Magliozzi and colleagues report that Aip5 and Bud6 directly interact to form a novel “composite” actin nucleator, used to assemble thick actin cables that direct intracellular transport. After nucleation, Aip5 remains bound to the pointed end of the filament, streaming from inward from the cell cortex with actin cable retrograde flow.
Abstract
In this study, we show that two fungal proteins, Aip5 (related to vertebrate SH3BGRL) and Bud6, directly interact to form a novel “composite nucleator,” in which the pointed end of a nascent actin seed is bound by the thioredoxin-related domain of Aip5. The Aip5-Bud6 complex assembles F-actin seeds with free barbed ends and recruits formins to processively elongate and protect these ends from capping protein. The nucleation activities of Aip5 and Bud6 are critical for maintaining proper thickness of actin cable bundles in vivo, which prevents premature cable detachment from the bud neck and secretory traffic defects, as revealed by live imaging. In vitro single-molecule imaging reveals that after actin nucleation, Aip5 remains associated with a pointed end of the filament, and in vivo Aip5 puncta are observed directionally streaming inward from polarity sites by actin cable retrograde flow. Our findings expand the known diversity of actin nucleation mechanisms and reveal that Aip5 functions as a pointed-end capper in vivo.
Introduction
Eukaryotic cells use actin nucleators to assemble a wide range of cellular actin structures that promote diverse functions, such as polarized cell growth, cell migration, intracellular transport, endocytosis, phagocytosis, and cytokinesis (Dominguez and Holmes, 2011; Svitkina, 2018; Moseley and Goode, 2006). Actin nucleators have been classified into three types (or “classes”): the Arp2/3 complex, formins, and type III nucleators (e.g., Spire, COBL, LMOD, and adenomatous polyposis coli [APC]) (Chesarone and Goode, 2009; Campellone and Welch, 2010). The Arp2/3 complex nucleates branched actin filaments at locations such as sites of endocytosis and the leading edge (Goley and Welch, 2006; Gautreau et al., 2022). Formins nucleate unbranched filaments and remain processively attached to their barbed ends, accelerating filament growth and helping protect barbed ends from capping proteins (Goode and Eck, 2007; Breitsprecher and Goode, 2013). Type III nucleators recruit and organize multiple actin monomers into a seed that nucleates polymerization of a filament with a free barbed end (Bradley et al., 2020; Quinlan et al., 2005; Okada et al., 2010; Breitsprecher et al., 2012; Ahuja et al., 2007; Sagot et al., 2002; Evangelista et al., 2002; Goley and Welch, 2006). Interestingly, many type III nucleators also directly collaborate with formins to assemble actin filaments in vitro and in vivo. Human APC interacts with formins to assemble filaments by a “rocket launcher” mechanism, in which the formin and APC separate after nucleation (Okada et al., 2010; Breitsprecher et al., 2012). Similarly, Spire and formins collaborate to assemble actin filaments, with Spire and formins separating after nucleation (Bradley et al., 2020; Quinlan et al., 2007; Sitar et al., 2011). Whereas animals express all three classes of actin nucleators, fungal and plant species have been thought to only have two: the Arp2/3 complex and formins. The absence of obvious type III nucleators in fungi and plants has been surprising given the critical roles they play in animal species in performing a range of actin-based functions.
Budding yeast have two formins, Bni1 and Bnr1, which assemble actin cables, the tracks for myosin-based transport of secretory vesicles to the bud tip (Evangelista et al., 2002; Sagot et al., 2002; Bretscher, 2003; Schott et al., 2002). Bni1 and Bnr1 assemble complementary sets of cables in the bud and mother cell compartments, respectively (Pruyne et al., 2004). Genetic loss of either formin alone is tolerated, whereas loss of both formins abolishes cable assembly and is lethal (Vallen et al., 2000). Yeast actin cables undergo continuous assembly and turnover, growing from polarity sites (bud tip and bud neck) where formins are activated, and extending toward the back of the mother cell at a rate of ∼0.4 µm/s (Yang and Pon, 2002; Yu et al., 2011). Each actin cable is itself a bundle, comprised of many shorter and overlapping actin filaments (∼0.5 µm long) cross-linked together, in which the barbed ends of the filaments are oriented toward the bud tip (Kamasaki et al., 2005). Thus, formation of a single cable involves multiple separate actin nucleation events, possibly one for each filament in a cable bundle. Yet, our understanding of how alterations in actin nucleation levels influence different parameters of cable architecture, such as cable number, length, and/or thickness (i.e., number of filaments in cross section), has remained limited.
In addition, we only have a limited understanding of how the filaments in actin cables, once nucleated, are stabilized so that they are able to grow to increased lengths (∼0.5 µm long) compared with the filaments in cortical actin patches nucleated by the Arp2/3 complex (∼0.05 µm long). The growth of actin filaments in the yeast cytosol is limited by the high concentrations of actin disassembly-promoting factors that it contains (Gonzalez Rodriguez et al., 2023), which rapidly disassemble unprotected filaments by a variety of mechanisms (Goode et al., 2023). While tropomyosin decorates the sides of the constituent actin filaments in cables and plays an instrumental role in protecting them from rapid disassembly, it has remained uncertain whether the filaments in cables are also capped at their ends to provide further stabilization. A recent live-imaging study in budding yeast could find no evidence for capping protein association with the barbed ends of filaments in actin cables (Wirshing et al., 2023), and no pointed-end capper has been identified in the fungal kingdom aside from the Arp2/3 complex, which is exclusively found in the branched actin networks of cortical patches, not in cables.
A number of formin-binding partners tune their actin nucleation and elongation activities in vivo. Among these, Bud6 and Aip5 each have been shown to enhance Bni1- and Bnr1-mediated actin nucleation (but not elongation) through their abilities to directly bind formins and actin monomers (Moseley et al., 2004; Graziano et al., 2011; Graziano et al., 2013; Moseley and Goode, 2005; Xie et al., 2019; Xie et al., 2022). Importantly, the fragments of Bud6 (489–788) and Aip5 (1,110–1,234) used in these biochemical studies exhibit little or no actin nucleation activity on their own at physiological concentrations. Thus, neither Aip5 nor Bud6 meets the criteria for being defined as actin nucleators. Although Bud6 and Aip5 directly interact in vitro and in vivo (Glomb et al., 2019), the fragments used in the in vitro studies above did not include the regions that mediate their interaction. This has left open the question of whether direct interactions between Aip5 and Bud6 might change their activities on actin.
Here, we purified full-length Bud6 (Bud6-FL) and a C-terminal fragment of Aip5 (Aip5C-L), which includes the Bud6-binding site (BBS) (Glomb et al., 2019). We show that Bud6-FL and Aip5C-L, when combined, robustly nucleate actin assembly, forming filament seeds with free barbed ends. Through a combination of mutagenesis and quantitative image analysis, we determine that the nucleation activities of Bud6 and Aip5 are critical for maintaining proper actin cable cross-sectional thickness. Further, using live imaging we show that proper cable thickness prevents aberrant detachment of cables from the bud neck leading to defects in secretory traffic. Using in vitro single-molecule imaging, we demonstrate that after actin nucleation Aip5 caps the pointed ends of filaments, and using in vivo live imaging, we show that Aip5 puncta remain associated with actin cables as they undergo retrograde flow from polarity sites (bud tip and neck) toward the back of the mother cell.
Results
Aip5 and Bud6 form a complex that potently nucleates actin assembly and recruits formins to elongate the filaments
Previous studies have shown that C-terminal fragments of Aip5 (1,110–1,234) and Bud6 (489–788) each enhance formin-mediated actin nucleation, but themselves lack appreciable actin nucleation activity and thus are not classified as actin nucleators (Glomb et al., 2019; Xie et al., 2019; Moseley et al., 2004). However, Aip5 and Bud6 also directly associate with each other in vitro and in vivo through domains that were not included in the studies above, leaving open the question of how direct interactions between Bud6 and Aip5 might affect their activities.
To address this open question, we purified a longer fragment of Aip5 (Aip5C-L), which includes its BBS, and full-length Bud6 (Bud6-FL), which includes its N-terminal Aip5-binding region (Glomb et al., 2019) (Fig. 1 A). We then tested the activities of Aip5C-L and Bud6-FL, individually and combined, at physiological concentrations (100 nM) (Xie et al., 2019; Xie et al., 2022) in bulk pyrene-actin assembly assays containing vertebrate muscle actin. In these assays, Aip5C-L and Bud6-FL individually lacked appreciable actin nucleation activity, but together, they robustly nucleated actin assembly both in the presence and in the absence of profilin (Fig. 1, B and C; and Fig. S1, A and B). On the other hand, nucleation was not observed when Aip5C-L and C-Bud6 (489–788) were combined (Fig. S1 C), suggesting that nucleation requires their direct interaction.
Figure 1.
Bud6 and Aip5 synergize to nucleate actin assembly. (A) Schematics of Bud6 and Aip5. The arrow indicates domains in Aip5 and Bud6 mediating their direct interaction (Glomb et al., 2019). BBS: Bud6-binding site. (B) Bulk actin assembly assays showing that Bud6-FL and Aip5C-L synergize to assemble F-actin. Reactions contain 2 μM G-actin (5% pyrene-labeled), 5 μM yeast profilin, and as indicated 100 nM Bud6-FL and/or 100 nM Aip5C-L. Data shown are from one of three independent experiments. (C) Quantification of assembly rates determined from the slopes of curves as in B. Data are the mean ± SD and are from three independent trials (n = 3). ****P ≤ 0.0001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (D) Fluorescence anisotropy assays using 40 nM labeled Bud6 (1–145)-KCK-Alexa488 and variable concentrations of Aip5C-L. Data were fit to a single-site–binding isotherm (cyan line). (E) Concentration-dependent effects of Aip5C-L on actin nucleation in the presence of 2 μM G-actin (5% pyrene-labeled), 5 μM yeast profilin, and 100 nM Bud6-FL. See Materials and methods for how rates of F-actin assembly were quantified and the data were fit (magenta line) to determine Kapp. (F) Representative TIRF images taken 600 s after initiation of actin polymerization. Reactions contain 1 μM G-actin (10% Alexa 488–labeled, 0.5% biotin-labeled), 5 μM yeast profilin, and as indicated 100 nM Bud6-FL and/or 100 nM Aip5C-L. Scale bar, 20 μm. Data shown are from one of three independent experiments. (G) Number of actin filaments nucleated per FOV, 600 s after initiation of polymerization. Data are the mean ± SD and are from three independent trials (n = 9 FOVs per condition). ****P ≤ 0.0001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (H) Actin filament elongation rates quantified from images as in F. Data are the mean ± SD and are from three independent trials (n = 15 filaments per condition). ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (I) Bulk actin assembly assays showing that Bud6-FL and Aip5C-L collaborate with C-Bni1 to assemble F-actin in the presence of profilin and capping protein. Reactions contain 2 μM G-actin (5% pyrene-labeled), 5 μM yeast profilin, 100 nM yeast capping protein (Cap1/2), and as indicated 10 nM C-Bni1 (FH1-C), 100 nM Bud6-FL, and 100 nM Aip5C-L. Data shown are from one of three independent experiments. (J) Quantification of assembly rates determined from the slopes of curves in I. Data are the mean ± SD and are from three independent trials (n = 3). ****P ≤ 0.0001, **P ≤ 0.01, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. FOV, field of view.
Figure S1.
Related to Fig. 1 : Supporting in vitro data on the combined actin nucleation activities of Bud6 and Aip5. (A) Bulk actin assembly assays showing that Bud6-FL and Aip5C-L synergize to assemble F-actin. Reactions contain 2 μM G-actin (5% pyrene-labeled) and as indicated 100 nM Bud6-FL and/or 100 nM Aip5C-L. Data shown are from one of three independent experiments. (B) Quantification of assembly rates determined from the slopes of curves as in B. Data are the mean ± SD and are from three independent experiments (n = 3). ****P ≤ 0.0001, **P ≤ 0.01, *P ≤ 0.05, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (C) Bulk actin assembly assays showing that Bud6C-L (488–789) and Aip5C-L fail to synergize in nucleating actin assembly. Reactions contain 2 μM G-actin (5% pyrene-labeled), and as indicated 100 nM Bud6-FL, 100 nM Bud6C-L, and/or 100 nM Aip5C-L. (D) Bulk assays showing that Bud6-FL and Aip5C-L synergize to nucleate yeast actin assembly. Reactions contain 2 μM yeast G-actin (with 5% pyrene-labeled vertebrate muscle actin), 5 μM yeast profilin, and as indicated 100 nM Bud6-FL and/or 100 nM Aip5C-L. Data shown are from one of three independent experiments. (E) Quantification of assembly rates determined from the slopes of curves as in D. Data are the mean ± SD and are from three independent experiments (n = 3). ****P ≤ 0.0001, ***P ≤ 0.001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (F) Bulk assays showing that Bud6-FL and Aip5C-L collaborate with C-Bni1 to assemble yeast actin in the presence of profilin and capping protein. Reactions contain 2 μM yeast G-actin (with 5% pyrene-labeled vertebrate muscle actin), 5 μM yeast profilin, 500 nM yeast capping protein (Cap1/2), and as indicated 10 nM C-Bni1 (FH1-C), 100 nM Bud6-FL, and 100 nM Aip5C-L.
Although the domains in Aip5 and Bud6 that mediate their direct interaction have been mapped (Glomb et al., 2019), the strength of the interaction has not been reported. Therefore, we used fluorescence anisotropy to quantify the interaction of a labeled fragment of Bud6 (1–145) with unlabeled Aip5C-L (Fig. 1 D). This revealed a relatively high-affinity interaction (KD = 85 ±33 nM). In addition, we tittered the nucleation effects of Bud6 (100 nM) with variable concentrations of Aip5C-L, which revealed a Kapp (71 nM) similar to their binding affinity (Fig. 1 E). Together, these observations show that the synergistic nucleation effects of Aip5 and Bud6 require their direct interaction, and that this collaboration is effective at concentrations similar to their known cellular concentrations (Xie et al., 2019; Xie et al., 2022).
These nucleation effects were confirmed and extended by direct visualization of actin assembly using total internal reflection fluorescence (TIRF) microscopy (Fig. 1 F and Video 1), where Aip5C-L and Bud6-FL together increased the number of new filaments assembled over time (Fig. 1 G). Importantly, the filaments nucleated by Aip5C-L and Bud6-FL grew at their barbed ends at a rate similar to spontaneously nucleated filaments in control reactions (Fig. 1 H). Thus, Aip5C-L and Bud6-FL form new actin filaments that grow from their barbed ends. Indeed, the further addition of capping protein to bulk actin assembly assays abolished the growth of filaments nucleated by Aip5C-L and Bud6-FL (Fig. 1 I).
Video 1.
Related to Fig. 1 F : Bud6-FL and Aip5C-L jointly promote actin nucleation in TIRF microscopy assays. Open-flow TIRF microscopy assays showing effects of 50 nM Bud6-FL and/or 50 nM Aip5C-L on F-actin assembly. All reactions contain 1 μM G-actin (10% Alexa 488–labeled). Video playback is 10 frames per second and is sped up 100-fold from real time, with a timestamp. Scale bars, 20 μm.
Since Aip5 and Bud6 each collaborate with formins in vivo to assemble actin cables (Graziano et al., 2011; Xie et al., 2019; Xie et al., 2022), we next considered whether the Aip5-Bud6 complex collaborates with formins by a mechanism related to APC–formin and Spire–formin partnerships. In these mechanisms, a single factor (APC or Spire) organizes actin monomers into an F-actin seed and directly recruits formins to processively cap the barbed end, enabling it to elongate in the presence of the capping protein (Okada et al., 2010; Breitsprecher et al., 2012; Bradley et al., 2020). To test whether the Aip5-Bud6 complex might be part of a similar mechanism, we compared the activities of Aip5 and Bud6, individually and combined, with yeast formins in bulk pyrene-actin assembly assays containing profilin and capping protein to mimic cytosolic conditions where spontaneous actin assembly is suppressed (Gonzalez Rodriguez et al., 2023). In these assays, we used C-Bni1, which includes the FH1 and FH2 domains, as well as the formin C-terminal tail that binds to Bud6 and Aip5 (Xie et al., 2019; Moseley and Goode, 2005). Under these conditions, Bud6-FL and Aip5C-L together showed minimal actin filament assembly activity, consistent with an inability to protect barbed ends from the capping protein (Fig. 1, I and J). C-Bni1 showed some activity on its own, as expected. However, combining C-Bni1 with both Bud6-FL and Aip5C-L led to robust filament assembly. These results support the model that Aip5 and Bud6 together nucleate the F-actin seed and recruit the formin to processively elongate the filament in the presence of the capping protein.
In the bulk and TIRF assays above, and elsewhere in this paper unless otherwise noted, we use vertebrate muscle actin, which is fairly similar in sequence (∼90% identical) and assembly properties to yeast actin. However, it is also important to validate the activities of yeast actin-binding proteins using yeast actin. To address this, in bulk actin assembly assays we verified that wild-type Aip5C-L and Bud6 together robustly nucleate yeast actin assembly (Fig. S1, D and E), similar to their combined effects on vertebrate actin. Furthermore, we verified that Aip5C-L and Bud6 collaborate with C-Bni1 in the presence of yeast actin, profilin, and capping protein (Fig. S1 F). Importantly, these assays also revealed that Aip5C-L on its own (without Bud6) has increased actin nucleation activity on yeast actin compared with vertebrate actin (Fig. S1 D). We speculate that this may be due to Aip5 having a higher binding affinity for the pointed ends of yeast actin filaments compared with vertebrate actin filaments (see below). Overall, these results validate the synergy of Aip5C-L and Bud6 in jointly promoting actin nucleation, and their synergy in collaborating with formins to assemble actin filaments in the presence of capping protein.
Loss of Aip5 and Bud6 in vivo leads to defects in actin cable architecture and premature detachment of cables from the bud neck
To gain a better understanding of how the nucleation activities of Aip5 and Bud6 contribute to actin cable formation in vivo, we compared cable architecture in the mother cell compartments of wild-type, bud6Δ, aip5Δ, and bud6Δ aip5Δ double-mutant cells. Previous studies on bud6Δ mutants have reported that they have reduced cable numbers and reduced overall cable network levels, but have not quantified cable length and/or thickness (Graziano et al., 2011, 2013; Glomb et al., 2019; Xie et al., 2022). Previous studies on aip5Δ mutants have reported reduced cable network levels and reduced cable lengths, but reached conflicting conclusions as to whether aip5Δ mutants have reduced cable number (Glomb et al., 2019; Xie et al., 2019; Xie et al., 2022). Collectively, these studies have provided a crucial foundation for our understanding of how cable networks are impaired in bud6Δ and aip5Δ mutants. However, to gain a more complete view of how actin cable networks are altered in bud6Δ and aip5Δ single mutants and bud6Δ aip5Δ double mutants, we decided to directly measure and quantitatively compare each of the key parameters (cable number, cable length, and cable cross-sectional fluorescence intensity) in each of these strains, analyzed in parallel in one study.
Wild-type, bud6Δ, aip5Δ, and bud6Δ aip5Δ cells were grown in parallel, fixed, stained with Alexa 488-phalloidin, and imaged by super-resolution microscopy (Fig. 2 A). Using max intensity projections, we measured the number of cables per mother cell and average cable length, and using sum intensity projections, we measured average cable cross-sectional fluorescence intensity (i.e., relative number of filaments in cable thickness). Interestingly, bud6Δ and aip5Δ single mutants had similar numbers of cables compared with wild-type cells, and bud6Δ aip5Δ double mutants had only slightly reduced cable numbers (Fig. 2 B). More strikingly, cable thickness was greatly reduced in each single mutant strain, and to a greater extent in the double mutants (Fig. 2 C). These changes were accompanied by a more modest reduction in cable length in all three mutant strains (Fig. 2 D). Together, our results show that the most pronounced defect in cables caused by loss of Aip5 and/or Bud6 is a reduction in cable thickness. This in turn suggests that high levels of actin nucleation, supported by Bud6 and Aip5, are required to maintain cable bundle thickness.
Figure 2.
Loss of BUD6 and AIP5 results in thinner and shorter actin cables. (A) Representative images of indicated wild-type and mutant yeast strains fixed and stained with Alexa 488-phalloidin. Scale bar, 5 μm. Data shown are from one of three independent experiments. (B) Graph comparing the number of actin cables in indicated yeast strains. N = 60 cells from three independent experiments. Data are the mean ± SD. **P ≤ 0.01, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (C) Graph comparing the cross-sectional fluorescence intensity of actin cables in indicated yeast strains. N = 60 cells from three independent experiments. Data are the mean ± SD. ****P ≤ 0.0001, ***P ≤ 0.001, **P ≤ 0.01, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (D) Graph comparing actin cable length. N = 60 cables from three independent experiments. Error bars, SD. ****P ≤ 0.0001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (E) (Top) Representative images of indicated yeast strains fixed and stained with Alexa 488-phalloidin. Scale bar, 2 μm. (Bottom) Schematic of Bud6 domains, highlighting point mutations that disrupt Bud6 interactions with formins (bud6-35) and G-actin (bud6-8). Data shown are from one of three independent experiments. (F) Graph comparing the cross-sectional fluorescence intensity of actin cables in indicated yeast strains. N = 60 cells from three independent experiments. Data are the mean ± SD. ****P ≤ 0.0001 by one-way ANOVA followed by Tukey’s HSD post hoc test.
In fission yeast, a homeostatic mechanism exists to balance F-actin distribution between actin patch networks and actin cable networks, such that genetic or pharmacological insults causing a reduction in one network result in more intense staining of the other network (Burke et al., 2014; Suarez et al., 2015). For this reason, we asked whether the reduction in F-actin cable staining in our aip5Δ and bud6Δ mutants in budding yeast changes either the total F-actin levels in cells (Fig. S2 A) or the intensity of actin patch staining (Fig. S2 B). Our data show that the total cellular F-actin levels are reduced in the mutants, but that actin patch intensities do not change. It was not possible to accurately quantify the number of patches per cell, so it remains possible that patch networks experience some increase when cables are diminished. However, our data demonstrate clearly that the mutants have reduced levels of total cellular F-actin, suggesting that the mechanisms for cable patch homeostasis in budding yeast may be different from fission yeast.
Figure S2.
Related to Fig. 2 : Supporting in vivo data on the effects of bud6 and aip5 mutants on cellular F-actin levels and actin cable architecture. (A) Quantification of whole-cell F-actin fluorescence intensities (by phalloidin staining). Each data point is one cell, in which its total fluorescence (raw integrated density) was divided by cell area. Data are the mean ± SD and are from three independent experiments (n = 30 cells per strain). **P ≤ 0.01, *P ≤ 0.05, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (B) Quantification of cortical patch F-actin fluorescence intensities. Data are the mean ± SD and are from three independent experiments (n = 150 patches per strain). Ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (C) Graph comparing the number of actin cables in indicated yeast strains. N = 60 cells from three independent experiments. Data are the mean ± SD. ****P ≤ 0.0001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (D) Graph comparing actin cable length. N = 60 cables from three independent experiments. Data are the mean ± SD. ****P ≤ 0.0001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test.
To determine whether the actin cable phenotypes that we observed in bud6Δ and bud6Δaip5Δ mutants depend on Bud6’s interactions with formins (bud6-35) and/or G-actin (bud6-8) (Fig. 2 E), we analyzed the cable architecture of these point mutations in vivo (Graziano et al., 2011). We found that bud6-35 and bud6-8 mutants caused similar cable defects to bud6Δ (in AIP5 or aip5Δ backgrounds), including markedly reduced cable thickness (cross-sectional fluorescence intensity), modestly reduced cable length, and no significant change in cable number compared with wild-type cells (Fig. 2 F; and Fig. S2, C and D). Overall, these results indicate that the ability of Bud6 to promote cable assembly in vivo depends not only on its actin nucleation effects, but also on its ability to recruit and collaborate with formins, which agrees with our in vitro assays (Fig. 1, I and J).
To gain insights into how the defects in actin cable architecture in bud6Δ and aip5Δ mutants might affect cable dynamics, we performed live-cell imaging. We integrated into each strain our recently described LifeAct-mNG cable marker, which has improved brightness and photostability in comparison with previous markers (Wirshing and Goode, 2024). In wild-type cells, we observed cables extending from the bud neck to the back of the mother cell, where they remained stabilized at that length for a minute or longer before abruptly disappearing (Fig. 3 A and Video 2). In bud6Δ aip5Δ cells (and to a lesser extent in the single mutants), the cables decorated by LifeAct-mNG appeared more faint, consistent with our reduced phalloidin staining in fixed cells. Even more strikingly however, in these mutants the extending cables often detached from the neck before reaching the back of the mother cell, then rapidly disappeared (Fig. 3, A and B). Given that cables serve as tracks for secretory vesicle transport to the bud tip (Pruyne et al., 1998; Johnston et al., 1991), we reasoned that these cable detachment events might compromise secretory traffic to the bud and used live imaging to compare levels of secretory vesicles (GFP-Sec4) in the mother cell compartment (Fig. 3 C and Video 3). In wild-type cells, secretory vesicles are rapidly transported from the mother cell to the bud tip, and therefore sparse in the mother cell compartment and highly enriched at the bud cortex. In contrast, we observed abnormally high numbers of secretory vesicles in the mother cell compartments of bud6Δ, aip5Δ, and bud6Δ aip5Δ cells (Fig. 3 D), suggesting a defect in vesicle traffic. The impaired transport of vesicles and accumulation in the mother cell compartment in these mutants correlate with the severity of their altered actin cable architecture and cable detachment events.
Figure 3.
BUD6 and AIP5 mutants assemble actin cables that prematurely detach from the bud neck, resulting in defective polarized secretion. (A) Representative SoRa images from live-cell imaging experiments showing LifeAct-mNG–labeled F-actin cables extending (blue arrows) and detaching (red arrowheads). Scale bar, 3 μm. Data shown are from one of three independent experiments. (B) Quantification of cable detachment events at the bud neck during a 60-s viewing period for indicated strains. Data from three independent trials (n = 60 cells per strain). ****P ≤ 0.0001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. Data are the Mean ± SD. (C) Representative images of GFP-Sec4 marked secretory vesicle localization for indicated strains. Scale bar, 5 μm. Data shown are from one of three independent experiments. (D) Quantification of the number of secretory vesicles (GFP-Sec4 puncta) in the mother cell compartments of the indicated yeast strains. Data are from three independent trials (n = 75 cells per strain). ****P ≤ 0.0001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. Data are the mean ± SD. SoRa, Super-Resolution by Optical Re-Assignment.
Video 2.
Related to Fig. 3 A : Loss of BUD6 and AIP5 leads to actin cables prematurely detaching from the bud neck. SoRa time-lapse imaging of LifeAct-mNG–labeled F-actin in the indicated yeast strains. Note the detachment of cables from the bud neck in mutant strains. Video playback is 10 frames per second and has been sped up sixfold from real time, with a timestamp. Scale bar, 2 μm.
Video 3.
Related to Fig. 3 C : Loss of BUD6 and AIP5 leads to aberrant accumulation of secretory vesicles in the mother cell compartment. SoRa time-lapse imaging of GFP-Sec4 marked secretory vesicles in the indicated yeast strains. Note the accumulation of vesicles in the mother cell compartment in mutant strains. Video playback is 10 frames per second and has been sped up fivefold from real time, with a timestamp. Scale bar, 2 μm.
Taken together, these results show that bud6Δ, aip5Δ, and bud6Δ aip5Δ mutants have aberrantly thin actin cables, which fail to maintain stable attachments to the bud neck cable, leading to defects in polarized secretory traffic.
Structural modeling and mutational analysis of the composite actin nucleator
To gain structural insights into how Aip5 and Bud6 nucleate an F-actin seed, we used AlphaFold3 to model the interactions of actin subunits with the actin-binding C-terminal fragments of Aip5 and Bud6. A consistent picture emerged from the models, in which the globular thioredoxin-related domain of Aip5 (1,129–1,233) contacts the final two actin subunits at the “pointed end” of an F-actin seed, possibly stabilizing a short-pitch contact between these two actin subunits (Fig. 4, A and B; and Fig. S3, A and B). This interaction of Aip5 with the pointed end is also supported by the evolutionary conservation of these actin-contacting surface residues (Fig. S3, A–C), and by a study showing that a triple mutation in Aip5 (L1150A, N1162A, and S1165A) that changes key actin-binding residues in our model disrupts binding of Aip5 to actin in vitro (Fig. 4 C; and Fig. S3, B and D) (Xie et al., 2019).
Figure 4.
Structural modeling of how Bud6 and Aip5 interact with actin and jointly nucleate actin filament assembly. (A) AlphaFold model of Bud6-Flank (699–786) and Aip5C-L (997–1,233) in a complex with three actin subunits. (B) Cartoon highlighting key features of the model. (C) AlphaFold model of Aip5C thioredoxin-like domain. The three residues mutated in 3M actin-binding deficient mutant (L1150, N1162, and S1165) are highlighted in orange space-filling spheres. Aip5 is in the same orientation as in A. (D) (Top) Aip5 domain schematics to indicate the following: 3M (L1150A, N1162A, and S1165A, G-actin–binding mutant), ΔBBS (truncation of putative BBS), Aip5C-L (Aip5 protein fragment containing BBS), Aip5C-S (Aip5 protein fragment lacking BBS), and Bud6flank-Aip5C-S chimera (fusion protein between G-actin–binding regions of Bud6 and Aip5). (Bottom) Schematic of Bud and Aip5 constructs used to test the structural model, including Bud6-ΔABS2 (residues 1–740), Bud6flank (residues 699–788), and a Bud6flank-Aip5C-S chimera. (E) Bulk actin assembly assays showing that Bud6-FL and Aip5C-L actin assembly is dependent on Aip5C G-actin and Bud6 binding. Reactions contain 2 μM G-actin (5% pyrene-labeled), 5 μM yeast profilin, and as indicated 100 nM Bud6-FL and/or 100 nM Aip5C constructs. Data shown are from one of three independent experiments. (F) Quantification of assembly rates determined from the slopes of curves as in B. Data are the mean ± SD and are from three independent experiments (n = 3). ****P ≤ 0.0001, ** P ≤ 0.01, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (G) Representative images of indicated yeast strains fixed and stained with Alexa 561-phalloidin to determine actin cable thickness (cross-sectional fluorescence intensity). Scale bar, 5 μm. Data shown are from one of three independent experiments. (H) Graph comparing the cross-sectional fluorescence intensity of actin cables in indicated yeast strains. N = 75 cells from three independent experiments. Error bars, SD. ****P ≤ 0.0001, ***P ≤ 0.001 by one-way ANOVA followed by Tukey’s HSD post hoc test. (I) Representative bulk assays showing that the ability of Bud6 and Aip5C-L to jointly promote actin nucleation depends on the second actin-binding site (ABS2) in Bud6. Reactions contain 2 μM G-actin (5% pyrene-labeled), 5 μM yeast profilin, and as indicated 100 nm Aip5C-L with 100 nM Bud6-FL or Bud6-ΔABS2. Control reactions contain only actin and profilin. Data shown are from one of three independent experiments. (J) Quantification of assembly rates determined from the slopes of curves as in I. Data are the mean ± SD and are from three independent experiments (n = 3). ***P ≤ 0.001, ** P ≤ 0.01, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (K) Representative images of indicated yeast strains fixed and stained with Alexa 561-phalloidin. Scale bar, 5 μm. Data shown are from one of three independent experiments. (L) Graph comparing the cross-sectional fluorescence intensity of actin cables in indicated yeast strains. N = 90 cells from three independent experiments. Error bars, SD. ****P ≤ 0.0001 by Welch’s unequal variance t test. (M) Bulk actin assembly assays showing that the Bud6-Flank and Aip5C-S fragments potently nucleate actin assembly when brought in close proximity using a short (GGS) linker.
Figure S3.
Related to Fig. 4 : Conservation in the thioredoxin-related domains of yeast Aip5 and vertebrate SH3BGRL2. (A) AlphaFold model of how Bud6 flank (699–788) and Aip5 (1,129–1,233) collaborate to seed an actin filament (left). Subunits are color-coded as in main figure. PE, pointed end. In the AlphaFold model, the structure of the Aip5 thioredoxin-related domain is highly consistent with its known crystal structure (Xie et al., 2019, PDB# 6ABR). The Aip5 thioredoxin-related domain alone was also rotated 90° relative to the model structure (right). (B–D) Surface representations of the Aip5 thioredoxin-related domain (1,129–1,233) highlighting key features. (B) Surface is highlighted in gray except where it appears to be in contact with actin PE1 or actin PE2 subunits, highlighted in magenta or pink, respectively. (C) Surface conservation mapped on the surface of Aip5 (1,129–1,233) (legend at bottom). (D) Three surface residues mutated in Aip5 3M (L1150, N1162, and S1165) highlighted in orange. Surface representations in B–D are in the same orientation as A (right). (E) Structural overlay of Aip5 (1,129–1,233; AlphaFold model, magenta) with cryo-EM structure of SH3BGRL2 (orange). (F) Structure of SH3BGRL2 bound to the pointed end of an actin filament (PDB# 8IAI) (top) compared with our model of Aip5 (1,129–1,233) bound to the pointed end of an actin filament (bottom). In E and F, note that secondary structures of Aip5 and SH3BGRL2 are highly similar except in the N-terminal α-helix of Aip5, which is absent in SH3BGRL2.
Our structural model further revealed that the G-actin–binding “flank” domain of Bud6 (699–788) contacts the sides of two adjacent subunits in the actin seed, possibly stabilizing the long-pitch contact between them (Fig. 4, A and B). In the model, the N-terminal portion of the Bud6 flank contains the known actin-binding site (which we refer to here as ABS1), consisting of two helices (αA and αB), and agrees with the known cocrystal structure of ABS1 bound to G-actin (Park et al., 2015). However, our model also suggests the existence of a second actin-binding site (termed ABS2), again consisting of two alpha helices (αC and αD), which closely resembles the structure of ABS1. An important implication of this model is that the Bud6 flank domain uses its ABS1 and ABS2 domains to recruit two actin monomers, which in turn interact longitudinally as part of the nucleation mechanism, while complementary interactions of Aip5 with the pointed end stabilize the actin seed.
We tested several key predictions of the model by generating a set of mutant and wild-type Aip5 and Bud6 polypeptides (Fig. 4 D). First, we asked how the previously described triple mutation (“3M”; L1150A, N1162A, and S1165A) (Xie et al., 2019), which disrupts key actin-binding surfaces in our model, impacts the ability of Aip5C-L to work with Bud6-FL to nucleate actin assembly. In bulk assays, mutant Aip5C-L(3M) showed greatly reduced nucleation activity compared with wild-type Aip5C-L (Fig. 4, E and F), consistent with our structural model. Similar effects were observed using yeast actin (Fig. S4 A). Second, we asked how direct interactions between Aip5C-L and Bud6-FL influence their nucleation activity. For this test, we truncated the BBS domain from Aip5C-L (997–1,234), producing a shorter fragment of Aip5C-S (1,110–1,234) (Fig. 4 D). In bulk assays, Aip5C-S combined with Bud6-FL produced a greatly reduced level of nucleation activity compared to Aip5C-L combined with Bud6-FL (Fig. 4, E and F). Thus, Aip5-Bud6 interactions are required for robust nucleation activity. In parallel, we dissected this model in vivo by asking how the 3M actin-binding mutation integrated into AIP5 (aip5-3M) or internally deleting the BBS (aip5ΔBBS) would affect actin cable organization (Fig. 4, D, G, and H). This analysis revealed that Aip5’s interactions with Bud6 and actin are both important for maintaining actin cable thickness (cross-sectional fluorescence intensity), but the interactions with actin are more critical than the interactions with Bud6, which mirrors our in vitro results (Fig. 4, E and F).
Figure S4.
Related to Fig. 4 : In vitro effects of the Aip5C-L(3M) mutant on yeast actin nucleation, and in vivo localization of Bud6-ΔABS2. (A) Bulk assays showing that the Aip5C-L(3M) mutant has impaired nucleation effects on yeast actin filament assembly, alone or together with Bud6-FL. Reactions contain 2 μM yeast G-actin (with 5% pyrene-labeled vertebrate muscle actin), 5 μM yeast profilin, and as indicated 100 nM Bud6-FL and/or 100 nM Aip5C-L or Aip5C-L(3M). (B) Representative images of the indicated yeast strains comparing the localization of wild-type Bud6-mNG and Bud6ΔABS2-mNG. Note that deletion of the ABS2 domain in Bud6 does not appear to change its localization in vivo. Scale bar, 5 μm.
As a third test of our structural model, we assessed the importance of the putative ABS2 sequences in Bud6 (741–788) for nucleation. Previous work showed that the flank domain of Bud6 contains a G-actin–binding site, which we refer to as ABS1 (699–740), and that it is critical for Bud6 functions in vitro and in vivo (Graziano et al., 2011; Park et al., 2015). However, our model predicts that there may be a second actin-binding site or ABS2 (741–788). Therefore, we generated a new construct (Bud6-ΔABS2) that truncates the ABS2 from Bud6-FL. In bulk assays containing Aip5C-L, Bud6-ΔABS2 failed to promote actin nucleation (Fig. 4, I and J). In vivo, the bud6-ΔABS2 mutant caused a severe reduction in actin cable thickness (Fig. 4, K and L) similar to bud6Δ (Fig. 2 C), even though its localization was similar to wild-type Bud6 (Fig. S4 B). These results, together with our analysis of bud6-8 (which targets ABS1), support the model that actin nucleation by Bud6 requires both actin-binding sites (ABS1 and ABS2), and that each is critical for Bud6 contributions to cable assembly in vivo.
As a final test of our model, we asked whether a Bud6-Aip5 chimera (Bud6flank-Aip5C-S), consisting of the actin-binding flank domain of Bud6 (699–788) connected by a short linker sequence (GGS) to the actin-binding thioredoxin-related domain of Aip5 (1,110–1,234), is sufficient to promote actin nucleation. Remarkably, the chimera strongly nucleated actin assembly, whereas the two separate halves of the chimera (Bud6 flank + Aip5C-S) combined in one reaction showed minimal nucleation activity in trans (Fig. 4 M). These results strongly support our structural model and demonstrate that robust actin nucleation by Aip5 and Bud6 requires both their interactions with each other and their individual interactions with actin.
Aip5 moves inward from the cell cortex with actin cable retrograde flow
To investigate where in cells Aip5, Bud6, and Bni1 collaborate to assemble actin cables, we compared their localization patterns and dynamics by live imaging. A previous study showed that Bni1-3xGFP forms puncta in the cytosol, which are transiently recruited to the bud cortex and activated to assemble actin cables, then move inward from the cortex passively with cable retrograde flow, presumably attached to filament barbed ends (Buttery et al., 2007). Further, this study showed that Bud6 exists in a distinct set of highly dynamic puncta, which overlap with Bni1 only at the bud tip, and do not move inward with cables. For improved brightness and photostability, we tagged the C termini of Aip5, Bud6, and Bni1 with 3xmNG and performed live imaging on each strain. Our results broadly agree with the study above, as we observed that Bni1 and Bud6 are in dynamic puncta enriched at the bud tip, and that Bni1 puncta (but not Bud6 puncta) move inward from the bud cortex in a manner consistent with passive transport by cable retrograde flow (Fig. 5 A, top row, and Video 4). In contrast to Bni1 puncta, Bud6 puncta moved in the opposite direction, toward the bud tip. These movements are consistent with the colocalization of Bud6 with secretory vesicles, which are trafficked by myosin along actin cables to the bud tip (Garabedian et al., 2018). When we treated cells with latrunculin B (LatB) to disassemble F-actin, Bni1 puncta accumulated at the bud cortex, consistent with the observations of Buttery and colleagues (Fig. 5 A, bottom row, and Video 5). Latrunculin treatment also led to a strong enrichment of Bud6 at the bud neck (red arrows, Fig. 5 A, bottom row, and Video 5), possibly mediated by its interactions with Bnr1, which is anchored to the bud neck independent of cable assembly (Buttery et al., 2007).
Figure 5.
Cortical Aip5 puncta move inward with actin cable retrograde flow. (A) Top panel: Representative images of endogenously tagged Aip5-3xmNG, Bni1-3xmNG, or Bud6-3xmNG in cells. Bottom panel: The same strains treated with 200 μM LatB to disassemble F-actin. Note the loss of Aip5-3xmNG puncta in the mother cell compartment upon LatB treatment. Scale bars, 5 μm. Data shown are from one of three independent experiments. (B) Histograms showing the distributions of the velocities of Bni1 and Aip5 puncta moving along actin cables. N = 60 puncta from three independent experiments. Puncta were traced for at least 1 μm. (C) Representative images of indicated yeast strains expressing wild-type, 3M, or ΔBBS Aip5-3xmNG constructs. Scale bar, 5 μm. (D) Representative images of indicated yeast strains expressing Aip5-3xmNG. Scale bar, 5 μm.
Video 4.
Related to Fig. 5 A (top row): Aip5-3xmNG and Bni1-3xmNG puncta (but not Bud6-3xmNG puncta) move inward from the cell cortex with actin cable retrograde flow. SoRa time-lapse imaging showing movements of Aip5-3xmNG, Bni1-3xmNG, and Bud6-3xmNG puncta. Aip5-3xmNG and Bni1-3xmNG puncta move inward from polarity sites (bud tip and bud neck) toward the back of the mother cell. In contrast, Bud6-3xmNG puncta move in the opposite direction, toward the bud tip, which is explained by their known association with secretory vesicles. Video playback is 15 frames per second and is sped up fourfold from real time, with a timestamp. Scale bar, 2 μm.
Video 5.
Related to Fig. 5 (bottom row): LatB treatment abolishes the directed movements of Aip5-3xmNG, Bni1-3xmNG, and Bud6-3xmNG. SoRa time-lapse imaging showing that treatment of cells with LatB for 10 min to disassemble F-actin attenuates the retrograde movements of Aip5-3xmNG and Bni1-3xmNG puncta, and the anterograde movements of Bud6-3xmNG puncta. Video playback is 15 frames per second and is sped up fourfold from real time, with a timestamp. Scale bar, 2 μm.
The Aip5-GFP signal is known to be concentrated at the bud tip and bud neck, and becomes more enriched at these sites after latrunculin treatment (Xie et al., 2019). Our live imaging of Aip5-3xmNG yielded similar results (Fig. 5 A, Videos 4, and 5). However, our use of an improved tag allowed us to also detect a new set of Aip5-3xmNG puncta, not previously reported, which directionally stream inward from polarity sites (bud cortex and bud neck) toward the back of the mother cell, i.e., in a manner consistent with passive transport by cable retrograde flow (Fig. 5 A and Video 4). Consistent with this view, the velocities of Aip5-3xmNG puncta were similar to actin cable extension rates, and to the velocities of Bni1-3xmNG passive movements on cables (Fig. 5 B) (Buttery et al., 2007; Yang and Pon, 2002). Interestingly, Aip5-3xmNG movements were readily visible in both the mother and bud compartments, whereas Bni1-3xmNG movements were primarily observed in the bud. This result raises the possibility of a compartment-specific Bni1 retrieval mechanism from barbed ends. Importantly, the retrograde movements of Aip5-3xmNG and Bni1-3xmNG puncta, and the anterograde movements of Bud6-3xmNG, were abolished by latrunculin treatment (Video 5), demonstrating that the directed movements of all three proteins are actin-dependent. Furthermore, the directed movement of Aip5 puncta on cables was abolished by the pointed-end actin-binding defective mutant Aip5-3M-3xmNG (Fig. 5 C and Video 6), suggesting that Aip5 interacts with the pointed ends of actin filaments in cables, and through this interaction is passively transported with cables by retrograde flow.
Video 6.
Related to Fig. 5 C : Retrograde movements of Aip5-3xmNG puncta are disrupted by an actin-binding deficient mutant. SoRa time-lapse imaging showing movements of Aip5-3xmNG puncta in the indicated strain backgrounds. “WT” is wild type, 3M is the actin-binding deficient triple mutation, and “ΔBBS” is an internal deletion of the Bud6-binding domain (997–1,110). Video playback is 15 frames per second and is sped up fourfold from real time, with a timestamp. Scale bar, 2 μm.
We also noticed that Aip5 levels appeared to be higher in the actin-binding deficient mutant Aip5-3M-3xmNG cells compared with wild-type Aip5-3xmNG or Aip5-ΔBBS-3xmNG cells (Fig. 5 C and Video 6). This hypothesis was confirmed quantitatively (Fig. S5 A), and suggests that the inability of Aip5 to bind actin and associate with cables may affect its in vivo expression and/or turnover rates.
Figure S5.
Related to Fig. 5 : In vivo disruption of Aip5–actin interactions (Aip5-3M) results in higher Aip5 protein levels compared with wild-type Aip5 or Aip5ΔBBS. (A) Quantification of Aip5-3xmNG whole-cell fluorescence intensities. Data are the mean ± SD and are from three independent experiments (n = 60 cells per strain). ****P ≤ 0.0001, ns, P > 0.05 by one-way ANOVA followed by Tukey’s HSD post hoc test. (B) Velocities of Aip5-3xmNG puncta streaming inward from polarity sites (bud tip and neck) toward the back of the mother cell in the indicated strains. Data are the mean ± SD and are from three independent experiments (n = 60 puncta per condition). ns, P > 0.05 by Welch’s unequal variance t test. (C) High-speed cosedimentation assays showing that Aip5C-S does not appreciably associate with the sides of actin filaments (2 µM). Although Aip5 binds to the pointed ends of the filaments, it is not detected in these assays because the concentration of pointed ends is very low (<1 nM). (D) Pointed-end depolymerization rates for yeast actin filaments in the presence or absence of the indicated Aip5 polypeptides. Reactions contain 2 µM yeast F-actin (with 10% pyrene-labeled vertebrate muscle actin) and 40 nM CapZ to cap the barbed ends. Depolymerization was induced at time 0 by the addition of 3 µM vitamin D–binding protein in the presence or absence of 25 nM of the indicated Aip5 constructs. Note that Aip5C-L(3M) fails to cap the pointed ends. Source data are available for this figure: SourceData FS5.
We next asked whether the retrograde movements of Aip5-3xmNG puncta depend on Aip5’s interactions with Bud6 and/or Bni1. In bni1Δ cells, Aip5-3xmNG movements were seen only in the mother cell (Fig. 5 D and Video 7), as expected since Bni1 is required for cable assembly in the bud (Pruyne et al., 2004). Further, Aip5-3mNG puncta moved at a similar velocity in wild-type and bni1Δ cells (Fig. S5 B), reinforcing the view that Aip5 movements on actin cables are independent of Bni1. Note that it is not surprising Aip5-3mNG and Bni1-3mNG puncta move on cables at the same rate, since each appears to be passively transported on cables. These results also demonstrate that the movements of Aip5-3xmNG puncta in the mother cell do not require interactions with Bni1. The retrograde movements of Aip5-3xmNG also were similar in bud6Δ and wild-type cells (Fig. 5 D and Video 7), consistent with the cable association of Aip5 but not Bud6 (Buttery et al., 2007; and our data above). Furthermore, the movements of Aip5-ΔBBS-3xmNG puncta (a mutant lacking the Bud6-binding domain) were similar to wild-type Aip5-3xmNG, demonstrating that Aip5 movements on cables are independent of interactions with Bud6 (Fig. 5 C and Video 6). Finally, Aip5-3xmNG movements were similar in bnr1Δ and wild-type cells (Fig. 5 D and Video 7), which is expected given that Bni1-assembled cables grow from the bud into the mother cell compartment in the bnr1Δ background (Pruyne et al., 2004).
Video 7.
Related to Fig. 5 D : Retrograde movement of Aip5-3xmNG puncta in bud6Δ, bni1Δ, and bnr1Δ strains. SoRa time-lapse imaging showing movement of Aip5-3xmNG puncta in the indicated strains. Video playback is 15 frames per second and is sped up fourfold from real time, with a timestamp. Scale bar, 2 μm.
Overall, from these results we conclude that Aip5 associates with the pointed ends of actin filaments in cables, and through these interactions (and independent of interactions with Bud6, Bni1, or Bnr1) is passively transported from polarity sites toward the back of the mother cell by cable retrograde flow.
Aip5 caps pointed ends of actin filaments in vitro
Our AlphaFold modeling suggests that the thioredoxin-related domain of Aip5 directly associates with the pointed end of the actin filament (Fig. 4 A). To further test this model, we asked whether the thioredoxin-related domain of Aip5 (Aip5C-S) is capable of attenuating pointed-end depolymerization in bulk assays using pyrene-labeled actin filaments with capped barbed ends. Consistent with pointed-end capping, Aip5C-S slowed pointed-end depolymerization in a concentration-dependent manner, with a Kapp of ∼120 nM (Fig. 6, A and B). Similar results were observed for Aip5C-S and Aip5C-L, which differ only by the inclusion of the BBS, and pointed-end capping was abolished in the actin-binding deficient Aip5C-L 3M mutant (Fig. 6, C and D). Aip5C-S also failed to cosediment with F-actin in high-speed copelleting assays (Fig. S5 C), further indicating that its ability to slow pointed-end depolymerization arises from an interaction with filament ends rather than sides. The effects of Aip5C on pointed-end depolymerization of yeast actin filaments were even stronger (Kapp ∼ 11 nM) (Fig. 6 E), consistent with the stronger nucleation effects of Aip5C using yeast actin (Fig. S1, D and E) and with the view that Aip5 binds with higher affinity to yeast actin than vertebrate actin. We also confirmed that the actin-binding deficient mutant Aip5C-L(3M) fails to block pointed-end depolymerization of yeast actin filaments (Fig. S5 D).
Figure 6.
Aip5 caps the pointed ends of actin filaments in vitro. (A) Kinetics of F-actin pointed-end depolymerization in bulk assays. Reactions contain 2 μM F-actin (10% pyrene-labeled) capped at their barbed ends with 40 nM CapZ. Depolymerization was induced at time 0 by the addition of 3 µM vitamin D–binding protein in the presence or absence of variable concentrations of Aip5C-S (colored solid lines). Control reactions contained no Aip5C-S. Depolymerization curves were each fit to a single exponential (dashed lines) to determine decay rates. Data shown are from one of three independent experiments. (B) Actin filament depolymerization decay rates. Data from multiple experiments (n = 3) were plotted as a function of Aip5C-S concentration and fit to a single-site–binding isotherm to determine Kapp (see Materials and methods). Error bars (SD) are present, but are very small and cannot be readily seen on this scale. (C) Pointed-end depolymerization rates for actin filaments, analyzed as in A, in the presence or absence of the indicated Aip5C polypeptides. Control reactions contained no Aip5 polypeptides. Data shown are from one of three independent experiments. (D) Mean decay rates from three independent experiments (n = 3) of various Aip5C constructs. Error bars, SD. Significant differences between all conditions were calculated using one-way ANOVA followed by Tukey’s post hoc test. All comparisons were significantly different from one another except for the two comparisons displayed in the figure, which were not significantly different, ns, P > 0.05. (E) Yeast actin filament depolymerization decay rates. Data are plotted as a function of Aip5C-S concentration and fit to a single-site–binding isotherm to determine Kapp. Reactions contain filaments polymerized from 2 µM yeast actin (with 10% pyrene-labeled vertebrate muscle actin) in the presence of 40 nM CapZ to cap barbed ends. Depolymerization of F-actin was induced at time 0 by the addition of 3 µM vitamin D–binding protein in the presence or absence of variable concentrations of Aip5C-S. (F) Single-molecule TIRF imaging shows that Aip5C-L-SNAP549 decorates the pointed ends of yeast actin filaments that elongate at their barbed ends. Reaction contains 1 μM yeast G-actin (with 10% Alexa 488–labeled and 0.5% biotin-labeled vertebrate muscle actin), 5 μM yeast profilin, and 5 nM Aip5C-L-SNAP549. Yellow arrowhead, barbed end, as inferred from the growth rate. Scale bar, 5 μm. Kymograph (to the right) shows Aip5C-L-SNAP549 remaining at the pointed end of a filament, while the barbed end grows over time. BE, barbed end. PE, pointed end. Data shown are from one of three independent experiments. Scale bar, 2 μm. (G) Three-color TIRF imaging demonstrates that Aip5C-L-SNAP549 (magenta) decorates the pointed ends of yeast actin filaments (green), while SNAP649-CapZ (blue) decorates the opposite (barbed) end. Filaments were first assembled using 1 μM yeast G-actin (with 10% Alexa 488–labeled and 0.5% biotin-labeled vertebrate muscle actin), 5 μM yeast profilin, and 5 nM Aip5C-L-SNAP549, and then, 5 nM CapZ was added. Scale bar, 1 μm.
Finally, we asked whether we could directly visualize labeled Aip5C-L-SNAP549 molecules at the pointed ends of actin filaments using in vitro single-molecule TIRF imaging. In these experiments, we used filaments assembled from yeast actin, because of the higher affinity of Aip5 for yeast actin. We found that filaments assembled in the presence of Aip5C-L-SNAP549 remained stably decorated at one end of the filament by Aip5C-L-SNAP549, while the opposite end of the filament elongated over time at a rate (∼6 subunits/s) consistent with barbed-end growth (Fig. 6 F and Video 8). In another set of experiments, if we first assembled filaments in the presence of Aip5C-L-SNAP549 and then flowed in SNAP649-CapZ to cap barbed ends, we observed filaments decorated at one end by Aip5C-L-SNAP549 and the other end by SNAP649-CapZ (Fig. 6 G).
Video 8.
Related to Fig. 6 F : Single-molecule TIRF imaging reveals labeled Aip5C-L-SNAP 549 remaining associated with the pointed ends of yeast actin filaments after nucleation, while barbed ends grow. Reaction contains 1 μM yeast G-actin (with 10% Alexa 488–labeled vertebrate muscle actin), 5 µM yeast profilin, and 5 nM Aip5C-L-SNAP549. Video playback is 10 frames per second, and is sped up 100-fold from real time, with a timestamp. Scale bar, 5 μm.
Discussion
Our findings reveal that Aip5 and Bud6 form a novel “composite” actin nucleator in budding yeast that collaborates with formins to assemble robust actin cables (see the working model in Fig. 7 A). Through a combination of structural modeling, biochemical analysis, and in vivo functional characterization, we dissected the roles of Aip5 and Bud6 in the composite nucleation mechanism. We found that the thioredoxin-related domain of Aip5 caps and stabilizes the pointed end of the actin seed, and remains associated with the pointed end of the filament as its barbed end grows, while Bud6’s C-terminal flank domain contains two actin-binding sites (ABS1 and ABS2) that each bind actin monomers and stabilize the side of the actin seed (Fig. 7 B). Aip5 and Bud6 also directly recruit formins to cap the barbed ends of the F-actin seeds they generate, promoting processive elongation of the barbed ends even in the presence of capping protein. Importantly, we have shown that the interactions of Aip5 and Bud6 with each other and with actin are all essential in vivo for maintaining normal actin cable thickness and preventing aberrant cable detachment from the bud neck to ensure proper vesicle trafficking (Fig. 7 C). Interestingly, the cable defects in aip5Δ bud6Δ double mutants are more severe than in aip5Δ or bud6Δ single mutants. This can be explained by the respective nucleation-promoting factor (NPF) activities of Aip5 and Bud6 on formins. Aip5 and Bud6 can each enhance actin nucleation in the presence of formins, and these NPF effects require their direct interactions with actin and with formins (Graziano et al., 2011, 2013; Glomb et al., 2019; Xie et al., 2019; Xie et al., 2022). The overall picture this paints is that the composite nucleation mechanism we have uncovered here involving Aip5-Bud6 interactions functions intimately with formins (as depicted in Fig. 7 B). As a result of this complex network of interactions, either component alone (Aip5 or Bud6) can promote actin nucleation in the presence of formins, and thus, loss of a single component (Aip5 or Bud6) results in less severe cable defects compared with loss of both components.
Figure 7.
Working model for actin cable formation by the Aip5-Bud6 composite nucleation mechanism and its collaboration with formins. (A) At polarity sites, actin filament assembly is initiated by the Aip5-Bud6 composite nucleator working in concert with formins Bni1 (bud tip) or Bnr1 (bud neck). The cables assembled at the bud tip are released from the cell cortex, with Bni1 on filament barbed ends and Aip5 on pointed ends. Cables assembled at the bud neck grow while attached to Bnr1 formins, which are anchored to septins at the bud neck. Cables in the mother cell compartment are decorated with Aip5 at their pointed ends. Cables undergo retrograde flow, moving from polarity sites toward the back of the mother cell at a rate of ∼0.4 µm/s. (B) Proposed molecular mechanism for Aip5, Bud6, and formins collaborating to assemble actin filaments, depicted in four steps (left to right): actin monomer recruitment, filament seeding, formin recruitment, and filament elongation and capping. Arrow in Step 1 indicates Bud6-Aip5 association. Arrow in Step 3 indicates formin FH2 domain associating with the barbed end of the F-actin seed nucleated by Aip5 and Bud6. The black arrow in Step 4 indicates Bud6 dissociation from the nucleated F-actin seed, while the larger gray arrow indicates the direction of filament polymerization. (C) Comparison of actin cable architecture and function in wild-type and aip5Δ bud6Δ cells. In wild-type cells, cables grow in a retrograde manner from the bud tip and bud neck toward the back of the mother cell at a rate of ∼ 0.4 µm/s, while secretory vesicles are transported by myosin V (Myo2) toward the bud tip at ∼3 µm/s. Cables in aip5Δ bud6Δ cells are much thinner compared with wild-type cells, and prematurely detach from the bud neck, leading to defects in vesicle transport and an accumulation of vesicles in the mother cell.
The composite nature of the Aip5-Bud6 nucleation mechanism distinguishes it from other known nucleators that recruit and organize actin monomers (Chesarone and Goode, 2009; Campellone and Welch, 2010; Dominguez, 2016). Unlike other type III nucleators (e.g., Spire, COBL, Lmod, and APC), which each nucleate autonomously, the Aip5-Bud6 nucleator requires two separate proteins to interact for effective nucleation. Interestingly, this two-component system is conceptually related to how the Arp2/3 complex and its WASP family cofactors nucleate actin assembly. In both systems, one component (Aip5 or Arp2/3) caps pointed ends, while the other component (Bud6 or WASP) uses WH2 or WH2-like domains to bind actin monomers. Furthermore, in both cases the non-capping component (Bud6 or WASP) has high affinity for actin monomers but does not bind F-actin, and thus dissociates after formation of the filament, whereas the capping component (Aip5 or Arp2/3) remains on the pointed end of the newly assembled filament. The primary difference is that whereas Aip5-Bud6 produces linear filaments, Arp2/3-WASP primarily produces branched networks. The bipartite architecture of the Aip5-Bud6 nucleation apparatus may allow for tighter spatial and temporal regulation of actin nucleation, as cells can control when and where Aip5 and Bud6 interact to form active nucleation complexes. Indeed, our localization studies show that Aip5 and Bud6 overlap primarily at the bud tip and bud neck, which is precisely where formins are recruited and actin cable assembly is initiated.
Our live-imaging experiments using an enhanced (3xmNG) tag revealed a previously unrecognized dynamic behavior for Aip5, in which Aip5 puncta stream directionally inward from polarity sites (bud cortex and bud neck) with actin cable retrograde flow. The passive transport on actin cables is abolished by latrunculin treatment, and requires direct binding of Aip5 to actin, as demonstrated by the Aip5-3M mutant that disrupts its pointed-end actin-binding interface. Further, the movements are independent of Aip5 interactions with Bud6, Bni1, or Bnr1, providing further support for the Aip5–actin cable association being direct. This association of Aip5 with the pointed ends of filaments could help stabilize cables during retrograde flow, perhaps protecting them from high concentrations of disassembly-promoting factors in the yeast cytosol (Gonzalez Rodriguez et al., 2023). Another consequence of the Aip5 association with cables may be its depletion from polarity sites, which could serve as a negative feedback mechanism for cable assembly. In this scenario, more nucleation events would lead to more Aip5 molecules incorporated into cables, depleting the pool of free Aip5 available to join Bud6 in nucleating actin assembly at polarity sites. Conversely, physiological conditions or fluctuations that reduce cable levels would release Aip5 and increase the frequency of nucleation events, promoting cable growth.
In contrast to Aip5, we found that Bud6 did not move inward with actin cables, in agreement with earlier studies (Buttery et al., 2007). These observations suggest a “nucleate and release” model where Bud6 dissociates from the F-actin seed upon nucleation, while Aip5 remains associated with the pointed end of the growing filament (Fig. 7 B). This model is further supported by our structural analysis showing that Bud6’s actin-binding interfaces would be partially displaced when G-actin polymerizes into F-actin. The first alpha helix in the WH2-related domain of Bud6 binds to the hydrophobic pocket between subdomains I and III in G-actin (Park et al., 2015), and would be displaced by the D-loop of the adjacent actin subunit when actin polymerizes into filament. This clash also explains why C-Bud6 (489–788) exhibits high affinity for G-actin but no detectable affinity for F-actin (Moseley et al., 2004; Graziano et al., 2011).
To our knowledge, Aip5 represents the first identified pointed-end capper in the fungal kingdom, aside from the Arp2/3 complex. A structural similarity search revealed that the actin-binding thioredoxin-related domain of Aip5 closely resembles that of vertebrate SH3BGRL2 (Fig. S3 E). Moreover, in our structural model Aip5 occupies a nearly identical position at the pointed end of the actin filament to the position of SH3BGRL2 at pointed ends observed in a recent cryo-EM study on cortical actin networks in red blood cells (Fig. S3 F) (Li et al., 2023). During the preparation of our manuscript, a preprint reported that SH3BGRL2 caps actin filament pointed ends in vitro (Heiringhoff et al., 2025, Preprint). This structural conservation from yeast to humans suggests an evolutionarily conserved role for at least a subset of thioredoxin-related domains in capping actin filament pointed ends across diverse cellular contexts. Interestingly, in the above-mentioned cryo-EM study (Li et al., 2023), SH3BGRL2 and tropomodulin (Tmod) were found together at the same pointed ends of filaments, and showed non-overlapping footprints with actin at the pointed end. This raises the possibility that other proteins might be capable of joining Aip5 at the pointed end in fungi, although there is no obvious Tmod homolog in fungal species.
The physiological significance of the Aip5-Bud6 composite actin nucleation mechanism is demonstrated by previous studies showing that aip5Δ and bud6Δ mutants have severely diminished actin cable networks (Graziano et al., 2011, 2013; Glomb et al., 2019; Xie et al., 2019; Xie et al., 2022). Our new results build upon these previous findings and demonstrate that the most pronounced defects in cables observed in aip5Δ and bud6Δ cells are reduced cross-sectional thickness of cables. Further, we use live imaging to show that these architectural defects in cables correlate with aberrant cable detachment from the bud neck and impaired secretory vesicle trafficking (Fig. 7 C). Thus, robust actin cable nucleation by Aip5 and Bud6 is required to maintain the normal density of actin filaments in a cable bundle. In turn, cable density (or thickness) appears to be critical for maintaining cable attachment to the bud neck, where the formin Bnr1 is anchored. Thinner cables may detach more readily as a result of having fewer filaments in them, and therefore, fewer attachment points to anchored Bnr1 at the bud neck. This might lead to cables detaching more readily under normal force loads, such as from myosin II (at the bud neck) or myosin V (along cables), and/or be prematurely severed or depolymerized.
In conclusion, our work identifies the Aip5-Bud6 complex as a new class of composite actin nucleator that directly collaborates with formins to assemble actin cables. Our work also reveals that Aip5 remains on the pointed ends of filaments as a capper and is passively transported toward the back of cells by actin cable retrograde flow. These findings expand the known diversity of actin nucleation mechanisms and establish a more direct evolutionary connection between yeast and metazoan cytoskeletal regulation than previously appreciated. Future studies will undoubtedly reveal more precisely how Aip5-Bud6 interactions are spatiotemporally regulated in cells and whether additional factors contribute to formin-mediated actin cable assembly in the absence of this composite nucleator.
Materials and methods
Plasmids and yeast strains
Yeast strain construction
All yeast strains used in this study were in the W303 background (his3-11,15; ura3-52; leu2-3,112; ade2-1; trp1-1; psi+) (listed in Table S1). Integrations were generated using the Longtine PCR method (Longtine et al., 1998). Aip5-mNG::HIS3 (BGY4427), aip5-3M-mNG::URA3 (BGY4428), aip5ΔBBS-mNG::URA3 (BGY4429), Aip5-3xmNG::HIS3 (BGY4432), aip5-3M-3xmNG::HIS3 (BGY4440), aip5ΔBBS-3xmNG::HIS3 (BGY4441), Bni1-3xmNG::URA3 (BGY4433), Bud6-3xmNG::HIS3 (BGY4434), Bud6-mNG::HIS3 (BGY4438), and Bud6Δ740-788-mNG::HIS3 (BGY4439) were integrated into wild-type W303 background cells. Single-gene deletion strains aip5Δ::URA3 (BGY4416), aip5Δ::HIS3 (BGY4417), and bud6Δ::TRP1 (BGY1418) were generated by placing the entire coding regions with indicated auxotrophic markers. BUD6 strains bud6-3xHA::HIS3 (BGY1410), bud6-35-3xHA::HIS3 (BGY1411), and bud6-8-3xHA::HIS3 (BGY1412) were constructed. We first integrated an in-frame 3xHA tag (marked with HIS3) at the 3′ end of the BUD6 open reading frame into a wild-type w303 yeast strain, generating BUD6-3xHA::HIS3 (BGY1410). Next, the BUD6-3xHA::HIS3 ORF plus 300 base pairs 5′ UTR and 200 base pairs 3′ UTR was PCR-amplified from genomic DNA and cloned into the pRS315 vector using XhoI and NotI sites. This plasmid was used as a template for site-directed mutagenesis to generate integration vectors for bud6-35 and bud6-8. For integrations, DNA fragments encoding for amino acids 550–788 and the 3xHA tag, along with 200 base pairs 3′ UTR and the marker, were amplified from the plasmids and transformed into BGY12, producing BGY1411 (bud6-35-3xHA::HIS3) and BGY1412 (bud6-8-3xHA::HIS3) (Graziano et al., 2011). Crosses and tetrad dissection yielded the following haploid strains: aip5Δ::URA3 bud6Δ::TRP1 (BGY4418), aip5Δ::HIS3 bud6Δ::TRP1 (BGY4419), aip5Δ::URA3 bud6-3xHA::HIS3 (BGY4420), aip5Δ::URA3 bud6-35-3xHA::HIS3 (BGY4421), aip5Δ::URA3 bud6-8-3xHA::HIS3 (BGY4422), Aip5-3xmNG::HIS3 bud6Δ::TRP1 (BGY4435), Aip5-3xmNG::HIS3 bni1Δ::TRP1 (BGY4436), and Aip5-3xmNG::HIS3 bnr1Δ::TRP1 (BGY4437). To visualize F-actin in live cells, LifeAct-mNG::HIS3 (BGY4346) was crossed to aip5Δ::HIS3 bud6Δ::TRP1 to yield the following haploid strains: LifeAct-mNG::HIS3 aip5Δ::HIS3 (BGY4423), LifeAct-mNG::HIS3 bud6Δ::TRP1 (BGY4424), LifeAct-mNG::HIS3 aip5Δ::HIS3 bud6Δ::TRP1 (BGY4425). For live imaging of secretory vesicles, we transformed yeast with CEN plasmids that express GFP-SEC4 (Rands and Goode, 2021).
Plasmid construction
All plasmids and DNA oligos used in this study are listed in Tables S2 and S3, respectively. To generate the aip5-3M-mNG and aip5-3M-3xmNG yeast strains, custom pFA6A plasmids pBG2682 (aip5-3M-mNG) and pBG2706 (aip5-3M-3xmNG) were made by inserting a gBlocks Gene Fragment (Integrated DNA Technologies) that encodes residues 1,000–1,234 of Aip5 with three point mutations L1150A, N1162A, and S1165A. To generate aip5ΔBBS-mNG and aip5ΔBBS-3xmNG yeast strains, custom pFA6A plasmids pBG2683 (aip5ΔBBS-mNG) and pBG2705 (aip5ΔBBS-3xmNG) were made by inserting a gBlocks Gene Fragment (Integrated DNA Technologies) containing the C-terminal fragment of Aip5 (941-end) with a flexible linker (GSGS) in place of Aip5 residues 1,000–1,125. For E. coli expression and purification of MBP-Bud6-FL-6xHis, we used a plasmid (pBG2343) generated by cloning the entire Bud6 coding sequence into a pMALc2 backbone with a C-terminal 6xHis tag (Magliozzi et al., 2024). For E. coli expression and purification of MBP-Bud6Δ740-788-6xHis, we generated a new plasmid (pBG2699) by cloning Bud6 residues 1–739 into a modified pMALc2 backbone with a C-terminal 6xHis tag. Bud6C-L (pBG811) was made by cloning Bud6 residues 489–788 into a pET-GST-TEV backbone (Moseley et al., 2004). Bud6flank (pBG1075) was made by cloning residues 699–788 into a modified pET-30a backbone with an N-terminal GST-fusion followed by TEV recognition site (Park et al., 2015). MBP-Bud6-FL-SNAP-6xHis (pBG2684) was made by cloning the full-length Bud6 coding sequence into a modified pMALc2 backbone with a C-terminal 6xHis tag. 6xHis-Bud6(1–145)-KCK (pBG2707) was made by inserting a gBlocks Gene Fragment (Integrated DNA Technologies) encoding Bud6 residues 1–145 followed by a KCK sequence into a pET28a backbone, introducing a single cysteine for fluorescent maleimide labeling. Plasmids for E. coli expression of Aip5C-L (997–1,234) (pBG2687), Aip5C-L-3M (pBG2692), and Aip5C-S (1,110–1,234) (pBG2688) were generated by inserting sequences into a modified pRSF-DUET-1 backbone with an N-terminal PreScission protease cleavable Strep-tag (Wirshing et al., 2023). The Aip5C-L-SNAP-6xHis expression plasmid (pBG2690) was created by inserting Aip5C-L (997–1,234) into a modified pET28a backbone such that only the C-terminal 6xHis tag was fused to Aip5. The Bud6flank-GGS-Aip5C-S chimera plasmid (pBG2700) was made by inserting a gBlocks Gene Fragment (Integrated DNA Technologies) composed of Bud6 residues 699–788, GGS as a flexible linker, and Aip5C-S (1,110–1,234) into a modified pET28a backbone such that the N-terminal cleavable 6xHis tag was fused to the chimera. The FH1-FH2-tail fragment of formin C-Bni1 (FH1-FH2-tail, 1,227–1,953) was subcloned into an E. coli expression vector with N-terminal hexahistidine (6xHis) followed by a small ubiquitin-related modifier (SUMO) tag in the pET-23b plasmid backbone, generating plasmid 6xHIS-SUMO-Bni1-(1,227–1,953) (pBG1936) (Wirshing et al., 2023). For yeast Cap1/2 expression in E. coli, we used an existing plasmid (pBG2643) (Wirshing et al., 2023). To generate this plasmid, CAP1 and CAP2 genes were PCR-amplified from the yeast genome using primers that introduce a Strep-tag and PreScission protease cleavage sequence at the N terminus of Cap2, and these DNA fragments were cloned into separate sites in the E. coli vector pRSFDuet-1.
Fixed-cell imaging and analysis
To analyze actin cable organization, yeast strains were grown in synthetic complete media (SCM) to OD600 0.4–0.6, fixed in 4% formaldehyde for 45 min at room temperature, and then pelleted and washed twice with 1× PBS (pH 7.3). Cells were resuspended in 1× PBS (pH 7.3) + 0.1% Triton X-100 buffer and stained overnight with Alexa Fluor 488 phalloidin or Alexa Fluor 549 phalloidin (Life Technologies). Next, cells were washed in 1× PBS (pH 7.3), resuspended with VectaShield mounting media (Vector Laboratories), mounted onto slides, and imaged at room temperature on a Nikon Ti2-E inverted confocal microscope equipped with a CSU-W1 SoRa (Yokogawa) and a Prime BSI sCMOS camera (Teledyne Photometrics) controlled by Nikon NIS-Elements Advanced Research software using a 60× oil objective (NA 1.4; Nikon Instruments). 3D stacks through the entire cell were acquired at 0.2 μm steps using an exposure time of 500 ms at 488-nm excitation and 40% laser power or 500 ms at 561-nm excitation and 50% laser power. Images were denoised using “Denoise.ai” plugin in NIS-Elements Advanced Research software (Nikon) and background-subtracted using the average mean intensity from five manually chosen areas in the image that do not contain cells.
For counting actin cable number, a “maximum intensity” projection was generated for each acquired Z-stack, and a straight line was drawn across the widest part of the mother cell. A line-scan measurement fluorescence profile of the line was quantified in FIJI (National Institutes of Health, Bethesda, MD, USA). The line-scan profile was saved and input into a Python script designed to count actin cables from the peaks of the line-scan “cable_enumeration.py” and analyzed using a peak height threshold of 0.3 to filter out indistinguishable peaks (McInally et al., 2024). Actin cable cross-sectional fluorescence (thickness) was measured from line scans of ”sum intensity” projections of the original Z-stacks. Line scans were analyzed using a Python script adapted from the cable counting script altered to calculate area under line scans (summing the total intensity of the line-scan fluorescence profile). For quantifying actin cable length, each individual cable was manually traced in FIJI (National Institutes of Health), from the bud neck to their terminus in the mother cell using maximum intensity projections (McInally et al., 2022). A 3D stack was used to differentiate between cables that overlapped and to precisely determine both the origins and distal tips of the cables. For cable length analysis, we included every detectable cable in the cell that extended from the bud neck to some endpoint in the mother cell compartment; the only cables excluded from the analysis were a minor fraction that became closely intertwined with other cables making it impossible to resolve their individual lengths or cables detached from the bud neck.
To compare whole-cell F-actin fluorescence intensities (phalloidin staining), the different strains were grown in parallel, fixed, stained with Alexa 488-phalloidin, and imaged. Cell outlines were manually traced, and the fluorescence levels for each cell (raw integrated densities) were divided by cell area (µm2) in FIJI (National Institutes of Health). For comparing cortical actin patch intensities, ROIs were drawn around each resolvable actin patch in a cell, and the fluorescence raw integrated densities were measured. Patches were manually traced with the circle function in FIJI (National Institutes of Health).
Live-cell imaging and analysis
For time-lapse imaging of Aip5-3xmNG, Bni1-3xmNG, and Bud6-3xmNG puncta, cells were grown to a log phase in SCM at 25°C. Cells were then mounted directly onto glass slides and immediately imaged at room temperature in SCM using a CSU-W1 SoRa (Yokogawa) and a Prime BSI sCMOS camera (Teledyne Photometrics) controlled by Nikon NIS-Elements Advanced Research software using a 100× oil objective (NA 1.4; Nikon Instruments), exposure time of 200 ms at 488-nm excitation, and 60% laser power. Single focal plane images were acquired for 60 s at 300-ms intervals using exposure times of 200 ms at 60% intensity (488-nm excitation). For quantifying the velocities of Bni1-3xmNG and Aip5-3xmNG retrograde movements, puncta were traced from the time of their detachment from the cell cortex and for at least 1 μm of retrograde movement from the bud tip toward the back of the mother cell. For acquiring still images of strains expressing Aip5-3xmNG, Bud6-3xmNG, or Bni1-3xmNG, cells were grown as above and 3D stacks through the entire cell were acquired at 0.3 μm steps using an exposure time of 500 ms at 488-nm excitation and 50% laser power. For the LatB treatment experiments, the indicated yeast strains were grown as above, then treated with either DMSO or 200 μM LatB in DMSO for 10 min at 25°C prior to imaging. Images were analyzed in FIJI (National Institutes of Health).
For imaging Bud6-mNG, Bud6Δ740-788-mNG, Aip5-mNG, Aip5-3M-mNG, and Aip5ΔBBS-mNG, cells were grown to a log phase in SCM at 25°C. Cells were then mounted directly onto glass slides and immediately imaged at room temperature in SCM using a CSU-W1 SoRa (Yokogawa) and a Prime BSI sCMOS camera (Teledyne Photometrics) controlled by Nikon NIS-Elements Advanced Research software using a 100× oil objective (NA 1.4; Nikon Instruments). 3D stacks through the entire cell were acquired at 0.3 μm steps using an exposure time of 500 ms at 488-nm excitation and 50% laser power. Aip5-3xmNG whole-cell fluorescence intensities were quantified by measuring fluorescence raw integrated densities. Cell outlines were manually traced, and the fluorescence levels for each cell were divided by its cell area (µm2) in FIJI (National Institutes of Health).
For live imaging of actin cables, cells expressing integrated LifeAct-mNG were grown to a log phase in SCM at 25°C. Cells were then mounted onto SCM 1% agarose pads and imaged at room temperature using a CSU-W1 SoRa (Yokogawa) and a Prime BSI sCMOS camera (Teledyne Photometrics) controlled by Nikon NIS-Elements Advanced Research software using a 100× oil objective (NA 1.4; Nikon Instruments), exposure time of 50 ms at 488-nm excitation, and 20% laser power. 3D stacks through the top half of the cell were acquired with 5 × 0.5 μm steps with no delay between acquisitions, and this cycle was repeated for a total duration of 60 s to measure cable extension. Cable detachment events were tracked manually using maximum intensity Z-projection image videos and scored based on observing the cable extending into the mother cell and then detaching from the bud neck. Images were analyzed in FIJI (National Institutes of Health).
For counting secretory vesicles in mother cell compartments, yeast transformed with a CEN plasmid expressing GFP-Sec4 were grown to a log phase in synthetic selective media at 25°C. Cells were then then mounted onto synthetic selective medium 1% agarose pads and imaged at room temperature on a Nikon Ti2-E inverted confocal microscope equipped with a CSU-W1 SoRa (Yokogawa) and a Prime BSI sCMOS camera (Teledyne Photometrics) controlled by Nikon NIS-Elements Advanced Research software using a 100× oil objective (NA 1.4; Nikon Instruments), using an exposure time of 500 ms at 488-nm excitation and 40% laser power. 3D stacks through the entire cell were acquired at 0.2 μm steps. FIJI was used to generate maximum projection images, and then, GFP-Sec4 vesicles in the mother cell compartment were manually counted.
Live time-lapse imaging of yeast expressing GFP-Sec4 was conducted by growing and mounting cells as mentioned above for counting secretory vesicles. Cells were imaged at room temperature on a Nikon Ti2-E inverted confocal microscope equipped with a CSU-W1 SoRa (Yokogawa) and a Prime BSI sCMOS camera (Teledyne Photometrics) controlled by Nikon NIS-Elements Advanced Research software using a 100× oil objective (NA 1.4; Nikon Instruments), using an exposure time of 100 ms at 488-nm excitation and 55% laser power. Single focal plane images were acquired at 0.2-s intervals. Images were analyzed in FIJI (National Institutes of Health).
Protein purification
Vertebrate muscle actin purification
Rabbit skeletal muscle actin was purified from acetone powder (Spudich and Watt, 1971) generated from frozen ground hind leg muscle tissue of young rabbits (Pel-Freez Biologicals). Lyophilized acetone powder stored at −80°C was mechanically sheared in a coffee grinder, resuspended in G-buffer (5 mM Tris-HCl, pH 8.0, 0.2 mM ATP, 0.5 mM dithiothreitol [DTT], 0.1 mM CaCl2), and then cleared by centrifugation for 20 min at 50,000 × g, 4°C. The supernatant was filtered through Grade 1 Whatman paper, and then, the actin was polymerized by the addition of 2 mM MgCl2 and 50 mM KCl to the filtrate and overnight incubation at 4°C with slow stirring. The next morning, KCl powder was added to a final 0.6 M, and stirring was continued for another 30 min at 4°C. F-actin was pelleted by centrifugation for 150 min at 120,000 × g, 4°C. The pellet was solubilized by Dounce homogenization and dialyzed against 1 liter of G-buffer at 4°C (three consecutive times at 12- to 18-h intervals). Monomeric actin was then precleared for 30 min at 435,000 × g, 4°C, and loaded onto a HiPrep 16/60 Sephacryl S-200 HR gel-filtration column (GE Healthcare). Peak fractions containing actin were stored at 4°C.
For the fluorescently labeled actin used in open-flow cell TIRF microscopy assays, actin was labeled on cysteine 374 (Kuhn and Pollard, 2005). Briefly, the F-actin pellet described above was Dounce-homogenized and dialyzed against G-buffer lacking DTT. Monomeric actin was then polymerized by adding an equal volume of 2× labeling buffer (50 mM imidazole, pH 7.5, 200 mM KCl, 0.3 mM ATP, 4 mM MgCl2). After 5 min, the actin was mixed with a fivefold molar excess of Alexa 488-maleimide (Thermo Fisher Scientific), resuspended in anhydrous dimethylformamide, and incubated in the dark for 15 h at 4°C. Labeled F-actin was pelleted as above, and the pellet was rinsed briefly with G-buffer, then depolymerized by Dounce homogenization, and dialyzed against G-buffer for 2 days at 4°C. Labeled, monomeric actin was purified further on a HiPrep 16/60 Sephacryl S-200 HR gel-filtration column as above. Alexa 488-actin was dialyzed for 15 h against G-buffer with 50% glycerol and stored at −20°C. The concentration and labeling efficiency were determined by measuring the absorbance at 280 and 494 nm, using these molar extinction coefficients: ε290 actin = 26,600 M−1 cm−1, ε494 Alexa 488 = 71,000 M−1 cm−1, and Alexa 488 correction factor at 290 = 0.11.
For preparing biotinylated actin used in open-flow cell TIRF microscopy assays, the F-actin pellet above was Dounce-homogenized and dialyzed against G-buffer lacking DTT. Monomeric actin was then polymerized by the addition of an equal volume of 2× labeling buffer (50 mM imidazole, pH 7.5, 200 mM KCl, 0.3 mM ATP, and 4 mM MgCl2). After 5 min, the actin was mixed with a fivefold molar excess of NHS-XX-Biotin (Merck KGaA) and incubated for 15 h at 4°C. The F-actin was pelleted as above, and the pellet was rinsed with G-buffer, then homogenized with a Dounce homogenizer, and dialyzed against G-buffer for 48 h at 4°C. Biotinylated monomeric actin was purified further on an S200 (16/60) gel-filtration column as above. Aliquots of biotin actin were snap-frozen in liquid N2 and stored at −80°C.
For bulk pyrene-actin assembly assays, actin was labeled with pyrenyl-iodoacetamide on cysteine 374 (Pollard and Cooper, 1984). Actin prepared as above, excluding the gel-filtration step, was dialyzed against pyrene buffer (25 mM Tris-HCl, pH 7.5, 100 mM KCl, 0.02% NaN3, 0.3 mM ATP, and 2 mM MgSO4) for 3 h, then diluted with pyrene buffer to 1 mg/ml (23.8 μM). A sevenfold molar excess of pyrenyl-iodoacetamide was added, and this actin solution was incubated overnight at 4°C. Next, large aggregates were cleared by low-speed centrifugation, and the supernatant (containing F-actin) was centrifuged for 3 h at 45,000 rpm in a rotor (Ti70; Beckman Coulter) at 4°C. The F-actin pellets were mechanically disrupted using a Dounce homogenizer, then dialyzed against G-buffer for 1–2 days, and finally gel-filtered on a 16/60 S200 column as above. Peak fractions were pooled, aliquoted, snap-frozen, and stored at −80°C until use.
Saccharomyces cerevisiae actin purification
Saccharomyces cerevisiae actin was purified from commercial baker’s yeast (Red Star Yeast) using DNase I affinity and Mono Q (GE Healthcare) chromatography as follows (Goode, 2002; Pollard et al., 2020; Wirshing et al., 2023). Briefly, wet yeast bricks (packed cells) were washed with water, flash-frozen in pellets in liquid nitrogen, and then mechanically lysed by grinding into a fine powder while submerged in liquid nitrogen using a Waring blender. Lysed cell powder was stored at −80°C until use. To initiate a preparation, yeast powder was mixed 1:1 (wt:vol) with room temperature G-buffer (5 mM Tris-HCl, pH 8.0, 0.2 mM ATP, 0.1 mM CaCl2, and 0.5 mM DTT) supplemented with protease inhibitors (0.5 μg/ml each of leupeptin, aprotinin, antipain, chymostatin, and pepstatin A, and 1 mM phenylmethylsulfonyl fluoride [PMSF]) and then centrifuged at 215,000 × g for 1 h in a Ti60 rotor (Beckman). Supernatants were loaded onto a 20 ml DNase I column. The DNase I column was washed with 10 column volumes of G-buffer, then 2 column volumes of G-buffer plus 10% formamide, then 2 column volumes of G-buffer plus 0.2 M ammonium chloride, and finally 2 column volumes of G-buffer alone. Actin was eluted from the column with 2 column volumes of G-buffer plus 50% formamide and immediately loaded onto a 1-ml Mono Q column on an AKTA fast protein liquid chromatography (FPLC) (GE Healthcare). The Mono Q column was washed with 5 ml of G-buffer, and then, actin was eluted using a 20-ml linear salt gradient (100–300 mM KCl) in G-buffer. Peak fractions containing actin (which eluted at ∼250 mM KCl) were concentrated to 40–45 μM, then dialyzed three times over a 48-h period, each time against 1 liter of G-buffer. Actin was clarified by centrifugation at 373,000 × g for 20 min, aliquoted, flash-frozen, and stored at −80°C.
Profilin purification
Untagged S. cerevisiae profilin (Pfy1) was purified from E. coli as follows (Wirshing et al., 2023; Kaiser et al., 1989). To generate a column, poly-L-proline (P3886; Sigma-Aldrich) was conjugated to CnBr-Sepharose (C9210; Sigma-Aldrich) by suspending 15 g CnBr-Sepharose in 200 ml 1 mM HCl and rocking for 15 min. Swelled CnBr-Sepharose was collected on a sintered glass filter and washed with 500 ml of 1 mM HCl followed by 500 ml of coupling buffer (0.1 M NaHCO3, 0.5 M NaCl). After rinsing, CnBr-Sepharose was resuspended in 100 ml coupling buffer and divided into 25 ml aliquots, and 125 mg poly-L-proline dissolved in coupling buffer was added to each aliquot. Poly-L-proline and CnBr-Sepharose were incubated together overnight at 4°C with rocking. Following incubation, poly-L-proline Sepharose beads were collected by centrifugation and washed with coupling buffer followed by 100 mM Tris-HCl (pH 8.0). Rinsed aliquots were resuspended in ∼40 ml of 100 mM Tris-HCl, pH 8.0, and incubated with rocking at 4°C for 2 h. Poly-L-proline Sepharose beads were then washed three times with 25 ml storage buffer (20 mM Tris-HCl, pH 8.0, 150 mM KCl, 0.2 mM DTT), pooled, and stored at 4°C in storage buffer with 0.01% sodium azide. Untagged Pfy1 was expressed from a plasmid in E. coli strain Rosetta (DE3) with the pRARE rare codon plasmid. Cells were grown to a log phase in Terrific Broth (TB) at 37°C, then induced with 1 mM IPTG for 4 h at 37°C. Cells were harvested by centrifugation, and pellets were stored at −80°C overnight. Cell pellets were resuspended in lysis buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA, 2 mM DTT) supplemented with protease inhibitor cocktail and mechanically lysed by passing through LM20 Microfluidizer (Microfluidics) two times at 18,000 PSI. Following lysis, PMSF (1 mM final) was added, and then, the lysate was clarified by centrifugation at 27,000 × g for 30 min at 4°C. The poly-L-proline Sepharose column was preequilibrated with five column volumes of lysis buffer, and the clarified cell lysate was passed over the column two times. Next, the column was washed two times with 10 column volumes of wash buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA, 2 mM DTT, 100 mM NaCl) and the profilin was eluted using two column volumes of elution buffer (wash buffer plus 8 M urea). Profilin was dialyzed four times sequentially over a 24-h period, each time against 1 liter of storage buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA, 2 mM DTT), then concentrated using an Amicon Ultra centrifugal filter device (3 kDa MWCO) (Sigma-Aldrich), flash-frozen in aliquots, and stored at −80°C.
Purification of Bud6 polypeptides
MBP-Bud6-FL-6xHis and MBP-Bud6Δ740-788-6xHis were expressed in Rosetta 2 BL21 (DE3) E. coli cells carrying the pRARE rare codon plasmid. Cells were grown to OD600 = 0.7–0.9 in TB supplemented with 0.2% glucose, as well as ampicillin and chloramphenicol to maintain selection of the expression plasmid and the pRARE plasmid, respectively. Expression was induced with 0.4 mM IPTG overnight at 18°C, and then, cells were pelleted and stored at −80°C. Cell pellets were thawed, resuspended in lysis buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 5 mM imidazole, pH 8.0, 0.2% Tween-20, and 1 mM DTT) with a protease inhibitor cocktail (0.5 µM each of pepstatin A, antipain, leupeptin, aprotinin, and chymostatin), and then lysed in LM20 Microfluidizer (Microfluidics) at 18,000 psi for 4 total passes of sample. Lysates were then cleared in a Ti60 rotor at 20,000 rpm for 20 min. The supernatant was mixed with 2 ml of Ni-NTA beads (Qiagen) and incubated for 1 h at 4°C with rotation. The beads were then washed three times with 10 ml Bud6 Ni-NTA wash buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 30 mM imidazole, pH 8.0, and 1 mM DTT) in a gravity column (Bio-Rad) at 4°C. MBP-Bud6-FL-6xHis was eluted from the beads using Bud6 Ni-NTA elution buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 300 mM imidazole, pH 8.0, and 1 mM DTT) and collected in 0.5 ml fractions. Peak fractions were pooled, diluted fourfold in Bud6 amylose buffer (50 mM Tris, pH 8.0, 500 mM NaCl, and 1 mM DTT), mixed with 1 ml of amylose beads (New England Biolabs), and rotated at 4°C for 2 h. The beads were then washed three times with 10 ml Bud6 amylose wash buffer (50 mM Tris, pH 8.0, 500 mM NaCl, and 1 mM DTT) in a gravity column at 4°C. MBP-Bud6-FL-6xHis or MBP-Bud6Δ740-788-6xHis was eluted from the beads using Bud6 amylose elution buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 50 mM maltose, and 1 mM DTT), then exchanged into HEKG10D buffer (20 mM HEPES, pH 7.5, 1 mM EDTA, 50 mM KCl, 10% glycerol, and 1 mM DTT) on a PD MiniTrap G-25 desalting column (Cytiva). Peak fractions were pooled, snap-frozen in liquid nitrogen, and stored at −80°C.
To purify and label MBP-Bud6-FL-SNAP647-6xHis, Rosetta 2 BL21 (DE3) E. coli cells carrying the pRARE rare codon plasmid were grown as above for unlabeled MBP-Bud6-FL-6xHis, and a lysate was generated and incubated with Ni-NTA resin as above. After washing the resin, beads were incubated with 10 μM SNAP-surface647 (New England Biolabs) for 18 h with rotation at 4°C and covered in aluminum foil to avoid light exposure. Next, the beads were loaded onto a gravity-flow column and washed with 10 column volumes of Bud6 Ni-NTA wash buffer. Protein was eluted with Bud6 Ni-NTA elution buffer. Peak fractions were pooled, diluted fourfold in Bud6 amylose buffer, and mixed with 1 ml of amylose beads (New England Biolabs) and rotated at 4°C for 2 h. Amylose beads were then washed, eluted, and exchanged in HEKG10D buffer as above. Peak fractions were then pooled, snap-frozen in liquid nitrogen, and stored at −80°C.
Bud6C-L (489–788) was expressed in BL21(DE3) E. coli and purified as follows (Graziano et al., 2013). Cells were grown in TB to a late log phase and induced using 0.4 mM IPTG for 3–4 h at 37°C. Cells were pelleted and frozen at −80°C. Frozen pellets were thawed, resuspended in lysis buffer (50 mM Tris, pH 8.5, 150 mM NaCl, 5 mM EDTA, 1.5% sarkosyl, 5 mM DTT, and standard protease inhibitors), and lysed with lysozyme treatment and sonication. Cell lysates were cleared by centrifugation at 12,000 rpm for 10 min in a Sorvall S600 rotor (Thermo Fisher Scientific). Triton X-100 (final concentration 3.3% [vol/vol]) was added to the supernatant, and the mixture was then mixed with 1 ml preswollen glutathione agarose in PBS (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, and 1.47 mM KH2PO4, pH 7.4). After incubation at 4°C for 3–4 h, beads were washed four times with PBS and then washed twice with HEKD (20 mM HEPES, pH 7.5, 1 mM EDTA, 50 mM KCl, and 1 mM DTT). Bud6C-L was cleaved from GST and released from beads by digestion with TEV protease for 2 h at room temperature, aliquoted, and snap-frozen.
The Bud6flank domain (699–788) was expressed as a GST-fusion in BL21 (DE3) E. coli cells carrying the pLys plasmid. Bacterial cells were grown in TB kanamycin and chloramphenicol to maintain selection of the expression plasmid and the pLys plasmid, respectively. Expression was induced with 0.4 mM IPTG overnight at 18°C, and then, cells were pelleted and stored at −80°C. Frozen pellets were thawed, resuspended in lysis buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 5 mM EDTA, 5 mM DTT, 0.2% TX-100) with a protease inhibitor cocktail (0.5 µM each of pepstatin A, antipain, leupeptin, aprotinin, and chymostatin), and then lysed in LM20 Microfluidizer (Microfluidics) at 18,000 psi for 4 total passes of sample. Cell lysates were cleared by centrifugation at 10,000 rpm for 10 min in a Sorvall S600 rotor (Thermo Fisher Scientific). The supernatant was mixed with 1 ml glutathione resin (Thermo Fisher Scientific) and incubated at 4°C for 2 h with rotation. The beads were then washed with 10 ml wash buffer (50 mM Tris, pH 8.0, 300 mM NaCl, 2 mM DTT) in a gravity column (Bio-Rad) at 4°C. Protein was then eluted with GST elution buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 1 mM EDTA, 1 mM DTT, and 20 mM glutathione), and the GST tag was cleaved by incubation with TEV protease overnight at 4°C. Untagged Bud6-Flank was then separated from free GST by size-exclusion chromatography using a Superdex 75 10/300 Increase GL column (GE Healthcare) equilibrated with HEKG10D buffer. Peak fractions were pooled and incubated for 1 h with fresh Ni-NTA resin (Qiagen) to deplete the His-tagged TEV protease. The protein was concentrated, aliquoted, snap-frozen in liquid nitrogen, and stored at −80°C.
For fluorescence anisotropy assays, a 6xHis-Bud6(1–145)-KCK probe was expressed in BL21 (DE3) E. coli cells carrying the pLys plasmid. Bacterial cells were grown in TB kanamycin and chloramphenicol to maintain selection of the expression plasmid and the pLys plasmid, respectively. Expression was induced with 0.4 mM IPTG overnight at 18°C, and then, cells were pelleted and stored at −80°C. Frozen pellets were thawed, resuspended in lysis buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 1 mM DTT, 20 mM imidazole, pH 8.0, and 0.2% TX-100) with a protease inhibitor cocktail (0.5 µM each of pepstatin A, antipain, leupeptin, aprotinin, and chymostatin), and then lysed in LM20 Microfluidizer (Microfluidics) at 18,000 psi for 4 total passes of sample. Cell lysates were cleared by centrifugation at 10,000 rpm for 10 min in a Sorvall S600 rotor (Thermo Fisher Scientific). The supernatant was mixed with 1 ml Ni-NTA beads (Qiagen) and incubated at 4°C for 1 h with rotation. The beads were then washed three times with 10 ml Bud6 Ni-NTA wash buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 30 mM imidazole, pH 8.0, and 1 mM DTT) in a gravity column (Bio-Rad) at 4°C. 6xHis-Bud6(1–145)-KCK was eluted from the beads using Bud6 Ni-NTA elution buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 300 mM imidazole, pH 8.0, and 1 mM DTT) and collected in 0.5 ml fractions. The peak elution fraction was cleared by centrifugation for 10 min in a microfuge at 15,000 rpm at 4°C and loaded onto a Superdex 75 10/300 Increase GL column (GE Healthcare) equilibrated in HEKG10D buffer. A fivefold molar excess of Alexa 488-maleimide was added to the peak gel-filtration fraction and incubated with rotation overnight at 4°C. The next day, the labeled protein fraction was diluted 10-fold in HEKG10D buffer and incubated with 0.5 ml Ni-NTA beads (Qiagen) at 4°C for 1 h with rotation. The beads were then loaded onto a gravity column (Bio-Rad) at 4°C and washed to remove free dye in the flow-through. The labeled protein was then eluted from the beads using Bud6 Ni-NTA elution buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 300 mM imidazole, pH 8.0, and 1 mM DTT). Protein was then exchanged into HEKG10D buffer (20 mM HEPES, pH 7.5, 1 mM EDTA, 50 mM KCl, 10% glycerol, and 1 mM DTT) on a PD MiniTrap G-25 desalting column (Cytiva). Peak fractions were pooled, snap-frozen in liquid nitrogen, and stored at −80°C.
Purification of Aip5 polypeptides
Unlabeled Aip5C constructs (Aip5C-L, Aip5C-L (3M), and Aip5C-S) were expressed in Rosetta 2 BL21 (DE3) E. coli cells carrying the pRARE rare codon plasmid. Cells were grown to OD600 = 1.0–1.5 in TB with ampicillin and chloramphenicol to maintain selection of the expression plasmid and the pRARE plasmid, respectively. Expression was induced with 0.4 mM IPTG overnight at 18°C, and then, cells were pelleted and stored at −80°C. Cell pellets were thawed, resuspended in Aip5 lysis buffer (50 mM Tris-HCl, pH 8.0, 300 mM NaCl, 1 mM EDTA, and 1 mM DTT) with a protease inhibitor cocktail (0.5 µM each of pepstatin A, antipain, leupeptin, aprotinin, and chymostatin), and then lysed in LM20 Microfluidizer (Microfluidics) at 18,000 psi for 4 total passes of sample. Lysates were cleared in a Ti60 rotor at 20,000 rpm for 20 min. The supernatant was mixed with 1 ml of Strep-Tactin Sepharose beads (IBA Lifesciences) and incubated for 2 h at 4°C with gentle mixing. The beads were then washed with 10 column volumes of Aip5 wash buffer (50 mM Tris, pH 8.0, 300 mM NaCl, and 1 mM DTT) in a gravity column (Bio-Rad) at 4°C. The beads were collected, resuspended in Aip5 storage buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 1 mM EDTA, and 1 mM DTT), and incubated overnight at 4°C with rotation with GST-fused PreScission protease to release Aip5 from the beads. The following day, the Aip5-containing supernatant was incubated with 1 ml glutathione beads (Thermo Fisher Scientific) for 2 h to remove PreScission protease. After the beads were pelleted, the Aip5-containing supernatant was concentrated, aliquoted, snap-frozen in liquid nitrogen, and stored at −80°C.
To purify and label Aip5C-L-SNAP549-6xHis, Rosetta 2 BL21 (DE3) E. coli cells carrying the pRARE rare codon plasmid cells were grown to OD600 = 1.0–1.5 in TB with kanamycin and chloramphenicol to maintain selection of the expression plasmid and the pRARE plasmid, respectively. Expression was induced with 0.4 mM IPTG overnight at 18°C, and then, cells were pelleted and stored at −80°C. For purification, pellets were thawed and resuspended in Aip5 Ni-NTA lysis buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 10 mM imidazole, pH 8.0, and 1 mM DTT) supplemented with a protease inhibitor cocktail (0.5 µM each of pepstatin A, antipain, leupeptin, aprotinin, and chymostatin) and lysed in LM20 Microfluidizer (Microfluidics) at 18,000 psi for 4 total passes of sample. Lysates were then cleared in a Ti60 rotor at 20,000 rpm for 20 min. The supernatant was mixed with 2 ml of Ni-NTA beads (Qiagen) for 1 h at 4°C, with gentle mixing. After washing the resin, beads were incubated overnight with 10 μM SNAP-Surface-549 (New England Biolabs) with rotation at 4°C covered in aluminum foil to avoid light exposure. The beads were then washed with 10 column volumes of Aip5 Ni-NTA wash buffer (50 mM Tris, pH 8.0, 500 mM NaCl, 20 mM imidazole, pH 8.0, and 1 mM DTT) in a gravity column (Bio-Rad) at 4°C. The labeled protein was eluted using Aip5 Ni-NTA elution buffer (50 mM Tris, pH 8.0, 500 NaCl, 500 mM imidazole, pH 8.0, and 1 mM DTT). The peak fraction was cleared by centrifugation for 10 min in a microfuge at 15,000 rpm at 4°C and loaded onto a Superdex 200 10/300 increase GL column (GE Healthcare) equilibrated in Aip5 storage buffer. Peak fractions were pooled, aliquoted, snap-frozen in liquid nitrogen, and stored at −80°C.
Purification of Bud6-Aip5 chimera
6xHis-Bud6Flank-GGS-Aip5C-S chimera was expressed in BL21 (DE3) E. coli cells carrying the pLys plasmid. Cells were grown to OD600 = 1.0–1.5 in TB with kanamycin and chloramphenicol to maintain selection of the expression plasmid and the pLys plasmid, respectively. Expression was induced with 0.4 mM IPTG overnight at 18°C, and then, cells were pelleted and stored at −80°C. Cell pellets were thawed, resuspended in lysis buffer (50 mM Tris-HCl, pH 8.0, 300 mM NaCl, 0.2% Tween-20, 20 mM imidazole, pH 8.0, and 1 mM DTT) with a protease inhibitor cocktail (0.5 µM each of pepstatin A, antipain, leupeptin, aprotinin, and chymostatin), and then lysed in LM20 Microfluidizer (Microfluidics) at 18,000 psi for 4 total passes of sample. Cell lysates were cleared by centrifugation at 10,000 rpm for 20 min in a rotor (Sorvall S600; Thermo Fisher Scientific). The supernatant was mixed with 1 ml of Ni-NTA beads (Qiagen) and incubated for 1 h at 4°C, with gentle mixing. The beads were then washed three times with 10 ml wash buffer (50 mM Tris, pH 8.0, 300 mM NaCl, 30 mM imidazole, pH 8.0, and 1 mM DTT) in a gravity column (Bio-Rad) at 4°C. Protein was eluted from the beads using elution buffer (50 mM Tris, pH 8.0, 300 mM NaCl, 300 mM imidazole, pH 8.0, and 1 mM DTT) and collected in 0.5 ml fractions. The peak eluate fraction was cleared by centrifugation for 10 min in a microfuge at 15,000 rpm at 4°C and loaded onto a Superdex 200 10/300 increase GL column (GE Healthcare) equilibrated in HEKG10D buffer. Peak fractions were pooled, aliquoted, snap-frozen in liquid nitrogen, and stored at −80°C.
Purification of Bni1 polypeptides
The C-Bni1 (FH1-FH2-tail) polypeptide was purified using a 6xHis-SUMO tag strategy as follows (Wirshing et al., 2023). E. coli Rosetta (DE3) cells carrying the pRARE rare codon plasmid and transformed with the formin constructs were grown to a log phase in TB at 37°C and induced with 1 mM IPTG for 16 h at 18°C. Cells were harvested by centrifugation, and pellets were stored at −80°C. Pellets were thawed and resuspended in lysis buffer (30 mM imidazole, pH 8.0, 0.5 mM DTT, and 2× PBS [40 mM sodium phosphate buffer and 200 mM NaCl, pH 7.4]) supplemented with 150 mM NaCl, 1% NP-40, and protease inhibitor cocktail. Next, cells were lysed by sonication on ice, clarified by centrifugation at 27,000 × g for 30 min at 4°C, and mixed with 1 ml Ni-NTA beads (Qiagen) for 1 h at 4°C with gentle mixing. The slurry was transferred to a gravity-flow column, and the agarose was allowed to settle by gravity. The column was washed twice with 10 column volumes of lysis buffer plus 350 mM NaCl and then twice with 10 column volumes of wash buffer (20 mM HEPES, pH 7.5, 100 mM NaCl, 5% glycerol). Formins were eluted using wash buffer plus 250 mM imidazole. The 6xHis-SUMO tag was removed by overnight digestion at 4°C with Ulp1 protease while dialyzing the reaction against 500 ml HEKG10 buffer (20 mM HEPES, pH 7.5, 1 mM EDTA, 50 mM KCl, 10% glycerol) with 2 mM DTT. Following dialysis, formins were further purified by gel filtration on a Superose 6 10/300 Gl column (GE Healthcare) equilibrated in HEKG10D. Peak fractions were pooled, concentrated, aliquoted, snap-frozen, and stored at −80°C.
Purification of yeast Cap1/2
To purify S. cerevisiae capping protein (Cap1/2), Rosetta 2 BL21 (DE3) E. coli cells carrying the pRARE rare codon plasmid cells were grown to OD600 = 1.0–1.5 in TB with ampicillin and chloramphenicol to maintain selection of the expression plasmid and the pRARE plasmid, respectively. Cells were grown to a log phase in TB at 37°C and induced with 1 mM IPTG for 4 h at 37°C. Cells were harvested by centrifugation, and pellets were stored at −80°C. Pellets were resuspended in buffer W (100 mM Tris-HCl, pH 8.0, 150 mM NaCl, and 1 mM EDTA) supplemented with 1 mM DTT and protease inhibitors, then chemically lysed by incubation with 0.1 mg/ml lysozyme on ice for 30 min with intermittent agitation. The cell lysate was clarified by centrifugation at 27,000 × g for 30 min at 4°C and mixed with 5 ml Strep-Tactin Sepharose resin (IBA Lifesciences) equilibrated with buffer W. The resin was transferred to an empty column and allowed to settle by gravity. The lysate was passed over the column three times, and then, the column was washed five times with 10 column volumes of buffer W. Finally, the resin was transferred to a tube with 5 ml of buffer W plus 1 mM DTT. The Strep-Tactin tag was removed by digestion with PreScission protease (GST-tagged) overnight at 4°C, releasing untagged Cap1/2. Following cleavage, the GST-PreScission protease was removed by incubation with 0.5 ml Pierce Glutathione Agarose (Thermo Fisher Scientific) at 4°C for 1 h with rocking. Cap1/2 was concentrated to 60–80 μM, dialyzed against 1 liter storage buffer (20 mM HEPES, pH 7.5, 1 mM EDTA, 50 mM KCl, 1 mM DTT), aliquoted, snap-frozen, and stored at −80°C.
Purification of vertebrate CapZ
A plasmid for expressing vertebrate CapZ was transformed into E. coli strain BL21 pLysS. Cells were grown in LB to a log phase, and then, expression was induced for 3 h at 37°C by the addition of 0.4 mM IPTG. Cells were collected by centrifugation, washed with 25 ml water, and resuspended in lysis buffer (20 mM Tris, pH 8.0, 1 mM EDTA, 0.1% Triton X-100, protease inhibitors) and lysed by lysozyme treatment and sonication. The cell lysate was clarified by centrifugation at 12,500 × g for 30 min at 4°C. Supernatants were loaded onto a 1 ml Q-HiTrap column (GE Healthcare) and eluted with a 45 ml salt gradient (0–500 mM KCl) in 20 mM Tris, pH 8.0. Peak fractions were pooled, concentrated to 3 ml, and loaded onto a 26/60 Superdex 75 gel-filtration column (GE Healthcare) equilibrated in 50 mM KCl, 20 mM Tris, pH 8.0. Peak fractions were pooled and loaded onto a 5 ml Mono Q column (GE Healthcare) and eluted with a 30 ml salt gradient (0–500 mM KCl) in 20 mM Tris, pH 8.0. Peak fractions were pooled, dialyzed overnight at 4°C into HEK buffer (20 mM HEPES, pH 7.4, 1 mM EDTA, 50 mM KCl), aliquoted, snap-frozen in liquid N2, and stored at 80°C.
SNAP649-CapZ was expressed in E. coli BL21 DE3 and purified as follows (Shekhar et al., 2019). Cells were grown to a log phase at 37°C in TB medium, and then, expression was induced using 1 mM IPTG at 18°C overnight. Cells were harvested by centrifugation, and pellets were stored at −80°C. Frozen pellets were resuspended in lysis buffer (20 mM NaPO4, pH 7.8, 300 mM NaCl, 1 mM DTT, 15 mM imidazole, 1 mM PMSF) supplemented with a protease inhibitor cocktail (0.5 µM each of pepstatin A, antipain, leupeptin, aprotinin, and chymostatin). Cells were lysed by sonication with a tip sonicator while keeping the tubes on ice. The lysate was cleared by centrifugation at 150,000 × g for 30 min at 4°C. The supernatant was then flowed through a HisTrap column connected to a FPLC system. The column with the bound protein was first extensively washed with the washing buffer (20 mM NaPO4, pH 7.8, 300 mM NaCl, 1 mM DTT, and 15 mM imidazole) to remove nonspecifically bound proteins. SNAP-CapZ was then eluted with a linear zero to 250 mM imidazole gradient in 20 mM NaPO4, pH 7.8, 300 mM NaCl, and 1 mM DTT. The eluted protein was concentrated and labeled with SNAP-Surface-649 (New England Biolabs) according to the manufacturer’s instructions. Free dye was removed using size-exclusion chromatography by loading the labeled protein on a Superose 6 gel-filtration column (GE Healthcare) eluted with 20 mM HEPES, pH 7.5, 150 mM KCl, 0.5 mM DTT. Fractions containing the protein were combined, concentrated, aliquoted, snap-frozen in liquid N2, and stored at −80°C.
TIRF microscopy and analysis
Chamber preparation
Glass coverslips (60 × 24 mm; Thermo Fisher Scientific) were first cleaned by incubating with 1% Hellmanex III cleaning solution (Hellma Analytics) for 1 h at 60°C. Coverslips were then rinsed thoroughly with Milli-Q ultrapure water (MQ) and then subsequently incubated in 180-proof ethanol for 5 min. Coverslips were then rinsed again in MQ and incubated in a beaker filled with 0.1M KOH for 10 min. Coverslips were then washed one more time with MQ and dried with a nitrogen gas stream. The cleaned, dried coverslips were then coated with 40 mg/ml methoxypolyethylene glycol (PEG)–silane MW 2,000 and 8 mg/ml biotin-PEG-silane MW 3400 (Laysan Bio) in 80% ethanol, pH 2.0, by incubation overnight at room temperature. The next day, coverslips were rinsed with MQ, dried with an N2 stream, and stored in 50-ml conical tubes at −80°C for up to 6 mo. Before use in TIRF experiments, flow cells were assembled by rinsing PEG-coated coverslips with water, drying with N2, and adhering them to μ-Slide VI 0.1 (0.1 × 17 × 1 mm) flow chambers (Ibidi) with double-sided tape (2.5 cm × 2 mm × 120 µm) and 5-min epoxy resin (Devcon). Before each reaction, the flow cell was incubated for 1 min with 1% bovine serum albumin (BSA) in HEK buffer, and then equilibrated with TIRF buffer (50 mM imidazole, pH 7.4, 50 mM KCl, 1 mM MgCl2, 1 mM ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA), 0.2 mM ATP, 10 mM DTT, 15 mM glucose, 20 µg/ml catalase, 100 µg/ml glucose oxidase) plus 0.5% methylcellulose (4,000 cP).
Sample preparation and imaging
Actin and other proteins (as specified in figure captions) were flowed into the chamber. Time-lapse TIRF imaging was performed at room temperature on a Nikon-Ti2000 inverted microscope equipped with a 150-mW argon laser (Melles Griot), a 60× oil TIRF objective with an NA of 1.49 (Nikon Instruments), and an electron-multiplying charge-coupled device camera (Andor iXon). Focus was maintained by the Perfect Focus system (Nikon Instruments). Single-wavelength and multiwavelength images were acquired every 10 s using Imaging Software Elements (Nikon Instruments). Images were analyzed in FIJI.
For actin filament nucleation and elongation TIRF assays, gel-filtered monomeric actin in G-buffer was cleared by ultracentrifugation for 1 h at 4°C at 350,000 × g in a TLA-100 rotor (Beckman Coulter), and the upper ∼70% of the supernatant was carefully recovered and used. For each reaction, 1 μM monomeric actin (10% Alexa 488–labeled), 5 μM yeast profilin, and the indicated proteins or control buffer were flowed into the reaction chamber. Actin filament assembly was monitored for 10 min, acquiring images every 10 s. For counting the number of filaments nucleated, FOVs were analyzed for each condition 600 s after initiation of F-actin polymerization, and the number of actin filaments in the FOV was counted using the cell counter function in FIJI. Filament elongation rates were measured by plotting filament length versus time, where the elongation rate is the slope. To express rates in actin subunits s−1, we used the conversion factor of 374 subunits per µm of F-actin. Differences in the elongation rates and in the number of filaments nucleated for different conditions were assessed by 1-way ANOVA followed by Tukey’s HSD post hoc test using Prism (GraphPad).
For single-molecule TIRF imaging of Aip5C-L-SNAP549 at the pointed ends of elongating yeast actin filaments, frozen yeast actin was thawed and cleared by ultracentrifugation for 20 min at 4°C at 350,000 × g in a TLA-100 rotor (Beckman Coulter), and the upper ∼80% of the supernatant was carefully recovered and used. For each reaction, 5 nM Aip5C-L-SNAP549, 1 μM yeast G-actin (with 10% Alexa 488–labeled and 0.5% biotin-actin vertebrate muscle actin), and 5 μM yeast profilin were flowed into the reaction chamber. Actin filament assembly was monitored for 15 min, acquiring images every 10 s. A two-color kymograph was generated by tracing an Aip5C-L-SNAP549-capped filament growing for 400 s using the segmented line tool and “kymograph builder” plugin (FIJI). For three-color experiments to visualize SNAP649-CapZ and Aip5C-L-SNAP549 at opposite ends of the actin filaments, yeast actin filaments were first assembled as above in the presence of Aip5C-L-SNAP549, and then, 5 nM SNAP649-CapZ was flowed into the reaction chamber. After five minutes, images were acquired to visualize Aip5C-L-SNAP549, SNAP649-CapZ, and Alexa 488–labeled yeast F-actin. For enhanced visualization of single molecules on filament ends, images were denoised using Denoise.ai plugin in NIS-Elements Advanced Research software (Nikon) and background-subtracted using the average mean intensity from five manually chosen areas in the image that do not contain cells.
Bulk pyrene-actin assembly assays
Gel-filtered monomeric vertebrate muscle actin and/or purified yeast actin in G-buffer was cleared by ultracentrifugation for 1 h at 4°C at 350,000 × g in a TLA-100 rotor (Beckman Coulter), and the upper ∼70% of the supernatant was carefully harvested. All reactions (60 μl) contained 2 μM G-actin (5% pyrene-labeled), which was converted to Mg2+-ATP-actin 2 min before use by the addition of 10 mM EGTA and 1 mM MgCl2. To initiate actin assembly, 42 μl of Mg2+-ATP-G-actin was mixed with 15 μl of proteins or control buffer and 3 μl of 20× initiation mix (40 mM MgCl2, 10 mM ATP, and 1 M KCl), then placed in a 3 × 3 mm High Precision Cell quartz microcuvette (Hellma Analytics). Pyrene-actin fluorescence was monitored at excitation 365 nm (5-nm bandpass) and emission 407 nm (5-nm bandpass) at room temperature for 1,000–2,000 s in a fluorimeter (Photon Technology International). Vertebrate muscle actin assembly rates were quantified by measuring the slopes of the reaction curves from time points 0–300 s (without profilin) or 200–500 s (with profilin). Yeast actin assembly rates were quantified by measuring the slopes of the reaction curves from time points 0–300 s (with profilin).
To determine the concentration-dependent effects (and KApp) of Aip5C-L in nucleating F-actin jointly with Bud6-FL, 60 μl bulk reactions were set up, as described above. Reactions contained 2 μM rabbit muscle G-actin (5% pyrene-labeled), 100 nM Bud6-FL, and variable concentrations of Aip5C-L (0–500 nM). Reactions were loaded into a 96-well, flat-black, half-area plate (Corning), and pyrene fluorescence was monitored at excitation 365 nm and emission 407 nm in a Tecan Infinite M200-PRO microplate reader. The resulting curves were each fit to an exponential decay curve (“one-phase association” in Prism) in Prism (GraphPad) to extract an estimate of the actin assembly rate. The calculated actin assembly rates were then plotted and fit to a quadratic solution of the single-site–binding isotherm also fitting the assembly rates in the presence of saturating Aip5C-L and absence of Aip5C-L, using the following equation (Tomasso et al., 2025):
where fbound follows a single-site–binding quadratic isotherm:
Fitting was performed in Prism (GraphPad), which yielded the parameters of rbound, rfree, and Kapp. In this equation, robs is the polymerization rate observed in the bulk pyrene-actin assembly assay, rbound is the actin assembly rate when Bud6-FL is saturated with Aip5C-L, rfree is the actin assembly rate in the absence of Aip5C-L. Kapp is the apparent Kd of Aip5C-L for Bud6-FL, Ltotal is the concentration of Aip5C-L, and Ptotal is the concentration of Bud6-FL in the reaction (100 nM). Kapp, rfree, and rbound were optimized to best fit the data using the Levenberg–Marquardt method. Kapp is reported with curve fit error (standard error of fit yielded by Prism).
Bulk pyrene-actin filament disassembly assays
To monitor depolymerization at the pointed ends of actin filaments assembled from vertebrate actin, we first polymerized 20 μM G-actin (10% pyrene-labeled) in the presence of 400 nM CapZ for 2 h at room temperature (∼23°C). This produced a stock of actin filaments capped at their barbed ends. Reactions were then set up, containing a final concentration of 2 μM F-actin and 40 nM CapZ, along with the indicated concentrations of Aip5C-S or Aip5C-L, in a 96-well, flat-black, half-area plate (Corning). Depolymerization was initiated at time 0 by the addition of 3 µM vitamin D–binding protein (Cell Sciences). Pyrene fluorescence was monitored at excitation 365 nm and emission 407 nm in a Tecan Infinite M200-PRO microplate reader. The resulting fluorescence data for each reaction were normalized and fit to a single exponential using Prism (GraphPad), yielding a depolymerization rate constant for each reaction. To determine the Kapp of Aip5C-S inhibition of depolymerization from pointed ends, the depolymerization rates for each Aip5C-S concentration were averaged and plotted with their standard deviation (n = 3). These data were fit in Prism (GraphPad) to the single-site–binding isotherm used to determine Kapp for Bud6-Aip5C-L promotion of actin assembly (see above). Here, robs is the depolymerization rate observed in the bulk assay, rbound is the depolymerization rate of barbed-end capped filaments when saturated by Aip5C-S, and rfree is the depolymerization rate of barbed-end capped filaments in the absence of Aip5C-S. Kapp is the apparent Kd of Aip5C-S for pointed ends, Ltotal is the concentration of Aip5C-S, and Ptotal is the concentration of pointed ends in the reaction, which we assume to be equal to the final concentration of capping protein (40 nM). Kapp is reported with curve fit error (standard error of fit yielded by Prism).
To monitor depolymerization at the pointed ends of actin filaments assembled from yeast actin, we first polymerized 20 µM yeast G-actin (with 5% pyrene-labeled vertebrate muscle actin) in the presence of 400 nM CapZ. These filaments were mixed with the indicated concentrations of Aip5C-S or Aip5C-L construct in a 384-well, flat-black plate (Corning). Depolymerization initiation and observation were conducted as stated above. Depolymerization rates yielded by single-exponential fits to each curve were plotted and fit to a single-site–binding isotherm as above, with the modification of using the Langmuir isotherm, in lieu of the quadratic isotherm, to determine fbound:
where Ltotal is the total concentration of Aip5C-S, and Kapp is the apparent Kd of Aip5C-S for pointed ends. As above, Kapp is reported with standard error of fit yielded by Prism.
Fluorescence anisotropy assays
Fluorescence anisotropy was used to quantify the binding affinity of a labeled reporter (Bud6-1-145-KCK-Alexa488) for unlabeled Aip5C-L. Bud6-1-145-KCK-Alexa488 was diluted into FA buffer (50 mM KCl, 1 mM EGTA, 1 mM MgCl2, 0.5 mM DTT, and 1 mg/ml BSA), and 40 nM of the reporter was mixed with variable concentrations of Aip5C-L (0–750 nM). The mixtures were incubated for 10 min at room temperature, then placed into a 10 mm × 10 mm cuvette in T-form PTI QuantaMaster Spectrofluorometer, equipped with Glan–Thompson polarizers. Fluorescence anisotropy of the reporter was measured at excitation 485 nm (5-nm bandpass) and polarized emission at 525 nm (5-nm bandpass). Emission intensity was averaged for 3 min and converted to anisotropy values, after correcting for background signal intensity and G-factor. Values were then plotted and fit to a quadratic solution of the single-site–binding isotherm to derive the KD.
F-actin copelleting assays
Cosedimentation of Aip5C-S with F-actin was measured by incubating 30 μl samples of 2 μM F-actin with different concentrations of Aip5C-S (0, 0.1, 0.5, 1, 2, or 5 µM) overnight at 4°C in Aip5 in this final buffer: 50 mM Tris, pH 8, 138 mM NaCl, 4 mM KCl, 16 μM CaCl2, 160 μM MgCl2, 56 μM ATP, 1 mM EDTA, and 1 mM DTT. The next day, samples were equilibrated at room temperature (∼23°C) for 1 h and then centrifuged at 90,000 rpm (∼312,000 × g) for 30 min at 25°C in a TLA-100 rotor (Beckman Coulter). Equal fractions of the supernatants and pellets were separated on a 10% SDS-PAGE, stained with Coomassie brilliant blue, and imaged using ChemiDoc Imaging System (Bio-Rad).
Structural modeling using AlphaFold
Reference sequences for S. cerevisiae Bud6 (Ref. Seq. #KAJ1542076), Aip5 (Ref. Seq. #NP_116671), and actin (Ref. Seq. #NP_116614) were obtained from NCBI. AlphaFold3 server (Abramson et al., 2024) was used to model a complex of Bud6 (residues 699–786), Aip5 (residues 997–1,233), and three copies of full-length actin, with default settings and a model seed of 1408571676. The model was assessed for pIDDT scores of at least 70 through the structured regions (disordered regions had lower scores and were not considered). In addition, the five models that were produced were aligned in PyMOL (2025) and showed very close agreement, with RMSD to the first model of <0.7 Å, after excluding disordered regions in Aip5. The entire Bud6flank domain (699–788) had a consistent structure, whereas only a portion of Aip5 (residues 1,129–1,233) had a consistent structure; only these consistent structures are shown in Fig. 4. Modeling was repeated with various numbers of actin subunits and different regions of Bud6 and Aip5. The model shown captures the most consistently observed features near the actin subunits.
Aspects of the model for which high-resolution structures were available were superposed to assess agreement. The modeled structure of the C-terminal thioredoxin-related domain in Aip5 agreed well with the existing crystal structure of this domain (PDB# 6ABR, chain A; Xie et al., 2019) with an RMSD of <0.3 Å. The modeled actins better approximate the flattened F-actin conformation at the subunit level, with each subunit agreeing with the structure of yeast actin in a filament (PDB# 9GO5) (Stevenson et al., 2025) with an RMSD of 0.4–0.8 Å, and with a gelsolin yeast actin monomer (PDB# 1YAG [Vorobiev et al., 2003]) to better than 0.9–1.3 Å. The organization of the actin trimer in the model is similar to that in F-actin, although with a slightly longer helical pitch. Thus, the model agrees with existing structural data. To assess the proposed Aip5 contact, we compared the actin contact surface on Aip5 (contact defined as residues with any atom within 4 Å of either pointed-end actin subunit) with surface conservation as determined using ConSurf-DB (Ben Chorin et al., 2020; Goldenberg et al., 2009), and with an existing point mutation (3M) (Xie et al., 2019).
To search for other proteins with 3D structural homology to the thioredoxin-related domain of Aip5, we used DALI (Holm et al., 2023) with the 6ABR chain A structure. This search revealed the parent structure for Aip5 thioredoxin/glutaredoxin domain, as well as a mutant structure, and that of the same domain from C. albicans. Furthermore, the search revealed structural similarity to the SH3 Domain Binding Glutamate-Rich Like domain family, including the SH3BGRL2 component of the spectrin–actin junctional complex from red blood cells (Li et al., 2023). Importantly, this structure was not part of the training set for AlphaFold3, as the structures were added to the PDB after the cutoff for training data.
Quantification and statistical analysis
All experiments were repeated multiple times and yielded similar results (see figure legends for repeat numbers for each experiment). All image intensity quantification was performed in FIJI. All statistical tests, nonlinear fitting, and calculations were performed in Prism 10 (GraphPad). Data distributions were assumed to be normal, but this was not formally tested. Statistical comparison between indicated conditions was conducted using two-sided Student’s t test, using Welch’s correction for unequal standard deviations where indicated, or one-way ANOVA followed by Tukey’s HSD post hoc, as indicated in figure legends and Methods subsections. Differences were considered significant if the P value was less than predetermined critical values: *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.
Online supplemental material
Fig S1 shows additional characterization of Bud6 and Aip5 actin nucleation. Bud6 and Aip5 potently synergize in promoting F-actin assembly in the absence of profilin with vertebrate muscle actin. In addition, Bud6 and Aip5 synergize in promoting the assembly of yeast actin, and collaborate with yeast formin Bni1 to assemble actin in the presence of profilin and capping protein. Fig S2 shows loss of BUD6 and AIP5 results in a decrease in whole-cell F-actin staining without affecting actin patch intensity. Additionally, point mutations in Bud6 that disrupt its interactions with formins and G-actin lead to similar defects in actin cable number and length as a full deletion of BUD6. Fig S3 uses AlphaFold3 modeling to show a structural view (different from the view in the main figure) of the Bud6-Aip5 actin nucleation mechanism, as well as a structural overlay between the thioredoxin-related domains of yeast Aip5 and vertebrate SH3BGRL2. Fig S4 shows that the Aip5C-L (3M) mutant disrupts Bud6-Aip5 nucleation effects on yeast actin. Also, Bud6 localization in vivo is unaffected by truncation of its second actin-binding site (ABS2). Fig S5 shows that cells expressing the integrated Aip5-3M mutant (tagged with 3xmNG) have elevated expression levels compared with cells expressing wild-type Aip5 (tagged with 3xmNG). Loss of BNI1 does not affect Aip5-3xmNG punctum velocities on actin cables. High-speed F-actin cosedimentation assays show that Aip5C-S does not bind the filament sides. Also, the Aip5C-L (3M) mutant disrupts pointed-end capping of yeast F-actin in vitro. Table S1 contains a list of yeast strains used in this study. Table S2 contains a list and descriptions of plasmids used in this study. Table S3 contains a list and description of DNA oligos used in this study. Video 1 shows Bud6-FL and Aip5C-L effects on actin filament assembly in TIRF assays. Video 2 shows live-cell imaging of actin cables (labeled with LifeAct-mNG) prematurely detaching from the bud neck in bud6Δ, aip5Δ, and bud6Δ aip5Δ mutants. Video 3 shows by live-cell imaging that loss of BUD6 and/or AIP5 leads to an abnormal accumulation of secretory vesicles (marked with GFP-Sec4) in the mother cell compartment. Video 4 shows that Aip5-3xmNG and Bni1-3xmNG puncta, but not Bud6-3xmNG puncta, undergo directed movements (retrograde flow), moving from the bud tip to the bud neck (Bni1-3xmNG puncta) or the bud tip to the back of the mother cell (Aip5-3xmNG). Video 5 shows that treating cells with 200 μM LatB abolishes the retrograde movements of Aip5-3xmNG and Bni1-3xmNG puncta. Video 6 shows that the Aip5-3M-3xmNG mutant, which is impaired in pointed-end binding in vitro, disrupts association of Aip5 puncta with actin cables. In contrast, the Aip5ΔBBS-3xmNG mutant, which impairs Aip5-Bud6 interactions, shows normal movement of puncta with actin cable retrograde flow. Video 7 shows that the in vivo movements of wild-type Aip5-3xmNG puncta on cables are independent of Bni1, Bnr1, or Bud6. Video 8 shows in vitro TIRF single-molecule imaging of labeled Aip5C-L-SNAP549 molecules associating with the pointed ends of yeast actin filaments, which are actively growing at their barbed ends.
Supplementary Material
contains a list of yeast strains used in this study.
contains a list and descriptions of plasmids used in this study.
contains a list and description of DNA oligos used in this study.
is the source file for Fig. S5.
Acknowledgments
We are grateful to members of the Goode lab for input throughout this project, and we especially thank Julian Eskin for producing the graphical models.
This work was funded by a grant from National Institute of General Medical Sciences (NIGMS) to Bruce L. Goode (R35 GM134895) and an NRSA postdoctoral fellowship from NIGMS to Joseph O. Magliozzi (F32 GM150204).
Author contributions: Joseph O. Magliozzi: conceptualization, data curation, formal analysis, funding acquisition, investigation, methodology, project administration, resources, software, supervision, validation, visualization, and writing—original draft, review, and editing. Lucas A. Runyan: formal analysis, investigation, methodology, validation, visualization, and writing—original draft, review, and editing. Adah Welsh: formal analysis and investigation. Shae B. Padrick: conceptualization, formal analysis, methodology, visualization, and writing—original draft, review, and editing. Bruce L. Goode: conceptualization, funding acquisition, project administration, resources, supervision, and writing—original draft, review, and editing.
Data availability
Data supporting the findings of this manuscript are available from the corresponding author upon request.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials
contains a list of yeast strains used in this study.
contains a list and descriptions of plasmids used in this study.
contains a list and description of DNA oligos used in this study.
is the source file for Fig. S5.
Data Availability Statement
Data supporting the findings of this manuscript are available from the corresponding author upon request.