Induction of Ca²⁺-driven apoptosis in chronic lymphocytic leukemia cells by peptide-mediated disruption of Bcl-2–IP₃ receptor interaction

The Bcl-2 protein contributes to the pathophysiology of cancer and the resistance of cancer to therapeutic agents by virtue of its ability to inhibit apoptosis.^1,2  The Bcl-2–positive lymphoid malignancies follicular lymphoma and chronic lymphocytic leukemia (CLL) are prime examples. They are associated with an elevation of Bcl-2 because of the t(14;18) chromosomal translocation in follicular lymphoma³  and the loss of miR-15a and miR16-1 in CLL.^4,5  Cure of these malignancies is infrequently achieved with current therapeutic modalities, and thus a major challenge remains to develop new treatment modalities based on an understanding of the fundamental disease mechanisms.⁶ 

Bcl-2 blocks apoptosis in part by binding its proapoptotic relatives, thus preserving mitochondrial integrity and preventing cytochrome c release.^7,8  Therefore, considerable investment has been made in the development of novel therapeutic agents, such as ABT-737, that disrupt the inhibitory interaction of Bcl-2 with its proapoptotic relatives.^1,2,9-11  Several of these agents are currently undergoing clinical testing. However, Bcl-2 also inhibits apoptosis by regulating the release of Ca²⁺ from the endoplasmic reticulum (ER).^12-14  This recently characterized mechanism involves a physical interaction of Bcl-2 with the inositol 1,4,5-trisphosphate receptor (IP₃R) Ca²⁺-release channel on the ER. Through this interaction, Bcl-2 prevents cytoplasmic Ca²⁺ elevation sufficient to trigger apoptosis. However, the potential contribution of the Bcl-2–IP₃R interaction to the survival of CLL has not been investigated, so the opportunity for targeting this interaction therapeutically has not yet been realized.

The IP₃R is an IP₃-gated Ca²⁺ channel that is highly conserved, represented by 3 isoforms, and present in virtually all cell types.^15,16  IP₃-dependent release of Ca²⁺ from the ER into the cytoplasm produces Ca²⁺ signals, generally in the form of Ca²⁺ oscillations, which govern diverse cellular functions including cell proliferation and survival.^17,18  Ca²⁺ oscillations support cell survival in part by positively regulating mitochondrial metabolism, but sustained high-amplitude elevations of Ca²⁺ induce mitochondrial Ca²⁺ overload and apoptosis.^19-21  Bcl-2 inhibits high-amplitude, proapoptotic Ca²⁺ elevation but does not interfere with physiologic Ca²⁺ oscillations.²²  In fact, under certain circumstances Bcl-2 and its homolog Bcl-xl enhance Ca²⁺ oscillations,^22-26  and through this mechanism are predicted to promote efficient mitochondrial bioenergetics.²⁷  Thus, Bcl-2 supports cell survival both by enhancing physiologic Ca²⁺ signals and by blocking proapoptotic Ca²⁺ elevation.

A major focus of our work has been to understand how Bcl-2 inhibits proapoptotic elevation of Ca²⁺ based on evidence that Bcl-2 binds to the IP₃R and thus inhibits ER Ca²⁺ release.^28-32  Although this interaction has been mainly detected in cell extracts by coimmunoprecipitation or blue native gel electrophoresis, we recently confirmed the Bcl-2–IP₃R interaction in living cells using fluorescence resonance energy transfer.³³  Furthermore, our findings indicate that the interaction involves direct binding of the BH4 domain of Bcl-2 to the regulatory and coupling domain of the IP₃R.^33,34  This region, located between the IP₃-binding domain at the N-terminus of the IP₃R and the transmembrane channel domain near the C-terminus, binds a variety of regulatory proteins and undergoes several secondary modifications, thus modulating channel opening in response to IP₃ binding.^15,16 

A major tool in our recent investigation of the Bcl-2–IP₃R interaction has been a synthetic peptide, previously referred to as peptide 2 and in the present study as the IP₃R-derived peptide (IDP).³³  The IDP corresponds to a 20–amino acid sequence within the Bcl-2–binding site on the IP₃R and functions as a competitive inhibitor of the Bcl-2–IP₃R interaction.^33,34  By disrupting this interaction, IDP reverses Bcl-2–mediated inhibition of both IP₃-dependent channel opening and IP₃-dependent ER Ca²⁺ release. Thus, when delivered into Bcl-2–positive T cells by fusion with the cell-penetrating peptide of HIV transactivator of transcription (TAT-IDP) or by interaction with Chariot peptide uptake reagent, the IDP reverses Bcl-2–mediated inhibition of IP₃-dependent Ca²⁺ elevation and apoptosis induced by anti-CD3 antibody.³³ 

We report the development of a TAT-IDP analog, TAT-IDP_DD/AA, which induces striking Ca²⁺ elevation in primary CLL cells by binding to the BH4 domain of Bcl-2 and disrupting Bcl-2–IP₃R interaction, thereby triggering Ca²⁺-dependent apoptosis. These findings suggest a novel approach for targeting Bcl-2 for therapeutic purposes. In addition, the discovery that disrupting the Bcl-2–IP₃R interaction is sufficient to induce IP₃R-mediated Ca²⁺ elevation suggests that Bcl-2 may contribute to the pathophysiology of CLL by suppressing proapoptotic Ca²⁺ signals downstream of constitutively active B-cell receptor (BCR) signals.

Fura-2 AM and Hoechst 33342 were from Invitrogen, pepstatin A from Roche, and ABT-737 from Abbott Laboratories. Peptides were from GenScript and were > 95% pure as shown by mass spectrometry and high-performance liquid chromatography.

WEHI7.2 cells were cultured and stably transfected with expression vectors encoding wild-type Bcl-2 and Bcl-2RS/GG, as described previously.^28,33  Jurkat and RS11846 cells were cultured in RPMI medium supplemented with 10% fetal bovine serum, l-glutamine (2mM), and nonessential amino acids (100μM). Cells were maintained in 5% carbon dioxide at high humidity. DT40 cells were cultured in RPMI medium with 10% fetal bovine serum, 1% chicken serum, and 2mM l-glutamine.

Primary lymphocytes were separated using a Ficoll Hypaque density gradient from the peripheral blood of normal adult volunteers or adult patients with CLL, and suspended in RPMI medium supplemented with 10% fetal bovine serum, l-glutamine (2mM), and nonessential amino acids (100μM) at a density of 2 million cells/mL. B and T cells were separated from normal mononuclear cells by magnetic separation using CD19 microbeads (Miltenyi Biotec). All procedures followed the guidelines and regulations in accordance with internal review board protocol ICC2902/11-02-28 of Case Western Reserve University Cancer Center/University Hospitals of Cleveland Ireland Cancer Center. The use of human subjects in this proposal was approved by the institutional review board of Case Western Reserve University School of Medicine.

Single-cell digital imaging of Fura-2 AM–loaded cells in extracellular buffer (130mM NaCl, 5mM KCl, 1.5mM CaCl₂, 1mM MgCl₂, 25mM HEPES [N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid], pH 7.5, 1 mg/mL of bovine serum albumin, and 5mM glucose) was described previously in detail.²²  Xestospongin C (Enzo Life Sciences International) and U73122 (Sigma), both dissolved in dimethylsulfoxide (DMSO), were added to cells 30 minutes before Ca²⁺ measurements.

Unidirectional ⁴⁵Ca²⁺-flux assay methods were described previously.^34,35  Briefly, this method involves loading nonmitochondrial Ca²⁺ stores in saponin-permeabilized mouse embryonic fibroblast (MEF) cells with ⁴⁵Ca²⁺. Saponin (20 μg/mL) selectively permeabilizes the plasma membrane by its preferential interaction with cholesterol, providing direct access to the intracellular Ca²⁺ stores, and azide prevents ⁴⁵Ca²⁺ uptake by the mitochondria, leaving only the ER as a Ca²⁺ store. IP₃ (3μM)–mediated ⁴⁵Ca²⁺ efflux was then measured as described previously.^34,35  BH4-Bcl2 peptide (40μM), in the presence or absence of the IDP_DD/AA peptide (40μM), was added from 4 minutes before IP₃ to 2 minutes after IP₃. Ca²⁺ release was plotted as fractional loss (percentage/2 minutes) as a function of time.

Western blotting was performed as described previously.³³  The following antibodies were used: anti–human Bcl-2 (551052; BD Biosciences), anti–mouse/human Bcl-2 (Santa Cruz Biotechnology), and anti-actin (Sigma). Glutathione S-transferase (GST) pull-down methods were described previously in detail,³³  and were used here with these modifications: 3xFLAG-Bcl-2 was expressed in COS-7 cells; mutant GST-IP₃R1 domain 3, in which Asp¹⁴⁰³Asp¹⁴⁰⁴ residues were mutated to Ala, was tested; and glutathione-Sepharose beads were washed 4 times with modified interaction buffer (150 instead of 300mM NaCl without bovine serum albumin), and GST-fusion proteins were eluted by incubating beads with 40 μL of LDS (Invitrogen) for 3 minutes at 95°C. 3xFLAG-tagged Bcl-2 was detected by Western blotting using a mouse monoclonal ANTI-FLAG M2-Peroxidase antibody (Sigma). Total protein content was visualized by GelCode blue (ThermoFisher Scientific) staining of the blot after film development. Quantification was done with ImageJ software Version 1.43u (National Institutes of Health; publisher, Wayne Rasband).

Binding of GST-Domain3, GST-Domain3DD/AA, and parental GST to the BH4 domain of Bcl-2 was analyzed by surface plasmon resonance (SPR) at 25°C using a Biacore 2000 instrument. Fusion proteins were affinity purified and dialyzed against phosphate-buffered saline (PBS) using a Slide-A-Lyzer with a cutoff of 3 kDa (Pierce) to minimize buffer effect. Fusion protein concentration was determined using bicinchoninic acid protein assay reagent (Pierce), and the quality and integrity were examined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and GelCode blue (Pierce). Equal amounts (200 ng or 58.2 pmol of biotinylated peptide) of > 80% pure biotinylated BH4-Bcl2 peptide (Biotin-RTGYDNREIVMKYIHYKLSQRGYEW) and BH4-Bcl2-scrambled peptide (Biotin-WYEKQRSLHGIMYYVIEDRNTKGYR) were immobilized on 2 different flow cells of a streptavidin-coated sensor chip (BR-1000-32; Biacore) using PBS supplemented with 0.005% P20 at pH 7.0. Measurements with GST-fusion proteins as analyte were performed in PBS at a flow rate of 30 μL/min. Different concentrations of the analyte were used in a random order to assess binding (injection volume, 120 μL). Bound peptide was removed by injection of 5 μL of regeneration buffer (25mM NaOH, 0.002% SDS) at 10 μL/min. Background signals were obtained from the reference flow cell containing the BH4–Bcl2-scrambled peptide, and were subtracted to generate response curves using Biaevaluation 3.0 software. Only the association phase of the binding curve was taken into account. Data from 2-3 different sensor grams for each condition were fitted by nonlinear regression analysis to a Hill-Langmuir binding isotherm using Origin 7.0 software.

Bcl-2–IP₃R coimmunoprecipitation was performed as described previously.³³  For biotin-streptavidin pull-down experiments, Bcl-2–overexpressing WEHI7.2 cells were incubated on ice for 30 minutes in 400 μL of CHAPS lysis buffer (50mM Tris-HCl, pH 7.5,100mM NaCl, 2mM EDTA,1% CHAPS, 50mM NaF, 1mM Na₃VO₄, and the protease inhibitor cocktail PhosSTOP). Cell lysates were centrifuged for 15 minutes at 20,000g at 4°C, and the supernatant (protein concentration, 5 mg/mL) was continuously rotated with 200μM biotin-Scr or biotin-IDP_DD/AA or without peptide at 4°C for 16 hours. Streptavidin-Sepharose beads (40 μL) (Pierce Pull-Down Biotinylated Protein:Protein Interaction Kit, ThermoFisher, Cat no. 21115) were washed 3 times with Tris-buffered saline (25mM Tris-HCl, 0.15M NaCl, pH 7) and then added to 400 μL of cell lysate, followed by rotation for 2 hours at 4°C. The beads were washed 4 times with Tris-buffered saline, centrifuged for 30-60 seconds each time, and then incubated for 3-5 minutes at room temperature with 50 μL of elution buffer provided by the Pierce pull-down kit, and centrifuged at 1250g for 30-60 seconds. The supernatant was boiled in sodium dodecyl sulfate-polyacrylamide gel electrophoresis sample buffer for 5 minutes, and Western blotting for Bcl-2 was performed as described in Western blotting section (above).

Trypan blue dye uptake was detected by light microscopy, and propidium iodide uptake was quantified by flow cytometry in association with the annexin V measurements using an FACSAria flow cytometer (Beckman Coulter). Flow cytometric data were analyzed using FlowJo Version 8.8.4 for Macintosh (TreeStar). Apoptotic nuclei were quantified in Hoechst 33342–stained cells by epifluorescence microscopy (Zeiss Axiovert S100) using a 63× fluorescent oil objective (Carl Zeiss) and a Hamumatsu Orca camera at room temperature. The activities of caspase-3 and caspase-7 were measured with the Caspase-Glo 3/7 luminescence assay (Promega) according to the manufacturer's protocol. To monitor mitochondrial membrane potential, cells were incubated with 20nM tetramethyl-rhodamine methyl ester for 20 minutes at 37°C, washed twice with extracellular buffer (see “CA²⁺ measurements” above), and resuspended in extracellular buffer before adding peptides. Confocal images were acquired with an UltraVIEW VoX spinning disc confocal system (Perkin-Elmer) mounted on a Leica DMI6000B microscope equipped with an HCX PL APO 100×/1.4 oil-immersion objective. Tetramethyl-rhodamine methyl ester fluorescence was excited at 561 nm, and its emission was collected using a 580-650 nm bandpass filter.

A Student t test was used to assess statistical differences between groups. A 2-tailed P value of .05 was the threshold for significance. Statistical tests were performed using Microsoft Excel 2004 for Macintosh.

Four cell lines, representing lymphoid malignancies of both T- and B-cell origin, were used as shown in Figures 1,Figure 2,Figure 3–4 to lay the foundation for studies using primary CLL cells shown in Figures 5 and 6. The sequence of TAT-IDP and its scrambled control analog, TAT-Scr, are shown in Figure 1A, together with a diagram pinpointing the IP₃R region from which the IDP sequence was derived. Also shown is an immunoblot documenting that Bcl-2 is virtually undetectable in the WEHI7.2 murine T-cell lymphoma line and that Bcl-2 levels are similar in WEHI7.2 cells expressing wild-type Bcl-2 or mutant Bcl-2RS/GG, in which arginine 6 and serine 7 are both converted to glycine (Figure 1B). We demonstrated previously that wild-type Bcl-2 interacts with the IP₃R in these cells, whereas the Bcl-2RS/GG does not.³³  In experiments summarized in Figure 1C, Ca²⁺ elevations were induced when Bcl-2–positive WEHI7.2 cells were treated with TAT-IDP, whereas Ca²⁺ elevations were very infrequent after treatment with TAT alone, IDP alone, or a mixture of TAT and IDP. IDP also induced Ca²⁺ elevations when introduced into the same cells with Chariot peptide uptake reagent (data not shown). The pattern of Ca²⁺ elevation induced by TAT-IDP in Bcl-2–positive WEHI7.2 cells was analyzed by single-cell digital imaging in a large number of experiments (Figure 1D). Two patterns were arbitrarily defined: 1 or 2 isolated Ca²⁺ oscillations were observed in 22% ± 2% of cells, whereas a sustained pattern of Ca²⁺ oscillations (3 or more peaks) was observed in 36% ± 7% of the cells. Thus, on average, more than 50% of Bcl-2–positive cells reproducibly displayed Ca²⁺ elevations in response to TAT-IDP, whereas TAT-Scr did not induce Ca²⁺ elevation to a significant degree. Cell viability was not lost after TAT-IDP treatment (data not shown). Moreover, TAT-IDP did not induce Ca²⁺ elevations to nearly the same degree in Bcl-2–negative cells or Bcl-2RS/GG–positive cells as in cells expressing wild-type Bcl-2 (Figure 1E-F). In summary, these findings indicate that IDP induced Ca²⁺ oscillations when introduced into cells either by fusion with the cell-penetrating peptide TAT or with a commercial peptide uptake reagent, and that this process was dependent on Bcl-2.

TAT-IDP also induces Ca²⁺ oscillations in the Jurkat human T-cell leukemia line (Figure 2A subpanel i), which natively expresses Bcl-2 and in which we have previously documented interaction of endogenous Bcl-2 with the IP₃R.³³  TAT-linked peptides are delivered into cells by macropinocytosis³⁶  and are therefore exposed to endocytic proteases, mainly cathepsins.³⁷  Moreover, the presence of an aspartyl protease cleavage site, PLDD, in IDP was predicted by 2 algorithms (CLC Main Workbench and the CASVM server³⁸ ). The possibility that TAT-IDP may be subject to degradation when delivered into Jurkat cells was also suggested by a slight, but not significant, increase in both the percentage of cells displaying peptide-induced Ca²⁺ elevations (Figure 2B subpanel i) and a significant enhancement of the amplitude of these Ca²⁺ elevations (Figure 2B subpanel ii) after pretreatment with the aspartyl protease inhibitor pepstatin A. Therefore, an IDP analog, TAT-IDP_DD/AA was generated with the goal of eliminating the predicted protease cleavage site (Figure 2C). Representative Ca²⁺ oscillations induced by TAT-IDP_DD/AA are shown in Figure 2A (subpanel ii). TAT-IDP and TAT-IDP_DD/AA induced Ca²⁺ oscillations in a similar percentage of cells (Figure 2D), but both the amplitude (Figure 2A and E) and frequency (Figure 2A and F) of Ca²⁺ oscillations were much higher with TAT-IDP_DD/AA than with TAT-IDP. Another peptide analog in which the predicted protease cleavage site is eliminated without altering overall peptide charge, TAT-IDP_DD/EE, also induced Ca²⁺ oscillations of higher amplitude and frequency, very similar to TAT-IDP_DD/AA (data not shown). In addition, the increased activity of TAT-IDP_DD/AA compared with TAT-IDP was observed in the RS11846 human B-cell lymphoma line, which has elevated Bcl-2 because of a t(14;18) chromosomal translocation typical of human follicular lymphoma³⁹  (Figure 2G-I). Thus, the induction of Ca²⁺ oscillations by these peptides is not unique to T lymphocytes.

Because TAT-IDP_DD/AA was significantly more effective at inducing Ca²⁺ oscillations than TAT-IDP, we investigated its mechanism of action at a biochemical level. As shown in GST pull-down experiments in Figure 3A, binding of 3xFLAG-Bcl-2 to GST-Domain3 was not significantly different from the binding to GST-Domain3DD/AA. Furthermore, the binding of these GST-tagged domains to Biotin-BH4-Bcl-2 (a biotin-tagged peptide corresponding to the BH4 domain of Bcl-2) was measured by SPR (Figure 3B). The results indicated that GST-Domain3 and GST-Domain3DD/AA bound specifically to Biotin-BH4-Bcl-2 with similar affinities. In addition, interaction of IDP_DD/AA with Bcl-2 was documented by biotin-streptavidin pull-down (Figure 3C), and disruption of the Bcl-2–IP₃R interaction by IDP_DD/AA was confirmed in coimmunoprecipitation experiments (Figure 3D). Consistent with these observations, IDP_DD/AA reversed the Bcl-2–imposed inhibition of IP₃-induced ⁴⁵Ca²⁺ efflux from the ER when added to cells after ⁴⁵Ca²⁺ loading and permeabilization with saponin, as was shown previously for IDP³⁴  (Figure 3E).

Finally, the IP₃R inhibitor xestospongin C and the phospholipase C inhibitor U73122 reduced the percentage of cells displaying peptide-induced Ca²⁺ oscillations, providing further evidence that Ca²⁺ responses to both TAT-IDP and TAT-IDP_DD/AA were IP₃R dependent (Figure 4A). In addition, each of these peptides induced Ca²⁺ oscillations in the DT40 B-cell lymphoma line, which expresses all 3 IP₃R isoforms (Figure 4B), but the induction of Ca²⁺ oscillations by these peptides was significantly less in triple IP₃R knockout DT40 cells (Figure 4C).

In summary, these findings suggest that the increase in Ca²⁺ responses induced in Bcl-2–positive cell lines by TAT-IDP_DD/AA compared with TAT-IDP was likely because of increased cellular uptake and stability as a result of reduced proteolytic cleavage rather than increased affinity of TAT-IDP_DD/AA for Bcl-2. In addition, these findings confirm that TAT-IDP_DD/AA functions as a competitive inhibitor of the Bcl-2–IP3R interaction, as was shown previously for TAT-IDP.³³ 

The effects of these peptides were next investigated in primary CLL cells, chosen for this purpose because CLL is invariably associated with an elevated level of Bcl-2 and because Bcl-2 plays an important role in apoptosis resistance in this disease. CLL cells were isolated from the peripheral blood of untreated patients. Malignant lymphocytes, which represented more than 80% of lymphocytes in peripheral blood samples, were separated by density gradient separation and used immediately in experiments without interval storage. The relatively high level of Bcl-2 in CLL cells compared with that in normal B lymphocytes is presented in Figure 5A and is representative of the CLL samples used here. The patterns of Ca²⁺ elevation induced by TAT-Scr, TAT-IDP, and TAT-IDP_DD/AA in CLL cells are illustrated by representative single-cell traces in Figure 5B-D and are summarized quantitatively in Figure 5E. Less than 10% of untreated cells (not shown) or TAT-Scr–treated cells had detectable Ca²⁺ elevations in recordings that were typically 90 minutes in duration. TAT-IDP induced Ca²⁺ elevation in only 26% ± 2% of CLL cells, corresponding to either single or dual spikes (14% ± 2%) or repetitive oscillations (≥ 3 successive Ca²⁺ spikes; 12% ± 2%). Thus, overall Ca²⁺ responses to TAT-IDP were much less than those observed in the cell lines investigated in this study. On the other hand, TAT-IDP_DD/AA induced Ca²⁺ elevation in 51% ± 5% of CLL cells isolated from 7 patients.

The response patterns and percentages of responses were remarkably similar among the individual patient samples. Several different patterns of Ca²⁺ elevation were observed in CLL cells after TAT-IDP_DD/AA addition (Figure 5D). One pattern, observed in only 6% ± 2% of cells, consisted of prolonged Ca²⁺ oscillations without baseline Ca²⁺ elevation, similar to the patterns observed previously in Bcl-2–positive cell lines described above in Figure 2. The more prominent patterns included 1 or 2 high-amplitude spikes associated with prolonged basal Ca²⁺ elevation (28% ± 5% of cells) or a relatively brief burst of Ca²⁺ oscillations, also associated with prolonged basal Ca²⁺ elevation (17% ± 4% of cells). Analysis of multiple experiments monitoring Ca²⁺ responses in a large number of cells documented that differences in the relative abilities of TAT-Scr, TAT-IDP, and TAT-IDP_DD/AA to induce Ca²⁺ elevations were highly significant (Figure 5E). In addition, the amplitude of Ca²⁺ peaks was significantly higher after TAT-IDP_DD/AA compared with TAT-IDP (Figure 5F). While differing in pattern and amplitude, the Ca²⁺ responses to both TAT-IDP (data not shown) and TAT-IDP_DD/AA were inhibited by xestospongin C, which is consistent with a role of IP₃R-mediated Ca²⁺ release in the initiation of Ca²⁺ elevations by both peptides (Figure 5G).

In contrast to peptide-induced Ca²⁺ elevation, the BH3-mimetic ABT-737 did not induce Ca²⁺ elevation in CLL cells, even though ABT-737 induced the death of 40% of CLL cells at 24 hours (Figure 5H). This latter finding underscores the difference in mechanism between ABT-737 and peptides designed to inhibit Bcl-2 binding to the IP₃R (see “Discussion”). In summary, the findings shown in Figure 5 indicate that TAT-IDP_DD/AA induces significant Ca²⁺ elevation in primary CLL cells, and that the pattern of Ca²⁺ elevation is different from that elicited by TAT-IDP_DD/AA in Bcl-2–positive cell lines. Two main differences were detected: (1) TAT-IDP_DD/AA induced only a brief period of Ca²⁺ spiking rather than prolonged Ca²⁺ oscillations, and (2) after TAT-IDP_DD/AA addition, the brief period of Ca²⁺ spiking in CLL cells was followed by a continuous cytoplasmic Ca²⁺ elevation that was not observed in the cell lines.

Because continuous cytoplasmic Ca²⁺ elevation can trigger apoptosis and necrosis, the effects of TAT-IDP_DD/AA on CLL-cell viability were tested. As shown in Figure 6, TAT-IDP_DD/AA-mediated Ca²⁺ elevation induced substantial apoptosis in CLL cells. This was observed consistently in CLL samples isolated from 8 patients. Cell death was documented by trypan blue dye uptake (Figure 6A) and propidium iodide uptake (Figure 6D), while apoptosis induction was documented by observing apoptotic nuclear morphology (Figure 6B-C) and by detecting phosphatidylserine exposure on the cell surface with annexin V (Figure 6E). Consistent with these findings, TAT-IDP_DD/AA treatment activated caspases (Figure 6F) and caused loss of mitochondrial membrane potential (Figure 6G). The induction of cell death was specific to TAT-IDP_DD/AA, because neither TAT-Scr nor TAT-IDP induced apoptosis in CLL cells. Moreover, apoptosis induction by TAT-IDP_DD/AA was inhibited by xestospongin C, confirming that TAT-IDP_DD/AA-induced apoptosis was IP₃R mediated (Figure 6H). Because IP₃R-mediated apoptosis is triggered by Ca²⁺ elevation,^19,20  we inferred that Ca²⁺ elevation is the initiating factor in TAT-IDP_DD/AA-induced apoptosis. Ideally, intracellular Ca²⁺ chelators (eg, BAPTA-AM) would be used to suppress Ca²⁺ elevation and thereby directly confirm the role of Ca²⁺ elevation in TAT-IDP_DD/AA-induced apoptosis. However, this is not possible because buffering intracellular Ca²⁺ with BAPTA-AM kills CLL cells (F. Z., unpublished observations), ostensibly by inhibiting the basal Ca²⁺ signals required for cell survival. CLL cells were more than 6-fold more sensitive to apoptosis induced by TAT-IDP_DD/AA than normal peripheral blood lymphocytes (Figure 7A-B), suggesting that TAT-IDP_DD/AA may selectively target leukemia cells that overexpress Bcl-2.

The findings reported here provide novel insight into the role of Bcl-2 in the pathogenesis of CLL, and suggest that it may be worthwhile to target the Bcl-2–IP₃R interaction therapeutically. The present undertaking was fostered by the unexpected discovery that IDP by itself (ie, without an external IP₃-generating stimulus such as anti-CD3–mediated T-cell receptor [TCR] activation) induces Ca²⁺ elevation when introduced in Bcl-2–positive leukemia/lymphoma cells. IDP induced Ca²⁺ elevation regardless of its mode of delivery into cells (ie, as a TAT fusion peptide or using Chariot peptide uptake reagent). The recognition of a potential aspartyl cleavage site within the IDP sequence prompted the modification of TAT-IDP to TAT-IDP_DD/AA. Both peptides induce IP₃R-dependent Ca²⁺ elevation by binding to the BH4 domain of Bcl-2 and thereby disrupting the Bcl-2–IP₃R interaction, but TAT-IDP_DD/AA induces Ca²⁺ elevations of higher frequency and amplitude than observed with IDP itself, ostensibly because it is less subject to degradation during endocytic uptake (in vitro biochemical studies indicate a similar affinity of IDP and TAT-IDP_DD/AA for Bcl-2).

Although peptide-mediated disruption of the Bcl-2–IP₃R interaction induced Ca²⁺ elevation both in cell lines and in primary CLL cells, the patterns of Ca²⁺ elevation were different. In each of the cell lines, the predominant pattern was oscillatory, composed of periodic Ca²⁺ spikes. A similar oscillatory pattern was induced in CLL cells by TAT-IDP, whereas in CLL cells, TAT-IDP_DD/AA initially induced high-amplitude Ca²⁺ elevations (either 1 or 2 transient elevations or a brief burst of Ca²⁺ oscillations), followed by a sustained elevation of Ca²⁺ above basal levels. These differences in the pattern of Ca²⁺ elevation were highly reproducible and correlated with different outcomes of peptide treatment. TAT-IDP and TAT-IDP_DD/AA did not induce cell death in the cell lines, nor did TAT-IDP induce cell death in primary CLL cells. In contrast, TAT-IDP_DD/AA induced apoptosis in primary CLL cells. These findings are reminiscent of the earlier observation that Ca²⁺ oscillations induced by TCR stimulation do not induce apoptosis, whereas apoptosis ensues after strong TCR activation sufficient to induce Ca²⁺ elevation that is of high amplitude and continuous rather than oscillatory.²²  Overall, these observations are consistent with the known prosurvival functions of Ca²⁺ oscillations, including stimulation of mitochondrial metabolism, whereas sustained high-amplitude elevations of Ca²⁺ trigger apoptosis, in part by overloading mitochondria with Ca²⁺ and thus decreasing mitochondrial membrane potential.^40-45  Further investigation will be required to fully understand the nature of these differences in the Ca²⁺-signaling pattern. Moreover, continued work will be required to fully understand the detailed mechanism by which TAT-IDP_DD/AA induces apoptosis in primary CLL cells. Nevertheless, this mechanism is likely to involve the intrinsic, Bax/Bak-dependent apoptotic pathway, because Ca²⁺ elevation is known to trigger this cell-death pathway by either activating or inducing the expression of Bad and Bim, BH3 members of the Bcl-2 protein family.^19,20 

Remodeling of Ca²⁺ homeostasis and signaling through a variety of mechanisms supports the survival and proliferation of cancer cells.²¹  The findings reported here suggest that the interaction of Bcl-2 with IP₃R may contribute to this remodeling process by repressing Ca²⁺ signals driven by tonic BCR signaling, a hallmark of CLL pathogenesis.^6,46-48  Therefore, because BCR signaling increases IP₃ synthesis by activating phospholipase C, exposure of CLL cells to TAT-IDP_DD/AA alone would be sufficient to expose these Ca²⁺ signals by displacing Bcl-2 from IP₃R. The capacity of TAT-IDP_DD/AA to induce Ca²⁺-mediated apoptosis raises the prospect of targeting the Bcl-2–IP₃R interaction for therapeutic purposes in CLL and other Bcl-2–positive malignancies. The concept of targeting the Bcl-2–IP₃R interaction for therapeutic purposes is supported by evidence that TAT-IDP_DD/AA induces apoptosis in primary CLL cells, but not in normal lymphocytes.

Based on the proposed key role of BCR signaling in the pathogenesis of CLL, novel therapeutic approaches are being directed at various elements of the BCR-signaling pathway, including BCR-associated kinases such as Lyn and ZAP-70 and BCR regulatory molecules such as CD19 and CD20.⁶  Bcl-2, which is almost invariably elevated in CLL, is a prime therapeutic target. A considerable amount of effort is currently being directed toward inhibiting Bcl-2, based on understanding the Bcl-2 mechanism at a structure-function level.^1,2,11  A major target is the hydrophobic groove formed by the BH1, BH2, and BH3 domains of Bcl-2, in which the BH3 domains of proapoptotic Bcl-2 family members bind. BH3-mimetics, such as ABT-737, are modeled to fit in this groove, thus liberating proapoptotic proteins from Bcl-2 and thereby inducing apoptosis.^9,49,50  Other Bcl-2 inhibitors trigger cell death but, unlike ABT-737, their mode of cell-death induction is independent of Bax/Bak activation.⁵¹  On the other hand, TAT-IDP_DD/AA binds to the BH4 domain, displacing Bcl-2 from IP₃R and inducing apoptosis by a Ca²⁺-dependent mechanism disparate from that of ABT-737, which does not elevate Ca²⁺. Therefore, the disruption of the Bcl-2–IP₃R interaction represents a unique strategy with which to target CLL therapeutically, and is different from approaches that inhibit upstream components of the BCR-signaling pathway and from strategies that interrupt the binding of proapoptotic proteins by Bcl-2.

We thank Stephen Tahir, Abbott Laboratories, for providing ABT-737, and Michael Berridge, Martin Bootman, Zhenghe John Wang, and Andrew Lavik for helpful suggestions regarding the manuscript.

This work was supported by National Institutes of Health grants RO1 CA085804 (to C.W.D.), T32 CA059366 (M.W.H.), T32 HL007147 (J.K.M.), T32 GM08803-02 (C.R.) and T32 GM007250 (to J.K.M. and C.R.). Research performed at the K.U. Leuven was supported by Research Program G.0604.07 of the Research Foundation-Flanders (FWO), by grant GOA/09/012 of the Concerted Actions Program of the K.U. Leuven (to J.B.P. and H.D.S.), and by Onderzoekstoelage grant STRT1/10/044 of the K.U. Leuven (to G.B.). G.M. is the recipient of a doctoral fellowship from the FWO.

Contributions: F.Z. performed research, analyzed and interpreted data; M.W.H. performed research, analyzed and interpreted data, and wrote the manuscript; G.B. performed research, analyzed and interpreted data, and wrote the manuscript; G.M. performed research, analyzed and interpreted data; J.B.P. provided valuable new reagents, analyzed and interpreted data, and wrote the manuscript; and H.D.S. provided valuable new reagents, analyzed and interpreted data, and wrote the manuscript; Y.R. performed research; J.K.M. performed research, analyzed and interpreted data; M.L. performed research; C.R. analyzed and interpreted data; S.M. analyzed and interpreted data; and C.W.D. designed research, analyzed and interpreted data, performed statistical analysis, and wrote the manuscript.

Conflict of interest disclosure: The authors declare no competing financial interests.

Correspondence: Clark W. Distelhorst, MD, Charles S. Britton II Professor of Hematology/Oncology, Case Western Reserve University School of Medicine, 10900 Euclid Ave, Cleveland, OH 44106; e-mail: cwd@case.edu.

National Institutes of Health