Photodynamic Therapy-Induced Apoptosis in Lymphoma Cells: Translocation of Cytochrome c Causes Inhibition of Respiration as Well as Caspase Activation

Biochemical and Biophysical Research Communications 255, 673– 679 (1999) Article ID bbrc.1999.0261, available online at http://www.idealibrary.com on

Photodynamic Therapy-Induced Apoptosis in Lymphoma Cells: Translocation of Cytochrome c Causes Inhibition of Respiration as Well as Caspase Activation Marie E. Varnes, 1 Song-Mao Chiu, Liang-Yan Xue, and Nancy L. Oleinick Department of Radiation Oncology, Case Western Reserve University, Cleveland, Ohio 44106-4942

Received January 21, 1999

L5178Y-R mouse lymphoma (LY-R) cells undergo rapid apoptosis when treated with photodynamic therapy (PDT) sensitized with the silicon phthalocyanine Pc 4. In this study we show that cytochrome c is released into the cytosol within 10 min of an LD 99.9 dose of PDT. Cellular respiration is inhibited by 42% at 15 min, and 60% at 30 min after PDT treatment, and caspase 3-like protease activity is elevated by 15 min post-PDT. In digitonin-permeabilized cells addition of cytochrome c to the respiration buffer reverses PDTinduced inhibition of state 3 respiration via Complex I by 40 – 60%, and via Complex III by 50 –90%. In contrast, extramitochondrial cytochrome c does not stimulate respiration in permeabilized control cells, and catalyzes only a low rate of oxygen consumption via electron transfer to cytochrome b5 on the outer mitochondrial membrane. These results demonstrate that PDTinduced inhibition of respiration is primarily due to leakage of cytochrome c into the cytosol rather than to damage to the major enzyme complexes of the electron transport chain. Whether or not inhibition of respiration influences the time course or extent of Pc 4-PDTinduced apoptosis in LY-R cells is not clear at the present time. © 1999 Academic Press Key Words: photodynamic therapy; cytochrome c; respiration; apoptosis; caspases.

Photodynamic therapy (PDT) is a treatment modality in which a photosensitive compound, accumulated 1 To whom correspondence should be addressed at Department of Radiation Oncology, Case Western Reserve University, School of Medicine, BRB, 10900 Euclid Avenue, Cleveland, Ohio 44106-4942. Fax: 216-368-1142. E-mail: [email protected]. Abbreviations: AMC, 7-amino-4-methyl-coumarin; Chaps, 3-[3cholanidopropyl)-dimethyl-ammonio] 1-propanesulfonate; cyto-c, cytochrome c; Pc 4, silicon phthalocyanine, [HOSiPcOSi(CH 3) 2(CH 2) 3 N(CH 3) 2]; PDT, photodynamic therapy; PMSF, phenyl-methanesulfonyl flouride; TMPD, N,N,N9,N9-tetramethyl-p-phenylenediamine; zVADfmk, benzyloxycarbonyl-Val-Ala-Asp(O-methyl)-fluoromethylketone.

in tumor cells or tissue, is activated by illumination with visible light. The interaction among photosensitizer, light and oxygen produces singlet oxygen and other reactive oxygen species which rapidly induce an intermitotic cell death (1, 2). PDT of mammalian cells in culture can induce apoptosis, necrosis, or cell death with characteristics of both processes, depending on the cell type, the nature of the photosensitizer and the severity of the treatment conditions (1– 6). Oleinick and colleagues have previously shown that PDT sensitized with either chloroaluminum phthalocyanine or the silicon phthalocyanine Pc 4 induces rapid, classical apoptosis in L5178Y-R (LY-R) mouse lymphoma cells (7–9). PDT induces a variety of signaling events that have been implicated in apoptosis (1, 2). Signals so far observed upon treatment of LY-R cells with Pc 4-PDT are (a) sphingomyelinase activation and ceramide release (9), (b) tyrosine phosphorylation of the HS1 subunit of a membrane-associated src family kinase (10), (c) activation of the stress kinases SAPK/JNK and p38/HOG1 (11), and (d) activation of caspase 3-like proteases (8). Which of these is the principal causative factor for apoptosis, or whether an interaction of several signaling pathways determines the fate of LY-R cells, is not yet clear. Recently our attention has focused on the role of the mitochondria in PDT-induced apoptosis, for several reasons. First, caspase 3 has been identified as a component of a mitochondrial apoptogenic pathway leading from release of cytochrome c (cyto-c) into the cytosol to activation of a cascade of proteases and subsequent activation of the endonuclease DFF-40 (12– 16). Secondly, Oleinick and colleagues 2 have recently found that Pc 4 accumulates in mitochondria and intracellular organelles but not in the plasma membrane. Kessel et al. (6, 17) showed that photosensitizers which localize specifically to the mitochondria induce apoptosis in murine leukemia cells, whereas photosen2

N. L. Oleinick, A. L. Nieminen, and N. Trivedi, unpublished observations. Mentioned in (1).

673

0006-291X/99 $30.00 Copyright © 1999 by Academic Press All rights of reproduction in any form reserved.

Vol. 255, No. 3, 1999

BIOCHEMICAL AND BIOPHYSICAL RESEARCH COMMUNICATIONS

sitizers which localize to lysosomes or plasma membranes kill the cells preferentially by necrosis. Thirdly, PDT is a well-known inducer of various kinds of mitochondrial damage including uncoupling of respiration and oxidative phosphorylation (18, 19), inhibition of the enzyme complexes of the electron transport chain (20 –22), inactivation of the permeability transition pore (23) and disruption of the transmembrane potential (6). Mitochondrial damage resulting in increased membrane permeability preceeds release of cyto-c and subsequent initiation of apoptosis (24). This study shows that Pc 4-PDT initiates a rapid release of cyto-c from the mitochondria into the cytosol, followed by partial inhibition of respiration and activation of caspase 3-like proteases. In digitoninpermeabilized cells respiratory inhibition can be significantly reversed by adding exogenous cyto-c to the respiration buffer. The latter results demonstrate that inhibition of respiration is a consequence of loss of cyto-c from the inner membrane space rather than of direct damage to the enzyme complexes of the respiratory chain. MATERIALS AND METHODS Materials. T h e p h t h a l o c y a n i n e p h o t o s e n s i t i z e r P c 4 [HOSiPcOSi(CH 3 ) 2 (CH 2 ) 3 N(CH 3 ) 2 ] (25) was supplied by Drs. Yingsyi Li and Malcolm Kenney of the Department of Chemistry, CWRU, and used as a 0.5 mM stock solution in dimethyl formamide. The solution was stored in the dark at 4°C, and was stable for up to one month after preparation. High purity digitonin (Calbiochem, La Jolla, CA) was prepared as a 5 mg/ml solution in DMSO and stored at 220°C until use. Cytochrome c (Sigma, St. Louis, MO) was prepared as a 5 mg/ml stock solution in Hepesbuffered PBS, pH 7.2. Other reagents were obtained from Sigma. The Bradford assay (Biorad, Richmond, CA) was used for protein determinations. Cell culture and photodynamic treatment. LY-R cells (26) were grown in Fisher’s medium supplemented with 10% heat-inactivated horse serum and 5 mM glutamine. For experiments, cells were seeded at about 2.0 3 10 5/ml and grown overnight in Fisher’s medium containing 1% serum (low serum medium, LSM). For PDT experiments cells in LSM were treated with 200 nM Pc 4 for 18 h before light exposure. Cells were irradiated using an EFOS LED array (EFOS, Mississauga, ONT, Canada) at a fluence of 45 mJ/cm 2 (1.1 mJ/cm 2/s, l max ; 670 – 675 nm). After light exposure the cells were kept in the dark at 37°C until use. Respiration and mitochondrial electron transport. Respiration in intact cells was measured using a Clark-type polarographic electrode and Model 53 Biological Oxygen Monitor (YSI, Inc., Yellow Springs, OH). Cells were pelleted, then resuspended at 5 3 10 6 cells/ml in bicarbonate-free tissue culture medium containing 10 mM Hepes and 5% heat-inactivated horse serum, pH 7.2. The cells were placed in the thermostated (37°C) electrode chamber and stirred for a few minutes before inserting the electrode. Respiratory rates were calculated from the initial, linear portion of the curves, as pmol O 2 consumed/min/10 6 cells. Respiration in permeabilized cells was measured as described by Hofhaus et al. (27). Control or PDT-treated cells were pelleted, then resuspended in sucrose buffer A (250 mM sucrose, 10 mM MgCl 2 and 20 mM Hepes, pH 7.1) containing 75 mg/ml digitonin, for 1 min at room temperature. The cells were washed once with buffer A, then resuspended in respiration medium (buffer A containing 1 mM ADP

and 2 mM potassium phosphate) at a density of 5 3 10 6 cells/ml and placed in the electrode chamber. After allowing a few minutes for endogenous substrates to be consumed, various substrates and inhibitors of Complexes I, III, and IV of the electron transport chain were introduced into the chamber using a long-needled Hamilton syringe (Hamilton Co., Reno, NV). Concentrations of the various substrates and inhibitors were as follows: malate, glutamate, glycerophosphate and succinate, each 5 mM; ascorbate, 10 mM; TMPD (N,N,N9,N9-tetramethyl-p-phenylenediamine), 200 nM; rotenone, 0.1 mM, antimycin A, 20 nM; KCN, 0.1 mM. Respiratory rates for each complex were calculated as pmol O 2 consumed/min/10 6 cells. Cyto-c release from the mitochondria. A modification of the procedure described by Yang et al. (15) was used. Cells were washed in cold sucrose buffer B (0.25 M sucrose, 10 mM Tris–HCl, 3 mM MgCl 2, pH 7.4,) and resuspended in 5 volumes of lysis buffer (250 mM sucrose, 10 mM KCl, 1.5 mM MgCl 2, 1.0 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, 0.1 mM PMSF, and 20 mM Hepes, pH 7.5). The cells were homogenized using 12 strokes of a fitted Teflon pestle. The homogenate was centrifuged at 750g for 10 min at 4°C, then the supernate was further centrifuged at 10,000g for 15 min. The resulting mitochondrial pellets were lysed and boiled for 5 min in 20 ml of protein gel loading buffer (28). The resulting solution was passed through a 0.2-mm filter, and then through a 0.1-mm Ultrafree-MC filter (Millipore Corp., Bedford, MA). The filtrate was precipitated with acetone. Equal amounts of mitochondrial or cytosolic protein were electrophoresed on 15% SDS–polyacrylamide (28). After transferring protein from the gels onto PVDF membranes, the membranes were probed with anti-cytochrome c (7H8.2C12, PharMingen, San Diego, CA) or anti-cytochrome oxidase (Mab anti subunit IV, Molecular Probes, Eugene, OR). The immune complexes were detected using the ECL system (Amersham, Arlington Heights, IL). For these experiments the cells were treated with PDT while in medium containing 10% serum, since cell fractionation procedures gave inconsistent results with cells maintained in the LSM. Caspase 3-like activity. Approximately 2–5 3 10 6 cells were collected by centrifugation and washed once with sucrose buffer B. The cells were resuspended in 200 ml of Buffer C (10 mM KCl, 1 mM EDTA, 1 mM EGTA, 1 mM PMSF, 100 mM pepstatin, 100 mM leupeptin, 0.5% Triton X-100 and 20 mM Hepes, pH 7.5), incubated on ice for 20 min, then sonicated and stored at 280°C until use. For assay of caspase 3 activity, aliquots containing 25 mg protein were incubated at 37°C for 1 h in 60 ml of caspase reaction buffer (10% sucrose, 0.1% Chaps,1 mM EGTA, 1 mM EDTA, 5 mM DTT, 1 mM PMSF, 100 mM pepstatin, 100 mM leupeptin, 50 mM DEVD-AMC (BIOMOL, Plymouth Meeting, PA), 25 mM Hepes, pH 7.4). The released fluorescence was measured in a Perkin–Elmer LS50 fluorometer (l ex380 nm and l em460 nm).

RESULTS When LY-R cells in medium containing 1% serum are pre-treated with 200 nM Pc 4 for 18 –24 h then subjected to 45 mJ/cm 2 red light (EFOS LED array), they undergo rapid apoptosis, with DNA ladders evident by 30 min after treatment and DNA degradation complete by 2 h post-treatment (9). The percent of the total cell population exhibiting apoptosis is 3.4, 27, 66, and 84.9% for 15 min, 30 min, 1 h, and 2 h after light exposure, respectively. Figure 1 shows that significant levels of cyto-c are released from the mitochondria into the cytosol of PDTtreated cells by 10 min after light exposure. Cytochrome oxidase staining was used to confirm that cytosolic extracts were not contaminated by mitochondria.

674

Vol. 255, No. 3, 1999

BIOCHEMICAL AND BIOPHYSICAL RESEARCH COMMUNICATIONS

FIG. 1. Release of cyto-c from the mitochondria of PDT-treated cells. LY-R cells were treated with 200 nM Pc 4 and 45 mJ/cm 2 red light, then returned to a 37°C incubator and kept in the dark. At the indicated times the cells were harvested, resuspended in lysis buffer, homogenized and subjected to differential centrifugation to separate the mitochondria from cytosolic proteins. Equal amounts of mitochondrial and cytosolic proteins were electrophoresed on a 15% SDS polyacrylamide gel then transferred to PVDF membranes. The membranes were probed with anti-cytochrome c and anti-cytochrome oxidase as described in the Methods, and the immune complexes were detected using the ECL system.

As explained in the Methods, these experiments were done with cells in medium containing 10% serum rather than in LSM. Under such conditions all the cells die via apoptosis, but the onset of caspase activation is about 5–10 min later than when cells are treated in LSM (S-M. Chiu, manuscript in preparation). Release of cyto-c from the mitochondria sometimes, but not always, causes inhibition of respiration (29). Figure 2 shows that respiration is inhibited in PDTtreated LY-R cells, by 42 6 7% at 15 min posttreatment and by 60 6 5% at 30 min posttreatment. The times indicated in Fig. 2 represent the interval between exposure to red light and harvesting of the samples for assay. About 10 min was required to wash the cells and allow temperature equilibration of the samples. We do not know if oxidative damage to the mitochondria may have continued during sample preparation; however, rates of respiration for both control and PDT-treated cells were linear for at least 8 min after initiating the analysis. When the plasma membrane of cells is selectively permeabilized with digitonin and the cells are resuspended in buffer containing ADP, phosphate and a respiratory substrate, the cells carry out state 3 rspiration according to the scheme outlined below (27, 30).

FIG. 2. Inhibition of respiration in LY-R cells after PDT treatment. The cells were treated with 200 nM Pc 4 and 45 mJ/cm 2 red light. At the times indicated, the cells were harvested by centrifugation, resuspended in bicarbonate-free medium containing 1% serum, then analyzed for the rate of oxygen consumption using a Clark-type oxygen electrode. The dashed line represents the percent of cells remaining viable, i.e., nonapoptotic, at each time point, as determined by Separovic et al. (7) using the terminal deoxynucleotide transferase assay and flow cytometry. The average control respiratory rate was 649 6 100 pmol O 2 consumed/min/10 6 cells, measured in four separate experiments.

Malate and glutamate, via their dehydrogenases, reduce NAD to NADH, which then transfers electrons in a series of steps through Complex I (NADH-Q reductase), ubiquinone (Q), Complex III (QH 2-cytochrome c reductase), cyto-c, Complex IV (cytochrome c oxidase), and finally to O 2. Succinate and glycerophosphate (G3P) feed into the transport chain at Q, and ascorbate and TMPD donate electrons directly to cyto-c. Using specific substrates for and inhibitors of complexes I, III, and IV, one can measure the integrity of electron flow through each of these three complexes. However, since the rate of oxygen consumption is the measured parameter, only the activity of complex IV can be assessed independently of the others (27). Figure 3 shows the effect of PDT on respiration in permeabilized cells, 20 min after light exposure. Each curve is representative of the results of more than three separate experiments. For untreated cells the

SCHEME 1 675

Vol. 255, No. 3, 1999

BIOCHEMICAL AND BIOPHYSICAL RESEARCH COMMUNICATIONS

FIG. 3. Respiration in untreated (control) and PDT-treated cells after permeabilization of the plasma membranes with digitonin. Control: Permeabilized cells were resuspended at 5 3 10 6 cells/ml in 2 ml of respiration medium, and the following reagents were added sequentially, as indicated by the arrows: GM, glutamate and malate; R, rotenone; GS, glycerophosphate and succinate; A, antimycin A; TA, TMPD and ascorbate; KCN. Concentrations of each reagent are given in the Methods. Rates of respiration for Complex I (C-1), Complex III (C-III) and Complex IV (C-IV) are given in parenthesis as pmol/O 2 consumed/min/10 6 cells. PDT-treated: Cells were treated with 200 nM Pc 4 and 45 mJ/cm 2 red light, and 20 min later were subjected to membrane permeabilization and analysis of respiration. Additions of substrates and inhibitors of each complex, and calculations of respiratory rates, are the same as for controls. The data are from a representative experiment.

average rates of oxygen consumption were 655 6 223, 763 6 260, and 2559 6 1043 pmol O 2/min/10 6 cells, for glutamate/malate, succinate/G3P and ascorbate/TMPD as substrates, respectively. Because of the variability of these rates among different cell preparations, respiration in PDT-treated cells was always compared to that for the controls assayed on the same day. In Figure 3, PDT treatment reduced oxygen consumption by 62, 34, and 23% for Complexes IV, III, and I, respectively. These percentages may reflect the observed order of rates of oxygen consumption in untreated cells, i.e., Complex IV . Complex III . Complex I. PDT is known to induce oxidation of membrane cholesterol (31). Thus PDT-treated cells might be especially sensitive to digitonin, which acts on cholesterol (27, 30). Over-treatment could cause partial permeabilization of the mitochondria such that the respiratory inhibition seen in Figure 3 could be due to an artifact of sample preparation. To test this, 1.25 3 10 5 untreated or PDT-treated cells were pelleted, resuspended in various concentrations of digitonin, and incubated for 1 min at room temperature. After being washed in sucrose buffer A, the cells were treated with 0.04% trypan blue for three min, then the percentage of permeabilized cells was scored. Figure 4 shows that control and PDT-treated cells respond similarly to digitonin treatment. The slightly elevated percentage of trypan blue positive cells at the lower digitonin concentrations reflects a slightly higher background in the PDT-treated cells. Lower concentrations of digitonin were used in

this experiment than for respiration measurements because the cell density was less (30). We found that respiration via Complexes I and III in PDT-treated permeabilized cells could be significantly restored by adding exogenous cyto-c to the respiration

FIG. 4. Effect of digitonin concentration on plasma membrane permeability of untreated and PDT-treated cells. A total of 1.25 3 10 5 cells were collected by gentle centrifugation and resuspended in 250 ml of sucrose buffer A (see Methods) containing the indicated concentrations of digitonin. The cells were incubated in digitonin solution for 1 min at room temperature, then washed two times in PBS and stained with trypan blue. Control: Untreated cells incubated in LSM for 18 –24 h before assay. PDT-treated: Treatment conditions were as described for Figures 1 and 2, and cells were harvested for assay 30 min after exposure to red light.

676

Vol. 255, No. 3, 1999

BIOCHEMICAL AND BIOPHYSICAL RESEARCH COMMUNICATIONS

FIG. 5. Effects of exogenous cyto-c on respiration in permeabilized cells. Upper panel: Cells were treated with 200 nM Pc 4 and 45 mJ/cm 2 red light, held in the dark for 20 min after PDT, then permeabilized with digitonin. The cells were resuspended in respiration medium at a density of 5.0 3 10 6/ml. (A) The respiratory rate with glutamate and malate (GM) as substrates was 56 pmol O 2 consumed/min/10 6 cells. Addition of 30 mg/ml cyto-c increased the rate to 438 pmol O 2/min. (B) The respiratory rate with succinate and glycerophosphate (GS) as substrates was 45 pmol O 2/min. Addition of 20 mg/ml cyto-c increased the rate to 377; it was not further increased by subsequent increases in cyto-c concentration to 40 and 60 mg/ml. Lower panel: Cells incubated in LSM for 18 h were harvested and resuspended in respiration medium as described above. (C) The respiratory rate with GM as substrates was 443 pmol O 2/min/10 6 cells. Cyto-c (30 mg/ml) added after rotenone, induced a slow rate of oxygen consumption, 38 pmol O 2/min. (D) The rate with GM as substrates was 457 pmol O 2/min, and was decreased slightly, to 422, by addition of cyto-c. There was a residual rate of 31 pmol/min after rotenone treatment. (E) The respiratory rate with GS as substrates was 435 pmol O 2/min. Addition of 30 mg/ml cyto-c, after inhibition of respiration with antimycin A, caused little oxygen consumption, 8 pmol O 2/min. The data are from representative experiments.

buffer (Figure 5, upper panel). In (A) addition of 30 mg/ml of cyto-c increased the respiratory rate by 7.8fold. In (B) the rate was increased by 8.4-fold when 20 mg/ml cyto-c was added, and was not increased further by higher concentrations of cyto-c. Rotenone (A) and antimycin A (B) inhibited respiration completely, in the presence or absence of cyto-c. In four separate experiments addition of 30 mg/ml cyto-c to PDT-treated cells resulted in 40 – 60% recovery of the respiratory rate when glutamate/malate were used as substrates, and 50 –90% recovery when succinate/G3P were used as substrates. Cyto-c had no effect on the respiratory rate of PDT-treated cells when ascorbate/TMPD were used as substrates (data not shown). KCN could not be used to inhibit complex IV when exogenous cyto-c was present. If the outer mitochondria of LY-R cells contains cytochrome b5, it is possible that exogenous cyto-c could accept electrons from b5 and transfer them to cytochrome oxidase, thus shunting electrons around the points of PDT-induced damage rather than truly reversing the damage (32–34). To test this, untreated permeabilized cells were stimulated with glutamate

and malate to obtain a baseline respiratory rate, then respiration was inhibited with rotenone. Cyt c (30 mg/ ml) was added after rotenone addition. Residual O 2 consumption would be due to b5 shunting, since rotenone completely inhibits electron flow from glutamate/ malate to O 2 (see diagram). Figure 5 (C) shows that cyto-c causes only a small (8.5%) increase in the rate of O 2 consumption when added to untreated cells after rotenone treatment. Thus b5 shunting is not a significant factor in the effect of cyto-c on respiration in PDT-treated cells. Cyto-c slightly inhibits respiration in control cells when added immediately after glutamate/malate (D) or succinate/G3P, and only slightly increases respiration after its inhibition with antimycin A (E). He et al. (8) previously showed that caspase 3-like protease activity is elevated when LY-R cells are treated with Pc 4-PDT. However these investigators used a different light source than used in the present study. Figure 6 shows that caspase 3-like enzymes are activated by 15 min after treatment of the cells in LSM with 45 mJ/cm 2 red light, and that activation is maximal by 30 min after light exposure.

677

Vol. 255, No. 3, 1999

BIOCHEMICAL AND BIOPHYSICAL RESEARCH COMMUNICATIONS

FIG. 6. Activation of caspase 3-like activity (DEVD-ase) in LY-R cells after PDT treatment. Treatment conditions were as described for the previous figures. At the times indicated, cells were collected and lysed, then incubated with DEVD-AMC at 37°C for 1 h. The fluorescence intensity due to release of AMC was measured at various times after treatment, and compared to that of cells which were not exposed to light (0 time). Data represent the means from at least four independent experiments, 6 SD.

DISCUSSION In the present study we demonstrate that an LD 99.9 dose of Pc 4-PDT induces loss of cyto-c from the mitochondria of LY-R cells, inhibits respiration and causes activation of caspase 3-like proteases, all within 15 min of light exposure. The studies of others strongly support the hypothesis that cyto-c leakage preceeds, and is causative for, activation of caspase 3-like proteases (12–16). Herein we have focused on the relationship between loss of cyto-c from the mitochondria and inhibition of respiration. In order to examine the rates of electron flow through each of the respiratory Complexes I, III, and IV, we permeabilized the plasma membranes of LY-R cells with digitonin, then investigated state 3 respiration using specific substrates for and inhibitors of each complex (Figs. 3 and 4). Respiration in untreated permeabilized cells was 655 6 223 pmol O 2 /min/10 6 cells, compared to 649 6 100 in untreated intact cells. Although the variability in respiratory rates among experiments was high for digitonin-treated cells, the extent of inhibition of respiration by PDT was also similar for intact and permeabilized cells (Figs. 1, 3, and 4). The major observation of this study is that addition of cyto-c to the medium of PDT-treated, permeabilized cells significantly reverses the inhibition of respiration. Reversal ranged from 40 – 60% with glutamate and malate as substrates, and from 50 –90% with succinate and glycerophosphate as substrates, relative to the corresponding controls, in four separate experiments. This leads us to hypoth-

esize that PDT-induced inhibition of respiration is primarily due to translocation of cyto-c from the mitochondria into the cytosol rather than to inactivation of the enzyme complexes of the respiratory chain. This study does not address the issue of whether or not Pc 4-PDT affects other mitochondrial components, such as the permeability transition pore or the F 0 F 1 ATPase proton pump (1, 2, 18 –23). The mechanism by which cyto-c is released from the mitochondria of Pc 4-PDT-damaged cells is currently under investigation. 2 Caspase activation is a principal determinant of Pc 4- PDT-induced apoptosis in LY-R cells, since apoptosis can be nearly completely blocked by incubation of the cells with the pan-caspase inhibitor zVAD-fmk (8). Whether or not inhibition of respiration plays a secondary role in the apoptotic process is not yet clear. Respiration and glycolysis together regulate ATP levels in most cells. Changes in ATP levels are known to affect the expression of apoptosis (24), but the relationship between ATP generation and mode of cell death varies with the cell type and the cytotoxic agent. For example, depletion of ATP in Jurkat cells to ,50% of controls converts staurosporine-induced death from apoptosis to necrosis, but ATP depletion to #70% is necessary to change the mode of aCD95-induced death (34). Kessel and Luo (6) used oligomycin in glucose-free medium to reduce ATP levels in P388 cells to ,5% of controls, and observed no effect on apoptosis induced by LD 90 doses of PDT with two mitochondria-localizing photosensitizers. We have not yet tested the effects of varying ATP levels on PDT-induced apoptosis in LY-R cells. Another way in which inhibition of respiration may affect the response to PDT is by altering the amount or distribution of reactive oxygen species generated by PDT treatment. Light activation of photosensitizers produces singlet oxygen and other reactive oxygen species in cells (1, 2, 36, 37). Gibson and Hilf (21) showed that the photosensitivity of cytochrome c oxidase in isolated mitochondria increases as the oxygen concentration is increased from 1 to 10%. Oxygen tension might become higher in the mitochondria of PDT-treated LY-R cells as respiration is progressively inhibited. If free radical waves continue to be generated for some time after the initial light exposure, the higher O 2 tension could increase radical production and thus exacerbate PDT damage. We plan to use fluorescent indicators of free radicals to test this hypothesis in the near future. ACKNOWLEDGMENTS The authors wish to thank Dr. Charles Hoppel (Department of Pharmacology, CWRU) for his very helpful discussions of this work and Dr. Anna-Liisa Nieminen (Department of Anatomy, CWRU) for her critical review of this manuscript. This work was supported by Grants P01 CA48735 and P30 CA 43703 from the National Cancer Institute, NIH.

678

Vol. 255, No. 3, 1999

BIOCHEMICAL AND BIOPHYSICAL RESEARCH COMMUNICATIONS

REFERENCES 1. Oleinick, N. L. and Evans, H. H. (1998) Rad. Res. 150 (Suppl.), S146-S156. 2. Dougherty, T. J., Gomer, C. J., Henderson, B. W., Jori, G., Kesse, D., Korbelik, M., Moan, J. and Peng, Q. (1998) J. Natl. Cancer Inst. 90, 889 –902. 3. He, J. and Oleinick, N. L. (1995) Proc. SPIE Int. Soc. Opt. Eng. 2371, 92–96. 4. Dellinger, M. (1996) Photochem. Photobiol. 64, 182–187. 5. Fisher, A. M. R., Danenberg, K., Banerjee, D., Bertino, J. R., Danenberg, P. and Gomer, C. J. (1997) Photochem. Photobiol. 66, 265–270. 6. Kessel, D. and Luo, Y. (1998) J. of Photochem. Photobiol. 42, 89 –95. 7. Agarwal, M. L., Larkin, H. E., Zaidi, S. I. A., Mukhtar, H. and Oleinick, N. L. (1993) Cancer Res. 53, 5897–5902. 8. He, J., Whitacre, C. M., Xue, L.-Y., Berger, N. A. and Oleinick, N. L. (1998) Cancer Res. 58, 940 –946. 9. Separovic, D., He, J. and Oleinick, N. L. (1997) Cancer Res. 57, 1717–1721. 10. Xue, L.-Y., He, J. and Oleinick, N. L. (1997) Photochem. Photobiol. 66, 105–113. 11. Oleinick, N. L., He, J., Xue, L.-Y. and Separovic, D. (1998) Proc. SPIE Int. Soc. Opt. Eng. 3247, 82– 88. 12. Liu, X., Zou, H., Slaughter, C. and Wang, X. (1997) Cell 89, 175–184. 13. Cohen, G. M. (1997) Biochem. J. 326, 1–16. 14. Zamzami, N., Hirsch, T., Dallaporta, B., Petit, P. X. and Kroemer, G. (1997) J. Bioenergetics and Biomembranes 29, 185–193. 15. Yang, J., Liu, X., Bhalla, K., Kim, C. N., Ibrado, A. M., Cai, J., Peng, T-I., Jones, D. P. and Wang, X. (1997) Science 275, 1129 –1132. 16. Green, D. R. and Reed, J. C. (1998) Science 281, 1309 –1312. 17. Kessel, D., Luo, Y., Deng, Y. and Chang, C. K. (1997) Photochem. Photobiol. 65, 422– 426. 18. Salet, C. and Moreno, G. (1981) Int.J. Radiat. Biol. 39, 227–230.

19. Salet, C., Moreno, G., Atlante, A. and Passarella, S. (1991) Photochem. Photobiol. 53, 391–393. 20. Musser, D. A. and Oseroff, A. R. (1994) Photochem. Photobiol. 59, 621– 626. 21. Gibson, S. L. and Hilf, R. (1985) Photochem. Photobiol. 42, 367– 373. 22. Hilf, R., Gibson, S. L., Murrant, R. S., Ceckler, T. L. and Bryant, R. G. (1987) Proc. SPIE Int. Soc. Opt. Eng. 847, 2–10. 23. Salet, C., Moreno, G., Riccheli, F. and Bernardi, P. (1997) J. Biol. Chem. 272, 21938 –21943. 24. Kroemer, G., Dallaporta, B. and Resche-Rigon, M. (1998) Annu. Rev. Physiol. 60, 619 – 642. 25. Oleinick, N. L., Antunez, A. R., Clay, M. E., Rihter, B. D. and Kenney, M. E. (1993) Photochem. Photobiol. 57, 242–247. 26. Evans, H. H., Ricanati, M. and Horng, M.-F. (1987) Proc. Natl. Acad. Sci. USA 84, 7562–7566. 27. Hofhaus, G., Shakeley, R. M. and Attardi, G. (1996) Methods in Enzymology 264, 476 – 483. 28. Xue, L.-Y., Agarwal, M. L. and Varnes, M. E. (1995) Photochem. Photobiol. 62, 135–143. 29. Reed, J. C. (1997) Cell 91, 559 –562. 30. Granger, D. L. and Lehninger, A. L. (1982) J. Cell Biol. 95, 527–535. 31. Korytowski, W., Bachowski, G. J. and Girotti, A. W. (1992) Photochem. Photobiol. 56, 1– 8. 32. Bernardi, P. and Azzone, G. F. (1981) J. Biol. Chem. 256, 7187– 7192. 33. La Piana, G., Fransvea, E., Marzulli, D. and Lofrumento, N. E. (1998) Biochem. Biophys. Res. Commun. 246, 556 –561. 34. Skulachev, V. P. (1998) FEBS Lett. 423, 275–280. 35. Leist, M., Single, B., Castoldi, A. F., Ku¨hnle, S. and Nicotera, P (1997) J. Exp.Med. 185, 1481–1486. 36. Rosenthal, I. (1991) Photochem. Photobiol. 53, 859 – 870. 37. Agarwal, R., Athar, M., Urban, S. A., Bickers, D. R. and Mukhtar, H. (1991) Cancer Lett. 56, 125–129.

679