Abstract
The ploidy cycle, integral to sexual reproduction, requires not only meiosis to halve the number of chromosomes, but also mechanisms that ensure zygotes are formed by exactly two partners1–5. During sexual reproduction of the fungal model organism Schizosaccharomyces pombe, haploid P- and M-cells normally fuse to form a diploid zygote that immediately enters meiosis6. Here, we reveal that fast post-fusion reconstitution of a bi-partite transcription factor actively blocks re-fertilization. We first identify mutants that undergo transient cell fusion involving cytosol exchange but not karyogamy, and show this drives distinct cell fates in the two gametes: The P-partner undergoes lethal, haploid meiosis while the M-cell persists in mating. Consistently, we find that the zygotic transcription that drives meiosis is initiated rapidly only from the P-parental genome, even in wild type cells. This asymmetric gene expression depends on a bi-partite complex formed post-fusion between the nuclear P-cell-specific homeobox protein Pi and a cytosolic M-specific peptide Mi7,8, which is captured by Pi in the P-nucleus. Zygotic transcription is thus poised to initiate in the P-nucleus as fast as Mi reaches it. The asymmetric nuclear accumulation is inherent to the transcription factor design, and is reconstituted by a pair of synthetic interactors, one localized to the nucleus of one gamete and the other in the cytosol of its partner. Strikingly, imposing a delay in zygotic transcription, by postponing Mi expression or deleting its transcriptional target in the P-genome, leads to zygotes fusing with additional gametes, thus forming polyploids and eventually aneuploid progeny. We further show that the signaling cascade to block re-fertilization shares components with, but bifurcates from, meiotic induction9–11. Thus, cytoplasmic connection upon gamete fusion leads to rapid reconstitution of a bi-partite transcription factor in one partner to block re-fertilization and induce meiosis, thus ensuring genome maintenance during sexual reproduction.
While studying mutants displaying cell fusion defects (ref.12), we discovered a striking post-fusion asymmetry between mating partners. Deleting the p21-activated kinase Pak2 impaired cell fusion, resulting in an increased number of unfused partners and extended lifetime of fusion focus components13 at the cell-cell contact site (Fig. 1A, S1A). Unexpectedly, pak2Δ mating also produced ∼10% of aberrant asci never seen in wildtype, which appeared to be derived from either three parental cells containing more than four spores (Fig. 1A, type IIIa), or a single cell that underwent sporulation (Fig. 1A, type IIIb). Three lines of evidence revealed that these asci result from meiosis and sporulation taking place in haploid cells.
First, live imaging showed that karyogamy did not occur between mating partners that produced aberrant asci, even though the spore-forming partner showed microtubule reorganization typical of meiosis: a long cytosolic bundle, characteristic of the meiotic ‘horsetail’ stage14–17 followed by two successive rounds of spindle formation, indicative of meiosis-I and meiosis-II (ref.18, Fig. 1B, S1B, Mov. S1). The other partner cell maintained interphase microtubule organization and ability to mate with an additional partner (thus yielding type IIIa asci). Second, labeling one chromosomal locus with lacO:GFP-NLS-LacI system19 revealed that the spore-forming cell contained only two foci distributed between up to four spores, indicating these spores are aneuploid (Fig. S1C). By contrast, wild-type asci formed four spores each with a single focus (ref.20, Fig. S1C). Third, flow cytometry analysis of DNA-stained spores from pak2Δ crosses indicated presence of spores with less then 1C content (Fig. 1C). We conclude that aberrant pak2Δ asci form upon meiosis and sporulation in a single, haploid cell.
Haploid meiosis was provoked by transient fusion between partners: pak2Δ cells that produced aberrant asci initially formed a fusion focus and fused, thus exchanging cytosolic GFP, but then re-sealed their fusion pore, as indicated by persistent unequal levels of GFP between partners (n>20, Fig. 1D, S1E, Mov. S2,3). Transient fusion was required to induce haploid meiosis since deleting the Fus1 formin, an essential component of the fusion machinery13,21, prevented formation of aberrant asci by pak2Δ cells (n>200, Fig. S1F). Remarkably, crosses between wild-type and fus1Δ mutant, in which fusion efficiency is reduced but not abrogated13,21, also resulted in transient fusion events each followed by haploid meiosis in one partner, whether this was fus1Δ or wildtype (Fig. 1E, S1F-G, Mov. S4). We conclude that transient cell-cell fusion, independently of genotype, induces meiosis in one partner cell.
Quantification of homothallic wildtype and pak2Δ mating mixtures for phenotypes described in the main text and represented in inset images. (B) Time-lapse of homothallic pak2Δ mating cells expressing GFP-α-tubulin in green and nuclear marker Uch2-mCherry in magenta. Note lack of karyogamy in the outlined pak2Δ mating pair even though one partner forms meiotic spindles (arrows) and spores (arrowhead), while the other partner maintains interphase microtubule organization. Fig. S1B shows wildtype control. (C) Flow cytometry analysis of DNA content in spores produced by wildtype and pak2Δ cells. (D) Time-lapse of pak2Δ mating cells expressing cytosolic GFP from the P-cell specific pmap3 promoter. Note the transient cell fusion, observed as exchange of GFP, between M- and P1-cells followed by sealing of the fusion pore (arrow) and spore formation (full arrowheads) in the P1-partner. Also note formation of the fusion focus (empty arrowhead), labeled by Myo52-GFP in the M-cell and Myo52-tdTomato in the P-cell. Fig. S1D reports wildtype control. (E) Time-lapse showing transient cell fusion between wildtype P- and fus1Δ M-cells expressing GFP from the M-cell specific pmam2 promoter. Persistent GFP signal difference between partners indicated sealing of the fusion pore. Note spore formation (arrowhead) only in the wildtype P-cell.
Zygotic transcription occurs first in the P-cell
Remarkably, all instances of transient cell fusion induced haploid meiosis strictly in the P-cell (n=64 in pak2Δ; n=9 between fus1Δ and wild-type), and the master meiotic regulator Mei311,22 was induced only in P-cells undergoing transient fusion. In transiently fusing pairs, tagging mei3 with a fast-folding GFP variant (sfGFP)23 in the P-genome resulted in strong fluorescent signal in the P-cell undergoing haploid meiosis, whereas labeling of M-genome mei3 did not lead to any noticeable fluorescence induction in either partner (Fig. 2A, 2B and Mov. S5, n>10 in each cross). We note that the M-cell encoded Mei3-sfGFP was expressed upon complete fusion that formed normal zygotes (Fig. 2B and Mov. S5, note Mei3-expression in zygote formed by M- and P2-cells). Importantly, mei3 was also asymmetrically expressed in wildtype zygotes: P-genome encoded Mei3-sfGFP expression was significantly more rapid (about 15 min earlier) than Mei3-sfGFP induction from the M-cell genome (Fig. 2C, 2E, S2A and Mov. S6). Thus, fission yeast zygotes express the meiotic inducer Mei3 first from the P-cell genome.
Mei3 expression is under the regulation of two cell type-specific factors, the P-cell-specific homeodomain protein Pi and the M-cell-specific 42 amino acid peptide Mi7,8. To test whether these govern the asymmetric induction of mei3 from parental genomes, we swapped the coding sequences of Pi and Mi. Without affecting mating and sporulation efficiencies nor spore viability (Fig. S2B-D), the Pi-Mi swap resulted in Mei3 being expressed first from the M-cell genome (Fig. 2D-E, S2E and Mov. S7), and transient fusion inducing haploid meiosis in M-cells (Fig. 2F, S2F-G and Mov. S8, 13 out of 14 instances). Thus, asymmetric zygotic transcription of mei3 expression is governed by the cell type-specific expression of Mi and Pi, with mei3 being induced first from the genome of the Pi-expressing cell.
Pi and Mi nuclear-cytosolic shuttling underlies their asymmetric activity
To explore how parental genomes are differentially regulated by Pi and Mi, we tagged each with sfGFP at their native genomic loci. Mi-sfGFP expression commenced during early mating, producing a faint cytoplasmic signal in M-cells. Unexpectedly, after fusion, Mi-sfGFP rapidly accumulated in the P-cell nucleus and only ∼5-minutes later in the M-cell nucleus (Fig. 3A, S3A and Mov. S9). In pak2Δ cells undergoing transient cell fusion Mi-sfGFP transferred from the M- to the P-cell and accumulated in the nucleus of the P-cell, which then underwent haploid sporulation (Fig. 3B, n>10). Importantly, Mi did not accumulate in the nucleus of the M-cell that underwent transient cell fusion. We conclude that Mi nuclear accumulation requires a P-cell-specific factor that remains asymmetrically distributed upon transient cell fusion.
Several lines of evidence indicate that this P-cell-specific factor is Pi. First, Pi and Mi form complexes in vivo (Fig. 3C). Second, Pi was localized to the nucleus: N-terminal tagging of Pi with sfGFP produced a very low-intensity signal in nuclei in both P-cells and zygotes (Fig. S3B and Mov. S10). This nuclear localization was more evident in fus1Δ cells, which attempt, and fail, to fuse for a long time (Fig. S3C, Mov. S10), or upon overexpression in interphase cells (Fig. 3D). Third, Mi-mCherry overexpressed in interphase cells was cytosolic but accumulated in the nucleus when sfGFP-Pi was co-overexpressed (Fig. 3D). Finally, deletion of Pi prevented Mi-sfGFP nuclear enrichment after fusion (Fig. 3E, Mov. S11).
These data are consistent with a model (Fig. S3D) where Pi accumulates in the P-gamete nucleus while Mi is present throughout the nucleo-cytoplasm of the M-gamete. Upon cell fusion the 5kDa Mi is rapidly exchanged between partners and trapped by Pi in the P-nucleus where both proteins form a complex to rapidly induce Mei3 expression. Expression of Mei3 from the M-genome is postponed due to delayed exchange of Pi between partners.
We considered several reasons for the observed rapid, asymmetric nuclear localization of Mi. The 5kDa Mi peptide may diffuse significantly more rapidly than its 19kDa partner Pi24. However, tagging Mi with 27kDa sfGFP did not abrogate the asymmetry, as shown above, suggesting the asymmetry does not simply rely on protein sizes. The observed reversal of asymmetry upon Mi-Pi swap however indicates that the asymmetry does not depend on other cell type-specific factors. We thus considered whether distinct subcellular localization of Pi and Mi to the nucleus and the cytosol underlie the asymmetry. To mimic Pi and Mi characteristics, we constructed an artificial system consisting of synthetic hetero-specific short coiled-coils SynZip3 and SynZip425, the former fused to mCherry and the latter to sfGFP and a nuclear localization signal (NLS), each expressed in one partner. Remarkably, upon cell-cell fusion, the cytosolic marker accumulated in the partner’s nucleus immediately, whereas the equilibration of the nuclear marker was delayed, thus mimicking the behavior of Mi and Pi (Fig. 3F, Mov. S12). The observed asymmetry was even more striking in instances of transient cell fusion (Fig. S3E, Mov. S12). When both proteins lacked an NLS (Fig. S3F, Mov. S12), asymmetric localization was not observed upon fusion. We conclude that the observed asymmetry is inherent to the distinct sub-cellular localization of Mi and Pi, a design that promotes a more rapid Mei3 transcription from the P-genome.
Rapid induction of Mei3 transcription prevents re-fertilization and polyploid formation
To address the physiological function of this fast Mei3 induction, we delayed Mei3 expression in two distinct ways. First, we placed Mi in M-cells under control of a P-cell specific pmap3 promoter, such that Mi (and consequently Mei3) becomes expressed only after successful fusion, once P-cell-specific transcription factors activate the pmap3 promoter in the M-genome. This delayed Mei3-sfGFP expression by ∼30 minutes (Fig. 4A, S4A and Mov. S13). Second, we simply deleted mei3 from the P-genome, such that Mei3 is expressed only from the M-genome, and thus delayed by ∼15 minutes (Fig. 2E). While neither approach affected mating or sporulation efficiencies upon complete fusions of otherwise wildtype cells (Fig. S4B, n=3×300), both methods fully prevented haploid sporulation upon transient fusion of pak2 mutants. Instead, we observed persistent mating behaviors with both partners repeatedly attempting to form a zygote (note the post-fusion growth of mating projections in Fig. 4B, 4C and Mov. S14, n=10). Deleting mei3 in pak2Δ M-cells, however, did not affect haploid sporulation in the P-cell (Fig. S4C and Mov. S14, n>10).
Remarkably, when Mei3 expression was delayed by either method in otherwise wildtype crosses, zygotes formed but then continued to engage additional partners (Fig. 4D and Mov. S15-16, n=18 for Mi expression delay and 18 for mei3 deletion in P-cells, out of 7’000 events each). Multi-partner zygotes entered meiosis, as shown by microtubule labeling, and formed four, likely aneuploid spores (Fig. 4E, S4E, Mov. S15-16). By contrast, we rarely observed triploid zygotes when mei3 was deleted from the M-cell genome (2 of n > 7’000 events). Polyploid zygotes were never observed in wildtype (n > 10’000). In sum our data reveal that the rapid onset of Mei3 expression from the zygotic P-cell genome functions to prevent formation of polyploid zygotes and consequently aneuploid spores.
Furthermore, approximately 1% of all zygotes formed by homothallic PiΔ, MiΔ or mei3Δ strains not only did not enter meiosis, but were seen fusing with an additional partner (Fig. S4G-I and Mov. S17, n>500). A much larger fraction (>30%) of zygotes exhibited growth suggestive of mating behavior (Fig. 4F, Mov. S17-18). The frequency of re-fertilization events appeared to increase further with time but could not be quantified due to deterioration of imaging chambers.
Mei3 alleviates repression of the master meiotic regulator, the RNA binding protein Mei210,26,27, which, in complex with meiRNA/sme29, promotes meiosis by indirectly allowing expression of downstream factors including the forkhead transcription factor Mei428,29. Both mei4Δ and sme2Δ zygotes arrested prior to spore formation but were never observed engaging additional partners (Fig. S4J-K). Conversely, nearly 10% of mei2Δ zygotes fused with additional partners (Fig. 4G and Mov. S18). Since a point mutation in Mei2 C-terminal RNA-recognition motif9 also resulted in ∼10% of multi-partner zygotes (Fig. 4H and Mov. S18), we conclude that the re-fusion block is reliant on Mei2 signaling, likely through RNA binding, but not in complex with meiRNA. These data establish that repression of mating in zygotes shares components with, but bifurcates from, meiotic induction.
Mechanisms preventing re-fertilization are present in evolutionarily divergent phyla to ensure ploidy maintenance across generations. Almost universally, these mechanisms involve rapid changes, be it through release of cortical granules1 and shedding of cell-surface receptors in mammals2, membrane depolarization in amphibians1,30, or degradation of pollen guidance cues in plants31. Our results now provide a first glimpse into the mechanism blocking re-fertilization in fungi, by showing that rapid activation of zygotic transcription promotes discontinuation of mating in yeast zygotes. The bi-partite design of the Mi-Pi transcription factor, reminiscent of that of hormone nuclear receptors and their ligands32, favors post-fusion activation speed, which is limited only by the rate at which the Mi peptide reaches the already nuclear-localized homeodomain Pi protein. Because this simple two-component system inherently leads to asymmetric zygotic expression, asymmetry may naturally follow from the selective pressure to rapidly block re-fertilization. The observation that even a transient cytoplasmic connection is sufficient for formation of an active complex raises the possibility that such transcription factor design may also be used to impart cell fate changes upon, or build sensors to monitor, other instances of transient cytoplasmic bridge formation.
Materials and Methods
Detailed materials and methods are provided in a supplemental file appended below.
Materials and Methods
Growth conditions
The growth conditions used for experiments are detailed in ref.1 and the overview of experimental procedure is presented in Fig. S5A. Briefly, freshly streaked cells were inoculated into MSL+N media2 and incubated overnight at 25°C with 200 rpm shaking to exponential phase. The following evening cultures were diluted to O.D.600nm=0.025 in 20 ml of MSL+N media and incubated overnight at 30°C with 200 rpm agitation to exponential phase. Experiments on exponentially growing cultures were performed at this point. For time-lapse imaging of mating, homothallic cells or 1:1 mixtures of heterothallic cells were pelleted for one minute at 1000g in microcentrifuge tubes and washed three times in 1ml of MSL-N media2. Cells were then diluted in 3ml of MSL-N media to final O.D.600nm=1.5 and incubated at 30°C with 200 rpm agitation for 4-6 hours. Finally cells were mounted onto MSL-N agarose pads, covered with a coverslip and the chamber was sealed using VALAP (Vaseline, Lanolin, Paraffin; 1:1:1).
For flow cytometry and quantifications of mating and sporulation efficiencies, ∼3×107 of MSL-N washed cells were re-suspended in 20 µl of MSL-N media and spotted onto MSL-N 2% agar plates and incubated at 25°C for the indicated number of hours. The number of unmated cells, unsporulated zygotes and sporulated zygotes was determined using transmitted light microscopy, and mating and sporulation efficiencies were calculated using the following formulas:
The reported results are from three replicates, error bars denote standard deviation and p-values were calculated using Welch’s t-test.
Flow cytometry
We collected the mated cell suspensions from MSL-N plates and resuspended samples in 1ml of MSL-N media containing 10µl of glusulase (NEE154001EA, Perkin-Elmer, Waltham, USA). After overnight incubation at 30°C we visually verified that all cells except spores had lysed upon glusulase treatment. The samples were washed three times with MSL-N and resuspended in 3ml of water containing 2-5 µg/ml of Hoechst 33342 DNA dye (B2261, Sigma-Aldrich, St. Louis, USA) and incubated for 15-30 minutes at room temperature. Samples were immediately analyzed on Becket-Dickson Fortessa with proprietor software platform using the 355nm laser line with 450/50 filter. The experiment was run in duplicate.
Strains
All strains are reported in Table S1. Standard fission yeast genetic and molecular biology tools were used in the study3. Sequences of mutants obtained in this study are provided as annotated, GenBank-formatted sequences (see Supplemental Sequences).
The mating loci of strains used in the study are illustrated in Fig. S5B. Mi and Pi deletions remove the ORF of the genes. sfGFP tagging of Pi and Mi was done at the N- and C-terminus respectively.
In homothallic strains, the mat1 locus determines the cell mating type and switches between sequences encoded by the silent mat2 and mat3 loci [reviewed in ref.4]. Importantly, a H1Δ17 mutation at the mat1 locus H1-homolgy box prevents mating type switching5. As depicted in the Fig. S5B, in the synthetic mat1 locus sequences, the wild-type H1-homology box was replaced with the one amplified from the PB9 strain carrying the H1Δ17 mutation (a kind gift from Benoit Arcangioli, Institut Pasteur, Paris, France). The recombinant DNA also contain a selection cassette and homology arms that targets the constructs to the mat1 locus of homothallic cells. The described sequences were cloned and amplified inside the pBlueScript plasmid. Finally the indicated restriction enzymes (see Supp.Seq.1-4) were used to extract the fragment of interest and transform it into h90 wild type cells. Correct integration was verified by PCR and/or inability to perform mating type switching.
mat2 and mat3 loci that reside in heterochromatic DNA and thus cannot be targeted by homologous recombination in wild-type cells were manipulated as described in ref.6. An overview of the experimental procedure is presented in Fig. S6A. The mat2 locus manipulations were performed in the TP1 strain (a kind gift from Genevieve Thon, University of Copenhagen, Copenhagen, Denmark), which lacks the clr3 gene, to allow recombination at the mat2 locus. Furthermore, the TP1 strain carries ade6+ and ura4+ cassettes in the vicinity of the mat2 locus. The TP1 strain was transformed with DNA fragments carrying the designed mat2 locus and flanking wild-type sequences. 5-FOA resistant, adenine auxotrophs were selected and tested for correct integration by PCR. The mat2 mutant locus was then introduced in otherwise wild-type cells via genetic crosses. mat3 manipulations were similarly performed in the PG3089 strain (a kind gift from Genevieve Thon), which lacks the clr4 gene and carries the ura4+ cassette in the proximity of the mat3 locus. The sequences of the mutant mat2 and mat3 loci are provided (see Supp.Seq.5-8).
For Pi overexpression we used the constitutive tdh1 promoter comprised of 1000bp upstream of the tdh1 start codon to drive expression of Pi N-terminally tagged with sfGFP. This construct was cloned into plasmids that contain the budding yeast ADH1 transcriptional terminator sequence, BleMX selection cassette and sequences targeting integration at either ura4 or his5 locus (Supp.Seq.9-10). AfeI digested ura4 targeting plasmid and StuI digested his5 targeting plasmid were transformed in ura4 and his5 mutant cells respectively. Prototrophic, zeocin-resistant clones were selected and checked for correct integration by PCR and genetic crosses.
For Mi overexpression we used either the tdh1 or the nmt41 promoter to drive expression of Mi that was C-terminally tagged with mCherry. These constructs were cloned into plasmids that target either ura4 or lys3 locus and contain kanMX or natMX selection cassettes respectively (Supp.Seq.11-12). AfeI digested ura4 targeting plasmid and BstZ17I digested lys3 targeting plasmid were transformed in corresponding auxotrophic strains and prototrophic clones with adequate antibiotic resistance were selected and checked for correct integration by PCR and genetic crosses.
mei3Δ mutant background was used to prevent induction of meiosis by co-expression of Mi and Pi7,8 as was the case for control strains expressing individual proteins.
For Mi expression from the P-cell-specific pmap3 promoter we used 2063bp upstream of the map3 start codon followed by the Mi ORF sequence, the budding yeast ADH1 transcriptional terminator sequence and BleMX selection. This construct was cloned inside the pJK148 plasmid9 that targets the construct to the leu1 locus after NdeI digestion (Supp.Seq.13). The linearized plasmid was transformed into h-cells lacking Mi and a G418 resistant transformant that efficiently sporulated when crossed with wildtype h+ cells was selected. The same transformation clone was used to introduce the pak2Δ::hphMX mutation as described below.
pak2Δ::ura4+ mutant strain was derived from ref10. For the pak2Δ::hphMX we cloned the 430bp of 3’UTRpak2 followed by 399bp of the 5’UTRpak2 into the pFA6a-hphMX vector which created an AfeI site between the two UTR’s. The obtained plasmid (Supp.Seq.14) was linearized with AfeI, transformed into cells and hygromycin-B resistant colonies were selected and tested for correct integration by PCR.
mei2Δ::kanMX and mei3Δ::kanMX mutant strains were derived from the S. pombe deletion library strains S7B02 and S20A08 respectively (Bioneer, Daejeon, Republic of Korea) and were verified by PCR.
sme2Δ::ura4+ and mei4Δ::ura4+ mutations were derived from Japan National BioResource Project strains FY7237 and FY7361, respectively.
fus1Δ::natMX was generated using methodology detailed in ref.11. Briefly, primers with 80bp homology to sequences immediately flanking fus1 ORF (see Table S2) were used to PCR amplify the natMX selection cassette from the pFA6a-natMX plasmid and transformed into cells. Nourseothricin resistant clones were selected and genotyped by PCR.
For construction of mei2 point mutant we first cloned the 3’ region (sequence from 313-672bp downstream mei2 stop codon) followed by the mei2 gene (sequences from 487bp upstream of the start codon to 312bp downstream of the stop codon), which creates an AfeI restriction enzyme between the two fragments. This construct was inserted in the pFA6a-hphMX plasmid and we introduced the F644A mutation by site-directed mutagenesis (Supp.Seq.15-16). Linearization of the plasmid with AfeI targets the construct to replace the genomic mei2 gene upon transformation. The hygromycin-B resistant clones were selected and the correct integration replacing the wild-type mei2 was confirmed by PCR.
The strains with a fluorescently labeled genomic locus12 were derived from FY13708 strain obtained from the Japan National BioResource Project. In these cells the lys1+ locus is genetically linked to the LacO sequence array that binds the GFP-LacI expressed from the dis1 promoter at the his7+ locus.
For microtubule visualization we used cells with SV40 promoter driven expression of GFP-α-tubulin13 derived from a FC1234 strain, a kind gift from Fred Chang’s group.
For mei3 fluorophore tagging, yeast codon optimized sfGFP (a kind gift from Michael Knop, Heidelberg University, Germany) was introduced into a pFA6a-natMX vector to obtain the pFA6a-sfGFP-natMX vector (Supp.Seq.17). The plasmid was then used as a template for PCR with primers that amplify sfGFP followed by natMX cassette and carry 80bp homology to sequences immediately flanking mei3 STOP codon (see Table S2). The PCR product was transformed into wild-type cells, and nourseothricin resistant clones were selected and genotyped by PCR. All mei3-sfGFP expressing strains used in the study were derived from the same transformation clone by genetic crosses.
For C-terminal mCherry tagging of nuclear marker uch2 and spindle pole body marker pcp1, we used the pFA6a-mCherry-natMX and pFA6a-mCherry-kanMX plasmids respectively. In both cases, we cloned the 3’UTR region of the gene followed by C-terminal fragment of the ORF keeping it in frame with the fluorophore (Supp.Seq.18-19). The obtained plasmids to tag uch2 and pcp1 were then treated with AfeI and AfeI+SnaBI restriction enzymes respectively, and the digested DNA was transformed into cells. Clones resistant to either nourseothricin or G418 were selected and genotyped by PCR.
myo52-GFP and myo52-tdTomato were introduced from strains FC85714 and ySM740 15 respectively.
The construct to express the green and red fluorophores from the P-cell-specific map3 promoter were previously described in ref.16. For the expression of GFP from the M-cell-specific mam2 promoter we cloned the 438bp of promoter upstream of the start site into a pRIP-GFP plasmid that contains the ura4+ selection cassette (Supp.Seq.20). Transformation of the ura4-mutant cells with the PmeI digested final plasmid targeted the construct to the native mam2 promoter and conferred growth in absence of uracil.
Sequences encoding small interacting SynZip peptides17 were a kind gift from Serge Pelet, University of Lausanne, Switzerland. The SynZip3 and SynZip4 peptides were cloned at the C-terminus of mCherry and sfGFP respectively. Expression of mCherry-SynZip3 was driven from the pact1 promoter consisting of 822bp upstream of the act1 start codon. The sfGFP-SynZip4 was driven from the ptdh1* promoter comprised of 896bp upstream of the tdh1 start codon. For the NLS-sfGFP-SynZip4 we introduced the SV40 nuclear localization signal at the sfGFP N-terminus and used the ptdh1* promoter comprised of 896bp upstream of the tdh1 start codon. All constructs were integrated in the plasmid that carries the ura4+ selection cassette and targets the ura4 locus. All constructs are detailed in Supp.Seq.21-23. The AfeI linearized final plasmids were transformed into ura4 mutants and uracil prototrophs were selected for further experiments.
Co-immunoprecipitation
We grew 50ml of mei3Δ mutant cell cultures overexpressing Mi-mCherry or sfGFP-Pi or both to exponential phase and collected cells by centrifugation for one minute at 1000 rcf. From this point the experiment was performed on ice. Cells were transferred to microcentrifuge tubes, washed twice with ice-cold CXS buffer (50mM HEPES,pH7.5; 20mM KCl; 2mM EDTA; 1mM MgCl2; protease inhibitors) and re-suspended in 300µl of the CXS buffer and ∼200µl of zirconium beads were added. Cells were lysed using the bead beater with 10 cycles of 20-second beating and 40 seconds cooling on ice. The beads were removed and samples transferred to a new tube where the cell debris was pelleted by centrifugation for 15 minutes at 13’000g. We collected the supernatant and determined the protein concentration using the Bradford assay. We adjusted the samples to 1mg of total protein in 500µl of CXS buffer and set aside 30µl for western blotting analysis (Fig. 3A labeled as INPUT). The samples were then incubated with 30µl of GFP-Trap beads (gtma-20, Chromotek, Planegg-Martinsried, Germany) prewashed in CXS buffer. After one hour incubation on a rotating stand at 4°C beads were washed three times in CXS buffer and re-suspended in Laemmli buffer. Samples were analyzed with standard western blotting protocol. For primary antibodies we used 1:3’000 dilution of anti-GFP antibody (Cat.No. 11814460001, Roche, Basel, Switzerland) anti-RFP antibody (6G6, Chromotek, Planegg-Martinsried, Germany) and TAT-1 antibody (a kind gift from Keith Gull, University of Oxford, UK). We used the infrared fluorophore coupled secondary antibodies at 1:5’00 dilution (R-05061 and R05054, Advansta, Menlo Park, USA) and visualized the blots on the Fusion FX (Vilber, Collégien, France). The experiment was reproduced in three independent replicates.
Microscopy and image analysis
To acquire images in Fig. 3D and S1C we used a spinning-disk confocal system that uses an inverted microscope (DMI6000B; Leica) equipped with an HCX Plan Apochromat 100x/1.46 NA oil objective and an UltraVIEW system (PerkinElmer; including a real-time confocal scanning head [CSU22; Yokagawa Electric Corporation], solid-state laser lines, and an electron-multiplying charge coupled device camera [C9100; Hamamatsu Photonics]). Stacks of z-series confocal sections were acquired at 0.3-µm intervals using Volocity software (PerkinElmer).
All other micrographs were obtained by wide-field microscopy performed on a DeltaVision platform (Applied Precision) composed of a customized inverted microscope (IX-71; Olympus), a UPlan Apochromat 100x/1.4 NA oil objective, a camera (CoolSNAP HQ2; Photometrics), and a color combined unit illuminator (Insight SSI 7; Social Science Insights). Images were acquired using softWoRx v4.1.2 software (Applied Precision). Principally we imaged a single Z-section with the exception of data presented in Fig. 1B and S1B where 6 sections with 0.5µm spacing were acquired and the presented images are a maximum projection of images deconvolved using softWoRx v4.1.2 inbuilt module. All conclusions based on imaging data were derived from at least three independent observations.
Image processing and fluorescence intensity based quantifications were performed in ImageJ (NIH, Bethesda, USA). Supplemental movies were converted from TIFF to MOV format using the inbuilt MPEG-4 compression. For overnight time-lapses that exhibited drifting, presented images were aligned using the MultiStackReg1.45 Plugin. All quantifications were performed on raw images.
The lifetime of the fusion focus was measured on time-lapse images as the interval between the fluorescently labeled marker proteins first focalizing16 and cell fusion. Results are reported with the box-and-whiskers plot where center lines show the medians, box limits indicate the 25th and 75th percentiles, whiskers extend 1.5 times the interquartile range from the 25th and 75th percentiles, outliers are represented by dots and the Kruskal-Wallis test p-value is reported.
To distinguish the expression of mei3 from P- and M-genome we performed crosses between heterothallic strains where only one partner carried the mei3-sfGFP allele, while the other partner had the unlabeled, wildtype gene. Quantification of Mei3-sfGFP fluorescence induction used cell fusion as time zero, evidenced as the first timepoint with exchange of cytosolic RFP between partners. The mean signal in individual partners prior to fusion, and thus prior to mei3 induction, was considered as background signal. Mean fluorescence of the whole zygote was recorded over time and the average of the indicated number of cells is reported with shaded regions denoting mean ± standard deviation. The post-fusion time for the mean zygotic intensity to reach 50 arbitrary units above background was recorded for each zygote and the results are reported with the box-and-whiskers plot where center lines show the medians, box limits indicate the 25th and 75th percentiles, whiskers extend 1.5 times the interquartile range from the 25th and 75th percentiles, outliers are represented by dots and the Kruskal-Wallis test p-value is reported.
For Mi-sfGFP nuclear entry dynamics we used the last time-point prior to exchange of Mi-sfGFP between partners as time zero. The nuclei of the zygote were clearly identifiable based on Mi-sfGFP signal alone, which allowed us to outline the nuclear region required in these quantifications. The signal in the P-cell prior to cell fusion was considered as background. Mean fluorescence of each nuclear region was recorded over time and the average of the indicated number of cells is reported with shaded regions denoting mean ± standard deviation and the Kruskal-Wallis test p-value is reported.
For quantification of dynamics of NLS-sfGFP-SynZip4, sfGFP-SynZip4 and mCherry-SynZip3 we used the last time-point prior to exchange of red fluorophore between partners as time zero. The nuclear region was outlined based on the accumulation of the fluorescent signals in the central cell region. The signal in the partner cell lacking the fluorophore prior to cell fusion was considered as background signal. Mean fluorescence of each nuclear region was recorded over time and the average of the indicated number of cells is reported with shaded regions denoting mean ± standard deviation and the Kruskal-Wallis test p-value is reported. Note that the mCherry-SynZip3-expressing cell also displays background vacuolar fluorescence, likely due to pre-fusion degradation of some mCherry-SynZip3.
Acknowledgements
We thank Benoit Arcangioli and Geneviève Thon for strains and advice, Magdalena Marek for help with flow cytometry, Felipe Bendezú and Serge Pelet for strains and reagents, and Richard Benton, Tonni Grube Andersen, Stefan Gruber, Sara Mitri, Jan-Willem Veening and Martin lab members for critical reading of the manuscript. This work was supported by an EMBO long-term fellowship to AV and an ERC Consolidator Grant (CellFusion) and a Swiss National Science foundation grant (31003A_155944) to SGM.