The poxvirus A39R protein is a member of the semaphorin family previously reported to bind plexin C1. We show that, in the mouse, plexin C1 is expressed on dendritic cells (DCs) and neutrophils and is the only receptor for A39R on these cells. The biological effects of a recombinant form of A39R were examined in vitro on mouse DCs derived from wild-type or plexin C1−/− mice. A39R binding to plexin C1 on DCs inhibited integrin-mediated adhesion and spreading in vitro. This phenomenon was accompanied by a decrease in integrin signaling, measured by focal adhesion kinase phosphorylation, and a rearrangement of the actin cytoskeleton, without inducing DC maturation or affecting their viability. The A39R effect on DC adhesion was blocked by a specific inhibitor of cofilin phosphorylation, suggesting that the regulation of F-actin turnover by plexin C1 was essential to induce cellular retraction. Furthermore, A39R binding to plexin C1 inhibited chemokine-induced migration of DCs in vitro, suggesting that plexins and semaphorins could be involved in the regulation of leukocyte movement.

Cytoskeleton is required for cells to move, polarize, change shape, engulf particles, or interact with other cells. Different families of proteins inducing cytoskeleton remodeling have been identified. Among these are semaphorins, which belong to a growing family of soluble and membrane bound proteins that can provide either repulsive or attractive guidance signals on a wide range of neurons (1, 2, 3, 4, 5). Developmental defects in several loss of function semaphorin mutants have revealed the crucial importance of these proteins in the guidance or migration of several cell types like neurons or endothelial cells. Semaphorins share a conserved, 500 amino acid residue domain near their amino terminus called the Sema domain. Currently, 25 members have been identified from invertebrates to human (2). A semaphorin homologue called A39R has also been identified as being encoded within the genomes of several poxviruses like vaccinia (6).

Two families of semaphorin receptors have been identified: plexins and neuropilins (1, 2, 3, 4, 5). Plexins are type I transmembrane proteins that also contain a Sema domain in their extracellular part and an intracellular domain called the SP (sex and plexins) domain (7, 8). They are divided into four subfamilies, plexin A to plexin D, according to the structure of their extracellular domain. Semaphorins signal through direct binding to plexins alone, or in some cases, in combination with neuropilins. The most remarkable consequence of the engagement of plexins by semaphorins in neurons is the local rearrangement of the actin cytoskeleton (3, 5, 9) through the regulation of the activity of small GTPases of the Rho and Rac families (9, 10) and their downstream effector actin-binding protein cofilin (11). Cofilin phosphorylation by Lin-11-Isl-1-Mec-3 kinase in response to semaphorin inhibits cofilin ability to bind and depolymerize pointed ends of F-actin, which inhibits F-actin turnover in neuron growth cones. Moreover, three recent reports indicate that integrin function is regulated by semaphorins (12, 13, 14) by an as yet unknown mechanism.

Emerging evidence points also to a role for semaphorins and plexins in the immune system (for a review, see Ref. (15). However, these two families of proteins have been studied independently and no semaphorin-plexin pair has been identified in the immune system. Some semaphorins have been shown to play roles in immune regulation, but they seem to function through unique receptors that are not used in the nervous system. Several plexins are also expressed by leukocytes, but their cellular ligands as well as the consequence of their engagement on cellular physiology and actin-based cytoskeleton in particular are unknown.

We report that plexin C1 is expressed on mouse dendritic cells (DCs)2 and neutrophils and is the only receptor for the viral semaphorin homologue A39R on these cells. We investigated the consequences of plexin C1 engagement by a recombinant form of A39R on mouse bone marrow-derived DCs. We found that in vitro binding of A39R to plexin C1 inhibited integrin-mediated DC adhesion and spread without affecting their maturation or viability. A39R induced a decrease of LPS-induced focal adhesion kinase (FAK) phosphorylation in DCs and the rearrangement of the actin cytoskeleton. Moreover, we found that a synthetic cell-permeable peptide containing a cofilin phosphorylation site suppressed A39R-mediated effects on DC adhesion. Finally, A39R treatment inhibited DC ability to migrate to various chemokines in vitro, indicating a potential role for semaphorins and plexins in the regulation of leukocyte movement.

C57BL/6 mice were obtained from The Jackson Laboratory and used at the age of 6–12 wk. The generation of plexin C1−/− mice has already been described (12). Plexin C1−/− mice were bred in house. All of the experimental procedures used in this study were approved by a local committee according to federal guidelines. SFHA39R (shorty-Flag-histidine or A39R) was expressed from Chinese hamster ovary cells and purified as described (7). All proteins were tested for endotoxin contents using a Limulus assay (Whittaker M.A. Bioproducts) and were shown to be <3 EU/mg. A39R was used at 1 μg/ml in every in vitro assay unless otherwise mentioned. Blocking Abs against β1 integrin (clone HA2/5) and β2 integrin (clone GAME-46) were obtained from BD Immunocytometry Systems) and used at 10 μg/ml. Isotype control (hamster IgM and rat IgG1; BD Biosciences) were used as controls. Synthetic peptides were obtained from Synpep. S3 and RV peptide sequences have been described elsewhere (11). S3 peptide contains the phosphorylation site of cofilin and the cell-permeable sequence motif of penetratin. Cofilin phosphorylation has been shown to be efficiently and specifically inhibited in cells cultured in the presence of this peptide. As a control, the RV peptide containing the same penetratin domain and the reverse sequence of the cofilin phosphorylation domain were used.

Bone marrow cells were isolated by flushing femurs with complete medium supplemented with 2% heat-inactivated FBS (Invitrogen Life Technologies). RBC were lysed. The cells were then resuspended in culture medium consisting of McCoy’s medium supplemented with essential and nonessential amino acids, 1 mmol/L sodium pyruvate, 2.5 mmol/L HEPES buffer (pH 7.4), vitamins, 5.5 × 10−5 mol/L 2-ME, 100 U/ml penicillin, 100 μg/ml streptomycin, 0.3 mg/ml l-glutamine, and 10% FBS (all medium reagents from Invitrogen Life Technologies). Bone marrow cells were cultured as previously described (16) for 9 days at 1 × 106 cells/ml, in tissue culture flasks (BD Falcon; BD Biosciences Discovery Labware) in culture medium supplemented with 200 ng/ml recombinant human Fms-like tyrosine kinase-3 ligand (Flt3L) (Chinese hamster ovary cell-derived; Amgen). After primary culture with Flt3L, DCs were subsequently cultured in IMDM (Invitrogen Life Technologies) supplemented with nonessential amino acids, 1 mmol/L sodium pyruvate, 2.5 mmol/L HEPES buffer (pH 7.4), 2-ME, 100 U/ml penicillin, 100 μg/ml streptomycin, 0.3 mg/ml l-glutamine, and 10% FBS (all medium reagents from Invitrogen Life Technologies). In vitro activation of the DCs was accomplished by the addition of 2 μg/ml phosphorothioate-modified oligodeoxynucleotides containing CpG motifs 1826 (TCCATGACGTTCCTGACGTT) (Proligo), 5 μg/ml CD40L trimer (Amgen), 20 ng/ml recombinant murine GM-CSF (R&D Systems) for the indicated times.

DCs were cultured in flat-bottom 96-well plates (BD Biosciences) at a concentration of 1 × 106/ml (100 μl/well). LPS (Sigma-Aldrich) was added at 1 μg/ml when indicated. When indicated, 96-well plates were also previously coated for 2 h at 37°C with bovine fibronectin (Sigma-Aldrich). After culture, medium was removed from the wells and nonadherent cells were gently washed away by pipetting up and down with PBS. PBS was then removed and plates were frozen at −80°C. Plates were then thawed and the number of cells that were present in the wells before freezing was evaluated by measuring the amount of DNA in each well using the Cyquant kit (Molecular Probes) according to the manufacturer’s instructions. The plates were read on a VictorII fluorescence reader (Wallac). The percentage of adherent cells was evaluated by measuring the amount of DNA of a standard dilution of cells using the same technique.

Cells were preblocked at 4°C for 20 min in FACS buffer (PBS containing 2% FBS, 2% normal rat serum, 2% normal hamster serum, 2% normal mouse serum, 10 μg/ml CD16/CD32 (2.4G2) anti-FcR mAb (BD Biosciences), and 0.02% sodium azide). All mAbs were purchased from BD Pharmingen except where noted. In addition to isotypes controls, the following mAbs (clone name given in parentheses) were used: CD3 (145-2C11), CD8 (53-6.7), CD11b (M1/70), CD11c (HL3), CD19 (1D3), CD40 (HM40-3), CD45R/B220 (RA3-6B2), CD86 (GL-1), IAb (AF6-120.1), Gr-1 (RB6-8C5), Ly6c (AL-21), and Pan-NK (DX5). PE-conjugated F4/80 (CI:A3-1) was purchased from Caltag Laboratories. One IgG1 mAb (M652) was obtained from a rat immunized with a fusion protein consisting of the first 950 amino acids (predicted to be extracellular) of mouse plexin C1 fused with the Fc part of human IgG1. A39R binding on DCs was detected using a biotinylated rat mAb against the Flag (Flag M1; Amgen). Streptavidin-PerCP (BD Biosciences) was used for secondary staining. Flow cytometric analyses were performed on a FACSCalibur with CellQuest software (both BD Biosciences).

For confocal analyses, DCs were cultured into eight chamber coverslips that were previously coated with bovine fibronectin. In some experiments, wild-type and plexin C1−/− DCs were labeled with cell tracker red and green (Molecular Probes), respectively, before culture according to manufacturer’s instructions. The distribution of F-actin was visualized using Alexa 488-conjugated phalloidin (Molecular Probes) on paraformaldehyde-fixed DCs that had been permeabilized in PBS 0.1% Triton X-100 BSA 0.5%. For live imaging, coverslips were placed on a 37°C heated stage. Samples were viewed and images acquired on a confocal laser scanning microscope (Nikon; Molecular Dynamics) using a 60 Å objective and appropriate filters.

The 35 million DCs/condition were cultured onto 150 × 25 mm Petri dishes in 20 ml of medium. At the end of the culture, plates were quickly transferred onto an ice bath. Floating cells were collected by quick centrifugation at 4°C and lysed together with adherent cells on the Petri dish in RIPA lysis buffer (150 mM NaCl, 1% Nonidet P-40, 5 mM Tris pH 8.0). Lysates were incubated for 1 h with 1 μg of anti-FAK (Santa Cruz Biotechnology) after preclearing and then incubated with 25 μl of protein A-agarose for 2 h. Agarose beads were washed three times with lysis buffer. Immunoprecipitates were separated on Novex precast gels (Invitrogen Life Technologies). Proteins were transferred to a polyvinylidene difluoride membrane (Invitrogen Life Technologies) and probed with anti-FAK Ab (Santa Cruz Biotechnology) or anti-phospho-FAK (Y397) Ab (Zymed Laboratories).

Levels of IL-1, IL-2, IL-4, IL-6, TNF-α, GM-CSF, IFN-γ, and IL-12p70 in DC culture supernatants were measured using the Beadlyte Mouse MultiCytokine Detection System (Upstate Biotechnology) and the Luminex100 plate reader (Luminex) according to the manufacturer’s instructions. Quantification of cytokines was performed by regression analysis from a standard curve generated using cytokine standards included in the kit. Only IL-6, TNF-α, and IL-12p70 were detected in DC culture supernatant.

Cells were labeled with calcein-AM dye (Molecular Probes) according to the manufacturer’s instructions. Recombinant mouse MIP-1α/CCL, (R&D Systems) was diluted in PBS 0.1% BSA at a final concentration of 100 ng/ml and was added to the bottom wells (30 μl) of Neuroprobe ChemoTX 96-well plates no. 101-3 (NeuroProbe) together or not with A39R. Pore size was 3 μm, and well diameter was 3.2 mm. Labeled cells were resuspended in RPMI 1640/10% FBS and added (2 × 104 cells in 25 μl) to the top filter sites of the ChemoTX system. The plates were then incubated at 37° and 5% CO2 for 60 min. After incubation, the cell droplets on the top of the plate were washed off thoroughly with PBS/0.1% BSA four to five times. Excess liquid was removed from the top of the filter. The plates were read on a Molecular Devices Gemini Spectramax XS reader at excitation 490 nm/emission 528 nm, with a cutoff of 515 nm. Values were expressed as mean fluorescent count fold increase by calculating as follows: (experimental fluorescent counts)/(spontaneous fluorescent counts).

All values for p were calculated with the unpaired Student’s t test assuming equal variances.

Plexin C1 was previously identified as a receptor for A39R in human and mouse but its expression has only been partially documented in the immune system (7). We therefore generated an Ab against mouse plexin C1 (see Materials and Methods) that allowed us to measure the expression of plexin C1 on a large panel of immune cells. Little to no expression was found on lymphocytes, NK cells, and macrophages (Fig. 1,A). By contrast, neutrophils, splenic DCs, skin Langerhans cells, as well as in vitro generated DCs obtained from cultures of bone marrow cells in the presence of Flt3L were all found to express detectable levels of surface plexin C1 (Fig. 1,A and data not shown). Importantly, the anti-plexin C1 Ab did not bind to plexin C1−/− DCs or to other plexin C1−/− cell types (Fig. 1,B and data not shown). Consistent with Ab staining, recombinant poxvirus A39R strongly bound to C57BL/6 bone marrow-derived DCs and to freshly isolated C57BL/6 splenic DCs but not to DCs, neutrophils, or any other leukocytes from plexin C1−/− deficient mice (Fig. 1 B and data not shown). Thus, these data demonstrate that plexin C1 is the only receptor for A39R on mouse leukocytes and that DCs and neutrophils express it.

FIGURE 1.

Plexin C1 is preferentially expressed on DCs and neutrophils and is the only receptor for A39R. A, Spleen, bone marrow cells (for neutrophils), or peritoneal exudate cells (for macrophages) were stained with various Abs combinations plus an Ab against plexin C1 for flow cytometry. Plexin C1 expression was then determined on the different subsets using the following analyze gates: T cells (CD3+CD19DX5), B cells (CD19+B220+), NK cells (DX5+CD3), macrophages (CD11bbrightF4/80+), neutrophils (GR1brightCD11bbright), DCs (CD11c+), CD8+ DCs (CD11c+CD8+), CD8 DCs (CD11c+CD8), plasmacytoid DCs (pDCs, CD19B220+Ly6C+). Flt3L (FL) bone marrow-derived DCs were stained only with the anti-plexin C1 Ab. Results are representative of three experiments. B, Wild-type and plexin C1−/− bone marrow-derived DCs were either stained with the anti-plexin C1 Ab (left panels) or incubated with A39R and then stained with an Ab against the Flag (right panels). T, T cells; B, B cells; Mφ, macrophages; Neutro, neutrophils; pDCs, plasmacytoid dendritic cells; BMDC, bone marrow-derived dendritic cells. Results are representative of three experiments.

FIGURE 1.

Plexin C1 is preferentially expressed on DCs and neutrophils and is the only receptor for A39R. A, Spleen, bone marrow cells (for neutrophils), or peritoneal exudate cells (for macrophages) were stained with various Abs combinations plus an Ab against plexin C1 for flow cytometry. Plexin C1 expression was then determined on the different subsets using the following analyze gates: T cells (CD3+CD19DX5), B cells (CD19+B220+), NK cells (DX5+CD3), macrophages (CD11bbrightF4/80+), neutrophils (GR1brightCD11bbright), DCs (CD11c+), CD8+ DCs (CD11c+CD8+), CD8 DCs (CD11c+CD8), plasmacytoid DCs (pDCs, CD19B220+Ly6C+). Flt3L (FL) bone marrow-derived DCs were stained only with the anti-plexin C1 Ab. Results are representative of three experiments. B, Wild-type and plexin C1−/− bone marrow-derived DCs were either stained with the anti-plexin C1 Ab (left panels) or incubated with A39R and then stained with an Ab against the Flag (right panels). T, T cells; B, B cells; Mφ, macrophages; Neutro, neutrophils; pDCs, plasmacytoid dendritic cells; BMDC, bone marrow-derived dendritic cells. Results are representative of three experiments.

Close modal

Semaphorins are known to induce axon collapse (1, 2, 3, 4, 5), a phenomenon that could involve the regulation of cell adhesion through the modulation of integrin activity (13). We asked whether A39R could have a similar effect on mouse DCs. To test this hypothesis, we took advantage of the fact that freshly isolated spleen DCs or bone marrow-derived DCs placed in culture transiently adhere to plastic (Fig. 2,A and data not shown). This adhesion occurs within 2 h and is enhanced by maturation-inducing agents like LPS (Fig. 2,A). This adhesion is dynamic as DCs are motile cells that, under those in vitro conditions, constantly extend and retract lamellipodia, spreading and stretching their cellular body from one adherence point to another. This makes them often multipolar and from time to time creates long “dendrites” (Fig. 2,B and Ref. (17). Strikingly, when A39R was added at the beginning of the culture together with LPS, it completely prevented the formation of membrane processes by DCs (Fig. 2,C) as well as their adhesion to plastic. This effect was dependent on plexin C1 as A39R did not affect plexin C1−/− DCs either cultured in separate wells (Fig. 2, D and E) or in the same wells as wild-type cells (Fig. 2, F and G, wild-type and plexin C1−/− cells discriminated by a prior labeling with green and red dye, respectively). A39R was effective on wild-type DCs at a wide range of concentrations (Fig. 2,H), and importantly, without affecting their viability (Fig. 2 I). A39R also inhibited DC adhesion when DCs were cultured on extracellular matrix substrates such as fibronectin, (see below) collagen IV, or vitronectin (data not shown), which suggested that A39R-induced signal inhibited DC adhesion mediated by integrins. To explore this possibility, the role of integrins in in vitro DC adhesion on plastic or on a physiologic substrate like fibronectin was studied.

FIGURE 2.

A39R prevents DC adhesion to plastic. A–H, Wild-type (WT) and plexin C1−/− bone marrow-derived DCs were cultured with medium alone, or medium supplemented with LPS or LPS+A39R, as indicated. DCs were cultured for different times (A), or 2 h (B–H). A and H, the percentage of adherent cells was measured at the end of the culture as described in Materials and Methods. B–G, Representative confocal microscopy images at the end of the 2 h cultures. F and G, Wild-type and plexin C1−/− DCs have been labeled with cell tracker red and cell tracker green, respectively, and then mixed up at a 1:1 ratio before the culture. H, Wild-type (□) and plexin C1−/− (•) DCs are represented. Mean adhesion (∗) shown atop bar is statistically different from mean adhesion in the control condition without A39R (p < 0.01). No symbol indicates no difference. I, Wild-type DCs were cultured with (□) or without (▪) A39R (1 μg/ml) and the percentage of viable cells was measured over time in culture by flow cytometry as the percentage of propidium iodide-negative cells. The two curves are not statistically different. Results are representative of at least three experiments.

FIGURE 2.

A39R prevents DC adhesion to plastic. A–H, Wild-type (WT) and plexin C1−/− bone marrow-derived DCs were cultured with medium alone, or medium supplemented with LPS or LPS+A39R, as indicated. DCs were cultured for different times (A), or 2 h (B–H). A and H, the percentage of adherent cells was measured at the end of the culture as described in Materials and Methods. B–G, Representative confocal microscopy images at the end of the 2 h cultures. F and G, Wild-type and plexin C1−/− DCs have been labeled with cell tracker red and cell tracker green, respectively, and then mixed up at a 1:1 ratio before the culture. H, Wild-type (□) and plexin C1−/− (•) DCs are represented. Mean adhesion (∗) shown atop bar is statistically different from mean adhesion in the control condition without A39R (p < 0.01). No symbol indicates no difference. I, Wild-type DCs were cultured with (□) or without (▪) A39R (1 μg/ml) and the percentage of viable cells was measured over time in culture by flow cytometry as the percentage of propidium iodide-negative cells. The two curves are not statistically different. Results are representative of at least three experiments.

Close modal

DC adhesion to plastic.

β2 integrins confer a general stickiness to polymorphonuclear granulocytes resulting in their strong adhesion to many substrates, including plastic (18). To determine the role of β2 integrins in DC adhesion to plastic, we cultured DCs on plastic, in the presence of blocking anti-β2, or control Abs (isotype control or blocking anti-β1), and measured their adhesion (Fig. 3 A, left). Results show that DC adhesion to plastic was dependent on β2 integrins as the addition of anti-β2 Ab reduced DC adhesion to plastic by 60%. By contrast, a blocking anti-β1 Ab had no effect on DC adhesion to plastic and a combination of anti-β2 and anti-β1 Abs had the same effect than the anti-β2 Ab alone.

FIGURE 3.

A39R inhibits integrin-mediated DC adhesion and inhibits FAK phosphorylation. A, Wild-type (WT) bone marrow-derived DCs were cultured in medium supplemented with LPS into uncoated (left) or fibronectin-coated (right) wells. A39R or blocking Abs against integrins were added, as indicated. After 2 h of culture, the percentage of adherent cells was measured, as described in Materials and Methods. Mean adhesion (∗) atop bar is significantly different from mean adhesion in the control condition (Ig ctrl) (p < 0.001). No symbol indicates no significant difference. B, FAK phosphorylation was analyzed in DCs at the end of the 2 h culture by immunoprecipitation of total FAK and immunoblotting with anti-phospho-FAK (Y397) Ab. Results in are representative of at least three experiments.

FIGURE 3.

A39R inhibits integrin-mediated DC adhesion and inhibits FAK phosphorylation. A, Wild-type (WT) bone marrow-derived DCs were cultured in medium supplemented with LPS into uncoated (left) or fibronectin-coated (right) wells. A39R or blocking Abs against integrins were added, as indicated. After 2 h of culture, the percentage of adherent cells was measured, as described in Materials and Methods. Mean adhesion (∗) atop bar is significantly different from mean adhesion in the control condition (Ig ctrl) (p < 0.001). No symbol indicates no significant difference. B, FAK phosphorylation was analyzed in DCs at the end of the 2 h culture by immunoprecipitation of total FAK and immunoblotting with anti-phospho-FAK (Y397) Ab. Results in are representative of at least three experiments.

Close modal

DC adhesion to fibronectin.

In vitro adhesion of mouse polymorphonuclear or human Langerhans cells to fibronectin occurs through β1 integrin receptors (19). The effect of blocking anti-β2, anti-β1, or control Abs on DC adhesion to fibronectin was measured (Fig. 3,A, right). DC adhesion to fibronectin-coated wells was reduced by 40% by the anti-β2 blocking Ab (Fig. 3 A, right), showing a residual β2 binding to plastic even in the presence of fibronectin. Anti-β1 Ab alone reduced DC adhesion to fibronectin by 40% as well. A combination of both Abs had an additive effect and reduced DC adhesion by 80%. These results show that in these culture conditions, DC adhesion to fibronectin-coated wells was mediated at 80% by β1 and β2 integrins.

Effect of A39R on integrin-mediated DC adhesion.

Results presented in Fig. 3 A show that DC adhesion to plastic or fibronectin was reduced by 80% by A39R. In particular, the effect of A39R on DC adhesion to fibronectin was similar than the combined effect of blocking Abs against β1 and β2 integrins. Thus, plexin C1 engagement by A39R inhibited DC adhesion mediated by integrins.

These results suggest that integrin function in DCs could be affected following A39R binding to plexin C1. To further test this hypothesis, we measured the effect of A39R on the level of FAK phosphorylation in DCs, as FAK phosphorylation is associated with integrin activation, i.e., cells expressing active integrins contain a high proportion of Y397-phosphorylated FAK (20, 21, 22, 23, 24). As shown in Fig. 3 B, a high proportion of FAK was phosphorylated on Y397 following a culture in the presence of LPS. By contrast, LPS/A39R-treated DCs displayed a more reduced proportion of phosphorylated FAK. Thus A39R-induced signal inhibits integrin signaling as measured by FAK phosphorylation in DCs, further supporting the inhibition of integrin function by A39R.

Strong evidence shows that semaphorin binding to plexins induces actin cytoskeleton rearrangement in neurons or cell lines (3, 5, 9), a phenomenon associated with cellular retraction. Therefore, we investigated the effect of A39R on the actin cytoskeleton of DCs. To better visualize this effect, A39R was added to DCs that had been previously left to adhere for 2 h. In these conditions, as monitored by time-lapse microscopy, A39R induced a progressive retraction of adherent DC membrane processes within 10 min of treatment (Fig. 4,A). Collapsing DCs displayed multiple spikes rather than the characteristic lamellipodia that progressively disappeared resulting in a round or loosely adherent cell (Fig. 4,A). This effect was plexin C1-dependent as plexin C1−/− DC morphology was not altered in response to A39R (data not shown). Detection of F-actin with phalloidin in spread, adherent DCs showed diffuse intracellular pattern and some more intense staining at the edge of the membrane (Fig. 4,B, before A39R). Upon treatment with A39R, the collapse of lamellipodia was associated with an increase in the intensity of the F-actin staining at the base of the spikes or within the spikes themselves, consistent with a local reorganization of the actin cytoskeleton (Fig. 4,B, after A39R, early, and intermediate stages). At a later stage, F-actin staining was restricted to the perinucleus region of the rounded cell (Fig. 4,B, after A39R, final stage). Upon treatment with A39R, FAK phosphorylation on Y397 progressively decreased (Fig. 4,C). After 30 min of treatment with A39R, this phosphorylation was barely detectable (Fig. 4,C) and DCs were detached (Fig. 4 D).

FIGURE 4.

A39R induces actin cytoskeleton rearrangement, retraction of membrane processes, and detachment of adherent DCs. A, Wild-type (WT) bone marrow-derived DCs were cultured for 2 h in medium supplemented with LPS onto fibronectin-coated coverslips that were placed on a 37 degrees heated stage mounted on the confocal microscope. Pictures were taken by confocal time-lapse microscopy at the indicated times after addition of A39R at a concentration of 1 μg/ml. B–D, Wild-type bone marrow-derived DCs were cultured for 2 h in medium supplemented with LPS onto fibronectin-coated coverslips (B), petri dishes (C), or 96-well plates (D) before the addition of A39R (1 μg/ml) for different times. B, The distribution of F-actin in DCs was visualized by phalloidin staining either before A39R treatment or 15 min after addition of A39R. After A39R treatment, three types of F-actin distribution were observed corresponding to three stages in the membrane retraction process: “early”, “intermediate,” and “final”. DCs in intermediate and final stages of membrane retraction were found as soon as 1 min after A39R addition and within the next 15 min after. After 20 min of A39R treatment, at least 95% of the DCs were in the final stages. C, FAK phosphorylation was analyzed in DCs at the indicated times after addition of A39R by immunoprecipitation of total FAK and immunoblotting with anti-phospho-FAK (Y397) Ab. D, The number of adherent DCs in each well was measured 30 min after the addition of different concentrations of A39R. Results are expressed as the mean percentage ± SD of adherent cells relative to the control condition without A39R. Mean adhesion (∗) shown atop bar is significantly different from mean adhesion in the control condition without A39R (p < 0.01). No symbol indicates no significant difference. Results are representative of at least three experiments.

FIGURE 4.

A39R induces actin cytoskeleton rearrangement, retraction of membrane processes, and detachment of adherent DCs. A, Wild-type (WT) bone marrow-derived DCs were cultured for 2 h in medium supplemented with LPS onto fibronectin-coated coverslips that were placed on a 37 degrees heated stage mounted on the confocal microscope. Pictures were taken by confocal time-lapse microscopy at the indicated times after addition of A39R at a concentration of 1 μg/ml. B–D, Wild-type bone marrow-derived DCs were cultured for 2 h in medium supplemented with LPS onto fibronectin-coated coverslips (B), petri dishes (C), or 96-well plates (D) before the addition of A39R (1 μg/ml) for different times. B, The distribution of F-actin in DCs was visualized by phalloidin staining either before A39R treatment or 15 min after addition of A39R. After A39R treatment, three types of F-actin distribution were observed corresponding to three stages in the membrane retraction process: “early”, “intermediate,” and “final”. DCs in intermediate and final stages of membrane retraction were found as soon as 1 min after A39R addition and within the next 15 min after. After 20 min of A39R treatment, at least 95% of the DCs were in the final stages. C, FAK phosphorylation was analyzed in DCs at the indicated times after addition of A39R by immunoprecipitation of total FAK and immunoblotting with anti-phospho-FAK (Y397) Ab. D, The number of adherent DCs in each well was measured 30 min after the addition of different concentrations of A39R. Results are expressed as the mean percentage ± SD of adherent cells relative to the control condition without A39R. Mean adhesion (∗) shown atop bar is significantly different from mean adhesion in the control condition without A39R (p < 0.01). No symbol indicates no significant difference. Results are representative of at least three experiments.

Close modal

Taken together, these results demonstrate that A39R induces a rearrangement of the actin cytoskeleton, and inhibits FAK phosphorylation in adherent plexin C1 expressing DCs, leading to a retraction of their membrane processes and inhibition of their adhesion.

Cofilin binds to and depolymerizes pointed ends of F-actin, thereby increasing monomeric actin intracellular concentration leading to an increased F-actin turnover (25, 26). Phosphorylation of cofilin by kinases like Lin-11-Isl-1-Mec-3 kinase dissociates it from F-actin and inhibits its F-actin depolymerization activity. Because Sema3A induces actin cytoskeleton rearrangement through cofilin phosphorylation in neurons (11), we tested the involvement of cofilin phosphorylation in A39R-induced detachment of DCs. DCs were cultured in the presence of S3, a synthetic cell-permeable peptide containing a cofilin phosphorylation site that prevents the phosphorylation of endogenous cofilin and its inactivation (11) or a negative control, scrambled, peptide (RV) before the addition of A39R or PBS, then DC adhesion was measured. As shown in Fig. 5, in the absence of peptide, the addition of A39R induced a detachment of ∼60% of DCs. Increasing concentration of RV peptide had no effect on this A39R-induced detachment. By contrast, micromolar concentrations of S3 peptide efficiently inhibited A39R-induced DC detachment, without increasing DC adhesion in the control PBS condition. These results suggest that cofilin phosphorylation is a key downstream event in A39R-induced plexin C1 signaling and further link plexin C1 with the regulation of actin cytoskeleton.

FIGURE 5.

A39R effect on DCs is blocked by a cofilin phosphorylation inhibitor. Wild-type bone marrow-derived DCs were cultured in medium supplemented with LPS for 1 h. RV (left) or S3 (right) peptides were then added at the indicated concentrations and DCs were cultured for one more hour. A39R or PBS was then added for 30 min, as indicated. The number of adherent DCs in each well was measured. Results are expressed as the mean percentage ± SD of adherent cells relative to the control condition without A39R and without peptide. Paired bars show that mean adhesions (∗) are significantly different (p < 0.001). NS, No statistical difference (p > 0.05).

FIGURE 5.

A39R effect on DCs is blocked by a cofilin phosphorylation inhibitor. Wild-type bone marrow-derived DCs were cultured in medium supplemented with LPS for 1 h. RV (left) or S3 (right) peptides were then added at the indicated concentrations and DCs were cultured for one more hour. A39R or PBS was then added for 30 min, as indicated. The number of adherent DCs in each well was measured. Results are expressed as the mean percentage ± SD of adherent cells relative to the control condition without A39R and without peptide. Paired bars show that mean adhesions (∗) are significantly different (p < 0.001). NS, No statistical difference (p > 0.05).

Close modal

As DC maturation and activation is associated with changes in adherence and morphology with reorganization of their cytoskeleton (27, 28, 29), we tested whether A39R-induced morphological changes were associated with DC maturation. First, DCs were cultured with A39R or medium and MHC class II and costimulatory molecules expression (Fig. 6,A) as well as cytokine production (Fig. 6,B) was measured. Fig. 6 shows that A39R did not induce DC maturation. Then, we cocultured DCs with or without A39R and stimuli known to induce DC maturation such as a combination of CpG oligonucleotides with CD40L and GM-CSF. Fig. 6 shows that A39R addition did not alter DC maturation induced by these stimuli. Thus, A39R does not induce maturation of DCs or affect the maturation induced by CpG/CD40L/GM-CSF.

FIGURE 6.

A39R does not induce maturation of DCs or affect the maturation induced by CpG oligonucleotides. Bone marrow-derived DCs were cultured for the indicated times (A) or for 24 h (B) in the indicated conditions. CpG/CD40L/GM-CSF mixture (CpG) is shown in A. CD40, CD86, and I-Ab expression was then measured by flow cytometry (A) or cytokine secretion was measured in the supernatant (B) as described in Materials and Methods. Results are representative of three experiments. A39R addition had no statistical effect on the parameters measured in A and B.

FIGURE 6.

A39R does not induce maturation of DCs or affect the maturation induced by CpG oligonucleotides. Bone marrow-derived DCs were cultured for the indicated times (A) or for 24 h (B) in the indicated conditions. CpG/CD40L/GM-CSF mixture (CpG) is shown in A. CD40, CD86, and I-Ab expression was then measured by flow cytometry (A) or cytokine secretion was measured in the supernatant (B) as described in Materials and Methods. Results are representative of three experiments. A39R addition had no statistical effect on the parameters measured in A and B.

Close modal

Leukocyte trafficking in homeostatic conditions and their recruitment to inflamed sites are regulated by different classes of chemoattractant molecules that regulate leukocyte actin cytoskeleton and their adhesion molecules such as integrins in a coordinated fashion. To investigate the effect of A39R semaphorin on DC migration to chemokines, an in vitro chemotaxis assay was used. Wild-type or plexin C1−/− DCs were placed in the upper chamber of a transwell system and medium or the CCL3 was added to the lower chamber. CCL3 is known to induce immature DC migration to inflamed sites through its binding to the chemokine receptor CCR5. When indicated, A39R, or as a negative control, heat-inactivated A39R were also added to the lower chamber at different concentrations, alone or in combination with CCL3. Results presented in Fig. 7 show that CCL3 induced DC migration to the lower chamber of the transwell system in comparison with medium alone or A39R alone. Strikingly, A39R, but not heat inactivated A39R, inhibited wild-type DC migration to CCL3 in a dose-dependent manner. By contrast, A39R had no effect on plexin C1−/− DC migration induced by CCL3. A similar inhibition of DC migration by A39R was also observed when the other CCR5 ligands CCL4 and CCL5 or the CXCR4 ligand stromal cell-derived factor-1 were used as chemoattractants (data not shown). Moreover, A39R effect was not directional as it had the same effect when added in the lower chamber with the chemokines than in the upper chamber with the cells, or in both chambers (data not shown). Thus, plexin C1 engagement by A39R semaphorin inhibits chemokine-induced migration of DCs in vitro.

FIGURE 7.

A39R inhibits chemokine-induced migration of DCs in vitro. In vitro wild-type (WT) (□) and plexin C1−/− (▪) DC migration analysis in a chemotaxis assay toward CCL3 is shown. When indicated A39R (left) or heat-inactivated A39R (A39RHI) (right) was also added to the lower chamber of the transwell system, together or not with CCL3. Values are expressed as follows: (experimental fluorescent counts)/(spontaneous fluorescent counts). Results are presented as the mean ± SD of triplicate wells, and the experiment was performed four times. Mean migration (∗) is significantly different (p < 0.001) from mean migration in the control condition (CCL3 without A39R or heat-inactivated A39R). No symbol indicates no statistical difference (p > 0.02).

FIGURE 7.

A39R inhibits chemokine-induced migration of DCs in vitro. In vitro wild-type (WT) (□) and plexin C1−/− (▪) DC migration analysis in a chemotaxis assay toward CCL3 is shown. When indicated A39R (left) or heat-inactivated A39R (A39RHI) (right) was also added to the lower chamber of the transwell system, together or not with CCL3. Values are expressed as follows: (experimental fluorescent counts)/(spontaneous fluorescent counts). Results are presented as the mean ± SD of triplicate wells, and the experiment was performed four times. Mean migration (∗) is significantly different (p < 0.001) from mean migration in the control condition (CCL3 without A39R or heat-inactivated A39R). No symbol indicates no statistical difference (p > 0.02).

Close modal

An increasing number of reports show semaphorin/plexin expression in immune cells (15). In particular, several semaphorins seem to function in the reciprocal stimulation of T cells and APCs through nonplexin receptors. These semaphorins were thus believed to act through mechanisms that are different from those that are used in the nervous system (15). We show in this study for the first time that semaphorin binding to a plexin can regulate actin cytoskeleton rearrangements as well as integrin function in immune primary cells, in a similar manner than what was described for neurons.

We found that, in the mouse, plexin C1 was expressed mainly on DCs and neutrophils. Very little expression of plexin C1 was measured on B cells, although human plexin C1 was cloned from a B cell line, indicating a possible difference in plexin C1 expression between human and mouse. A39R did not bind to B cells, DCs, or to any other leukocytes from plexin C1−/− deficient mice (our study and data not shown) indicating that plexin C1 is the unique receptor for A39R on mouse leukocytes.

The inhibitory effect of A39R on DC adhesion was not due to a down-regulation of either β1 or β2 integrin expression, measured by flow cytometry (data not shown). Rather, this inhibition of adhesion was correlated with a dephosphorylation of FAK, a crucial mediator of integrin signaling, suggesting the inhibition of integrin function by A39R. We also found that A39R induced a plexin C1-dependent local increase in F-actin in adherent DCs, accompanied by the retraction of membrane processes and DC detachment. This effect, reminiscent of semaphorin-induced neuron growth cone collapse, was blocked by a cell permeable peptide (S3) that inhibits cofilin phosphorylation. Cofilin, a well-known mediator of F-actin depolymerization, is active in the unphosphorylated state. Thus, by triggering plexin C1, A39R could promote the phosphorylation of cofilin, which in turn, would reduce F-actin depolymerization. Aizawa et al. (11), the originators of the S3 peptide, showed a similar phenomenon in Sema3A-treated neurons, suggesting that cofilin phosphorylation is a central event in semaphorin-induced signaling in different cell types. It is, however, unclear how cofilin phosphorylation and the reduction of F-actin turnover may lead to lamellipodia/growth cone retraction. Barberis et al. (14) recently showed that Sema4D induced disassembly of focal adhesions in plexin B1 transfected COS cells. Interestingly, they found that this disassembly occurred before the actin cytoskeleton rearrangement suggesting that integrin inhibition and the resulting loss of adherence was the initial event induced by plexin signaling, causing membrane retraction. It is possible that cofilin phosphorylation occurs as a consequence of the regulation of integrin function by semaphorins. A direct link between plexins and integrins remains however to be identified.

We found that plexin C1 engagement by A39R could prevent chemokine-induced migration of DCs. Similarly, a previous study reported that CD100 and Sema3A semaphorins inhibited chemokine-induced migration of human monocytes in vitro through binding to an unknown receptor (30). This inhibition is not due to an interference with chemokine receptor signaling. Indeed we found that A39R did not alter the calcium response induced by chemokines (data not shown). Instead, the mechanism of A39R-induced inhibition of migration likely involves the inhibition of integrin function that is critical for leukocyte migration. Thus, in a simple model, integrin activity could be the molecular switch used by different guidance molecules either to attract or repel cells. Similarly, following early studies in neurons revealing the repellent activity of many semaphorins on cultured neurons in vitro, it was believed that semaphorins were guiding axons by providing them with “stop” signals in areas where they should not grow. These signals were thought to be complementary to attractant, chemokine signals to guide the growing axons. Recent studies in the field of endothelial cells refined this view. Indeed, Serini et al. (13) showed that autocrine loops of Sema3 chemorepellents exert an essential permissive role in the execution of vasculature remodelling by inhibiting integrin-mediated adhesion of endothelial cells to the extracellular matrix, allowing the de-adhesion necessary for vascular remodelling. They proposed that such a fine-tuning of integrin-mediated adhesion to the extracellular matrix allows a graded control of endothelial cell migration rate and redirectioning during migration. Thus, semaphorin signals could in fact help cells to move along a chemokine gradient, provided that both signals are not delivered in an antagonistic manner, by contributing to the dynamic of integrin activity. More experiments using semaphorin and plexin-deficient animals are required to test whether this model could apply to leukocyte migration. In the case of plexin C1, the identification of its cognate cellular ligand will also be a determinant to study its precise function in the regulation of leukocyte movement. Based on in vitro binding studies and homology with A39R it was suggested that Sema7A is the ligand for plexin C1 (8). However, our own attempts as well as others (31) have failed to show any binding of recombinant Sema7A on primary plexin C1 expressing leukocytes. Moreover, Sema7A has been shown to promote axon outgrowth in neurons in a plexin C1-independent manner (12).

A39R semaphorin is produced by many strains of poxviruses and seems to be secreted at very high amounts by virus-infected cells. A study by Gardner et al. (32) specifically addressed the role of A39R using different recombinant strains of vaccinia virus in a model of intradermal infection in mice. The authors reported that the forced expression of A39R by vaccinia strains that do not normally express it increased the severity and the persistence of skin lesions. It was, however, unclear whether this phenomenon was due to an increased immunopathology or to an increased virulence of the strain. The authors proposed that A39R might have intrinsic proinflammatory properties in this mouse system. This possibility was supported by the observation by Comeau et al. (7) that plexin C1 engagement by A39R induced a modest increase in the production of some proinflammatory cytokines in human monocytes. However, we did not detect any production of IL-1, IL-2, IL-4, IL-6, TNF-α, GM-CSF, IFN-γ and IL-12p70 by various mouse leukocytes (spleen cells, bone marrow cells, DCs, neutrophils, B cells) upon treatment with A39R in vitro (our study and data not shown), suggesting that a proinflammatory activity for A39R in mice is unlikely. Thus, we rather favor the hypothesis that A39R secretion might somehow alter the movement of plexin C1-expressing cells (DCs, monocytes, granulocytes) in the proximity of infected cells. Indeed, the uncontrolled expression of a semaphorin is predicted to have important consequences on the movement of responsive cells based on their activity on integrin function, as illustrated by our in vitro experiments. This effect could benefit the virus by inhibiting the recruitment of immune effector cells at the site of virus infection. Another possibility could be that regulating cell cytoskeleton and shape of neighboring cells could help the virus to infect them, as recently shown for human T cell leukemia virus (33).

In conclusion, we found that A39R binding to plexin C1 on primary mouse DCs induces actin cytoskeleton rearrangement, inhibits integrin-mediated adhesion and FAK phosphorylation, and impairs chemokine-induced migration in vitro. These data identify for the first time semaphorins and plexins as potent regulators of the actin cytoskeleton in the immune system and suggest that they could be involved in the regulation of leukocyte movement.

We especially thank Dr. M. K. Spriggs who initiated the work on semaphorins. We also thank Dr. J. Peschon who generated plexin C1-deficient mice, S. Wong Madden for the production of A39R protein, K. Schooley, for technical advice, J. Bradshaw, D. Kaufman, L. Strockbine for technical help, G. Carlton for graphics assistance, and Drs. D. Fitzpatrick, C. Maliszewski, J. Vakili, and A. Youakim for critical reading of the manuscript.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

2

Abbreviations used in this paper: DC, dendritic cell; FAK, focal adhesion kinase; Flt3L, Fms-like tyrosine kinase-3 ligand.

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