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Infections That Induce Autoimmune Diabetes in BBDR Rats Modulate CD4⁺CD25⁺ T Cell Populations^1,2

Danny Zipris, Jan-Luuk Hillebrands^¶, Raymond M. Welsh^*,, Jan Rozing^¶, Jenny X. Xie, John P. Mordes, Dale L. Greiner^*, and Aldo A. Rossini^3,*,,

Programs in ^* Immunology and Virology and Molecular Medicine and Departments of Medicine and Pathology, University of Massachusetts Medical School, Worcester, MA 01655; and ^¶ Division of Cell Biology, Section of Immunology, Faculty of Medical Sciences, University of Groningen, Groningen, The Netherlands

	Abstract

Top Abstract Introduction Materials and Methods Results H-1 infection also induces... Infection of BBDR rats... Discussion References

 
Viruses are believed to contribute to the pathogenesis of autoimmune type 1A diabetes in humans. This pathogenic process can be modeled in the BBDR rat, which develops pancreatic insulitis and type 1A-like diabetes after infection with Kilham’s rat virus (RV). The mechanism is unknown, but does not involve infection of the pancreatic islets. We first documented that RV infection of BBDR rats induces diabetes, whereas infection with its close homologue H-1 does not. Both viruses induced similar humoral and cellular immune responses in the host, but only RV also caused a decrease in splenic CD4⁺CD25⁺ T cells in both BBDR rats and normal WF rats. Surprisingly, RV infection increased CD4⁺CD25⁺ T cells in pancreatic lymph nodes of BBDR but not WF rats. This increase appeared to be due to the accumulation of nonproliferating CD4⁺CD25⁺ T cells. The results imply that the reduction in splenic CD4⁺CD25⁺ cells observed in RV-infected animals is virus specific, whereas the increase in pancreatic lymph node CD4⁺CD25⁺ cells is both virus and rat strain specific. The data suggest that RV but not H-1 infection alters T cell regulation in BBDR rats and permits the expression of autoimmune diabetes. More generally, the results suggest a mechanism that could link an underlying genetic predisposition to environmental perturbation and transform a "regulated predisposition" into autoimmune diabetes, namely, failure to maintain regulatory CD4⁺CD25⁺ T cell function.

	Introduction

Top Abstract Introduction Materials and Methods Results H-1 infection also induces... Infection of BBDR rats... Discussion References

 
It is generally agreed that environmental perturbants, specifically viral infection, play a role in the expression of type 1 diabetes mellitus in genetically susceptible individuals (1). Unfortunately, although data from epidemiological studies and the analysis of discordant monozygotic twins strongly support the concept, little is known with certainty regarding underlying mechanisms. Human studies of the interaction of environment and genetic predisposition are exceedingly difficult. The most widely used animal model of autoimmune diabetes, the nonobese diabetic (NOD)⁴ mouse, models the process poorly because the overwhelming majority of perturbants reduce the frequency of diabetes and often prevent it entirely (2). Among murine viruses, encephalomyocarditis virus (3), mouse hepatitis virus (4), lymphocytic choriomeningitis virus (5), and others reduce the frequency of disease (6). Only pancreatotropic Coxsackie virus has been documented to accelerate disease in these mice (7), but this murine infection is associated with a significant degree of inflammation and exocrine pancreatic necrosis (8) not characteristic of the prediabetic state in humans who develop type 1A diabetes (9, 10).

Recent attempts to address the problem have relied on transgenic mouse systems. For example, using a tetracycline-inducible promoter for TNF- and CD80 expression, Green et al. (11) were able to synchronize environmental perturbation with the expression of diabetes. Using this system, they defined periods of immunoregulation and anti-islet aggression and suggested that failure to maintain regulatory CD4⁺CD25⁺ T cell function plays a critical role in disease progression and expression.

To study the problem of environmental factors in diabetes induction and the possible role of regulatory CD4⁺CD25⁺ T cells in a less artificial system, we have used the BBDR rat. Spontaneous type 1 diabetes does not occur in viral Ab-free BBDR rats (12), but the disorder can be induced in about one-third of these animals by infection with the UMass strain of rat virus (RV-UMass, formerly designated Kilham’s rat virus) (13, 14). Induction of diabetes is not associated with infection of the islets of Langerhans (15) and the combination of RV-UMass infection plus injection of the immune activator poly(I:C) leads to insulitis and diabetes in nearly all treated animals (16).

RV is a parvovirus, a ssDNA virus that infects several animal species including rodents (17) and humans (18). RV, H-1, and the recently described rat parvovirus-1 (RPV-1, formerly designated orphan parvovirus) are the three autonomous parvoviruses that infect rats (17). RV encodes three overlapping structural proteins, VP1, VP2, and VP3, and two overlapping nonstructural proteins, NS1 and NS2 (17). RV is most homologous with H-1; RV and H-1 VP proteins are 80% homologous and their NS proteins are 100% homologous.

The mechanism by which RV-UMass infection leads to the expression of autoimmune diabetes in the BBDR rat is not yet fully understood. The virus infects lymphoid organs and endothelial cells but not insulin-producing cells (15). It was initially hypothesized that islet-specific T cells in the BBDR rat were cross-reactive with a RV peptide and that diabetes was induced in infected rats by classical molecular mimicry. This hypothesis, however, could not be confirmed in studies of BBDR rats injected with viral vectors expressing RV proteins (19). Diabetes failed to occur despite the generation of robust cellular and humoral immune responses. The authors of that study documented the expansion of CD8⁺ T cells in infected animals, but did not demonstrate virus-specific CD8⁺ T cell responses. They suggested that a virus-induced imbalance between Th1-like CD4⁺CD45RC⁺ and Th-2-like CD4⁺CD45RC^- T cells led to disease expression in these genetically susceptible animals.

To test further the hypothesis that failure to maintain regulatory T cell function may be a critical determinant of disease expression in type 1A diabetes, we conducted additional analyses of the BBDR rat, exploiting the fact that it provides a clearly defined period of normal "regulation" that is transformed into "aggression" in a characteristic time frame by RV infection. Specifically, we analyzed the humoral and cellular immune responses of BBDR and normal WF rats to infection with either RV or its homologue H-1, which does not cause diabetes. We observed cross-reactivity of the cellular but not humoral immune responses to these infectious agents, but a differential effect on the proportion of CD4⁺CD25⁺ T cells in lymphoid tissues. Given the immunoregulatory role of these cells in the pathogenesis of autoimmunity in general (20, 21, 22) and autoimmune diabetes in particular (11, 23), our results suggest that one mechanism of virus-induced autoimmune diabetes may involve failure to maintain CD4⁺CD25⁺ T regulatory cell function in a genetically susceptible background.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results H-1 infection also induces... Infection of BBDR rats... Discussion References

 
Animals, viruses, and cell lines

Viral Ab-free BBDR/Wor rats of either sex were obtained from BRM (Worcester, MA) and housed in viral Ab-free quarters. Animals from this vendor are certified to be free of Sendai virus, pneumonia virus of mice, sialodacryoadenitis virus, rat corona virus, Kilham’s rat virus, H-1, GD7, Reo-3, Mycoplasma pulmonis, lymphocytic choriomeningitis virus, mouse adenovirus, Hantaan virus, and Encephalitozoon cuniculi. Viral Ab-free WF rats of either sex were purchased from Harlan Sprague Dawley (Indianapolis, IN).

RV-UMass and NRK cells were obtained from stocks maintained in our laboratories. RV was propagated in NRK cells grown in DMEM. Toolan’s H-1 virus was obtained from American Type Culture Collection (ATCC, Manassas, VA) and was propagated in Chang liver cells, also obtained from ATCC.

Treatment protocols

Rats were infected with 1 x 10⁷ PFU RV or H-1 i.p. in a volume of 1 ml when 22–28 days old as described previously (13, 16). Poly(I:C) was purchased from Sigma-Aldrich (St. Louis, MO), dissolved in Dulbecco’s PBS (1 mg/ml), sterile filtered, and stored at -20°C until used. The concentration of contaminating endotoxin was determined commercially (Charles River Endosafe, Charleston, SC) and was uniformly <10 U/mg (24). Rats were injected i.p. with poly(I:C) (5 µg/g body weight on 3 consecutive days) when 21–25 days of age. Poly(I:C)-treated rats were either given no further treatment or infected with RV or H-1 on the day after the last injection. In experiments designed to measure the frequency of diabetes induction, all treated rats were screened for glycosuria twice weekly until diabetes onset or until day 40 after infection or the last injection of poly(I:C). Diabetes was diagnosed on the basis of a plasma glucose concentration >250 mg/dl (11.1 mM). In some experiments, pancreas specimens were obtained from animals that were not diabetic at the conclusion of the experiment, fixed in 10% buffered Formalin, stained with H&E, and examined by a pathologist not aware of the treatment status of the specimens.

Plaque assay

NRK cells (10⁵ cells/well) were incubated overnight in six-well plates at 37°C in 4 ml of medium in a humidified atmosphere of 95% air and 5% CO₂. The next day, fresh medium was added, and a series of 10-fold dilutions of a stock virus culture or serum from experimental animals was added to the cells. Two hours later, cells were overlaid with 0.5% agarose gel and cultures were incubated for 5 days. Agarose plugs were then removed, and NRK cells were stained with crystal violet for 5 min and washed. Plaques were counted manually by light microscopy.

Cell isolation procedures and flow microfluorometry

Spleen cells and heparinized blood were obtained from control and infected rats at various time points. Single-cell suspensions were prepared and erythrocytes were lysed with NH₄Cl as described elsewhere (25). Cells were washed twice with PBS and resuspended in high-glucose DMEM containing 10% heat-inactivated FBS, 1 mM sodium pyruvate, 100 U/ml penicillin, 0.1 mg/ml streptomycin, and 50 mM 2-ME (Life Technologies, Grand Island, NY).

Abs to the TCR (clone R73, mouse IgG1), IL-2R -chain (CD25, clone OX-39, mouse IgG1), CD4 (clone OX-35, mouse IgG1), CD8 chain (clone OX-8, mouse IgG1), anti-CD62L (anti-L-selectin, clone HRL1, Armenian hamster IgG), PE-conjugated anti-NK (NKR-P1A, clone 10/78, mouse IgG1), appropriate isotype control Abs, and CyChrome-conjugated streptavidin were purchased from BD PharMingen (San Diego, CA). Biotinylated anti-ART2.1 (rat IgG2b) was produced from the DS4.23 hybridoma maintained in our laboratory. Abs were used either directly conjugated with FITC or PE or were used as biotin conjugates followed by CyChrome-streptavidin.

Two- and three-color flow microfluorometry analyses of spleen and blood cells were performed as described previously (26). Briefly, 0.5 x 10⁶ viable cells were reacted with optimal concentration of Abs for 30 min at 4°C. Cells were stained with directly conjugated Abs, washed, and fixed with 1% paraformaldehyde. Cells labeled with biotinylated Abs were washed, incubated with streptavidin-CyChrome for 30 min at 4°C, washed, and then fixed in a final concentration of 1% paraformaldehyde in PBS. Cells were analyzed using a FACScan instrument (BD Biosciences, Mountain View, CA). A minimum of 30,000 events was acquired for each analysis. The lymphocyte fraction was gated on the basis of forward and side scatter.

Detection of anti-RV and anti-H-1 Abs

Anti-RV and anti-H-1 Abs were detected using 96-well ELISA plates coated with intact RV or H-1 virus (Charles River Breeding Laboratories, Wilmington, MA). Wells were blocked with 5% Blotto (Bio-Rad, Hercules, CA) in PBS containing 0.05% Tween 20 (Sigma-Aldrich) for 1 h at 37°C and then washed twice with PBS-Tween 20. Serial dilutions of sera from control or RV- and H-1-infected rats were then added in triplicate to wells for 1 h at room temperature, followed by three washes with PBS-Tween 20. Biotinylated goat anti-rat IgG (H and L chain specific; Jackson ImmunoResearch, West Grove, PA) was then added to the wells. After incubation for 1 h at room temperature, wells were washed three times and then incubated with streptavidin-HRP (Jackson ImmunoResearch) for 30 min. Wells were then washed five times and HRP was detected with o-phenylenediamine dihydrochloride (Sigma-Aldrich). Color development was terminated with 5 N HCl and plates were read at 492 nm using an Emax ELISA plate reader (Molecular Devices, Sunnyvale, CA).

Assay for virus-specific CD8⁺IFN-⁺ T cells

We developed a new system for the detection of virus-specific CD8⁺ T cell responses in the rat based on a published murine assay (27) that uses intracellular IFN- expression as a surrogate marker of CTLs. Y3-Ag 1.2.3 cells (Y3 cells; ATCC) were infected with 0.5 x 10⁶ PFU RV or H-1 in 4 ml of complete DMEM for 18 h and used as virus-infected APCs. Y3 cells express the same RT1^u MHC haplotype as do both BBDR and WF rats. One million BBDR or WF spleen cells or 100 µl of peripheral blood leukocytes were prepared as described above and incubated in the presence of 5 x 10⁵ virus-infected Y3 APCs in 200 µl of complete DMEM supplemented with 1 µl/ml brefeldin A (Golgiplug; BD PharMingen) and 20 ng/ml human rIL-2 (BD PharMingen) in 96-well plates (BD Falcon, Franklin Lakes, NJ). Cells were cultured at 37°C in an atmosphere of 95% air and 5% CO₂ for 5 h and then stained with PE-conjugated anti-rat CD8 mAb. Cells were then fixed, permeabilized, and reacted with FITC-conjugated anti-rat IFN- mAb (Clone DB-1; BD PharMingen) or its isotype control (mouse IgG1; BD PharMingen), using the Cytofix/Cytoperm kit (BD PharMingen). Cells were analyzed by flow microfluorometry as described above.

5-Bromo-2'-deoxyuridine (BrdU) labeling and detection by flow microfluorometry

Groups of 25- to 32-day-old BBDR rats were untreated or injected i.p. with 10⁷ PFU of RV. Spleen and pancreatic lymph nodes were recovered from the untreated rats and RV-infected BBDR rats 3 and 4 days after infection. Both groups were injected i.p. with BrdU (100 mg/kg body weight; Sigma-Aldrich) 24 and 12 h before tissue recovery. BrdU was dissolved in PBS and 0.1 N NaOH, pH 8 for injection. Single-cell suspensions were prepared and stained with anti-CD4 and anti-CD25 mAbs. Incorporated BrdU was detected as described elsewhere (28). Briefly, after surface labeling, cells were fixed in ice-cold 95% ethanol for 30 min, washed with PBS, and permeabilized in PBS containing 1% paraformaldehyde and 0.01% Tween 20 for 30 min at room temperature. Cells were then treated with 50 U of DNase I (Sigma-Aldrich) and subsequently stained with FITC-conjugated anti-BrdU mAb (clone 3D4; BD PharMingen) (28). BrdU incorporation in CD4⁺CD25⁺ cells was measured by flow microfluorometry.

Purification of CD4⁺CD25⁺ and CD4⁺CD25^- T cell subsets

CD4⁺CD25^- responder T cells and CD4⁺CD25⁺ regulatory T cells were obtained from pooled single-cell suspensions of cervical and mesenteric lymph nodes of 4- to 5-wk-old BBDR rats. T cells were purified using nylon wool fiber columns (Polysciences, Warrington, PA). T cell purity after nylon wool separation was measured by flow cytometric analysis of TCR expression and was consistently >95%. Enriched T cell populations were reacted with anti-CD8 (OX8, mouse IgG1) for 50 min on ice. After two washes, the anti-CD8 mAb-labeled T cells were incubated for 20 min with rat anti-mouse IgG1 Microbeads (Miltenyi Biotec, Auburn, CA) at 4°C. After two washes, bound cells were separated using LD depletion columns placed in a strong magnetic field (MidiMACS; Miltenyi Biotec). The purity of the unbound (CD4⁺ T cells) was determined by flow cytometry analysis following staining with FITC-conjugated anti-CD4 mAb (clone OX35) and was consistently >98%. CD4⁺CD25⁺ T cells were then enriched from the purified CD4⁺ T cells by incubation with PE-conjugated anti-CD25 mAb (clone OX39) for 30 min on ice. After two washes, cells were incubated for 20 min with anti-PE Microbeads (Miltenyi Biotec) at 4°C. After washing, bound cells were separated using MS columns placed in a strong magnetic field (MiniMACS; Miltenyi Biotec). The purity of the bound and unbound populations of CD4⁺ T cells was determined by flow cytometry after staining with PE-conjugated anti-CD25 mAb. The percentage of CD4⁺CD25⁺ T cells in the bound fraction was 80–90% and <3% in the unbound fraction.

APC preparation

APCs were isolated from BBDR splenocytes by plastic adherence. Briefly, single-cell suspensions were prepared and adjusted to a concentration of 5 x 10⁶/ml after lysis of RBCs in NH₄Cl buffer. Cells were plated in 15 x 100-mm Petri dishes (5 x 10⁷/dish) and incubated for 1 h at 37°C in an atmosphere of 5% CO₂ and 95% air. Unbound cells were removed by washing twice with warm (37°C) PBS. Adherent cells were detached using a cell scraper, adjusted to 3 x 10⁷/ml, and exposed to 30 Gy gamma-irradiation. Irradiated APCs were recounted and adjusted to a concentration of 1 x 10⁶/ml in culture medium (see below).

Proliferation assays

Cultures were performed in RPMI 1640 supplemented with 10% heat-inactivated FBS, penicillin (100 U/ml), streptomycin (100 µg/ml), 2 mM glutamine, 0.1 M nonessential amino acids, 1 mM sodium pyruvate, and 55 µM 2-ME (all from Gibco, Grand Island, NY). To analyze CD4⁺CD25⁺ T cell-mediated suppression, 5 x 10⁴ CD4⁺CD25^- (responder) T cells were cultured in 96-well plates with 5 x 10⁴ irradiated APCs, Con A (2.5 µg/ml) and 1 x 10⁵ CD4⁺CD25⁺ (suppressor) T cells for 72 h at 37°C in the presence of 5% CO₂ and 95% air. To correct for absolute cell numbers in coculture experiments in which CD4⁺CD25⁺ cells were added to a constant number of CD4⁺CD25^- cells, CD4⁺CD25^- cells were also cocultured using 1 x 10⁵ 30 Gy-irradiated nylon wool-purified T cells in place of CD4⁺CD25⁺ T cells. For each stimulation assay, CD4⁺CD25^- cells alone and CD4⁺CD25^- plus CD4⁺CD25⁺ cells were assayed at the same time in quadruplicate. Cultures were pulsed with 1 µCi [³H]thymidine/well for the last 14 h of culture. Proliferative responses in each independent experiment were determined by averaging the cpm of each quadruplicate assay. The relative proliferation of CD4⁺CD25^- cells in each of the two cultures was calculated as a ratio, and the proliferation of CD4⁺CD25^- cells alone was arbitrarily set to 100%. The assay was performed five times.

Statistics

Parametric data are presented as the mean ± 1 SD. Comparisons of two means were made with the unpaired t test with the Bonferroni adjustment as required for multiple comparisons (29). Comparison of proliferative responses in suppressor cell assays was made using the paired t test. Differences among groups of rats with respect to diabetes-free survival were analyzed by the method of Kaplan and Meier using the log rank statistic (30).

	Results

Top Abstract Introduction Materials and Methods Results H-1 infection also induces... Infection of BBDR rats... Discussion References

 
RV but not H-1 induces autoimmune diabetes in BBDR rats

We first established that the combination of RV infection (1 x 10⁷ PFU) and a brief course of treatment with poly(I:C) (5 µg/g body weight x 3) induced diabetes in 100% of treated BBDR rats (n = 6; Table I). In addition, we confirmed that RV alone at a dose of 1 x 10⁷ PFU induced diabetes in 40% of 21- to 25-day-old BBDR rats (n = 27), a rate comparable to that reported previously (13) but significantly lower than that in rats treated with RV plus poly(I:C). Combination treatment with both RV and poly(I:C) also induced diabetes more rapidly (10–14 days) than did treatment with RV alone (16–27 days, p < 0.001). In contrast, infection of BBDR rats with H-1 virus (1 x 10⁷ PFU) failed to induce any diabetes, either alone (n = 6) or in combination with poly(I:C) (n = 10; Table I). Light microscopic examination of the pancreata of six of these animals revealed no evidence of insulitis. Control rats (n = 5) treated with poly(I:C) alone also failed to develop diabetes.

To begin to dissect the mechanism by which RV but not H-1 induces autoimmune diabetes in the BBDR rat, we studied the classical host humoral immune response to viral infection. As shown in Fig. 1 and consistent with previous observations (17), BBDR rats infected with either RV or H-1 (neither treated with poly(I:C)) generated a robust humoral immune response to the intact virus by day 14. Infection with H-1 generated Abs that cross-reacted only minimally with RV, and infection with RV generated Abs with no detectable cross-reactivity with H-1. The data suggest that infection with 10⁷ PFUs of RV or H-1 generates comparable virus-specific humoral immune responses in the BBDR rat.

View larger version (18K): [in this window] [in a new window]   FIGURE 1. Antibody responses. Two groups of three BBDR rats each were infected with 107 PFUs of RV or H-1. Serum from infected animals was recovered 14 days later and assayed for the presence of virus-specific Abs as described in Materials and Methods. Each bar represents the mean ± SD.

Having characterized the humoral immune response, we next investigated the cellular immune response to infection with these two agents. To quantify the CD8⁺ T cell response in the rat, we developed a rat assay system using intracellular IFN- as a marker for virus-specific CD8⁺ T cells; the assay was based on a similar system validated in the mouse (27).

RV infection induces virus-specific CD8⁺ IFN-⁺ T cells in BBDR rats.

BBDR rats were infected with RV and the presence of CD8⁺ IFN--producing cells in spleens was quantified by flow microfluorometry. We observed that CD8⁺ IFN-⁺ cells were present in spleens of all infected animals by day 11 and they remained readily detectable on days 14 and 21 (Table II). A representative set of flow cytometric profiles are shown in Fig. 2, left column. The specificity of RV-induced CD8⁺IFN-⁺ cells was established by incubating spleen cells from infected animals in the presence of uninfected APCs and in the absence of APCs. In both assays, very few CD8⁺IFN-⁺ cells (<0.2%) were detected.

View this table: [in this window] [in a new window]   Table II. CD8+IFN-+ T lymphocytes in RV- and H-1-infected ratsa

View larger version (35K): [in this window] [in a new window]   FIGURE 2. Percentage of CD8+IFN-+ cells in RV-infected BBDR rats. BBDR rats 21–25 days of age were randomized and either left untreated or infected with 107 PFU of RV. On the days indicated, spleen cells from individual animals were recovered and incubated in the presence of RV-infected Y3 APCs as described in Materials and Methods. After 5 h, spleen cells were recovered and processed for surface staining with Ab directed against CD8 and intracellular staining with Ab directed against IFN- as described in Materials and Methods. Horizontal (IFN-) and vertical (CD8) axes indicate fluorescence intensity. The percentage of CD8+ cells that were also IFN-+ is indicated in the upper right quadrant of each panel. The percentage of total CD8+ cells ranged from 6 to 43%. The percentage of cells that reacted with the CD8 isotype control Ab was <0.3% and the percentage of CD8+ cells that reacted with the IFN- isotype control Ab was <0.1%. Shown is a representative example of five independent trials.

View larger version (36K): [in this window] [in a new window]   FIGURE 3. Phenotype of CD8+IFN-+ cells. BBDR rats 21–25 days of age were infected with 107 PFU of RV on day 0. On day 14, the animals were killed and spleen cells were incubated in the presence of RV-infected Y3 APCs as described in Materials and Methods. After 5 h, spleen cells were recovered and processed for intracellular staining with Ab directed against IFN- and for surface staining with Abs directed against CD4, CD8, TCR, or NKR-P1A. A, Horizontal (IFN-) and vertical (CD8) axes indicate fluorescence intensity. B–D, Histograms indicative of the number of cells in the gate of A that stained positively for expression of TCR (B), NKR-P1A (C), and CD4 (D). Horizontal axes present fluorescence intensity and the vertical axis indicates cell number. For comparison, the insets in each panel show the fluorescence intensity profile of the total cell population in A stained to detect the same surface Ag. In the insets, the dotted lines depict the fluorescence intensity of cells stained with isotype control Abs. The percentage of cells that reacted with the CD8 isotype control Ab was <0.3% and the percentage of CD8+ cells that reacted with the IFN- isotype control Ab was <0.1%. Shown is a representative set of histograms from two similar replicates.

RV infection induces a cellular immune response in diabetes-resistant WF rats that is comparable to that observed in the BBDR rat.

To determine whether the CD8⁺ T cell response to RV in the BBDR rat is comparable to that of a normal, diabetes-resistant rat, we quantified the number of CD8⁺IFN-⁺ cells in RV-infected WF rats. The data show that RV-specific CD8⁺IFN-⁺ splenic T cells became detectable on day 11 after infection, remained elevated on day 14, and declined by day 21 (Table II). Flow cytometric profiles were comparable to those obtained using BBDR spleen cells (data not shown). Similar analyses were performed on PBMC from RV-infected WF rats. Again, RV-specific CD8⁺IFN-⁺ PBMC became detectable on day 11 after infection, remained elevated on day 14, and declined by day 21 (Table II).

	H-1 infection also induces a CD8+IFN-+ T cell response in BBDR rats

Top Abstract Introduction Materials and Methods Results H-1 infection also induces... Infection of BBDR rats... Discussion References

 
We next quantified the cellular immune response of BBDR rats infected with H-1 virus. In preliminary analyses, we observed that spleen cells from H-1-infected BBDR rats that were incubated in the presence of H-1-infected Y3 cells expressed IFN- at the same level as did control spleen cells from uninfected rats. We therefore could not directly test for the presence of H-1-specific CD8⁺ T cells. However, because GenBank data indicate that H-1 and RV share 77% capsid protein (VP1 and VP2) sequence homology and 100% nonstructural protein (NS1 and NS2) nucleotide sequence identity, we hypothesized that it would be possible to detect H-1-specific CD8⁺ T cell responses using RV-infected Y3 APCs.

Using this approach, we documented that CD8⁺IFN-⁺ spleen cells were detectable on day 11 after H-1 infection and remained elevated on day 14 (Table II). Representative flow cytometric profiles are shown in Fig. 4. Similar analyses were performed on PBMC and we observed that CD8⁺IFN-⁺ T cells became detectable on day 11 and remained elevated on day 14 (Table II).

View larger version (38K): [in this window] [in a new window]   FIGURE 4. Percentage of CD8+IFN-+ cells in H-1-infected BBDR rats. BBDR rats 21–25 days of age were infected with 107 PFU of H-1 on day 0. Spleen cells were obtained on days 8, 11, and 14 and incubated in the presence of Y3 cells that had been infected with RV as described in Materials and Methods. After 5 h, spleen cells were recovered and the percentage of CD8+IFN-+ cells was quantified as in Fig. 2. Horizontal (IFN-) and vertical (CD8) axes indicate fluorescence intensity. The percentage of CD8+ IFN-+ cells is indicated in the upper right quadrant of each panel. The percentage of CD8+ cells ranged from 8 to 25%. The percentage of cells that reacted with the CD8 isotype control Ab was <0.2% and the percentage of CD8+ cells that reacted with the IFN- isotype control Ab was <0.1%. Shown is a representative example of four similar independent trials.

	Infection of BBDR rats with RV but not H-1 alters the frequency of CD4+CD25+ T cells in spleen and pancreatic lymph nodes

Top Abstract Introduction Materials and Methods Results H-1 infection also induces... Infection of BBDR rats... Discussion References

 
The above data document that RV and H-1 induced similar humoral and cellular immune responses in BBDR rats. To further dissect the mechanism by which RV but not H-1 induces autoimmune diabetes in the BBDR rat, we next hypothesized that these viruses differentially modified the immune system of the infected host. We began testing this hypothesis by immunophenotyping hosts at different time points after infection.

RV induces similar changes in frequencies of peripheral CD4⁺ and CD8⁺ T cells in diabetes-susceptible BBDR and normal WF rats.

BBDR rats were infected with RV, and spleen cells were isolated 4, 8, 11, 14, and 21 days after infection and analyzed by flow cytometry. As shown in Fig. 5A, we observed that as early as 4 days after infection, the frequency of CD4⁺TCR⁺ T cells was less in infected rats than in uninfected controls. The percentage of CD4⁺TCR⁺ T cells in infected rats remained lower than in controls until day 8 after infection and was essentially identical in the two groups by day 21. As also shown in Fig. 5B, we observed that on day 4 after infection the frequency of CD8⁺TCR⁺ T cells was less in infected rats than in uninfected controls. However, as would be expected in virally infected animals, the percentage of these CD8⁺TCR⁺ T cells thereafter rose and was higher in infected animals than in uninfected controls on days 14 and 21 after infection.

View larger version (27K): [in this window] [in a new window]   FIGURE 5. Frequency of CD4+TCR+ and CD8+TCR+ spleen cells in BBDR and WF rats infected with RV or H-1. Groups of BBDR (A, B, F, and G) rats were either untreated or infected with RV or H-1. Groups of WF rats (C and D) were either untreated or infected with RV. On the days indicated, animals were killed and spleen cells were analyzed for the percentage of CD4+TCR+ and CD8+ TCR+ cells by flow microfluorometry. Each data point represents the mean ± SD of 2–19 measurements, as indicated in the legends in each panel.

H-1 infection does not induce changes in frequencies of peripheral CD4⁺ or CD8⁺ T cells in BBDR rats.

We next determined the percentages of CD4⁺TCR⁺ and CD8⁺TCR⁺ spleen cells in control and H-1-infected BBDR rats. As shown in Fig. 5, the frequency of both CD4⁺TCR⁺ (E) and CD8⁺TCR⁺ (F) T cells was similar in control and infected rats on days 4, 11, and 14 after infection.

RV but not H-1 or poly(I:C) alters the frequency of CD4⁺CD25⁺ T cells in both spleen and pancreatic lymph nodes of BBDR rats.

Because CD4⁺CD25⁺ T cells may play a major role in regulating autoreactive diabetogenic T cells in mice and rats (11, 20, 22, 23), we next tested the hypothesis that RV induces autoimmune diabetes by altering the proportion of these cells. BBDR rats were infected with RV and their spleen cells were isolated and analyzed by flow microfluorometry 3 to 5 days after infection.

As shown in Table III, infection with RV was associated with a reduction in the proportion of CD4⁺ and CD8⁺ T cells in the spleen of BBDR rats. These data are consistent with the day 4 data shown in Fig. 5. RV infection was also associated with a reduction in the percentage of CD4⁺, but not CD8⁺ splenic T cells in WF rats (Table III). RV infection did not, however, alter the frequency of CD8⁺ T cells in pancreatic lymph nodes in either strain, or of CD4⁺ T cells in the BBDR rat. RV infection did appear to reduce the frequency of CD4⁺ pancreatic lymph node T cells in WF rats, but this reduction did not achieve statistical significance when adjusted for multiple comparisons.

View larger version (38K): [in this window] [in a new window]   FIGURE 6. Frequency of CD4+CD25+ in RV-infected BBDR and WF rats and characterization of CD4+CD25+ cells. BBDR and WF rats were either untreated or infected with RV. On days 4 and 5, pancreatic lymph node cells were reacted with Abs directed against TCR, CD4, CD25, and CD62L. Representative dot plots demonstrating the presence of CD4+CD25+ cells in the pancreatic lymph nodes in uninfected and RV-infected animals are shown as indicated in this figure. The number above the arrow in each panel in the left column indicates the percentage of CD4+ cells that express CD25+ cells in the pancreatic lymph nodes. In the right column are histograms demonstrating the number of BBDR and WF pancreatic lymph node cells (vertical axis) in the CD4+CD25+ gate that stained positively for expression of CD62L (horizontal axis). The insets in the right column present the fluorescence intensity profile of total cells that expressed CD62L (solid lines) and isotype controls (dotted lines).

Unlike RV, neither H-1 infection nor treatment with poly(I:C) altered the percentages of CD4⁺, CD8⁺, or CD4⁺CD25⁺ T cells in either the spleen or pancreatic lymph nodes of BBDR rats. Treatment with poly(I:C) was associated with a reduction in the percentage of splenic CD4⁺ T cells in the BBDR rat, but this reduction did not achieve statistical significance.

RV infection increases the frequency of CD4⁺CD25⁺ T cells in pancreatic lymph nodes of BBDR rats but not their incorporation of BrdU. The increase of CD4⁺CD25⁺ T cells in the pancreatic lymph nodes of RV-infected BBDR rats could be due to activation of CD4⁺ T cells, leading to their expansion by proliferation or an increase in the accumulation of nonproliferating CD4⁺CD25⁺ T cells with regulatory characteristics. The expression of CD62L^high on the CD4⁺CD25⁺ T cells suggests that they have anergic properties, but it is difficult to distinguish between activated and anergic CD4⁺CD25⁺ T cells based solely on phenotypic markers (32).

To determine whether the pancreatic lymph node CD4⁺ T cells expressing CD25 were activated or hyporesponsive, we assessed their proliferation by analysis of in vivo incorporation of BrdU into CD4⁺CD25⁺ in uninfected and RV-infected BBDR rats. As expected, the percentage of CD4⁺ that express CD25⁺ was increased (11.8 ± 2.1%, n = 8) in the pancreatic lymph nodes 3–4 days after RV infection as compared with uninfected BBDR rats (8.1 ± 2.1%, n = 4, p < 0.025). The percentage of CD4⁺ cells that were CD25⁺ and incorporated BrdU in RV-infected (1.5 ± 0.6%, n = 8) and uninfected (0.8 ± 0.3%, n = 4) BBDR rats was statistically similar (p = NS). In addition, the percentage of CD4⁺ cells that were CD25⁺ but did not incorporate BrdU in RV-infected (9.6 ± 1.5%) and uninfected (7.6 ± 2.6%) BBDR rats was statistically similar (p = NS). The data show that only 16 and 11% of CD4⁺ cells that were CD25⁺ in the pancreatic lymph nodes of RV-infected and uninfected BBDR rats, respectively, incorporated BrdU. Representative flow cytometric profiles of CD4⁺ cells that are CD25⁺ and incorporated BrdU are shown in Fig. 7.

View larger version (50K): [in this window] [in a new window]   FIGURE 7. Percentage of CD4+CD25+ cells that incorporate BrdU in the pancreatic lymph nodes of uninfected and RV-infected BBDR rats. BBDR rats were uninfected or infected with 107 PFU of RV and injected with BrdU 24 and 12 h before recovery of pancreatic lymph node cells as described in Materials and Methods. Cells were recovered from RV-infected rats on days 3 and 4 after infection. Cells from uninfected and RV-infected BBDR rats were labeled with anti-CD4, anti-CD25, and anti-BrdU mAbs as described in Materials and Methods. Samples were analyzed by flow microfluorometry for the percentage of CD4+CD25+ that had incorporated BrdU. In the left column are representative dot plots demonstrating the presence of CD4+CD25+ cells in pancreatic lymph nodes in uninfected BrdU isotype control (top row), uninfected (middle row), and in day 3 RV-infected (bottom row) BBDR rats. The CD4+ cell compartment is identified with the solid line rectangular gate; the dashed line represents the gating used to identify the CD4+CD25+ cells. Numbers indicate the percentage of CD4+ T cells that express CD25. In the right column are dot plots gated on total CD4+ T cells (the entire solid line rectangular gate in the left column) that express CD25 (horizontal axis) and label with anti-BrdU mAb (vertical axis). The numbers give the percentage of total CD4+ T cells in each quadrant. The two control panels in the top row indicate the percentage of CD4+CD25+ T cells in the pancreatic lymph nodes of uninfected BBDR rats (left column) and the percentage of total CD4+ cells that are labeled with CD25 and the anti-BrdU isotype control Ab (right column). Shown are representative histograms drawn from four to eight independent trials; the overall averages are given in Results.

To determine whether the CD4⁺CD25⁺ cell phenotype in BBDR rats identifies a regulatory cell population that can suppress the proliferation of CD4⁺CD25^- cells, we purified BBDR CD4⁺CD25⁺ lymph node cells and cocultured them with CD4⁺CD25^- cells in the presence of Con A plus irradiated splenic APCs. As shown in Fig. 8, addition of purified CD4⁺CD25⁺ cells to cultures of CD4⁺CD25^- cells stimulated with Con A in the presence of APCs inhibited their proliferation by >50% (n = 5 independent trials). To control for cell density and "crowding," we performed additional cultures in which we added irradiated T cells in place of CD4⁺CD25⁺ cells. This did not cause any decrease in the proliferation of CD4⁺CD25^- cells (data not shown).

View larger version (13K): [in this window] [in a new window]   FIGURE 8. BBDR CD4+CD25+ lymph node cells exhibit suppressive activity in vitro. CD4+CD25- and CD4+CD25+ T cells were purified from pools of cervical and mesenteric lymph nodes from 30- to 35-day-old BBDR rats as described in Materials and Methods. Purified CD4+CD25+ T cells were cocultured for 72 h with CD4+CD25- T cells ("responders") at a ratio of 2:1 in the presence of irradiated APCs and Con A (). Control cultures consisted of CD4+CD25- T cells stimulated with Con A in the presence of irradiated APCs (). Cell proliferation was measured by incorporation of [3H]thymidine during the final 14 h of culture. Proliferative responses are presented as the percentage of proliferation detected under coculture conditions relative to the proliferation of CD4+CD25- responder T cells (normalized to 100%). Each bar represents the results of five independent experiments, each performed in quadruplicate shown as the mean ± 1 SD.

	Discussion

Top Abstract Introduction Materials and Methods Results H-1 infection also induces... Infection of BBDR rats... Discussion References

We first documented that RV-UMass infection of BBDR rats induces diabetes, but infection with its close homologue H-1 does not. Both viruses induced humoral and cellular immune responses. RV infection also caused a substantial decrease in splenic CD4⁺CD25⁺ T cells in both BBDR rats and diabetes-resistant WF rats. In contrast, RV infection increased the levels of CD4⁺CD25⁺ T cells in the pancreatic lymph nodes of BBDR rats but had no effect on this population in WF rats. The data suggest that RV but not H-1 infection alters T cell regulation in the BBDR rat and permits the expression of diabetes. Consistent with this interpretation, we also documented that the CD4⁺CD25⁺ phenotype identifies a rat T cell population that can suppress Con A-induced proliferation of CD4⁺CD25^- cells. In their aggregate, the results imply that the reduction in splenic CD4⁺ CD25⁺ cells observed in RV-infected animals is virus specific, whereas the increase in pancreatic lymph node CD4⁺CD25⁺ cells is both virus- and rat strain specific. More generally, the results suggest a mechanism that could link an underlying genetic predisposition with an environmental perturbant that can transform a "regulated predisposition" into autoimmune diabetes. That mechanism is the failure to maintain regulatory CD4⁺CD25⁺ T cell function.

Previous studies have suggested that CD4⁺ regulatory T cells and not molecular mimicry may be important for the expression of BB rat diabetes after RV infection (19), and the present studies are consistent with this view. We began by identifying a nondiabetogenic, highly homologous parvovirus, H-1, and then using it to define the specific immune responses associated with progression to autoimmunity in RV-infected animals. After documenting that RV but not H-1-induced diabetes (with or without poly(I:C)), we analyzed the humoral and cellular immune responses to both pathogens.

We observed that both RV-UMass and H-1 elicited strong Ab responses in BBDR rats. The magnitude and kinetics of the humoral responses to both viruses were similar. Interestingly, there was no evidence of cross-reactivity between the two Ab responses to intact virus. This assay, however, likely detects only capsid VP proteins and not the NS proteins that are essentially identical in RV and H-1 (17).

We next observed that both RV-UMass and H-1 infection induced antiviral CD8⁺ T cell responses. As expected, the level of CD8⁺ T cells in the circulation of both BBDR and WF rats rose shortly after infection with RV. Some of these CD8⁺ T cells were shown to be RV-reactive CD8⁺IFN-⁺ T cells, the majority of which expressed the NK cell marker NKR-P1A. These are characteristics of the activated virus-specific CD8⁺ T cells (33, 34). In contrast, we did not detect an increase in the total CD8⁺ T cell population in the spleens of rats infected with H-1. Importantly, however, H-1-infected rats also generated antiviral CD8⁺ IFN-⁺ T cells. In contrast to the humoral immune responses, we observed the cellular immune responses to be cross-reactive.

One possible explanation for the differences between H-1 and RV with respect to diabetes induction could be differences in cell- and tissue-specific tropism. RV infects lymphohemopoietic cells including T cells and B cells (17). RV also infects endothelial cells and megakaryocytes (15). In contrast, H-1 predominantly infects nonlymphohemopoietic organs such as the kidney (35). It is not yet clear whether either RV or H-1 infects macrophages or dendritic cells. These differences in tropism, with RV targeted more toward the lymphohemopoietic cells than H-1, might position RV to modulate the immune system of infected animals in ways that H-1 could not.

Our data support strongly the hypothesis that RV but not H-1 infection selectively alters the levels of CD4⁺CD25⁺ regulatory T cells in the draining pancreatic lymph nodes of BBDR rats. We documented that alteration of CD4⁺CD25⁺ T cells in the spleen of BBDR and WF rats by RV was similar. In addition, we documented that RV but not H-1 reduced the frequency of splenic CD4⁺CD25⁺ T cells. The crucial observation providing insight into the potential mechanism by which RV may selectively induce diabetes in BBDR rats was its association with an increase CD4⁺CD25⁺ T cells in the draining pancreatic lymph nodes of BBDR but not WF rats.

We recognize that attribution of regulatory function to these CD4⁺CD25⁺ T cells on the basis of phenotype is suggestive but not definitive (32). To test directly whether CD4⁺CD25⁺ T cells in BBDR rats are capable of regulatory function, we performed coculture experiments. We documented that CD4⁺CD25⁺ T cells can inhibit the proliferation of Con A-stimulated CD4⁺CD25^- T cells in vitro. Our observation that there was no difference in the incorporation of BrdU by CD4⁺CD25⁺ T cells in uninfected and infected rats suggests that the increased levels of CD4⁺CD25⁺ T cells in the pancreatic lymph nodes of RV-infected BBDR rats is not due to proliferation of activated CD4⁺ T cells. CD25 and CD62L^high expression in the absence of proliferation are characteristics associated with anergic (regulatory) CD4⁺CD25⁺ T cells. In their aggregate, these three characteristics, CD4⁺CD25⁺CD62L^high phenotype, in vitro suppressive activity, and low level of proliferative activity in vivo, strongly suggest that RV infection induces the accumulation of suppressor cell populations in the draining pancreatic lymph nodes of animals genetically predisposed to autoimmune diabetes.

The regulatory role of CD4⁺CD25⁺ T cells has been documented in many experimental systems (11, 20, 21, 22, 23). Involvement of CD4⁺CD25⁺ T cells in the regulation of autoimmunity was suggested in early studies in which neonatal thymectomy of mice led to a reduction in the frequency of peripheral CD4⁺CD25⁺ T cells and the development of multiple autoimmune disorders (21, 36). Adoptive transfer of CD4⁺CD25⁺ into these thymectomized mice prevented development of autoimmunity.

In another model of autoimmune diabetes, evidence has accumulated that CD4⁺CD25⁺ regulatory T cell defects may be involved in disease expression in NOD mice. In normal mice, CD4⁺CD25⁺ T cells that express the naive cell marker CD62L represent 5–10% of spleen CD4⁺ cells (31, 37). CD4⁺CD25⁺ T cell levels are reduced in NOD mice (23), and diabetes is prevented in NOD mice by the adoptive transfer of CD4⁺CD25⁺ cells to prediabetic animals (37). CD4⁺CD25⁺ T cells are also reduced in prediabetic CD80/CD86 and CD28-deficient NOD mice and the transfer of CD4⁺CD25⁺ T cells to these mice prevents diabetes (23).

Our data document that CD4⁺CD25⁺ T cells in the BBDR rat have suppressive activity in vitro, and in preliminary experiments, we have determined that purified populations of CD4⁺CD25⁺ T cells can also delay the expression of spontaneous diabetes in BBDP rats (J.-L. Hillebrands, unpublished observations). We further note that some of the earliest evidence that regulatory T cells can control autoimmune disease expression was generated in studies of the BB rat (12). BBDP rats spontaneously develop diabetes and adoptive transfer of a CD4⁺ART2⁺ T cell population prevents diabetes expression. (ART2 is a rat surface alloantigen that marks a population of regulatory T cells (38).) Conversely, depletion of ART2⁺ T cells in diabetes-resistant BBDR rats in combination with injection of an immune system activator such as poly(I:C) or RV infection induces diabetes in close to 100% of these animals (12). More recently, we have observed that the majority of CD4⁺CD25⁺ T cells express ART2 on their surface and that depletion of ART2⁺ cells leads to elimination of essentially all of the CD4⁺CD25⁺ T cells in the periphery (D. Zipris, unpublished observations). Similar observations have been made in another rat model of diabetes. In the thymectomized, irradiated PVG rat, it has been shown that adoptive transfer of CD4⁺CD25⁺CD45RC^- T cells prevents diabetes (37, 39, 40).

Our documentation of an increase in CD4⁺CD25⁺ T cells in the pancreatic lymph nodes of RV-infected BBDR rats is reminiscent of a recent report that CD4⁺CD25⁺ T cells accumulate preferentially in pancreatic lymph nodes and islets in a transgenic mouse model in which diabetes is induced by regulated expression of TNF- and CD80 in pancreatic islets (11). The basis for this increase in CD4⁺CD25⁺ T cells in the lymph nodes draining the pancreas during the prediabetic phase of disease in NOD mice and in BBDR rats is not known.

In conclusion, our data suggest that RV-infected BBDR rats can be used to identify mechanisms that link underlying genetic predisposition to autoimmune diabetes (BBDR vs WF) with environmental perturbation (RV vs H-1). More generally, modulation of CD4⁺CD25⁺ T cells may be an important component of the mechanism by which environmental agents induce type 1A diabetes in genetically susceptible individuals.

	Acknowledgments

 
We thank Michael Bates for technical assistance.

	Footnotes

 
¹ This work was supported in part by Grants DK49106, DK36024, and DK25306 and Center Grant DK32520 from the National Institutes of Health. J.-L.H. was supported by the Ter Meulen Fund, Royal Netherlands Academy of Arts and Sciences (Project 6419) and the Dutch Diabetes Foundation (Project 2001.05.003).

² The contents of this publication are solely the responsibility of the authors and do not necessarily represent the official views of the National Institutes of Health.

³ Address correspondence and reprint requests to Dr. Aldo A. Rossini, Diabetes Division, University of Massachusetts Medical School, 373 Plantation Street, Suite 218, Worcester, MA 01605. E-mail address: aldo.rossini{at}umassmed.edu

⁴ Abbreviations used in this paper: NOD, nonobese diabetic; RV-UMass, UMass strain of rat virus; RPV-1 rat parovirus 1; BrdU, 5-bromo-2'-deoxyuridine.

Received for publication April 24, 2002. Accepted for publication January 30, 2003.

	References

Top Abstract Introduction Materials and Methods Results H-1 infection also induces... Infection of BBDR rats... Discussion References

 

1. Yoon, J. W., H.-S. Jun. 2000. Role of viruses in the pathogenesis of type 1 diabetes mellitus. D. LeRoith, and S. I. Taylor, and J. M. Olefsky, eds. Diabetes Mellitus: A fundamental and Clinical Text 2nd Ed.419. Lippincott Williams & Wilkins, Philadelphia.
2. Atkinson, M., E. H. Leiter. 1999. The NOD mouse model of insulin dependent diabetes: as good as it gets?. Nature Med. 5:601.[Medline]
3. Hermitte, L., B. Vialettes, P. Naquet, C. Atlan, M.-J. Payan, P. Vague. 1990. Paradoxical lessening of autoimmune processes in non-obese diabetic mice after infection with the diabetogenic variant of encephalomyocarditis virus. Eur. J. Immunol. 20:1297.[Medline]
4. Wilberz, S., H. J. Partke, F. Dagnaes-Hansen, L. Herberg. 1991. Persistent MHV (mouse hepatitis virus) infection reduces the incidence of diabetes mellitus in non-obese diabetic mice. Diabetologia 34:2.[Medline]
5. Oldstone, M. B. A.. 1990. Viruses as therapeutic agents. I. Treatment of nonobese insulin-dependent diabetes mice with virus prevents insulin-dependent diabetes mellitus while maintaining general immune competence. J. Exp. Med. 171:2077.[Abstract/Free Full Text]
6. Leiter, E. H.. 1998. NOD mice and related strains: origins, husbandry, and biology. E. H. Leiter, and M. A. Atkinson, eds. NOD Mice and Related Strains: Research Applications in Diabetes, AIDS, Cancer and Other Diseases 1. R. G. Landes Co., Austin, TX.
7. Serreze, D. V., E. W. Ottendorfer, T. M. Ellis, C. J. Gauntt, M. A. Atkinson. 2000. Acceleration of type 1 diabetes by a Coxsackievirus infection requires a preexisting critical mass of autoreactive T-cells in pancreatic islets. Diabetes 49:708.[Abstract]
8. Horwitz, M. S., T. Krahl, C. Fine, J. Lee, N. Sarvetnick. 1999. Protection from lethal Coxsackievirus-induced pancreatitis by expression of interferon. J. Virol. 73:1756.[Abstract/Free Full Text]
9. Imagawa, A., T. Hanafusa, S. Tamura, M. Moriwaki, N. Itoh, K. Yamamoto, H. Iwahashi, K. Yamagata, M. Waguri, T. Nanmo, et al 2001. Pancreatic biopsy as a procedure for detecting in situ autoimmune phenomena in type I diabetes: close correlation between serological markers and histological evidence of cellular autoimmunity. Diabetes 50:1269.[Abstract/Free Full Text]
10. LeCompte, P. M., W. Gepts. 1977. The pathology of juvenile diabetes. B. W. Volk, and K. F. Wellmann, eds. The Diabetic Pancreas 325. Plenum, New York.
11. Green, E. A., Y. Choi, R. A. Flavell. 2002. Pancreatic lymph node-derived CD4⁺CD25⁺ T_reg cells: highly potent regulators of diabetes that require TRANCE-RANK signals. Immunity 16:183.[Medline]
12. Mordes, J. P., R. Bortell, H. Groen, D. L. Guberski, A. A. Rossini, D. L. Greiner. 2001. Autoimmune diabetes mellitus in the BB rat. A. A. F. Sima, and E. Shafrir, eds. Animal Models of Diabetes: A Primer 1. Harwood Academic Publishers, Amsterdam.
13. Guberski, D. L., V. A. Thomas, W. R. Shek, A. A. Like, E. S. Handler, A. A. Rossini, J. E. Wallace, R. M. Welsh. 1991. Induction of type 1 diabetes by Kilham’s rat virus in diabetes resistant BB/Wor rats. Science 254:1010.[Abstract/Free Full Text]
14. Thomas, V. A., B. A. Woda, E. S. Handler, D. L. Greiner, J. P. Mordes, A. A. Rossini. 1991. Altered expression of diabetes in BB/Wor rats by exposure to viral pathogens. Diabetes 40:255.[Abstract]
15. Brown, D. W., R. M. Welsh, A. A. Like. 1993. Infection of peripancreatic lymph nodes but not islets precedes Kilham rat virus-induced diabetes in BB/Wor rats. J. Virol. 67:5873.[Abstract/Free Full Text]
16. Ellerman, K. E., C. A. Richards, D. L. Guberski, W. R. Shek, A. A. Like. 1996. Kilham rat virus triggers T-cell-dependent autoimmune diabetes in multiple strains of rat. Diabetes 45:557.[Abstract]
17. Jacoby, R. O., L. J. Ball-Goodrich, D. G. Besselsen, M. D. McKisic, L. K. Riley, A. L. Smith. 1996. Rodent parvovirus infections. Lab. Anim. Sci. 46:370.[Medline]
18. Cherry, J. D.. 1999. Parvovirus infections in children and adults. Adv. Pediatr. 46:245.[Medline]
19. Chung, Y. H., H. S. Jun, M. Son, M. Bao, H. Y. Bae, Y. Kang, J. W. Yoon. 2000. Cellular and molecular mechanism for Kilham rat virus-induced autoimmune diabetes in DR-BB rats. J. Immunol. 165:2866.[Abstract/Free Full Text]
20. McHugh, R. S., E. M. Shevach, A. M. Thornton. 2001. Control of organ-specific autoimmunity by immunoregulatory CD4⁺CD25⁺ T cells. Microbes Infect. 3:919.[Medline]
21. Sakaguchi, S., T. Takahashi, S. Yamazaki, Y. Kuniyasu, M. Itoh, N. Sakaguchi, J. Shimizu. 2001. Immunologic self tolerance maintained by T-cell-mediated control of self-reactive T cells: implications for autoimmunity and tumor immunity. Microbes Infect. 3:911.[Medline]
22. Stephens, L. A., D. Mason. 2001. Characterisation of thymus-derived regulatory T cells that protect against organ-specific autoimmune disease. Microbes Infect. 3:905.[Medline]
23. Salomon, B., D. J. Lenschow, L. Rhee, N. Ashourian, B. Singh, A. Sharpe, J. A. Bluestone. 2000. B7./CD28 costimulation is essential for the homeostasis of the CD4⁺CD25⁺ immunoregulatory T cells that control autoimmune diabetes. Immunity 12:431.[Medline]
24. Foy, T. M., D. M. Shepherd, F. H. Durie, A. Aruffo, J. A. Ledbetter, R. J. Noelle. 1993. In vivo CD40-gp39 interactions are essential for thymus-dependent humoral immunity. II. Prolonged suppression of the humoral immune response by an antibody to the ligand for CD40, gp39. J. Exp. Med. 178:1567.[Abstract/Free Full Text]
25. Whalen, B. J., D. L. Greiner, J. P. Mordes, A. A. Rossini. 1994. Adoptive transfer of autoimmune diabetes mellitus to athymic rats: synergy of CD4⁺ and CD8⁺ T cells and prevention by RT6⁺ T cells. J. Autoimmun. 7:819.[Medline]
26. Iwakoshi, N. N., D. L. Greiner, A. A. Rossini, J. P. Mordes. 1999. Diabetes prone BB rats are severely deficient in natural killer T cells. Autoimmunity 31:1.[Medline]
27. Selin, L. K., M. Y. Lin, K. A. Kraemer, D. M. Pardoll, J. P. Schneck, S. M. Varga, P. A. Santolucito, A. K. Pinto, R. M. Welsh. 1999. Attrition of T cell memory: selective loss of LCMV epitope-specific memory CD8 T cells following infections with heterologous viruses. Immunity 11:733.[Medline]
28. Tough, D. F., J. Sprent. 1994. Turnover of naive- and memory-phenotype T cells. J. Exp. Med. 179:1127.[Abstract/Free Full Text]
29. Glantz, S. A.. 1981. Primer of Biostatistics 63. McGraw-Hill, New York.
30. Kaplan, E. L., P. Meier. 1958. Nonparametric estimation from incomplete observations. J. Am. Statist. Assoc. 53:457.
31. Herbelin, A., J. M. Gombert, F. Lepault, J. F. Bach, L. Chatenoud. 1998. Mature mainstream TCR⁺CD4⁺ thymocytes expressing L-selectin mediate "active tolerance" in the nonobese diabetic mouse. J. Immunol. 161:2620.[Abstract/Free Full Text]
32. Wu, A. J., H. Hua, S. H. Munson, H. O. McDevitt. 2002. Tumor necrosis factor- regulation of CD4⁺C25⁺ T cell levels in NOD mice. Proc. Natl. Acad. Sci. USA 99:12287.[Abstract/Free Full Text]
33. Kambayashi, T., E. Assarsson, J. Michaelsson, P. Berglund, A. D. Diehl, B. J. Chambers, H. G. Ljunggren. 2000. Emergence of CD8⁺ T cells expressing NK cell receptors in influenza A virus-infected mice. J. Immunol. 165:4964.[Abstract/Free Full Text]
34. Slifka, M. K., J. L. Whitton. 2000. Activated and memory CD8⁺ T cells can be distinguished by their cytokine profiles and phenotypic markers. J. Immunol. 164:208.[Abstract/Free Full Text]
35. Besselsen, D. G., C. L. Besch-Williford, D. J. Pintel, C. L. Franklin, R. R. Hook, Jr, L. K. Riley. 1995. Detection of H-1. parvovirus and Kilham rat virus by PCR. J. Clin. Microbiol. 33:1699.[Abstract]
36. Asano, M., M. Toda, N. Sakaguchi, S. Sakaguchi. 1996. Autoimmune disease as a consequence of developmental abnormality of a T cell subpopulation. J. Exp. Med. 184:387.[Abstract/Free Full Text]
37. Lepault, F., M. C. Gagnerault. 2000. Characterization of peripheral regulatory CD4⁺ T cells that prevent diabetes onset in nonobese diabetic mice. J. Immunol. 164:240.[Abstract/Free Full Text]
38. Bortell, R., T. Kanaitsuka, L. A. Stevens, J. Moss, J. P. Mordes, A. A. Rossini, D. L. Greiner. 1999. The RT6. (Art2) family of ADP-ribosyltransferases in rat and mouse. Mol. Cell. Biochem. 193:61.[Medline]
39. Saoudi, A., B. Seddon, D. Fowell, D. Mason. 1996. The thymus contains a high frequency of cells that prevent autoimmune diabetes on transfer into prediabetic recipients. J. Exp. Med. 184:2393.[Abstract/Free Full Text]
40. Seddon, B., A. Saoudi, M. Nicholson, D. Mason. 1996. CD4⁺CD8^- thymocytes that express L-selectin protect rats from diabetes upon adoptive transfer. Eur. J. Immunol. 26:2702.[Medline]

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				M. J. Hessner, X. Wang, L. Meyer, R. Geoffrey, S. Jia, J. Fuller, A. Lernmark, and S. Ghosh Involvement of Eotaxin, Eosinophils, and Pancreatic Predisposition in Development of Type 1 Diabetes Mellitus in the BioBreeding Rat J. Immunol., December 1, 2004; 173(11): 6993 - 7002. [Abstract] [Full Text] [PDF]

				I. I. Mendez, Y.-H. Chung, H.-S. Jun, and J.-W. Yoon Immunoregulatory Role of Nitric Oxide in Kilham Rat Virus-Induced Autoimmune Diabetes in DR-BB Rats J. Immunol., July 15, 2004; 173(2): 1327 - 1335. [Abstract] [Full Text] [PDF]

				L. Wen, J. Peng, Z. Li, and F. S. Wong The Effect of Innate Immunity on Autoimmune Diabetes and the Expression of Toll-Like Receptors on Pancreatic Islets J. Immunol., March 1, 2004; 172(5): 3173 - 3180. [Abstract] [Full Text] [PDF]

				A. A. Rossini Autoimmune Diabetes and the Circle of Tolerance Diabetes, February 1, 2004; 53(2): 267 - 275. [Abstract] [Full Text]

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