic meiosis was 1.5 times longer in mcp5Δ cells than in WT cells (Fig. 1 C) and that the timing with which cells containing two or four nuclei peaked was similar in azygotic meiosis in both strains of pat1 background (Fig. S2 F). These results suggest that the delay of prophase I in the zygotic meiosis of mcp5Δ cells is due to a delay in the progression of karyogamy. Both phenotypes of deficiency in nuclear movement and delay in karyogamy of mcp5Δ are similar to those of the dhc1 mutant (Yamamoto et al., 1999).

We examined the subcellular localization of Mcp5 by constructing a GFP-Mcp5–expressing strain in the h⁹⁰ genetic background. Because the frequency of recombination of the gfp-mcp5 cells was at the WT level (unpublished data), we judged the function of GFP-Mcp5 to be intact in this strain. The cells were induced to enter zygotic meiosis, observed by living and methanol fixation, and stained with Hoechst 33342 to detect the nucleus and with anti–α-tubulin antibody to detect the microtubules. As shown in Fig. 2 A (top), no GFP signal was detected during the vegetative phase. During the horsetail phase, however, the GFP-Mcp5 signals appeared in the cytoplasm with a gradient pattern, namely, seeping from the arm to the middle and from the arm to the cell end. Furthermore, some bright dots were detected at the cell cortex. Significantly, astral microtubules were observed to be elongating in the direction of some GFP-Mcp5 dots (Fig. 2 B). Notably, dynein heavy chain (Dhc) 1, one of the +TIPs (microtubule plus-end tracking proteins), colocalized with Mcp5 at a point of cell cortex during horsetail phase (Fig. 2 C). These cortical dots disappeared after the horsetail phase, and the signal was not detected again through meiosis I and II (Fig. 2 A). When Mcp5-GFP was overproduced in vegetative cells, Mcp5-GFP localized at the cell cortex, suggesting that other meiosis-specific proteins are not required for the cell cortex localization of Mcp5 (Fig. 2 D).

To examine the effect of Mcp5 on the sliding of microtubules at the cell cortex during the horsetail phase, h⁹⁰ WT (YY105) and h⁹⁰mcp5Δ (ST218-1) cells were chemically fixed 6 h after meiosis was induced by nitrogen starvation and then immunostained with anti-Sad1 antibody, and the tubulin-GFP image was observed under a microscope (Fig. 3, A and B). We counted only the cells at horsetail phase, at which point the microtubules exhibit a filamentous structure that extends from the SPB to the edge of the cell. We denoted the microtubules as “sliding” or “not sliding,” depending on whether the microtubules curved or did not curve along the cell cortex, respectively. This analysis revealed that ∼50% of WT cells (Fig. 3, A [top] and B [left]) but only 12.8% of mcp5Δ cells (Fig. 3, A [bottom] and B [right]) exhibited sliding microtubules. This indicates that mcp5Δ cells are defective in the cortical sliding of their microtubules. It is known that Mcp6/Hrs1 plays an important role in organizing astral microtubule arrays at the SPB during the horsetail phase (Saito et al., 2005; Tanaka et al., 2005). However, mcp5Δ cells show normal localization of Mcp6/Hrs1 (unpublished data). Thus, it appears that although Mcp5 is not required for the proper formation of astral microtubules, it is needed for the proper sliding of the microtubules along the cell cortex.

For Dhc1 to generate pulling force on the microtubule tip and so pull the SPB to the cortex, it must be fixed at the cell cortex. Dynein has both microtubule-binding and minus end–directed motor activity, and Yamamoto et al. (2001) proposed that a dynein anchor is required for dynein to accomplish these functions. To assess the role Mcp5 may play in this process, we investigated the localization of Dhc1-GFP during the horsetail phase in live observations (Fig. 3 C). In WT cells, the Dhc1-GFP dot was observed at the SPB and cell cortex along with the microtubules (Fig. 3, C [top] and D [left]; and Video 9). In contrast, most mcp5Δ cells did not exhibit an intense Dhc1-GFP dot at the cell cortex (Fig. 3, C [bottom] and D [right]; and Video 10). When we counted the number of cortical Dhc1-GFP dots, we found that 81% of WT but only 14% of mcp5Δ cells harbored these dots (Fig. 3 D, colored bars). Moreover, these dots localized more frequently at the leading edge (57%) than on the tail side (24%) in WT cells, whereas the mcp5Δ cells showed a reversed trend (3 vs. 11%). These results suggest that horsetail movement is linked to the accumulation of cortical Dhc1-GFP dots and indicate that Mcp5 is required for anchoring dynein at the cell cortex, thus allowing the proper U-turn movement of the nucleus.

To determine the function of the PH domain of Mcp5, we constructed the PH domain deletion mutant Mcp5 (PHΔ). When we overproduced the Mcp5 (PHΔ)-GFP fusion protein in h⁹⁰ WT cells (CRL266) using high-copy plasmid pREP1 driven by its nmt1 promoter (Okuzaki et al., 2003), the cortical localization of Mcp5 was lost (Fig. 4 A). These results indicate that the PH domain is required for the cortical localization of Mcp5. Interestingly, most of the Mcp5 (PHΔ)-GFP signals were detected in the nucleus, and only a small amount of signal was observed along the microtubule (like Dhc1; Fig. 3 C) during both karyogamy and the horsetail phase (Fig. 4 A). It is possible that the PH-deleted Mcp5 mutant is transported to the nucleus through its nuclear localization signal (NLS). Furthermore, the Dhc1-like linear localization of Mcp5 (PHΔ) could signify that Mcp5 (PHΔ) is transported to the microtubules as a cargo protein of dynein.

To determine the role of the coiled-coil motif of Mcp5 plays, we prepared a mutant strain that expresses Mcp5 protein lacking two coiled-coil motifs, namely, Mcp5 (C-CΔ). When we overproduced the Mcp5 (C-CΔ)–GFP fusion protein in h⁹⁰ WT cells (CRL266), it localized at the cell cortex (Fig. 4 B). These results indicate that the coiled-coil domain is not required for the cortical localization of Mcp5. We next observed the phenotypes of the mcp5 (c-cΔ) strain during meiosis and found that it showed aberrant nuclear oscillation (Fig. 4 E), reducing the recombination rate (Fig. S2 C) and the abnormal spore formation (not depicted). These results indicate that the coiled-coil domain of Mcp5 plays a pivotal role in its full function and suggest that an unidentified partner that associates via the coiled-coil domain is involved in regulating nuclear movement.

Mcp5 localizes at the cell cortex via its PH domain (Fig. 4 A) and colocalizes with dynein (Dhc1) or tubulin-like Num1 of S. cerevisiae (Farkasovsky and Küntzel, 2001). Given that Num1 forms a transient complex in vivo with dynein and tubulin (Farkasovsky and Küntzel, 2001), it may be that Mcp5 also associates transiently with dynein and tubulin in vivo.

Fig. 5 illustrates our working hypothesis about the role Mcp5 plays in anchoring dynein and thereby generating pulling force. When the SPB (Fig. 5, gray circle) and the horsetail nucleus reaches at one end of the cell cortex, the microtubules carrying the SPB and nucleus extend toward the opposite end (Fig. 5 A). Pairing of homologous chromosomes may occur inefficiently in the round nucleus at the cell end. When the dynein complex (Fig. 5 A, green triangle) on the microtubules is trapped and anchored by Mcp5 (red circle), the minus-end motor activity may be stimulated (Fig. 5 B). Thus, the dynein starts to pull microtubules by its minus-end motor activity, which causes the cortical sliding of the microtubules. Then, the nucleus turns around and is pulled back toward the opposite cell end. Homologous chromosome pairing may be achieved widely in the elongated nucleus, which may induce the recombination of the homologous chromosomes. When the SPB arrives at the cell cortex where the dynein is anchored by Mcp5, the pulling force is gradually weakened and the microtubules behind the horsetail nucleus will extend to the opposite cell end, returning to stage A (Fig. 5 C). These stages are repeated several times during 2–3 h of prophase I. The magnified view of the cortical site on stage B is shown in Fig. 5 D. Like Num1 (Yu et al., 2004), Mcp5 is localized at the cell cortex, probably through its PH domain–mediated interaction with phosphatidylinositol (4,5) bisphosphate. Mcp5 on the membrane helps the cortical sliding of microtubules through anchoring the dynein–dynactin complex.

In conclusion, the homologous proteins Mcp5 and Num1 play similar roles in dynein-dependent nuclear migration (for review see Bloom, 2001; Yamamoto and Hiraoka, 2003), even though they are used in different cellular events, namely, Mcp5 in meiosis during prophase I and Num1 in mitosis during the budding (cell division) phase (Heil-Chapdelaine et al., 2000).

The S. pombe strains used in this study are listed in Table S1 . The complete media YPD or YE, the synthetic Edinburgh minimal medium 2 (EMM2), and the sporulation medium molt extract or EMM2-nitrogen (EMM2-N; 1% glucose) were used. We used high-copy plasmid pREP1 driven by its nmt1 promoter for overproduction experiments (Okuzaki et al., 2003).

For time-lapse observations, cells expressing GFP-tagged Polα (CRL026-1, ST205, and ST349) or lacI-NLS-GFP and the lacO repeat integrated at the lys1 locus (AY174-7B and ST240) or cut3 locus (NP43-20 and ST268) were cultured in 10 ml EMM2 + supplements until they reached midlog phase at 28°C. They were then induced to enter meiosis by incubation in EMM2-N at 28°C. After 5 h of nitrogen starvation, the cells were put on a glass-bottomed dish whose surface was coated with 0.2% concanavalin A, and images under a fluorescence microscope (IX71; Olympus) were recorded every 2.5 min (1 s of exposure time) after the initiation of karyogamy. For observation of lacI-GFP dots, images were taken with a 0.3-s exposure at 5-min intervals with 10 optical sections made at 0.5-μm intervals for each time point. Projected images obtained with MetaMorph software (Universal Imaging Corp.) were analyzed.

For NH₂-terminal tagging to generate GFP-tagged mcp5⁺, we synthesized the following six oligonucleotides and used them as primers: MCP5-5F, 5′-AAAACTGCAGCTACGGCGGATTAAGGGTAG-3′; MCP5-5R, 5′-GGG-TTTCATATGGAAGGATCAGTATATGTTTTG -3′; Xho-mcp5N, 5′-CCGCTCGAGCGGATGGAGAAAAAACAAGATAACG-3′; Sma-mcp5C, 5′-TCCCCCGGGTTAAATGCCGGGCTTAGTATC-3′; MCP5-3F, 5′-TCCCCCGGGGCATGTTTCAGTCATTGC-3′; and MCP5-3R, 5′-CTCGAGGAGCTCCTTACAAAATAAAAATGCTG-3′. The underlined sequences denote the artificially introduced restriction enzyme sites for PstI, NdeI, XhoI, SmaI, SmaI, and SacI, respectively. These PCR products were inserted into the pRGT1 vector via the PstI–NdeI, XhoI–SmaI, and SmaI–SacI sites, respectively. This plasmid construct was digested with PstI and SacI, and the resulting construct was introduced into the mcp5⁺∷ura4⁺ strain ST256. After growth on EMM2 + 5-fluoroorotic acid plates, the ura transformants were screened by PCR analysis to identify the (mcp5⁺∷ura4⁺)∷(GFP-mcp5⁺) replacement.

For COOH-terminal tagging to generate FLAG-tagged mcp5⁺, we followed the previously described method (Saito et al., 2004). Thus, we performed PCR using the WT (TP4-5A) genome as a template and obtained a DNA fragment carrying the ORF and the 3′ downstream region of the mcp5⁺ gene. For this purpose, we synthesized the following two oligonucleotides and used them as primers: MCP5N, 5′-CGGCGCGCCGCATATGGAGAAAAAACAAGATAA-3′, and MCP5C, 5′-GTACTCGAGGCGGCCGCGGAATGCCGGGCTTAGTATCAA-3′. The underlined sequences denote the artificially introduced restriction enzyme sites for NdeI and NotI, respectively. To obtain the 3′ downstream region, we used the primers described in the previous paragraph that produce the Smal–Sacl fragment. These PCR products were inserted into the pTT (FLAG)-Lys3 vector (Saito et al., 2005), which is designed to allow one-step integration. This plasmid construct was digested with NsiI. The resulting construct was introduced into HM105 (h⁻ lys3). We then screened the Lys+ transformants and confirmed the precise integration of the constructs by PCR.

Fluorescent microscopic observations were performed as described previously (Saito et al., 2005). Cells from a single colony were cultured in 10 ml EMM2 with supplements until they reached midlog phase at 28°C. The cells were collected by centrifugation, washed three times with 1 ml EMM2-N, and induced to enter meiosis by incubation in EMM2-N at 28°C for 6 h.

For immunofluorescence experiments, meiotic cells were fixed after the procedure using methanol or glutaraldehyde. For methanol fixation, cells were collected by aspiration through a glass filter that traps cells. The cells were immediately immersed into methanol at –80°C and left overnight to fix the cells. The cells were then washed off the glass filter by distilled water and collected by centrifugation, and the pellet was washed three times with PBS. For glutaraldehyde fixation, 10 ml of cultured cells were incubated in ∼3% paraformaldehyde and 0.25% glutaraldehyde for 1 h.

In indirect immunofluorescence microscopy, microtubules were stained with the anti–α-tubulin antibody TAT1 (after methanol fixation; a gift from K. Gull, University of Manchester, Manchester, UK) and the SPB was stained with the anti-Sad1 antibody (after glutaraldehyde fixation; a gift from O. Niwa, Kazusa DNA Research Institute, Kisarazu, Japan). Subsequently, we added an Alexa 594–conjugated goat anti–mouse antibody (Invitrogen) for TAT1 or an Alexa 594–conjugated goat anti–rabbit antibody (Invitrogen) for the anti-Sad1 antibody, respectively. In both cases, the samples were then stained with 0.2 mg/ml Hoechst 33342 in PBS for 1 min and mounted with antifade mounting medium containing p-phenylenediamine. Fluorescence images of these cells were observed using a fluorescence microscope (BX51; Olympus) with a charge-coupled device camera (CoolSNAP; Roper Scientific). Immunofluorescence images were acquired using Photoshop 7.0 (Adobe).

Fig. S1 shows that Mcp5 is a meiosis-specific coiled-coil protein that belongs to Num1 family proteins found specifically among fungi. Fig. S2 shows characterization of mcp5Δ cells. Video 1 shows a wild-type cell expressing GFP-labeled DNA polymerase α during zygotic meiosis. Video 2 shows an mcp5Δ cell expressing GFP-labeled DNA polymerase α during zygotic meiosis. Video 3 shows a wild-type cell expressing GFP-labeled DNA polymerase α during azygotic meiosis. Video 4 shows an mcp5Δ cell expressing GFP-labeled DNA polymerase α during azygotic meiosis. Video 5 shows a wild-type cell whose lys1 loci are labeled by GFP-LacI-NLS fusion protein. Video 6 shows an mcp5Δ cell whose lys1 loci are labeled by GFP-LacI-NLS fusion protein. Video 7 shows a wild-type cell whose cut3 loci are labeled by GFP-LacI-NLS fusion protein. Video 8 shows an mcp5Δ cell whose cut3 loci are labeled by GFP-LacI-NLS fusion protein. Video 9 shows a wild-type cell expressing GFP-labeled Dhc1 during horsetail phase. Video 10 shows an mcp5Δ cell expressing GFP-labeled Dhc1 during horsetail phase. Table S1 shows a list of strains used in this study.

We are obliged to Drs. O. Niwa and K. Gull for antibodies; A. Miyawaki for SECFP; and G.R. Smith, Y. Hiraoka, A. Yamamoto, C. Shimoda, and M. Yamamoto for S. pombe strains. We also thank Drs. K. Nabeshima and T. Tougan for technical advice.

This work was supported by grants-in-aid for Scientific Research on Priority Areas (16013226), Scientific Research (S) (15101006), Exploratory Research and Science, and Technology Incubation Program in Advanced Regions from the Ministry of Education, Science, Sports, and Culture in Japan. T.T. Saito is a Research Fellow of the Japan Society for the Promotion of Science.