ABSTRACT

Phagosome formation is a complicated process that requires spatiotemporally regulated actin reorganization. We found that RhoC GTPase is a critical regulator of FcγR-mediated phagocytosis in macrophages. Our live-cell imaging revealed that RhoC, but not RhoA, is recruited to phagocytic cups engulfing IgG-opsonized erythrocytes (IgG-Es). RhoC silencing through RNAi, CRISPR/Cas-mediated RhoC knockout, and the expression of dominant-negative or constitutively active RhoC mutants suppressed the phagocytosis of IgG-Es. Moreover, RhoC-GTP pulldown experiments showed that endogenous RhoC is transiently activated during phagosome formation. Notably, actin-driven pseudopod extension, which is required for the formation of phagocytic cups, was severely impaired in cells expressing the constitutively active mutant RhoC-G14V, which induced abnormal F-actin accumulation underneath the plasma membrane. mDia1 (encoded by DIAPH1), a Rho-dependent actin nucleation factor, and RhoC were colocalized at the phagocytic cups. Similar to what was seen for RhoC, mDia1 silencing through RNAi inhibited phagosome formation. Additionally, the coexpression of mDia1 with constitutively active mutant RhoC-G14V or expression of active mutant mDia1-ΔN3 drastically inhibited the uptake of IgG-Es. These data suggest that RhoC modulates phagosome formation be modifying actin cytoskeletal remodeling via mDia1.

INTRODUCTION

Phagocytosis – a specialized form of endocytosis that permits the uptake of large particles (>0.5 μm in diameter) – plays an essential role in the host defense mechanism and in tissue remodeling. Professional phagocytes, such as macrophages and neutrophils, recognize, internalize and dispose of foreign particles, invading microorganisms and apoptotic bodies, thus contributing to the resolution of infections and the clearance of senescent or damaged cells. Phagocytosis of a particle is initiated by the binding of that particle to a specific cell surface receptor, including Fcγ receptors (FcγRs), complement receptors (CRs) and scavenger receptors (Freeman and Grinstein, 2014). Phagocytosis via FcγRs, the best-characterized pathway of phagocytosis, entails actin polymerization and reorganization, which drive the extension of pseudopods around IgG-opsonized particles to form phagocytic cups (Araki et al., 1996, 2003; Groves et al., 2008; Swanson, 2008). Pseudopod extension is accompanied by the formation of branched F-actin networks generated by the Arp2/3 complex (Campellone and Welch, 2010). During phagocytic cup formation, the actin-nucleating activity of Arp2/3 is stimulated via binding to the Wiskott-Aldrich syndrome protein (WASP) and WASP-family verprolin-homologous protein (WAVE) family members (Lorenzi et al., 2000; May et al., 2000; Park and Cox, 2009; Tsuboi and Meerloo, 2007). Arp2/3-mediated actin-filament assembly has been proposed as the driving force for the formation of phagocytic cups (May et al., 2000). Importantly, to complete phagosome formation, actin disassembly at the base of phagocytic cups needs to occur concurrently with the maximal extension of pseudopods around particles (Egami et al., 2011; Greenberg et al., 1993). After the closure of the phagocytic cups, the newly formed phagosomes mature via a series of interactions with endocytic compartments and finally fuse with lysosomes for particle degradation (Downey et al., 1999; Fairn and Grinstein, 2012).

Rho family GTPases are small GTP-binding proteins that primarily function as molecular switches by cycling between their active GTP- and inactive GDP-bound forms. The GTP-bound forms interact with their downstream effectors and regulate cytoskeletal dynamics, thereby affecting cell polarity and motility. To date, 20 genes encoding Rho family members have been identified in the human genome (Vega and Ridley, 2007). During FcγR-mediated phagocytosis, several Rho family GTPases (e.g. Rac1, Rac2, Cdc42 and RhoG) are recruited to phagocytic cups and modulate phagosome formation (Beemiller et al., 2010; Caron and Hall, 1998; Cox et al., 1997; Hoppe and Swanson, 2004; Ikeda et al., 2017; Massol et al., 1998; Tzircotis et al., 2011). Among these GTPases, the GTP-bound forms of Cdc42 and Rac1 have been shown to stimulate the Arp2/3 complex via WASP and WAVE family proteins (Chen et al., 2010; Eden et al., 2002; Lorenzi et al., 2000; Park and Cox, 2009; Tsuboi and Meerloo, 2007), primarily resulting in the formation of a branched F-actin network at the phagocytic cup. However, the roles of other Rho family members – such as RhoA, RhoB and RhoC, which promote the polymerization of unbranched F-actin – in phagosome formation have not been fully examined. These GTPases share 88% amino acid sequence identity (Wheeler and Ridley, 2004) but appear to have different functions (Vega et al., 2011). RhoA and RhoC are localized to the plasma membrane and cytoplasm, while RhoB is observed in the endosomal membranes and controls endosomal trafficking (Adamson et al., 1992; Heasman and Ridley, 2008). Several lines of evidence indicate that RhoA, but not RhoC, is essential for CR3 (integrin αMβ1)-mediated phagocytosis in macrophages (Caron and Hall, 1998; Kim et al., 2012; Tzircotis et al., 2011; Wiedemann et al., 2006). RhoB has been reported to be expressed in macrophages and be involved in mannose receptor-mediated phagocytosis (Zhang et al., 2005). Importantly, the involvement of RhoA and RhoC in FcγR-mediated phagocytosis remains ill defined.

Mammalian diaphanous-related formins act as Rho GTPase effectors and induce de novo formation of unbranched actin filaments. The formin proteins mDia1/2 (encoded by DIAPH1 and DIAPH2, respectively) and FMNL1 are involved in several types of phagocytosis (Brandt et al., 2007; Colucci-Guyon et al., 2005; Naj et al., 2013; Seth et al., 2006). During FcγR-mediated phagocytosis, FMNL1 is recruited to phagocytic cups in a Cdc42-dependent manner and regulates phagosome formation (Seth et al., 2006). In contrast, mDia1/2 – major downstream effectors of RhoA, RhoB and RhoC – are primarily implicated in CR3-mediated phagocytosis in macrophages (Colucci-Guyon et al., 2005). Intriguingly, in neutrophils, mDia1 has been shown to be essential for both CR3- and FcγR-mediated phagocytosis (Shi et al., 2009). Therefore, the participation of mDia1 in FcγR-mediated phagocytosis in macrophages should be investigated.

In the present study, by using live-cell imaging, we found that RhoC, but not RhoA, is transiently recruited to actin-enriched phagocytic cups during FcγR-mediated phagocytosis. Furthermore, the expression of RhoC or mDia1 mutants, RNAi-based knockdown and CRISPR/Cas-mediated knockout (KO) analyses revealed that RhoC regulates phagosome formation via mDia1 in macrophages. Our study provides novel insight into the importance of unbranched F-actin remodeling via RhoC/mDia1 signaling in the uptake of particles during FcγR-mediated phagocytosis.

RESULTS

RhoC, but not RhoA, is recruited to phagocytic cups during FcγR-mediated phagocytosis

Previous reports have shown that several Rho GTPases (Cdc42, Rac1, Rac2 and RhoG) are involved in FcγR-mediated phagocytosis (Cox et al., 1997; Hoppe and Swanson, 2004; Ikeda et al., 2017; Massol et al., 1998; Tzircotis et al., 2011). However, the involvement of two other Rho family members, RhoA and RhoC, during FcγR-mediated phagocytosis has not been fully investigated. RhoA is expressed in RAW264 macrophages and bone marrow-derived macrophages (BMMs) (Kim et al., 2012; Nakaya et al., 2006), while RhoC is highly expressed in RAW264 cells and is detectable at low levels in BMMs by western blotting (data not shown). To examine the spatiotemporal dynamics of Rho GTPases during FcγR-mediated phagocytosis, RAW264 macrophages coexpressing GFP–RhoA and TagRFP–RhoC were allowed to phagocytose IgG-opsonized erythrocytes (IgG-Es) and analyzed by live-cell imaging with a confocal laser microscope. Prior to phagocytosis, RhoA and RhoC were mainly found in the cytosol, as previously reported in other cell types (Adamson et al., 1992). After the binding of IgG-Es to the cells, RhoC, but not RhoA, was recruited to the membranes of the phagocytic cups extending along the surface of the IgG-Es (Fig. 1A, t=2 min; Movie 1). RhoC subsequently dissociated from the membranes of the nascent phagosomes (Fig. 1A, t=8 min). Time-lapse imaging showed that RhoC was localized in the membranes of linear and circular ruffles (precursor forms of macropinosomes), regardless of the addition of IgG-Es to the cells, as previously reported in other cell species (Zawistowski et al., 2013). We confirmed that the expression of RhoA does not affect the localization of RhoC (Fig. 1A; Fig. S1). To ascertain the specific recruitment of RhoC to the phagocytic cups, we performed line-scan analysis of the fluorescence intensities of GFP–RhoA and TagRFP–RhoC and quantified their fluorescence intensities. As shown in Fig. 1B,C, RhoC clearly accumulated in the membranes of phagocytic cups, in contrast with RhoA.

Fig. 1.

RhoC, but not RhoA, accumulates in the phagocytic cups during FcγR-mediated phagocytosis. (A) Live RAW264 cells coexpressing GFP–RhoA (green) and TagRFP–RhoC (red) were put into contact with IgG-Es and observed by confocal laser microscopy. Phase-contrast images are shown in the top panels. The elapsed time is indicated at the top left. The binding of IgG-Es to the cell surface is set as time 0. The insets show higher magnification images of the indicated areas of the cells. TagRFP–RhoC was recruited to the membranes of the phagocytic cups, whereas GFP–RhoA did not accumulate. Representative images from three independent experiments are shown. The corresponding movie is Movie 1. Scale bar: 5 μm. (B) Line-scan analysis performed using MetaMorph software shows the fluorescence intensities of GFP–RhoA (green) and TagRFP–RhoC (red) at the position of the line in the enlarged image of the boxed region in Fig. 1A, 2 min. Strong fluorescent signals of TagRFP–RhoC were detected at the phagocytic cups (arrowheads). (C) Quantitation of the accumulation levels of RhoC and RhoA at the phagocytic cup. Maximum fluorescence intensity values were measured at the phagocytic cup in each cell coexpressing TagRFP–RhoC and GFP–RhoA or expressing TagRFP (Mock). The fluorescence intensity of TagRFP–RhoC, GFP–RhoA or TagRFP was normalized to that of a region in the cytoplasm. Values represent the means±s.e.m. of three independent replicates (n=3; 30 phagocytic cups in more than five cells in each condition were assessed per replicate). *P<0.05 (one-way ANOVA followed by Tukey's test). (D) RAW264 macrophages were incubated with 2-μm hIgG-coated beads for 10 min before fixation and immunostained with an anti-RhoC antibody, an anti-RhoA antibody or an isotype-matched control antibody. Phase-contrast images are shown. The insets are magnifications of the boxed areas. Note the recruitment of endogenous RhoC to the phagocytic cups. Scale bars: 5 μm.

Fig. 1.

RhoC, but not RhoA, accumulates in the phagocytic cups during FcγR-mediated phagocytosis. (A) Live RAW264 cells coexpressing GFP–RhoA (green) and TagRFP–RhoC (red) were put into contact with IgG-Es and observed by confocal laser microscopy. Phase-contrast images are shown in the top panels. The elapsed time is indicated at the top left. The binding of IgG-Es to the cell surface is set as time 0. The insets show higher magnification images of the indicated areas of the cells. TagRFP–RhoC was recruited to the membranes of the phagocytic cups, whereas GFP–RhoA did not accumulate. Representative images from three independent experiments are shown. The corresponding movie is Movie 1. Scale bar: 5 μm. (B) Line-scan analysis performed using MetaMorph software shows the fluorescence intensities of GFP–RhoA (green) and TagRFP–RhoC (red) at the position of the line in the enlarged image of the boxed region in Fig. 1A, 2 min. Strong fluorescent signals of TagRFP–RhoC were detected at the phagocytic cups (arrowheads). (C) Quantitation of the accumulation levels of RhoC and RhoA at the phagocytic cup. Maximum fluorescence intensity values were measured at the phagocytic cup in each cell coexpressing TagRFP–RhoC and GFP–RhoA or expressing TagRFP (Mock). The fluorescence intensity of TagRFP–RhoC, GFP–RhoA or TagRFP was normalized to that of a region in the cytoplasm. Values represent the means±s.e.m. of three independent replicates (n=3; 30 phagocytic cups in more than five cells in each condition were assessed per replicate). *P<0.05 (one-way ANOVA followed by Tukey's test). (D) RAW264 macrophages were incubated with 2-μm hIgG-coated beads for 10 min before fixation and immunostained with an anti-RhoC antibody, an anti-RhoA antibody or an isotype-matched control antibody. Phase-contrast images are shown. The insets are magnifications of the boxed areas. Note the recruitment of endogenous RhoC to the phagocytic cups. Scale bars: 5 μm.

To further examine the localization of endogenous RhoC in RAW264 macrophages during FcγR-mediated phagocytosis, immunocytochemical analysis using a monoclonal antibody against RhoC was performed. First, we validated the specificity of the RhoC antibody in an immunofluorescence study (Fig. S2). RAW264 cells were first incubated with human IgG (hIgG)-coated beads for 10 min, and then fixed and immunostained with the anti-RhoC antibody. Immunofluorescence staining demonstrated that endogenous RhoC localized to the phagocytic cups extending along the surfaces of the hIgG-coated particles (Fig. 1D). In contrast, endogenous RhoA was scarcely observed at the forming phagosomes.

RhoC is transiently activated during FcγR-mediated phagocytosis

To quantitatively monitor the activation levels of endogenous RhoC during FcγR-mediated phagocytosis, we took advantage of the GTP-dependent interaction of RhoC with the Rhotekin Rho-binding domain (RBD) (Ren et al., 1999). We performed a GST pulldown assay based on GST-fused RBD to measure the amount of the GTP-bound form of RhoC present during phagocytosis in RAW264 macrophages. As shown in Fig. 2, the activation of RhoC was readily detected and reached a maximum at ∼2 min after incubation with IgG-Es. The amount of the GTP-bound form of RhoC then returned to its basal level within 30 min. Although RhoA was activated during phagocytosis of IgG-Es, the activation of RhoA occurred slightly later than did that of RhoC (Fig. S3). These findings indicate that endogenous RhoC is transiently activated during phagosome formation.

Fig. 2.

RhoC is transiently activated during phagosome formation. RAW264 macrophages were incubated with or without (0 min) IgG-Es for various times at 37°C. Cell lysates were prepared and incubated with GST or GST–RBD. The proteins associated with GST (top row, right-most lane) or GST–RBD (top row, other lanes) were pulled down by using glutathione–Sepharose beads and analyzed by western blotting for RhoC. The lane for GST is a negative control for the pulldown assay. The lane labeled GST-RBD next to GST is a positive control. The middle rows show aliquots of the total cell lysates. Ponceau S staining was used to visualize GST–RBD and GST (bottom rows). Note the increase in RhoC-GTP levels during FcγR-mediated phagocytosis. The intensity of the protein bands was measured by densitometry quantitative analysis of RhoC-GTP and is shown as mean±s.e.m. (n=3) in the corresponding bar graph. The protein band intensity of RhoC-GTP was normalized to that of total RhoC (GTP-bound plus GDP-bound forms). **P<0.01 compared to time 0 (one-way ANOVA followed by Dunnett's test).

Fig. 2.

RhoC is transiently activated during phagosome formation. RAW264 macrophages were incubated with or without (0 min) IgG-Es for various times at 37°C. Cell lysates were prepared and incubated with GST or GST–RBD. The proteins associated with GST (top row, right-most lane) or GST–RBD (top row, other lanes) were pulled down by using glutathione–Sepharose beads and analyzed by western blotting for RhoC. The lane for GST is a negative control for the pulldown assay. The lane labeled GST-RBD next to GST is a positive control. The middle rows show aliquots of the total cell lysates. Ponceau S staining was used to visualize GST–RBD and GST (bottom rows). Note the increase in RhoC-GTP levels during FcγR-mediated phagocytosis. The intensity of the protein bands was measured by densitometry quantitative analysis of RhoC-GTP and is shown as mean±s.e.m. (n=3) in the corresponding bar graph. The protein band intensity of RhoC-GTP was normalized to that of total RhoC (GTP-bound plus GDP-bound forms). **P<0.01 compared to time 0 (one-way ANOVA followed by Dunnett's test).

Expression of RhoC mutants or depletion of endogenous RhoC suppresses FcγR-mediated phagocytosis

The localization of RhoC and the transient activation of this GTPase at the early stage of phagocytosis strongly imply that RhoC is involved in phagosome formation. To test the functional contribution of RhoC to the uptake of IgG-Es, we examined the effects of RhoC mutant expression on the binding of IgG-Es to the cell surface and on their internalization into phagosomes. We transiently transfected RAW264 cells with wild-type (wt) RhoC-wt, the constitutively active mutant RhoC-G14V or the dominant-negative mutant RhoC-T19N. Quantitative assays of the binding and phagocytosis of IgG-Es were then performed in cells expressing each RhoC allele. As shown in Fig. 3, the expression of GFP–RhoC-G14V slightly decreased the binding of IgG-Es to the cells (P<0.05), whereas the binding of IgG-Es was not affected by the expression of RhoC-wt or RhoC-T19N. Importantly, the expression of either GFP–RhoC-G14V or GFP–RhoC-T19N inhibited the phagocytosis of IgG-Es, in contrast to expression of the GFP (mock) control.

Fig. 3.

Effect of constitutively active and dominant-negative RhoC mutant expression on FcγR-mediated phagocytosis and the binding of IgG-Es. To quantify FcγR-mediated phagocytosis, RAW264 macrophages transfected with GFP (mock), GFP-RhoC-wt, GFP-RhoC-G14V or GFP-RhoC-T19N were incubated with IgG-Es for 20 min at 37°C. The cells were fixed after disruption of the extracellularly exposed IgG-Es. The efficiency of phagosome formation (gray bars) was calculated based on 50 transfected cells and 50 non-transfected cells. The results are expressed as a percentage of control (non-transfected) cells. The means±s.e.m. of three independent experiments are plotted. For the binding assay, RAW264 cells transfected with each indicated construct were incubated with IgG-Es for 30 min at 4°C. After a brief washing, the cells were fixed. The efficiency of IgG-Es binding to the cells (open bars) was calculated based on 50 transfected cells and 50 non-transfected cells. The data are expressed as the means±s.e.m. of three independent experiments. *P<0.05; **P<0.01 compared to GFP-transfected cells (mock) (one-way ANOVA followed by Dunnett's test).

Fig. 3.

Effect of constitutively active and dominant-negative RhoC mutant expression on FcγR-mediated phagocytosis and the binding of IgG-Es. To quantify FcγR-mediated phagocytosis, RAW264 macrophages transfected with GFP (mock), GFP-RhoC-wt, GFP-RhoC-G14V or GFP-RhoC-T19N were incubated with IgG-Es for 20 min at 37°C. The cells were fixed after disruption of the extracellularly exposed IgG-Es. The efficiency of phagosome formation (gray bars) was calculated based on 50 transfected cells and 50 non-transfected cells. The results are expressed as a percentage of control (non-transfected) cells. The means±s.e.m. of three independent experiments are plotted. For the binding assay, RAW264 cells transfected with each indicated construct were incubated with IgG-Es for 30 min at 4°C. After a brief washing, the cells were fixed. The efficiency of IgG-Es binding to the cells (open bars) was calculated based on 50 transfected cells and 50 non-transfected cells. The data are expressed as the means±s.e.m. of three independent experiments. *P<0.05; **P<0.01 compared to GFP-transfected cells (mock) (one-way ANOVA followed by Dunnett's test).

To further substantiate the significance of RhoC GTPase in phagosome formation, we adopted the RNAi approach to knockdown endogenous RhoC in RAW264 cells. The cells were transiently transfected with plasmids coding for several short hairpin RNAs (shRNAs). As shown in Fig. 4A, the expression of each RhoC shRNA (GI557339 or GI557340) decreased RhoC protein levels, as assessed by immunostaining with the anti-RhoC antibody. We then performed a quantitative phagocytosis assay on these transfectants. The efficiency of phagocytosis in cells expressing RhoC shRNAs was compared with that in non-transfected control cells. The depletion of endogenous RhoC by shRNA inhibited phagocytosis to a degree comparable with that observed in cells expressing dominant-negative mutant RhoC-T19N (Fig. 4B and Fig. 3, respectively). Similar results were obtained upon siRNA-mediated RhoC silencing in RAW264 cells (Fig. S4). In contrast, siRNA-mediated knockdown of RhoA expression had no observable effect on phagosome formation (Fig. S5).

Fig. 4.

Effect of RhoC depletion on FcγR-mediated phagocytosis. (A) RAW264 macrophages expressing each indicated shRNA were fixed and immunostained with the antibody against RhoC (red). Cells transfected with shRNA constructs were identified by GFP fluorescence (green). Note the marked decrease of RhoC immunoreactivity in cells expressing RhoC shRNAs. Phase-contrast images are also shown. Scale bars: 5 μm. (B) The efficiency of IgG-E uptake was calculated based on 50 cells transfected with RhoC shRNA or control shRNA construct (mock) and 50 non-transfected cells. The values expressed as a percentage of control (non-transfected) cells represent the means±s.e.m. of three independent experiments. *P<0.05 compared to mock-transfected cells (one-way ANOVA followed by Dunnett's test). (C) RAW264 cells transfected with pSpCas9(BB)-2A-puro plasmid (mock) or pSpCas9(BB)-2A-puro-RhoC encoding Cas9 protein and gRNA for RhoC were diluted and selected for single clones with puromycin. Each clone was then expanded without puromycin. The knockout of RhoC protein was verified by western blotting using the antibody against RhoC. RhoA and GAPDH were used as internal controls. (D) The efficiency of IgG-E phagocytosis was calculated based on 50 cells knocked out for RhoC and 50 cells transfected with pSpCas9(BB)-2A-puro plasmid (mock) or 50 non-transfected (control) cells. The results are expressed as the mean±s.e.m. percentage compared with control cells for four independent experiments. *P<0.05 compared to mock transfected cells (one-way ANOVA followed by Dunnett's test).

Fig. 4.

Effect of RhoC depletion on FcγR-mediated phagocytosis. (A) RAW264 macrophages expressing each indicated shRNA were fixed and immunostained with the antibody against RhoC (red). Cells transfected with shRNA constructs were identified by GFP fluorescence (green). Note the marked decrease of RhoC immunoreactivity in cells expressing RhoC shRNAs. Phase-contrast images are also shown. Scale bars: 5 μm. (B) The efficiency of IgG-E uptake was calculated based on 50 cells transfected with RhoC shRNA or control shRNA construct (mock) and 50 non-transfected cells. The values expressed as a percentage of control (non-transfected) cells represent the means±s.e.m. of three independent experiments. *P<0.05 compared to mock-transfected cells (one-way ANOVA followed by Dunnett's test). (C) RAW264 cells transfected with pSpCas9(BB)-2A-puro plasmid (mock) or pSpCas9(BB)-2A-puro-RhoC encoding Cas9 protein and gRNA for RhoC were diluted and selected for single clones with puromycin. Each clone was then expanded without puromycin. The knockout of RhoC protein was verified by western blotting using the antibody against RhoC. RhoA and GAPDH were used as internal controls. (D) The efficiency of IgG-E phagocytosis was calculated based on 50 cells knocked out for RhoC and 50 cells transfected with pSpCas9(BB)-2A-puro plasmid (mock) or 50 non-transfected (control) cells. The results are expressed as the mean±s.e.m. percentage compared with control cells for four independent experiments. *P<0.05 compared to mock transfected cells (one-way ANOVA followed by Dunnett's test).

We have shown the key role of RhoC in phagosome formation through experiments where the dominant-negative mutant RhoC-T19N was overexpressed or endogenous RhoC was knocked down. To further confirm our findings and to exclude the possibility that phagocytosis was inhibited by undesired off-target effects, we generated RhoC-deficient RAW264 cells by using the CRISPR/Cas system. Two guiding RNA (gRNA) sequences were tested and shown to successfully deplete RhoC protein. As shown in Fig. 4C, two KO clones did not show any detectable RhoC protein. The expression of RhoA was not affected in these RhoC-KO cells. We also performed immunocytochemical analysis with the monoclonal antibody against RhoC. No reactivity was observed in RhoC-deficient cells (Fig. S6). RhoC-KO cells showed an increase in cell spreading area compared to that in (control) wild-type cells. Predictably, quantitative analysis of phagocytosis showed that the uptake of IgG-Es was inhibited in the RhoC-deficient cells (Fig. 4D). These results suggest that RhoC is an important component of FcγR-mediated phagocytosis.

RhoC regulates the remodeling of the actin cytoskeleton during phagosome formation

Rho GTPases such as Rac1 and Cdc42 are known to spatiotemporally remodel the actin cytoskeleton during FcγR-mediated phagocytosis (Ikeda et al., 2017; Swanson, 2008). Previous reports have shown that RhoC regulates actin-rich lamellipodial protrusion during tumor cell invasion, which requires a dynamic reorganization of the actin cytoskeleton (Donnelly et al., 2014). These facts prompted us to examine the involvement of RhoC in the remodeling of the actin cytoskeleton during phagocytosis. We first analyzed the spatiotemporal relationship between RhoC and actin dynamics in live RAW264 cells coexpressing GFP–RhoC and TagRFP–actin during FcγR-mediated phagocytosis. Time-lapse imaging showed that both proteins were colocalized and accumulated at the phagocytic cups at the same time (Fig. 5A,B; Movie 2). Subsequently, the proteins were relocated from the membranes of the newly formed phagosomes to the cytosol at almost the same time (Fig. 5A). Similar results were obtained when we used cells expressing GFP–RhoC and TagRFP–LifeAct, which labels F-actin (data not shown). To further dissect the functional localization of RhoC and actin at the phagocytic cups, RAW264 cells expressing GFP–RhoC were incubated with IgG-Es and stained with Rhodamine–phalloidin to visualize F-actin structures. Consistent with our live-cell imaging findings, GFP–RhoC was localized to F-actin-rich phagocytic cups during FcγR-mediated phagocytosis (Fig. 5C, arrows).

Fig. 5.

Confocal imaging showing the spatiotemporal relationship between RhoC and actin during phagosome formation. (A) Live RAW264 cells coexpressing GFP–RhoC (green) and TagRFP–actin (red) were allowed to interact with IgG-Es and observed by confocal laser microscopy. Phase-contrast images are also shown (top panels). The elapsed time is indicated at the top left. Note the colocalization of RhoC and actin at the phagocytic cups. Representative images from three independent experiments are shown. The corresponding movie is Movie 2. Scale bar: 5 μm. (B) Quantification of the levels of RhoC and actin accumulation at the phagocytic cup. Maximum fluorescence intensity values were measured at the phagocytic cup in each cell coexpressing GFP–RhoC and TagRFP–Actin or expressing GFP (mock). The fluorescence intensity of GFP–RhoC, TagRFP–Actin or GFP was normalized to that of a region in the cytoplasm. Values represent the means±s.e.m. of three independent replicates (n=3; 30 phagocytic cups in more than 5 cells in each condition were assessed per replicate). *P<0.05; **P<0.01 (one-way ANOVA followed by Tukey's test). (C) RAW264 macrophages transfected with GFP–RhoC (green) were incubated with IgG-Es for 10 min at 37°C, fixed and stained with Rhodamine–phalloidin (red). Consistent with the above finding, GFP–RhoC was localized to F-actin-rich phagocytic cups (arrows). Scale bar: 5 μm.

Fig. 5.

Confocal imaging showing the spatiotemporal relationship between RhoC and actin during phagosome formation. (A) Live RAW264 cells coexpressing GFP–RhoC (green) and TagRFP–actin (red) were allowed to interact with IgG-Es and observed by confocal laser microscopy. Phase-contrast images are also shown (top panels). The elapsed time is indicated at the top left. Note the colocalization of RhoC and actin at the phagocytic cups. Representative images from three independent experiments are shown. The corresponding movie is Movie 2. Scale bar: 5 μm. (B) Quantification of the levels of RhoC and actin accumulation at the phagocytic cup. Maximum fluorescence intensity values were measured at the phagocytic cup in each cell coexpressing GFP–RhoC and TagRFP–Actin or expressing GFP (mock). The fluorescence intensity of GFP–RhoC, TagRFP–Actin or GFP was normalized to that of a region in the cytoplasm. Values represent the means±s.e.m. of three independent replicates (n=3; 30 phagocytic cups in more than 5 cells in each condition were assessed per replicate). *P<0.05; **P<0.01 (one-way ANOVA followed by Tukey's test). (C) RAW264 macrophages transfected with GFP–RhoC (green) were incubated with IgG-Es for 10 min at 37°C, fixed and stained with Rhodamine–phalloidin (red). Consistent with the above finding, GFP–RhoC was localized to F-actin-rich phagocytic cups (arrows). Scale bar: 5 μm.

Fig. 3 shows that the expression of constitutively active mutant RhoC-G14V or dominant-negative mutant RhoC-T19N inhibits FcγR-mediated phagocytosis. We next addressed the role of RhoC in the regulation of actin cytoskeletal remodeling. RAW264 cells were transfected with RhoC-G14V, RhoC-wt or RhoC-T19N and then stained with rhodamine phalloidin. Confocal microscopy imaging revealed that the expression of constitutively active mutant RhoC-G14V induces an abnormally thickened layer of cortical F-actin underneath the plasma membrane (Fig. 6A, arrows), whereas the expression of RhoC-wt or dominant-negative mutant RhoC-T19N does not (Fig. 6A, arrowheads). Intriguingly, the actin-driven pseudopod extension required to form phagocytic cups was severely impaired in cells expressing the constitutively active RhoC mutant (Fig. 6B, arrows). In contrast, phagocytic cup formation was frequently observed in cells expressing RhoC-wt or the dominant-negative RhoC mutant (Fig. 6B, arrowheads). These data suggest that RhoC modulates the actin cytoskeletal remodeling required to form phagosomes during FcγR-mediated phagocytosis.

Fig. 6.

Effect of RhoC mutant expression on actin polymerization and phagocytic cup formation. (A) RAW264 cells expressing GFP–RhoC-G14V, GFP–RhoC-wt, GFP–RhoC-T19N or GFP (green) were fixed and stained with Rhodamine–phalloidin (red) to visualize F-actin. The expression of constitutively active mutant RhoC-G14V increased cortical F-actin levels (arrows), whereas the expression of RhoC-wt, the dominant-negative mutant RhoC-T19N or the GFP control had no effect on the cellular content of polymerized F-actin (arrowheads). Scale bars: 5 μm. (B) RAW264 macrophages transfected with each indicated construct were incubated with IgG-Es for 10 min at 37°C, fixed and stained with rhodamine-phalloidin. Although phagocytic cup formation was severely impaired in cells expressing GFP-RhoC-G14V (arrows), pseudopod extension along the surfaces of IgG-Es was occasionally observed in cells expressing GFP-RhoC-wt, GFP-RhoC-T19N or GFP control (arrowheads). Scale bars: 5 μm.

Fig. 6.

Effect of RhoC mutant expression on actin polymerization and phagocytic cup formation. (A) RAW264 cells expressing GFP–RhoC-G14V, GFP–RhoC-wt, GFP–RhoC-T19N or GFP (green) were fixed and stained with Rhodamine–phalloidin (red) to visualize F-actin. The expression of constitutively active mutant RhoC-G14V increased cortical F-actin levels (arrows), whereas the expression of RhoC-wt, the dominant-negative mutant RhoC-T19N or the GFP control had no effect on the cellular content of polymerized F-actin (arrowheads). Scale bars: 5 μm. (B) RAW264 macrophages transfected with each indicated construct were incubated with IgG-Es for 10 min at 37°C, fixed and stained with rhodamine-phalloidin. Although phagocytic cup formation was severely impaired in cells expressing GFP-RhoC-G14V (arrows), pseudopod extension along the surfaces of IgG-Es was occasionally observed in cells expressing GFP-RhoC-wt, GFP-RhoC-T19N or GFP control (arrowheads). Scale bars: 5 μm.

mDia1 functions as a downstream effector of RhoC and controls phagosome formation during FcγR-mediated phagocytosis

mDia1, mDia2 and mDia3 are major effectors of Rho GTPases (e.g. RhoA, RhoB and RhoC) and are involved in unbranched F-actin nucleation and elongation (Kühn and Geyer, 2014). Previous reports have shown that mDia1 specifically interacts with and is activated by RhoA, RhoB and RhoC. In contrast, mDia2 and mDia3 are activated by other Rho family members, such as Cdc42 (Young and Copeland, 2010). In macrophages, mDia1 primarily participates in CR3-dependent phagocytosis (Colucci-Guyon et al., 2005). Interestingly, mDia1 has been reported to be required for both FcγR- and CR3-mediated phagocytosis in neutrophils (Shi et al., 2009). Therefore, we examined the possible role of mDia1 as a downstream effector of RhoC in regulating FcγR-mediated phagocytosis in macrophages. Time-lapse observations of live cells coexpressing GFP–mDia1 and TagRFP–RhoC showed that mDia1 and RhoC are distributed throughout the cytosol before the onset of phagocytosis. After the binding of IgG-Es to the cell, both proteins accumulated at the same time and were colocalized at the phagocytic cups (Fig. 7A; Movie 3). Line-scan analysis found that the RhoC and mDia1 fluorescence intensities were correlated (Fig. 7B). Importantly, the recruitment of mDia1 to phagocytic cup formation sites was enhanced by the coexpression of mDia1 with RhoC (Fig. 7A,C,D; Fig. S7). In contrast, the coexpression of mDia1 with RhoA did not promote the relocation of mDia1 to the phagocytic cups. After closure of the phagocytic cups, mDia1 and RhoC dissociated simultaneously from the nascent phagosomes.

Fig. 7.

Time-lapse images showing localization of mDia1 and RhoC during FcγR-mediated phagocytosis. (A) Live RAW264 cells coexpressing GFP–mDia1 (green) and TagRFP–RhoC (red) were allowed to contact IgG-Es and observed by confocal laser microscopy. Note the colocalization of mDia1 and RhoC at the phagocytic cups. The corresponding movie is Movie 3. Scale bar: 5 μm. (B) A line-scan analysis performed with MetaMorph software shows the fluorescence intensities of GFP–mDia1 (green) and TagRFP–RhoC (red) at the position of the line in the enlarged image of the boxed region (Fig. 7A, 3 min). (C) RAW264 macrophages coexpressing TagRFP–mDia1 (red) and GFP–RhoA (green) were incubated with IgG-Es and observed by confocal laser microscopy. Scale bar: 5 μm. (D) Quantification of the recruitment levels of mDia1 to the phagocytic cup. Maximum TagRFP–mDia1 fluorescence intensity values were measured at the phagocytic cup in cells coexpressing GFP–RhoC, GFP–RhoA or GFP. The fluorescence intensity of TagRFP–mDia1 was normalized to that of a region in the cytoplasm. Values represent the means±s.e.m. of three independent replicates (n=3; 30 phagocytic cups in more than five cells in each condition were assessed per replicate). *P<0.05 (one-way ANOVA followed by Tukey's test).

Fig. 7.

Time-lapse images showing localization of mDia1 and RhoC during FcγR-mediated phagocytosis. (A) Live RAW264 cells coexpressing GFP–mDia1 (green) and TagRFP–RhoC (red) were allowed to contact IgG-Es and observed by confocal laser microscopy. Note the colocalization of mDia1 and RhoC at the phagocytic cups. The corresponding movie is Movie 3. Scale bar: 5 μm. (B) A line-scan analysis performed with MetaMorph software shows the fluorescence intensities of GFP–mDia1 (green) and TagRFP–RhoC (red) at the position of the line in the enlarged image of the boxed region (Fig. 7A, 3 min). (C) RAW264 macrophages coexpressing TagRFP–mDia1 (red) and GFP–RhoA (green) were incubated with IgG-Es and observed by confocal laser microscopy. Scale bar: 5 μm. (D) Quantification of the recruitment levels of mDia1 to the phagocytic cup. Maximum TagRFP–mDia1 fluorescence intensity values were measured at the phagocytic cup in cells coexpressing GFP–RhoC, GFP–RhoA or GFP. The fluorescence intensity of TagRFP–mDia1 was normalized to that of a region in the cytoplasm. Values represent the means±s.e.m. of three independent replicates (n=3; 30 phagocytic cups in more than five cells in each condition were assessed per replicate). *P<0.05 (one-way ANOVA followed by Tukey's test).

Based on the above data, we validated the effects of mDia1 and RhoC expression on FcγR-mediated phagocytosis and the binding of IgG-Es. In a quantitative assay of phagocytosis, the expression of GFP–mDia1-wt alone had no effect on the uptake of IgG-Es (Fig. 8A). Importantly, both the coexpression of mDia1-wt with the constitutively active mutant RhoC-G14V and the expression of mDia1-ΔN3 (a constitutively active mutant of mDia1) drastically inhibited phagosome formation. In addition, the efficiency of particle binding was slightly reduced in cells coexpressing mDia1-wt with RhoC-G14V or expressing mDia1-ΔN3; in contrast, the expression of mDia1-wt alone did not affect the binding of IgG-Es to the cell. Confocal microscopic imaging showed that the expression of the constitutively active mutant of mDia1 promotes a thickened layer of cortical F-actin (Fig. S8, arrows), whereas the expression of mDia1-wt does not (Fig. S8, arrowheads). We next addressed the role of endogenous mDia1 in FcγR-mediated phagocytosis by using an siRNA approach. As shown in Fig. 8B, the transfection of mDia1 siRNA decreased in its expression in RAW264 macrophages. Notably, the efficiency of IgG-E uptake was reduced in mDia1-knockdown cells, whereas that of IgG-Es binding was not affected (Fig. 8C). Moreover, treatment with SMIFH2, an inhibitor of formin-mediated actin assembly, also inhibited phagosome formation (Fig. 8D). Taken together, these findings indicate that mDia1 – a downstream effector of RhoC – is an important component of FcγR-mediated phagocytosis and regulates phagosome formation.

Fig. 8.

Effect of mDia1 expression and downregulation on FcγR-mediated phagocytosis and the binding of IgG-Es. (A) RAW264 macrophages expressing the GFP–mDia1 construct and/or TagRFP–RhoC-G14V were incubated with IgG-Es for 20 min at 37°C. The efficiency of phagosome formation (gray bars) and of IgG-E binding (open bars) was calculated based on 50 transfected cells and 50 non-transfected cells. Cdc42-G12V (constitutively active mutant) was used as a positive control for the assay. The results are expressed as a percentage of that in control (non-transfected) cells. The data represent the means±s.e.m. of three independent experiments. *P<0.05; ***P<0.001 versus corresponding mock-transfected cells. ##P<0.01; ###P<0.001 versus corresponding cells transfected with mDia1-wt. P<0.05 versus corresponding cells transfected with Cdc42-G12V. P<0.05 versus corresponding cells transfected with mDia1-wt and Cdc42-G12V (one-way ANOVA followed by Tukey's test). (B) RAW264 cells were transfected with mDia1 siRNA or mock siRNA. After 48 h, the cells were collected. The knockdown of mDia1 protein was verified by western blotting using the antibody specific for mDia1. mDia2 levels were used as an inter2nal control. (C) The efficiency of IgG-Es phagocytosis (gray bars) and of IgG-E binding to the cells (open bars) was calculated based on 50 cells transfected with mDia1 siRNA and 50 cells transfected with mock siRNA or 50 non-transfected (control) cells. The results are expressed as a percentage of that in control (non-transfected) cells. The means±s.e.m. of four independent experiments are plotted. **P<0.01 compared to cells transfected with mock siRNA (Student's t-test). (D) The efficiency of IgG-E phagocytosis (gray bars) and of IgG-E binding to the cells (open bars) was calculated based on 50 cells treated with 10 μM SMIFH2 and 50 cells treated with DMSO (mock) or 50 untreated (control) cells. The results are expressed as a percentage of that in control (untreated) cells. The means±s.e.m. of four independent experiments are plotted. **P<0.01 compared to cells treated with DMSO (Student's t-test).

Fig. 8.

Effect of mDia1 expression and downregulation on FcγR-mediated phagocytosis and the binding of IgG-Es. (A) RAW264 macrophages expressing the GFP–mDia1 construct and/or TagRFP–RhoC-G14V were incubated with IgG-Es for 20 min at 37°C. The efficiency of phagosome formation (gray bars) and of IgG-E binding (open bars) was calculated based on 50 transfected cells and 50 non-transfected cells. Cdc42-G12V (constitutively active mutant) was used as a positive control for the assay. The results are expressed as a percentage of that in control (non-transfected) cells. The data represent the means±s.e.m. of three independent experiments. *P<0.05; ***P<0.001 versus corresponding mock-transfected cells. ##P<0.01; ###P<0.001 versus corresponding cells transfected with mDia1-wt. P<0.05 versus corresponding cells transfected with Cdc42-G12V. P<0.05 versus corresponding cells transfected with mDia1-wt and Cdc42-G12V (one-way ANOVA followed by Tukey's test). (B) RAW264 cells were transfected with mDia1 siRNA or mock siRNA. After 48 h, the cells were collected. The knockdown of mDia1 protein was verified by western blotting using the antibody specific for mDia1. mDia2 levels were used as an inter2nal control. (C) The efficiency of IgG-Es phagocytosis (gray bars) and of IgG-E binding to the cells (open bars) was calculated based on 50 cells transfected with mDia1 siRNA and 50 cells transfected with mock siRNA or 50 non-transfected (control) cells. The results are expressed as a percentage of that in control (non-transfected) cells. The means±s.e.m. of four independent experiments are plotted. **P<0.01 compared to cells transfected with mock siRNA (Student's t-test). (D) The efficiency of IgG-E phagocytosis (gray bars) and of IgG-E binding to the cells (open bars) was calculated based on 50 cells treated with 10 μM SMIFH2 and 50 cells treated with DMSO (mock) or 50 untreated (control) cells. The results are expressed as a percentage of that in control (untreated) cells. The means±s.e.m. of four independent experiments are plotted. **P<0.01 compared to cells treated with DMSO (Student's t-test).

DISCUSSION

The present study provides the first evidence that RhoC is involved in FcγR-mediated phagocytosis. Our live-cell imaging and immunocytochemical analysis showed that RhoC is recruited to the membranes of phagocytic cups and then dissociates from the membranes of nascent phagosomes. Importantly, our RhoC-GTP pulldown assay demonstrated that RhoC is transiently activated during phagosome formation. Furthermore, we found that the expression of constitutively active mutant RhoC-G14V or dominant-negative mutant RhoC-T19N suppresses FcγR-mediated phagocytosis. We therefore postulate that the activation–inactivation cycling of RhoC is required for the uptake of IgG-opsonized particles. In contrast to RhoC, RhoA did not accumulate at the phagocytic cups. Moreover, the silencing of RhoA had no effect on phagosome formation. These data suggest that RhoA is not involved in FcγR-dependent phagosome formation. Intriguingly, we found that both RhoA and RhoC are activated during phagocytosis. It is noteworthy that the activation of RhoA occurs slightly later than that of RhoC. Previous reports have shown that RhoA is activated during FcγR-mediated phagocytosis and regulates superoxide formation (Kim et al., 2004; Li et al., 2012). Therefore, RhoA may play a major role in the regulation of superoxide formation during FcγR-mediated phagocytosis. RhoB activation has been reported as being required for mannose receptor-mediated phagosome formation (Zhang et al., 2005). However, the precise localization of RhoB during FcγR-mediated phagocytosis remains unclear. Investigating the role of RhoB in phagocytosis should be the subject of further research.

The functional molecule regulating the spatiotemporal translocation of RhoC to the phagocytic cups remains to be identified. Our confocal microscopy observations revealed that RhoC, but not its close homolog RhoA, accumulates in the membranes of phagocytic cups and readily dissociates from the internalized phagosomes. Furthermore, the pulldown assay for RhoC-GTP demonstrated that RhoC is predominantly activated during phagosome formation. These results imply that specific guanine nucleotide exchange factors (GEFs) and GTPase-activating proteins (GAPs) for RhoC are involved in FcγR-mediated phagocytosis. However, these upstream regulators of RhoC in the process of phagosome formation are currently unknown. Further studies should identify the specific GEFs and GAPs for RhoC to better understand the molecular details of the signal transduction pathway during phagosome formation. Recently, Patel et al. have reported that RhoC shows a higher degree of membrane association than does RhoA (Patel et al., 2016). In their study, these authors demonstrated that arginine 188 of RhoC promotes membrane binding. In contrast with RhoC, RhoA has a serine residue at position 188 in place of an arginine residue. During phagosome formation, protein kinase A (PKA) is targeted to nascent phagosomes (Pryzwansky et al., 1998). Importantly, RhoA phosphorylation on serine 188 by PKA has been shown to induce RhoA translocation from membranes to the cytosol (Forget et al., 2002; Lang et al., 1996). PKA is thus thought to phosphorylate serine 188 of RhoA in the process of FcγR-mediated phagocytosis, thereby inhibiting RhoA recruitment to the phagocytic cups.

The local remodeling of the actin cytoskeleton during phagosome formation is regulated by the recruitment and activation of the actin-nucleating activity of Arp2/3. The activation of Cdc42 and Rac1 has been shown to stimulate the Arp2/3 complex via WASP and WAVE family proteins to form a branched F-actin network at the phagocytic cup (Cox et al., 1997; Lorenzi et al., 2000; Massol et al., 1998; May et al., 2000; Park and Cox, 2009; Tsuboi and Meerloo, 2007). Interestingly, the GTP-bound form of Cdc42 also functions as an upstream regulator of the formin protein FMNL1, which promotes the formation of unbranched actin filaments during FcγR-mediated phagocytosis (Otomo et al., 2005; Romero et al., 2004; Seth et al., 2006). Until now, the modulation of unbranched actin polymerization has been comparatively less studied than the regulation of the branched F-actin network at the phagocytic cup in FcγR-mediated phagocytosis. We found that the actin-driven pseudopod extension required to form phagocytic cups is severely inhibited in cells expressing the constitutively active mutant RhoC-G14V, which induces an abnormal cortical F-actin layer and potentially promotes the polymerization of unbranched actin filaments. Moreover, the expression of RhoC-G14V slightly reduces the binding of IgG-Es to the cell. These data suggest that the remodeling of unbranched cortical F-actin underneath the plasma membrane is required for the process of particle binding and subsequent phagocytic cup formation. At present, the significance of cortical F-actin remodeling during the initial stage of phagocytosis remains unknown. This remodeling may release monomeric G-actin from cortical F-actin for incorporation into new filaments to form phagocytic cups. Alternatively, in the process of phagocytic target capture, the remodeling of cortical F-actin may promote the mobility of FcγRs for particle binding (Jaumouille and Grinstein, 2011; Mao et al., 2009).

It is important to determine both the downstream effector of RhoC during FcγR-mediated phagocytosis and how RhoC mechanistically regulates phagosome formation. Our time-lapse imaging demonstrated that RhoC and mDia1, one of the major effectors of Rho GTPases, are colocalized at the phagocytic cups. Moreover, a quantitative assay of phagocytosis found that both the coexpression of mDia1-wt with the GTP-locked mutant RhoC-G14V and the expression of activated mutant mDia1-ΔN3 result in a remarkable decrease in the rate of IgG-Es uptake. A previous report has indicated that the expression of activated mutant mDia1-ΔN3 induces a dramatic increase in the basal level of polymerized F-actin (Vicente-Manzanares et al., 2003). In this study, we revealed that the expression of constitutively active mutant RhoC-G14V, which activates mDia1, facilitates the formation of an abnormal F-actin layer underneath the plasma membrane. Similar to what was seen with RhoC-G14V, mDia1-ΔN3 expression induced an abnormal F-actin layer in RAW264 macrophages, whereas the expression of mDia1-wt did not. Collectively, these findings indicate that mDia1 is activated at the phagocytic cups in a RhoC-GTP-dependent manner and that it functions as a downstream effector of RhoC during FcγR-mediated phagocytosis. We observed that RhoC depletion increases the cell spreading area in RAW264 macrophages. A previous report has shown that RhoC knockdown facilitates cell spreading and induces Rac1 activation around the periphery in the lamellipodia of PC3 cells (Vega et al., 2011). Recently, we have demonstrated that the expression of a constitutively active Rac1 mutant suppresses FcγR-mediated phagocytosis (Ikeda et al., 2017). RhoC may also modulate Rac1 activity, thereby regulating phagosome formation.

In conclusion, although RhoC may have multiple downstream effectors involved in FcγR-mediated phagocytosis, our study emphasizes the importance of RhoC in the modulation of phagosome formation by activating mDia1, which can regulate the remodeling of cortical F-actin.

MATERIALS AND METHODS

Reagents

Bovine serum albumin (BSA) and Dulbecco's modified Eagle's medium (DMEM) were purchased from Sigma (St. Louis, MO). Fetal bovine serum (FBS) was obtained from BioSolutions International (Melbourne, Australia). Rabbit monoclonal anti-RhoC antibody (D40E4) (3430, Cell Signaling Technology, Danvers, MA), mouse monoclonal anti-RhoA antibody (ARH04, Cytoskeleton, Denver, CO), mouse monoclonal anti-mDia1 antibody (clone 51) (610849, BD Biosciences, San Jose, CA), rabbit polyclonal anti-mDia2 antibody (C-terminal region) (DP3491, ECM Biosciences, Versailles, KY), mouse monoclonal anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) antibody (AM4300, Ambion, Huntingdon, UK), rabbit anti-sheep erythrocyte IgG (ICN55806, Organo Teknika-Cappel, Durham, NC), goat anti-rabbit-IgG conjugated to Alexa Fluor 594 (A11037, Molecular Probes, Eugene, OR), goat anti-mouse IgG conjugated to Alexa Fluor 488 (A11029, Molecular Probes, Eugene, OR), anti-mouse- and anti-rabbit-IgG conjugated to horseradish peroxidase (HRP) (W4021 and W4011, Promega, Madison, WI), purified human IgG (I4506, Sigma, St. Louis, MO), monoclonal rabbit IgG isotype control (DA1E) (3900, Cell Signaling Technology, Danvers, MA), monoclonal mouse IgG isotype control (G3A1) (5415, Cell Signaling Technology, Danvers, MA), Rhodamine-conjugated phalloidin (Molecular Probes, Eugene, OR), SMIFH2 (Merck Millipore, Nottingham, UK) and 2-μm-diameter polystyrene microspheres (Polysciences, PA) were commercially obtained. Other reagents were purchased from Wako Pure Chemicals (Osaka, Japan) or Nakalai Tesque (Kyoto, Japan) unless otherwise indicated.

Cell culture

Mouse macrophage RAW264 cells (RCB0535, Tsukuba, Japan) were cultured in DMEM supplemented with 10% heat-inactivated FBS, 100 U/ml penicillin and 100 μg/ml streptomycin, as described in the manuals of the cell line bank (ATCC , TIB-71) (growth medium). Before the experiments, the culture medium was replaced with Ringer's buffer (RB) consisting of 155 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 2 mM Na2HPO4, 10 mM D-glucose, 10 mM HEPES-NaOH (pH 7.2) and 0.5 mg/ml BSA.

DNA constructs and transfection

The full-length cDNA coding region of mouse RhoC was amplified by PCR. The fragment was cloned into the EcoRI and BamHI restriction sites of the pEGFP-C1 vector (Clontech, Palo Alto, CA). pEGFP-RhoC-T19N (dominant-negative mutant) and pEGFP-RhoC-G14V (constitutively active mutant) were generated by using the QuikChange II site-directed mutagenesis kit (Stratagene, La Jolla, CA). pTagRFP-RhoC, pTagRFP-RhoC-T19N and pTagRFP-RhoC-G14V were generated by the replacement of EGFP with TagRFP. pTagRFP-Actin was purchased from Evrogen (Moscow, Russia). pcDNA3-EGFP-RhoA-wt was Addgene plasmid #12965 (deposited by Gary Bokoch; Subauste et al., 2000). YFP-Cdc42 (V12) was kindly provided by Dr. Joel A. Swanson (University of Michigan, Ann Arbor, MI). pEGFP-Cdc42-G12V (constitutively active mutant) was generated by replacing EYFP with EGFP. The full-length cDNA coding region of mouse Dia1 (mDia1) and the region encoding the constitutively activate mutant (amino acids 543–1182) of mouse Dia1 (mDia1-ΔN3) were amplified by PCR. The fragments were cloned into the BglII and SalI restriction sites of the pEGFP-C1 and pTagRFP-C (Evrogen) vectors. All constructs were verified by sequencing prior to use. Transfection of the plasmids into RAW264 cells was performed by using the Neon Transfection System (Invitrogen, Carlsbad, CA) according to the manufacturer's instructions. The transfected cells were seeded onto 25-mm coverslips and maintained in growth medium. Experiments were performed 12–24 h after transfection.

Silencing of endogenous RhoC through shRNA

Short hairpin RNAs (shRNAs) were used to knockdown RhoC expression in RAW264 cells. RhoC shRNAs cloned into the pGFP-V-RS vector were purchased from Origene (Rockville, MD). Two different RhoC shRNAs (GI557339 and GI557340) and a control shRNA (TR30008, negative control shRNA pGFP-V-RS non-effective tGFP plasmid) were used for transfection. The transfectants were cultured for 6 days, tested for the expression of RhoC through immunocytochemistry and used for the phagocytosis assay.

siRNA transfection

RAW264 cells were transfected with siRNA duplexes specific for RhoC, RhoA, mDia1 or bock siRNA (MISSION siRNA Universal Negative Control, Sigma) by using Viromer Blue (Lipocalyx Halle, Germany) according to the manufacturer's instructions. Transfection of RhoC siRNA was performed twice. At 48 h after the initial transfection with RhoC siRNA or mock siRNA, the cells were transfected again with the same siRNA and maintained in growth medium for 72 h. The cells were then analyzed by western blotting and used for the phagocytosis assay. The cells transfected with RhoA, mDia1 or mock siRNA were cultured for 48 h and used for subsequent experiments. The siRNA target sequences were: RhoC siRNA 1, 5′-CAUCCUCAUGUGUUUCUCCAUUGAC-3′; RhoC siRNA 2, 5′-AGGAUCAGUGCCUUUGGCUACCUCG-3′; RhoA siRNA 1, 5′-GACAUGCUUGCUCAUAGUCUUC-3′ (Guilluy et al., 2011); RhoA siRNA 2, 5′-GAAGUCAAGCAUUUCUGUC-3′ (Yoon et al., 2016); and mDia1 siRNA, 5′-GCGACGGCGGCAAACAUAAGAAAUU-3′ (Yamana et al., 2006).

CRISPR/Cas-mediated knockout of RhoC

The pSpCas9(BB)-2A-puro (PX459) plasmid was Addgene plasmid #48139 (deposited by Feng Zhang; Ran et al., 2013). For knockout of RhoC in RAW264 macrophages, the following gRNA sequences were tested: 5′-GGCTGCGATCCGAAAGAAGC-3′ and 5′-CTATATAGCCGACATCGAAG-3′. After ligation of the synthesized sequences into pSpCas9(BB)-2A-puro, the pSpCas9(BB)-2A-puro-RhoC constructs were verified by sequencing. Plasmid transfection was performed by using the Neon Transfection System. At 12 h after transfection, the cells were selected with 5 μg/ml puromycin for 24 h. Subsequently, a limiting dilution of the surviving cells was made. The resultant single colonies were then expanded without puromycin. RhoC-knockout clones were confirmed by both DNA sequencing and western blotting.

Protein expression and purification

pGEX-2T-RBD encoding the GST-Rho-binding domain (RBD) (amino acids 7–89) of Rhotekin was Addgene plasmid #15247 (deposited by Martin Schwartz; Ren et al., 1999). We expressed GST or GST–RBD protein in Escherichia coli [BL21(DE3)] using methods similar to those described previously (Egami et al., 2015). Cells grown in LB medium were incubated in the presence of 0.1 mM isopropyl-1-thio-β-D-galactopyranoside (15 h at 25°C). All subsequent purification steps were performed at 4°C. After centrifugation, the resulting cell pellets were resuspended in a buffer (20 mM Tris-HCl pH 7.5, 1 mM EDTA, 1 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride, 50 units/ml aprotinin, 2 μg/ml leupeptin and 2 μg/ml pepstatin A) and subjected to sonication. The cell debris was removed by centrifugation, and the resultant supernatant was used as an E. coli lysate. Purification of the GST-fusion proteins to near homogeneity was achieved by using glutathione-Sepharose 4B affinity chromatography (GE Healthcare, Piscataway, NJ). The purity of the samples was at least 80%, as confirmed by Coomassie Brilliant Blue staining of SDS-PAGE gels.

GST pulldown assay and western blotting

RAW264 cells were washed with ice-cold phosphate-buffer saline (PBS) and suspended in lysis buffer containing 25 mM Tris-HCl pH 7.2, 150 mM NaCl, 5 mM MgCl2, 1% NP-40, 5% glycerol and protease inhibitor cocktail (Nacalai Tesque, Kyoto, Japan). The cell lysates were briefly sonicated at 4°C and separated from the pellets after centrifugation at 12,100 g for 15 min. Protein concentrations were estimated with the BCA protein assay reagent. Glutathione–Sepharose beads coupled to GST or GST–RBD were incubated for 2 h at 4°C with 500 μg of the cell lysates. After the beads were washed four times with lysis buffer, the proteins bound to the beads were analyzed on 12.5% SDS-PAGE gels followed by western blotting. The samples were subjected to SDS-PAGE and transferred to a polyvinylidene difluoride (PVDF) membrane (Bio-Rad, Richmond, CA). Western blotting was conducted using the ECL Prime Western Blotting detection system (GE Healthcare, Piscataway, NJ). The membrane was blocked with 5% nonfat dried milk in PBS containing 0.1% Tween 20 for 30 min at room temperature and probed with anti-RhoC antibody (1:2000), anti-RhoA antibody (1:1000) or anti-GAPDH antibody (1:10,000) at 4°C overnight. After washing, the membrane was incubated with HRP-conjugated anti-rabbit-IgG or anti-mouse-IgG secondary antibody (dilution 1:10,000) for 2 h at room temperature, developed using an ECL Prime regent and exposed to Hyperfilm (GE Healthcare, Piscataway, NJ). GST and GST–RBD was stained with Ponceau S. The intensity of the protein bands was measured using Photoshop CS software.

Phagocytosis assay

Sheep erythrocytes were opsonized with rabbit anti-sheep erythrocyte IgG (1:200, Organo Teknika-Cappel) and resuspended in PBS as described previously (Araki et al., 1996). For the quantitative assay of phagocytosis, IgG-opsonized erythrocytes (IgG-Es) were added to adherent RAW264 macrophages. After 20 min of incubation with IgG-Es at 37°C, the cells on the coverslips were dipped into distilled water for 20 s to disrupt the extracellularly exposed IgG-Es, then fixed with 4% paraformaldehyde and 0.1% glutaraldehyde for 15 min. The number of internalized IgG-Es was counted in 50 cells randomly chosen under phase-contrast and fluorescence microscopy. The phagocytic index (i.e. the mean number of IgG-Es taken up per cell) was then calculated. The index obtained for the transfected cells was divided by the index obtained for the non-transfected (control) cells and expressed as a percentage of that found for the control cells. For the binding assay, RAW264 cells were incubated with IgG-Es for 30 min at 4°C, briefly washed in ice-cold PBS to remove the unbound IgG-Es and fixed. The number of cell-bound IgG-Es was then counted, and the binding index (i.e. the mean number of bound IgG-Es per cell) was calculated. The binding index was expressed as a percentage of that in non-transfected (control) cells. For immunocytochemistry, 106 polystyrene beads were washed in PBS and incubated in ∼10 mg/ml human IgG (hIgG) for 1 h at 37°C. After washing with PBS, hIgG-opsonized beads were added to the adherent RAW264 cells. The cells were then incubated for 10 min at 37°C and fixed with 4% paraformaldehyde.

Live-cell imaging and data analysis

RAW264 macrophages were cultured onto 25-mm circular coverslips. Each coverslip was assembled into an RB-filled chamber on the thermocontrolled stage (Tokai Hit, Shizuoka, Japan). Phase-contrast and fluorescence images of live cells were sequentially acquired with an Axio Observer Z1 inverted microscope equipped with a laser scanning unit (LSM700, Zeiss) and Plan-Apochromat 63× NA 1.4 lens under the control of ZEN2009 software (Zeiss), as previously described (Egami et al., 2015). Time-lapse images of phase-contrast and fluorescence microscopy were taken at 15 s intervals and assembled into QuickTime movies. At least three examples were observed in each experiment. MetaMorph 7.8 and Photoshop CS5 software were used to process images subsequent to data acquisition. To quantify the accumulation levels of the proteins, the maximal values of TagRFP and/or GFP signal intensity at the phagocytic cup were measured by using MetaMorph software. The intensity of the TagRFP and/or GFP signal was normalized to the fluorescence intensity of a region in the cytoplasm. The mean±s.e.m. values for three independent experiments were plotted.

Immunostaining

For immunostaining with anti-RhoC antibody, RAW264 cells grown on coverslips were fixed in 4% paraformaldehyde for 15 min, rinsed three times with PBS and permeabilized with 0.1% Triton X-100 in PBS for 2 min. Fixed samples were blocked with 1% BSA in PBS, then incubated twice for 1 h, first with anti-RhoC antibody (diluted 1:500 in 1% BSA in PBS) and then with goat anti-rabbit-IgG conjugated to Alexa Fluor 594 (1:500). To visualize F-actin, cells were fixed in 4% paraformaldehyde and 0.1% glutaraldehyde for 15 min, briefly rinsed with PBS and permeabilized with 0.1% Triton X-100 in PBS for 2 min. Specimens were then stained with Rhodamine-conjugated phalloidin (1.25 U/ml final concentration) for 30 min.

Statistical analysis

Two-tailed Student's t-tests or one-way ANOVA followed by a Tukey's test or Dunnett's test were performed. All P-values were considered significant at P<0.05.

Acknowledgements

The authors would like to thank Dr. Katsuya Miyake for his helpful discussion, as well as Mr. Kazuhiro Yokoi and Ms. Yukiko Iwabu for their skillful assistance.

Footnotes

Author contributions

Conceptualization: Y.E., N.A.; Methodology: Y.E., K.K.; Software: Y.E.; Validation: Y.E., N.A.; Formal analysis: Y.E.; Investigation: Y.E.; Resources: Y.E., K.K., N.A.; Data curation: Y.E., N.A.; Writing - original draft: Y.E., N.A.; Writing - review & editing: Y.E., N.A.; Visualization: Y.E.; Supervision: Y.E., N.A.; Project administration: Y.E., N.A.; Funding acquisition: Y.E., K.K., N.A.

Funding

This study was supported by the Japan Society for the Promotion of Science (JSPS) (KAKENHI grant number 16K08468 to Y.E., and was also supported in part by JSPS KAKENHI grant number 26670094 to N.A. and grant number 26860136 to K.K.). The work was partially funded by Kagawa University Scientific Research Encourages Research Funding 2015.

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Competing interests

The authors declare no competing or financial interests.

Supplementary information