1. Cell Biology

Nim1-related kinases regulate septin organization and cytokinesis by modulating Hof1 at the cell division site

  1. Bindu Bhojappa
  2. Anubhav Dhar
  3. VT Bagyashree
  4. Jayanti Kumari
  5. Freya Cardozo
  6. Vaseef Rizvi
  7. Deepthi Guturu
  8. Saravanan Palani  Is a corresponding author
  1. Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, India
https://doi.org/10.7554/eLife.106366.3
Share this article
Cite this article
  1. Bindu Bhojappa
  2. Anubhav Dhar
  3. VT Bagyashree
  4. Jayanti Kumari
  5. Freya Cardozo
  6. Vaseef Rizvi
  7. Deepthi Guturu
  8. Saravanan Palani
(2026)
Nim1-related kinases regulate septin organization and cytokinesis by modulating Hof1 at the cell division site
eLife 14:RP106366.
https://doi.org/10.7554/eLife.106366.3
  2. Download BibTeX
  3. Download .RIS

eLife Assessment

In this valuable study, Bhojappa et al. investigate the function of four septin-associated kinases (Elm1p, Gin4p, Hsl1p and Kcc4p) in controlling septin ring stability, actomyosin ring organization and constriction, and cytokinesis in budding yeast. The evidence supporting the conclusions is convincing. The results are a meaningful addition to previously published work and provide a clearer picture of a complex mechanism involving proteins of partially redundant functions.

https://doi.org/10.7554/eLife.106366.3.sa0

Abstract

The septin scaffold recruits and organizes actomyosin ring (AMR) components; thus, ensuring faithful cytokinesis. The septin-associated kinases – Elm1, Gin4, Hsl1, and Kcc4 are thought to stabilize and regulate the septin architecture at the bud neck, but the underlying mechanisms remain largely unknown. Here, we present a comprehensive, quantitative analysis of these four septin-associated kinases and reveal major roles for Elm1 and Gin4 in septin stability and architectural transitions during the cell cycle. We find that Elm1 and Gin4 play a previously overlooked role in AMR organization and constriction during cytokinesis. We report that the Gin4 kinase interacts directly with the AMR component and F-BAR protein Hof1 via its C-terminal membrane-binding kinase associated-1 (KA1) domain, and is likely involved in the proper organization and anchoring of Hof1 at the bud neck, representing an unappreciated mode of regulation during cytokinesis. We further show that Gin4 controls septin organization and AMR constriction in a kinase-independent manner, similar to Elm1. Using an extensive GFP-GBP-based tethering assay in elm1Δ and gin4Δ cells, we identify an important role for Hsl1 in maintaining septin organization and cell shape in coordination with Elm1, Gin4, and Kcc4, independent of its role in the morphogenetic checkpoint. Furthermore, our data indicate that Hsl1 acts downstream of Elm1, with its membrane-binding KA1 domain being critical for its function. Together, these findings reveal new insights into the modes by which the kinases Gin4 and Elm1 regulate cytokinesis, highlight a redundant role for Hsl1 in controlling septin organization and cytokinesis, and uncover the inherent redundancy and adaptability of the septin kinase network in Saccharomyces cerevisiae.

Introduction

Septins, the fourth cytoskeletal element (Mostowy and Cossart, 2012), are an evolutionarily conserved family of GTP-binding proteins that assemble into filamentous structures on cell membranes (Bridges et al., 2014; Beber et al., 2019). Although septins belong to the P-loop GTPases (Leipe et al., 2002), recent studies have revealed that some septin proteins have lost the ability to hydrolyze bound GTP during evolution (Delic et al., 2024; Hussain et al., 2023). They play critical roles in various fundamental processes such as cytokinesis, ciliogenesis, spermatogenesis, and phagocytosis (Hartwell, 1971; Ihara et al., 2005; Kanamaru et al., 2022; Huang et al., 2008). Septins polymerize into hetero-oligomeric complexes that further assemble into higher-order structures, such as filaments, rings, bundles, and gauzes (Beber et al., 2019; Bertin et al., 2008; Bertin et al., 2012; Byers and Goetsch, 1976; Ong et al., 2014; McMurray et al., 2011). These structures function as scaffolds that recruit and organize proteins required for cytokinesis and cell division across eukaryotes (Spiliotis and Nelson, 2006; Wloka et al., 2011). Septins act as a diffusion barrier at the cell division site in fungi and animal cells, contributing to the assembly, maturation, and constriction of the contractile actomyosin ring (AMR) machinery during cytokinesis (Bi et al., 1998; Dobbelaere and Barral, 2004; Gladfelter et al., 2001; Lippincott and Li, 1998a; Barral and Mansuy, 2007). However, studies have suggested that redundant mechanisms can compensate for the septin diffusion barrier, rendering it dispensable for AMR constriction and cytokinesis, indicating that septins may not be strictly essential for AMR function in all contexts (McMurray et al., 2011; Wloka et al., 2011). The budding yeast Saccharomyces cerevisiae is an excellent model system for studying the relationship between septin organization and cytokinesis. In this organism, five mitotic septins Cdc3, Cdc12, Cdc10, Cdc11, and Shs1 (Hartwell, 1971; Ford and Pringle, 1991; Haarer and Pringle, 1987; Kim et al., 1991; Mino et al., 1998) assemble into an octameric complex that subsequently polymerizes into septin filaments or rods capable of organizing diverse higher-order architectures at the plasma membrane (Bridges et al., 2014; Beber et al., 2019; Cannon et al., 2019).

The terminal subunits of the octameric complex influence the architecture of the septin structure: Cdc11 promotes septin filament bundling, whereas Shs1 favors the formation of ring-like structures (Garcia et al., 2011; Weems and McMurray, 2017). Septin filaments assemble at the mother-bud neck to form a stable ‘hourglass’ structure that acts as a scaffold for proteins involved in cytokinesis, including components of the AMR complex (Ong et al., 2014; Wloka et al., 2011; Bi et al., 1998). At mitotic exit, this hourglass structure undergoes dynamic remodeling into a double-ring structure, marking the onset of cytokinesis (Tamborrini et al., 2018; Lippincott et al., 2001; Cid et al., 2001). This transition allows proper anchoring of the AMR to the plasma membrane and leads to efficient ring constriction and centripetal septum deposition between the dividing cells (Bi et al., 1998; Bi, 2001; Schmidt et al., 2002; Cheffings et al., 2016; Foltman and Sanchez-Diaz, 2024). The mitotic exit network (MEN) is thought to play a crucial role in initiating this transition (Tamborrini et al., 2018; Lippincott et al., 2001). Despite its importance, the precise molecular mechanisms regulating the hourglass to double-ring (HDR) transition remain incompletely understood.

Several kinases associate with septin structures and regulate septin dynamics during different phases of the cell cycle. These septin-associated kinases localize to the septin hourglass at the beginning of the G1 phase and dissociate from the bud neck at the onset of cytokinesis, coincident with the start of the HDR transition (Perez et al., 2016; Marquardt et al., 2024; Marquardt et al., 2020; Diaz et al., 2025). In S. cerevisiae, four conserved septin-associated kinases – Elm1 (Marquardt et al., 2024; Marquardt et al., 2020; Sreenivasan and Kellogg, 1999; Szkotnicki et al., 2008; Kang et al., 2016), Gin4 (Marquardt et al., 2024; Diaz et al., 2025; Altman and Kellogg, 1997; Carroll et al., 1998; Dobbelaere et al., 2003; Buttery et al., 2012; Jasani et al., 2020; Mortensen et al., 2002; Longtine et al., 1998), Hsl1 (Finnigan et al., 2016; King et al., 2012; Sakchaisri et al., 2004; Hanrahan and Snyder, 2003; Theesfeld et al., 2003; Crutchley et al., 2009), and Kcc4 (Moravcevic et al., 2010; Okuzaki et al., 2003) are thought to exert distinct and overlapping regulatory effects on septin stability and organization (Marquardt et al., 2024; Marquardt et al., 2020; Longtine et al., 1998; Barral et al., 1999; Longtine et al., 2000). Elm1 and Gin4 deletion significantly destabilizes the septin architecture (Marquardt et al., 2024; Marquardt et al., 2020; Sreenivasan and Kellogg, 1999; Barral et al., 1999), while loss of Hsl1 and Kcc4 does not have any effect (Barral et al., 1999). Several hypotheses have been proposed to describe the functions of these kinases during cytokinesis. Previous studies suggested that the LKB1-like kinase Elm1 and the Nim1-related kinases (Gin4, Hsl1, and Kcc4) function in parallel pathways to regulate septin architecture at the bud neck (Barral et al., 1999; Bouquin et al., 2000). More recently, it was shown that Elm1 acts as the most upstream kinase in this septin kinase network by regulating the activation and phosphorylation of the Nim1-related kinases (Marquardt et al., 2020; Asano et al., 2006). In addition, Elm1 is known to regulate septin organization by binding to its substrate Bni5 independently of its kinase activity (Marquardt et al., 2020). Gin4, on the other hand, is thought to regulate septin architecture by phosphorylating Shs1 (Mortensen et al., 2002). Hsl1 is best known for its role in negatively regulating Swe1 in the morphogenetic checkpoint pathway by forming a complex with Hsl7 (King et al., 2012; Theesfeld et al., 2003; Barral et al., 1999). How Hsl1 and Kcc4 regulate septin organization remains unknown, and the mechanisms by which these kinases coordinate septin organization and cytokinesis remain incompletely understood.

Despite significant progress in understanding septin architecture and its regulation, the coordinated effects of septin-associated proteins on this process are only beginning to be understood, and several important questions remain open in the field of septin biology. In particular, it is unclear how septin-associated kinases contribute to the organization and remodeling of septin structures, and how these regulatory pathways are coordinated with AMR assembly and cytokinesis. Furthermore, whether these kinases influence cytokinesis through direct interactions with components of the cytokinetic machinery or indirectly through effects on septin organization remains largely unexplored. In this study, we investigate the roles of the septin-associated kinases Elm1, Gin4, Hsl1, and Kcc4 in regulating septin organization and cytokinesis in S. cerevisiae, with the goal of understanding how this kinase network coordinates septin dynamics with AMR function during cell division.

Results

Dynamics of septin-associated kinases during the cell cycle and defects associated with their deletions

S. cerevisiae contains four major septin-regulatory kinases Elm1, Gin4, Hsl1, and Kcc4 that are known to localize to the mother-bud neck and contribute to septin stability and the morphogenesis checkpoint during the cell cycle (Marquardt et al., 2020; Barral et al., 1999). Three of these kinases, commonly referred to as Nim1-related kinases (Barral et al., 1999): Gin4, Hsl1, and Kcc4 contain a membrane-binding kinase associated-1 (KA1) domain at the C-terminus (Figure 1A). The KA1 domain shows affinity for acidic phospholipids (e.g., phosphatidylserine) on the plasma membrane and is required for the bud neck targeting (Buttery et al., 2012; Jasani et al., 2020; Finnigan et al., 2016; Moravcevic et al., 2010). To characterize their temporal kinetics and localized accumulation at the bud neck septin scaffold, we performed time-lapse imaging of endogenously GFP-tagged septin kinases in cells co-expressing the core septin Cdc3 tagged with mCherry. All four kinases showed simultaneous, gradual recruitment to the septin hourglass during bud emergence and exhibited a sequential pattern of disappearance during the onset of cytokinesis, consistent with previous observations (Marquardt et al., 2024; Marquardt et al., 2020; Figure 1—figure supplement 1A–G, Figure 1—video 1). Elm1 and Kcc4 exhibited lower fluorescence intensities at the bud neck prior to septin remodeling, suggesting a lower abundance at this site at the defined time period (Figure 1—figure supplement 1E–G), compared with the higher abundance of Gin4 and Hsl1, which exhibited increased fluorescence intensities prior to septin remodeling (Figure 1—figure supplement 1E–G). At the onset of cytokinesis, when the septin scaffold undergoes remodeling from the hourglass structure to a double-ring structure (HDR transition), all four kinases left the bud neck just before or during the HDR transition in a sequential manner (Figure 1—figure supplement 1E–G, Figure 1—video 1). Interestingly, fluorescence intensities of Elm1 and Kcc4 began to decrease ~4–6 min before the HDR transition (Figure 1—figure supplement 1E–G). In contrast, Gin4 and Hsl1 fluorescence signal intensities began to decrease just before or during the HDR transition, suggesting that they may serve as a trigger or key molecular cue for septin remodeling that subsequently crosstalks with downstream events of cytokinesis (Figure 1—figure supplement 1E–G). These observations suggest a defined temporal order of removal of these kinases from the bud neck during cytokinesis, which may directly rely on the interdependency of their localizations on one another. Overall, these data show that the four septin kinases are associated with the septin hourglass structure with similar patterns of recruitment but distinct removal kinetics, suggesting that they may play both overlapping and non-overlapping roles in septin hourglass stability and the HDR transition.

Figure 1 with 5 supplements see all
Download asset Open asset
Septin organization and actomyosin ring (AMR) dynamics are defective in elm1Δ and gin4Δ cells.

(A) Domain-level architecture of Elm1, Gin4, Kcc4, and Hsl1 kinases. The KA1 domain represents the membrane-binding domain. (B) Representative time-lapse images showing Cdc3-mCherry dynamics in septin-kinase deletion strains, captured at 2-min time intervals (t0 = spindle breakpoint). (C) Normalized fluorescence intensity profile showing the temporal kinetics of Cdc3-mCherry in the indicated strains shown in (B) (wild-type: n = 41, elm1Δ: n = 29, gin4Δ: n = 33, hsl1Δ: n = 34, kcc4Δ: n = 36 cells). (D) Representative time-lapse images of Inn1-mNG constriction captured at 2-min time intervals in septin-kinase deletion strains (t0 = appearance of Inn1-mNG at the bud neck). (E) Quantification of the residence time of Inn1-mNG during the onset of cytokinesis in the indicated strains of (D).Kruskal–Wallis nonparametric test (****p < 0.0001, ns: p > 0.05). Whiskers represent mean ± 1.5 × standard deviation (SD). (F) Normalized fluorescence intensity graph showing the kinetics of Inn1-mNG in the indicated strains of (D). A population of elm1Δ and gin4Δ cells exhibiting asymmetric constriction is included in the plot (wild-type: n = 21, elm1Δ: n = 13, gin4Δ: n = 23, hsl1Δ: n = 21, and kcc4Δ: n = 18 cells).

Figure 1—source data 1

Numerical data underlying the quantitative analyses represented in Figure 1.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig1-data1-v1.xlsx

Septin-associated kinases Elm1 and Gin4 majorly regulate septin stability, HDR transition, and cellular morphology

Septin rearrangement during cytokinesis is believed to be partly governed by post-translational modifications (PTMs), including phosphorylation, ubiquitination, acetylation, and SUMOylation (Perez et al., 2016; Carroll et al., 1998; Hernández-Rodríguez and Momany, 2012; Johnson and Blobel, 1999; Marquardt et al., 2019; McMurray and Thorner, 2009; Mitchell et al., 2011; Takahashi et al., 1999). Previously, different studies have highlighted the role of kinase-mediated modifications in septin remodeling, but a comprehensive quantitative analysis for all the four septin-associated kinases under similar conditions is not present (Marquardt et al., 2020; Mortensen et al., 2002; Longtine et al., 1998; Schmidt et al., 2003; McQuilken et al., 2017). Thus, to understand the extent to which different septin kinases affect cell physiology and septin organization, we characterized cell morphology, cell growth, and septin organization in wild-type cells and in cells harboring deletions of the septin-regulatory kinases – Elm1, Gin4, Hsl1, and Kcc4. We observed that cells exhibited elongated and clumped morphologies in the elm1Δ and gin4Δ backgrounds, respectively, at all three temperatures (23, 30, and 37°C) tested, consistent with previous observations (Marquardt et al., 2024; Sreenivasan and Kellogg, 1999; Barral et al., 1999; Blacketer et al., 1993; Figure 1—figure supplement 2A, B). Quantitative analysis revealed elm1Δ cells displayed a more severe defect, with nearly 100% of cells showing elongated morphology that failed to undergo isometric growth, whereas gin4Δ cells exhibited clumped morphology in over 73.33% of the cellular population (Figure 1—figure supplement 2A, B). While hsl1Δ cells largely showed round morphology similar to wild-type at 23 and 30°C, a fraction of cells exhibited mild morphological defects at 37°C (Figure 1—figure supplement 2A, B). In contrast, no alterations in morphology were observed in kcc4Δ cells (Figure 1—figure supplement 2A, B). Growth assays of these kinase deletions showed a similar trend: elm1Δ and gin4Δ cells showed growth sensitivity at both 23 and 30°C, which was heightened at 37°C for elm1Δ, while hsl1Δ exhibited growth sensitivity only at the higher temperature 37°C, in contrast to kcc4Δ cells, which grew identically to the wild-type strain (Figure 1—figure supplement 2C).

Next, we sought to systematically assess the effects of the absence of these kinases on septin architecture and dynamics. The core septin Cdc3 fused to mCherry was used to visualize the septin structures during the cell cycle (Fang et al., 2010). Cdc3-mCherry localized to the emerging bud site in wild-type, elm1Δ, gin4Δ, hsl1Δ, and kcc4Δ cells but was subsequently mislocalized to the bud cortex, consistent with previous findings (Marquardt et al., 2020; Sreenivasan and Kellogg, 1999; Longtine et al., 1998; Bouquin et al., 2000) in ~100% of elm1Δ cells and 80.48% of gin4Δ cells (Figure 1—figure supplement 2E, G; Figure 1—video 2). In wild-type cells, septin formed a stable hourglass structure at the bud neck, which subsequently split into double rings coinciding with spindle breakage and mitotic exit (Figure 1—figure supplement 2D, F). The septin hourglass architecture appeared abnormal in elm1Δ and gin4Δ cells and exhibited a defective HDR transition (Figure 1B, C; Figure 1—video 2). The elm1Δ cells showed a stronger defect in septin organization during bud emergence and the HDR transition compared with gin4Δ cells (Figure 1B, C; Figure 1—figure supplement 2E, G). This suggests a more prominent role for Elm1 in the maintenance of septin filament architecture, consistent with previous literature reporting a role for Elm1 in stabilizing the daughter-side of the septin hourglass structure by regulating septin filament pairing (Marquardt et al., 2020). Cells lacking the Hsl1 and Kcc4 kinases showed normal septin organization during bud emergence and HDR transition at the onset of cytokinesis, similar to wild-type cells (Figure 1B, C; Figure 1—figure supplement 2E, G). However, approximately 7.14% of hsl1Δ cells displayed septin mislocalization to the bud cortex during bud emergence (Figure 1—figure supplement 2E, G), indicating that Hsl1 may also play a minor or supporting role in septin dynamics through a compensatory mode of regulation. These results therefore demonstrate a major requirement for Elm1 and Gin4 in proper septin arrangement and dynamics at the cell division site, while also hinting at a parallel minor role for Hsl1.

Elm1 and Gin4 regulate AMR organization and constriction

Given that our data suggest major defects in septin regulation occur in the absence of Elm1 and Gin4, we asked whether these defects also translate into effects on AMR-mediated cytokinesis. We therefore investigated the roles of Elm1 and Gin4 in AMR organization at the cell division site and in constriction dynamics during cytokinesis. The AMR machinery generates the force required to drive plasma membrane ingression for successful cell separation (Bi et al., 1998; Foltman et al., 2016). In budding yeast, the AMR machinery consists primarily of actin filaments, tropomyosin, formins, Myo1, Hof1, Iqg1, Cyk3, and Inn1, along with many other regulatory proteins (Bi et al., 1998; Lippincott and Li, 1998a; Meitinger et al., 2013; Sanchez-Diaz et al., 2008; Nishihama et al., 2009). We used endogenously labeled Myo1-mNeonGreen (mNG), the sole non-muscle myosin-II in S. cerevisiae, to determine AMR organization during bud emergence. Previous studies have shown that Myo1 recruits to the bud neck in a bi-phasic manner – via Bni5 prior to cytokinesis and via Iqg1 during cytokinesis (Fang et al., 2010; Wloka et al., 2013). Consistent with its interaction with septin-binding protein Bni5 (Fang et al., 2010), we observed that, similar to Cdc3-mCherry, Myo1-mNG was mislocalized to the bud cortex during bud emergence in 75.86% and 30% of elm1Δ and gin4Δ cells, respectively (Figure 2—figure supplement 1A, B). We then used Inn1, which arrives 2 min after the start of AMR constriction and aids in plasma membrane ingression by coupling AMR constriction with primary septum (PS) formation as a marker to determine AMR constriction dynamics during cytokinesis (Sanchez-Diaz et al., 2008; Nishihama et al., 2009). In wild-type cells, Inn1-mNG arrived at the bud neck just prior to spindle breakage and constricted symmetrically as part of the AMR complex with a residence time of ~5–6 min, and eventually disappeared from the bud neck (Figure 1D–F). In elm1Δ and gin4Δ cells, Inn1-mNG was recruited normally to the cell division site but displayed an abnormal constriction pattern with an increased duration of constriction (~8–10 min) (Figure 1D–F, Figure 1—video 3). By contrast, in hsl1Δ and kcc4Δ cells, Inn1-mNG was recruited normally and constricted similarly to the wild-type strain (Figure 1D–F, Figure 1—video 3). These results agree with the regulation of these kinases on septin dynamics, suggesting that Elm1 and Gin4 may directly or indirectly regulate AMR constriction and cytokinesis.

Inn1 interacts with Hof1, an F-BAR protein known to associate with the septin hourglass, transit to the AMR during the split-ring trigger (Foltman et al., 2016; Meitinger et al., 2013; Garabedian et al., 2020; Lippincott and Li, 1998b; Vallen et al., 2000; Young et al., 2010; Varela Salgado et al., 2024), and sense membrane curvature (Meitinger et al., 2013; Nishihama et al., 2009; Varela Salgado et al., 2024). Inn1 also functions in parallel with Cyk3; Hof1, Inn1, and Cyk3 form a tripartite complex referred to as the Ingression Progression Complex (IPC) (Nishihama et al., 2009; Onishi et al., 2013; Jendretzki et al., 2009; Devrekanli et al., 2012). This complex then activates the primary component of the PS, Chitin Synthase II (Chs2) (Nishihama et al., 2009; Onishi et al., 2013; Jendretzki et al., 2009; Devrekanli et al., 2012). Thus, we monitored the constriction pattern and dynamics of Chs2-mNG and observed an increased residence time for Chs2 at the bud neck in both elm1Δ and gin4Δ cells (Figure 2—figure supplement 1C–E). Our observations suggest that loss of the septin kinases Elm1 and Gin4 may result in improper organization and constriction dynamics of the AMR, and possibly impaired IPC formation, resulting in abnormal cytokinesis.

To test the organization of the AMR/IPC at the bud neck, we assessed the organization of the F-BAR protein Hof1. Deletion of Hof1 has been shown to result in asymmetric AMR constriction and improper chitin deposition during PS synthesis (Nishihama et al., 2009). We then analyzed the residence time of Hof1-mNG at the bud neck after the spindle breakpoint during cytokinesis. Hof1-mNG exhibited a residence time of 10–12 min after the mitotic spindle break in wild-type cells, which increased to >15 min in both elm1Δ and gin4Δ cells (Figure 2A, B; Figure 2—video 1). By contrast, we did not observe any significant change in the recruitment and temporal kinetics of Hof1-mNG in elm1Δ and gin4Δ cells compared with wild-type during the mitotic spindle break (Figure 2C). Next, we analyzed the organization of Hof1-mNG at the cell division site. In wild-type cells, Hof1-mNG appeared as a continuous, uniform ring at the bud neck in 93.05% of the cellular population, whereas discontinuous or fragmented Hof1-mNG rings were observed in 82.69% and 71.07% of elm1Δ and gin4Δ cells, respectively (Figure 2D, E). These observations suggest that Elm1 and Gin4 regulate the proper spatial assembly of various AMR components to ensure robust cytokinesis. The effect on AMR dynamics observed in elm1Δ and gin4Δ cells may be an indirect consequence mediated through septins, because septin organization is significantly defective in these backgrounds. However, a previous study has shown that disruption of the septin scaffold does not affect cytokinesis, suggesting that Elm1 and Gin4 could regulate AMR organization and constriction independently of their effects on septin organization (Wloka et al., 2011). Together, these results reveal an unappreciated role for Elm1 and Gin4 in AMR assembly, organization, and constriction, opening avenues for future investigation.

Figure 2 with 3 supplements see all
Download asset Open asset
Gin4 regulates Hof1 organization and dynamics through interaction with the N-terminal membrane-binding F-BAR domain.

(A) Representative time-lapse images of Hof1-mNG in wild-type, elm1Δ, and gin4Δ cells imaged at 2-min time intervals (t0 = spindle breakpoint). (B) Quantification of the residence time of Hof1-mNG after spindle breakpoint in the indicated strains shown in (A).Kruskal–Wallis nonparametric test (**p < 0.01, ns: p > 0.05). Whiskers represent mean ± 1.5 × SD. (C) Normalized fluorescence intensity profile for the temporal kinetics of Hof1-mNG during cytokinesis in the indicated strains shown in (A), (wild-type: n = 33, elm1Δ: n = 21, gin4Δ: n = 23 cells). (D) Representative images showing the organization of Hof1-mNG at the bud neck in the indicated strains shown in (A). Scale bar: 5 µm. (E) Bar graph showing the percentage of cells exhibiting misorganization of Hof1 at the cell division site in the indicated strains shown in (D) .One-way ANOVA with Tukey’s multiple-comparison test (****p < 0.0001, ns: p > 0.05). Error bars indicate + SD from the mean. p-values correspond to the population of cells exhibiting disorganized Hof1-mNG at the bud neck (N = 3, wild-type: n = 518, elm1Δ: n = 225, gin4Δ: n = 384 cells). (F) Representative images from the Yeast Two-Hybrid assay depicting the interaction of full-length Gin4, Gin4-KID, and Gin4-KA1 with full-length Hof1. (G) Representative Yeast Two-Hybrid images showing the interaction between Gin4-KA1 (1003–1142 aa) and the N-terminal F-BAR containing domain of Hof1 (1–350 aa). (H) Immunoblot showing the in vitro binding assay for the interaction between 6xHis-bdSUMO-Gin4KA1 and GST-Hof1F-BAR fragments. GST-tagged Hof1 fragment is immobilized on glutathione resin, and bound 6xHis-bdSUMO-tagged Gin4 fragments were analyzed by SDS–PAGE. Input (top) and pull-down fractions (middle) were probed with anti-His antibody, while GST-Hof1 fragments (bottom) were detected using anti-GST antibody, (+) present, (−) absent. (I) Proposed model depicting the possible interplay between Gin4 and Hof1 at the plasma membrane interface. Created with BioRender.com.

Figure 2—source data 1

Numerical data underlying the quantitative analyses represented in Figure 2.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig2-data1-v1.xlsx
Figure 2—source data 2

PDF containing the uncropped western blots corresponding to the in vitro binding assay shown in Figure 2H, with the relevant bands and regions clearly indicated.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig2-data2-v1.zip
Figure 2—source data 3

Original files of the full, uncropped, unedited western blots corresponding to the in vitro binding assay shown in Figure 2H.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig2-data3-v1.zip

Gin4 directly interacts with the N-terminal F-BAR domain of Hof1 to regulate proper cytokinesis

To understand how the Elm1 and Gin4 kinases affect AMR dynamics, we performed a Yeast Two-Hybrid (Y2H) assay to identify novel interacting partners of these proteins. We found that two Gin4 fragments – Kinase Independent Domain (KID; 290–1142 aa) and kinase associated-1 (KA1; 1003–1142 aa), but not the full-length protein, interacted with the full-length Hof1 protein (Figure 2F). Further, the N-terminal Hof1 fragment (1–350 aa), which contains its F-BAR domain, but not the other fragments, interacted with the Gin4-KA1 domain (Figure 2G, Figure 2—figure supplement 1H). By contrast, neither full-length Elm1 nor its C-terminal fragment (421–640 aa) showed any interaction with Hof1 in our Y2H assay (Figure 2F). However, the Y2H assay for detecting protein-protein interactions has inherent limitations, including a bias toward high-affinity interactions and susceptibility to false-positive detection (Brückner et al., 2009). As a result, biologically relevant interactions may be missed, particularly when one or both interacting partners are membrane-associated, self-activating, require PTMs, or interact indirectly through adaptor proteins (Brückner et al., 2009). Given these limitations of the Y2H approach, we performed an in vitro binding assay between the Gin4KA1 domain and the N-terminal region of Hof1N (1–350 aa) to independently validate the direct interaction between these proteins.

We purified 6His-bdSUMO-Gin4-KA1 (1003–1142 aa) and GST-Hof1-N-terminus (1–350 aa) fragments from E. coli. Strikingly, our results revealed that the N-terminal, membrane-binding, F-BAR-containing domain of Hof1 (1–350 aa) interacts strongly with the C-terminal membrane-binding KA1 domain of Gin4 (1003–1142 aa) (Figure 2H). The Hof1 N-terminal F-BAR fragment did not pull down the control 6His-bdSUMO fragment but specifically pulled down 6His-bdSUMO-Gin4KA1 under identical experimental conditions (Figure 2H), confirming the specificity of the interaction between Hof1 F-BAR domain and Gin4-KA1.

Overall, these results reveal a direct physical link between the Gin4 and the AMR component Hof1, suggesting a novel and direct mode of regulation of cytokinesis by the septin kinase network in S. cerevisiae. The involvement of the membrane-binding interfaces of these proteins also suggests a possible coordinative mechanism in which Gin4 may regulate Hof1 to control its proper assembly and anchoring to the membrane prior to cytokinesis (Figure 2I). This is supported by the disorganized and irregular Hof1 ring at the cell division site observed in gin4Δ cells (Figure 2D and E). Moreover, gin4Δ cells also show asymmetric AMR constriction (Figure 2A), which is a known characteristic phenotype of hof1Δ cells (Nishihama et al., 2009). Thus, our data, combined with this line of evidence, strongly suggest that Gin4 exerts direct control over AMR dynamics by physically interacting with Hof1.

To further understand the direct role of Gin4’s membrane-binding KA1 domain in cytokinesis and Hof1 dynamics, we generated an endogenously tagged Gin4-ka1Δ-GFP strain and compared its localization kinetics with those of Gin4-GFP during mitotic spindle breakdown. We observed a marked reduction in Gin4-ka1Δ-GFP localization at the bud neck during cytokinesis, suggesting that the KA1 domain contributes to the recruitment of Gin4 (Figure 2—figure supplement 2A–C). In addition, we analyzed the temporal kinetics of Hof1-mNG at the bud neck in both gin4Δ and Gin4-ka1Δ cells. Consistent with previous findings by Moravcevic et al., 2010, the Gin4-ka1Δ mutant phenocopied the gin4Δ deletion, exhibiting a clumped morphology (Moravcevic et al., 2010). In both gin4Δ and Gin4-ka1Δ cells, Hof1-mNG exhibited a comparable constriction pattern after mitotic spindle breakdown, extending beyond 20 min (Figure 2—figure supplement 2D, E). Additionally, Hof1-mNG exhibited similar localization kinetics during cytokinesis in gin4Δ and Gin4-ka1Δ cells (Figure 2—figure supplement 2D, F). Taken together, these results strongly suggest that the Gin4-KA1 domain is indispensable for Gin4 function and may directly regulate AMR maturation and constriction by influencing Hof1 dynamics at the cell division site.

Gin4 exerts its control over septin organization and AMR components in a kinase-independent manner

Elm1 kinase activity is essential for its localization to the bud neck but is dispensable for its role in septin stability (Marquardt et al., 2020). Bni5 is a myosin-recruiting protein that stabilizes and crosslinks septin filaments by binding to the terminal septin Cdc11 (Fang et al., 2010; Booth et al., 2016; Okada et al., 2025; Patasi et al., 2015). Elm1 regulates the septin hourglass structure and stability through its binding partner Bni5, independent of its kinase activity (Marquardt et al., 2020). We therefore asked whether Gin4 kinase activity is required for its functions in the regulation of septin organization and cytokinesis. To address this, we complemented gin4Δ cells with the kinase-dead gin4-K48A allele (Marquardt et al., 2024; Altman and Kellogg, 1997; Meitinger and Pereira, 2017) (henceforth referred to as Gin4KD), integrated into the leu2 locus. Interestingly, Gin4KD-GFP localized to the bud neck, similar to endogenously tagged Gin4 and Gin4FL-GFP, and complemented the septin mislocalization defects during bud emergence in 78% of total gin4Δ cells (Figure 3A, B). Additionally, expression of Gin4KD-GFP in gin4Δ cells restored the residence time of Inn1-mNG at the bud neck to ~5–7 min and the temporal kinetics during cytokinesis, similar to the wild-type strain (Figure 3C–E). Furthermore, Gin4KD restored Hof1 misorganization defects at the cell division site when expressed in gin4Δ cells in 77.14% of the cellular population (Figure 2—figure supplement 1F, G). Gin4 kinase is known to stabilize septins by phosphorylating its substrate, Shs1 (Mortensen et al., 2002). Hof1 has been shown to exhibit synthetic lethality with Gin4, possibly by regulating septins in a parallel pathway (Meitinger et al., 2013). To test whether this synthetic lethality depends on the kinase activity of Gin4, we expressed Gin4FL and Gin4KD in cells carrying the gin4Δ hof1Δ double deletion (Figure 2—figure supplement 1I). Interestingly, Gin4KD rescued the synthetic lethality, similar to Gin4FL, suggesting that the kinase activity of Gin4 is not required to restore viability in the absence of Hof1 (Figure 2—figure supplement 1I). These findings provide strong evidence that Gin4 kinase activity is largely dispensable for its localization and for its functions in septin organization and cytokinesis, addressing an important aspect of how Gin4 may perform these functions in the absence of catalysis while raising several new questions.

Figure 3
Download asset Open asset
Gin4 controls septin organization and AMR dynamics independently of its kinase activity.

(A) Representative images showing septin defects or mislocalization during bud emergence in gin4Δ cells and rescue of the mislocalization by Gin4FL-GFP and Gin4KD-GFP constructs expressed under pTEF promoter. White arrows indicate mislocalized Cdc3-mcherry in the represented strains. Scale bar: 5 µm. (B) Quantitation of septin mislocalization and its rescue in the indicated strains shown in (A) (wild-type: n = 38, gin4Δ: n = 32, gin4Δ-Gin4FL-GFP: n = 32, gin4Δ-Gin4KD-GFP: n = 36 cells). (C) Representative time-lapse images showing Inn1-mNG dynamics acquired at 2-min time intervals in gin4Δ cells expressing Gin4FL and Gin4KD constructs cloned under endogenous promoter (t0 = appearance of Inn1-mNG at the bud neck). (D) Plot of normalized fluorescence intensity of Inn1-mNG in indicated strains shown in (C). A population of gin4Δ-Empty vector cells exhibiting asymmetric constriction is plotted in the graph (t0 = spindle breakpoint) (wild-type: n = 38, gin4Δ-Empty vector: n = 21, gin4Δ-Gin4FL: n = 39, gin4Δ-Gin4KD: n = 31 cells). (E) Quantification of the residence time of Inn1-mNG during cytokinesis in indicated strains of (C).Kruskal–Wallis nonparametric test (****p < 0.0001, ns: p > 0.05). Whiskers represent mean ± 1.5 × SD.

Figure 3—source data 1

Numerical data underlying the quantitative analyses represented in Figure 3.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig3-data1-v1.xlsx

Hsl1 kinase can partially compensate for Elm1 function upon forced tethering to the bud neck

Marquardt et al., 2024 showed that Gin4 and Elm1 exhibit reciprocal regulation during different cell cycle stages, and that their localization to the bud neck is interdependent (Marquardt et al., 2024). The study also reported that Gin4 recruitment and localization to the bud neck is hampered as the cell cycle progresses in elm1Δ cells (Marquardt et al., 2024). We independently corroborated these results and observed that Gin4 was mislocalized to the bud cortex during bud emergence (Figure 4—figure supplement 1A) and exhibited defective localization to the bud neck during later cell cycle stages in elm1Δ cells (Figure 4—figure supplement 1B–D). Next, we asked whether restoring Gin4 localization to the bud neck could rescue the elongated morphology in cells lacking Elm1. To test this hypothesis, we performed an artificial tethering assay using the GFP-GBP strategy (Figure 4A). The GFP-binding protein (GBP) is a ~13 kDa soluble protein derived from the heavy chain of camelid (llama) antibodies (Rothbauer et al., 2006). This single-domain antibody binds GFP-tagged proteins with very high affinity, in the picomolar range. GBP also binds other GFP variants, such as Yellow Fluorescent Protein (YFP), with a strict 1:1 stoichiometric ratio (Schornack et al., 2009). By contrast, GBP does not bind to Cyan Fluorescent Protein (CFP) or to derivatives of DsRed, including monomeric Red Fluorescent Protein (mRFP) and mOrange (Schornack et al., 2009). Owing to its high binding affinity and specificity, the GFP-GBP system was initially developed for the purification of GFP-tagged proteins (Rothbauer et al., 2006; Schornack et al., 2009). More recently, this strategy has been adapted to artificially redirect proteins to non-canonical subcellular locations (Marquardt et al., 2024; Rothbauer et al., 2006; Schornack et al., 2009). Based on this approach, we tagged the terminal septin Shs1 (Mortensen et al., 2002; Egelhofer et al., 2008) and the bud neck-localizing septin-associated proteins (SAPs) the anillin-like protein – Bud4 (Chen et al., 2020), Bni5 (Marquardt et al., 2020), and Hsl1 with GBP at their C-termini. These GBP-tagged proteins were then used to redirect mislocalized Gin4-GFP to the bud neck, allowing us to assess whether forced relocalization of Gin4 could rescue the elongated morphology exhibited by elm1Δ cells.

Figure 4 with 1 supplement see all
Download asset Open asset
Artificial tethering of Gin4-GFP to the bud neck via Hsl1-GBP restores functionality in elm1Δ cells in a Gin4 kinase-independent manner.

(A) Schematic representation of GFP-GBP artificial tethering strategy, in which the protein of interest is targeted to the bud neck by C-terminal GFP tagging, while the binding partner is tagged with GBP. Created with BioRender.com. (B) Representative images showing artificial tethering of Gin4-GFP with Hsl1-GBP in elm1Δ cells. Scale bar: 5 µm. (C) Bar graph showing the percentage of cells exhibiting round morphology in the indicated strains shown in (B).One-way ANOVA with Tukey’s multiple-comparison test (**p < 0.01, ****p < 0.0001).Error bars indicate + SD from the mean (N = 3, wild-type: n = 407, elm1Δ: n = 508, elm1Δ-Hsl1-GBP: n = 400 cells). (D) Quantification of aspect ratios in the indicated strains shown in (B).Kruskal–Wallis nonparametric statistical test (****p < 0.0001). Whiskers represent mean ± 1.5 × SD (N = 3, n > 150 cells/strain). (E) Graph depicting the raw fluorescence intensity of Gin4-GFP at the large bud in the indicated strains shown in (B).Kruskal–Wallis nonparametric statistical test (***p < 0.001, ****p < 0.0001). Whiskers represent mean ± 1.5 × SD (N = 3, wild-type: n = 125, elm1Δ: n = 125, elm1Δ-Hsl1-GBP: n = 118 cells). (F) Representative images showing the localization of Gin4FL-GFP and Gin4KD-GFP constructs expressed under the pTEF promoter and their tethering to the bud neck with Hsl1-GBP in elm1Δ gin4Δ double-deletion strains. Scale bar: 5 µm. (G) Stacked column graph showing the percentage of cells exhibiting round and elongated/clumped morphologies in the indicated strains shown in (F).One-way ANOVA with Tukey’s multiple-comparison test (****p < 0.0001, ns: p > 0.05). Error bars indicate + SD from the mean. p-values correspond to the population of cells exhibiting round morphology (N = 3, elm1Δ gin4Δ: n = 339, elm1Δ gin4Δ-Empty vector: n = 327, elm1Δ gin4Δ-Gin4FL: n = 173, elm1Δ gin4Δ-Gin4KD: n = 236, elm1Δ gin4Δ-Empty vector-Hsl1-GBP: n = 356, elm1Δ gin4Δ-Gin4FL-Hsl1-GBP: n = 365, elm1Δ gin4Δ-Gin4KD-Hsl1-GBP: n = 459 cells). (H) Quantification of aspect ratios in the indicated strains shown in (F).Kruskal–Wallis nonparametric statistical test (****p < 0.0001, ns: p > 0.05).Whiskers represent mean ± 1.5 × SD (N = 3, n > 90 cells/strain). (I) Representative images showing artificial tethering of Gin4-GFP with Hsl1-ka1Δ-GBP in elm1Δ cells. Scale bar: 5 µm. (J) Stacked column graph showing the percentage of cells exhibiting round and elongated/clumped morphologies in the indicated strains shown in (I).One-way ANOVA with Tukey’s multiple comparison test (****p < 0.0001, ns: p > 0.05).Error bars indicate + SD from the mean. p-values correspond to the population of cells exhibiting round morphology (N = 3, wild-type: n = 674, Hsl1-ka1Δ-GBP: n = 419, elm1Δ: n = 572, elm1Δ Hsl1-ka1Δ-GBP: n = 337 cells). (K) Quantification of aspect ratios in the indicated strains shown in (I).Kruskal–Wallis nonparametric statistical test (****p < 0.0001, ns: p > 0.05). Whiskers represent mean ± 1.5 × SD (N = 3, n > 150 cells/strain).

Figure 4—source data 1

Numerical data underlying the quantitative analyses represented in Figure 4.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig4-data1-v1.xlsx

Forced tethering of Gin4-GFP to the bud neck via its related kinase Hsl1-GBP restored normal cellular morphology in 63.75% of the population in elm1Δ cells (Figure 4B, C, Table 1). To quantitatively assess this rescue, we also analyzed the aspect ratio, measured as the ratio of the major axis to the minor axis of the cell. As expected, the mean aspect ratio was higher in elm1Δ cells than in wild-type cells, reflecting their elongated morphology (Figure 4D), and decreased in cells where Gin4-GFP was tethered to the bud neck with Hsl1-GBP, corroborating the rescue of cell morphology observed after restoring Gin4-GFP localization to the bud neck (Figure 4D). We measured fluorescence intensities of Gin4-GFP with and without tethering to the bud neck via Hsl1-GBP in large-budded elm1Δ cells, which validated that Gin4-GFP was indeed tethered to the bud neck (Figure 4E). In contrast to the rescue observed after tethering with Hsl1-GBP, artificial tethering of Gin4-GFP via other bud neck proteins Shs1, Bni5, or Bud4 did not rescue the elongated phenotype of elm1Δ cells, despite restoration of bud neck localization (Figure 4—figure supplement 1E–G). These results suggest that the observed rescue may be due to two underlying causes: (1) restoration of Gin4 protein at the bud neck and (2) tethering of Gin4 to Hsl1.

Table 1
Table summarizing the rescue of elongated or clumped morphology upon redirecting the Nim1-related kinases to the bud neck in elm1Δ cells and Elm1-GFP in gin4Δ cells using GFP-GBP artificial tethering strategy.
Protein tagged with GFPProtein tagged with GBPGenetic backgroundNumber of cells exhibiting round morphologyNumber of cells exhibiting elongated/clumped morphologyTotal number of cellsPercentage of cellular population exhibiting round morphologyRescue of elongated or clumped morphology
Gin4Shs1elm1Δ244644884.91No
Gin4Bud4elm1Δ204514714.24No
Gin4Hsl1elm1Δ25514540063.75Yes
Gin4Hsl1-ka1Δelm1Δ63313371.78No
Gin4Bni5elm1Δ174314483.79No
Gin4-Full lengthHsl1elm1Δ gin4Δ2828336577.26Yes
Gin4-Kinase deadHsl1elm1Δ gin4Δ30115845965.57Yes
Hsl1Shs1elm1Δ26446272636.36Yes
Hsl1Bud4elm1Δ11060771715.34No
Hsl1Bni5elm1Δ43543581.11No
Hsl1Gin4elm1Δ52123375469.09Yes
Hsl1Kcc4elm1Δ49836486257.77Yes
Kcc4Shs1elm1Δ215665873.57No
Kcc4Bud4elm1Δ606036639.04No
Kcc4Hsl1elm1Δ51619270872.88Yes
Kcc4Bni5elm1Δ65635691.05No
Kcc4Gin4elm1Δ11936448324.63No
Elm1Bud4gin4Δ61627088669.52Yes
Elm1Hsl1gin4Δ39928968857.99Yes
Elm1Hsl1-ka1Δgin4Δ13947060922.82No
Elm1Shs1gin4Δ49930180062.37Yes

  1. Bold text indicates conditions in which artificial tethering resulted in rescue of the elongated or clumped cellular morphology, as indicated in the final column.

To test whether Gin4 kinase activity is required for the observed rescue in cellular morphology upon artificial tethering of Gin4-GFP to the bud neck with Hsl1-GBP. We tethered Gin4FL-GFP and Gin4KD-GFP, expressed under the pTEF promoter, via Hsl1-GBP in gin4Δ elm1Δ cells, and observed that both constructs restored cellular morphology and aspect ratio to wild-type levels in 77.26% and 65.57% of cells, respectively (Figure 4F–H, Table 1). These results suggest that Gin4 tethering to the bud neck via Hsl1 may compensate for Elm1 function in a kinase-independent manner, further reinforcing the kinase-independent nature of Gin4 function. Because Hsl1 also contains a membrane-binding C-terminal KA1 domain (Finnigan et al., 2016; Moravcevic et al., 2010), we tested its requirement for the observed functional rescue in elm1Δ cells and found that artificial tethering of Gin4-GFP via Hsl1-ka1Δ-GBP did not rescue the cellular morphology of elm1Δ cells (Figure 4I–K). Together, these observations suggest that (1) restoring Gin4 localization to the bud neck via Hsl1 is necessary to partially rescue the morphological defects and cell shape exhibited by elm1Δ cells; (2) Gin4 might form a complex with Hsl1, which becomes essential in the absence of Elm1, indicating that Elm1 might play a significant role in regulating the Gin4–Hsl1 complex; and (3) proper membrane tethering of Hsl1 via its KA1 domain may be essential for its function in elm1Δ cells.

Septin and cytokinesis defects are rescued by artificial tethering of Gin4-GFP to the bud neck via Hsl1-GBP in elm1Δ cells

Rescue of cell shape alone does not necessarily imply restoration of underlying physiological processes such as septin organization, AMR dynamics, or cytokinesis. To address this, we analyzed septin dynamics using Cdc3-mCherry in elm1Δ cells with and without artificial tethering of Gin4-GFP to the bud neck via Hsl1-GBP. Specifically, we examined the initial recruitment of septins during bud emergence. Consistent with previous results (Marquardt et al., 2024; Marquardt et al., 2020) and our observations, 100% of elm1Δ cells exhibited mislocalization of Cdc3-mCherry to the bud cortex during bud emergence (Figure 5A, B). By contrast, only 35.71% of cells showed septin mislocalization when Gin4-GFP was artificially tethered to the bud neck via Hsl1-GBP (Figure 5A, B). We also analyzed the localization kinetics of Gin4-GFP in these strains and found that nearly 100% of elm1Δ cells exhibited Gin4-GFP mislocalization to the bud cortex in the absence of tethering, whereas this was reduced to 42.85% upon tethering via Hsl1-GBP (Figure 5A, C). The extent of rescue in Cdc3-mCherry mislocalization (64.28%) and Gin4-GFP mislocalization (57.14%) in elm1Δ cells closely correlated with the fraction of cells that also exhibited rescue of cellular morphology upon tethering with Hsl1-GBP (Figures 4B and C and 5A-C).

Figure 5
Download asset Open asset
Redirecting Gin4 to the bud neck via Hsl1 in elm1Δ cells rescues septin organization and AMR dynamics.

(A) Representative time-lapse montages showing the rescue of mislocalization of both Gin4-GFP and Cdc3-mCherry upon artificial tethering with Hsl1-GBP in elm1Δ cells. Scale bar: 5 µm. (B) Stacked column graph showing the percentage of cells exhibiting normal septin localization or septin mislocalization to bud cortex during the bud emergence in the indicated strains shown in (A) (wild-type: n = 39, Hsl1-GBP: n = 39, elm1Δ: n = 39, elm1Δ-Hsl1-GBP: n = 42 cells). (C) Quantification of Gin4-GFP localization rescue in the indicated strains shown in (A) (wild-type: n = 39, Hsl1-GBP: n = 39, elm1Δ: n = 39, elm1Δ-Hsl1-GBP: n = 42 cells). (D) Representative montages of the bud neck showing the constriction kinetics of Myo1-ymScarletI in elm1Δ strains in which Gin4-GFP is artificially tethered to the bud neck via Hsl1-GBP. (E) Quantitative analysis for normal localization and mislocalization of Myo1-ymScarletI in the indicated strains shown in (D) (wild-type: n = 41, Hsl1-GBP: n = 37, elm1Δ: n = 39, elm1Δ-Hsl1-GBP: n = 44 cells). (F) Graph showing the constriction dynamics of Myo1-ymScarletI in the indicated strains shown in (D). Disappearance of Gin4 from the bud neck is used to normalize the initial time point for Myo1 constriction. Kruskal–Wallis nonparametric statistical test (****p < 0.0001, ns: p > 0.05). Whiskers represent mean ± 1.5 × SD (wild-type: n = 39, Hsl1-GBP: n = 30, elm1Δ: n = 32, elm1Δ-Hsl1-GBP: n = 39 cells).

Figure 5—source data 1

Numerical data underlying the quantitative analyses represented in Figure 5.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig5-data1-v1.xlsx

To determine whether AMR defects were similarly rescued, we analyzed the dynamics of Myo1 C-terminally tagged with ymScarletI during cytokinesis. In elm1Δ cells, 76.92% of cells displayed Myo1 mislocalization to the bud cortex during its recruitment, consistent with previous reports (Marquardt et al., 2020). Upon artificial tethering of Gin4-GFP to the bud neck via Hsl1-GBP, the proportion of cells exhibiting Myo1 mislocalization was reduced to 11.36% (Figure 5D, E). Additionally, we quantified the residence time of Myo1 at the bud neck by defining the disappearance of Gin4 as t0, corresponding to the onset of constriction, and measuring the time intervals until complete loss of the Myo1 signal from the bud neck. In wild-type cells, Myo1 constriction was completed within approximately 4 min, whereas in elm1Δ cells this process was prolonged to nearly 10 min (Figure 5D, F). The residence time was rescued to ~4 min upon tethering Gin4-GFP to the bud neck with Hsl1-GBP in elm1Δ cells (Figure 5D, F).

Taken together, these results indicate that (1) rescue of cellular morphology correlates with restoration of both Gin4 and septin localization dynamics as the cell cycle progresses, suggesting a direct mode of regulation of Gin4 and Hsl1 downstream of Elm1 and (2) restoration of AMR dynamics in elm1Δ cells upon artificial tethering of Gin4 to the bud neck via Hsl1 suggests an unconventional role for Hsl1 in the absence of Elm1.

Rescue of septin localization in elm1Δ cells upon redirecting Gin4-GFP to the bud neck via Hsl1-GBP is largely independent of the morphogenetic checkpoint

Swe1 is a checkpoint kinase that regulates the morphogenetic signaling pathway and plays a key role in controlling cell shape by regulating isometric cell growth (Sreenivasan and Kellogg, 1999; Barral et al., 1999; Bouquin et al., 2000). Previous studies have shown that Elm1, Gin4, Hsl1, and Kcc4 negatively regulate Swe1 through inhibitory phosphorylation, thereby promoting isometric growth and maintaining a round cell morphology (Sreenivasan and Kellogg, 1999; Barral et al., 1999). It has also been reported that deletion of Swe1 in combination with septin kinase deletions (elm1Δ swe1Δ, gin4Δ swe1Δ, or hsl1Δ swe1Δ) rescues the elongated and clumped morphologies associated with the corresponding individual kinase deletions (Sreenivasan and Kellogg, 1999; Bouquin et al., 2000). However, this rescue is specific to cell shape and does not restore septin localization or cytokinesis defects (Sreenivasan and Kellogg, 1999; Bouquin et al., 2000). Based on these observations, we asked whether the rescue of cellular morphology and aspect ratio observed in elm1Δ cells upon artificial redirection of Gin4-GFP to the bud neck via Hsl1-GBP is mediated through negative regulation of Swe1 during mitotic spindle breakdown, or instead represents a direct consequence of restoring Gin4 and Hsl1 localization.

To address this, we deleted Swe1 in elm1Δ cells in which Gin4-GFP was either untethered or artificially tethered to the bud neck using Hsl1-GBP. Consistent with Bouquin et al., 2000, deletion of Swe1 alone rescued the elongated morphology in 60.66% of the elm1Δ population when Gin4-GFP was not tethered to the bud neck (Figure 6—figure supplement 1A–C). Strikingly, deletion of Swe1 did not alter the proportion of round cells in elm1Δ cells in which Gin4-GFP was redirected to the bud neck via Hsl1-GBP (Figure 6—figure supplement 1A–C).

We further analyzed septin organization using Cdc3-mCherry in elm1Δ and elm1Δ swe1Δ cells, with and without artificial tethering of Gin4-GFP to the bud neck via Hsl1-GBP. Although cell shape was rescued in both elm1Δ swe1Δ control cells and elm1Δ swe1Δ cells with Gin4-GFP tethered via Hsl1-GBP, only 23.63% of cells in the elm1Δ swe1Δ control strain exhibited rescue of septin mislocalization during bud emergence (Figure 6A, B). This partial rescue by Swe1 deletion also correlated with restoration of mislocalized Gin4 during bud emergence in elm1Δ cells (Figure 6C, D). Notably, more than 70% of cells exhibited rescue of both Gin4 and Cdc3 mislocalization in elm1Δ swe1Δ cells when Gin4-GFP was artificially tethered to the bud neck via Hsl1-GBP, in contrast to the rescue exhibited by only ~23% of cells in which Gin4-GFP was not tethered (Figure 6A–D).

Figure 6 with 1 supplement see all
Download asset Open asset
Gin4 tethering to the bud neck via Hsl1 bypasses the requirement for Elm1 independently of the morphogenetic checkpoint kinase Swe1.

(A) Time-lapse montages representing the dynamics of Cdc3-mCherry in both elm1Δ and elm1Δ swe1Δ strains upon artificial tethering of Gin4-GFP to bud neck with Hsl1-GBP (t0 = bud emergence). Scale bar: 5 µm. (B) Stacked column graph showing the percentage of cells exhibiting normal localization and mislocalization of Cdc3-mCherry in the indicated strains shown in (A) (wild-type: n = 55, swe1Δ: n = 49, elm1Δ: n = 58, elm1Δ swe1Δ: n = 51, elm1Δ-Gin4-GFP: n = 55, elm1Δ swe1Δ-Gin4-GFP: n = 55, elm1Δ-Gin4-GFP-Hsl1-GBP: n = 58, elm1Δ swe1Δ-Gin4-GFP-Hsl1-GBP: n = 52 cells). (C) Representative time-series montages showing the localization of Gin4-GFP in both elm1Δ and elm1Δ swe1Δ strains upon artificial tethering of Gin4-GFP to the bud neck via Hsl1-GBP (t0 = bud emergence). Scale bar: 5 µm. (D) Quantification graphs depicting the normal localization and mislocalization of Gin4-GFP at the bud cortex during bud emergence in the indicated strains shown in (C) (wild-type: n = 57, swe1Δ: n = 52, Hsl1-GBP: n = 54, Hsl1-GBP swe1Δ: n = 51, elm1Δ-Gin4-GFP: n = 55, elm1Δ swe1Δ-Gin4-GFP: n = 55, elm1Δ-Gin4-GFP-Hsl1-GBP: n = 58, elm1Δ swe1Δ-Gin4-GFP-Hsl1-GBP: n = 52 cells).

Figure 6—source data 1

Numerical data underlying the quantitative analyses represented in Figure 6.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig6-data1-v1.xlsx

Collectively, these findings suggest that (1) the rescue of elongated cell morphology to normal round cell morphology in elm1Δ cells upon artificial tethering of Gin4-GFP to the bud neck via Hsl1-GBP may depend on Hsl1-mediated regulation of the morphogenesis checkpoint; however, (2) the observed rescue in septin and Gin4 localization clearly does not depend on the morphogenesis checkpoint and reflects direct regulation of septin organization by Gin4 and Hsl1.

Intrigued by the rescue in cellular morphology observed only after tethering Gin4-GFP to Hsl1-GBP and not to other bud neck proteins, we extended our assay to its related kinases and tethered Hsl1-GFP and Kcc4-GFP via various bud-neck proteins in elm1Δ cells (Figure 7A–C and Figure 7—figure supplement 1A–C). Our results revealed that Hsl1-GFP restored normal cell morphology in 36.36% of cells in elm1Δ when tethered with Shs1-GBP, while tethering with Gin4-GBP or Kcc4-GBP resulted in significantly higher rescue of 69.09% and 57.77%, respectively, suggesting possible coordination among these kinases (Figure 7A–C, Table 1). Strikingly, like Gin4-GFP, Kcc4-GFP tethering also restored cell morphology similar to wild-type in 72.88% of the population, only when tethered to the bud neck with Hsl1-GBP (Figure 7—figure supplement 1A–C, Table 1). However, Kcc4-GFP tethering via different bud neck proteins did not rescue the phenotype, although a minor rescue in 24.63% of cells was observed upon tethering with Gin4-GBP, suggesting possible crosstalk between Kcc4 and Gin4 (Figure 7—figure supplement 1A–C, Table 1).

Figure 7 with 2 supplements see all
Download asset Open asset
Restoring Hsl1-GFP localization to the bud neck via septins or its related kinases can bypass the requirement of Elm1.

(A) Representative images showing artificial tethering of Hsl1-GFP to the bud neck via Shs1-GBP, Bud4-GBP, Kcc4-GBP, Bni5-GBP, and Gin4-GBP in elm1Δ cells. Scale bar: 5 µm. (B) Bar graph showing the percentage of cells exhibiting round morphology in the indicated strains shown in (A).One-way ANOVA with Tukey’s multiple-comparison test (****p < 0.0001, ns: p > 0.05). Error bars indicate + SD from the mean (N = 3, wild-type: n = 724, elm1Δ: n = 458, elm1Δ-Shs1-GBP: n = 726, elm1Δ-Bud4-GBP: n = 717, elm1Δ-Kcc4-GBP: n = 862, elm1Δ-Bni5-GBP: n = 358, and elm1Δ-Gin4-GBP: n = 754 cells). (C) Quantification of aspect ratios in the indicated strains shown in (A).Kruskal–Wallis nonparametric statistical test (***p < 0.001, ****p < 0.0001). Whiskers represent mean ± 1.5 × SD (N = 3, n > 135 cells/strain). (D) Representative montages of the bud neck depicting the localization of Hsl1-GFP in elm1Δ and gin4Δ cells during cytokinesis. DC* = differential contrast. (E) Plot of raw fluorescence intensity of Hsl1-GFP in the indicated strains shown in (D) (t0 = spindle breakpoint) (wild-type: n = 29, elm1Δ: n = 24, gin4Δ: n = 38 cells). (F) Plot of normalized fluorescence intensity of Hsl1-GFP in the indicated strains shown in (D) (t0 = spindle breakpoint).

Figure 7—source data 1

Numerical data underlying the quantitative analyses represented in Figure 7.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig7-data1-v1.xlsx

To support these results, we analyzed the localization of Hsl1 and Kcc4 in elm1Δ and gin4Δ cells. As reported previously (Marquardt et al., 2020), Kcc4 exhibited mislocalization to the bud cortex during the initial phases of the cell cycle and defective localization at the onset of cytokinesis (Figure 7—figure supplement 2B–E). Recruitment and localization of Hsl1 to the bud neck were also severely impaired in elm1Δ cells (Figure 7D–F, Figure 7—figure supplement 2A), suggesting a role for Elm1 in maintaining Hsl1 localization at the bud neck. By contrast, recruitment and localization of Hsl1 and Kcc4 were similar to wild-type in gin4Δ cells (Figure 7D–F, Figure 7—figure supplement 2A–E). These data suggest that loss of Hsl1 at the bud neck may contribute significantly to the elongated phenotype observed in elm1Δ cells and, consistent with this idea, restoration of Hsl1 localization rescues the observed morphological defects in our experiments. The degree of phenotypic rescue appears to depend on the presence of Hsl1 at the bud neck, and higher rescue percentages are observed when Hsl1 is tethered with another kinase (Gin4 or Kcc4), suggesting coordinated effects among these kinases, and that Hsl1 may act as a downstream effector in the septin kinase network.

Our results reveal that Hsl1 regulates normal cell morphology, possibly through downstream signaling events triggered by the presence of Elm1 in wild-type cells, in coordination with either Gin4 or Kcc4. Thus, Elm1 may act upstream to control the localization and activation of the Nim1-related kinases (Hsl1, Gin4, and Kcc4) at the septin scaffold in a precise, spatiotemporal manner, within a network with multiple feedback regulatory links. Overall, the ability to restore normal function by simply rewiring the septin kinase network demonstrates the inherent redundancy and robustness of this network in S. cerevisiae.

Hsl1 and its membrane-binding ability are required for the rescue of gin4Δ cells by artificially tethered Elm1

Elm1 does not localize to the bud neck in the absence of Gin4 and can rescue the elongated/clumped phenotype exhibited by gin4Δ cells upon artificial tethering to the bud neck septin scaffold (Marquardt et al., 2024). We corroborated these findings independently and found that Elm1 is not recruited to the bud neck in gin4Δ cells (Figure 8—figure supplement 1A–D). We found that Elm1-GFP tethering via Shs1-GBP, Bud4-GBP, or Hsl1-GBP could restore normal cell morphology and aspect ratio in gin4Δ cells (Figure 8—figure supplement 1E–G). We therefore asked whether the presence of Hsl1 is necessary for the observed rescue upon Elm1 tethering to the bud neck in gin4Δ cells. To test this, we artificially tethered Elm1-GFP to the bud neck via Shs1-GBP in gin4Δ and gin4Δ hsl1Δ cells. While 62.37% of gin4Δ cells showed restoration of cell morphology and aspect ratio, no rescue was observed in the gin4Δ hsl1Δ double deletion strain upon tethering Elm1 to the bud neck, indicating that Hsl1 is required downstream of Elm1 tethering in gin4Δ cells (Figure 8A, C, D, Table 1). Additionally, we observed rescue of septin misorganization upon tethering Elm1-GFP to the bud neck with Shs1-GBP in gin4Δ cells in 82.92% of the cellular population, which was lost in gin4Δ hsl1Δ cells in >75% of the population (Figure 8A, E). Redirected Elm1-GFP also showed mislocalization to the bud cortex during bud emergence in gin4Δ hsl1Δ cells (Figure 8F). A loss of rescue was observed when Elm1-GFP was tethered to the bud neck in gin4Δ cells via Hsl1-ka1Δ-GBP, suggesting that Hsl1 acts downstream of Elm1 and that its membrane-binding KA1 domain is required for its function (Figure 8—figure supplement 1H–J). These data support the idea that Hsl1 acts downstream of Elm1 and Gin4 in the septin kinase network to control septin stability.

Figure 8 with 2 supplements see all
Download asset Open asset
A non-canonical role for Hsl1-Kinase in regulating septin organization and AMR dynamics.

(A) Representative images showing Cdc3-mCherry mislocalization during bud emergence in gin4Δ and gin4Δ hsl1Δ cells in which Elm1-GFP is artificially tethered to bud neck via Shs1-GBP. White arrows indicate mislocalized Cdc3-mCherry in the represented strains. Scale bar: 5 µm. (B) Representative time-lapse montages of the bud neck showing the constriction profile of Inn1-3xmCherry in gin4Δ and gin4Δ hsl1Δ cells in which Elm1-GFP is artificially tethered to bud neck via Shs1-GBP. (C) Bar graph showing the percentage of cells exhibiting round and elongated morphologies in the indicated strains shown in (A).One-way ANOVA with Tukey’s multiple-comparison test (**p < 0.01, ***p < 0.001, ns: p > 0.05). Error bars indicate + SD from the mean. p-values correspond to population of cells exhibiting round morphology (N = 3, gin4Δ: n = 689, gin4Δ-Shs1-GBP: n = 639, gin4Δ hsl1Δ: n = 492, gin4Δ hsl1Δ-Shs1-GBP: n = 590 cells). (D) Quantification of aspect ratios in the indicated strains shown in (A). Kruskal–Wallis nonparametric statistical test (****:p < 0.0001, ns: p > 0.05). Whiskers represent mean ± 1.5 × SD (N = 3, n > 165 cells/strain). (E) Stacked column graph representing the percentage of cells exhibiting normal septin localization and septin mislocalization in the indicated strains of (A) (gin4Δ: n = 44, gin4Δ-Shs1-GBP: n = 41, gin4Δ hsl1Δ: n = 38, gin4Δ hsl1Δ-Shs1-GBP: n = 44 cells). (F) Quantification of normal localization and mislocalization of Elm1-GFP in the indicated strains shown in (A) (gin4Δ-Shs1-GBP: n = 37, gin4Δ hsl1Δ-Shs1-GBP: n = 44 cells) (G) Quantification of the residence time of Inn1-3xmcherry during cytokinesis in the indicated strains shown in (B). Kruskal–Wallis nonparametric statistical test (*p < 0.05, **p < 0.01, ****p < 0.0001, ns: p > 0.05). Whiskers represent mean ± 1.5 × SD (N = 2, n > 60 cells/strain).

Figure 8—source data 1

Numerical data underlying the quantitative analyses represented in Figure 8.

https://cdn.elifesciences.org/articles/106366/elife-106366-fig8-data1-v1.xlsx

Next, we asked whether Elm1-GFP tethering to the bud neck in gin4Δ cells could rescue other observed cellular defects related to AMR dynamics during cytokinesis. To test this, we first assessed the recruitment and localization of the Myo1 during bud emergence. We observed that Myo1-3xmCherry exhibited mislocalization to the bud tip in 12.82% gin4Δ cells during bud emergence (Figure 8—figure supplement 2A, B). However, Myo1-3xmCherry localization was restored to the bud neck upon artificial tethering of Elm1-GFP in gin4Δ cells. The cellular population with mislocalized Myo1-3xmCherry increased further upon deletion of Hsl1 in gin4Δ cells, even when Elm1-GFP was artificially tethered to the bud neck with Shs1-GBP, reaching up to 87.17% of the cellular population (Figure 8—figure supplement 2A, B). We also assessed the residence time of Inn1-3xmCherry in these cells to measure constriction dynamics of the AMR (Sanchez-Diaz et al., 2008; Nishihama et al., 2009). We found that the Inn1-3xmCherry residence time was similar to that of wild-type cells when Elm1-GFP was tethered via Shs1-GBP in gin4Δ cells, but was significantly increased when Elm1-GFP was tethered via Shs1-GBP in gin4Δ hsl1Δ cells (Figure 8B, G). These results suggest that (1) Elm1 tethering to the bud neck restores normal Myo1 localization and AMR constriction dynamics in the absence of Gin4, (2) Hsl1 is required for Elm1 to exert its influence over AMR organization and dynamics after bud neck tethering in gin4Δ cells, and (3) the membrane-binding KA1 domain of Hsl1 is required for this activity downstream of Gin4.

Taken together, our results suggest that Hsl1 functions both downstream of and in coordination with Elm1 and Gin4 to maintain septin stability and ensure proper cytokinesis (Figure 9). In wild-type cells, Hsl1 likely acts downstream of Elm1 and Gin4 and localizes normally to the bud neck, where it may help coordinate septin architecture with AMR constriction during cytokinesis (Figure 9). In the absence of Elm1, localization of both Gin4 and Hsl1 becomes defective during later cell cycle stages, disrupting the potential crosstalk between Gin4 and Hsl1. This disruption may lead to septin mislocalization and disorganization of the AMR machinery (Figure 9). Notably, restoring Hsl1 localization using a GFP-GBP tethering strategy strengthens this link, suggesting that Elm1 plays an important role in recruiting or stabilizing Hsl1 at the bud neck. These observations indicate that Hsl1 acts downstream of Elm1 in this pathway, with partially overlapping functions. Furthermore, the requirement for Hsl1 in the rescue observed upon Elm1 tethering in gin4Δ cells supports the idea that Hsl1 mediates downstream effects of Elm1 in regulating septin stability and cytokinesis. Therefore, we propose that Hsl1, through a compensatory pathway, facilitates functional crosstalk between the septin kinase network and septin/AMR components, which is essential for proper septin organization and symmetric AMR constriction, ultimately ensuring faithful cytokinesis.

Figure 9
Download asset Open asset
Representative model for the role of Hsl1 kinase in septin organization and AMR constriction downstream of Gin4 and Elm1.

Created with BioRender.com.

Discussion

How septin architecture remodeling crosstalk with the AMR machinery and cytokinesis remains a dynamic area of investigation, as these processes are spatiotemporally controlled by septin-interacting proteins and PTMs (Perez et al., 2016; Dobbelaere et al., 2003; Hernández-Rodríguez and Momany, 2012; Johnson and Blobel, 1999; Marquardt et al., 2019; McMurray and Thorner, 2009; Mitchell et al., 2011; Takahashi et al., 1999). Yet the regulatory mechanisms governing septin remodeling and AMR constriction during the cell cycle in yeast and higher eukaryotes remain incompletely understood. In S. cerevisiae, four septin-associated kinases Elm1, Gin4, Hsl1, and Kcc4 have been reported to primarily regulate septin-dependent processes (Marquardt et al., 2024; Marquardt et al., 2020; Barral et al., 1999; Bouquin et al., 2000). However, the precise mechanisms by which these kinases influence septin architecture and coordinate cytokinesis remain poorly understood. In this study, we identify critical roles for Elm1 and Gin4 in maintaining proper septin organization and AMR dynamics during cytokinesis. Our results further reveal that Gin4 exerts its functions in septin stability and AMR dynamics in a kinase-independent manner, challenging the prior view of kinase-dependent functions. We also identify a direct physical interaction between Gin4 and the AMR protein Hof1, providing new insights into how septin regulatory pathways directly crosstalk with the cytokinetic machinery.

Septin remodeling marks the onset of AMR constriction and cytokinesis (Wloka et al., 2011; Bi et al., 1998; Tamborrini et al., 2018; Varela Salgado et al., 2024), but the molecular mechanisms coordinating these processes remain largely unclear. We observed that septins are severely mislocalized to the bud cortex in elm1Δ and gin4Δ cells, supporting previous reports that these kinases are required for proper septin organization at the bud neck (Marquardt et al., 2024; Marquardt et al., 2020; Barral et al., 1999). Consistent with these defects, the dynamics of the AMR complex proteins Inn1 and Hof1 were significantly perturbed, leading to delayed constriction in the absence of Elm1 and Gin4 (Figures 1D–F–2A–E). These observations suggest that septin-associated kinases regulate intermediate signaling events that coordinate septin architecture with the assembly and stability of the cytokinetic machinery, and they highlight an underappreciated mode of regulation of AMR organization and dynamics.

A key question arising from these observations is whether the crosstalk between the septin kinase network and AMR dynamics is direct and septin-independent, or indirect and septin-dependent. Previous studies have shown that AMR constriction can proceed in certain septin mutants that disrupt septin organization, suggesting that septin kinases may also exert direct regulatory effects on the cytokinetic machinery (Wloka et al., 2011), further supporting a direct contribution of Elm1 and Gin4 to the dysregulated cytokinetic events observed in this study. Supporting this idea, we found that Gin4 interacts with the F-BAR protein Hof1, as demonstrated by Y2H and in vitro binding assays (Figure 2F–H). Notably, the membrane-binding domains of Gin4 and Hof1 were sufficient for this interaction, suggesting that Gin4 activation at the membrane (Jasani et al., 2020; Moravcevic et al., 2010) may directly influence Hof1 organization and function during AMR maturation (Figure 2H). These findings suggest a potential mechanism for anchoring Hof1 to the plasma membrane prior to cytokinesis (Figure 2I). Interestingly, previous work has shown that gin4Δ and hof1Δ are synthetic lethal and that Hof1 exhibits mislocalization in gin4Δ cells (Meitinger et al., 2013). These results also point to a coordinative mechanism in which Gin4 directly crosstalks with an AMR component, Hof1, to regulate AMR organization and constriction. To our knowledge, this represents the first evidence physically linking the septin kinase network to AMR proteins and highlights a novel regulatory input on AMR assembly in S. cerevisiae. Future work is required to delineate the molecular mechanisms underlying this regulation.

Interestingly, our results further demonstrate that the kinase activity of Gin4 is not required for its role in septin organization, AMR constriction (Figure 3A–E), or cell viability in the absence of Hof1 (Figure 2—figure supplement 1I). This highlights an important aspect of Gin4 function in cells and is similar to the kinase-independent function reported for Elm1 (Marquardt et al., 2020). These observations raise an important question regarding the functional significance of kinase activity within the septin kinase network. Our results, together with previous studies (Marquardt et al., 2024; Barral et al., 1999), establish that built-in functional redundancies exist among the septin kinases (Elm1, Gin4, Hsl1, and Kcc4), and that their recruitment to the bud neck is also interdependent. Kinase activities may play an important role in regulating the signaling that coordinates the precise arrival and spatiotemporal activation of these kinases at the bud neck. These results also raise important questions about post-translational regulatory mechanisms of septin filament organization, as a number of studies have shown that these kinases directly phosphorylate septins (Mortensen et al., 2002). Thus, the role of post-translational regulation of septins will be better understood with future work in this direction.

Although these kinases exhibit distinct functions, they also display mutual interdependence (Marquardt et al., 2024; Asano et al., 2006). Recent studies have proposed a new perspective regarding this crosstalk, revealing reciprocal regulation between Gin4 and Elm1 through direct binding and phosphorylation at different stages of the cell cycle (Marquardt et al., 2024). The existence of redundancy in the septin kinase network explains the longstanding differences in the phenotypes observed in elm1Δ and gin4Δ cells. The elm1Δ cells show more pronounced abnormalities in cell shape and septin organization than gin4Δ cells, even though Elm1 is known to be absent from the bud neck in gin4Δ cells (Marquardt et al., 2024; Sreenivasan and Kellogg, 1999; Barral et al., 1999). Our analysis of the temporal localization dynamics of Hsl1 and Kcc4 suggests a potential explanation for this discrepancy. Specifically, the recruitment of Hsl1 and Kcc4 to the bud neck was strongly perturbed in elm1Δ cells but remained largely intact in gin4Δ cells (Figure 7D–F, Figure 7—figure supplement 2A–E). These findings suggest that the localization of Hsl1 and Kcc4 to the bud neck is crucial for their normal function and may explain the less prominent phenotype observed following loss of Gin4, emphasizing the redundant roles of Hsl1 and Kcc4 in septin organization.

Consistent with this interpretation, our GFP-GBP tethering experiments demonstrated that restoring localization of Hsl1 in elm1Δ cells, together with Gin4 and Kcc4, can bypass the requirement for Elm1 during the cell cycle, independent of its function in the morphogenetic checkpoint pathway (Figures 4B–E–6A–D, 7A–C and Figure 7—figure supplement 1A–C). These findings highlight a previously underappreciated level of functional redundancy among the Nim1-related kinases. Our results also suggest that Hsl1 can play a non-canonical role in regulating septin organization and AMR dynamics downstream of Elm1 and Gin4 (Figures 5A–F–8A–G and Figure 8—figure supplement 2A, B). We therefore speculate that synergy exists among Elm1, Gin4, and Hsl1 to regulate downstream signaling pathways that control septin organization and cytokinesis. Investigating the requirement for Hsl1 kinase activity in the absence of Elm1 or Gin4 may provide further insight into the molecular mechanisms underlying Hsl1 regulation. Additionally, elucidating the phosphorylation-dependent modifications involved, along with identifying the specific regulatory residues in future studies, will help clarify the molecular mechanisms by which these kinases coordinate with one another to ensure the precise execution of cytokinesis.

Strikingly, deletion of the membrane-binding KA1 domain of Hsl1 abolished its ability to rescue the elongated defect exhibited in the absence of Elm1 (Figure 4I–K and Figure 8—figure supplement 1H–J), emphasizing the importance of membrane association for the proper function of these kinases (Moravcevic et al., 2010). Membrane binding may facilitate kinase activation or enable interactions with septin filaments and other regulatory proteins at the bud neck. However, the precise mechanisms by which membrane association contributes to Hsl1 function remain unclear and will require further investigation. The underlying molecular mechanisms and actual cause-and-effect relationships by which Hsl1 and its related kinases regulate septin organization at the filament-level remain to be understood. Approaches such as platinum-replica electron microscopy and in vitro reconstitution assays may prove instrumental in dissecting how these kinases influence septin filament organization and higher-order assembly in future studies (Cannon et al., 2019; Marquardt et al., 2020; Varela Salgado et al., 2024). Such studies will provide critical insights into the modes of regulation of septins through the coordinated action and signaling inputs from multi-septin kinase modules across eukaryotes.

In summary, our study provides a comprehensive quantitative analysis of septin-regulatory kinases in controlling septin organization and cytokinesis. We identify Elm1 and Gin4 as key regulators of septin architecture and uncover previously uncharacterized roles for these kinases in AMR organization and constriction. These findings pave the way for future investigations into the intricate molecular mechanisms and cross-regulatory interactions within the septin-kinase network and with AMR proteins during the cell cycle in S. cerevisiae and other related organisms, where septins and their associated proteins play key roles in processes such as cell division.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain backgroundEscherichia coli-DH5aNew England BiolabsCatalog # C2987I
Strain, strain backgroundEscherichia coli-Top10Thermo Fisher ScientificCatalog number: C404003
Strain, strain backgroundEscherichia coli-BL21(DE3)New England BiolabsCatalog #
C2527
Strain, strain backgroundSaccharomyces cerevisiae S288C-background strainsThis paperYeast strains used in this study are listed in Supplementary file 1. Both previously reported and newly generated strains, together with their respective references, are indicated therein. All strains are available from the corresponding author upon request.
Chemical compound, drugG418-DisulphateSisco Research Laboratories Pvt. Ltd.Cat#:58327
Chemical compound, drugHygromycin BSisco Research Laboratories Pvt. Ltd.Cat#:67317
Chemical compound, drugNourseothricinJena BioscienceCat#:AB-102
Chemical compound, drugAmpicillinG-BiosciencesCat#AB1005Lot#201512
Chemical compound, drugKanamycinG-BiosciencesCat#AB1025Lot#230201
Commercial assay or kitNEBuilder HiFi DNA Assembly Master MixNew England BiolabsCat#:E262L
Commercial assay or kitGET Plasmid MiniprepG-BiosciencesCat#:786-361
Commercial assay or kitQIAquick Gel Extraction KitQIAGENCat#:28704
Commercial assay or kitGeneJET PCR Purification KitThermo Fisher ScientificCat#:K0701
Sequence-based reagentPrimers used in this studyThis paperOligonucleotide sequences used in this study are provided in Supplementary file 2.
Recombinant DNA reagent/ PlasmidsPlasmids used in this studyThis paperSupplementary file 3 contains a list of all plasmids used in this study, including newly generated constructs and plasmids described previously, along with their respective references. These plasmids will be made available by the corresponding author upon request.
Software, algorithmFijiFijiRRID:SCR_002285https://imagej.net/software/fiji/
Software, algorithmOriginPro Pro (2015, Sr2, 69.2.272)OriginLab CorporationRRID:SCR_014212https://www.originlab.com/
Software, algorithmCellSens Dimension 3.1OlympusRRID:SCR_014551https://www.olympus-lifescience.com/en/software/cellsens/
Software, algorithmGraphPad Prism 6.04GraphPad SoftwareRRID:SCR_002798https://www.graphpad.com/
Software, algorithmAndor FusionOxford InstrumentsRRID:SCR_023609https://andor.oxinst.com/downloads/view/fusion-release-2.3
Software, algorithmInkscapeInkscape ProjectRRID:SCR_014479Used for figure preparation
Software, algorithmBioRenderBioRender.comRRID:SCR_018361Used for figure preparation (Figures 2I, 4A, and 9)
Software, algorithmAdobe IllustratorAdobeRRID:SCR_010279Used for figure preparation (Figure 2F, G, Figure 2—figure supplement 1H)
Antibodyanti-His (H-3), mouse monoclonal IgG1Santa Cruz BiotechnologyCat#:sc-8036, RRID:AB_627727WB; (1:2000); Lot#:C0421
Antibodyanti-GST (B-14), mouse monoclonal IgG1Santa Cruz BiotechnologyCat#:sc-138, RRID:AB_627677WB; (1:2000); Lot#:K1020
Antibodyanti-mouse IgG-HRPCell Signaling TechnologyCat#:7076S, RRID:AB_330924WB; (1:3000): Lot#:36
OtherConcanavalin A, type VISigma-AldrichCat#:C2010
OtherConfocal dish (35 mm)ibidi GmbHCat#:81218-200
OtherConfocal dish (35 mm)CellvisCat#:D35C4-20-1.5-N
OtherNi-NTA agarose resinG-BiosciencesCat#:786-940Affinity resin used for purification of 6His-bdSUMO-Gin4KA1
OtherGlutathione Sepharose 4B resinGE HealthcareCat#:GE17-0756-01Affinity resin used for purification/immobilization of GST-Hof1N-ter
OtherPoly-Prep chromatography columnsBio-Rad LaboratoriesCat#:731-1550Used for affinity purification
OtherPD Midi Trap G-10 columnsGE HealthcareCat#:GE28-9180-11Used for protein buffer exchange

Strain construction

Request a detailed protocol

All yeast strains in this study were constructed using the S. cerevisiae wild-type strains ESM356 and YPH499, derived from the S288C genetic background. The strain number and the genotype are listed in Supplementary file 1. C-terminal epitope tagging and endogenous gene deletions were carried out based on the PCR-based integration strategy (Janke et al., 2004; Knop et al., 1999). All strains were cultured at 23°C with shaking at 180 rpm. Synthetic complete (S.C.) media lacking tryptophan, uracil, leucine, and histidine were used to select auxotrophic recombinants. Antibiotic selections for endogenous gene manipulation were done on YPD agar plates supplemented with 200 µg/ml G418, 100 µg/ml hygromycin-B, and 100 µg/ml nourseothricin. To visualize septin dynamics, Cdc3-mCherry, YIP211-cdc3-mCherry (Fang et al., 2010; Caviston et al., 2003), and YIP204-cdc3-mCherry (Wloka et al., 2013) plasmids were cut with BglII (Catalog No. R0144S; New England Biolabs) and integrated into the CDC3 locus. Cell cycle dynamics were visualized using a yeast expression vector with mRuby2-Tub1 under the His3 promoter (pHIS3p:mRuby2-Tub1+3″UTR::Ura3) (Markus et al., 2015) and pAFS125-GFP-TUB1 (Straight et al., 1997), which were cut with ApaI (Catalog No. R0114S; New England Biolabs) and StuI (Catalog No. R0187S; New England Biolabs), respectively, and integrated into the ura3 loci. Yeast transformation for epitope tagging and endogenous gene deletions was performed using a lithium acetate (LiOAc)-based protocol as described (Sikorski and Hieter, 1989).

Oligonucleotides and plasmids

Request a detailed protocol

All oligonucleotides used for endogenous gene tagging and deletion in this study are listed in Supplementary file 2. The plasmids used and generated for this study are listed in Supplementary file 3. For the generation of clones used in Figure 3A, B, Gin4FL was amplified from the endogenous locus and integrated into pRS305 under the pTEF promoter. The ligation reaction was performed using NEB Hifi-Builder. Catalytically dead Gin4KD(K48A) was constructed based on the site-directed mutagenesis strategy using the oligos (P855, P856, P857, and P1024). The kinase-dead mutant was confirmed correct by sequencing. To construct Gin4FL and Gin4KD used in Figure 3C–E, Figure 2—figure supplement 1F, G, and 1I, the Gin4 gene fragment was amplified from the genomic loci with the endogenous promoter using PCR and was ligated into a pRS305 integration vector cut with BamHI-HF (Catalog No. R3136S; New England Biolabs).

Live-cell imaging

Request a detailed protocol

Cells were inoculated in S.C. media and incubated overnight at 23°C. The secondary culture was diluted to log phase (optical density – 0.4–0.6) from the overnight saturated culture. 6% Concanavalin A type VI was coated onto a confocal dish and was incubated at 23°C. The secondary culture cells were pelleted at 3200 rpm for 3 min. The cells were washed in S.C. media to remove non-adherent cells and subjected to live-cell imaging.

The laser point scanning confocal microscope Olympus FV-3000, equipped with high-sensitivity GaAsP photomultiplier tube detectors and solid-state lasers (488 and 561 nm), was used to acquire time-lapse imaging for Figure 1—figure supplement 1A–D and Figure 1—figure supplement 2A. Images were acquired at a 1-min time interval with 0.5 µm step-size and seven z-slices for 2–3 hr. The raw images were three-dimensionally (3D) deconvolved in Olympus CellSens Dimension (3.1) software. Maximum-intensity projection images were used for representation.

The Olympus IX83 widefield microscope equipped with a CoolLED PE-4000 LED light source and Prime-BSI sCMOS camera was used for acquiring images shown in Figures 1B and D—3C and Figure 2—figure supplement 1C. All images were acquired with a 100x oil-immersion objective with 1.45 N.A., 1 µm step-size, and three z-slices at a 2-min time interval for 60–150 min. The raw images were processed for 3D deconvolution using Olympus CellSens Dimension (3.1) software. Maximum-intensity projection images were used for representation.

The images for Hof1 misorganization and artificial tethering experiments in Figure 2D; Figure 2—figure supplement 1A, F; Figure 4B, I; Figure 7A; Figure 4—figure supplement 1E; Figure 7—figure supplement 1A; and Figure 8—figure supplement 1E, H were acquired using the Oxford Andor Dragonfly 502 spinning disc confocal system equipped with a fully motorized Dmi8 inverted setup. The raw images were captured with a 100x oil immersion objective with 0.5 µm step-size, 11 z-slices, in an Andor Sona sCMOS camera, and deconvolved using Andor Fusion software. The 3D-deconvolved and max-projected images were used for representation.

The Olympus IX-83 Laser scanning spinning disc Yokogawa-W1 microscope was used to capture Figures 3A, 4F and 8A, Figure 6—figure supplement 1A, and Figure 8—figure supplement 2A images. Raw images were acquired with a 100x oil-immersion objective with 1.45 N.A., using 1 µm step-size and five z-slices. The time-lapse imaging for Figure 1—figure supplement 1E; Figure 1—figure supplement 2D; Figure 2—figure supplement 2A, D; Figure 5A, D; Figure 4—figure supplement 1A, B; Figure 6A, C; Figure 7D; Figure 8B; Figure 7—figure supplement 2A–C; and Figure 8—figure supplement 1A, B was acquired at a 1-min time interval with 60x objective with 1.42 N.A. The images and time-lapse montages were 3D-deconvolved in Olympus CellSens Dimension (3.1) software. Maximum-intensity projection images were used for representation.

Protein accumulation kinetics quantification and analysis

Request a detailed protocol

The live-cell imaging data were opened in Fiji (ImageJ2) image analysis software (Schindelin et al., 2012) using the protocol established (Okada et al., 2021). In brief, a region of interest (ROI) was drawn around the bud neck of the sum-intensity projected image. The extracted fluorescent intensity values were corrected using background values. All the individual fluorescent intensity values obtained in an Excel sheet at each time point were normalized using the minimum and maximum values according to the formula below:

Normalized value=(Raw value−minimum value)/(maximum value−minimum value)

The data points ranging between the –20 and +20 time points were separately extracted for the ‘n’ number of cells indicated with each graph, respectively. The mean and standard deviation were calculated. A line plot was generated using Origin-Pro image analysis software to represent the temporal accumulation kinetics of the protein of interest. The small bud is considered as t0 for quantifications involving bud emergence. The complete drop in Cdc3-mCherry intensity is considered as the HDR transition in Figure 1—figure supplement 1E. The spindle breakpoint, where the spindle splits during cytokinesis, is considered as t0 or the reference time-point for measuring Hof1, Inn1, and Chs2 temporal dynamics during cytokinesis.

Quantitative analysis of septin and Hof1 misorganization was performed manually by counting the number of cells exhibiting septin mislocalization to the bud cortex during bud emergence and the defective/discontinuous Hof1 ring at the bud neck. The residence time of AMR components at the bud neck during cytokinesis was calculated from the time-lapse movies as the time interval between the first appearance of the signal and its complete disappearance from the bud neck.

Quantitative analysis of cellular morphology

Request a detailed protocol

Brightfield images were acquired using a 100x oil-immersion objective. Cells containing a large bud were selected for analysis. Cell boundaries were manually drawn by selecting a polygonal ROI in Fiji. The marked ROIs were analyzed to obtain measurements of cell area, major axis length, minor axis length, and aspect ratio. The aspect ratio, defined as the ratio of the cell’s major axis to the minor axis, was used as a quantitative parameter to distinguish between round and elliptical cell shapes.

Y2H assay

Request a detailed protocol

Genes or gene fragments of interest were amplified and cloned into NotI restriction enzyme (NEB; Catalog No. R3189S)-cut pMM5S and pMM6S plasmids consisting of the DNA binding domain of LexA (LexDBD) and activation domain of Gal4 (Gal4AD), respectively. The clones were transformed into the yeast strains SGY37 (MATa) and YPH500 (MATalpha). The transformants were selected on S.C. media lacking histidine and leucine, respectively. Mating was performed on YPD plates and was incubated at 30°C for 2 days, followed by replica plating on SC-His-Leu double selection plates. The plates were incubated for 2 days at 30°C, then overlaid with freshly prepared X-Gal mix (500 mM sodium phosphate buffer (pH 7.0), 10% SDS, 1 M KCl, 1 M MgCl2, 0.04% X-gal, and 0.4% low-melting agarose) for detection of β-galactosidase activity (Geissler et al., 1996; Palani et al., 2012). Interaction of LexADBD-Protein of interest A with Gal4AD-Protein of interest B resulted in expression of the lacZ gene encoding β-galactosidase, which converted X-gal to a product with blue color (Gyuris et al., 1993). The plates were scanned and documented after 14–16 hr of incubation with overlay mix.

Protein expression, purification, and in vitro binding assay

Request a detailed protocol

The N-terminal fragment of Hof1 containing the F-BAR domain (1–350 aa) and KA1 fragment of Gin4 (1003–1142 aa) were PCR-amplified from S. cerevisiae genomic DNA and cloned into pGEX-5X1 cut with EcoRI restriction enzyme (Catalog No. R3101S) and pET28a-6His-bdSUMO vector cut with NdeI restriction enzyme (Catalog No. R0111S), respectively, using NEB Hifi Builder. Both plasmids were transformed into BL21-DE3 E. coli competent cells and were selected on Luria Bertani (LB) plates containing Kanamycin (50 mg/ml) for pGEX-5X1-Hof1N (1-350) and Ampicillin (100 mg/ml) for pET28a-6His-bdSUMO-Gin4KA1. LB broth with Kanamycin or Ampicillin was used to grow primary cultures overnight at 37°C with constant shaking at 200 rpm. The primary culture was diluted to 0.1 O.D. in 500 ml LB broth with Kanamycin or Ampicillin. Protein expression was induced with 0.5 mM isopropyl β-D-1-thiogalactopyranoside once the O.D. reached log phase (0.4–0.5). The culture was incubated at 30°C with shaking at 200 rpm for 3–4 hr. The cells were pelleted at 4°C, 7000 rpm, for 20 min. Media were aspirated and the pellets were washed with cold phosphate-buffered saline (PBS) containing 1 mM phenylmethylsulfonyl fluoride and were stored at –80°C.

Purification of 6His-bdSUMO-Gin4KA1

Request a detailed protocol

Pellets were kept on ice for 10 min. and were subjected to lysis with the lysis buffer (50 mM Tris pH 8, 200 mM NaCl, 10% glycerol). The lysed pellets were clarified at 4°C and 15,000 × g. The clarified lysate was mixed with Ni-NTA agarose resin, washed with wash buffer (50 mM Tris pH 8, 200 mM NaCl, 10 mM imidazole), and was incubated at 4°C for 2 hr. The lysate with Ni-NTA resins was then transferred to poly-prep chromatography columns, and was subjected to gradient elution with increasing concentrations of imidazole. The eluted protein fractions were buffer-exchanged with storage buffer (50 mM Tris-HCl pH 7.4, 2000 mM NaCl, 1 mM DTT, and 10% glycerol) using PD Midi Trap G-10 Sephadex columns. The purified proteins were flash-frozen in liquid nitrogen and were stored at –80°C. The protein concentration was estimated using a spectrophotometer at 280 nm and was analyzed, along with known concentrations of bovine sesum albumin standards, on sodium dodecyl-sulfate–polyacrylamide gel electrophoresis.

Purification of GST-Hof1N (1–350)

Request a detailed protocol

Pellets were kept on ice for 10 min. and were subjected to lysis with the lysis buffer (PBS, 0.5 mM EDTA, 1 mM DTT, 1 mg/ml lysozyme, and complete mini-EDTA-free protease inhibitor cocktail tablets). The clarified lysate was incubated with 1% Triton X-100 for 20 min. on ice and centrifuged at 4°C, 22,000 × g, for 30 min. The lysate was then transferred to a tube containing glutathione sepharose-4B resin, washed with wash buffer (5x) (PBS, 0.5 mM EDTA, and 1 mM DTT), and was incubated at 4°C for 2–3 hr. The lysate with glutathione Sepharose resins was then transferred to poly-prep chromatography columns after incubation, washed with wash buffer, and was stored at 4°C.

In vitro binding assay

Request a detailed protocol

GST or GST-Hof1N (1-350) (~3 µg) bound to glutathione beads were mixed with ~1 µg 6His-bdSUMO-Gin4KA1 and incubated in binding buffer (PBS, 0.2% NP-40) for 1 hr at 4°C. The beads were washed three to five times with wash buffer (PBS, 1% NP-40) and were analyzed by western blot using Anti-His (H-3) mouse monoclonal IgG1, Anti-GST (B-14) mouse monoclonal IgG1, and Anti-mouse IgG-HRP.

Spot assay

Request a detailed protocol

The yeast strains were grown in liquid YPD broth overnight at 23°C. The cultures were diluted to 1.0 O.D. and were serially diluted in the order 100, 10–1, 10–2, 10–3, and 10–4. These dilutions were spotted on three YPD plates and incubated at 23, 30, and 37°C. Growth was monitored, and plates were scanned after 48 hr of incubation.

The strains used for rescue of gin4Δ hof1Δ synthetic lethality (Figure 2—figure supplement 1I) were grown overnight in S.C. media lacking Uracil and Leucine. The culture was diluted to an O.D. of 1.0 and was serially diluted in the order 100, 10–1, 10–2, 10–3, and 10–4. These dilutions were spotted on three SC-Ura-Leu plates and were incubated at 23°C for 2 days. Subsequently, the colonies were patched onto 5-FOA plates (1 mg/ml) to select for Ura- cells.

Statistical analysis

Request a detailed protocol

All experiments were performed independently three times. Statistical significance for bar graphs and stacked column plots was determined using one-way ANOVA with Tukey’s multiple-comparison test, whereas residence-time plots were analyzed using the Kruskal–Wallis nonparametric test. All statistical analyses were performed using in-built functions of GraphPad Prism (v.6.04). A p-value of p < 0.05 was considered statistically significant (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, ns: p > 0.05).

Lead contact and material availability

Request a detailed protocol

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Dr. Saravanan Palani (spalani@iisc.ac.in). Plasmids and yeast strains generated in this study are available from the lead contact upon request.

Data availability

All data generated or analysed during this study are included in the manuscript and its supporting files. Source data files containing the numerical data underlying the figures and figure supplements, as well as uncropped images underlying the protein interaction experiments, have been provided.

References

Article and author information

Author details

  1. Bindu Bhojappa

    Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, Bangalore, India
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8864-2651

  2. Anubhav Dhar

    Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, Bangalore, India
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Writing – original draft, Writing – review and editing
    Competing interests
    No competing interests declared

  3. VT Bagyashree

    Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, Bangalore, India
    Contribution
    Resources, Data curation, Formal analysis, Validation, Investigation, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared

  4. Jayanti Kumari

    Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, Bangalore, India
    Contribution
    Resources, Data curation, Formal analysis, Validation, Investigation, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared

  5. Freya Cardozo

    Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, Bangalore, India
    Contribution
    Formal analysis, Validation, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared

  6. Vaseef Rizvi

    Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, Bangalore, India
    Contribution
    Formal analysis, Validation, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared

  7. Deepthi Guturu

    Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, Bangalore, India
    Contribution
    Formal analysis, Investigation, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared

  8. Saravanan Palani

    Department of Biochemistry, Division of Biological Sciences, Indian Institute of Science, Bangalore, India
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing
    For correspondence
    spalani@iisc.ac.in
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1893-6777

Funding

Wellcome Trust/DBT India Alliance (IA/I/21/1/505633)

  • Saravanan Palani

Anusandhan National Research Foundation (SRG/2021/001600)

  • Saravanan Palani

Anusandhan National Research Foundation (ANRF/ARG/2025/004009/LS)

  • Saravanan Palani

The funders had no role in study design, data collection, and interpretation, or the decision to submit the work for publication. For the purpose of Open Access, the authors have applied a CC BY public copyright license to any Author Accepted Manuscript version arising from this submission.

Acknowledgements

We thank the DST-FIST microscope facility of the Department of Biochemistry, Indian Institute of Science. We also thank the Bioimaging facility, Division of Biological Sciences. We thank Prof. Erfei Bi (University of Pennsylvania) and Prof. Gislene Pereira (Centre for Organismal Studies (COS) Heidelberg) for the plasmids. We are grateful for the help of Ms. Spoorthi Hirematt in image acquisition at the DST-FIST microscope facility. We thank Prof. PN Rangarajan, Prof. Franz Meitinger, and Prof. Ramanujam Srinivasan for their valuable suggestions on the manuscript. Schematic diagrams were created in BioRender. BB acknowledges the DST-INSPIRE Fellowship. AD and JK acknowledge the IISc PhD fellowship from IISc. FC acknowledges the ICMR fellowship.

Version history

  1. Preprint posted:
  2. Sent for peer review:
  3. Reviewed Preprint version 1:
  4. Reviewed Preprint version 2:
  5. Version of Record published:

Cite all versions

You can cite all versions using the DOI https://doi.org/10.7554/eLife.106366. This DOI represents all versions, and will always resolve to the latest one.

Copyright

© 2025, Bhojappa et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

Metrics

  • 2,283
    views
  • 61
    downloads
  • 0
    citations

Views, downloads and citations are aggregated across all versions of this paper published by eLife.

Download links

A two-part list of links to download the article, or parts of the article, in various formats.

Downloads (link to download the article as PDF)

Open citations (links to open the citations from this article in various online reference manager services)

Cite this article (links to download the citations from this article in formats compatible with various reference manager tools)

  1. Bindu Bhojappa
  2. Anubhav Dhar
  3. VT Bagyashree
  4. Jayanti Kumari
  5. Freya Cardozo
  6. Vaseef Rizvi
  7. Deepthi Guturu
  8. Saravanan Palani
(2026)
Nim1-related kinases regulate septin organization and cytokinesis by modulating Hof1 at the cell division site
eLife 14:RP106366.
https://doi.org/10.7554/eLife.106366.3

Share this article

https://doi.org/10.7554/eLife.106366

Categories and tags

Research organism