Abstract
The spindle assembly checkpoint (SAC) is a conserved mitotic regulator that preserves genome stability. Despite its central role in preserving the fidelity of mitosis, the strength of the SAC varies widely between cell types. How the SAC is adapted to different cellular contexts remains largely unknown. Here we show that both cell size and cell fate impact SAC strength. While smaller cells have a stronger SAC, cells with a germline fate show increased SAC activity relative to their somatic counterparts across all cell sizes. We find that enhanced SAC activity in the germline blastomere P1 requires proper specification of cell fate downstream of the conserved PAR polarity proteins, supporting a model in which checkpoint factors are distributed asymmetrically during early germ cell divisions. Our results indicate that size scaling of SAC activity is modulated by cell fate and reveal a novel interaction between asymmetric cell division and the SAC.
Introduction
The fidelity of mitosis depends upon equal partitioning of the replicated genome between daughter cells. During mitosis, sister chromatid pairs connect to the mitotic spindle via kinetochore-microtubule attachments. Stable attachment of sister chromatids to opposite spindle poles (bi-orientation) ensures that, upon chromatid separation, one copy segregates to each daughter cell. Attachment of sister chromatids to the mitotic spindle is an inherently stochastic process of variable duration (1). Thus to safeguard against chromosome segregation errors, the spindle assembly checkpoint (SAC) monitors kinetochore-microtubule attachments and prevents anaphase onset until stable bi-orientation has been achieved. Weakening of the SAC can lead to aneuploidy and has been associated with tumour development in both model systems and human cancers (2, 3). Accordingly, a major class of anti-mitotic chemotherapeutics (spindle poisons) depends upon robust SAC activity to disrupt mitosis in cancer cells (4).
Interfering with the formation of stable kinetochore-microtubule attachments prevents cells from satisfying the SAC and leads to prolonged mitotic delays. However, the SAC cannot block anaphase indefinitely, and most cells, even in the complete absence of spindle microtubules, will eventually exit mitosis (5). The duration of mitotic arrest under conditions that preclude satisfaction of the SAC is often used as an indication of the “strength” of the SAC. The strength of the SAC varies widely between different cell types (6, 7), cells from different organisms (5) and cells at different developmental stages (8-10). While variation in the strength of the SAC is widespread, an understanding of how this variation is achieved is currently lacking.
The core SAC module consists of the proteins Bub1, Bub3, BubR1, Mad1 and Mad2, which are recruited to unattached kinetochores in a stepwise fashion to promote assembly of the mitotic checkpoint complex (MCC), which acts as a diffusible “wait-anaphase” signal (11, 12). The MCC consists of Mad2, BubR1 and Bub3 bound to the anaphase-promoting complex/cyclosome (APC/C) cofactor Cdc20 (13). The MCC binds to the APC/C (14, 15), blocking degradation of APC/C substrates, namely Cyclin B (16) and securin (17), thereby maintaining a mitotic state. Once all kinetochores are stably bound by microtubules, disassembly of the MCC (18, 19), dephosphorylation of SAC proteins (20) and stripping of Mad1 and Mad2 from all kinetochores (21) effectively silences the SAC and permits anaphase onset and mitotic exit.
Mitotic exit, despite failure to satisfy the SAC, occurs when activity of Cyclin B/Cdk1 falls below the threshold necessary to maintain a mitotic state (5). In mammalian cells, this mitotic “slippage” occurs because the active cytoplasmic pool of MCC is not sufficient to completely inhibit the APC/C, and the progressive degradation of Cyclin B eventually enables mitotic exit (22). The steady-state concentration of the MCC pool is influenced by the rate of new, kinetochore-catalyzed MCC generation and disassembly of existing cytoplasmic MCC (23, 24), the former being intimately related to the severity of the spindle defect (25-27). Thus variation in the strength of the SAC, as assayed by the duration of mitotic delay following spindle perturbation, may be linked to differences in MCC production, activity or stability.
One factor that contributes to SAC strength in certain circumstances is cell size. In vitro experiments using Xenopus laevis egg extracts suggested that an increased nuclear to cytoplasmic ratio, as would be found in smaller cells, could increase SAC activity (28). Recent work in C. elegans embryos has shown that the strength of the SAC indeed scales with cell size, with smaller cells exhibiting longer SAC-dependent delays upon spindle perturbation (29). However, in other organisms, the SAC remains inactive until the mid-blastula transition and acquisition of SAC activity is neither accelerated by decreasing cell volume (Xenopus laevis, (8, 9)) nor delayed by increasing cell volume (Danio rerio, (10)), indicating that SAC activity can also be developmentally regulated independently of changes in cell volume. How this developmental regulation of the SAC is achieved is unknown.
We have recently reported that the adult germline stem cells (GSCs) of C. elegans exhibit a stronger SAC relative to early embryonic cells (30) providing a tractable model in which to examine SAC adaptability. Here we use an inducible monopolar spindle assay to investigate the developmental origins of enhanced germline SAC activity. In agreement with Galli and Morgan (29), we find that the duration of SAC-dependent mitotic delays increases as cell size decreases during embryogenesis. However, the relationship between cell size and SAC activity is strongly influenced by cell fate, with cells in the germline lineage displaying a stronger SAC relative to their cell size than cells with a somatic fate. At the 2-cell stage, we find that differential SAC activity in the somatic AB versus germline P1 blastomere requires the highly conserved PAR polarity proteins, supporting a model in which asymmetric partitioning of a checkpoint-enhancing factor increases the strength of the SAC in germline cells.
Results
Germline blastomeres are more sensitive to spindle perturbations
C. elegans GSCs are derived from a single founder cell (P4), which is specified during embryogenesis by a series of asymmetric cell divisions (31). As the C. elegans embryonic lineage is invariant and fully mapped, each cell can be identified by its position within the embryo and its cell cycle characteristics (Figure 1A, (31, 32)). We were therefore able to ask whether the embryonic precursors to GSCs, cells in the germline P lineage, also exhibited an enhanced SAC, by comparing the duration of mitosis between each of the founding cell lineages, in cells with or without spindle perturbations. As C. elegans embryos are largely refractory to treatment with small molecule spindle poisons without physical or genetic manipulations to permeabilize the egg shell (33, 34), we opted for a genetic method to induce spindle perturbations with temporal control. We utilized temperature sensitive alleles of genes involved in spindle formation, combined with fluorescent markers to visualize the mitotic spindle (β-tubulin fused to GFP, hereafter β-tubulin::GFP) and/or the nucleus/chromatin (histone H2B fused to mCherry, hereafter H2B::mCH). This allowed us to disrupt spindle formation with a simple temperature shift using a standard temperature-controlled imaging chamber and to follow mitotic progression by monitoring changes in nuclear and chromosome morphology.
We used a recessive temperature-sensitive allele of the gene encoding the polo-related kinase ZYG-1, which prevents centriole duplication and produces cells with monopolar spindles (zyg-1(or297), hereafter zyg-1(ts), (35, 36)). Monopolar spindles can generate mitotic delays that are indistinguishable from Nocodazole treatments that either partially (6, 7) or completely (25) inhibit microtubule formation, suggesting a robust SAC response, and are an established means for SAC studies in C. elegans (37). zyg-1(ts) is fast-acting (36) and readily permitted the induction of monopolar spindles in all embryonic cells up to the 16-cell stage. Following NEBD, condensed chromosomes spread radially around the single spindle pole and remained dispersed until the start of decondensation (Figures 1B and 1C, Movie S1). In later stage embryos, centrosome duplication was unaffected by temperature shift and all divisions were bipolar (data not shown).
In agreement with previous reports, we found that the duration of mitosis in cells without spindle perturbations was invariant across embryonic stages and cell lineages (Figure 1D, (29, 38)). In cells in which spindle formation was perturbed, we noticed significant differences in the duration of mitosis between cells at different embryonic stages and in different lineages. Notably, cells in the germline P lineage remained in mitosis for significantly longer than both their immediate somatic siblings and somatic cells at later embryonic stages (Figures 1E). A similar result was obtained using a temperature sensitive allele of the microtubule subunit β-tubulin (tbb-2(or362), (39)), which alters microtubule dynamics and/or stability, to disrupt spindle formation (Figure S1 and Movie S2)
The difference in mitotic delay between germline and somatic cells is not solely due to cell size
Specification of the germline is achieved via a series of asymmetric cell divisions, such that, the germline blastomere is always smaller than its immediate somatic sibling (31). As the strength of the SAC negatively correlates with cell size during C. elegans embryogenesis (29), the increased duration of monopolar mitoses in germline blastomeres could be due to their relatively small size. Thus, we considered cell size in two ways. We used nuclear area, which scales with cell size in many organisms including C. elegans (40, 41), and which we could measure for the same cells in which we assayed mitotic delay, as a proxy for cell size (Figure S2A and S2B). In both the germline P and somatic AB lineages, the duration of monopolar mitoses negatively correlated with nuclear area, supporting the idea that smaller cells possess a stronger SAC (r = -0.73, p < 0.001 and r = -0.61, p < 0.001, respectively). However, germline cells exhibited longer mitotic delays than somatic AB lineage cells with similar nuclear areas (Figure 2A), suggesting that, between comparably sized cells, the SAC is stronger in germline cells.
Nuclear size reached an upper limit and did not reflect differences in cell volume in larger cells (Figure S2B and S2F). A similar phenomenon has been described for the size scaling of both individual chromosomes and the mitotic spindle (42, 43). Thus, in a separate experiment, we calculated the average volume for each cell in the different lineages relative to the volume of the founding P0 blastomere (Figure S2C-E). In both lineages, mitotic delay increased as cell volume decreased (Figure 2B, r = -0.62, p < 0.001 for the AB lineage; r = -0.72, p < 0.001 for the P lineage). However, the relationship between cell volume and the duration of monopolar mitoses differed significantly between the two lineages (AB versus P regression slope: p = 0.03). Here again, germline cells experienced longer mitotic delays relative to their cell volume than somatic AB cells, suggesting that SAC strength is also shaped by cell lineage.
Both cell size and lineage-specific differences in the duration of monopolar mitoses are checkpoint dependent
The increased duration of monopolar mitoses in germline blastomeres could be due to a stronger SAC; alternatively, checkpoint-independent factors could contribute to a delayed mitotic exit in these cells. To discriminate between these possibilities, we examined the duration of monopolar mitoses in the germline P and somatic AB lineages when checkpoint activity was eliminated using RNAi depletion of the C. elegans ortholog of Mad1, mdf-1(RNAi) (Figure 2C and 2D, (44)), or a null allele of the C. elegans ortholog of Mad2, mdf-2(tm2190) (Figure 2E, (45, 46)). In the absence of checkpoint activity, cells with monopolar spindles rapidly exited mitosis, with chromosome decondensation evident within 4 minutes of NEBD (Figure 2C and Movie S3). Plotting the duration of monopolar mitoses in the germline P and somatic AB lineages relative to cell size (as approximated by nuclear area; see Figure S2 and preceding section) revealed that all cells, irrespective of their size or lineage, exited mitosis with the same timing (Figure 2D and 2E). The duration of checkpoint deficient monopolar mitoses was comparable to that of normal bipolar divisions (3.3±0.5 versus 3.4±0.3 minutes, respectively). Thus monopolar spindle-induced mitotic delays are entirely checkpoint dependent and differences in the duration of these delays are likely to reflect differences in the strength of the SAC.
SAC activity may also be subject to developmental regulation
The strength of the checkpoint increases in both germline and somatic AB cells as cell size decreases. If cell size affected SAC strength according to a universal size-scaling relationship, the linear regression lines for mitotic delay versus cell volume for each lineage should be parallel. Instead, SAC activity increases more rapidly with decreasing cell volume in the germline P lineage (Figure 2B and S3A), suggesting that either cell size impacts SAC activity differently in the two lineages or that SAC activity is progressively modified during lineage development. To distinguish between these possibilities, we modified embryo size experimentally and measured the duration of monopolar mitoses at a single developmental stage, in the somatic AB and germline P1 blastomeres of 2-cell stage embryos.
We used RNAi depletion of ANI-2 and PTC-1 to generate small embryos, and C27D9.1 to generate large embryos (Figure 3A and data not shown, (41, 47-49)). Altering embryo size by these means gave us a 5-fold range of embryo volumes, from 1.2 × 104 μm3, slightly smaller than a control P1 cell, to 6.1 × 104 μm3, about twice the size of an average control embryo, without disrupting basal mitotic timing and other defining features of AB and P1, such as cell cycle asynchrony and cell size asymmetry (Figure S3B-D). Hereafter cell volume is expressed as the radius of the equivalent sphere (), which we will call “cell size”.
Cell size and the duration of monopolar mitoses were negatively correlated in both AB and P1 blastomeres (Figure 3B, r = -0.86, p < 0.001 and r = -0.83, p < 0.001, respectively), indicating that cell size can impact SAC activity irrespective of developmental stage. While the rate at which the duration of monopolar mitoses changed relative to cell size was similar for AB and P1 (Figure 3B, AB versus P1 regression slope: p = 0.12), the average duration of monopolar mitoses was always higher in P1 cells (Figure 3B, AB versus P1 regression y-intercept: p = 0.008), indicating that P1 cells have a stronger SAC regardless of cell size. These results suggest that cell identity sets a baseline of SAC activity, which is then scaled according to cell size similarly in AB and P1 and argue against a cell fate-specific difference in SAC size scaling during development of the AB and P lineages. Thus we favor the alternative hypothesis, that checkpoint activity is modified within each lineage as development progresses. Altogether our results demonstrate that SAC activity is determined by a combination of cell size, cell fate and potentially developmental stage in the early C. elegans embryo.
PAR protein-mediated cytoplasmic asymmetries are required for differential SAC activity in AB versus P1
The pattern of checkpoint activity that we have observed in 2- to 16-cell stage embryos mirrors the asymmetric inheritance of certain germline factors, raising the possibility that a determinant of SAC activity could be similarly regulated. For example, several germline proteins are initially present at low levels in the somatic siblings of germline blastomeres, but absent in their progeny (50-53), and germline-enriched maternal transcripts are initially symmetric during cell division and subsequently lost from somatic cells and/or their progeny (54). Correspondingly, the sibling of the germline blastomere in 4- and 8-cell stage embryos (EMS and C, respectively) exhibited monopolar mitoses that were intermediate in duration to those in germline P and somatic AB lineage cells, and, in the progeny of EMS (MS and E), these longer monopolar mitoses were lost (Figures 1E and 2B). The asymmetric inheritance of germline determinants is regulated by the highly conserved PAR proteins, which lead us to ask whether differences in SAC activity also require PAR proteins.
The polarized, cortical distribution of PAR proteins regulates both cell size and the cytoplasmic distribution of cell fate determinants during the division of P0 (55). PAR-6, PAR-3 and the atypical protein kinase C PKC-3 localize to the anterior cortex, while PAR-2 localizes to the posterior cortex. The asymmetric cortical distribution of PAR proteins depends on mutual antagonism between the anterior and posterior PARs. Consequently, in the absence of anterior PARs, posterior PARs move into the anterior, and vice versa (55). The PAR-1 kinase is also enriched in the posterior and is both cytoplasmic and cortical (56). Activity of PAR-1 regulates the asymmetric segregation and/or degradation of cytoplasmic cell fate determinants, leading to the enrichment of germline factors in P1 and somatic factors in AB (57). PAR-1 plays a minimal role in the regulation of cortical PAR protein asymmetries (58-60), but is absolutely required for downstream cytoplasmic asymmetries (56, 61, 62).
To determine how much of the difference in SAC strength between AB and P1 was due to cell size, we compared the average difference in mitotic delay between AB and P1 cells that were the same size versus AB and P1 cells where the normal cell size ratio was preserved. Monopolar mitoses were 4.3±1.8 minutes longer, on average, in P1 cells when AB was larger than P1, irrespective of embryo volume, and 2.0±1.9 minutes longer, on average, in P1 when AB and P1 were the same size (Figure 3C and 3D). When we compared sibling AB/P1 pairs from control embryos versus embryos in which AB and P1 were the same size but maintained somatic versus germline fate (gpr-1/2(RNAi) (63-65)), the difference in the duration of monopolar mitoses was reduced from 4.4±1.1 to 2.7±1.0 minutes (Figure 3E and 3F and Movie S4). Thus, in 2-cell stage embryos, approximately half of the difference in checkpoint strength between AB and P1 is due to their size difference.
We next asked whether the activity of PAR proteins was responsible for the remaining difference in mitotic delay between equally sized AB and P1 cells. While disrupting PAR proteins in P0 will change the fate of the resulting cells, for simplicity we will continue to refer to the “anterior” blastomere as AB and the “posterior” blastomere as P1. RNAi depletion of PAR-6 resulted in sibling AB and P1 cells that were the same size, similar to gpr-1/2(RNAi) embryos (Figure 3E and 3F). However, in par-6(RNAi) embryos the difference in duration of monopolar mitoses was further reduced to 1.0±0.9 minutes, with several AB/P1 sibling pairs exiting mitosis synchronously (Figure 3G, Movie S5). Similarly, RNAi depletion of PAR-1 reduced the difference in the duration of monopolar mitoses between AB/P1 sibling pairs to 0.34±1.6 minutes (Figure 3G). RNAi depletion of additional PAR proteins (PKC-3 and PAR-2) gave similar results (Figure S4A and S4B). Thus PAR protein mediated asymmetries account for much of the remaining difference in checkpoint activity when AB and P1 are the same size. Depletion of PAR-6 and PAR-1 diminished the difference in mitotic delay between AB and P1 sibling pairs to a similar extent. As asymmetries in the cortical localization of PAR proteins are largely maintained in the absence of PAR-1 (58-60), while cytoplasmic asymmetries are lost entirely (56, 61, 62), our results suggest that PAR-1-dependent cytoplasmic asymmetries, but not cortical polarity per se, drive differential checkpoint activity in AB and P1.
Loss of either anterior or posterior PAR proteins increases SAC activity in AB
PAR proteins regulate the levels and localization of asymmetrically distributed cellular components largely via the posterior enrichment of PAR-1. Removing anterior PAR proteins permits PAR-1 activity in the anterior, resulting in two cells with germline P1 traits. Conversely, eliminating posterior PAR proteins allows somatic AB traits to emerge in both cells (55). To ask whether SAC activity was subject to similar regulation, we measured the absolute duration of monopolar mitoses in PAR protein-depleted AB and P1 cells. The mean duration of monopolar mitoses relative to mean cell size for par(RNAi) AB and P1 cells (excepting par-2(RNAi)), more closely resembled the predicted values for P1 than AB control cells; whereas values for gpr-1/2(RNAi) embryos, in which polarity is normal, were similar to the corresponding control cell (Figure 4A). RNAi depletion of PAR proteins impacts the size of AB and P1, generating AB cells that are slightly smaller and P1 cells that are slightly larger than control AB and P1 cells, respectively. Thus, to confirm this trend, we utilized data from Figure 3B (hereafter referred to as “control” cells), binned cells by size and compared the duration of monopolar mitoses in par(RNAi) AB and P1 cells to comparably-sized, but normally polarized and fated, control AB and P1 cells.
RNAi depletion of the anterior PAR-6 and PKC-3 proteins increased the duration of monopolar mitoses in AB cells, such that mitotic delays in depleted AB cells closely resembled those in control P1 cells (Figure 4B). Surprisingly, however, RNAi depletion of the posterior PAR-1 protein also increased the overall duration of monopolar mitoses in AB, while depletion of the posterior PAR-2 protein gave an intermediate phenotype (Figure 4B). AB and P1 cells from gpr-1/2(RNAi) embryos, showed mitotic delays that were indistinguishable from comparably sized control AB and P1 cells, respectively (Figure S4C). All par(RNAi) embryos displayed the expected phenotypes with respect to cell cycle duration and asynchrony (66) (Figure S5A) indicating that our depletions were efficient.
We confirmed these results using a kinase-dead allele of par-1 (it51 (56)) and a near-null allele of par-2 (lw32 (58, 67)), with their respective genetic controls (see Table S1 for complete genotypes). AB cells in par-1(it51) embryos had monopolar mitoses similar in duration to control P1 cells and significantly longer than control AB cells (Figure 4C). In par-2(lw32) AB and P1 cells, the mean duration of monopolar mitoses was largely unchanged. RNAi depletion of PAR-1 in par-2(lw32) embryos eliminated the difference in mitotic delay between sibling AB/P1 pairs almost entirely (0.06 ± 0.9 minutes), and phenocopied par-1(it51) and par-1(RNAi) alone, with unaffected P1 cells and longer monopolar mitoses in AB cells (Figure 4CD). par-1(it51), par-2(lw32) and par-2(lw32); par-1(RNAi) embryos all exhibited changes in cell cycle duration and asynchrony consistent with their genotype (Figure S5B). Under all conditions, par-1(-) AB cells were significantly smaller than control AB cells and comparable in size to control P1 cells (Figure 4C and 4D); however, in light of our AB/P1 size scaling and gpr-1/2(RNAi) results (Figure 3), small AB cells should have shorter monopolar mitoses than comparably sized control P1 cells. As monopolar mitoses in par-1(-) AB cells are largely indistinguishable in duration from control P1 cells, we conclude that the impact of losing PAR-1 is not due exclusively to its effect on cell size. Overall, our genetic and RNAi data are closely aligned and strongly suggest that PAR-1 activity is required for lower SAC activity in AB relative to P1.
Discussion
We have shown that, during C. elegans embryogenesis, both cell size and cell fate impact the strength of the SAC. Spindle perturbations produce longer SAC-dependent mitotic delays in smaller cells, supporting the long-standing hypothesis that the strength of the SAC can be influenced by cell size (28, 29), while cells in the germline P lineage delay for longer than comparably-sized somatic cells, indicating that SAC strength is further subject to cell fate specific regulation. Variation in the strength of the SAC could arise at many points in the checkpoint signaling mechanism. Conceptually, we envision two primary possibilities: variation at the level of signal generation or variation in the rate of signal degradation. The former encompasses changes in the efficacy of MCC generation at the kinetochore or the number of MCC-generating kinetochores. The latter is a catchall class for changes that occur away from the kinetochore, including differences in the rate of cytoplasmic MCC disassembly or the efficiency of APC/C inhibition. A third possibility could arise downstream of the SAC, if, for example, cell size or cell fate changed the threshold of APC/C substrates at which cells exit mitosis. We think this unlikely, as the timing of mitotic exit is invariant across cells of different sizes and fates when SAC regulation is removed and APC/C activity is unimpeded.
Changes in cell size have been proposed to change the strength of the SAC via the first mechanism, increased signal generation. Smaller cells have a higher ratio of DNA to cytoplasm (28) and, effectively, a higher number of kinetochores per unit cytoplasm, thereby favoring MCC production (29). In cells with monopolar spindles, germline cell fate could act to increase the number of signal generating kinetochores. Cells with monopolar spindles fail to satisfy the SAC due to the persistence of monotelic and syntelic kinetochore-microtubule attachments (68). The presence of these attachments, albeit erroneous, may reduce the proportion of checkpoint-signaling kinetochores, as compared to Nocodazole treated cells, in which microtubule formation is largely blocked (26). Accordingly, longer monopolar mitoses in germline-fated cells could be achieved by further reducing kinetochore-microtubule attachment, thereby increasing the proportion of MCC-producing kinetochores. Alternatively, a microtubule-dependent checkpoint silencing mechanism functions at the kinetochore in C. elegans embryos (69) and rendering this mechanism more or less robust could also affect the proportion of checkpoint-signaling kinetochores in cells with monopolar spindles. While we cannot exclude either of these possibilities, we note that germline blastomeres may also exhibit longer mitotic arrests than somatic cells following Nocodazole treatment, in which very few kinetochore-microtubule attachments are made (29). Thus we favor the hypothesis that cell size sets the kinetochore to cytoplasmic ratio and cell fate acts in parallel, either at the level of signal generation, by modifying MCC production per kinetochore, or downstream of the kinetochore by modulating the cytoplasmic activity of the checkpoint.
Our results indicate that, at the 2-cell embryonic stage, PAR protein mediated cytoplasmic asymmetries underlie differential checkpoint strength between the somatic AB and germline P1 cell. PAR proteins regulate the levels and localization of certain asymmetrically distributed cellular components in 2-cell stage embryos largely via the posterior enrichment of PAR-1, which, in turn, drives anterior localization of two closely related proteins, MEX-5 and MEX-6 (hereafter MEX-5/6), and inhibits their enrichment and activity in the posterior (59). MEX-5/6 anchor and enrich certain factors in the anterior (66, 70) and promote the degradation of germline determinants in AB and/or its progeny (71). Removing anterior PAR proteins permits PAR-1 activity in the anterior, resulting in global inhibition of MEX-5/6 and two cells with certain germline P1 traits. Conversely, eliminating posterior PAR proteins allows MEX-5/6 activity in the posterior and certain somatic AB traits emerge in both cells (55).
Our results raise the question of why removing PAR-1, which is usually undetectable in AB, increased the duration of monopolar mitoses in AB, while depleting PAR-2 had a minimal effect. This result stands in contrast to a well-characterized asymmetric cellular behavior regulated by PAR proteins, cell cycle asynchrony (66, 70, 72, 73). If checkpoint activity were regulated in an analogous fashion, we would expect loss of par-1 and par-2 to result in shorter mitotic delays in P1, leaving AB largely unaffected. Instead, our results are consistent with a model in which a checkpoint-promoting factor (“factor X”) is enriched in P1 via a PAR-1-dependent mechanism. In the absence of PAR-2, anterior PAR proteins expand into the posterior, but are present in a graded fashion, and the posterior enrichment of some germline factors (e.g. P granules) is often maintained (58, 74). In addition, although PAR-1 is no longer asymmetric in par-2(-) embryos, its activity is not lost (58, 75, 76). As differential mitotic delays between AB and P1 cells in par-2(lw32) embryos were entirely dependent on PAR-1, we conclude that cytoplasmic asymmetries downstream of PAR-1, and partially independent of PAR-2, are largely responsible for posterior enrichment of factor X. In the absence of PAR-6, PKC-3 or PAR-1, this factor X is equally inherited by AB and P1, thereby increasing checkpoint activity in AB while maintaining it in P1 (Fig. 4e). This model requires that levels of factor X are not limiting for checkpoint activity and that factor X is not degraded in the absence of PAR-1, at least at the 2-cell stage. As in par-1 mutants, degradation of germline factors is delayed such that germplasm proteins persist in all cells until the 4-cell stage (52), this latter condition is not without precedent.
Altogether, our results suggest that a checkpoint-regulating factor (or factors) is partitioned during the asymmetric division of germline blastomeres, downstream of PAR-mediated cell polarity, such that germline cells possess a stronger SAC relative to their nuclear to cytoplasmic ratio than their somatic siblings. SAC activity may be further tuned within each lineage as development progresses. Future work to identify the relevant molecular asymmetries between somatic and germline cells driving differential SAC activity will permit further investigation of this model and identification of where PAR-mediated cell polarity and checkpoint regulation intersect.
Materials and Methods
C. elegans strains and culture
All strains were maintained at 15°C on NGM plates, seeded with E. coli bacteria (OP50) following standard procedures (77). All genotypes are listed in Supplementary Table S1. L4 stage larvae were transferred to fresh OP50 plates at 15°C for 2 days, after which time embryos were harvested from gravid adults for imaging. RNAi depletions were performed at 15°C by feeding (78) as follows. Synchronized L1 larvae were obtained by sodium hypochlorite treatment (1.2% NaOCl, 250mM KOH) and were plated onto NGM plates containing 1.5mM IPTG and 25μg/ml Carbenicillin, seeded with HT115 bacteria containing the empty feeding vector L4440. Three days later early L4 larvae were transferred to fresh RNAi feeding plates seeded with bacteria expressing doubled-stranded RNA for gene inactivation or the empty vector L4440 for controls. Embryos were collected for imaging three days later. To isolate homozygous par-2(lw32) animals, L1 larvae on L4440 plates were shifted to 25°C for 24 hours, which allowed identification of homozygous animals based on the linked, recessive, temperature-sensitive allele unc-45(e286). Unc larvae were transferred to RNAi feeding plates, returned to 15°C and imaged three days later. Control genotypes (UM471 or UM628, see Supplementary Table S1) were similarly treated. The following clones from the Arhinger RNAi library were used: sjj_C50F4.11 (mdf-1/Mad1), sjj_K10B2.5 (ani-2), sjj_ZK675.1 (ptc-1), sjj_C38C10.4 (gpr-1/2), sjj_T26E3.3 (par-6), sjj_F09E5.1 (pkc-3), sjj_H39E23.1 (par-1), and sjj_F58B6.3 (par-2). The clone targeting C27D9.1 was kindly provided by Dr. B. Lacroix. All clones were verified by sequencing.
Embryo mounting and live-imaging
Embryos were harvested by cutting open gravid hermaphrodites in M9 buffer using two 25-gauge needles. Embryos were transferred by mouth pipet to a 2% agarose pad, positioned using an eyelash and covered with a coverslip. The chamber was backfilled with M9 and sealed using VaLaP (1:1:1 Vaseline, lanolin and paraffin). Embryos were mounted at room temperature in solutions that had been cooled to 15°C. Slides were transferred to a temperature-controlled imaging chamber set to 26°C and time-lapse images were acquired using either a Cell Observer SD spinning disc confocal (Zeiss; Yokogawa) equipped with a stage-top incubator (Pecon) or a DeltaVision inverted microscope (GE Healthcare Life Sciences) equipped with a WeatherStation environmental chamber (PrecisionControl). Spinning disc images were captured using a AxioCam 506 Mono camera (Zeiss) and a 63x/1.4 NA Plan Apochromat DIC oil immersion objective (Zeiss) in Zen software (Zeiss). DeltaVision images were captured using a Coolsnap HQ2 CCD camera (Photometrics) and a 60x/1.42 NA Plan Apo N oil immersion objective (Olympus) in softWoRx software (GE Healthcare Life Sciences). Time-lapse acquisitions were 45 to 60 minutes in duration, with approximately 30 second time sampling. At each time point a roughly 20μm thick z-stack was acquired, using either 1 or 1.5μm sectioning.
Image processing and measurements
Image processing and analysis was carried out in ImageJ (NIH). The duration of bipolar, monopolar and tbb-2(ts) mitoses were determined manually by monitoring H2B::mCH fluorescence. NEBD was defined as the first frame in which non-incorporated H2B::mCH was lost from the nuclear area. DECOND was defined as the first frame in which the distribution of H2B::mCH shifted from bright and compact to fainter and diffuse. Maximal z-stack projections of unprocessed image files were scored. Single z-slices were examined if projected images were ambiguous. For all strains in which β-tubulin::GFP was also present (all experiments excepting par-1(it51) and its associated controls), the timing of NEBD was confirmed by the appearance of microtubules in the nuclear space and the start of DECOND was concomitant with the start of spindle disassembly and shrinking of the spindle pole. To exclude potentially confounding effects of accumulated mitotic errors due to any partial conditionality of our temperature sensitive alleles, only cells in which the preceding parental division was observed to be normal were analyzed. AB and P1 cell cycle duration was measured as the time from P0 metaphase to NEBD in each cell respectively, by manually monitoring H2B::mCH fluorescence. P0 metaphase was defined as the last time point prior to visible chromosome separation. NEBD was defined as above. To measure nuclear area, a sum projection of the central 3 z-slices of the nucleus of interest was processed and segmented. The average of the dimensions of a bounding box fit to the segmented nucleus was taken as representative of nuclear diameter and nuclear area was then calculated as the area of the corresponding circle (Supplementary Fig. S2a,b). Reported values reflect the average of measurements made at three time points, 1 to 2 minutes prior to NEBD. To determine the relative volume of cells in each lineage from the 2 to 16-cell stage, the position of the mitotic spindle midpoint along the division axis, relative to the midpoint of the dividing cell was used to determine what proportion of each dividing cell was allocated to each daughter (Supplementary Fig. S2c,d). The Plot Profile tool in ImageJ was used to generate signal intensity profiles along a line drawn parallel to the division axis, through the segregating sister nuclei, in cells in which the cell membrane was labeled with mNeonGreen::PH (mNG::PH) and nuclei were marked by H2B::mCH. Spindle displacement was defined as the offset of the center point between the two membrane mNG::PH peaks versus the center point between the two H2B::mCH peaks. When segregating sister nuclei were not in the same xy plane (i.e. the cell division axis was tilted relative to the plane of imaging), the Stack Reslice tool in ImageJ was used to construct an image of the long axis of the dividing cell from the encompassing z slices. Measurements were made at three time points during anaphase, after the start of membrane ingression. Nearly identical volume relationships were obtained when cell volume was calculated from cross-sectional area and cell height measurements using the mNG::PH membrane signal to manually outline cells (data not shown). The cell volume of AB and P1 was calculated by combining measurements of spindle displacement in P0, with measurements of embryo volume. Spindle displacement was measured using the Plot Profile tool in ImageJ to generate signal intensity profiles along the anterior to posterior axis of the embryo. The spindle midpoint was defined as the center point between the two peaks of β-tubulin::GFP fluorescence, corresponding to the spindle poles, and/or the two peaks of H2B::mCH fluorescence, corresponding to the segregating sister nuclei. The position of the spindle midpoint relative to the embryo midpoint was used as an indication of what proportion of the zygote would be inherited by each cell. Embryo volume was measured by manually outlining each embryo using the Polygon Selection Tool in ImageJ and fitting an ellipse to the resulting area. Embryo volume was then calculated as an ellipsoid of the same dimensions. Final values reflect the average of three measurements made at three different time points during anaphase in P0.
Graphing and statistical analysis
Graphs were generated in Microsoft Excel or MATLAB (MathWorks). Statistical analysis was carried out in MATLAB. Comparison of multiple means was performed using an Anova1 with a Tukey-Kramer post-hoc test. Linear least squares regression models were calculated in MATLAB, and Pearson’s linear correlation coefficient (r) was reported, with statistical significance assessed using Student’s t-distribution. Regression coefficients were compared by bootstrapping, using a custom MATLAB script. For data presented in Fig. 3c,d, bootstrap analysis was performed manually in Excel, by calculating the difference in the duration of monopolar mitoses for all possible pairwise comparisons within a given size range. All possible outcomes were displayed as a boxplot graph and means were compared by an Anova1 with Tukey-Kramer post-hoc test. “Small” embryos are those in which AB is within ± 1 standard deviation (s.d.) of the size of a Control(RNAi) P1 cell. “Large” embryos are those in which P1 is within ± 1 s.d. of the size of a Control(RNAi) AB cell. “Normal” embryos are those in which AB is within ± 1 s.d. of the size of a Control(RNAi) AB cell. “P1” cell size compares AB and P1 cells that are both within ± 1 s.d. of the size of a Control(RNAi) P1 cell. “AB” cell size compares AB and P1 cells that are both within ± 1 s.d. of the size of a Control(RNAi) AB cell. For Fig. 3f,g, only gpr-1/2(RNAi), par-6(RNAi) and par-1(RNAi) embryos in which the position of the mitotic spindle in P0 was roughly centered (50±2% of embryo length) were considered. For data presented in Fig. 4b, for each RNAi treatment, AB and P1 cells were pooled and only cells within ±1 standard deviation of the average cell size for that depletion were considered. This same size range was applied to the total data set of normally polarized and fated AB and P1 cells (Control(RNAi), ani-2(RNAi), ptc-1(RNAi), C27D9.1(RNAi), gpr-1/2(RNAi)). Thus all cells, both depleted and control, AB and P1, analyzed for a given RNAi treatment are approximately the same size. For par-1(RNAi), the small size of the data set required increasing the size range to within ±2 standard deviations of the mean. For all Figures n.s. = p>0.05, * = p<0.05, ** = p<0.01 and *** = p<0.001.
Author Contributions
A.R.G. designed, and carried out all experiments, with the exception of the par-1(it51) experiment which was carried out by V.P. A.R.G. analyzed the data, and wrote the manuscript with input from J-C.L. and P.S.M.
Supplemental Movies
All time-lapse movies were acquired on a Cell Observer SD spinning disc confocal (Zeiss; Yokogawa) equipped with a stage-top incubator (Pecon) set to 26°C, using an AxioCam 506 Mono camera (Zeiss), with 4x4 binning, and a 63x/1.4 NA Plan Apochromat DIC oil immersion objective (Zeiss) in Zen software (Zeiss). Images were acquired every 30 seconds, with the following exceptions: Movie S3 (mdf-1(RNAi)) every 33 seconds; Movie S4 (gpr-1/2(RNAi)), every 38 seconds. Images were processed using ImageJ (NIH). Maximum intensity projections of 18 z-slices (1 μm sectioning) centered on the embryo of interest are shown. Images were adjusted for brightness and contrast and scaled 4 times to adjust for file compression during .avi conversion. H2B::mCH is shown in cyan and β-tubulin::GFP is shown in red. All embryos are oriented with anterior to the left. Scale bar = 10 μm. Time stamp is in min:sec. Movies play at 4 frames per second, excepting Movie S4 (gpr-1/2(RNAi)), which plays at 3 frames per second.
Movie S1. zyg-1(ts)-induced monopolar spindles in 2- and 4-cell stage embryos.
Movie S2. tbb-2(ts)-induced spindle defects in 2- and 4-cell stage embryos.
Movie S3. zyg-1(ts)-induced monopolar spindles in 2- and 4-cell stage embryos following RNAi depletion of MDF-1.
Movie S4. zyg-1(ts)-induced monopolar spindles in a 2-cell stage embryo following RNAi depletion of GPR-1/2.
Movie S5. zyg-1(ts)-induced monopolar spindles in a 2-cell stage embryo following RNAi depletion of PAR-6.
Acknowledgements
We thank A. Golden, B. Goldstein, D. Dickinson and the Caenorhabditis Genetics Center for strains, C. Charbonneau of IRIC’s Bio-imaging facility for technical assistance and E. D. Salmon and members of the Labbé and Maddox labs for helpful discussions. This work was funded by a grant from the CIHR (MOP-115171) to J-C.L. and P.S.M. J-C.L. holds the Canada Research Chair in Cell Division and Differentiation. P.S.M. is supported as a William Burwell Harrison Fellow of Biology. IRIC is supported in part by the Canada Foundation for Innovation and the Fonds de Recherche du Québec – Santé.
References
- 1.↵
- 2.↵
- 3.↵
- 4.↵
- 5.↵
- 6.↵
- 7.↵
- 8.↵
- 9.↵
- 10.↵
- 11.↵
- 12.↵
- 13.↵
- 14.↵
- 15.↵
- 16.↵
- 17.↵
- 18.↵
- 19.↵
- 20.↵
- 21.↵
- 22.↵
- 23.↵
- 24.↵
- 25.↵
- 26.↵
- 27.↵
- 28.↵
- 29.↵
- 30.↵
- 31.↵
- 32.↵
- 33.↵
- 34.↵
- 35.↵
- 36.↵
- 37.↵
- 38.↵
- 39.↵
- 40.↵
- 41.↵
- 42.↵
- 43.↵
- 44.↵
- 45.↵
- 46.↵
- 47.↵
- 48.
- 49.↵
- 50.↵
- 51.
- 52.↵
- 53.↵
- 54.↵
- 55.↵
- 56.↵
- 57.↵
- 58.↵
- 59.↵
- 60.↵
- 61.↵
- 62.↵
- 63.↵
- 64.
- 65.↵
- 66.↵
- 67.↵
- 68.↵
- 69.↵
- 70.↵
- 71.↵
- 72.↵
- 73.↵
- 74.↵
- 75.↵
- 76.↵
- 77.↵
- 78.↵