Shigella flexneri evades septin-mediated cell-autonomous immunity via protein ADP-riboxanation

Domain: neuroscience
Published: 2026-01-27
Editorial outcome: major_revision

Abstract
Cell-autonomous immunity represents evolutionarily conserved defense mechanisms present in both immune and non-immune cells. One of such mechanisms is mediated by cytoskeletal septins that entrap cytosolic bacterial pathogens within cage-like structures. To promote infection, Shigella flexneri delivers effector proteins directly into host cells via a type III secretion system. Here we demonstrate that OspC effectors enable Shigella flexneri to evade septin cage entrapment. Mechanistically, OspC catalyzes ADP-riboxanation of SEPT9 at Arg561, a site essential for stabilizing septin hetero-oligomers. Notably, Arg561 ADP-riboxanation impairs septin polymerization and hence assembly of higher-order structures, including filaments and cage-like structures. Furthermore, we provide evidence that OspC effectors act synergistically with OspG to antagonize septin cage entrapment via two distinct post-translational modifications, thereby facilitating cell-to-cell spread and intracellular replication. Overall, our work reveals the elegant strategies of bacterial pathogens to evade septin-mediated cell-autonomous immunity and offers avenues for therapeutic intervention.

Introduction
Shigella spp. are Gram-negative human pathogens that cause bacillary dysentery by invading the colonic epithelium1. Shigella continues to be a major public health concern with over 160 million shigellosis cases worldwide annually, including up to 100,000 fatalities especially for children under 5 in developing countries2. Among different Shigella species, Shigella flexneri (S. flexneri) is the most common cause of shigellosis and also the most studied in research laboratories worldwide1. S. flexneri pathogenesis is largely dictated by its type III secretion system (T3SS) encoded on a large (~200 kb) virulence plasmid. The T3SS is a syringe-like molecular apparatus that functions as a channel and allows direct delivery of virulence factors (known as effector proteins) into host cells. These effectors (~30 proteins) have the capacity to subvert diverse cellular processes in a highly orchestrated manner, thereby promoting bacterial invasion as well as subsequent dissemination to neighboring host cells. Once internalized in host epithelial cells, S. flexneri actively breaks the initial phagocytic vacuole and gains direct access to its cytosolic replicative niche.

To counteract pathogen replication, host innate immune defenses and other antimicrobial processes, including cell-autonomous immunity, are activated with the ultimate goal of clearing infection. Cell-autonomous immunity refers to evolutionarily conserved defense pathways present in both professional immune cells and non-immune cells (e.g., epithelial cells) to defend against invading microbial pathogens3. Recent studies have shown that S. flexneri is strikingly adept at evading cell-autonomous immunity via dedicated T3SS effectors4. Notably, the S. flexneri effector IpaH9.8 subverts guanylate-binding protein (GBP)-mediated immunity through a mechanism dependent on its E3 ubiquitin ligase activity and proteasome-mediated degradation of GBPs5,6.

Septins are a family of highly conserved cytoskeletal proteins that can form filaments and interact with cellular membranes, actin filaments, and microtubules7,8. Human cells encode 13 septin paralogs, which are categorized into four distinct subgroups based on amino acid sequence identity, including SEPT2 (SEPT1, SEPT2, SEPT4, SEPT5), SEPT3 (SEPT3, SEPT9, SEPT12), SEPT6 (SEPT6, SEPT8, SEPT10, SEPT11, SEPT14), and SEPT78–10. Subunits from each of the four septin subgroups assemble into hetero-oligomeric complexes in a specific order, including hexamers (SEPT2–SEPT6–SEPT7–SEPT7–SEPT6–SEPT2) and octamers (SEPT2–SEPT6–SEPT7–SEPT9–SEPT9–SEPT7–SEPT6–SEPT2), with alternating G-interface (consisting of the GTP-binding domain) and NC-interface (comprising the amino- and carboxy-terminal regions) along the linear hetero-oligomeric complex8,11. In addition to their classical role in cell division, recent studies have also revealed septins as crucial players in host-pathogen interactions. During S. flexneri infection, septins are recruited to IcsA-mediated actin polymerization and form cage-like structures around cytosolic bacteria. These septin cages counteract S. flexneri actin-based motility and recruit the autophagic adaptor p62 and NDP52 to target bacteria for autophagic destruction12–14.

Recently, we uncovered a S. flexneri kinase effector OspG that modulates host ubiquitin signaling to evade septin cage entrapment. Mechanistically, OspG phosphorylates a master regulator of host cullin-RING E3 ligases (CRLs) and promotes ubiquitination of septins, thereby leading to cage destruction. As a result, significantly more cytosolic bacteria were caged by septins in the case of an ospG-deficient S. flexneri mutant, as compared to its parental WT strain15. Nevertheless, even for the ΔospG strain, the majority of intracellular bacteria were not entrapped by septin cages in infected host cells. We therefore hypothesized that S. flexneri may have evolved additional strategies to target septins and subvert cage assembly.

Here we show that the S. flexneri OspC family of T3SS effectors directly modifies septins via protein ADP-riboxanation, a recently identified derivative of classical ADP-ribosylation16,17. Furthermore, we demonstrate that SEPT9 is a physiological substrate of OspC effectors during S. flexneri infection. Notably, OspC effectors target Arg561 of SEPT9, a critical residue for stabilizing the NC-interface to maintain inter-subunit contacts within septin octamers, thereby preventing the assembly of septin cages. These findings reveal bacterial effectors that have the capacity to directly target septins and thus facilitate pathogen escape from cell-autonomous immunity.

Results
Septins are conserved substrates of the S. flexneri OspC family of effectors
The OspC family of effectors includes OspC1, OspC2, OspC3, and OspC4. These effectors all contain a C-terminal ankyrin-repeat domain (ARD) and an N-terminal ADP-riboxanase domain, overall exhibiting more than 60% sequence identity (Figure S1a). Previously we have shown that OspC3, but not OspC1/2, inhibits host pyroptosis via ADP-riboxanation of caspase-11/417, while all three effectors (OspC1, OspC2 and OspC3) are able to modify the eukaryotic translation initiation factor 3 (eIF3) complex, leading to protein translational arrest and stress granule formation16. In addition, the OspC effectors, primarily OspC1 and OspC3, inhibit the expression of interferon-inducible genes18. To further compare the substrate spectrum of OspC effectors, we catalogued potential OspC3 targets via ADP-ribosylome profiling (Figure S1b, see methods for more experimental details) and analyzed them together with those of OspC1. In total, we ended up with 88 candidates of ADP-riboxanated substrates, of which 41 proteins were shared by both OspC3 and OspC1. Our data also suggested that OspC3 likely has substantially more exclusive targets than OspC1 (38 versus 9, see Fig. 1a and Supplementary Data 1). In agreement, immunoblotting data of OspC-expressing cells (probed with an antibody specific to ADP-ribose) revealed that OspC3 samples have more intense bands than OspC1 (Fig. 1a). Gene ontology (GO) analyses suggested that core targets of OspC1 and OspC3 are likely to be evolutionarily conserved, whereas divergent functions may have evolved for individual effectors, such as double-strand break repair for OspC3 and vesicle targeting for OspC1 (Fig. 1b). Next, we focused our attention on conserved cellular pathways that are targeted by both effectors. Consistent with our previous study, the eIF3 complex represents the most enriched GO term in OspC3 and OspC1 datasets. Notably, ‘septin complex’ and ‘septin ring’ appear as the next highly enriched terms (Fig. 1b). Indeed, three out of four septin octamer subunits (i.e., SEPT2, SEPT7 and SEPT9, Figure S1c) exhibited higher mass spectrometry signals in cells expressing catalytically active OspCs (Fig. 1c).

Ectopic expression of OspC effectors promotes ADP-riboxanation of septins in host cells
Considering that proteins identified in our ADP-ribosylome datasets are not necessarily modified (i.e., they may be enriched due to interaction with bonafide ADP-riboxanated substrates), we next sought to determine exactly which septins are post-translationally modified by OspC effectors. We individually expressed a panel of FLAG-tagged septin subunits (SEPT2, SEPT6, SEPT7, SEPT9, SEPT10, and SEPT11) in 293T cells together with either WT OspC3 or its catalytically dead mutant (Fig. 1d). Next, immunoprecipitated septins were probed with an ADP-ribose-specific antibody, which also recognizes protein ADP-riboxanation. Immunoblotting data demonstrated that all tested septins, except for SEPT7, were readily ADP-riboxanated by OspC3, but not the catalytically dead mutant. The inability of detecting modified SEPT7 is likely due to its lower expression levels in our transfection assays (relative to other septins). When assayed alone or with a longer exposure time, SEPT7 was indeed detected with clear modification signals (Figs. 1d, 1g and S2e). Consistent with our proteomic data, these septins were also ADP-riboxanated when they were co-expressed with the other two paralogues, OspC1 and OspC2, in host cells in an enzymatic activity-dependent manner (Fig. 1e–h).

Given that OspC3 exhibited the strongest ADP-riboxanase activity (Fig. 1a), we used it as a representative effector of the OspC family (unless otherwise specified). Previous studies have shown that the ARD domain of OspC3 is essential for caspase-4/11 recognition17. Therefore, we examined whether this is also the case for OspC3 to target septins. SEPT9 co-expressed with either WT OspC3 or its variants was assayed by immunoblotting for potential ADP-riboxanation. Similar to CASP4, SEPT9 was readily modified by full-length OspC3, whereas modification signals dropped substantially with the N-terminal domain truncated (i.e., the Nt mutant) (Figure S2a and S2b). The ARD domain of OspC3 was essential for targeting SEPT2, SEPT6, and SEPT7 (Figure S2c–e). Consistent with this, co-IP experiments demonstrated that full-length OspC3 or its ARD domain, but not the N-terminal enzymatic domain, immunoprecipitated endogenous SEPT2, SEPT6, SEPT7, and SEPT9 (Figure S2f). Together, these results indicate that SEPT9 modification depends on OspC’s catalytic N-terminal domain, the efficiency of which is enhanced by septin-binding via the C-terminal ARD domain.

OspC3 induces ADP-riboxanation of multiple residues on septins
Next, we sought to identify the modification sites of septins by tandem mass spectrometry (MS/MS) analyses. As a representative member, SEPT9 co-expressed with OspC3 exhibited slower migration on SDS-PAGE, indicating potential post-translational modifications (Fig. 2a). MS/MS analyses of digested SEPT9 revealed at least four ADP-riboxanated sites, including Arg17, Arg46, Arg134, and Arg561 (Fig. 2a). While the first three sites all come from the N-terminal extension, Arg561 is located in the C-terminal α6 helices, which participate in forming salt bridges to stabilize the NC-interface. To help prioritize functional studies of these modified residues, we further analyzed the extent of ADP-riboxanation with respect to individual sites; we extracted such information from corresponding peptide intensities observed in mass spectromtery experiments (details are provided in the Methods section). For instance, the peptide T HMQNIKDITSSIHFEAYR *VKR containing ADP-riboxanated Arg561 was only detected in samples from cells expressing WT OspC3 but not its catalytically dead mutant EH/AA (Fig. 2b, c). In contrast, the unmodified peptide T HMQNIKDITSSIHFEAYR was detected in both samples, while its signal dropped sharply in SEPT9 co-expressed with WT OspC3 (Fig. 2b). We estimated a modification rate of 80% based on the extent of such signal attenuation. In comparison, ADP-riboxanation rates at R17, R46, and R134 were markedly lower than that of R561 (Figure S3 and Table 1). Together, these findings suggest that R561 is the major site of ADP-riboxanation.

In line with MS/MS analyses, successive mutation of the identified sites to lysine (1RK to 4RK) resulted in a progressive decrease in modification levels, although residual signals were still visible in immunoblotting assays (Fig. 2d). Indeed, further MS/MS analysis of SEPT9-4RK identified six additional ADP-riboxanation sites (Fig. 2a, right). Nonetheless, a SEPT9 variant with substitution of these six sites to lysine (10RK) was still modifiable, though the signal was further reduced (Fig. 2e). Similarly, we identified at least three modified residues in SEPT2 (Arg29, Arg138, and Arg306) and SEPT6 (Arg299, Arg328, and Arg421), respectively (Figures S4 and S5). These results demonstrate that OspC3 catalyzes multiple modifications on septins.

OspCs directly ADP-riboxanate SEPT9 in vitro and during bacterial infection
To address whether septins are direct substrates of OspC ADP-riboxanase activities, OspC3 and septins were co-expressed in E. coli cells. Immunoblotting assays suggested that OspC3 ADP-riboxanated SEPT9 in vitro in a manner dependent on both its enzymatic activity and the eukaryotic co-factor, calmodulin (CaM)19 (Fig. 2f). Furthermore, in an in vitro reaction using purified recombinant proteins, we found that the OspC3-CaM complex modified SEPT9 in an NAD+-dependent manner (Fig. 2g, lanes 1–5). Consistent with previous reports that stimulation of OspC3 ADP-riboxanase activity requires Ca2+-free CaM, high levels of calcium ions, but not the ion-chelating reagent EDTA, abolished the modification reaction (Fig. 2g, lanes 6–7)19,20. We also analyzed the same sets of SEPT9 samples prepared from E. coli co-expression (Fig. 2f) and in vitro reactions (Fig. 2g) by tandem mass spectrometry analyses. Unexpectedly, only three out of four sites that were previously identified in 293T cells (i.e., Arg17, Arg46, and Arg134) were detected in their modified forms, while Arg561 was not detectably modified (Figures S6a and S6b). Given Arg561 as the primary modification site in 293T cells (Table 1), we speculate that ADP-riboxanation of SEPT9 Arg561 may require spatial orientation of an intact septin complex.

To further determine whether septins are the physiological targets of OspC effectors, we carried out S. flexneri infection of HeLa cells expressing FLAG-tagged septin subunits (i.e., SEPT2, SEPT6, SEPT7, and SEPT9). Immunoprecipitated subunits were probed for potential ADP-riboxanation and SEPT9 was the sole protein that exhibited detectable signals of modification (Fig. 2h). These findings are in line with our previous co-expression data, in which SEPT9 was most modified in 293T cells relative to other septin subunits. Intriguingly, SEPT9 was also modified in uninfected control cells, though this signal was significantly elevated during infection (Fig. 2h). Given the absence of known ADP-riboxanases in eukaryotic cells (at least not found until now), we reasoned that basal levels of SEPT9 modification are likely due to classical ADP-ribosylation.

To further confirm the role of T3SS-delivered OspC effectors in mediating ADP-riboxanation of SEPT9, we constructed a series of S. flexneri mutants missing either individual or a combination of several ospC genes. Successive knock out of ospC genes led to a progressive decrease in ADP-riboxanation levels of SEPT9 during bacterial infection. In particular, simultaneous deletion of ospC1 and ospC3 resulted in substantial loss of modification signals (Fig. 2i and S6c). As expected, minimal signals were observed upon deletion of all four ospC homologs. Complementation of this strain with both OspC1 and OspC3 restored modification at levels higher than wild-type S. flexneri infection (Fig. 2i), likely due to higher expression levels of plasmid-borne effectors (Figure S6d). Furthermore, SEPT9 with substitution of Arg561 to either Ala or Lys failed to undergo OspC-mediated ADP-riboxanation during bacterial infection (Fig. 2i), although the 4RK/10RK mutants still exhibited detectable modifications when co-expressed with OspC3 in 293T cells (Fig. 2e). Immunoblotting analyses of endogenous SEPT9 revealed OspC-dependent ADP-riboxanation during S. flexneri infection (Fig. 2j). In agreement, mass spectrometry analyses showed that ~4% of host SEPT9 was modified in cells infected by WT S. flexneri (Figure S6e). As expected, we observed higher modification rates (~12%) for infection with the WT OspC complementation strain (Table 2). Together, these results suggest that SEPT9 is the preferred substrate of OspC effectors during S. flexneri infection with Arg561 as the primary site of ADP-riboxanation.

OspC-mediated ADP-riboxanation destabilizes the septin complex
To interrogate the potential impact of OspC-catalyzed ADP-riboxanation on septin functions, we next asked whether these modifications would disrupt the interactions between different septin subunits as such associations are thought to play an important role in the assembly of hetero-oligomeric complexes21. Using SEPT9 as a surrogate, we immunoprecipitated SEPT9-associated proteins from cells expressing either wild-type OspC3 or its catalytically dead mutant. Subsequent quantitative mass spectrometry analyses revealed that 8 proteins showed reduced associations with SEPT9, while 15 proteins exhibited enhanced interactions in cells producing WT OspC3 (Fig. 3a, Supplementary Data 2). Importantly, we found that half of the reduced proteins were septin subunits including SEPT6, SEPT7, SEPT8, and SEPT11 (Fig. 3a). In line with these data, catalytically active OspC3 also reduced the interactions between ectopically expressed FLAG-SEPT9 and HA-SEPT2/6 (Fig. 3b, c). Further network analyses of the SEPT9-interactome data clearly revealed a cluster of proteins with markedly reduced interactions as the septin complex (Fig. 3d). Of note, signals derived from ubiquitin was enriched in the interactome of modified SEPT9 (Fig. 3a), indicating potential ubiquitination of SEPT9 or its associated proteins. Additional GO analysis indicated that processes such as septin ring formation and cytoskeleton-dependent cytokinesis were less represented, whereas cytoplasmic translational initiation and nuclear import signal receptor activity were enriched (Figure S7a).

To further support that septin ADP-riboxanation disrupts interactions of different subunits, we co-expressed EGFP-OspC3 and FLAG-SEPT9 in 293T cells. Immunoprecipitated SEPT9 was probed with antibodies specific for endogenous septin subunits. Consistent with previous mass spectrometry data, we found reduced association of SEPT9 with other octamer subunits (i.e., SEPT2, SEPT6, and SEPT7) in an ADP-riboxanation-dependent manner (Fig. 3e). Unexpectedly, we found that the SEPT9 mutants unable to undergo ADP-riboxanation (i.e., 4RK and 10RK) lost their regular subcellular localization and instead exhibited nuclear distribution (Figure S7b). This altered localization may, at least partially, account for the weakened interactions between these mutants and other septin subunits (Fig. 3e). In agreement, in vitro pull-down assays using recombinant proteins purified from E. coli also demonstrated that ADP-riboxanated SEPT9 bound less efficiently to SEPT7 in comparison to its unmodified form. Importantly, SEPT9-SEPT7 interaction was dependent on the modification sites, as further supported by data from 4RK and 10RK mutants (Fig. 3f). These residues may play a critical role in mediating interactions with actin filaments and microtubules, which ultimately determine the subcellular localization of SEPT98.

ADP-riboxanation of SEPT9 at Arg561 destabilizes septin octamer and obstructs septin cage assembly
Next, we focused our attention on the functional roles of ADP-riboxanation of SEPT9 occurring at Arg561 given that it is the primary site of modification induced by OspC effectors during Shigella infection. Importantly, residues from the C-terminal α6 helices (Glu558 and Arg561) and the loops following α2 (Glu394, Arg399) form salt bridges and hence play a critical role in stabilizing the SEPT9-SEPT9 NC-interface22 (Fig. 4a). Given negative charges introduced by ADP-riboxanation17, we reasoned that modification of Arg561 likely destabilizes septin octamers by disrupting salt bridges and the NC-interface.

To test this hypothesis, we constituted SEPT9-containing octamers in vitro by using recombinant proteins purified with a two-tag purification scheme (Figure S8a) as previously described23. Subsequent SDS-PAGE and mass photometry analyses confirmed the stoichiometric assembly of recombinant septin octamers (Fig. 4b and S8b). Of note, SEPT9-WT formed both tetramers (~200 kDa) and octamers (~400 kDa), with the latter being the major form. In contrast, SEPT9-NCmut1 (I281D and M288D) and SEPT9-R561E formed only tetramers, due to the impaired NC-interface as previously reported21,24. Notably, SEPT9 samples co-expressed with WT OspC3 (and CaM), but not its catalytically dead mutant, formed less octamers and more tetramers instead (Fig. 4b), indicating disruption of the SEPT9-SEPT9 NC-interface. Purified heteromeric complexes are capable of self-assembling into filaments under conditions such as low-salt buffers (e.g., 50 mM KCl)7. Consistent with mass photometry analysis, SEPT9-WT octamers, but not complexes containing SEPT9 mutants or ADP-riboxanated proteins, successfully polymerized under low-salt conditions (Fig. 4c). Together, these data support our conclusion that ADP-riboxanation of SEPT9 at Arg561 destabilizes septin octamers by a mechanism involving disruption of salt bridges at the NC-interface.

To determine whether Arg561 modification has any impact on the assembly of septin cages during S. flexneri infection, we transfected FLAG-SEPT9 or its variants in HeLa cells at levels close to endogenous septins. Immunostaining analyses revealed that WT, but not SEPT9-NCmut1, was efficiently incorporated into septin cages following S. flexneri infection (Figure S8c). Interestingly, SEPT9-NCmut2 was still able to assemble septin cages, albeit with lower efficiency, likely due to partial destruction of salt bridges. Similarly, SEPT9 R561K, but not R561A and R561E mutants, formed cage-like structures (Fig. 5a). We further quantified fluorescence intensities of SEPT9 surrounding intracellular bacteria. In agreement with analyses of septin cage assembly, analyses of transfected SEPT9 variants indicated that both R561 and the NC-interface are required for SEPT9 recruitment to intracellular S. flexneri (Fig. 5b and S8d). By comparison, fluorescence intensities of total S. flexneri-associated SEPT9 (including both transfected and endogenous proteins) were not significantly different among the same set of transfected samples (Fig. 5c and S8e). In addition, ADP-riboxanated SEPT9 (induced by ectopic expression of GFP-OspC3, but not the GFP-vector or inactive mutant) failed to assemble into septin cages during bacterial infection (Figure S8f). Quantitative assessment revealed that ~14% of cytosolic bacteria were entrapped by SEPT9-positive cages. Expression of WT OspC3, but not the EH/AA mutant, significantly reduced cage assembly to ~4% (Fig. 5d, e). Collectively, these data suggest that OspC-mediated ADP-riboxanation of SEPT9 at Arg561 impairs salt bridges and disrupts the SEPT9-SEPT9 NC-interface, thereby preventing septin recruitment and S. flexneri cage assembly.

S. flexneri evades septin cage-based cell-autonomous immunity during infection in an OspC-dependent manner
Cytosolic bacterial pathogens, including Shigella, are entrapped by septin cage-like structures to restrict their intracellular mobility and facilitate their targeting to antibacterial autophagy12,14. We next investigated whether T3SS-delivered OspC effectors can antagonize septin cage assembly and hence facilitate bacterial escape during S. flexneri infection. We challenged HeLa cells with WT bacteria and stained for endogenous septins to visualize cage-like structures at 2 h 40 min post infection. Quantitative microscopy revealed that ~11% of cytosolic S. flexneri were compartmentalized in septin cages (Fig. 6a, b). As we previously reported, when cells were infected by an ospG-deletion (ΔospG) strain, septin cages formed at a higher percentage (21%)15. By comparison, HeLa cells challenged with a S. flexneri strain missing all ospC genes (ΔospC) exhibited similar rates of cage assembly (~20%). In addition, reintroduction of both WT OspC1 and OspC3, but not their catalytically dead mutants, into the ΔospC strain restored the ability to block septin cage assembly (Fig. 6c, d). Given that both OspC and OspG effectors target septins via distinct post-translational modifications, we constructed a double-knockout mutant (ΔospCΔospG). Notably, septin cage formation was increased to an even higher level (~31%) in ΔospCΔospG infected cells, thereby indicating that OspCs and OspG work in concert to inhibit cage assembly (Fig. 6a, b).

To further investigate the roles of OspC and OspG in bacterial pathogenesis, we assessed intracellular proliferation and cell-to-cell spread of S. flexneri using gentamicin protection and plaque formation assays, respectively. Notably, simultaneous deletion of both ospC and ospG genes nearly abolished S. flexneri intercellular spread, whereas single deletion mutants (i.e, ΔospC or ΔospG) exhibited partial phenotypes (Fig. 6e). In addition, complementation of the double-knockout strain with either wild-type OspC or OspG, but not the inactive EH/AA mutant, restored bacterial spreading ability (Fig. 6f). Similar data were recorded for intracellular replication, though growth defects of the double mutant were not as severe as cell-to-cell spread (Fig. 6h, i). In all cases, bacterial invasion was unaffected by these deletions (Fig. 6g). Collectively, these findings suggest that OspCs act synergistically with OspG and liberate S. flexneri from cage entrapment by targeting septins through different post-translational modifications (Fig. 6j).

Methods
Plasmids and bacterial strains
DNAs for ospC1, ospC2, and ospC3 were amplified from S. flexneri 2a strain 2457T (kindly provided by Dr. Feng Shao from National Institute of Biological Sciences, Beijing). Complementary DNAs (cDNA) for human SEPT2, SEPT6, SEPT7, SEPT9 (isoform 1, the longest isoform), SEPT10, and SEPT11 were described previously15. DNAs were ligated into pCS2 vectors for transient expression in mammalian cells. For recombinant protein expression in E. coli, DNAs were inserted into pGEX-6p-2, pET28a-6×His, pET28a-6×His-SUMO, pACYCDuet-1 and pET-MCN series vectors, such as pnEA-vH (pET-15b backbone) and pnCS (pCDFDuet backbone) vectors37. All truncations and point mutations were generated by the standard polymerase chain reaction (PCR) cloning method. All plasmids were verified by DNA sequencing.

S. flexneri was freshly streaked from glycerol stocks onto tryptone soy agar supplemented with 0.02% (w/v) Congo red, and subsequently a single red colony was selected and inoculated into tryptone soy broth (TSB) with appropriate antibiotics. For efficient infection of host cells, all S. flexneri strains used in this study were transformed with a plasmid expressing afimbrial adhesin (Afa). The S. flexneri ospC knockout strain was generated using the CRISPR/Cas9 system as described previously38. Briefly, single-guide RNAs (sgRNAs) targeting the coding sequence (CDS) or promoter region of the gene of interest were designed using the CRISPOR website and introduced into the pTarget plasmid via PCR. Notably, the CDSs of ospC2, ospC3, and ospC4 are highly similar, which poses significant challenges for the sgRNA design. It is crucial to avoid targeting multiple ospC genes simultaneously, as this may result in multiple double-strand breaks and cell death. Additionally, the homologous arms should be designed to exclude sequences that could be targeted by sgRNAs used in subsequent rounds of gene editing. Specifically, ospC genes were knocked out in a sequential order: ospC1, ospC3, ospC4, and ospC2, resulting in a strain with all ospC homologs deleted. Sequences of all primers and gRNAs used for Shigella gene deletion are listed in Supplementary Data 3. After obtaining the ospC quadruple mutant, ospG was further knocked out to generate the ΔospCΔospG strain.

For complementation, Strep-tag-II C-terminally-tagged DNAs for ospC1, C-terminal HA-tagged DNAs for ospC3 or their mutants were cloned into the pBBR1-tet vector (generated by replacing the lac promoter and lac operator of pBBR1MCS2 with the tet promoter) for constitutive expression. To visualize S. flexneri in immunofluorescence experiments, the DNA encoding EGFP with a ribosome binding site (RBS) sequence was cloned into the pBBR1-tet vector. All mutant strains were confirmed by PCR using at least two different pairs of primers, and PCR products were further validated by DNA sequencing.

Cell culture, transfection and immunoprecipitation
293T (Cat No. CRL-3216) and HeLa cells (Cat No. CRM-CCL-2) were obtained from American Type Culture Collection (ATCC) and grown in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (TRANS, FS301-02) and 2 mM L-glutamine. Cell monolayers were maintained at 37°C in a humidified 5% CO2 incubator. All cells were tested for mycoplasma using the standard PCR method. Transient transfection was performed using polyethylenimine (PolySciences) or jetPRIME (Polyplus-transfection) following the manufacturer’s instructions.

For immunoprecipitation, 293T cells (~1×107) grown to 80% confluence in 10 cm dishes were transfected with indicated plasmids. Twenty-four hours after transfection, cell pellets were harvested and lysed in the TNT buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, and 1% Triton X-100) supplemented with protease inhibitor cocktail (Selleck, B14002) for 30 min, followed by centrifugation at 4°C (16,000×g for 10 min). The clarified lysates were incubated with pre-washed anti-FLAG M2 affinity beads (Sigma-Aldrich, A2220) at 4°C with gentle rotation for 4 h. The beads were washed four times with the lysis buffer, and bound proteins were eluted with FLAG peptides (MedChemExpress, HY-P0319A) and run on SDS-PAGE followed by either Western blot or LC-MS analyses.

Western blots and antibodies
Protein samples were separated by SDS-PAGE and proteins in the gel were transferred onto PVDF membranes, followed by blocking with non-fat milk for 1 h at room temperature and incubation with specific primary antibodies (diluted at 1:1000 to 1:5000) overnight at 4°C. Afterward, the membranes were extensively washed (for 4 times) with TBST buffer (composed of 25 mM Tris pH 7.5, 140 mM NaCl, 3 mM KCl, and 0.1% Tween-20) and incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies (1:5000). Following another round of washes, proteins of interest were visualized by using a chemiluminescence imaging system (Tanon 5200).

The following primary antibodies were used in this study: anti-poly/mono-ADP ribose (1:2000, Cell Signaling Technology, 83732), anti-GFP (1:2000, Proteintech, 66002-1-Ig), anti-GAPDH (1:5000, Proteintech, 60004-1-Ig), anti-β-actin (1:5000, Cell Signaling Technology, 4967), anti-FLAG for Western blot (1:5000, Zen Bioscience, R24091), anti-FLAG for immunofluorescence (1:300, Beyotime Biotechnology, AF519), anti-HA (1:5,000, Invitrogen, 26183), anti-GST (1:1000, Zen Bioscience, 390028), anti-His (1:1000, CWBIO, CW0286M), anti-S-tag (1:2,500, Sino Biological, 101290-T38), anti-SEPT2 (1:2,500, Proteintech, 11397-1-AP), anti-SEPT6 (1:1000, Santa Cruz, sc-514781), anti-SEPT7 (1:2,000, Proteintech, 13818-1-AP), anti-SEPT9 (IF 1:500, WB 1:2500, Atlas Antibodies, HPA042564), and anti-Strep-Tag II (1:2,500, Abmart, M40014S). The secondary antibodies include HRP-conjugated anti-mouse IgG (1:5000, Jackson ImmunoResearch Laboratories, 115-035-003), HRP-conjugated anti-rabbit IgG (1:5000, Jackson ImmunoResearch Laboratories, 111-035-003), Alexa Fluor 488-labeled anti-mouse IgG (1:500, Jackson ImmunoResearch Laboratories, 115-545-003), Cyanine3 (Cy3)-conjugated anti-rabbit IgG (1:500, Jackson ImmunoResearch Laboratories, 111-165-003), and Alexa Fluor 647-labeled anti-mouse IgG (1:500, Beyotime Biotechnology, A0473).

Bacterial infection of host cells
S. flexneri strains carrying an Afa-encoding plasmid were used for infection experiments in this study. A red colony of S. flexneri, grown on Congo red agar plates, was inoculated into TSB and cultured for approximately 16 h at 37°C with constant shaking. On the following day, overnight bacterial cultures were diluted 1:50 into fresh pre-warmed TSB, and further cultured until an OD600 of 0.6 was reached. Cell monolayers were equilibrated with pre-warmed DMEM for 30 minutes and infected with S. flexneri at indicated multiplicity of infection (MOI).

For detection of septin ADP-riboxanation during infection, HeLa cells transfected with 3×FLAG-septin in 6-well plates for 24 h were infected with relevant S. flexneri strains at an MOI of 50. After 45 min of infection, infected cell monolayers were washed three times with pre-warmed DMEM and further incubated with fresh DMEM containing 10% FBS and 50 μg/mL gentamicin at 37°C and 5% CO2 for 2 h to kill extracellular bacteria. At indicated time points, cell monolayers were collected, lysed, and followed by anti-FLAG immunoprecipitation. Eluted samples were collected for SDS-PAGE and Western blot analyses. For detection of endogenous SEPT9, bacterial infections were performed as described above and cell lysates were subjected to immunoprecipitation with an anti-SEPT9 antibody or an isotype control IgG pre-immobilized on AminoLink Coupling Resin (Thermo Fisher Scientific). Bound proteins were eluted using 0.1 M glycine–HCl (pH 2.7) and immediately neutralized by adding 1/10 volume of 2 M Tris–HCl (pH 8.0).

For microscopy analyses of septin cages, HeLa cells were seeded onto glass coverslips in 24-well plates for 36 h before bacterial infection. For experiments involving mutation of Arg561 or residues at the NC-interface of SEPT9, pre-seeded HeLa cells were further transfected with indicated plasmids for 20 h before infection. Cells were infected with S. flexneri at an MOI of 6 (unless otherwise specified) for 45 min, washed with fresh media and further incubated in fresh DMEM containing 10% FBS and 50 μg/mL gentamicin for 2 h 40 min. Cells were washed twice with PBS and subsequently fixed with 4% paraformaldehyde (PFA) for 15 min. The fixed cells were stored in PBS until further immunofluorescence analysis.

To measure intracellular replication, HeLa cells cultured to 80% confluence in 6-well plates were infected with S. flexneri at an MOI of 20. Bacterial infections were performed as described above. After infection, cell monolayers were incubated with fresh DMEM containing 10% FBS and 100 μg/mL gentamicin at 37°C with 5% CO2 for 30 min to eliminate extracellular bacteria. Subsequently, the gentamicin concentration was reduced to 25 μg/mL. At each time point (0.5, 2, and 4 h), cells were lysed in cold PBS containing 0.1% Triton X-100, and colony-forming units (CFUs) were determined by serial dilution and plating on TSB agar plates containing 30 μg/mL streptomycin.

For investigation of intercellular spread, plaque formation assays were performed as described previously39. Briefly, HeLa cells cultured to 100% confluence in 6-well plates were incubated with S. flexneri lacking a fimbrial adhesin at an MOI of 0.005 for 2 h to facilitate invasion. After incubation, cells were washed three times with pre-warmed DMEM and subsequently overlaid with 3 mL of agarose overlay medium (DMEM containing 5% FBS, 50 μg/mL gentamicin, and 0.5% (w/v) agarose). Following 48 h of incubation, the agarose overlay was removed, and infected HeLa cells were subjected to Giemsa staining for plaque visualization. The plaque area (A) was calculated using the formula A=π(d/2)2, where d represents the diameter of a plaque measured using the ImageJ (v1.48) software. Average plaque areas were determined by analyzing 30 plaques per group.

Purification of recombinant proteins
The E. coli BL21 (DE3) strains transformed with indicated plasmids were used for protein expression and purification. Protein expression was induced by adding 0.4 mM isopropyl β-d-1-thiogalactopyranoside (IPTG) when the OD600 of cell cultures reached approximately 0.8. The bacterial cells were subsequently cultured at either 22°C for 12 h or 18°C for 16 h, depending on the stability and/or solubility properties of target proteins. Proteins tagged with GST, His6, or His6-SUMO were purified using glutathione resins (Sangon Biotech, C600031) or HisTrap HP affinity chromatography (GE Healthcare Life Sciences). If necessary, the GST or SUMO tags were removed by PreScission or Ulp1 protease digestion, respectively. Further purification was achieved through ion-exchange chromatography, followed by size-exclusion chromatography. For downstream applications, buffer exchange was performed by using HiTrap Desalting chromatography (GE Healthcare Life Sciences). Purified proteins were then concentrated and stored at −80°C in a buffer containing 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 2 mM dithiothreitol, and 5% glycerol.

Recombinant human septin complexes, including the SEPT2-msfGFP-SEPT6-SEPT7 hexamer and the SEPT2-msfGFP 8mer-9_i1, were purified as previously described23,29,40. Briefly, E. coli BL21 (DE3) strains were transformed with compatible bicistronic plasmids expressing the respective units, and induced for protein expression. Bacterial lysates were subjected to a two-step affinity purification scheme. First, HisTrap affinity chromatography was used to capture complexes containing N-terminal His6-tagged SEPT2. Next, Strep-Tactin affinity chromatography was employed to isolate hexameric complexes (via the C-terminal Strep-tag II on SEPT7) or octameric complexes (via the C-terminal Strep-tag II on SEPT9). The purified complexes were dialyzed to remove imidazole and reconstituted in stabilization buffer (50 mM Tris-HCl pH 8.0, 300 mM KCl, 5 mM MgCl₂, and 1 mM DTT). This method yielded stoichiometric septin complexes to homogeneity suitable for downstream analyses.

In vitro SEPT9 pull-down assays
Recombinant GST-Strep-tag II-SEPT9 proteins or corresponding 4RK/10RK mutants were incubated with GST-fused SEPT7 at 4°C for 2 h. The mixture was then incubated with Strep-Tactin resins at 4°C for 30 min. The resins were subsequently washed three times with the TNT buffer and boiled in 2× SDS loading buffer. All samples were analyzed via SDS-PAGE followed by Western blot.

Immunofluorescence and microscopy
Cells were fixed with 4% PFA after S. flexneri infection, and washed three times with PBS (5 min per wash). Permeabilization was performed with 0.1% Triton X-100 in PBS for 5 min, and washed again with PBS for three times. Subsequently, cells were blocked with 5% goat serum in PBS for 30 min at room temperature and incubated overnight at 4°C with primary antibodies diluted in PBS containing 1% BSA and 0.05% Tween-20. Next morning, samples were washed three times with PBS and incubated with secondary antibodies for 1 h at room temperature. Cells were then washed with PBS, stained with DAPI for 5 min, and washed three times again with PBS. Slides were mounted with antifade mounting medium (Beyotime biotechnology, P0126). Fluorescence images were captured using a Nikon Eclipse Ti microscope, while confocal images were acquired with a Leica TCS SP8 STED or Leica STELLARIS 8 microscope. Image analysis was performed using LAS X software (v3.5.6) for Leica systems and ImageJ software (v1.48) for further processing and quantification. For quantification of fluorescence intensities of S. flexneri-associated SEPT9, bacteria were first detected by setting an intensity threshold in dedicated channels. Using bacteria as a mask, fluorescence intensities were extracted from corresponding regions in other channels. These intensities were then normalized to the total mean fluorescence intensity of individual cells.

Mass spectrometry analyses
ADP-ribosylome enrichment was performed as previously described16. Briefly, recombinant His6-eAf1521 was expressed, purified, and covalently coupled to NHS-resins. Clarified cell lysates (from ~2×107 293T cells overexpressing OspC) were incubated with NHS-resins coated with His6-eAf1521 at 4°C for 2 h with gentle rotation. The enrichment process was repeated twice, and pooled beads were washed four times with the TNT buffer. Captured proteins were eluted in 2× SDS loading buffer, separated by SDS-PAGE and stained with Coomassie blue for further MS analyses. The entire lane was excised into several fractions and subjected to in-gel digestion with sequencing-grade trypsin (Promega, USA) as described previously41. Protein bands of interest were reduced and alkylated using dithiothreitol and iodoacetamide, respectively. Protein samples were digested overnight with trypsin. Tryptic peptides were extracted and dried on a SpeedVac vacuum concentrator. The resulting peptides were dissolved in HPLC-grade water prior to LC-MS/MS analyses.

For label-free proteomics analyses, nanoflow reversed-phase liquid chromatography (EASY-nLC, Thermo Scientific) coupled with a hybrid ion trap-Orbitrap mass spectrometer (LTQ Orbitrap Velos, Thermo Scientific) was employed for peptide separation and detection. A homemade capillary column (75 μm×150 mm) with a laser-pulled electrospray tip (Model P-2000, Sutter Instruments) was packed with 5 μm, 120 Å Xtimate C18 silica-based particles (Welch Materials Inc.). The mobile phases consisted of solvent A (97% water, 3% acetonitrile, 0.1% formic acid) and solvent B (80% acetonitrile, 0.1% formic acid). The LC separation was performed at a flow rate of 300 nL/min with the following gradient: solvent B was increased from 5% to 40% over 40 min, then rapidly raised to 95% in 2 min and maintained for 10 min. Finally, the column was equilibrated with 100% solvent A. Eluted peptides were electrosprayed directly into the mass spectrometer for MS and MS/MS analyses in a data-dependent acquisition mode. One full MS scan (60,000 resolution, m/z 400–1200) was acquired and the 10 most intense ions were selected for fragmentation under collision-induced dissociation (CID) in the ion trap with the following parameters: ≥+2 precursor ion charge, 2 Da precursor ion isolation window, and 35 normalized collision energy. Dynamic exclusion was set with repeat duration of 30 s and exclusion duration of 12 s.

To determine modification sites, immunoprecipitated septins from 1×107 293T cells expressing OspC3 or its catalytically inactive mutant were separated by SDS-PAGE. Protein bands of interest were excised and subjected to in-gel digestion as described above. Resulting peptide samples were analyzed using a hybrid ion trap-Orbitrap mass spectrometer (LTQ-Orbitrap Velos, Thermo Fisher Scientific) in data-dependent acquisition mode. LC-MS parameters were similar to those described above, except that the m/z range of the MS scan was set to 400–1500. Raw MS files were searched against a septin database with Mascot (version 2.3.02, Matrix Science). Key search parameters included a mass tolerance of 20 ppm for precursor ions, 0.8 Da for MS/MS ions, a maximum of four missed cleavages, carbamidomethylation of Cys (+57.021 Da) as a static modification, and Met oxidation (+15.995 Da) and ADP-riboxanation of Arg (+524.03 Da) as dynamic modifications. Spectra of all potentially modified peptides were manually inspected, and their fragment ions were assigned for determination of modification sites. Corresponding peptide peaks were extracted using Thermo Xcalibur (Thermo Fisher Scientific, v2.2), and modification rates of ADP-riboxanated septins were calculated as percentage decrease of normalized peak areas for corresponding unmodified peptides.

In vitro ADP-riboxanation reactions
For in vitro reactions, 20 μg of purified GST-SEPT9 proteins were incubated with 1 μg of GST-OspC3 or its EH/AA mutant and 0.5 μg of His-Sumo-CaM in 40 μL of a reaction buffer containing 50 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM DTT, 2 mM MgCl2, and 1 mM NAD+. To assess the effect of calcium ions on the modification process, 5 mM CaCl2 or 5 mM EDTA (as a calcium chelator) was added to the reaction mixture. The reaction was carried out at 37°C for 60 min and terminated by boiling in the SDS loading buffer at 95°C for 10 min. The samples were analyzed by SDS-PAGE and subsequently subjected to either Western blot analyses or Coomassie blue staining. The stained gels were excised for further quantitative LC/MS analyses of ADP-riboxanated peptides derived from SEPT9.

Self-assembly of septin complexes and mass photometry analysis
Self-assembly of septin complexes was performed by diluting stored septin complexes from a high-salt buffer (50 mM Tris-HCl pH 8, 300 mM KCl, 5 mM MgCl₂, and 1 mM DTT) into a low-salt buffer (50 mM Tris-HCl pH 8, 50 mM KCl, 5 mM MgCl₂, and 1 mM DTT) with a final protein concentration of 400 μM. The complexes were incubated at room temperature (RT) for 4 h in the dark, and the reconstituted septin assemblies were imaged using a custom-built flow cell. Briefly, 2 μL of this dilution was placed on a clean glass slide and a coverslip was mounted on top with double-sided tape serving as a spacer. The samples were incubated for 20 min before fluorescence imaging. The images were processed and quantified using ImageJ software.

Mass photometry experiments were conducted using a TwoMP mass photometer (Refeyn Ltd, UK). Movies were recorded at 475.2 Hz with an exposure time of 2.06 ms, and optimized for camera counts to prevent saturation. The instrument was calibrated prior to measurements using BSA (66 kDa, 132 kDa) and thyroglobulin (660 kDa). Focus was achieved by adding a high-salt buffer (50 mM Tris-HCl pH 8.0, 300 mM KCl, 5 mM MgCl₂, and 1 mM DTT) into an 18 μL well, followed by identifying and locking the focal position with the instrument’s autofocus. For each acquisition, 2 μL of septin hetero-oligomer was added at a final concentration of 10 nM. All experiments were performed with at least three replicates. Data were analyzed using DiscoverMP software (v2023 R2).

Label-free protein quantification and bioinformatic analyses
For differential ADP-ribosylome analysis, spectral counting-based quantification was utilized due to its efficiency in identifying the most robust proteomic changes during the initial proteomics screen. Raw MS files were processed using Mascot Daemon (v2.3.02, Matrix Science). Tandem mass spectra were searched against a non-redundant human protein database (UP000005640, version 2021_04, containing 20,388 entries) downloaded from UniProt. Key search parameters included a mass tolerance of 20 ppm for precursor ions, 0.8 Da for MS/MS ions, and a maximum of two missed cleavages. Carbamidomethylation of cysteine residues (+57.021 Da) was set as a static modification, while methionine oxidation (+15.995 Da) was chosen as a variable modification. False discovery rates (FDRs) of peptides and proteins were set at below 1%. Downstream bioinformatic analyses were performed in R (R Foundation for Statistical Computing) using RStudio (an integrated development environment for R). At least two valid quantification values in any group were required, and missing values were imputed with a constant low value to enable calculation of fold changes.

For SEPT9 IP-MS experiments, intensity-based label-free quantification was employed due to its higher accuracy and sensitivity in detecting modest differences. Raw data were analyzed using MaxQuant (v1.5.5.1)42 with default settings. The ‘match between runs’ and label-free quantification (LFQ) features were enabled. The resulting data table was uploaded to the LFQ-Analyst website43 for statistical analysis. Protein-protein interactions (PPI) were analyzed using Metascape44 and densely connected network components were identified with Molecular Complex Detection (MCODE) algorithm45. The resulting network was visualized using Cytoscape (v3.10.3). A heatmap was generated using the pheatmap R package (v1.0.12). Gene Ontology (GO) term enrichment was conducted using the clusterProfiler R package (v4.14.4), and GO terms with adjusted P-values < 0.05 were considered significantly enriched. Bar plots and scatter plots were created using the ggplot2 R package (v3.5.1). A Venn diagram was generated using the VennDiagram R package (v1.7.3).

Statistics and reproducibility
All samples were included from the analysis without randomization or blinding procedures. No statistical methods were used to pre-determine sample size. Experiments were independently repeated at least three times to achieve statistical significance. Data were presented as mean ± SEM, and statistical significance was determined by the unpaired two-tailed Student’s t test, or the ANOVA test followed by Tukey’s post hoc test or Holm-Sidak post hoc test as indicated in figure legends. Representative results were shown. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. All statistical analyses were performed using GraphPad Prism 9.4 software.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this work, Tang et al. employed multiple strategies—including western blotting, mass spectrometry, and microscopy—to\ndemonstrate that OspC effectors enable S. flexneri to evade septin cage entrapment. Mechanistically, they show that OspC\ncatalyzes ADP-riboxanation of SEPT9 at Arg561, a critical residue essential for stabilizing septin hetero-oligomers.\nThe writing style and scope align well with the aims of Nature Communications. Therefore, the journal should consider this\nmanuscript for publication. I have some minor to moderate comments to help further improve the manuscript.\n1. Data Availability: The authors should provide a link to access the raw mass spectrometry (MS) data,for example,\nProteomeXchange.\n2. Modified Site Assignment: For the ADP-riboxanation sites identified via MS/MS, did the authors manually inspect the\nspectra and assign fragment ions for all potential modified peptides? Please clearly describe the procedure for modified site\nassignment in the Methods section.\n3. The rational of using both spectral counting and MaxLFQ (maxquant) for label-free quantification needs to be discussed.\nUsually, intensity-based label-free quantification will be more accurate than the spectral counting method.\n4. For Table 1, how is the modification rate calculated? Some brief information here will be helpful. Because the modification\nwill impact the ionization efficiency of the peptides strongly, it will be hard to calculate the modification rates just based on\nthe peptide intensity information.\n5. Residue Specificity: ADP-ribosylation can also occur on glutamic acid residues. For the peptide sequence shown in\nFigure 2C, is it possible that the modification occurs on glutamic acid rather than arginine? Please clarify.\n6. Experimental Details: a) For each experiment—such as immunoprecipitation (IP) or MS analysis—the authors should\nspecify the number of cells used.\nb) The flow rate used in the nanoLC-MS analysis should be provided.\nc) The authors should specify the number of biological and technical replicates used for the\nMS quantification analysis.\nd) For label-free data analysis, please specify the download date and the number of protein entries in the FASTA file used.", "reviewer_2": "The study by Tang et al. explores how the bacterial pathogen Shigella flexneri evades host defense mechanisms by\ntargeting septins, a family of proteins that form cage-like structures to entrap cytosolic bacteria. The key findings are as\nfollows: S. flexneri uses its Type Three Secretion System (T3SS) effector proteins from the OspC family to modify septins via\nADP-riboxanation, a form of post-translational modification closely related to ADP-ribosylation. SEPT9 is identified as a\nprimary target among septin proteins. OspC effectors target SEPT9 and modify it at Arginine 561 (Arg561), disrupting the\nNC-interface between adjacent SEPT9 subunits. This disruption leads to disassembly of septin octamers, thereby inhibiting\nseptin cage formation.\nAdditionally, both OspC and another effector protein, OspG, act in concert to inhibit septin cage assembly. Mutant strains\nlacking these effectors show increased septin cage entrapment of bacteria, indicating a cooperative role in evading host\ndefense. This study reveals that S. flexneri has evolved two distinct mechanisms to counteract septin-mediated cell-\nautonomous immunity.\nThis study offers novel insights into the mechanisms by which S. flexneri evades host cell defenses through targeted\nmodifications of septins. Overall, the experimental rigor of the study is commendable. This robustness is not surprising given\nthat two labs with a successful track record in studying host-pathogen interactions—specifically, shigella cage entrapment\nby septins and effector-based modification of cellular targets—collaborated on this research.\nThe experiments presented are well-designed and executed to a high standard. The experimental setup largely builds on\nthe authors' previous work on eIF3 and caspase modifications.\nThe relevance of the study lies in the fact that it identifies a direct modification of septins by a bacterial toxin or effector (“an\noverdue finding”). OspCs are phylogenetically and functionally conserved across multiple bacterial pathogens, which further\nunderscores the significance of this finding.\nHowever, the authors face an inherent challenge: they are investigating a highly complex system involving multiple effectors,\nsome of which are partially redundant or dispensable for the biological outcome—Shigella infection. Furthermore, they\nreport promiscuous substrate activity of their effector, adding another layer of complexity: OspCs modify eIF3, caspase-11\nand now SEPT9. A previously identified additional effector OspG targets septins indirectly by ubiquitination via CAND1\nphosphorylation .\nIt is quite common for two or more effectors from the same bacterial species to target the same host protein or pathway\n(functional redundancy or effector convergence). Likewise, it is also possible for individual effectors to target multiple\nsubstrates. However, these substrates are often functionally and/or structurally related.\nThus, further investigation into the biological significance of the septin modification by the authors would be beneficial.\nMajor points:\n#1\nLine 143: “SEPT9 was post-translationally modified in a manner dependent on both the enzymatic activity and ARD domain\nof OspC3 (Fig. S2A and S2B).”\nThis interpretation might be misleading and requires further clarification. It suggests that both, the enzymatic N-terminal\ndomain and the ARD (C-terminal domain) are required for the modification, which is not fully supported by the data. In\nFigures S2A and S2B (Flag-IP, ADPr), a band is still visible for the N-terminal domain (enzymatic domain) alone—albeit\nstrongly reduced compared to the full-length protein. Taken together with the results in Figure S2F, this indicates that the\npost-translational modification of SEPT9 depends on its catalytic N-terminal domain. However, the modification is strongly\nenhanced by septin-binding mediated by the C-terminal ARD.\nPlease clarify.\n#2 Fig. The results from Fig. 2D+E clearly indicate that OspCs exhibit a high level of promiscuity concerning modified\nresidues and substrate specificity. This is a significant problem of the study. Therefore, all findings must be interpreted with\ngreat caution. In the majority of their experiments, the authors overexpress the effector OspC along with the target or\nsubstrate protein under investigation using vectors with strong promoters ensuring high expression levels. However, high\nconcentrations of OspC can lead to reduced specificity. It is important to note that physiologically, the number of effector\nmolecules delivered by a T3SS is very limited. High septin concentrations may mask the modification of other preferred\n(potentially more physiological) substrates due to the excessive supply.\nExperiments employing bacterial infection and endogenous proteins are therefore eligible.\nIn Fig. 2H+I+J, the authors perform cell infections (still with Flag-SEPT9 overexpression).\n#2.1 In Fig. 2H, the ADP-ribosylation of SEPT9 without infection is an interesting finding. Why does this ADP-ribosylation\nalmost disappear in Fig. 2I and 2J? Please clarify.\n#2.2 Flag-tagged SEPT9 always runs between 72 and 100 kDa, but in Fig. 2H, it appears closer to 100 kDa. Please clarify.\n#2.3 As mentioned above, infection experiments with endogenous protein levels would significantly enhance the\nsignificance of the observations.\n-The authors should make efforts to infect cells without overexpression of septins, perform\nimmunoprecipitation (IP) of endogenous septins, and probe for the studied modification.\n-A classical yet informative assay could be employed to determine the extent of endogenous SEPT9 modification during\ninfection. Specifically, lysates from infected cells—either following immunoprecipitation of SEPT9 or directly from whole-cell\nextracts—could be subjected to an in vitro post ADP-ribosylation reaction using recombinant OspC and labeled NAD⁺ (e.g.,\nbiotinylated or radiolabeled). If SEPT9 is already efficiently ADP-ribosylated during infection with wild-type Shigella,\nminimal to no additional labeling would be detected. In contrast, infection with a catalytic-dead Shigella or KO mutant strain would\nleave SEPT9 unmodified, resulting in a strong signal upon in vitro labeling.\nThis approach could provide insights into the efficiency of SEPT9 modification in vivo and could help estimate substrate\npreferences of OspC under near-physiological conditions. Moreover, when using total lysates, other OspC substrates will\nalso be labeled, enabling comparison of modification levels and preferences between SEPT9 and alternative targets.\nWhile the presence of multiple ADP-ribosylation sites on OspC substrates might complicate signal interpretation, this assay\nremains valuable for evaluating the physiological relevance of OspC substrate specificity and activity during infection.\n#3 – Fig. 4D and Fig. S8D (Fluorescence Imaging Quality):\nThe fluorescence images presented in Fig. 4D (and S8D) are currently not informative to the reader. In particular,\nbacteria are difficult to detect in the DAPI channel due to the weak staining color and overlap with host nuclei. Given the authors’\nimaging capabilities, it is recommended to acquire 4-channel fluorescence images. Labeling bacteria with a distinct\nfluorophore (e.g., GFP or a green dye), SEPT9 in red, and Flag in far-red (as done in Fig. 4E) would significantly enhance\nclarity and interpretability. This approach would allow clear spatial distinction.\n#4 -Fig. 4D and Fig. S8C (Individual Line Scans)\nThe individual line scans shown in Fig. 4D and Fig. S8C exhibit relatively noisy profiles and offer limited informational value\nto the reader. Since single line scans are inherently qualitative and not suited for statistical evaluation, they do not robustly\nsupport conclusions about fluorescence intensity differences between strains or conditions.\nA more informative and quantitative approach would involve using intracellular bacteria as a mask to extract fluorescence\nintensities from the corresponding regions in other channels (e.g., SEPT9 and Flag). This would allow for the rapid\nquantification of signal intensities across hundreds of bacteria, enabling meaningful statistical comparisons.\nImplementing such an analysis would significantly enhance the quantitative rigor and clarity of the presented data.\n#5 – Fig. 4E (Image Selection and Magnification):\nThe magnified region in Fig. 4E is not optimally chosen. The selected area (EH/AA) includes a large oversaturated region\nthat provides limited information about individual bacteria. Other fields of view within the same dataset appear to better\nillustrate the phenotype, similar to vector control conditions, where single, clearly caged bacteria are visible. A more\nrepresentative and well-balanced zoomed area would substantially improve the figure’s impact.\n#6 – Fig. 5B (Quantitative Validation of Phenotype):\nFig. 5B effectively demonstrates the individual and combined roles of OspC and OspG in bacterial caging. However, to\nprovide a more complete picture of their contribution to Shigella pathogenesis, a complementary CFU assay using the same\nbacterial strains would be highly beneficial. A side-by-side comparison of intracellular survival or replication across the\ndifferent effectors would clarify the impact of both effectors on overall infection.\n#7 – Disruption of the Septin Octamer\nThe authors propose that OspC effector proteins destabilize SEPT9-containing septin octamers via ADP-ribosylation,\nleading to a shift toward tetramers. This hypothesis is supported by co-expression experiments in E. coli (Fig. 4B) and also\noverexpression experiments in 293T cells (Fig. 3E) look promising.\nHowever, there is a possibility to test the proposed model in the context of Shigella infection. The Mavrakis lab has\ndeveloped a toolbox (available via Addgene) that enables the investigation of specific septin–septin interactions (Martins et\nal. 2023). Using the splitGFP-SEPT9 construct, the GFP signal is expected to be lost during infection with wild-type Shigella\ndue to OspC activity, whereas the signal should be maintained in infections with a catalytic-dead OspC or KO mutant, where\nno modification occurs.\nMinor point:\nLine 127: “catalytic mutant”. I prefer the terms 'catalytically dead/inactive' or 'enzymatically inactive', as they more clearly\nindicate the loss of enzymatic function. That said, this remains a matter of personal preference.\nSummary:\nThe authors present a highly interesting and timely study with an overall solid methodological framework. This reviewer\nconsiders the findings to be a long-awaited contribution to both the host–pathogen interaction and septin biology fields —\nspecifically, the demonstration of direct septin modification by a bacterial effector or toxin. With the addition of some further\nexperimental evidence supporting the specificity and function of this modification in a more physiological context, the\nmanuscript will be suitable for publication.\nRecommendation: Major Revision", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_4": "Bacteria can be trapped in cage-like structures containing septin, key components of the cytoskeleton. These septin cages represents an important aspect of cell-autonomous immunity. In this manuscript the authors examine how a facultative intracellular bacterial pathogen, Shigella flexneri, can evade septin cages during infection. Building on prior work, they show that type 3 secreted proteins of the OspC family can suppress septin cages through a chemical modification (ADP-riboxanation) of septin 9 at a critical amino acid required for cage assembly. Their findings provide a new mechanism for evasion of septin cages by a bacterial pathogen. Furthermore, their findings show how OspC acts cooperatively with OspG to evade septin cages, thus underlining the importance of this cell autonomous immune defense. The paper is well written and the data and interpretations quite clear and convincing. I am in support of publication and have only minor comments for the authors consideration.\nMinor comments:\n-line 26 and line45, use either type 3 secretion or type III secretion consistently throughout\n-line 257 “Since ADP-riboxanation of arginine basically converts it to a negatively charged residue…”. This is not really\naccurate and could be worded better.\n-line 306, “a double-knockout mutant (dospCdospG). Notably, septin cage formation was increased to an even higher level\n(31%). This is an exciting finding . The authors should comment on whether the more prevalent septin cages appear the\nsame (morphologically, position within the cell, etc), or perhaps distinct, from cages observed in WT infected cells or\nsingle mutant infected cells.\n-is bacterial intracellular growth, or cell-to-cell spread, altered in the double-knockout mutant (dospCdospG)?\n-Figure 5A, D. It is very difficult to see bacteria within septin cages. The use of insets with higher magnification would likely\nhelp.\n-Figure S7B. Where is the 10RK mutant?"}, "author_response": "We thank the editor and reviewers for their constructive feedback and positive comments.\nWe are grateful for their overall interest in our study. As suggested by reviewers, we carried\nout further investigations and made substantial improvements to significantly strengthen our\nmanuscript message. We updated all figure panel labels to lowercase to comply with journal\npublication guidelines. Detailed point-to-point responses are provided below and all edits to\nthe manuscript are highlighted in yellow for ease of reference.\nREVIEWER COMMENTS\nReviewer #1:\nIn this work, Tang et al. employed multiple strategies—including western blotting,\nmass spectrometry, and microscopy—to demonstrate that OspC effectors enable S. flexneri to\nevade septin cage entrapment. Mechanistically, they show that OspC catalyzes\nADP-riboxanation of SEPT9 at Arg561, a critical residue essential for stabilizing septin\nhetero-oligomers.\nThe writing style and scope align well with the aims of Nature Communications. Therefore,\nthe journal should consider this manuscript for publication. I have some minor to moderate\ncomments to help further improve the manuscript.\nResponse: We thank the reviewer for their positive feedback and support for publication.\n1. Data Availability: The authors should provide a link to access the raw mass spectrometry\n(MS) data, for example, ProteomeXchange.\nResponse: We deposited raw MS data to ProteomeXchange and included a “Data Availability”\nsection, which provides details of our proteomic datasets. The processed data (mostly in\nExcel format) have also been included as supplemental materials (see Supplemental Data\n1-2).\n2. Modified Site Assignment: For the ADP-riboxanation sites identified via MS/MS, did the\nauthors manually inspect the spectra and assign fragment ions for all potential modified\npeptides? Please clearly describe the procedure for modified site assignment in the Methods\nsection.\nResponse: We manually inspected spectra for all modified peptides. To ensure clarity and\ntransparency, we provided a detailed description of the procedure in the Methods section\n(see lines 604-619).\n3. The rational of using both spectral counting and MaxLFQ (maxquant) for label-free\nquantification needs to be discussed. Usually, intensity-based label-free quantification will be\nmore accurate than the spectral counting method.\nspectral counting. While the intensity-based approach is superior for detecting subtle\ndifferences, spectral counting is efficient for identifying the most robust proteomic changes.\nTherefore, for the initial proteomic screen to identify potentially modified targets (Fig. 1A), we\nused spectral counting for rapid assessment to tease out the most significant substrates.\nWhen these hits undergo further testing, the likelihood of not dropping out is usually higher.\nIn contrast, when we evaluated impact of ADP-riboxanation on interaction strength between\ndifferent septin subunits, intensity-based quantification was used because differences were\nrelatively modest (by approximately 50%, see Fig. 3E). As suggested, we clarified our\nrationale in the revised manuscript (see lines 650-652 and 664-665).\n4. For Table 1, how is the modification rate calculated? Some brief information here will be\nhelpful. Because the modification will impact the ionization efficiency of the peptides strongly,\nit will be hard to calculate the modification rates just based on the peptide intensity\ninformation.\nResponse: We agree that modifications often change ionization efficiency of peptides.\nTherefore, intensity readouts of modified and unmodified peptides cannot be compared\ndirectly (i.e., they are somewhat analogous to different currencies). To circumvent this\nproblem, modification rate was calculated as percentage decrease of peptide intensities for\nunmodified peptides only. We have two experimental conditions (i.e., septins co-expressed\nwith either wild-type OspC or catalytically inactive OspC, see the figure below) and compare\ndifferences of unmodified signals (denoted by black chromatographic peaks) between these\ntwo cases. For clarity we included description of this calculation process in the Methods\nsection (lines 616-619).\n5. Residue Specificity: ADP-ribosylation can also occur on glutamic acid residues. For the\npeptide sequence shown in Figure 2C, is it possible that the modification occurs on glutamic\nacid rather than arginine? Please clarify.\nbroader range of residues, including glutamic acid. However, ADP-riboxanation is distinct\nfrom canonical ADP-ribosylation in terms of its modification chemistry. As demonstrated in\nour previous study (Li et al., Nature 2021), this modification occurs specifically on arginine\nresidues. This specificity is derived from additional deamination of arginine’s guanidinium\ngroup, thus yielding an oxazolidine ring structure in modified substrates. Glutamic acids, in\nprinciple, are not able to undergo such a unique chemical reaction. Additionally, mutational\nstudies with substitution of modified Arg561 to either Ala or Lys further support the proposed\nmodification site (new Fig. 2i). We clarified this with text updates in our revised manuscript\n(lines 611-614).\n6. Experimental Details: a) For each experiment—such as immunoprecipitation (IP) or MS\nanalysis—the authors should specify the number of cells used.\nb) The flow rate used in the nanoLC-MS analysis should be provided.\nc) The authors should specify the number of biological and technical replicates used for the\nMS quantification analysis.\nd) For label-free data analysis, please specify the download date and the number of protein\nentries in the FASTA file used.\nResponse: These experimental details have been included in the revised manuscript.\na) 1-2 × 107 293T cells were used for IP (line 445) and MS analysis (line 577), respectively.\nb) The flow rate of nanoLC-MS was set at 300 nL/min (line 594).\nc) Two and four biological replicates were used for ADP-ribosylome (Fig. 1c and line 825) and\ninteractome (Fig. 4a and line 881) analyses, respectively.\nd) The FASTA file was downloaded in April 2021, containing 20,388 entries with the identifier\nUP000005640 (line 654).\nReviewer #2 (Remarks to the Author):\nThe study by Tang et al. explores how the bacterial pathogen Shigella flexneri evades host\ndefense mechanisms by targeting septins, a family of proteins that form cage-like structures\nto entrap cytosolic bacteria. The key findings are as follows: S. flexneri uses its Type Three\nSecretion System (T3SS) effector proteins from the OspC family to modify septins via\nADP-riboxanation, a form of post-translational modification closely related to\nADP-ribosylation. SEPT9 is identified as a primary target among septin proteins. OspC\neffectors target SEPT9 and modify it at Arginine 561 (Arg561), disrupting the NC-interface\nbetween adjacent SEPT9 subunits. This disruption leads to disassembly of septin octamers,\nthereby inhibiting septin cage formation. Additionally, both OspC and another effector protein,\nOspG, act in concert to inhibit septin cage assembly. Mutant strains lacking these effectors\nshow increased septin cage entrapment of bacteria, indicating a cooperative role in evading\nhost defense. This study reveals that S. flexneri has evolved two distinct mechanisms to\ncounteract septin-mediated cell-autonomous immunity.\nThis study offers novel insights into the mechanisms by which S. flexneri evades host cell\n3\ndefenses through targeted modifications of septins. Overall, the experimental rigor of the\nstudy is commendable. This robustness is not surprising given that two labs with a successful\ntrack record in studying host-pathogen interactions—specifically, shigella cage entrapment by\nseptins and effector-based modification of cellular targets—collaborated on this research.\nThe experiments presented are well-designed and executed to a high standard.\nThe experimental setup largely builds on the authors' previous work on eIF3 and caspase\nmodifications. The relevance of the study lies in the fact that it identifies a direct modification\nof septins by a bacterial toxin or effector (“an overdue finding”). OspCs are phylogenetically\nand functionally conserved across multiple bacterial pathogens, which further underscores\nthe significance of this finding.\nResponse: We thank the reviewer for their highly positive comments on our work, in terms of\nboth scientific significance and quality. It is encouraging to read that experts in the field\nconsider this work (i.e., direct septin modification by bacterial proteins) as an “overdue”\nfinding.\nHowever, the authors face an inherent challenge: they are investigating a highly complex\nsystem involving multiple effectors, some of which are partially redundant or dispensable for\nthe biological outcome—Shigella infection. Furthermore, they report promiscuous substrate\nactivity of their effector, adding another layer of complexity: OspCs modify eIF3, caspase-11\nand now SEPT9. A previously identified additional effector OspG targets septins indirectly by\nubiquitination via CAND1 phosphorylation.\nIt is quite common for two or more effectors from the same bacterial species to target the\nsame host protein or pathway (functional redundancy or effector convergence). Likewise, it is\nalso possible for individual effectors to target multiple substrates. However, these substrates\nare often functionally and/or structurally related.\nThus, further investigation into the biological significance of the septin modification by the authors would be beneficial.\nResponse: We acknowledge the complexity of bacterial infection involving multiple effectors,\nsuch as functional redundancy and substrate promiscuity. In our study we clearly show that\nOspCs and OspG work together to counteract septin cage entrapment via two distinct\nmodifications, thereby facilitating bacterial cell-to-cell spread and replication (see results of\nour newly conducted experiments in Fig. 5e-i).\nMajor points:\n#1 Line 143: “SEPT9 was post-translationally modified in a manner dependent on both the\nenzymatic activity and ARD domain of OspC3 (Fig. S2A and S2B).”\nThis interpretation might be misleading and requires further clarification. It suggests that both,\nthe enzymatic N-terminal domain and the ARD (C-terminal domain) are required for the\nmodification, which is not fully supported by the data. In Figures S2A and S2B (Flag-IP, ADPr),\na band is still visible for the N-terminal domain (enzymatic domain) alone—albeit strongly\nreduced compared to the full-length protein. Taken together with the results in Figure S2F,\nthis indicates that the post-translational modification of SEPT9 depends on its catalytic\nN-terminal domain. However, the modification is strongly enhanced by septin-binding\nmediated by the C-terminal ARD.\nPlease clarify.\nResponse: We agree with the reviewer’s interpretation of our data. Indeed the modification\ndepends on the catalytic domain and can be strongly enhanced by ARD domain-mediated\ninteraction. We revised the original claim in the text accordingly (see lines 143-145 and\n149-151).\n#2 Fig. The results from Fig. 2D+E clearly indicate that OspCs exhibit a high level of\npromiscuity concerning modified residues and substrate specificity. This is a significant\nproblem of the study. Therefore, all findings must be interpreted with great caution. In the\nmajority of their experiments, the authors overexpress the effector OspC along with the target\nor substrate protein under investigation using vectors with strong promoters ensuring high\nexpression levels. However, high concentrations of OspC can lead to reduced specificity. It is\nimportant to note that physiologically, the number of effector molecules delivered by a T3SS\nis very limited. High septin concentrations may mask the modification of other preferred\n(potentially more physiological) substrates due to the excessive supply. Experiments\nemploying bacterial infection and endogenous proteins are therefore eligible. In Fig. 2H+I+J,\nthe authors perform cell infections (still with Flag-SEPT9 overexpression).\nResponse: We agree that overexpression of either effectors or substrates can lead to\nmodification results that are sometimes unlikely to occur in physiologically relevant conditions\n(e.g., bacterial infection). As suggested, we repeated infection-based experiments and\nexamine modification status of endogenous septins (at least for SEPT9). Below, we address\neach of the reviewer’s specific comments.\n#2.1 In Fig. 2H, the ADP-ribosylation of SEPT9 without infection is an interesting finding. Why\ndoes this ADP-ribosylation almost disappear in Fig. 2I and 2J? Please clarify.\nResponse: We acknowledge the observed discrepancy in levels of endogenously\nADP-ribosylated SEPT9 between Fig. 2H and Fig. 2I/J. To clarify this, we repeated\nexperiments under identical conditions and ensure reproducibility with multiple replicates\n(new Fig. 2i). Previous differences may result from repeated use of α-ADPr antibody,\nleading to reduced sensitivity for detecting weak endogenous modifications of SEPT9.\n#2.2 Flag-tagged SEPT9 always runs between 72 and 100 kDa, but in Fig. 2H, it appears\ncloser to 100 kDa. Please clarify.\nResponse: From the original uncropped blot the gel ran unevenly, thereby making SEPT9\nappear higher than expected (see Rebuttal Figure 1).\n#2.3 As mentioned above, infection experiments with endogenous protein levels would\nsignificantly enhance the significance of the observations.\n-The authors should make efforts to infect cells without overexpression of septins, perform\nimmunoprecipitation (IP) of endogenous septins, and probe for the studied modification.\nResponse: We immunoprecipitated endogenous SEPT9 and probed with α-ADPr antibody\n(new Fig. 2j). Immunoblotting data revealed modification of endogenous SEPT9 during\nShigella infection in an OspC-dependent manner, and this modification was confirmed by\nmass spectrometry analysis (new Fig. S6e).\n-A classical yet informative assay could be employed to determine the extent of endogenous\nSEPT9 modification during infection. Specifically, lysates from infected cells—either following\nimmunoprecipitation of SEPT9 or directly from whole-cell extracts—could be subjected to an\nin vitro post ADP-ribosylation reaction using recombinant OspC and labeled NAD (e.g.,\nbiotinylated or radiolabeled). If SEPT9 is already efficiently ADP-ribosylated during infection\n⁺\nwith wild-type Shigella, minimal to no additional labeling would be detected. In contrast,\ninfection with a catalytic-dead Shigella or KO mutant strain would leave SEPT9 unmodified,\nresulting in a strong signal upon in vitro labeling.\nThis approach could provide insights into the efficiency of SEPT9 modification in vivo and\ncould help estimate substrate preferences of OspC under near-physiological conditions.\nMoreover, when using total lysates, other OspC substrates will also be labeled, enabling\ncomparison of modification levels and preferences between SEPT9 and alternative targets.\nWhile the presence of multiple ADP-ribosylation sites on OspC substrates might complicate\nsignal interpretation, this assay remains valuable for evaluating the physiological relevance of\nOspC substrate specificity and activity during infection.\nprinciple, can be informative in determining extent of modifications. The concept of analyzing\nthe proportion of unmodified septins during infection as an indirect readout of modification\nrate is similar to the approach we described in response to reviewer #1 (see point #4).\nHowever, the success of such assays relies on a sizeable fraction of substrates (i.e., septins)\nbeing modified prior to in vitro reactions. In reality, we learned from years of experience that\nrates of effector-catalyzed modifications during infection are generally very low (e.g., <5% of\ntotal protein pools). Such low rates are largely due to the very limited number of effector\nmolecules delivered by the T3SS, a point raised by the reviewer (see point #2). Considering\nthis, we would need to distinguish 95% unmodified from 100% unmodified by in vitro NAD+\nlabeling and immunoblotting assays, which is far beyond quantification accuracy of Western\nblotting.\nAnother complication may arise from the fact that some modifications (i.e., SEPT9 at R561)\nmay require presence of septin complexes or higher-order assemblies, which are likely\ndisrupted during cell lysis. In support of this, R561 modification was not detected in\nprokaryotic co-expression systems or in vitro ADP-ribosylation reactions (see Fig. S6A and\nS6B).\nAs an alternative to in vitro labeling experiments, we directly measured the percentage of\nunmodified septins under infection conditions by mass spectrometry (see response to point\n#4 from reviewer #1 for methodological details, and refer to newly added Fig. 2j, Fig. S6e,\nand Table 2 for corresponding data). In general, quantification accuracy of mass spectrometry\nis much higher than that of immunoblotting assays. Mass spectrometry analysis revealed that\nmodification rates were approximately 4% in cells infected by wild type Shigella and 12% with\nthe complementation strain (ΔospC+pOspC1/C3).\n#3 – Fig. 4D and Fig. S8D (Fluorescence Imaging Quality):\nThe fluorescence images presented in Fig. 4D (and S8D) are currently not informative to the\nreader. In particular, bacteria are difficult to detect in the DAPI channel due to the weak\nstaining color and overlap with host nuclei. Given the authors’ imaging capabilities, it is\nrecommended to acquire 4-channel fluorescence images. Labeling bacteria with a distinct\nfluorophore (e.g., GFP or a green dye), SEPT9 in red, and Flag in far-red (as done in Fig. 4E)\nwould significantly enhance clarity and interpretability. This approach would allow clear\nspatial distinction.\nResponse: We repeated experiments using GFP-expressing bacteria (new Fig. 4d and S8f),\nwhich indeed substantially improved clarity and interpretability of our data.\n#4 -Fig. 4D and Fig. S8C (Individual Line Scans)\nThe individual line scans shown in Fig. 4D and Fig. S8C exhibit relatively noisy profiles and\noffer limited informational value to the reader. Since single line scans are inherently\nqualitative and not suited for statistical evaluation, they do not robustly support conclusions\n7\nabout fluorescence intensity differences between strains or conditions.\nA more informative and quantitative approach would involve using intracellular bacteria as a\nmask to extract fluorescence intensities from the corresponding regions in other channels\n(e.g., SEPT9 and Flag). This would allow for the rapid quantification of signal intensities\nacross hundreds of bacteria, enabling meaningful statistical comparisons. Implementing such\nan analysis would significantly enhance the quantitative rigor and clarity of the presented data.\nResponse: We appreciate the suggestion. By using bacteria as a mask, extracted\nfluorescence intensities were further normalized to the total mean fluorescence intensity of\nthe respective individual cells. Statistical analysis revealed that both R561 and the\nNC-interface are required for FLAG-SEPT9 recruitment to Shigella (new Fig. 4d–e and S8c–\nd). In comparison, fluorescence intensities of total Shigella-associated SEPT9 (including both\ntransfected and endogenous proteins) were not significantly different among the same set of\ntransfected samples (new Fig. 4f and S8e). We updated text in the Methods (line 569-573)\nand Results section (line 288-298) accordingly.\nDue to overwhelming signal of host nuclei in the DAPI channel (as mentioned in point #3), we\nreplaced DAPI with GFP-labeled bacteria to improve visualization (new Fig. 4d and S8c).\n#5 – Fig. 4E (Image Selection and Magnification):\nThe magnified region in Fig. 4E is not optimally chosen. The selected area (EH/AA) includes\na large oversaturated region that provides limited information about individual bacteria. Other\nfields of view within the same dataset appear to better illustrate the phenotype, similar to\nvector control conditions, where single, clearly caged bacteria are visible. A more\nrepresentative and well-balanced zoomed area would substantially improve the figure’s\nimpact.\nResponse: We replaced this magnified region with a more representative and well-balanced\nzoomed area to support our conclusions (new Fig. 4g).\n#6 – Fig. 5B (Quantitative Validation of Phenotype):\nFig. 5B effectively demonstrates the individual and combined roles of OspC and OspG in\nbacterial caging. However, to provide a more complete picture of their contribution to Shigella\npathogenesis, a complementary CFU assay using the same bacterial strains would be highly\nbeneficial. A side-by-side comparison of intracellular survival or replication across the\ndifferent effectors would clarify the impact of both effectors on overall infection.\nResponse: We conducted CFU assays as suggested. Though single deletion mutants did not\nshow any growth defects intracellularly, we did observe reduced replication of the double\nmutant (see lines 331-334 and new Fig. 5g–i). Given that septin cages restrict intracellular\nmobility and cell-to-cell spread of cytosolic bacteria, we also carried out a plaque formation\nassay (see comments of reviewer #4). These new results revealed collaborative contribution\nof these two effectors to Shigella pathogenesis (new Fig. 5e–i, lines 327-338).\n8\n#7 – Disruption of the Septin Octamer\nThe authors propose that OspC effector proteins destabilize SEPT9-containing septin\noctamers via ADP-ribosylation, leading to a shift toward tetramers. This hypothesis is\nsupported by co-expression experiments in E. coli (Fig. 4B) and also overexpression\nexperiments in 293T cells (Fig. 3E) look promising.\nHowever, there is a possibility to test the proposed model in the context of Shigella infection.\nThe Mavrakis lab has developed a toolbox (available via Addgene) that enables the\ninvestigation of specific septin–septin interactions (Martins et al. 2023). Using the\nsplitGFP-SEPT9 construct, the GFP signal is expected to be lost during infection with\nwild-type Shigella due to OspC activity, whereas the signal should be maintained in infections\nwith a catalytic-dead OspC or KO mutant, where no modification occurs.\nResponse: We agree that direct visualization of SEPT9–SEPT9 interactions during infection\nwith the splitGFP-SEPT9 construct would further validate our hypothesis within a\nphysiologically relevant context. Nonetheless, we need to consider that minimal amounts of\neffectors are delivered by T3SSs during infection (see point #2). Together with modification\nrate of septins as low as <5% during Shigella infection, it is very likely that the suggested\nassay will not be sensitive enough to detect impact of OspC on septin-septin interactions.\nMinor point:\nLine 127: “catalytic mutant”. I prefer the terms 'catalytically dead/inactive' or 'enzymatically\ninactive', as they more clearly indicate the loss of enzymatic function. That said, this remains\na matter of personal preference.\nResponse: We revised the terms to 'catalytically dead/inactive' throughout the manuscript.\nSummary:\nThe authors present a highly interesting and timely study with an overall solid methodological\nframework. This reviewer considers the findings to be a long-awaited contribution to both the\nhost–pathogen interaction and septin biology fields — specifically, the demonstration of direct\nseptin modification by a bacterial effector or toxin. With the addition of some further\nexperimental evidence supporting the specificity and function of this modification in a more\nphysiological context, the manuscript will be suitable for publication.\nRecommendation: Major Revision\nResponse: We thank the reviewer for their positive evaluation and suggestions to strengthen\nour manuscript message.\nReviewer #3 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This\n9\nis part of the Nature Communications initiative to facilitate training in peer review and to\nprovide appropriate recognition for Early Career Researchers who co-review manuscripts.\nResponse: Thank you, we are grateful for ECRs to evaluate our work.\nReviewer #4 (Remarks to the Author):\nBacteria can be trapped in cage-like structures containing septin, key components of the\ncytoskeleton. These septin cages represents an important aspect of cell-autonomous\nimmunity. In this manuscript the authors examine how a facultative intracellular bacterial\npathogen, Shigella flexneri, can evade septin cages during infection. Building on prior work,\nthey show that type 3 secreted proteins of the OspC family can suppress septin cages\nthrough a chemical modification (ADP-riboxanation) of septin 9 at a critical amino acid\nrequired for cage assembly. Their findings provide a new mechanism for evasion of septin\ncages by a bacterial pathogen. Furthermore, their findings show how OspC acts\ncooperatively with OspG to evade septin cages, thus underlining the importance of this cell\nautonomous immune defense. The paper is well written and the data and interpretations quite\nclear and convincing. I am in support of publication and have only minor comments for the\nauthors consideration.\nResponse: We thank the reviewer for their enthusiasm.\nMinor comments:\n-line 26 and line45, use either type 3 secretion or type III secretion consistently throughout\nResponse: We revised text to type III secretion and ensure consistency throughout the\nmanuscript.\n-line 257 “Since ADP-riboxanation of arginine basically converts it to a negatively charged\nresidue…”. This is not really accurate and could be worded better.\nResponse: We modified text for improved clarity (lines 267-269).\n-line 306, “a double-knockout mutant (dospCdospG). Notably, septin cage formation was\nincreased to an even higher level (31%). This is an exciting finding. The authors should\ncomment on whether the more prevalent septin cages appear the same (morphologically,\nposition within the cell, etc), or perhaps distinct, from cages observed in WT infected cells or\nsingle mutant infected cells.\nResponse: We agree that a bacterial strain triggering >30% septin cage entrapment can help\nto transform the community. We did not observe significant morphological or positional\ndifferences in septin cages under experimental conditions we tested. To address this in depth\nfuture studies will be required by employing super-resolution and/or high-content microscopy.\n10\n-is bacterial intracellular growth, or cell-to-cell spread, altered in the double-knockout mutant\n(dospCdospG)?\nResponse: We performed both plaque formation and CFU assays. Overall, the double mutant\nexhibited stronger phenotypes/defects in these new assays (new Fig. 5e–i, lines 325-338).\n-Figure 5A, D. It is very difficult to see bacteria within septin cages. The use of insets with\nhigher magnification would likely help.\nResponse: We included insets with higher magnification (new Fig. 5a, 5c).\n-Figure S7B. Where is the 10RK mutant?\nResponse: We incorporated the 10RK panel (new Fig. S7b).\nfor your support as well as appreciation of our work!"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors did an excellent job in addressing our concerns and we do not have any comments.", "reviewer_2": "The authors have made sufficient efforts to address the reviewer’s main point of criticism -namely, to demonstrate the modification of endogenous SEPT9 in the context of infection at endogenous protein levels. The quality of several immunofluorescence images has also been significantly improved. Additional infection experiments further support the physiological relevance of the effector. I therefore support the publication of the presented work.\nSince SEPT9 has been identified as a new target of a bacterial effector, the presented study represents an important contribution to host–pathogen interaction and septin research.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_4": "The authors have addressed my comments."}, "author_response": ""}]

Decision letter