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Endocrinology Vol. 139, No. 9 3752-3762
Copyright © 1998 by The Endocrine Society

ARTICLES

Prolonged Depletion of Guanosine Triphosphate Induces Death of Insulin-Secreting Cells by Apoptosis1

Guodong Li, Venkatesh Babu G. Segu, Mary E. Rabaglia, Rui-Hua Luo, Anjaneyulu Kowluru and Stewart A. Metz2

Medical Service, Section of Endocrinology, Middleton Veterans Administration Hospital (G.L., V.B.G.S., M.E.R., A.K., S.A.M.), Madison, Wisconsin 53705; and the Department of Medicine, Division of Endocrinology, University of Wisconsin Medical School, Madison, Wisconsin 53792; and National University Medical Institutes, National University of Singapore (G.L., R.H.L.), S-119260 Singapore

Address all correspondence and requests for reprints to: Dr. Guodong Li, National University Medical Institutes, National University of Singapore, MD 11 #02–01, 10 Kent Ridge Crescent, S-119260 Singapore. E-mail: nmiligd{at}med2.nusstf.nus.sg


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Inhibitors of IMP dehydrogenase, such as mycophenolic acid (MPA) and mizoribine, which deplete cellular GTP, are used clinically as immunosuppressive drugs. The prolonged effect of such agents on insulin-secreting ß-cells (HIT-T15 and INS-1) was investigated. Both MPA and mizoribine inhibited mitogenesis, as reflected by [3H]thymidine incorporation. Cell number, DNA and protein contents, and cell (metabolic) viability were decreased by about 30%, 60%, and 80% after treatment of HIT cells with clinically relevant concentrations (e.g. 1 µg/ml) of MPA for 1, 2, and 4 days, respectively. Mizoribine (48 h) similarly induced the death of HIT cells. INS-1 cells also were damaged by prolonged MPA treatment. MPA-treated HIT cells displayed a strong and localized staining with a DNA-binding dye (propidium iodide), suggesting condensation and fragmentation of DNA, which were confirmed by detection of DNA laddering in multiples of about 180 bp. DNA fragmentation was observed after 24-h MPA treatment and was dose dependent (29%, 49%, and 70% of cells were affected after 48-h exposure to 1, 3, and 10 µg/ml MPA, respectively). Examination of MPA-treated cells by electron microscopy revealed typical signs of apoptosis: condensed and marginated chromatin, apoptotic bodies, cytosolic vacuolization, and loss of microvilli. MPA-induced cell death was almost totally prevented by supplementation with guanosine, but not with adenosine or deoxyguanosine, indicating a specific effect of GTP depletion. An inhibitor of protein isoprenylation (lovastatin, 10–100 µM for 2–3 days) induced cell death and DNA degradation similar to those induced by sustained GTP depletion, suggesting a mediatory role of posttranslationally modified GTP-binding proteins. Indeed, impeding the function of G proteins of the Rho family (via glucosylation using Clostridium difficile toxin B), although not itself inducing apoptosis, potentiated cell death induced by MPA or lovastatin. These findings indicate that prolonged depletion of GTP induces ß-cell death compatible with apoptosis; this probably involves a direct impairment of GTP-dependent RNA-primed DNA synthesis, but also appears to be modulated by small GTP-binding proteins. Treatment of intact adult rat islets (the ß-cells of which replicate slowly) induced a modest, but definite, death by apoptosis over 1- to 3-day periods. Thus, more prolonged use of the new generation of immunosuppressive agents exemplified by MPA might have deleterious effects on the survival of islet or pancreas grafts.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GTP IS required for many biological activities in the cell, e.g. synthesis of DNA, RNA, and proteins; nutrient metabolism; and cell signaling. It is well established that GTP-binding proteins play a diversity of roles as switches in cell growth, receptor activation, exocytosis, ion channel conductivity, and change in cell shape (1, 2).

Over the past several years, the roles of GTP and GTP-binding proteins in the physiology of the normal pancreatic ß-cell have been studied in our laboratory using specific inhibitors of inosine monophosphate dehydrogenase, e.g. mycophenolic acid (MPA) and mizoribine (MZ), as tools (3, 4, 5, 6, 7, 8). These agents deplete cellular GTP by blockade of the conversion of IMP to GMP, the precursor to the synthesis of GDP and GTP. In these studies, short term (18-h) exposure to MPA or MZ inhibited the production of guanine nucleotides and, concomitantly, potently inhibited nutrient-induced insulin secretion in isolated islets (3, 4, 7). Subsequently, we also observed that depletion of GTP using MPA or MZ inhibited Ca2+-stimulated insulin secretion from cloned ß-cells (HIT-T15 and INS-1) (9). The effects of MPA or MZ were specific for the depletion of guanine nucleotides, as their effects were totally reversed by provision of guanine or guanosine, but not by adenine or adenosine. Thus, MPA appears to have short term inhibitory effects on ß-cell function, as manifested by insulin secretion.

MPA is the active compound of mycophenolate mofetil, a new immunosuppressive agent widely used in clinical therapy, including pancreas transplantation (10, 11). However, potentially harmful effects of this drug on islet cells have been observed in vitro, including impediment of insulin secretion and reduction of the DNA content of rat islets (12, 13). The MPA concentrations (between 2–25 µg/ml) used in our short term in vitro studies are within the range of MPA levels achievable in humans (14), rodents, and primates (14, 15) in vivo during therapy. The objective of the current study was to characterize the subacute effects of MPA on the mitogenesis and survival of insulin-secreting ß-cells. We found that 1–4 days of treatment of ß-cells with GTP-depleting agents blocked mitogenesis, followed by progressive cell killing in a manner of cell death characteristic of apoptosis (programed cell death), as evidenced by both ultrastructural and biochemical parameters. The cell death induced by GTP depletion might be modulated or mediated by GTP-binding proteins, as restraint of function of certain GTP-binding proteins (by blockade of their posttranslational isoprenylation with lovastatin treatment or exposure to Clostridium difficile toxin B) also induced or potentiated apoptosis of insulin-secreting cells.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
MPA, propidium iodide, Hoechst 33258, and deoxyribonuclease-free ribonuclease A (RNase A) were purchased from Sigma (St. Louis, MO); [3H]thymidine was obtained from DuPont-New England Nuclear (Boston, MA); RPMI 1640, FCS, and Tris-boric acid-EDTA buffer were purchased from Life Technologies (Gaithersburg, MD). MZ was a gift from Dr. N. Kazmatani (Tokyo Women’s Medical College, Tokyo, Japan). Lovastatin was a gift from Dr. A. W. Alberts (Merck, Sharpe, and Dohme Research laboratories, Rahway, NJ) and was transformed to its sodium salt before use. C. difficile toxin B was a gift from Dr. K. Aktories of Albert-Ludwigs-University (Freiburg, Germany). The cell death detection enzyme-linked immunosorbent assay (ELISA) kit was purchased from Boehringer Mannheim (Mannheim, Germany).

Cell culture
Insulin-secreting HIT-T15 (passages 73–80; provided by Drs. R. P. Robertson and H.-J. Zhang, University of Minneapolis, Minneapolis, MN) and INS-1 (passages 60–88; a gift from Dr. C. B. Wollheim, University of Geneva, Switzerland) cells were cultured in RPMI 1640 supplemented with 10% FCS as described previously (16, 17).

Intact rat islets were isolated from Sprague-Dawley rats, as previously described (3, 4, 5, 6, 7, 8).

[3H]Thymidine incorporation
HIT cells were cultured in 96-well plates and treated with test agents for various periods in RPMI 1640. [3H]Thymidine (0.5 µCi/well) was included during the last 3 h unless specified. Cells were disrupted and collected with a cell harvester (PHD, Cambridge Technology, Watertown, MA); samples were washed through glass-fiber membranes on which DNA was retained. The membranes were counted using a ß-scintillation counter.

3-(4,5)-Dimethylthiazol-2-yl)-5-(3-carboxy-methoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) assay
The metabolic viability of the cells was monitored (18) using a MTS assay kit (CellTiter 96) developed by Promega (Madison, WI). The principle of this assay is similar to that of the older, widely used 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT) assay that measures colored formazan, produced in a reduction reaction driven by nutrient metabolism; formazan can be easily quantified spectrophotometrically. The MTS assay has a major advantage over the MTT assay, as the formazan formed during the former is soluble and exits the cells (19), and the formazan produced by the latter is crystalline and must be solubilized (which requires disrupting the cells) (20).

HIT cells were seeded onto 24- or 96-well plates and cultured in the presence of test agents for various periods. A mixture of MTS and phenazine methosulfate (an electron-coupling reagent; final concentrations, 333 and 25 µg/ml, respectively) was added, and cells were incubated for 30 min at 37 C. The reaction was stopped by addition of 10% SDS if samples were not immediately subjected to determination of absorbance. Formazan formed from reduction of MTS (18, 19) was quantified by measurement of absorbance of the medium at 490 nm using a microplate reader. All data have been corrected for background signals.

DNA staining
HIT or INS-1 cells seeded on glass coverslips were cultured in RPMI 1640 and treated with test agents for the times specified. Cells were then fixed in 4% paraformaldehyde in PBS for 15 min and dried in air. The cells were left in 70% ethanol for 10 min at -20 C. Thereafter, the cells on coverslips were incubated in PBS containing 4 µg/ml propidium iodide and 100 µg/ml DNA-free RNase A for 30 min at 37 C. After three washes with PBS, the coverslips were mounted and examined by inverted fluorescence microscopy (DIAPHOT-TMD, Nikon, Melville, NY) using a filter set (emission at 510 nm) for fluorescein. Quantitative analysis of the data was carried out by counting cells (>300 in each case) in randomly selected fields on the photographs taken under the microscope.

Electron microscopy
HIT cells were seeded on glass coverslips and treated with or without MPA in culture. The cells were fixed with 2% glutaraldehyde and postfixed with 1% OsO4 in 0.1 M phosphate buffer. The samples were embedded in Spurrs resin, and the sections were examined using a JEOL 100Cx electron microscope (JEOL, Peabody, MA). Quantitative analysis of the data was carried out by counting all cells on the photographs nonselectively taken under the microscope.

DNA laddering
HIT cells in culture dishes (10-cm diameter) were treated with test agents for 2 days. The nonadherent cells were collected by centrifugation and pooled with the attached cells, which were scraped in lysis buffer (10 mM Tris, 10 mM EDTA, and 1% Nonidet P-40, pH 7.5). After centrifugation at 13,000 x g, the supernatant was first extracted with phenol and then with phenol-chloroform-isoamyl alcohol (25:24:1) to remove proteins and lipids. DNA in the supernatants was then precipitated in ethanol overnight at -20 C and pelleted at 14,000 x g. After washing with 70% ethanol, the pellet was resuspended in Tris-EDTA buffer. The samples were incubated with deoxyribonuclease-free RNase A at 37 C for 1 h. Equal fractions of extraction volumes from each experimental condition were loaded onto 2% agarose gel and run in Tris-boric acid-EDTA buffer under 95 V. DNA was stained with ethidium bromide, and photographs were taken under UV light.

Detection of cell apoptosis by ELISA
The mono- and oligonucleosomes due to DNA fragmentation induced during apoptosis were also detected with a commercially available ELISA kit (from Boehringer Mannheim, catologue no. 1544-675). HIT or INS-1 cells were seeded on multiwell plates and cultured in RPMI 1640 containing various agents for 48 h. Nonadherent cells were collected and extracted together with attached cells in the lysis buffer containing 0.3% Triton X-100 or using lysis solution provided by the kit for 20 min on ice. The cell extracts were centrifuged for 10 min at 14,000 x g. The supernatants containing mono- and oligonucleosomes were equally diluted for all samples and assayed according to the kit instructions. In brief, the ELISA plate was coated with antihistone. After blocking the wells using a buffer provided by the kit, samples were added and incubated for 90 min. Then anti-DNA conjugated with peroxidase was introduced. Color was developed by adding the substrate of peroxidase (ABTS), and optical density was measured at 405 nm with a plate reader. The results are expressed as enrichment factor (the ratio of the OD readings from treated cells and the OD values from corresponding control cells.

Studies of normal, intact pancreatic islets
For studies of normal rat islets, 50–100 islets/plate were cultured for 18–67 h in the presence or absence of MPA (25 µg/ml) or diluent as described above. NaF (2.5–5 mM) or, in one study, S-nitrosoglutathione (500 µM) was added to some plates as a positive apoptosis control (21). Islets were disrupted in lysis buffer (0.5 ml 10 mM Tris-HCl, containing 0.3% Triton X-100 and 10 mM EDTA, pH 7.5) at 4 C for 30 min, with gentle vortexing every 10 min. After centrifugation (14,000 rpm for 10 min), the lysates were transferred to a fresh tube and then analyzed by the cell death detection ELISA method, generally using a 1:10 dilution of cell extract in the assay.

Miscellaneous
Cell number was determined using a cell-counting chamber after detachment of cells with gentle trypsinization (0.025% trypsin for 2–3 min). Protein content, DNA content, and insulin content were determined by Bio-Rad assay (Bio-Rad, Richmond, CA), the DNA-binding fluorescent probe Hoechst 33258, and RIA, respectively.

Data are expressed as the mean ± SE, and statistical analysis was performed using Student’s two-tailed unpaired t test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Inhibition of mitogenesis by depletion of GTP
The effect of GTP depletion on mitogenesis was examined by measurement of DNA synthesis using [3H]thymidine incorporation. Figure 1Go shows that [3H]thymidine incorporation into HIT-T15 cell DNA was very sensitive to inhibition by MPA treatment (Fig. 1BGo), with inhibition (-35%; P < 0.01) seen as early as 1 h at 1 µg/ml MPA (Fig. 1AGo). At 1 µg/ml MPA, [3H]thymidine incorporation was reduced by 67% after 3 h of treatment and was almost completely abolished after 6- to 24-h exposure to MPA. This effect was also dose dependent; after 6-h treatment with MPA, [3H]thymidine incorporation was reduced by 39%, 80%, and 98% at 0.03, 0.1, and 1 µg/ml MPA, respectively (Fig. 1BGo). MZ, another structurally dissimilar (but mechanistically identical), GTP-depleting agent, also inhibited [3H]thymidine incorporation, although less potently than MPA (Fig. 1BGo), similar to GTP depletion (3). At the 3 h point, the inhibitory effect of MPA could be completely prevented by coprovision of 500 µM guanosine even though guanosine itself diminished [3H]thymidine incorporation in control cells (Fig. 1AGo). With treatments longer than 6 h, coprovision of 500 µM (but not 100 µM) guanosine could prevent, albeit only partially, the MPA effect (Fig. 1CGo). Deoxyguanosine (30 µM) was also able to partially prevent the MPA effect, whereas adenosine had no such effect and was, in fact, inhibitory by itself (Fig. 1CGo).



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  		Figure 1. Effects of MPA and MZ on [3H]thymidine incorporation in HIT cells. HIT cells were cultured in 96-well plates and were exposed to different concentrations of MPA or MZ alone or combined with other agents in culture medium for 6 h (B and C). [3H]Thymidine was included in the last 3 h, except in A, where [3H]thymidine was added simultaneously with test substances. Labeled DNA was collected with a cell harvester and counted in a scintillation counter. Values are the mean ± SE of 5–10 observations.

 
Cell death induced by prolonged treatment with GTP-depleting agents
Treatment of HIT or INS-1 cells with MPA for 1–4 days caused cell death, as assessed by cell number, trypan blue exclusion test, and MTS test. Cell number was decreased after long term (2 days) MPA treatment (Fig. 2Go). This mainly resulted from cell death and quantitatively could be only partially explained by a blockade of cell proliferation, as the number of control cells increased only slightly (10–20%) during 2 days. The latter is due to the fact that cells had already approached confluence at the time when MPA was added. MPA-treated cells appeared rounded and became easily detached after more than 2 days of treatment, resulting in some cells floating in the culture medium, most of which were stained by trypan blue; this effect displayed dose and time dependence. The cells remaining attached excluded the dye.



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  		Figure 2. Prolonged MPA treatment induces cell death. A, HIT cells on multiwell plates were treated with 3 µg/ml MPA for 48 h. Protein content, DNA content, and insulin content were determined by Bio-Rad assay, the DNA-binding fluorescent probe Hoechst 33258, and RIA, respectively, in parallel in the same experiments. Cell viability was examined by MTS test. B, HIT cells were treated with different concentrations of MPA for various time periods. Guanosine or adenosine was also included (in experiments using 1 µg/ml MPA for 48 h). Cell viability was examined by MTS test. All results are expressed as a percentage of the control value for comparison of the determinations between wells. Values are the mean ± SE from three or four experiments.

 
Total cell number was decreased by about 70% after 2-day treatment with 3 µg/ml MPA, with a comparable drop in contents of DNA, protein, and insulin (Fig. 2AGo). A sensitive assay of metabolic viability (the reduction of a tetrazolium salt to colored formazan) was also used. Formazan production from MTS was reduced to the same extent as cell number (Fig. 2AGo), whereas MTS reduction in those cells that were still viable (checked by trypan blue exclusion) was only decreased by 9% compared with the data from control cells with respect to actual cell number. Therefore, the MTS test was employed to monitor the cell death induced by prolonged MPA treatment at various doses and for different periods. Formazan production was not altered by 12-h treatment with 0.1–10 µg/ml MPA and was slightly decreased (~20%) after 24-h treatment at 3 and 10 µg/ml MPA (Fig. 2BGo). After 48-h treatment, formazan generation was reduced by 30% at 0.3 µg/ml and by 75% at 3–10 µg/ml MPA (Fig. 2BGo). Four-day MPA (1 µg/ml) treatment decreased formazan formation by 80%. The effects of prolonged MPA treatment could be prevented by inclusion of guanosine (Fig. 2BGo) in a dose-dependent manner (Table 1Go), whereas neither adenosine (up to 500 µM) nor deoxyguanosine (30–500 µM) had a preventive effect (Fig. 2BGo and Table 1Go). In preliminary studies, noxious effects of MPA were also seen after 4 days of culture at 4.4 mM glucose, similar to those seen after culture at 11.1 mM glucose.


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  		Table 1. Effects of nucleosides and other agents on metabolic viability in control and MPA-treated HIT cells

 
MZ also caused the death of HIT cells. Treatment of the cells for 4 days decreased formazan formation by 55%, 64%, and 81%, respectively, at 12.5, 25, and 75 µg/ml, accompanied by a parallel reduction in cell number and DNA content. Similar effects of prolonged exposure to MPA on induction of cell death were also seen in INS-1 cells, which possess properties more similar to those of primary ß-cells (17) than to those of HIT-T15 cells. However, this effect on INS-1 cells required higher doses of MPA (25 µg/ml) to both deplete GTP (9) and kill cells, as reflected by the loss of cells and the appearance of condensed nuclear staining by the DNA-binding dye propidium iodide (data not shown). The effects of MPA in INS-1 cells on GTP (9) and cell death were prevented by coprovision of 100 µM guanine, but not by adenine (data not shown), which are readily salvaged into purine nucleotides in these cells (9).

Induction of apoptosis in ß-cells by treatment with GTP-depleting agents
The nature of cell death induced by prolonged treatment with GTP-depleting agents was examined. During these studies, we observed that the dying cells were avidly stained by the DNA-binding dye propidium iodide after fixation (Fig. 3Go), suggesting the presence of programed cell death or apoptosis (1). In control HIT cells (Fig. 3aGo), staining by the DNA-binding dye was homogeneous. In MPA-treated cells, strong localized and condensed staining appeared (indicated by arrows in Fig. 3Go). Some small, strongly stained spots could be seen, presumably reflecting apoptotic bodies. The MPA effect was dose dependent; after 2-day treatment with 1 (Fig. 3bGo), 3 (Fig. 3cGo), or 10 µg/ml MPA (Fig. 3dGo), 29%, 49%, and 70% of the cells were affected, as judged by counting cells on photographs taken under fluorescence microscopy. This type of condensed and fragmented staining, seen after 24-h MPA treatment, was not observed after 6- and 12-h treatment. In INS-1 cells, similar results were seen after 25 µg/ml MPA treatment for 2 days (data not shown). The MPA effect was prevented by coprovision of 500 µM guanosine (Fig. 3fGo), whereas the effect of a lower dose (200 µM) of guanosine was incomplete. Adenosine (500 µM) again did not prevent MPA effect (Fig. 3eGo).



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  		Figure 3. Staining by DNA-binding dye after prolonged MPA treatment of HIT cells. HIT cells seeded on glass coverslips were fixed and permeabilized by ethanol. Subsequently, cells were incubated with the DNA-binding dye propidium iodide. Some MPA-treated cells were lost during washing. The samples were examined by a fluorescence microscope. a, Control; b–d, treated with 1, 3, and 10 µg/ml MPA for 48 h; e, treated with 1 µg/ml MPA plus 500 µM adenosine; f, treated with 1 µg/ml MPA plus 500 µM guanosine. Bar = 100 µm. Similar results were obtained in three experiments.

 
The effect of prolonged treatment of MPA on HIT cells was also examined via electron microscopy. In control cells (Fig. 4aGo), the chromatin is floccular and dispersed throughout nuclei; the nuclear membrane is intact. Different intact cellular organelles are visible, including mitochondria, and microvilli are visible at the plasma membrane. Little or no vacuolization of cytosol was seen. In MPA (3 µg/ml for 48-h)-treated cells (Fig. 4Go, b and c), the typical morphological changes of apoptosis were seen (1). The early changes included chromatin condensation and margination under the nuclear envelope with damage to the nuclear membrane (Fig. 4bGo). The mitochondria remained normal. Some dilated endoplasmic reticulum and cytoplasmic vacuoles were seen. Microvilli have disappeared. In Fig. 3cGo, one cell (open arrow) was in the early stages of apoptosis (condensed and marginated chromatin and also condensed cytoplasm), and two cells (solid arrow) were in the later stage of apoptosis. In these cells, nuclei and cytoplasm were further condensed, and apoptotic bodies were formed. When these results from electron microscopic observations were quantitatively analyzed, only 4% (1 of 27 cells) of control cells displayed the apoptotic changes. In MPA-treated cells, 25% (17 of 67) displayed chromatin condensation/margination, and 16% (11 of 67) had the appearance of apoptotic bodies.



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  		Figure 4. Prolonged MPA treatment induced apoptotic changes in HIT cells. After treatment with 3 µg/ml MPA for 48 h, HIT cells cultured on glass coverslips were fixed and examined by electron microscopy. a, Control; b and c, treated with 3 µg/ml MPA for 2 days. In c, one cell (open arrow) was at the early stage of apoptosis, and two cells (solid arrow) were at the later stage of apoptosis. Bars in a and b = 1 µm; bar in c = 2 µm.

 
During apoptosis, DNA is often degraded by endonucleases, and DNA fragments in multiples of approximately 180 bp (oligonucleosomes) are produced (22). These small, soluble DNA fragments can be extracted from the supernatants after high speed centrifugation (14,000 x g) and can be detected on 2% agarose gel by electrophoresis. Treatment of HIT cells with 1, 3, and 10 µg/ml MPA for 2 days resulted in a DNA laddering of multiples of about 180 bp in a dose-dependent manner (Fig. 5AGo). This effect of MPA on DNA degradation was mostly prevented by guanosine, but was unaffected by adenosine (Fig. 5BGo), indicating a specific effect of GTP depletion. Similar results were observed in another series of experiments using an ELISA kit detecting apoptosis (Fig. 6Go). The amounts of oligonucleosomes in both MPA-treated HIT and INS-1 cells were about 3 times their control values. Inclusion of guanosine, but not adenosine, prevented the MPA effect (Fig. 6Go). Thus, both ultrastructural and biochemical evidence indicates that prolonged depletion of GTP induces apoptosis of insulin-secreting cells.



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  		Figure 5. DNA degradation caused by prolonged MPA treatment. HIT cells in culture dishes were treated with test agents for 48 h. After washing, cells were scraped and lysed. Soluble DNA fragments were extracted with phenol. An equal fraction of extraction samples under different experimental conditions was loaded on a 2% agarose gel. After separation by electrophoresis, DNA was stained with ethidium bromide, and photographs were taken under UV light. A: Lane 1, Laddering of DNA (100 bp) standards (low mol wt fragments move further toward the bottom of the gel); lane 2, control cells; lanes 3–5, treated with 1, 3, and 10 µg/ml MPA, respectively. B: Lane 1, Laddering of 100 bp DNA; lane 2, control cells; lane 3, treated with 3 µg/ml MPA; lane 4, treated with 3 µg/ml MPA plus 500 µM adenosine; lane 5, treated with 3 µg/ml MPA plus 500 µM guanosine. Similar results were obtained in four experiments.

 


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  		Figure 6. Detection of apoptotic death of HIT and INS cells by an ELISA. HIT or INS cells in multiwell culture plates were treated with test agents for 48 h. After washing, cells were lysed, and extracts were centrifuged at 14,000 rpm for 10 min. The oligonucleosomes in supernatants were assayed using an ELISA kit. The concentrations of agents (where present) were: MPA, 1 µg/ml; adenosine, 500 µM; and guanosine, 500 µM. Values are the mean ± SE of three or four observations. *, P < 0.05 compared with MPA-treated alone.

 
In preliminary studies, MPA treatment was applied to intact rat islets for 18–67 h, and apoptosis was monitored by ELISA. MPA induced a small, but consistent, degree of apoptosis. In four studies of islets cultured for 18–42 h in 25–50 µg/ml MPA, indexes of apoptosis rose by 40% from 0.10 ± 0.03 to 0.14 ± 0.035 (P = 0.012 by paired t test). In a preliminary study over a more extended period of 67 h, apoptosis was increased by MPA treatment more markedly (3-fold) from 0.03 ± 0.01 to 0.09 ± 0.01 (P = 0.003, by t test). By way of comparison, after 18 h, S-nitrosoglutathione treatment (500 µM) produced values that were 167% of the control level, and NaF treatment (5 mM) produced values that were 208% of the control level. At 42 h, the values with 2.5 mM NaF increased further to 615% of the control level.

Ca2+-activated endonuclease(s) have been implicated in the DNA degradation of apoptosis (23). However, neither the endonuclease inhibitor (24) aurintricarboxylic acid (0.1–0.5 mM) nor the Ca2+ entry blocker Ni2+ (1 mM) affected MPA-induced cell death (Table 1Go), suggesting that a step(s) other than Ca2+-activated endonucleases may be the main mediator(s) in this event. It has been suggested that activation of protein kinase C mitigates apoptosis (25), possibly by counteracting the proapoptotic effects of ceramides. However, phorbol myristate acetate (a strong protein kinase C activator) had no apparent effect on cell death due to prolonged MPA treatment (Table 1Go). Lastly, the effect of inhibition of messenger RNA synthesis was examined, because there is evidence that synthesis of new proteins is involved in apoptosis (22). Actinomycin D (0.5–4 µM) alone reduced formazan production, but did not alter the effect of MPA, suggesting that new protein synthesis may not be necessary for MPA-induced cell death, or alternatively, that MPA by itself inhibited the synthesis of a protein that retards apoptosis.

Apoptosis induced by prolonged lovastatin treatment
Many ras-related small molecular GTP-binding proteins (SMGs) and other signaling proteins, such as {gamma}-subunits of trimeric GTP-binding proteins, undergo a cascade of posttranslational modifications required for full biological function (26). We previously observed (27) that GTP depletion inhibits the posttranslational activation of at least one SMG (namely Cdc42 in intact HIT cells). Similar findings have recently been observed to extend to the methylation of {gamma}-subunits of trimeric G proteins (28). Therefore, to investigate the possibility that cell death induced by GTP depletion involves dysfunction of GTP-binding proteins, the effect of lovastatin, an inhibitor of isoprenylation of GTP-binding proteins in ß-cells (5, 29, 30), on cell death was studied. As shown in Fig. 7AGo, 72-h treatment with 10–100 µM lovastatin reduced cell number in a dose-dependent manner, accompanied by a parallel reduction in formazan production. The cellular contents of DNA, protein, and insulin were also markedly decreased. All effects of lovastatin on these parameters were prevented by coprovision of mevalonate, which bypasses the blockade of 3-hydroxy-3-methylglutaryl coenzyme A reductase induced by lovastatin (26). When cell death was monitored by MTS assay (Fig. 7BGo), it was found that 24-h treatment with lovastatin was not sufficient to cause cell death; 48 h were required for induction of this event. Biochemical evidence also indicated that 72-h treatment with 10–100 µM lovastatin resulted in apoptosis, as indicated by DNA laddering of multiples of 180 bp (Fig. 8Go). Mevalonate (100 µM) prevented this DNA degradation to a large extent. These studies using GTP depletion or inhibition of isoprenoid precursors are together compatible with (but do not prove) an involvement of inhibitory G proteins (either SMGs or {gamma}-subunits of trimeric G proteins) in the induction of apoptosis.



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  		Figure 7. Cell death induced by long term lovastatin treatment. A, HIT cells on multiwell plates were treated with 10–100 µM lovastatin for 72 h. Mevalonate was also included where indicated. Protein, DNA, and insulin contents were determined by Bio-Rad assay, the DNA-binding fluorescent probe Hoechst 33258, and RIA, respectively. Cell viability was examined by the MTS test. B, HIT cells were treated with different concentrations of lovastatin for 24, 48, or 72 h. Cell viability was examined by MTS test. All results are expressed as a percentage of the control value for comparison of the determinations between wells. Values are the mean ± SE from two experiments.

 


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  		Figure 8. DNA degradation caused by long term lovastatin treatment. HIT cells were treated with test agents in culture medium for 72 h. Lane 1, Laddering of 100 bp DNA; lane 2, control cells; lanes 3–5, treated with 10, 30, and 100 µM lovastatin, respectively; lane 6, treated with 100 µM lovastatin and 100 µM mevalonate. Similar results were obtained in three experiments.

 
It has been reported that Rho proteins (a family of Ras-related GTP-binding proteins including Rho, Rac, and Cdc42) may play a modulatory role, either inhibitory or stimulatory (30, 31, 32, 33, 34, 35), in the induction of apoptosis in lymphoma cells and fibroblasts. To examine whether MPA- or lovastatin-induced death of insulin-secreting cells was mediated or modulated by Rho proteins (RhoA, Rac1, and Cdc42), the effect of C. difficile toxin B was studied. This toxin blocks the function of these SMGs via their glucosylation (36). HIT cells treated with the toxin alone (5–1000 ng/ml) for up to 72 h did not display cell death, as assessed by MTS assay; however, these cells did exhibit morphological changes (rounding-up), indicating that Rho family functions on the actin cytoskeleton (37) had been blocked in these intact cells. Thus, inactivation of Rho proteins alone was not capable of initiating apoptosis. However, toxin B significantly enhanced the cell death induced by either MPA or lovastatin (Table 2Go), suggesting a modulatory role of Rho proteins.


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  		Table 2. Effect of Clostridium difficile toxin B on cell death induced by MPA and lovastatin

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
There are two major types of cell death: necrosis and apoptosis (programed cell death) (1, 2). In contrast to necrosis, in which the cell (including cellular organelles) swells and releases cellular components, resulting in chemotaxis of neutrophils and the induction of inflammation, the typical morphological changes of apoptosis include blebbing of the cell surface, cell shrinkage, loss of microvilli, condensation and margination of chromatin, formation of apoptotic bodies, and engulfment by macrophages or adjacent cells without release of cellular components or inflammation.

In the present study, MPA or MZ impeded [3H]thymidine incorporation (mitogenesis) after as little as 1 h of treatment and even at low concentrations of MPA. Longer durations (1–4 days) and greater degrees of depletion of cellular GTP induced by MPA or MZ caused death with characteristics of apoptosis in either of two insulin-secreting ß-cell lines (HIT-T15 and INS-1), as indicated by several lines of evidence. First, cell number was decreased with parallel reductions in DNA, protein, and insulin contents. Second, condensation and fragmentation of nuclear DNA were demonstrated morphologically by fluorescent staining with a DNA-binding dye (propidium iodide) after fixation and permeabilization of cells (38). Third, biochemical evidence of apoptosis was provided by detection of the characteristic, soluble DNA fragments, which form a DNA ladder of multiples of about 180 bp (oligonucleosomes) (39). This latter finding was confirmed using a cell death detection assay. Finally, electron microscopic examination revealed most of the typical changes of cell apoptosis listed above.

We have previously documented in detail the effects of MPA on ATP, ATP/ADP, GTP, and GTP/GDP in both cell lines (9). In HIT-T15 cells, the contents of GTP and ATP fell by 71–74% and 30–43%, respectively, and the GTP/GDP ratio declined by 43–46% without significant alteration of ATP/ADP ratio after 18-h treatment with 1.6–6.3 µg/ml MPA (9). Linkage of apoptotic cell death to depletion of guanine nucleotides is clear, as all changes induced by MPA were prevented virtually completely by coprovision of guanosine (HIT cells) or guanine (INS-1 cells) [but not adenosine (HIT cells) or adenine (INS-1 cells)] at concentrations that restored the GTP content and GTP/GDP ratio to control levels (9). These findings were confirmed by the identical results seen using a second, structurally dissimilar (but mechanistically similar) immunosuppressive compound, MZ. Very recently, it has been reported that another IMP dehydrogenase (IMPDH) inhibitor, tiazofurin, induces apoptosis (in erythroleukemia cells) (40). It is possible that depletion of deoxy (d)-GTP plays a contributory role; indeed, a reduction in GTP will be accompanied by a decrease in dGTP due to its interconversion by ribonucleotide reductase (41). Depletion of dGTP may impair DNA synthesis (41) and thereby predisposes to apoptosis (42, 43). However, depletion of dGTP does not appear to be the main mediator of MPA-induced cell toxicity (41, 44), because either low (30 µM) or high (500 µM) concentrations of exogenous deoxyguanosine, which restore the dGTP content of cells even at 25 µM (41, 43, 44), failed to prevent the effect of MPA on the cell death.

There is evidence for the involvement of Ca2+-activated endonuclease in apoptotic cell death (23). Our preliminary data suggest that there occurs a small increase in basal level of cytosolic free Ca2+ after depletion of GTP by MPA (unpublished data). However, neither inhibition of Ca2+ entry using a non-selective Ca2+ channel blocker (Ni2+), nor an inhibitor of Ca2+-activated endonuclease, aurintricarboxylic acid (24) altered apoptosis induced by MPA, suggesting that GTP-depletion causes DNA degradation and the induction of apoptosis via other mechanisms. Rather, very recent studies in leukemia cells, reported while our studies were in progress, suggest that GTP depletion, induced pharmacologically, can directly impair DNA synthesis and induce apoptotic cell death probably by inhibiting RNA-primed DNA synthesis (45). Interestingly, the inhibition of inosine monophosphate induced by exogenous MPA may be mimicked by p53 (46, 47), an endogenous peptide inhibitor of IMPDH which acts as a tumor suppressor and promotes apoptosis (46, 47, 48). Indeed, pharmacologic inhibition of IMPDH may unmask the biological effects of p53 on growth, an effect reversed by overexpression of IMPDH (46, 47, 48). Therefore, our studies using MPA may transcend pharmacologic relevance and provide, in addition, insights into the regulation of growth and induction of apoptosis by physiologic molecules.

Depletion of cellular GTP by MPA and MZ also impeded mitogenesis, as assessed by [3H]thymidine incorporation. (We have previously observed a similar effect of MPA in neonatal (nontransformed) ß-cells, as assessed by bromodeoxyuridine labeling) (49). This effect in the current study occurred quite early (1 h); this is a time point at which GTP is significantly reduced (9), albeit to only a limited degree (to 39 ± 2% of the control value with 1 µg/ml MPA; n = 6; P < 0.001). The GTP/GDP ratio also falls to 46 ± 1% of the control value (n = 6; P < 0.001) under the same conditions. Thus, mitogenesis was very sensitive to depletion of GTP (and possibly dGTP). Indeed, it has been postulated that inhibition of a small (or compartmentalized) pool of nuclear GTP that rapidly turns over will block mitogenesis (41, 50). Similar observations have been reported in other cells (11, 45), including pancreatic islets (13). In contrast to induction of cell death and DNA degradation (see above and Refs. 44, 45), this effect seems to be partially dependent on depletion of dGTP, as it could be partially prevented by via coprovision of deoxyguanosine at a short time period (6 h). Alternatively, it is possible that deoxyguanosine acted solely via repletion of cellular GTP content (41, 44). Parenthetically we note that [3H]thymidine incorporation was inhibited by adenosine or guanosine alone (i.e. in the absence of MPA). This might be due to interference with the transport of nucleosides (i.e. [3H]thymidine), as similar concentrations of the nucleosides compete with each other for their transport and inhibit [3H]thymidine incorporation in some (but not all) cell systems (51). In addition, accumulation of purine nucleotides might inhibit ribonucleotide reductase and cause a depletion of pyrimidine deoxynucleotides (52), leading to inhibition of thymidine incorporation.

In contrast to mitogenesis, induction of cell death required a longer time (24 h) and a higher dose (>0.3 µg/ml) of MPA and a more prolonged inhibition of DNA synthesis, suggesting that other cellular events must occur to trigger apoptosis after a sustained depletion of GTP. Therefore, we have investigated additional mechanisms by which prolonged GTP depletion might induce apoptosis, in particular the possible involvement of trimeric or SMGs. Recent studies (30, 31, 32, 33, 34, 35, 53, 54) suggest that GTP-binding proteins may modulate the process of apoptosis. For instance, mastoparan3 (a potent stimulator of trimeric G proteins) can induce apoptotic changes (53), whereas stimulation of the muscarinic receptor (m3 subtype) blocked apoptosis, probably via a G protein, in neurons (54). Protein kinase C, which is one of the signaling pathway downstream of certain G proteins, also has been reported to influence apoptosis (25, 55), generally in an inhibitory way. Both Rho (30, 32, 33) and Ras (32, 55) have been implicated as modulators of apoptosis. We have observed that GTP depletion induced by MPA or MZ inhibits G protein-mediated signal transduction in normal rat islets, in association with inhibition of phospholipase C (8); this might be predicted to modify cell survival by the loss of activation of protein kinase C. However, the failure of phorbol myristate acetate to modulate MPA-induced apoptosis suggests that an inhibition of phospholipase C, with a concurrent reduction in diglyceride production and activation of protein kinase C, is not the mechanism of action of GTP depletion (45).

We have also carried out preliminary studies on a novel GTP protein (Gh) that has bifunctional properties; in its GTP-bound form it activates phospholipase C, whereas it acts as a tissue transglutaminase when GTP levels are low and the cytosolic Ca2+ concentration is high (56). It has been reported that an increase in transglutaminase activity might promote apoptosis (57). We did find both transglutaminase activity and the presence of tissue transglutaminase (also called transglutaminase II) by immunoblotting in islets and also in HIT cells (Kowluru, A., G. Li, and S.A. Metz, unpublished data). However, additional results (data not shown) suggest that this pathway may not significantly contribute to apoptosis induced by GTP depletion. First, monodansylcadaverine (100–300 µM) or glycine methylester (5 mM), two relatively specific transglutaminase inhibitors (56, 58), did not prevent MPA-evoked cell death; secondly, GTP depletion was not accompanied by any clear increase in total transglutaminase activity in ß-cells (data not shown).

We therefore turned to ras-related SMGs, which can modulate apoptosis in both a positive and a negative direction (30, 31, 32, 33, 34, 35, 53, 59, 60). It is possible that depletion of guanine nucleotides alters the function of one (or more) SMG that normally suppress programed cell death and thereby elicits apoptotic cell death. Indeed, our recent studies indicated, for the first time, that a marked depletion of ß-cell GTP content (and/or the GTP/GDP ratio) achieved by use of MPA or MZ impedes the function of at least one SMG of the Rho family, Cdc42 (27), as well as the activation of Ras (49) and may perturb the function of certain isoforms of subunit of the trimeric G proteins (28). Similar findings have been observed for Ras activation in K562 cells treated with tiazofurin (61), another inhibitor of GTP synthesis. Furthermore, one of the molecular targets of Cdc42/Rac guanosine triphosphatases (PAK) has been implicated as an inhibitor of apoptosis (62). Two of the experiments in the present study also suggest a possible link between GTP depletion-induced apoptosis and specific GTP-binding protein(s). The first involves lovastatin, a drug that inhibits the isoprenylation of SMGs by blockade of synthesis of mevalonate, the precursor of isoprenoids. It is well established that most SMGs (as well as the {gamma}-subunits of heterotrimeric G proteins) are isoprenylated at their C-termini; this isoprenylation is required for their proper function (26). Our previous work established that lovastatin (in short term treatment) inhibited isoprenylation of several small G proteins in islets or ß-cells; this effect was accompanied by redistribution of several small GTP-binding proteins (especially Cdc42) from their membrane sites to the cytosol and by aberrant insulin secretory responses (5, 29). In the present study, treating cells for 3 days with lovastatin induced apoptotic cell death, which was prevented by the coprovision of mevalonic acid. It has been recently reported that lovastatin also induced apoptosis in HL-60 cells, lymphocytes, glioma cells, or prostate cells (35, 63, 64, 65, 66, 67, 68, 69). We hypothesize that this effect represents at least in part blockade of the prenylation of (one or more) SMGs inhibitory to apoptosis. This might be analogous to observations that deletion of the bcl-2 gene can unmask apoptosis by removing inhibitory influences (55). However, we cannot exclude an alternative possibility, namely that the effects of lovastatin were mediated either by the reduction in synthesis of other derivatives of mevalonate or by inhibition of the prenylation of proteins other than G proteins (e.g. nuclear lamins). Therefore, the possible involvement of Rho proteins was specifically studied. Glucosylation of Rho proteins (RhoA, Rac1, and Cdc42) using C. difficile toxin B abolishes the function of these proteins (36). Although this treatment did not itself induce apoptosis, it significantly enhanced cell death induced by MPA or lovastatin. These results suggest that Rho proteins may possess properties that protect against apoptosis in insulin-secreting cells, as found in some other cells (30, 31, 62).

A large amount of experimental and clinical studies indicate that mycophenolate, when provided as its prodrug mycophenolate mofetil, is an immunosuppressive drug with considerable promise in prevention or reversal of rejection of transplanted grafts, probably due its relatively specific toxicity to lymphocytes (10, 11). In some studies of relatively short duration, gross negative effects of mycophenolate on ß-cell function (i.e. hyperglycemia) were not reported (70); however, systematic long term studies of glucose tolerance have not been performed. Our observations in this study indicate that prolonged treatment of MPA can induce a modest degree of apoptosis of normal (i.e. nontransformed) islet cells, even over short periods (2–3 days). Normal ß-cells do have a modest capacity for proliferation (71), and other ß-cell toxins, such as streptozotocin, have rapidly induced apoptotic cell death in intact normal rodent islets (72). Furthermore, even a very small amount of apoptotic ß-cell death can impair the ability of normal compensatory mechanisms to maintain an intact ß-cell mass (73), thereby predisposing toward diabetes. It has been reported that MPA has a harmful effect on glucose tolerance in animals (12). Therefore, it may be that the GTP depletion induced by prolonged therapy with mycophenolate mofetil will also impair ß-cell survival in humans either by inhibition of RNA-primed DNA synthesis (45) and/or activating p53 (74). Our results indicate that more studies in vivo are required to determine whether, like antecedent generations of immunosuppressives, MPA is toxic to ß-cells and could induce ß-cell depletion, which, in turn, might lead to a recurrence of an insulin-dependent state over time in patients with insulin-dependent diabetes who received a pancreas or islet graft. Furthermore, interleukin-1ß (which is widely accepted as a cytokine involved with ß-cell death in type 1 diabetes) profoundly reduces GTP in ß-cells (75) and induces apoptotic death (76). The current studies suggest that these events might be integrally related.


   Acknowledgments
 
We are grateful to Dr. M. Meredith for measuring purine nucleotides, and to Jim Stevens for technical assistance. We also thank Drs. R. P. Robertson and C. B. Wollheim for providing HIT-T15 and INS-1 cells, respectively, and Dr. K. Aktories for providing toxin B.


   Footnotes
 
1 This work was supported by the Office of Research and Development, Medical Research Service, the Department of Veterans Affairs (to S.A.M. and A.K.), NIH Grant DK-37312, the American Diabetes Association, a gift from the Rennebohm Foundation, and National Medical Research Council of Singapore Grant 6600010 (to G.L.). Back

2 Present address: Pacific Northwest Research Foundation, 720 Broadway, Seattle, Washington 98122. Back

3 However, in our hands, mastoparan (3 µM, 48 h) failed to induce cell death; 30 µM mastoparan induced a 32% reduction of formazan production with a 20% decrease in cell number. Back

Received May 28, 1998.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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