CD4 T cell priming in dendritic cell-deficient mice

Paola Castiglioni1, Christina Lu1, David Lo2, Michael Croft3, Pierre Langlade-Demoyen4, Maurizio Zanetti1 and Mara Gerloni1

1 Department of Medicine and Cancer Center, University of California San Diego, 9500 Gilman Drive, La Jolla CA 92093-0837, USA 2 Digital Gene Technologies, La Jolla, CA 92037, USA 3 Division of Immunochemistry, La Jolla Institute for Allergy and Immunology, La Jolla, CA 92121, USA 4 Laboratoire d’Immunite Cellulaire Anti-Virale, Institut Pasteur, 75724 Paris, France

Correspondence to: M. Zanetti; E-mail:mzanetti{at}ucsd.edu
Transmitting editor: S. M. Hedrick


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Bone marrow (BM) chimeras (BMC) generated from mice carrying a null (–/–) mutation in the relB gene of the NF-{kappa}B family represent an ideal model for in vivo studies on the role of dendritic cells (DC) in the adaptive immune response. The spleen and lymph nodes (LN) of relB–/– BMC contain a small number of residual DC, mainly CD8{alpha}+, that fail to up-regulate MHC class II and co-stimulatory molecules after stimulation in vitro. Moreover, residual spleen DC of relB–/– BMC have a 4-fold decrease in the ability to uptake and process soluble model antigen, ovalbumin (OVA), and failed to prime CD4 and CD8 T cells in vitro and in vivo. In addition, they also failed to present OVA peptide to OT-II transgenic T lymphocytes at a normal 1:10 (stimulator:responder) cell ratio. In spite of these multiple DC defects, relB–/– BMC immunized with plasmid DNA targeted to the spleen as the site of immune induction develop a specific CD4+ T cell response comparable to that of relB competent mice. These data demonstrate that CD4 + T cells can be primed in the absence of functional DC and suggest that relB may gauge the T cell response in vivo.

Keywords: dendritic cell, in vivo transgenesis, relB, T cell immunity


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
During the past 15 years it has become apparent that dendritic cells (DC) play a key role in the initiation of the adaptive immune response (1). Their extraordinary capacity to activate T cells is explained in part by their high expression of MHC class II and co-stimulatory molecules, and in part by their localization within secondary lymphoid organs (the marginal zone and T cell-rich areas) which gives them a strategic advantage for the interception and presentation of antigen (2).

DC patrol the organism by virtue of a widespread distribution in lymphoid and non-lymphoid tissues, and by their ability to promptly migrate from non-lymphoid tissues to draining lymph nodes (LN) (3). Migration together with a rapid transition from immature to mature phenotype under phlogogenic stimulation (microbial and inflammatory products) (4) render DC the ideal antigen-presenting cells (APC) for T cell responses (5). Upon phlogogenic stimuli DC up-regulate and stabilize expression of MHC class II molecules and co-stimulatory molecules (CD40, CD80 and CD86), and produce IL-12 for a brief time to enable Th1 responses (6). Activated DC control the emerging immune response through dynamic and molecular adaptations possibly unique among immune cells. However, in spite of abundant in vitro and ex vivo (711) evidence, the extent to which DC play a role in vivo has been probed only in a few instances (12,13) and no studies exist on their role during systemic immunity initiating in the spleen as the site of immune induction.

The spleen is the largest secondary lymphoid organ and the site where adaptive immunity likely develops in response to antigens trapped from the blood stream. Numerous antigens fall in this category, including those associated with infections transmitted by arthropods (bacteria, viruses and protozoa), blood transfusion, tumor cells, certain bacterial infections (e.g. pneumococcal bacteremia), virus-infected cells and ecotropic viruses associated with autoimmune diseases. There exists enough suggestive evidence that in mammals the spleen is not only an organ with hemocatheretic functions, but also a site of primary importance in the development of the adaptive immune response.

To probe the role of splenic DC in CD4 T cell priming we used mice that lack functional bone marrow (BM)-derived DC as a result of a null (–/–) mutation in the relB gene which codes for a component of the NF-{kappa}B complex (14). Homozygous relB–/– mice have an atrophic thymic medulla, possess no LN and lack BM-derived DC (15). However, homozygous relB–/– mice possess CD8{alpha}+ lymphoid DC in the spleen (16). BM chimeras (BMC) generated by transferring homozygous –/– relB BM cells into lethally irradiated (1100 rad) hemizygous (+/–) relB recipients carry the same DC defect as relB–/– mice, but have a longer lifespan (14). In contrast to the severe deficit in DC, B cells are apparently normal. Experiments in vitro show that B lymphocytes of relB–/– mice proliferate to lipopolysaccharide (LPS) (our unpublished results), undergo isotype switch (17) and express normal levels of chemokine receptors (18). Experiments in vivo show that following immunization with virus or DNA relB–/– BMC produce antibodies and form germinal centers (14,19). Thus, relB–/– BMC constitute an ideal animal model to test the role of spleen DC in the generation of a T cell response.

In light of the foregoing we decided to study the ability of residual spleen DC in relB–/– BMC to prime T cell responses to a model antigen, ovalbumin (OVA), and compare these results with those obtained in response to DNA immunization where antigen is synthesized directly within the spleen. In this latter approach, termed somatic transgene immunization (20), immunity ensues as a result of direct intraspleen inoculation of plasmid DNA (21) coding for an immunoglobulin heavy (Ig H) chain gene. Previously, we showed that plasmid DNA is taken up by B lymphocytes that remain in the spleen (22) and, since the transgene is under the control of a B cell-specific promoter, B lymphocytes begin synthesis of transgenic Ig. By using Ig H chain genes coding for heterologous T cell epitopes in the complementarity determining regions of the variable domain (23) it has been possible to trigger specific CD4 (24) T cell responses. The process of in vivo transgenesis is also followed by secretion of sizeable amounts (up to 70 ng/ml) of transgenic Ig, and these are thought to reinforce immunity locally and spread it to secondary lymphoid organs systemically (24).


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Mice and generation of BMC
C57BL/6 mice (8–10 weeks old) were purchased from the Jackson Laboratories (Bar Harbor, ME). Homozygous relB –/– mice were bred in the animal facility of the University of California San Diego. BMC were generated by injecting i.v. 5 x 10 6 BM cells from relB–/– (15) or C57Bl/6 mice into lethally irradiated (1100 rad) hemizygous relB+/– or C57BL/6 recipients respectively. Mice were used 5–6 weeks after BM transfer. OT-I (25) and OT-II (26) TCR transgenic mice were the kind gift of Drs M. Croft and S. Schoenberger (La Jolla Institute for Allergy and Immunology).

Plasmid DNA, proteins and synthetic peptides
Plasmid {gamma}1NV2NA3 was engineered as described in (23). Plasmid DNA was purified using a Qiagen Megaprep kit (Qiagen, Chatsworth, CA). Purified plasmid was stored at –20°C until use. OVA (grade VII) was purchased from Sigma (St Louis, MO). FITC-conjugated OVA was purchased from Molecular Probes (Eugene, OR). Synthetic peptides NANPNVDPNANP, (–NVDP–), its control NANPNANPNANP, (NANP)3, and OVA peptide ISQAVHAAHAEINEAGR (amino acids 323–339) were synthesized in the Peptide Chemistry Facility of the California Institute of Technology (Pasadena, CA).

DC isolation and culture
DC were isolated from spleens and LN of relB –/– BMC, C57Bl/6 BMC, relB–/– and C57Bl/6 mice respectively. Spleens and LN were digested with collagenase D (0.5 mg/ml) and DNase I (0.1 mg/ml) (Sigma) for 25 min at room temperature with frequent pipetting to break up fragments. The undigested fibrous material was removed by filtration through a nylon mesh. Samples of 108 spleen cells and 107 LN cells were cultured in six-well plates with IMDM (Gibco/BRL, Grand Island, NY) supplemented with 10% heat-inactivated FCS (HyClone, Logan, UT), recombinant mouse granulocyte macrophage colony stimulating factor (GM-CSF, 1000 U/ml; PharMingen, San Diego, CA) and recombinant mouse IL-4 (4 ng/ml; R & D Systems, Minneapolis, MN). Cultures were split and the medium was changed when necessary. On day 4 LPS (Escherichia coli, 10 µg/ml; Sigma) was added to cultures for an additional 24 h. For in vitro or in vivo experiments with OVA and OVA peptide, DC from spleens of relB–/– BMC, C57Bl/6 BMC and C57Bl6 mice were additionally purified using a discontinuous gradient. Briefly, after tissue digestion spleen cells were resuspended in PBS (Gibco/BRL) and OptiPreo (3:1 v/v; Accurate, Westbury, NY) giving a 1.085 g/ml solution. This suspension was over-layered with 5 ml of a 1.065 g/ml iodixanol solution and 3 ml of PBS, and centrifuged at 1600 r.p.m. for 15 min. The low-density fraction was collected and depleted of B cells using magnetic beads coated with a mAb against mouse CD19 (Miltenyi Biotech, Auburn, CA) followed by sorting on a MACS separation column (Miltenyi Biotech). CD19 cells were additionally enriched in DC using CD11c MicroBeads (Miltenyi Biotech). Splenic DC were 85–90% pure.

T cell priming in vitro
The ability of residual spleen DC of relB–/– BMC to prime naive T lymphocytes was assessed in an in vitro culture system using naive C57Bl/6 CD4 and CD8 T lymphocytes and OVA (Sigma) as the antigen. C57Bl/6 spleen CD4 T lymphocytes were isolated using magnetic beads coated with a mAb against mouse CD4 (Miltenyi Biotech) followed by sorting on a MACS magnetic separation column (Miltenyi Biotech). The CD4 T cells obtained were 90% pure as determined by FACS analysis. CD8 T lymphocytes were enriched using magnetic beads coated with mAb against mouse CD4, CD19 and CD11c (Miltenyi Biotech). The CD8 T cell population was >60% pure. Purified spleen DC from relB –/– BMC, C57Bl/6 BMC or C57Bl/6 were plated on 96-well round-bottomed plates at 0–2 x 104 cells/well and incubated overnight with OVA (0–500 µg/ml final concentration). Antigen-pulsed DC were irradiated (3000 rad) and syngeneic naive CD4 or CD8 T lymphocytes were added (5 x 104 cells/well) (27). Super natants were harvested on day 5 for cytokine detection and the proliferative response was measured by the uptake of [3H]Thymidine added (1 µCi/well) on day 6 for 18–20 h. The assay was performed as detailed below. In additional experiments we pulsed purified spleen DC from relB–/– BMC and C57 Bl/6 BMC with OVA323–339 peptide (1 µg/ml) and used them (0–5 x 104 cells/well) to stimulate OT-II CD4 T lymphocytes (105 cell/well). OT-II splenocytes were depleted of APC and CD8 T cells using a cocktail of mAb and rabbit complement. The mAb were M5114 anti-I-A, CA4 anti-class II, RA3.6.B2 anti-B220, PK136 anti-NK, M1/70 anti-CD11b and 3.155 anti-CD8 (kindly provided by Stephen Schoenberger, La Jolla Institute for Allergy and Immunology).

In vivo immunization procedures
DNA immunization
Mice were inoculated intraspleenically with 100 µg of plasmid DNA in 50 µl of sterile saline solution as previously described (21).

Immunization with OVA
The ability of residual spleen DC in relB–/– BMC to uptake and process OVA was tested by i.v. injection of soluble OVA (3 mg/mouse) dissolved in PBS. After 20 h DC from pools of two to four spleens were isolated as described above and cultured (0–6000/well) with 30,000 OVA-specific CD8 T cells from OT-I mice (>60% purity). Purified DC were also cultured (0–105/well) with 105 OVA-specific CD4 T cells from OT-II mice (> 50% purity). Culture medium contained murine recombinant IL-2 (50 U/ml). Supernatants for cytokine detection were harvested after 40 h and the T cell proliferation response measured as detailed below. PBS alone was injected in C57Bl/6 mice to control for the response of T cells to DC in the absence of OVA (background response). In uptake studies FITC-conjugated OVA was injected i.v. (2 mg/mouse). Spleen DC were isolated 1 h later from a pool of three or four spleens. Non-fluorescent native OVA was injected into control mice to provide the DC background for FITC labeling (28).

[3H]Thymidine incorporation assay
Fourteen days after DNA inoculation mice were sacrificed and the spleens were removed to prepare single-cell suspensions. Red blood cells were eliminated with lysis buffer (Sigma). Lymphocytes were cultured (106 cells/ml) in RPMI 1640 medium (Irvine, Santa Ana, CA) supplemented with 7.5% FCS and 50 µM 2-mercaptoethanol, in the presence or absence of synthetic peptide –NVDP– or control peptide (NANP)3 (50 µg/ml) in triplicate wells by incubation at 37°C in 10% CO 2 for 3 days. [3H]Thymidine was added at 1 µCi/well and the cells were incubated for 16–18 h at 37°C. Cells were harvested onto glass fiber filter mats using a Tomtec cell harvester and the radioactivity was measured in a liquid scintillation counter (Betaplate; Wallac, Turku, Finland). Results are expressed as means ± SD of c.p.m. of triplicate cell cultures.

Detection of cytokines
Culture supernatants were harvested 40 h after initial seeding (or 5 days after, for in vitro T cell priming experiments) and were stored at –20°C. The supernatants from three separate triplicate cultures were pooled for each mouse. IL-2 activity was determined either through a bioassay utilizing the IL-2- and IL-4-dependent NK.3 cells in the presence of anti-IL-4 (purified from the 11B11 cell line; ATCC, Rockville, MD) as described previously (24) or using the OptEIA mouse IL-2 set (PharMingen, San Diego, CA) (Figs 3 and 5). IL-4 and IFN-{gamma} were measured in the same culture supernatants by ELISA as described previously (24). Standard curves were constructed with purified IL-2, IL-4 and IFN-{gamma}. Tests were performed in duplicate.



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Fig. 3. Spleen DC of relB –/– BMC fail to prime CD4 T cell responses against OVA. (A) OVA-pulsed spleen DC from relB–/– BMC or C57Bl/6 BMC were used to prime naive CD4 T lymphocytes from the spleen of C57Bl/6 mice. CD4 T cells (5 x 104) were cultured with different concentrations of DC as indicated in each panel. The experiments shown were performed pulsing DC with OVA at 500 µg/ml. For IL-2 detection culture supernatants were harvested on day 5. [3H]Thymidine incorporation was measured on day 6. The inset in the right panel shows that non-pulsed DC (2 x 104) do not elicit any proliferative response. (B) relB–/– BMC and C57Bl/6 BMC were injected i.v. with OVA (3 mg). At 20 h after injection, DC were purified from a pool of two to four spleens and cultured at different concentrations (0–105 cells/well) with 105 OT-II CD4 T lymphocytes. [3H]thymidine incorporation was measured after 72 h. Proliferation assays were run in triplicate. Results are from a single experiment representative of two experiments with a total of five relB–/– BMC examined.

 
Phenotype analysis
The phenotyping of DC and their quantitation were performed on collagenase-treated spleen and LN cells ex vivo and after in vitro culture. Single-cell suspensions from relB –/–BMC, C57Bl/6 BMC, relB–/– and C57Bl/6 were incubated with magnetic beads coated with a mAb against mouse CD11c (Miltenyi Biotech) and then sorted on a MACS separation column (Miltenyi Biotech). Cells selected on the basis of CD11c expression were then stained with biotin-conjugated mAb against mouse CD11c (clone HL3) (PharMingen). After 20–30 min at 4°C, cells were washed with PBS containing 0.5% BSA and 0.05% NaN3, and stained with CyChrome–streptavidin and one of the following phycoerythrin-conjugated mAb: anti-I-Ab (clone AF6-120.1), anti-CD80 (clone 16-10A1), anti-CD40 (clone 3/23), anti-Db (clone KH95) and anti-CD8 (clone 53-6.7) (PharMingen). Spleen DC from FITC–OVA-injected mice were isolated and double-stained with CD11c (clone HL3) mAb. Cells were analyzed by flow cytometry on a FACSCalibur (Becton Dickinson) and gates were set to select for viable DC. For DC quantitation cells were gated on CD11c/ I-Ab double-positive cells.


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Phenotypic characterization of residual DC in relB–/– BMC
The phenotypic characteristics of residual DC in the spleen of relB–/– BMC were analyzed and compared with those of C57Bl/6 BMC, relB–/– and C57Bl/6 mice. Analyses were performed ex vivo (i.e. immediately after tissue harvest) and in vitro following culture with IL-4 (4 ng/ml) and GM-CSF (1000 U/ml) for 96 h plus LPS (10 µg/ml) during the last 24 h. FACS analyses were carried out on CD11c+ cells double-stained for MHC class II (I-Ab), CD40 or CD80. Splenic DC from both homozygous relB–/– mice and relB–/– BMC (Fig. 1 A) have a much reduced expression of these cell surface molecules compared with C57Bl/6 mice or their BMC. An analysis of LN DC from relB –/– BMC showed a very similar defect (Fig. 1 B). Importantly, in relB–/– BMC and relB–/– mice up-regulation of MHC class II and CD40 or CD80 after in vitro culture and stimulation with LPS was minimal if any. In addition, even though CD11c+ cells were counted ex vivo in both relB–/– BMC and relB–/– mice, no increase in number was noted after in vitro culture and LPS stimulation (Table 1). Consequently, lack of survival and maturation in culture made it impossible to profile CD40 and CD80 on these cells. Taken together these results suggest that CD11c+ cells in relB–/– BMC have little if any constitutive expression of MHC class II and co-stimulatory molecules that fail to up-regulate after in vitro culture and LPS stimulation.



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Fig. 1. Surface phenotype analysis and quantitation of DC isolated from spleens (A) of C57Bl/6, C57Bl/6 BMC, relB –/– and relB–/– BMC, and LN (B) of C57Bl/6, C57Bl/6 BMC and relB–/– BMC. CD11c+ cells were analyzed for the expression of MHC class II (I-Ab), CD80 and CD40 surface molecules. The results are shown as histograms with fluorescence intensity on the x -axis and cell number on the y-axis. Dotted lines represent ex vivo staining of spleen cells, whereas thick lines represent staining after 4 days in culture with GM-CSF, IL-4 and LPS which was added during the following 24 h as described in Methods. Solid histograms refer in each instance to the background (i.e. autofluorescence).

 

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Table 1. DC in the spleen and LN of relB–/– BMC
 
Splenic DC of relB –/– BMC fail to prime T cells to soluble antigen
We tested the ability of residual splenic DC of relB –/– BMC to process and present a model antigen (OVA) in vitro and in vivo. Both approaches required the isolation and enrichment (~85–90% purity) of DC from the spleen of relB–/– BMC, C57Bl/6 BMC and C57Bl/6 mice as controls (Fig. 2).



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Fig. 2. Surface phenotype analysis of DC purified from spleen cell suspensions of C57Bl/6 (a and b), C57 Bl/6 BMC (c and d) and relB–/– BMC (e and f). After purification DC were stained with an anti-CD11c, anti-MHC class II (I-Ab) and anti-CD8{alpha} mAb. The percentage of CD11c/MHC class II/CD8{alpha}-positive cells is shown in each panel. Results shown are representative of three to five independent purifications of pools of two to four mice in each instance.

 
An initial in vitro experiment showed that purified spleen DC from relB–/– BMC pulsed overnight with OVA at 37°C failed to prime naive C57Bl/6 CD4 T lymphocytes. In fact, after a 6-day culture CD4 T cell proliferation occurred in cultures seeded with antigen-pulsed DC from C57Bl/6 BMC but not relB –/– BMC (Fig. 3A, left panel). A syngeneic mixed lymphocyte reaction can be ruled out in view of the fact that 2 x 104 DC/well in the absence of OVA did not induce any proliferation (see inset). Lack of CD4 T cell priming in cultures where relB–/– BMC DC served as the APC was mirrored by the absence of IL-2 production (Fig. 3A, right panel). Similarly, no T cell activation was found when antigen uptake and processing was first allowed to occur in vivo. To this end, relB–/– BMC, C57Bl/6 BMC and C57Bl/6 mice, were injected i.v. with OVA (3 mg/mouse) to enable antigen uptake by splenic DC (29). At 20 h after injection, DC were purified from the spleens and used as APC to activate naive, OVA-specific OT-II CD4 T lymphocytes (26). As shown in Fig. 3(B), CD4 T cell proliferation was observed in cultures seeded with DC from C57Bl/6 BMC, but not with DC from relB–/– BMC or from mice injected with PBS as controls (data no shown).

As an additional proof of the profound deficit in T cell priming by residual spleen DC of relB–/– BMC, new experiments were performed to explore the ability of OVA-pulsed DC to prime naive C57Bl/6 CD8 T cells. IL-2 and IFN-{gamma} were detected only in cultures seeded with in vitro-pulsed DC purified from C57Bl/6 and C57Bl/6 BMC, but not relB–/– BMC (Fig. 4A). Similarly, in experiments in which uptake of OVA was performed in vivo followed by in vitro presentation to OVA-specific OT-I CD8 T lymphocytes (25), proliferation was observed only in cultures seeded with DC from C57Bl/6 and C57Bl/6 BMC, but not relB–/– BMC (Fig. 4B, right panel). No IFN-{gamma} was detected in cultures seeded with DC from OVA-injected relB –/– BMC (Fig. 4B, left panel).



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Fig. 4. Spleen DC of relB –/– BMC fail to prime CD8 T cell responses against OVA. (A) OVA-pulsed spleen DC from relB–/– BMC, C57Bl/6 BMC and C57Bl/6 were used to prime naive CD8 T lymphocytes from the spleen of C57Bl/6 mice. CD8 T cells (5 x 104) were cultured with different concentrations of DC (2 x 104, solid bars; 104, open bars). For IL-2 and IFN-{gamma} detection culture supernatants were harvested on day 5. The experiments shown were performed pulsing DC with OVA at 500 µg/ml. (B) relB –/– BMC, C57Bl/6 BMC and C57Bl/6 were injected with OVA (3 mg) i.v. At 20 h after injection, DC were purified from a pool of two to four spleens and cultured at different concentrations (6 x 103, solid bars; 3 x 103, open bars) (left panel) with 3 x 10 4 OT-I CD8 T lymphocytes. The supernatants for IFN-{gamma} detection were collected after 40 h and [3H]thymidine incorporation was measured after 72 h. C57Bl/6 mice injected with PBS were used as a control. Cytokine detection assays were run in duplicate. Proliferation assays were run in triplicate.

 
Collectively, these results show that residual spleen DC in relB–/– BMC are unable to prime T cell responses whether antigen uptake is allowed to occur in vitro or in vivo.

Deficit in antigen uptake and presentation by residual spleen DC in relB–/– BMC
Lack of T cell priming could be due to a deficit of antigen uptake and/or lack of presentation to T cells. To decide among these possibilities two experiments were performed. First, we assessed the ability of splenic DC from relB–/– BMC to capture FITC-conjugated soluble OVA. Within 1 h of injection spleen DC purified from C57Bl/6 mice and C56Bl/6 BMC showed extensive uptake of OVA (>60%) (Fig. 5 A). In contrast, OVA uptake by DC purified from relB –/– BMC was much lower (14.8%). Thus, residual spleen DC in relB–/– BMC have a much-reduced (4-fold) ability to capture soluble antigen relative to relB competent mice. Second, we tested the ability of residual spleen DC from relB–/– BMC to present the OVA 323–339 peptide to OT-II CD4 T lymphocytes (Fig. 5 B). Not surprisingly at a standard (1:10) stimulator (DC):responder (OT-II cells) ratio DC of relB–/– BMC failed to induce T cell priming, whereas DC from control C57Bl/6 BMC showed a vigorous response. DC from relB–/– BMC promoted T cell proliferation only when added in higher number/well that also altered the stimulator to responder ratio from 1:10 to 1:2. At this latter ratio DC from relB–/– BMC induced a response that was ~3-fold lower than DC from C57Bl/6 BMC (left panel). The deficit in T cell priming was mirrored by a very low detection of IL-2 in cultures seeded with DC from relB–/– BMC (right panel). Collectively, it appears that the defect in APC function by residual spleen DC in relB–/– BMC is a complex one, including a much-reduced capacity to capture soluble antigen and an impaired ability to present processed antigen peptide.



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Fig. 5. Antigen uptake and presentation by residual spleen DC in relB–/– BMC. (A) Mice were injected i.v. with 2 mg of FITC-conjugated OVA. DC were isolated 1 h later from pools of three to four spleens, stained with an anti-CD11c and anti-CD40 mAb, and analyzed by flow cytometry. The percentage of FITC fluorescence within the DC is shown in each panel. (B) OT-II CD4 T lymphocytes were stimulated at different ratios with OVA323–339 peptide pulsed spleen DC from relB–/– BMC and C57Bl/6 BMC. DC pulsed with –NVDP– peptide were used as control. [3H]Thymidine incorporation (left panel) was measured after 72 h. Proliferation assays were run in triplicate. For IL-2 detection (right panel) supernatants were harvested after 40 h of culture. Tests were run in duplicate.

 
CD4 T cell priming in the spleen of rel B–/– BMC
The above defects in APC function by spleen DC in relB –/– BMC resulted in their inability to prime T cells against soluble antigens. Thus it became interesting to see if a CD4 T cell response could be induced in vivo using somatic transgene immunization (21) model DNA vaccination. Experiments were performed using a plasmid {gamma}1NV2NA3 comprising an Ig H chain gene whose V domain is engineered to code for two dodecapeptides from the circumsporozoite protein of Plasmodium falciparum malaria parasite: the Th cell determinant –NVDP– in the second complementarity determining region (CDR2) and the B cell epitope (NANP)3 in the third complementarity determining region (CDR3) (23). The Th cell determinant and the B cell epitope differ by only 2 amino acid residues (A -> V and N -> D) at positions 5 and 6, but this is sufficient to account for absolute specificity of the T cell response in C57Bl/6 mice (24).

Spleen cells were harvested 14 days after intraspleenic DNA inoculation, a time corresponding to the peak of the CD4 T cell response (24). In three independent experiments we found that all immunized relB–/– BMC responded with activation of CD4 T cells specific for the Th cell determinant (Table 2 ). Overall, the response in relB–/– BMC was weaker than in unmanipulated C57Bl/6 mice, but comparable to C57Bl/6 BMC used as controls, reflecting possible effects by lethal irradiation/BM reconstitution. No proliferation was observed using spleen lymphocytes from a naive C5Bl/6 mouse, indicating that DNA immunization specifically primed naive T cells rather than expanding memory precursor T cells. The magnitude and specificity of the proliferative response by spleen lymphocytes were mirrored by the levels of IL-2 in the corresponding culture supernatants (Table 3). IL-2 was detected in the supernatants of all cultures irrespective of the experimental category. IFN-{gamma} and IL-4 were also detected, albeit in different amounts, in agreement with the fact that somatic transgene immunization expands mainly uncommitted (Th0) CD4 T cells (24). Thus, the deficit in BM-derived DC in relB –/– BMC did not affect the magnitude or the type of CD4 T cell response following immunization via intraspleenic inoculation of DNA. From the foregoing, we conclude that following immunization by directly targeting the spleen as the site of immune induction, a specific CD4 T cell response can be induced in the absence of functional DC.


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Table 2. CD4 T cell proliferative response in the spleen of relB–/– BMC
 

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Table 3. Cytokine analysis in the spleen
 

    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
DC are considered a key player in T cell activation and are possibly the most potent regulators of the immune response (1). DC express a variety of surface receptors for the internalization of antigens (4) and go through the maturation process quite rapidly (30). In vivo clustering and interaction between DC and T cells have been visualized in draining LN, and found to be rapid (~24 h) (12). Immunization with protein antigen in Freund’s’ adjuvant showed that DC and not B cells present antigen to CD4 T cells in LN (31). However, after i.v. injection of soluble antigen, DC are only as good as B cells in capturing antigen and expressing MHC–peptide complex (13). Thus, whether DC or B cells serve as the APC for CD4 T cells seems to depend on the characteristics of the immunization (antigen depot in immunological adjuvant versus soluble antigen). Here we used rel B–/– BMC whose spleen DC, as demonstrated here, are unable to prime T cells to soluble antigen or processed peptide, to see whether T cell priming in these mice is still possible following DNA immunization targeted to the spleen as the site of immune induction (22).

In the mouse, splenic DC of BM origin have been distinguished into two broad categories on the basis of surface expression of CD8{alpha} (32). These two phenotypically different categories have different localization in lymphoid tissues (2) and possibly different immunological functions (33). Homozygous relB–/– mice and relB–/– BMC (–/– -> C57Bl/6) lack CD8a DC, but have residual CD8{alpha}+ DC (16). In our hands the absolute number of total splenic DC is low (Table 1). Moreover, these cells have a much-impaired ability to undergo maturation in vitro (Fig. 1). If compared with the findings of Wu et al. (16), our results show comparable expression of MHC class II molecules on residual spleen CD11c+ cells of relB –/– BMC (Fig. 2). However, the severely compromised APC functions of residual spleen DC of relB –/– BMC documented here are in apparent contrast with the reported ability of these mice to induce an allo mixed lymphocyte reaction (16). There may be several reasons for this discrepancy. (i) Our BMC are generated using as recipients irradiated heterozygous relB (+/–) mice and not C57Bl/6 mice. (ii) We used an 85–95% pure DC population, whereas in the mixed lymphocyte culture experiment DC were purified by a different method and were used at a non-specified level of purity. (iii) More importantly, an allo mixed lymphocyte reaction reflects stimulation of T cells by DC, but does not imply processing and presentation of MHC antigens by self-MHC molecules. Therefore, while the two studies are not comparable, our findings establish that spleen DC of relB–/– BMC have a severely impaired APC function vis-à-vis nominal soluble antigen.

We documented multiple defects in APC function of residual spleen DC in relB–/– BMC. Injection of the FITC-conjugated OVA disclosed a markedly reduced ability to capture circulating soluble antigen in vivo. However, this alone cannot explain why relB–/– DC failed to present OVA after in vitro pulsing because this experiment was performed at a high antigen concentration (500 µg/ml). This also argues against a role of residual spleen DC during somatic transgene immunization where the concentration of the transgene product at peak time is <=70 ng/ml (21), i.e. at least 4 x 103-fold less than in the in vitro pulsing experiment. As to their ability to present peptide antigen, our data show that residual spleen DC from relB –/– BMC cannot prime OVA-specific CD4 T cells unless used in high number. Even so, their antigen-presenting capacity is ~3-fold lower than that of relB competent spleen DC.

The experiments presented here are the first direct evidence that residual DC in relB–/– BMC are functionally defective, and de facto unable, to induce antigen-specific T cell responses. Whether the defect is intrinsic to the residual CD8a + subset or secondary to the lack of relB-competent CD8a DC cannot be established. The fact that both DC types appear to originate from a common myeloid progenitor (34,35) and the relB –/– BM fails to generate functional DC in hemizygous recipients suggest that the immunological incompetence of CD8{alpha}+ DC documented here may be part of the same genetic defect.

If residual DC cannot prime T cells, how are T cells primed following somatic transgene immunization? Several considerations point to a direct involvement of in vivo transgenic B lymphocytes. We have already documented that the transgene persists in spleen B lymphocytes for a protracted period of time (22). Notably, the Ig H transgene is under control of a B cell-specific promoter. Also, in our hands, spleen-derived DC are not transfected by the plasmid DNA used in the present study (our unpublished data). Finally, the amount of secreted transgenic product (<=70 ng/ml) is too low to enable uptake by relB–/– BMC DC and T cell priming. In addition, the possibility of T cell activation by cross-priming (36), albeit plausible, can be ruled out based on the fact that relB–/– BMC fail to induce a cytotoxic T lymphocyte response against a model tumor antigen via cross-priming (37). Taken together, the scenario we favor at this point is that after DNA injection into the spleen of relB–/– BMC, antigen synthesis, processing and presentation occur in B lymphocytes as the default pathway. Whatever the mechanisms might be, our results provide evidence that CD4 T cell priming in the spleen as the site of immune induction can occur in the absence of functional DC. While residual splenic DC of relB–/– BMC failed to prime T cells against the model antigen OVA in vitro and in vivo, specific T cell priming in these mice was demonstrated following somatic transgene immunization. Thus, transgenic B lymphocytes may be the key APC in our experiments since B lymphocytes can present their own Ig V region peptides to CD4 T cells in vitro (38) and the expansion of T cells in vivo can be driven by activated (3941) or naive (42) B lymphocytes. Preliminary experiments show that that B lymphocytes transgenic for a plasmid coding for OVA323–339 CD4 peptide can prime naive CD4 T cells from OT-II mice in the absence of DC (our unpublished data), strengthening the findings of this paper.

In conclusion, we demonstrate that CD4 T cell priming during an adaptive immune response originating in the spleen as the site of immune induction can occur without any manifest role for resident DC. Our findings are relevant to the mechanisms of antigen presentation during a systemic adaptive immune response where the initial interaction between antigen, B lymphocytes as the APC and T cells occurs in the spleen. While bringing new elements to our understanding on the function of DC and B cells in the local splenic environment, and their respective role in the initiation of T cell immunity, these findings begin to elucidate the role played by the relB gene in gauging the adaptive T cell response in vivo through its control of DC function.


    Acknowledgements
 
The authors thank Drs Ralph Steinman, Stephen Schoenberger and Javier Hernandez for constructive criticism and the generous supply of OT-II mice (Stephen Schoenberger). This work was supported in part by NIH grants RO1 CA 77427 and R21 AI49774.


    Abbreviations
 
APC—antigen-presenting cell

BM—bone marrow

BMC—bone marrow chimera

DC—dendritic cell

GM-CSF—granulocyte macrophage colony stimulating factor

OVA—ovalbumin


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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