Skip to main content
NCBI home page
Proceedings of the National Academy of Sciences of the United States of America logoLink to Proceedings of the National Academy of Sciences of the United States of America
. 2006 Apr 3;103(15):5787–5792. doi: 10.1073/pnas.0600103103

The expression level of the voltage-dependent anion channel controls life and death of the cell

Salah Abu-Hamad 1, Sara Sivan 1, Varda Shoshan-Barmatz 1,*
PMCID: PMC1458651  PMID: 16585511

Abstract

Mitochondria not only generate cellular energy, but also act as the point for cellular decisions leading to apoptosis. The voltage-dependent anion channel (VDAC), as a major mitochondrial outer-membrane transporter, has an important role in energy production by controlling metabolite traffic and is also recognized as a key protein in mitochondria-mediated apoptosis. In this study, the role of VDAC1 in regulating cell survival and death was investigated by silencing endogenous human (h)VDAC1 expression by using a short hairpin RNA (shRNA)-expressing vector. The shRNA effectively down-regulated the expression in human T-REx-293 cells of hVDAC1 but not murine (m)VDAC1. Cells in which hVDAC1 expression was decreased by ≈90% proliferated extremely slowly. Normal growth was, however, restored upon expression of mVDAC1 in a tetracycline-regulated manner. Although low tetracycline concentrations promoted cell growth, high concentrations induced mVDAC1 overexpression, leading to cell death. Cells with low levels of VDAC1 showed 4-fold-lower ATP-synthesis capacity and contained low ATP and ADP levels, with a strong correlation between ATP levels and cell growth, suggesting limited metabolite exchange between mitochondria and cytosol. The possibility of suppressing endogenous hVDAC1 expression and introducing native and mutated mVDAC1 is used to further explore the involvement of VDAC1 in apoptosis. Cells suppressed for hVDAC1 but expressing either native mVDAC1 or an E72Q mutant underwent apoptosis induced by various stimuli that can be inhibited by ruthenium red in the native cells but not in the mutated cells, suggesting that VDAC1 regulates apoptosis independent of the apoptosis-inducing pathway.

Keywords: apoptosis, mitochondria, short hairpin RNA

Mitochondria play a critical role in cell life, including ATP synthesis, calcium homeostasis, and cell signaling (1). Effective exchange of metabolites between mitochondria and the cytoplasm is essential for normal cell physiology. Such exchange requires transport across both the outer and the inner mitochondrial membranes (OMM and IMM, respectively). In mammalian mitochondria, metabolite transport across the IMM is carried out by ≈50 transport processes catalyzed by >30 carriers (2), whereas metabolite traffic across the OMM is supported by the voltage-dependent anion-selective channel (VDAC) (3, 4).

VDAC plays an important role as a controlled passage for adenine nucleotides (5), Ca2+ (6), and other metabolites (7) into and out of mitochondria. Indeed, it is now recognized that OMM permeability is regulated by various ligands and soluble proteins, as a result of VDAC activity modulation (3). VDAC, moreover, functions as a docking site for cellular kinases, such as hexokinase, providing the enzyme with preferential access to ATP derived from oxidative phosphorylation (3, 8). Finally, along with being an important site for the regulation of cellular energy metabolism, VDAC also serves as a site for apoptotic signaling (3, 9, 10).

Overexpression of murine or rat (11) and human or rice (12) VDAC1 in a variety of cells induced apoptotic cell death, pointing to VDAC1 as a conserved mitochondrial element of the death machinery (11–15). Such cell death could, however, be blocked by the antiapoptotic protein Bcl-2, by 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid (12), ruthenium red (RuR) (6), and hexokinase (HK) (16), all also shown to inhibit VDAC-channel activity (3, 17, 18). Moreover, the antiapoptotic effects of RuR and HK were not observed in cells expressing E72Q-mutated VDAC1 (11). Taken together, these results suggest that VDAC functions as a gatekeeper in mitochondria-mediated apoptosis. Thus, VDAC expression levels may serve as a crucial factor in the process of mitochondria-mediated apoptosis. Because VDAC controls the transport of ATP, ADP, and other metabolites between the cytosol and mitochondria, down-regulation of VDAC expression should lead to disrupted energy production.

In mammalian cells, gene-expression silencing by sequence-specific degradation of mRNA can be achieved by using small inhibitory (si)RNAs or short hairpin (sh)RNA acting through RNA interference (RNAi) (19, 20).

In this study, we have used shRNA to silence VDAC1 expression. We demonstrate that diminished human (h)VDAC1 expression resulted in growth inhibition and decreased ATP synthesis and cytosolic ADP and ATP levels. The effects produced by shRNA could, however, be prevented by expressing murine (m)VDAC1 native or mutated VDAC1 and are used to further explore the involvement of VDAC1 in apoptosis.

Results

Suppression of VDAC1 Expression and Cell Proliferation by shRNA.

RNAi has been used as a tool to control the expression of specific genes in numerous organisms (19). To interfere with the expression of endogenous VDAC1 in transformed primary human embryonal kidney T-REx-293 cells, the RNAi was performed by using the shRNA approach. Accordingly, a single shRNA targeting a coding region of hVDAC1 that differs in three nucleotides from the same region of mVDAC1 was designed (Table 1). As an initial validation, the ability of this shRNA to limit VDAC1 expression in T-REx-293 cells expressing high levels of hVDAC was tested. Several stable clones of T-REx-293 cells transfected with a plasmid encoding hVDAC1-shRNA were analyzed by Western blotting using anti-VDAC1 antibodies. Endogenous hVDAC1 expression was suppressed by 82–96% (Fig. 1A). Colony 1 was selected for further experiments. The dramatic decrease in VDAC1 expression is also clearly illustrated in representative confocal images of native and hVDAC1-shRNA-expressing cells immunostained with anti-VDAC1 antibody (Fig. 1B). The distribution of VDAC1, as visualized by confocal microscope, was punctuate in control cells and hVDAC1-shRNA-T-REx cells expressing mVDAC1, suggesting that both native hVDAC1 and recombinant mVDAC1 were mostly localized to mitochondria.

Table 1.

shRNA target nucleotide sequence in hVDAC1 and homologous sequences in mVDAC1 and in the hVDAC2 and hVDAC3 isoforms

VDAC isoform and species Sequence
hVDAC1 157-AAAGTGACGGGCAGTCTGGAA-177
mVDAC1 157-AAAGTGAACGGCAGCCTGGAA-177
hVDAC2 190-AAAGTTACTGGGACCTTGGAG-210
hVDAC3 157-AAAGCATCAGGCAACCTAGAA-177
Open in a new tab

The nucleotides that differ from those in hVDAC1 are presented in bold and underlined letters. The numbers in each sequence indicate positions in the coding sequence.

Fig. 1.

Fig. 1.

Open in a new tab

Reduction of hVDAC1 expression and cell growth in shRNA-T-REx-293 cells. T-REx-293 cells were transfected with 19 base pairs of the hVDAC1 sequence to suppress endogenous hVDAC1 expression. (A) Immunoblot analyses of hVDAC1 and actin expression in control and various stable hVDAC1-shRNA-T-REx-293 colonies were performed by using anti-VDAC1 or anti-actin antibodies. (B) Immunocytochemical analysis of VDAC1 expression in control, hVDAC1-shRNA-T-REx-293, and hVDAC1-shRNA-T-REx-293 cells expressing mVDAC1. Immunostaining using anti-VDAC1 antibodies and Alexa Fluor 488-conjugated anti-mouse antibodies as a secondary antibody was monitored by confocal microscopy. (Scale bars, 20 μm.) (C) Quantitative analysis of cell growth rates of control (•) and hVDAC1-shRNA-T-REx-293 cells (○), followed by trypan-blue staining. The results represent the mean ± SEM of four different experiments carried out with different cell cultures.

The hVDAC1-shRNA-expressing cells showing a low level of VDAC1 expression proliferated extremely slowly in comparison with normal cells (Fig. 1C). Because the hVDAC1-shRNA used is specific for VDAC1, and specific anti-VDAC1 antibodies were used (21, 22), the results suggest that VDAC1 is required for normal cell growth.

Decreased Cell Growth Is Restored by mVDAC1.

Next, it was verified whether the dramatically reduced cell growth observed upon RNAi is due to specific suppression of VDAC1 expression by the hVDAC1-shRNA used rather than being the result of interference with the expression of other proteins. Because the shRNA sequence used was designed to specifically inhibit the expression of hVDAC1 but not mVDAC1 (see Table 1), the hVDAC1-shRNA-T-REx-293 cells were transfected to express mVDAC1 under the control of an inducible tetracycline-dependent human cytomegalovirus promoter (Fig. 2). Transfecting the T-REx-293 cells with a plasmid-based tetracycline-inducible mVDAC1 expression system restored cell growth in a time- and tetracycline-concentration-dependent manner (Fig. 2 A and B). Low tetracycline concentrations (<1 μg/ml) promoted cell growth (Fig. 2 A and B) and mVDAC1 expression (Fig. 2 C and D). At the optimal tetracycline concentration (1 μg/ml), the growth rate of the cells heterologously expressing mVDAC1 was the same as that of the control T-REx-293 cells expressing native hVDAC1 (Fig. 2A). The decrease in cell growth observed at high concentrations of tetracycline was due to apoptotic cell death induced by mVDAC1 overexpression (see below).

Fig. 2.

Fig. 2.

Open in a new tab

mVDAC1 expression in stably expressing shRNA-T-REx-293 cells restores cell growth. (A) Growth of control cells (•) and hVDAC1-shRNA-T-REx-293 cells transfected with mVDAC1, grown without tetracycline (○) or with 0.2 μg/ml (■) or (□) 1 μg/ml tetracycline, was monitored as a function of time. (B) Cell growth of hVDAC1-shRNA-T-REx-293 cells transfected with mVDAC1 as a function of tetracycline concentration. Cell viability and growth rates were followed by trypan-blue staining. (C and D) The VDAC1 expression level on the 6th day of growth was analyzed in cell extracts (30 μg of protein) by using anti-VDAC1 or anti-actin antibodies as a function of the indicated tetracycline concentration (C) or as a function of time (D). (E) Quantitative analysis of immunoblots representing VDAC1 expression level as a function of tetracycline concentration or of growth time and presented as percentage of the endogenous hVDAC1 in the control cells. The results represent the mean ± SEM of four to seven different experiments carried out with different cell cultures.

The expression of mVDAC1 in these cells is clearly shown in their immunostaining with anti-VDAC1 antibodies (Fig. 1B) and by Western blot (Fig. 2 Cand D). Analysis of the Western blots demonstrated that restoring cell growth was accompanied by progressive increase in the expression level of mVDAC1. Moreover, an exponential relationship between cell growth restoration and increase in mVDAC1 expression was obtained (Fig. 2E), suggesting that cell growth required a certain minimal level of VDAC1.

ATP Synthesis and Cellular Levels Are Decreased in hVDAC1-shRNA-Expressing Cells.

To ascertain that the decreased expression of hVDAC1 leading to inhibition of cell proliferation acts through a disruption of energy production, rates of ATP synthesis by mitochondria isolated from control, hVDAC1-shRNA-T-REx-293 cells, and from the same cells expressing mVDAC1 were compared. For all cell types, a half-maximal rate of ATP synthesis was obtained at ≈100 μM ADP. However, the steady-state level of ATP synthesis by mitochondria isolated from hVDAC1-shRNA-T-REx-293 cells was 4-fold lower than that of mitochondria isolated from control cells or from hVDAC1-shRNA cells transfected to also express mVDAC1 (Fig. 3A).

Fig. 3.

Fig. 3.

Open in a new tab

Cytosolic ATP levels and mitochondrial ATP-synthesis rates are decreased in shRNA-T-REx-293 cells, a correlation between cell growth and ATP levels. Mitochondria were isolated from control, VDAC1-shRNA-T-REx-293, and VDAC1-shRNA-T-REx-293 cells expressing mVDAC induced by tetracycline (1 μg/ml) as described in ref 41. (A) ATP synthesis by control (•), VDAC1-shRNA-T-REx-293 (▴), and VDAC1-shRNA-T-REx-293 cells expressing mVDAC (○), as a function of ADP concentration, was assayed as described in Materials and Methods. ATP (black) and ADP (gray) content, determined by using luciferin/luciferase (B) and the citrate synthase activity (C) of cell extracts were assayed as described in Materials and Methods. (D) ATP levels (•) and cell growth (○) were analyzed in VDAC1-shRNA-T-REx-293 cells expressing mVDAC under the control of different concentrations of tetracycline. (Inset) The same results presented as cell growth as a function of the cellular ATP level. The results represent the mean ± SEM of four different experiments carried out with different mitochondrial preparations.

Because VDAC provides the major pathway for nucleotide movement across the outer mitochondrial membrane, the 4-fold decrease in the steady-state level of ATP synthesized may be due to limited transport of ADP and/or synthesized ATP in and out of the mitochondria. Therefore, the next step was to measure the levels of ATP and ADP in control and hVDAC1-shRNAT-REx-293 cells (Fig. 3B). The results clearly showed a decrease of ≈40% in the levels of ATP or ADP in the hVDAC1-shRNA T-REx-293 cells. The levels of ATP and ADP were, however, restored when the cells were transfected to express mVDAC1. Thus, the decrease in total ATP and ADP in the hVDAC1-shRNA-T-REx-293 cells may explain the slow growth of these cells.

To eliminate the possibility that decreased mitochondrial numbers were responsible for the observed decrease in ATP and ADP levels, the activity of citrate synthase, a marker of mitochondrial mass (23), was assayed. No difference between control and hVDAC1-shRNA cells in the content of the mitochondrial matrix enzyme citrate synthase was observed (Fig. 3C).

The relationship between restoring cell growth due to mVDAC1 expression and ATP levels in the cells (Fig. 3D) clearly indicates a tight correlation between the two, and suggests that VDAC1 controls nucleotide fluxes into and out of the mitochondria.

Apoptotic Cell Death Is Induced by Tetracycline-Regulated VDAC1 Overexpression.

It has been demonstrated that VDAC1 overexpression resulted in apoptotic cell death (11, 12). In control and hVDAC1-shRNA T-REx-293 cells transfected to express mVDAC1 under the control of low concentrations of tetracycline (1 μg/ml), only ≈5% of the cells showed nuclear fragmentation. However, at high concentrations of tetracycline (2.5 μg/ml), hVDAC1-shRNA T-REx-293 cells transfected to express mVDAC1 grew for 3 days and then started to die, with all cells dead by the 5th day (Fig. 4A). Apoptotic cell death was reflected by enhanced nuclear fragmentation (Fig. 4B). Quantitative analysis of apoptosis in the different cells showed that the exposure of cells to 2.5 μg/ml tetracycline resulted, after 4–5 days, in apoptotic cell death (70–90%). In contrast, control or hVDAC1-shRNA T-REx-293 cells expressing mVDAC1 in the presence of 1 μg/ml tetracycline showed ≈5–8% apoptotic cell death (Fig. 4C).

Fig. 4.

Fig. 4.

Open in a new tab

Tetracycline-induced mVDAC1 overexpression induces apoptotic cell death. (A) Cell growth rates were measured by trypan-blue staining of control (•) and hVDAC1-shRNA-T-REx-293 cells expressing mVDAC1 induced by tetracycline at 1 (▴) or 2.5 (○) μg/ml. (B) Cell viability of control and hVDAC1-shRNA-T-REx-293 cells expressing mVDAC1 induced by tetracycline (1 or 2.5 μg/ml) was analyzed by acridine orange/ethidium bromide staining. Arrow and arrowhead indicate cells in an early and late apoptotic state, respectively. [Scale bars, 50 μm (day 3) and 10 μm (day 5).] (C) Quantitative analysis of cell viability involves three independent experiments, as does that in B, in which early and late apoptotic cells were counted. Quantitative analysis of apoptosis in the different cells was performed by ANOVA and t test. Data show the mean ± SEM (n = 3). (D) Flow-cytometric analysis of apoptosis in control cells or hVDAC1-shRNA-T-REx-293 cells expressing mVDAC1 induced by 1 or 2.5 μg/ml tetracycline, as determined by using Annexin V-PE and 7-AAD. The percentages presented are as follows: The left corner lower regions indicate viable cells, which exclude 7-AAD and are negative for Annexin V-PE; left upper corners indicate cells which are Annexin V-PE positive but impermeable to 7-AAD; and right lower corners indicate nonviable, necrotic, or late apoptotic cells, which are positive for both Annexin V-PE staining and for 7-AAD uptake. Results are representative of three independent experiments.

To confirm that mVDAC1 overexpression mediated cell death by induction of apoptosis, flow-cytometric analysis of the cells was carried out. Phosphatidylserine, an early marker of apoptosis detected by staining with annexin-V and reduced DNA content, a marker for late apoptosis/necrosis detected by staining with 7-aminoactinomycin D (7-AAD), were analyzed (Fig. 4D). In hVDAC1-shRNA T-REx-293 cells, induction of mVDAC1 overexpression by a high concentration of tetracycline (2.5 μg/ml) resulted in an increase of ≈3-fold in both annexin V-phycoerythrin (PE)-positive/7-AAD-negative (i.e., early apoptotic cells) and annexin V-PE-positive/7-AAD-positive cells (i.e., late apoptotic cells) was obtained, as compared with control or with mVDAC1-hVDAC1-shRNA cells in the presence of a low tetracycline concentration (1 μg/ml), (Fig. 4D). The results in Fig. 4 thus point to apoptosis as the likely mechanism of cell death induced upon overexpression of mVDAC1.

RuR Did Not Protect Against Cell Death Induced by Various Stimuli in VDAC1-shRNA-T-REx-293 Cells Expressing E72Q-mVDAC1.

In a previous study, we demonstrated that RuR was unable to interact with E72Q-mVDAC1, and, hence, cells expressing the mutant protein could not be protected from apoptotic cell death by RuR (11). This finding is now confirmed for the hVDAC1-shRNA-T-REx-293 cells expressing mVDAC1 and the mutant E72Q-mVDAC1 as well (Fig. 5A). Immunoblot analysis of the level of VDAC1 in control and in hVDAC1-shRNA-T-REx-293 cells expressing either native or E72Q-mVDAC1 showed a decrease in the level of VDAC1 of 85–90% upon overexpression (2.5 μg/ml tetracycline). However, cell death was prevented by RuR only in those cells expressing native VDAC1 but not in E72Q-mVDAC1-expressing cells (Fig. 5B), indicating that the apoptotic cell death induced by VDAC1 overexpression leads to VDAC1 degradation that was prevented upon RuR protection against apoptosis (Fig. 5A). This finding suggests that it is not the increase in the total amount of VDAC1 but, rather, the increase in its functionality that is responsible for the apoptotic cell death.

Fig. 5.

Fig. 5.

Open in a new tab

RuR inhibits apoptotic cell death and VDAC degradation in native, but not E72QmVDAC1-overexpressing, cells. (A) Control, VDAC1-shRNA-T-REx-293, and VDAC1-shRNA-T-REx-293 cells expressing mVDAC or E72Q-mVDAC induced by tetracycline (1 or 2.5 μg/ml) were exposed to RuR (2 μM) 56 h after transfection. Twenty-two hours later, cell viability was analyzed by acridine orange/ethidium bromide staining as in Fig. 4C. (B) Immunoblot analysis of whole-cell extracts (30 μg of protein) was carried out by using anti-VDAC or anti-actin antibodies 22 h after exposure to RuR.

The RuR-insensitive VDAC1 mutant allows us to test for VDAC1 involvement in apoptosis induced by various stimuli by examining the protective action of RuR in cells expressing native in comparison to the RuR-insensitive mutant. Accordingly, hVDAC1-shRNA-T-REx-293 cells expressing mVDAC1 or its mutant E72Q-mVDAC1 (under the control of 1 μg/ml tetracycline) were challenged with various apoptosis-inducing agents acting by different mechanisms, i.e., As2O3 (24), curcumin (13), and staurosporine (STS) (11), and the effect of RuR on cell death as induced by these different stimuli was analyzed (Table 2). STS, curcumin, and As2O3 induced cell death in hVDAC1-shRNA-T-REx-293 cells expressing either native or E72Q-mutated mVDAC1. RuR protected against the apoptotic cell death as induced by STS, curcumin, or As2O3 of cells expressing native VDAC1, as reported for other cell lines (11–15) but not the E72Q mutant. Given the RuR-insensitive behavior of E72Q-mVDAC1, as reflected by the inability of RuR to inhibit channel activity in this mutant (11), it appears that RuR protection against apoptosis is exerted through its direct interaction with VDAC. These results suggest, therefore, that VDAC1 is a key player in apoptosis, regardless of the inducer and the mechanism by which it induces cell death.

Table 2.

RuR protects against apoptosis induced by various reagents in cells expressing mVDAC1 but not expressing E72Q-mVDAC1

Apoptosis inducer Apoptotic cell death, %
mVDAC1
 E72Q-mVDAC1
−RuR +RuR −RuR +RuR
None 5.2 ± 0.8 3.2 ± 0.2 6.0 ± 1.6 3.7 ± 0.7
Curcumin 42.7 ± 1.5 11.9 ± 1.2 48.3 ± 5.3 32.4 ± 1.4
As2O3 41.7 ± 1.9 11.4 ± 4.0 38.4 ± 0.3 33.7 ± 3.6
STS 48.7 ± 1.6 13.8 ± 2.4 50.5 ± 2.0 50.3 ± 1.1
mVDAC1 overexpression 71.3 ± 2.1 32.3 ± 2.3 75.8 ± 4.5 76.0 ± 5.4
Open in a new tab

hVDAC1-shRNA-T-REx-293 cells were transfected with a plasmid encoding native or E72Q-mutated mVDAC1. mVDAC1 expression was controlled by added tetracycline (1 μg/ml in experiments 1 to 4 or 2.5 μg/ml for experiment 5). Three days after tetracycline addition, cells were exposed for 5 h to STS (1.25 μM) or for 48 h to curcumin (120 μM) or As2O3 (6 μM) in the absence or the presence of 2 μM RuR. Cell viability was analyzed by acridine orange/ethidium bromide staining as described in Fig. 4. Data shown are the mean ± SEM (n = 3).

Discussion

In this study, we have established a hVDAC1-expression silencing system using a highly specific hVDAC1-shRNA. In this system, the level of hVDAC1 expression was dramatically decreased by 90%, indicating the effectiveness of the selected shRNA sequence. Treated cells proliferated extremely slowly in comparison with normal hVDAC1-expressing cells, but normal growth rates were restored upon expression of mVDAC1. Thus, the results suggest that inhibited cell growth by the shRNA is directly related to the absence of VDAC1.

Given the role of VDAC in energy production through controlling metabolite traffic, it is likely that the decrease in energy production observed upon down-regulation of VDAC1 expression is responsible for growth inhibition, as reflected in the strong relationship between growth and cellular ATP level (Fig. 3D). The decrease in ATP and ADP levels may reflect impaired translocation of ADP to the mitochondria and of the mitochondrially synthesized ATP to the cytosol.

The reduced growth, ATP synthesis rates, and ATP and ADP content of the hVDAC1-shRNA-TREx 293 cells could be restored to normal rates by introducing mVDAC1 through an inducible expression vector. Overexpression of mVDAC1, however, initiated a mitochondrial death cascade in these cells. Such cell death is not restricted to tetracycline-induced overexpression of mVDAC1, because the same phenomenon was observed in cell lines expressing mVDAC1-GFP, rat VDAC1 E72Q-mVDAC1 (11), or plant VDAC (12). These findings suggest that VDAC1 plays a key role in the regulation of mitochondria-mediated apoptosis.

Interestingly, RuR was found to protect against cell death induced by VDAC1 overexpression as well as by various chemical reagents (11–15). We explored the fact that RuR-mediated protection against apoptosis was not observed with E72Q-mutated VDAC1 to verify the involvement of VDAC1 in apoptosis induced by various stimuli. Our results show that RuR protection against cell death induced by STS, curcumin, or As2O3 is lost in E72Q-mVDAC1-expressing hVDAC1-shRNA-TREx 293 cells. STS, curcumin, and As2O3 induce apoptosis via different pathways. STS-induced apoptosis might be engaged in various cellular events, including suppression of p38 phosphorylation and activation of JNK (25). However, it is accepted that STS-mediated apoptosis ultimately occurs via a mitochondria-related mechanism. Curcumin has been described as an antitumoral, antioxidant, and antiinflammatory agent capable of inducing apoptosis in numerous cellular systems, mainly involving the mitochondria-mediated pathway (26), although other mechanisms have been proposed (27–29). Cell death as induced by As2O3 coincides with cytochrome c release from mitochondria inhibited by Bcl-x(L) (30).

The protective effect of RuR against cell death as induced by the various stimuli, acting through different mechanisms, suggests that RuR targets a common step in the apoptotic effect of these compounds. The inability of RuR to protect against the apoptotic cell death induced by the various stimuli in cells expressing E72Q-mVDAC1 suggests, therefore, that RuR exerts its antiapoptotic effect by direct interaction with VDAC1 (6). Thus, VDAC1 is involved in the pathways used by these apoptosis-inducing agents. How the early events, induced by STS, curcumin, or As2O3, are transmitted to mitochondria and how they affect VDAC1 activity in apoptosis is not clear. Translocation of Bax to the mitochondria (31), dissociation of mitochondrial-bound hexokinase-1 (32), Bcl-2 (33), or VDAC oligimerization may be involved, as suggested for As2O3 (24).

In conclusion, our results show that, upon shRNA silencing of VDAC1 expression, cells proliferate extremely slowly, most likely because of limited exchange of ATP/ADP between the cytosol and the mitochondrion, indicating that VDAC1 is necessary for normal cell growth. Thus, the use of hVDAC1-shRNA to interfere with VDAC1 expression constitutes a potential therapeutic measure for inhibiting cell growth. Recently, the therapeutic potential of siRNA has been recognized, particularly in areas of infectious diseases and cancer (34, 35). Silencing of Bcl-2 induced massive p53-dependent apoptosis (36), and reducing the level of the androgen receptor in prostate cancer cells led to apoptosis by disrupting the Bcl-xL-mediated survival signal (37). It should be noted that the prosurvival Bcl2 family of proteins act through mitochondria-mediated apoptosis (38) and most likely involves interaction with VDAC (39). Thus, if hVDAC1 expression is targeted by shRNA, it can disrupt the survival of high-energy-demanding cancer cell and, thus, may serve as an agent of tumor suppression. Moreover, the overexpression of VDAC1 that induced apoptotic cell death offers an alternative route of cancer therapy.

Materials and Methods

Materials.

Most reagents were purchased from Sigma. Monoclonal anti-VDAC antibodies (clone 173/045) came from Calbiochem–Novobiochem (Nottingham, U.K.). Monoclonal antibodies against actin were from Santa Cruz Biotechnology. Horseradish peroxidase-conjugated anti-mouse antibodies were obtained from Promega. Alexa Fluor-488-conjugated goat anti-mouse antibodies were from Molecular Probes. Annexin V-PE came from BD Biosciences Pharmingen.

Construction of Plasmids.

Construction of plasmid pSUPERretro encoding shRNA targeting hVDAC1.

Specific silencing of the endogenous hVDAC1 was achieved by using a shRNA-expressing vector. Nucleotides 159–177 of the hVDAC1 coding sequence were chosen as target for shRNA. This sequence is presented in Table 1, as are the homologous sequences of mVDAC, hVDAC2, and hVDAC3. The hVDAC1-shRNA-encoding sequence was created by using the two complimentary oligonucleotides indicated below, each containing the 19 nucleotides target sequence of hVDAC1 (159–177), followed by a short spacer and an antisense sequence of the target: Oligonucleotide 1, AGCTTAAAAAAGTGACGGGCAGTCTGGAA TCTCTTGAA TTCCAGACTGCCCGTCACTG and oligonucleotide 2, GATCCAGTGACGGGCAGTCTGGAATTCAAGAGATTCCAGACTGCCCGTCACTTTTTTA.

The hVDAC1-shRNA-encoding sequence was cloned into the BglII and HindIII sites of the pSUPERretro plasmid (OligoEngine, Seattle, WA), containing a puromycin-resistance gene. Transcription of this sequence by RNA-polymerase III produces a hairpin (hVDAC1-shRNA).

Construction of a plasmid for tetracycline-regulated expression of mVDAC1.

The mVDAC1 coding sequence was cloned into the BamH1 and EcoRV restriction sites of the pcDNA4/TO vector (Invitrogen) containing the zeocin-resistance gene and two tetracycline operator sites within the human cytomegalovirus immediate-early promoter to allow for tetracycline-regulated expression of the mVDAC1 in transfected cells (40).

Cell Culture.

T-REx-293 cells.

A transformed primary human embryonal kidney cell line (Invitrogen) grown under an atmosphere of 95% air and 5% CO2 in DMEM supplemented with 10% FCS, 2 mM l-glutamine, 1,000 units/ml penicillin, 1 mg/ml streptomycin, and 5 μg/ml blasticidin. Other cell lines used are stably transfected derivatives of T-REx-293 that express the tetracycline repressor.

hVDAC1-shRNA T-REx-293 cells.

T-REx-293 cells, stably transfected with the pSUPERretro plasmid encoding shRNA targeting hVDAC1 were grown with 0.5 μg/ml puromycin and 5 μg/ml blasticidin.

pc-mVDAC1-hVDAC1-shRNA T-REx-293 cells.

hVDAC1-shRNA-T-REx-293 cells were transfected with plasmid mVDAC1- or E72Q-mVDAC1-pcDNA4/TO, expressing mVDAC1 or E72Q-mVDAC1 under the control of tetracycline. Cells were grown with 200 μg/ml zeocin, 0.5 μg/ml puromycin, and 5 μg/ml blasticidin.

Transfection and Selection of Stable Transformants.

Cell transfection with plasmid pSUPERretro-shRNA-hVDAC1 by calcium phosphate.

Logarithmically growing T-REx-293 cells were transfected with plasmid pSUPERretro-shRNA-hVDAC1. Plasmid DNA (5 μg), in a solution of 125 mM CaCl2, 25 mM Hepes, 140 mM NaCl, and 0.75 mM Na2HPO4, pH 7.1, was added to each 4 ml of cell culture (≈7 × 105 cells per ml). After 4 h, 4 ml of fresh DMEM supplemented with 10% FCS, 2 mM l-glutamine, 1,000 units/ml penicillin, 1 mg/ml streptomycin, and 5 μg/ml blasticidin were added, and the cells wee incubated overnight at 37°C. The medium was then replaced, and, 48 h later, 0.5 μg/ml puromycin was added for selection of transfected cells. Growth was monitored for two weeks, with medium being refreshed every 48 h. Colonies were analyzed separately for hVDAC1 level by immunoblot using monoclonal anti-VDAC1 antibodies. A clone expressing ≈10% of normal hVDAC1 level was selected for further experiments.

Cell transfection with plasmid pcDNA4/TO encoding native or mutated mVDAC1 by calcium phosphate.

Logarithmically growing hVDAC1-shRNA-T-REx-293 cells were transfected with plasmid pcDNA4/TO-mVDAC1. Linearized NruI cut-plasmid DNA was transfected into these cells as described above, and, 48 h later, 200 μg/ml zeocin was added for selection of transfected cells. After selection, transformed cells, referred to as mVDAC-hVDAC-shRNA-REx-293 cells, containing the two plasmids, i.e., pSUPERretro-expressing shRNA and pcDNA4/TO-mVDAC1, were obtained. The selected cells were grown with 200 μg/ml zeocin, 0.5 μg/ml puromycin, and 5 μg/ml blasticidin.

Tetracycline-Induced mVDAC1 Expression.

Induction of mVDAC1 expression in hVDAC1-shRNA-T-REx-293 cells was accomplished by exposing cells to 200–2,500 ng/ml tetracycline. Cell growth rates were monitored by using trypan-blue staining. The expression levels of mVDAC1 were followed by Western blot analysis of cell extracts using monoclonal anti-VDAC1 antibodies (21, 22) and quantified by densitometry. As a control for protein amount in all samples, blotting with anti-actin antibodies was performed.

Acridine Orange/Ethidium Bromide Staining of Cells.

Cell viability was analyzed by staining with acridine orange and ethidium bromide in PBS as described in ref. 11. To record images, fluorescence microscopy (Olympus IX51) and an Olympus DP70 camera were used.

Flow-Cytometry Analysis.

Control and hVDAC1-shRNA-expressing cells (1 × 106 cells per flask) were cultured without or with tetracycline (1 or 2.5 mg/ml) for 3–5 days at 37°C in a 5% CO2 atmosphere. Cultures were exposed to trypsin (0.5% in PBS containing 0.1% EDTA) and harvested in complete DMEM. Detached cells were analyzed for early apoptotic cells by exposing to Annexin V-PE. Apoptotic cells were stained with a 7-aminoactinomycin D red fluorescent probe. Apoptotic cells were analyzed by using a FACScan cytometer (Beckton-Dickinson, Franklin Lakes, NJ). A total of 10,000 cells were counted per sample.

Light-Microscope Immunocytochemistry.

Cells cultured on cover slips in a 24-well dish were fixed by using 4% paraformaldehyde and preincubated with a blocking solution containing 5% normal goat serum, 1% BSA, and 0.2% Triton X-100 for 30 min and then incubated with anti-VDAC antibodies diluted in blocking solution containing 1% normal goat serum for 2 h at room temperature. After three washes with PBS, cells were incubated for 1 h with Alexa Fluor-488-conjugated antibody. Immunofluorescent signals were monitored by using a Zeiss LSM 510 confocal microscope.

ATP Synthesis by Isolated Mitochondria.

Mitochondria were isolated from control, hVDAC1-shRNA-T-REx-293, and hVDAC1-shRNA-T-REx-293 cells transfected to express mVDAC1 (1μg/ml tetracycline) as described in ref. 41. ATP synthesis was assayed by using an enzymatic assay coupled to NADP+ reduction.

Citrate Synthase, ATP, and ADP Levels.

Citrate synthase activity was determined in cell extracts (1 × 107 cells per ml) obtained by sonication using the coupled reaction among oxaloacetate, acetyl-CoA, and 5,5′-dithiobis (2,4-nitrobenzoic acid) (42) as monitored at 412 nm (molar coefficient = 14,150). ATP concentrations were determined by the luciferin/luciferase reaction (43). Cells (3 × 107 cells per ml) were centrifuged and resuspended in PBS, and perchloric acid was added to a final concentration of 6%. The mixture was centrifuged, and the pellet was saved for protein determination (44). The neutralized supernatant was assayed for ATP and ADP levels. ADP was measured by converting ADP to ATP with 4 mM creatine phosphate and 5 units of creatine kinase.

Acknowledgments

We thank Dr. S. Lavi for advice in establishing the shRNA system and M. Hershfinkel for help with the confocal microscope. This research was supported by a grant from the Israel Science Foundation.

Abbreviations

7-AAD

7-aminoactinomycin D

PE

phycoerythrin

RNAi

RNA interference

RuR

ruthenium red

shRNA

short hairpin RNA

siRNA

small interfering RNA

STS

staurosporine

VDAC

voltage-dependent anion channel

hVDAC

human VDAC

mVDAC

murine VDAC.

Footnotes

Conflict of interest statement: No conflicts declared.

References

  • 1.Brown G. C. Biochem. J. 1992;284:1–13. doi: 10.1042/bj2840001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Passarella S., Atlante A., Valenti D., de Bari L. Mitochondrion. 2003;2:319–343. doi: 10.1016/S1567-7249(03)00008-4. [DOI] [PubMed] [Google Scholar]
  • 3.Shoshan-Barmatz V., Gincel D. Cell Biochem. Biophys. 2003;39:279–292. doi: 10.1385/CBB:39:3:279. [DOI] [PubMed] [Google Scholar]
  • 4.Colombini M. Mol. Cell. Biochem. 2004;256–257:107–115. doi: 10.1023/b:mcbi.0000009862.17396.8d. [DOI] [PubMed] [Google Scholar]
  • 5.Rostovtseva T., Colombini M. J. Biol. Chem. 1996;271:28006–28008. doi: 10.1074/jbc.271.45.28006. [DOI] [PubMed] [Google Scholar]
  • 6.Gincel D., Zaid H., Shoshan-Barmatz V. Biochem. J. 2001;358:147–155. doi: 10.1042/0264-6021:3580147. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Hodge T., Colombini M. J. Membr. Biol. 1997;157:271–279. doi: 10.1007/s002329900235. [DOI] [PubMed] [Google Scholar]
  • 8.Cesar Mde C., Wilson J. E. Arch. Biochem. Biophys. 1998;350:109–117. doi: 10.1006/abbi.1997.0497. [DOI] [PubMed] [Google Scholar]
  • 9.Crompton M. Biochem. J. 1999;341:233–249. [PMC free article] [PubMed] [Google Scholar]
  • 10.Kroemer G., Zamzami N., Susin S. A. Immunol. Today. 1997;18:44–51. doi: 10.1016/s0167-5699(97)80014-x. [DOI] [PubMed] [Google Scholar]
  • 11.Zaid H., Abu-Hamad S., Israelson A., Nathan I., Shoshan-Barmatz V. Cell Death Differ. 2005;12:751–760. doi: 10.1038/sj.cdd.4401599. [DOI] [PubMed] [Google Scholar]
  • 12.Godbole A., Varghese J., Sarin A., Mathew M. K. Biochim. Biophys. Acta. 2003;1642:87–96. doi: 10.1016/s0167-4889(03)00102-2. [DOI] [PubMed] [Google Scholar]
  • 13.Bae J. H., Park J. W., Kwon T. K. Biochem. Biophys. Res. Commun. 2003;303:1073–1079. doi: 10.1016/s0006-291x(03)00479-0. [DOI] [PubMed] [Google Scholar]
  • 14.Baek J. H., Lee Y. S., Kang C. M., Kim J. A., Kwon K. S., Son H. C., Kim K. W. Int. J. Cancer. 1997;73:725–728. doi: 10.1002/(sici)1097-0215(19971127)73:5<725::aid-ijc19>3.0.co;2-4. [DOI] [PubMed] [Google Scholar]
  • 15.Ding W. X., Shen H. M., Ong C. N. Biochem. Biophys. Res. Commun. 2001;285:1155–1161. doi: 10.1006/bbrc.2001.5309. [DOI] [PubMed] [Google Scholar]
  • 16.Azoulay-Zohar H., Israelson A., Abu-Hamad S., Shoshan-Barmatz V. Biochem. J. 2004;377:347–355. doi: 10.1042/BJ20031465. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Shafir I., Feng W., Shoshan-Barmatz V. J. Bioenerg. Biomembr. 1998;30:499–510. doi: 10.1023/a:1020598315287. [DOI] [PubMed] [Google Scholar]
  • 18.Shoshan-Barmatz V., Hadad N., Feng W., Shafir I., Orr I., Varsanyi M., Heilmeyer L. M. FEBS Lett. 1996;386:205–210. doi: 10.1016/0014-5793(96)00442-5. [DOI] [PubMed] [Google Scholar]
  • 19.Mello C. C., Conte D., Jr Nature. 2004;431:338–342. doi: 10.1038/nature02872. [DOI] [PubMed] [Google Scholar]
  • 20.Meister G., Tuschl T. Nature. 2004;431:343–349. doi: 10.1038/nature02873. [DOI] [PubMed] [Google Scholar]
  • 21.Winkelbach H., Walter G., Morys-Wortmann C., Paetzold G., Hesse D., Zimmermann B., Florke H., Reymann S., Stadtmuller U., Thinnes F. P., et al. Biochem. Med. Metab. Biol. 1994;52:120–127. doi: 10.1006/bmmb.1994.1042. [DOI] [PubMed] [Google Scholar]
  • 22.Babel D., Walter G., Gotz H., Thinnes F. P., Jurgens L., Konig U., Hilschmann N. Biol. Chem. Hoppe-Seyler. 1991;372:1027–1034. doi: 10.1515/bchm3.1991.372.2.1027. [DOI] [PubMed] [Google Scholar]
  • 23.Moraes C. T., Ciacci F., Bonilla E., Jansen C., Hirano M., Rao N., Lovelace R. E., Rowland L. P., Schon E. A., DiMauro S. J. Clin. Invest. 1993;92:2906–2915. doi: 10.1172/JCI116913. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Zheng Y., Shi Y., Tian C., Jiang C., Jin H., Chen J., Almasan A., Tang H., Chen Q. Oncogene. 2004;23:1239–1247. doi: 10.1038/sj.onc.1207205. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Zhang H., Vollmer M., De Geyter M., Durrenberger M., De Geyter C. Int. J. Oncol. 2005;26:1575–1580. [PubMed] [Google Scholar]
  • 26.Duvoix A., Blasius R., Delhalle S., Schnekenburger M., Morceau F., Henry E., Dicato M., Diederich M. Cancer Lett. 2005;223:181–190. doi: 10.1016/j.canlet.2004.09.041. [DOI] [PubMed] [Google Scholar]
  • 27.Kwon Y. K., Jun J. M., Shin S. W., Cho J. W., Suh S. I. Int. J. Oncol. 2005;26:1597–1603. [PubMed] [Google Scholar]
  • 28.Bharti A. C., Donato N., Aggarwal B. B. J. Immunol. 2003;171:3863–3871. doi: 10.4049/jimmunol.171.7.3863. [DOI] [PubMed] [Google Scholar]
  • 29.Liu H. L., Chen Y., Cui G. H., Zhou J. F. Acta Pharmacol. Sin. 2005;26:603–609. doi: 10.1111/j.1745-7254.2005.00081.x. [DOI] [PubMed] [Google Scholar]
  • 30.Scholz C., Wieder T., Starck L., Essmann F., Schulze-Osthoff K., Dorken B., Daniel P. T. Oncogene. 2005;24:1904–1913. doi: 10.1038/sj.onc.1208233. [DOI] [PubMed] [Google Scholar]
  • 31.Chandra D., Choy G., Daniel P. T., Tang D. G. J. Biol. Chem. 2005;280:19051–19061. doi: 10.1074/jbc.M501391200. [DOI] [PubMed] [Google Scholar]
  • 32.Pastorino J. G., Shulga N., Hoek J. B. J. Biol. Chem. 2002;277:7610–7618. doi: 10.1074/jbc.M109950200. [DOI] [PubMed] [Google Scholar]
  • 33.Chow S. K., Chan J. Y., Fung K. P. J. Cell. Biochem. 2004;93:173–187. doi: 10.1002/jcb.20102. [DOI] [PubMed] [Google Scholar]
  • 34.Tong A. W., Zhang Y. A., Nemunaitis J. Curr. Opin. Mol. Ther. 2005;7:114–124. [PubMed] [Google Scholar]
  • 35.Ichim T. E., Li M., Qian H., Popov I. A., Rycerz K., Zheng X., White D., Zhong R., Min W. P. Am. J. Transplant. 2004;4:1227–1236. doi: 10.1111/j.1600-6143.2004.00530.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Jiang M., Milner J. Genes Dev. 2003;17:832–837. doi: 10.1101/gad.252603. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Liao X., Tang S., Thrasher J. B., Griebling T. L., Li B. Mol. Cancer Ther. 2005;4:505–515. doi: 10.1158/1535-7163.MCT-04-0313. [DOI] [PubMed] [Google Scholar]
  • 38.Gross A., McDonnell J. M., Korsmeyer S. J. Genes Dev. 1999;13:1899–1911. doi: 10.1101/gad.13.15.1899. [DOI] [PubMed] [Google Scholar]
  • 39.Shimizu S., Narita M., Tsujimoto Y. Nature. 1999;399:483–487. doi: 10.1038/20959. [DOI] [PubMed] [Google Scholar]
  • 40.Yao F., Svensjo T., Winkler T., Lu M., Eriksson C., Eriksson E. Hum. Gene Ther. 1998;9:1939–1950. doi: 10.1089/hum.1998.9.13-1939. [DOI] [PubMed] [Google Scholar]
  • 41.Bourgeron T., Chretien D., Rotig A., Munnich A., Rustin P. Biochem. Biophys. Res. Commun. 1992;186:16–23. doi: 10.1016/s0006-291x(05)80769-7. [DOI] [PubMed] [Google Scholar]
  • 42.Bond D. R., Mester T., Nesbo C. L., Izquierdo-Lopez A. V., Collart F. L., Lovley D. R. Appl. Environ. Microbiol. 2005;71:3858–3865. doi: 10.1128/AEM.71.7.3858-3865.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43.Tam K. F., Ng T. Y., Liu S. S., Tsang P. C., Kwong P. W., Ngan H. Y. Gynecol. Oncol. 2005;96:765–770. doi: 10.1016/j.ygyno.2004.11.025. [DOI] [PubMed] [Google Scholar]
  • 44.Lowry O. H., Rosebrough N. J., Farr A. L., Randall R. J. J. Biol. Chem. 1951;193:265–275. [PubMed] [Google Scholar]

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences

RESOURCES