The stomatin-like protein StlP organizes membrane microdomains to govern polar growth in filamentous actinobacteria under hyperosmotic stress

Zhong, Xiaobo; Baur, Sarah S. M.; Ongenae, Veronique M. A.; Guerrero Egido, Guillermo; Shitut, Shraddha; Du, Chao; Vijgenboom, Erik; van Wezel, Gilles P.; Carrion Bravo, Victor; Briegel, Ariane; Bramkamp, Marc; Claessen, Dennis

doi:10.1038/s41467-025-58093-x

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Published: 18 March 2025

The stomatin-like protein StlP organizes membrane microdomains to govern polar growth in filamentous actinobacteria under hyperosmotic stress

Nature Communications volume 16, Article number: 2669 (2025) Cite this article

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Abstract

The cell wall represents an essential structure conserved among most bacteria, playing a crucial role in growth and development. While extensively studied model bacteria have provided insights into cell wall synthesis coordination, the mechanism governing polar growth in actinobacteria remains enigmatic. Here we identify the stomatin-like protein StlP as a pivotal factor for orchestrating polar growth in filamentous actinobacteria under hyperosmotic stress. StlP facilitates the establishment of a membrane microdomain with increased membrane fluidity, a process crucial for maintaining proper growth. The absence of StlP leads to branching of filaments, aberrant cell wall synthesis, thinning of the cell wall, and the extrusion of cell wall-deficient cells at hyphal tips. StlP interacts with key components of the apical glycan synthesis machinery, providing protection to filaments during apical growth. Introduction of StlP in actinobacteria lacking this protein enhances polar growth and resilience under hyperosmotic stress, accompanied by the formation of a membrane microdomain. Our findings imply that stomatin-like proteins, exemplified by StlP, confer a competitive advantage to actinobacteria encountering hyperosmotic stress. Given the widespread conservation of StlP in filamentous actinobacteria, our results propose that the mediation of polar growth through membrane microdomain formation is a conserved phenomenon in these bacteria.

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Introduction

The cell wall is considered an essential structure in bacteria that protects cells from environmental stresses^1,2. To enable bacterial growth, the cell wall needs to be expanded, which involves inserting new cell wall material at the sites of growth. Elongation of rod-shaped cells typically occurs in two distinct manners³. Some rod-shaped bacteria, such as Escherichia coli and Bacillus subtilis elongate by incorporating new cell wall material in a rather diffuse manner in the cylindrical part of the cell⁴. By contrast, other bacteria grow by inserting new cell wall material at the cell poles, referred to as polar growth, which is widespread in actinobacteria^5,6. This mode-of-growth has been well studied in actinomycetes, which are filamentous bacteria that form branched mycelial networks in soil environments. In their natural environment, actinomycetes are often confronted with suboptimal conditions, such as fluctuations in water availability causing dramatic osmotic imbalances⁷. Paradoxically, we recently showed how conditions of hyperosmotic stress causes shedding of the cell wall in several actinomycetes, implying that such conditions interfere with the process of cell wall growth⁸.

Polar growth in actinomycetes is guided by the cytoskeletal protein DivIVA, which localizes in actively growing tips acting as a scaffold for other proteins involved in organizing tip growth, such as the coiled-coil protein Scy and the intermediate filament-like protein FilP^9,10,11. Unlike Scy and FilP, DivIVA plays an essential function in polar growth, by directly interacting with the machinery involved in synthesis of peptidoglycan, a major constituent of the cell wall^12,13. Both partial depletion and overproduction of DivIVA significantly altered cell shape and caused irregular branching⁹. Scy is an unusual long coiled-coil protein that co-localizes with DivIVA at hyphal tips. Scy was suggested to form higher order assemblies and thereby serves as a hub to stabilize the tip-organizing center, which also includes cell division proteins and proteins involved in chromosome segregation^11,14,15. Collectively, these proteins facilitate that cell wall synthesis is coordinated with chromosome segregation to ensure proper growth and development of the mycelium.

DivIVA also interacts with the putative cellulose synthase CslA¹⁶. The glycan produced by CslA is thought to provide protection to the tips, which are constantly remodeled during growth, in particular under conditions of osmotic stress¹⁶. Synthesis of the cellulose-like glycan not only depends on the synthase CslA, but also on a range of other proteins that are all encoded in a conserved gene cluster (Supplementary Fig. 1A). Together with CslA, the radical copper oxidase GlxA are the key proteins responsible for synthesis and likely modification of the glycan^17,18. Following synthesis of the glycan, the lytic polysaccharides monooxygenase LpmP and endoglucanase CslZ facilitate deposition of the glycan chain at the cell surface, possibly by creating a passage through the thick peptidoglycan (PG) layer¹⁹. The cooperation of CslA/GlxA and CslZ/LpmP implies that a multicomplex is established at the tip related to the proper synthesis and secretion of the cellulose-like glycan.

One of the proteins in the cellulose biosynthetic gene cluster that has not been studied encodes a stomatin/prohibitin/flotillin/HflK/C (SPFH)-domain protein, hereinafter referred to as StlP (for stomatin-like protein). The SPFH-superfamily proteins facilitate formation of lipid rafts, which in eukaryotes often contain protein complexes that collectively carry out important biological processes, such as ion channel regulation and touch sensation^20,21. Notably, prokaryotic SPFH proteins, such as flotillins, also form comparable structures known as functional membrane microdomains (FMMs)^22,23. These FMMs were shown to be involved in cell wall biosynthesis in B. subtilis²⁴ and Staphylococcus aureus²⁵. Additionally, it was revealed that, in B. subtilis, flotillins play a direct role in controlling membrane fluidity homeostasis²⁶ with their expression intriguingly regulated by stress-specific signals²⁷.

In this study, we reveal the pivotal role of StlP in coordinating the creation of a microdomain crucial for sustaining tip growth in Streptomyces coelicolor during hyperosmotic stress conditions. The absence of StlP leads to anomalous hyphal shape alterations and, notably, the expulsion of cells lacking their cell wall. StlP undergoes polymerization into oligomers and localizes within the membrane, culminating in the formation of a membrane region characterized by heightened fluidity. This phenomenon is essential for harmonizing the growth of both membrane and cell wall during tip extension. Significantly, the ectopic expression of StlP in actinomycetes, known for naturally extruding wall-deficient cells, effectively prevents such extrusion. These findings collectively underscore the significance of StlP in establishing a membrane microdomain at hyphal tips, thereby playing a critical role in facilitating proper cell wall assembly under hyperosmotic stress conditions in filamentous actinobacteria.

Results

StlP is a stomatin-like protein in Streptomyces coelicolor

StlP is encoded in the conserved cellulose biosynthesis gene cluster in streptomycetes and located downstream of lpmP¹⁹ (Supplementary Fig. 1A). Analysis of protein domains with InterPro and structure prediction via AlphaFold reveal that StlP contains three domains: a disordered region (aa 1-100), followed by a transmembrane hairpin composed by two helices (aa 106-126 and 149-168), and a SPFH (stomatin, prohibitin, flotillin, HflK/C) domain (aa 209-311) (Supplementary Fig. 1B, C).

Considering the diverse membrane topologies observed in members of the SPFH superfamily²⁸, we examined the membrane topology of StlP to determine its classification within the family of SPFH-containing proteins. In silico analyses predicted that the C-terminal SPFH domain of StlP is in the cytoplasm (Supplementary Fig. 1D). To validate this prediction, we employed a β-lactamase assay in which this enzyme was fused to the C-terminus of StlP as described²⁹. E. coli cells expressing functional StlP-BlaM_NS fusions demonstrate sensitivity to ampicillin, in contrast to cells expressing BlaM_FL with its original signal peptide (Supplementary Fig. 1E–G). This result confirms that the C-terminal SPFH domain of StlP is situated in the cytoplasm, consistent with the observed membrane topology in podocin/stomatin family proteins²⁰ and its N-terminal transmembrane hairpin. In contrast to StlP, other SPFH-like proteins from S. coelicolor have a significantly different predicted membrane topology (Supplementary Fig. 2). A phylogenetic analysis, utilizing amino acid sequences from well-known prokaryotic and eukaryotic SPFH proteins, revealed that S. coelicolor StlP and Mouse stomatin STOML-1 form a monophyletic clade, indicating that StlP resembles a stomatin-like protein (Supplementary Fig. 3A). Further sequence alignment of StlP with other stomatins indicates that Streptomyces StlP contains the signature proline³⁰ residue required for membrane hairpin formation and a comparable domain arrangement (Supplementary Fig. 3B). This alignment suggests that StlP shares more similarities with eukaryotic stomatins than prokaryotic stomatins. Indeed, structural superposition shows that the stomatin domain (StlP_SD) of StlP aligns better with Mouse stomatin (MmStomatin) than the bacterial stomatin FliL (Supplementary Fig. 3C, D). In summary, these findings collectively establish StlP as a novel stomatin-like protein, containing a classic N-terminal transmembrane hairpin and stomatin domain, within the prokaryotic family of SPFH proteins.

StlP is important for morphogenesis under hyperosmotic stress conditions

To investigate the function of StlP, a stlP mutant (∆stlP) was created that contained a Tn5062 transposon insertion, positioned upstream of the stomatin domain. This constructed stlP mutant was used to study its phenotype in various conditions. The stlP mutant displayed significantly reduced growth rates in both TSBS and LPB liquid medium, which contain 10 and 22% sucrose, respectively (Fig. 1A and Supplementary Fig. 4A). On MS agar medium (without sucrose), no apparent differences in growth were observed between the parent and the stlP mutant (Supplementary Fig. 4B). By contrast, when grown on LPMA agar medium (containing 22% sucrose), the average diameter of individual colonies of the stlP mutant (1.4 ± 0.3 mm) was significantly reduced compared to the wild-type strain (3.8 ± 0.6 mm) (Fig. 1B, C), suggesting that growth was severely hampered. Furthermore, we noticed that excess membrane was extruded from hyphal tips of the stlP mutant, as evident in the FM5-95 panels comparing M145 and ∆stlP in Fig. 1C. Additionally, we observed many DNA-containing vesicles present in the medium (Fig. 1D) that were reminiscent of cell wall-deficient cells extruded by several filamentous actinobacteria. These vesicles were absent in the parental strain, consistent with earlier findings⁸. Time-lapse imaging revealed that the vesicles were extruded from hyphal tips (Fig. 1E and Supplementary Movies 2). Quantification revealed that the mutant formed 2.5 × 10⁵ vesicles ml⁻¹, while none were found in the parental strain (Supplementary Fig. 5). Importantly, the significant reduced colony and formation of vesicles were also observed in the full-length stlP mutant (∆stlP_FL) generated via homologous recombination (Supplementary Fig. 6). To confirm that these phenotypes were caused by the absence of StlP, we introduced plasmid pXZ15 in the mutant, in which stlP is expressed from the constitutive gapAp promoter. Reintroduction of this plasmid partially restored the growth speed in liquid media (Fig. 1A and Supplementary Fig. 4A) and the average colony diameter on LPMA medium (2.5 ± 0.5 mm) (Fig. 1B, C), while extrusion of membranes and DNA-containing vesicles was also reduced by 80% (Fig. 1D and Supplementary Fig. 5).

Fig. 1: The stomatin-like protein StlP is important for morphogenesis in *S. coelicolor.*

Microscopy analysis also indicated that the hyphal branching pattern was affected by the deletion of stlP (Fig. 1F, Supplementary Fig. 4D and Supplementary Movies 1 and 2). More specifically, the number of branches was dramatically increased in the stlP mutant when grown under hyperosmotic stress conditions (LPMA medium) (Fig. 1F). A similar effect was observed when the sucrose in LPMA (0.64 M) was replaced with NaCl (0.32 M, Supplementary Fig. 7). To quantitatively compare branching between the strains, we measured the distance from the tip to the proximal branching point in hyphae (Supplementary Figs. 4D and 7). Deletion of stlP changed the distribution of the tip-to-branch distances significantly. We found that more than 60% of all hyphae in ∆stlP had a proximal branch within the first 5 μm from the tip (Fig. 1G), as compared to approximately 15% for the parent and the complemented mutant. Also, no hyphae of the stlP mutant branched further than 35 μm from the tip. Additionally, when grown on MS medium, hyphae with a tip-to-branch distance less than 10 μm accounted for 3.6% and 8.9% of the parent and mutant strain, respectively (Supplementary Fig. 4E). This suggests a mild impact on the branching pattern due to the deletion of stlP under normal growth conditions. Taken together, these findings underscore that the absence of StlP significantly affects polar growth and causing an increase in the emergence of lateral branches in S. coelicolor, particularly manifesting under conditions of hyperosmotic stress.

StlP is important for spatially confining cell wall synthesis to hyphal tips

To investigate the localization of StlP, we introduced pXZ16 into S. coelicolor M145, thereby constitutively expressing a C-terminal mCherry fusion to StlP. Foci of StlP-mCherry mostly localized at growing tips and emerging branches (Fig. 2A). We also localized the polar growth determinant DivIVA in the presence and absence of StlP by expressing a DivIVA-mCherry fusion from the constitutive gapAp promoter. Interestingly, in the absence of StlP, DivIVA-mCherry was not only localized to hyphal tips but was also found in numerous foci along the cylindrical part of the filaments (Fig. 2B), suggesting diffusion of cell wall synthesis from the apex. In agreement, nascent PG was incorporated at multiple sites along the filament in the stlP mutant (Fig. 2C), coinciding with an increase in the average diameter of the hyphae (Fig. 2D). Using calcofluor white staining, we observed that multiple synthesis foci of the cellulose-like glycan produced by CslA appeared at both established and emerging hyphal tips in the stlP mutant, in contrast to the parental strain (Fig. 2E). Furthermore, quantitative analysis revealed a significant decrease (approximately 65%) in the accumulated glycan at the hyphal tips of the stlP mutant. Consistent with other mutants affected in apical glycan synthesis¹⁹, we indeed noticed that the reduced glycan levels made the stlP mutant sensitive to lysozyme (Fig. 2G). Glycan deposition and lysozyme resistance were restored when the complementation plasmid pXZ15 was introduced in the stlP mutant (Fig. 2E–G).

**Fig. 2: StlP localizes to hyphal tips and affects cell wall synthesis.**

To accurately evaluate the effect of the absence StlP on cell wall thickness of S. coelicolor, we conducted a thorough examination by preparing and imaging sacculi of M145 and the ∆stlP strains using cryo-electron tomography (cryo-ET). The analysis of cell wall thickness measurement revealed a significant reduction in the overall cell wall thickness upon stlP deletion compared to the parental strain. Specifically, the thickness in the apical region decreased from 35.5 ± 1.7 nm to 6.4 ± 0.6 nm, while the thickness in the subapical region was reduced from 37.1 ± 2.3 nm to 9.9 ± 0.6 nm (Fig. 2H, I and Supplementary Movies 3 and 4). These results demonstrate the importance of StlP for cell wall synthesis and thickness of S. coelicolor.

To establish how StlP contributes to delocalized cell wall synthesis, we tested interactions of StlP with the proteins involved in tip growth and cellulose biosynthesis. To this end, constructs were generated that produced C-terminal fusions of StlP, LpmP, SCO2835, CslA, GlxA, CslZ, DivIVA, Scy and FilP to either the T25 or T18 fragments of the adenylate cyclase, respectively. Co-transformation of these constructs in E. coli BTH101 revealed that StlP did not interact with DivIVA, Scy or FilP. However, StlP robustly interacts with LpmP, SCO2835, CslA and CslZ, but also with itself and weakly interacts with GlxA (Supplementary Fig. 8A, B). Pull-down experiments using E. coli BL21 cells expressing a C-terminal 8×His-tagged StlP (StlP-8His), a C-terminal Flag-tagged CslA (CslA-Flag), and GlxA confirmed that StlP-8His effectively pulled down CslA-Flag and GlxA, as demonstrated by size exclusion chromatography and immunoblotting analyses (Supplementary Fig. 8D, E). Additionally, Native-PAGE analysis revealed the presence of the StlP-8His/CslA-Flag/GlxA complex with an approximate molecular weight of 480 kDa (Supplementary Fig. 8F). To validate these findings, we investigated the interactions between StlP, CslA, and GlxA in Streptomyces, and performed bimolecular fluorescence complementation (BiFC) experiments. To this end, eGFP was split into two fragments, eGFP_N and eGFP_C, which were fused to either the N- or C-terminus of StlP, CslA, or GlxA. For GlxA, the eGFP_C fragment was specifically inserted between its signal peptide and its N-terminal transmembrane anchor, ensuring that the eGFP signal remained localized to the inner membrane following GlxA secretion. Constructs were generated using apramycin-resistant pSET152 and thiostrepton-resistant pWHM3-oriT backbones, with expression driven by the constitutive gapAp promoter. Various constructs pairs were conjugated into the wild-type strain M145, and protein interactions were assessed based on detectable eGFP fluorescence at hyphal tips, resulting from the reconstitution of eGFP fragments (eGFP_N and eGFP_C) through complementation. Controls included strains expressing individual protein fusions. Results showed that eGFP_N fused to the C-terminus of StlP complemented eGFP_C fragments fused to either the N- or C-terminus of CslA, indicating robust StlP-CslA interactions (Fig. 3A and Supplementary Fig. 9). No fluorescence was observed between StlP and GlxA constructs (Fig. 3B and Supplementary Fig. 9). eGFP_C fused to the N-terminus of GlxA complemented eGFP_N fused to the C-terminus of CslA, demonstrating a strong interaction between CslA and GlxA (Fig. 3 and Supplementary Fig. 9). These findings establish that StlP directly interacts with components of the cellulose biosynthesis complex and spatially confines cell wall synthesis to hyphal tips in S. coelicolor.

Fig. 3: BiFC analysis of StlP, CslA, and GlxA interactions in *Streptomyces.*

StlP oligomerizes and forms membrane microdomain at hyphal tips of S. coelicolor

Bacterial two-hybrid analysis suggested that StlP interacts with itself but also with components of the machinery involved in glycan synthesis (Supplementary Fig. 8A). To test if StlP interacts with itself via the stomatin domains, the corresponding domain (aa 204-326; referred to as StlP_SD) was expressed in E. coli BL21(DE3). The StlP_SD monomer was expressed, showing a predicted molecular mass of 19.6 kDa (Fig. 4A). When 20 µg of concentrated StlP_SD protein (1 mg ml⁻¹) were loaded in a SDS-PAGE gel, a ladder of oligomers was found, suggesting that monomers readily self-assemble and are resistant to SDS (Fig. 4A). Notably, analysis of 20 µg of concentrated StlP_SD protein using Native-PAGE revealed a predominance of heptamers, along with smaller amounts of decamers and trimers (Fig. 4A). To obtain more insight into self-assembly of StlP, we predicted the partial structure of StlP (aa 106-326), encompassing the two N-terminal TMHs (aa 106-126 and aa 149-168) and the stomatin domain (aa 209-311) using AlphaFold 2.0. When six StlP monomers were used as input, an ordered hexamer and partial ring-like structure was obtained, while ten monomers assembled into a symmetric ring (Fig. 4B and Supplementary Fig. 10). These results suggest that StlP can assemble into oligomers via its stomatin domain at high concentrations, which in the full-length protein would probably result in formation of a membrane microdomain via its two N-terminal transmembrane helices.

**Fig. 4: StlP oligomerizes and controls membrane fluidity at hyphal tips.**

In bacteria, SPFH proteins are involved in membrane fluidity homeostasis, evidenced by FloT and FloA that fluidize the membrane and absence of them leads to an overall rigidification^24,26. We therefore predicted that the oligomerization of StlP leads to the formation of a local membrane region with increased fluidity (RIF) at hyphal tips. To test this, we stained mycelia of S. coelicolor with DilC₁₂, a lipid dye with high specificity for fluid membranes due to its short hydrocarbon tail^31,32. We observed bright DilC₁₂ staining RIFs at hyphal tips, and also along hyphae grown for 16 h in LPB medium (Fig. 4C). The signal corresponding to DilC₁₂ staining was absent from hyphal tips of the stlP mutant and restored in the complemented strain (Fig. 4C). These results suggest that hyphal tips possess a RIF, which is dependent on the presence of StlP. To substantiate the presence of RIFs at hyphal tips, we conducted a quantitative assessment of membrane fluidity using the membrane-intercalating dye Laurdan. This dye exhibits a shift in fluorescence emission wavelength based on membrane fluidity^31,33. After culturing mycelia of M145 and its stlP mutant in LPB medium for 16 h, we stained them with Laurdan and then calculated the general polarization (GP) value at the tip region of each hypha (see “Materials and methods”). In the absence of stlP, the average GP value (−0.09 ± 0.08) increased compared to its parental strain (−0.12 ± 0.13), inferring a decreased membrane fluidity in the stlP mutant (Fig. 4D, E). Furthermore, the apical membrane fluidity of the stlP mutant could be restored by raising the growth temperature to 37 °C (Fig. 4F), which aligns with the idea that elevated temperatures contribute to membrane phase transitions, leading to an increased fluidity³⁴. Altogether, these results reveal that StlP oligomerizes and forms membrane domains with enhanced membrane fluidity at hyphal tips of S. coelicolor.

Morphogenesis controlled by StlP is conserved in filamentous actinobacteria

Our results indicate that StlP contributes to proper cell wall synthesis and membrane organization under hyperosmotic stress, and thereby controls morphogenesis of S. coelicolor. To see how prevalent StlP is, we combined PSI-BLAST and TMHMM prediction to identify stomatin homologs of StlP in the dataset of 15045 RefSeq representative bacteria and archaea. This analysis showed that the majority of StlP homologs are present in actinobacteria, while a few are also present in Rhizobium (Fig. 5). Furthermore, inside the filamentous actinobacteria, StlP orthologs are present in genera including Streptomyces, Kitasatospora and Streptacidiphillus (Fig. 5). In some clades of these bacteria, all members have an orthologue of StlP, while in others StlP is less common or virtually absent. Notably, Kitasatospora viridifaciens DSM40239 is among the clades that lack StlP (Fig. 5).

**Fig. 5: Phylogenetic analysis of the distribution of StlP proteins in Proteobacteria.**

To see how widespread the function of StlP is in other actinobacteria, the construct pXZ15, wherein the stlP was expressed from the constitutive gapAp promoter, was introduced into Kitasatospora viridifaciens DSM40239 via conjugation, which is known to extrude wall-deficient cells under hyperosmotic stress^8,35. Importantly, constitutive expression of stlP induced the formation of a fluid membrane microdomain at hyphal tips in K. viridifaciens, as evidenced by DilC₁₂ staining (Fig. 6A). Furthermore, constitutive expression of stlP significantly increased the average membrane fluidity of hyphae (Fig. 6B, C) and allowed colonies to cope much better with hyperosmotic stress, as shown by the strongly increased colony diameter of the strain expressing stlP (2.75 ± 0.6 mm) as compared to the parental strain (1.25 ± 0.9 mm) (Fig. 6D, E) and by the reduced lateral branching of K. viridifaciens (Fig. 6F, G). Furthermore, constitutive expression of stlP largely abolished the extrusion of wall-deficient cells in K. viridifaciens (Fig. 6H, I). This response to induced StlP expression was specific, as the constitutive expression of the endogenous HflK/C protein, encoded by BOQ63_030050 and sharing 21% sequence identity with StlP, was unable to prevent the extrusion of cell wall-deficient cells in K. viridifaciens (Supplementary Figs. 11 and 12). Taken together, these results demonstrate that the stomatin-like protein StlP controls growth of filamentous actinobacteria under hyperosmotic stress by the localized control of membrane fluidity, and this phenomenon is well-conserved in filamentous actinobacteria.

**Fig. 6: Enhanced robustness of *K. viridifaciens* growth under hyperosmotic stress conditions through constitutive StlP expression.**

Discussion

Stomatin-like proteins are ubiquitous in all domains of life. A universal feature of these proteins is to form functional nanoscale microdomains in biological membranes. In turn, these microdomains serve as platforms to locate protein complexes involved in important biological processes, such as transducing mechanosensory signals in mice and modulation of ion channels in mammalian cells^36,37,38. The recent structural analysis of the stomatin-like protein FliL from Vibrio alginolyticus showed that FliL shares some structural elements with eukaryotic stomatins, suggesting that stomatin-like proteins are conserved from mammals to bacteria³⁸. Here, we identified the stomatin-like protein StlP as a novel polar growth determinant in filamentous actinobacteria.

Establishing cell polarity and ensuring robust polar growth are important for actinobacteria to cope with environmental fluctuations. Our work shows that the presence of a microdomain, coordinated by the stomatin-like protein StlP, contributes to survival in conditions of hyperosmotic stress. The presence of this domain ensures that the synthesis of cell wall components remains localized at hyphal tips, which is pivotal for radial expansion of the colony. Indeed, the deletion of stlP leads to reduced expansion of the colony, coinciding with the formation of numerous small branches. We believe that this altered branching pattern has no direct benefit: escape from this stress, by radial growth, is hampered. However, under these conditions we see that hyphae also extrude wall-deficient cells, presumably due to the imbalance between membrane and cell wall synthesis. The formation of such cells may offer an alternative dispersal advantage for survival under hyperosmotic stress for other filamentous actinobacteria that lack StlP.

Stomatins oligomerize in cell membranes through interactions between their stomatin-domains, thus providing a scaffold for numerous other proteins. This functionality has been demonstrated for several stomatins, including mouse stomatin³⁷, Pyrococcus horikoshii stomatin³⁹ and the stomatin FliL of Vibrio alginolyticus³⁸. Cross-linking studies and structure predictions demonstrate that the stomatin domain of StlP oligomerizes and that StlP interacts with the machinery involved in synthesis of a cellulose-like glycan. One of these proteins is the cellulose synthase-like protein CslA, which in turn directly interacts with DivIVA¹⁶. Interestingly, PG synthesis of actinomycetes is directed at hyphal tips in a DivIVA-dependent manner. More specifically, DivIVA recruits penicillin-binding proteins (PBPs) during polarized growth, as deduced from the detected interaction between DivIVA and PBP3 in M. tuberculosis^11,12. StlP seems to contribute to establishing a localized region in the cellular membrane carrying crucial components of the so-called tip organizing center, which controls cell wall synthesis during the polar growth in filamentous actinobacteria (Fig. 7). Indeed, the observation of diffused PG and surface cellulose synthesis foci and dramatically reduced cell wall thickness of the stlP mutant indicated that PG synthesis is significantly weakened upon the deletion of stlP (Fig. 2). We thus propose a model for the cell wall synthetic machinery at hyphal tips when filamentous actinobacteria grow in hyperosmotic stress conditions. In this model, DivIVA guides the machinery responsible for peptidoglycan (PG) and surface cellulose-like glycan synthesis to the poles through interactions with CslA and PBPs. At these poles, the systems are restricted by the formation of StlP rings, which, in turn, induce localized membrane fluidization (Fig. 7). Here, the StlP ring enables these filamentous actinobacteria to produce robust cell walls during polarized growth under conditions of hyperosmotic stress. Without StlP, the membrane fluidity in the tip region decreases. Combined with the loss of localized cell wall synthesis, this leads to the extrusion of wall-deficient cells. Reversely, constitutive expression of StlP in species that naturally lack this protein prevents the typical extrusion of wall-deficient cells. Taken together, this work provides a plausible mechanism how wall-deficient cells are extruded in polar-growing actinobacteria. By interrupting cell wall synthesis, by exposing the cells to hyperosmotic stress or antibiotics, the coordinated balance between membrane and cell wall synthesis is lost. In turn, this leads to shedding of excess membranes from the polar growth sites and is facilitated by the localized weakening of the cell wall.

**Fig. 7: Proposed model for tip growth under conditions of hyperosmotic stress.**

In summary, our work for the first time characterizes a stomatin-like protein that regulates tip growth in conditions of hyperosmotic stress. The strong phenotype associated with its absence make this also an interesting candidate to target in pathogenic bacteria that grow from the cell poles. In this context, it is noteworthy that mycobacteria have the ability to adopt a wall-deficient lifestyle⁴⁰, and it’s worth highlighting that Mycobacterium tuberculosis possesses a StlP homolog.

Our study also advances the understanding of the functional divergence of the SPFH protein family, emphasizing their evolutionary specialization across taxa. Although conserved in both prokaryotes and eukaryotes, SPFH proteins have adapted to distinct roles. For example, while the stomatin-like protein StoA in filamentous fungi affects hyphal morphology without being essential for apical membrane microdomain formation⁴¹, StlP in Streptomyces is critical for microdomain formation at hyphal tips. Interestingly, in bacteria, SPFH proteins appear to universally contribute to membrane fluidity and cell wall biosynthesis, with examples such as flotillin in Bacillus subtilis, which regulates MreB dynamics²⁴, and in Staphylococcus aureus, where it tethers PBP2a to facilitate peptidoglycan assembly²⁵. Our findings reveal that StlP in Streptomyces not only regulates localized membrane fluidity but also impacts cellulose-like glycan biosynthesis, a key cell wall component. These observations suggest that SPFH proteins engage in analogous biological processes—such as cell wall construction and membrane dynamics—by interacting with different client proteins across bacterial species. Recent structural studies of SPFH proteins across domains of life reveal a remarkable ability to form cage-like architectures, albeit with varying cage sizes and membrane topologies^22,28. These structural differences likely dictate protein localization, client interactions, and functional specialization. Collectively, our findings imply the evolutionary plasticity of SPFH proteins, where conserved structural principles are repurposed to meet the distinct physiological demands of prokaryotic and eukaryotic systems. This functional and mechanistic diversity not only highlights the adaptability of the SPFH protein family but also sets the stage for further exploration of their roles in cellular organization and dynamics.

Methods

Strains and growth conditions

Strains used in this study are listed in Supplementary Table 1. Solid MS (Mannitol Soy flour) medium⁴² was used for collection of Streptomyces spores and for conjugation experiments, while MYM medium⁴³ was used for obtaining Kitasatospora spores. To compare colony sizes and observe the release of cell wall-deficient (CWD) cells under hyperosmotic stress, solid L-phase medium agar (LPMA) medium⁸ or a modified LPMA medium (in which the 0.64 M sucrose was replaced with 0.32 M NaCl) were used. TSBS⁴² medium was used to grow Streptomyces in liquid medium without hyperosmotic stress. For quantification of the number of CWD cells, as well as measurement of hyphal diameters and membrane fluidity, liquid L-phase broth (LPB) was used⁸. Briefly, 10⁶ CFU ml⁻¹ spores were inoculated in 20 ml LPB in 50 ml flasks without coil while shaking at 100 rpm ml⁻¹. All Streptomyces and Kitasatospora strains were grown at 30 °C. For hyphal branching detection, spores were inoculated onto cellophane membranes overlaying LPMA plates, which were then incubated at 30 °C for 16 h prior to analysis. Lysozyme sensitive assay were performed essentially as described¹⁹. Briefly, approximately 1000 spores of each strain were plated on Difco nutrient agar plates either or not supplemented with 0.25 mg ml⁻¹ lysozyme (from chicken egg white, ≥40,000 units mg⁻¹, Sigma). After 48 h of growth, the total number of colonies was counted. For every strain, percentage of the number of colonies on the plate with lysozyme relative to the number of colonies on the plate without lysozyme as an estimate for lysozyme sensitivity.

E. coli DH5α⁴⁴ was used for cloning and β-lactamase experiments. E. coli BL21 (DE3)⁴⁵ and BTH101⁴⁶ were used in protein expression and bacterial two-hybrid assays, respectively. For conjugation, E. coli ET12567 harboring pUZ8002 was used. All E. coli strains were grown in LB medium at 37 °C with appropriate antibiotics, if needed.

Plasmid construction and transformation

Plasmids and primers used in this study are listed in Supplementary Tables 2 and 3, respectively.

To create the ∆stlP_FL mutant (i.e., the stlP mutant in which the entire coding sequence of stlP was removed), the unstable multi-copy vector pWHM3-oriT was used as described⁴⁷. The 1.2 kb of upstream and downstream regions of the stlP were amplified from genomic DNA of S. coelicolor with primer pairs LArm-F/LArm-R and RArm-F/RArm-R, respectively. The apramycin resistance cassette aac(3) IV flanked by loxP sites was amplified from plasmid pGWS728⁴⁸ using primers Apra-F/Apra-R. Amplified PCR products were Gibson assembled with XbaI-linearized pWHM3-oriT, yielding plasmid pXZ44.

For complementation of the stlP mutant, the coding sequence of stlP (SCO2834) was amplified from genomic DNA of S. coelicolor with primers stlP-F/stlP-R1. The PCR product was ligated as an NdeI/BamHI fragment into plasmid hpXZ2¹⁹ harboring the constitutive gapAp prompter of SCO1947, yielding plasmid pXZ15.

For localization of StlP and DivlVA, the mCherry reporter was fused to the C-terminus of these two proteins. Coding sequences for stlP and divIVA were amplified from genomic DNA of S. coelicolor with primers stlP-F/stlP-R2 and 2077-F/2077-R, respectively. The gene encoding for mCherry was amplified from plasmid pGWS791²⁹ using primers mCh-F/mCh-R. After digestions with NdeI/HindIII (for stlP and divlVA) and HindIII/XbaI (for mCherry), combinations of stlP/mCherry and divIVA/mCherry were ligated with plasmid hpXZ2 that was cut with NdeI/XbaI, generating plasmid pXZ16 and pXZ17, respectively.

To constitutively express the HflK/C-like protein BOQ63_030050 in Kitasatospora viridifaciens and the S. coelicolor stlP mutant, the coding sequence of BOQ63_030050 was amplified from genomic DNA of K. viridifaciens DSM40239 with primers 0050-F/0050-R. The PCR product was ligated as an NdeI/XbaI fragment into plasmid hpXZ2 harboring the constitutive gapAp prompter, yielding plasmid pXZ41.

To express the stomatin-encoding domain of StlP (StlP_SD) in E. coli, nucleotides 610-1107 of stlP were amplified from S. coelicolor genomic DNA with primers 2834_sto-F/2834_sto-R and ligated into pET28a plasmid as a NdeI and HindIII fragment, yielding plasmid pXZ18. Consequently, StlP_SD was expressed carrying a N-terminal Histidine-tag (6xHis).

For construction of the plasmids used in the pull-down assays, nucleotides 298-1152 of stlP were amplified with primers StlP-Fc/StlP-Rc and ligated into pACYCDuet-1 plasmid via NcoI and NdeI, yielding plasmid pXZ42. The coding sequence of cslA (SCO2836) and glxA (SCO2837) were amplified form S. coelicolor M145 genomic DNA with primer combinations CslA-F/CslA-R and GlxA-F/GlxA-R, respectively. The amplified fragments were sequentially ligated into pETDuet-1 using NcoI/HindIII (for inserting cslA) and NdeI/XhoI (for inserting glxA), yielding pXZ43. Using these plasmids, StlP and CslA are expressed with a C-terminal 8×Histidine-tag and Flag-tag, respectively, while GlxA was expressed without any tags.

To construct plasmids used in the biofluorescence complementation assay, enhanced GFP (eGFP) was split into two fragments as previously described⁴⁹. the N-terminal fragment (eGFP_N), comprising amino acid residues 1–158, and the C-terminal portion (eGFP_C), consisting of residues 159–239. The eGFP_N and eGFP_C fragments were then fused to either the N- or C-terminus of the target proteins (StlP, CslA and GlxA) via a flexible linker sequence, GSGGSG. To this end, combinations of PCR products were assembled using Gibson assembly into the pSET152 or pWHM3-oriT vector that have been digested with BamHI, which generates plasmids expressing various proteins fusions, under the control of constitutive gapAp promoter (see Supplementary Table 4 for details).

All plasmids were introduced into E. coli and Streptomyces via heat-shock transformation⁵⁰ and conjugation⁴², respectively.

Inactivation of stlP in Streptomyces

For inactivation of stlP in S. coelicolor, we used cosmid StE20 carrying the Tn5062 transposon inserted after nucleotide position 892 relative to the start site of stlP (kindly provided by Prof. Paul Dyson, see Supplementary Table 2). This cosmid was introduced into S. coelicolor via conjugation using ET12567/pUZ8002⁴², after which exconjugants were screened as described⁵¹. Colonies that were kanamycin-sensitive and apramycin-resistant were selected and used for further analysis. Mutants carrying the expected phenotype were verified by sequencing.

Growth curve generation

For preparing germinated spores, spores of different strains were resuspended in double-strength germination medium⁴² at a final concentration of 10⁶ CFU ml⁻¹ and incubated at 30 °C for 6–8 h while shaking at 200 rpm min⁻¹. For generating the biomass growth curve, the RoboLector L-4-BL-II equipped with a parallelized shaken cultivation device was used⁵². Briefly, the germinated spores were centrifuged and resuspended in either TSBS or LPB media before being distributed (1 ml per well) into a 48-well FlowerPlates (Basesweiler Germany). The temperature and humidity were set at 30 °C and 85%, respectively. The biomass was collected automatically and measured in arbitrary units. All measurements were performed in triplicate.

Bacterial 2-hybrid analysis

To assess interactions of StlP with other tip-localizing proteins, the bacterial hybrid assay was used⁴⁶. Therefore, lpmP (SCO2833), stlP (SCO2834), sco2835, cslA (SCO2836), glxA (SCO2837), cslZ (SCO2838), divIVA (SCO2077), filP (SCO5396) and scy (SCO5397) were amplified using primers Th2833-F/Th2833-R, Th2834-F/Th2834-R, Th2835-F/Th2835-R, Th2836-F/Th2836-R, Th2837-F/Th2837-R, Th2838-F/Th2838-R, divlVA-F/divlVA-R, filP-F/filP-R and scy-F/scy-R, respectively. All amplified DNA fragments were cloned into the pKT25 and pUT18C plasmids using EcoRI and XbaI, yielding plasmids pXZ19 (pUT18C + lpmP), pXZ20 (pKT25 + stlP), pXZ21 (pUT18C + stlP), pXZ22 (pKT25 + SCO2835), pXZ23 (pUT18C + SCO2835), pXZ24 (pKT25 + cslA), pXZ25 (pUT18C + cslA), pXZ26 (pKT25 + glxA), pXZ27 (pUT18C + glxA), pXZ28 (pKT25 + cslZ), pXZ29 (pKT25 + divIVA), pXZ30 (pUT18C + divIVA), pXZ31 (pKT25 + filP), pXZ32 (pUT18C + filP), pXZ33 (pKT25 + scy) and pXZ34 (pUT18C + scy) (see Supplementary Table 2).

E. coli BT101 carrying combinations of these constructs were used in bacterial 2-hybrid experiments to evaluate protein interactions as described⁵³.

Topology determination of StlP

To study the transmembrane topology of StlP, the β-lactamase-encoding gene blaM without its signal sequence was fused to the 3’ end of stlP, as described previously²⁹. Only if BlaM is secreted, cells will be resistant to ampicillin. To this end, a blaM variant lacking the region encoding the signal sequence for secretion (blaM_NS) was amplified from plasmid pHJL401⁵⁴ with primers blaM-F/blaM-R. In parallel, full length blaM (including the region for the signal sequence, hereinafter referred to as blaM_FL) was amplified from the same plasmid using the blaM_FL-F/blaM-R primers. The stlP gene was amplified from S. coelicolor genomic DNA using primers stlP-F/stlP-R3. Subsequently, the amplified products were cut using restriction enzymes NdeI-HindIII (stlP), HindIII-EcoRI (blaM_NS) and NdeI-EcoRI (blaM_FL), and digested combinations of stlP/blaM_NS, and blaM_FL were separately ligated into pXZ2¹⁹ that was cut with NdeI-EcoRI, yielding pXZ35 and pXZ36 (see Supplementary Table 2).

E. coli DH5α harboring plasmid pSET152, pXZ35 and pXZ36 were used to assess the membrane topology of StlP, which were performed as described previously²⁹. Full-length BlaM (BlaM_FL) expressed from pXZ36 served as a control for β-lactamase activity in the medium.

Quantification of CWD cells

Culturing and filtration of CWD cells of Streptomyces and Kitasatospora strains was essentially performed as described⁸, with the exception that S. coelicolor strains were grown for 16 h, while K. viridifaciens strains were grown for 40 h. Quantification of CWD cells was performed with a Bright-Line^TM Hemocytometer (Merck), as described⁵⁵. Briefly, 10 μl of filtered culture supernatant were loaded into the counting chamber, after which cells were quantified under a Zeiss Axio microscope equipped with an Axiocam 105 camera. CWD cell numbers were counted manually, and the density was calculated. For each strain, the measurements were performed in triplicate.

Protein expression and purification

To purify the stomatin-encoding domain of StlP (StlP_SD), E. coli BL21 (DE3) cells harboring plasmid pXZ17 were cultured at 37 °C to an OD₆₀₀ of 0.8 in LB medium containing 50 mg ml⁻¹ kanamycin. Then, 0.5 mM isopropyl β-D-thiogalactopyranoside was added to induce protein expression, after which cells were grown at 30 °C for 18 h. The induced cells were subsequently lysed by sonication in binding buffer (50 mM Tris–HCl, 200 mM NaCl, pH 8.0), and after centrifugation, the lysate was loaded on a Ni²⁺-chelating column equilibrated with binding buffer. Ten column volumes of washing buffer (50 mM Tris–HCl, 200 mM NaCl, 0.1 mM imidazole, pH 8.0) and 5 mL of elution buffer (50 mM Tris–HCl, 200 mM NaCl, 10 mM imidazole, pH 8.0) were used to wash and elute StlP_SD, respectively. Finally, the protein was purified by gel filtration using a Hiload 16/600 Superdex 200 pg column (GE Healthcare) equilibrated with buffer (50 mM Tris–HCl, 100 mM NaCl, pH 8.0). Sample fractions were analyzed on a 12.5% SDS-PAGE gel. Fractions were stored directly at −80 °C or first concentrated to 1 mg ml⁻¹ with the 3 kDa molecular weight cutoff concentrator (Millipore) and then stored.

Pull-down experiments

The pull-down assay of StlP-8His with CslA-Flag and GlxA was performed with heterologous E. coli BL21 (DE3) cells harboring plasmid pXZ42 and pXZ43. Cells were cultured at 37 °C in Terrific Broth (TB) medium containing 50 mg ml⁻¹ ampicillin and 25 mg ml⁻¹ chloramphenicol. The subsequent protein purification was largely performed as previously described¹⁷, with some modifications. Briefly, the induced cells were collected and then lysed by sonication. Subsequently, the crude membranes were collected and solubilized. After removing the insoluble membrane fraction by centrifugation, the soluble membrane fraction was incubated overnight at 4 °C with equilibrated Ni²⁺ resin. The resin was packed in a gravity flow chromatography column and washed with 50 mL modified WB1 buffer containing 50 mM phosphate buffer, pH 7.2, 200 mM NaCl, 8% glycerol, 60 mM imidazole and 0.02% DDM (n-Dodecyl-β-D-Maltoside, Sigma). Proteins were eluted with 15 mL elution buffer containing 50 mM phosphate buffer, pH 7.2, 200 mM NaCl, 8% glycerol, 500 mM imidazole and 0.02% DDM.

For SEC analysis, the eluted protein complexes were concentrated using 100-kDa cut-off Amicon® ultra centrifugal filters (Millipore) before being loaded into the Superdex 200 Increase 10/300 GL column, which has been equilibrated with buffer containing 50 mM phosphate buffer, pH 7.2, 200 mM NaCl, 8% glycerol and 0.02% DDM.

SDS-/Native-PAGE and immunoblotting assay

SDS- and Native-PAGE analyses were performed using the Bio-Rad Mini-PROTEIN system with manually-made 7.5% gels and 12% Mini-PROTEAN® TGX™ Precast Protein Gels (BIO-RAD), respectively. Coomassie Brilliant Blue (Sigma) was used to visualize protein bands. The proteins were detected by immunoblotting, which was essentially performed as described¹⁸. For the detection of the StlP-8×His, CslA-Flag and β-lactamase, the His-tag and DYKDDDDK tag monoclonal antibodies (Proteintech) and β-lactamase monoclonal antibody (Thermo Fisher) were used as the primary antibodies, respectively. Subsequently, anti-Mouse IgG-Alkaline Phosphatase (Sigma) was used as the secondary antibody. The primary and secondary antibodies used for detection of GlxA were identical to those described¹⁸. Images were collected using an Epson Perfection V37 scanner.

Microscopy

To visualize the emergence of CWD cells, spores of the wild-type strain and the ∆stlP mutant were pre-germinated in double strength germination medium⁴². Then 10 μl of germlings were used for live-imaging, for which an ibiTreat 35 mm low imaging dish (ibidi) and a LPMA-pad was used as before⁸. Live imaging was carried out using a Zeiss LSM900 Airyscan 2 microscope. If necessary, Z-Stack acquisitions were used. For visualization of membrane and nucleic acids, 0.05 mg ml⁻¹ FM5-95 and 0.5 μM SYTO-9 (Sigma) were used, respectively.

The detection of nascent peptidoglycan was done by using BODIPY-FL vancomycin (Sigma), essentially as described⁵⁶. Briefly, after growing Streptomyces strains in LPB medium for 16 h, mycelia were collected (3300 × g, 10 min) and resuspended in 200 μl of fresh LPB medium containing 1 μg ml⁻¹ BODIPY-FL vancomycin and 1 μg ml⁻¹ vancomycin. After 10 min incubation at 30 °C, the mycelia were washed 3 times with PBS. Five μl of the washed mycelia was used for microscopy analysis using a Zeiss LSM900 Airyscan 2 microscope.

For visualizing fluid membrane microdomains, mycelia were stained with DilC₁₂^31,32. Briefly, DilC₁₂ was dissolved in DMSO and 10 mg ml^_1 stock was prepared. For sample preparation, spores of each strain were inoculated in LPB medium at a final concentration of 10⁶ CFU ml⁻¹. After 16 h of growth, mycelia were collected by centrifuge (3300 × g, 10 min) and resuspended in prewarmed fresh LPB medium supplemented with 100 µg ml⁻¹ DilC₁₂, followed by growth for 3 h at 30 °C while shaking at 100 rpm min⁻¹. Then, mycelia were collected, washed 3 times with prewarmed LPB supplemented with 1% DMSO, and resuspended in the same wash buffer. DilC₁₂ signal was detected via a Cy3 filter (535 nm excitation and 590 nm emission) using a Zeiss LSM900 Airyscan 2 microscope, wherein the cultivation chamber had been set 30 °C to avoid temperature changes during imaging.

For measurement of membrane fluidity, samples were prepared essentially as described²⁶. Briefly, Laurdan (6-Dodecanoyl-N, N-dymethyl2-naphthylamine, Sigma) was dissolved in dimethylformamide (DMF) and a 10 mM stock was prepared. For preparation of mycelia for Laurdan staining, 20 ml 16 h-old mycelia were collected (3300 × g, 10 min) and resuspended in 1 ml 30 °C pre-warmed LPB medium containing 1 mM Laurdan. For sample preparation of the stlP mutant, the culture was filtrated through a 100 μm cut-off filter (Falcon Cell Strainer 100 μm Nylon) to remove the majority of the CWD cells. The filtered mycelium was resuspended in 1 ml 30 °C pre-warmed LPB medium and used for Laurdan staining. After 10 min incubation in the dark at 30 °C, stained mycelia were collected again and washed twice in 30 °C pre-warmed PBS buffer supplemented with 20% sucrose and 1% DMF and finally resuspended in 200 μl pre-warmed PBS buffer supplemented with 20% sucrose. Fluorescent intensities were measured at 435 and 490 nm, following excitation at 350 nm using a Zeiss LSM900 Airyscan 2 microscope, wherein the cultivation chamber had been set 30 °C to avoid temperature changes during imaging. To calculate the membrane fluidity at hyphal tips, the tip region was cropped as a square (1 μm × 1 μm) from the image and the corresponding generalized polarization (GP) value was determined using ImageJ software in combination with the macro tool CalculateGP designed by Norbert Vischer (https://sils.fnwi.uva.nl/bcb/objectj/examples/CalculateGP/MD/gp.html) as described⁵⁷.

For measurement of hyphal diameters, 16 h-old mycelia were collected (3300 × g, 10 min) and resuspended in fresh LPB medium containing 0.05 mg ml⁻¹ FM5-95. The distance between the stained membranes was used to measure the hyphal diameters by averaging the diameter at 3 distinct spots in the hypha.

The cellulose-like glycan at hyphal tips was visualized by calcofluor white (Sigma) staining and quantified as previous described¹⁹. Briefly, 10⁶ CFU ml⁻¹ spores were inoculated in 20 ml LPB in 50 ml flasks without coil while shaking at 100 rpm ml⁻¹. After 16 h of growth, mycelia were collected (3300 × g, 10 min) and resuspended in 200 μl fresh LPB medium containing 100 μg ml⁻¹ Calcofluor white (Thermo Fisher, catalog R21507). After staining for 10 min at room temperature, 5 μl of the stained mycelia was used for microscopy analysis using a Zeiss LSM900 Airyscan 2 microscope. For quantitatively comparing fluorescence, the measure region with the size of 15 μm × 15 μm squares at hyphal tips was used.

For analysis of hyphal branching patterns, mycelium was grown from single spore and imaged using a Zeiss Axio microscope equipped with an Axiocam 105 camera as described previously¹⁹. The distance from the tip to the proximal branch point was measured. A proximal branch was defined as having a length of 1–4 μm as previous described⁵⁸.

For measurement of colony sizes, strain were grown on LPMA medium in petri dishes with a 9 cm diameter aiming for ±100 colonies per plate. After growing them for 5–7 days at 30 °C, plates were scanned with Epson Perfection V37 scanner and colony size was measured subsequently.

For biofluorescence complementation assay, spores of each strain (M145 + pXZ45, M145 + pXZ46, M145 + pXZ47, M145 + pXZ48, M145 + pXZ49, M145 + pXZ50, M145 + pXZ51, M145 + pXZ45 + pXZ47, M145 + pXZ45 + pXZ48, M145 + pXZ46 + pXZ47, M145 + pXZ46 + pXZ48, M145 + pXZ45 + pXZ49, M145 + pXZ46 + pXZ49, M145 + pXZ50 + pXZ49, M145 + pXZ51 + pXZ49) were inoculated in LPB medium at a final concentration of 10⁶ CFU ml⁻¹. After 16–24 h of growth (depends on the growth speed), mycelia were collected by centrifuge (3300 × g, 10 min) and signal of eGFP at hyphal tips were detected using the Zeiss LSM900 Airyscan 2 microscope. All measurement and images processing were executed with ImageJ software (version 2.0.0/1.53c/Java 1.8.0_172/64-bit).

Sacculus isolation and cryo-electron tomography

Isolation of sacculi of S. coelicolor M145 and the stlP mutant was essentially performed as described⁵⁹, except that 16 h-old liquid cultures were used and the step of removing teichoic acids was neglected.

Sample preparation for cryo-electron tomography (cryo-ET) was performed as described⁵⁹. Briefly, after adding the colloidal gold beads, sacculi solutions were vitrificated and applied on the EM grids. Grids were examined using a 120 kV Talos TEM (FEI/Thermo Fisher) and cryo-ET data were collected using a Titan Krios instrument (Thermo Fisher Scientific). The measurement of cell wall thickness was performed as described⁵⁹.

Bioinformatic analysis

Protein domains and protein structures were predicted by InterPro (https://www.ebi.ac.uk/interpro/) and AlphaFold 2.0⁶⁰. The prediction of protein membrane topology was performed by TransMembrane Hidden Markov Model (TMHMM) (Version 2.0) (https://services.healthtech.dtu.dk/service.php?TMHMM-2.0). Alignment of protein structures was done by PyMOL software (Version 2.5). Amino acids sequence alignment was done by ESPript 3.0 (https://espript.ibcp.fr/ESPript/cgi-bin/ESPript.cgi).

To phylogenetically compare StlP with other SPFH proteins, MEGA 7 was used. Amino acid sequences of all SPFH proteins were downloaded from the UniProt database. For phylogenetic analysis of the distribution of StlP, the amino acid sequence of StlP was used to run Position-Specific Iterative (PSI)-BLAST to find homologs in the dataset of 15405 reference sequence database (RefSeq) representative bacteria and archaea. The homologs with a bitscore >130 were chosen and subsequently each hit was subjected to membrane topology prediction using TMHMM server (Version 2.0). Only hits with an identical membrane topology were considered valid StlP homologs. The phylogenetic tree was annotated using iTOL (https://itol.embl.de/).

Statistical analyses and reproducibility

For statistical analyses, GraphPad Prism software (version 8.0.2) was used. p < 0.05 was considered as significance. All experiments were repeated at least three times with similar results.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The predicted protein structures have been deposited in Figshare (https://figshare.com/articles/media/Predicted_StlP_structures/26161969). Source data are provided with this paper.

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Acknowledgements

This work was funded by a Vici grant (VI.C.192.002) from the Dutch Research Council to Dennis Claessen and a grant from the Deutsche Forschungsgemeinschaft (BR2815/7-1) to S.S.M.B.

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Ariane Briegel
Present address: Integrative Structural Cell Biology Unit, Department of Structural Biology and Chemistry, CNRS UMR 3528, Institut Pasteur, 75724, Paris, France

Authors and Affiliations

Molecular Biotechnology, Institute of Biology, Leiden University, Leiden, The Netherlands
Xiaobo Zhong, Veronique M. A. Ongenae, Guillermo Guerrero Egido, Shraddha Shitut, Chao Du, Erik Vijgenboom, Gilles P. van Wezel, Ariane Briegel & Dennis Claessen
Institute for General Microbiology, Christian-Albrechts-University of Kiel, Kiel, Germany
Sarah S. M. Baur & Marc Bramkamp
Departamento de Microbiología, Facultad de Ciencias, Campus Universitario de Teatinos s/n, Universidad de Málaga, 29010, Málaga, Spain
Victor Carrion Bravo

Authors

Xiaobo Zhong
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Sarah S. M. Baur
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Veronique M. A. Ongenae
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Guillermo Guerrero Egido
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Shraddha Shitut
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Chao Du
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Erik Vijgenboom
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Gilles P. van Wezel
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Victor Carrion Bravo
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Ariane Briegel
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Marc Bramkamp
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Dennis Claessen
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Contributions

X.Z. conducted the majority of the experiments. S.M.B. and S.S. assisted with the membrane fluidity analysis. V.O. carried out the sacculi imaging using Cryo-ET. G.G.E. and C.D. performed the phylogenetic analyses. E.V., G.P.W., V.C.B., A.B., and M.B. provided guidance and manuscript editing. D.C. oversaw the project. X.Z. and D.C. prepared the manuscript.

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Correspondence to Dennis Claessen.

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Zhong, X., Baur, S.S.M., Ongenae, V.M.A. et al. The stomatin-like protein StlP organizes membrane microdomains to govern polar growth in filamentous actinobacteria under hyperosmotic stress. Nat Commun 16, 2669 (2025). https://doi.org/10.1038/s41467-025-58093-x

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Received: 18 January 2024
Accepted: 07 March 2025
Published: 18 March 2025
DOI: https://doi.org/10.1038/s41467-025-58093-x

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