Microtubules that assemble the mitotic spindle are generated by centrosomal nucleation, chromatin-mediated nucleation, and nucleation from the surface of other microtubules mediated by the augmin complex. Impairment of centrosomal nucleation in apical progenitors of the developing mouse brain induces p53-dependent apoptosis and causes non-lethal microcephaly. Whether disruption of non-centrosomal nucleation has similar effects is unclear. Here, we show, using mouse embryos, that conditional knockout of the augmin subunit Haus6 in apical progenitors led to spindle defects and mitotic delay. This triggered massive apoptosis and abortion of brain development. Co-deletion of Trp53 rescued cell death, but surviving progenitors failed to organize a pseudostratified epithelium, and brain development still failed. This could be explained by exacerbated mitotic errors and resulting chromosomal defects including increased DNA damage. Thus, in contrast to centrosomes, augmin is crucial for apical progenitor mitosis, and, even in the absence of p53, for progression of brain development.
Spindle assembly crucially depends on microtubule nucleation by the γ-tubulin ring complex (γTuRC). During mitosis, γTuRC generates microtubules through three different pathways: centrosomal nucleation, chromatin-mediated nucleation, and nucleation from the surface of other microtubules (Meunier and Vernos, 2016; Petry, 2016; Prosser and Pelletier, 2017). The latter mechanism is mediated by the augmin complex and has been referred to as a microtubule amplification mechanism (Goshima et al., 2008; Goshima and Kimura, 2010; Lawo et al., 2009; Uehara et al., 2009). Augmin binds to the lattice of microtubules generated by the centrosome- and chromatin-dependent pathways and, through recruitment of γTuRC, promotes nucleation of additional microtubules that grow as branches from these sites (Alfaro-Aco et al., 2020; Kamasaki et al., 2013; Petry et al., 2013; Tariq et al., 2020). The existence of multiple nucleation pathways may provide some level of redundancy to spindle assembly, but concerted action by multiple nucleation mechanisms has also been described (Hayward et al., 2014; Prosser and Pelletier, 2017). While functional studies in Xenopus egg extract and cultured cell models have generated a wealth of information regarding the types of spindle defects that occur when specific nucleation pathways are compromised, how these defects impinge on cell fate and development remains poorly defined.
Gene mutations that cause functional or numerical centrosome aberrations are associated with primary microcephaly, a developmental disorder that results in the reduced thickness of the cerebral cortex. Depletion of apical progenitors following abnormal mitoses has been identified as a pathogenic mechanism (Jayaraman et al., 2018; Marthiens and Basto, 2020; Nano and Basto, 2017). Apical progenitors of the developing cerebral cortex are highly polarized cells. Their cell bodies are positioned in the ventricular zone (VZ), while their apical and basal processes contact the ventricular surface (VS) and basal lamina, respectively (Arai and Taverna, 2017; Chenn et al., 1998; Chou et al., 2018). Prior to mitosis, the nucleus migrates apically and mitotic chromosome segregation occurs near the apical surface. Early during cortical development, apical progenitors divide symmetrically, expanding the progenitor pool. At later stages they switch to self-renewing asymmetric mitoses, producing a neuron or intermediate progenitor in each division. Centrosomal microtubules were proposed to be at the core of these fate decisions, by controlling the distribution of cell fate determinants through correct positioning of the mitotic spindle (Homem et al., 2015; Taverna et al., 2014; Uzquiano et al., 2018). Recent work showed that progenitor fate is strongly impacted by mitotic duration. Mitotic delay results in more neurogenic divisions and an increased percentage of progenitors undergoing p53-dependent apoptosis, depleting the progenitor pool (Mitchell-Dick et al., 2020; Pilaz et al., 2016). Consistently, mitotic delay, premature differentiation, and apoptosis have all been observed for centrosome defects in mouse models of primary microcephaly (Insolera et al., 2014; Lin et al., 2020; Marjanović et al., 2015; McIntyre et al., 2012; Novorol et al., 2013). Interestingly, in cases where it has been tested, such as Cenpj- or Cep63-deficient mice, the reduced cortical thickness was fully rescued by co-deletion of Trp53, identifying p53-dependent apoptotic cell death as main driver of microcephaly in these models (Insolera et al., 2014; Marjanović et al., 2015). Recently, it was shown that this response involves the USP28-53BP1-p53-p21-dependent mitotic surveillance pathway, which is triggered by prolonged mitosis resulting from centrosome loss (Phan et al., 2021). Depletion of progenitors by apoptosis may be less important in human microcephaly, where organoid models have revealed premature differentiation as the main response (Gabriel et al., 2016; Lancaster et al., 2013).
The roles of chromatin-mediated nucleation and augmin-dependent amplification in this context are less clear. Mouse embryos deficient for Tpx2, a spindle assembly factor that functions in chromatin-mediated nucleation, abort development after a few rounds of highly abnormal mitotic divisions (Aguirre-Portolés et al., 2012). Similar observations were made for mouse embryos lacking the expression of the augmin subunit Haus6 (Watanabe et al., 2016). However, since early mouse development occurs in the absence of centrosomes (Gueth-Hallonet et al., 1993), the embryos in the above studies lacked two of the three mitotic nucleation pathways.
Early functional studies by augmin knockdown in cell lines described mitotic defects that ranged from relatively mild for Drosophila cells (Goshima et al., 2008; Meireles et al., 2009) to more severe for human cells (Lawo et al., 2009), suggesting cell type- or organism-specific differences. Consistent with this, the knockout of augmin in Aspergillus has no obvious phenotype (Edzuka et al., 2014), Drosophila augmin mutants are viable with mild mitotic defects observed in only some cell types (Meireles et al., 2009; Wainman et al., 2009), and a zebrafish mutant is also viable but displays defects in the expansion and maintenance of the hematopoietic stem cell pool (Du et al., 2011). A more recent inducible knockout of the augmin subunit HAUS8 in non-transformed human RPE1 cells caused mild spindle defects before cells underwent p53-dependent G1 arrest, but co-deletion of Trp53 exacerbated the mitotic phenotype (McKinley and Cheeseman, 2017). This response may involve the USP28-53BP1-p53-p21-dependent mitotic surveillance pathway, which is triggered by centrosome loss or prolonged mitosis (Fong et al., 2016; Lambrus et al., 2016; Meitinger et al., 2016), but this was not directly tested.
To uncover the specific role of augmin-mediated microtubule amplification in mitotic spindle assembly and cell fate determination, we sought to study augmin deficiency in centrosome-containing cells in vivo. To this end, we conditionally knocked out Haus6 in proliferating apical progenitors in the embryonic mouse brain using nestin promotor-driven Cre expression. We found that augmin is essential for brain development, promoting mitotic progression, and preventing p53-dependent apoptosis in neural progenitors. Intriguingly, while the absence of p53 promoted growth in Haus6 knockout brains, this was accompanied by exacerbated mitotic errors and disruption of tissue integrity. Our results show that contrary to centrosomal microtubule nucleation, the augmin-dependent pathway is essential for apical progenitor mitotic progression and survival, and thus for brain development.
Previous work has shown that the augmin complex is composed of eight subunits and that depletion of any subunit interferes with augmin assembly and function (Goshima et al., 2008; Lawo et al., 2009; Uehara et al., 2009). Mouse embryos that completely lack expression of the augmin subunit Haus6 do not survive the blastocyst stage (Watanabe et al., 2016). In order to test the specific requirement for augmin in proliferating neural progenitors, we obtained floxed Haus6 mice in which exon 1 of the Haus6 gene is flanked by loxP sequences (Watanabe et al., 2016). To generate Haus6 conditional knockout (Haus6 cKO) mice for the current study, we removed the neomycin cassette that was present in the original strain adjacent to exon 1 (see Materials and methods for details). We then crossed these mice with mice expressing Cre recombinase under the control of the Nestin promoter, to induce Haus6 knockout in apical progenitors starting around day E10.5 (Figure 1a; Figure 1—figure supplement 1a; Graus-Porta et al., 2001; Tronche et al., 1999). In contrast to the full knockout (Watanabe et al., 2016), Haus6 cKO mice passed through all developmental stages and at E13.5 we observed efficient deletion of Haus6 in the brain (Figure 1—figure supplement 1b). Whereas mice with a heterozygous Haus6 deletion developed normally and were fertile, homozygous Haus6 cKO mice died around birth. Analysis of Haus6 cKO animals at E17.5 showed severe defects in brain development, whereas overall body development appeared normal (Figure 1b,c; Figure 1—figure supplement 1c). Histopathology analysis revealed a strong disruption or absence of different forebrain structures (cortex, thalamus, and hypothalamus) and of the cerebellum (Figure 1c; Figure 1—figure supplement 1c). To evaluate whether this was due to agenesis or tissue loss during development, we analyzed embryos at E13.5. Even at this earlier stage, brains in Haus6 cKO embryos displayed severe defects compared to control embryos. Lateral cortexes in Haus6 cKO embryos were almost completely absent and thalamus structures, while partially formed, displayed a strong reduction in radial thickness (Figure 1d,f). Moreover, spaces between tissue structures were filled with cellular debris. These data suggest that, in Haus6 cKO brains at early developmental stages, formation of structures that would give rise to the cortex, thalamus, and hypothalamus is initiated but not completed, leading to tissue loss and abortion of brain development at later stages.
To analyze defective brain development in Haus6 cKO animals at E13.5 at the cellular level, we focused on the thalamus, which was at least partially preserved. We co-stained brain sections with antibodies against PAX6 and βIII-tubulin to label apical progenitors and neurons, respectively. In Haus6 cKO embryos, we observed that the reduced radial thickness in the thalamus was due to a striking thinning of the neuronal layer by ~90% when compared to controls (Figure 1e,f), indicating severely impaired neurogenesis. In some parts, where tissue organization appeared to be disrupted, we also observed neurons that were misplaced in apical regions (Figure 1e). To directly test if augmin deficiency impaired mitoses, we identified and quantified mitotic cells in the thalamus using Ser10-phospho-Histone H3 (pH3-Ser10) staining. In Haus6 cKO embryos, we observed a ~4-fold increase in the number of mitotic cells in the region closest to the VS compared to controls, whereas there were no significant differences in more basal regions (Figure 2a,b). The percentage of Haus6 cKO mitotic cells in prometaphase was strongly increased, whereas metaphases and ana/telophases were reduced relative to controls (Figure 2c; Figure 2—figure supplement 1a). This increase in early and decrease in later mitotic figures were consistent with a delay in spindle assembly. Taken together, these observations suggest that augmin deficiency in progenitors of the thalamus leads to a defect in progression to metaphase, causing mitotic delay.
To analyze cortical progenitors and since there were no intact cortical structures in Haus6 cKO brains at E13.5, we analyzed embryos at E11.5. At this stage, cortical structures were present suggesting that, as for the thalamus, cortical tissue is originally formed but lost at later stages. Similar to the situation in the thalamus at E13.5, in Haus6 cKO cortexes at E11.5 the percentage of mitotic progenitors was increased when compared to controls and this occurred specifically in the apical region and not in more basal layers. Again, this increase in mitotic cells was due to accumulation in prometaphase (Figure 2—figure supplement 1b–e). Taken together, the data show that augmin plays an important role in allowing the timely mitotic progression of apical progenitors in different regions of the developing mouse brain.
To test if augmin-deficient progenitors displayed spindle defects, we analyzed brain sections with antibodies against γ-tubulin and α-tubulin (Figure 2d–i). Mitotic apical progenitors in the thalamus of control animals displayed strong, centrosomal staining of γ-tubulin at spindle poles and more diffuse γ-tubulin signals along spindle microtubules. In Haus6 cKO embryos, γ-tubulin could not be detected on spindle microtubules. Moreover, in ~50% of cells, the staining of γ-tubulin at spindle poles was dispersed into multiple smaller foci (Figure 2d,e). Some of these foci were not associated with centrioles, as revealed by centrin staining, suggesting that they resulted from PCM fragmentation rather than centrosome amplification (Figure 2—figure supplement 1f). Consistent with this, centriole numbers in Haus6 cKO cells were not increased compared to controls (Figure 2—figure supplement 1g). Similar observations were previously made by knockdown of augmin subunits in cell lines (Lawo et al., 2009). Labeling of microtubules by α-tubulin antibodies revealed spindle abnormalities in about half of the mitotic progenitors in Haus6 cKO animals (Figure 2f,g). This included cases where spindle microtubules could not be detected (Figure 2h,i), suggesting decreased stability as previously reported (Goshima et al., 2008; Lawo et al., 2009; Zhu et al., 2008). Defective spindles in Haus6 cKO cells lacked the bipolar configuration with two robust and focused microtubule asters typically seen in controls. Instead, spindle microtubules were associated with multiple, scattered γ-tubulin foci, resulting in spindles that appeared disorganized, sometimes with multiple poles (Figure 2f). However, bipolar configurations including at ana/telophase were also observed and cell divisions occurred in Haus6 cKO progenitors, suggesting that mitosis was not completely blocked.
Considering that augmin-deficiency caused pole fragmentation, we wondered whether this affected spindle positioning. We measured spindle angles relative to the VS in dividing apical progenitors in the thalamus and in the cortex of E13.5 and E11.5 Haus6 cKO embryos, respectively. We found that in both cases the majority of spindles axes were oriented horizontally similar to spindles in control cells (Figure 2—figure supplement 2h–j). This is consistent with results from previous work showing that the presence of multiple spindle poles in progenitors due to extra centrosomes does not significantly affect spindle orientation (Marthiens et al., 2013).
In summary, augmin deficiency in apical progenitors disrupts the recruitment of γ-tubulin to spindle microtubules, causes pole fragmentation, and interferes with bipolar spindle assembly and mitotic progression.
We sought to determine the fate of progenitors undergoing abnormal mitoses after the loss of augmin. We probed thalamus and cortex of E13.5 and E11.5 Haus6 cKO embryos, respectively, for p53 induction and the presence of the apoptotic marker cleaved caspase-3. Indeed, p53 and cleaved caspase-3 were strongly upregulated in both brain regions (Figure 2j,k; Figure 2—figure supplement 2a), whereas cells positive for these markers were barely found in the corresponding tissues of control embryos. To reveal the identity of cells overexpressing p53, we performed a triple staining with antibodies against p53, the neuronal marker βIII-tubulin, and the apical progenitor marker PAX6 (Figure 2l). This experiment showed that in the Haus6 cKO thalamus ~87% of the p53-positive cells were also positive for PAX6 and only a minor fraction (~5%) for βIII-tubulin (Figure 2m). Moreover, we observed that PAX6-positive progenitors displaying p53 induction were exclusively interphase cells, based on the presence of intact nuclei. We concluded that p53 induction occurred specifically in augmin-deficient progenitors, after exit from abnormal mitoses. Interestingly, some cells in the thalamus of Haus6 cKO embryos also displayed upregulated expression of the cell cycle inhibitor p21 (Figure 2—figure supplement 2b,c).
Taken together, the data suggests that mitotic spindle defects in Haus6 cKO progenitors are not catastrophic per se, but efficiently trigger cell cycle arrest and apoptotic cell death upon completion of mitosis.
Since massive apoptosis in Haus6 cKO brains was correlated with p53 induction, we wondered whether cell death was p53-dependent and the cause of aborted brain development. To address this, we crossed Haus6 cKO mice with Trp53 KO mice (Figure 3a). Strikingly, at E13.5, a stage at which Haus6 cKO brains displayed massive apoptosis, lacked cortical structures, and had a poorly developed thalamus, Haus6 cKO Trp53 KO brains showed only minimal signs of apoptosis and there was some growth in the regions where cortex and thalamus would be expected to form (Figure 3b–d). Consistent with this, there was also no upregulation of p21 (Figure 3—figure supplement 1a–d). Tissue growth was enhanced when compared to the single Haus6 cKO brains, but seemed to lack the layered organization observed in control brains at this stage (Figure 3b). At E17.5, however, when thalamus and cortex were well formed in controls, in Haus6 cKO Trp53 KO embryos cortex and thalamus structures appeared thin and undeveloped (Figure 3e). Moreover, as observed for Haus6 cKO embryos, Haus6 cKO Trp53 KO animals were not viable and died around birth.
In summary, massive apoptosis and cell cycle arrest in Haus6 cKO brains are rescued in Haus6 cKO Trp53 KO brains, promoting growth in the affected brain regions, but this growth is not productive for proper brain development.
Next, we examined how co-deletion of Haus6 and Trp53 affected mitosis in proliferating progenitors. Similar to Haus6 cKO alone (Figure 2), Haus6 cKO Trp53 KO embryos also had an increased density of mitotic cells in the cortex and in the thalamus as revealed by Ser10-phospho-Histone H3 staining (Figure 4a–c). The majority of these cells were in prometaphase (Figure 4d) and had disorganized spindles with fragmented spindle poles (Figure 4—figure supplement 1a–d). While these defects were overall similar to those observed in Haus6 cKO brains, we also observed some differences. Centrin staining showed that ~30% of mitotic Haus6 cKO Trp53 KO cells had an increased number of centrioles, indicating the presence of extra centrosomes (Figure 4—figure supplement 1e,f). Mitotic cells with extra centrosomes had a ~2-fold increased size compared to cells with normal centrosome number (Figure 4—figure supplement 1g), suggesting that these cells had previously failed cytokinesis, as observed in augmin-depleted cultured cells (Uehara et al., 2009). Consistent with abnormal cell divisions, we also observed various abnormalities in post-metaphase cells. Compared to Trp53 KO control littermates there was a strong increase in the number of defective anaphases and telophases including multipolar spindle configurations, lagging chromosomes, and micronuclei formation (Figure 4e–g). We also noticed that a fraction of Haus6 cKO progenitors displayed enlarged nuclei in interphase (Figure 4h), suggesting aneuploidy/polyploidy triggered by abnormal chromosome segregation and/or failed cytokinesis. Considering that there were very few apoptotic cells in the double KO brains (Figure 3c), we speculated that continued proliferation may exacerbate mitotic defects. We analyzed multipolar metaphases and abnormal anaphase and telophases in Haus6 cKO and Haus6 cKO Trp53 KO embryos at E13.5 in the thalamus, a structure that was present in embryos of both genotypes at this stage. We found that mitotic defects were more severe in the Haus6 cKO Trp53 KO brains when compared to Haus6 cKO brains (Figure 4i,j).
Since mitotic errors can cause DNA breaks (Quignon et al., 2007), we probed brain tissue of Haus6 cKO Trp53 KO embryos for the presence of γH2AX foci, a marker of an active DNA damage response. Indeed, at E13.5 the percentage of cells with interphase nuclei displaying DNA damage was strongly increased in both the cortex and thalamus when compared to controls (Figure 5a–d). Side-by-side comparison of γH2AX staining in E13.5 thalamus of Haus6 cKO and Haus6 cKO Trp53 KO embryos showed that augmin deficiency led to increased DNA damage relative to controls and that absence of p53 further increased this effect (Figure 5c,d). Thus, the extent of mitotic defects that we observed in Haus6 cKO and Haus6 cKO Trp53 KO embryos was correlated with a concomitant increase in DNA damage.
Centrosome defects result in premature differentiation in human cerebral organoid models (Gabriel et al., 2016; Lancaster et al., 2013). We wondered whether premature differentiation may contribute to the defects observed in augmin-deficient mouse brains. To address this, we labeled embryonic apical progenitors in S-phase by BrdU injection into pregnant mice at E12.5 and sacrificed the embryos for analysis 24 hr later (Figure 6a). We then determined among the BrdU-positive cells the proportion that had exited the cell cycle (negative for Ki67 staining) or that underwent neuronal differentiation (negative for PAX6 staining, positive for βIII-tubulin staining) in cortex and thalamus (Figure 6b–e). We observed that compared to controls the proportion of BrdU-positive, βIII-tubulin expressing cells was reduced in both Haus6 cKO and Haus6 cKO Trp53 KO brains. This result suggested that mitotic defects caused by augmin deficiency did not result in premature differentiation but rather interfered with neurogenesis, and that this was not rescued by co-deletion of Trp53.
Apart from the aberrant mitoses in Haus6 cKO Trp53 KO progenitors, the distribution of mitotic figures within the tissue was also highly abnormal. Whereas in control and Haus6 cKO brains, the vast majority of mitotic figures with condensed chromosomes were observed in the apical region, near the VS (Figure 2a,b; Figure 2—figure supplement 1b,d), in Haus6 cKO Trp53 KO brains most of the mitotic figures were distributed throughout the tissue including more basal regions (Figure 4a–c).
The presence of large numbers of basally positioned mitotic figures in the cortex and thalamus of Haus6 cKO Trp53 KO embryos could indicate that apical progenitors had delaminated, that their nuclei did not migrate to the apical region prior to division, or that the cells displaying mitotic defects in basal layers were not apical progenitors. The latter possibility was tested by PAX6 staining (Figure 7a). Whereas in the cortex of Trp53 KO controls PAX6-positive cells were confined to the VZ, well separated from more basally positioned neurons labeled by βIII-tubulin staining, in Haus6 cKO Trp53 KO cortex PAX6-positive cells localized indiscriminately in basal and apical regions of the cortex, largely overlapping with regions populated by βIII-tubulin-positive neurons (Figure 7a,d). Interestingly, TBR2-positive intermediate progenitors, residing in the subventricular zone in control sections, had also lost this confined localization in Haus6 cKO Trp53 KO cortexes (Figure 7b,e). During development, apical progenitors in interphase maintain a bipolar structure with their centrosomes lining the VS, a configuration that is readily visualized by γ-tubulin staining in control embryos (Figure 7c). In Haus6 cKO Trp53 KO embryos, apical centrosome localization was strongly reduced and sometimes completely lost (Figure 7c,f). Instead, clusters of γ-tubulin foci were observed in subventricular regions, where they were never observed in controls (Figure 7c). Centrin staining indicated the presence of many centrioles, confirming that these were clustered centrosomes rather than PCM fragments (Figure 7—figure supplement 1a). Taken together, these observations suggested that progenitors in Haus6 cKO Trp53 KO cortexes were not only incorrectly positioned, but had also lost their polar organization. To assess this more directly, we stained for nestin, an intermediate filament protein specifically expressed in apical progenitors. In the cortex of control embryos, nestin-stained progenitors displayed a highly polarized, apicobasal morphology and a laterally aligned arrangement within the tissue (Figure 7—figure supplement 2a). In contrast, polarized morphology and lateral alignment were completely disrupted in progenitors of Haus6 cKO Trp53 KO embryos (Figure 7—figure supplement 2a). Consistent with these observations, staining with α-tubulin antibodies revealed that microtubules displayed apicobasal organization in control cells, running along the length of the highly polarized cell bodies (Figure 7—figure supplement 2b,c). In contrast, microtubules in Haus6 cKO Trp53 KO progenitors lacked apicobasal orientation and appeared disorganized (Figure 7—figure supplement 2b,c).
Taken together, these data suggest that in Haus6 cKO Trp53 KO embryos apical progenitors had lost their polarized organization and divided ectopically. As a result, neuroepithelium integrity was severely disrupted.
The mitotic spindle serves to segregate the replicated chromosomes faithfully into two daughter cells. This task is carried out by spindle microtubules and a multitude of proteins that nucleate, organize, and remodel these microtubules during mitotic progression. Here, we have analyzed the contribution of one of three different microtubule nucleation pathways, augmin-mediated microtubule amplification, to mitotic spindle assembly in proliferating neural progenitor cells during mouse brain development. Previous work found that impairment of centrosomal microtubule nucleation in apical progenitors slowed mitotic spindle assembly and progression, leading to p53-dependent apoptosis and causing microcephaly (Insolera et al., 2014; Lin et al., 2020; Marjanović et al., 2015; McIntyre et al., 2012; Novorol et al., 2013). Similarly, we found that augmin-deficiency also impaired spindle assembly, delayed mitosis, and induced p53-dependent apoptosis. In agreement with previous functional studies in cell lines (Lawo et al., 2009), augmin-deficient progenitors displayed fragmented spindle poles, but this did not significantly impair spindle positioning. The most important outcome of these defects was cell death. Our finding that the large majority of cells positive for expression of p53 and the apoptotic marker cleaved caspase-3 were PAX6-positive interphase cells, suggests that cell death occurred after completion of abnormal mitoses. Despite the similarities with centrosome defects, the Haus6 conditional knockout phenotype is much more severe. Rather than leading to microcephaly, augmin deficiency completely aborted brain development. To our knowledge, this has not been reported for any other microtubule regulator affecting mitotic spindle assembly and progression. How can this be explained? While mitotic defects and apoptosis were also observed after loss of centrioles by conditional CenpJ/Cpap/Sas4 knockout (Insolera et al., 2014) and amplification of centrosome number by PLK4 overexpression (Marthiens et al., 2013), the specific spindle defects caused by augmin deficiency may be a more potent trigger of apoptotic cell death than defects resulting from centrosome abnormalities. It should be noted that a more recent Cenpj conditional knockout mouse model displayed more severe disruption of forebrain structures, causing lethality a few weeks after birth (Lin et al., 2020). Still, these defects seem less severe than what we observed after augmin knockout. One may expect that preventing cell death in augmin-deficient progenitors would, at least to some degree, rescue brain development. Co-deletion of Trp53 in Haus6 cKO mice largely rescued apoptosis, revealing that cell death was p53-dependent, but did not rescue brain development and lethality. In the absence of apoptosis, augmin-deficient progenitors likely underwent repeated cycles of abnormal mitoses, leading to increasingly severe mitotic abnormalities. This behavior has recently been described after the induced knockout of the augmin subunit HAUS8 in the RPE1 cell line. Whereas HAUS8 knockout in a TRP53 wild-type background only mildly impaired mitosis before cells arrested in G1, co-deletion of TRP53 eliminated cell cycle arrest and exacerbated mitotic defects (McKinley and Cheeseman, 2017). Consistent with this possibility, Haus6 cKO Trp53 KO progenitors had more severe mitotic defects than Haus6 cKO cells, including lagging chromosomes and multipolar spindles at post-metaphase stages, and displayed increased DNA damage. We have not formally tested whether cell death in augmin-deficient progenitors involves the recently described, USP28-53BP1-p53-p21-dependent mitotic surveillance pathway (Lambrus and Holland, 2017). However, our results show that during brain development cells that have undergone erroneous mitosis are efficiently eliminated in a p53-dependent manner, and that this occurs independently of whether the cause is centrosomal or non-centrosomal. The situation may be different in human brain development, where premature differentiation rather than apoptosis was shown to be the main response to centrosome defects in microcephaly organoid models (Gabriel et al., 2016; Lancaster et al., 2013). How human brain development would be affected by augmin deficiency is unclear. However, considering the severity of the Haus6 KO phenotype in mice, augmin deficiency may also be lethal in humans.
The pole-fragmentation phenotype in augmin-deficient mitotic progenitors may be comparable to mitoses in the presence of extra centrosomes, as described in mice overexpressing PLK4 (Marthiens et al., 2013). In these animals co-deletion of Trp53 also exacerbated mitotic defects and aneuploidy, but the outcome was still a microcephalic brain (Marthiens et al., 2013). In contrast, in the case of Haus6 cKO Trp53 KO progenitors in our study, continued proliferation was not productive for brain development. While some cortical structures were present at E13.5, they lacked a pseudostratified epithelial organization. Progenitors had lost their characteristic, highly polarized morphology and formed a disorganized cell mass that was intermingled with βIII-tubulin-positive differentiated neurons, in both apical and basal regions. Considering that the polarized apical progenitor morphology is integral to the organization of the neuroepithelium, providing scaffold function and guidance for translocating basal progenitors and migrating neurons, it is not surprising that these defects lead to abortion of brain development.
Exacerbated mitotic errors and DNA damage as a result of continued proliferation are a reasonable explanation for the severely disrupted tissue integrity in Haus6 cKO Trp53 KO brains. However, we cannot exclude that additional roles of augmin contribute to this phenotype. For example, augmin may promote progenitor polarity by generating and/or maintaining the apicobasal interphase microtubule array. Recent work has shown that experimentally altered spindle positioning in progenitors can lead to loss of apical membrane. This can be compensated for by re-extension of the apical process and re-integration of the apical foot at the VS (Fujita et al., 2020). Assuming a role of augmin in progenitor polarity, this process may be impaired in augmin-deficient cells. Consistent with this possibility, microtubules in Haus6 cKO Trp53 KO progenitors appeared disorganized, lacking the apicobasal alignment that is observed in control cells. However, it is unclear whether this is cause or consequence of the loss of polarized cell morphology. It should also be noted that augmin nucleates microtubules in post-mitotic neurons, affecting their morphogenesis and their migration (Cunha-Ferreira et al., 2018; Sánchez-Huertas et al., 2016), which could contribute to tissue disruption in Haus6 cKO Trp53 KO brains.
In summary, our work shows that, in contrast to centrosomal nucleation, augmin-mediated microtubule amplification in neural apical progenitors is essential for brain development and cannot be compensated for by the chromatin- and centrosome-dependent nucleation pathways. As in the case of progenitors lacking centrosomal nucleation, mitotic delay caused by augmin deficiency triggers p53-dependent apoptosis. While cell death can be prevented by co-deletion of Trp53, the specific defects that result from the loss of augmin are sufficient to completely abort brain development, independent of p53 status.
Nestin-Cre Haus6 cKO were obtained by crossing Haus6 floxed (Haus6fl) mice with B6.Cg-Tg(Nes-cre)1Kln/J mice. Haus6 floxed Neo mice (Haus6fl-Neo) (Accession no. CDB1218K, http://www2.clst.riken.jp/arg/mutant%20mice%20list.html) were generated as described (Watanabe et al., 2016). To generate Haus6 floxed mice (Haus6fl) (RBRC09630, Accession no. CDB1354K, http://www2.clst.riken.jp/arg/mutant%20mice%20list.html, Haus6fl-Neo mice were crossed with C57BL/6-Tg(CAG-flpe)36Ito/ItoRbrc (RBRC01834) (Kanki et al., 2006). The resultant mice without the PGK-neo cassette (Haus6 flox mice) were maintained by heterozygous crossing (C57BL/6N background). B6-Tg(CAG-FLPe)36 was provided by the RIKEN BRC through the National Bio-Resource Project of the MEXT, Japan. B6.Cg-Tg(Nes-cre)1Kln/J) mice were a gift from Maria Pia Cosma (CRG, Barcelona, Spain) and previously purchased from Jackson Laboratories. To obtain Nestin-Cre Haus6 cKO Trp53 KO mice, mice carrying the floxed Haus6 (Haus6fl) and Nestin-Cre alleles were crossed with mice lacking p53. p53-deficient mice (B6.129S2-Trp53tm1Tyj/J) were purchased from Jackson Laboratories. All the mouse strains were maintained on a mixed 129/SvEv-C57BL/6 background in strict accordance with the European Community (2010/63/UE) guidelines in the specific-pathogen-free animal facilities of the Barcelona Science Park (PCB). All protocols were approved by the Animal Care and Use Committee of the PCB/University of Barcelona (IACUC; CEEA-PCB) and by the Departament de Territori I Sostenibilitat of the Generalitat de Catalunya in accordance with applicable legislation (Real Decreto 53/2013). All efforts were made to minimize use and suffering.
Genotyping was performed by polymerase chain reaction (PCR) using genomic DNA extracted from tail or ear biopsies. Biopsies were digested with Proteinase-K (0.4 mg/ml in 10 mM Tris-HCl, 20 mM NaCl, 0.2% SDS, and 0.5 mM EDTA) overnight at 56°C. DNA was recovered by isopropanol precipitation, washed in 70% ethanol, dried, and resuspended in H2O. To detect Haus6 wt (800 bp), Haus6 floxed (1080 bp), and Haus6 KO (530 bp) alleles by PCR the following pair of primers were used: mAug6KO_FW (5′-CAACCCGAGCAACAGAAACC-3′) and mAug6KO_Rev (5′-CCTCCCACCAACTACAGACC-3′). These PCRs were run for 35 cycles with an annealing temperature of 64.5°C. To detect the transgenic Cre-recombinase allele in Nestin-Cre cKO mice (100 bp) primers olMR1084 (5′-GCGGTCTGGCAGTAAAAACTATC-3’) and olMR1085 (5′-GTGAAACAGCATTGCTGTCACTT-3′) were used. For this PCR, primers olMR7338 (5′-CTAGGCCACAGAATTGAAAGATCT-3′) and olMR7339 (5′-GTAGGTGGAAATTCTAGCATCATCC-3′) were used as internal control (324 bp). These PCRs were run for 35 cycles with an annealing temperature of 51.7°C.
Pregnant females with embryos at E12.5 were injected intraperitoneally with 5-Bromo-2′-deoxyuridine (BrdU) (B5002; Sigma-Aldrich) diluted in phosphate-buffered saline (PBS) at a final concentration of 120 mg per kg of animal weight. After 24 hr, embryonic brain tissue was processed for histopathology analysis as described in the next section.
For histopathology analysis of mouse embryos, timed pregnant female mice were euthanized and embryos were removed. Following euthanasia, embryo heads were fixed in 4% PFA diluted in PBS overnight at 4°C, followed by cryoprotection in increasing concentration of sucrose in PBS (first 15%, then 30%, with a 24 hr incubation at 4°C for each sucrose concentration), followed by overnight incubation in a 1:1 solution of 30% sucrose and OCT (Tissue-Tek). Tissues were then embedded in OCT and frozen in liquid nitrogen-cooled isopentane. For tissue histological analysis, 10-µm-thick cryosections were prepared, placed on glass slides, and processed for either hematoxylin/eosin staining using standard protocols or for immunofluorescence staining. For immunofluorescence staining, cryosections were thawed at room temperature, washed with PBS, and subjected to heat-mediated antigen retrieval in citrate buffer (10 mM citric acid) at pH 6, as required. Tissue sections were permeabilized with PBS containing 0.05% TX100 (PBS-T 0.05%) for 15 min and blocked with blocking solution (10% goat serum diluted in PBS-T 0.1%). Sections were then incubated overnight at 4°C with primary antibodies diluted in blocking solution. The next day, after washing with PBS-T 0.05%, sections were incubated for 60 min with Alexa-Fluor conjugated complementary secondary antibodies and DAPI to stain DNA. Sections were again washed with PBS-T 0.05% and mounted with Prolong Gold antifading reagent (Thermo Fisher Scientific).
For immunofluorescence stainings after BrdU incorporation sections were dried and fixed with neutral buffer formalin (HT501128-4L, Sigma-Aldrich) for 10 min. Antigen retrieval was performed using citrate buffer pH 6 for 20 min at 97°C using a PT Link (Dako-Agilent). Quenching of endogenous peroxidase was performed by incubation for 10 min with Peroxidase-Blocking Solution (S2023, Dako-Agilent). Unspecific unions were blocked using 5% of goat normal serum (16210064, Life Technologies) with 2.5% BSA (10735078001, Sigma-Aldrich) for 60 min. Blocking of unspecific endogenous mouse Ig staining was also performed using Mouse on Mouse (M.O.M) Immunodetection Kit – (BMK-2202, Vector Laboratories). Primary antibodies were diluted in EnVision FLEX Antibody Diluent (K800621, Dako-Agilent) and incubated overnight at 4°C. Secondary antibodies were diluted at 1:500 and incubated for 60 min. Samples were stained with DAPI (D9542, Sigma-Aldrich) and mounted with Fluorescence mounting medium (S3023, Dako-Agilent). Specificity of staining was confirmed by staining with rabbit IgG, polyclonal Isotype control (ab27478, Abcam), mouse IgG1, Kappa Monoclonal (NCG01) Isotype Control (ab81032, Abcam), or a mouse IgG2a kappa Isotype Control (eBM2a) (14-4724-82 IgM, Invitrogen).
Immunohistochemistry (IHC) was performed using 7 µm cuts. Prior to IHC, antigen retrieval was performed using Tris-EDTA buffer pH 9 for 20 min at 97°C using a PT Link (Dako-Agilent). Quenching of endogenous peroxidase was performed by a 10 min incubation with Peroxidase-Blocking Solution (Dako REAL S2023). Blocking was done in M.O.M. blocking reagent (MKB-2213, Vector Laboratories), 5% of goat normal serum (16210064, Thermo Fisher Scientific) mixed with 2.5% BSA diluted in Envision Flex Wash buffer (K800721, Dako-Agilent) and with Casein solution (ref: 760-219, Roche) for 60 min and 30 min, respectively. Primary and secondary antibodies were diluted with EnVision FLEX Antibody Diluent (K800621, Dako-Agilent) and incubated for 120 min. Antigen–antibody complexes were revealed with 3-3′-diaminobenzidine (K3468, Dako-Agilent). Sections were counterstained with hematoxylin (S202084, Dako-Agilent) and mounted with Mounting Medium, Toluene-Free (CS705, Dako-Agilent) using a Dako CoverStainer. Specificity of staining was confirmed by using a mouse IgG1 isotype control (ab81032, Abcam).
All antibodies are listed in the key resources table.
Histology sections stained with hematoxylin/eosin (Figure 1c,d; Figure 1—figure supplement 1c; Figure 3b,e) or used for IHC (Figure 5a,c; Figure 2—figure supplement 2b; Figure 3—figure supplement 1a,c) were imaged with the digital slide scanner Nanozoomer 2.0 HT from Hamamatsu and processed with NDP.view two software from Hamamatsu. Immunofluorescence labeled histology sections (Figure 1e; Figure 2a,d,h,j,l; Figure 2—figure supplement 1a,b,c,f,h; Figure 2—figure supplement 2a; Figure 3c; Figure 4a,b; Figure 4—figure supplement 1a,c,e; Figure 6; Figure 7; Figure 7—figure supplement 1; Figure 7—figure supplement 2) were imaged with a Leica TCS SP5 laser scanning spectral confocal microscope. Confocal Z-stacks were acquired with 0.5 µm or 1 µm of step size depending on the experiment and using laser parameters that avoided the presence of saturated pixels. Immunofluorescence-labeled histology sections shown in Figure 2f and Figure 4e were imaged with a Zeiss 880 confocal microscope equipped with an Airyscan. In the images shown in Figure 2f, for the Superresolution Airyscan mode a 63× magnification, 1.4 NA oil-immersion lens with a digital zoom of 1.8× was used. The z-step between the stacks was set at 0.211 µm. In the images shown in Figure 4e, for the Fast Airyscan mode a 40× magnification 1.2 NA multi-immersion lens with a digital zoom of 1.8× was used. The z-step between the stacks was set at 0.5 µm. XY resolution was set at 1588×1588. Airyscan raw data were preprocessed with the automatic setting of Zen Black. Additional image processing and maximum intensity z-projections were done in ImageJ software. In each experiment, serial brain sections from multiple animals per genotype were analyzed (details in figure legends).
Radial thickness of the thalamus was measured with ImageJ as the distance between the VS and the basal surface of this brain region in E13.5 embryos. In the same regions, radial thickness of the area occupied by PAX6 and βIII-tubulin cell populations was measured.
For mitotic density, cell counts of the thalamic/cortical wall were divided into 30 µm thick bins from the apical to basal surfaces. The number of mitotic phospho-Histone H3 positive cells was counted in each bin and normalized to the column width of the region analyzed. Mitotic density in each bin was expressed as the number of mitotic cells per 100 µm of column. Centrosome integrity in mitotic cells dividing close to the apical surface of the thalamus/cortex was analyzed by quantifying the percentage of cells displaying unfocused/fragmented spindle poles, each composed of multiple γ-tubulin dots. Mitotic spindle integrity was analyzed in cells dividing close to the apical surface of the thalamus and the percentage of cells displaying abnormal, non-bipolar organized spindles were quantified.
To evaluate p53 expression, cell death, DNA damage, cell cycle exit, and neurogenesis in the embryonic forebrain, representative images of the thalamus/cortex containing the entire apicobasal axis of the tissue were selected. The number of p53 and cleaved caspase-3 positive cells was counted and divided by the area of the selected region. To evaluate the cell population overexpressing p53 in the thalamus, coronal sections were co-stained against p53, PAX6, and βIII-tubulin. Cells in which the p53-positive nucleus was costained with PAX6 were counted as PAX6-positive. Cells with the p53-positive nucleus that did not stain for PAX6 and were surrounded by a cytoplasmic βIII-tubulin signal were considered as βIII-tubulin-positive. To evaluate the expression of phosphorylated Histone H2AX, image files obtained with the Nanozoomer 2.0 HT slide scanner were opened with the image analysis software QuPath (Bankhead et al., 2017). The number of phosphorylated Histone H2AX-positive cells was divided by the total amount of hematoxylin-stained cells in the specific tissue, counted using the QuPath software. To evaluate cell cycle exit and neurogenesis, embryonic tissue sections obtained from pregnant females injected with BrdU were co-stained with BrdU and Ki67 antibodies (for cell cycle exit analysis) or BrdU, PAX6, and βIII-tubulin antibodies (for neurogenesis analysis). To evaluate co-expression of the different markers, image files obtained with the Nanozoomer 2.0 HT slide scanner were opened with the image analysis software QuPath and the ‘Positive Cell Detection tool was used’. Cell cycle exit was analyzed by determining the number of BrdU-positive cells that did not stain for the cell cycle marker Ki67 relative to the total number of BrdU-positive cells in the respective tissue. Neurogenesis was evaluated by determining the number of BrdU-positive cells that were also positive for βIII-tubulin staining but negative for PAX6 staining relative to the total number of BrdU-positive cells in the respective tissue. In all experiments, for each brain, at least two coronal tissue sections were quantified.
To measure interphase nucleus size in cortical neural progenitors (Figure 4h), tissue sections were immunostained with PAX6 antibodies and DAPI to label DNA. The area of nuclei in PAX6-positive cells in the cortex was measured in z-stack images using the ‘Positive cell detection’ tool of QuPath software. Mitotic cells were excluded from this analysis.
To quantify the distribution of neural progenitors within the cortex, cryosections providing lateral views of the cortex were immunostained against PAX6 or TBR2. In both cases, the ‘Plot Analysis’ tool of ImageJ was used to measure signal intensity along the apicobasal axis of the cortex. Measurements were grouped into 9.8-µm-wide bins and the average value for each bin was plotted as the percentage of the sum of all bin intensities.
For analysis of mitotic spindle orientation, cryosections providing coronal views of the thalamus/cortex were immunostained with DAPI and the mitotic DNA marker phosphorylated-Histone H3 and the centrosome/spindle pole marker γ-tubulin. The orientation of the mitotic spindle was then determined by measuring the angle between the pole-to-pole axis and the ventricular lining.
All graphs with error bars are presented as means with standard deviation. To determine statistical significance between samples, an unpaired two-way Student’s t-test was used. Statistical calculations and generation of graphs were performed in Excel or Graphpad Prism6 (ns=not significant, *p<0.05, **p<0.01, ***p<0.001).
All data generated or analyzed during this study are included in the manuscript and supporting files.
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Joseph G GleesonReviewing Editor; Howard Hughes Medical Institute, The Rockefeller University, United States
Anna AkhmanovaSenior Editor; Utrecht University, Netherlands
In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.
[Editors' note: this paper was reviewed by Review Commons.]
This manuscript describes a role for augmin complex during brain development. Augmin complex recruits γ-tubulin ring complex (γTuRC) to microtubule lattices to nucleate microtubule branches. The authors show how loss of Haus6, a part of the augmin complex, in neural progenitors, leads to elevated p53 activity and apoptosis, with severe consequences on overall brain development. In particular, augmin-deleted neural progenitors display spindle abnormalities and mitotic delay, which induce DNA damage accountable for p53-induced apoptosis.https://doi.org/10.7554/eLife.67989.sa1
We thank all three reviewers for their very useful and constructive comments. Below is our point-by-point response.
Evidence, reproducibility and clarity:
The manuscript by Viais R et al. describes a novel role for augmin complex in apoptosis prevention during brain development. Augmin complex recruits g TuRC to microtubule lattices to nucleate microtubule branches. The authors show how -in its absence- neural progenitors have elevated p53 activity and apoptotic rate, with severe consequences on overall brain development. In particular, augmin-deleted neural progenitors display spindle abnormalities and mitotic delay, which induce DNA damage accountable for p53-induced apoptosis.
One point that I personally found very interesting is the role of augmin-dependent MT nucleation depletion in interphase. The authors mention (line 152) that at stage E13.5, besides the number of neurons being reduced, a few neurons were misplaced in the apical region, indicating a role for augmin-driven MT nucleation in cell migration. Moreover, the authors showed that p53 genetic deletion in the Haus6 cKO rescues the apoptosis phenotype but not the tissue disorganisation, suggesting that augmin-dependent microtubule might play a role in tissue polarity. While this is well presented in the discussion, the title in line 268 narrowly refers to mitotic augmin roles. I would like here to see the authors referring to putative roles for augmin-mediated MT nucleation in interphase, by toning down the title in line 268.
We note that severe loss of tissue integrity is evident in the p53 KO background. In this background cells are allowed to repeatedly undergo defective cell divisions with aberrant chromosome segregation, producing increasingly abnormal daughter cells that may eventually fail to support epithelial integrity. Regarding possible neuronal migration defects, this has been previously observed in a study by the Hoogenraad group (Cunha-Ferreira et al., 2018) and this is mentioned in our discussion. To account for the possibility that augmin may have roles beyond mitosis, we have changed the heading to a more neutral statement, not specifically referring to proliferation/mitosis: “Loss of augmin in p53 KO brains disrupts neuroepithelium integrity”.
Overall, the text is well written and flows easily. Figures are clear and legends provide sufficient information on experimental conditions, number of replicates and scale bars. I noticed that, although the number of repeats is specified, the number of cells scored per experiment is not always included. In my comments below I highlight cases where this missing information should be added.
1. In the Cep63 KO (Marjanovic et al., 2015) and the CenpJ KO mice (Insolera et al., 2014), as well as other recently published papers (e.g. Phan TP et al., EMBO Journal, 2020) part of the phenotypical characterisation of the KO mice displays pictures of the overall brain dissected from the mice. Could the author show these images?
The main difference between the cited studies (including our own, Marjanovic et al., on the role of CEP63 in brain development) and our current study is that in the previous studies brains are microcephalic but essentially intact, whereas in our current study brain development was aborted and accompanied by cell death and severe tissue disruption. As a result, in many cases these brains are very fragile and difficult/impossible to isolate. An additional challenge is the fact that brain disruption occurs at a very early developmental stage (before E13.5), where dissection is more difficult than at later stages. We note that all the brains presented in the above cited studies were from later embryonic stages or newborn/adult mice. Therefore, instead of dissecting brains, we decided to present encephalic coronal and sagittal sections as shown in Figure 1c, d, e, Figure 1-supplement 1c, and Figure 3b, e to show the overall impact of Haus6 cKO and Haus6 cKO p53 KO on embryonic brain morphology at E13.5 and E17.5.
2. Fig2d: do the insets correspond to higher magnification images? What is the zoom factor? I could not find it in the legend.
The zoom factor is 1.4 – we have added this information to the figure legend.
3. Fig2E,I and K graphs: how many cells were quantified here over how many experiments? I could not find information in the figure legend.
For all quantitative data, we have added information regarding the number of embryos and counted cells to the figure legends.
4. The impact of Haus6 on mitotic spindle needs further clarification:
– Fig2F: here, the authors show quantification for abnormal and multipolar spindle together. Later on, the abnormal spindle phenotype is no longer discussed (Figure 4). I was wondering what is the individual contribution of abnormal and multipolar spindle, separately. Which one of the two is more frequent? Could the authors explain in the text how they define an abnormal spindle? Is it the lack of MT with the condensed chromosome area?
We agree that our previous classification was somewhat confusing. The spindle defects in Haus6 cKO cells are directly linked to the spindle pole fragmentation phenotype shown in Figure 2d, e. Association of spindle microtubules with these scattered PCM fragments causes spindles to appear overall disorganized. In some cases, multiple smaller asters are present, which is what we had termed “multipolar”. However, this does not always involve multipolar DNA configurations, which we separately quantify in Figure 4. To avoid confusion, we now classify spindle morphologies based on tubulin staining simply as “normal” (bipolar configuration, two robust and focused asters) or “disorganized” (lack of bipolar configuration, in some cases multiple smaller asters). We have also included a better description of this classification (lines 204-212).
– Could it be that augmin deletion induce an instability in MTs within the mitotic spindle, leading to the "empty" or with very few MTs spindles? Or could it be that more cold-sensitive MTs are affected by fixation? What is the percentage of the spindle with no MT in control?
Yes, it is likely that augmin-deficient spindles are less well preserved during fixation due to compromised spindle microtubule stability. Indeed, in tissue culture cells augmin deficient spindle microtubules are more depolymerization-sensitive than controls (Goshima et al., 2008; Lawo et al., 2009; Zhu et al., 2008). We have quantified this effect and found that the percentage of mitotic cells lacking spindle microtubule staining is indeed increased in Haus6 cKO brains (Figure 2h, i).
– Did the authors quantify anaphase/telophase phenotypes as they did in Fig4f?
Yes, this quantification was already included in Figure 4j, where we compared abnormal chromosome configurations between Haus6 cKO and Haus6 cKO p53 KO.
– How do authors explain PCM fragmentation here? Could this phenotype be due to an initial cytokinesis defect which led the cells to accumulate extra centrosomes? Or could this maybe be a product of aberrant PCM maturation/centrosome duplication? Could the authors add here a line to discuss the possible origin of pole fragmentation?
The PCM fragmentation phenotype has previously been described after augmin RNAi in cultured cells (Lawo et al., 2009). We refer to this result and we have added the above reference, to emphasize this point. The authors showed that this phenotype does not involve amplification of centriole number, but is caused by an imbalance in microtubule-dependent forces acting on the PCM and leading to its fragmentation. Thus, the extra poles were formed by acentriolar PCM fragments. We have clarified this issue by quantifying centriole numbers in mitotic cells (when centriole duplication is complete) in control and Haus6 cKO brains. This confirmed the data previously obtained in cell lines and showed that the fragmented spindle poles after Haus6 cKO are not due to extra centrioles (Figure 2-supplement 1f, g) (see also below).
Apart from the PCM fragmentation phenotype that does not involve changes in centriole number, previous work in cultured cells also described cytokinesis defects (Uehara et al., 2009). Failed cytokinesis would indeed lead to increased centriole number. However, it would also increase DNA content, which would be visible by an increase in the size of interphase nuclei. We observed this in Haus6 cKO p53 KO cells, which can undergo repeated divisions in the absence of HAUS6. These data were presented in our previous manuscript version (Figure 4h). We have now quantified centriole numbers and found that these were increased in a subset of mitotic cells in Haus6 cKO p53 KO brains. As predicted and consistent with cytokinesis failure, these cells have an increased size compared to controls with normal centriole number. The new data are presented in Figure 4-supplement 1e,f,g.
5. Figure 4 Did the authors quantify centrosome fragmentation and abnormal spindle here? As they characterised them for the Haus6 cKO mouse, it would be preferable to maintain the same characterisation for the Haus6 cKO p53KO.
We have quantified pole fragmentation and spindle defects as shown for Haus6 cKO in Figure 2 also for Haus6 cKO p53 KO. The new data are presented in Figure 4-supplemement 1.
6. Figure 4C and d: how many replicates were done to obtain these graphs? I think the authors forgot to add this information in the figure legend.
This information has been included in the figure legend.
7. Fig4f,g, I and J: how many cells were counted per experiment? I appreciate the authors writing the n of experiments performed.
We have added this information to the figure legend.
8. Fig5d: how many cells were counted per experiment?
We have added this information to the figure legend.
While it was already known that mitotic delay affects the neuronal progenitor pool through activation of p53-dependent apoptosis (Pilaz L-J, Neuron 2016; Mitchell-Dick A, Dev Neurosci 2020), and that this can be triggered by depletion of centrosomal proteins as Cenpj and Cep63, the role of surface-dependent microtubule nucleation was not identified so far. Some insights come from a Haus6-KO mouse model which dies during blastocyst stage after several aberrant mitosis (Watanabe S, Cell Reports, 2016). In parallel, McKinley KL et al. showed that Haus8 depletion in human cells (RPE1cells) triggered p53-dependent G1 arrest following mitotic defects (McKinley KL, Developmental Cell, 2017). Building on the Hause6 KO mouse and human cell line data, here Viais R et al. discover a novel role for the augmin-mediated MT nucleation in neural progenitor growth and brain development in vivo, through prevention of p53-induced apoptosis.
Specifically, Viais R et al. show that:
1. Surface-dependent microtubule nucleation depletion severely impacts brain development, disrupting partly or completely forebrain domains and cerebellum;
2. Surface-dependent microtubule nucleation depletion induce spindle abnormalities, resulting in mitotic delay in apical progenitors;
3. Mitotic delay results in DNA breaks, p53 activation and p53-induced apoptosis.
This is a tidy, well-executed study with good quality data. These findings propose a novel mechanism that results essential for neural progenitor and overall brain development.
In my opinion, a large audience will benefit from these discoveries: from developmental biologists to cell biologists focused on microtubule dynamics, cell cycle, differentiation, stem cells and cell polarity.
Key works describing my area of expertise: microtubule dynamics, centrosome function, cell cycle regulation and cell polarity.
Evidence, reproducibility and clarity:
Viais, Lüders and colleagues here present an analysis of augmin's roles in neural stem cell development. They describe a dramatic impact of the conditional ablation of Haus6 on embryonic brain development in the mouse, with mitotic problems that lead to greatly-increased levels of apoptosis. The rescue of this apoptosis by mutation of the gene that encodes p53 did not restore brain development, which was still aberrant, due to mitotic errors.
The paper is clearly written, with well-designed and controlled experiments. Its conclusions are well supported by the data presented. I have few comments on the technical aspects of the work- it appears very solid to me.
1. Clearer explanation of the mouse strains used should be provided. The section describing the generation of the Haus6 conditional on p.5 should specify that this is the same as was already published in the 2016 Watanabe paper (this is in the Materials and methods), but this should be more clearly specified. More specific details of the p53 knockout mice from Jackson should be included in the Materials and methods.
We have included additional information describing the generation of the Haus6 cKO mice in the text (lines 135-138). It is not exactly the same as described in the Watanabe et al. paper. The previously published strain (Watanabe et al., 2016) contained a floxed Haus6 cKO allele with a flanking neomycin cassette. For the current study the neomycin cassette was removed. Details are described in the method section and also shown in Figure S1a. Specific information regarding the p53 KO strain has been added to the method section.
2. Figure 1a contains minimal information on the Haus6 locus. More detail should be included for information, if this Figure is to remain (although reference to the targeting details in the original description would be sufficient). It is unclear what the timeline diagram is to convey and it should be improved or deleted. A similar comment applies for the details in Figure 3a, although the colour scheme for the different genotypes is useful.
More detailed information on the Haus6 locus is shown in the schematic of Figure 1-supplement 1a and in the referenced study (Watanabe et al., 2016). Since the targeting of Haus6 exon1 was previously described, we believe that including this information as a supplementary figure and referring to the previous study is appropriate.
Regarding the schematics in Figure 1a and Figure 3a, we have improved these. The timeline shows the time points of Cre expression and of obtaining embryos for analysis.
3. The important PCR controls in Figure S1b have an unexplained 1000 bp band that appears only in the floxed heterozygote. It would be helpful if the authors explained this in the relevant Figure legend.
This band is an artefact and likely represents heteroduplexes of floxed (1080 bp) and wild type (530 bp) DNA strands due to extended regions of complementary. We have explained this in the figure legend.
4. Assuming the putative centrosome 'clusters' in Figure 6c are similar to the fragmented structures seen in thalamus in Figure 2d, a different description should be used to avoid confusion with multiple centrosomes, which is not a phenotype here. It is not clear how the loss of centrosomes from the ventricular surface was scored, whether it was based on total γ-tubulin signal or individual centrosomes; how fragmented poles would affect that is unclear, so the legend and relevant details should clarify this point.
The fragmented spindle poles shown in Figure 2d are different from the centrosome clusters in Figure 6c (now Figure 7c). The fragmented poles are fragments of PCM rather than extra centrosomes. Fragmentation is specific to mitosis, involving forces exerted by spindle microtubules (Lawo et al., 2009). In contrast, the centrosome clusters that we observed in Haus6 cKO p53 KO apical progenitors represent centrosomes from multiple cells in interphase, most likely as part of apical membrane patches that have delaminated form the ventricular surface. In the intact epithelium of controls these centrosomes line the ventricular surface. To avoid confusion, we now indicate in the text and legend that these centrosome clusters involve interphase cells. In addition, using centrin staining, we now show that these clusters contain multiple centrioles (Figure 7-supplement 1), in contrast to the acentriolar PCM fragments in mitotic cells.
5. Phospho-histone H2AX should be referred to as a marker of activation of the DNA damage response, rather than DNA repair.
We have changed the text accordingly.
i. Figure 1b should include a scale bar.
We have added the scale bar.
ii. The labelling of Figure 1f should be revised.
The labels have been fixed.
iii. Figure 2k is not labelled in this Figure.
This has been fixed.
iv. Scale bars should be included in the blow-ups in Figure 6c.
We have added the scale bars.
While it is striking that they see complete disruption of brain development, rather than microcephaly, arguably the mechanistic novelty of the findings is moderate, in that the impacts of Haus6 deficiency on mitotic spindle assembly are well established. The authors only allude to potential additional and novel activities of augmin (in neural progenitors, potentially) that might explain this possibly-unexpected outcome of this study.
The topic is likely to be of interest to people in the field of mitosis, genome stability and brain development.
My expertise is cell biology/ mitosis, less so on murine brain development.
Evidence, reproducibility and clarity:
Jens Lüders and Co demonstrates the essential role of Augmin-mediated MT is critical for proper brain development in mice. The most striking point is that even p53 is eliminated, the microcephaly phenotypes of Haus6 KOs were not rescued. This could mean that the Augmin-mediated MT process is critical to cellular functions that are independent of p53. The authors claim that there are increased DNA damage and excessive mitotic errors. In these aspects, the current work is fascinating.
Nevertheless, what causes massive damage to the neural epithelial tissues in the double mutant is not well explained or examined. Few questions appear in mind before I go into the detail. Are these animals still harbor functional centrosomes and their numerical status?
This is an important point that was also raised by the other reviewers. Based on previous work in cells lines (Lawo et al., 2009), we do not expect that loss of augmin directly impairs centrosome number or MTOC activity. Indeed, Lawo et al. showed that centriole number was unaffected. The only centrosome defect that the authors observed was fragmentation of the PCM during mitosis, but this was shown to be due to imbalanced forces exerted by spindle microtubules: fragmentation could be rescued by microtubule depolymerization or depletion of the cortical microtubule tethering factor NUMA. We have now examined this issue also in our mouse model by staining and counting of centrioles in mitotic apical progenitors of control and Haus6 cKO embryos. This confirmed that centriole number is not increased by Haus6 cKO (Figure 2-supplement 1f, g).
The microcephaly part of the introduction needs some more work. In particular, the authors need to explain apical progenitors' depletion, possibly the correct mechanisms in causing microcephaly. By saying cortical progenitors, it becomes vague. Indeed, there would also be cortical progenitors depleted. But, the fundamental mechanisms are the depletion of apical progenitors lined up at VZ's lumen. Two works in this connection generated brain tissues from microcephaly patients carrying mutations in CenpJ and CDK5RAP2 (Gabriel and Lancaster et al). Authors should cite their work and relate their findings to mouse brain data.
We have introduced text changes in the introduction to indicate the specific role of apical progenitor depletion in microcephaly and the differences in the underlying mechanism between mouse and human organoid models (line 61; lines 87-90). In this context we also cite the Gabriel et al. and Lancaster et al. studies.
-What makes me worry is, looking at figure 1E, there is pretty much no brain, and of course, authors have analyzed what is left over. How could one distinguish reduced PAX6 area and TUJ1 area is due to the gross defects in brain development. Clearly, Haus6 KO causes a severe defect in brain development. Thus, deriving a conclusion from the damaged brain can be misleading. One way to circumvent this problem is to perform 2D experiments with isolated cell types (let us say NPCs and testing if they can spontaneous differentiate).
We note that overall brain structures are only lost by E17.5, but brain structures (albeit defective) are still present at E13.5. Indeed, all of our quantifications were done at E13.5 or earlier stages. That being said, we understand the concern that quantifications in defective brain structures may be misleading. However, 2D cultures, for which cells are removed from their tissue context, may have similar issues. For this reason, we have performed BrdU injection experiments. 24 hours after incorporation of the label by proliferating apical progenitors during S phase, we fixed embryos and determined the proportion of BrdU-positive cells that had stopped cycling (Ki67 negative), expressed the progenitor marker PAX6, or were positive for the neuronal marker βIII-tubulin. The data show that there is no significant fraction of cells undergoing premature differentiation (new Figure 6).
Independently of the BrdU experiments, we have also stained brain sections with antibodies against the cell cycle inhibitor p21. Expression of p21 was increased in a fraction of cells in Haus6 cKO brains (Figure 2-supplement 2b,c) and this was rescued in Haus6 cKO p53 KO brains (Figure 3-supplement 1a-d). Together the data confirm apoptosis or cell cycle arrest, but not premature differentiation, as main responses to mitotic errors after Haus6 cKO.
Figure 2: A nice illustration that Hau6 KO animals harbor many mitotic figures. The quantifications lack how many slices and how many cells were analyzed. Simply n=4 does not say much. 4 animals were considered but how many cells/slices would help identify mitotic cells/animals' distribution. A simple bar diagram does not tell a lot.
We have added this information for all quantifications to the figure legends.
As a minor point, how did the authors unambiguously scored prometaphase cells and other mitotic figures? Representative figures will help. Besides, what is the meaning of many prometaphase cells? At least a discussion would help.
This is a good suggestion and we have provided examples of the mitotic figures that we scored in Figure 2-supplement 1a. We now explain the meaning of the increase in prometaphase cells in the description of this result (lines 178-179).
Can the authors probe centrosomes (not by using γ-tubulin) and relate their presence or absence to p53 upregulation? This is an important point because a complete loss of centrosome is known to trigger p53 upregulation. This may be different in Haus6 KO. This could mean (i.e, centrosomes are normal in numbers or increase in numbers), p53 upregulation is regardless of centrosomes loss.
Indeed, we believe that p53 upregulation in Haus6-deficient brains is not caused by loss of centrosomes. Instead, our data suggest, as explained in the discussion, that mitotic delay caused by augmin deficiency is sufficient for p53 upregulation. We can now further support this conclusion by showing that centriole numbers are similar after Haus6 cKO (Figure 2-supplement 1f, g).
I have a hard time to ascertain how the authors scored interphase cells that enriched with p53. Some representative images with identity markers will help.
Scoring p53-positive interphase cells is relatively straightforward since the p53 signal is nuclear and not observed in mitotic apical progenitors. We have included a magnified region of the tissue shown in Figure 2l, displaying PAX6/p53-positive nuclei of individual cells.
Looking at the p53 status in Haus6 KO animals, it is intriguing that p53 upregulation is not unique to centrosome loss. At this point, it becomes essential to thoroughly analyze the centrosome status to cross-check if Haus6 loss abrogates centrosomes; if so, how much.
Since centrosome number is linked to centriole number, we have addressed this point by quantifying centriole numbers by centrin staining in mitotic apical progenitors (see above) (Figure 2-supplement 1f, g).
Double KO could subside the cell death, but not tissue growth is impressive. So what is going on there? Is there a premature differentiation that leads to NPCs depletion? I believe the authors should generate 2D experiments with cells derived from these double KO animals compared to Haus6 KO and test if there is a premature differentiation that can lead to malformation of the forebrain. Here staining for the forebrain progenitor markers will additionally help (Perhaps FOXG1).
As already explained in more detail above, BrdU injection prior to fixation, followed by staining with cell cycle and cell type-specific markers did not reveal any evidence for premature differentiation after Haus6 cKO.
Looking at Figure 6, it becomes clear that the double KOs have severe issues in maintaining the apical progenitors suggesting that they undergo premature differentiation before attaining a sufficient pool of NPCs. Testing this will bridge the paper between descriptive findings to mechanisms.
This point relates to the reviewer’s previous point: do Haus6 cKO p53 KO apical progenitors prematurely differentiate? We believe that cell loss, tissue disruption, and aborted development may also be explained without premature differentiation. In the absence of p53, repeated abnormal mitoses (Figure 4, Figure 4-supplement 1), and the resulting increasingly severe chromosomal aberrations including DNA damage (Figure 5) may produce cells that eventually won’t be able to proliferate and function properly. This interpretation is also supported by the new BrdU injection experiments that did not detect evidence for premature differentiation.
The Discussion section is excellent, but it should add some human relevance. in particular, are there p53 dependent cell deaths that have been described in human tissues. In my opinion, it seems specific in the mouse brain. The discussion can also have statements about why the human brain is so sensitive even for mild mutations. I am not sure if those human mutations can cause similar defects in the mouse brain. Most of the mice based studies have been focusing on eliminating complete genes of interest.
We have included a section in the discussion to relate our findings to human brain development and the differences with results obtained in mouse models regarding the role of apoptosis (lines 421-424).
Overall, this is a very well done work but requires some more experiments for mechanisms understanding. Addressing those will make the paper fit to get published.
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- Jens Lüders
- Jens Lüders
- Jens Lüders
- Jens Lüders
- Ricardo Viais
- Marcos Fariña-Mosquera
- Marina Villamor-Payà
- Sadanori Watanabe
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
The authors are grateful to Gohta Goshima (Nagoya University, Japan) for generously providing floxed Haus6 mice that were generated in his laboratory and for feedback on the manuscript. The authors acknowledge excellent support by the IRB Barcelona Histopathology and Advanced Digital Microscopy core facilities for help with sample preparation and analysis, and by the Mouse Mutant Core facility for deriving floxed Haus6 mice from sperm samples. The authors thank Travis Stracker (NIH-NCI, Bethesda) for mouse cage space and discussion, Eduardo Soriano and Antoni Parcerisas (University of Barcelona, Spain) for technical help and discussions, Andrew Holland (Johns Hopkins University, Baltimore) for anti-centrin antibodies, Pia Cosma (CRG, Barcelona, Spain) for providing Nestin-Cre mice, and Irina Matos (The Rockefeller University, New York) for helpful comments on the manuscript.
Animal experimentation: All the mouse strains were maintained on a mixed 129/SvEv-C57BL/6 background in strict accordance with the European Community (2010/63/UE) guidelines in the Specific-Pathogen Free (SPF) animal facilities of the Barcelona Science Park (PCB). All protocols were approved by the Animal Care and Use Committee of the PCB/University of Barcelona (IACUC; CEEA-PCB) and by the Departament de Territori I Sostenibilitat of the Generalitat de Catalunya in accordance with applicable legislation (Real Decreto 53/2013). All efforts were made to minimize use and suffering.
- Anna Akhmanova, Utrecht University, Netherlands
- Joseph G Gleeson, Howard Hughes Medical Institute, The Rockefeller University, United States
© 2021, Viais et al.
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.
Dynamic regulation of transcription is crucial for the cellular responses to various environmental or developmental cues. Gdown1 is a ubiquitously expressed, RNA polymerase II (Pol II) interacting protein, essential for the embryonic development of metazoan. It tightly binds Pol II in vitro and competitively blocks the binding of TFIIF and possibly other transcriptional regulatory factors, yet its cellular functions and regulatory circuits remain unclear. Here, we show that human GDOWN1 strictly localizes in the cytoplasm of various types of somatic cells and exhibits a potent resistance to the imposed driving force for its nuclear localization. Combined with the genetic and microscope-based approaches, two types of the functionally coupled and evolutionally conserved localization regulatory motifs are identified, including the CRM1-dependent nucleus export signal (NES) and a novel Cytoplasmic Anchoring Signal (CAS) that mediates its retention outside of the nuclear pore complexes (NPC). Mutagenesis of CAS alleviates GDOWN1’s cytoplasmic retention, thus unlocks its nucleocytoplasmic shuttling properties, and the increased nuclear import and accumulation of GDOWN1 results in a drastic reduction of both Pol II and its associated global transcription levels. Importantly, the nuclear translocation of GDOWN1 occurs in response to the oxidative stresses, and the ablation of GDOWN1 significantly weakens the cellular tolerance. Collectively, our work uncovers the molecular basis of GDOWN1’s subcellular localization and a novel cellular strategy of modulating global transcription and stress-adaptation via controlling the nuclear translocation of GDOWN1.
Axon degeneration contributes to the disruption of neuronal circuit function in diseased and injured nervous systems. Severed axons degenerate following the activation of an evolutionarily conserved signaling pathway, which culminates in the activation of SARM1 in mammals to execute the pathological depletion of the metabolite NAD+. SARM1 NADase activity is activated by the NAD+ precursor nicotinamide mononucleotide (NMN). In mammals, keeping NMN levels low potently preserves axons after injury. However, it remains unclear whether NMN is also a key mediator of axon degeneration and dSarm activation in flies. Here, we demonstrate that lowering NMN levels in Drosophila through the expression of a newly generated prokaryotic NMN-Deamidase (NMN-D) preserves severed axons for months and keeps them circuit-integrated for weeks. NMN-D alters the NAD+ metabolic flux by lowering NMN, while NAD+ remains unchanged in vivo. Increased NMN synthesis, by the expression of mouse nicotinamide phosphoribosyltransferase (mNAMPT), leads to faster axon degeneration after injury. We also show that NMN-induced activation of dSarm mediates axon degeneration in vivo. Finally, NMN-D delays neurodegeneration caused by loss of the sole NMN-consuming and NAD+-synthesizing enzyme dNmnat. Our results reveal a critical role for NMN in neurodegeneration in the fly, which extends beyond axonal injury. The potent neuroprotection by reducing NMN levels is similar to the interference with other essential mediators of axon degeneration in Drosophila.