Nascent RHOH acts as a molecular brake on actomyosin-mediated effector functions of inflammatory neutrophils

RHOH expression is hardly detectable in freshly isolated human and mouse neutrophils; however, its expression is up-regulated by GM-CSF stimulation in less than 3 h (S1A Fig), confirming previously published work [27,38]. High expression of RHOH is also detected in neutrophils from Coronavirus Disease 2019 (COVID-19) patients [39] and in tumor-infiltrating neutrophils in mice [40] as assessed by analyzing publicly accessible single-cell RNA sequencing (scRNA-seq) datasets (GEO GSE158055 and ArrayExpress E-MTAB-8832, respectively) (S1B and S1C Fig), suggesting that RHOH represents a shared transcriptional adaptation conferred to neutrophils by inflammatory microenvironments in distinct neutrophilic diseases.

To assess the impact of RHOH in neutrophil effector function, we isolated blood neutrophils from healthy donors (HD) and CF patients with signs of systemic inflammation (Fig 1A) and confirmed the previously reported high RHOH expression in CF neutrophils (Fig 1B) [27]. There was no obvious difference in ROS production between activated HD and CF neutrophils, as assessed by flow cytometry (Fig 1C). However, degranulation was impaired in CF neutrophils compared to HD neutrophils, as shown by reduced cell surface expression of CD63 and CD66b (surrogate marker for azurophilic granules and specific granules, respectively) and elastase activity (Fig 1A and 1E). Moreover, neutrophils isolated from CF patients failed to form detectable extracellular DNA fibers upon activation (Fig 1F). Consistently, we observed a significant decrease in the amount of double-stranded DNA (dsDNA) release in the supernatants of activated CF neutrophils relative to HD neutrophils (Fig 1G). As a consequence, CF neutrophils generated no detectable NETs, as demonstrated by colocalization of extracellular DNA fibers and neutrophil elastase (NE) (Fig 1H).

To confirm that the observed reduced functionality of CF neutrophils was secondary owing to the inflammatory microenvironment, we incubated human neutrophils from HDs with serum from CF patients. In contrast to HD serum, CF serum significantly elevated the protein level of RHOH (S2A Fig). Moreover, following activation, neutrophils treated with CF serum exhibited a dramatic decrease in degranulation (S2B Fig), dsDNA release (S2C Fig), and in the formation of NETs (S2D Fig), as compared with neutrophils treated with HD serum. These transcriptional and functional changes, which mirrored the phenotype of CF neutrophils, pointed to the hypothesis that inflammatory signals regulate neutrophil effector functions with an intracellular pathway that involves RHOH.

Considering the heterogeneity of CF neutrophils due to disease severity and treatments, we further investigated the functional role of RhoH in purified neutrophils from bone marrow of wild-type (WT) and Rhoh^-/- mice (Fig 2A, S3A and S3B Fig). Rhoh^-/- neutrophils were able to degranulate (Fig 2B) and extrude their dsDNA (Fig 2C and 2D) as efficiently as WT neutrophils in response to C5a following GM-CSF priming. Similar to human neutrophils, primary WT mouse neutrophils expressed barely detectable RhoH protein but up-regulated their RhoH expression following GM-CSF stimulation for 3 h (Fig 2A). Intriguingly, the 3-h stimulated WT neutrophils displayed impaired degranulation (Fig 2B) and dsDNA release (Fig 2C and 2D), while Rhoh^-/- neutrophils treated equally did not show such changes (Fig 2A–D). Other neutrophil agonists known to induce NET formation [35,36], such as PMA, showed similar results (S3C and S3D Fig).

Fig 2. Induction of RhoH expression by GM-CSF stimulation or overexpression of HA-RhoH impairs neutrophil degranulation and dsDNA release.

(A) Neutrophils purified from bone marrow of WT and Rhoh^-/- mice were incubated with GM-CSF for the indicated times followed by immunoblotting. (B–D) Neutrophils from WT and Rhoh^-/- were pretreated with GM-CSF for the indicated times followed by activation with C5a for 15 min. (B) Neutrophil degranulation was determined by flow cytometry. (C) Extracellular DNA fibers were analyzed by confocal microscopy. Scale bars, 10 μm. (D) Quantification of released dsDNA in the culture supernatants. (E) Expression of RhoH in the indicated HoxB8 neutrophils was analyzed by immunoblotting. (F–H) Mature HoxB8 neutrophils were treated with the indicated stimuli. (F) Neutrophil degranulation was determined by flow cytometry. (G) Extracellular DNA fibers were analyzed by confocal microscopy. Scale bars, 10 μm. (H) Quantification of released dsDNA in culture supernatants. (A, C, E, G) Similar results were obtained from 3 independent experiments. (B, D, F, H) Each symbol represents a single experiment. Values are means ± SD. (B, D) Two-way ANOVA with Tukey’s multiple comparisons test. (F, H) Two-way ANOVA with Šídák’s multiple comparisons test. The underlying data for Fig 2B, 2D, 2F, and 2H can be found in S1 Data. The underlying data for Fig 2A and 2E can be found in S1 Raw images. dsDNA, double-stranded DNA; GM-CSF, granulocyte/macrophage colony-stimulating factor; WT, wild-type.

https://doi.org/10.1371/journal.pbio.3001794.g002

We next generated stem cell factor (SCF)-dependent, conditional HoxB8-immortalized (henceforth HoxB8) myeloid progenitor cells from WT and Rhoh^-/- mice. Neutrophils differentiated from these cells have been validated as a useful model for functional studies [35,36]. To determine whether the defect in NET formation was directly related to RhoH, we transduced HoxB8 Rhoh^-/- cells with lentiviral constructs carrying triple hemagglutinin-tagged Rhoh (Gene ID: 74734) (HA–Rhoh) or the corresponding empty vector (EV). After 5 days of differentiation, the re-expressed RhoH in HoxB8 Rhoh^-/- neutrophils exhibited the typical segmented nucleus (S4A Fig) and increased surface expression of neutrophil marker Ly6G (S4B Fig). In addition, the stable expression of HA-RhoH was confirmed by immunoblotting (Fig 2E). The subsequent functional assays revealed that reconstitution of RhoH protein in Rhoh^-/- neutrophils markedly reduced degranulation as compared to HoxB8 EV and non-transduced WT neutrophils (Fig 2F). Moreover, decreased amounts of released dsDNA and extracellular DNA fibers were also observed in HoxB8 RhoH reconstituted neutrophils (Fig 2G and 2H). Taken together, in both human and mouse neutrophils, elevated RHOH expression reduced neutrophil effector functions.

To dissect the underlying molecular mechanism involved in the inhibitory effects of RhoH on neutrophil effector functions, we sought to identify RhoH-interacting proteins in mouse neutrophils by performing HA-RhoH pulldown assay. As shown by Coomassie blue staining, a candidate protein approximately 230 kilo Daltons (kD) in size selectively bound to HA-RhoH upon GM-CSF priming and C5a activation, while this association was weak in activated EV or unstimulated HA-RhoH neutrophils (Fig 3A, top). Mass spectrometry analysis revealed this protein as non-muscle myosin heavy chain IIA (NMHC IIA) (Accession: NP_071855), which is also known as Myh9 or myosin-9, with 45.1% coverage (Fig 3A, bottom). We further verified the increased interaction between RhoH and NMHC IIA in activated mouse neutrophils (Fig 3B). Moreover, this association was also detected in human neutrophils after induction of RHOH by GM-CSF stimulation (Fig 3C).

Fig 3. RhoH interacts with NMHC IIA.

(A) Coomassie blue staining of HA pulldown proteins in HoxB8 neutrophils after stimulation with the indicated triggers. The proteins interacting with RhoH were identified by mass spectrometry analysis. (B, C) Immunoblot analysis after Co-IP with anti-HA antibody in Hoxb8 neutrophils (B) and anti-NMHC IIA antibody in human neutrophils (C). Quantification of the precipitated NMHC IIA (B) or RHOH (C) was shown below the representative images. (D–F) Pretreated human neutrophils with vehicle control or myosin IIA inhibitors for 30 min were subsequently activated with GM-CSF in combination with C5a. (D) Neutrophil degranulation was determined by flow cytometry. (E) Quantification of dsDNA released into the supernatants. (F) NET formation indicated by the colocalization of NE (green) with released DNA (PI, red) was analyzed by confocal microscopy. Scale bars, 10 μm. (G) Protein expression of NMHC IIA in mature HoxB8 neutrophils treated with control (Ctrl) or Myh9 shRNA was analyzed by immunoblot. (H, I) Mature HoxB8 neutrophils were treated as indicated. (H) Neutrophil degranulation was determined by flow cytometry. (I) Quantification of released dsDNA in culture supernatants. (J) Extracellular DNA fibers were analyzed by confocal microscopy. Scale bars, 10 μm. (A–C, F, G, J) Three independent experiments were performed. (B–E, H, I) Values represent means ± SD. (B–E) One-way ANOVA with Dunnett’s multiple comparisons test. (H, I) Two-way ANOVA with Šídák’s multiple comparisons test. The underlying numerical data for Fig 3B–E, 3H, and 3I can be found in S1 Data. The uncropped immunoblots for Fig 3A–C and 3G can be found in S1 Raw images. Co-IP, co-immunoprecipitation; dsDNA, double-stranded DNA; GM-CSF, granulocyte/macrophage colony-stimulating factor; NE, neutrophil elastase; NET, neutrophil extracellular trap; NMHC IIA, non-muscle myosin heavy chain IIA; PI, propidium iodide.

https://doi.org/10.1371/journal.pbio.3001794.g003

Among 3 isoforms of non-muscle myosin II heavy chain (NMHC), termed IIA, B, and C, neutrophils express only NMHC IIA [41]. NMHC IIA functions in the form of myosin IIA and is tightly regulated at the level of folding, myosin filament assembly, actin binding, ATPase, and motor activity, as well as by interactions with other proteins [42]. The fundamental role of myosin IIA in neutrophil migration has been extensively studied in vitro and in vivo [43], yet the involvement of myosin IIA in neutrophil effector function is not fully understood. To examine the potential role of myosin IIA in neutrophil functions, we pretreated human neutrophils with either ML-7, which selectively blocks myosin light chain kinase (MLCK)-mediated activation of myosin II, or (S)-4′-nitro-blebbistatin (nBleb), a photostable and non-fluorescent blebbistatin derivative which inhibits the ATPase activity and sequesters myosin II in a low-affinity state for actin [44,45]. Upon GM-CSF priming and subsequent C5a activation, both inhibitors decreased, in a concentration-dependent manner, neutrophil degranulation (Fig 3D) and dsDNA release (Fig 3E), and, as a consequence, severely impaired the formation of extracellular traps (Fig 3F).

In addition, we reduced NMHC IIA expression with 2 different shRNA targeting Myh9 (Gene ID: 17886) in HoxB8 cells. After differentiation into mature mouse neutrophils (S4B and S4C Fig), successful reduction of NMHC IIA expression was confirmed by immunoblotting (Fig 3G and S5A Fig). NMHC IIA-deficient cells showed defects in NET formation as evidenced by a reduction of degranulation (Fig 3H and S5B Fig) and dsDNA release (Fig 3I and 3J, and S5C Fig).

Given the regulatory role of myosin IIA in actin reorganization, we evaluated the potential inhibition of actin dynamics by myosin IIA inhibitors. F-actin was in a ring-like manner close to the cell membrane in resting human neutrophils but accumulated asymmetrically in the cytosol after neutrophil activation (S6A Fig). ML-7 treatment caused no significant changes in the morphology of resting neutrophils but greatly reduced neutrophil spreading and F-actin formation upon activation (S6A Fig). Consistent with the previous report [43], nBleb treatment caused an increased cell area in resting neutrophils. Subsequent activation of these cells by GM-CSF plus C5a did not induce significant cell spreading additionally (S6A Fig). Short-term application of nBleb did not affect actin polymerization upon neutrophil activation (S6A Fig). In contrast to ML-7 that reduced activation of regulatory light chain (RLC), nBleb had no effects on the phosphorylation of RLC (S6B Fig). In addition, neither ML-7 nor nBleb treatment interfered with ROS production (S6C Fig).

In natural killer (NK) cells, myosin IIA is associated with lytic granules to facilitate the interaction of granules with F-actin at the immunologic synapse for exocytosis [46,47]. Mechanistically, the phosphorylation of NMHC IIA at serine 1943 (S1943) in the tailpiece enables its association with granules and act as a single molecule actin motor [48]. To ascertain if this also applied to neutrophils, we first examined the phosphorylation of NMHC IIA at S1943 by immunoblotting. Interestingly, comparable level of phosphorylation was detected in resting and activated human neutrophils (Fig 4A). We further isolated granules from resting and activated human neutrophils using density gradient ultracentrifugation and equally collected into 10 fractions and detected no traces of actin, indicating the purity of granule fractions (Fig 4B). Granule proteins were mainly detected in fractions 7 to 10 (Fig 4B), as verified by elastase activity (Fig 4C) and capability to kill E. coli-GFP (Fig 4D). NMHC IIA was identified in the postnuclear lysate (PNL) and partial co-fractionation with granule protein MPO, lactoferrin, and MMP-9 (Fig 4B). Consistent with the similar amount of phosphorylation at S1943 in neutrophils before and after activation (Fig 4A), the presence of NMHC IIA in granules also did not change upon neutrophil activation (Fig 4B).

Fig 4. NMHC IIA associates with neutrophil organelles and mediates their binding to actin filaments.

(A) Phosphorylation of myosin IIA in human neutrophils treated as indicated was analyzed by immunoblotting. Threonine 18 (T18); Serine 19 (S19); Serine 1943 (S1943). (B–D) The PNL from unstimulated or GM-CSF primed and C5a-activated human neutrophils was isolated and collected equally into 1–10 fractions, from top to bottom. (B) Immunoblot analysis of density gradient fractions. (C) NE activity of density gradient fractions. (D) Bacteria killing mediated by density gradient fractions. (E) Immunoblot analysis of NMHC IIA in cytosolic and mitochondrial fractions from control and activated human neutrophils. C, cytosolic fraction. M, mitochondrial fraction. (F) Granular or mitochondrial localization of NMHC IIA in unstimulated and activated human neutrophils was analyzed by confocal microscopy. Scale bars, 10 μm. Numerical analysis was performed on 10 cells in each group and the Pearson’s coefficient in colocalized volume between NMHC IIA and CD63 or MTO was calculated using Imaris software. (G) Enriched granules and mitochondria from mature HoxB8 neutrophils treated with control (Ctrl) or Myh9 shRNA were incubated for 30 min in the presence or absence of F-actin. The pellet (P) and supernatant (S) were evaluated by immunoblotting (left). The ratios of F-actin in the pellet (P) to the supernatant (S) were quantified (right). (A, B, E–G) Data are representative of 3 independent experiments. (C, D, F, G) Values are means ± SD. (F, G) Unpaired 2-tailed Student t test was applied. The underlying numerical data for Fig 4C, 4D, 4F, and 4G can be found in S1 Data. The uncropped immunoblots for Fig 4A, 4B, 4E, and 4G can be found in S1 Raw images. GM-CSF, granulocyte/macrophage colony-stimulating factor; NE, neutrophil elastase; NMHC IIA, non-muscle myosin heavy chain IIA; PNL, postnuclear lysate.

https://doi.org/10.1371/journal.pbio.3001794.g004

Beyond granules, NMHC IIA has also been shown to interact with mitochondria membranes independently of F-actin as demonstrated previously in platelets [49]. Therefore, resting and activated human neutrophils were subjected to cell fractionation, and the mitochondrial fraction was separated from the cytosolic fraction and analyzed by immunoblotting. We detected NMHC IIA mainly in the cytosol and only traces in the isolated mitochondrial fraction, together with other known mitochondrion-localizing proteins, such as mitofusin-2 and TOMM20 (Fig 4E). We compared the distribution of NMHC IIA in the mitochondrial fraction between resting and activated human neutrophils and failed to observe any significant differences (Fig 4E). Moreover, the Pearson’s coefficient value between the colocalized volume of NMHC IIA and CD63 or MTO indicated similar colocalization in resting and activated neutrophils (Fig 4F). Taken together, these results pointed to an interaction between myosin IIA and neutrophil organelles that is constantly present in resting and activated human neutrophils.

To test whether NMHC IIA enabled the loading of the associated organelles to F-actin for intracellular trafficking, we performed an actin-binding in vitro assay as previously described [47]. Isolated mouse neutrophil granules and mitochondria were incubated with exogenous F-actin and then subjected to centrifugation at a speed which is sufficient to pellet organelles, but not F-actin alone. The actin crosslinking protein, α-actinin, was used as a positive control to aggregate F-actin (Fig 4G, upper panel). As expected, the addition of granules and mitochondria co-sedimented the F-actin in the pellet and reduced the free F-actin in the supernatant of HoxB8 neutrophils treated with control (Ctrl) shRNA (Fig 4G, lower panel). The amount of NMHC IIA associated with the granules and mitochondria decreased and therefore the percentage of sedimented F-actin in the pellet was significantly reduced in Myh9 shRNA compared to Ctrl treated mouse neutrophils (Fig 4G, lower panel). These findings, together with results obtained from NET formation assays (Fig 3H–3J), suggested that NMHC IIA enables the binding of granules and mitochondria to F-actin for intracellular organelle trafficking.

To understand the cellular alterations underlying the interaction between RhoH and NMHC IIA upon neutrophil activation, we next evaluated the intracellular localization of NMHC IIA. Similar amounts of NMHC IIA were detected in granule and mitochondrial fractions from EV and HA-RhoH expressing neutrophils (S7A and S7B Fig). Intriguingly, RhoH was also found to localize at granules and mitochondria (S7A and S7B Fig), presumably owing to its interaction with NMHC IIA. Moreover, overexpression of RhoH reduced the ability of neutrophil granules and mitochondria to interact with F-actin (Fig 5A), corresponding with the impaired degranulation and dsDNA release seen in these cells (Fig 2E–2H).

Fig 5. RhoH inhibits NMHC IIA-mediated granules and mitochondria binding to F-actin.

(A) Enriched granules and mitochondria from mature HoxB8 neutrophils expressing HA-RhoH or the corresponding EV were incubated for 30 min in the presence or absence of F-actin. After centrifugation, the pellet (P) and supernatant (S) were evaluated by immunoblotting (left). The ratios of F-actin in the pellet (P) to the supernatant (S) were quantified (right). (B) Docking analysis for predicting the binding sites between RHOH and NMHC IIA (upper). Sequence alignment of RAC1, CDC42, RHOA, and RHOH (lower). (C) Immunoblot analysis of Co-IP with anti-HA antibody in cell lysates from activated HoxB8 neutrophils expressing WT or mutated HA-RhoH. Quantification of the precipitated NMHC IIA was shown below the representative images. (D) Enriched granules and mitochondria were incubated for 30 min in the presence or absence of F-actin. The pellet (P) and supernatant (S) were evaluated by immunoblotting (left). The ratios of F-actin in the pellet (P) to the supernatant (S) were quantified (right). (E–G) Mature HoxB8 neutrophils expressing WT or mutated HA-RhoH or EV were treated as indicated. (E) Neutrophil degranulation was determined by flow cytometry. (F) Extracellular DNA fibers were analyzed by confocal microscopy. Scale bars, 10 μm. (G) Quantification of dsDNA released into the supernatants. (A, C, D, F) Data are representative of 3 independent experiments. Values are means ± SD. Two-tailed Student t test (A, D); 1-way ANOVA with Dunnett’s multiple comparisons test (C); 2-way ANOVA with Tukey’s multiple comparisons test (E, G). The underlying numerical data for Fig 5A, 5C–E, and 5G can be found in S1 Data. The uncropped immunoblots for Fig 5A, 5C, and 5D can be found in S1 Raw images. Co-IP, co-immunoprecipitation; dsDNA, double-stranded DNA; EV, empty vector; NMHC IIA, non-muscle myosin heavy chain IIA; WT, wild-type.

https://doi.org/10.1371/journal.pbio.3001794.g005

Molecular docking analysis suggests that RHOH spatially interacted with NMHC IIA by forming hydrogen bonds with several amino acid residues (Fig 5B). Among which, alanine (Ala) 655 and Ala 659 were present at the actin-binding motifs (residues 654–676) of NMHC IIA [50]. RHOH residue lysine (Lys) 34 (K34), which belongs to the switch I domain [51], was located at the interface with NMHC IIA (Fig 5B). Sequence alignment of RHOH and the classical RHO GTPases RAC1, CDC42, RHOA indicated that K34 (depicted in red) is a RHOH-specific amino acid, which might reflect its indispensability for RhoH function (Fig 5B). We therefore proposed that RHOH may engage the actin-binding interface in NMHC IIA to impair NMHC IIA-actin interaction and induce molecular motor dysfunction for neutrophil organelle transport.

To this end, we designed specific point mutations in mouse RhoH (Accession: NP_001074574.1) and transduced HoxB8 cells with the corresponding lentiviral constructs. Upon differentiation into mature neutrophils, a point mutation of Lys 34 to Ala (K34A) in RhoH (RhoH^K34A) displayed decreased ability to interact with NMHC IIA (Fig 5C), allowing the binding of granules and mitochondria to F-actin in an actin-binding in vitro assay (Fig 5D). These cells also elicited efficient degranulation and dsDNA release following activation, in contrast to WT RhoH overexpressing HoxB8 neutrophils (Fig 5E–5G). According to the molecular docking results, the neighboring residue tyrosine (Tyr) 33 in RhoH was not involved in binding to NMHC IIA and therefore used as a mutant control (Fig 5B). Unexpectedly, the Tyr 33 to phenylalanine (Phe) (RhoH^Y33F) mutant and the combined double mutant (RhoH^{Y33F and K34A}) failed to stably express at protein level despite profound mRNA expression (Fig 5C and S8A Fig). Lower expression of RhoH^Y33F or RhoH^{Y33F and K34A} mutants appeared to be due to lysosomal degradation since the lysosomal proton pump inhibitor bafilomycin A1 (Baf A1) could block RhoH degradation (S8B Fig), as it has been previously reported in T cells [22]. Collectively, these results demonstrated that RhoH inhibits myosin IIA-mediated granules and mitochondria transport by blocking their binding to F-actin.

Given that RHO GTPases are key regulators of the cell cytoskeleton, we further evaluated the effects of RhoH on F-actin and microtubule cytoskeleton before and after neutrophil activation. We did not observe significant difference in microtubule network between EV and HA-RhoH expressing neutrophils (S9A Fig). However, overexpression of RhoH reduced F-actin formation upon neutrophil activation as compared to EV (Fig 6A), without interfering with ROS production (S9B Fig), which plays an important role in regulating F-actin polymerization. Defective F-actin formation was reversed by K34A mutation in RhoH (Fig 6A). Moreover, RhoH had no effects on the phosphorylation of RLC and NMHC IIA of myosin IIA upon neutrophil activation (Fig 6B), indicating that other factors are involved in the regulation of F-actin formation by RhoH.

Fig 6. RhoH impairs actin rearrangement presumably by modulating Rac1 activity.

(A, B) Mature HoxB8 neutrophils expressing WT or mutated HA-RhoH or EV were treated as indicated. (A) F-actin distribution was analyzed by confocal microscopy (left). Scale bars, 10 μm. Quantification of F-actin fluorescence intensity was performed by automated analysis of microscopic images using Imaris software. The 25 images (each containing 8–12 cells) from 3 independent experiments were included for each condition (right). (B) Phosphorylation of myosin IIA was analyzed by immunoblotting. (C, D) The levels of GTP-bound and total Rac1 and Rac2 proteins in the indicated mouse bone marrow (BM) neutrophils (C) and HoxB8 neutrophils (D) were examined by immunoblotting following pulldown assay. All data are representative of 3 independent experiments. Values are means ± SD. One-way ANOVA with Tukey’s multiple comparisons test was applied. The underlying numerical data for Fig 6A–D can be found in S1 Data. The uncropped immunoblots for Fig 6B–D can be found in S1 Raw images. EV, empty vector; WT, wild-type.

https://doi.org/10.1371/journal.pbio.3001794.g006

RHOH has previously been reported to modulate the activities of other RHO GTPases such as RAC, CDC42, and RHOA [52,53], therefore, we next determined the activation of these RHO GTPases by effector pulldown assays. Similar amounts of GTP-bound Cdc42 (S9C Fig) and GTP-bound RhoA (S9D Fig) were observed upon stimulation of neutrophils expressing HA-RhoH and EV. Rhoh deficiency did not cause significant changes in Rac1 and Rac2 activation (Fig 6C). RhoH overexpression reduced Rac1 activity, which was abolished by K34A mutation in RhoH (Fig 6D). The inhibitory effect was not observed on Rac2 activation (Fig 6D). Thus, RhoH inhibits F-actin formation presumably by modulating Rac1 activity.

In order to verify the regulatory role of RhoH in vivo, we used an acute murine peritonitis model in which a low intraperitoneal dose of GFP-E. coli is applied [54]. Consistent with our results obtained in in vitro stimulation experiments (Fig 2A), RhoH was up-regulated in peritoneal cells in a time-dependent manner, with abundant expression observed at 8 h following E. coli challenge (Fig 7A). Compared to WT, bacterial loads at 8 h after infection were significantly reduced in Rhoh^-/- mice (Fig 7B). NETs were likely to be involved in the observed bacteria killing, as large amount of dsDNA (Fig 7C) and NE (Fig 7D) were detected in peritoneal lavage fluid (PLF) supernatants. In addition, higher levels of dsDNA and NE were detected in PLF obtained from Rhoh^-/- mice (Fig 7C and 7D).

Fig 7. Rhoh^-/- neutrophils display enhanced NET formation and bacteria killing in experimental peritonitis.

(A) Peritoneal cells from WT mice challenged intraperitoneally with E. coli was analyzed by immunoblot. Protein lysates of bone marrow (BM) neutrophils from WT and Rhoh^-/- mice were loaded as negative control. Data were representative of 3 independent experiments. (B–I) WT and Rhoh^-/- mice were challenged intraperitoneally with E. coli and sacrificed after 8 h; n = 10 for each group unless mentioned otherwise. (B) Bacteria burden was quantified as CFU. (C) Quantification of released dsDNA in PLF supernatants. (D) Quantification of NE activity in PLF supernatants. (E) Total peritoneal cell count. (F) Infiltrated peritoneal neutrophils were defined as CD45⁺ and Ly6G⁺. Left, representative original flow cytometry data. Right, proportion of peritoneal neutrophils. (G) Number of peritoneal neutrophils. (H) Protein level of RhoH in peritoneal cells was analyzed by immunoblot (n = 5). (I) Representative confocal images of peritoneal cells stained with MitoSOX Red and Hoechst 33342, left. Scale bars, 10 μm. Quantification of cells releasing dsDNA fibers was performed in 2 randomly selected images from each mouse, right. Values are means ± SD. Statistical analyses were performed using unpaired 2-tailed Student t test. The underlying data for Fig 7B–E, 7G, and 7I can be found in S1 Data. The underlying data for Fig 7A and 7H can be found in S1 Raw images. CFU, colony-forming unit; dsDNA, double-stranded DNA; NE, neutrophil elastase; NET, neutrophil extracellular trap; PLF, peritoneal lavage fluid; WT, wild-type.

https://doi.org/10.1371/journal.pbio.3001794.g007

Enhanced bacteria killing and increased accumulation of NETs components may result from more neutrophils recruited to the inflamed peritoneal cavities of Rhoh^-/- mice. However, although E. coli injection resulted in abundant accumulation of immune cells (Fig 7E) and neutrophils represented the dominant cell type (Fig 7F), we failed to observe significant differences in the absolute number of neutrophils between WT and Rhoh^-/- mice (Fig 7G). Since we detected up-regulated RhoH expression in infiltrating cells in WT mice (Fig 7H), we speculated that perturbed neutrophil activity may be the reason. Indeed, Rhoh^-/- neutrophils displayed enhanced ability in forming extracellular dsDNA fibers (Fig 7I). Collectively, RhoH knockout results in enhanced NET formation and host defense against bacteria, confirming RhoH as a molecular brake on neutrophil effector functions under in vivo conditions.

Rhoh^-/- mice with a C57BL/6J background were generated and provided by Dr. C. Brakebusch (Department of Molecular Pathology, University of Copenhagen, Copenhagen, Denmark) [18]. WT mice on C57BL/6J background were bred germfree at the Clean Mouse Facility, University of Bern, Switzerland. Mice were housed in specific-pathogen-free (SPF) conditions and were euthanized using carbon dioxide gas before material harvest. Sex- and age-matched mice were used in our experiments. Mouse studies were approved by the Cantonal Veterinary Office of Bern, Switzerland (license number 40/09) and carried out in accordance with the Swiss federal legislation on animal welfare.

Human blood neutrophils from healthy individuals and CF patients were isolated by Ficoll-Hypaque centrifugation as described previously [35]. Written, informed consent was obtained from all blood donors and this study was approved by the Ethics Committee of Canton Bern (approval number 18/2001). The purified populations contained >95% neutrophils as assessed by staining with the Hematocolor Stain Set (Merck Millipore) followed by light microscopic analysis.

Primary mouse bone marrow neutrophils were isolated from WT and Rhoh^-/- mice with a negative selection technique using the EasySep Mouse Neutrophil Enrichment Kit (Stemcell Technologies) [35]. Neutrophil purity was always higher than 85% as assessed by Diff-Quik staining and light microscopy.

Murine HA-tagged Rhoh (HA-Rhoh) cDNA containing vector EX-Mm24061-Lv118 and the corresponding EV EX-NEG-Lv1118 were purchased from GeneCopoeia, (Rockville, Maryland, USA). To generate point mutations in HA-RhoH, the QuikChange II XL Site-Directed Mutagenesis Kit (Cat # 200521, Agilent Technologies, Basel, Switzerland) was used. Mutation primers were as follows: HA-RhoH_Tyr33 forward: 5′-CCT TCC CGG AGG CCT TCA AAC CCA CGG TGT-3′; HA-RhoH_Tyr33 reverse: 5′-ACA CCG TGG GTT TGA AGG CCT CCG GGA AGG-3′; HA-RhoH_Lys34 forward: 5′-TCC CGG AGG CCT ACG CAC CCA CGG TGT ACG-3′; HA-RhoH_Lys34 reverse: 5′-CGT ACA CCG TGG GTG CGT AGG CCT CCG GGA-3′; HA-RhoH_Tyr33Lys34 forward: 5′-GAC CTT CCC GGA GGC CTT CGC ACC CAC GGT GTA-3′; and HA-RhoH_Tyr33Lys34 reverse: 5′-GTA CAC CGT GGG TGC GAA GGC CTC CGG GAA GGT C-3′. All mutations were introduced by PCR following the manufacturer’s instructions. The resulting constructs were verified by sequence analysis. Sequencing primers: forward: 5′-CGG TGG GAG GTC TAT ATA AGC AG-3′, reverse: 5′-ATT GTG GAT GAA TAC TGC C-3′. Bacterial glycerol stock harboring sequence-verified shRNA of mouse Myh9 lentiviral plasmid vector: shRNA Myh9 (1), sequence: CGG TAA ATT CAT TCG TAT CAA; shRNA Myh9 (2), sequence: GCC ATA CAA CAA ATA CCG CTT or the control pLKO.1-puro vector were obtained from Sigma-Aldrich (Mission shRNA). Virus was produced by transfecting HEK-293T cells with the plasmid of interest using a calcium phosphate transfection method [68]. The supernatants were collected after 24 h, filtered through a 0.22 μm filter (Merck Millipore) and stored at −80°C before use.

SCF-dependent, conditional HoxB8-immortalized myeloid progenitors were generated from WT and Rhoh^-/- mice as previously described [69]. HoxB8 cells were passaged in RPMI 1640/GlutaMAX with 10% FCS, 100 U/ml penicillin, 100 μg/ml streptomycin, 5% SCF (CHO/SCF cell conditioned medium), and 100 nM 4-OHT. To transfer the gene of interest, 1 ml of viral supernatant was added to 5 × 10⁵ HoxB8 cells in the presence of 16 μg of Polybrene (Sigma-Aldrich) and 100 nM 4-OHT. Transduced cells were selected with antibiotic starting 48 h after transduction and transfection efficiency was evaluated by immunoblot.

Total RNA was extracted from neutrophils using the Quick-RNA MicroPrep Kit (Zymo Research, Irvine, California, USA) and reverse transcribed to cDNA with SuperScript III Reverse Transcriptase Kit (Thermo Fisher Scientific). Quantitative PCR was carried out with the iTaq Universal SYBR Green Supermix (Bio-Rad Laboratories, Cressier, Switzerland) using the CFX Connect Real-Time PCR Detection system (Bio-Rad Laboratories). The following primers were used [27]: mouse Rhoh forward: 5′-GAC CTT CCC GGA GGC CTA CA-3′ and mouse Rhoh reverse: 5′-TGC CGG CAG TGT CCC AGA GA-3′; mouse 18S forward: 5′-ATC CCT GAG AAG TTC CAG CA-3′ and mouse 18S reverse:5′-CCT GGG TCA TCA TCA GGC TT-3′. Primers were synthesized by Microsynth AG, Balgach, Switzerland.

Neutrophils were washed twice with cold PBS and lysed in lysis buffer containing 50 mM Tris (pH 7.4), 150 mM NaCl, 10% glycerol, 1% Triton X-100, 2 mM EDTA, 10 mM sodium pyrophosphate, 50 mM sodium fluoride, and 200 μm sodium orthovanadate. Shortly before use, 100 μm PMSF, a protease inhibitor cocktail and phosphatase inhibitors (Sigma-Aldrich) were freshly added into the lysis buffer. After incubation in lysis buffer on ice for 30 min, cell lysates were centrifugated at 13,000 rpm for 10 min at 4°C and the supernatant was collected. Thereafter, the protein concentration was measured with the BCA Protein Assay Kit (Thermo Fisher Scientific). Samples were heated at 95°C for 5 min. Proteins were separated by SDS-PAGE and transferred onto PVDF membranes (Immobilion-P; Merck Millipore). Membranes were incubated with a blocking buffer consisting of Tris-buffered saline (pH 7.4) with 0.1% Tween 20 (TBST) and 5% non-fat dry milk at room temperature for 1 h and then incubated with primary antibodies at 4°C overnight. The primary antibodies used as follows: anti-RhoH [26,27], anti-HA (1:1,000, Roche Diagnostics, Rotkreuz, Switzerland); anti-p-RLC (T18/S19) (1:500, Cell Signaling Technology, Danvers, Massachusetts, USA); anti-RLC (1:1,000, Cell Signaling Technology); anti-NMHC IIA (1:2,000, Abcam); anti-p-NMHC IIA (S1943) (1:2,000, Cell Signaling Technology); anti-MPO (1:1,000, Thermo Fisher Scientific); anti-Lactoferrin (1:500, Santa Cruz Biotechnology, Dallas, Texas, USA); anti-MMP9 (1:500, Santa Cruz Biotechnology); anti-mitofusin-2 (1:1,000, Cell Signaling Technology), anti-TOMM20 (1:1,000; Novus Biologicals); anti-VDAC (1:1,000; Merck Millipore); anti-OPA1 (1:1,000, Sigma-Aldrich); anti-Rho GDI (1:1,000, Santa Cruz Biotechnology); anti-actin (1:10,000, Cytoskeleton distributed by LuBioScience GmbH) and anti-GAPDH (1:2,000; Merck Millipore). In the next morning, membranes were washed in TBST for 30 min at room temperature and incubated with the corresponding HRP-conjugated secondary antibody (GE Healthcare Life Sciences, Little Chalfont, UK) in TBST with 5% non-fat dry milk at room temperature for 1 h. After washing, membranes were developed with LI-COR Odyssey imaging system and signals were quantified using Image Studio software (LI-COR Biosciences, Lincoln, Nebraska, USA).

Subcellular fractionation was performed as previously described [70]. Briefly, neutrophils were lysed in cytosolic extraction buffer (CEB) (20 mM HEPES (pH 7.2), 250 mM sucrose, 10 mM KCl, 1.5 mM MgCl₂, and 2 mM EDTA) with digitonin (0.625 mg/ml) on ice for 20 min. Lysed cells were centrifuged at 700 g at 4°C for 10 min. The supernatant was collected and centrifuged again at 7,000 g for 30 min at 4°C. The supernatant was then transferred to a new tube for ultracentrifugation at 21,000 g at 4°C for 60 min. The supernatant after ultracentrifugation contains the cytosolic fraction. The pellet after centrifugation at 7,000 g contains the mitochondrial fraction was washed 2 times with cold PBS. Thereafter, the cell pellet was lysed with CEB buffer containing 1% SDS by boiling at 95°C for 5 min. The lysate was centrifuged at 14,000 rpm at 4°C for 5 min. The supernatant after centrifugation at 14,000 rpm at 4°C for 5 min that contained the mitochondrial fraction was collected for immunoblot analysis.

Neutrophil granules were isolated by density gradient ultracentrifugation using Lysosomal Enrichment Kit (Sigma-Aldrich) as previously described [47]. In brief, neutrophils were homogenized using a Dounce homogenizer and cell lysates were centrifuged at 500 g for 10 min to remove the nuclei. The postnuclear supernatant was layered onto a discontinuous Optiprep gradient (8% to 27%) and centrifuged at 150,000 g at 4°C in a 50 TI fixed angle rotor in Beckman Optima L-90K Ultracentrifuge. The 10 fractions were equally collected from top to bottom and diluted with cold PBS. Optiprep was removed by centrifugation of each fraction at 21,000 g for 30 min. The separated subsets were used for further analysis.

The protein structures of RhoH and NMHC IIA were built by using SWISS-MODEL (https://swissmodel.expasy.org). The SWISS-MODEL template library (SMTL version 2020-08-05, PDB release 2020-07-31) was searched with BLAST and HHBlits for evolutionary related structures matching the target sequence. Models were built based on the target-template alignment using ProMod3. The best models referred to reported crystal structures of CDC42 (6SIU) and Myosin 2 heavy chain (3JAX) in the PDB database as templates were picked out for the prediction of the binding sites by performing ZDOCK SERVER (https://zdock.umassmed.edu). The top dock poses in the largest cluster were analyzed and visualized by PyMOL (https://pymol.org).

E. coli-GFP were prepared as described above, and 50 × 10⁶E. coli in 150 μl PBS were injected intraperitoneally into mice to induce peritonitis as previously reported [54]. Serial dilutions of the final bacterial inoculum were distributed on agar plates and incubated overnight at 37°C to verify the number of viable bacteria injected. At designated time points after E. coli challenge, mice were anesthetized and sacrificed. The peritoneal cavity was flushed using 2 ml PBS containing 2.5 mM EDTA. The harvested peritoneal wash was distributed on agar plates in triplicate. After overnight incubation at 37°C, the number of colony-forming unit (CFU) was counted on the second day. The remaining peritoneal wash was centrifuged, and the supernatant was collected as PLF and aliquoted for further measurements. The peritoneal cells in the pellet were washed and subsequently stained with MitoSOX Red (5 μm) and 1 μg/ml Hoechst 33342 to visualize NETs or incubated with blocking buffer prior to cell surface staining with the following antibodies from BioLegend: PerCP-conjugated anti-CD45 and APC/cy7-conjugated anti-Ly6G antibodies. All samples were analyzed by flow cytometry (FACSVerse, BD Biosciences) followed by quantification using FlowJo software. Additionally, proteins were extracted from peritoneal cells and analyzed for RhoH protein expression by immunoblotting.