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Previous Article | Next Article 
Molecular and Cellular Biology, May 2005, p. 3620-3629, Vol. 25, No. 9
0270-7306/05/$08.00+0 doi:10.1128/MCB.25.9.3620-3629.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.
Department of Medical Biophysics,1 The Campbell Family Institute for Breast Cancer Research,4 Ontario Cancer Institute, Departments of Medicine and PhysiologyUniversity of Toronto,2 The Advanced Medical Discovery Institute (AMDI),3 St. Michael's Hospital,5 McLaughlin Centre for Molecular Medicine, Toronto, Ontario, Canada6
Received 16 August 2004/ Returned for modification 1 November 2004/ Accepted 7 January 2005
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ABSTRACT |
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INTRODUCTION |
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Caspases are evolutionarily conserved cysteine-aspartyl specific proteases that play a key role in apoptosis. In mammals, there are over 14 caspases, of which some are involved in apoptosis and others in cytokine activation (10, 44, 53). There are two main apoptotic pathways: the extrinsic or cell death receptor pathway involving the tumor necrosis factor superfamily of receptors and the intrinsic or mitochondrial pathway (46, 53). Caspase-8 and caspase-9 are the upstream caspases involved in the extrinsic and intrinsic pathways, respectively. Caspase-3, -6, and -7 are effector caspases downstream of both pathways. Gene targeting strategies have provided valuable tools to study the physiologic function of individual caspases in vivo and have shown their roles not only in apoptosis but also in other fundamental cellular processes. Caspase-3 and -9 have been shown to play a critical role in developmental neuronal apoptosis (14, 21, 54). Caspase-3 is required in Fas-mediated cell death in activated peripheral T cells and hepatocytes but not in immature T cells from the thymus (14, 21, 51, 54). In B lymphocytes, caspase-3 plays a negative regulatory role in cell cycle progression (52); in keratinocytes, caspase-3 is critical in cellular differentiation (35). Caspase-8 is required for cardiac development and T-cell homeostasis (41, 42, 49). Collectively, these studies highlight the tissue- and context-specific functions of individual caspases and their intricate regulation in all facets of cellular function.
Several in vitro studies have suggested that caspase-dependent apoptotic pathways are essential for ß-cell apoptosis (12, 25, 26). Cultured human islet cells were shown to undergo apoptosis in response to elevated glucose concentrations by upregulating Fas, a receptor which activates the extrinsic apoptotic pathway (26). Furthermore, ß-cell expression of c-Flip, a molecule known to inhibit caspase-8-mediated apoptosis, was shown to inhibit Fas-induced apoptosis and induce ß-cell proliferation (25). In vivo studies of nonobese diabetic (NOD) mice and diabetes-prone BB/S rats also demonstrate presence of islet apoptosis by terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) (6, 23, 32). In 2003, Reddy et al. examined pancreatic islets of NOD mice during cyclophosphamide-accelerated diabetes, which showed caspase-3 immunoreactivity predominantly in intraislet macrophages but not islet cells, suggesting that apoptotic ß cells are cleared rapidly after diabetes onset (37). Even though islet apoptosis has been shown to be an essential event in both type 1 and type 2 diabetes (6, 27, 32), the specific in vivo role of individual caspases in ß-cell apoptosis and disease progression in diabetes models remains to be elucidated.
Here, we use a genetic approach to study the role of caspase-3 in ß-cell apoptosis, using Caspase-3 knockout (Casp3–/–) mice in the multiple-low-dose streptozotocin (MLDS) autoimmune diabetes model. Streptozotocin (STZ) is a glucose analog that selectively destroys pancreatic ß cells (11, 38). High-dose administration of STZ has been shown to directly cause DNA damage, resulting in massive necrosis of ß cells (40). However, MLDS has been shown to cause selective ß-cell destruction that in turn induces immune reactions against islets (24). Nude athymic mice do not develop diabetes with STZ treatment, suggesting that the STZ mechanism relies on T-cell functions to mediate its effects on islets (29).
Our results show that Casp3–/– mice are protected from developing diabetes and that their islets are resistant to apoptosis, following MLDS administration. Casp3+/– mice exhibit insulitis following MLDS, contributing to autoimmune diabetes initiation. In contrast, Casp3–/–mice do not develop insulitis. This key finding has led to examining the role of caspase-3 in the antigen cross-presentation pathway. For this purpose, we introduced a Casp3 null mutation into the RIP-GP/P14 double-transgenic model that expresses a defined ß-cell antigen and T-cell receptor specific for this defined antigen. These mice express a glycoprotein from lymphocytic choriomeningitis virus (LCMV) under the control of the rat insulin promoter (RIP) and is specifically recognized by CD8 T cells (P14) in the islets (34, 36). Using these mice, we show that caspase-3 is required for inducing activation and proliferation of ß-cell-specific T cells in the local PDLN. We show that caspase-3-dependent apoptosis is essential for initiating autoimmune diabetes by facilitating ß-cell-antigen presentation for T-cell activation and invasion of the islets.
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MATERIALS AND METHODS |
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Induction of diabetes and glucose monitoring. The mice were injected intraperitoneally with STZ (40 mg of STZ/kg of body weight) for 5 consecutive days (40). Tail vein glucose was monitored using One Touch Ultra (Life Scan; Johnson & Johnson).
Histology. Paraffin-embedded pancreatic tissue was stained with hematoxylin and eosin (H&E) and examined by light microscopy. To prepare specimen for TUNEL staining, paraffin sections were dewaxed and equilibrated in phosphate-buffered saline (PBS; pH 7.4), followed by incubation in 20 µg of proteinase K (Roche Biochemicals)/µl in 10 mM Tris-HCl (pH 7.5) for 15 min at 37°C. Following washing, the reaction mixture containing terminal deoxynucleotidyltransferase, labeled nucleotides, and DNA polymerase was then applied to sections in a humidified chamber for 90 min at 37°C, according to the manufacturer's instructions (Roche Biochemicals). Some sections were then washed in PBS and prepared for visualization with the fluorescent microscope. Anti-fluorescein antibody peroxidase conjugate was added to other sections for 30 min at 37°C. Sections were washed and incubated with 3,3-diaminobenzidine (Sigma) substrate and visualized by light microscopy.
Islet isolation. To isolate the islets, 2 ml of 2-mg/ml collagenase (type IV; Sigma) was injected into the common bile duct for pancreatic digestion. The digested pancreas was removed and incubated at 37° for 20 to 28 min. The digest was washed twice by resuspension in Hank's balanced salt solution (HBSS) and centrifuged at 400 x g for 1 min at 4°C. The pellet was resuspended in HBSS and filtered through gauze. Islets were then handpicked in HBSS. Islets were maintained in suspension culture in RPMI 1640 supplemented with 10% fetal bovine serum, 100 U of penicillin/ml, and 100 µg of streptomycin/ml and incubated at 37°C and 5% CO2.
In vitro STZ treatment. STZ was dissolved in 0.1 mM sodium citrate (pH 4.5) at 4°C and was added to each well of 96-well tissue culture plates containing 30 to 40 islets to obtain the required final STZ concentrations (0, 0.25, and 0.5 mM). Islet were incubated overnight (18 h) at 37°C in humidified air and 5% CO2.
Islet morphology and viability analysis of STZ-treated islets. Islet morphology in culture was assessed with a Zeiss inverted microscope. To assess viability, STZ-treated islets were trypsinized to generate single-cell suspensions. For the annexin V-7-amino-actinomycin D (7-AAD) binding assay, cells were resuspended with annexin V binding buffer and incubated with 1 µl of annexin V/100 µl of binding buffer and 0.5 µm of 7-AAD for 15 min at room temperature in the dark. Cells were washed again with annexin V binding buffer, and the percentage of 7-AAD+ and annexin V+ cells was determined by flow cytometry with a FACSCalibur (Becton Dickinson).
Immunohistochemistry.
Pancreatic tissue was fixed overnight in a solution of freshly prepared 4% paraformaldehyde in 0.1 M PBS, pH 7.4, at 4°C. Samples were dehydrated and prepared as paraffin blocks and stained with H&E as mentioned above. For detection of insulin, glucagon, and CD3
on sections, monoclonal anti-insulin, anti-glucagon, and anti-CD3 antibodies (1:100; DAKO), respectively, were used. The appropriate primary antibody was added in blocking buffer and incubated overnight at 4°C. Sections were washed and incubated in biotinylated secondary antibody at 1:2,000 for 2 h at room temperature, followed by washing and incubation with avidin biotin complex (Vectastain Elite ABC kit; Vector Laboratories, Burlingame, Calif.) at 1:100 for 1 h at room temperature. Sections were washed and incubated with 3,3-diaminobenzidine (Sigma).
Islet insulin and glucagon content. Insulin and glucagon composition per islet were measured with Image-Pro Plus software (Media Cybernetics, Silver Spring, Md.). Slides immunostained for insulin and glucagon were scanned with a Super Coolscan 8000 ED (Nikon, Tokyo, Japan). Representative pancreas sections for each mouse genotype were chosen for analysis. The Image-Pro Plus system was used to quantify the insulin- and glucagon-stained area and the total islet area to determine the percentage of islets stained for insulin and glucagon, respectively.
T-cell activation analyses.
Lymph nodes were isolated from RIP-GP/P14/Casp3+/– and RIP-GP/P14/Casp3–/– mice 4 days post-MLDS administration. Isolated PDLNs and pooled nondraining lymph nodes (NDLNs) (inguinal and axillary) were prepared into single-cell suspensions. Lymphocytes were stained with antibodies recognizing CD8
and CD69 (BD PharMingen). Detection of biotin-conjugated antibodies was performed using streptavidin-conjugated red 670 (GIBCO BRL). Live events were collected based on forward and side scatter profiles on a FACScan flow cytometer (Becton Dickinson) and analyzed using CELLQuest software (Becton Dickinson). The percentage of CD69-positive cells was determined by gating on the CD8+ cells.
Bone marrow chimeras.
Casp3+/+, Casp3+/–, and Casp3–/– donor mice were intravenously injected with CD4+ and CD8+ T-cell-depleting antibodies 3 days and 1 day prior to bone marrow transfer. Recipient mice received 9 Gy of 
irradiation just prior to bone marrow transfer. Bone marrow from donor mice was harvested from femurs and washed in HBSS. A total of 107 bone marrow cells in 200 µl of HBSS were transferred intravenously into sex-matched recipients. Eight weeks later, peripheral blood was stained for Thy 1.1 (donor) and Thy 1.2 (recipient) to determine bone marrow reconstitution.
Peptide gp33 immunization. Mice were immunized with an intravenous administration of 10 µg of the LCMV glycoprotein peptide p33 (KAVYNFATM) in HBSS (36).
Adoptive transfer of CFSE-labeled P14 TCR transgenic cells.
Single-cell suspensions of spleen cells from P14 TCR transgenic mice were prepared, and CD8+ T cells were purified using anti-CD8-coated beads (Miltenyi Biotech) and a magnetic activated cell sorter (Miltenyi Biotech). The purified CD8+ T cells were then labeled with 5- and 6-carboxyfluorescein diacetate succinimidyl ester (CFSE), washed in serum-free RPMI 1640 (GIBCO BRL), and resuspended in serum-free medium at 106 cells per 200 µl containing 5 µM CFSE (Molecular Probes). The purified CD8+ T cells were incubated for 10 min at 37°C in an atmosphere containing 5% CO2 and then washed in RPMI 1640 containing 10% fetal calf serum (Sigma-Aldrich). A total of 5 x 106 CD8-purified, CFSE-labeled cells were then transferred intravenously into nonirradiated RIP-GP/Casp3+/– and RIP-GP/Casp3–/– mice 48 h post-MLDS. Recipients were sacrificed, and PDLNs and NDLNs were harvested and stained with an antibody recognizing V2 (BD PharMingen) for fluorescence-activated cell sorter analysis. The lymphocytes from PDLNs and NDLNs were gated on V2+ lymphocytes, and CFSE profiles were determined.
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RESULTS |
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cells were confined to the rim of the islets, suggesting that the architecture of Casp3–/– islets was normal. In contrast, islet cells from Casp3+/– mice were architecturally distorted, containing significantly fewer insulin-positive cells and proportionally more glucagon-positive cells (Fig. 2A). Quantitative image analysis was used to assess the proportion of insulin and glucagons positive cells per islet in Casp3+/– and Casp3–/– pancreatic sections. The percentage of Casp3+/– and Casp3–/– islets stained for insulin were 58.48% ± 3.10% and 71.62% ± 2.35% (P = 0.002), respectively, and 58.63% ± 3.36% and 27.22% ± 3.65% (P = 0.001), respectively, for glucagon (Fig. 2B). Together, these findings further support that Casp3–/– islets are protected from destruction.
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staining, in keeping with MLDS leading to autoimmune destruction of the islets (Fig. 3A). Intriguingly, no lymphocyte infiltration was detected in islets of Casp3–/– mice at any of the time points examined (11, 14, 17, and 120 days post-MLDS). In Casp3+/– mice, the mean percentage of islets that displayed infiltration per pancreas was 45%. The severity of lymphocyte infiltration in the islets of Casp3+/– mice was scored by the following grade: grade 0, normal; grade I, periductal lymphocyte infiltrate; grade II, peri-insulitis; grade III, insulitis (lymphocytic infiltrate invading islets); and grade IV, severe insulitis (massive lymphocytic infiltrate with islet destruction) (8). Most of the islets showing infiltration were high grade, between grades III and IV (Fig. 3B).
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Casp3+/+ and Casp3+/– mice reconstituted with Casp3+/+, Casp3+/–, or Casp3–/– mouse bone marrow developed diabetes with high blood glucose levels 2 weeks post-MLDS and displayed lymphocyte infiltration in their islets in a time frame and severity similar to those in the original experiments performed with Casp3+/– mice (Fig. 3A; Table 1). In contrast, Casp3–/– mice reconstituted with either Casp3+/+ or Casp3–/– bone marrow were protected from diabetes development. These mice did not show evidence of any insulitis, much like Casp3–/– mice without reconstitution as described above.
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Caspase-3 is required for cross-priming of ß-cell-specific T cells in PDLNs in the MLDS diabetes model. The lack of islet infiltration in caspase-3 null islets led us to hypothesize that caspase-3-dependent islet apoptosis may be important for priming ß-cell-specific T cells. To this end, the Casp3 null mutation was introduced into the RIP-GP/P14 transgenic system to generate RIP-GP/P14/Casp3+/– and RIP-GP/P14/Casp3–/– mice, which enabled us to examine the effect of caspase-3 on islet apoptosis and subsequent cross-presentation of a nominal ß-cell-specific antigen to antigen-specific T cells.
MLDS in the RIP-GP/P14/Casp3+/– and RIP-GP/P14/Casp3–/– mice led to a rise in blood glucose levels similar to that in Casp3+/– mice and Casp3–/– mice, respectively. Analysis of PDLNs isolated from MLDS-treated RIP-GP/P14/Casp3+/– mice showed that 32% of CD8+ T cells were CD69hi, compared to 9% in NDLNs (Fig. 5A). In contrast, for CD8+ T cells isolated from RIP-GP/P14/Casp3–/– PDLNs, only 3% were CD69hi, similar to that of NDLNs (5%) (Fig. 5A). There was no difference in the percentage of CD69hi CD4+ T cells isolated from either RIP-GP/P14/Casp3+/– or RIP-GP/P14/Casp3–/– mice in both the PDLNs and NDLNs, since CD4+ T cells do not recognize the predominant transgenic antigen present in this mouse model.
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These results show that caspase-3 is required for cross-presentation of ß-cell antigen leading to antigen-specific T-cell activation in the local draining lymph nodes. Importantly, the lack of T-cell activation in the caspase-3-deficient mice is not due to an intrinsic defect of the T cells but rather to the absence of caspase-3-dependent islet apoptosis and cross-presentation in the PDLNs.
In vivo proliferation of antigen-specific T cells. To further characterize the proliferative response of ß-cell-specific T cells in the PDLNs subsequent to MLDS, in vivo, we adoptively transferred CFSE-labeled P14 transgenic T cells into RIP-GP/Casp3+/– and RIP-GP/Casp3–/– mice 2 days after the last injection of the MLDS regimen. CFSE labeling allows tracking of subsequent cellular divisions in vivo. Four days after the adoptive transfer, mice were sacrificed, and lymphocytes from PDLNs and NDLNs were isolated. Since all adoptively transferred lymphocytes that have caspase-3 present are P14 cells, these adoptive transfer experiments further supported the importance of caspase-3 in the islets, as opposed to the potential effect of caspase-3 in the lymphocytes. We observed that PDLNs of RIP-GP/Casp3+/– mice contained primarily CFSE-low cells, suggesting that the majority of adoptively transferred P14 T cells had undergone cell division (62%). In contrast, PDLNs of RIP-GP/Casp3–/– mice still maintained a CFSE-high peak, reflecting fewer cells that have undergone proliferation (45%) (Fig. 5B). Control NDLNs of both RIP-GP/Casp3+/– and RIP-GP/Casp3–/– mice contained CFSE-high cells, reflecting a lack of cell division, consistent with the absence of cross-presented antigens in these lymph nodes. In addition to the CFSE analysis, the same adoptively transferred CD8+P14 T cells were analyzed for their state of activation. In keeping with the higher levels of proliferation found in the PDLN of RIP-GP/Casp3+/– mice, 72% of the CD8+P14 T cells were CD69hi, in contrast to 29% of the adoptively transferred CD8+P14 T cells which were CD69hi in PDLNs of RIP-GP/Casp3–/– mice (Fig. 5C). These results show that caspase-3-dependent ß-cell apoptosis is required for activation of antigen-specific T cells and initiation of autoimmune diabetes.
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DISCUSSION |
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Although cross-presentation of autoantigens has generally been thought to induce T-cell tolerance (1, 19, 22), immunity has also been shown to occur in some experimental models (30, 31, 55). Activation of antigen-specific T cells is thought to be promoted through interactions with activated, mature APCs. We show that the caspase-3-dependent ß-cell apoptosis provides a source of antigen for cross-presentation and activation of ß-cell-specific T cells. Consistent with our results, recent studies demonstrate that apoptotic tumor cells can cross-prime host tumor-specific CD8+ T cells (30, 31, 55). In this way, the significant degree of cell death that occurs in proliferating tumors provides appropriate signals to APCs to promote immunity. In addition, using MLDS in NOD mice expressing transgenic TCRs, Zhang and colleagues showed the STZ-induced ß-cell damage can enhance priming of ß-cell-specific T cells (56). These studies further support that exaggerated ß-cell damage can induce activation of ß-cell-specific T cells. In addition, NOD mice exhibit a defect in the clearance of apoptotic ß cells (33). Therefore, the apoptotic cells may be a critical determinant contributing to the initiation of autoimmunity by having the capacity to instruct APCs to modulate immune responses so that the outcome is T-cell activation (2, 43).
When would such priming occur in type 1 diabetes? Theoretically, any destruction causing antigen to be shed and presented by nearby APCs may provide an opportunity for ß-cell-specific T cells to become activated in predisposed individuals. For example, viruses, toxins, and abnormal physiological tissue turnover have been shown to be possible initiators for diabetes development. Previous studies suggest that virus-mediated destruction of islets can trigger autoimmune diabetes (18). For example, coxsackie virus infection has been associated with autoimmune diabetes in both humans and animal models (20). In keeping with our results, it has been shown that virus-mediated bystander tissue damage can induce immune activation against ß cells, much like STZ-induced autoimmune destruction of the islets (5, 17, 18, 20).
Another opportunity where ß-cell-specific T cells may encounter excess antigen is during the neonatal period. ß cells were once thought to be terminally differentiated adynamic cells. However, mathematical models and enhanced technology have shown that ß cells are dynamic throughout the life of the organism (13, 47). In particular, during the neonatal period there is enhanced remodeling associated with ß-cell apoptosis. In autoimmune-prone mice such as NOD mice, ß-cell death is exaggerated compared to mice that are not prone to spontaneous diabetes (47). It has been speculated that this period may be an opportunity where ß-cell antigen is cross-presented to initiate autoimmune diabetes. Consistent with this hypothesis, studies of spontaneously occurring diabetes models such as BDC2.5 transgenic mice and 8.3 NOD mice have shown ß-cell-specific T-cell activation in the PDLNs occurring in a synchronous manner following the neonatal remodeling period (16, 50, 56). This event precedes insulitis and diabetes development in these animals. Recently, using the BDC2.5 transgenic model, Turley et al. (2003) showed that the enhanced wave of neonatal ß-cell death precipitates the activation of ß-cell-specific T cells in the PDLNs (48).
Together, our genetic models show the importance of caspase-3-dependent ß-cell apoptosis in initiating T-cell priming, a prerequisite event in the pathogenesis of type 1 diabetes. Understanding the initiating events in diabetes induction may lead to novel therapeutic interventions for the prevention of diabetes and may have implications in enhancing islet survival in islet transplantation. Therapeutically targeting caspase-3 activity in ß cells may be relevant for preventing the initiation of autoimmune diabetes.
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ACKNOWLEDGMENTS |
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We thank Kelvin So and Michelle Sleiman for their technical assistance. We also thank Laurent Sabbagh, Jacinth Abraham, Mark Cattral, Kinh Tung Nguyen, and Paul Doherty for critical review of the manuscript.
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FOOTNOTES |
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