help button home button Endocrine Society Endocrinology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Purchase Article
Right arrow View Shopping Cart
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Huo, J.
Right arrow Articles by Li, G.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Huo, J.
Right arrow Articles by Li, G.
Endocrinology Vol. 143, No. 5 1695-1704
Copyright © 2002 by The Endocrine Society

INTRACELLULAR SIGNAL SYSTEMS

Activation of Caspase-2 Mediates the Apoptosis Induced by GTP-Depletion in Insulin-Secreting (HIT-T15) Cells*

JianXin Huo, Rui-Hua Luo, Stewart A. Metz and GuoDong Li

Cardiovascular Research Institute (J.H., R.-H.L., G.L.), National University Medical Institutes, National University of Singapore, Singapore 117597; and Pacific Northwest Research Institute (S.A.M.), Seattle, Washington 98122

Address all correspondence and requests for reprints to: Guodong Li, National University Medical Institutes, MD11 02-01, 10 Medical Drive, Singapore 117597, Singapore. E-mail: . nmiligd{at}nus.edu.sg


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
This study investigated the possible involvement of a specific caspase(s) (a family of aspartate-specific cysteine proteases) in programmed cell death of islet ß-cells due to sustained GTP depletion. Treatment (up to 48 h) with 3 µg/ml mycophenolic acid (MPA), which specifically depletes intracellular guanine nucleotides, reduced cell-cycle progression from G1 phase into S and G2/M phases (as assessed by flow cytometry) and, subsequently, induced apoptosis of HIT-15 cells (transformed pancreatic ß-cells). The latter was accompanied by a marked increase of caspase-2 activity (+343%) and moderate activation of caspase-9 (+150%) and caspase-3 (+145%). Importantly, only caspase-2 activation preceded induction of apoptosis. There was no change in activity of caspase-1, -4, -5, -6, and -8. Release of the mitochondrial protein cytochrome c into cytosol was also observed at a late stage. Cotreatment of cells with a permeable pan-caspase inhibitor (Z-VAD-FMK) blocked GTP depletion-induced cell death in a dose-dependent manner. A specific caspase-2 inhibitor (Z-VDVAD-FMK), but not a caspase-3 inhibitor (DEVD-CHO), was also capable of restoring cell viability. Interestingly, activation of caspase-2 leads to caspase-3 activation because the caspase-2 inhibitor abrogated caspase-3 activity. Our results indicate that, while activation of multiple caspases are involved in the execution phase of GTP depletion-induced apoptosis, caspase-2 appears to play the major role in the initiation of this program. This study revealed a novel, caspase-2 mediated form of apoptosis that may be consequent to impaired mitogenesis.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
INOSINE 5'-MONOPHOSPHATE dehydrogenase (IMPDH, EC 1.1.1.205), which catalyzes the conversion of IMP into GMP, is a key enzyme in the biosynthesis of GTP, deoxy-GTP, and other guanine nucleotides (GNs) (1, 2). These nucleotides are not only essential for the construction of DNA and RNA, but also important for the regulation of many cellular signaling pathways such as G proteins and gene expression (1, 2, 3).

Specific inhibitors of IMPDH have been used to treat cancers and psoriasis and more recently to prevent transplantation rejection (4). These inhibitors interfere with cell proliferation and differentiation (1, 3) and may induce apoptosis of several types of cells (2, 5, 6). Mycophenolic acid (MPA), among the IMPDH inhibitors, is widely used as an immunosuppressive drug (4). MPA depletes cellular GNs and this effect can be reversed entirely by provision of guanine or guanosine but not adenine or adenosine (5, 7). We and others have demonstrated that short-term depletion of GTP achieved using MPA altered membrane ion fluxes and inhibited insulin secretion in islet ß-cells (1, 5, 7, 8, 9, 10, 11); the latter effect was probably due in part to the interference with PLC activation, and interference with the activation of specific G proteins (12). Furthermore, we also found that depletion of GTP by MPA or mizoribine (a structurally distinct inhibitor of IMPDH) blocked DNA synthesis whereas longer GN depletion (>24 h) induced death of these cells (both islets and ß-cell lines) with characteristics of apoptosis as assessed morphologically by electron microscopic observations and biochemically by DNA fragmentation (laddering) (5). The apoptosis by MPA was prevented by guanosine (which restores intracellular levels of guanine nucleotides), but not by adenosine (5). The underlying mechanism(s) whereby GN depletion inhibits mitogenesis and induces apoptosis are unclear, although it might be due to the combined effects of inhibition of primer RNA biosynthesis (13), reduction in availability of guanine deoxyribonucleotide, and to the interference with the function of some small G proteins (5, 12). In addition, IL-1ß caused a reduction of GNs (14) and apoptosis (15) in islet ß-cells.

Caspases, a family of aspartate-specific cysteine proteases, plays a critical role as either initiators or effectors in the execution of apoptotic cell death (16, 17). At least 14 members have been identified. Based on the size of the prodomain, caspases can be classed into two groups possessing different functions; large prodomain caspases (initiator) receiving apoptotic signals function as upstream signal transducers (e.g. caspase-1, -2, -8, -9), whereas short prodomain caspases (effector or executor) are activated by the formers and operate as downstream amplifiers that cleave death substrates (e.g. caspase-3, -6, and -7) (16, 17). Different subcellular compartment, such as plasma membrane, mitochondrion, endoplasmic reticulum and nucleus are implicated in apoptotic execution under various stimuli (17, 18, 19, 20). Two major pathways of caspase-mediated apoptosis have been well established (16, 17). One involves the membrane death receptor [such as Fas antigen (CD95 or APO-1) and TNF receptor (TNFR)] operated apoptotic death with caspase-8 acting as the upstream initiator in the activation of caspases (16, 17). Another cascade (initiated by caspase-9) of caspase activation is involved in the apoptosis due to mitochondria damage (releasing cytochrome c) such as that caused by radiation and toxic drugs (18, 21, 22). More recently it has been reported that caspase-12 mediated the apoptosis induced by endoplasmic reticulum stress by the agents that release calcium from intracellular stores (19), whereas caspase-11 is initially activated in the inflammatory LPS-induced cell death (20).

It is not entirely clear how the caspases with a large prodomain are activated, but their binding to adapter molecules, which results in their oligomerization and a series of autocatalysis and heteroproteolysis (16, 17), is a critical step. Activation of different caspases by various apoptotic signals may be mediated by a specific adapter, e.g. FADD (interacting with Fas and TNFR) for caspase-8, Apaf (binding to cytochrome c) for caspase-9, CARDIAK (interacting with the TNFR-associated factors) for caspase-1 and RAIDD [receptor-interacting protein (RIP)-associated ICH-1/CED-3-homologous protein with a death domain] for caspase-2 (16, 17, 23, 24). These adapters possess distinct domains responsible for their interactions with the upstream apoptotic molecules and with the prodomain of initiator caspases (16, 17), and thus may well be the targets to be regulated by other cellular signaling pathways.

It was unclear whether any caspase is implicated in GN depletion-induced apoptosis in insulin-secreting cells. The present study was undertaken to address this question by measuring the activities of caspase-1 through caspase-10 and by use of selective inhibitors of their activity. Our findings indicate that GN depletion activates multiple caspases and causes cytochrome c release in transformed pancreatic ß-cells (HIT-T15). Furthermore, caspase-2 (also called ICH-1 or Nedd2) activity was increased before other caspases and also to a larger extent. A specific caspase-2 inhibitor was able to prevent the cells from apoptosis due to MPA treatment, suggesting that caspase-2 plays a critical role in mediating this type of cell death.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Most of reagents and chemicals used in this study were purchased from the companies listed below in detail. Roswell Park Memorial Institute (RPMI) 1640 medium, MPA, phenazine methosulfate (PMS), Hoechst 33258, DNA standard (sodium salt, from calf thymus), guanosine, adenosine, EGTA, EDTA, HEPES, 3-(3-cholamidopropyl)dimethylammonio-1-propane sulfonate, dithiothreitol (DTT), phenylmethylsulfonylfluride (PMSF), pepstatin A, aprotinin, and monoclonal antibody of ß-tubulin were from Sigma (St. Louis, MO); 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) were from Promega Corp. (Madison, WI); FCS was from Life Technologies, Inc. (Rockville, MD); BSA and leupeptin were from Roche Molecular Biochemicals (Mannheim, Germany); nitrocellulose membrane (0.45 µm), polyvinylidine difluoride (PVDF) membrane, caspase substrates and some caspase inhibitors were supplied by Bio-Rad Laboratories, Inc. (Hercules, CA); caspase-3 inhibitor (DEVD-CHO) was from Calbiochem (La Jolla, CA); purified anti-cytochrome c antibody (7H8.2C12, mouse IgG2b) was from PharMingen (San Diego, CA); and RAIDD antibody, antirabbit IgG-HRP and antimouse IgG-HRP was from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA).

Cell culture
Insulin-secreting HIT-T15 cells (passage 76–83) were maintained on Falcon dishes in RPMI 1640 supplemented with 10% decomplemented FCS (vol/vol), 100 IU penicillin/ml and 100 µg streptomycin/ml. Cells were seeded at a density of 1.0 x 106/ml in culture dishes or multiwell plates, and the medium was changed every 48 h. MPA and other test agents were added to cells when they reached more than 80% confluence.

Flow cytometric analysis for DNA fragmentation and cell cycle
Prolonged treatment of HIT cells with MPA inhibited DNA synthesis and caused DNA fragmentation (DNA laddering), leading to apoptosis (5). In the present study, DNA fragmentation and cell cycle were determined by propidium iodide (PI) staining and flow cytometry as described by Nicoletti et al. (25). This assay measures fragmented nuclei and thus detects the subdiploidy apoptosis (25). Briefly, HIT cells were cultured in six-well plates with 3 µg/ml MPA for 8–48 h. After one gentle rinse by PBS, the cells were collected by trypsinization and centrifugation. Then the cell pellets were gently resuspended thoroughly in 0.5 ml PBS followed by 2-h fixation (at 4 C) in 4.5 ml 70% ethanol. Cells suspended in 70% ethanol were stored at -20 C if PI staining was not performed immediately. For PI staining, the ethanol suspended cells were centrifuged for 5 min at 200 x g to remove cell debris. The cell pellets were resuspended in 5 ml PBS and centrifuged for 5 min at 200 x g. Afterward, the cells pellets were incubated in 1 ml PI/Triton X-100 staining solution (containing 20 µg/ml PI, 0.1% Triton X-100 and 0.2 mg/ml RNase A in PBS) for 30 min at room temperature. Flow cytometric analysis was carried out using a fluorescence-activated-cells-sorter (EPICS Elite ESP, Beckman Coulter, Inc., Hialeah, FL). An excitation source of 488 nm was achieved using a 15 mW air-cooled argon-ion laser. Fluorescence emission was collected through a 610 nm band pass filter for PI. PI fluorescence data were collected on a linear scale. Ten thousand cells were evaluated for each sample. The flow rate was kept at 200–300 cells/sec. The data were processed and analyzed using WinMDI software (Scripps Institute, La Jolla, CA).

The number of floating and rinsed cells not included in the flow cytometric analysis and other 2 apoptotic assays below (MTS test and DNA measurements) was various depending on the time periods of MPA treatments (5.7%, 7.3%, 8.3%, 9.3%, 12.8%, 19%, and 33.9% at 0, 8, 16, 24, 32, 40, and 48 h, respectively; n = 3). Almost all of these cells (>94%) were dead cells or apoptotic cells as assessed by PI staining and flow cytometry.

Induction of apoptosis by GTP depletion
MPA-induced apoptosis of HIT cells was also evaluated by MTS test and DNA content (5). Our previous studies have demonstrated that these assessments are good markers to monitor GN depletion-induced apoptotic cell death, which had been formally established by more specific means such as electron microscopic examination and measurements of DNA laddering (5). MPA stock solution (2 mg/ml) was prepared in ethanol and added into cell culture medium to a final concentration of 3 µg/ml up to 48 h.

Cell viability was monitored by MTS test, which detects reduction of a tetrazolium salt into a colored and water-soluble formazan in intact cells (5, 26). Dehydrogenases in metabolically active cells accomplish the conversion of MTS into the formazan using PMS as the electron-coupling reagent (26). We have confirmed that the intensity of absorbance from formazan product is proportional to the number of control living cells added into the wells. HIT cells cultured on 96-well plates were treated with test agents for various periods of time. After one gentle wash to discard culture medium and any floating cells, a mixture of MTS and PMS in fresh RPMI 1640 was added and the plates were incubated for 30 min. The absorbance of the formazan at 490 nm in these wells was measured in 96-well plates using a plate reader.

Cell death was also monitored by determination of DNA contents of cells seeded in microtiter plates with a fluorescent DNA-binding dye, Hoechst 33258 (27). After washes to remove growth medium and any floating cells, the attached cells were lyzed by sonication in distilled water. Aliquots of the cell lysates were reacted with the dye in 96-well plates. The fluorescence was measured by a fluorescent plate reader (SPECTRAmax GEMINI, from Molecular Devices, Sunnyvale, CA) at excitation and emission wavelengths of 356 nm and 448 nm, respectively. DNA contents in the wells were calculated by comparison with the standard curves.

The raw values from MTS test and DNA measurements were affected by many factors such as cell numbers, probe concentrations, and equipment function. Indeed, there is variance in the cell numbers used between experiments, as well as between time points for controls due to proliferation and for treated cells due to cell loss. As a consequence, it is difficult to compare the raw data between individual experiments (including the different time points of treatment within one experiment) and to pool the data for statistical analyses. Therefore, we always measured the samples of both treated cells and control cells at any time point (they were seeded at the same time in triplicate each) and calculated the mean values for each condition. Afterwards, the ratio of mean values of treated samples was expressed as percentage of control. In this way, the change due to a given treatment (dose or time) could be assessed more precisely. Because the floating and rinsed cells were excluded, the percentages of apoptotic cells, expressed as percentage of total cells originally plated, would be overestimated by the measurements of MTS test and DNA content in particular at the late time points of MPA treatment in this study. However, any such overestimation would be greatly minimized if the detached cells were to be included in the analyses because almost all of them were apoptotic.

Caspase activity measurement
Measurements of caspase activity are simplified by using fluorogenic synthetic oligopeptide substrates (28). These amino acid sequences are identical or similar to those found in full-length protein substrates, and where differences occur, the optimized synthetic substrates appear to be cleaved as efficiently or better than the full-length proteins. Substrates are synthesized by modifying the caspase cleavage site (C-terminal aspartic acid) with 7-amino-4-trifluoromethyl coumarin (AFC). When liberated from the peptide, AFC produces an optical change resulting in emissions of fluorescence.

HIT cells were cultured in six-well plates. For each independent experiment, duplicates of both control and treated wells were applied for any time point of treatment. Activity of individual caspase in samples from control and treated cells at any time point of treatment was measured and compared. The results were expressed as percentage of control. After decanting medium and rinsing cells with cold PBS, 100 µl lysis buffer [10 mM HEPES, pH 7.4; 2 mM EDTA; 0.1% 3-(3-cholamidopropyl)dimethylammonio-1-propane sulfonate; 5 mM DTT; 350 µg/ml PMSF; 10 µg/ml pepstatin A; 10 µg/ml aprotinin; and 20 µg/ml leupeptin] was added to each well and cells were scraped with a cell lifter and transferred to clean tubes. Afterwards the tubes were frozen and thawed for 4 cycles to lyze the cells by transferring from a -20 C freezer to a 37 C water bath. Lysates were centrifuged (Eppendorf, Hamburg, Germany) at 4 C for 30 min at full speed. The supernatants were transferred to clean tubes and kept on ice if the assay was to be performed within 1 h or, otherwise, stored at -70 C. The caspase activity assay was performed according to a protocol provided by the supplier. In brief, the reaction components containing caspase substrates were thoroughly mixed with the blank or samples in a 96-well microtiter plate. After incubation at 37 C for 3 h, the plates were scanned by a fluorescent plate reader to measure the AFC fluorescence at excitation and emission wavelengths of 395 nm and 525 nm, respectively. The rate of AFC fluorescence signal change was proportional to the enzyme activity. All values were corrected by blank readings and the enzyme activity was normalized to a fixed protein concentration.

Protein contents were measured by using Bradford (Bio-Rad Laboratories, Inc.) assay. The color change of Coomassie brilliant blue G-250 dye (shifted from 465 nm to 595 nm when binding to protein) was detected by a plate reader, and compared with a standard curve.

Isolation of fractions enriched in mitochondria and cytosol
All following steps were performed at 4 C as described previously (29). After washing with cold PBS, HIT cells were detached by scraping and suspended in homogenization buffer containing 20 mM Tris-HCl (pH 7.4), 0.5 mM EDTA, 0.5 mM EGTA, 250 mM sucrose, 1 mM DTT, and the following protease inhibitors (in µg/ml): 10 leupeptin, 4 aprotinin, 2 pepstatin, and 100 PMSF. The cells were first subjected to cavitation in a nitrogen bomb (from Parr) under 10 bars for 30 min and then the achieved cell suspensions were further homogenized by using a Dunce tissue grinder (from Kontes) with the tight pestle. The homogenates were centrifuged at 900 x g for 10 min to remove unbroken cells and nuclei. The supernatants were then centrifuged at 5500 x g for 20 min. The pellets (rich in mitochondria) were resuspended in homogenization buffer and sonicated for 3 times (5 sec each). Cytosol was achieved by centrifuging the supernatant at 100,000 x g for 60 min. Protein concentrations were determined and samples were aliquoted and kept at -70 C.

Immunoprecipitation of RAIDD and caspase-2
HIT cells cultured in six-well plates were treated by MPA (3 µg/ml) for indicated time. All procedures below were performed at 4 C. After washing in cold PBS buffer, the cells were lysed in cold immunoprecipitation buffer (1% Triton X-100, 150 mM NaCl, 1 mM EDTA, 10 mM Tris pH 7.4, 1 mM EGTA pH 8.0, 0.2 mM sodium ortho-vanadate, 0.2 mM PMSF, 0.5% NP-40) for 30 min. Cell homogenates scraped from the plates were then passed 10 times through a 27-gauge needle to disperse any large aggregates, and centrifuged (16,000 x g) for 15 min to get whole cell lysates. For initiation of immunoprecipitation, 2 µg (10 µl) anti-RAIDD antibody was added into each Eppendorf tube containing 1 ml whole cell lysate (total 100 µg protein) and incubated in constant agitation for 1 h. Afterwards, 10 µl of 50% agarose-immobilized protein A was added to the mixture and incubated in agitation for 30 min. After centrifuging for 4 min (16,000 x g), the pellets were rinsed by cold immunoprecipitation buffer for three times and resuspended in 30 µl double-concentrated electrophoresis sample buffer (125 mM Tris, pH 6.8; 4% SDS; 10% glycerol; 0.006% bromophenol blue; 2% ß-mercaptoethanol). The samples were boiled and subject to electrophoresis and Western blotting for detection of RAIDD and caspase-2 as described below.

Western blotting of caspase-2, RAIDD and cytochrome c
For detection of caspase-2 in cell homogenates, aliquots of samples (20 µg protein each) were boiled in a loading buffer (100 mM Tris-HCl, pH 6.8; 200 mM DTT; 20% glycerol; 4% SDS; and 0.2% bromophenol blue) for 5 min and centrifuged at 9000 x g for 5 min. These samples or aforementioned immunoprecipitation samples were subjected to a 10% SDS-PAGE and proteins were then electrotransferred to PVDF membranes. The membranes were blocked in Tris-buffered saline buffer with Tween-20 (TBST) (20 mM Tris-HCl, pH 7.5; 150 mM NaCl; and 0.2% Tween-20) containing 5% dried nonfat milk for 1 h at room temperature. Subsequently, membranes were blotted with the antibodies against caspase-2 (1:250) or RAIDD (1:1000) for 1 h at room temperature. After three washes in TBST (5 min each), the membranes were incubated with horseradish peroxidase conjugated secondary antibodies (antirabbit IgG or antimouse IgG as appropriate) for 60 min. The membranes were washed again (3 x 15 min) in TBST and chemiluminescent signals were detected using an ECL system (from Pierce Chemical Co., Rockford, IL) according to the manufacturer’s instructions.

For detection of cytochrome c, aliquots of cytosol (40 µg protein) and mitochondrial lysate (20 µg protein) were subjected to 15% SDS-PAGE, and proteins then transferred to PVDF membranes. The membranes were blocked with dried nonfat milk and subsequently blotted with cytochrome c antibody with a 1:1000 dilution. Afterwards, the immunodetection was performed as described above. ß-tubulin in the samples was also detected as a loading control. The immunospots on the membranes were scanned, analyzed and semiquantified by densitometry.

Statistical analysis
Data were expressed as mean ± SE and analyzed by ANOVA and t test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Induction of cell death by GN depletion with MPA treatment
Sustained (24–48 h) GN depletion using either MPA (or mizoribine, a structurally dissimilar inhibitor of GTP synthesis) induced death of insulin-secreting cells (HIT, INS-1, and rat islets) with characteristics of apoptosis as described in an earlier study (5), including typical apoptotic morphological changes, chromosome condensation/margination, and DNA fragmentation (DNA laddering). In the current study, HIT cells were treated with 3 µg/ml MPA for up to 48 h. Flow cytometric analysis of PI-stained cells indicated that MPA induced cell arrest in G1 (diploid peak) phase (Fig. 1AGo and Table 1Go). The cells at S and G2/M phases were significantly reduced after MPA treatment at 16 h and beyond. Meanwhile, MPA increased number of cells with fragmented nuclei; the DNA content of these cells was lower than that at G1 phase (in subG1 fraction) (Fig. 1AGo and Table 1Go). These subdiploidy cells with fragmented nuclei thus were undergoing apoptosis (25). This apoptotic effect of MPA was time dependent and occurred significantly after 24-h treatment with MPA (Fig. 1BGo), confirming our previous findings obtained from using different methods (5).



View larger version (24K):
[in this window]
[in a new window]
  		Figure 1. Time-course of cell death induced by MPA treatment. HIT cells seeded on multiwell plates were treated by MPA (3 µg/ml) in culture media for up to 48 h. A, Cells were stained with PI and subject to flow cytometric analysis of MPA effects on cell growth/death. The graph is a typical representative of three independent experiments. The cells whose DNA content was lower than G1 phase contained fragmented nuclei and were undergoing apoptosis. B, Percentages of MPA-induced subdiploidy apoptotic cells analyzed by flow cytometry. Values are mean ± SE from three independent experiments. C, Cell viability was evaluated both by formazan production in MTS test and by determination of DNA content in the wells under different conditions. Corrections were made for the influence of a slight increase (15–20% during 48 h) in the cell number of control wells in all data (both MTS test and DNA content) at indicated time points. Values are mean ± SE from seven independent experiments. *, P < 0.05; and **, P < 0.01 vs. zero time point.

 

View this table:
[in this window]
[in a new window]
  		Table 1. Effects of MPA treatment on cell cycle of HIT cells

 
Cell viability was also assessed by MTS test and DNA content [changes in which were correlated with other typical markers of apoptosis induced by GN depletion (5)] every 8 h during treatment. MPA led to a dose-dependent decreases both of formazan production (MTS test) and DNA content (data not shown). The time-courses of alterations of these two parameters following MPA treatment of HIT cells were generally parallel, although the decease in DNA occurred slightly before the drop in MTS signal (Fig. 1CGo), possibly because GTP reduction directly inhibits DNA synthesis (3, 5). Significant reductions (-15%, P < 0.01) of DNA were observed at 24-h treatment, a time point when clear apoptotic alterations (chromosome condensation/fragmentation, DNA laddering) occurred (5). Both formazan production and DNA content were deceased by about 60% after 48-h exposure to the GN-depleting agent. The time course of changes in MTS test and DNA content after MPA treatment was correlated to the percentages of apoptotic cells (Fig. 1BGo) as assessed by flow cytometry under the same conditions.

Activation of caspases and release of cytochrome c by MPA treatment
The activity of caspase-1 through caspase-10 in HIT cells after MPA treatment was measured by using their respective, specific substrates (30) (Fig. 2Go). Six such synthetic oligopeptide substrates were used: Ac-WEHD-AFC for caspase-1, -4, and -5; Ac-VDVAD-AFC for caspase-2; Ac-DEVD-AFC for caspase-3, -7 and -10; Ac-VEID-AFC for caspase-6; Ac-LETD-AFC for caspase-8; and Ac-LEHD-AFC for caspase-9. Cell homogenates were prepared after MPA treatment (3 µg/ml) at various time intervals. Activity of several caspases was increased subsequent to sustained GN depletion. Caspase-2 was clearly activated at 16 h, earlier than other caspases and before the onset of apoptosis, and maximal activity (+343%) was seen at 32 h, before the maximal levels of apoptosis achieved. In another study measuring caspase-2 activity in shorter intervals (every 2 h) during MPA treatment (0–12 h), the protease activity was significantly increased by 105% (n = 3; P < 0.001) at 12 h. Caspase-3 (and perhaps also -7 and -10) and caspase-9 were only activated at 24 h MPA treatment and their activity was increased maximally by 145% and 150%, respectively at 32 h (Fig. 2Go).



View larger version (41K):
[in this window]
[in a new window]
  		Figure 2. Activity of caspases in HIT cells during MPA treatment. Cells seeded in six-well plates were treated with 3 µg/ml MPA in culture media for 8–48 h. Cell homogenates were prepared as described in details in the Methods section. Caspase activity was determined by measurement of the cleavage of the fluorogenic, specific substrates. Values are mean ± SE from at least four independent experiments. *, P < 0.05; and **, P < 0.01 vs. control.

 
Caspase-2 activation was also confirmed by detection of its cleavage (31) by Western blotting (Fig. 3Go). Inactivated caspase-2 in control HIT cells displayed a band of 48 kDa on the SDS-polyacrylamide gel. This caspase isoform was dominantly present in cytosol and faintly detected in mitochondria (Fig. 3AGo). The 48-kDa inactive caspase-2 in cytosol was slightly elevated with the time of culture both in control and MPA-treated cells. After MPA treatment, a band of 35 kDa in cytosol appeared at 16 h and reached maximal at 32 h (Fig. 3BGo), a time course in close parallel with the increase of its activity assessed by substrate cleavage (see Fig. 2Go). No change of caspase-2 in mitochondria was found. In addition, we also investigated the distribution of RAIDD and its interaction with caspase-2 by immunoprecipitation. RAIDD in HIT cells was mainly located in cytosol (data not shown). Using a Protein A-conjugated antibody, caspase-2 could be pulled down together with RAIDD (Fig. 3CGo). Moreover, MPA treatment slightly reduced the amount (a decease of about 20%; P < 0.05; n = 4) of caspase-2 (inactive form) coprecipitated with RAIDD.



View larger version (55K):
[in this window]
[in a new window]
  		Figure 3. Cleavage of caspase-2 by MPA treatment and interaction of RAIDD with caspase-2. A and B, HIT cells were treated with 3 µg/ml MPA for 8–48 h. Cell lysates were subjected to 10% SDS-PAGE and caspase-2 cleavage was detected by immunoblotting using an antibody reacting with both the enzyme (48 kDa) and its large cleavage product (35 kDa). No difference in the ß-tubulin content (loading control) was found between samples. The results are the representative of five independent experiments. C, HIT cell homogenates (100 µg protein each) were incubated with anti-RAIDD antibody followed by addition of agarose-immobilized protein A. After washing and centrifuging, the immunoprecipitation pellets were resuspended in electrophoresis buffer and boiled. The samples were subject to electrophoresis and Western blotting for detection of RAIDD and caspase-2. Data are the representative of four independent experiments.

 
Caspase-9 can be activated by cytochrome c (18, 21); therefore, it is possible that GN depletion by MPA treatment is capable of causing the release of cytochrome c from mitochondria into cytosol. This was shown to be true because cytochrome c in cytosol was increased while that in the mitochondria fraction was decreased in MPA-treated HIT cells (Fig. 4Go). By analyzing the data from Western blotting with densitometry, the cytochrome c content in cytosol was elevated by 57% (P < 0.05) at 24 h, and 69% and 174% (both P < 0.01) after 32 and 48 h in corresponding to 18% (P < 0.05) and 61% (P < 0.01) reduction of its content in mitochondria. No significant release was observed at 8- and 16-h treatment. Thus, these changes occurred after increase in active caspase-2 but before apoptotic changes (Fig. 1Go).



View larger version (58K):
[in this window]
[in a new window]
  		Figure 4. Release of cytochrome c from mitochondria into cytosol during MPA treatment. HIT cells were treated with 3 µg/ml MPA for the indicated periods. Mitochondria fraction and cytosol were obtained by subcellular fractionation. They were subjected to 15% SDS-PAGE and cytochrome c was determined by immunoblotting. There was no difference in the ß-tubulin content between samples. The results are the representative of four independent experiments.

 
There was no apparent increase in the activity of caspase-1, -4, -5, -6, and -8 during MPA treatment (Fig. 2Go).

Blockade of MPA-induced apoptosis by caspase inhibitors
Although GN depletion activates several caspases, it was unclear whether the latter mediate entirely the apoptosis seen under the same conditions, in view of the reports that apoptosis could occur without involvement of caspases (32, 33, 34). Therefore, caspase inhibitors were used to address this question. All cell-permeable caspase inhibitors used below were nontoxic and had no effects on both MTS test and DNA content in HIT cells by themselves under the concentrations applied.

A pan-caspase inhibitor, Z-VAD-FMK that suppresses a broad of the enzymes in the group (35), was first used to examine its effects on GN depletion-induced cell death at various concentrations and at different time of treatment (Fig. 5Go). The decreases in both formazan production (MTS test) and DNA content due to 48-h MPA treatment were slightly prevented by 10 µM and were completely restored to normal by 100 µM of the inhibitor (Fig. 5AGo). When Z-VAD-FMK (100 µM) was added together with MPA to the cells for various periods, the changes both in formazan production (Fig. 5BGo) and DNA content (Fig. 5CGo) by GN depletion was completely blocked at all examined time points.



View larger version (18K):
[in this window]
[in a new window]
  		Figure 5. Blockade of GN depletion-induced cell death by a pan-caspase inhibitor. HIT cells in multiwell plates were treated with MPA (3 µg/ml) in the presence or absence of a pan-caspase inhibitor (Z-VAD-FMK) in culture media for various periods. Cell death was assessed by both MTS test and the determination of DNA content in the wells. Values are means ± SE of at least three independent experiments. A, Dose-dependent prevention of GN depletion-induced apoptosis by Z-VAD-FMK at 48 h of MPA treatment; B and C, time-course of the protective effects of the pan-caspase inhibitor (100 µM) on cell viability monitored by MTS test and the DNA content. Values are mean ± SE from four to five independent experiments. *, P < 0.05, and **, P < 0.01 vs. MPA treatment alone.

 
It is possible that the protective effect of caspase inhibitors on GN depletion-induced apoptosis was mediated via an action on restoration of cellular levels of GTP. The results from two series of experiments, however, are against this notion. First, the pan-caspase inhibitor (Z-VAD-FMK, 100 µM) was unable to reverse the inhibition of stimulated insulin secretion in HIT cells treated with MPA for 6 h (data not shown), a time when levels of GTP are reduced by more than 80% (8, 9). By contrast, addition of guanosine was able to restore the GN levels and also reverse the inhibitory effect of MPA on insulin secretion (8, 9). Second, GN depletion by MPA activated tissue transglutaminase in HIT cells in a dose- and time-dependent manner (our manuscript in preparation). This transglutaminase is an enzyme whose activity is suppressed by micromolar levels of GTP and may be involved in apoptotic process, in particular the morphological changes. We found that the pan-caspase inhibitor was able to reverse MPA-induced apoptosis but failed to block the MPA-enhanced tissue transglutaminase activity (data not shown). However, cotreatment with guanosine could block both MPA-induced apoptosis and activation of tissue transglutaminase. Therefore, our results clearly indicate that reversal of GN depletion-induced HIT cell death by caspase inhibitors is not trivially due to prevention of the GTP reduction.

To define whether a particular caspase(s) plays the critical role in GN depletion-induced apoptosis, specific caspase inhibitors were also used. The caspase-2 inhibitor, Z-VDVAD-FMK (30, 31, 36), protected the cells from death due to MPA treatment (Fig. 6Go). At 100 µM, the caspase-2 inhibitor restored the formazan production almost completely to control values, similar to the action by the pan-caspase inhibitor described above. In addition, the reduction of DNA content could be restored to more than 80% of control by the caspase-2 inhibitor (this residual reduction, which was still significantly lower than control, may be due to direct inhibition of DNA synthesis by GTP depletion; see above). On the contrary, the caspase-3 inhibitor, DEVD-CHO (100 µM) that blocks apoptosis in other occasions (37, 38), was only able to partially restore DNA content by 35% and did not restore the cell viability at all during 48-h MPA treatment (Fig. 7Go). The caspase-3 inhibitor also failed to reverse the morphological changes due to MPA (data not shown). These findings by using caspase inhibitors suggest that, although activated caspase-3 is partially responsible for only the DNA degradation, caspase-2 is a major mediator in the induction of apoptotic death of insulin-secreting cells induced by GN depletion.



View larger version (18K):
[in this window]
[in a new window]
  		Figure 6. Blockade of GN depletion-induced cell death by a specific caspase-2 inhibitor. HIT cells grown in multiwell plates were treated with MPA (3 µg/ml) in the presence or absence of a specific caspase-2 inhibitor (31 ) (Z-VDVAD-FMK) for various periods. Apoptosis was monitored by both MTS test and the determination of DNA content. Results are means ± SE of at least four independent experiments. A, Dose-dependent blockage of GN depletion-induced apoptosis by Z-VDVAD-FMK at 48 h of MPA treatment; B and C, time-course of the effects of the caspase-2 inhibitor (100 µM) on cell viability assessed by MTS test and the DNA content. Values are mean ± SE from three to four independent experiments. *, P < 0.05, and **, P < 0.01 vs. MPA treatment alone.

 


View larger version (32K):
[in this window]
[in a new window]
  		Figure 7. Partial prevention by a caspase-3 inhibitor of reduction of DNA content induced by GN depletion; comparison to absence of effect on cell viability (MTS test). HIT cells were treated with 3 µg/ml MPA and a caspase-3 inhibitor (DEVD-CHO, 100 µM) for 48 h. Results are means ± SE of three independent experiments. **, P < 0.01 vs. MPA treatment alone.

 
Activation of caspase-2 upstream of caspase-3 and caspase-9
To further address the critical role of caspase-2 in GN depletion-induced apoptosis, the activity of caspase-2, caspase-3, and caspase-9 in the cell lysates after MPA treatment in the presence of the caspase-2 inhibitor was examined. The results revealed that 100 µM Z-VDVAD-FMK abolished entirely the activation of all three caspases in HIT cells treated with 3 µg/ml MPA for 32 h (Fig. 8Go). This indicates that activation of caspase-2 occurs before the activation of caspase-3 and caspase-9 in the scenario of the cell death investigated.



View larger version (33K):
[in this window]
[in a new window]
  		Figure 8. Blockade of activation of caspase-2, -3, and -9 by a caspase-2 inhibitor. HIT cells were treated with 3 µg/ml MPA and 100 µM caspase inhibitor (Z-VDVAD-FMK) in culture media for 32 h. Activity of caspase-2, caspase-3 and caspase-9 in cell lysates was measured by using their specific, fluorogenic substrates. Values are means ± SE from three independent experiments. *, P < 0.05; and **, P < 0.01 vs. MPA treatment alone.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
This is the first study reporting that multiple caspases (including at least caspase-2, -3, and -9) were activated and the release of mitochondrial cytochrome c occurred during apoptosis of ß-cells induced by GTP deletion with MPA. Importantly, caspase-2 like protease(s) was activated earlier and more dramatically than other caspases. Furthermore, GTP depletion-induced apoptosis and caspase activation were blocked by a pan-caspase inhibitor and, significantly, protected by a specific caspase-2 inhibitor as well, pointing a major role of the latter in the initiation of apoptotic death of insulin-secreting HIT cells. Thus, our study revealed a novel, caspase-2-mediated form of apoptosis that may be consequent to impaired mitogenesis as demonstrated by flow cytometry in the current study and by thymidine/5-bromo-2'-deoxyuridine incorporation previously (3, 5).

Our data strongly suggest that HIT cell apoptosis due to GN depletion was mediated by activation of at least two initiators (caspase-2 and caspase-9) and at least one effector (caspase-3). In addition, caspase-3 activation occurred later than caspase-2 and a specific caspase-2 inhibitor also blocked caspase-3 activation, indicating that the latter is activated subsequent to the former. Caspase-3 is a downstream effector directly killing cells in some circumstances (37, 38), but there is increasing evidence that caspase-3 inhibition fails to block apoptosis in other systems (39, 40). In our study, a caspase-3 inhibitor failed to prevent the ß-cell from death induced by GTP-depletion; however, it exerted some protective effect on DNA reduction which may suggest some role in the latter process. This also suggests that other effector caspases play important roles, along with caspase-3, in the final execution of cell death.

We found that caspase-2 in HIT cells was mainly localized in cytosol and only small portion was detected in mitochondria-rich fraction. It has been proposed that caspase-2 and -9 zymogens and cytochrome c essentially localized in mitochondria may be released and become activated, eventually triggering of apoptosis (18, 21, 22, 37, 41). Thus, the integrity of mitochondria may be critical for subcellular redistribution and activation of these caspases. In this study, we could not find detectable change of caspase-2 content in mitochondria after GN depletion, suggesting that the activated caspase-2 is not mainly derived from mitochondria. Although we did observe the leakage of cytochrome c from mitochondria and modest activation of caspase-9 (indicating a damage of the organelles) occurred in GN-depleted ß-cells, these events took place later than the increase of caspase-2 activity. In addition, a caspase-2 inhibitor was able to suppress the activation of both caspase-2 and caspase-9. All of these results suggest that cytochrome c release and activation of caspase-9 are delayed or secondary phenomenon and the damage of mitochondria might not be the essential event in the triggering of apoptosis induced by sustained GN depletion.

Caspase-2 is involved in the apoptosis occurred in various circumstances. It is activated during cell death induced by TNFR in B-lymphocytes and other cells (42, 43), but the major mediator of TNFR is caspase-8 as the deletion of RAIDD (caspase-2 adapter) failed to block TNF-induced apoptosis (23). Caspase-2 activation is also required for the apoptosis induced by withdrawal of trophic support from PC12 cells and sympathetic neurons (44, 45) and caspase-3-like activity is not required in this paradigm. TGFß-evoked apoptosis of a variety of cells also involves caspase-2 activation (46). Thus, it is highly possible that the antecedent blockade of mitogenesis (Fig. 1AGo and Table 1Go) subsequently induces apoptosis (Fig. 1Go, B and C) to prevent accumulation of DNA-aberrant cells in G1/S phases. It is interesting to note that caspase-2, similar to its effect in GN depletion-elicited HIT cell death, plays a major role in apoptosis in B-lymphocytes induced by B cell receptor cross-linking (36). In the latter system, caspase-2 was activated early and markedly while caspase-9 activity was moderately enhanced without the involvement of caspase-8. In addition, the cell death could be prevented by pan-caspase inhibitors or specific caspase-2 inhibitor, resembling the findings in GN depletion-induced apoptosis of HIT cells. Thus caspase-2 activation may represent another major class of apoptotic pathways distinct from that mediated by caspase-8 and caspase-9 (16, 17).

How does GN depletion activate caspase-2 in ß-cells? Although it might involve the inhibition of primer RNA biosynthesis (13), reductions in mitogenic competence, and/or the interference with the function of some small G proteins (5, 12), the exact nature of the upstream mediators leading to the observed activation of caspase-2 is unclear. It has been proposed that RAIDD may be an upstream regulator of caspase-2 because an interaction between the two proteins occurs both in the cell and in cell-free system (23, 42). Endogenous RAIDD is mainly localized in the cytoplasm and to some extent in the nucleus, where its interaction with caspase-2 may occur (42). We also found that RAIDD interacts with caspase-2 in HIT cells, as assessed by immunoprecipitation. It appeared this interaction was slightly reduced by GN depletion. What importance of this change to GN depletion-induced apoptosis is unclear in the moment because there is no study that reported an alteration of RAIDD-caspase-2 interaction during apoptosis.

Our other preliminary studies may shed some light on the mechanism underlying caspase-2 activation by GN depletion. It is known that the molecules involved in the regulation of cyclin-dependent kinases (CDKs) (47), such as p53 and p21waf1/cip1, play an important role in the control of cell survival and death (2, 48). p53 is a tumor suppressor that regulates p21waf1/cip1 and its mass is often increased in apoptotic cells (48, 49). Cdk4 is particular important for islet ß-cell growth and postnatal survival (50). Induction of p21waf1/cip1 is observed in the islets under growth arrest and death due to exposure to oxygen radicals (51). Similar increases in p21waf1/cip1 and also p53 occur in amylin-induced ß-cell death (48). It is of particular interest that p53 may regulate IMPDH (and vice versa) (52), the key enzyme in GTP biosynthesis that is targeted by MPA. Nonetheless, p53 seems unlikely to play a major role in GN depletion-induced apoptosis as its mass was reduced after MPA treatment1. Interestingly, however, our unpublished preliminary results2 indicated that p21waf1/cip1 mass was up-regulated in close parallel with caspase-2 activation, suggesting that p21waf1/cip1 might be involved in the initiation of cell death due to GN depletion. Moreover, the increase in p21waf1/cip1 during GN depletion could not be blocked by caspase inhibitors that prevented ß-cells from death, suggesting that p21waf1/cip1 might be an upstream signal for caspase-2 activation. The possible detailed relationship between the induction of p21waf1/cip1 and the activation of caspase-2 during GN depletion requires additional study.

Both clinical and experimental studies indicates that a failure of islet ß-cell function, arrest of cell growth, and promotion of cell death by apoptosis occur during challenge with cytokines, glucose toxicity, or lipotoxicity (53, 54, 55). Cytokines such as IL-1ß, TNF, and interferon induce autoimmune damage (53, 56) and cause ß-cell death probably mediated by FAS and TNFR (15, 57, 58). Similar apoptotic effects on ß-cells are also seen subsequent of the stimulation of amyloid (48) or free radicals such as nitric oxide generated from streptozotocin (56). However, the information on caspases in the death of islet ß-cells occurred in above circumstances is very fragmentary. Our unpublished data3 indicate that caspase-2 may be not the critical protease in all forms of programmed cell death because the caspase-2 inhibitor (Z-VDVAD-FMK) could not prevent the apoptotic death of HIT cells induced by streptozotocin and only partially (by ~30%) reversed cytokine (IL-1ß plus TNF{alpha})-induced HIT cell death. It has been reported that IL-1ß lowered GTP by approximately 60% in islet ß-dells (14). Other studies reported the activation of caspase-1 and -3 by cytokines in ß-cells (58, 59). These observations suggest again that caspase-2 activation may play a specific, critical role in the apoptosis induced by GN depletion with MPA, due to impairment of mitogenesis. Little is known about the role of caspases in ß-cell apoptosis caused by glucose toxicity and lipid toxicity. Our preliminary results indicated that high concentrations of FFA increased the activity of both caspase-2 and caspase-3 in insulin-secreting INS-1 cells4. The present study on ß-cell growth and death during GTP-depletion may enrich our knowledge in this aspect.


   Acknowledgments
 
We are grateful to Dr. R. P. Robertson for providing HIT-T15 cells.


   Footnotes
 
Portions of this work were presented at the 60th Annual Scientific Meetings of American Diabetes Association, San Antonio, Texas, June 2000. This work was supported by the National Medical Research Council of Singapore (R-364-000-004-213 and NMRC/0540/2001 (to G.D.L.) and the National Institutes of Health (DK-37312, to S.A.M.). J.H. was a recipient of a Research Scholarship from the National University of Singapore, and his work in this paper is partly to meet the requirements for completing the Ph.D. study.

Abbreviations: AFC, 7-Amino-4-trifluoromethyl coumarin; DTT, dithiothreitol; GN, guanine nucleotide; IMPDH, inosine 5'-monophosphate dehydrogenase; MPA, mycophenolic acid; MTS, 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium; PI, propidium iodide; PMS, phenazine methosulfate; PMSF, phenylmethylsulfonyl fluoride; PVDF, polyvinylidine difluoride; RAIDD, RIP-associated ICH-1/CED-3-homologous protein with a death domain; RIP, receptor-interacting protein; TBST, Tris-buffered saline buffer with Tween-20; TNFR, TNF receptor.

1 By assessment with Western-blotting, p53 was reduced by 17% and 50% after treatment with 3 µg/ml MPA for 16 and 48 h, respectively. Back

2 p21waf1/cip1 was increased by 23%, 159%, and 220% following 8, 24, and 32 h treatment with 3 µg/ml MPA in HIT cells. Back

3 The DNA content of HIT cells (per well) was 81.1 ± 4.1% and 83.0 ± 3.4% of control (n = 3 each) after 24-h treatment with 10 mM streptozotocin alone or plus 100 µM Z-VDVAD-FMK. The DNA content of HIT cells (per well) was 57.8 ± 1.8% of control after 24-h treatment with cytokines (50 ng/ml IL-1ß and 50 ng/ml TNF{alpha}) and 71.4 ± 2.4% of control (P < 0.01 vs. cytokines alone; n = 6 each) when coexposed to 100 µM Z-VDVAD-FMK. Back

4 The activity of caspase-2 and caspase-3 was increased by 160% and 389% in INS-1 cells after 32-h exposure to 2 mM fatty acids. Back

Received April 30, 2001.

Accepted for publication January 22, 2002.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Metz SA, Kowluru A 1999 Inosine monophosphate dehydrogenase: a molecular switch integrating pleiotropic GTP-dependent ß-cell functions. Proc Assoc Am Physicians 111:335–346[CrossRef][Medline]
  2. Jayaram HN, Cooney DA, Grusch M 1999 Consequences of IMP dehydrogenase inhibition, and its relationship to cancer and apoptosis. Curr Med Chem 6:561–574[Medline]
  3. Metz S, Holland S, Johnson L, Espling E, Rabaglia M, Segu V, Brockenbrough JS, Tran PO 2001 Inosine-5'-monophosphate dehydrogenase is required for mitogenic competence of transformed pancreatic ß cells. Endocrinology 142:193–204[Abstract/Free Full Text]
  4. Fulton B, Markham A 1996 Mycophenolate mofetil. A review of its pharmacodynamic and pharmacokinetic properties and clinical efficacy in renal transplantation. Drugs 51:278–298[Medline]
  5. Li GD, Segu VB, Rabaglia ME, Luo RH, Kowluru A, Metz SA 1998 Prolonged depletion of guanosine triphosphate induces death of insulin-secreting cells by apoptosis. Endocrinology 139:3752–3762[Abstract/Free Full Text]
  6. Cohn RG, Mirkovich A, Dunlap B, Burton P, Chiu SH, Eugui E, Caulfield JP 1999 Mycophenolic acid increases apoptosis, lysosomes and lipid droplets in human lymphoid and monocytic cell lines. Transplantation 68:411–418[Medline]
  7. Metz SA, Rabaglia ME, Pintar TJ 1992 Selective inhibitors of GTP synthesis impede exocytotic insulin release from intact rat islets. J Biol Chem 267:12517–12527[Abstract/Free Full Text]
  8. Li GD, Luo RH, Metz SA 2000 Effects of inhibitors of guanine nucleotide synthesis on membrane potential and cytosolic free Ca2+ levels in insulin-secreting cells. Biochem Pharmacol 59:545–556[CrossRef][Medline]
  9. Meredith M, Li GD, Metz SA 1997 GTP depletion impedes calcium-induced insulin secretion from intact HIT-T15 or INS-1 ß cells. Biochem Pharmacol 53:1873–1882[CrossRef][Medline]
  10. Detimary P, Xiao C, Henquin JC 1997 Tight links between adenine and guanine nucleotide pools in mouse pancreatic islets: a study with mycophenolic acid. Biochem J 324 (Pt 2):467–471
  11. Komatsu M, Noda M, Sharp GW 1998 Nutrient augmentation of Ca2+-dependent and Ca2+-independent pathways in stimulus-coupling to insulin secretion can be distinguished by their guanosine triphosphate requirements: studies on rat pancreatic islets. Endocrinology 139:1172–1183[Abstract/Free Full Text]
  12. Kowluru A, Seavey SE, Li G, Sorenson RL, Weinhaus AJ, Nesher R, Rabaglia ME, Vadakekalam J, Metz SA 1996 Glucose- and GTP-dependent stimulation of the carboxyl methylation of CDC42 in rodent and human pancreatic islets and pure ß cells: evidence for an essential role of GTP-binding proteins in nutrient-induced insulin secretion. J Clin Invest 98:540–555[Medline]
  13. Catapano CV, Dayton JS, Mitchell BS, Fernandes DJ 1995 GTP depletion induced by IMP dehydrogenase inhibitors blocks RNA-primed DNA synthesis. Mol Pharmacol 47:948–955[Abstract]
  14. Meredith M, Rabaglia M, Corbett JA, Metz SA 1996 Dual functional effects of interleukin-1ß on purine nucleotides and insulin secretion in rat islets and INS-1 cells. Diabetes 45:1783–1791[Abstract]
  15. Iwahashi H, Hanafusa T, Eguchi Y, Nakajima H, Miyagawa J, Itoh N, Tomita K, Namba M, Kuwajima M, Noguchi T, Tsujimoto Y, Matsuzawa Y 1996 Cytokine-induced apoptotic cell death in a mouse pancreatic ß-cell line: inhibition by Bcl-2. Diabetologia 39:530–536[Medline]
  16. Kumar S 1999 Mechanisms mediating caspase activation in cell death. Cell Death Differ 6:1060–1066[CrossRef][Medline]
  17. Earnshaw WC, Martins LM, Kaufmann SH 1999 Mammalian caspases: structure, activation, substrates, and functions during apoptosis. Annu Rev Biochem 68:383–424[CrossRef][Medline]
  18. Green DR, Reed JC 1998 Mitochondria and apoptosis. Science 281:1309–1312[Abstract/Free Full Text]
  19. Nakagawa T, Zhu H, Morishima N, Li E, Xu J, Yankner BA, Yuan J 2000 Caspase-12 mediates endoplasmic-reticulum-specific apoptosis and cytotoxicity by amyloid-ß. Nature 403:98–103[CrossRef][Medline]
  20. Wang S, Miura M, Jung YK, Zhu H, Li E, Yuan J 1998 Murine caspase-11, an ICE-interacting protease, is essential for the activation of ICE. Cell 92:501–509[CrossRef][Medline]
  21. Li P, Nijhawan D, Budihardjo I, Srinivasula SM, Ahmad M, Alnemri ES, Wang X 1997 Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade. Cell 91:479–489[CrossRef][Medline]
  22. Ito A, Uehara T, Tokumitsu A, Okuma Y, Nomura Y 1999 Possible involvement of cytochrome c release and sequential activation of caspases in ceramide-induced apoptosis in SK-N-MC cells. Biochim Biophys Acta 1452:263–274[Medline]
  23. Duan H, Dixit VM 1997 RAIDD is a new 'death’ adaptor molecule. Nature 385:86–89[CrossRef][Medline]
  24. Thome M, Hofmann K, Burns K, Martinon F, Bodmer JL, Mattmann C, Tschopp J 1998 Identification of CARDIAK, a RIP-like kinase that associates with caspase-1. Curr Biol 8:885–888[CrossRef][Medline]
  25. Nicoletti I, Migliorati G, Pagliacci MC, Grignani F, Riccardi C 1991 A rapid and simple method for measuring thymocyte apoptosis by propidium iodide staining and flow cytometry. J Immunol Methods 139:271–279[CrossRef][Medline]
  26. Cory AH, Owen TC, Barltrop JA, Cory JG 1991 Use of an aqueous soluble tetrazolium/formazan assay for cell growth assays in culture. Cancer Commun 3:207–212[Medline]
  27. Rago R, Mitchen J, Wilding G 1990 DNA fluorometric assay in 96-well tissue culture plates using Hoechst 33258 after cell lysis by freezing in distilled water. Anal Biochem 191:31–34[CrossRef][Medline]
  28. Komoriya A, Packard BZ, Brown MJ, Wu ML, Henkart PA 2000 Assessment of caspase activities in intact apoptotic thymocytes using cell-permeable fluorogenic caspase substrates. J Exp Med 191:1819–1828[Abstract/Free Full Text]
  29. Li GD, Kowluru A, Metz SA 1996 Characterization of prenylcysteine methyltransferase in insulin-secreting cells. Biochem J 316:345–351
  30. Talanian RV, Quinlan C, Trautz S, Hackett MC, Mankovich JA, Banach D, Ghayur T, Brady KD, Wong WW 1997 Substrate specificities of caspase family proteases. J Biol Chem 272:9677–9682[Abstract/Free Full Text]
  31. Li H, Bergeron L, Cryns V, Pasternack MS, Zhu H, Shi L, Greenberg A, Yuan J 1997 Activation of caspase-2 in apoptosis. J Biol Chem 272:21010–21017[Abstract/Free Full Text]
  32. Borner C, Monney L 1999 Apoptosis without caspases: an inefficient molecular guillotine? Cell Death Differ 6:497–507[CrossRef][Medline]
  33. Lorenzo HK, Susin SA, Penninger J, Kroemer G 1999 Apoptosis inducing factor (AIF): a phylogenetically old, caspase-independent effector of cell death. Cell Death Differ 6:516–524[CrossRef][Medline]
  34. Deas O, Dumont C, MacFarlane M, Rouleau M, Hebib C, Harper F, Hirsch F, Charpentier B, Cohen GM, Senik A 1998 Caspase-independent cell death induced by anti-CD2 or staurosporine in activated human peripheral T lymphocytes. J Immunol 161:3375–3383[Abstract/Free Full Text]
  35. MacFarlane M, Cain K, Sun XM, Alnemri ES, Cohen GM 1997 Processing/activation of at least four interleukin-1ß converting enzyme-like proteases occurs during the execution phase of apoptosis in human monocytic tumor cells. J Cell Biol 137:469–479[Abstract/Free Full Text]
  36. Chen W, Wang HG, Srinivasula SM, Alnemri ES, Cooper NR 1999 B cell apoptosis triggered by antigen receptor ligation proceeds via a novel caspase-dependent pathway. J Immunol 163:2483–2491[Abstract/Free Full Text]
  37. Banki K, Hutter E, Gonchoroff NJ, Perl A 1999 Elevation of mitochondrial transmembrane potential and reactive oxygen intermediate levels are early events and occur independently from activation of caspases in Fas signaling. J Immunol 162:1466–1479[Abstract/Free Full Text]
  38. Soane L, Rus H, Niculescu F, Shin ML 1999 Inhibition of oligodendrocyte apoptosis by sublytic C5b-9 is associated with enhanced synthesis of bcl-2 and mediated by inhibition of caspase-3 activation. J Immunol 163:6132–6138[Abstract/Free Full Text]
  39. Tepper AD, Cock JG, de Vries E, Borst J, van Blitterswijk WJ 1997 CD95/Fas-induced ceramide formation proceeds with slow kinetics and is not blocked by caspase-3/CPP32 inhibition. J Biol Chem 272:24308–24312[Abstract/Free Full Text]
  40. Bras A, Ruiz-Vela A, Gonzalez dB, Martinez A 1999 Caspase activation by BCR cross-linking in immature B cells: differential effects on growth arrest and apoptosis. FASEB J 13:931–944[Abstract/Free Full Text]
  41. Susin SA, Lorenzo HK, Zamzami N, Marzo I, Brenner C, Larochette N, Prevost MC, Alzari PM, Kroemer G 1999 Mitochondrial release of caspase-2 and -9 during the apoptotic process. J Exp Med 189:381–394[Abstract/Free Full Text]
  42. Shearwin-Whyatt LM, Harvey NL, Kumar S 2000 Subcellular localization and CARD-dependent oligomerization of the death adaptor RAIDD. Cell Death Differ 7:155–165[CrossRef][Medline]
  43. Xu Y, Bialik S, Jones BE, Iimuro Y, Kitsis RN, Srinivasan A, Brenner DA, Czaja MJ 1998 NF-{kappa}B inactivation converts a hepatocyte cell line TNF-{alpha} response from proliferation to apoptosis. Am J Physiol 275:C1058–C1066
  44. Stefanis L, Troy CM, Qi H, Shelanski ML, Greene LA 1998 Caspase-2 (Nedd-2) processing and death of trophic factor-deprived PC12 cells and sympathetic neurons occur independently of caspase-3 (CPP32)- like activity. J Neurosci 18:9204–9215[Abstract/Free Full Text]
  45. Haviv R, Lindenboim L, Yuan J, Stein R 1998 Need for caspase-2 in apoptosis of growth-factor-deprived PC12 cells. J Neurosci Res 52:491–497[CrossRef][Medline]
  46. Choi KS, Lim IK, Brady JN, Kim SJ 1998 ICE-like protease (caspase) is involved in transforming growth factor ß1-mediated apoptosis in FaO rat hepatoma cell line. Hepatology 27:415–421[CrossRef][Medline]
  47. Matsumoto Y, Hayashi K, Nishida E 1999 Cyclin-dependent kinase 2 (Cdk2) is required for centrosome duplication in mammalian cells. Curr Biol 9:429–432[CrossRef][Medline]
  48. Zhang S, Liu J, Saafi EL, Cooper GJ 1999 Induction of apoptosis by human amylin in RINm5F islet ß-cells is associated with enhanced expression of p53 and p21WAF1/CIP1. FEBS Lett 455:315–320[CrossRef][Medline]
  49. Wu M, Bellas RE, Shen J, Sonenshein GE 1998 Roles of the tumor suppressor p53 and the cyclin-dependent kinase inhibitor p21WAF1/CIP1 in receptor-mediated apoptosis of WEHI 231 B lymphoma cells. J Exp Med 187:1671–1679[Abstract/Free Full Text]
  50. Rane SG, Dubus P, Mettus RV, Galbreath EJ, Boden G, Reddy EP, Barbacid M 1999 Loss of Cdk4 expression causes insulin-deficient diabetes and Cdk4 activation results in ß-islet cell hyperplasia. Nat Genet 22:44–52[CrossRef][Medline]
  51. Kaneto H, Kajimoto Y, Fujitani Y, Matsuoka T, Sakamoto K, Matsuhisa M, Yamasaki Y, Hori M 1999 Oxidative stress induces p21 expression in pancreatic islet cells: possible implication in ß-cell dysfunction. Diabetologia 42:1093–1097[CrossRef][Medline]
  52. Sherley JL 1991 Guanine nucleotide biosynthesis is regulated by the cellular p53 concentration. J Biol Chem 266:24815–24828[Abstract/Free Full Text]
  53. Kay TW, Thomas HE, Harrison LC, Allison I 2000 The ß cell in autoimmune diabetes: many mechanisms and pathways of loss. Trends Endocrinol Metab 11:11–15[CrossRef][Medline]
  54. Shimabukuro M, Zhou YT, Levi M, Unger RH 1998 Fatty acid-induced ß cell apoptosis: a link between obesity and diabetes. Proc Natl Acad Sci USA 95:2498–2502[Abstract/Free Full Text]
  55. Pick A, Clark J, Kubstrup C, Levisetti M, Pugh W, Bonner-Weir S, Polonsky KS 1998 Role of apoptosis in failure of ß-cell mass compensation for insulin resistance and ß-cell defects in the male Zucker diabetic fatty rat. Diabetes 47:358–364[Abstract]
  56. Kaneto H, Kajimoto Y, Miyagawa J, Matsuoka T, Fujitani Y, Umayahara Y, Hanafusa T, Matsuzawa Y, Yamasaki Y, Hori M 1999 Beneficial effects of antioxidants in diabetes: possible protection of pancreatic ß-cells against glucose toxicity. Diabetes 48:2398–2406[Abstract]
  57. Delaney CA, Pavlovic D, Hoorens A, Pipeleers DG, Eizirik DL 1997 Cytokines induce deoxyribonucleic acid strand breaks and apoptosis in human pancreatic islet cells. Endocrinology 138:2610–2614[Abstract/Free Full Text]
  58. Stephens LA, Thomas HE, Ming L, Grell M, Darwiche R, Volodin L, Kay TW 1999 Tumor necrosis factor-{alpha}-activated cell death pathways in NIT-1 insulinoma cells and primary pancreatic ß cells. Endocrinology 140:3219–3227[Abstract/Free Full Text]
  59. Karlsen AE, Pavlovic D, Nielsen K, Jensen J, Andersen HU, Pociot F, Mandrup-Poulsen T, Eizirik DL, Nerup J 2000 Interferon-{gamma} induces interleukin-1 converting enzyme expression in pancreatic islets by an interferon regulatory factor-1-dependent mechanism. J Clin Endocrinol Metab 85:830–836[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 J. Immunol.Home page
B. Chaigne-Delalande, G. Guidicelli, L. Couzi, P. Merville, W. Mahfouf, S. Bouchet, M. Molimard, B. Pinson, J.-F. Moreau, and P. Legembre
The Immunosuppressor Mycophenolic Acid Kills Activated Lymphocytes by Inducing a Nonclassical Actin-Dependent Necrotic Signal
J. Immunol., December 1, 2008; 181(11): 7630 - 7638.
[Abstract] [Full Text] [PDF]

Home page
 J EndocrinolHome page
M Asfari, S Berta, S Coffy, M Kergoat, C Charon, and A Audet
Short-term treatment with mycophenolic acid and tacrolimus is tolerogenic for INS-1 cell clone transplantation and the deleterious effects of the drugs are limited: in vivo and in vitro studies
J. Endocrinol., July 1, 2005; 186(1): 213 - 220.
[Abstract] [Full Text] [PDF]