Non-centrosomal microtubule organizing centers (ncMTOCs) are found in most differentiated cells, but how these structures regulate microtubule organization and dynamics is largely unknown. We optimized a tissue-specific degradation system to test the role of the essential centrosomal microtubule nucleators γ-tubulin ring complex (γ-TuRC) and AIR-1/Aurora A at the apical ncMTOC, where they both localize in Caenorhabditis elegans embryonic intestinal epithelial cells. As at the centrosome, the core γ-TuRC component GIP-1/GCP3 is required to recruit other γ-TuRC components to the apical ncMTOC, including MZT-1/MZT1, characterized here for the first time in animal development. In contrast, AIR-1 and MZT-1 were specifically required to recruit γ-TuRC to the centrosome, but not to centrioles or to the apical ncMTOC. Surprisingly, microtubules remain robustly organized at the apical ncMTOC upon γ-TuRC and AIR-1 co-depletion, and upon depletion of other known microtubule regulators, including TPXL-1/TPX2, ZYG-9/ch-TOG, PTRN-1/CAMSAP, and NOCA-1/Ninein. However, loss of GIP-1 removed a subset of dynamic EBP-2/EB1–marked microtubules, and the remaining dynamic microtubules grew faster. Together, these results suggest that different microtubule organizing centers (MTOCs) use discrete proteins for their function, and that the apical ncMTOC is composed of distinct populations of γ-TuRC-dependent and -independent microtubules that compete for a limited pool of resources.
Described nearly 50 years ago, microtubule organizing centers (MTOCs) generate specific spatial patterns of microtubules as needed for cell function [1]. The best-studied MTOC is the centrosome, a non-membrane bound organelle that organizes microtubules into a radial array from its pericentriolar material (PCM) or from subdistal appendages attached to the mother centriole. However, in many types of differentiated cells, microtubules are organized at non-centrosomal sites to accommodate diverse cell functions. In animal cells, these non-centrosomal MTOCs (ncMTOCs) can be found in the axons and dendrites of neurons, around the nuclear envelope of skeletal muscle cells, at the apical surface of epithelial cells, and at the Golgi complex [2–8]. ncMTOCs can promote nonradial arrangements of microtubules, such as the linear arrays of microtubules present along the apicobasal axis in epithelial cells. How ncMTOCs are established and whether they are composed of the same proteins that impart MTOC activity at the centrosome are largely unknown. In general, MTOCs can be defined as cellular sites that nucleate, anchor, and stabilize microtubules; however, the molecular basis for these functions has been elusive [9,10]. Because of the inherent structural and chemical polarity of microtubule polymers, microtubules are nucleated and anchored at their minus ends. Thus, a defining feature of MTOCs is that they interact with microtubule minus ends. The first microtubule minus-end protein described was γ-tubulin, which, together with GCP2 and GCP3, forms the γ-tubulin small complex (γ-TuSC) [11]. In some organisms, additional γ-tubulin complex proteins (GCPs) combine with the γ-TuSC to form the larger γ-tubulin ring complex (γ-TuRC); organisms lacking these additional GCPs are thought to oligomerize γ-TuSCs into similar ring complexes [12,13]. To date, only the γ-TuSC components TBG-1/γ-tubulin, GIP-1/GCP3, and GIP-2/GCP2 have been identified in C. elegans, suggesting that C. elegans γ-TuRC may share the yeast γ-TuRC composition. As our experiments do not distinguish between γ-TuSC and γ-TuRC, we will use the term γ-TuRC for simplicity. A putative C. elegans ortholog of mitotic spindle-organizing protein associated with a ring of γ-tubulin (MOZART1), MZT1, a γ-TuRC-interacting protein and proposed γ-TuRC component, was identified based on sequence homology to the Arabidopsis ortholog, but its function has not been investigated [14]. γ-TuRC has microtubule nucleation capacity and can also cap the minus ends of microtubules, preventing minus-end growth or depolymerization [15,16]. Whether γ-TuRC predominantly functions as a nucleator or as a minus-end cap or anchor in vivo is a matter of debate. Although γ-TuRC is essential in organisms ranging from yeast to humans, γ-TuRC depletion does not result in the elimination of all microtubules from the cell, suggesting that other mechanisms exist to grow and anchor microtubules at MTOCs. γ-TuRC removal in vivo has severely deleterious effects on mitosis, but microtubules are still present [17–19]. The presence of microtubules in dividing C. elegans embryonic cells appears to rely on both γ-TuRC function and the mitotic kinase AIR-1/Aurora A [20,21], as only depletion of both TBG-1 and AIR-1 from dividing cells results in the elimination of centrosomal microtubules. Whether γ-TuRC and AIR-1, or other essential centrosomal MTOC proteins, function redundantly to build microtubules at ncMTOCs in animal cells is unknown, most notably because of the early requirement of these proteins in mitosis that prohibits an assessment of any later roles during differentiation. In C. elegans embryonic intestinal cells, MTOC function is reassigned from the centrosome during mitosis to the apical surface as cells begin to polarize [6], thereby establishing an apical ncMTOC in each cell. Intestinal cells all derive from the “E” blastomere, undergoing four rounds of division before polarizing at the “E16” stage, when the intestinal primordium is comprised of 16 epithelial cells (S1A Fig, [22]). Shortly after the E8–E16 division, the E16 cells follow a stereotypical pattern of polarization and establish their apical surfaces facing a common midline [6], the eventual site of the lumen of the epithelial tube. ncMTOCs positioned along these apical surfaces nucleate and organize microtubules into fountain-like arrays emanating away from the midline on either side [6,23,24]. Intriguingly, many centrosomal MTOC proteins, including AIR-1 and GIP-1, also localize to this ncMTOC [6]. Because the earliest stages of ncMTOC formation can be easily tracked, the embryonic intestinal primordium provides an ideal system in which to test the role of specific proteins in ncMTOC establishment in vivo. Here, we test the hypothesis that the proteins required to build microtubules at the centrosome play a similar role at ncMTOCs. To do this, we optimized an existing tissue-specific degradation strategy to test the role of GIP-1 and AIR-1 at ncMTOCs in C. elegans embryonic intestinal cells. As at the centrosome, we find that GIP-1 is required to localize the other γ-TuRC members, TBG-1 and GIP-2. Additionally, we show that a predicted ortholog of the γ-TuRC protein MZT1 is essential in C. elegans and colocalizes with γ-TuRC in all contexts but is uniquely required for localization of γ-TuRC to the PCM, and not to the centriole or to the apical ncMTOC. This differential requirement for proteins at the centrosome versus the apical ncMTOC was a common trend, as AIR-1 was also only required to localize GIP-1 and TAC-1 to the PCM, but not to the apical ncMTOC. In addition to GIP-1 and AIR-1, we assessed the requirement of other known microtubule regulators, including ZYG-9/chTOG, PTRN-1/CAMSAP, NOCA-1/Ninein, and TPXL-1/TPX2. Surprisingly, we found that, overall, the depletion of these proteins did not disrupt apical microtubule organization. Furthermore, removal of GIP-1 had only a minor effect on microtubule dynamics at the apical ncMTOC; a subset of microtubules was perturbed, as indicated by a change in EBP-2/EB1 localization and dynamics. These results highlight the differences between the centrosome and other MTOCs and suggest that ncMTOCs are composed of at least two populations of microtubules, γ-TuRC-dependent and γ-TuRC-independent.
To test the role of γ-TuRC and AIR-1 at ncMTOCs, we needed a strategy to deplete proteins essential in the early embryo (early essential proteins) at later stages of development. For example, we had previously been unable to assess the function of γ-TuRC components in differentiated cells in vivo, as their depletion causes severe mitotic defects that result in early embryonic lethality. Tissue-specific degradation strategies have provided a means to deplete such early essential proteins [25,26]. We therefore optimized an existing tissue-specific degradation system (Fig 1A and 1B). The germline cell fate determinant PIE-1 is degraded in somatic cells in the early embryo ([27], Fig 1C). This degradation requires a zinc finger domain 1 (ZF) on PIE-1 and the SOCS-box protein ZIF-1, which targets PIE-1 for degradation by an E3 ubiquitin ligase [27]. Previous reports found that the ZF domain could be added to any protein of interest and that protein is degraded in somatic cells by ZIF-1 ([28], Fig 1E and 1G). However, endogenous ZIF-1 is only expressed in the early embryo, so degradation of targets later in development requires exogenous expression of ZIF-1 [29]. The major drawback of this system is that degradation of early essential proteins by endogenous ZIF-1 leads to an early arrest. We found that a zif-1 deletion mutant is viable (92% ± 8. 4% embryonic viability in zif-1 (gk117) worms compared to 99% ± 1. 9% in N2 worms) despite the apparent loss of ZIF-1 activity (Fig 1C and 1D). Using a zif-1 mutant background (“zif-1 (−) ”), we can tag any gene with the ZF domain using CRISPR/Cas9 and the resulting ZF-tagged protein is not degraded (Fig 1F, 1H, 1I, 1L and 1O), thus allowing for normal development. We then express ZIF-1 under the control of a tissue-specific promoter to degrade ZF-tagged targets. Using this strategy, we tagged the γ-TuRC component GIP-1/GCP3, the predicted MZT1 ortholog W03G9. 8, which we hereafter refer to as MZT-1 (see below), and the mitotic kinase AIR-1/Aurora A with ZF: : GFP, allowing us to monitor protein expression and localization and to degrade each protein with exogenous ZIF-1 expression (Fig 1I–1Q). As expected, GIP-1 localized to the apical ncMTOC in intestinal cells and AIR-1 decorated microtubules (Fig 1I and 1O, [6]). ZIF-1 was then expressed using the promoter for the elt-2 gene, which is expressed exclusively in the intestine starting at intestinal stage E2 (S1A Fig). ZIF-1 expression led to intestine-specific removal of GIP-1, MZT-1, or AIR-1 (“GIP-1gut (−), ” “MZT-1gut (−), ” “AIR-1gut (−) ”), as demonstrated by the loss of both apical and cytoplasmic GFP signal (Fig 1I–1Q, S1B–S1E Fig, S2A–S2D’ Fig). We quantified this intestine-specific depletion in two ways. First, we measured the total amount of reduction of GFP signal in the intestinal primordium of “gut (−) ” embryos as compared to “gut (+) ” siblings that lacked the ZIF-1-expressing array (percent GFP depletion: GIP-1gut (−), 93. 1%; MZT-1gut (−), 92. 1%; AIR-1gut (−), 82. 1%; S3A Fig). This is likely an underestimate that is due to the out-of-focus light contributed by non-degraded ZF: : GFP in non-intestinal cells that complicates this assessment, especially when highly expressed genes like air-1 were tagged. We also took line scans across the midline of the intestinal primordium in both gut (−) and gut (+) embryos to measure apical enrichment (Fig 1K, 1N and 1Q, S3B and S3D Fig). We see no significant apical enrichment of GFP signal in GIP-1gut (−), MZT-1gut (−), or AIR-1gut (−) embryos and a significant reduction in both apical and cytoplasmic GFP intensity, as compared to gut (+) control embryos. This degradation strategy can also be used to co-deplete AIR-1 and GIP-1 ([GIP-1; AIR-1]gut (−), S1D Fig, S2A–S2D’ Fig, Materials and methods). Thus, the zif-1 mutant coupled with the ZIF-1/ZF degradation system provides an effective tool for depleting early essential proteins in a tissue-specific manner. To test the role of γ-TuRC and AIR-1 in establishing the apical ncMTOC, we needed to effectively remove these proteins from intestinal cells prior to the E16 stage, when cells reassign MTOC function to the apical membrane. Intestinal differentiation in C. elegans proceeds in the absence of cell division [30], suggesting that mitotic defects in intestinal cells per se would not affect their ability to build an ncMTOC. Thus, we could begin degradation of the desired targets during the intestinal divisions to ensure they would be effectively cleared by the time cells began to polarize and establish the apical ncMTOC. In C. elegans, loss of maternal AIR-1 results in severe mitotic defects, including multinucleate cells, polyploidy, disorganized microtubules, and failed centrosome separation [31,32]; the additional removal of γ-TuRC components results in monopolar spindles and loss of centrosomal microtubules in the first cell division [20,21]. We thus used mitotic defects in intestinal cells as a phenotypic readout for effective degradation of GIP-1 and AIR-1 prior to polarization. The E blastomere undergoes four rounds of division to generate the polarized 16-cell primordium (E16, Fig 2A). As ZIF-1 was expressed from an early promoter (elt-2p) that is active beginning around E2–E4, we expected that successful removal of γ-TuRC and AIR-1 should result in polarized intestinal primordia with between 2 and 16 cells. Indeed, we found that degradation of GIP-1, MZT-1, or AIR-1 resulted in embryos with 8. 6 ± 2. 3,7. 6 ± 1. 8, or 9. 2 ± 1. 5 intestinal nuclei, respectively, and that degradation of both AIR-1 and GIP-1 resulted in embryos with 4. 0 ± 0. 0 intestinal nuclei (Fig 2A and 2B). Expression of ZIF-1 from a promoter that is active around E8 (ifb-2p) also significantly reduced the number of intestinal nuclei, although to a lesser extent (Fig 2A and 2B, S1A Fig). As further proof that we were effectively depleting the desired targets, we found that embryos with decreased or no zygotic air-1, and with only a maternal supply of ZF: : GFP-tagged AIR-1 (“AIR-1*”), had intestinal nuclear numbers indistinguishable from AIR-1gut (−) embryos (see Materials and methods, 9. 1 ± 2. 2, Fig 2B). We frequently observed mitotic defects in AIR-1gut (−) and GIP-1gut (−) embryos, such as scattered condensed chromosomes, binucleate cells, and abnormal mitotic spindles (S2I Fig, S1 Movie). These results are consistent with the reported role for γ-TuRC and AIR-1 in mitosis and suggest that GIP-1 and AIR-1 are effectively depleted from intestinal cells beginning at approximately E4. We next tested whether intestinal cells can polarize and differentiate in the absence of GIP-1 or AIR-1. In intestinal cells depleted of both GIP-1 and AIR-1, the apical polarity protein PAR-3, whose localization to the apical surface is a hallmark of apicobasal polarity [33], was unperturbed (S2E–S2H Fig). Similarly, we observed intestine-specific lysosome-like organelles known as gut granules (Fig 3D), which are hallmarks of intestinal differentiation [23,30]. Together, these results confirmed that we could use tissue-specific degradation to deplete GIP-1 and AIR-1 prior to apical ncMTOC formation without any dramatic effects on intestinal differentiation. MZT proteins have been shown to recruit γ-TuRC to spindle poles in plants, fungi, and human cell culture lines but have not been characterized in vivo in animal cells [14,34–38]. A previous study of the Arabidopsis homolog of the small protein MZT1 (GIP1) identified the uncharacterized gene W03G9. 8 (hereafter called mzt-1) by sequence homology as a potential MZT1 ortholog in C. elegans (Fig 3A, [14]). We tagged the endogenous locus of mzt-1 with ZF: : GFP to monitor endogenous MZT-1 localization and assess its function. In all embryonic, larval, and adult tissues examined, we observed consistent colocalization of ZF: : GFP: : MZT-1 with tagRFP: : GIP-1, including at centrosomes in the early embryo, the apical membrane of the intestinal and pharynx primordia, the cell junctions of seam cells, and the plasma membrane and centrioles of the germline (Fig 3B). In the intestinal primordium, MZT-1 localized to centrioles and the apical ncMTOC in an identical pattern to all three γ-TuRC components GIP-1/GCP3, GIP-2/GCP2, and TBG-1/γ-tubulin (Fig 3C, 3F, 3I and 3L). Like other γ-TuRC members tbg-1 and gip-1 [17,39], we find that mzt-1 is also required for embryonic viability. In a zif-1 (+) background, ZF: : GFP: : MZT-1 is degraded ubiquitously in somatic cells by endogenous ZIF-1, and we observed fully penetrant maternal effect lethality (0/420 embryos hatched). By contrast, in control embryos (zif-1 (+); GFP: : MZT-1), endogenous GFP: : MZT-1 is not degraded by ZIF-1 and the embryos are viable (340/344 hatched and grew to adulthood). Based on sequence homology, colocalization with GIP-1, and embryonic lethality, our results suggest that W03G9. 8/mzt-1 is the C. elegans ortholog of MZT1. Our ability to deplete GIP-1 (Fig 3G) and MZT-1 (Fig 3E) in intestinal cells afforded us the opportunity to test their role in the recruitment of each other and of other γ-TuRC components to the apical ncMTOC. At the centrosome, γ-TuRC components exhibit interdependent localization [17]. We see a similar requirement for GIP-1 in localizing MZT-1, GIP-2, and TBG-1 to the apical membrane (compare Fig 3C, 3I, 3L to 3D, 3J and 3M), suggesting that, as at the centrosome, these proteins localize to the apical ncMTOC as a complex. Additionally, the disrupted localization of other γ-TuRC components upon GIP-1 depletion further confirmed that we had successfully perturbed GIP-1 function. In contrast to GIP-1gut (−) embryos, GIP-1, GIP-2, and TBG-1 all localized apically in MZT-1gut (−) embryos (Fig 3H, 3K and 3N), suggesting that MZT-1 is not required to recruit γ-TuRC to the apical ncMTOC. The surprising finding that γ-TuRC still localizes to the apical ncMTOC upon MZT-1 depletion led us to further investigate the role of MZT-1 in γ-TuRC recruitment to different sites. γ-TuRC localizes to three distinct locations around the time of polarization. During the divisions that precede intestinal polarization, γ-TuRC accumulation at the PCM is coupled with the cell cycle. GIP-1 is recruited to the PCM of the centrosome as cells enter mitosis (Fig 4A). Following mitosis, the centrosome migrates to the lateral membrane and the PCM is greatly reduced. In this lateral configuration, we frequently observe two γ-TuRC positive puncta per cell, which, by electron microscopy, are centrioles associated with a small shell of PCM (“paired centrosomes” [6], Fig 4B, “interphase” inset, blue double arrows). The paired centrosomes then migrate to the apical surface, where they appear as naked centrioles that completely lack PCM by electron microscopy [6]. At this stage, GIP-1 localizes to both the centrioles and the apical ncMTOC (Fig 4C). By contrast, MZT-1gut (−) embryos localized GIP-1 to their centrioles but failed to recruit GIP-1 to the PCM during mitosis (compare Fig 4A to 4D). We frequently saw inappropriate numbers of centrioles in MZT-1gut (−) embryos (number of GIP-1 positive foci in control: 1. 8 ± 0. 5, n = 64 cells; MZT-1gut (−): 3. 0 ± 1. 0, n = 134 cells; two-tailed t test: p = 4. 4 × 10−24), which may be an indirect consequence of earlier mitotic defects and which appeared to result in multipolar spindles (Fig 4D). Following mitosis in MZT-1gut (−) embryos, GIP-1 remained associated with centrioles and accumulated at the apical ncMTOC, as in control embryos (Figs 3H, 4B, 4C, 4E and 4F). To our knowledge, these results provide the first in vivo role for MZT-1/MZT1 in animal cells, suggesting that MZT-1 is a PCM-specific linker for γ-TuRC in dividing cells. Consistent with these findings, human MZT1 appears to promote the targeting and activation of an intact γ-TuRC to the centrosome in human tissue culture cells [38]. In C. elegans, MZT-1 localization tracks with γ-TuRC localization at the centriole, the PCM, and the apical ncMTOC. However, MZT-1 is not required to target γ-TuRC to the apical ncMTOC but requires GIP-1 to localize there, suggesting that MZT-1 is stably associated with the complex even when not playing a targeting role. Many microtubule minus-end proteins are found at MTOCs, reflecting the defining ability of MTOCs to nucleate and stabilize microtubule minus ends. To date, only a handful of these proteins have been identified [5], including γ-TuRC and the microtubule-stabilizing protein PTRN-1/Patronin/CAMSAP [11,15,40–43]. Additionally, NOCA-1/Ninein is often found to colocalize with the minus ends of microtubules, although it has never been shown to directly bind to minus ends [44]. A previous report found that γ-TuRC and NOCA-1 function together in parallel with PTRN-1 to maintain non-centrosomal microtubule arrays in C. elegans larval and adult skin [45]. Furthermore, the NOCA-1 h-isoform appears to localize to the membrane ncMTOC in the C. elegans adult germline using a palmitoylation site [45]. In the absence of this site, γ-TuRC is required to target NOCA-1 to the ncMTOC. We found that both PTRN-1 and NOCA-1 localize to the apical ncMTOC in wild-type embryonic intestinal cells (S4A, S4B and S4E Fig). We therefore tested the roles of GIP-1 and MZT-1 in the apical localization of PTRN-1 and NOCA-1. PTRN-1 and NOCA-1 (d- and e-isoforms) appeared to localize normally in GIP-1gut (−) or MZT-1gut (−) embryos (S4C, S4D, S4F and S4G Fig). The NOCA-1 d- and e-isoforms lack the NOCA-1h region containing the characterized palmitoylation site; however, we did not rule out the use of alternative palmitoylation sites, which the d-isoform is predicted to contain [46]. AIR-1/Aurora A is a mitotic kinase that helps activate MTOC function at the centrosome, in part by driving PCM accumulation of targets required for microtubule nucleation and polymerization, such as γ-TuRC and TAC-1/TACC3 [32,47–50]. The phosphorylated, kinase-active form of AIR-1 localizes to the apical ncMTOC along with GIP-1 and TAC-1 [6], suggesting that AIR-1 could similarly regulate the accumulation of these targets at the apical ncMTOC. We first asked whether AIR-1 is required for normal GIP-1 and TAC-1 accumulation at the mitotic centrosome of dividing E8 cells; as predicted from previous studies in other cell types and organisms, average GFP: : GIP-1 centrosomal fluorescence was significantly reduced in AIR-1gut (−) cells compared with control cells (532. 7 ± 172. 8 versus 1,194. 6 ± 329. 9, p = 1. 35 × 10−7, Fig 5A–5C, see Materials and methods). We note that centriolar GIP-1 signal appeared to remain upon AIR-1 depletion (Fig 5B). We observed a similar reduction in GFP: : TAC-1 levels at the mitotic centrosome of AIR-1gut (−) cells compared to control cells (501. 9 ± 91. 7 versus 972. 0 ± 463. 6, p = 2. 88 × 10−6, Fig 5D–5F), suggesting that AIR-1 can be efficiently depleted prior to ncMTOC formation. In contrast to the E8 mitotic centrosomes, GFP: : GIP-1 and GFP: : TAC-1 were still recruited to the apical ncMTOC in control and AIR-1gut (−) embryos (Fig 5G–5J). These results indicate that AIR-1 is required for GIP-1 and TAC-1 recruitment to intestinal mitotic centrosomes, as is known in other cell types and organisms [48,51], but that AIR-1 is not required for their localization to the apical ncMTOC, further distinguishing the centrosome from the apical ncMTOC. γ-TuRC and AIR-1 are required to nucleate microtubules at the centrosome in C. elegans [20]. We used GIP-1gut (−), MZT-1gut (−), AIR-1gut (−), and [GIP-1; AIR-1]gut (−) embryos to test whether these proteins are similarly required to build microtubules at the apical ncMTOC. To monitor microtubules upon depletion, we simultaneously expressed intestine-specific mCherry: : TBA-1/α-tubulin along with “early” ZIF-1 (Fig 6A and 6B) or we expressed “late” ZIF-1 and labeled microtubules ubiquitously with mCherry: : TBA-1 from a maternally expressed integrated transgene (Fig 6E). In control embryos, microtubules emanate from the apical surfaces of intestinal cells (Fig 6B and 6E), an observation that is visualized by plotting mCherry: : TBA-1 intensity along a line drawn across the apical midline and quantified by measuring apical enrichment (Fig 6C–6G, S3 Fig). Despite severe mitotic defects in GIP-1gut (−) cells revealed by live imaging, microtubules still localize robustly to the apical ncMTOC (Fig 6A, S1 Movie). We see apical microtubule enrichment in MZT-1gut (−), GIP-1gut (−), AIR-1gut (−), and [GIP-1; AIR-1]gut (−) embryos that is not significantly different (Fig 6A–6F), and in one case significantly higher (Fig 6G), than in control embryos. The surprising finding that known microtubule nucleators are not required to build the majority of microtubules at the apical ncMTOC indicates that other mechanisms or molecular players are required to perform this task. We investigated other known microtubule regulators—the anchoring protein NOCA-1/Ninein, stabilizers PTRN-1/CAMPSAP and TPXL-1/TPX2, and the polymerase ZYG-9/chTOG [18,52,53]—to determine if they are required to organize microtubules apically. We found that apical microtubule enrichment was slightly but significantly decreased only upon depletion of ZYG-9, but that, even then, microtubules remained apically enriched (Fig 6B, 6D and 6F). We were particularly surprised to see grossly normal apical microtubule organization in [GIP-1; NOCA-1]gut (−); ptrn-1 (0) triple mutant embryos, as noca-1 and ptrn-1 are required in parallel to maintain the organization of non-centrosomal microtubule arrays in hypodermal epithelial cells [45]. These results suggest that different MTOCs, and even different ncMTOCs, have distinct molecular and genetic requirements to generate specific microtubule arrays, and that more mechanisms remain to be identified. While we found that overall apical organization of microtubules was intact, we explored whether microtubule dynamics were altered upon depletion of microtubule regulators. One possibility is that a majority of microtubules at the apical ncMTOC are stable, persisting from the mitotic divisions prior to polarization. We tested this possibility in two ways. First, we found that upon nocodazole treatment (10 μg/mL and 30 μg/mL), apical microtubule enrichment was significantly reduced both over time and compared to control-treated embryos (Fig 6H–6J), indicating that apical microtubules can be destabilized. This experiment also demonstrates that our measurement methods (line intensity profiles and apical enrichment) are sensitive enough to detect differences in varying amounts of apical microtubules. Second, we probed microtubule dynamics at the apical ncMTOC by examining the localization of EBP-2/EB1, a microtubule plus-end-binding protein that associates with growing microtubule plus ends. To do this, we tagged the endogenous ebp-2 locus with GFP using CRISPR/Cas9 to visualize endogenous EBP-2 comets. In control embryos, EBP-2 accumulated at the apical surface and moved along the apical surface and out along lateral microtubule tracks toward the basal part of the cell, consistent with microtubules growing from the apical ncMTOC (Fig 7A–7C, S2 Movie). By contrast, EBP-2 enrichment at the apical surface in GIP-1gut (−) embryos was decreased compared to controls (1. 51- versus 1. 67-fold enriched, p = 0. 02; Fig 7C). Additionally, we saw a significant decrease in EBP-2 enrichment in [GIP-1; AIR-1]gut (−) compared to AIR-1gut (−) embryos (1. 47- versus 1. 64-fold enriched, p = 0. 001, Fig 7C), suggesting that the loss of GIP-1 primarily causes the decrease in apical EBP-2 enrichment. Consistent with this observation, we found a significant decrease in the number of EBP-2 comets in [GIP-1; AIR-1]gut (−) compared to AIR-1gut (−) embryos, and in pooled embryos with GIP-1-depleted genotypes compared to pooled embryos with GIP-1 (+) genotypes (Fig 7E, Materials and methods). These results suggest that, unlike at the centrosome, γ-TuRC and AIR-1 are not required to build microtubules at the ncMTOC. Instead, γ-TuRC is required to build a subset of dynamic microtubules alongside other γ-TuRC-independent microtubules at the apical ncMTOC. We next measured the speed of the EBP-2 comets coming from the apical ncMTOC to determine if the dynamics of microtubule growth were altered (Fig 7D and 7F). In control embryos, apically derived comets had an average speed of 0. 558 μm/second, which was significantly slower than the reported speeds for comets originating from centrosomes in the early embryo that had been previously labeled with overexpressed EBP-2: : GFP [18] or for endogenous early embryo centrosomal comet speeds we measured (“2-cell, ” 0. 8884 μm/second, p = 7. 68 × 10−13, two-tailed t test, Fig 7D and 7F). We also measured comet speeds from centrosomes in the E8–E16 division (“E8, ” 0. 563 μm/second), which had similar speeds to apically derived comets and were also significantly slower than comets from centrosomes in the 2-cell embryo (Fig 7D and 7F), suggesting that cell type, cell size, or centrosome size may influence comet speed [54]. Surprisingly, we found that the speed of apically derived comets was significantly increased relative to controls in GIP-1gut (−) (0. 721 μm/second), [GIP-1; AIR-1]gut (−) (0. 726 μm/second), and [GIP-1; NOCA-1]gut (−); ptrn-1 (0) (0. 706 μm/second) embryos (p < 0. 0005 for all comparisons, Fig 7D and 7F). This increase in speed was not observed in AIR-1gut (−) (0. 572 μm/second, p = 0. 66) or MZT-1gut (−) embryos (0. 585 μm/second, p = 0. 33), and comets in GIP-1gut (−) embryos were significantly faster than in MZT-1gut (−) embryos (p = 0. 002). This lack of an increase in apically derived comet speed following AIR-1 or MZT-1 depletion argues that changes in intestinal morphology caused by too few intestinal cells, which should be nearly identical in AIR-1gut (−), MZT-1gut (−), and GIP-1gut (−) embryos, cannot account for the observed increase in comet speeds in GIP-1gut (−) embryos. These results further demonstrate that MZT-1 is not required for γ-TuRC function at the apical ncMTOC. The presence of fewer, faster dynamic microtubules following the depletion of GIP-1 suggests that a limiting microtubule growth factor is normally present at the apical ncMTOC and that the loss of γ-TuRC-based microtubules frees up that limiting factor, permitting faster growth of γ-TuRC-independent microtubules (Fig 7G).
Using a tissue-specific protein degradation system, we tested the role of factors essential for building microtubules at the centrosome in building microtubules at an ncMTOC. These studies reveal two important findings (Fig 7G, S5 Fig): (1) all MTOCs are not equivalent, with different MTOCs requiring distinct proteins to build and localize microtubules and microtubule regulators, and (2) ncMTOCs can be composed of discrete populations of γ-TuRC-dependent and -independent microtubules. We demonstrate that our adapted ZIF-1/ZF degradation system is a robust method for depleting endogenous proteins in a specific tissue of interest (the primordial intestinal epithelium), thereby allowing us to probe the function of early essential genes in differentiating tissues. We monitored and characterized the effectiveness and efficiency of endogenous protein depletion by adding both GFP and ZF via CRISPR to genes of interest. Degradation of many of these critical centrosomal proteins during intestinal divisions caused mitotic defects, confirming that targeted proteins were depleted before the apical ncMTOC formed. With this adapted method now validated, future studies can omit the GFP and use ZF-tagged CRISPR alleles, which will permit a broader range of quantitative analyses of GFP markers. A consequence of early depletion of important centrosomal proteins was fewer intestinal cells, causing architectural defects in the intestinal primordium, such as overall shorter apical midlines. However, the overall reduced midline surface, especially in [GIP-1; AIR-1]gut (−) embryos, does not explain the general upward trend in apical enrichment of α-tubulin we observed; we found no evidence of a correlation between midline length and α-tubulin enrichment (see Materials and methods). In addition to changes in intestinal geometry, early depletion of centrosomal proteins likely also caused changes in ploidy. While these changes may have impacted zygotic gene expression, they cannot explain, for example, the observed differences among MZT-1gut (−), GIP-1gut (−), and AIR-1gut (−) embryos, which all have similar nuclear numbers and thus likely similar ploidy defects. In sum, differences in microtubule dynamics do not appear to correlate with ploidy or architecture defect severity (Figs 2 and 7), indicating that these secondary defects alone cannot account for the changes in microtubule dynamics we observe. Using the ZIF-1/ZF system, we characterized the predicted C. elegans ortholog of MZT1, presenting the first in vivo characterization of a MZT1 ortholog in animal development to our knowledge. As in many systems, MZT-1 colocalizes with other γ-TuRC components, is required for γ-TuRC localization to the mitotic spindle pole, and is essential for viability. Surprisingly, we found that γ-TuRC does not require MZT-1 for its localization to the centrioles and apical MTOC. In addition to localizing γ-TuRC, MZT1 is important for nucleation activity of γ-TuRC in Candida and in human tissue culture cells [38,55]. However, we found that only intestinal GIP-1 depletion, and not MZT-1 depletion, impacted comet number and dynamics, suggesting that the MZT-1 found at the apical MTOC is not important for γ-TuRC activity and may simply be a nonfunctional component of the γ-TuRC complex at these non-PCM sites. Strikingly, we found that dynamic microtubules were still observed growing from the apical ncMTOC following depletion of GIP-1 and AIR-1, which are essential for centrosomal MTOC activity [20]. This finding indicates that additional mechanisms for generating dynamic microtubules must exist. One exciting possibility is that additional, as yet undiscovered nucleators exist in the cell. Based on previous studies on centrosomal microtubules, these hypothetical molecules might be unique for building microtubules at ncMTOCs. Rather than additional nucleators, another possible mechanism could be through the action of microtubule-stabilizing and -anchoring proteins, as has been seen for other ncMTOCs. In fact, the exact role of γ-TuRC in vivo is not known. The relatively poor nucleation capacity of γ-TuRC in vitro suggests that factors that activate its nucleation capacity at MTOCs exist in vivo [12,56]. Alternatively, the primary function of γ-TuRC might not be nucleation, as is suggested by imaging studies of centrosomes from γ-tubulin-depleted C. elegans embryos [57]; a large number of microtubules are still found associated with the centrosome following γ-tubulin depletion, but are disorganized relative to the centrioles. These data raise the possibility that γ-TuRC functions in anchoring microtubules onto the PCM and that perhaps dynamic microtubules at the apical ncMTOC are generated from many different types of stabilized microtubule seeds. For example, proteins like PTRN-1/Patronin/CAMSAP and NOCA-1/Ninein could protect and anchor small microtubule seeds that grow in parallel to the γ-TuRC-based microtubules. Evidence for this type of model has been seen in Drosophila oocytes and C. elegans skin cells [45,58]. However, our analysis of GIP-1gut (−); NOCA-1gut (−); ptrn-1 (0) triple mutants suggests that in embryonic intestinal cells, the ncMTOC does not require PTRN-1 and NOCA-1/Ninein, even in parallel with GIP-1. Furthermore, microtubules remained organized at the apical MTOC upon depletion of the microtubule polymerase ZYG-9/chTOG and the spindle assembly factor TPXL-1/TPX2, suggesting that additional microtubule regulators remain to be discovered. A final possibility is that MTOCs could facilitate microtubule growth not by localizing nucleators but instead by increasing the local tubulin heterodimer concentration. Microtubules can be nucleated in vitro in the absence of any additional molecules, depending on the concentration of tubulin. Recent studies of in vitro reconstituted PCM suggest that the centrosome might build microtubules in part through the selective concentration of tubulin [53]. ncMTOCs might similarly concentrate tubulin, leading to localized microtubule growth. Two of our findings are consistent with this possibility. First, we observed α-tubulin enrichment at the apical MTOC following microtubule depolymerization with nocodazole, although we cannot distinguish between free tubulin heterodimers and small protected microtubule seeds. Second, we found a small but significant decrease in apical microtubule enrichment in ZYG-9gut (−) embryos, which can concentrate tubulin in vitro at SPD-5 condensates [53] and could perhaps play a similar role at the apical ncMTOC. Our finding that microtubules have increased growth speeds following GIP-1 depletion suggests that a limiting growth factor is normally present at the apical ncMTOC. This limiting factor could be sequestered by γ-TuRC itself, or it could be limiting because of competition for it among the large number of growing microtubules. In the first case, we would expect loss of γ-TuRC to release this factor and cause both increased microtubule growth speed and comet number. However, we observed fewer, faster comets and decreased apical EBP-2 enrichment upon GIP-1 depletion. These observations are more consistent with a model in which loss of γ-TuRC leads to fewer growing microtubules, thereby increasing the availability of a limiting factor to growing microtubules and allowing faster growth (Fig 7G). This limiting factor could be tubulin heterodimers themselves; however, the mechanisms for concentrating a pool of tubulin at a membrane are completely unknown. Finally, we found that γ-TuRC and AIR-1 are not required to form the majority of apical microtubules, raising the question of why these proteins so specifically localize there as a new ncMTOC is being established. One possibility is that the main function of localizing γ-TuRC and AIR-1 to the apical ncMTOC is to effectively remove them from the centrosome at the end of mitosis as centrosomal MTOC function is attenuated. We hypothesize that the ability to remove microtubules from the centrosome is an important step in mitotic exit, as hyperactive MTOC function at the centrosome has been linked to cancer [59–61]. Creating a sink for centrosomal microtubule regulators at an alternative site in the cell would provide a quick and effective way of maintaining the inactivation of MTOC function at the centrosome. Different cell types require a large variety of specific patterns of microtubule organization, and future work will be critical to discover the additional molecular players and mechanisms that contribute to the formation and function of different types of MTOCs.
Nematodes were cultured and manipulated as previously described [62]. Experiments were performed using 1- or 2-day-old adults. The strains used in this study are listed in Table 1. Two CRISPR editing methods were used to generate knock-in insertions of the ZF: : GFP or GFP tag. gip-1 (wow3[gfp: : gip-1], gip-1 (wow5[zf: : gfp: : gip-1]) and ptrn-1 (wow4[ptrn-1: : gfp]) were constructed using repair templates with short homology arms (SHAs) and PCR-based screening to test for insertions into the endogenous loci [63]. All other CRISPR-based insertions were achieved using a modified self-excising cassette (SEC), into which the ZF-tag had been inserted N-terminal to GFP (plasmid JF250) [64]. The SEC-derived CRISPR alleles thus also contain additional tags: 3×FLAG for GFP alleles and 3×Myc for TagRFP-T alleles. Cas9 and sgRNAs were delivered using plasmid pDD162, into which the appropriate sgRNA sequence had been added with the Q5 Site-Directed Mutagenesis Kit (NEB). The Cas9/sgRNA plasmid, repair template, and appropriate selection markers were injected into N2 or zif-1 (gk117) mutant 1-day-old adult worms. Worms were recovered and processed according to published protocols [63,64]. Successfully edited worms were outcrossed at least two times before being used for subsequent experiments. sgRNA and homology arm sequences are listed in S1 Table. To assess embryonic viability, 20–30 young adult hermaphrodites of each genotype were singled onto small plates and allowed to lay for 4 hours at 20°C, and then adults were removed and eggs were counted. After 3 days at 20°C, the number of surviving worms present on each plate was counted, and viability for each plate was calculated as the total number of L4s and adults divided by the number of eggs. When comparing N2 and zif-1 (−) (JLF155) lethality, two N2 plates had more surviving worms at day 3 than the number of eggs initially counted, and those plates were omitted. The ZF/ZIF-1 system was executed as previously described, with the following modifications. ZF: : GFP tags were inserted into endogenous loci in a zif-1 (gk117) mutant background using CRISPR editing techniques (see above). Exogenous ZIF-1 was expressed from either extrachromosomal or integrated arrays (Table 2), which were generated by injecting 50 ng/μL of each plasmid and of a co-injection marker (either pRF4[rol-6 (d) ] at 100 ng/μL or myo-2p: : mcherry at 2. 5 ng/μL) into zif-1 (gk117) mutant animals, with pBS to inject at a minimum concentration of 150 ng/μL. For comparisons across multiple genotypes, the same array was introduced by mating. To test the efficiency of AIR-1 depletion—specifically, to test whether zygotic expression of AIR-1: : ZF: : GFP provided AIR-1 activity in AIR-1gut (−) embryos—we generated air-1 (0) embryos with only maternally provided AIR-1: : ZF: : GFP, which was degraded in the intestinal primordium by ZIF-1 expression. Our genetic strategy was as follows: We used the air-1 (ok571) allele, a deletion that removes a large portion of the AIR-1 kinase domain and is a putative null allele. We crossed zif-1 (gk117) stIs10220[end-1p: : his1: : mcherry]; air-1 (ok571) /oxTi878[vha-6p: : gfp]; wowEx10[elt-2p: : zif-1; end-1p: : mcherry: : tba-1] hermaphrodites with zif-1 (gk117) stIs10220; air-1 (wow14[air-1: : zf: : gfp]) males to generate F1 “air-1: : zf: : gfp/ (0) ” hermaphrodites, and scored their F2 embryos. We had no markers to distinguish embryonic genotypes, so we scored all progeny and assumed Mendelian segregation of genotypes: 25% air-1: : zf: : gfp homozygotes, 50% air-1: : zf: : gfp/ (0) heterozygotes, and 25% air-1 (0) homozygotes, all with maternally expressed AIR-1: : ZF: : GFP. Consistent with this predicted distribution, we saw many larvae with perturbed gonad development, a hallmark of the air-1 (0) phenotype [65]. We denote these pooled embryos as AIR-1* in Fig 2. If zygotic expression of AIR-1 was providing activity, we would expect to observe a change in the distribution of intestinal nuclear number in AIR-1gut (−) compared with AIR-1gut (−) *. However, intestinal nuclear numbers in AIR-1gut (−) * and AIR-1gut (−) embryos were indistinguishable, suggesting that zygotic expression of AIR-1: : ZF: : GFP provides little or no AIR-1 rescuing activity and that AIR-1: : ZF: : GFP depletion is highly efficient. We used α-GFP immunostaining to test how completely ZF: : GFP-tagged proteins are degraded in the intestinal primordium (see S2 Fig). We stained [GIP-1; AIR-1]gut (−) embryos (JLF232), in which heterozygous ifb-2p: : zif-1 (wowIs3) causes intestine-specific degradation of both proteins in embryos that inherit the transgene. GFP fluorescence survives the staining protocol, so we asked how the perduring GFP signal correlates with Cy3 signal from α-GFP staining. Most JLF232 embryos fell into two categories: strong Cy3 and GFP signal at centrosomes or the apical membrane (n > 40), or no localized Cy3 nor GFP signal in the intestinal primordium (n = 30). As α-GFP antibody staining is highly sensitive, this result suggests that GIP-1 and AIR-1 are both degraded quickly (prior to GFP maturation) and efficiently, and that the loss of visible GFP in live embryos accurately indicates efficient ZF-tagged protein degradation. We did not find a balancer that allowed us to maintain an EBP-2: : GFP; [GIP-1; AIR-1]gut (−) strain. We thus used the following cross strategy to generate these embryos: ebp-2 (wow47[ebp-2: : gfp]); zif-1 (gk117) /eT1[nIs267]; wowEx10[elt-2p: : zif-1] hermaphrodites were crossed with JLF298 males to generate ebp-2: : gfp; zf: : gfp: : gip-1 zif-1 (gk117) /eT1[nIs267]; air-1: : zf: : gfp/eT1; wowEx10 F1 hermaphrodites. F1 cross progeny were identified by PCR genotyping for air-1: : zf: : gfp (air-1 F: GAACGTCTCCCACTTGTTGACATC, ZF R: TTTTTCTACCGGTACCCTCGG). F2 progeny with the wowEx10 array and without the eT1 balancer were selected (ebp-2: : gfp; zf: : gfp: : gip-1 zif-1 (gk117); air-1: : zf: : gfp; wowEx10). zf: : gfp: : gip-1 was frequently heterozygous, likely due to recombination between eT1 and zf: : gfp: : gip-1 in the F1 mother. Thus, to ensure we were looking at EBP-2: : GFP; [GIP-1; AIR-1]gut (−) embryos, we only scored embryos with four intestinal nuclei (see Fig 2). JLF36 embryos were treated with nocodazole, as has been previously described [6]. Briefly, trypan blue–coated embryos were affixed to poly-lysine-coated coverslips and submerged in embryonic growth medium (EGM) [66] containing either 0. 1% DMSO or 10 or 30 μg/mL nocodazole. E16-stage embryos with their dorsal surfaces facing the coverslip were selected for analysis. The eggshell and vitelline membrane were punctured using a Micropoint nitrogen dye laser (Andor Technology, Belfast, United Kingdom), allowing the EGM + DMSO or nocodazole to reach the embryo. Embryos were imaged immediately following puncturing (T1,10–45 seconds later) and then again after 10 minutes (T2). Embryos were fixed and stained as previously described [23]. Briefly, embryos of the appropriate stage were collected and adhered to a poly-lysine-coated slide with a Teflon spacer and covered with a coverslip. Embryos were fixed by freeze-crack followed by 100% MeOH for 5 minutes. Embryos were rehydrated in PBS and incubated with rabbit α-GFP primary antibody (Abcam, Cambridge, United Kingdom, 1/200) or mouse α-PAR-3 [28] either overnight at 4°C or for 1 hour at 37°C. Embryos were washed in PBT and incubated with the appropriate CY3-conjugated secondary antibodies (Jackson ImmunoResearch Laboratories, West Grove, PA, 1/200) and 100 ng/mL DAPI (Sigma-Aldrich, St. Louis, MO) overnight at 4°C or for 1 hour at 37°C and mounted under coverslips in Vectashield (Vector Laboratories, Burlingame, CA). At least 20 mutant embryos were scored for analysis of mutant phenotypes. Embryos for microscopy were dissected from gravid hermaphrodites incubated for 4–4. 5 hours in M9 (at 20°C for EBP-2: : GFP comet speed, at 25°C for EBP-2: : GFP enrichment, and at room temperature for all other experiments). For live imaging, samples were mounted on a pad made of 3% agarose dissolved in M9. Live imaging was performed on a Nikon Ti-E inverted microscope (Nikon Instruments, Melville, NY) using a 60× Oil Plan Apochromat (NA = 1. 4) or 100× Oil Plan Apochromat (NA = 1. 45) objective and controlled by NIS Elements software (Nikon). Images were acquired with an Andor Ixon Ultra back thinned EM-CCD camera using 488 nm or 561 nm imaging lasers and a Yokogawa X1 confocal spinning disk head equipped with a 1. 5× magnifying lens. For live imaging, images were taken at a sampling rate of 0. 5 μm. Images were processed in NIS Elements, the Fiji distribution of ImageJ (“Fiji”) [67,68], or Adobe Photoshop. Fixed images were obtained using a 60× Oil Plan Apochromat objective (NA = 1. 4) on either the above system or a Nikon Ni-E compound microscope with an Andor Zyla sCMOS camera.
Eukaryotic cells require specific arrangements of microtubules to carry out diverse functions, including cell division and intracellular transport. In dividing animal cells, microtubules are arranged radially around two centrosomes, promoting the correct distribution of DNA into daughter cells. As cells differentiate, microtubules become organized at non-centrosomal sites, often yielding decentralized microtubule arrays. Although a large body of work has focused on understanding how microtubules are grown from and organized by the centrosome, very little is known about how these activities are performed at non-centrosomal sites. We optimized a technique to degrade proteins in C. elegans differentiated cells, specifically testing the role of essential centrosome proteins in building non-centrosomal microtubules, which emanate from the apical membrane in embryonic intestinal epithelial cells. Surprisingly, we found fundamental differences between the centrosome and membrane, both in the mechanisms that recruit microtubule regulators to each site, and in the proteins that are required to build and organize microtubules. In addition, we found that when fewer microtubules grew from membranes, the remaining microtubules grew faster, suggesting competition for limited materials. Together, our study highlights differences in how centrosomes and membranes grow and organize microtubules, and that multiple pathways contribute membrane-organized microtubules.
invertebrates
medicine and health sciences
centrosomes
microtubules
caenorhabditis
cell cycle and cell division
cell processes
astronomical sciences
green fluorescent protein
animals
comets
animal models
developmental biology
caenorhabditis elegans
luminescent proteins
model organisms
experimental organism systems
embryos
cellular structures and organelles
cytoskeleton
digestive system
research and analysis methods
embryology
proteins
gastrointestinal tract
biochemistry
celestial objects
eukaryota
cell biology
anatomy
nematoda
biology and life sciences
physical sciences
organisms
Tissue-specific degradation of essential centrosome components reveals distinct microtubule populations at microtubule organizing centers