Cytokinesis requires activation of the GTPase RhoA. ECT-2, the exchange factor responsible for RhoA activation, is regulated to ensure spatiotemporal control of contractile ring assembly. Centralspindlin, composed of the Rho family GTPase-activating protein (RhoGAP) MgcRacGAP/CYK-4 and the kinesin MKLP1/ZEN-4, is known to activate ECT-2, but the underlying mechanism is not understood. We report that ECT-2-mediated RhoA activation depends on the ability of CYK-4 to localize to the plasma membrane, bind RhoA, and promote GTP hydrolysis by RhoA. Defects resulting from loss of CYK-4 RhoGAP activity can be rescued by activating mutations in ECT-2 or depletion of RGA-3/4, which functions as a conventional RhoGAP for RhoA. Consistent with CYK-4 RhoGAP activity contributing to GEF activation, the catalytic domains of CYK-4 and ECT-2 directly interact. Thus, counterintuitively, CYK-4 RhoGAP activity promotes RhoA activation. We propose that the most active form of the cytokinetic RhoGEF involves complex formation between ECT-2, centralspindlin and RhoA.https://doi.org/10.7554/eLife.08898.001
Cell division is a process in which a cell splits to form two daughter cells. In most cases, the cell first duplicates its genetic material and then the two copies are pulled to opposite ends of the cell. A ring of protein filaments—called the contractile ring—then assembles to form a band around the cell at the site of the division. This ring contracts and the force generated separates the cells in a step known as cytokinesis.
A protein belonging to the Rho family, called RhoA, is essential for cytokinesis because it controls the formation of the contractile ring. Rho proteins are switched on by the activities of other proteins called guanine nucleotide exchange factors. Another group of proteins known as ‘GTPase activating proteins’ (or GAPs for short) generally act to promote the ability of Rho proteins to turn themselves off.
In animals and other multicellular organisms, a GAP called CYK-4 largely concentrates on the spindle midzone, but some of the protein also moves to part of the cell membrane near the future site of cell division. It binds to a guanine nucleotide exchange factor called ECT-2 to switch RhoA on, which in turn promotes the formation of the contractile ring. However, it is not clear why a protein that activates RhoA is also able to trigger its inactivation.
In this study, Zhang and Glotzer studied cell division in a roundworm called Caenorhabditis elegans. The experiments show that cells that lacked the GAP activity of CYK-4 were unable to complete cytokinesis because RhoA was not fully switched on. This requirement could be bypassed in cells with mutant forms of ECT-2 that were overactive. Therefore, an activity that was thought to inactivate RhoA actually promotes its activation. Further experiments show that the section (or ‘domain’) of CYK-4 that has GAP activity interacts directly with the guanine nucleotide exchange domain of ECT2. Zhang and Glotzer suggest that this interaction stimulates ECT2 and thereby promotes the activation of RhoA.
Further experiments will reveal how CYK-4 stimulates ECT-2. In addition, it will be important to determine whether other proteins with GAP domains also work in this unconventional way.https://doi.org/10.7554/eLife.08898.002
Cell division involves the myosin-mediated contraction of an actin-based contractile ring. In metazoans, contractile ring assembly involves activation of the GTPase RhoA. RhoA directly activates formin-mediated actin polymerization and indirectly promotes myosin activation. As the contractile ring must assemble at the correct position and at the correct time, between the segregating chromosomes in anaphase, RhoA activation is subject to multiple regulatory mechanisms (see Green et al., 2012 for review).
The primary direct activator of RhoA during cytokinesis is the RhoGEF ECT-2. ECT-2 contains N-terminal BRCT domains and a C-terminal RhoGEF domain (Kim et al., 2005; Zou et al., 2014). Presumably as a consequence of being autoinhibited, ECT-2 function depends on activators. One of the important activators of ECT-2 during cytokinesis is the centralspindlin complex, which is a heterotetramer containing a dimeric kinesin, ZEN-4 (aka MKLP1, Pavarotti) and dimeric CYK-4 (aka MgcRacGAP, Tum/RacGAP50C) (Mishima et al., 2002). Centralspindlin organizes the spindle midzone and directly recruits numerous regulators of cytokinesis, including ECT-2, to this location (Burkard et al., 2009; Wolfe et al., 2009). ECT-2 and centralspindlin are conserved among—and restricted to—metazoans (Frédéric et al., 2013) (unpublished results). Though these proteins are conserved, their names are distinct in each organism. For simplicity, Caenorhabditis elegans names will be used throughout this manuscript with the exception that we will refer to RHO-1 with the more common name RhoA.
Recruitment of ECT-2 to the spindle midzone involves regulated binding between ECT-2 and CYK-4. The BRCT domains of ECT-2 bind to CYK-4 phosphorylated by PLK-1 (Burkard et al., 2009; Wolfe et al., 2009). CYK-4 phosphorylation occurs in a cell cycle and microtubule-regulated manner. Furthermore, CDK-1 phosphorylation of ECT-2 inhibits the ECT-2-CYK-4 interaction during metaphase and inactivates a membrane binding motif within ECT-2 (Yüce et al., 2005; Su et al., 2011). The phosphorylation-dependent interaction between ECT-2 and centralspindlin is required for RhoA activation during cytokinesis in human cells (Burkard et al., 2007; Wolfe et al., 2009).
Centralspindlin also localizes in trace, but biologically relevant, amounts on the cell membrane. Centralspindlin accumulation to the midzone and the membrane are independently regulated by its oligomerization, which is inhibited by a 14-3-3 protein and promoted by the chromosome passenger complex (Douglas et al., 2010; Basant et al., 2015).
The CYK-4 subunit of centralspindlin contains an evolutionarily conserved Rho family GTPase-activating protein (RhoGAP) domain (Jantsch-Plunger et al., 2000). The function of this domain has been examined in a number of different contexts. In vitro, the GAP domain of CYK-4 efficiently activates the GTPase activity of the Rho-family GTPases, CED-10/Rac1 and CDC-42. CYK-4 also has GAP activity towards RhoA, but it is far less active towards RhoA as compared to Rac1 and CDC-42 (Touré et al., 1998; Jantsch-Plunger et al., 2000; Bastos et al., 2012). Despite extensive effort, there is no consensus for the biological role of this GAP activity (see White and Glotzer, 2012 for review). In some cell types, the GAP activity appears dispensable (Goldstein et al., 2005; Yamada et al., 2006), in others it appears to be important to negatively regulate Rac1 (D'Avino et al., 2004; Canman et al., 2008; Bastos et al., 2012), whereas in yet others it appears to promote RhoA activation (D'Avino et al., 2004; Zavortink et al., 2005; Loria et al., 2012). System-specific differences may underlie some of these diverse results. However, these studies differ in the mutations used to assess the function of the GAP domain, which is likely to affect the results. In addition, some of the controversy may be due to misinterpretation of indirect effects.
The function of the RhoGAP domain has been examined in C. elegans embryos in some detail. These studies have focused largely on a temperature-sensitive, separation-of-function substitution mutation, E448K, that lies in the RhoGAP domain of CYK-4, cyk-4(or749ts) (Canman et al., 2008). The mutant protein can complex with the centralspindlin kinesin, ZEN-4, and bundle microtubules in the central spindle. However, cytokinesis does not proceed to completion in these embryos. Interestingly, depletion of CED-10/Rac1 or the actin nucleator subunit ARP-2 enables these embryos to complete cytokinesis (Canman et al., 2008). These genetic interactions have been interpreted to indicate that the GAP domain of CYK-4 is important to keep CED-10/Rac1 inactive and prevent the accumulation of branched actin in the equatorial region (Canman et al., 2008). However, subsequent analysis demonstrated that cyk-4(or749ts); ced-10(−) embryos are phenotypically abnormal (Loria et al., 2012). In particular, cyk-4; ced-10 mutant embryos have reduced accumulation of RhoA effectors as compared to ced-10 mutants alone (Loria et al., 2012). These results suggest that this mutation in CYK-4 affects more than the RhoGAP activity of CYK-4 or that CED-10/Rac1 is not the relevant target of the GAP domain, or both.
Analysis of centralspindlin function in C. elegans embryos is impeded by the existence of a second, parallel pathway that promotes RhoA activation in the early embryo. Upon fertilization, embryos exhibit RhoA-dependent contractility that promotes embryo polarization and culminates in the formation of a transient furrow known as the pseudocleavage furrow. This wave of contractility requires a poorly conserved protein known as NOP-1 and is largely independent of centralspindlin (Tse et al., 2012). Cytokinetic contractility, on the other hand, involves both NOP-1 and centralspindlin. However, nop-1 is nonessential; loss of function mutants are viable and fertile (Rose et al., 1995). Cytokinesis proceeds to completion in the absence of NOP-1, although furrow initiation is slightly delayed and RhoA accumulates to lower levels at the cleavage furrow (Tse et al., 2012). Due to its role in RhoA activation, polarization in NOP-1-deficient embryos is also delayed. Mutational inactivation of NOP-1 permits direct analysis of centralspindlin-dependent furrow formation. Notably, when NOP-1 is inactivated, cyk-4(or749ts) embryos are completely defective in RhoA activation (Tse et al., 2012).
The CYK-4 GAP domain is adjacent to a C1 domain that mediates membrane localization of centralspindlin (Lekomtsev et al., 2012). Given that the cyk-4(or749ts) substitution renders the protein thermosensitive, it is possible that these phenotypes are not the sole consequence of loss of GAP activity; this mutation could affect other functions of the GAP domain, the adjacent C1 domain may also be affected. To clarify these issues, we performed a targeted structure-function analysis of the C1 and GAP domains of CYK-4. We demonstrate that the cyk-4(or749ts) allele indeed affects its ability to associate with the membrane and show that this activity contributes to RhoA activation. We further show that the active site of the GAP domain contributes to the accumulation of downstream effectors of RhoA and RhoA-dependent contractility. Furthermore, we find that the catalytic domains of CYK-4 and ECT-2 directly interact in vitro. Finally, we show that hypomorphic defects in CYK-4-mediated RhoA dependent contractility can be suppressed by either loss of the RhoGAP activity provided by RGA-3/4 or by either of two activating mutations in ECT-2. These activating mutations in ect-2 rescue cyk-4(or749ts) and GAP-deficient CYK-4. Our results indicate that CYK-4 GAP activity is involved in ECT-2-mediated RhoA activation.
We sought to conduct a structure-function analysis of CYK-4 to determine the individual contributions of the GAP and C1 domains of CYK-4. We established a rescue assay based on single copy integrants of GFP-tagged CYK-4 transgenes driven by the cyk-4 promoter inserted at a defined locus in the C. elegans genome using Mos1-mediated integration (Figure 1—figure supplement 1) (Frøkjaer-Jensen et al., 2012). The transgenes were expressed at consistent levels (Figure 1—figure supplement 2). The transgenes were rendered resistant to an RNAi construct that could effectively deplete endogenous CYK-4 by targeting the 3′ UTR and portions of the coding sequence (Figure 1—figure supplement 2). By combining the appropriate mutant transgene with RNAi to specifically deplete endogenous CYK-4, we obtained embryos that express CYK-4MUT. In this and all subsequent experiments, when a given variant is assayed, endogenous CYK-4 is depleted by RNAi; these will be referred to as cyk-4mut embryos. We first sought to validate the rescue assay, by assaying cyk-4E448K embryos and found that they closely phenocopy cyk-4(or749ts) embryos in which endogenous CYK-4 has the E448K substitution (Figure 1—figure supplement 3). This phenocopy indicates functional depletion of endogenous CYK-4. Additionally, a wild-type transgene was fully functional as it could complement a large deletion in CYK-4 to viability and fertility (Figure 6—figure supplement 1).
Consistent with the C1 domain promoting membrane association, CYK-4∆C1 does not accumulate on ingressing cleavage furrows (Figure 1A,B). As cyk-4(or749ts) is temperature sensitive, we considered the possibility that the thermosensitivity also destabilizes the C1 domain that lies adjacent to the CYK-4 GAP domain (Figure 1—figure supplement 1). Indeed, at the restrictive temperature, CYK-4E448K exhibits a similar defect in localization as CYK-4∆C1 (Figure 1A,B). CYK-4 also associates with the membrane in the germline and indirect evidence suggests that this localization is compromised in embryos expressing CYK-4E448K (Zhou et al., 2013). We generated strains in which both the endogenous cyk-4 and the GFP-tagged transgene contained the E448K mutation. At the permissive temperature, CYK-4E448K membrane recruitment was readily detected in the germline, however this localization was lost as animals were shifted to the restrictive temperature (Figure 1—figure supplement 4). Similarly, the C1 domain is required for membrane localization in the gonad (Figure 1—figure supplement 4). Thus, CYK-4E448K impairs membrane localization, and it is likely to impair the function of both its GAP and C1 domains.
We compared the progression of cytokinesis in embryos expressing CYK-4WT, CYK-4∆C1, and CYK-4E448K. CYK-4∆C1 embryos exhibit a cytokinetic defect that is largely similar to that of cyk-4E448K embryos. In particular, furrow ingression is slow and incomplete (Figure 1C,F). However, subtle differences were detected; the onset of significant furrow ingression is delayed relative to controls in cyk-4E448K embryos but not in cyk-4∆C1 embryos. This suggests that the CYK-4E448K phenotype could reflect a compound defect, rather than a sole defect in the ability to associate with the membrane.
Cleavage furrow ingression in C. elegans depends on the combined action of centralspindlin and a non-essential protein, NOP-1 (Tse et al., 2012). To determine whether the C1 domain is essential for centralspindlin-dependent furrow ingression, we expressed CYK-4WT, CYK-4∆C1, and CYK-4E448K in embryos that lack the NOP-1-dependent pathway for furrow ingression. As expected, CYK-4WT supports full furrow ingression in this sensitized background. In stark contrast, neither CYK-4∆C1 nor CYK-4E448K support detectable furrow ingression in the absence of NOP-1 activity (Figure 1D,G).
Previous studies demonstrated that loss of function mutations in ced-10/rac-1 or depletion of the protein partially suppress the cytokinesis defect in cyk-4(or749ts) embryos (Figure 1E,H). However, as described above, it is important to examine the extent of suppression in the absence of the parallel, NOP-1-dependent, pathway. We therefore examined whether ced-10/rac-1 loss of function mutations could suppress the cytokinesis defect in cyk-4∆C1. Although cleavage furrows in cyk-4∆C1; ced-10(n1993) embryos ingress somewhat more deeply than cyk-4∆C1 embryos, they do not complete cytokinesis (Figure 1E,H). The simplest interpretation of these results is that although CYK-4E448K diminishes membrane association of CYK-4, it may retain some function at the restrictive temperature such that it facilitates the abscission step in cyk-4(or749ts); ced-10(n1993) embryos, as CYK-4-mediated membrane association is essential for completion of cytokinesis in cultured human cells (Lekomtsev et al., 2012). Importantly, when NOP-1 activity is compromised, inactivation of CED-10/Rac1 does not suppress the defect in furrow ingression caused by CYK-4E448K (Figure 1J,I), suggesting that CYK-4 does not act directly on CED-10.
To begin to determine the role of the catalytic activity of CYK-4 during cytokinesis, we first studied the consequence of mutating the highly conserved catalytic arginine that stabilizes the transition state during GTP hydrolysis. Substitution of the catalytic arginine with alanine strongly attenuates GAP activity against Rac in a variety of CYK-4 orthologs and is widely used to inactivate Rho family GAPs (Rittinger et al., 1997; Yamada et al., 2006; Miller and Bement, 2009; Bastos et al., 2012; Zanin et al., 2013). As expected, CYK-4R459A GAP domain retains the ability to bind to RhoA•GTP, demonstrating that the protein is well folded in vitro (Figure 2—figure supplement 1). While CYK-4 GAP exhibits GAP activity towards both RhoA and CED-10/Rac1, CYK-4R459A GAP lacks detectable GAP activity towards either GTPase (Figure 2—figure supplement 2). In order to determine if catalytic activity is required for viability, we used a strain heterozygous for a deletion mutant of CYK-4, cyk-4(ok1034). One quarter of the embryos from these heterozygous hermaphrodites contain maternally provided CYK-4 and lack zygotic CYK-4. These zygotic null embryos fail to hatch and arrest with a variety of terminal phenotypes (Figure 2A, left). Many, but not all, embryos contain muscle tissue and have undergone partial morphogenesis. The embryos also contain enlarged cells, likely due to defects in cytokinesis (Sugimoto et al., 2001). We introduced the GAP-defective transgene into this strain. Remarkably, cyk-4(ok1034); cyk-4R459A animals hatch and develop to adulthood. However, these animals are sterile (Figure 2A, middle). Thus, the GAP activity of CYK-4 is not essential for post embryonic development but it has an important role in gonad development, likely due to a requirement for post-embryonic cell proliferation in the germline. As a consequence, it is not possible to use classical genetic tools to obtain embryos in which CYK-4R459A is the sole form of CYK-4.
To study the role of the CYK-4 GAP activity during embryogenesis, we used the aforementioned assay (Figure 1—figure supplement 1). Embryos, expressing only CYK-4R459A, are largely normal during the initial stages of the first cell cycle. They undergo pseudocleavage, mitotic spindle assembly, chromosome segregation, central spindle assembly during anaphase, and CYK-4R459A becomes highly enriched on the spindle midzone (Figure 2B). Cleavage furrow initiation occurs and the furrow ingresses at near wild-type rates to near completion. However, cytokinesis does not complete and the furrow ultimately regresses; this phenotype was fully penetrant (Figure 2C); these embryos also fail to complete cytokinesis following meiosis II (data not shown). These results suggest that a late step in cytokinesis is most sensitive to loss of CYK-4 GAP activity. This phenotype is distinct from that of cyk-4(or749ts) embryos–cleavage furrows in cyk-4R459A embryos ingress more rapidly and more deeply than cyk-4E448K embryos (Figure 1C). We assessed the ability of CYK-4R459A to associate with membrane during furrow ingression (Figure 2B,D). CYK-4R459A hyper accumulates on the membrane as compared to WT CYK-4; this localization suggests that CYK-4R459A is well folded in vivo. Therefore, the cytokinetic defect in this strain is unlikely to be an indirect consequence of a failure of CYK-4 to localize to the membrane.
To extend these results, confirm that CYK-4 must interact with Rho family GTPases during cytokinesis, and eliminate the possibility that the phenotype of CYK-4R459A is due to enhanced binding of CYK-4 to active RhoA, we engineered mutations in CYK-4 that reduce its binding to RhoA and other GTPases (Rittinger et al., 1997; Sekimata et al., 1999). Two conserved, surface exposed, basic residues in the RhoA interface (K495, R499) (Figure 3—figure supplement 1) were charge reversed to glutamic acid, generating CYK-4EE, and characterized in the transgenic rescue assay. Embryos expressing only CYK-4EE, like those expressing CYK-4R459A and CYK-4∆C1, exhibit fully penetrant embryonic lethality (Figure 3—figure supplement 1). CYK-4EE exhibits reduced binding to RhoA in vitro (Figure 2—figure supplement 1), and it does not exhibit membrane hyperaccumulation in vivo (Figure 3A,B). Interestingly, cyk-4EE embryos exhibit a stronger furrow ingression defect than cyk-4R459A embryos, as furrow ingression is slower and less complete (Figure 3C,D). Importantly, NOP-1 depletion from cyk-4EE embryos largely eliminates furrow ingression (Figure 3C,D). Thus, Rho GTPase binding by CYK-4 is essential for centralspindlin-mediated cytokinetic ingression.
We next sought to determine the Rho family GTPase to which CYK-4 must bind to fulfill its function in vivo. If furrow formation is dependent on CYK-4 binding to either CED-10/Rac1 or CDC-42 to generate a positive regulatory complex, then inactivation of these GTPases would be predicted to cause a phenotype at least as severe as a mutation that weakens the GTPase binding site of CYK-4. However, mutation of CED-10/Rac1, or depletion of CDC-42, does not affect the rate of cleavage furrow ingression (Jantsch-Plunger et al., 2000; Loria et al., 2012), even when combined with mutations in NOP-1 (Figure 3E,F). Indeed, cytokinesis occurs efficiently and proceeds to completion in embryos in which NOP-1, CED-10/Rac1, and CDC-42 are simultaneously inactivated (Figure 3E,F). We infer, therefore, that RhoA is the relevant GTPase that CYK-4 binds to promote cleavage furrow formation. Due to its direct role in furrow ingression, it is not possible to test RhoA in the same manner.
We next sought to determine how the GAP activity of CYK-4 promotes cytokinesis. Previous studies proposed at least three models for the phenotype seen in cyk-4R459A embryos. First, the GAP domain could function as canonical GAP that acts on RhoA, causing CYK-4 GAP-deficient embryos fail to complete cytokinesis because of a requirement for RhoA inactivation at late cytokinesis. Second, CED-10/Rac1 could be an important target of CYK-4 GAP activity, causing CYK-4 GAP deficient embryos to accumulate ectopic Rac1 activity that interferes with cytokinesis. Third, although it is counterintuitive, CYK-4 GAP activity could somehow promote RhoA activation, and therefore the CYK-4 GAP deficient embryos may fail cytokinesis due to incomplete RhoA activation. We sought to distinguish between these alternatives.
The first and third models make opposite predictions for the outcome of experiments in which RhoA levels are perturbed (Figure 4A). If the failure to complete cytokinesis in cyk-4R459A embryos is due to hyperactivation of RhoA, as would be predicted from the canonical model for the function of a RhoA GAP, then a reduction in active RhoA levels might ameliorate the defect. Conversely, if the GAP active site promotes RhoA activation, then the reduction of RhoA activity would be predicted to exacerbate the phenotype of cyk-4R459A. To distinguish between these models, we reduced RhoA levels by mutationally inactivating NOP-1. As expected, all control embryos (nop-1(it142); cyk-4WT) complete cytokinesis (Figure 4Bi). Surprisingly, nop-1(it142); cyk-4R459A mutant embryos exhibit extremely weak furrow ingression; furrows in these embryos ingressed less than ∼10% of egg width (Figure 4Biii,C). This result supports models in which CYK-4 GAP activity is involved in RhoA activation.
Previous studies have implicated CED-10/Rac1 as a target of CYK-4 GAP activity, although these studies utilized the cyk-4(or749ts) mutation that impairs membrane localization of CYK-4 (Figure 1A,B, Figure 1—figure supplement 4A). Therefore, we addressed whether loss of function mutations in ced-10 affect cytokinesis in cyk-4R459A embryos. Interestingly, we found that all ced-10; cyk-4R459A embryos fully ingress and 75% complete cytokinesis (Figure 4Biv,C), suggesting significant, albeit incomplete rescue. Depletion of ARX-2, a component of the Arp2/3 complex, a downsteam effector of Rac GTPases, provides similar rescue as mutation in ced-10 (Figure 4—figure supplement 1). Two other Rac related proteins, RAC-2 and MIG-2, could, in principle, be additional targets of the CYK-4 GAP domain. However, depletion of RAC-2 does not rescue completion of cytokinesis in cyk-4R459A embryos (Figure 4—figure supplement 1). Furthermore, gain of function mutations in mig-2 (Zipkin et al., 1997) do not cause cytokinesis defects, even in sensitized genetic backgrounds (Figure 4—figure supplement 2).
Mutations in ced-10 also slightly increase the extent of furrow ingression in cyk-4∆C1 embryos (Figure 1E,H). To more stringently test whether the GAP activity of CYK-4 is linked with CED-10/Rac1 inactivation, we assessed the progression of cytokinesis in embryos that lack NOP-1 function. Crucially, nop-1; ced-10 embryos complete cytokinesis (Figure 4Bvi). If CED-10/Rac1 inactivation is the primary function of the CYK-4 GAP domain, then CYK-4 GAP activity would be predicted to be dispensable in nop-1; ced-10 embryos. However, in stark contrast to this prediction, nop-1; ced-10; cyk-4R459A embryos fail to form ingressing cleavage furrows altogether (Figure 4Bv,C). Significant furrow ingression is not restored by depletion of either RAC-2 or ARX-2 in nop-1; ced-10; cyk-4R459A embryos, suggesting that the cytokinesis defect is not due to activation of other Rac-family proteins (Figure 4—figure supplement 3). These data demonstrate that the catalytic activity of the CYK-4 GAP domain must have a function that is distinct from maintaining CED-10/Rac1 in an inactive state.
To further test models in which CYK-4 RhoGAP catalytic activity is important to either promote RhoA activation or to promote RhoA inactivation, we examined the consequence of depletion of the predominant RhoA GAP in the early embryo, RGA-3/4 (Schmutz et al., 2007; Schonegg et al., 2007). As previously shown, depletion of RGA-3/4 causes cortical hypercontractility in otherwise wild-type embryos, during both pseudocleavage and cytokinesis, and results in embryonic lethality (Figure 5A) (Schmutz et al., 2007; Schonegg et al., 2007). When RGA-3/4 is depleted from cyk-4R459A embryos, all embryos complete cytokinesis (Figure 5B, Figure 5—figure supplement 1A), further suggesting that the GAP activity of CYK-4 promotes, rather than counteracts, RhoA activation.
To test this model more stringently, we asked whether cytokinesis also completes when RGA-3/4 is depleted from cyk-4R459A embryos also lacking NOP-1 (i.e., nop-1(it142); rga-3/4(RNAi); cyk-4R459A embryos). Remarkably, although furrows in cyk-4R459A; nop-1 embryos barely ingress, when RGA-3/4 is depleted, furrow ingression is completed in 100% of embryos (Figure 5B,C, Figure 5—figure supplement 1B). This result also rules out the possibility that RGA-3/4 depletion allows completion of cytokinesis because it stabilizes RhoA that was activated in a NOP-1-dependent manner. Depletion of RGA-3/4 did not significantly modify the cytokinetic phenotype of nop-1; cyk-4(RNAi) embryos (Figure 5C, Figure 5—figure supplement 1B), demonstrating CYK-4 dependence to this suppression. Furthermore, complete suppression was specific to cyk-4R459A embryos, depletion of RGA-3/4 induced deeper but still incomplete ingression in nop-1; cyk-4EE and nop-1; cyk-4∆C1 embryos. These strains formed an allelic series in order of decreasing extents of ingression: cyk-4R459A > cyk-4EE > cyk-4∆C1 ∼ cyk-4(RNAi) (Figure 5C, Figure 5—figure supplement 1B).
RhoA is a dose dependent regulator of cleavage furrow formation (Loria et al., 2012) and CYK-4 is involved in RhoA activation by relieving autoinhibition of ECT-2 (Kim et al., 2005; Yüce et al., 2005). We therefore assayed whether CYK-4 GAP domain mutations affect accumulation of RhoA effectors. Because the RhoA biosensor and the CYK-4 transgenes are both GFP-tagged and integrated at the same position of the genome, we assayed the accumulation of RFP-tagged non-muscle myosin, NMY-2, a key effector of RhoA, as a proxy for RhoA activation. To validate that NMY-2::mRFP is a valid proxy for RhoA activity levels, we compared the accumulation of these two markers to the cleavage furrow during cytokinesis when co-expressed. The recruitment of NMY-2::mRFP and the RhoA biosensor are highly correlated in space, time, and intensity (Figure 5—figure supplement 2A,B). In addition, the correlation between these markers remains strong when either NOP-1, CYK-4, or RGA-3/4 are depleted, despite the significant changes in the extent of recruitment caused by these perturbations. Thus NMY-2::mRFP provides a reliable proxy for RhoA activation.
We assayed NMY-2::mRFP levels in CYK-4WT, CYK-4∆C1, CYK-4R459A, and CYK-4EE embryos during anaphase. Mutations in CYK-4 that reduce the rate and extent of cleavage furrow also reduce NMY-2::mRFP accumulation (Figure 5D, top row, Figure 5E). The defect in myosin accumulation caused by mutations in the GAP domain of CYK-4 is far more severe and apparent in NOP-1-depleted embryos (Figure 5D, bottom row, Figure 5E). Conversely, depletion of RGA-3/4 increases myosin accumulation in CYK-4R459A and CYK-4EE embryos, both in the presence and absence of NOP-1. These data support models in which the catalytic activity of the CYK-4 GAP domain contributes to RhoA activation.
To obtain additional insight into the mechanism by which CYK-4 promotes cytokinesis, we took an unbiased genetic suppression approach. We mutagenized cyk-4(or749ts) animals, grew the mutagenized animals at the permissive temperature for two generations to allow potential suppressors to become homozygous and shifted them to 25°C to select for suppressors. We isolated three strong suppressors out of a total of ∼10^5 mutagenized F1 genomes. Suppressor strains were subjected to sequencing of the cyk-4 locus to identify potential intragenic suppressors. One strain contained a substitution mutation in CYK-4, H485Y, relatively close to the or749ts substitution E448K (Figure 6A). We also isolated two strong extragenic suppressors, xs110 and xs111, that rescue cyk-4(or749ts) to viability at the restrictive temperature. The suppressed strains complete cytokinesis with high efficiency (>90%) and support high viability (Figure 6—figure supplement 1).
Candidate extragenic suppressors were genetically mapped to chromosome II near the ect-2 locus. Substitution mutations in the ect-2 locus were identified in both suppressor strains (Figure 6A). ect-2(xs110) contained a single nucleotide change in the PH domain, resulting in a G707D substitution (Figure 6A, Figure 6—figure supplement 2A). In a related RhoGEF for which there is a co-crystal structure with RhoA (PDZRhoGEF), the residue analogous to G707 lies in an α helix in the PH domain that comes into close proximity to the α3 helix of RhoA (Figure 6—figure supplement 2B) (Chen et al., 2010).
We recorded the progression of cytokinesis in cyk-4(or749ts); ect-2(xs110) embryos and found that the embryos not only complete cytokinesis as expected but also the delay in furrow initiation and the slow furrow ingression phenotypes characteristic of cyk-4(or749ts) embryos were largely corrected (Figure 6B,Cii). Thus, unlike ced-10(n1993), ect-2(xs110) suppresses the primary defect of the cyk-4(or749ts) mutation.
To confirm that the ect-2(xs110) substitution was causative, we used the CRISPR-associated nuclease Cas9 to re-create this mutation (Zhang and Glotzer, 2014). We injected cyk-4(or749ts) animals with a plasmid that expresses both Cas9 and a sgRNA designed to create a double strand break near E705 and provided an oligonucleotide repair template containing the G707D substitution. The animals were maintained at the permissive temperature for two generations before shifting to the restrictive temperature. We were able to isolate a strain that was viable and fertile. The ect-2 locus was sequenced and de novo generation of the G707D substitution was confirmed. This mutation in ect-2 therefore suppresses all the essential functions affected by the cyk-4(or749ts) allele.
We next investigated whether this mutation causes a detectable phenotype when separated from cyk-4(or749ts). Interestingly, ect-2(xs110) embryos exhibit hypercontractility during both pseudocleavage and cytokinesis (Figure 6Bi); this hypercontractility is associated with enhanced cortical accumulation of myosin II (Figure 6—figure supplement 3). Hypercontractility is also observed in embryos from ect-2(xs110)/+ hermaphrodites, indicating ect-2(xs110) is a dominant, gain of function allele (Figure 6—figure supplement 4). The hypercontractility is reduced in ect-2(xs110); cyk-4(or749ts) embryos (ii), indicating that ECT-2G707D hyperactivity is partially dependent on CYK-4 and that cyk-4(or749ts) and ect-2(xs110) exhibit mutual suppression. Comparison of ect-2(xs110); cyk-4(or749ts) embryos to ect-2(xs110); cyk-4(or749ts); nop-1(RNAi) (Figure 6ii vs Figure 6iii) embryos reveals that NOP-1 also contributes to contractility in ECT-2G707D embryos.
An unusual phenotype was observed in ect-2(xs110) embryos. Following anaphase, the cleavage furrow frequently initiates from a site significantly anterior to the midpoint of the anaphase spindle (Figure 6D–F). As the furrow ingresses, it undergoes a dramatic repositioning so that it ultimately bisects the anaphase spindle. Nevertheless, the ect-2(xs110) strain is viable and fertile despite exhibiting hypercontractility during polarization and cytokinesis (Figure 6—figure supplement 1).
The second suppressor allele, ect-2(xs111), also contains a substitution mutation in ECT-2. This mutation is located in the linker region between the cryptic BRCT0 domain and BRCT1 (Zou et al., 2014) (Figure 6A). Several criteria indicate that this mutation is also causal. First, SNP mapping placed suppressor activity near the ect-2 locus. Second, the suppressor was analyzed by one step mapping and whole genome sequencing (Doitsidou et al., 2010). ect-2 is the only gene in the candidate region that contained a non-silent mutation that has any role in cytokinesis. Third, biochemical data indicate that this mutation relieves ECT-2 autoinhibition (see below). The ect-2(xs111) gain of function allele exhibited similar overall characteristics as ect-2(xs110) (Figure 6B,C), although the spindle positioning defect was less severe (not shown). The one remarkable difference was that ect-2(xs111); cyk-4(or749ts); nop-1(RNAi) (Figure 6Bviii) embryos fully ingressed during cytokinesis, though they do not form pseudocleavage furrows. The ability of these embryos to complete cytokinesis depends upon residual activity from CYK-4E448K, as depletion of CYK-4 by RNAi prevents completion of cytokinesis in ect-2(xs111) embryos (Figure 6Bx). As complete furrow ingression is not seen in comparable ect-2(xs110) embryos (Figure 6Biii), ect-2(xs111) may be more strongly activated than ect-2(xs110).
This genetic screen demonstrates that only rare mutations suppress cyk-4(or749ts) and that the essential function of CYK-4 that is inactivated by CYK-4E448K is the ability to activate RhoA. Note that while ced-10(n1993) can partially suppress cytokinesis defects in CYK-4E448K expressing embryos (cytokinesis remains delayed and slow in the double mutant; and only ∼67% of embryos complete division), ced-10(n1993) does not rescue cyk-4(or749ts) to viability (Figure 6—figure supplement 1).
CYK-4R459A causes a less severe phenotype than CYK-4E448K, therefore we predicted that ect-2(xs110) and ect-2(xs111) could also suppress CYK-4R459A. We used CRISPR/Cas9 to introduce the R459A mutation into the endogenous cyk-4 gene and crossed it into both ect-2 hyperactive mutants. We were able to isolate strains in which the sole source of CYK-4 lacks the critical arginine in the active site (Figure 6Biv,ix). The resulting strains exhibited high viability and fertility (Figure 6—figure supplement 1). This finding provides independent confirmation that the sole essential function of the RhoGAP active site of CYK-4 is to stimulate ECT-2-mediated RhoA activation.
These genetic and cell biological results demonstrate that the GAP activity of CYK-4 contributes to RhoA activation. As ECT-2 is required for all RhoA activity during cytokinesis, the CYK-4 GAP domain is likely to serve this role by modulating ECT-2. Given that the canonical function of a RhoGAP domain is to inhibit RhoA activity, it is surprising that a protein containing a RhoGAP domain enhances RhoA activation. However, CYK-4 and ECT-2 form a protein complex through their regulatory N-termini (Burkard et al., 2007; Wolfe et al., 2009), therefore the C-terminal GAP domain of CYK-4 will be in the vicinity of the ECT-2 RhoGEF domain.
We therefore hypothesized that the interactions between CYK-4 and ECT-2 are not limited to their N-termini. To test this, we purified the C-terminal domains of CYK-4 and ECT-2 (Figure 7A) and performed binding assays. We found that the catalytic C-termini of CYK-4 and ECT-2 directly interact (Figure 7B, Figure 7—figure supplement 1); a similar complex is also found with human orthologs (data not shown). We assayed for activation of the ECT-2 GEF activity by the CYK-4 GAP domain in vitro. However, we have not yet been able to detect stimulation of GEF activity (data not shown). This negative result could be due to missing components, a requirement for the context provided by the full length, oligomerization competent proteins (Basant et al., 2015), or the absence of the plasma membrane to which CYK-4 must bind in vivo in order to activate ECT-2.
As ect-2(xs110) and ect-2(xs111) suppress the phenotypes caused by mutations in the CYK-4 GAP domain, we sought to understand the biochemical basis of activation by the proteins they encode, ECT-2G707D and ECT-2E129K, respectively. Given that the E129K mutation lies near the N-terminal BRCT domain (Figure 7A), we hypothesized that it could interfere with ECT-2 autoinhibition. To test this possibility, we assayed for binding between the N- and C- termini of ECT-2. Wild-type N- and C- termini form a complex that is readily detected in vitro. However, the E129K substitution, but not G707D, significantly reduces binding of the ECT-2 N- and C-termini (Figure 7C, Figure 7—figure supplement 1), suggesting that this allele functions by relieving autoinhibition.
The G707D mutation in ECT-2 is located in the PH portion of the RhoGEF domain (Figure 7A, Figure 6—figure supplement 2). In principle, this mutation could promote RhoA activation by a number of mechanisms including activation of the GEF domain, relieving autoinhibition of ECT-2, stabilizing the interaction with CYK-4 and/or stabilizing the interaction of ECT-2 with the plasma membrane. We did not observe a change in the association of ECT-2_N with ECT-2_CG707D, suggesting that the mutation doesn't relieve autoinhibition. However, as the mutated residue maps to a helix that lies near RhoA in a co–crystal structure of a related RhoGEF (Figure 6—figure supplement 2), we tested whether it activates RhoGEF activity. We assayed ECT-2 GEF activity in an in vitro exchange assay. The ECT-2G707D variant exhibits a modest increase in GEF activity compared to wild-type ECT-2 over a range of concentrations (Figure 7D), perhaps by increasing the affinity of ECT-2 for RhoA.
Diverse mechanisms ensure that the cytokinetic contractile ring assembles at the cell equator following chromosome segregation. These regulatory mechanisms converge to promote local accumulation of active RhoA at the cell equator which is an essential prerequisite for contractile ring assembly. Whereas it is widely accepted that the RhoGEF ECT-2 is the primary activator of RhoA and the centralspindlin component CYK-4 contributes to RhoA activation, the mechanism(s) by which CYK-4 promotes RhoA activation have been rather unclear. Here, we demonstrate that CYK-4 has multiple functional domains that are required for it to promote RhoA activation. In addition to the previously characterized binding interaction with ECT-2, we show that both the C1 domain and the catalytic activity of the RhoGAP domain of CYK-4 are also required for full activation of ECT-2. Furthermore, our results indicate in order for the CYK-4 GAP domain to promote RhoA activation, it has to act catalytically on RhoA•GTP. This implies that RhoA plays a role in promoting its own activation.
Four results suggest that CYK-4 GAP activity promotes RhoA activation. First, CYK-4 GAP activity is required for the completion of cytokinesis and embryos lacking this activity exhibit reduced levels of RhoA effectors. Second, when the NOP-1-dependent, parallel pathway for RhoA activation is eliminated, the requirement for CYK-4 GAP activity for furrow formation and effector recruitment is greatly enhanced. Third, we have demonstrated a biochemical interaction between the GAP domain of CYK-4 and the GEF domain of ECT-2. Fourth, the requirement for CYK-4 GAP activity can be alleviated by three independent perturbations that each increase RhoA activity levels.
One of the perturbations that suppresses the CYK-4 GAP-deficient phenotype is depletion of the primary RhoA GAP, RGA-3/4. Embryos defective in RGA-3/4 alone exhibit hypercontractility and are largely inviable; these phenotypes are consistent with the canonical function of a RhoA GAP. However, loss of CYK-4 GAP activity and loss of RGA-3/4 counterbalance each other during cytokinesis. Furthermore, suppression by RGA-3/4 depletion is potent, it can restore cytokinesis in embryos deficient in both NOP-1 and CYK-4 GAP activity in which furrows otherwise barely ingress.
A large, unbiased, genome-wide screen for suppressors of cyk-4(or749ts) corroborates the model that CYK-4 GAP activity promotes RhoA activation. We identified two strong, extragenic, gain of function suppressor mutations in the RhoGEF ECT-2 (Figure 6). Because these suppressors rescue cyk-4(or749ts) and cyk-4R459A to viability, the essential function of the CYK-4 GAP activity must be to promote RhoA activation.
These results raise the fundamental question: by what mechanism does the GAP activity of CYK-4 contribute to RhoA activation? We propose a working model in which the most active form of the ECT-2 RhoGEF is a complex containing ECT-2 and CYK-4 with a molecule of RhoA•GDP bound to the GAP active site (Figure 7E).
The GTPase bound to CYK-4 is likely to be RhoA, rather than CED-10/Rac1 or CDC-42. If CYK-4 had to bind CED-10/Rac1 or CDC-42, then depletion of those GTPases should impair cytokinesis as severely as a mutation that attenuates GTPase binding by the CYK-4 GAP domain. However, neither CED-10/Rac1 nor CDC-42 is required for cytokinesis, even in NOP-1-defective embryos (Figure 3), whereas weakening GTPase binding by the CYK-4 GAP domain strongly impacts cytokinesis. The model has a further implication: to form the most active ECT-2 GEF complex, CYK-4 GAP binds RhoA•GTP. Therefore, RhoA•GTP plays a role in RhoGEF activation, suggesting the presence of a positive feedback loop during cytokinesis.
This working model is supported by the finding that full activation of RhoA and cytokinesis requires that the CYK-4 GAP domain both bind a Rho family GTPase (Figure 3) and activate its ability to hydrolyze GTP (Figures 2, 4, 5); indeed mutations in the CYK-4 GAP domain that diminish GTPase binding exhibit a stronger defect in RhoA activation than mutation of the GAP active site (Figure 5). Our mutational analysis has trapped ECT-2 in four distinct states that form an allelic series (Figure 7—figure supplement 2A). We propose that the least active form of ECT-2 is not bound to CYK-4 and has little GEF activity. Once CYK-4 is phosphorylated, it can be bound by the ECT-2 N-terminal BRCT domains rendering it weakly activated (equivalent to CYK-4EE). This form may also exhibit some interactions between the GEF domain of ECT-2 and the GAP domain of CYK-4, as these domains can interact in vitro without RhoA present. If CYK-4 can bind to RhoA•GTP, it induces a higher activity state, as evidenced by the increased activity of CYK-4R459A, which is sufficient for zygotic development. Finally, if CYK-4 can induce GTP hydrolysis by RhoA, this results in the fully active ECT-2/CYK-4/RhoA•GDP or ECT-2/CYK-4/RhoA•GDP + Pi state populated by the wild-type protein. We speculate that this complex results in full relief from autoinhibition within the ECT-2 GEF domain.
Not only does this model explain why CYK-4 retains GAP activity towards RhoA, it also explains why its ability to inactivate RhoA is attenuated relative to Rac and Cdc42. High turnover rates of RhoA•GTP induced by CYK-4 might rapidly consume RhoA•GTP at the site of production, yielding a futile cycle of RhoA activation and inactivation. However, this working model must be tested by structural studies and biochemical reconstitution assays that reflect the in vivo situation. Accurate reconstitutions will need to account for the facts that cytokinetic RhoA activation involves the CYK-4 C1 domain (Figure 1C), the ability of CYK-4 to bind to ZEN-4, and the ability of ZEN-4 to oligomerize (Basant et al., 2015).
We tested the model that GAP activity of CYK-4 is important to maintain CED-10/Rac1 in an inactive state. Some of our results do support this model, as the failure to complete the first cytokinesis in embryos lacking CYK-4 GAP activity can be partially restored by a loss of function mutation in ced-10/Rac1 (Figure 4B). We therefore tested whether inactivation of CED-10/Rac1 suppresses loss of CYK-4 GAP activity in NOP-1-deficient embryos. GAP-deficient CYK-4 does not promote significant ingression of the cleavage furrow in embryos lacking NOP-1, irrespective of the presence or absence of CED-10/Rac1 (Figure 4Biii,v). Finally, mutations in CED-10/Rac1 do not suppress the lethality of a temperature sensitive mutation in cyk-4 (Figure 6—figure supplement 1).
Thus, because the active site of the CYK-4 GAP domain is required in the absence of CED-10/Rac1, CED-10/Rac1 inactivation cannot be the primary function of the CYK-4 GAP domain. Rather, these results suggest a model in which loss of CED-10/Rac1 function causes a reduction in overall cortical tension which, in turn, allows an increase in the extent of NOP-1-dependent furrow ingression (Loria et al., 2012).
The experiments presented here demonstrate that CYK-4 GAP activity promotes RhoA activation and that this function is essential in early C. elegans embryos and in the adult germline (Figures 2A, 4B). Our experiments also addressed the function of CYK-4 GAP activity post-embryonically. We find that whereas zygotic cyk-4 null embryos die during embryogenesis, expression of catalytically inactive CYK-4 provides significant rescue, supporting development into viable, albeit sterile, adults (Figure 2A). Thus, while early embryos require the GAP activity of CYK-4, this requirement is relaxed post-embryonically. The requirement for CYK-4 GAP activity can be experimentally eliminated by hyperactivation of ECT-2 or depletion of the RhoA GAP RGA-3/4 (Figures 5B, 6B). Interestingly, RGA-3/4 is primarily expressed in the germline and in early embryos (NextDB, cited in Schmutz et al., 2007 and data not shown), thus regulated RGA-3/4 expression could contribute to the tissue specific requirements.
These findings allow us to reconcile many previous results on the role of the CYK-4 GAP domain during cytokinesis. Some studies provided evidence that the GAP activity is dispensable (Goldstein et al., 2005; Yamada et al., 2006), whereas others suggested it is required for Rac1 inactivation (D'Avino et al., 2004; Canman et al., 2008; Bastos et al., 2012), RhoA inactivation (Miller and Bement, 2009), or RhoA activation (D'Avino et al., 2004; Zavortink et al., 2005; Loria et al., 2012). The first set of results is consistent with the results presented here, as some cell types may not require the GAP activity of CYK-4 for cytokinesis, as seen in post embryonic cells in C. elegans. As numerous studies have shown that active RhoA can indirectly inhibit Rac1 (see Guilluy et al., 2011 for review), some of the results that point to a role for CYK-4 GAP activity in attenuating Rac1 levels (Bastos et al., 2012) may be indirectly caused by a reduction RhoA activation or by indirectly controlling cortical tension. Thus, many previous results can be explained without proposing that the CYK-4 GAP domain performs different functions in different organisms or cell types. We do not rule out the possibility that, in certain contexts, CYK-4 or its orthologs negatively regulate Rac or Cdc42. Recent evidence indicates that the Xenopus ortholog of CYK-4 concentrates at cell–cell junctions and negatively regulates GTPases at that site (Breznau et al., 2015). Further work is required to resolve why CYK-4 acts as a positive regulator of RhoA in C. elegans embryos and a negative regulator in Xenopus embryos (Miller and Bement, 2009; Breznau et al., 2015).
The signaling mechanisms we have discovered in cytokinesis have analogies in other signaling pathways. Our favored model, in which RhoA promotes its own activation, is reminiscent of the positive feedback in Cdc42 activation during yeast budding (Howell and Lew, 2012) and the activation of the SOS1 RasGEF domain by a molecule of Ras•GTP that serves as an allosteric activator (Gureasko et al., 2008). Interestingly, ECT-2 has also been implicated in SOS regulation (Canevascini et al., 2005). Likewise, CYK-4 is not the only protein with a GTPase activating domain that plays a role in promoting GEF activity. A similar function, in cis, has been seen in p115 RhoGEF, which is activated by Gα13 (Chen et al., 2012). In this case, the binding of a molecule of Gα13 to an allosteric site on the RhoGEF domain of p115 is stabilized by p115's N-terminal RGS domain (Figure 7—figure supplement 2B). RGS domains accelerate GTP hydrolysis by Gα, that is, they are Gα GAPs (Tesmer et al., 1997). As RhoGEF activation is not an obvious function for a RhoGAP domain, additional cases may have gone undetected.
RhoA activation is controlled by multiple layers of regulation during cytokinesis. In addition to cell-cycle regulated changes in the phosphorylation state of CYK-4 and ECT-2 that control their binding and localization (Yüce et al., 2005; Su et al., 2011; Zou et al., 2014), full activation of RhoA also involves membrane binding by CYK-4 (Figure 1) which, in turn, requires centralspindlin oligomerization (Basant et al., 2015). Like the requirement for CYK-4 GAP activity, the requirement for the C1 domain of CYK-4 is context dependent. Whereas the C1 domain makes a major contribution to furrow ingression in C. elegans embryos, studies in Hela cells demonstrate that the C1 domain contributes to RhoA activation, but it is not essential (Lekomtsev et al., 2012). Thus, centralspindlin has several domains that contribute to maximal activation of ECT-2. However, not all cell types may require maximal activation of ECT-2 either because of physical properties of the cell (cell size, cortical tension, and tissue organization) or because of their biochemical properties (expression of RGA-3/4 orthologs). Nevertheless, the evolutionary conservation of all of these functions suggests that they play critical roles during some stage(s) of metazoan development.
Animals were grown at 20°C on standard nematode growth media (NGM) plates seeded with OP50 Escherichia coli. Some strains were provided by the Caenorhabditis Genetics Center. All strains used in this study are listed in Supplementary file 1.
RNAi was administered by feeding nematodes with E. coli expressing the appropriate double-stranded RNA (dsRNA) (Timmons and Fire, 1998). HT115 bacterial cultures were grown in Luria broth with 100 μg/ml ampicillin overnight at 37°C. Cultures (250 μl) were seeded on NGM plates containing 100 μg/ml ampicillin and 1 mM IPTG and incubated at room temperature for 16 hr. RNAi plasmids were obtained from the library produced by Kamath et al. (2003). Young L4 hermaphrodites were picked onto the plates for feeding at 25°C at least 24 hr prior to dissection. For RNAi depletion of temperature-sensitive alleles, L4 larvae were fed for 48 hr at 16°C, then shifted to 25°C for at least 1 hr before imaging.
For experiments where two genes were simultaneously knocked down by RNAi, bacterial cultures of E. coli expressing the appropriate dsRNA were mixed in a 1:1 ratio seeded onto NGM plates as described above. If stronger depletion of one of the two genes was desired, embryos were first hatched onto feeding plates targeting the gene. L4 worms were transferred to fresh plates with bacteria expressing dsRNA against both genes.
Young gravid hermaphrodites were transferred to fresh seeded NGM plates in triplicate. Remove worms from plates after ∼8 hr of egg laying. The eggs laid on plates were scored manually under dissecting microscope. To determine unhatched embryos, embryos remaining on plates were scored 1 day after the parents were removed. The embryonic lethality percentage is calculated as the number of unhatched embryos divided by the total egg production.
To generate CYK-4::GFP MosSCI constructs, ∼2 kb sequences upstream of cyk-4, cyk-4 genomic DNA tagged with C-terminal GFP coding sequences, and pie-1 3′ UTR sequences were generated by overlapping PCR and inserted to pCFJ150 by SLiCE (Zhang et al., 2012). cyk-4 genomic sequences between BamHI and AvrII were recoded to generated RNAi resistant alleles. To introduce cyk-4 mutations, sequences covering mutations were generated by overlapping PCR using pCJF150-cyk-4-gfp as template and the appropriate primers (see primer sequences in Supplementary file 2): MG4199/MG4276 and MG4200/MG4277 for E448K; MG4199/MG4202 and MG4200/MG4201 for R459A; MG4199/MG4489 and MG4200/MG4488 for K459E/R499E(EE); MG4199/MG4070 and MG4200/MG4071 for ∆C1. Overlapping PCR products were inserted into pCFJ15-cyk-4-gfp linearized with NaeI by SLiCE. All constructs were sequence verified.
Cas9/sgRNA plasmids were derived from pDD162 vector (Dickinson et al., 2013). ect-2 sgRNA target sequences were generated by overlapping PCR using pDD162 as PCR template and the appropriate primers (see primer sequences in Supplementary file 2), MG4735/MG4773 and MG4774/MG4736. Overlapping PCR products were inserted into pDD162 linearized with SpeI/BsrBI by SLiCE.
Transgenic lines expressing single copy CYK-4::GFP or mutant CYK-4::GFP were generated by integrating constructs into the Mos1 element ttTi5605 on chromosome II using the MosSCI method (Frøkjaer-Jensen et al., 2008).
For oligonucleotide templates (ODNs) based CRISPR experiments (Zhang and Glotzer, 2014; Zhao et al., 2014), microinjection was performed by injecting DNA mixture into gonad arms of cyk-4(or749ts) young gravid hermaphrodites. Injected cyk-4(or749ts) were maintained at 16°C for 3–4 days then shifted to 25°C until starvation. Viable worms were isolated and subjected to single worm PCR to identify desired mutations. The injection mixture consists of Cas9/sgRNA plasmids and ODNs. The final concentrations of plasmids and ODNs are Cas9/ect-2 sgRNA vector at 50 ng/μl and ect-2 ODN (MG4801 5′-TTGTATGGTGCCTGATTCATCGTGACGAGCAAGATGGTGACATTGACACAGTCTTCGAAT-3′) at 50 ng/μl.
To prepare one-cell embryos for imaging, gravid hermaphrodites were dissected into egg salt buffer (HEPES pH 7.4 5 mM, NaCl 118 mM, KCl 40 mM, MgCl2 3.4 mM, CaCl2 3.4 mM) on coverslips, mounted onto 2.5% agar pads and sealed with vaseline. For Nomarski imaging, embryos were observed with a Zeiss (Thornwood, NY) Axioplan II with a 100×/1.3 Plan-Neofluar objective. Images were captured with a charge-coupled device (CCD) camera (Imaging Source, Charlotte, NC) controlled by Gawker (gawker.sourceforge.net). Images were acquired every 5 s and processed with ImageJ (http://rsbweb.nih.gov/ij). For confocal imaging, embryos were imaged with a 63×/1.4 oil-immersion lens on (1) a Zeiss Axiovert 200M equipped with a Yokogawa CSU-10 spinning-disk unit (McBain, Simi Valley, CA) and illuminated with 50-mW, 473-nm and 20-mW, 561-nm lasers (Cobolt, Solna, Sweden), or (2) a Zeiss Axioimager M1 equipped with a Yokogawa CSU-X1 spinning-disk unit (Solamere, Salt Lake City, UT) and illuminated with 50-mW, 488-nm and 50-mW, 561-nm lasers (Coherent, Santa Clara, CA). Images were captured on a Cascade 1K EM-CCD camera or a Cascade 512BT (Photometrics, Tucson, AZ) controlled by MetaMorph (Molecular Devices, Sunnyvale, CA). Image processing was performed with ImageJ. Time-lapse acquisitions were assembled into movies using Metamorph and ImageJ.
To measure furrow ingression kinetics, a single central plane image of GFP::PH or mCherry::PH was acquired at 10 s intervals starting at anaphase as assessed by the CYK-4::GFP or mCherry::HIS-58 signal. The position of the furrow was assessed in each frame by manual tracking of GFP::PH or mCherry::PH signal. The extent of ingression in each frame was calculated as d/w, where w is the total width of the embryo and d is the distance between the furrow tips. To determine whether furrow ingression kinetics were statistically significant different between multiple genotypes, data sets of normalized cortical distance from 100 s to 410 s after anaphase onset were analyzed with a Kruskal–Wallis non-parametric one-way analysis of variance (ANOVA) using Dunnett's multiple comparisons test.
To quantitate the abundance of NMY-2::mRFP at the equatorial region, a stack of five planes spanning 2.5 μm was captured every 10 s. The Z-stacks were projected using a maximum intensity projection algorithm and corrected for photobleaching. Using custom ImageJ macros, the background signal was measured in a remote region of each frame. A region of fixed size in the equatorial region was thresholded with a minimal value of 1.25× background and the total thresholded signal in the region was integrated and normalized to the background. The total value of intensity was summed for a defined number of planes after anaphase onset.
To quantitate the abundance of CYK-4::GFP at the furrow tip, a stack of five planes spanning 2.5 μm was captured every 10 s. The frame in which furrow ingressed to half of the egg width or the deepest was chosen. The Z-stacks were projected using a maximum intensity projection algorithm and corrected for photobleaching. The background was measured as integrated intensity of a square adjacent to the furrow tip. The GFP intensity was measured as integrated intensity of a square with the same area covering the furrow tip and normalized to the background by subtracting background intensity (see Figure 1).
The coding sequences for CYK-4 C1 and RhoGAP domain (342-681aa), ECT-2 DH/PH domain (356-792aa), CED-10/Rac1 and RhoA (C.e.) were cloned into the GST expression vector pGEX-4T-tobacco etch virus (TEV), and the coding sequences for ECT-2 BRCT domain (1-363aa) were cloned into MBP expression vector pMAL-c2-TEV. GST- and MBP-tagged proteins were expressed in E. coli strain BL21 by adding 0.3 mM IPTG at OD600 reached 0.5–0.7 at 25°C. Cells were grown for another 4 hr at 25°C and collected. Frozen cells were thawed in lysis buffer (50 mM Tris pH 7.5, 150 mM NaCl, 5 mM MgCl2, 10% Glycerol, 1 mM PMSF, 1 μg/ml leupeptin, 1 μg/ml pepstatin A, 0.1% Triton X-100, 1 mM DTT, 0.5 mg/ml lysozyme) and lysed by sonication. The bacterial lysate was centrifuged at 40,000×g at 4°C for 30 min.
For GST-tagged proteins, glutathione-Sepharose 4B beads (bioWORLD) were added to supernatant and incubated at 4°C for 4 hr. The beads were washed 3× with 50 mM Tris pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1 mM DTT. Protein-bound beads in were either stored in 50 mM Tris pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1 mM DTT, 50% glycerol at −20°C, or cleaved from beads by incubation with His-tagged TEV protease at 4°C overnight. TEV protease was removed by incubation with TALON beads (Clontech). Cleaved fusion proteins were stored in 10% glycerol at −80°C.
For MBP-tagged proteins, amylose resin (New England Biolabs) was added to supernatant and incubated at 4°C for 4 hr. Beads were placed in a poly-prep chromatography column (Bio-Rad), and washed with 12 column volumes of 50 mM Tris pH 7.5, 150 mM NaCl, 5 mM MgCl2, and 1 mM DTT. Fusion proteins were eluted with 50 mM Tris pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1 mM DTT, 10 mM maltose, and stored in 10% glycerol at −80°C.
C.e. GST-RhoA was loaded with GDP (Self and Hall, 1995). Beads were washed with low magnesium buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM DTT) and 1 mM GDP was added. Beads were incubated with shaking at room temperature for 15 min, placed on ice, and 20 mM MgCl2 was added and incubated on ice for 5 min. Beads were washed three times with 50 mM Tris pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1 mM DTT. GDP-loaded RhoA was cleaved from beads with TEV protease, glycerol added to 10%, flash frozen and stored at −80°C.
Fluorescence-based kinetic assays were performed in HORIBA FluoroLog-3 Spectrofluorometer, with fluorescence analog of GTP, mant-GTP (AnaSpec). All nucleotide exchange assays were performed in the presence of 1 μM RhoA-GDP, 200 nM mant-GTP, the indicated concentration of ECT-2 DH/PH domain in 20 mM Tris pH 7.5, 50 mM NaCl, 10 mM MgCl2, 1 mM DTT, 50 μg/ml BSA, 1% glycerol. The relative fluorescence was monitored for 90 s before adding mant-GTP, and for 510 s after adding mant-GTP; measurements were taken every 15 s. The reaction rate, v, is defined as ∆F/∆t, where F = fluorescence, t = time.
CYK-4 GAP (final concentration from 0 to 800 nM) and RhoA or Rac1 (final concentration 9 μM) were mixed in 1× reaction buffer (50 mM Tris pH 7.5, 50 mM NaCl, 5 mM MgCl2, 1 mM DTT, 1% glycerol), then GTP was added to 1 mM to start the reaction. After 30 min, inorganic phosphate was assayed using a malachite green-based assay (Kodama et al., 1986); absorbance was measured with a NanoDrop 2000 spectrophotometer (Thermo Scientific). For time course experiments, 200 nM CYK-4 GAP was added into the reaction and, at the indicated time points, aliquots of the reaction were removed to assess free phosphate.
For each binding experiment, purified fusion proteins were added to the protein-bound glutathione-sepharose beads and incubated for 1 hr at 4°C. After three washes in cold wash buffer (50 mM Tris pH 7.5, 150 mM NaCl, 5 mM MgCl2, 1 mM DTT), proteins were eluted into loading buffer, separated by SDS-PAGE, and detected by coomassie blue staining.
CYK4 inhibits Rac1-dependent PAK1 and ARHGEF7 effector pathways during cytokinesisThe Journal of Cell Biology 198:865–880.https://doi.org/10.1083/jcb.201204107
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Mohan BalasubramanianReviewing Editor; University of Warwick, United Kingdom
eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.
Thank you for submitting your work entitled “The RhoGAP activity of CYK-4/MgcRacGAP functions non-canonically by promoting RhoA activation during cytokinesis” for peer review at eLife. Your submission has been favorably evaluated by Randy Schekman (Senior Editor) and three reviewers, one of whom, Mohan Balasubramanian, is a member of our Board of Reviewing Editors.
The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this decision to help you prepare a revised submission.
As you see, the views were varied (but all supportive), and Reviewer 3 has raised a number of technical issues all of which seem reasonable and I would like you to consider and respond to these comments. In particular: 1) please provide in vivo data to address key points of Reviewer 3 (such as potential effect of remaining Rac activities; RAC-2 or MIG-2), and 2) please provide proper controls for in vitro experiments raised by Reviewer 3.
As for Reviewer 2, although the biochemical experiments are ultimately required to advance your model further, this may not be accomplished in a single paper and may not be possible within the timeframe of the revision. If you have made some progress since submission of your paper and if they address the mechanism of GAP activation of ECT2, you may add it.
Please also address the points of Reviewer 1, most of which seem to be aimed at improving the readability as well as placing the paper in the right context through further discussion. Please see below the comments of the three reviewers.
This is an interesting paper by Glotzer and colleagues in which they have investigated the function of CYK-4 GAP protein in C. elegans cytokinesis. Through structure-function analysis of CYK-4 GAP they have identified that the GAP activity of CYK-4 is essential for cytokinesis, and performs at least 2 different functions. They make the important discovery that the GAP activity is crucial for nucleotide exchange on RhoA, the GTPase that activates both actin polymerization and myosin II activation. The work is very thorough and I particularly like the experiment where, through an unbiased screen for suppressors of CYK-4 GAP mutants, they identify new alleles of the RhoA exchange factor ECT2 itself. They further go on to show that CYK-4 physically interacts with ECT-2 and that an Ect2 mutant they isolated has higher nucleotide exchange activity (albeit a modest one).
I think the paper is appropriate for publication in eLife. However, I would like to see the biochemistry developed better. In particular, I am surprised that none of the GAP mutants identified in the work have been characterized in the biochemical studies described in the latter part of the paper. The two key experiments are: 1) how do the GAP mutants behave in the physical interaction experiments with ECT2 and 2) how does the presence of wild-type or GAP mutants of CYK-4 affects the ECT2-mediated nucleotide exchange on RhoA. These experiments are crucial if they are to advance the model in Figure 7.
I think some more discussion is also warranted on how mutations in the PH domain potentially affect the GEF activity. Molecular dynamics may help, although I realize this can be a new foray altogether.
Zhang and Glotzer provide an extensive and compelling analysis of the controversial role of the C. elegans RhoGAP CYK-4 during cytokinesis in animal cells. Their data strongly support the conclusion that the primary role of CYK-4/RhoGAP is to activate RhoA through its RhoGEF ECT-2. This is a counter-intuitive finding, as normally one thinks of GAPs as negative regulators of GTPases, not activators. The authors provide a rationale for this surprising role, and their evidence certainly supports the conclusion and also can to a large extent explain alternative views (invoking either indirect effects on other GTPases, or indirect effects from depletion of other GTPases to explain results suggesting that either the GAP activity of this family of RhoGAPs is not necessary or that it acts on other GTPases, in particular Rac as reported by Canman et al., 2008). The authors provide an elegant and impressive analysis of this issue, using a powerful combination of mutational analysis, live cell imaging and biochemistry. This manuscript provides the most extensive and compelling analysis of how CYK-4 influences cytokinesis via the RhoGEF ECT-2 and the RhoA GTPase, and should go a long ways toward resolving this complex issue.
While the manuscript in my opinion clearly warrants publication in eLife, the authors first need to address the following concerns that, while perhaps relatively minor, nevertheless could possibly improve the clarity of their arguments.
1) In the Introduction, subsection “The role of the GAP activity of CYK-4 GAP has been controversial”, the authors point out that CYK-4 has GAP activity toward RhoA but much more toward Rac1 and CDC-42. The authors should directly address how these observations relate to their conclusions concerning their model for how CYK-4 functions as an activator of RhoA.
2) With respect to Figure 1, the authors should discuss why there is some furrow ingression in embryos expressing CYK-4 lacking the C1 domain, and lacking nop-1 and ced-10 (in contrast to CYK-4E448K).
3) In two places (subsection “The cyk-4(or749ts) allele, E448K, exhibits defects in membrane association” and “CYK-4 GAP domain mutations that prevent RhoA binding are highly defective in RhoA activation”), the authors conclude results paragraphs by stating that an observation is “important” without explaining why it is important. Doing so might help clarify their conclusions.
4) In the subsection “Mutations in the active site of CYK-4 can be suppressed by RGA-3/4 depletion”, the authors refer to hyper-contractility after RGA-3/4 depletion and imply (and to some extent document) that this phenotype is suppressed by CYK-4 mutants. The authors should more explicitly state if this is an accurate conclusion.
5) In the subsection “Mutations in the active site of CYK-4 can be suppressed by RGA-3/4 depletion”, the authors refer to an allelic series of CYK-4 mutants. It might be helpful to explicitly state why the different mutants exhibit this gradation in effect.
6) In the first paragraph of the Discussion, the authors state that their data indicate that CYK-4 has to act catalytically on RhoA•GTP. The authors need to explain how their data support this conclusion.
7) Is it possible that the suppressor mutations in ECT-2 (xs110 and xs111) affect other functions than GEF activity? It might be worth discussing this possibility.
8) In the subsection “Mechanism of ECT-2 activation and Positive Feedback”, the authors refer first to Figure 3, and then to Figures 2, 4, 5 as supporting two conclusions. It would help if the authors elaborate to explicitly state what data in these figures support these conclusions.
9) The authors should explicitly state what they think the GAP active site does to influence ECT-2 GEF activity.
10) In the subsection “Mechanism of ECT-2 activation and Positive Feedback”, the authors state that CYK-4 GAP binds RhoA•GTP, but the model in Figure 7E in fact shows it binding RhoA:GDP. This should be explained.
11) The model diagram (Figure 7E) did not really do much to help me understand how they authors think CYK-4 activates RhoA.
CYK-4 plays a crucial role in cytokinesis as a non-motor subunit of centralspindlin, which bundles microtubules to form the central spindle and midbody. However, the role of the RhoGAP domain of CYK-4 remains unclear or controversial as the authors nicely describe in the Introduction. The cell biological and genetical data that the Glotzer lab has been providing in a recent series of papers are indeed very difficult to explain with Rac or Cdc42 as sole targets of CYK-4 but are consistent with the positive role of CYK-4 on RhoA activation. A major difficulty of this theory is that a GAP usually promotes conversion of a GTPase from GTP-form to GDP-form, which is in general an inactive form, by stimulating the GTP hydrolysis. A clear molecular mechanism for CYK-4 to activate RhoA has been a huge missing link in the file of cytokinesis.
In this manuscript, the authors examined various mutations in the GAP domain of CYK-4 including the E448K mutation that had previously been identified as a temperature-sensitive lethal allele (or749ts) and can be suppressed by loss-of-function of the CED-10/Rac or its effectors. By combination with the loss-of-function of NOP-1, a mysterious activator of Ect2 RhoGEF specific for nematodes, and RGA-3/4, a well conserved GAP that specifically inactivates Rho, the authors have strengthen their previous conclusion “that the RhoGAP domain of CYK-4 has an essential role in RhoA activation” (this is the title of Loria, 2012) except for a concern about the presence of other Rac proteins in addition to CED-10 (see below), which was not considered. Very interestingly, a screen for suppressors of cyk-4(or749ts) identified two novel alleles of Ect2 RhoGEF that cause its hyper activation, consistently with the proposed role of CYK-4 in RhoA activation. Finally, the authors tried to reveal biochemical mechanisms of RhoA activation by CYK-4 and found the direct binding between the GEF domain of ECT-2 and the C-terminal fragment of CYK-4 containing C1 and RhoGAP domains.
These are all interesting findings. However, it is questionable whether these are sufficient to support the authors’ rather strong claim that the non-canonical activity of CYK-4 RhoGAP domain promotes RhoA activation though its interaction with the GEF domain of ECT-2. The novel interaction between CYK-4 GAP and ECT-2 GEF domains relies solely on a single binding experiment in Figure 7B (7C and 7D are just unconvincing attempts to explain the hyper activation of ECT-2 by the new ect-2 alleles suggested by the in vivo phenotype). I really appreciate the amount of work necessary for the cell biology and genetics experiments. However, these are just strengthening their own already published claim. To propose a molecular mechanism of RhoA activation by the CYK-4 RhoGAP domain, stronger in vitro data should be provided at lease those obvious and feasible ones such as the influences of all the GAP mutations studied in vivo on the CYK-4-ECT-2 interaction.
Another concern is that the authors' logics are sometimes inconsistent. For example, as the explanations for the similar weaker cortical accumulation of the E448K and EE mutants, which show very similar in vivo phenotypes (slow ingression and regression at ∼50% ingression around 300s, weaker cortical accumulation, no furrowing upon nop-1 loss-of-function), they employed different reasonings, indirect effect on the function of C1 domain and the weakened interaction with RhoA, respectively. I agree with the authors on that mutations inside a protein fold might make the folding less stable more frequently than surface mutations and understand that a concern about this possibility prompted the current studies. However, this is an issue to be answered by proper biochemical/biophysical assays such as circular dichroism spectroscopy, which was undertaken in a recent paper from the Glotzer lab (White et al., 2013 http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3707682/). Otherwise, based on the phenotypic similarity, it would be more reasonable to explain the phenotype of the E448K mutation is also caused by the reduced affinity to Rho (or other GTPases). Anyway, there are different possible mechanisms for the cortical recruitment of CYK-4 that are not necessarily exclusive to each other; C1-phosphoinositide interaction, GAP-Rho interaction, interaction with astral microtubule gathered by the ingressing furrow. This should be explained more explicitly rather than picking up a convenient one for individual cases.
1) A table of the C. elegans strains used in this study with strain names and genotypes should be provided.
2) The following statements are ambiguous: “The RhoGAP activity of CYK-4/MgcRacGAP functions non-canonically” (title), “serve as a RhoGAP” and “consistent with CYK-4 RhoGAP activity” (Abstract). It should be clarified what the authors mean by “RhoGAP”. ‘RhoGAP activity’ could be any kind of activity that the RhoGAP domain has, the GAP activity against any Rho-family GTPase or a GAP activity specifically for Rho (but not for other GTPases such as Rac or Cdc42). It might be helpful to use “RhoGAP” for a domain name and “Rho-GAP”, “Rac-GAP”, “Rho-family-GAP” etc. for referring.
3) In the subsection “RhoA activation during cytokinesis”, the authors state: “The direct activator of RhoA during cytokinesis is the RhoGEF ECT-2”. There are other RhoA activators during cytokinesis such as GEF-H1 (Birkenfeld J 2007 Dev. Cell http://www.ncbi.nlm.nih.gov/pubmed/17488622). “A major activator of RhoA…” would be reasonable.
4) In the subsection “RhoA activation during cytokinesis”, you claim that “recruitment of ECT-2 to the spindle midzone involves regulated binding between ECT-2 and CYK-4…”. So far, the regulation of the interaction between CYK-4 and ECT-2 by phosphorylation by PLK-1 has only been shown in mammalian cells although there is no strong reason to imagine that the same regulation might not work in C. elegans.
5) Please clarify the following reference: “…it appears to be required to activate RhoA (D'Avino 2004)”. Although D'Avino et al. claimed “these observations are consistent with RacGAP50C inhibiting Rac and promoting Rho activity” and put RacGAP50C upstream to Rho in their model, this does not necessarily mean RacGAP50C is “required” for RhoA activation.
6) “These results suggest that this mutation in CYK-4 affects more than the RhoGAP activity of CYK-4 or that CED-10/Rac1 is not the relevant target of the GAP domain, or both” (Introduction). This is neglecting another obvious possibility that there might be another Rac that works redundantly with CED-10. C. elegans has two additional Rac-related genes, RAC-2 and MIG-2.
7) “…Subsequent analysis demonstrated that cyk-4(or749ts) does cause a phenotype in ced-10-embryos (Loria et al., 2012)” (Introduction). This might be due to elevated RAC-2 and MIG-2 activities by the cyk-4 mutation, which still remained after the depletion of CED-10. Imperfect restoration of the wild-type behavior does not discredit the importance of the suppression of Rac and downstream effectors.
8) Figures 1A, 1B, 3A, 3B: Why is there a big difference between the central spindle signal of CYK-4∆C1::GFP and CYK-4E448K::GFP? Five slices with 2.5 µm intervals should cover a range of 10 µm, which covers more than two thirds of the thickness of the 50%-ingressed cleavage furrow. For quantification, it is not clear how the usage of mCherry::PH as a standard is justified. The PH domain labels the plasma membrane by binding to phosphoinositides, which might be regulated by Rho-fmaily GTPases (http://www.ncbi.nlm.nih.gov/pubmed/24914539, http://www.ncbi.nlm.nih.gov/pmc/articles/PMC4114633/). Indeed, mCherry::PH signal at the 50%-ingresssed furrow is stronger in cyk-4R459A::gap embryo (Figure 4B ii) than in cyk-4WT embryos (Figure 1C the first row, Figure 4B i). The intensities of mCherry::PH used for standardization must be presented.
9) Another issue related to these experiments is that how CYK-4 accumulates at the furrow cortex is not clear. Although the authors mention it as accumulation on the plasma membrane, it might be reflecting the interactions between the C1 domain and phosphoinositides or between RhoGAP-Rho, or the compaction of the equatorial asters by the ingressing furrow.
10) How about the temperature sensitivity of EE and R459A?
11) In the subsection “The cyk-4(or749ts) allele, E448K, exhibits defects in membrane association”, the authors state: “the onset of furrow ingression is delayed relative to controls in cyk-4E448K embryos, but not in cyk-4∆C1 embryos”. In Figure 1C, the deformation of the plasma membrane in the E448K embryo starts at the similar timing to the WT one (∼ at the 9th time point) while it becomes debatable in the ∆C1 embryo in the 11th or 12th time point. From the graph in Figure 1F, no clear difference is detected in the timing of the furrow initiation while the ingression is clearly slower in the E448K embryos.
12) Please clarify the following passage: “…such that it facilitates the abscission step in cyk-4(or749ts); ced-10(n1933) embryos”. It is not clear why the abscission was suddenly mentioned while no experiment has been performed to specifically examine it.
13) Figure 2–figure supplement 1: Controls are missing to prove that the bands in the bead-bound fractions represent the specific binding to RhoA instead of beads or GST. Binding to the GST-alone beads needs to be performed in parallel. How is the binding affected by the E448K mutation?
14) In the subsection “Mutants in the active site of the CYK-4 GAP domain are cytokinesis-defective”, the authors claim: “Thus, the GAP activity of CYK-4 is not essential for post embryonic development but…”. This is superficially true. However, there is another possibility that the mutant message/product might delay the consumption of the wild-type message/product derived from the heterozygous mother and kept the level of functional CYK-4 enough for post embryonic development (but not for maturation of germ line). The sentences “Thus, while early embryos require the GAP activity of CYK-4, this requirement is relaxed post embryonically” and “…as some cell types may not require the GAP activity of CYK-4, as seen in post embryonic cells in C. elegans” (subsection “GEF activation model accounts for previous results”) should also be reconsidered.
15) “CYK-4R459A hyper accumulates on the membrane as compared to WT CYK-4; this localization suggests that CYK-4R459A is well folded in vivo” – is this logical? In general, partial misfolding might result in tighter membrane binding by forming small aggregates.
16) The sentence: “If furrow formation is dependent on CYK-4 binding to either CED-10/Rac1 or CDC-42, then inactivation of these GTPases would be predicated to cause a phenotype at least as severe as a mutation that weakens the GTPase binding site of CYK-4”. This inexplicitly assumes that these GTPases are positive regulators of the furrow formation. Thus, the logical conclusion of experimental observations is that furrow formation is independent of CYK-4 binding to them or that these are not positive regulators of the furrow formation.
17) Figure 1–figure supplement 4 and 5: Is the structure on which CYK-4E448K::GFP was detected really the gonad membrane? The DIC image is not clear which structure is stained. Only after comparison with the images in Figure1–figure supplement 5 can readers understand what the structure labeled with the CYK-4 mutant in Figure 1–figure supplement 4 is.
18) Please clarify this statement: “These data demonstrate that the catalytic activity of the CYK-4 GAP domain must have a function that is distinct from maintaining CED-10/Rac1 in an inactive sate”. This assumes that CED-10 is the only GTPase to be inactivated by CYK-4.
19) “However, CYK-4 and ECT-2 form a protein complex through their regulatory N-termini, therefore the C-terminal RhoGAP domain of CYK-4 will be in the vicinity of the ECT-2 RhoGEF domain” (subsection “cyk-4 suppressor mutations activate ect-2). This is not true. Whether the C-termini are in the vicinity is nothing to do with the fact that they bind through the N-termini.
20) In the same subsection: “…the E129K substitution, but not G707D, significantly reduces binding of the ECT-2 N- and C-terminal (Figure 7C)…”. As the MBP-ECT-2_N wild type and E129K proteins are the different protein samples prepared independently, strictly, the data provided can't exclude a possibility that MBP-ECT-2_N wild-type shows stronger non-specific binding to the GSH-beads or GST than the E129K mutant. Control experiments with GST instead of GST-ECT-2_DHPH should be performed in parallel.
21) In the Results: “The ECT-2G707D variant exhibits a modest increase in GEF activity compared to wild-type ECT-2 over a range of concentrations (Figure 7D).” This is not really convincing. The same amount of the wild-type and mutant proteins are supposed to have been loaded on the gel. However, there is a small but clear difference in the intensity of the bands, especially the ∼50 kDa minor ones. I am afraid that there might also be a similar difference between the main bands. The difference between 1 µg and 1.2 µg is barely detectable by SDS-PAGE and Coomassie staining. In general, it is not trivial to quantify protein concentration with the precision of less than 10% error. Anyway, in the first place, it is questionable whether such a minor difference in the GEF activity can explain the dramatic in vivo phenotype.
22) The references in this sentence are misleading: “Accurate reconstitutions will need to account for the facts that cytokinetic RhoA activation involves the CYK-4 C1 domain, the ability of CYK-4 to bind to ZEN-4, and the ability of ZEN-4 to oligomerize (Figure 1C) (Basant, et al., 2015).” Figure 1C does not test the oligomerization. Defect in the oligomerization does not prevent RhoA activation (Hutterer, 2010 http://www.ncbi.nlm.nih.gov/pubmed/19962307, Dr. Glotzer is one of the authors of this paper). Basant et al. (2015) demonstrated elevated plasma membrane association of the centralspindlin mutant defective for 14-3-3-binding and thus promoted for oligomerization (zen-4(S682A)). However, it is not clear whether the rather global cortical contractility caused by this mutation is due to activation/inactivation of RhoA or other GTPases.
23) “RhoA inactivation (Miller and Bement, 2009)” (subsection “GEF activation model accounts for previous results”) – this is misleading, as the major claim of this paper is the importance of the flux of RhoA turnover facilitated by combination of GEF and GAP for the formation of a sharp zone of the active form of RhoA.
24) Legend for Figure 6–figure supplement 1: What was the temperature?
25) In the subsection “Image quantification”, it is not clear how the time-series data sets of the normalized cortical distance were analyzed by the Kruskal-Wallis test. Were the rates of ingression calculated and then tested? Or does the test directly compare the shape of the curves? Please provide details and proper references. Referring to the Kruskal-Wallis test by “ANOVA” (legends for Figure 1F-I, 2C, 3D, 4C) is confusing since ANOVA is usually used to refer to (parametric) analysis of variance, which requires the normality of the data, and different from the Kruskal-Wallis test, which is based on ranks and does not make any assumption about the probability distributions of the variables.https://doi.org/10.7554/eLife.08898.030
- Michael Glotzer
- Donglei Zhang
- Michael Glotzer
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
This work was supported by the NIGMS (GM085087), a Chicago Postdoctoral Fellowship to DZ, and NCATS (UL1 TR000430). We thank Benjamin Wolfe, Andy Loria, Agnieszka Grzegorzewska, and Yael Feinstein for their contributions to early stages of this project. We also thank Bruce Bowerman, Douglas Bishop, and Angika Basant for helpful comments on the manuscript. We acknowledge the support of the C. elegans stock center which provided some strains.
- Mohan Balasubramanian, Reviewing Editor, University of Warwick, United Kingdom
© 2015, Zhang and Glotzer
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.