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The Journal of Immunology, 2007, 179, 3821-3830
Copyright © 2007 by The American Association of Immunologists, Inc.
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The Role of B Cells in the Development of CD4 Effector T Cells during a Polarized Th2 Immune Response1

Qian Liu*, Zhugong Liu*, Cristina T. Rozo*, Hossein A. Hamed*, Farhang Alem*, Joseph F. Urban, Jr.{dagger} and William C. Gause2,*

* Department of Medicine, New Jersey Medical School, University of Medicine and Dentistry of New Jersey, Newark, NJ 07103; and {dagger} Nutrient Requirements and Functions Laboratory, Beltsville Human Nutrition Research Center, U.S. Department of Agriculture, Beltsville, MD 20705-2350


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Previous studies have suggested that B cells promote Th2 cell development by inhibiting Th1 cell differentiation. To examine whether B cells are directly required for the development of IL-4-producing T cells in the lymph node during a highly polarized Th2 response, B cell-deficient and wild-type mice were inoculated with the nematode parasite, Nippostrongylus brasiliensis. On day 7, in the absence of increased IFN-{gamma}, IL-4 protein and gene expression from CD4 T cells in the draining lymph nodes were markedly reduced in B cell-deficient mice and could not be restored by multiple immunizations. Using a DO11.10 T cell adoptive transfer system, OVA-specific T cell IL-4 production and cell cycle progression, but not cell surface expression of early activation markers, were impaired in B cell-deficient recipient mice following immunization with N. brasiliensis plus OVA. Laser capture microdissection and immunofluorescent staining showed that pronounced IL-4 mRNA and protein secretion by donor DO11.10 T cells first occurred in the T cell:B cell zone of the lymph node shortly after inoculation of IL-4–/– recipients, suggesting that this microenvironment is critical for initial Th2 cell development. Reconstitution of B cell-deficient mice with wild-type naive B cells, or IL-4–/– B cells, substantially restored Ag-specific T cell IL-4 production. However, reconstitution with B7-1/B7-2-deficient B cells failed to rescue the IL-4-producing DO11.10 T cells. These results suggest that B cells, expressing B7 costimulatory molecules, are required in the absence of an underlying IFN-{gamma}-mediated response for the development of a polarized primary Ag-specific Th2 response in vivo.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Naive CD4 T cells are thought to first interact with dendritic cells (DC),3 which express MHC class II-peptide complexes that engage the TCR, in the T cell zone of peripheral lymphoid tissues (1, 2). Expression of costimulatory molecules and cytokines by DC, as well as cognate interactions, can influence the capacity of CD4 T cells to undergo division (2). Once activated, CD4 T cells migrate toward the B cell zone where they form conjugates with B cells at the junction of the T and B cell regions (3). Activated B cells are competent to act as APCs (4, 5) and in this capacity can also express both cytokines (6, 7, 8, 9) and costimulatory molecules (8, 10). B cells may be necessary to support continued CD4 T cell expansion, which is required for a sustained response leading to effector function and memory (11, 12). B cells may also preferentially favor the development of IL-4-producing Th2 cells, with some studies suggesting that B cells may down-regulate the level of IL-12 released by DC, thereby inhibiting the development of IFN-{gamma}-producing Th1 cells (13). Other studies have suggested that the prolonged stimulation required to support the multiple rounds of T cell division necessary for polarized Th2 cell differentiation may require APC interactions with B cells as well as DC (8, 12).

Most studies of B cell knockout mice have examined the response to Ags that evoke mixed Th1/Th2 cytokine patterns (14, 15, 16, 17, 18). In studies with Trichuris muris, the host protective mucosal Th2 response was inhibited in B cell-deficient mice, whereas a Th1 response was sustained. Administration of anti-IL-12 Ab restored the resistance, suggesting that B cells blocked IL-12 thereby promoting the Th2 response (19). Similar results were obtained following parenteral immunization of B cell-deficient mice with Schistosoma mansoni eggs, in that immune deviation from a Th2 to a Th1 response was observed (17). In another series of studies, immunization with alum-precipitated OVA and Bordetella pertussis in B cell-deficient mice was also associated with the preferential development of Th1 cells, although in this system anti-IL-12 Ab administration did not favor the Th2 response and a Th1 response was sustained (12). Because of the mixed Th1/Th2 nature of these immune responses that have been examined in B cell-deficient mice, it has been difficult to exclude the possibility that the main Th2 cell promoting effect of B cells was their suppression of Th1 cell differentiation.

The immune response to the trichostrongylid nematode parasite, Nippostrongylus brasiliensis (Nb), is unusual in that it elicits a highly polarized Th2 response associated with marked elevations in CD4 T cell-derived IL-4 and IL-13, but undetectable elevations in IFN-{gamma} (20). Furthermore, blockade of costimulatory molecules does not lead to increased IFN-{gamma} production (21); in contrast, Th1 responses are often observed following costimulatory blockade during mixed Th1/Th2 responses (22, 23, 24). Previous studies have further shown that Nb can drive the rapid differentiation of Th2 cells in lymph nodes draining the site of intracutaneous inoculation in the ear, providing a useful model system for examining Th2 cell differentiation in vivo (21). In this system, it was further demonstrated that Nb can act as an adjuvant to drive the differentiation of nonparasite Ag-specific naive T cells to Th2 effector cells in vivo (21).

Specific microenvironments in the lymph node where cytokine-producing T cells first develop may play an essential role in naive Th cell activation and differentiation. Identification of these microenvironments may provide important insights into the factors that determine Th effector cell cytokine expression patterns. Previous studies have suggested that the T:B zone is critical for optimal T cell proliferation and that ~70% of Ag-specific T cells interact with B cells at the T:B zone on day 2 postimmunization (3). A more recent study using laser capture microdissection (LCM) has shown that Th2 cytokines are expressed preferentially in the B cell zone (25), although in the latter study, the cell source was not identified. As yet, few studies have examined in which lymph node microenvironment T cells first express IL-4 mRNA or protein.

In this study, we have used Nb infection and an Ag-specific T cell transfer system to examine the role of B cells in the development of Th2 cells in vivo during a polarized Th2 response. Our studies show that B cells are not required for the initial activation of T cells but are required for their subsequent proliferation and differentiation into Th2 cells, even in the absence of an underlying IFN-{gamma} response; furthermore, for the first time, we show that IL-4 expressing Ag-specific T cells first develop in the T:B zone of the lymph node.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

B cell-deficient mice (BALB/c Jhdtm1) from Taconic Farms, IL-4-deficient mice (IL-4–/–; purchased from The Jackson Laboratory), B7-1/B7-2-deficient mice, and DO11.10 TCR transgenic mice were on an inbred BALB/c background. The DO11.10 mice contain a large population of CD4 T cells that express a TCR specific for chicken OVA peptide 323–339-I-Ad complexes. This TCR is uniquely recognized by the KJ1-26 anti-clonotypic mAb (26). All the mice were maintained and bred in a specific pathogen-free facility during the experiments at the New Jersey Medical School-University of Medicine and Dentistry of New Jersey (Newark, NJ) research animal facility. The studies have been reviewed and approved by Institutional Animal Care and Use Committee at New Jersey Medical School. The experiments in this study were conduced according to the principles set forth in the Guide for the Care and Use of Laboratory Animal, Institute of Animal Resources, National Research Council, Department of Health, Education and Welfare (publication no. NIH 78-23).

Parasite infection and OVA immunization

Mice were inoculated intracutaneously in the ear with 300 infective Nb third-stage larvae (L3). HPLC-purified OVA peptide 323–339 with the sequence I-S-Q-A-V-H-A-A-H-A-E-I-N-E-A-G-R-COOH was synthesized by Molecular Resource Facility at New Jersey Medical School. In some experiments, Nb L3 and 30 µg of OVA peptide (Nb plus OVA) were injected intracutaneously in the ears of DO11.10 T cell transfer recipient mice.

ELISPOT

ELISPOT assays were done as previously described (21). Briefly, single-cell lymph node suspensions were prepared in RPMI 1640 containing 10% heat-inactivated FCS, 100 U/ml penicillin, 100 µg/ml streptomycin, and 2 mM L-glutamine (all from Invitrogen Life Technologies). A total of 0.5 x 106 cells were seeded into each well of an anti-IL-4 (clone BVD4-1D11.2), a gift from Dr. F. D. Finkelman (University of Cincinnati College of Medicine Cincinnati, OH), coated Immulon IV 96-well microtiter plate (Microtiter). After overnight culture, the plate was washed several times with PBS followed by washes with PBS/Tween 20. Secondary biotinylated anti-IL-4 Ab was diluted in PBS/0.05% Tween 20/5% FCS, added at 100 µl/well, and incubated overnight at 4°C. Plates were then washed, and a 1/2000 dilution of Streptavidin-alkaline phosphatase (Jackson ImmunoResearch Laboratories) was added. Plates were developed and results were quantitated.

Cytokine gene expression by RT-PCR

For RT-PCR, total RNA was extracted from tissue and then reverse transcribed as previously described (27). Real-time PCR kits (PE Applied Biosystems), specific for individual cytokines or rRNA, were used to quantitate differences in gene expression, and all data were normalized to constitutive rRNA values. The PE Applied Biosystems 7700 sequence detector was used for amplification of target mRNA, and quantification of differences between treatment groups was calculated according to manufacturer’s instructions.

Adoptive transfers

Peripheral lymph nodes and spleens were harvested from DO11.10 TCR transgenic mice that were age- and sex-matched to the adoptive transfer recipients. Single-cell suspensions were prepared by pressing tissue through a nylon strainer (BD Biosciences). The DO11.10 T cells were incubated with anti-CD4 beads and were further purified by passing through an LS+ column (Miltenyi Biotec). In some experiments, purified CD4+ T cells (with purity around 99% as determined by FACS) were resuspended at 5 x 107 cells/ml in PBS containing 0.1% BSA. A final concentration of 10 µM CFSE fluorescent dye (Molecular Probes) was added and incubated for 10 min at 37°C. The labeled cells were washed twice in cell culture medium containing 10% FCS (Invitrogen Life Technologies) before transfer. A total of 5 x 106 OVA-specific CD4+ T cells were injected i.v. into recipient mice. B cells were negatively purified from untreated wild-type (WT) BALB/c mice or from IL-4- or B7-1/B7-2-deficient mice by using the B cell isolation kit (Miltenyi Biotec). The B cells were over 90% pure. In adoptive B cell transfer experiments, 2.5–3 x 107 purified B cells were injected i.v. into recipient mice.

Flow cytometry

Lymph node cells were harvested, and 1 x 106 cells were blocked with Fc Block (BD Pharmingen) and then incubated with anti-CD4-CyChrome (BD Pharmingen), KJ1-26-FITC (Caltag Laboratories), and anti-CD44, anti-CD69, or anti-CD62 ligand (anti-CD62L) PE (BD Pharmingen). After washes, cells were fixed with 1% paraformaldehyde (Fisher Scientific) and analyzed by flow cytometry using an EPICS XL-MCL (Beckman Coulter). For CFSE-labeled cells, anti-CD4-CyChrome or anti-CD4-PerCp and KJ1-26-PE (Caltag Laboratories) were used to distinguish the DO11.10 T cells. Cell cycle progression was monitored by measuring sequential reductions in CFSE fluorescence of KJ1-26+ CD4+ cells, and the proliferation index (PI) was calculated using ModFit software (Verity Software House).

Ex vivo intracellular cytokine measurement

For intracellular cytokine detection, 5 x 106 cells from draining cervical lymph nodes (CLNs) were incubated for 6 h with 10 µg/ml OVA peptide. GolgiStop (BD Pharmingen) was added to the culture for the last 4 h. Lymphocytes were harvested and incubated with Fc Block (2.4G2; BD Pharmingen) plus 10% rat serum (Sigma-Aldrich) for 20 min at room temperature. Cell surface markers were stained by anti-CD4-PerCp (BD Pharmingen) and KJ1-26-PE mAb (Caltag Laboratories). Cells were fixed in 4% paraformaldehyde (Fisher Scientific) and permeabilized in 0.5% saponin (Sigma-Aldrich) before staining with allophycocyanin-conjugated rat anti-mouse IL-4 or anti-IFN-{gamma} mAb (BD Pharmingen). Over 350,000 lymphocyte-gated events were collected, to obtain over 2000 KJ1-26+ CD4+-gated events, and analyzed by flow cytometry using a FACSCalibur (BD Biosciences).

Immunohistological staining

The draining CLNs were harvested from individual mice and frozen in liquid nitrogen, and 8-µm tissue sections were obtained from near the center of the CLN using a HM505E cryostat (Richard-Allan Scientific). Tissue sections were allowed to dry at room temperature for 30 min, fixed in cold acetone for 10 min, and stored at –80°C. Tissue sections of CLN were then dual-stained with KJ1-26-PE (Caltag Laboratories) and Alexa Fluor 647 anti-B220 (Caltag Laboratories), and cover slipped using Fluoromount G (Southern Biotechnology Associates). The surface of the entire CLN section was mapped and scanned using Image-ProPlus software (Media Cybernetics). Mapped regions of the lymph node were then individually photographed at a magnification of x200 using a Leica DFC350 FX camera (Leica Microsystems) mounted on a Leica DM 6000B computerized fluorescence microscope, and tiled using Image-ProPlus software (Media Cybernetics). Each fluorescent channel was photographed separately, and the three sets of x200-magnified images were merged using Image-ProPlus software to create the final picture of the whole CLN section.

Sorting of CD4+ T cells or DO11.10 T cells after immunization

For CD4 T cell sorting, single-cell suspensions of draining lymph nodes were prepared from Nb-inoculated mice and incubated with anti-CD4 microbeads (Miltenyi Biotec). CD4+ T cells were purified by positive selection. The purities were >98%. For cell sorting of OVA-specific DO11.10 T cells, CLN cells were stained with KJ1-26-PE mAb, and then labeled with anti-PE beads (Miltenyi Biotec). Labeled cells were passed through two MS+ columns (Miltenyi Biotec) according to the protocol provided by the manufacturer. The KJ1-26+ population was positively selected and assessed for purity using FACS analysis. The purities were 85–90% in all sorts. Sorted cells were centrifuged, and pellets were dissolved in RNA isolation denaturing solution. RNAs were extracted from sorted cells using an RNA isolation kit (Stratagene Cloning Systems).

LCM analysis

The CLNs, taken from DO11.10 T cell-transferred IL-4–/– mice either untreated or inoculated with Nb, were frozen on dry-ice acetone and stored at –80°C. The 6-µm tissue sections were cut from frozen blocks using an HM505E cryostat (Richard-Allan Scientific), dehydrated, and stained for H&E (Sigma-Aldrich). LCM analysis of stained sections was performed on a PixCell II LCM system (Arcturus Engineering), as previously described (28). Briefly, cells were captured from T zones, B zones, and T:B zones using CapSure LCM Caps (Arcturus Engineering), which contain 20 µl of RNA isolation denaturing solution. The LCM cap was inserted into a 0.5-ml microcentrifuge tube containing 180 µl of RNA isolation denaturing solution, and total RNA was extracted using an RNA isolation kit (Stratagene Cloning Systems).

IL-4 in situ immunofluorescent staining

In situ IL-4 protein staining was as previously described (29). Briefly, CLNs were collected, frozen in cold acetone and stored at –80°C. The 6-µm tissue sections were obtained and after fixation with cold acetone, the tissues were blocked with 0.1% BSA-PBS containing 10% rat sera and 1 µg/ml blocking anti-Fc Ab (2.4G2; BD Pharmingen) for 45 min at room temperature. After incubation with Alexa Fluor 488-11B11, (BD Pharmingen), Alexa Fluor 488-BDV6-24G2, and Alexa Fluor 647 anti-B220, as well as KJ1.26-PE Abs, the tissues were washed with PBS, H2O and covered with a slipcover by using Fluoromount G. Slides were examined and digitally imaged at x1000 magnification using a Leica DM 6000B immunofluorescent microscope with automated stage and Leica DFC 350 FX camera (Leica Microsystems). We photographed each fluorescent channel separately and then merged them together. Alexa Fluor 488-rat IgG1 (BD Pharmingen) was used as an isotype control.

Statistical analysis

Statistical difference at a level of p < 0.05 between groups was assessed using ANOVA and Fisher’s protected least significant difference (LSD) test for pairwise comparisons. The software program SigmaStat (Jandel) was used for all statistical analyses.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
The Th2 immune response is impaired in B cell-deficient mice and cannot be restored by multiple immunizations

Previous studies have suggested that Th2 responses are preferentially inhibited in B cell-deficient mice; however, in most cases these studies involved responses that also had substantial Th1 cell activation (8, 14, 15, 16, 17, 18, 19). It is still unknown whether B cells can also exert direct promoting effects on the development of Th2 cells rather than indirectly influencing Th2 cell differentiation by inhibiting Th1 cells. Our previous studies have shown that we could successfully establish a highly polarized Th2 response with no detectable elevation of IFN-{gamma} in the draining lymph node after immunization with Nb in the ear, even following B7 blockade (21, 30). This model can thus be useful in examining a potential Th1 cell-independent effect of B cells on Th2 cell development.

To examine the role of B cells in Nb-induced Th2 cell development in this readily manipulated nonmucosal environment, 300 Nb larvae were injected intracutaneously in the ear. Seven days after inoculation, mice were killed and ear CLNs were collected for analysis. As shown in Fig. 1A after Nb infection, a pronounced number of IL-4-secreting cells was observed in the CLN from WT mice (p < 0.001), but this number was markedly reduced in Nb-inoculated B cell-deficient mice (p < 0.001). Previous studies have shown that besides Th2 cells, other cell types such as B cells can also produce IL-4 (8). To address whether this defective IL-4 production is due to reduced development of Th2 cells in the absence of B cells, CD4+ T cells were purified from CLN for determination of IL-4 protein and gene expression by ELISPOT and quantitative fluorogenic RT-PCR, respectively. Consistently, the number of IL-4-secreting cells (Fig. 1B) and IL-4 (Fig. 1C) and IL-13 (Fig. 1D) mRNA from sorted CD4+ T cells was all markedly decreased in B cell-deficient mice compared with WT mice following Nb inoculation (p < 0.001 for Fig. 1B). IFN-{gamma} mRNA from CD4+ T cells remained low in both Nb-immunized WT and B cell-deficient mice (Fig. 1E). These results demonstrated that following immunization with Nb, Th2 cell development in the draining lymph nodes is impaired in B cell-deficient mice and the decreased Th2 cell development in B cell-deficient mice does not promote Th1 cell development, indicating that B cells can promote a polarized Th2 response directly.


Figure 1
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  		FIGURE 1. B cell blockade inhibits Nb-induced Th2 response without triggering Th1 response. A–E, WT and B cell-deficient mice were immunized intracutaneously in the ear with 300 third-stage Nb larvae (L3). Seven days later, cells from CLNs were collected. The total number of IL-4-secreting cells per 106 CLN cells (A) and the number of IL-4-secreting CD4 T cells per 106 sorted CD4+ T cells (B) were assessed by ELISPOT. IL-4 mRNA (C), IL-13 mRNA (D) and IFN-{gamma} mRNA (E) from sorted CD4 T cells in the CLN at day 7 after Nb inoculation were assessed by quantitative real-time RT-PCR. F–H, WT and B cell-deficient mice were intracutaneously immunized with Nb in the ears. Four days later, mice were again challenged, and 7 days after primary immunization, cells from CLNs were collected and CD4 T cells were positively sorted by magnetic CD4-beads for quantitative real-time RT-PCR analysis of IL-4 (F), IL-13 (G) and IFN-{gamma} (H) mRNA. Results represent the mean and SE of each treatment group (four to five animals per treatment group), and similar results were obtained in two experiments. **, p < 0.001. Sorted cells were obtained from pooled samples from each treatment group.

 
Previous studies demonstrated that repeated immunization of WT mice with radiation-attenuated cercariae of S. mansoni increased resistance to S. mansoni infection; however, it failed to increase the defective protection in B cell-deficient mice (31). Consistently, the other study also showed that boosting B cell-deficient mice with Ag does not restore the impaired expansion of CD4 cells (11). To determine whether the reduced Th2 cells in B cell-deficient mice could be overcome by multiple immunizations in the context of Nb infection, WT and B cell-deficient mice were given Nb larvae on day 0. Four days later, some of the mice were administered a secondary inoculation with Nb larvae again in the same ear. At day 7, CD4+ T cells were purified from draining lymph nodes for cytokine mRNA expression. As shown in Fig. 1, F and G, multiple immunizations of B cell-deficient mice with Nb failed to increase CD4+ T cell IL-4 (Fig. 1F) and IL-13 (Fig. 1G) mRNA expression to the levels observed in CD4+ T cells from similarly immunized WT mice, although levels were increased compared with CD4+ T cells from B cell-deficient mice given a one-time immunization. IFN-{gamma} mRNA remained low in all treatment groups (Fig. 1H). These results thus indicate that multiple immunizations could not restore the defective Th2 cell development in B cell-deficient mice.

Ag-specific Th2 cell differentiation is inhibited in B cell-deficient mice

Our studies have shown that transferred naive OVA-specific DO11.10 T cells can differentiate into Th2 cells in vivo after the recipient mice are immunized with the combination of OVA peptide and Nb, but not OVA or Nb alone (21). This ability of Nb to act as an adjuvant to induce nonparasite Ag-specific Th2 cell differentiation makes it a highly useful tool to study the role of B cells in the development of highly polarized Ag-specific Th2 cells from a starting naive CD4 T cell population in vivo. A total of 4–5 x 106 sorted naive DO11.10 CD4+ T cells were transferred to WT or B cell-deficient BALB/c recipients (all mice in a given experiment received the same number of transferred cells). Two days after adoptive transfer, recipient mice were intracutaneously immunized in the ear with 300 Nb L3 plus 30 µg OVA peptide, doses previously used to induce a strong Ag-specific Th2 response in the draining CLN (21). At day 7 postimmunization, mice were killed and individual CLNs were collected and cells were prepared for cell sorting and flow cytometric analysis. As shown in Fig. 2A, compared with untreated mice, after immunization with Nb plus OVA peptide, the total number of OVA-specific T cells recovered from WT mice was markedly increased (>15-fold increase). In contrast, the number of KJ1-26+ cells recovered from treated B cell-deficient mice was increased only 2- to 3-fold. These data suggest that B cells are required for Ag-specific Th2 cell expansion where Nb acts as a Th2 adjuvant.


Figure 2
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  		FIGURE 2. B cells are required for Ag-specific Th2 cell development in vivo. WT and B cell-deficient mice were adoptively transferred with naive DO11.10 T cells on day 0. Two days later, these mice were intracutaneously immunized with Nb plus OVA peptide and 7 days after immunization, cells from CLNs were collected. A, The total number of DO11.10 T cells (CD4+ KJ1-26+) per CLN is shown. B, CLN cells were restimulated with OVA peptide in vitro and then stained for cytoplasmic IL-4. C and D, KJ1-26+ cells were positively selected by magnetic bead cell sorting and IL-4 and IFN-{gamma} mRNA from KJ1-26+ cells were determined by quantitative real-time RT-PCR. The experiment was repeated three times with similar results.

 
DO11.10 T cell cytoplasmic IL-4 protein levels were also assessed. On day 7 postimmunization, CLN cells were restimulated with OVA peptide, surface stained with anti-CD4-PerCp and anti-KJ1-26-PE followed by intracellular staining with anti-IL-4-allophycocyanin. Cells were gated on the CD4+ KJ1-26+ population. As shown in Fig. 2B, a pronounced increase in the percentage of IL-4 producing DO11.10 T cells was obtained in WT mice inoculated with Nb and OVA. In contrast, the percentage of IL-4-producing DO11.10 T cells remained very low in immunized B cell-deficient mice, suggesting that B cells are required for the development of Ag-specific Th2 cells from naive cells in vivo. To further confirm this suggestion, at day 7 postinoculation, DO11.10 T cells were positively sorted from draining lymph nodes for IL-4 and IFN-{gamma} gene expression, as described in Materials and Methods. As shown in Fig. 2C, following Nb plus OVA peptide immunization, a dramatically elevated IL-4 mRNA level from sorted DO11.10 T cells was seen in WT mice. However, IL-4 mRNA from sorted DO11.10 T cells in B cell-deficient mice was markedly reduced. IFN-{gamma} mRNA remained at baseline levels in all treatment groups (Fig. 2D). These results thus confirm and extend the findings described in Fig. 1, demonstrating that B cells do play an essential role in the development of Ag-specific polarized Th2 cells in vivo. These findings thus show that the adjuvant properties of Nb that drive Ag-specific Th2 effector cell development in vivo require B cells. Furthermore, this B cell requirement occurs independently of an underlying IFN-{gamma} response.

B cells are required for optimal Ag-specific T cell proliferation but not activation

To more specifically investigate the effects of B cells on the activation and development of Th2 cells in vivo, we examined Ag-specific T cell activation and proliferation. As described, DO11.10 T cells were transferred to recipient mice, inoculated with Nb plus OVA, and at day 7 postinoculation mice were sacrificed and ear CLNs were collected for analysis. Lymph node cell suspensions were stained for CD4 KJ1-26 and CD69, CD44, or CD62L, all of which are used to detect T cell activation, the former two being elevated, whereas CD62L is decreased before cytokine expression. Flow cytometry analysis of gated CD4+ KJ1-26+ cells (Fig. 3A) showed pronounced increases of CD44 (Fig. 3B) and CD69 (Fig. 3C) and decreases in CD62L (Fig. 3D) expression on DO11.10 T cells from Nb-inoculated WT and B cell-deficient recipient mice, consistent with an activated T cell phenotype even in the absence of B cells. These results indicate that compared with WT mice, DO11.10 T cell activation was generally sustained in B cell-deficient mice, suggesting that the defect of Th2 development in B cell-deficient mice was subsequent to early T cell activation.


Figure 3
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  		FIGURE 3. B cell blockade partially inhibits expansion but not activation of transferred Ag-specific T cells following Nb plus OVA inoculation. Following DO11.10 T cell transfer, WT and B cell-deficient recipient mice were immunized with Nb plus OVA as previously described. At day 7 postimmunization, CLN cells were collected from recipient mice, and cell suspensions were stained for CD4-CyChrome, KJ1-26-FITC, and CD44-PE, CD69-PE, or CD62L-PE. The gated CD4+ KJ1.26+ population (A) was analyzed for CD44 (B), CD69 (C), and CD62L (D). E, naive DO11.10 T cells were labeled with CFSE and transferred into B cell-deficient and WT BALB/c mice. Two days later, the mice were immunized with Nb plus OVA. At day 7 postimmunization, the CLN cells were collected. Cell cycle progression of donor DO11.10 T cells was determined according to the dilution of CFSE fluorescence. The experiment was repeated three times with similar results.

 
Previous studies indicated that DNA synthesis and entry into the cell cycle correlates with the development of Th2 cells (32, 33). To examine whether DO11.10 T cell cycle progression was intact in immunized B cell-deficient recipient mice, 5 x 106 sorted CD4 DO11.10 T cells were labeled with CFSE, and transferred to BALB/c WT and B cell-deficient recipients. Two days later, mice were inoculated intracutaneously in the ear with Nb plus OVA. Seven days after inoculation, mice were sacrificed and lymph node cell suspensions from ear CLNs were stained for KJ1-26 and CD4 and cell cycle progression was monitored by measuring sequential reductions in CFSE fluorescence of CD4+ KJ1-26+ cells. As shown in Fig. 3E, the majority (>90%) of transferred DO11.10 T cells remained in the first generation in unimmunized WT and B cell-deficient recipient mice, indicating that nonspecific activation did not occur in either recipient strain. However, at day 7 postimmunization, extensive cell cycling was observed in WT recipient mice inoculated with Nb plus OVA. Almost 50% of the transferred DO11.10 T cells had cycled nine or more generations. In contrast, although cell cycling did occur in B cell-deficient recipients, <20% of the DO11.10 T cells showed this degree of cycling in these mice. These findings combined with the DO11.10 T cell recovery results seen in Fig. 2A suggest that the presence of B cells drives Th2 cell proliferation and expansion in vivo.

Ag-specific Th2 effector cells first develop in the T:B zone

Our findings indicated that during Nb infection, B cells exert direct and potent effects on CD4 T cell proliferation, expansion, and differentiation into effector Th2 cells. These results suggest that, in the context of the lymph node microenvironment, B cells may provide an optimal milieu for the development of IL-4-secreting Th2 effector cells. If this is the case, the development of IL-4-secreting Th2 effector cells may first occur in the regions near B cells, such as B cell zones or T:B zones where B cells can effectively interact with developing T cells. To address this possibility, we needed to use the recently developed technique of LCM to isolate IL-4 mRNA expressing cells from the lymph node microenvironment. We also took advantage of our previous finding that autocrine IL-4 is sufficient to support Th2 cell differentiation in IL-4–/– recipient mice (34). In this model system, naive WT DO11.10 T cells are transferred to IL-4–/– mice. Because the only cells expressing IL-4 in IL-4–/– recipients are the transferred DO11.10 T cells, the expression of IL-4 mRNA in laser dissected tissues is derived only from the transferred WT DO11.10 T cells. After days 2, 3, and 4 postinoculation with Nb plus OVA, CLNs were removed and frozen sections were stained with Alexa Fluor 647-conjugated anti-mouse B220 and PE-conjugated KJ1-26 for B cells and DO11.10 T cells, respectively. As shown in Fig. 4A, specific regions of the lymph node were then laser microdissected, including the T zone, B zone, and T:B zone. RNA was extracted for IL-4 gene expression. As shown in Fig. 4B, IL-4 mRNA was markedly elevated at day 4 after inoculation in all zones of the lymph node, and at day 3 a similar pattern was observed but with decreased elevations. Intriguingly at day 2 after inoculation, increases in IL-4 mRNA were still detected but were localized to the T:B zone. These studies are the first to localize where Ag-specific Th2 cell IL-4 mRNA is initially elevated in the lymph node. They demonstrate that Ag-specific Th2 cells first appear at the interface of the T:B zone as early as 2 days after Nb plus OVA inoculation.


Figure 4
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  		FIGURE 4. IL-4 message expressed by OVA-specific Th2 cells is first elevated in T:B zone on day 2 postimmunization. Naive DO11.10 T cells were adoptively transferred into IL-4–/– mice. Two days later, recipient mice were intracutaneously immunized in the ears with Nb plus OVA (Nb+OVA). At days 2, 3, and 4 postimmunization, CLNs were collected. A, Tissue sections from CLNs were stained for KJ1-26-PE and B220-Alexa Fluor 647 at each time point and whole lymph node images obtained as described in Materials and Methods. B, Tissue samples from T zone, B zone, and T:B zone were collected by LCM and assessed for IL-4 mRNA by quantitative real-time RT-PCR.

 
To examine IL-4 protein in situ, we applied a recently developed IL-4 protein immunofluorescent staining method to localize IL-4 protein production by transferred naive WT DO11.10 T cells in draining lymph nodes from Nb-inoculated IL-4–/– recipient mice (29). CLNs were collected on days 3, 4, 5, and 7 after Nb+OVA inoculation of IL-4–/– recipient mice with transferred DO11.10 T cell after Frozen tissues were sectioned and stained for B cells, DO11.10 T cells, and IL-4 protein as described in Materials and Methods. In situ IL-4 protein is detected by using two Alexa Fluor 488-conjugated anti-IL-4 Abs, which recognize distinct epitopes of IL-4. As shown in Fig. 5, IL-4 protein was first detected in the T:B zone but not the B cell zone or the T cell zone at day 3 after inoculation (Fig. 5, D–F). By day 4, IL-4 was also detectable in the T cell zone (Fig. 5, G–I) and at days 5 and 7, IL-4 protein was distributed in all three regions (Fig. 5, J–O). Similar staining of untreated controls (Fig. 5, A–C) and isotype control Ab on day 4 (Fig. 5, P–R) showed minimal nonspecific staining. These results confirm the gene expression studies, as seen in Fig. 4, and suggest that IL-4-secreting Th2 effector cells first develop in the T:B zone microenvironment of the lymph node. It should be noted that we did not detect IL-4-producing cells at days 3 or 4 after Nb inoculation in the blood or spleen, consistent with other studies suggesting that the initial activation and proliferation of Ag-specific T cells occurs in the draining lymphoid tissues (35, 36).


Figure 5
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  		FIGURE 5. IL-4 protein produced by OVA-specific DO11.10 Th2 cells is first detectable in T:B zone by day 3 postimmunization. Naive DO11.10 T cells were transferred to IL-4–/– recipients, which were intracutaneously immunized with Nb plus OVA 2 days later. At days 3, 4, 5, and 7 postinoculation, CLNs were frozen, sectioned, and stained with PE-conjugated KJ1-26 Ab (DO11.10 T cells, red) and Alexa Fluor 647-conjugated anti-mouse B220 (B cells, blue). IL-4 protein (green) was detected by using two Alexa Fluor 488-conjugated anti-IL-4 mAbs, which recognize distinct epitopes of IL-4, as described in Materials and Methods. The B zone, T:B zone, and T zone microenvironments were then digitally imaged at a magnification of x1000. Representative samples from untreated (A–C), day 3 (D–F), day 4 (G–I), day 5 (J–L), and day 7 (M–O) are shown. Alexa Fluor 488-conjugated rat IgG1 was used as an isotype control for IL-4 staining, with day 4 shown (P–R). To facilitate comparison, individual panels (for each given day) were normalized for brightness and contrast. S, Enlarged image of day 3 T:B zone (E).

 
Reconstitution with naive WT B cells completely restores Ag-specific Th2 response in B cell-deficient mice

Our results indicate that in the context of Nb infection, the absence of B cells abrogates Ag-specific polarized Th2 cell development in vivo without promoting an IFN-{gamma} response, and that IL-4-secreting Ag-specific Th2 effector cells first develop in the T:B zone. These findings suggest that restoration of B cell-deficient mice with WT B cells should at least partially restore Th2 cell cytokine expression in vivo. Previous studies also suggested that B cell-deficient mice may have other deficiencies besides an absence of B cells. In particular, DC may not develop normally in the absence of B cells particularly in the spleen (37). Indeed, reconstitution with LPS-activated B cells has been shown to rescue the OVA-specific Th2 response in mice immunized with pertussis toxin and alum-precipitated OVA (8). However, LPS is a very potent B cell stimulant and its effects may have little relevance to B cell activation in the context of an immune response not involving endotoxin. To avoid in vitro B cell activation and its associated artifacts, B cells were purified from WT untreated mice as described in Materials and Methods, and 2.5–3 x 107 purified naive B cells were i.v. transferred together with CFSE-labeled naive DO11.10 T cells to recipient B cell-deficient mice. Two days later, B cell-deficient mice and WT controls were inoculated intracutaneously in ears with Nb plus OVA peptide. Seven days later, draining CLNs were collected.

Cytoplasmic IL-4 staining was performed following a 6-h restimulation with OVA peptide in vitro as described in Materials and Methods. The cells were gated on the CD4+ KJ1-26+ subpopulation. As shown in Fig. 6A, compared with untreated WT recipient mice, DO11.10 T cells transferred into WT mice subsequently inoculated with Nb plus OVA showed pronounced increases in IL-4 protein expression. In contrast, IL-4 production by DO11.10 T cells transferred into B cell-deficient mice remained at baseline levels postimmunization. Reconstitution of B cell-deficient recipient mice with WT untreated B cells completely restored IL-4 production by DO11.10 T cells to the level in WT recipient mice in the context of Nb infection.


Figure 6
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  		FIGURE 6. Reconstitution of B cell-deficient mice with untreated WT B cells restores Ag-specific T cell expansion and IL-4 production after Nb plus OVA inoculation. CFSE-labeled naive DO11.10 T cells were adoptively transferred to WT and B cell-deficient mice. Some B cell-deficient mice also received WT naive B cells. Two days later, recipient mice were inoculated with Nb plus OVA. A, At day 7 postimmunization, CLN cell suspensions were surface stained with anti-CD4-PerCp and KJ1-26-PE, and stained intracellularly with anti-IL-4-allophycocyanin. B, Cell division of transferred DO11.10 T cells was analyzed by flow cytometric analysis of CFSE fluorescence. The experiment was repeated twice with similar results.

 
In addition, cell division was analyzed as CFSE fluorescence gating on CD4+ KJ1-26+ donor cells. In Fig. 6B, after immunization, donor DO11.10 T cells underwent pronounced cell cycling with over 70% of DO11.10 T cells present in the last generation (PI = 26.03). In contrast, markedly decreased cell division was seen in DO11.10 T cells from B cell-deficient mice (PI = 13.39) because only 34% of DO11.10 T cells were in the last generation. Restoration of B cell-deficient mice with WT naive B cells significantly increased cell cycle progression of DO11.10 T cells to levels comparable to that in WT mice (PI = 23.92) with 69% of the cells in the last generation. Taken together, these results demonstrate that transfer of untreated WT B cells restored cell division and IL-4 production of donor DO11.10 T cells in B cell-deficient mice, demonstrating that B cells are required for the development of the Ag-specific Th2 response in the context of Nb infection.

B7, but not IL-4, expression by B cells is required for optimal Th2 cell development in vivo

Our studies indicated that B cells were required for the development of Ag–specific Th2 cells in vivo. We next examined possible mechanisms by which B cells affect Th2 cell development in vivo. Previous studies have suggested that IL-4 produced by B cells may be important in the development of the Th2-type response (7). We first examined whether B cells could express IL-4 after Nb inoculation. BALB/c mice were inoculated in the ear with Nb larvae and 1 day later, cell suspension from draining CLNs were collected. B cells were isolated, and 6-fold elevations in IL-4 mRNA were observed compared with IL-4 mRNA levels in B cells from CLNs of untreated mice (data not shown). These findings suggested that B cell-derived IL-4 might contribute to Th2 cell development in vivo. To further address whether B cells promote Th2 cell development via producing IL-4, we purified B cells from either untreated WT or IL-4–/– mice, and transferred these cells along with naive DO11.10 T cells into B cell-deficient mice and monitored OVA-specific Th2 responses in draining CLNs after inoculation with Nb plus OVA. As shown in Fig. 7A, reconstitution of B cell-deficient mice with either IL-4–/– B cells or WT B cells rescued the development of IL-4-producing DO11.10 T cells, suggesting that IL-4 produced by B cells was not required for B cell-mediated Th2 cell development.


Figure 7
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  		FIGURE 7. B7, but not IL-4, expression by B cells is important in mediating Th2 cell development in vivo. A and B, Naive DO11.10 T cells were adoptively transferred to WT and B cell-deficient mice. Some B cell-deficient mice also received IL-4–/– B cells (A) or B7-1/B7-2-deficient B cells (B), which were compared with B cell-deficient mice receiving WT B cells. C, Naive DO11.10 T cells were transferred to WT and B7-1/B7-2-deficient mice. Some B7-1/B7-2-deficient mice also received WT naive B cells. In all experiments, at day 2 after cell transfer mice (five mice per treatment group) were inoculated with Nb and OVA and at day 7 postimmunization, CLN cell suspensions were surfaced stained with anti-CD4-PerCp and KJ1-26-PE, and stained intracellularly with anti-IL-4-allophycocyanin. These experiments were repeated twice with similar results.

 
Previous studies showed that activated B cells express B7-1/B7-2 costimulatory molecules, which can provide second signals during naive T cell differentiation (38) and which may also be important in sustaining activated T cell responses (12, 38). This response together with our findings that both B7-1 and B7-2 were elevated on B cells during Nb infection suggested that B7 costimulation may play a role in mediating the development of the Th2-type response in vivo. To address whether B7 expressed on B cells promotes Th2 development in vivo, B7-1/B7-2-deficient or WT B cells, along with naive DO11.10 T cells, were transferred into B cell-deficient mice. The mice were immunized with Nb plus OVA peptide and at day 7 postimmunization, the development of IL-4–producing DO11.10 T cells was monitored by intracellular staining. As shown in Fig. 7B, reconstitution of B cell-deficient mice with B7-1/B7-2-deficient B cells failed to restore the DO11.10 T cell IL-4 response to Nb plus OVA, compared with transferred WT B cells. However, the frequency of IL-4-producing DO11.10 T cells in the B cell-deficient mice receiving B7-1/B7-2-deficient B cells was still higher than that in B cell-deficient mice without any B cell transfer. Thus our data suggest that B7 costimulation is at least partially responsible for B cell-mediated Th2 cell development.

To examine whether B7 costimulatory signals from B cells may be sufficient to at least partially drive the Th2-type response, B7-1/B7-2-deficient and WT mice were reconstituted with WT B cells along with DO11.10 T cells and 2 days later, immunized with Nb plus OVA. At day 7 postimmunization, IL-4-producing DO11.10 T cells in draining CLNs were examined by intracellular staining. As shown in Fig. 7C, compared with that in WT mice, after Nb plus OVA immunization, the development of IL-4-producing DO11.10 T cells was completely inhibited in B7-1/B7-2-deficient mice, consistent with a previously published study (30). However, after WT B cell transfer, the development of IL-4-producing DO11.10 T cells was partially rescued in B7-1/B7-2-deficient mice. These findings indicate that B7 expression by B cells alone is sufficient to promote the development of IL-4-producing T cells in vivo.


   Discussion
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
In this study, we examined the role of B cells in the development of IL-4-secreting CD4 Th effector cells during a polarized Th2 response. Our studies show that Th2 cell differentiation is largely dependent on B cells even in the absence of the development of an alternative IFN-{gamma} response and that the T:B zone is the lymph node microenvironment where IL-4 expressing Ag-specific Th2 effector cells first develop. Our studies further suggest that B7 expression by B cells plays an important role in Th2 cell development in vivo.

Previous studies in mixed Th1/Th2 responses have shown that in B cell-deficient mice the Th2 response shifts toward a Th1 response. In these studies, the underlying Th1 response apparently becomes dominant during B cell blockade and inhibits Th2 cell differentiation (14, 15, 16, 17, 18). In experiments where pertussis toxin is used as an adjuvant, B cell deficiency deviates the Th2 response toward a Th1 cytokine pattern and the ability of B cells to promote Th2 cell differentiation is OX40 ligand-dependent (12). In contrast, Nb promotes a polarized Th2 response in the absence of an underlying Th1 response (21). Our results clearly show that B cell blockade inhibits the Th2 response to Nb in the absence of corresponding increases in IFN-{gamma}, indicating that B cells can directly promote Th2 responses through IFN-{gamma}-independent mechanisms. It should be noted that we expect that other resistant strains, e.g., the BL/6 mouse, would also show a similar requirement for B cells in Th2 cell differentiation, as the highly polarized Th2-type response similarly occurs in these strains following Nb inoculation (39).

Our findings indicated that in the absence of B cells, Ag-specific T cells still differentiate to cleave CD62L and express the activation markers CD69 and CD44, all of which are associated with early stages of T cell activation following TCR signaling (40, 41, 42). These results are consistent with a previous study indicating that T cell priming can occur in B cell-deficient mice (43). We also determined that cell division of donor DO11.10 T cells was much reduced in B cell-deficient mice compared with WT mice, although a significant number of cells still progressed through at least 10 cell cycles. These results are consistent with earlier findings that T cells are initially activated in the T zone and then migrate to the follicle border to form tight physical contact with B cells for maximal proliferation (3). This mechanism suggests that T cell cognate interactions with B cells are not necessary for T cell activation, initial proliferation, and migration to the T:B zone but are required for optimal T cell proliferation. In contrast, the development of IL-4-producing T cells in B cell-deficient mice was largely abrogated. Previous studies have suggested that DNA synthesis and cell proliferation correlate with CD4 T effector cell differentiation (32, 44). We have previously shown that when autocrine IL-4 was inhibited, cell cycling and IL-4 production were separable, suggesting that cell cycling alone is not sufficient to support Th2 cell differentiation (33, 45). In this study, although significant DO11.10 T cell division occurred in B cell-deficient mice, the absence of IL-4 production by these cycling cells indicates that Th2 cell differentiation requires additional B cell-dependent signals besides those required for cell cycle progression.

Our finding that B cells are required for the development of Th2 cells in vivo was further confirmed by B cell reconstitution experiments. The transfer of WT untreated B cells to B cell-deficient mice with donor DO11.10 T cells completely restored cell cycle progression and IL-4 production of DO11.10 T cells after immunization with Nb and OVA. Previous studies have reconstituted B cell-deficient mice with B cells activated in vitro with endotoxin. This method is a particular problem when studying the Th2 response because LPS can stimulate Th1 or Th2 cell differentiation, depending on the particular conditions (46, 47, 48, 49), thereby potentially influencing the development of the Th2 response. Our studies thus substantially extend these previous findings by showing that transfer of untreated naive B cells is sufficient to fully restore Ag-specific T cell cell cycle progression and IL-4 production in B cell-deficient mice in the context of Nb infection, an effect perhaps partly due to the particularly potent adjuvant capability of this nematode parasite to drive the Th2 response.

We also examined the mechanism by which B cells promote the development of Th2 cells. IL-4 is a central cytokine for Th2 differentiation and previous studies have shown that B cells can produce IL-4 (7). Although we also detected increased B cell IL-4 gene expression following Nb inoculation, our findings indicate that B cell-derived IL-4 does not play an essential role in Th2 cell development, as reconstitution of B cell-deficient mice with IL-4–/– or WT B cells similarly restored the IL-4 response. These results are consistent with our previous studies indicating that in the absence of host IL-4 production, autocrine IL-4 is sufficient to support Ag-specific Th2 cell development (34). In contrast, our findings that transferred B cells required B7 expression for optimal restoration of Th2 responses in B cell-deficient mice indicate B cells play an important role in providing costimulatory signals through B7 in vivo.

We also observed that Ag-specific T cells first expressed IL-4 mRNA and protein, within 2–3 days after Nb inoculation, in the T:B zone microenvironment of the lymph node. Previous studies from our laboratory showed that autocrine IL-4 is sufficient for the development of Ag-specific Th2 cells from naive T cells in the context of Nb infection because IL-4–/– recipient mice supported transferred WT DO11.10 T cell differentiation following immunization with Nb plus OVA (34). This model system has the advantage in that donor DO11.10 T cells are the only cells capable of expressing IL-4, and thus when used in combination with LCM or in situ IL-4 protein staining, can facilitate localization of the lymph node microenvironment in which Ag-specific T cells first differentiate into Th2 cells in vivo. Previous studies have implicated the role of the T:B zone as important in Th2 cell differentiation. In one study, activated Th2 cell clones formed rings at the periphery of the T zones near B cell follicles (50). In another study, Th2 cytokines were expressed throughout the lymph nodes but especially in the B cell zones, although the cell source was not identified (25). Our studies localize for the first time the site of initial Th2 cell differentiation from naive precursors, as detected by increased IL-4 mRNA and protein at days 2 and 3, respectively, in the lymph node. The observation that IL-4 protein was sometimes not immediately surrounding donor T cells is consistent with previous observations that Ag-specific T cells are frequently moving, interacting with neighboring cells (51), and presumably secreting IL-4 at different sites. These results confirm earlier findings indicating that T cell proliferation requires T:B cell interaction in the follicular border (3) starting on day 2 and peaking on days 3 and 4, and significantly extend these findings by showing that Ag-specific Th2 cells first develop in the same region at the same time. Thus, our studies support the concept that there may be two stages in the process of the development of Th2 cells. The first stage is B cell-independent: after immunization with Ag, naive T cells interact with Ag-presenting DCs where they are activated, undergo minimal proliferation, and migrate to the T:B areas. In contrast, the second stage is B cell-dependent: after T cells migrate into the T:B zone, they undergo optimal proliferation and expansion, and in this microenvironment gain the ability to differentiate into Th2 cells.

Although we have shown an important role for B7 costimulatory signals at this second stage, future studies should further examine characteristics of this microenvironment and especially the phenotype and the functions of B cells in this region which might favor this Th effector cell differentiation pathway. It should be noted that previous studies have shown that OX40 interactions (52) by B cells are not required for the Nb adjuvant-driven development of Ag-specific Th2 cells (34, 53). However, recently B cells have been shown to have a number of functions besides their ability to produce cytokines or express costimulatory molecules. For example, B cells play a role in lymph node lymphangiogenesis and affect DC mobilization into a draining lymph node (54), and enhance DC maturation by promoting expression of CD86 (55). In addition, absence of B cells may alter T zone architecture in lymphoid tissue, particularly the spleen (37). Future studies should address these alternative mechanisms through which B cells may promote Th2 cell differentiation in vivo in the absence of an underlying Th1 response.

In summary, our experiments suggest that the T:B zone is the initial site of Th2 cell differentiation in the lymph node during the primary immune response to a potent Th2 cell inducing Ag and that B cells, at least partly through their expression of B7, play a crucial role in this microenvironment, sustaining the expansion and particularly the differentiation of Ag-specific Th2 cells.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest. The opinions or assertions contained within are the private views of the authors and should not be construed as official or necessarily reflecting the views of the University of Medicine and Dentistry of New Jersey or the Department of Agriculture.


   Footnotes
 
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.

1 This work was supported by Grant AI31678 from the National Institutes of Health. Back

2 Address correspondence and reprint requests to Dr. William C. Gause, Department of Medicine, New Jersey Medical School, University of Medicine and Dentistry of New Jersey, 185 South Orange Avenue, Newark, NJ 07103. E-mail address: gausewc{at}umdnj.edu Back

3 Abbreviations used in this paper: DC, dendritic cell; LCM, laser capture microdissection; CLN, cervical lymph node; PI, proliferation index; WT, wild type; Nb, Nippostrongylus brasiliensis; CD62L, CD62 ligand. Back

Received for publication June 28, 2007. Accepted for publication July 16, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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