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Departments of
*
Molecular Microbiology and Immunology and
Physiology and Biophysics, University of Southern California Keck School of Medicine, Los Angeles, CA 90033;
Institute of Immunology, University Hospital, Hamburg, Germany; and
The Jackson Laboratory, Bar Habor, ME 04609
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Several hypotheses have emerged to explain actions of ecto-NAD on T cells. One hypothesis is that it serves as a substrate for an arginine-specific cell surface ADP-ribosyltransferase (ART-2),3 which by ADP-ribosylating cell surface proteins causes the observed effects (1). Several observations support this hypothesis; anti-CD3 activation of murine T cells releases ART-2 and causes the cells to become resistant to NAD (5, 6), and using recombinant means, deletion of murine T cell ART-2 isoforms (ART2a and ART2b) inhibits cell surface ADP-ribosylation (7). Several of the proteins incorporating ADP-ribosyl groups have been identified, including LFA-1, CD27, CD43, CD45, and CD8 (8, 9), raising questions as to whether modification of one or more of these proteins induces cell death. It has been proposed that effects of ecto-NAD may arise from an activating ADP-ribosylation of purinergic receptors (10). It has also been proposed that ecto-NAD leads to an inactivating ADP-ribosylation of CD38, a cell surface NAD-glycohydrolase that can synthesize cyclic ADP-ribose (ADPR) (3). Yet another hypothesis arises from the observation that ecto-NAD inhibits mitogen-stimulated proliferation of rat T cells, which express little or no ART-2, and that, for rat T cells, adding ADPR is as effective as adding NAD. Because rat T cells degrade NAD to ADPR, the hypothesis arose that ADP-ribosylation may not be involved in the effects arising from the addition of NAD to cultured T cells but rather that NAD breakdown products act as ligands for purinergic receptors (11).
The data reported herein prompt a unifying model consistent with the above observations. The data demonstrate that adding NAD to T cells induces both rapid and slow death involving different sets of mechanisms. Rapid death involves pore formation and signaling through P2X7 receptors while slower death requires caspase activation but not P2X7 receptors. It is suggested that P2X7 engagement of ADP-ribosyl groups, attached by action of ART-2 to cell surface proteins, triggers receptor function and rapid cell death. This mechanism may provide a means by which NAD released during trauma controls T cell functions.
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Materials and Methods |
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Pathogen-free female C57BL/6 (B6), and BALB/c mice, 6–8 wk of age were obtained from The Jackson Laboratory. NOD.129S4(B6)-Art2aTm1FknArt2bTm1Fkn (henceforth denoted as ART-2–/– mice) were produced at The Jackson Laboratory by outcross to a previously described B6 stock carrying targeted mutations in the tandem genes encoding the ART-2.1 and ART-2.2 ectoenzymes (7). Following nine backcrosses to NOD/Lt (N10) and verifying presence of NOD alleles at all known "Idd" loci, intercrosses were initiated, and ART2–/– homozygous breeding stock were received and bred at the University of Southern California animal facility (Los Angeles, CA). B6 P2X7–/– mice were kindly provided by Dr. C. Gabel (Ann Arbor, MI) and Pfizer and were bred at the University of Southern California animal facility (12). T cells were purified from spleen cells by nylon wool nonadherent (NWNA) cells and cultured in complete RPMI 1640 medium, containing 10% FBS (2). By FACS analysis, 85–90% of the NWNA cells were CD3 positive. For FACS analysis, T cells were preincubated with anti-mouse CD16/CD32 (2.4G2) mAb from BD Biosciences to block Fc
Rs, and then incubated with various mAbs for 30 min at 4°C (2). Phenotypic analysis was performed using PE Cy5-conjugated anti-mouse CD3 (145-2C11) (BD Biosciences). To monitor induction of cell death, cells were stained with the Annexin V-FITC Apoptosis Detection Kit I (BD Biosciences). To demonstrate DNA fragmentation, cells were assayed using the TUNEL APO-Direct Apoptosis Assay kit (BD Biosciences). To quantitate cell surface ADP-ribosylation, NWNA cells (4 x 106 cells/ml in RPMI 1640) were incubated with etheno-NAD (Sigma-Aldrich), followed by incubation with etheno-ADPR-specific Ab 1G4, kindly provided by Dr. R. Santella (Mailma School of Public Health of Columbia University, New York, NY) (13) and a FITC goat anti-mouse Ig (BD Biosciences). FACS analysis was performed on a FACSCalibur (BD Biosciences).
Quantification of NAD metabolites
NWNA B6, BALB/c, and ART2–/– T cells (2 x 106) were incubated in 50 µl of serum-free PBS containing 200 µM NAD (Sigma-Aldrich) and 0.1 µCi [32P]NAD (Amersham) at 37°C. After centrifugation, supernatant from each sample (2 µl) was removed and applied to a 20 x 20 cm PEI-cellulose F glass-backed TLC plate (EMD Chemicals). TLC was developed in 0.3 M LiCl (Sigma-Aldrich) and exposed to a phosphor screen (Amersham) for 3 h at 25°C. NAD and ADPR were visualized using a PhosphorImager 445SI (Amersham), and radioactivity was quantified by ImageQuant 5.0 (Amersham) (11).
Western blot analyses
NWNA cells were incubated in RPMI 1640 with or without etheno-NAD (300 µM) for 30 min at 37°C. After washing twice in ice cold PBS, cells were re-suspended in lysis buffer (108 cells/ml PBS containing 1% Igepal, 1 mM AEBSF) and incubated for 30 min at 4°C. Insoluble material was removed by centrifugation (15 min, 10,000 x g), and supernatants were mixed with sample buffer, heated for 8 min at 95°C, and then loaded on SDS-PAGE gels. After electrophoresis, proteins were transferred onto PVDF membrane (Bio-Rad). Membranes were blocked with blocking buffer (LI-COR) for 1 h at room temperature. Blots were then incubated with 1G4 Ab (13) (ascites 1/500 dilution in TBS, 1% BSA, 0.05% Tween 20) for 2 h at room temperature. Membranes were washed with 0.05% Tween 20 in TBS and incubated with a 1/5000 dilution of Alexis Fluor 680-conjugated secondary anti-mouse IgG Abs (Molecular Probes) for 1 h. Protein bands were visualized by an infrared image system (LI-COR).
Cell proliferation and cell death assays and inhibition of caspases and P2X7 receptor function
To assay cell proliferation, NWNA cells were cultured on anti-CD3-coated tissue culture plates (2). Plates were incubated with a 1/1000 dilution of anti-CD3 Ab (500AA2 ascites) overnight. After washing the plates, T cells (5 x 105/well) were added in complete RPMI 1640 medium containing 10% FBS, and incubation was continued for 48 h. [3H]Thymidine (0.5 µCi/well) was added during the last 18 h of culture.
To assay induction of cell death, NWNA T cells were incubated with or without NAD, etheno-NAD, ADPR, or ATP (Sigma-Aldrich) for various times and assayed for propidium iodide (PI) uptake, annexin V, or TUNEL staining. To assay short-term effects of NAD (
5 min), a 10-fold excess ice-cold PBS was added to the cell suspension to dilute NAD. Following centrifugation, cells were washed twice in ice-cold PBS before culture in complete RPMI 1640 medium. Assays up to 2 h were done in complete RPMI 1640 medium without FBS, while those for 24 h contained 10% FBS. Caspase inhibitor Z-VADfmk (BD Biosciences) was used at 50 µM and added to cell cultures 1 h before addition of NAD (14). To inhibit P2X7 receptors, 20 µM KN-62 (Sigma-Aldrich) dissolved in 0.01% DMSO were added 10 min before addition of NAD or ATP (15).
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Results |
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Adding NAD to either B6 or BALB/c T cells leads to cell surface ADP-ribosylation (2, 4) and increases annexin V staining, but there are significant differences. B6 T cells undergo more extensive cell surface ADP-ribosylation than BALB/c T cells (1, 4), but a higher proportion of BALB/c T cells is induced for cell death (4). These differences become evident when purified T cells are incubated with ART-2 substrate etheno-NAD and then stained with etheno-ADPR-specific Ab 1G4 (13) (Fig. 1A). BALB/c T cells show staining when compared with cells from ART-2 gene-deleted mice (7), but it is much less pronounced then that of B6 T cells. Concordant results are seen in immunoblots. Cell extracts from BALB/c show barely detectable bands, whereas those from B6 T cells show strong bands (Fig. 1B). These differences between B6 and BALB/c T cells provide an experimental system to study the pathways responsible for NAD-induced cell death.
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Next, the ability of NAD to induce T cell death at 24 h was assayed. Fig. 2 shows that 100 µM NAD increases annexin V staining in the PI-negative B6 T cell population from 5.8 to 81.4%, but there is no significant change in the percentage of PI staining cells. For BALB/c T cells, 100 µM NAD increases annexin V staining from 2.2 to 42.6% in the PI-negative population, but in contrast to B6 T cells, there is an increase from 48.9 to 82% PI staining cells. Therefore, NAD inhibits T cell proliferation and induces annexin V staining in both B6 and BALB/c T cells, but whereas NAD induces PI permeability in BALB/c T cells, this is not the case in B6 T cells.
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Rat T cells hydrolyze NAD within 10 min of incubation (11) suggesting mouse T cells might do the same. Fig. 3 shows that hydrolysis to ADPR does occur but does not quite reach 50% by 2 h. Therefore, NAD hydrolysis by mouse T cells is slower than that by rat T cells. Also of note is that there are no differences between B6, BALB/c, and ART-2–/– T cells, supporting the notion that CD38 and not ART-2 is the principal NAD hydrolyzing cell surface enzyme (16). The finding that NAD is degraded during the time frame of our assays raises the possibility that NAD metabolites cause the effects ascribed to NAD.
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Therefore, it was important to examine effects of ADPR on T cell survival. Fig. 4 shows that 1000 µM ADPR increases annexin V staining in the PI-negative B6 T cell population from 4.3 to 20.7% and PI staining from 44.7 to 78.6% at 24 h. This result is quite different from that seen with NAD in Fig. 2 in which there was a dramatic increase in annexin V staining but no increase in PI staining. In BALB/c T cells, ADPR fails to increase annexin V staining, and the PI staining cell population increases by only 12%. Again, this result contrasts to the dramatic effects of NAD in BALB/c T cells (Fig. 2). Therefore, it appears that while high concentrations of ADPR inhibit cell proliferation in both BALB/c and B6 T cells, only in B6 T cells does ADPR induce a significant increase in annexin V and PI staining. Thus, suppression of cell proliferation by ADPR does not correlate with cell death, and the effects of NAD on T cells cannot be solely explained by actions of its metabolite ADPR.
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The finding that adding NAD and ADPR to B6 T cells is associated with different degrees of annexin V and PI staining at 24 h (Figs. 2 and 4) suggests that the two molecules promote cell death by different mechanisms. To examine this, the time course of cell death was examined in B6 T cells after the addition of either NAD or ADPR. Fig. 5 shows that 2 min after adding NAD, annexin V and PI staining reached a plateau. In contrast, attempts at demonstrating cell death by ADPR at these early time points were unsuccessful (data not shown). Therefore, NAD-induced cell death appears to be rapid while that induced by ADPR takes many hours. Results with BALB/c T cells confirm that NAD-induced cell death is exceedingly rapid as annexin V and PI staining plateau as early as 30 s after addition of NAD (Fig. 5). These results strongly suggest that NAD and ADPR induce cell death by different mechanisms and that B6 and BALB/c T cells differ in their sensitivity to these two molecules.
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In light of the above results, it was of interest to identify mechanisms involved in inducing rapid and slow cell death. Cell death often proceeds by apoptotic mechanisms, and a hallmark of apoptosis is the activation of caspases. Therefore, we tested whether inhibition of caspases interferes with rapid or slow induction of cell death. B6 T cells were first incubated with caspase inhibitor Z-VADfmk and then different concentrations of NAD were added. Fig. 6A shows that after 24 h, compared with no addition, adding NAD increases the PI-negative, annexin V-positive cell population from 5.1 to 81.2%, and that Z-VADfmk reduces staining to 26.7%. Therefore, the mechanism by which NAD induces slow cell death appears to involve the action of caspases. Because slow cell death is also seen with ADPR, we assayed effects of Z-VADfmk on ADPR-induced cell death. Fig. 6A shows ADPR increases the PI-negative annexin V-positive population from 5.1 to 19.9%, which is suppressed by the inhibitor. ADPR also increases the PI staining population, i.e., from 45.6 to 78%, which is decreased to 52.2% by Z-VADfmk (Fig. 6A). These results suggest that the slow induction of cell death by both NAD and ADPR involves caspases.
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To further test the hypothesis that apoptosis is not involved in the rapid induction of death, arising from the addition of NAD, T cells were assayed for DNA fragmentation. Fig. 7 shows that 2 h after the addition of NAD or ADPR there is no increase in DNA fragmentation assayed by TUNEL staining in either B6 or BALB/c T cells. During this time period, annexin V staining increased from 3.5 to 19.7% in B6 cells and from 3.2 to 58.9% in BALB/c T cells. Therefore, cell death in the first 2 h after addition of NAD does not involve DNA fragmentation. In contrast, TUNEL staining increased significantly in BALB/c and B6 T cells 24 h after the addition of NAD (Fig. 7). Adding ADPR instead of NAD produced similar but less dramatic effects after 24 h (Fig. 7). Therefore, both NAD and ADPR induce slow cell death by an apoptotic pathway, associated with DNA fragmentation.
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The finding that NAD but not ADPR induces rapid cell death is consistent with the notion that ADP-ribosylation is involved. However, because NAD also induces slow cell death, the question arises whether ADP-ribosylation contributes to the slow induction of cell death.
To find out, cell death was examined in T cells from wild-type NOD and NOD ART-2 gene-deleted mice. Fig. 8A shows that 2 h after adding NAD to NOD ART-2–/– T cells, there is no increase in annexin V staining shown in the dot plots or PI staining represented by percentages below the plots. In contrast, NAD effectively induces staining in T cells from wild-type NOD mice. Therefore, rapid cell death is not observed in T cells lacking ART-2, consistent with the requirement of ADP-ribosylation.
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Purinergic receptor P2X7 is required for NAD to induce rapid but not slow cell death
The finding that NAD degradation product ADPR mimics some of the effects induced by NAD, points to the involvement of purinergic receptors. Among the family of purinergic receptors, P2X7 is widely expressed on cells of the hemopoietic cell lineage (17, 18, 19, 20). Recent data also suggests that P2X7, whose well-established ligand is ATP, expresses low affinity for ADP (21). Moreover, inhibitors of P2X7 receptors can block induction of cell death by NAD (10). Taken together, these observations make P2X7 receptors a prime candidate to mediate effects of NAD and ADPR. To examine whether the rapid or slow mechanism of cell death, induced by NAD or ADPR, involve the P2X7 receptor, use was made of the finding that KN-62 inhibits within minutes the signaling function of this receptor (15, 22). BALB/c T cells were incubated with KN-62 for 10 min, followed by addition of NAD; 15 min later, T cells were stained for annexin V and PI. Fig. 9A shows that KN-62 completely inhibits NAD-induced annexin V and PI staining in this short-term assay. Similar results were obtained with B6 T cells (data not shown). However, we were not able to test KN62 in the slow induction of death because of its toxicity (data not shown).
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P2X7 receptor function is not impaired in etheno-NAD-treated cells
Two hypotheses arise to explain how cell surface ADP-ribosylation induced by NAD leads to rapid cell death. One is that the ADP-ribosylation of a protein modulates its function. The other is that ADP-ribosyl groups serve as ligands for a receptor. To test these hypotheses, we assessed effects of ADP-ribosylation on P2X7 receptor-triggered cell death by its ligand ATP. As previously reported (13), incubating T cells with etheno-NAD leads to etheno-ADP-ribosylation (Fig. 1, A and B) but not cell death. In fact, etheno-ADP-ribosylation renders T cells resistant to actions of NAD (10), presumably by filling the sites that are ADP-ribosylated upon the addition of NAD. Fig. 10 shows that while incubation of BALB/c T cells with etheno-NAD does not induce annexin V or PI staining, it does block the action of a subsequent addition of NAD. In contrast, etheno-NAD only slightly blocks rapid induction of annexin V by ATP and PI staining. These results are consistent with the notion that cell surface ADP-ribosyl groups act as ligands for P2X7 receptors, but not with the possibility that ADP-ribosylation acts by modifying protein function.
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Discussion |
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Another question is whether induction of death arises from NAD itself or its breakdown products. The results above confirm the observation (11) that T cells substantially hydrolyze NAD to ADPR, and that upon prolonged incubation ADPR exerts effects similar but not identical to those induced by NAD. It should also be noted that ADPR is metabolized to adenosine (25), which could act on T cells via action on purinergic receptors. Indeed effects of adenosine on T cells via action on A2A receptors are well documented in the literature (for review see Ref.26). Therefore, effects induced by the addition of NAD, particularly in incubations lasting 24 h, may very well result from the combined effects of NAD and its catabolic derivatives.
This report demonstrates that NAD can induce annexin V and PI staining in T cells by different mechanisms that lead to either rapid or slow cell death. Rapid cell death is not inhibited by caspase inhibitor Z-VADfmk, and therefore, is reminiscent of necrosis. Slow cell death is associated with DNA fragmentation and is inhibited by caspase inhibitor Z-VADfmk, and therefore, has features of apoptosis. It should be noted that while both NAD and ADPR slowly induce apoptotic death, they do not appear to act by identical mechanisms. This again may be due to action of NAD metabolites on purinergic receptors such as A2A receptors or others yet to be identified (26)
It is interesting that adding NAD to rat T cells leads to autoADP-ribosylation of cell surface ART-2 but apparently no other proteins (27, 28). Although adding either NAD or ADPR to rat T cells inhibits their proliferation, NAD does not induce a rapid increase in annexin V staining (28). Therefore, it appears that autoADP-ribosylation of cell surface ART-2 is not sufficient to induce rapid T cell death.
An enigmatic question is how NAD inhibits T cell functions and induces cell death. Initially, it appeared that effects observed after adding NAD to T cells correlated with cell surface ADP-ribosylation (2, 4). Subsequent data showed this correlation is not a tight one. B6 T cells undergo substantially more ADP-ribosylation than BALB/c T cells, yet NAD is less effective in promoting rapid cell death in B6 cells. An important difference between BALB/c and B6 mice is the differential expression of the two ART-2 genes ART-2a and ART-2b, which could be responsible for this effect. BALB/c mice express both ART-2a and ART-2b, whereas B6 mice express ART-2b only, because of a mutation in ART-2a (4). This raises the possibility that ADP-ribosylation by the ART-2a coded enzyme is more efficient in cell death induction then that by the ART-2b enzyme. However, this does not appear to be the case because NZW mice expressing only ART-2a possess T cells even less sensitive to NAD than B6 mice T cells (4). Therefore, it was important to demonstrate that ADP-ribosylation is critical for the induction of cell death. We show that deletion of the ART-2 genes in NOD mice blocks the rapid induction of cell death by NAD, consistent with results generated previously in a different mouse strain (7).
We present experimental evidence that deleting expression of the P2X7 gene blocks rapid cell death induction by NAD. These data prompt a plausible explanation for the apparent lack of correlation between the extent of T cell ADP-ribosylation and induction of rapid cell death in T cells from B6 and BALB/c mice. BALB/c T cells express the wild-type P2X7 receptor, which mediates efficient and rapid cell death upon binding ATP. In contrast, the B6 P2X7 receptor has a mutation in the TNFR1 deathlike domain resulting in much less efficient induction of cell death (20). Therefore, it appears that reduced induction of cell death by NAD in B6 compared with BALB/c T cells arises from the mutation altering the function of the P2X7 receptor in B6 mice.
Fig. 11 shows a model for the role of P2X7 receptors in mediating the effects arising from addition of NAD to cultured T cells. Using NAD, ART-2 attaches ADP-ribosyl groups to a cell surface protein near the P2X7 receptor or to the receptor itself. The ligand-binding site of the receptor engages the ADP-ribosyl groups, rapidly inducing cell death by pore formation. The data presented show that the death signal is induced within 30 s of NAD contact. Given the evidence that induction of the death signal involves activation of P2X7 receptors, it is likely that the formation of membrane pores is preceded by calcium fluxes. Indeed, incubation of mouse T cells with NAD induces calcium fluxes (10). Similar results were recently reported with human monocytes in which both NAD and ADPR induce calcium fluxes, but it was not determined which receptors may be involved (29). But, as previously demonstrated, the P2X7 receptor is also capable of inducing a slower, caspase-dependent apoptotic cell death (30, 31). Attachment of etheno-ADP-ribosyl groups fails to trigger rapid death via P2X7 activation due to the structural specificity of the P2X7 binding site, and by filling potential sites for ADP-ribosylations, etheno-NAD blocks the action of subsequently added NAD. However, after etheno-ADP-ribosylation, P2X7 receptors remain functional as ATP can still induce rapid cell death. In addition, metabolites of NAD and ADPR may react with other receptors that lead to slow cell death. Exposure of phosphatidyl serine, and thus binding of annexin V, characterizes the induction of both rapid and slow cell death.
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The question arises as to whether the model proposed above explains observations in earlier reports. Consistent with our model may be the observation that depleting rats or mice of T cells expressing ART-2 accelerates development of autoimmune diabetes and lupus erythematosus (34, 35). This could be explained by a mechanism in which NAD released from cells controls effector T cells. However, we have observed that deletion of the tandem ART-2 genes in the NOD mouse strain used in the above experiments (36) has no effect on diabetes progression (data not shown), even though lymphocytes from the parent NOD strain are particularly sensitive to signaling through P2X7 receptors (37). Taken together, these observations indicate the need to further study the role of ecto-NAD and its metabolites in the induction of cell death.
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Acknowledgments |
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Footnotes |
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1 This work was supported by Public Health Service Grants AI 40038 and AI 43954. 
2 Address correspondence and reprint requests to Dr. Gunther Dennert at the current address: University of Southern California/Norris Comprehensive Cancer Center, P.O. Box 33800, 1441 Eastlake Avenue, M/S 73, Los Angeles, CA 90033-0800. E-mail address: dennert{at}usc.edu
3 Abbreviations used in this paper: ART, ADP-ribosyltransferase; NWNA, nylon wool nonadherent; ADPR, ADP-ribose; PI, propidium iodide.
Received for publication September 21, 2004. Accepted for publication November 18, 2004.
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2 domain of murine major histocompatibility complex class I that specifically kills activated lymphocytes and blocks liver damage in the concanavalin A hepatitis model. J. Exp. Med. 198:497.
and IL-18 requires priming by lipopolysaccharide and is independent of caspase-1 cleavage. J. Biol. Chem. 276:3820. release from microglial cells through activation of ATP-primed P2X7 receptor channels. J. Neurosci. 22:3061. release. J. Immunol. 170:5728.This article has been cited by other articles:
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