Our earlier sequence-profile analysis along with site directed mutagenesis had shown SpoIVA to be a member of the TRAFAC class of GTPases with Era as its closest GTPase relative (Castaing et al., 2013). Since SpoIVA is strictly restricted to the clade of Firmicutes that form endospores we reasoned that SpoIVA was a relatively late innovation that evolved via gene duplication and rapid divergence from Era. Given that Era is a functional GTPase and SpoIVA shows a clear preference for ATP over GTP (Ramamurthi and Losick, 2008) we sought to identify the potential changes in the active site that might have led to this shift in specificity. For relatively closely related proteins that diverged from a common ancestor, reconstruction of an ancestral state may be achieved based on phylogeny and using statistical methods across the entire length of the proteins (Hochberg and Thornton, 2017). However, the phyletic patterns strongly indicate that the ultimate ancestor of SpoIVA was Era itself. Moreover, SpoIVA and Era have diverged too far from each other to successfully employ such an approach. Beyond myriad amino acid substitutions and insertions, SpoIVA has even acquired two C-terminal domain fusions that are not present in Era (Figure 1C; Castaing et al., 2014), which is reminiscent of other examples where the acquisition of large appendages to ancestral proteins have generated novel functions beyond the ancestral function (Escudero et al., 2020; Farr et al., 2017). Thus, the number of variables involved when all residues are considered would result in too vast of a parameter space to analyze using a common ancestor reconstruction method. We therefore focused on the highly conserved N-terminal TRAFAC NTPase domain of SpoIVA and computed a sequence logo for SpoIVA, Era and various other families of the TRAFAC class of GTPases, especially those of the GIMAP-Septin-Dynamin clade which show a comparable tendency for forming oligomers or polymers (Figure 1A). Next, the amino acid conservation pattern of SpoIVA was superimposed on a topology diagram of the ancestral core TRAFAC GTPase domain (Figure 1B). This allowed us to identify those conserved active site positions which were retained in the ancestral state in SpoIVA and those that were altered with respect to bona fide GTPases (Figure 1C).

The first three motifs (G1–3) respectively correspond to: the Walker A motif which binds the triphosphate of the NTP substrate; the sensor T which discriminates the GTP-bound state from the GDP-bound state; and the Walker B motif which chelates the catalytic Mg²⁺ and senses the bound triphosphate along with the sensor T (G2) motif (Figure 1A,B). In SpoIVA, these three motifs are retained in the ancestral state indicating that SpoIVA binds and senses the triphosphate moiety of the NTP similar to the ancestral GTPases (Castaing et al., 2013; Figure 1B–E).

In contrast, notable changes are seen in the G4 and G5 motifs of SpoIVA. Of these, G4 is comprised of the final residue of strand 6 of the core GTPase domain and a characteristic single-turn helix that follows it (Figure 1B,C). Among the GTPases closely related to SpoIVA, the G4 motif is of the form NKxD (where ‘x’ is any amino acid), for example, in Era and Eng (Figure 1A). The first position of this motif is typically either N or T in most families of the entire GTPase superfamily. Thus, SpoIVA retains the ancestral state in this position. This residue forms the ‘lower wall’ of the base-binding pocket of the active site and by itself does not appear to discriminate between the purines (Figure 1D,E). The next position is a K in most families of bona fide GTPases (Figure 1A) and the extended sidechain of this lysine forms the ‘lateral wall’ of the base-binding pocket (Figure 1D). Strikingly, this K is consistently substituted by an alcoholic (S/T) residue in the SpoIVA family. The next conserved position in G4 is the D, which is the most important determinant of guanine specificity (Figure 1D). In SpoIVA it is again mostly substituted by either of several polar residues, such as K, H, N, or R. Notably, unlike in bona fide GTPases this position is poorly constrained in SpoIVA, suggesting a relaxation of selection, which might have allowed the emergence of ATP selectivity.

The G5 motif follows immediately after strand-7 of the core GTPase domain and typically displays the motif SAx in classical TRAFAC GTPases. This region forms the wall of the base-binding pocket opposite to that formed by the conserved lysine in G4 (Figure 1D). In the case of SpoIVA the position corresponding to the conserved S in the G5 motif is less constrained and is usually either N, D, or S. The next residue is usually a cysteine in SpoIVA. Hence, this residue is likely to be similar to A with respect to its hydrophobicity and is not situated close to the distinguishing atoms of the purine base of the bound nucleotide. Thus, it is unlikely to have a major effect on base selectivity. Two residues downstream of S, there is a position that contributes to the wall of the base-binding pocket. This position is not particularly conserved in TRAFAC GTPases as a whole but in SpoIVA is either acidic (D/E) or Q/N in 41% of the orthologs. Together, these observations suggested that the changes in the G4 (NKxD) and G5 (SxE) regions relative to the bona fide GTPases may have contributed to the emergence of ATP-specificity in SpoIVA (Figure 1E).

To determine the relative contributions of the altered NKxD and SxE motifs in G4 and G5, respectively, on nucleotide hydrolysis specificity of SpoIVA, we first substituted different residues in each motif, either individually or in several combinations. Next, we overproduced and purified the variants from E. coli and tested the efficiency of each variant in hydrolyzing ATP and GTP in vitro. As a control, we compared these activities to that of B. subtilis Era GTPase that we purified using a similar protocol. We measured nucleotide hydrolysis for each variant at increasing nucleotide concentrations to produce saturation curves that revealed the substrate turnover rate (k_cat) and nucleotide concentration that produced half-maximal enzymatic activity (K_m) (Figure 2—figure supplement 1, Supplementary file 2). We then calculated the catalytic efficiency for each reaction (k_cat/K_m) which reflects how likely the forward reaction (hydrolysis of the bound nucleotide) will proceed (Figure 2A, Supplementary file 2). Wild-type (WT) SpoIVA displayed a catalytic efficiency of 4.2 ± 0.9 min⁻¹ mM⁻¹ for ATP, compared to just 1.3 ± 0.8 min⁻¹ mM⁻¹ for GTP, indicating that the protein hydrolyzed ATP approximately threefold more efficiently than GTP (Figure 2B). By comparison, Era did not display appreciable basal ATPase or GTPase activity, but upon incubation with an RNA oligonucleotide corresponding to the 16S rRNA sequence to which Era binds and which reportedly stimulates the enzymatic activity of Era (Meier et al., 2000; Tu et al., 2011), Era hydrolyzed GTP with a catalytic efficiency of 0.7 ± 0.2 min⁻¹ mM⁻¹, similar to the reported activity of E. coli Era (Tu et al., 2011). However, Era did not specifically hydrolyze ATP, as evidenced by the failure of the reaction to reach saturation and display Michaelis–Menten kinetics (Figure 2—figure supplement 1J, Figure 2A, ‘I.D.’ for ‘indeterminable’). Restoring the Asp in the degenerate NKxD motif of SpoIVA did not significantly change the catalytic efficiencies of ATP or GTP hydrolysis but restoring either the Lys or full NKxD motif in SpoIVA resulted in a ~2.5-fold increase in the catalytic efficiency for ATP and ~3.5-fold of that for GTP hydrolysis. Restoring either the single Lys or the full NKxD motif resulted in decreased preference for ATP (Figure 2B). Curiously, these variants also displayed an unusually high turnover rate that was ~10-fold higher for both NTPs than that displayed by WT SpoIVA (Figure 2—figure supplement 1C,D).

We next investigated if altering the SxE sequence in SpoIVA would reduce the unusually high enzymatic activity resulting from restoration of the NKxD motif. Substituting both the Ser and Glu with Ala (to disrupt both positions with an amino acid with a short sidechain that is unlikely to perturb overall SpoIVA structure) in the context of the restored NKxD motif resulted in lowered catalytic efficiencies, similar to WT SpoIVA (Figure 2A,C: ‘NKxD AxA’). This change also increased the catalytic efficiency of the enzyme for GTP, resulting in drastically reduced specificity for ATP relative to WT SpoIVA (Figure 2B). Changing the SxE sequence alone to AxA had a similar effect (Figure 2A, ‘NSxR AxA’). Substituting the Glu alone in the SxE sequence with Ala (‘NSxR SxA’) mimicked the NSxR AxA variant with respect to nucleotide specificity (Figure 2B), but further lowered the catalytic efficiency of the enzyme (Figure 2A). Finally, combining a restoration of just the Asp residue of the degenerate NKxD motif with disruption of just the Glu of the SxE sequence, resulted in an enzyme that displayed a similar catalytic efficiency to that of Era (Figure 2A, ‘NSxD SxA’; hereafter referred to as ‘SpoIVA^GTPase’; Figure 2C) with a slight preference for GTP over ATP (Figure 2B).

To test if altering the G4 NKxD motif and introducing the SxE sequence in G5 were sufficient for the emergence of preferential ATPase activity, we changed the NKxD motif in Era to NSxR, to resemble the G4 sequence in SpoIVA, and substituted a glutamate at the end of the SAx sequence in Era to produce ‘SAE’ (thereby introducing an SxE motif) and tested the nucleotide hydrolysis activity of the evolved variant. Similar to WT Era, the evolved Era (Era^ATPase) did not exhibit a basal NTPase activity, but upon stimulation with the 16S rRNA fragment, Era^ATPase hydrolyzed ATP with a catalytic efficiency of 0.2 ± 0.1 min⁻¹ mM⁻¹ (Figure 2A), but more curiously failed to specifically hydrolyze GTP (Figure 2—figure supplement 1K).

The key alterations in shifting the NTPase activity of SpoIVA toward that of Era were to restore the Asp of the NKxD and replace the Glu of the SxE (Figure 2C). Conversely, disrupting the NKxD motif of Era and introducing a Glu to create an SxE motif were sufficient to drive the preferential hydrolysis of ATP over GTP. The mutational analyses therefore indicate that the Asp of the NKxD motif contributes to nucleotide specificity and that the substitution of Asp to Arg seen in the extant B. subtilis SpoIVA contributes to altering the nucleotide-binding pocket to discriminate against GTP in favor of ATP. This is consistent with reported crystal structures of TRAFAC GTPases with a bound GTP (Knihtila et al., 2015) that show that the Asp of the NKxD motif can interact with the base of the bound nucleotide. In addition, another report showed that changing the Asp to another residue can alter the nucleotide-binding preference of Ras GTPase from GTP to xanthine triphosphate (XTP) (Kang et al., 1994; Weijland et al., 1994; Zhong et al., 1995). The Glu in the SxE sequence further contributes to ATP hydrolysis specificity and likely stabilizes the binding of ATP over GTP.

To ensure that the amino acid substitutions introduced to restore ancestral SpoIVA GTPase activity did not completely abrogate protein function, we tested the ability of the different variants to complement the sporulation defect caused by a deletion of the spoIVA gene in B. subtilis. Deletion of spoIVA resulted in a >10⁸-fold and a ~10⁶-fold reduction in the production of heat resistant spores and lysozyme-resistant spores, respectively (Roels et al., 1992, Figures 3A and Figure 3—figure supplement 1A, Supplementary file 2), which could be complemented in trans by the introduction of WT spoIVA at an ectopic chromosomal locus. In contrast, while expression of the NSxD variant (which displayed similar enzymatic activity as WT SpoIVA in vitro; Figures 2A and Figure 2—figure supplement 1B, Supplementary file 2) complemented the spoIVA deletion, complementation by the hyperactive NKxR or NKxD variants resulted in >10⁸-fold and ~10⁴-fold decreases in the production of heat-resistant spores, respectively. Similarly, mutants that harbored a spoIVA allele containing a full substitution of the SxE sequence (resulting in AxA) also largely failed to sporulate when the G4 motif also harbored alterations (Figures 3A and, Supplementary file 2; NKxD AxA and NKxR AxA). Immunoblot analysis of extracts prepared from sporulating cells revealed that these SpoIVA variants were produced at levels similar to WT SpoIVA (Figure 3—figure supplement 1B). In an otherwise WT background, though, the AxA substitution (NSxR AxA) sporulated at near WT levels. Interestingly, SpoIVA^GTPase (NSxD SxA), which showed reduced but similar hydrolysis of ATP and GTP (Figure 2), supported sporulation at near WT levels (Figures 3A and Figure 3—figure supplement 1A, Supplementary file 2). Although we cannot determine which nucleotide SpoIVA^GTPase utilized in vivo, we can conclude that this disruption of the nucleotide-binding pocket did not result in either a large-scale structural defect in the protein that catastrophically affected its function or reduced its accumulation in vivo.

We next examined the subcellular localization in vivo during sporulation of each variant fused to green fluorescent protein, expressed from an ectopic chromosomal locus under control of the native spoIVA promoter. WT GFP-SpoIVA localized to the surface of the forespore (Figure 3B,B’), as did the NSxD variant which sporulated at near-WT levels (Figure 3C,C’). However, restoring only the Lys of the degenerate G4 motif or restoring the entire NKxD motif in an otherwise WT SpoIVA resulted in the mis-localization of the variant as a focus near the surface of the forespore. Ala substitution of the SxE sequence along with the NKxD motif resulted in a similar mis-localization pattern (Figure 3D–F,D’–F’). In contrast, various disruptions to the SxE sequence alone did not abrogate localization of the variant (Figure 3G–I,G’–I’). Finally, SpoIVA^GTPase localized similar to WT (Figure 3J–J’), consistent with its ability to support sporulation at a near-WT level. Thus, disruption of the nucleotide-binding pocket of SpoIVA to permit the slightly preferential hydrolysis of GTP over ATP resulted in a protein that largely retained proper function in vivo.

Although the sporulation efficiency and subcellular localization data indicated that SpoIVA^GTPase was largely functional, it was difficult to infer if this variant used ATP or GTP to perform its function in vivo. We therefore monitored the nucleotide hydrolysis-driven conformational change and polymerization of SpoIVA in vitro in the presence of either nucleotide.

Structural changes in SpoIVA may be monitored by limited trypsin proteolysis (Castaing et al., 2013). We therefore incubated purified WT SpoIVA or variants with a low concentration of trypsin and assessed the extent of proteolysis at different time points by separating the reaction by Coomassie-stained SDS-PAGE. Importantly, the experiment was performed using 2 µM SpoIVA, which is below the critical concentration for SpoIVA polymerization (Figure 5—figure supplement 1A; Castaing et al., 2013), and therefore reflective of polymerization-independent conformational changes in the protein. In the absence of nucleotide, SpoIVA was rapidly degraded, resulting in a characteristic banding pattern (Figure 4A). In contrast, co-incubation with either ATP or GTP resulted in a different banding pattern: most noticeably, the full-length protein was considerably resistant to degradation even after 10 min, suggestive of a massive conformational change in the protein upon hydrolysis of either nucleotide. The disappearance of full-length SpoIVA was quantified to produce a decay rate which indicated that a conformational change in WT SpoIVA could be achieved at an approximately similar rate by hydrolysis of either by ATP or GTP (Figure 4B). In contrast, restoring the Lys alone, the entire NKxD motif alone, or in combination with the entirely disrupted SxE sequence (to AxA) resulted in a rapid decay rate in the presence of either ATP or GTP, indicating that these variants were unable to achieve the characteristic nucleotide-dependent conformational change, consistent with the observed in vivo defects of these variants (Figure 3, Supplementary file 2). However, variants harboring substitution of either the Ser or Glu singly or together in the SxE sequence with Ala or SpoIVA^GTPase displayed a decay rate in the presence of either nucleotide that was more similar to WT SpoIVA, suggesting that that these variants were able to utilize the energy released from hydrolysis of either ATP or GTP to drive the conformational change in the protein that is a prerequisite for polymerization (Castaing et al., 2013).

To further understand the nucleotide hydrolysis requirement for driving the conformational change in SpoIVA, we employed the limited trypsin digestion assay with SpoIVA and key SpoIVA variants using different nucleotides: ATP-γ-S and GTP-γ-S, which are non-hydrolyzable analogs of ATP and GTP; ADP and GDP; and ADP-AlF_x and GDP-AlF_x, which are nucleotide analogs that mimic the transition state of ATP and GTP in the hydrolysis reaction (Chen et al., 2007; Coleman and Sprang, 1999). In the presence of ATP-γ-S or GTP-γ-S, WT SpoIVA displayed an intermediate conformational change suggesting that the protein bound, but did not hydrolyze, the nucleotide (Figure 2—figure supplement 1L, M), whereas SpoIVA harboring a Walker A disruption (SpoIVA^A*, which prevents ATP binding [Ramamurthi and Losick, 2008]) did not undergo a similar conformational change (Figures 4C,D and Figure 4—figure supplement 1A-B). Consistent with this result, the SpoIVA variant harboring a Sensor T disruption (SpoIVA^T*, which binds, but does not hydrolyze ATP [Castaing et al., 2013]) displayed a similar conformational change as WT SpoIVA in the presence of ATP-γ-S and GTP-γ-S (Figures 4D and Figure 4—figure supplement 1C), and SpoIVA^T* binding to ATP. Together, this suggested that this intermediate conformational change is likely due to nucleotide binding and not due to slow hydrolysis of the bound nucleotide. Interestingly, the NKxD variant of SpoIVA, which exhibited elevated ATP and GTP hydrolysis levels (Figure 2A), displayed a conformational change in the presence of ATP-γ-S and GTP-γ-S similar to SpoIVA^A* (Figures 4D and Figure 4—figure supplement 1D), suggesting that it did not stably undergo the initial conformational change that occurs upon binding the nucleotide. WT SpoIVA and SpoIVA^T* exhibited a similar intermediate conformational change when incubated with ADP or GDP, but not the full conformational change that occurred upon nucleotide hydrolysis (Figure 4E). A similar intermediate conformational change upon binding GDP relative to binding GTP has been reported for other TRAFAC GTPases (Coleman and Sprang, 1999). Similar to the non-hydrolyzable nucleotides, SpoIVA^T* and the NKxD variant did not display any conformational change when incubated with ADP or GDP (Figure 4E). Curiously, incubation of WT SpoIVA with either ADP-AlF_x or GDP-AlF_x (Figure 4F) produced a conformational change that was similar to that produced upon full hydrolysis of the nucleotide (Figure 4B,C). This suggested that the transition state mimics the activated state facilitated by ATP hydrolysis that is responsible for the conformational change seen in Figure 4A. As controls, incubation of the SpoIVA^T*, which is devoid of the alcoholic residue found to be critical for stabilizing the transition state of the protein-nucleotide complex in TRAFAC GTPases (Coleman and Sprang, 1999; Leipe et al., 2002), SpoIVA^A*, or the NKxD variant with ADP-AlF_x or GDP-AlF_x did not exhibit a similar conformational change (Figure 4F). Taken together, the results suggest that simply binding to ADP or GDP is insufficient to produce the full conformational change in SpoIVA required for polymerization. Instead, hydrolysis of an NTP molecule, while bound to the active site, is required for rearranging SpoIVA into a polymerization-competent state. The data are also consistent with a model in which rapid turnover of the bound NTP is incompatible with producing such a conformational change.

Next, we tested the nucleotide-dependent polymerization rates of the SpoIVA variants by measuring the size distribution of polymerized SpoIVA molecules over time using dynamic light scattering (DLS). Incubation of purified WT SpoIVA, above the critical concentration for polymerization, with ATP, but not GTP, resulted in a steady increase in hydrodynamic radius (Rh) over a 5 hr period, consistent with ATP-dependent polymerization and what we previously observed (Castaing et al., 2013; Figure 5—figure supplement 1A). The initial slope of the polymerization reaction in the presence of nucleotide was quantified and reported relative to the initial slope of the reaction in the absence of nucleotide to yield polymerization rates for SpoIVA (Figure 5A, Supplementary file 2). This ratio revealed a greater than ~30% increase in polymerization rate with ATP than GTP (Figure 5A, Supplementary file 2), surprisingly indicating that the conformational change in SpoIVA driven by GTP hydrolysis (Figure 4A,B,E), which appeared similar to the conformational change driven by ATP hydrolysis, did not yield isomers of SpoIVA that were capable of polymerization.

We next examined the ability of SpoIVA variants in polymerizing with ATP and GTP. Restoring the Asp in the G4 motif, which did not display any obvious defect in the other assays, only slightly lowered the polymerization rate with ATP (Figures 5A and Figure 5—figure supplement 1B; Supplementary file 2). However, all variants that harbored a restoration of the Lys in the G4 motif were unable to polymerize with either nucleotide (Figures 5A and Figure 5—figure supplement 1C-E; Supplementary file 2), suggesting that substitutions leading to elevated nucleotide hydrolysis were unable to (1) induce a conformational change, (2) were defective in vivo, and (3) were also unable to promote SpoIVA polymerization. Disruptions to the SxE sequence lowered, but did not abolish, SpoIVA polymerization in the presence of ATP, but none of these variants polymerized in the presence of GTP (Figure 5A, Figure 5—figure supplement 1F-H; Supplementary file 2). Likewise, SpoIVA^GTPase, which was functional in vivo (Figure 3), and could use either ATP or GTP to induce a similar conformational change after nucleotide hydrolysis (Figure 4A,B), only polymerized in the presence of ATP (Figures 5A and Figure 5—figure supplement 1I; Supplementary file 2), suggesting a specific requirement for the nucleotide base, not simply the energy released from nucleotide hydrolysis, for its function.

Reported crystal structures of the eukaryotic septin GTPases, which also belong to the TRAFAC class of P-loop GTPases, show a dimer in which each monomer binds to a molecule of GDP that is stabilized by contacts from the other monomer with the guanosine base (Zeraik et al., 2014). Fully polymerized SpoIVA is devoid of any bound nucleotide (Castaing et al., 2013), but we wondered if SpoIVA would form an intermediate multimeric complex en route to polymerization whose formation would be dependent specifically on ADP binding before the hydrolyzed nucleotide was released. To test this, we first incubated 2 µM purified SpoIVA, below the threshold concentration for polymerization (Figure 5—figure supplement 1A), either in the absence of nucleotide or in the presence of ATP or GTP, and then separated the products by size exclusion chromatography (SEC). In the absence of nucleotide or the presence of GTP, SpoIVA migrated as a single peak (Figure 5B, top left; pink and gray traces). Quantitative mass imaging of this peak by employing interferometric scattering mass spectrometry (iSCAMS) (Young et al., 2018) revealed a mass of 103 ± 12 kDa (Figure 5—figure supplement 2A), similar to a predicted mass for a SpoIVA dimer of 114 kDa. Diluting the peak into a low salt buffer revealed an additional peak at 59 ± 8 kDa, similar to the predicted mass for a SpoIVA monomer of 57 kDa, along with the presumed dimeric peak of 109 ± 14 kDa corresponding to a SpoIVA dimer (Figure 5—figure supplement 2B). Addition of ATP resulted in shifting most of the protein to the void volume of the SEC column, indicating the formation of a larger complex (Figure 5B, top left; blue trace), even though the experiment was performed using a SpoIVA concentration that was below its critical concentration for polymerization (Figure 5—figure supplement 1A). In contrast, neither SpoIVA^T* in the presence ATP or GTP, nor WT SpoIVA in the presence of either ATP-γ-S or ADP, formed a larger complex, indicating that ATP, but not GTP, hydrolysis is required for producing the species in the void volume (Figure 5B, top right and bottom left). Interestingly, incubation of SpoIVA^GTPase with ATP, but not GTP, partially shifted a population into the void volume (Figure 5B, bottom right), indicating that the reengineered SpoIVA^GTPase retained its specific dependence on ATP instead of GTP in forming the polymerization-competent intermediate, despite being able to hydrolyze both NTPs. Since the mass of the void volume peak was varied and too large to examine using iSCAMS, we employed size exclusion with multi-angle light scattering (SEC-MALS) analysis (Some et al., 2019), which revealed a range of molecular weights ranging from 2 × 10³ kDa to 10⁶ kDa, suggesting multimers containing at least ~36 monomers of SpoIVA (Figure 5—figure supplement 2C). Examination of this void volume by negative stain and transmission electron microscopy revealed a species with a distinct structure that extensively self-interacted (Figure 5D). To check if this larger SpoIVA species may harbor an associated nucleotide, we examined the same peaks that eluted from the SEC column using 254 nm wavelength. Incubating WT SpoIVA or SpoIVA^GTPase with GTP did not reveal a significant absorbance at 254 nm for fractions containing protein (Figure 5C, top left and bottom right; pink traces), but the protein peaks of the sample in the void volume when incubated with ATP displayed significant absorbance at 254 nm (Figure 5C, blue traces), suggesting the presence of nucleotide in this fraction. Neither SpoIVA^T* incubated with ATP nor WT SpoIVA incubated with ATP-γ-S or ADP displayed significant absorbance at 254 nm (Figure 5C, top right and bottom left), suggesting that the nucleotide present in the void volume multimer of WT SpoIVA is likely ADP retained in the active site after hydrolysis. To confirm this, we extracted the nucleotide from the void volume fraction by denaturing the protein. Separation of the extracted material by SEC revealed that it eluted at a similar volume as ADP, not ATP (Figure 5—figure supplement 3).

We next tested if the SpoIVA-ADP complex in the void volume observed in Figure 5B is a functional intermediate that can subsequently polymerize once its concentration exceeds the threshold concentration for polymerization. We therefore first incubated purified SpoIVA at low concentration (2 µM, below the threshold concentration for polymerization; Figure 5—figure supplement 1A) in the presence and absence of ATP. Half of each sample was then subjected to buffer exchange by SEC to remove free nucleotide. The desalted protein was then concentrated 20-fold using pressure dialysis after which polymerization was assayed by the formation of insoluble SpoIVA in the pellet fraction after ultracentrifugation (Figure 5E). In parallel, select supernatant and pellet fractions were separated by SEC to detect ATP and ADP (Figure 5F–J). Only samples that were incubated with ATP and whose concentration was increased displayed appreciable SpoIVA in the pellet fraction (Figure 5E). Importantly, removing free ATP prior to concentrating the sample also resulted in SpoIVA polymerization, suggesting that pre-incubation with ATP produces a functional ADP-bound SpoIVA multimeric intermediate that can polymerize once its threshold concentration for polymerization is subsequently achieved. Interestingly, while the protein-containing pellet fraction did not contain any bound nucleotide (Figure 5J; Castaing et al., 2013), the supernatant fraction contained ATP and ADP (Figure 5I), likely from ATP-bound SpoIVA that had not polymerized and consistent with the release of ADP upon SpoIVA polymerization.

Taken together, the results are consistent with a model in which SpoIVA, at a low concentration that does not promote polymerization, hydrolyzes ATP to undergo a conformational change and subsequently assembles into heterogeneous high molecular weight multimers that retain the ADP product of hydrolysis. Upon an increase in concentration, this functional SpoIVA multimeric intermediate releases the bound ADP and forms a nucleotide-free mature polymer. In contrast, while hydrolysis of GTP drove a conformational change in SpoIVA similar to what was achieved with ATP hydrolysis (Figure 4A,B), GTP hydrolysis did not permit formation of the high molecular weight intermediate (and therefore did not permit polymerization), nor did the protein retain GDP after hydrolysis (Figure 5B,C).

As sporulation proceeds, intracellular ATP levels 2 hr after the induction of sporulation were reported to reach a high of ~1.5 mM while the level of GTP drops to a low of <0.06 mM (Lopez et al., 1981; Lopez et al., 1979; Ochi et al., 1982; Ochi et al., 1981). This relative abundance of ATP could therefore explain the evolutionary pressure that drove the switch in nucleotide-binding preference from GTP to ATP in SpoIVA. However, the time point at which these measurements were performed is before SpoIVA exerts its function during sporulation. We therefore harvested sporulating cells at various time points by vacuum filtration, extracted total nucleotides using organic solvent, and employed liquid chromatography-mass spectrometry (LC-MS) to quantify the relative abundance of individual nucleotides (Figure 6A–D). Immediately after induction of sporulation, ATP levels remained relatively constant (Figure 6A; blue trace, compare ‘pre-induction’ to 0 hr; Supplementary file 3), but increased almost twofold by the first hour, before returning to pre-sporulation levels in the second hour. At t = 3.5 hr, when SpoIVA is actively assembling the spore coat basement layer (Peluso et al., 2019; Price and Losick, 1999), ATP levels were ~70% of pre-sporulation levels; by t = 5 hr, ATP levels were less than 35% of pre-sporulation levels. Assuming a B. subtilis cell volume of 2.38 fL and using calculated LC-MS detection efficiencies for ATP and GTP (Fung et al., 2020), this corresponds to a pre-induction intracellular concentration of ATP of 2.3 mM ±0.89 mM in casein hydrolysate media; the concentration of ATP 3.5 hr after the induction of sporulation corresponds to 1.6 mM ± 0.25 mM, and 0.81 mM ± 0.084 mM at t = 5 hr, after achieving a concentration of 4.5 mM ± 0.044 mM at t = 1 hr. In contrast, GTP, which was present initially at 0.65 mM ± 0.33 mM plummeted to 0.071 mM ±0.034 mM immediately upon induction of sporulation (Figure 6A, pink trace; Supplementary file 3). Over the next 3.5 hr, GTP levels rose approximately threefold, and ended up twofold higher than at t = 0 after 5 hr. This corresponded to an approximately eightfold excess of ATP over GTP at t = 3.5 hr (intracellular concentration of 0.20 mM ±0.034 mM GTP), and an approximately fivefold excess of ATP at t = 5 hr (intracellular concentration of 0.16 mM ±0.012 GTP). The relative amounts of CTP and UTP were lower than that of ATP during the first 5 hr of sporulation (Figure 6A), but slightly higher than that of GTP. Consistent with the increase in ATP level in the first hour of sporulation, the levels of ADP and AMP increased approximately twofold immediately upon induction of sporulation (Figure 6B,C, Supplementary file 3). Curiously, not only did the nucleotide alarmone ppGpp increase ~30-fold immediately upon induction of sporulation (Figure 6D, orange trace; Supplementary file 3) to 0.11 mM ± 0.088, similar to what was previously reported (Ochi et al., 1982), but the newly discovered guanosine nucleotide alarmone pGpp, which is produced by hydrolysis of (p)ppGpp (Yang et al., 2020), also increased ~30-fold immediately (0.19 mM ± 0.10 mM at t = 0 hr) after sporulation was induced.

Since SpoIVA^GTPase functioned similar to the extant (WT) SpoIVA in vivo with respect to sporulation efficiency and localization (Figures 3 and Figure 3—figure supplement 1A; Supplementary file 2), we wondered if the extant SpoIVA evolved to more efficiently utilize ATP at physiological nucleotide concentrations, in a way that was not evident by measuring sporulation efficiency by heat and lysozyme resistance (Figures 3A and Figure 3—figure supplement 1A). Both SpoIVA and SpoIVA^GTPase failed to undergo nucleotide hydrolysis-mediated structural changes in the presence of 0.06 mM GTP at sub-polymerization levels of the protein but did so in the presence of 1.5 mM ATP (Figure 6E,F). However, when we examined SpoIVA polymerization at different concentrations of ATP, we observed that while neither SpoIVA nor SpoIVA^GTPase were able to polymerize in the presence of 0.06 mM ATP (Figure 6G), WT SpoIVA, but not SpoIVA^GTPase, was able to efficiently polymerize in the presence of an intermediate concentration of ATP (1.5 mM; Figure 6G), similar to what was observed for WT SpoIVA using 4 mM ATP. Thus, although SpoIVA^GTPase promiscuously hydrolyzed ATP and GTP (Figure 2A, Figure 2—figure supplement 1J) and could polymerize with excessive ATP (albeit at a slower rate; Figures 5A and 6G), it was unable to polymerize under limiting concentration of ATP (Figure 6G). The limiting amounts of ATP (1.5 mM, the approximate amount of intracellular ATP between 3.5 hr and 5 hr of sporulation) therefore suggests a selective pressure that could have driven the initial amino acid substitutions required to switch nucleotide preference from GTP to ATP in SpoIVA and subsequent substitutions to enhance polymerization activity.

Starting sets of members of each GTPase family were collected by running BLASTP searches (Altschul et al., 1997; Aravind and Koonin, 1999) against a database of 4440 complete genomes and 2983 metagenomes (coding for a total of 21,646,808 proteins) obtained from the genomes division of Genbank (ftp://ftp.ncbi.nlm.nih.gov/genomes/). These were then filtered by similarity-based clustering with the BLASTCLUST program (https://www.ncbi.nlm.nih.gov/Web/Newsltr/Spring04/blastlab.html) to obtain the representative sets for each family. They were then aligned using the Kalign program (Lassmann et al., 2009; Lassmann and Sonnhammer, 2005) and further improved by examining GTPase structures. The multiple sequence alignments thus obtained were used to compute the sequence logos for each family. The logos were computed using the codebase obtained from RWeblogo (https://CRAN.R-project.org/package=RWebLogo) with the residue size scaled as per the probability of their occurrence in a column of the alignment. The NTPase active site pockets were drawn using the MarvinSketch program (https://chemaxon.com/products/marvin).

His₆-tagged SpoIVA was overproduced in E. coli BL21(DE3) from plasmid pJP120 (WT SpoIVA) (Castaing et al., 2013) or pJP120 derivatives (SpoIVA variants constructed via QuikChange kit, Agilent) and purified using Ni²⁺ affinity chromatography (Qiagen) and subsequently by ion-exchange chromatography (Mono Q; Pharmacia) as described previously (Wu et al., 2015). Briefly, 4 × 500 ml cultures of BL21(DE3) harboring pJP120 or derivatives [pIL48 (spoIVA^S189K), pIL49 (spoIVA^R191D), pIL50 (spoIVA^S189K,R191D), pJH25 (spoIVA^{S189K,R191D,E218A}), pJH26 (spoIVA^{S189K,R191D,S216A}), pJH27 (spoIVA^{S189K,R191D,S216A,E218A}), pTU141 (spoIVA^S216A,E218A), pTU142 (spoIVA^S216A), pTU143 (spoIVA^E218A), and pTU220 (spoIVA^R191D,E218A)] were grown at 37°C in Terrific Broth (Fisher Scientific) containing 50 mg/ml kanamycin for plasmid maintenance to mid-logarithmic phase (~2.5 hr). Isopropyl-β-D-thiogalactopyranoside (Calbiochem, Millipore) was added to 1 mM final concentration to induce protein production and each culture was grown for 4 hr at 37°C. Harvested cells (which could be stored at −80°C) were resuspended in 25 ml of ice-cold Buffer A (50 mM Tris at pH 7.5, 150 mM NaCl) and disrupted by French Pressure Cell Press (SLM Aminco) at 12,000 psi. All subsequent steps were performed on ice. Unbroken cells and cell debris were removed by centrifugation at 35,000 rpm for 1 hr at 4°C and the cleared lysate was loaded on a single gravity column containing 3 ml of Ni-NTA agarose (QIAGEN), pre-equilibrated with ice-cold Buffer A, and incubated on ice for 30 min. Upon flow-through of the clarified lysate, the column was washed with 50 ml Wash Buffer I (Buffer A containing 20 mM imidazole), followed by 4 ml Wash Buffer II (Buffer A containing 80 mM imidazole). Protein was eluted with 10 ml ice-cold Elution buffer (Buffer A containing 250 mM imidazole). Imidazole was removed from eluted fractions using a PD-10 desalting column (GE Healthcare; 3.3 ml eluate/PD-10 desalting column) and eluted using 4 ml Buffer A. Peak fractions were identified using NanoDrop A₂₈₀ (ND-1000, Thermo Scientific), pooled, separated by ion exchange chromatography (Mono Q 5/50, GE Healthcare), and then eluted with a step-wise gradient of 150–1000 mM NaCl; His₆-SpoIVA routinely eluted at 0.4 M NaCl. Purified protein was stored at 4°C and was used in less than 48 hr after purification due to precipitation of the protein upon prolonged storage. For long-term storage, samples were flash-frozen on dry ice and stored at −80°C. To assess the multimerization of SpoIVA and variants, 2 µM purified WT or T* SpoIVA variant (Castaing et al., 2013) was incubated with no NTP, 4 mM ATP, or 4 mM GTP in Buffer B (50 mM Tris at pH 7.5, 400 mM NaCl, 5 mM MgCl₂) for 4 hr at 37°C. 700 µl of each sample was then passed through a 0.22 µm filter and separated using a Superose 6 Increase 10/300 GL size exclusion column (GE Healthcare) with Buffer B at a flow rate of 0.2 ml/min. For assessing the presence of ATP and ADP, where indicated, samples were heated at 95°C for 20 min to release protein bound nucleotide, centrifuged to remove insoluble material, and the supernatant separated using Superdex 30 Increase 10/30 GL size exclusion column (GE Healthcare) with 50 mM Tris at pH 7.5 at a flow rate of 0.2 ml/min. Retention volumes were compared to that of free ATP and ADP standards (Sigma). his₆-tagged era gene was PCR-amplified using primers (5’-GGGGAATTGTGAGCGGATAACAATTC which abutted an XbaI restriction site, and 5’-GCTTGTCGACGGAGCTCGAATTCGGATCTTAATATTCGTCCTCTTTAAAGCCAAAATC which abutted a BamHI restriction site and six histidine codons) from B. subtilis PY79 chromosomal DNA and cloned into vector pET28a strain to generated plasmid pJH17C. His₆-tagged Era was overproduced in E. coli BL21(DE3) from plasmid pJH17C or pTU281 (harboring His₆-tagged Era^{K125S, D127R, L156E} variant constructed via QuikChange site-directed mutagenesis kit, Agilent) and purified using Ni²⁺ affinity chromatography (Qiagen) and subsequently by ion-exchange chromatography similar to purification of SpoIVA. However, since Era is positively charged at neutral pH, imidazole elutions were further separated by ion exchange chromatography using Mono S column (Mono S 5/50, GE Healthcare) instead of Mono Q. Purified protein was either stored at −80°C, after quick freeze on dry-ice, for long-term storage, or stored at 4°C and used within 48 hr.

Different concentrations (ranging from 0 to 4 mM) of ATP or GTP (Sigma) were incubated with 0.3 μM purified His₆-SpoIVA or SpoIVA variants, or His₆-Era or Era variants in 50 μl Buffer B for 1 hr at 37°C. For reactions with Era and Era variants, 1 mM of the oligonucleotide rAUCACCUCCUUUCUA (corresponding to B. subtilis 3’ end of the 16S rRNA) was added to the reaction prior to the addition of NTPs in order to stimulate Era hydrolysis (Tu et al., 2011). Concentration of released inorganic phosphate was determined using Malachite Green Phosphate Assay kit (BioAssay Systems) according to manufacturer’s protocol. Briefly, reactions were stopped by the addition of 950 μl of water; 80 μl of diluted reaction was added to a single well of a flat-bottom 96-well plate (Costar). 20 μl of Malachite Green working reagent was added to each well and the reaction was incubated at room temperature for 30 min. Absorbance at 620 nm (Spark 10M plate reader, Tecan) of each reaction was compared to absorbances of known concentrations of phosphate standards. Absorbances from control reactions performed in the absence of SpoIVA for each NTP concentration were subtracted from absorbances of the respective reactions with SpoIVA to eliminate background hydrolysis. Hydrolysis rates for each NTP concentration were plotted using GraphPad Prism 7; V_max and K_m values were determined by fitting the data to Michaelis–Menten equation using best-fit values.

Limited proteolysis of His₆-SpoIVA and variants by partial trypsin digest was conducted as previously described (Castaing et al., 2013). Briefly, 2 μM His₆-SpoIVA was incubated in 100 μl of Buffer B supplemented with 4 mM NTP for 4 hr at 37°C. After addition of 1 μg/ml of trypsin (Sigma; diluted in 20 mM MgCl₂, 1 mM HCl), 15 μl of the reaction was removed and added at the indicated time points (0, 2, 5, and 10 min) to 5 μl of 4× LDS Sample Buffer (Invitrogen) containing beta-mercaptoethanol (Sigma) and heated at 95°C for 30 min to arrest proteolysis. 10 μl of each sample was separated by SDS-PAGE and stained with Coomassie blue. The intensity of the full-length His₆-SpoIVA band in each lane was quantified using ImageJ software (NIH), plotted as a function of time, and fitted to single-phase exponential decay using GraphPad Prism 7; reaction rates from each His₆-SpoIVA variant were normalized to the reaction rate of WT protein.

Fluorescence microscopic images of WT and mutant B. subtilis were taken as previously described (Ebmeier et al., 2012). Briefly, overnight cultures of B. subtilis grown in casein hydrolysate (CH) media at 22°C were diluted 1:20 into 20 ml CH and grown at 37°C for 2 hr. Sporulation was induced via resuspension method (Sterlini and Mandelstam, 1969) in A+B media supplemented with 80 μg/ml threonine (Sigma) at 37°C. After 3.5 hr, cells were harvested and resuspended in PBS (KD Medical) containing 1 µg/ml FM4-64 (Invitrogen) to visualize membranes, then placed on lysine-coated glass bottom dish (MatTek Corp.) under a 1% agarose pad. Cells were viewed with a DeltaVision Core microscope system (Applied Precision) equipped with an environmental control chamber. Images were captured with a Photometrics CoolSnap HQ2 camera. Seventeen planes were acquired every 0.2 μm at 22°C, and the data were deconvolved using SoftWorx software (GE Healthcare). At the sporulation time points that we examined, phase bright forespores had not yet developed; thus, the autofluorescence of forespores was not higher than background fluorescence. Additionally, control experiments with sporulating strains that did not harbor a gfp fusion indicated that the level of GFP fluorescence from fusions to SpoIVA was well above the limited background fluorescence of the cells.

To determine sporulation efficiencies, WT and mutant B. subtilis cells were grown in Difco Sporulation Medium for at least 24 hr at 37°C. Cultures were then exposed to 80°C for 20 min to kill non-sporulating cells. Surviving cells were enumerated by serial dilution and plating on LB agar. Viable spores were counted as colony forming units (CFUs); sporulation efficiencies were reported as a ratio to CFUs recovered from a parallel experiment using WT B. subtilis.

Steady state levels of SpoIVA and variants were assessed via immunoblotting as previously described (Tan et al., 2015). Briefly, B. subtilis cells were induced to sporulate via resuspension as described above. Sporulating cells were harvested and resuspended in 500 μl protoplast buffer (0.5 M sucrose, 10 mM K₂PO₄, 20 mM MgCl₂, and 0.1 mg/ml lysozyme [Sigma]) and incubated at 37°C for 30 min with shaking at 300 rpm. Protoplasts were harvested by centrifugation and lysed by resuspension in 200 μl PBS buffer. 15 μl of the sample was combined with 5 μl of 4× LDS sample buffer (NuPAGE), separated by SDS-PAGE, and transferred to PVDF membranes (Novex) using iBlot (Invitrogen). Blots were blocked in 5% skim milk (Carnation) in Tris-buffered saline (TBS)/Tween (TBS + 1% Tween 20; Sigma) overnight at 4°C with gentle shaking. Blots were incubated for 1 hr with antiserum raised against purified SpoIVA and detected using anti-rabbit IgG StarBright (Bio-Rad) with a ChemiDoc MP imager (BioRad).

Mass Photometry (MP, iSCAMS) experiments were carried out on a OneMP instrument (Refeyn, Oxford, UK) at room temperature. Rectangular 24 × 50 mm coverslips (#12544E, Fisher Scientific) and square 24 × 24 mm coverslips (#1405–10, Globe Scientific) were prepared by rinsing with water, ethanol, and isopropanol, and dried with clean nitrogen gas (Young et al., 2018) Approximately 10 µl of protein was loaded into the channel formed by stacked coverslips. MP signals were recorded for 100 s to allow detection of at least 2 × 10³ individual protein molecules. Raw MP data were processed in DiscoverMP software (Refeyn, Oxford, UK) and plotted as molar mass distribution histograms. For SEC-MALS, experiments were performed on a Agilent Series 1100 System (Agilent) with Superdex200 Increase 10/300 GL column (GE Healthcare), Helleos-II in-line multi angle light scattering detector (Wyatt Technology), and Optilab T-rEX refractive index detector (Wyatt Technology). SEC column was equilibrated with Buffer B until a stable refractive index baseline was reached. For sample analysis, 100 µl of SpoIVA at 0.94 mg/ml concentration in the presence of 4 mM ATP was injected at the 0.5 ml/min flow rate. All experiments were performed at room temperature, with MALS and RI detectors equilibrated at 20°C. Chromatograms were analyzed in ASTRA (V7.1, Wyatt Technology), and refractive index increment of 0.185 ml/g was used to determine the protein concentration.

Negative staining of the protein samples was performed on glow-discharged carbon-coated grids. For each condition, 3.5 µl sample was applied to a grid and incubated for 40 s. Excess sample was blotted away using a filter paper. The grid was then stained with 3.5 µl 1% uranyl acetate solution for 1 min and air-dried for imaging. Digital micrographs were collected using a 2 k CCD camera on a Hitachi 7650 electron microscope at an accelerating voltage of 80kV.

Cells were grown in CH media to OD_600nm ~0.5 and induced to sporulate via the resuspension method as described above. Metabolite extraction was performed as described previously (Yang et al., 2020). Briefly, 10 ml culture were sampled and harvested by filtration through PTFE membrane (Sartorius) at time points before and after resuspension in A+B media. Pellets on the PTFE membranes were soaked in 3 ml extraction solvent mix (on ice 50:50 [v/v] chloroform/water) and then vortexed to quench metabolism and extract metabolites. Cell extracts were centrifuged at 5000 × g for 10 min to remove the organic phase, and then centrifuged at 20,000 × g for 10 min to remove cell debris. Samples were frozen at −80°C if not analyzed immediately. Samples were analyzed using LC-MS and the metabolites were quantified as described previously (Fung et al., 2020; Yang et al., 2020), using an HPLC-MS system consisting of a Vanquish UHPLC system linked to electrospray ionization (ESI, negative mode) to a Q Exactive Orbitrap mass spectrometer (Thermo Scientific) operated in full-scan mode to detect targeted metabolites based on their accurate masses. LC was performed on an Acquity UPLC BEH C18 column (1.7 μm, 2.1 × 100 mm; Waters). Total run time was 30 min with a flow rate of 0.2 ml/min, using Solvent A (97:3 [v/v] water/methanol, 10 mM tributylamine, and 10 mM acetic acid) and acetonitrile as Solvent B. The gradient was as follows: 0 min, 5% B; 2.5 min, 5% B; 19 min, 100% B; 23.5 min 100% B; 24 min, 5% B; 30 min, 5% B. Quantification of metabolites from raw LC-MS data was performed by using the MAVEN software (Clasquin et al., 2012). Normalized ion count was defined and calculated as the ion count per OD_600nm per unit volume (5 ml) of the culture.