Abstract
Malformations of cortical development (MCD) of the human brain are a likely consequence of defective neuronal migration, and/or proliferation of neuronal progenitor cells, both of which are dictated in part by microtubule-dependent transport of various cargoes, including the mitotic spindle. Throughout the evolutionary spectrum, proper spindle positioning depends on cortically anchored dynein motors that exert forces on astral microtubules emanating from spindle poles. A single heterozygous amino acid change, G436R, in the conserved TUBA1A α-tubulin gene was reported to account for MCD in patients. The mechanism by which this mutation disrupts microtubule function in the developing cerebral cortex is not understood. Studying the consequence of tubulin mutations in mammalian cells is challenging partly because of the large number of α-tubulin isotypes expressed. To overcome this challenge, we have generated a budding yeast strain expressing the mutated tubulin (Tub1G437R in yeast) as one of the main sources of α-tubulin (in addition to Tub3, another α-tubulin isotype in this organism). Although viability of the yeast was unimpaired by this mutation, they became reliant on Tub3, as was apparent by the synthetic lethality of this mutant in combination with tub3Δ. We find that Tub1G437R assembles into microtubules that support normal G1 activity, but lead to enhanced dynein-dependent nuclear migration phenotypes during G2/M, and a consequential disruption of spindle positioning. We find that this mutation impairs the interaction between She1 – a negative regulator of dynein – and microtubules, as was apparent from a yeast two-hybrid assay, a co-sedimentation assay, and from live cell imaging. We conclude that a weaker interaction between She1 and Tub1G437R-containing microtubules results in enhanced dynein activity, ultimately leading to the spindle positioning defect. Our results provide the first evidence of an impaired interaction between microtubules and a dynein regulator as a consequence of a tubulin mutation, and sheds light on a mechanism that may be causative of neurodevelopmental diseases.
Introduction
Malformations of cortical development (MCD) are severe brain malformations associated with intellectual disability and infantile refractory epilepsy. MCD include lissencephaly, pachygyri, and polymicrogyria, brain malformations characterized by alterations in cortical gyration and sulcation. These diseases have a strong genetic basis involving the α- and β-tubulin subunits of microtubules, as well as various microtubule associated proteins (MAPs) and effectors. For example, mutations in LIS1 – an effector of microtubule based transport – account for ∼65% of classic lissencephaly (Kumar et al., 2010). LIS1 affects microtubule-based transport by activating the motility of the molecular motor cytoplasmic dynein-1 (Marzo et al., 2020; Mohamed M. Elshenawy, 2020; Zaw Min Htet, 2020), which is also found mutated in MCD patients (Laquerriere et al., 2017; Poirier et al., 2013; Vissers et al., 2010; Willemsen et al., 2012). Recently, a patient with pachygyria and severe microcephaly associated with postural delay and poor communication abilities was shown to possess a single de novo heterozygous mutation in the α-tubulin gene, TUBA1A, that results in a glycine to arginine substitution at position 437 (Bahi-Buisson et al., 2008). Although currently unclear, the cellular basis for disease in this patient may be defects in mitotic spindle orientation, which is a critical process during asymmetric cell division that is required for cellular differentiation and determination of daughter cell fate (Cabernard and Doe, 2009; Das and Storey, 2012; Williams et al., 2011; Wu et al., 2010). Defects in this process can limit the number of progenitor cells, and ultimately the neuronal mass of the developed brain (Bershteyn et al., 2017). This is apparent in mouse neuronal progenitor cells in which dynein dysregulation by Lis1 inhibition impairs microtubule dynamics, results in a mitotic spindle mispositioning phenotype, and leads to a consequential decrease in the number of progenitors during early development (Tsai et al., 2005; Yingling et al., 2008).
We hypothesized that the TUBA1A G436R mutation might cause disease by leading to defective spindle positioning. This may be due to: (1) alteration of microtubule dynamics and/or structure; or, (2) impairment of motor or MAP function due to disrupted microtubule binding. Distinguishing between these possibilities and deciphering the precise molecular defects arising from tubulin mutations is not a trivial task. Studying tubulin mutations in mammalian cells is complicated by the fact that numerous isoforms of α and β-tubulin are present in the human genome (9 α-tubulin, and 10 β-tubulin isoforms) (Findeisen et al., 2014; Khodiyar et al., 2007). Each of these has a distinct expression pattern, and thus every cell’s tubulin content is a composite mixture of these many variants. In contrast to higher eukaryotes, things are much simpler in the budding yeast Saccharomyces cerevisiae in which ∼70-90% of α-tubulin is expressed from the essential TUB1 gene, with the remaining ∼10-30% arising from the TUB3 gene (Bode et al., 2003; Gartz Hanson et al., 2016; Schatz et al., 1986b). In addition to its simplicity, the mechanisms and effectors of spindle orientation (e.g., dynein, Lis1) and microtubule dynamics and function (e.g., tubulin, EB1, CLIP-170, ChTOG) are highly conserved between humans and budding yeast. In this organism, it is mandatory that the mitotic spindle is correctly positioned along the mother-daughter cell axis and in close proximity to the bud neck prior to mitotic exit, otherwise cell viability is compromised. The reliance on this process for viability permitted the use of genetic screens that revealed the presence of two distinct pathways that can effect this process: namely, the Kar9/actomyosin and dynein pathways (Miller and Rose, 1998). The Kar9/actomyosin pathway relies on a microtubule guidance mechanism, whereby a microtubule plus end-associated myosin (Myo2) orients the mitotic spindle along the mother-daughter axis (Hwang et al., 2003; Lee et al., 2000; Yin et al., 2000). Myo2 is recruited to microtubule plus ends by the concerted effort of Kar9 (homolog of human adenomatous polyposis coli tumor suppressor, APC) and the autonomous microtubule plus end-tracking protein, Bim1 (homolog of human EB1). Recently, we have modeled the disease-correlated TUBB2B F265L β-tubulin mutation in budding yeast, and found that this mutation specifically compromised the Kar9/actomyosin pathway by disrupting the plus end localization of Bim1 (Denarier et al., 2019).
Cytoplasmic dynein, on the other hand, functions from the cell cortex, from where Num1-anchored motors walk along microtubules emanating from spindle pole bodies (the equivalent of centrosomes), which results in the positioning of the spindle at the mother-bud neck (Carminati and Stearns, 1997; Heil-Chapdelaine et al., 2000; Li et al., 1993). Dynein is delivered to Num1 receptor sites at the bud cortex by a two-step “offloading” mechanism: (1) microtubule plus end-associated Bik1 (homolog of human CLIP170) recruits dynein-Pac1 (homolog of human Lis1) complexes to dynamic plus ends (Badin-Larcon et al., 2004; Caudron et al., 2008; Lee et al., 2003; Sheeman et al., 2003); (2) plus end-associated dynein, which appears to be inactive (Lammers and Markus, 2015) is delivered, or “offloaded” to cortical Num1 receptor sites along with its effector complex, dynactin (Markus and Lee, 2011). The extent of dynein activity is largely governed by its localization to these sites; however, as in higher eukaryotes (Tan et al., 2019), at least one known MAP can also regulate dynein activity in cells: She1. The precise mechanism by which it does so in cells is currently unclear; however, in vitro studies show that She1 can reduce dynein velocity through simultaneous interactions with both microtubules and dynein (Ecklund et al., 2017), whereas live cell studies have shown that She1 plays a role in polarizing dynein-mediated spindle movements toward the daughter cell (Markus et al., 2012), perhaps in part by tuning dynactin recruitment to plus end-associated dynein (Markus et al., 2011; Woodruff et al., 2009).
To gain insight into the role and importance of α-tubulin G436 (hereafter referred to as G437, due to its position in yeast α-tubulin), and how it might affect the above processes, we produced S. cerevisiae yeast strains in which the native TUB1 locus was replaced with the G437R mutant allele (tub1G437R). Our results show that this mutation leads to alterations in microtubule dynamics, and a spindle positioning defect that is likely due to dysregulated dynein function. The dynein dysfunction phenotype is not a consequence of its mislocalization, but is more likely due to a reduced association of She1 with the mutant microtubules. Although there is no clear She1 homolog in human cells, we propose that this mutation might similarly interfere with dynein function by disrupting the microtubule binding behavior of a regulatory MAP, thus leading to neuronal physiological deficits, and a consequent disruption of cerebral cortex development.
Results
TUB1 G437R mutagenesis leads to an enhancement in microtubule dynamics during G2/M phase
Glycine 436 of TUBA1A α-tubulin, mutation of which is highly correlated with a developmental human brain disease, is conserved among α-tubulins from numerous organisms, including budding yeast (TUB1; overall 74.1% identity between human TUBA1A and yeast Tub1). Glycine 436 (Fig. 1A, red sphere) is one of three highly conserved small, hydrophobic residues in α-tubulin (Fig. 1B, red box) that immediately precede the disordered carboxy-terminal tail of α-tubulin. This region of α-tubulin partly constitutes the external surface of the microtubule to which MAPs and motor proteins bind (Fig. S1). Although some MAPs (i.e., Tau and Tpx2) bind proximal to G437, this residue does not appear to encompass the kinesin and dynein binding interfaces (see Fig. S1) (Alushin et al., 2014; Lowe et al., 2001; Nogales et al., 1998). Modeling an arginine into position 436 of porcine α-tubulin (pdb 3J6G; using UCSF Chimera (Pettersen et al., 2004)) revealed that such a mutation may potentially disrupt the terminal helix of α-tubulin (note the steric clash between the modeled arginine and helix 11 in rotational isomers 1 and 4; Fig. S2). To assess the phenotypic consequences of this mutation, we engineered yeast strains to express Tub1G437R, in which the mutation was introduced at the native TUB1 locus.
Heterozygous diploid cells (TUB1/tub1G437R) were sporulated, and the resulting haploid tub1G437R mutant cells were recovered at the expected frequency. The mutants exhibited no growth defects when cultured in nutrient-rich media (YPAD; see Fig. 1E) indicating that the mutation does not compromise yeast cell viability. Using a chromosomally-integrated RFP-tub1G437R (expressed in the presence of the untagged tub1G437R allele), we found that the mutant tubulin incorporates into spindle and cytoplasmic microtubules during all phases of the yeast cell cycle (G1, G2/M; Fig 1C).
In addition to TUB1, budding yeast possess a second α-tubulin gene encoded by TUB3. Whereas TUB1 is the major α-tubulin isotype and is required for cell viability, cells tolerate deletion of TUB3, which possesses 91% identity and 95% similarity with TUB1 (Schatz et al., 1986a). To determine if cells could tolerate expressing only Tub1G437R, we generated heterozygous TUB1/tub1G437R TUB3/tub3Δ diploid cells, sporulated them, and assessed viability of the recovered single and double mutant progeny. Although the single mutants exhibited relatively normal colony morphology, none of the double mutants were viable (6 out of 6 expected double mutants were inviable), revealing that expression of only the mutant G437R α-tubulin leads to cell death (Fig. 1D). Although the reason for the inviability of the double mutants is unclear, it suggests that either microtubules may not be assembled from only the mutant α-tubulin protein, or microtubules assembled from only the mutant α-tubulin protein do not support proper function. As a consequence of the inviability of the double mutants, we focused the remainder of our study on the tub1G437R single mutants (i.e., in the presence of wild-type TUB3).
Increased sensitivity of cells to the microtubule depolymerizing drug benomyl is a common phenotype of strains with α-tubulin and β-tubulin mutations (Richards et al., 2000). We assessed benomyl sensitivity of wild-type (TUB1) and tub1G437R cells by spotting a dilution series of each on solid media containing different concentrations of the drug (10 and 15 μg/ml; Fig. 1E) and examining cell growth. In the presence of benomyl, cell growth was markedly impaired for tub1G437R cells compared to wild-type cells, indicating an enhanced sensitivity to the drug as a consequence of the mutant tubulin. This suggests that the G437R mutant could be altering microtubule stability or dynamics.
To determine if this was the case, we measured microtubule dynamics parameters by tracking the movement of microtubule plus ends with Bik1-GFP, the homolog of human CLIP-170 (see Fig. 3A). While we did not observe any notable differences in microtubules dynamics parameters between wild-type and tub1G437R strains in G1 cells, we did note several differences during the G2/M phase of the cell cycle (Fig. 2). In particular, we noted an increase in the rates of polymerization (1.4 µm/min in wild-type, versus 1.7 µm/min in mutant cells) and depolymerization for microtubules in tub1G437R cells (1.6 µm/min versus 2.4 µm/min). We also observed a significant increase in the fraction of time the microtubules spent in their growth phase, and a concomitant reduction in the relative fraction of time spent in pause in the mutant cells. This resulted in an overall increase in microtubule dynamicity (Toso et al., 1993) (Fig. 2), which may account for the enhanced sensitivity of the mutant cells to the depolymerizing agent, benomyl. Finally, although the mean microtubule length did not significantly differ between the two strains (Fig. S3A), we noted that the mutant cells exhibited a larger fraction of long microtubules (38% of microtubules were ≥ 7µm in tub1G437R cells, versus 10% in wild-type cells; Fig. S3B and C), many of which extended from one cell compartment to the other (also see below).
The G437R mutation leads to increased spindle dynamics and impaired spindle positioning
Since the G437R mutation specifically affects microtubule dynamics during G2/M phase, during which astral microtubules effect mitotic spindle movements, we sought to assess the consequence of mutagenesis on mitotic spindle dynamics during this phase using Bik1-GFP as a fluorescent reporter (Fig. 3A). In wild-type cells, spindles sampled a relatively small area near the bud neck in the mother cell, and the majority of them were oriented along the mother-bud axis (Fig. S4, and Fig. 3A and B, top). Although the majority of spindles in tub1G437R cells were also oriented along the mother-bud axis (Fig. S4), they exhibited highly dynamic behavior; specifically, we observed numerous instances of the spindle oscillating back and forth between the mother and daughter cell compartments (Fig. 3A and B, bottom). To quantitate this phenomenon, we measured the total distance over which the mitotic spindle moved per minute within each cell. Compared with wild-type cells (TUB1), the spindles in tub1G437R cells moved a significantly longer distance (0.7 µm versus 1.2 µm; Fig. 3C). These observations indicate that excessive forces emanating from the mother and bud cortex are exerted upon the mitotic spindle in tub1G437R cells. We also noted that the spindle movements in tub1G437R cells occurred coincidently with microtubule “sliding” events, during which the plus end of the astral microtubule contacts the cell cortex and then curls along it, all the while maintaining lateral contact (see Fig. 3A, bottom, 3’ thru 6’; also see Video S1). Quantitation of these movements – which are characteristic of dynein-mediated spindle movement events (Adames and Cooper, 2000) – revealed an approximate two-fold increase in their frequency in tub1G437R cells with respect to wild-type cells (Fig. 3D).
A recent study found that dynein-mediated microtubule sliding events are often followed by a microtubule catastrophe (also mediated by dynein), which plays a role in attenuating the spindle movement events (Estrem et al., 2017). We noted that these particular microtubule catastrophe events (i.e., those following sliding events) are greatly reduced in tub1G437R cells (29% of sliding events are followed by catastrophe in tub1G437R cells, versus 85% in wild-type cells; Fig. 3E), suggesting that the G437R mutation reduces dynein’s ability to induce a catastrophe. Also consistent with these data, a larger fraction of mutant cells appeared to exhibit events in which very long microtubules that extended from one compartment to the other (i.e., mother to daughter, or vice versa) underwent characteristic dynein-mediated sliding between the two cellular compartments (Fig. 3F; also see Video S1).
The main function of cytoplasmic microtubules in vegetative yeast cells is to orient the mitotic spindle along the mother-daughter axis (by the Kar9/actomyosin pathway), and localize it proximally to the bud neck (by the dynein pathway) such that at the time of anaphase onset, the chromosomes are divided equally between the mother and daughter cells (Carminati and Stearns, 1997; Hwang et al., 2003; Li et al., 1993; Liakopoulos et al., 2003; Yin et al., 2000). Although we found that orientation of the spindle along the mother-daughter axis was not compromised in tub1G437R cells (Fig. S4) – suggesting the Kar9/actomyosin pathway is not compromised – we did note that the spindle was more frequently localized to the apical regions of the mother or daughter cells in the tub1G437R strain (26% of tub1G437R cells, versus 9% of wild-type cells; Fig. 3G). Taken together, these observations suggest that the G437R mutation leads to increased dynein-mediated spindle movements, and yet reduced dynein-mediated microtubule catastrophe events (following sliding events), which ultimately leads to a spindle mislocalization phenotype.
Enhanced spindle dynamics in tub1G437R mutant cells are dynein-dependent
As noted above, the dynein pathway effects microtubule “sliding” events that result in translocation of the mitotic spindle throughout the cell. Given our observations noted above, we sought to determine whether the tub1G437R mutant phenotypes are a consequence of hyperactive dynein. Consistent with the notion that dynein is responsible for the observed spindle behavior in tub1G437R cells, the increased spindle displacement phenotype in tub1G437R cells was eliminated by deletion of DYN1 (which encodes for the dynein heavy chain), but not by deletion of KIP3, a kinesin that has been implicated in regulating microtubule length and spindle movements (Fukuda et al., 2014; Gupta et al., 2006).
We next asked whether the increased dynein activity is due to enhanced targeting of dynein to microtubule plus ends or the cell cortex, which can be causative of increased cellular dynein activity (Markus et al., 2011). To this end, we imaged Dyn1-3GFP in cells also expressing RFP-Tub1 (or RFP-Tub1G437R) in wild-type or tub1G437R cells (Fig. 4C - E). We found no significant difference in either the frequency of Dyn1-3GFP targeting, or the fluorescence intensity values for Dyn1-3GFP at microtubule plus ends, spindle pole bodies (SPBs; equivalent of centrosomes), or the cell cortex (Fig. 4D and E). Thus, the increased dynein activity in tub1G437R cells is likely not a consequence of increased localization to any of these sites.
G437R microtubules exhibit reduced interaction with the dynein regulator She1
We wondered whether the apparent increase in dynein activity in tub1G437R cells could be a consequence of reduced microtubule binding by the microtubule associated protein (MAP) She1, a dynein inhibitor (Markus et al., 2012; Markus et al., 2011; Woodruff et al., 2009). Since She1 inhibitory activity requires its microtubule binding activity (Ecklund et al., 2017), we first tested the effect of G437R on the interaction between She1 and tubulin. A She1-Gal4 activation domain (AD) fusion was tested for a two-hybrid interaction with either Tub1 or Tub1G437R, the latter of which were fused to the LexA DNA-binding domain (LexADBD). Bim1, which is known to interact with α-tubulin in a two-hybrid assay (Krogan et al., 2006; Schwartz et al., 1997) was used as positive control, as was the kinesin Kip3. As expected, Bim1, Kip3 and She1 all showed a two-hybrid interaction with the Tub1 bait (Fig. 5A; positive interactions are apparent by growth on media lacking histidine, “-HIS”). Interestingly, the interaction between She1 and Tub1 – but not between Tub1 and either Bim1 or Kip3 – was reduced to background levels by the G437R mutation (Fig. 5A).
Like many MAPs, the interaction between She1 and microtubules requires the disordered C-terminal tails of tubulin (Ecklund et al., 2017; Markus et al., 2012). Thus, to further confirm the importance of G437 in the She1-tubulin interaction, we performed a pull-down assay in which this interaction was competitively inhibited by addition of a peptide encompassing the C-terminus of Tub1 (amino acids 415-447; both She1 and tubulin were used at 5 µg/ml or below, well below the critical concentration required for microtubule assembly). To this end, a 6His-She1 C-terminal fragment (She1Cterm; residues 194 to 338; which is sufficient for microtubule binding (Zhu et al., 2017)) was incubated with tubulin (see Fig. 5B for specificity of tubulin-She1Cterm interaction) in the absence or presence of a peptide corresponding to the C-terminus of either wild-type or G437R tubulin (see Methods). The relative extent to which the NiNTA-bead-immobilized She1Cterm pulled down tubulin was subsequently assessed by SDS-PAGE and immunoblot (using an antibody against a-tubulin). This revealed that the She1Cterm-tubulin interaction was strongly competed by the wild-type peptide, but much less so by the same peptide with the G437R mutation (Fig. 5C and D).
Next, we measured the extent to which She1 localizes to microtubules in cells by comparing the relative recruitment of full-length She1 to spindle microtubules (where She1 fluorescence is most prominent) in either wild-type or tub1G437R cells. Consistent with the two-hybrid and in vitro data described above, we found that spindle-localized She1 fluorescence was 55.7% lower in G437R mutant cells than in wild-type cells (Fig. 6A and B). Interestingly, we also noted that the relative fluorescence intensity of mRuby2-Tub1G437R in tub1G437R cells was reduced to a similar extent (by 41.9%) with respect to mRuby2-Tub1 in wild-type cells (Fig. 6A and B). We reasoned this was due to one of two possible scenarios: (1) that the number of microtubules in the mitotic spindle is reduced as a consequence of the G437R mutation, or (2) that the wild-type copy of Tub3 – the less prevalent α-tubulin in budding yeast (see above) – is compensating for a reduced capacity of Tub1G437R to incorporate into spindle microtubules. Since a reduced overall microtubule mass (scenario 1 above) would likely lead to severe consequences (e.g., chromosome segregation defects, reduced viability) that were not apparent by live cell imaging (see Fig. 1C) or apparent cell viability (see Fig. 1E, “YPAD”), we focused on the latter possibility. To this end, we measured the spindle-localized fluorescence intensity of mRuby2-Tub3 in either wild-type or tub1G437R cells and found a 92.3% increase in the extent to which Tub3 is incorporated into the spindles of mutant cells (Fig. 6A and B). Thus, the reduction in Tub1G437R incorporation into spindle microtubules is entirely offset by a compensatory incorporation of Tub3 (Fig. 6D). These data indicate that the Tub1G437R mutant tubulin is less able to incorporate into microtubules, and also explains the reliance of tub1G437R mutant cells on TUB3 (see Fig. 1D).
Finally, we used the relative Tub1/Tub3 spindle intensity values to calculate corrected spindle-localized intensity values for She1. The purpose of this correction was to normalize the degree of apparent She1-spindle binding to the relative microtubule mass. For instance, when She1 intensity values were normalized to Tub1 intensities – both of which were reduced in the mutant cells – the relative degree of She1-microtubule binding was only minimally reduced (Fig. 6C, left). However, when these normalized values were further corrected to account for the increased spindle incorporation of Tub3 into the spindles in the mutant cells, this revealed that She1 binding to Tub1G437R-containing microtubules is reduced by approximately 60% (Fig. 6C, right). Given the fact that the microtubules in the mutant cells are comprised of a roughly equal mixture of Tub1G437R and Tub3 (see Fig. 6D, and Discussion), the reduced spindle intensity of She1 in the mutant cells is likely underestimating the degree by which the mutant tubulin reduces She1-microtubule binding. Taken together, these findings indicate that She1 binding to Tub1G437R-containing microtubules is severely compromised, thus accounting for the hyperactive dynein phenotype in these cells.
Discussion
In summary, we have used budding yeast to characterize the consequences of the G437R α-tubulin mutation (equivalent to G436R in TUBA1A), which is likely causative of MCD in a human patient. Our results indicate that this mutation retains the ability to assemble into microtubules, albeit to a lesser extent than wild-type α-tubulin, which is apparent by the reduction of mRuby2-Tub1G437R fluorescence in spindle microtubules. The reduced incorporation of the mutant tubulin into microtubules appears to be compensated by a corresponding increase in Tub3 incorporation in the tub1G437R mutant, and explains the reliance of tub1G437R mutant cells on Tub3 expression. Previous studies estimated that Tub3 accounts for roughly ∼10-30% of the cell’s α-tubulin content (Bode et al., 2003; Gartz Hanson et al., 2016). From the respective 42% reduction and 92% increase in Tub1G437R and Tub3 microtubule incorporation in the mutant cells, we estimate that the relative abundance of Tub3:Tub1 in mutant microtubules is shifted such that Tub3 now accounts for 55-65% of the cell’s α-tubulin content (Fig. 6D). It is interesting to note that one of these studies (Gartz Hanson et al., 2016) observed a ∼17% increase in the rate of microtubule polymerization that was dependent on the presence of a wild-type copy of Tub3. This suggests that the increased proportion of Tub3 in the mutant microtubules may be partially responsible for the altered dynamicity observed here (16% increase in polymerization rate; Fig. 2).
It is interesting to note that the effects of G437R on dynamic instability were only significant during the G2/M phase of the cell cycle, at which point the growth and shrinkage rates, and the overall dynamicity increase significantly as a consequence of the mutation. Although the reasons for the cell cycle-dependent differences are unclear, they may be due to an inability (or increased ability) of a G2/M-specific factor to bind to and affect microtubule dynamics (either from the plus end, or along the lattice). Given the importance of the C-terminal tail in microtubule binding by numerous factors, and the proximity of G437 to the C-terminal tail of α-tubulin (see Fig. 1A and B), the possibility that this mutation is in fact impacting the structure/function of this region is a likely scenario (see Figs. S1 and S2). Although it is unclear how She1 affects microtubule dynamics, it’s reduced binding affinity for the mutant microtubules raises the possibility that the altered dynamics are a direct consequence of disrupted She1-microtubule binding, or an indirect cause of She1’s inability to modulate dynein or dynein-dynactin (see below).
The most notable phenotype in tub1G437R cells was the dramatic increase in spindle translocation events throughout the mother and daughter cells (see Fig. 3 and Video S1). Spindle movements in budding yeast occur coincidentally with nuclear movement (due to the closed mitosis that takes place in this organism) and involves (1) alignment of the spindle along the mother-bud axis (a Kar9/actomyosin-mediated process), and (2) nuclear migration toward, and into the bud (a dynein-mediated process). Motor proteins are key effectors of this process since they can directly modulate microtubule dynamics (e.g., Kip2, Kip3), and generate pushing and pulling forces via astral microtubules (Carvalho et al., 2004; Chen et al., 2019; Fukuda et al., 2014). In tub1G437R cells, orientation of the mitotic spindle along the mother-bud axis (by the Kar9/actomyosin pathway) was not apparently disrupted (see Fig. S4), whereas dynein-mediated microtubule pulling was dramatically enhanced. Several lines of evidence point to She1 as the main molecular effector of this phenotype. To begin with, G437 is situated on a region of the microtubule surface that is proximal to known contact points for several MAPs (e.g., Tau, Tpx2; see Fig. S1). Our data demonstrate that She1’s interaction with mutated microtubules or tubulin is impaired in vivo, and in two hybrid and pull-down assays. Finally, tub1G437R cells exhibit an increase in dynein-mediated spindle movements in a manner similar to what has been observed in she1Δ cells (Markus et al., 2012). She1 exhibits the unique ability to specifically regulate dynein by reducing dynein’s microtubule dissociation rate, and consequently reducing its rate of motility (Ecklund et al., 2017). Therefore, it is logical that reducing the affinity of She1 for microtubules (as noted in tub1G437R cells) would lead to an enhancement of dynein-mediated pulling forces similar to what was observed in she1Δ cells.
Also of interest is the apparent change in microtubule dynamics we observed specifically during and subsequent to dynein-mediated microtubule sliding events (see Fig. 3E). This decreased frequency of catastrophe events resulted in a greater proportion of long microtubules in the tub1G437R strain, which may also partly account for the increase in dynein activity. It has been shown that increased microtubule lengths directly correlate with enhanced dynein activity in cells (Estrem et al., 2017). Although the role of She1 in affecting microtubule dynamics is not clear, it is possible that dynein-mediated depolymerizing activity – as has been noted in vivo (Estrem et al., 2017) – requires microtubule-bound She1. In particular, Estrem et al. (2017) recently showed that microtubules undergo a catastrophe event coincident with a dynein-mediated sliding event. They proposed that offloading of dynein-dynactin (the latter of which is a critical regulator of dynein activity) from microtubule plus ends to the cell cortex shifts the balance such that dynactin – which presumably stabilizes microtubules – is depleted from plus ends, while sufficient levels of dynein – which destabilizes microtubules – remain plus end-associated. In addition, previous studies found that She1 plays a role in precluding dynein-dynactin interaction at microtubule plus ends (Markus et al., 2011; Woodruff et al., 2009). Thus, either dynein-mediated microtubule destabilization or dynactin-mediated microtubule stabilization might be enhanced or reduced, respectively, by microtubule-bound She1. Although it is unclear if She1 needs to bind microtubules to affect dynein-dynactin interaction, it is possible that the Tub1G437R-mediated reduction in She1-microtubule binding might also enhance dynein-mediated recruitment of dynactin to plus ends, which would presumably provide a microtubule stabilization effect (due to the increased presence of dynactin). These models are not mutually exclusive, and may in fact both be acting to affect microtubule length during dynein-mediated spindle movement events.
Given the consequences on apparent brain development in the patient with TUBA1AG436R (pachygyria, and severe microcephaly associated with postural delay and poor communication abilities), and the strong link between mutations in dynein activity and motor neuron diseases and developmental brain disorders (Bahi-Buisson et al., 2014; Laquerriere et al., 2017; Marzo et al., 2019; Vissers et al., 2010; Willemsen et al., 2012), our data linking disrupted dynein activity with this mutation are not entirely surprising. For example, dynein activity is critical for various aspects of early neuronal development, in part by promoting interkinetic nuclear migration in neuronal progenitors, and in the subsequent migration of the resulting postmitotic neurons (Del Bene et al., 2008; Hu et al., 2013; Tsai et al., 2010). Moreover, by effecting retrograde transport in neurons throughout their developmental progression, dynein activity is crucial for the maintenance of neuronal health, especially in motor neurons, in which cargoes must be transported over very long distances (≤ 1 m) (Bowman et al., 2000; Fu and Holzbaur, 2013; He et al., 2005; Hendricks et al., 2010; Rao et al., 2017; Shah et al., 2000; Wagner et al., 2004). However, of note, our findings indicate that dynein itself is unaffected by the mutation; rather, dynein activity is indirectly affected by the reduced microtubule binding affinity of a key regulatory MAP, She1. To date, a clear functional homolog of She1 in humans has not been identified. However, there indeed exists a myriad and complex network of MAPs in higher eukaryotes that may play similar roles to She1. For instance, the mammalian tau-related MAP4 protein, which binds in close proximity to G436 (Shigematsu et al., 2018), has been implicated in the control of dynein-mediated spindle orientation during mitosis in mammalian cells (Samora et al., 2011). MAP4 was also shown to physically interact with dynein-dynactin in vivo and to inhibit dynein-mediated microtubule gliding in vitro (Samora et al., 2011). MAP4 has also been shown to shorten dynein-dependent runs of melanosomes in Xenopus melanophores (Semenova et al., 2014). Another potential functional homolog of She1 is MAP9 (also known as ASter-Associated Protein, or ASAP), depletion of which disrupts spindle organization (Saffin et al., 2005; Venoux et al., 2008) and was recently shown to inhibit processive motility of purified dynein-dynactin complexes by specifically precluding microtubule binding by dynactin (Monroy et al., 2020). Thus, it will be important to determine how TUBA1AG436R affects binding of these important neuronal MAPs.
Material and methods
Plasmids, yeast strain growth, and genetic manipulation
Strains used in this study were isogenic to either BY4742 (for Figure 1C and E, Figure 2, and Figure 3; MATα; ura3Δ0 leu2Δ0 his3Δ1 lys2Δ0; provided by euroscarf http://www.euroscarf.de), or YEF473 (for Figure 1D, and Figure 6; ura3-52 lys2-801 leu2-Δ1 his3-Δ200 trp1-Δ63). The TUB1 integrating plasmid, pCR2-TUB1 consists of the region of the TUB1 locus from the intron (situated close the 5’ end of the gene) to 385 bp after the stop codon cloned into the pCR2 vector (Invitrogen). The HIS3 gene expression cassette was ligated into the BsrGI site within the 3’ untranslated region of the TUB1 sequence within pCR2 (pCR2-TUB1). The G437R mutation was subsequently introduced into pCR2-TUB1 by PCR, generating pCR2-tub1G437R. For integration into the native TUB1 locus, pCR2-TUB1 (either wild-type or mutant) was digested with SphI, transformed into yeast using the lithium acetate method, and transformants were selected on media lacking histidine. All transformants were confirmed by PCR and sequencing.
RFP-TUB1 is derived from pAF125 as previously described (Caudron et al. 2008). In addition to this construct, we also generated pHIS3p:mRuby2-tub1G437R+3’UTR::LEU2 to visualize microtubules in mutant cells. To this end, we engineered the G437R point mutation into pHIS3p:mRuby2-TUB1+3’UTR::LEU2 (Markus et al., 2015) using traditional molecular biological methods. For comparison of relative a-tubulin incorporation into mitotic spindles, we used yeast strains with similarly integrated mRuby2-a-tubulins (pHIS3p:mRuby2-TUB1+3’UTR::LEU2, or pHIS3p:mRuby2-tub1G437R+3’UTR::LEU2). To assess relative incorporation of Tub3 into the mitotic spindle, we replaced the TUB1+3’UTR cassette in pHIS3p:mRuby2-TUB1+3’UTR::TRP1 (Markus et al., 2015) with the TUB3 genomic sequence, including 150 bp of the 3’UTR. This plasmid, pHIS3p:mRuby2-TUB3+3’UTR::TRP1, was digested with BbvCI, transformed into yeast using the lithium acetate method, and transformants were selected on media lacking tryptophan. pGFP-Bik1 (Lin et al., 2001) was kindly provided by D. Pellman.
For the two-hybrid experiments, the TUB1 or tub1G437R open reading frames from the respective pCR2 vectors (see above) were cloned into the pLexA vector (Addgene) to produce LexADBD-Tub1 (or LexADBD-Tub1G437R) fusion proteins with a-tubulin and the DNA-binding domain of LexA (LexADBD). Bim1, Kip3, and She1 were amplified from genomic DNA and cloned into the pGADT7 vector (Invitrogen) to produce fusion proteins with the GAL4 activating domain (GAL4AD).
The 6His-tagged-She1Cter expression plasmid for was constructed in pet28 vector (Novagen). The She1 C-terminal part coding for amino-acid 194 to 338 was cloned between the NdeI and XhoI site of the vector downstream of 6His and thrombin site.
To generate a GFP11x7-She1-expressing yeast strain (She1 fused to 7 copies of strand 11 of GFP) (Kamiyama et al., 2016), the GFP11x7 cassette was PCR amplified from pACUH:GFP11×7-mCherry-beta-tubulin (Addgene, plasmid # 70218), and integrated at the 5’ end of the SHE1 open reading frame using the sequential URA3 selection/5-FOA counterselection method. To separately express strands 1 thru 10 of the GFP barrel (GFP1-10, which is required to reconstitute fluorescence), we generated a plasmid with an expression cassette encoding GFP1-10 under the control of the strong TEF1 promoter (TEF1p), as well as the TRP1 selectable marker (pRS304:TEF1p:GFP1-10). We PCR amplified the TEF1p:GFP1-10::TRP1 cassette and integrated it into the lys2-801 locus using homologous recombination into the GFP11×7-She1-expressing yeast strain.
To assess viability on solid media (Figure 1E), serial dilutions of fresh overnight cultures of wild-type or tub1G437R cells (three different haploid clones for each strain) were spotted onto solid YPAD media with or without benomyl, as indicated. Plates were incubated for two days at 30°C.
Image acquisition and analysis
Cell imaging was performed on either a Zeiss Axiovert microscope equipped with a Cool Snap ES CCD camera (Ropper Scientific; for Figures 1 thru 4), or Nikon Ti-E microscope equipped with a 1.49 NA 100X TIRF objective, a Ti-S-E motorized stage, piezo Z-control (Physik Instrumente), an iXon DU888 cooled EM-CCD camera (Andor), a stage-top incubation system (Okolab), and a spinning disc confocal scanner unit (CSUX1; Yokogawa) with an emission filter wheel (ET525/50M for GFP, and ET632/60M for mRuby2; Chroma; Figure 6). For Figures 1 thru 4, 11 Z-planes with 0.3 μm spacing were captured using 2×2 binning (the exposure time varied between experiments). For Figure 6, 11 Z-planes with 0.3 μm spacing were captured.
For microtubule dynamics measurements, maximum intensity projection images of Bik1-GFP-expressing cells were used. Microtubule lengths at each time point were measured manually from maximum intensity projections, and microtubule dynamic parameters were calculated as described (Kosco et al., 2001), using an in-house Visual Basic macro in Excel (Caudron et al., 2008). We measured background-corrected She1, Tub1, or Tub3 spindle-localized fluorescence from maximum intensity projections using ImageJ (NIH). Tub1-corrected She1 intensities were calculated by normalizing the fluorescence intensity values of each (She1, Tub1 and Tub3) to 1 (by dividing the raw mean background corrected values for that measured in wild-type or tub1G437R cells by the wild-type value), and then dividing the resulting normalized She1 values by the normalized Tub1 value for wild-type and tub1G437R, respectively. To additionally correct for Tub3 intensities, these Tub1-corrected values were then divided by the normalized Tub3 values (as calculated above; in which wild-type = 1, and tub1G437R = 1.91).
To assess spindle dynamics parameters, we determined spindle position in cells over time by clicking the center of a preanaphase spindle in each frame, and calculating the displacement between frames using an in-house developed ImageJ macro. The spindle position with respect to the cell boundaries (Figure 3G) was determined for spindles with a pole-to-pole length of 0.8–1.2 μm from the first frames of movies of G2/M cells. The quantification was done blind to the genotype. At least 52 preanaphase spindles from 3 independent clones were measured for each strain, as described (DeZwaan et al., 1997).
Production of recombinant protein
The 6His-She1Cter plasmid was transformed into Rosetta pLysS bacteria. For protein production, bacterial cultures grown in M9 medium were induced by addition of 1 mM IPTG overnight at 18°C. The clarified lysate was separated on a cation exchange column (SP GE healthcare), and bound protein was eluted in cation elution buffer (500 mM NaCl, 40 mM Tris pH 9.2), and then applied to a NiNTA column (Qiagen) after addition of 20 mM imidazole. After elution in NiNTA elution buffer (300 mM NaCl, 20mM Tris pH 7.5, 250mM imidazole), the protein was simultaneously concentrated and buffer-exchanged (with a Vivaspin Turbo 4; Sartorius) into PM buffer (80 mM Pipes pH 6.8, 1mM MgCl2, 0.2% NP40).
Pull-down experiments
For She1Cterm-tubulin pull-down experiments, 2.5 µg of purified tubulin (from cow brain) was incubated with or without 2.5 µg of 6His-She1Cter in 500 µl of PM buffer. After 30 minutes at room temperature, 10 µl of NiNTA beads were added (Thermo Fisher). After a 10 minute incubation, the beads were washed three times with PM buffer, and then boiled for 5 minutes in 60 µl of sample buffer before SDS-PAGE analysis. For peptide competition experiments, 50 µg of each peptide (dissolved in PM buffer; wild-type : EEGEF TEARE DLAAL ERDYI EVGAD SYAEE EEF; G437R: EEGEF TEARE DLAAL ERDYI EVRAD SYAEE EEF) were added to the mixture of She1 (0.25 µg) and tubulin (1.25 µg) and allowed to bind for 30 min at room temperature. Then, 10 µl of NiNTA beads were added (Thermo Fisher), incubated for 10 minutes, washed three times with PM buffer, and then boiled for 5 minutes in 60 µl of sample buffer prior to SDS-PAGE and immunoblot blot analysis. For immunblotting, the gel was transferred to a PVDF membrane (BioRad TransBlot Turbo), blocked with PBS supplemented with 5% milk for 30 minutes, and then probed with anti-α-tubulin (YL1/2 1:10,000) followed by Cy5 anti-rat secondary antibody (Jackson ImmunoResearch, 1:2000). To quantify the tubulin signals, the local background was subtracted and signals were normalized such that tubulin-bound 6His-She1Cter without peptide was equal to 100. The average of two replicates is presented in the graph (Figure 5D).
Structural analyses and mutation modeling
Molecular graphics and analyses were performed with the UCSF Chimera package. The rotamers function was used to model an arginine substitution into position 436 of the α-tubulin structure (pdb 3J6G). The top 4 rotamers were selected for visualization (see Fig. S2). Chimera is developed by the Resource for Biocomputing, Visualization, and Informatics at the University of California, San Francisco (supported by NIGMS P41-GM103311).
Acknowledgements
We would like to thank Jeff Moore for kindly providing a tub3Δ yeast strain. This work was funded by IINCA (TetraTips PLBIO10-030 to A.A. and C.B.) and the NIH/NIGMS (GM118492 to S.M.M.).