Dendritic cell function in mice lacking Plexin C1

Thierry Walzer, Laurent Galibert and Thibaut De Smedt1

Amgen Inc., Department of Immune Regulation, 1201 Amgen Court West Seattle, WA 98119, USA
1 Present address: Apoxis S.A., 18-20 Avenue de Sevelin, CH-1004 Lausanne, Switzerland

Correspondence to: T. Walzer; E-mail: twalzer2002{at}yahoo.fr


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Semaphorins are secreted or transmembrane proteins that provide essential repulsive guidance cues to growing axons or endothelial cells through their receptors of the Plexin and Neuropilin family. Semaphorins and Plexins are also expressed in the immune system where their function remains elusive. In particular, Plexin C1 is expressed by mouse dendritic cells (DCs) and is the receptor for the poxvirus semaphorin homolog A39R. We previously found that Plexin C1 engagement by A39R inhibits integrin-mediated DC adhesion and chemokine-induced migration. Here, we show that a cellular ligand for Plexin C1 is expressed both by activated T cells and DCs, suggesting that Plexin C1 might be engaged on DCs both in cis and in trans. We used Plexin C1–/– mice to explore the role of Plexin C1 in DC function. DC development is unaffected in these mice. In two different in vivo assays, Plexin C1–/– DC migration to lymph nodes (LNs) was lower than that of wild-type (WT) DC but this difference was not statistically significant. Plexin C1–/– bone marrow-derived DCs induced normal in vitro T cell responses but reduced in vivo T cell responses when injected subcutaneously to WT mice. Finally, in vivo T cell responses to ovalbumin peptide and contact hypersensitivity to dinitrofluorobenzene were slightly decreased in Plexin C1–/– mice. These results suggest a role for Plexin C1 in DC migration or mobility within the LNs.

Keywords: antigen presentation, guidance, motility, semaphorins, trafficking


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
In the nervous system, axonal pathways are shaped by repulsive cues provided by ligands of the semaphorin family that are sensed by migrating neuronal growth cones through neuropilin and plexin receptors (14). Semaphorins share a conserved, 500-amino acid residue domain near their amino terminus called the sema domain. Currently, 25 members have been identified and divided into eight classes according to species origin and membrane anchorage. A semaphorin homolog called A39R has also been identified as being encoded within the genomes of several poxviruses such as vaccinia (5). Plexins are type-I transmembrane proteins that also contain a sema domain in their extracellular part and share a domain called the ‘sex and plexins’ domain in their intracellular part (6). They are divided into four subfamilies, Plexins A–D, according to the structure of their extracellular domain.

Semaphorins signal through direct binding to plexins, alone or in combination with neuropilins for the class 3 semaphorins. The typical cellular response to plexin engagement by semaphorins is the retraction or collapse of membrane processes such as axons for neurons or pseudopodia for migrating cells. Several reports indicated that plexin engagement by semaphorins regulates integrin-mediated adhesion (79). A direct link between plexins and integrins has been recently identified. Indeed, Oinuma et al. found that the semaphorin 4D receptor Plexin B1 has a GTPase-activating protein activity for the Ras family GTPase R-Ras (10), known to play a key role in cell adhesion by activating integrins (11).

Plexins and semaphorins are expressed in the immune system. In particular, Plexin A1 and Plexin C1 are both expressed at high levels on mouse dendritic cells (DCs) where their function remains elusive (9, 12). Small interfering RNA-based knock down of Plexin A1 expression in the mouse DC line 3B11 impairs its capacity to activate antigen-specific T cells both in vitro and in vivo (12). The authors proposed that Plexin A1 could contribute to the formation of the immunological synapse between DCs and T cells. The poxvirus semaphorin A39R was found to bind with high affinity to Plexin C1 (13) on the surface of DCs and neutrophils (9). We have shown that Plexin C1 engagement by A39R inhibits integrin-mediated DC adhesion and in vitro DC chemotaxis, suggesting a role for Plexin C1 in DC migration (9).

Sema7A is the only known glycosyl-phosphatidylinositol (GPI)-linked cellular semaphorin and has been proposed to be a cellular ligand for Plexin C1 on the basis of sequence similarity between Sema7A and A39R and of recombinant Sema7A–AP binding on Plexin C1-transfected cell lines (6, 14). However, our own attempts as well as others (15) have failed to detect any binding of recombinant Sema7A–Fc on primary Plexin C1-expressing leukocytes. Moreover, Sema7A has been found to have a Plexin C1-independent neurotrophic activity, suggesting, at least, the existence of an alternate counter-structure for Sema7A on neurons. Here, to get further insight into Plexin C1 function, we measured the pattern of expression of Plexin C1-ligand in the immune system using a Plexin C1–Fc fusion protein. We found that this ligand is linked to the cell surface through a GPI link and is expressed on the surface of activated DCs and T-lymphocytes. We hypothesized that Plexin C1 engagement by this endogenous ligand either in cis or in trans could influence DC migration and/or antigen presentation. We took advantage of Plexin C1-deficient mice to test this hypothesis.


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Mice
All mice were used at 7–10 weeks of age. Female C57BL/6 and BALB/C mice were purchased from Taconic (Germantown, NY, USA). The generation of Plexin C1–/– mice has already been described (16). Plexin C1–/– mice were bred at Taconic. Female OT-2 TCR-transgenic (Tg) mice crossed with congenic B6.PL-Thy1a/Cy mice [Thy1.1 obtained from The Jackson Laboratory (Bar Harbor, ME, USA)] specific for chicken ovalbumin peptide (OVA) 323–339 (OT-2p) in the context of I-Ab were bred at Amgen Inc. (Seattle, WA, USA). OT-I TCR-Tg mice specific for chicken OVA 357–364 (OT-Ip) in the context of H-2Kb were purchased from The Jackson Laboratory. All mice were maintained at Amgen Inc. under specific pathogen-free conditions and an institutional animal care and use committee approved all experiments according to federal guidelines.

DC cultures
Bone marrow-derived dendritic cells (BMDCs) were obtained by culturing bone marrow cells with Flt3L (Amgen Inc.) as previously described (17). This procedure leads to the generation of classical CD11c+B220 DCs and of plasmacytoid CD11c+B220+ DCs (10–20% of CD11c+ cells). The percentage of plasmacytoid DCs was similar in wild-type (WT) and Plexin C1–/– bone marrow cultures. In vitro activation of the DCs was accomplished by the addition of 2 µg ml–1 phosphorothioate-modified oligodeoxynucleotides containing CpG motifs 1826 (TCCATGACGTTCCTGACGTT) (Proligo, Boulder, CO, USA), 5 µg ml–1 CD40L trimer (Amgen Inc.), 20 ng ml–1 recombinant murine granulocyte macrophage colony-stimulating factor (GM-CSF) (R&D Systems, Minneapolis, MN, USA), for 24 h.

Construction and expression of recombinant proteins
Plexin C1–Fc fusion protein contains the extracellular domain of Plexin C1 fused to a modified human IgG1 Fc region (18) after amino acid 950 and was sub-cloned into the mammalian expression vector pDC409 (19). Plasmids encoding the Fc fusion protein were transfected into CV-1/EBNA (American Type Tissue Collection CRL-10478) cells, and the fusion proteins were purified from culture supernatants by chromatography on a protein A-Poros column (PerSeptive Biosystems, Framingham, MA, USA) as described (20). A control–Fc fusion protein, p7.5-Fc, has been previously described (21).

Phosphatidylinositol-specific phospholipase C treatment
To obtain evidence for GPI linkage of Plexin C1, activated DCs were treated with 2 U ml–1 phosphatidylinositol-specific phospholipase C (PI-PLC) (Sigma, St Louis, MO, USA) at 37°C for 1 h. Subsequently, cells were washed with PBS and stained on ice by indirect immunofluorescence using Plexin C1–Fc as described above.

Chemotaxis assay
The procedure used has been described previously (17). Briefly, cells were labeled with Calcein-AM dye (Molecular Probes, Eugene, OR, USA) according to the manufacturer's instructions. Recombinant chemokines (R&D Systems) were diluted in PBS 0.1% BSA at a final concentration of 100 ng ml–1 and were added to the bottom wells of Neuroprobe ChemoTX 96-well plates #101–3 (NeuroProbe, Gaithersburg, MD, USA). Pore size was 3 µm, and well diameter was 3.2 mm. Labeled cells were re-suspended in RPMI 1640 10% FBS and added (2 x 104 cells in 25 µl) to the top filter sites of the ChemoTX system. The plates were then incubated at 37°C and 5% CO2 for 60 min. The plates were read on a Molecular Devices (Sunnyvale, CA, USA) Gemini Spectramax XS reader at excitation 490 nm/emission 528 nm. Values were expressed as mean fluorescent count fold increase by calculating as follows: experimental fluorescent counts/spontaneous fluorescent counts.

In vivo DC migration
BMDC migration.
WT and Plexin C1–/– BMDCs from a 9-day culture were labeled with 0.5 µM of the vital dye 5(and 6)-carboxyfluorescein diacetate succinimidyl ester (CFSE), mixed isomer (Molecular Probes Inc.). A total of 5 x 105 labeled cells were injected subcutaneously (s.c.) in the hind leg footpad. Popliteal lymph nodes (LNs) were recovered 24 h later and treated with 200 U ml–1 collagenase (Worthington, Lakewood, NJ, USA) in HBSS with Ca2+ and Mg2+ for 30 min at 37°C. The LNs were then mashed and further dissociated in Ca2+-free medium in the presence of 2 mmol l–1 EDTA. The number of cells was counted and CFSE fluorescence was measured by flow cytometry.

FITC painting.
FITC (FITC isomer I, Sigma) was dissolved in a 50 : 50 (vol/vol) acetone–dibutylphthalate mixture just before application. Mice were painted on the shaved back with 0.5% FITC. After 24 h, inguinal LNs were treated with 200 U ml–1 collagenase in HBSS with Ca2+ and Mg2+ for 30 min at 37°C. LNs were then mashed and further dissociated in Ca2+-free medium in the presence of 2 mmol l–1 EDTA. The number of cells was counted and cells were stained for I-Ab expression before flow cytometry acquisition.

Measurement of CD8+ and CD4+ T cell priming capacities by BMDCs.
DCs were incubated overnight with 0.5 mg ml–1 OVA-protein (Albumin, Chicken Egg, 5X Crystalline; Calbiochem-Novabiochem, San Diego, CA, USA) in the presence or absence of the maturation cocktail CpG/CD40L/GM-CSF (see above). Naive CD8+ or CD4+ or total T cells were enriched from spleen and peripheral LNs of OT-I, OT-2 or BALB/C mice, respectively, by immunomagnetic cell separation using negative selection with the StemSep Enrichment mixture kits for CD8+, CD4+ or total T cells (StemCell Technologies, Vancouver, British Columbia, Canada). Naive T cells at 1 x 104 cells per well were seeded into 96-well plates (Costar Corning, Cambridge, MA, USA) and cultured with varying numbers of protein-pulsed BMDCs in 200 µl of IMDM (Life Technologies, Carlsbad, CA, USA) plus additives per well. Cultures were maintained for 72 h at 37°C. The proliferation was assayed by pulsing the cells with 0.5 µCi of [3H]thymidine overnight and harvesting them the next day. At 72 h, culture supernatants were collected and tested in triplicate for IFN-{gamma} by ELISA (BD PharMingen, San Jose, CA, USA).

Flow cytometry
The cells were stained in FACS buffer [PBS containing 2% FBS, 1% normal rat serum, 1% normal hamster serum, 1% normal mouse serum and 10 µg ml–1 2.4G2 (a rat anti-mouse FcR) mAb]. The following mAbs (clone name given in parentheses) were used: CD4 (GK1.5), CD11b (M1/70), CD11c (HL3), CD19 (1D3), CD45R/B220 (RA3-6B2), CD90.1 (OX-7), I-Ab (AF6-120.1), Ly-6C (AL-21), Gr-1 (RB6-8C5), V{alpha}2 (B20.1) and Vß5 (MR9-4); all were from BD Biosciences (San Diego, CA, USA). F4/80 (CI:A3-1) was purchased from Caltag (Burlingame, CA, USA). CCR3 (83103) was purchased from R&D Systems. Biotinylated mAb binding was detected with streptavidin–PerCp (BD Biosciences). Flow cytometric analyses were performed on a FACSCalibur with CellQuest software (both BD Biosciences).

Immunization with OVA/Ribi
Mice were immunized with 100 µg OVA emulsified in Ribi adjuvant (Sigma) in the hind footpads. Draining popliteal LNs were harvested 6 days after immunization and LNs cells were re-stimulated in vitro with different doses of OVA as previously described (22). Proliferation and IFN-{gamma} secretion were measured as described above.

Contact hypersensitivity
Mice were sensitized by epicutaneous application of 25 µl of 0.2% dinitrofluorobenzene (DNFB) onto 2 cm2 of fur-shaved ventral skin. Five days later, animals were challenged by the application of 5 µl of 0.2% DNFB onto each side of the right ear. Ear thickness measurements were taken before hapten application with a spring-loaded micrometer. Measurements were repeated once daily after challenge. Ear swelling was calculated by subtracting the initial value from the value recorded on the corresponding day.


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
A ligand for Plexin C1 is expressed on activated DCs and T cells
We previously found that Plexin C1 is mainly expressed on DCs. To investigate Plexin C1 function on these cells, we first set out to determine the pattern of expression of Plexin C1-ligand. A soluble fusion protein composed of the extracellular domain of Plexin C1 fused with the Fc portion of human IgG1 was produced. Plexin C1–Fc did not significantly label freshly isolated spleen, LN or bone marrow cells—including DCs—as assessed by flow cytometry (Fig. 1A and B and data not shown). In contrast, after overnight activation with CpG oligonucleotides and CD40L, BMDCs or primary spleen DCs could be labeled with Plexin C1–Fc (Fig. 1C and data not shown). Plexin C1–Fc binding to activated DCs was mediated by the Plexin C1 part of the protein, as addition of an excess of recombinant A39R, completely abrogated labeling (Fig. 1C). Plexin C1–Fc binding to activated DCs was also strongly inhibited by PI-PLC pre-treatment, thus suggesting that the Plexin C1 counter-structure on activated DCs is linked to the cell surface through a GPI-bound protein (Fig. 1D). Finally, Plexin C1–Fc bound to T cells activated by a 24-h culture on plate-bound anti-CD3 antibody in the presence of anti-CD28 antibodies but not to resting T cells (Fig. 1E).



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Fig. 1. A GPI-linked ligand for Plexin C1 is expressed on activated DCs and T cells. Plexin C1–Fc (white histograms) or Ctrl–Fc (gray histograms) binding on resting splenocytes (A), immature BMDCs (B), activated BMDCs (C–D) or resting or anti-CD3-activated T cells (E) were measured by flow cytometry. (C, right panel) an excess of rA39R protein was added during the incubation with Plexin C1–Fc. In (D), activated DCs were treated with medium or PI-PLC, as indicated, before measurement of Plexin C1–Fc binding. Results in Fig. 1 are representative of three independent experiments.

 
Thus, activated DCs and T-lymphocytes express a surface GPI-bound protein that binds Plexin C1 extracellular domain. Plexin C1 expressed on DCs might therefore be engaged by this putative ligand either in cis or in trans in the context of a DC–T cell interaction.

DC development and distribution in Plexin C1–/– mice
Semaphorin–Plexin interactions have been shown to be crucial for the development of the neural and the cardiovascular tissues (7, 2327). Current models propose that semaphorin signals given either in a paracrine or autocrine fashion are essential to guide cells to their proper localization. We compared the percentage of the major DC subsets in lymphoid organs of WT and Plexin C1–/– mice by flow cytometry. Results presented in Fig. 2(A) show that the percentage of MHC-IIint-resident DCs and MHC-IIhigh skin-emigrant DCs are similar in skin-draining LNs of WT and Plexin C1–/– mice. The percentage of CD8{alpha}+ and CD8{alpha} DCs was comparable in the spleen of both strains of mice (Fig. 2B). Total cell numbers in the lymphoid organs as well as the percentage of the other leukocyte subsets (T cells, B cells, monocytes, macrophages, neutrophils, eosinophils) were also normal in Plexin C1–/– mice (data not shown). Finally, we compared the network of Langerhans cells in the epidermis of both strains of mice by staining epidermal sheets for DEC 205 expression. Results presented in Fig. 2(C and D) show that there is a little but significant increase in the number of Langerhans cells in the skin of Plexin C1–/– mice compared with WT mice. Langerhans cells morphology appeared similar in both strains of mice (data not shown).



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Fig. 2. DC development and localization in tissues are normal in Plexin C1–/– mice. (A) MHC-II and CD11c expression by inguinal LN cells was measured by flow cytometry. (B) CD8 and CD11c expression by spleen cells was measured by flow cytometry. (C–D) Epidermal sheets from WT and Plexin C1–/– mice were prepared and stained for DEC 205 expression as described in Methods. (C) Representative fields from WT and Plexin C1–/– skins as indicated. DEC 205 staining is in red. (D) Average number of DEC 205-positive cells (Langerhans cells) per field; *P < 0.05.

 
Thus, development and number of DCs in lymphoid organs of Plexin C1–/– mice are normal while skin Langerhans cells numbers are slightly increased in comparison to WT mice.

Plexin C1–/– DCs migration properties
Next we evaluated Plexin C1–/– DC ability to migrate in vitro or in vivo. Immature DCs are known to express chemokine receptors such as CCR1 and CCR5, which recognize inflammatory chemokines such as CCL3, whereas maturation down-regulates the expression of these receptors and up-regulates the expression of CCR7, ligands of which are CCL19 and CCL21 (28, 29). We used an in vitro chemotaxis assay to measure the ability of immature or mature BMDCs to migrate across a polycarbonate filter in response to CCL3, CCL19 or CXCL12. As expected, immature DCs migrated poorly to CCL19 but responded efficiently to CCL3 and CXCL12 (Fig. 3A). By contrast, mature DCs were not responsive to CCL3 but migrated vigorously to the CCR7 ligand CCL19 and also responded, albeit less well, to stromal-derived factor 1 (SDF-1) (Fig. 3A). No significant difference of migration was observed between WT and Plexin C1–/– DCs at all tested chemokine concentrations (Fig. 3A and data not shown).



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Fig. 3. Plexin C1–/– DCs migration properties. (A) WT and Plexin C1–/– BMDCs were cultured overnight in medium (immature) or medium supplemented with a CpG/CD40L/GM-CSF maturation cocktail (mature). Their in vitro migration to CCL3, CCL19 or CXCL12 was then measured as described in the experimental procedures. Results are representative of three independent experiments. (B) WT and Plexin C1–/– BMDCs were labeled with CFSE and injected s.c. in the rear footpads of C57BL/6 recipient mice together with 5 µg LPS. Twenty-four hours after the injection, the number of migrated DCs in the popliteal LNs was counted. Left panel: Plexin C1–Fc (white) or control–Fc (gray) binding to migrated DCs. Right panel: mean number of WT and Plexin C1–/– migrated DCs in pooled popliteal LNs (mean ± SD of five independent mice per group). Results are representative of three independent experiments. (C–D) Skin DC migration to LN after painting of shaved abdomens of WT and Plexin C1–/– mice with 1% FITC. (C) Representative flow cytometry profiles of WT mice treated with FITC or vehicle only. Migrated FITC+ MHC-IIhigh DCs are in the upper right quadrant. (D) Left panel: Plexin C1–Fc (white) or control–Fc (gray) binding to migrated FITC+ DCs. Right panel: relative number of WT and Plexin C1–/– migrated FITC+ DCs in pooled inguinal LNs (mean ± SD of eight independent experiments with five mice in each group).

 
Next, immature DCs from Plexin C1–/– or WT mice were labeled with CFSE and injected into the footpad of WT recipient animals together with LPS to induce their in vivo maturation and migration to LNs. Importantly, 24 h after injection, migrated CFSE+ DCs in the popliteal LNs expressed Plexin C1-ligand (Fig. 3B). Figure 3(B) shows that the mean number of migrated Plexin C1–/– BMDCs in this experiment was 20% lower but not statistically different from that of WT BMDCs.

Application of FITC to the skin of mice induces contact sensitivity resulting from the migration of FITC+ epidermal DCs to the draining LN (30). In this experimental model, migrated DCs can be identified in the skin-draining LNs by their FITC fluorescence and their high MHC-II expression (Fig. 3C). These FITC+/MHC-IIhigh DCs expressed Plexin C1 (data not shown) and low levels of Plexin C1-ligand (Fig. 3D). Figure 3(D) shows that the mean number of FITC+/MHC-IIhigh in the LNs of FITC-sensitized Plexin C1–/– mice was 20% lower but, again, not statistically different from that of WT mice.

Thus, Plexin C1–/– and WT DC migration to LNs were not statistically different although Plexin C1–/– tended to migrate slightly less than their WT counterpart.

Plexin C1–/– BMDCs antigen-presentation capabilities
Next, we compared WT and Plexin C1–/– DCs antigen-presentation properties. BMDCs were cultured overnight with OVA in the presence or absence of a maturation-inducing cocktail and then cultured at different T cell : DC ratios with OVA-specific H-2b-restricted naive TCR-Tg OT-I CD8 T cells (Fig. 4A), OT-2 CD4 T cells (Fig. 4B) or allogenic BALB/C H-2d-restricted T cells (Fig. 4C). T cell proliferation and IFN-{gamma} level in the supernatant were measured. Results presented in Fig. 4(A–C) show that both immature and mature DCs induced strong T cell activation. As expected, mature DCs induced a higher proliferation and IFN-{gamma} secretion by T cells. However, no significant difference was observed between the capacity of WT and Plexin C1–/– DCs, either immature or mature, to induce T cell activation in these in vitro conditions.



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Fig. 4. Plexin C1–/– BMDCs have normal in vitro antigen-presentation capabilities. OVA-pulsed immature (squares) and mature (circles) WT (filled symbols) and Plexin C1–/– (open symbols) BMDCs were cultured at different DC : T cell ratios with purified OT-I CD8 T cells (A), OT-2 CD4 T cells (B) or allogenic BALB/C T cells (C). T cell proliferation and IFN-{gamma} secretion were measured.

 
We then tested the ability of BMDCs to prime naive T cells in vivo. Congenic Thy1.1 + OT-2 CD4 T cells were labeled with CFSE and adoptively transferred to C57BL/6 mice. OVA-pulsed BMDCs from WT or Plexin C1–/– mice were then injected together with LPS to induce their maturation, either intra-footpad (Fig. 5A) or intravenously (i.v.) (Fig. 5B), to adoptively transferred mice. OT-2 T cell activation in the popliteal LNs or in the spleen was assessed by counting the number of Thy1.1 + CD4+ cells and by measuring their CFSE fluorescence, representative of their proliferation, 4 days after DC injection. Results presented in Fig. 5(A and B, left panels) show that OT-2 CD4 T cells strongly expanded in DC-injected mice compared with unimmunized control animals. Plexin C1–/– DC induced a 20% lower OT-2 T cell expansion than WT DCs when injected s.c. (Fig. 5A, P < 0.02). Accordingly, the CFSE profiles of activated OT-2 T cells revealed a slightly higher percentage of non-divided cells in the Plexin C1–/– DC-immunized animals compared with WT DC-immunized animals (Fig. 5A). By contrast, when injected i.v., WT and Plexin C1–/– DCs induced a similar proliferation/expansion of OT-2 T cells (Fig. 5B).



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Fig. 5. Comparison of in vivo WT and Plexin C1–/– BMDC antigen-presentation capabilities. OVA-pulsed WT or Plexin C1–/– BMDCs were injected either s.c. in the rear footpads (A) or i.v. (B) to C57BL/6 mice that had been previously adoptively transferred with CFSE-labeled naive TCR-Tg OT-2 CD4 T cells. CD4 T cell expansion and CFSE fluorescence were measured in the popliteal LNs (A) or in the spleen (B) 4 days after DC injection; *P < 0.02.

 
Altogether, these data suggest that Plexin C1–/– DC have normal antigen-presentation capabilities but a slightly defective ability to reach draining LNs when injected s.c.

In vivo T cell responses to exogenous antigens in Plexin C1–/– mice
We compared the ability of WT and Plexin C1–/– mice to mount T cell responses in vivo. First, WT and Plexin C1–/– mice were injected s.c. with OVA in Ribi adjuvant. T cell responses against s.c. injected antigens are known to be dependent on the ability of skin DCs to capture the antigen, migrate to LNs and prime antigen-specific naive T cells (31). Draining LNs were recovered 6 days after the OVA injection and LNs cells were cultured with OVA in vitro to re-stimulate OVA-specific T cells. Proliferation and IFN-{gamma} secretion were measured. Results presented in Fig. 6(A) show that LNs cells from OVA-immunized but not from control animals displayed strong proliferation and IFN-{gamma} production. Cell proliferation was similar in cultures of WT and Plexin C1–/– LNs. However IFN-{gamma} production was modestly but significantly decreased in Plexin C1–/– LN cell cultures (Fig. 6B).



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Fig. 6. Comparison of in vivo T cell responses in WT and Plexin C1–/– mice. (A–B) WT (filled squares) and Plexin C1–/– (open circles) mice were injected s.c. in the rear footpads with 100 µg OVA in Ribi adjuvant. Six days later draining LN cells were cultured with graded doses of OVA. Cell proliferation and IFN-{gamma} secretion were measured; open triangles, unimmunized control mice. (C) CHS to DNFB was compared in groups of five C57BL/6 and Plexin C1–/– mice as described in Methods. Results are expressed as the mean ear swelling (in micrometers) at different time points after challenge. Results are representative of six independent experiments; *P < 0.05.

 
Next, WT and Plexin C1–/– mice contact hypersensitivity (CHS) responses to DNFB were compared. CHS to haptens like DNFB has been shown to be dependent on hapten-specific cytotoxic CD8 T cells. Previous reports have also showed that hapten-specific T cell priming is dependent on the migration of DCs from the skin to the draining LNs (32). As shown in Fig. 6(C) CHS response was significantly reduced in Plexin C1–/– mice compared with WT mice, as measured by ear swelling in response to DNFB challenge.

Thus, Plexin C1–/– mice display slight defects in their in vivo T cell responses.


    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
We previously showed that poxvirus semaphorin A39R binding to Plexin C1 inhibited integrin-mediated adhesion of mouse DCs and impaired their chemokine-induced in vitro migration (9). The effect of A39R on DC migration was presumably due to an interference with integrin function, which is required during cell migration. These results established Plexin C1 as a potent regulator of DC adhesion/migration. This study was designed to further explore the role of Plexin C1 in DC biology. We first sought to determine where and when the cellular ligand for Plexin C1 is expressed. To this end, we used a Plexin C1–Fc fusion protein to measure its expression. We found that Plexin C1-ligand expression was restricted (in the immune system) to activated DCs and activated T-lymphocytes and identified the Plexin C1-ligand as a GPI-linked membrane. As Sema7A is the only known cellular semaphorin to be GPI linked, these results strongly suggest that Sema7A is the actual Plexin C1-ligand in the immune system. Another supportive evidence comes from microarray and northern blot analyses showing that Sema7A mRNA is strongly up-regulated upon maturation in DCs [(33), L.G., unpublished observations] and upon activation in T cells (34).

We next compared WT and Plexin C1–/– DC migration both in vitro and in vivo. DCs are highly motile cells that migrate from non-lymphoid organs to lymphoid organs upon maturation stimuli. Within lymphoid organs, they also constantly move in T cell areas, presumably to favor encounter with antigen-specific T cells. Finally, immature DCs get recruited to inflammation sites in response to different chemokines (28). Plexin C1–/– DC in vitro migration to CCL3, CXCL12 and CCL19 appeared normal. Plexin C1–/– DC in vivo migration was, on average, 20% lower than WT DCs migration in two different assays. Given the intrinsic high variability of these in vivo techniques, such a difference was not statistically significant. However, it correlates first, with a significantly higher number of Langerhans cells in the skin of Plexin C1–/– mice compared with control animals, which could be explained by their defective migration to draining LNs. Second, it also correlates with defective T cell responses in Plexin C1–/– mice that could be explained by reduced DC migration/mobility as discussed below.

We found that Plexin C1–/– mice had lowered DNFB-induced hypersensitivity responses and slightly decreased T cell responses (IFN-{gamma} secretion) to OVA administered s.c. These impaired T cell responses were unlikely to be due to an intrinsic defect of Plexin C1–/– T cells because, first, Plexin C1 is not expressed in T cells, second, T cell development and surface phenotype are normal in Plexin C1–/– mice (data not shown) and third, in vitro proliferation and cytokine secretion of purified Plexin C1–/– T cells induced by anti-CD3 antibody is normal (data not shown). Rather, our results point to a defect at the level of DCs. Indeed, Plexin C1–/– BMDCs induced a lower OVA-specific proliferative T cell response than WT BMDCs when they were injected s.c into WT mice adoptively transferred with OT-2 TCR-Tg T cells. Plexin C1–/– BMDC defect was not at the level of their antigen-presentation capabilities because these cells could induce normal in vitro CD4, CD8 and allogenic T cell responses. Moreover, Plexin C1–/– and WT BMDCs induced a similar transgenic T cell response when injected i.v. into WT mice adoptively transferred with TCR-Tg T cells. Thus, our results suggest a slightly impaired ability of Plexin C1–/– DCs to migrate from non-lymphoid tissues such as the skin or the lung to the draining LNs. This little defect in DC migration to LNs might be coupled to a defective mobility of these cells within the LNs, which would explain why DNFB-induced hypersensitivity responses are clearly defective in Plexin C1–/– mice while DC migration from skin to LNs is only mildly affected. More studies will be necessary to clarify this point.

It is not clear why Plexin C1–/– BMDC in vitro migration and adhesion (this report and reference 9) are normal while our results suggest a defect in their in vivo migration properties. Compensatory or redundancy mechanisms might hide Plexin C1–/– DC phenotype, particularly in vitro. This phenomenon clearly limits our understanding of the role of this protein in DC function. Based on our previous findings that A39R inhibits integrin-mediated adhesion in DCs (9), it can be speculated that Plexin C1 plays a role in the complex regulation of integrin function that occurs when DCs migrate. Interestingly, we found that the kinetics of appearance of Plexin C1-ligand on DCs following activation correlates with those of their detachment from the substrate in vitro (data not shown). Although these two events appeared not to be linked in vitro, it is possible that in vivo engagement of Plexin C1 on DCs in tissues like the skin might participate in the release of their adhesion to local extracellular matrix and neighboring cells that precedes their migration. Another possibility would be that Plexin C1–Plexin C1-ligand interactions contribute to the regional control of integrin activity that allows cell motility.

While DNFB-induced hypersensitivity responses are clearly defective in Plexin C1–/– mice, Plexin C1–/– BMDCs injection leads to normal T cell responses when injected i.v. and only slightly defective T cell responses when injected s.c. Although the basis of this phenomenon is not known, it is possible that adoptively transferred Plexin C1–/– DCs in conjunction with LPS promote the activation of host WT DCs which might be able to present exosomes secreted by Plexin C1–/– DCs to host T cells (35). In that regard, it could be of interest to perform the experiment in MHC class II-deficient mice with adoptively transferred OT-2 and Plexin C1–/– DCs.

In conclusion, we found that activated DCs expressed both Plexin C1 and its ligand and that Plexin C1–/– mice mount mildly defective T cell responses in vivo. Our results suggest a role for Plexin C1 in DC migration and/or mobility within the LNs. The absence of strong phenotype in these mice could be explained by the existence of compensatory/redundant mechanisms.


    Abbreviations
 
BMDC   bone marrow-derived dendritic cell
CFSE   5(and 6)-carboxyfluorescein diacetate succinimidyl ester
CHS   contact hypersensitivity
DC   dendritic cell
DNFB   dinitrofluorobenzene
GM-CSF   granulocyte macrophage colony-stimulating factor
GPI   glycosyl-phosphatidylinositol
i.v.   intravenously
LN   lymph node
OVA   ovalbumin peptide
PI-PLC   phosphatidylinositol-specific phospholipase C
s.c.   subcutaneously
SDF-1   stromal-derived factor 1
SE   staphylococcal enterotoxin
Tg   transgenic
WT   wild type

    Notes
 
Transmitting editor: E. Vivier

Received 23 March 2005, accepted 26 April 2005.


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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