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Transcriptional Program of Apoptosis Induction following Interleukin 2 Deprivation: Identification of RC3, a Calcium/Calmodulin Binding Protein, as a Novel Proapoptotic Factor

Howard Hughes Medical Institute, Programs in Gene Function and Expression and Molecular Medicine, University of Massachusetts Medical School, Worcester, Massachusetts 01605

Received 17 October 2002/ Returned for modification 28 January 2003/ Accepted 7 April 2003

	ABSTRACT

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Apoptosis of mature T lymphocytes preserves immune system homeostasis by counteracting transient increases in T-cell number. This process is regulated, at least in part, by the cytokine interleukin 2 (IL-2): T cells deprived of IL-2 undergo apoptosis. The mechanism of apoptosis induction by IL-2 deprivation remains to be determined but is known to require RNA synthesis, implying the existence of transcriptionally activated genes whose products induce cell death. To identify such genes, we have performed expression profiling in IL-2-dependent T cells following cytokine deprivation. Our results reveal an intricate transcriptional program entailing the induction of known proapoptotic factors and the simultaneous repression of known antiapoptotic factors. Surprisingly, one gene whose transcription substantially increased was RC3 (also called neurogranin), which encodes a calmodulin binding protein thought to be a neural-specific factor involved in learning and memory. We show that ectopic expression of RC3 in IL-2-dependent T cells increases the intracellular Ca²⁺ concentration and induces apoptosis even in the presence of cytokine. Buffering the Ca²⁺ increase with the cytoplasmic Ca²⁺ chelator BAPTA-AM [1,2-bis(2-aminophenoxy)ethane-N,N,N1,N-tetraacetic acid] blocks RC3-induced apoptosis, indicating that the rise in intracellular Ca²⁺ is required for apoptotic death. RC3 mutants unable to bind calmodulin fail to increase intracellular Ca²⁺ levels and to induce apoptosis. Based upon these results, we propose that IL-2 deprivation raises the level of RC3 and other apoptotic factors, which induce apoptosis by increasing the intracellular Ca²⁺ concentration.

	INTRODUCTION

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A complex network of cytokines controls the development, maturation, homeostasis, and responses of the immune system. An immune response typically involves a large increase in activated T cells, which must be eliminated following termination of the response to preserve homeostasis. This is achieved by apoptosis (also called programmed cell death), which can result from either cytokine deprivation or T-cell receptor activation (19). The cytokine interleukin 2 (IL-2) is produced and secreted by activated T cells and controls lymphocyte survival and cell cycle progression (17), and paradoxically it can also facilitate a particular apoptotic program in T lymphocytes termed activation-induced cell death (25). When deprived of IL-2, cycling T cells cease to proliferate and rapidly undergo apoptosis (19).

A key component of the apoptotic machinery is a proteolytic system involving an evolutionarily conserved family of cysteine aspartate proteases called caspases. Although this machinery is common to many forms of apoptosis, the precise mechanisms by which it becomes activated can vary considerably. Two pathways leading to caspase activation have been characterized: the extrinsic pathway, which involves so-called death receptors, and the intrinsic pathway, which involves the release of proapoptotic proteins from the mitochondria. In addition, some apoptotic pathways are transcription dependent, whereas others entail a purely posttranscriptional signal transduction cascade.

There have been several proposals for the mechanism by which IL-2 deprivation leads to apoptosis, including regulation of Bcl-2 family members, modulation of second messengers, and alteration of the activity of protein kinases or phosphatases (1, 5, 11). However, none of these regulatory events explain the requirement of de novo RNA or protein synthesis for induction of apoptosis following IL-2 deprivation (10). Here we use expression profiling to identify death-promoting genes that are transcriptionally activated in lymphocytes following IL-2 deprivation.

	MATERIALS AND METHODS

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Plasmids and cell culture. RC3 cDNA was hemagglutinin tagged at the 3' end and cloned into pcDNA3.1+ and pcDNA3.1- vectors (Invitrogen). IL-2-dependent murine HT-2 (T helper) cells, CTLL-2 (cytotoxic T) cells, myeloid 32D cells, HEK-293 cells, and human T-cell leukemia Jurkat cells were all obtained from the American Type Culture Collection; IL-3-dependent murine FL5.12 cells (pro-B lymphocytes) were provided by Craig Thompson (University of Pennsylvania). HT-2 and CTLL-2 cells were maintained in RPMI 1640 medium (GIBCO-BRL) supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine (GIBCO-BRL), 1 mM sodium pyruvate (GIBCO-BRL), 100 U of penicillin (Sigma) per ml, 100 µg of streptomycin (Sigma) per ml, 0.05 mM 2-mercaptoethanol (GIBCO-BRL), and 200 U of recombinant IL-2 (PharMingen) per ml or 10% rat T-STIM factor with concanavalin A (Becton Dickinson). FL5.12 and 32D cells were maintained in RPMI medium supplemented with 10% FBS, 1 mM L-glutamine, 100 U of penicillin per ml, 100 µg of streptomycin per ml, 0.05 mM 2-mercaptoethanol, and 3 ng of recombinant IL-3 (PharMingen) per ml. HEK-293 cells were maintained in Dulbecco's modified Eagle's medium supplemented with 10% FBS, 1 mM L-glutamine, 100 U of penicillin per ml, and 100 mg of streptomycin per ml. Jurkat cells were maintained in RPMI medium supplemented with 10% FBS, 1 mM L-glutamine, 100 U of penicillin per ml, and 100 mg of streptomycin per ml.

Human peripheral blood lymphocytes from healthy donors were initially cultured in RPMI medium supplemented with 10% FBS, 1 mM L-glutamine, and 5 µg of phytohemagglutinin (Sigma) per ml for 2 days. Cells were then collected and placed in RPMI containing 10% FBS and 10% human IL-2 (Advanced Biotechnologies) for 4 days with a medium change every 48 h. Staurosporine (Sigma) and thapsigargin (Calbiochem) were added to the media to final concentrations of 10 and 2 µM, respectively. BAPTA-AM [1,2-bis(2-aminophenoxy)ethane-N,N,N1,N-tetraacetic acid] (Sigma) was added to the media to a final concentration of 25 µM.

Microarray analysis. HT-2 cells were cultured in the presence of IL-2 and subjected to IL-2 deprivation as described previously (10). Poly(A)⁺ mRNA was isolated at 8 h following IL-2 withdrawal by using an Oligotex direct mRNA isolation kit (Qiagen). The integrity of mRNA was monitored by a Northern blot assay with a glyceraldehyde-3-phosphate dehydrogenase gene (GAPDH) probe. Two micrograms of mRNA was used to generate a cDNA library by using an oligo(dT) T7 primer and Superscript cDNA synthesis system (GIBCO-BRL) according to the manufacturer's instructions. The cDNA library was in vitro transcribed with biotinylated nucleotides by using the T7 Megascript kit (Ambion) according to the manufacturer's instructions. The resulting cRNA was fragmented by heating at 94°C for 30 min. GeneChip Murine 19K and 11K Affymetrix oligonucleotide arrays representing 30,000 known genes or expressed sequence tags were hybridized with fragmented cRNA overnight at 37°C. Following washing, the chips were scanned and analyzed with Affymetrix microarray suite software. The complete data set is available upon request.

Transient transfections and apoptotic assays. HT-2, CTLL-2, and FL5.12 cells were transfected with plasmids by using Superfect transfection reagent (Qiagen) according to the manufacturer's instructions. Cells were collected 24 h after transfection and stained with annexin V-fluorescein isothiocyanate (FITC)-propidium iodide (Oncogene). For cotransfection experiments, HT-2 cells were cotransfected with 1 µg of enhanced green fluorescent protein (GFP) plasmid (Clontech) and 4 µg of pcDNA3.1+/RC3 (sense orientation) or pcDNA3.1-/RC3 (antisense orientation). Cotransfected cells were stained with annexin V-phycoerythrin (PE) (PharMingen) and analyzed by fluorescence-activated cell sorting (FACS) analysis. At least two independent experiments were performed, and the results are presented as the averages ± standard deviations.

For HEK-293 fibroblasts, cells were seeded into a Lab-Tek II chamber slide (Nunc) at 6 x 10⁵ cells per slide and cotransfected with 0.5 µg of ß-galactosidase reporter plasmid (Clontech) and 4 µg of expression plasmid (as indicated), using Superfect transfection reagent. Following transfection, cells were washed three times with phosphate-buffered saline, fixed with 0.5% glutaraldehyde, and stained with 0.2% 5-bromo-4-chloro-3-indolyl-ß-D-galactoside (X-Gal) (Promega) as described previously (20). The percentage of transfected (blue) cells with apoptotic morphology was determined. At least two independent experiments were performed, and the results are presented as the averages ± standard deviations.

For DNA fragmentation analysis, DNA from 2 x 10⁶ cells was isolated by phenol extraction 24 h following transfection and analyzed on a 1% agarose gel as previously described (32).

Fluorospectrometric analysis of intracellular calcium concentration. To measure intracellular Ca²⁺ concentrations, 5 x 10⁶ HT-2 cells were stained with 2 mM fura-2 AM (Molecular Probes), a fluorescent Ca²⁺ indicator, in 1 ml of saline containing 1 mM CaCl₂. Cells were stained for 30 min at 37°C and for 10 min at room temperature. The cells were washed two times with saline containing 1 mM CaCl₂ and resuspended at 10⁶ cells/ml in the same solution. Spectrometric analysis was performed at 380 and 340 nm as described previously (36). The Ca²⁺ ionophore ionomycin was added to a final concentration of 10 mM at the end of each experiment to determine whether the limits of saturation of fura-2 AM had been reached. At least two independent experiments were performed, and the results are presented as the averages ± standard deviations.

Single-cell Ca²⁺ imaging. Intracellular Ca²⁺ was measured in primary human T lymphocytes and in HT-2 cells transfected with the RC3 expression plasmid. HT-2 cells were cotransfected with 5 µg of either pcDNA3.1+/RC3 or pcDNA3.1 and a plasmid expressing the truncated human CD4 molecule as a marker to select transfected cells. Transfected cells were then isolated from untransfected cells by using the MACSelect-transfected cell isolation kit (Miltenyi Biotech) according to the manufacturer's instructions. Transfected cells were then stained with 3 µM fura-2 AM in RPMI for 20 min at room temperature. Cells were washed three times with RPMI and plated onto poly-D-lysine-coated glass-bottom dishes (Mattek Corporation) for 10 min at room temperature. Cells were washed once with Ringer lactate solution. Images were collected at 340- and 380-nm wavelengths. Single-cell measurements of intracellular Ca²⁺ ion concentration were calculated from the 340 nm/380 nm ratios (generated from background-corrected 340 nm/380 nm images) by using the equation of Grynkiewicz et al. (18) and a K_d of 250 nM for fura-2 AM. At least two independent experiments were performed, and the results are presented as the averages ± standard deviations.

	RESULTS

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Identification of genes transcriptionally activated in T cells following IL-2 deprivation. The mouse HT-2 T helper cell line requires IL-2 for growth, and in its absence HT-2 cells undergo apoptosis (Fig. 1A) (34). For expression profiling, HT-2 poly(A)⁺ mRNA was isolated 8 h after IL-2 withdrawal and used to interrogate Affymetrix DNA chips that contained 30,000 genes or expressed sequence tags. Comparison of the transcription profiles of cells grown in the presence or absence of IL-2 revealed that 97% of the genes were unaffected by cytokine deprivation (Fig. 1B). Of the genes whose transcription was altered by cytokine deprivation, surprisingly, approximately three-quarters were stimulated and one-quarter were repressed (Fig. 1C).

View larger version (14K): [in this window] [in a new window]   FIG. 1. A transcriptional program of apoptosis induction following IL-2 deprivation. (A) Time course of apoptosis following IL-2 deprivation. The percentage of viable HT-2 cells was determined by annexin V-FITC staining. (B) Summary of microarray analysis. Affected is defined as a twofold or greater change following IL-2 deprivation. (C) Graphical representation of the fold changes of genes stimulated or repressed.

The finding that RC3 was transcriptionally activated in lymphocytes was surprising, because the RC3 protein was previously characterized as a neurally specific factor involved in synaptic plasticity and spatial learning (29, 37). Therefore, RC3 represented a putative novel gene involved in lymphocyte apoptosis, and we sought to test this hypothesis and study the mechanism of RC3 action. Northern blot analysis confirmed the results of the DNA microarray experiments: IL-2 withdrawal resulted in transcriptional activation of RC3 and repression of ATFx (Fig. 2A). A time course analysis revealed that RC3 transcriptional induction preceded the onset of apoptosis (Fig. 2B). Immunoblot analysis showed that like the mRNA levels, RC3 protein levels were also up-regulated following IL-2 withdrawal (Fig. 2C).

View larger version (31K): [in this window] [in a new window]   FIG. 2. Induction of RC3 transcription following IL-2 deprivation. (A) Transcriptional activation of RC3 upon IL-2 deprivation. Expression of RC3, ATFx, and GAPDH (as a loading control) in HT-2 cells was assessed by Northern blot analysis with 1 µg of poly(A)+ mRNA or 10 µg of total RNA 8 h after IL-2 deprivation. (B) Time course of RC3 induction. RC3 mRNA levels in HT-2 cells were analyzed at the indicated times following IL-2 deprivation. The Northern blot was stripped and reprobed with a 28S rRNA probe as a loading control. (C) Immunoblot analysis of whole-cell extracts (Lysate) or culture supernatants (Supt) from HT-2 cells grown in the presence or absence of IL-2. The same blot was stripped and reprobed with an -tubulin antibody (Sigma) as a loading control. The exposure time was 10 s.

View larger version (29K): [in this window] [in a new window]   FIG. 3. Cell specificity of RC3 transcription activation. (A) RC3 is transcriptionally activated in CTLL-2 cells upon IL-2 deprivation. Expression of RC3 and GAPDH (as a loading control) was assessed by Northern blot analysis with 1 µg of poly(A)+ mRNA or 10 µg of total RNA 8 h after IL-2 removal. (B) Northern blot analysis of poly(A)+ mRNA prepared from primary human T lymphocytes 12 to 14 h after IL-2 removal. (C) Northern blot analysis of poly(A)+ mRNA prepared from FL5.12 cells 8 h after IL-3 removal. The same blot was sequentially stripped and reprobed with 24p3 (as an induction control), ATFx (as a repression control), and GAPDH probes. (D) Northern analysis of poly(A)+ mRNA from 32D cells. The blot was probed sequentially with RC3 and GAPDH probes. (E) Immunoblot analysis of whole-cell extracts from Jurkat cells after staurosporine (STS) treatment, which induces apoptosis. The same blot was stripped and reprobed with a caspase-3 antibody (Santa Cruz Biotechnology) as a control; STS treatment results in proteolytic activation of caspase 3.

View larger version (44K): [in this window] [in a new window]   FIG. 4. Expression of RC3 induces apoptosis in HT-2 and CTLL-2 cells. (A) HT-2 and CTLL-2 cells were transiently transfected with 5 µg of either pcDNA3.1 (empty vector) or pcDNA3.1/RC3 or left untransfected (control). One day after transfection, cells were collected and stained with annexin V-FITC and analyzed by FACS. PI, propidium iodide. (B) Extracts were prepared from a portion of the transfected cells and analyzed by immunoblotting with an antihemagglutinin (anti-HA) monoclonal antibody or an -tubulin antibody. (C) HT-2 cells were cotransfected with 1 µg of GFP plasmid and 4 µg of either pcDNA3.1 (empty vector), pcDNA3.1+/RC3 (sense orientation), or pcDNA3.1-/RC3 (antisense orientation). Cotransfected cells were stained with annexin V-PE and analyzed by FACS. The percentage of GFP-positive cells that were also positive for annexin V-PE was quantitated and plotted. Error bars indicate standard deviations. (D) HT-2 and CTLL-2 cells transfected with empty vector plasmid (pcDNA3.1) or an RC3 expression plasmid were subjected to DNA fragmentation analysis.

A functional RC3 CaM binding domain is required for induction of apoptosis. Alterations in intracellular ion levels are a feature of several apoptotic pathways, and the involvement of Ca²⁺ has been well documented. For example, Ca²⁺ ionophores, which directly mobilize Ca²⁺ from internal stores, induce apoptosis in diverse cell types (27). Conversely, agents that decrease intracellular Ca²⁺ concentration can inhibit apoptosis; for example, one of the many activities of the antiapoptotic Bcl-2 protein is to prevent the loss of Ca²⁺ from intracellular stores, thereby blocking apoptosis (2).

Many of the physiological effects of Ca²⁺ are mediated by CaM, a small, ubiquitous protein that regulates a wide variety of cellular processes by activating Ca²⁺/CaM-dependent enzymes that, in turn, modulate signaling cascades. Like intracellular Ca²⁺, CaM may also regulate apoptosis; for example, increased CaM expression has been correlated with apoptosis (9), and, conversely, CaM antagonists can interfere with apoptosis (26).

RC3 contains a well-characterized 20-amino-acid sequence known as the IQ motif (3), which functions as a binding site for CaM (12). Specific single amino acid substitutions in the IQ domain eliminate binding of RC3 to CaM in vivo (31). In the brain, CaM appears to be the major protein that interacts with RC3, and the RC3-CaM interaction is believed to be essential for RC3 function (12, 31). To investigate whether the ability of RC3 to induce apoptosis correlated with its ability to bind CaM, we analyzed the apoptotic activity of RC3 derivatives bearing mutations in the CaM binding domain by using two different assays. In the first assay, HT-2 cells cultured in the presence of IL-2 were transfected with plasmids expressing either wild-type RC3 or an RC3 mutant, and the percentage of apoptotic cells was determined by annexin V-FITC staining. Figure 5A shows that mutation of Ile-33 to Gln (I33Q), which completely inhibits RC3 binding to CaM (31), also eliminated its proapoptotic activity. Similarly, mutation of either Ser-36 to Asp (S36D) or Arg-38 to Gln (R38Q), which weakens the RC3-CaM interaction (31), also reduced the proapoptotic activity of RC3. In contrast, the Ser-36-to-Ala mutation (S36A), which does not affect the RC3-CaM interaction (31), induced apoptosis at levels similar to those for wild-type RC3.

View larger version (64K): [in this window] [in a new window]   FIG. 5. A functional RC3 CaM binding domain is required for induction of apoptosis. (A) HT-2 cells grown in the presence of IL-2 were transfected with 4 µg of empty vector plasmid (pcDNA3.1), wild-type RC3 (WT), or RC3 mutants. Cells were collected 24 h after transfection, stained with annexin V-FITC, and subjected to FACS analysis to monitor cell viability. The protein sequence of the 20-amino-acid CaM binding domain (IQ motif) of RC3 is shown; mutated residues are indicated by arrows. Error bars indicate standard deviations. (B) HEK-293 cells were transiently cotransfected with a ß-galactosidase (ß-gal) reporter plasmid (0.5 µg) and the experimental plasmid (4 µg), as indicated. The cells were washed 24 h after transfection, fixed, and stained with 0.2% X-Gal. (C) Quantitative analysis of the experiment for panel B. Four hundred ß-galactosidase-positive cells were analyzed, and the number of blue cells displaying an apoptotic morphology was assessed.

Both IL-2 deprivation and RC3 expression increase intracellular calcium concentration. An increased intracellular Ca²⁺ concentration is known to induce apoptosis. We therefore reasoned that increased RC3 levels that occur during IL-2 deprivation might trigger apoptosis by sequestering CaM, thereby releasing Ca²⁺ and increasing the intracellular Ca²⁺ concentration. As an initial test of this hypothesis, we measured intracellular Ca²⁺ levels in HT-2 cells following IL-2 deprivation. As a positive control, HT-2 cells were treated with thapsigargin, which releases Ca²⁺ from intracellular pools and induces apoptosis. Figure 6A shows that thapsigargin treatment increased the intracellular Ca²⁺ concentration 10-fold, whereas IL-2 deprivation increased the intracellular Ca²⁺ concentration 5-fold. We conclude that IL-2 deprivation increases intracellular Ca²⁺ concentration.

View larger version (29K): [in this window] [in a new window]   FIG. 6. Both IL-2 deprivation and RC3 expression increase intracellular Ca2+ concentration. (A) HT-2 cells grown in the presence or absence of IL-2 for 12 h were stained with the fluorescent Ca2+ indicator fura-2 AM, and the intracellular Ca2+ concentration was measured (see Materials and Methods for details). As a control, HT-2 cells were treated with 2 µM thapsigargin (TSG), a Ca2+ store depletor, for 1 min before staining with fura-2 AM. Error bars indicate standard deviations. (B) HT-2 cells were transfected with a plasmid expressing RC3 or an RC3 mutant and analyzed as described for panel A. WT, wild type. (C) HT-2 cells were transfected with either pcDNA3.1 or an RC3 expression plasmid; transfected cells were isolated from untransfected cells (see Materials and Methods for details) and, in parallel with controls with or without IL-2, stained with fura-2 AM. The left panels show the pseudocolor images of intracellular Ca2+ within fura-2-loaded cells; high Ca2+ levels are shown in red, and low Ca2+ levels are shown in blue or violet. Cells cultured in the absence of IL-2 or expressing RC3 display an apoptotic morphology. The average intracellular Ca2+ concentration of the cells shown in each field of view was calculated from background corrected images (see Materials and Methods) and is shown in the panel on the right. (D) Primary human T lymphocytes were grown in the presence or absence of IL-2 for 12 to 14 h and stained with fura-2 AM. The left panels show the pseudocolor images, and the right panel shows the average intracellular Ca2+ concentration as described for panel C.

To directly quantify the Ca²⁺ increase induced by RC3 expression, we performed single-cell imaging experiments. HT-2 cells transfected with either an RC3 expression plasmid or pcDNA3.1 were isolated and subjected to single-cell intracellular Ca²⁺ imaging analysis. The pseudocolor image in Fig. 6C shows that RC3 expression resulted in an increased intracellular Ca²⁺ concentration. Quantitation revealed that RC3 expression increased the intracellular Ca²⁺ concentration approximately eightfold, comparable to the approximately ninefold increase observed upon IL-2 withdrawal in the same assay, confirming the results of Fig. 6A and B. Finally, a similar analysis for IL-2-dependent primary human T lymphocytes again indicated that RC3 expression increased the intracellular Ca²⁺ concentration (Fig. 6D).

An increase in intracellular calcium is required for induction of apoptosis. To determine if the Ca²⁺ increase that occurs following IL-3 withdrawal or RC3 expression is directly responsible for triggering apoptosis, we asked whether depletion of intracellular Ca²⁺ could protect cells from apoptotic death. Incubation in the presence of BAPTA-AM, an intracellular Ca²⁺ chelator, inhibited apoptosis induced by cytokine deprivation in IL-2-dependent HT-2 cells (Fig. 7A) and in IL-2-dependent primary human T lymphocytes (Fig. 7B). Similarly, BAPTA-AM treatment abolished the apoptotic effects of RC3 expression (Fig. 7C). These results indicate that intracellular free Ca²⁺ accumulation plays a critical role in apoptosis induced by either IL-2 deprivation or RC3 expression.

View larger version (16K): [in this window] [in a new window]   FIG. 7. An increase in intracellular Ca2+ is required for induction of apoptosis. (A) HT-2 cells grown in the presence or absence of IL-2 were incubated with BAPTA-AM, and the number of cells undergoing apoptosis was quantitated by annexin V-FITC staining and FACS analysis. Error bars indicate standard deviations. (B) Primary human T lymphocytes grown in the presence or absence of IL-2 were incubated with BAPTA-AM, and the number of cells undergoing apoptosis was quantitated as described for panel A. (C) HT-2 cells transfected with either pcDNA3.1 or an RC3 expression plasmid were treated with BAPTA-AM, and the number of cells undergoing apoptosis was analyzed as described for panel A.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

Although the mechanism by which IL-2 promotes transcription has been well characterized (reviewed in reference 15), the pathway by which IL-2 deprivation increases transcription of RC3 and other genes remains to be elucidated. One potential clue has come from studies with IL-2-dependent mouse lymphoid cells, which show that expression of the AP-1 transcription factors c-Fos and c-Jun is rapidly induced following growth factor deprivation (7). The RC3 promoter contains putative AP-1 binding sites (22), suggesting a possible mechanism for transcriptional activation of RC3 following IL-2 deprivation.

The proapoptotic activity of RC3 requires a functional CaM binding domain, indicating that the ability of RC3 to induce apoptosis is related to its role in modulating intracellular CaM and/or Ca²⁺ levels. Recent computer-based modeling of calpacitin-CaM interactions predicts that increased RC3 levels would lead to dissociation of the Ca²⁺-CaM complex, thereby increasing intracellular Ca²⁺ levels (14). Consistent with this prediction, we found that overexpression of RC3 resulted in an increased intracellular Ca²⁺ concentration (Fig. 6).

Based on the findings presented in this report and previous studies on calpacitin regulation and function, we propose the following model for the induction of apoptosis by RC3 in lymphoid cells. IL-2 deprivation induces RC3 expression, and the elevated levels of RC3 sequester CaM and stabilize it in its Ca²⁺-free form, which, as described above, contributes to an increase in intracellular Ca²⁺ and triggers apoptosis. In support of this model, antisense-mediated blocking of the Ca²⁺ channel renders T cells resistant to apoptosis, indicating that intracellular Ca²⁺ levels play a role in T-cell apoptosis (23).

IL-2 deprivation may also decrease protein kinase C (PKC) activity (17). PKC phosphorylates RC3 and blocks the RC3-CaM interaction (12). Thus, a reduction in PKC phosphorylation should stabilize the Ca²⁺-free form of CaM, which would also increase the intracellular Ca²⁺ concentration. Finally, RC3 could also increase intracellular Ca²⁺ and induce apoptosis by regulating the association of Ca²⁺/CaM with various proteins, thereby modulating other CaM-dependent pathways. One such protein may be the CaM-dependent plasma membrane Ca²⁺-ATPase; a decreased efflux of intracellular Ca²⁺ could contribute to the Ca²⁺ increase.

The ability of a CaM binding protein to induce apoptosis when overexpressed has also been demonstrated with two proapoptotic serine/threonine kinases: DAPK and DRP-1. Intriguingly, like that of RC3, expression of DRP-1 is also increased following IL-2 deprivation (Table 1). DRP-1 has been shown to induce apoptosis when expressed in a variety of cell lines (21), and both RC3 and DRP-1 have similar IQ motif CaM binding domains. It therefore appears that IL-2 deprivation increases the level of multiple CaM-interacting proteins that activate parallel apoptotic pathways, thus ensuring efficient onset of apoptosis.

	ACKNOWLEDGMENTS

 
We thank J. G. Sutcliffe for the RC3 cDNA and antibody, D. Storm for RC3 mutants, C. Sagerstorm for help with the microscopy, A. Earhardt and D. Lambright for help with the spectrofluorometry, the University of Massachusetts Medical School Center for AIDS Research (CFAR) for human peripheral blood lymphocytes, P. Furcinitti for help with the Ca²⁺ single-cell imaging, members of our laboratory for helpful discussions and comments on the manuscript, and S. Evans for editorial assistance.

This work was supported in part by an NIH grant (to M.R.G.) and by the Leukemia and Lymphoma Society (to L.R.D.). M.R.G. is an investigator of the Howard Hughes Medical Institute.

	FOOTNOTES

 
* Corresponding author. Mailing address: Howard Hughes Medical Institute, Programs in Gene Function and Expression and Molecular Medicine, University of Massachusetts Medical School, 364 Plantation St., Worcester, MA 01605. Phone: (508) 856-5331. Fax: (508) 856-5473. E-mail: michael.green{at}umassmed.edu.

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