Cellular uptake, localization and photodynamic effects of haematoporphyrin derivative in human glioma and squamous carcinoma cell lines

Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120 www.elsevier.com / locate / jphotobiol

Cellular uptake, localization and photodynamic effects of haematoporphyrin derivative in human glioma and squamous carcinoma cell lines a a, b c Seema Gupta , B.S. Dwarakanath *, K. Muralidhar , V. Jain a

Department of Biocybernetics, Institute of Nuclear Medicine and Allied Sciences, Brig SK Mazumdar Road, Timarpur, Delhi 110054, India b Department of Zoology, University of Delhi, Delhi 110007, India c Wallace-Kettering Neuroscience Institute, Kettering Medical Center and Department of Emergency Medicine, Wright State University, Dayton, OH 45429, USA Received 22 February 2002; received in revised form 16 October 2002; accepted 20 November 2002

Abstract Uptake, intracellular concentration, localization and photodynamic effects of a haematoporphyrin derivative (HpD, Photosan-3) were compared in human glioma (BMG-1, wild-type p53) and squamous carcinoma (4451, mutated p53) cell lines. Concentration and time dependence of cellular uptake of HpD was assayed from methanol extracts and whole cell suspension spectroscopy, while localization was studied by fluorescence microscopy-based image analysis. Colony-forming ability, apoptosis, cell-cycle progression and cytogenetic damage (micronuclei formation) were investigated as parameters of photodynamic response following irradiation with red light. BMG-1 cells were more sensitive to the photodynamic treatment than 4451 cells, although the 4451 cells accumulated a higher amount of HpD and did not differ significantly from BMG-1 cells with respect to intracellular localization. Photodynamically-induced cytogenetic damage and apoptosis were considerably higher in BMG-1 cells as compared to 4451 cells. The present results strongly suggest that manifestation of the photodynamically-induced lesions in the form of cytogenetic damage and apoptosis are among the important determinants of cellular sensitivity to HpD–PDT besides the photodynamic dose (intracellular concentration of the photosensitizer and the light dose).  2002 Elsevier Science B.V. All rights reserved. Keywords: Haematoporphyrin derivative; Photodynamic therapy; Intracellular localization; Cytogenetic damage; Apoptosis; p53

1. Introduction Photodynamic diagnosis (PDD) and photodynamic therapy (PDT) are promising new modalities shown to be effective in the early detection and treatment of several types of malignant and non-malignant diseases [1]. Both these technologies depend on the preferential accumulation of a light-activated dye (photosensitizer) in the pathologic lesion / cells. The fact that certain porphyrin-based compounds accumulate preferentially in tumors and can therefore be used for the detection and therapy of tumors has been known for several decades [2,3]. In recent years, a large number of new photosensitizers (second generation) have been shown to preferentially accumulate in hyperproliferative and malignant lesions and are currently under investigation in basic and preclinical studies. A few are being evaluated in clinical studies. The haematoporphyrin derivative (HpD), initially prepared by Lipson, now avail*Corresponding author. Fax: 191-11-391-9509. E-mail address: [email protected] (B.S. Dwarakanath).

able commercially in a partially purified form (dihaematoporphyrinester / ether) under the trade name Photofrin has been approved for specific clinical applications in several countries in Europe, America and Asia. Though Photofrin, being a complex mixture containing monomers, dimers and oligomers, is not an ideal photosensitizer since it may cause skin phototoxicity in some patients lasting several weeks, it is being used at present in the management of cancers of lung, esophagus, bladder and skin. The use of porphyrin-based photosensitizers in the diagnosis and therapy of malignant tumors of the brain and head and neck region is under active investigation [4,5]. To promote judicious and optimal clinical applications of PDD and PDT, the complex mechanisms underlying these promising techniques need to be better understood at various levels. Upon irradiation with light matching its absorption characteristics, the photosensitizer is excited to a higher energy singlet state. From the excited singlet energy state, the return to ground state can occur by the emission of light (fluorescence) or by dissipation of energy as heat. In PDD, emission of fluorescence is utilized to image and

1011-1344 / 02 / $ – see front matter  2002 Elsevier Science B.V. All rights reserved. doi:10.1016 / S1011-1344(02)00408-6

108

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

localize the tumor [2]. Clinically useful applications of PDD include intra-operative fluorescence guided surgical removal of malignant cells [6,7] besides early detection of premalignant and malignant lesions in the bladder, breast, gastro-intestinal tract, head and neck, lung and skin [8,9] using microscopic and endoscopic techniques. In contrast to PDD, photophysical and photochemical processes involving interactions of the excited photosensitizer with the surrounding molecules are responsible for PDT. In PDT, the excited singlet energy state of the photosensitizer comes down to the longer lived triplet state by intersystem crossing. Subsequent energy / charge transfer processes from the triplet state to the surrounding molecules can induce photosensitized chemical reactions. In the presence of oxygen, reactive oxygen species (ROS) such as singlet oxygen and hydroxyl radicals are produced, which can damage the biomolecules, cause alterations in functions of cell constituents and eventually lead to cell death through a network of biochemical and metabolic pathways. The effects produced by photosensitized reactions involving oxygen have been termed as photodynamic action [10]. The mechanisms underlying photodynamic action in cells and tissues are complex and depend upon multiple factors. Uptake and intracellular distribution of the photosensitizer in addition to its photophysical and photochemical characteristics are important in determining the efficacy of both PDD and PDT. Presence of the optimal amount of sensitizer in the vicinity of the sensitive target in a cell is an important determinant of photodynamic efficacy since singlet oxygen, the most important ROS generated by the interaction of sensitizer and light has a short life-time in which it can travel only a short distance [11]. The uptake and intracellular localization of the sensitizer are influenced by the physico-chemical properties of the sensitizer (charge distribution, lipophilicity, aggregation state etc.) as well as the cell type and its physiological state and microenvironment [12–16]. Depending upon the photosensitizer used, photodynamic treatment has been shown to induce lesions in membranes [17–19], mitochondria [20–22], lysosomes [23–25] and / or other essential biomolecules or organelles including DNA [26,27]. Cell nucleus, however, is a relatively less important target for photodynamic cytotoxicity than membranes [28]. Even though specific damage to cellular structures or functions has been observed at various intracellular sites, a clear causal relationship between the lesions and cell death has not emerged so far [29]. The cellular sensitivity to PDT differs a great deal between various cell types. Cells differing in their origin and radiosensitivity have been shown to respond differently to PDT [28,30]. Cellular and tissue responses to PDTinduced oxidative stress involve complex signal transduction pathways and alterations in gene expression leading either to repair of induced lesions, altered cell functions or to cell death. Cell death may occur either by apoptosis

and / or necrosis depending on the cell type, the target for photosensitization, concentration and intracellular distribution of the sensitizer [31], the incubation conditions and the light dose [19]. PDT has been shown to induce cellcycle arrest and apoptosis in a variety of cells [32–34]. Tumor suppressor gene p53 is known to influence both these processes and the treatment responses of tumors. Conflicting results have been, however, reported in recent studies designed to investigate the correlations between the p53 status and PDT-mediated apoptosis and treatment responses in cancer cells [35–37]. The relative contributions of the various factors in the manifestation of the photodynamic response remain yet to be clearly elucidated [29]. In the present study, we have investigated the relationships between cellular uptake, localization and various photodynamic effects (on cell growth, colony forming ability, cytogenetic damage and apoptosis) induced by a haematoporphyrin derivative, HpD, in two human tumor cell lines differing in their origin viz. a squamous cell carcinoma and a malignant glioma which also differ in their p53 status. Important results from these studies have been presented [38,39].

2. Materials and methods

2.1. Tumor cell lines Human cerebral glioma cell line (BMG-1; DNA index5 0.95; wild-type p53), established from a mixed glioma [40] and a human squamous carcinoma cell line (4451; DNA index51.50; mutated p53) established from an oral cavity lesion [41] were used in the present studies. Monolayer BMG-1 cells were grown in DMEM with 5% fetal calf serum (FCS) and 4451 cells with DMEM containing 10% FCS, penicillin (100 units / ml), streptomycin (50 mg / ml) and nystatin (2 mg / ml). Stock cultures were passaged every third day (BMG-1) or second day (4451) after harvesting the cells with 0.05% trypsin and seeding 8310 3 cells / cm 2 in tissue culture flasks to maintain the cells in the exponential phase. All experiments were carried out in exponentially growing cells.

2.2. Chemicals The haematoporphyrin derivative (HpD) used in these studies was Photosan-3 (PS-3), a commercial preparation (SeeLab, Germany). Hank’s Balanced Salt Solution (HBSS), Dulbecco’s modified Phosphate Buffered Saline (PBS), Dulbecco’s Modified Eagle’s Medium (DMEM), fetal calf serum (FCS), (N-[2-hydroxyethyl] piperazine-N9[2-ethanesulfonic acid]) (HEPES) buffer, propidium iodide (PI), 4,6-diamidino-2-phenyl indole (DAPI), Ribonuclease-A (RNase-A), Hoechst-H33342 (bis benzimide (29[4-ethoxyphenyl]-5-[4-methyl-1-piperazinyl]-2,59-bi-1H-

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

benzimidazole) trihydrochloride) and trypsin were obtained from Sigma, USA. Calf thymus DNA was obtained from E-Merck; Annexin-V-FITC from PharMingen, USA. All other chemicals used in the present study were of analytical grade from BDH, Glaxo laboratories (Qualigens), SRL, and E-Merck, India.

2.3. Estimation of intracellular concentration of the photosensitizer Monolayer cells were incubated in HBSS containing different concentrations of the sensitizer (2.5–10 mg / ml) and incubated at 37 8C in dark for various time intervals (0.5, 1, 2, 4 h). Cellular concentration of HpD was estimated by methanol extraction essentially according to Hilf et al. [42] and whole cell suspension spectroscopy.

2.3.1. Methanol extraction At the end of the incubation, HBSS was removed and cells were washed twice with PBS. Methanol was added to the attached cells and incubated for 1 h at 37 8C. Fluorescence spectra of the methanol extractable HpD was obtained with an excitation wavelength of 405 nm and emission was collected in the range of 550–750 nm using a Jobin Yvon fluorimeter (Model JY3C, France). Quantitation was carried out using the calibration curves generated with various concentrations of HpD in methanol by exciting at 405 nm and measuring the fluorescence at 630 nm. To determine the cell numbers in the culture, cells were scraped from flasks and homogenized using a hand homogenizer. The cell number was estimated from the DNA content of the homogenate using an AT-specific DNA fluorochrome, bisbenzimidazole (Hoechst-33342) and a calibration curve generated with calf thymus DNA [43]. 2.3.2. Fluorescence spectroscopic measurements in cell suspension Cells were trypsinized and incubated in the dark for various time intervals with HpD in HBSS at 37 8C. Fluorescence spectra (exc. 405 nm, em. 550–750 nm) of cells and supernatant before and after washing were obtained. Cellular uptake was calculated using standard calibration curves of the photosensitizer in HBSS. Intracellular concentration of HpD was estimated from the average cell volume. The cell volume (v) was determined by measuring the cell diameter (d) with the help of a micrometer, assuming spherical cell shape and using the formula v 5 p d 3 / 6. 2.4. Intracellular distribution of HpD using a fluorescence image analysis system Intracellular distribution of the sensitizer was studied by fluorescence microscopy using an image analysis system

109

(Olympus, BX60, Japan) equipped with a monochrome ¨ CCD camera (Grundig, FA87, Germany). The CCD camera contained a monochrome frame grabber of 24 Bit (Bit flow, USA) and the spectrum range was 400–1000 nm with an image sensor of 596 (V)3795 (H) total pixels. Cells were grown on coverslips for these studies. After incubation with HpD, coverslips were washed in PBS, mounted on slides and examined under the fluorescence microscope using a UV excitation filter (300–400 nm) and emission recorded in the red region of the spectrum (400– 800 nm). Images were acquired and stored in a digital pentium-based computer (166 MHz) and analyzed using the software provided by Optimas, USA. Cytoplasmic and nuclear localization of HpD was estimated by analyzing the images using area morphometry by marking the appropriate regions of interest (ROI).

2.5. Photodynamic treatment Cells were incubated in HBSS at 37 8C for 4 h with varying concentrations (1.25–20 mg / ml) of HpD. After the incubation, cells were washed with HBSS and exposed to red light (power53 W/ cm 2 ) using an optical filter with a cut-off at 610 nm from a high power (1000 W) xenon arc lamp (Oriel, USA). Optical power at the cell surface was measured using a radiometer (Model 1400 A, International Radiometer, USA) having a detector head (SL021 / FQ) with a flat response in the spectral range 400–1000 nm. Cells were incubated for another 2 h at 37 8C in HBSS before assay of cell response to treatment.

2.6. Cellular response to photodynamic treatment 2.6.1. Clonogenic survival assay Nearly 150 cells were plated in growth medium (DMEM110% FCS) after the treatment (as described above) and incubated in the dark under a humidified, 5% CO 2 atmosphere at 37 8C for 8–10 days to allow colony formation. Colonies were fixed with methanol and stained with 1% crystal violet. Colonies having more than 50 cells were counted and plating efficiency (PE) and surviving fraction (S.F.) were calculated. 2.6.2. Cell proliferation kinetics After photodynamic treatment, cells were incubated in growth medium for varying intervals of time, harvested by trypsinization and counted using a hemocytometer (adherent1floating). Flow cytometric measurements of cellular DNA content were performed with the ethanol (70%) fixed cells using an intercalating DNA fluorochrome, propidium iodide (PI) as described earlier [44]. Briefly, the cells (|0.5–1 million) were washed in PBS after removing ethanol and treated with Ribonuclease-A (200 mg / ml) for 30 min at 37 8C. Subsequently, cells were stained with PI (50 mg / ml) in PBS. Measurements were made with a laser-based (488 nm) flow cytometer (Facs

110

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

Calibur; Becton and Dickinson, USA) and data acquired using the Cell Quest software (Becton and Dickinson, USA). Cell cycle analysis was performed using the Modfit program.

2.6.3. Micronuclei formation Air-dried slides containing acetic acid–methanol (1:3, v / v) fixed cells were stained with 2-aminophenylindoledihydrochloride (DAPI) (10 mg / ml in citric acid (0.01 M), disodium phosphate (0.45 M) buffer containing 0.05% Tween-20 detergent) as described earlier [45]. Slides were examined under a fluorescence microscope using UV excitation filter and fluorescing nuclei were visualized using a blue emission filter. Cells containing micronuclei were counted from .1000 cells by employing the criteria of Countrymen and Heddle [46]. The fraction of cells containing micronuclei, called the M-fraction (%) was calculated as follows: M-fraction (%) 5 Nm /Nt 3 100, where Nm is the number of cells with micronuclei and Nt is the total number of cells analyzed. Since micronuclei formation is linked to cell proliferation, the micronuclei frequencies were normalized with respect to the cell numbers [40].

2.6.4. Detection and assay of apoptotic cells Percentage of cells undergoing apoptosis was determined microscopically and by flow cytometry using PI and Annexin-V-FITC labeled cells. Morphologically, marked condensation and margination of chromatin, fragmentation of nuclei and cell shrinkage characterize apoptotic cells and a good correlation between these morphological changes and the DNA ladder (one of the hallmarks of cells undergoing apoptosis) has been demonstrated [47]. At least 1000 cells were counted and percent apoptotic cells determined from slides prepared as described under Section 2.6.3 for micronuclei formation. In flow cytometric DNA analysis, the presence of hypodiploid (sub G 0 / G 1 ) population (with PI stained cells, as described for cell cycle analysis) is indicative of the apoptotic cell population. Cells undergoing apoptosis generally shrink and also show changes in internal structure, which is reflected in the alterations of light scatter. Therefore, treatment-induced changes in forward and side scatter of incident light were investigated by collecting these signals in the list mode using cell-quest software (Becton and Dickinson, USA). Analysis of light scatter was performed by off-line gating using appropriate windows created with untreated cells. Apoptotic cells were also detected by labeling of externalized phosphatidylserine using Annexin-V-FITC in unfixed cells [48]. For these measurements, cells were harvested by trypsinization and aliquots of 1310 5 cells resuspended in 100 ml binding buffer (10 mM HEPES /

NaOH, pH 7.4; 140 mM NaCl; 2.5 mM CaCl 2 ) and 5 ml of Annexin-V-FITC and 10 ml of PI (50 mg / ml) added. After 15 min at room temperature, 400 ml of binding buffer were added to each sample and analyzed by flow cytometry. The percentages of Annexin 1ve and 2ve cells were estimated by applying appropriate gates and using the regional statistics analysis facility provided in the Cell Quest software (Becton and Dickinson, USA).

3. Results

3.1. Kinetics of cellular uptake of HpD The kinetics of uptake as estimated by methanol extractable HpD in BMG-1 cells was observed to be biphasic, as it accumulated rapidly up to 1 h, with a subsequent decrease in the rate of uptake (Fig. 1a). These observations are similar to the earlier results reported for BHK-21 and other cell lines [49–51]. A qualitatively similar pattern of uptake was obtained from cell suspension spectroscopy (Fig. 1a) and image analysis (Fig. 1b).

Fig. 1. Time-dependent uptake of HpD in exponentially growing BMG-1 cells as determined by (a) methanol extraction, cell suspension fluorescence spectroscopy (conc. 10 mg / ml) and (b) image analysis (20 mg / ml) (50–75 cells were analyzed from 3–4 experiments).

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

However, with the image analysis system, a saturation in uptake was observed at longer incubation times. Similar uptake kinetics of hypocrellin B (HB) in EMT6 / Ed murine tumor cells has been reported using semiquantitative confocal microscopy and radiolabeled HB [52].

3.2. Intracellular concentration of HpD The kinetics of HpD uptake, studied by cell suspension spectroscopy in 4451 cells was similar to BMG-1 cells (Fig. 2a). However, the intracellular concentration of HpD was 5–7 times higher in 4451 cells as compared to BMG-1 cells at all the time points studied (Fig. 2a), which could be due to differences in the characteristics of these cells pertaining to uptake of HpD such as the density of LDL receptors, peripheral benzodiazepine receptors (PBR), cell cycle time, cell cycle distribution or cell volume [12– 15,53]. Indeed, exponentially growing BMG-1 cells were found to be larger than 4451 cells with mean volumes of 1.7760.028 pl and 1.0360.001 pl, respectively. Significant differences could not be observed, however, in cell cycle distribution. A linear increase in the intracellular concentration of HpD was observed up to a concentration of 10 mg / ml in the incubating medium (Fig. 2b) in both the cell lines.

3.3. Intracellular distribution Intracellular distribution of the sensitizer was studied by a fluorescence microscopy-based image analysis system (FMIA) using cells grown on coverslips. Images acquired at different time points were analyzed by marking either the whole cell or the nucleus as the region of interest (ROI). Most of the HpD fluorescence (red) was observed in the

111

cell membrane after 30 min of incubation; however, by 1–2 h the red fluorescence increased in the cytoplasm and in the perinuclear region. A careful examination of the images revealed that at 4 h 75–90% of the sensitizer was localized in the cytoplasm of both cell lines (Fig. 3). Porphyrins, which accumulate mainly via passive diffusion in cells, are known to localize in the cytomembranous structures such as mitochondria, lysosomes, endoplasmic reticulum etc. giving diffuse fluorescence in the cytoplasm [54]. No major differences were observed in the intracellular distribution of the sensitizer in the two cell lines (Fig. 3). Fluorescence microscopy coupled to an image analysis system (FMIA) provides a semi-quantitative method for measuring the uptake of sensitizer in addition to its distribution in a cell. Other methods like methanol extraction or whole cell suspension spectroscopy provide only an average value of the uptake in a cell population, while FMIA provides information on the variations in the amounts of uptake among a cell population. This method is more relevant in studies on photodynamic therapy of tumors as tumors contain heterogeneous cell populations, with uptake as well as localization varying among different types of cells. Further, the extraction methods lead to monomerization of the photosensitizers and increased fluorescence [55,56], while direct fluorescence measurements can provide the information about the state of the sensitizer inside the cell.

3.4. Cellular response to photodynamic treatment Treatment-induced growth inhibition, cell death (macrocolony assay), cytogenetic damage and apoptosis were investigated to study the photodynamic effects in both the cell lines.

Fig. 2. Time (a) and concentration (b) dependent uptake of HpD in exponentially growing BMG-1 and 4451 cells studied by cell suspension fluorescence spectroscopy. The concentration of HpD was 10 mg / ml in (a) and the incubation time was 4 h in (b).

112

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

Fig. 3. Intracellular distribution of HpD (20 mg / ml) showing cytoplasmic and nuclear (N) localization as a function of time in exponentially growing BMG-1 and 4451 cells.

3.4.1. Cell proliferation BMG-1 cells were incubated for 4 h with different concentrations of HpD in HBSS and effects on cell growth were studied following the photodynamic treatment. A concentration-dependent reduction in cell proliferation was observed (Fig. 4a). At a concentration of 2.5 mg / ml of HpD, a cytostatic effect was observed up to 40 h, followed by recovery leading to an increase in the cell number. At 5 mg / ml, however, the static phase was followed by a decrease in the cell number. A further increase in con-

centration of HpD resulted in an extensive cell death leading to a profound decrease in the cell numbers (data not shown). In 4451 cells, a longer lag period followed by slow increase in cell numbers was observed up to a concentration of 5 mg / ml, while no increase in cell numbers could be observed at higher concentrations (Fig. 4b). Cell cycle distribution after photodynamic treatment was analyzed from flow cytometric measurements of cellular DNA content. The fraction of 4451 cells in S-phase was

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

113

BMG-1 and 4451 cells (Fig. 6). Thus, at a concentration of 2.5 mg / ml, the SF value for BMG-1 cells was 0.0015 while it was 0.05 for 4451 cells implying a 30-fold difference in the photodynamic sensitivity between the two cell lines. A further increase in the concentration of HpD did not result in a decrease in the cell survival at light dose of 450 J / cm 2 in both the cell lines (Fig. 6). This may be partly due to relocalization of the photosensitizer to less sensitive targets during photoirradiation [29], but needs further investigation.

3.4.3. Cytogenetic damage Micronuclei expressed in post mitotic cells following treatment, arise from the chromosomal damage and cells with micronuclei are associated with loss of reproductive capacity [57,58]. The kinetics of micronuclei induction studied as a function of post-irradiation time varied both with the light dose and the concentration of the sensitizer (data not shown). At 2.5 mg / ml, the fraction of BMG-1 cells with micronuclei was highest at 40 h post-treatment and decreased thereafter (Fig. 7). Due to extensive damage (interphase death), micronuclei expression could not be analyzed in photodynamically-treated cells at higher HpD concentrations (10 and 20 mg / ml). Photodynamically-induced cytogenetic damage (micronuclei formation) was also found to vary as a function of HpD concentration in 4451 cells similar to the observations in BMG-1 cells. However, the fraction of cells expressing micronuclei (normalized with respect to cell proliferation) was nearly twofold higher in BMG-1 than in 4451 cells (Fig. 7).

Fig. 4. Effects of photodynamic treatment on the proliferation of exponentially growing (a) BMG-1 and (b) 4451 cells. Cells were incubated with HpD (2.5 and 5 mg / ml) for 4 h in HBSS and irradiated with red light (power53 W/ cm 2 ; light dose5450 J / cm 2 ).

not significantly altered following PDT, while a clear increase in G 2 1M fraction could be observed (Fig. 5). In BMG-1 cells, however, the fraction of S-phase cells were always lower in the photodynamically-treated groups as compared to untreated group at all the time points studied and G 2 block could not be observed (Fig. 5).

3.4.2. Clonogenic cell survival Effects of photoirradiation on cell survival at various intracellular concentrations of HpD were studied by macrocolony assay. The fraction of surviving cells, referred to as the surviving fraction (SF) decreased exponentially with increasing concentration of HpD up to 2.5 mg / ml in both

3.4.4. Apoptosis In the present studies, characteristic morphological features [47], presence of hypodiploid cell population (sub G 0 / G 1 ), alterations in light scatter and phosphatidylserine externalization (Annexin-V binding) were analyzed to identify the presence of apoptotic cells. HpD–PDT induced a significantly higher level of apoptosis in BMG-1 cells as compared to 4451, which was dependent on the concentration of HpD as well as post-irradiation time (PIT) in both the cell lines. Results obtained by various methods in these two cell lines are compared in Table 1. Morphological analysis showed that the frequency of apoptotic cells was highest at 18 h after photo-irradiation, which reduced following a recovery in the cell proliferation (Fig. 8). The fraction of BMG-1 cells undergoing apoptosis increased with concentration of HpD up to 5 mg / ml (.80%) but decreased on increasing the concentration to 10 mg / ml, possibly due to increased necrosis [59]. In 4451 cells, on the other hand, the increase in apoptosis could be observed even up to 10 mg / ml of HpD (60%). These differences in induction of apoptosis were more clearly evident at 2.5 mg / ml of HpD, where the

114

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

Fig. 5. Flow cytometric DNA histograms showing cell cycle distribution and apoptosis (arrow) observed in exponentially growing BMG-1 and 4451 cells following photodynamic treatment (HpD52.5 mg / ml; power53 W/ cm 2 ; light dose5450 J / cm 2 ; PIT540 h).

fraction of BMG-1 cells undergoing apoptosis was nearly 55% while it was less than 10% in 4451 cells. Essentially similar results were obtained by flow-cytometric analysis of light scatter (forward scatter related to size and side

Fig. 6. Response of exponentially growing BMG-1 and 4451 cells to photodynamic treatment studied as a function of concentration of HpD. Cells were incubated for 4 h in HBSS containing HpD and irradiated with red light (.610 nm; power53 W/ cm 2 ; light dose5450 J / cm 2 ). Two hours after irradiation cells were plated for macrocolony assay (n53).

scatter related to internal structure of cells), phosphatidylserine externalization (Fig. 9 and Table 1) as well as hypodiploid cell population (Fig. 5).

Fig. 7. Micronuclei frequency observed as a function of time in BMG-1 and 4451 cells following photodynamic treatment. Cells were incubated with HpD (2.5 mg / ml) for 4 h in HBSS and irradiated with red light (.610 nm; power53 W/ cm 2 ; light dose5450 J / cm 2 ).

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120 Table 1 Apoptosis studied by various methods in exponentially growing BMG-1 and 4451 cells observed at 40 h following photodynamic treatment (HpD52.5 mg / ml, 4 h in HBSS; light dose5450 J / cm 2 ) Method

Morphological Sub G 0 / G 1 Forward scatter Annexin-V-FITC 1ve

Apoptotic cells (%) BMG-1

4451

5562 2862 6165 7065

,10 ,2 562 ,2

4. Discussion In the present study, important differences and lack of correlation in the cellular uptake of the photosensitizer HpD (Photosan-3), its intracellular concentration and

Fig. 8. Treatment-induced apoptosis studied as a function of post-irradiation time (2.5–10 mg / ml) in BMG-1 and 4451 cells. Cells were incubated with HpD for 4 h in HBSS and irradiated with red light (.610 nm; power53 W/ cm 2 ; light dose5450 J / cm 2 ). Two hours after irradiation, cells were allowed to grow in growth medium and DAPI stained slides were observed for apoptotic cells.

115

photodynamic responses have been observed between the malignant glioma and oral squamous carcinoma cell lines. Lack of correlation between cellular uptake, intracellular concentration and photodynamic efficiency in cells from tumors of different origins as well as in cells with different histologies has been reported earlier [28,60]. Possible mechanisms underlying the present observations are briefly discussed. The oral squamous carcinoma cells accumulated significantly higher amounts of HpD than glioma cells (Table 2) but were less sensitive to the photodynamic treatment (Fig. 6). The cellular uptake, intracellular distribution and retention of HpD in cells depend on micro-environmental factors and cell characteristics including cell volume, state of proliferation, the capacity and affinity of the intracellular target sites to bind HpD and cell type [61–63]. The average volume of BMG-1 cells being larger than that of 4451 cells (Table 2) and the state of proliferation of both the cell populations being similar, the present results imply that there could be major differences in the binding sites of HpD in the two cell lines. However, significant differences in the gross localization pattern were not apparent in these two cell lines (Fig. 3). Possibly, quantitative differences in the HpD localization at different sites in the cytoplasm could not be resolved by the image analysis system used in the present study. Peripheral benzodiazepine receptors (PBR) located in the plasma membranes and outer membranes of mitochondria are considered to be among the important binding sites for porphyrins [53]. The number of mitochondria in 4451 cells is likely to be higher than in BMG-1 cells, since 4451 cells show lower rates of glucose usage and glycolysis [58]. Several types of cancer cells including malignant glioma have been reported to have a high density of PBR [64], which have been observed to correlate well with the rate of proliferation, grade of malignancy and prognosis [65–67]. The affinity of binding of porphyrins to PBR correlates with the photodynamic inactivation of cells suggesting that PBR form important subcellular targets for PDT [68]. This, however, requires further investigations since multiple binding sites of porphyrins with differences in binding affinities on PBR could exist [69]. Variations in the intracellular localization and density of PBR in plasma membrane, mitochondria, perinuclear regions and nucleus in different cell lines have been reported [66,70]. In 4451 cells, a larger fraction of HpD could be present in de-aggregated monomeric ionic forms, which have a higher quantum yield for fluorescence but photodynamically less efficient. The dynamic equilibrium between the various aggregated and ionic forms of porphyrins in HpD is known to be very sensitive to the microenvironment [71,72] and therefore constituents of HpD are likely to be present in different forms in different cell types. Further work to identify the aggregation state and binding of HpD in the two cell lines is, therefore, necessary. Besides the uptake and localization of the sensitizer

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

116

Fig. 9. Bivariate plots of forward and side scatter signals (observed at 18 h) and frequency distribution of Annexin-V binding (showing externalization of phosphatidylserine observed at 40 h) measured by flow cytometry showing PDT (HpD, 2.5 mg / ml) induced apoptosis in BMG-1 and 4451 cells.

(which may determine the types and numbers of primary photodynamically-induced lesions), the photodynamic efficacy is profoundly influenced by biological processes mediating and regulating the manifestation of induced lesions. For example, competitions between processes of fixation and repair of lesions (DNA and non-DNA) and between different pathways leading to cell death (necrosis and apoptosis), could play important roles in determining responses to PDT [73]. Intracellular localization studies showed that about 15% of HpD was localized in the nucleus of both the cell lines. Since, chromatin is bound firmly to the inner nuclear membrane [74], singlet oxygen generated at or near the nuclear membrane may damage the chromatin attached to the inner nuclear membrane [75]. Differences in the distribution of HpD in the perinuclear region were not apparent in the two cell lines in the present study. Interestingly, however, cytogenetic damage assayed by micronuclei frequency was significantly higher in BMG-1 cells (Table 2 and Fig. 7). Treatment-induced DNA damage (strand breaks) manifests in the form of chromo-

somal aberrations at mitosis mainly in the form of acentric fragments. Most of the acentric fragments fail to get incorporated into the daughter nuclei and would be found as micronuclei in the cytoplasm of daughter cells [46]. Cells with micronuclei are known to be associated with the loss of reproductive ability [57]. Centromere breaks and double minutes are the most frequent aberrations induced by HpD and light [75]. Since, repair of treatment-induced DNA and chromosomal damage is facilitated by delay in cell cycle progression due to checkpoints at G 1 -S and G 2 -M transition [76], a G 2 block observed in 4451 cells (Fig. 5) could be partly responsible for the enhanced repair of lesions and consequently for lower micronuclei frequencies and relative resistance to PDT in these cells as compared to BMG-1 cells. PDT-induced apoptosis, observed in several cell lines, has been considered as one of the important determinants of sensitivity [19,32–34,77]. The higher level of apoptotic death observed in BMG-1 (wild-type p53) as compared to 4451 cells (mutated p53; Tables 1 and 2) could be another reason for higher sensitivity of BMG-1 cells to PDT. In the

Table 2 Intracellular concentration, localization and photodynamic responses in BMG-1 and 4451 cells (HpD52.5 mg / ml, 4 h in HBSS; light dose5450 J / cm 2 ) Parameters Cell line

Average volume (pl)

p53 status

Intracellular conc. (mg / ml)

Localization

S.F.

M-fraction (%)a

Apop. (%)a

Normalized G 2 1M (% control)

BMG-1 4451

1.7760.028 1.0360.001

Wild-type Mutated

1.260.3 7.261.3

M and Cyt. M and Cyt.

0.0015 0.05

2162 1362

5562 ,2

120 166

M, Plasma membrane; Cyt., cytoplasm. a 40 h post irradiation (morphological criteria).

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

present work, four different assays / parameters were used for detection and quantitative estimate of the apoptotic index. All the methods used indicated a higher frequency of apoptotic cells in BMG-1 cells as compared to 4451 cells (Table 1). The percentage of apoptotic cells estimated from hypodiploid DNA content was, however, considerably lower in both the cell lines as compared to estimates of apoptotic cells by cell volume (forward scatter), PS externalization (Annexin-V binding) and morphological features (Table 1). Absence of DNA ladder in cells undergoing apoptosis revealed by different techniques has been reported earlier [78]. Differences in the estimates of apoptotic fractions measured by methods based on different parameters could be associated with different phases of the apoptotic process. However, such differences could also arise due to a number of other reasons viz. negligible cleavage of DNA at internucleosomal regions with cleavage at higher levels of chromatin organization, or the differences in the preparatory method used for flow cytometric DNA analysis, where an extraction buffer was not used which facilitates resolution of sub G 0 / G 1 phase from G 1 cells [79]. Apoptotic cell death can be initiated by a variety of metabolic and genetic stress signals. Several pathways that mediate apoptotic cell death are regulated in a complex manner by numerous genes and their products; p53 and Bcl family of proteins appear to be the most important ones [35–37,80–83]. Since, photodynamic treatment can induce cell membrane, mitochondrial, cytoskeletal and DNA damage, multiple independent as well as interdependent pathways could be involved in photodynamicallyinduced apoptosis. Further, the expression of death receptors like CD95 / Fas /Apo-1, TNF and TRAIL [84–87] and the release of apoptogenic factors such as cytochrome c, Smac / DIABLO through mitochondrial membrane permeability transition (MMPT) as well as the expression of Bcl family of genes are regulated by p53, although the exact mechanisms remain yet to be completely elucidated [88–90]. Since, significant differences in the levels of anti-apoptotic (Bcl-2, Bcl XL ) and pro-apoptotic (Bax) proteins could not be observed in these two cell lines (data not shown), it appears that apoptosis can indeed be induced both in wild type and mutated p53 gene carrying cell types albeit at different photodynamic doses. It is pertinent to note that although a positive correlation between the p53 status and photodynamic sensitivity has been found in several studies through transfection of wild type p53 gene in p53 mutant cell lines [37,91], recent studies have however shown that apoptosis can occur also through caspase and p53 independent pathways, for example, through the apoptosis-inducing factor (AIF), a mitochondrial flavoprotein [92] and its recently discovered homologous molecule designated as AMID (AIF-homologous mitochondrion-associated inducer of death) [93]. AIF has been shown to translocate from mitochondria to nucleus

117

following apoptotic stimulation inducing chromosome condensation and DNA fragmentation. AMID-induced apoptosis is however different as compared to the classic apoptotic pathways as it is delayed and not inhibited by Bcl-2. Further, association of AMID with the outer membrane of mitochondria appears to be critical although the exact mechanisms involved are not known. These pathways could also contribute significantly to the HpD–PDT induced apoptosis, particularly in 4451 cells with mutated p53 and warrant further investigations. Taken together, the present studies suggest that differences in the accumulation and distribution of HpD between different cells could arise from the variations in the frequency and affinity of HpD binding sites (mainly PBR located in mitochondria) as well as the aggregation and ionic states of intracellular HpD. Cellular responses to PDT crucially depend also on the processing of the PDTinduced lesions by competing repair / fixation pathways and therefore on the cellular context viz. the genotype and phenotypic / metabolic characteristics of cells besides the uptake and localization of the sensitizer. Since cell death following PDT can be induced by necrosis and multiple apoptotic pathways (p53 dependent as well as independent) regulated in a complex way by interactions of numerous genes and gene products, the differences in the PDT responses of various cell types cannot be predicted only on the basis of the uptake / intracellular concentration of the photosensitizer or status of a single gene such as p53.

Acknowledgements Thanks are due to Dr. N.K. Chaudhury, Mr. J.S. Adhikari and Dr. Sudhir Chandna for help in spectroscopic, flow cytometric and image analysis studies. We are grateful to Maj. Gen. T. Ravindranath, Director, INMAS and Dr. T. Lazar Mathew, former Director for their interest and support for this work. S.G. was a recipient of a research fellowship from the University Grants Commission and CSIR, Govt. of India.

References [1] T.J. Dougherty, C.J. Gomer, B.W. Henderson, G. Jori, D. Kessel, M. Korbelik, J. Moan, Q. Peng, Photodynamic therapy, J. Natl. Cancer Inst. 90 (1998) 889–905. [2] R.L. Lipson, E.J. Baldes, A.M. Olsen, Hematoporphyrin derivative: a new aid for endoscopic detection of malignant disease, J. Thorac. Cardiovasc. Surg. 42 (1961) 623. [3] T.J. Dougherty, G.B. Grindey, R. Field, K.R. Weishaupt, D.G. Boyle, Photoradiation therapy II, cure of animal tumors with hematoporphyrin and light, J. Natl. Cancer Inst. 55 (1975) 115–121. [4] P.J. Muller, B.C. Wilson, Photodynamic therapy for recurrent supratentorial gliomas, Semin. Surg. Oncol. 11 (1995) 346–354. [5] S. Krishnamurthy, S.K. Powers, P. Witmer, T. Brown, Optimal light

118

[6]

[7]

[8]

[9]

[10] [11]

[12]

[13]

[14]

[15]

[16]

[17]

[18]

[19]

[20]

[21]

[22]

[23]

[24]

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120 dose for interstitial photodynamic therapy in treatment for malignant brain tumors, Lasers Surg. Med. 27 (2000) 224–234. W. Stummer, A. Novotny, H. Stepp, C. Goetz, K. Bise, H.J. Reulen, Fluorescence-guided resection of glioblastoma multiforme by using 5-aminolevulinic acid-induced porphyrins: a prospective study in 52 consecutive patients, J. Neurosurg. 93 (2000) 1003–1013. A. Zimmermann, M. Ritsch-Marte, H. Kostron, mTHPC-mediated photodynamic diagnosis of malignant brain tumors, Photochem. Photobiol. 74 (2001) 611–616. R. Baumgartner, H. Fisslinger, D. Jocham, H. Lenz, L. Ruprecht, H. Stepp, E. Unsold, A fluorescence imaging device for endoscopic detection of early stage cancer—instrumental and experimental studies, Photochem. Photobiol. 46 (1987) 759–763. K. Svanberg, I. Wang, S. Colleen, I. Idvall, C. Ingvar, R. Rydell, D. Jocham, H. Diddens, S. Bown, G. Gregory, S. Montan, S. Andersson-Engels, S. Svanberg, Clinical multi-colour fluorescence imaging of malignant tumours—initial experience, Acta Radiol. 39 (1998) 2–9. H.F. Blum, Photodynamic Action and Disease caused by Light, Hofnov, New York, 1964. J. Moan, K. Berg, The photodegradation of porphyrins in cells can be used to estimate the lifetime of singlet oxygen, Photochem. Photobiol. 53 (1991) 549–553. G. Oenbrink, P. Jurgenlimke, D. Gabel, Accumulation of porphyrin in cells: influence of hydrophobicity, aggregation and protein binding, Photochem. Photobiol. 48 (1988) 451–456. K.W. Woodburn, N.J. Vardaxis, J.S. Hill, A.H. Kaye, D.R. Philips, Subcellular localization of porphyrins using confocal laser scanning microscopy, Photochem. Photobiol. 54 (1991) 725–732. K.W. Woodburn, N.J. Vardaxis, J.S. Hill, A.H. Kaye, J.A. Reiss, D.R. Phillips, Evaluation of porphyrin characteristics required for photodynamic therapy, Photochem. Photobiol. 55 (1992) 697–704. F. Khanum, V. Jain, Studies concerning the role of energy metabolism in the photodynamic therapy, Assoc. Med. Phys. Bull. 12 (1987) 113–124. M. Soncin, L. Polo, E. Reddi, G. Jori, M.E. Kenney, G. Cheng, M.A. Rodgers, Effect of axial ligation and delivery system on the tumor-localizing and -photosensitizing properties of Ge(IV)-octabutoxy-phthalocyanines, Br. J. Cancer. 71 (1995) 727–732. D. Kessel, D. Woodburn, C.J. Gomer, N. Jagerovic, K.M. Smith, Photosensitization with derivatives of chlorin p 6 , J. Photochem. Photobiol. B: Biol. 28 (1995) 13–18. D.K. Gaffney, S.L. Schober, F. Sieber, Merocyanine 540-sensitized photoinactivation of leukemia cells: role of oxygen and effects of plasma membrane integrity and mitochondrial respiration, Exp. Hematol. 18 (1990) 23–26. M. Dellinger, Apoptosis or necrosis following photofrin photosensitization: influence of the incubation protocol, Photochem. Photobiol. 64 (1996) 182–187. H. Dummin, T. Cernay, H.W. Zimmerman, Selective photosensitization of mitochondria in HeLa cells by cationic Zn(II) phthalocyanines with lipophilic side chains, J. Photochem. Photobiol. B: Biol. 37 (1997) 219–229. A.D. Munday, A. Sriratana, J.S. Hill, S.B. Kahl, P. Nagley, Mitochondria are the functional intracellular target for a photosensitizing boronated porphyrin, Biochim. Biophys. Acta 1311 (1996) 1–4. J.S. Modica-Napolitano, J.S. Joyal, G. Ara, A.R. Oseroff, J.R. Aprille, Mitochondrial toxicity of cationic photosensitizes for photochemotherapy, Cancer Res. 50 (1990) 7876–7881. K. Berg, J. Moan, Lysosomes and microtubules as targets for photochemotherapy of cancer, Photochem. Photobiol. 63 (1997) 403–409. M. Ambroz, A.J. MacRobert, J. Morgan, G. Rumbles, M.S.C. Foley, D. Phillips, Time resolved fluorescence spectroscopy and intracellular imaging of disulphonated aluminum phthalocyanine, J. Photochem. Photobiol. B: Biol. 22 (1994) 105–117.

[25] C.W. Lin, J.R. Shulock, S.D. Kirley, L. Cincotta, J.W. Foley, Photodynamic destruction of lysosomes mediated by Nile Blue photosensitizers, Photochem. Photobiol. 58 (1993) 81–91. [26] J. Moan, H. Waksvik, T. Christensen, DNA single strand breaks and sister chromatid exchanges induced by treatment with hematoporphyrin and light or by X-rays in human NHIK 3025 cells, Cancer Res. 40 (1980) 2915–2918. [27] L. Hong, D.S. Shin, Y.E. Lee, D.C. Nguyen, T.C. Trang, A.H. Pan, S.L.J. Huang, D.H. Chong, M.W. Berns, Subcellular phototoxicity of 5-aminolevulinic acid (ALA), Lasers Surg. Med. 22 (1998) 14–24. [28] D.J. Gomer, N. Ruckter, A.L. Murphree, Differential cell photosensitivity following porphyrin photodynamic therapy, Cancer Res. 48 (1988) 4539–4542. [29] J. Morgan, W.R. Potter, A.R. Oseroff, Comparison of photodynamic targets in a carcinoma cell line and its mitochondrial DNA-deficient derivative, Photochem. Photobiol. 71 (2000) 747–757. [30] L.C. Penning, J.W.M. Lagerberg, J.H. VanDierendonck, C.J. Cornelisse, T.M.A.R. Dubbleman, J. VanSteveninck, The role of DNA damage and inhibition of poly (ADP-ribosyl) lation in loss of clonogenicity of murine L929 fibroblasts, caused by photodynamically induced oxidative stress, Cancer Res. 54 (1994) 5561–5567. [31] D. Kessel, Y. Luo, Mitochondrial photodamage and PDT-induced apoptosis, J. Photochem. Photobiol. B: Biol. 42 (1998) 89–95. [32] M.L. Agarwal, M.E. Clay, E.J. Harvey, H.H. Evans, A.R. Antunez, N.L. Oleinick, Photodynamic therapy induces rapid cell death by apoptosis in L5178Y mouse lymphoma cells, Cancer Res. 51 (1991) 5993–5996. [33] Y. Luo, C.K. Chang, D. Kessel, Rapid initiation of apoptosis by photodynamic therapy, Photochem. Photobiol. 63 (1996) 528–534. [34] V. Carre, C. Jayat, R. Granet, P. Krausz, M. Guilloton, Chronology of apoptotic events induced in the K562 cell line by photodynamic treatment with hematoporphyrin and monoglucosylporphyrin, Photochem. Photobiol. 69 (1999) 55–60. [35] A.M.R. Fisher, N. Rucker, S. Wong, C.J. Gomer, Differential photosensitivity in wild-type and mutant p53 human colon carcinoma cell lines, J. Photochem. Photobiol. B: Biol. 42 (1998) 104–107. [36] A.M.R. Fisher, A. Ferrario, N. Rucker, S. Zhang, C.J. Gomer, Photodynamic therapy sensitivity is not altered in human tumor cells after abrogation of p53 function, Cancer Res. 59 (1999) 331–335. [37] Z. Tong, G. Singh, A.J. Rainbow, The role of p53 tumor suppressor in the response of human cells to photofrin-mediated photodynamic therapy, Photochem. Photobiol. 71 (2000) 201–210. [38] S. Gupta, V. Jain, K. Muralidhar, Differences in photodynamic effects of hematoporphyrin derivative in human squamous carcinoma and glioma cell lines, in: M. Fahim (Ed.), Trends in Physiological Sciences: Cells to Systems, Serials Publications, Delhi, 2002, pp. 225–247. [39] S. Gupta, K. Muralidhar, V. Jain, Experimental studies to improve photodynamic therapy of tumors, in: Radiobiology 2000-International Conference on Radiation Biology, Trivandrum, India, Feb. 17– 19, 2000. [40] B.S. Dwarakanath, V. Jain, Energy linked modifications of the radiation response in a human glioma cell line, Int. J. Radiat. Oncol. Biol. Phys. 17 (1989) 1033–1040. [41] F. Zolzer, S. Hillebrandt, C. Streffer, Radiation-induced G 1 block and p53 status in six human cell lines, Radiother. Oncol. 37 (1995) 20–28. [42] R. Hilf, P.B. Leakey, S.J. Sollot, S.L. Gibson, Photodynamic inactivation of R3230AC mammary carcinoma in vitro with hematoporphyrin derivative: effects of dose, time and serum on uptake and phototoxicity, Photochem. Photobiol. 37 (1983) 633–642. [43] C.F. Brunk, K.C. Jones, T.W. James, Assay for nanogram quantities of DNA in cellular homogenates, Anal. Biochem. 92 (1979) 497– 500. [44] B.S. Dwarakanath, J.S. Adhikari, V. Jain, Hematoporphyrin derivatives potentiate the radiosensitizing effects of 2-deoxy-D-glucose in cancer cells, Int. J. Radiat. Oncol. Biol. Phys. 43 (1999) 1125–1133.

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120 [45] B.S. Dwarakanath, V. Jain, Enhancement of the radiation-induced damage by 2-deoxy-D-glucose in organ cultures of brain tumors, Indian J. Med. Res. 82 (1985) 266–268. [46] P.I. Countryman, J.A. Heddle, Production of micronuclei from chromosome aberrations in irradiated cultures of human lymphocytes, Mutat. Res. 41 (1976) 321–332. [47] K. Yanagihara, M. Nii, K. Numoto et al., Radiation-induced apoptotic cell death in human gastric epithelial tumor cells: correlation between mitotic death and apoptosis, Int. J. Radiat. Biol. 77 (1995) 677–685. [48] I. Vermes, C. Haanen, H. Steffens-Nakken, C. Reutelingsoerger, A novel assay for apoptosis. Flow cytometric detection of phosphatidylserine expression on early apoptotic cells using fluorescein labeled Annexin-V, J. Immunol. Methods 184 (1995) 39–51. [49] F. Khanum, V. Jain, Cellular accumulation and biological activity of hematoporphyrin derivative (L) in comparison with photofrin II, Ind. J. Exp. Biol. 35 (1997) 348–355. [50] D. Kessel, Transport and binding of HpD and relative porphyrins by murine leukemia L1210 cells, Cancer Res. 41 (1981) 1318–1323. [51] R.M. Bohmer, G. Morstyn, Uptake of hematoporphyrin derivative by normal and malignant cells: effects of serum, pH, temperature and cell size, Cancer Res. 45 (1985) 5328–5334. [52] G.G. Miller, K. Brown, R.B. Moore, Z.J. Diwu, J. Liu, L. Huang, J.W. Lown, D.A. Begg, V. Chlumecky, J. Tulip, M.S. McPhee, Uptake kinetics and intracellular localization of hypocrellin photosensitizers for photodynamic therapy: a confocal microscopy study, Photochem. Photobiol. 61 (1995) 632–638. [53] A. Verma, S.H. Snyder, Peripheral type benzodiazepine receptors, Annu. Rev. Pharmacol. Toxicol. 29 (1989) 307–322. [54] R. Hilf, D.B. Small, R.S. Murant, P.B. Leaky, S.L. Gison, Hematoporphyrin derivative sensitivity of mitochondrial succinic dehydrogenase and selected cytosolic enzymes of R3230AC mammary adenocarcinoma of rats, Cancer Res. 44 (1984) 1483–1488. [55] R. Cubeddu, R. Ramponi, G. Bottiroli, Time resolved fluorescence spectroscopy of hematoporphyrin in micelles, Chem. Phys. Lett. 128 (1986) 439–442. [56] R. Cubeddu, R. Ramponi, G. Bottiroli, Disaggregation effects on hematoporphyrin derivative in the presence of surfactants at different concentrations: temperature dependence, SPIE 701 (1987) 316– 319. [57] J. Midander, L. Revesz, The frequency of micronuclei as a measure of cell survival in irradiated cell populations, Int. J. Radiat. Biol. 38 (1980) 237–242. [58] B.S. Dwarakanath, F. Zolzer, S. Chandna, T. Bauch, J.S. Adhikari, W. Muller, C. Streffer, V. Jain, Heterogeneity in 2-deoxy-D-glucose induced modification of radiation response in human tumor cell lines, Int. J. Radiat. Oncol. Biol. Phys. 50 (2001) 1051–1061. [59] B.B. Noodt, G.H. Rodal, M. Wainwright, Q. Peng, R. Horobin, J.M. Nesland, K. Berg, Apoptosis induction by different pathways by methylene blue derivative and light from mitochondrial sites in V79 cells, Int. J. Cancer 75 (1998) 1–8. [60] R.R. Perry, W. Matthew, J.B. Mitchell, A. Russo, S. Evans, H.I. Pass, Sensitivity of different human lung cancer histologies to photodynamic therapy, Cancer Res. 50 (1990) 4272–4276. [61] T. Christensen, J. Moan, Binding of hematoporphyrin to synchronized cells from the line NHIK 3025, Cancer Lett. 9 (1980) 105–110. [62] V. Jain, F. Khanum, New horizons in the development of photodynamic therapy of cancer, in: V. Jain, H. Goel, W. Pohlit (Eds.), Recent Advances in Radiation Oncology, Indian Council of Medical Research, New Delhi, 1990, pp. 275–294. [63] A. Verma, S.L. Facchina, D.J. Hirsch, S.Y. Song, L.F. Dillahey, J.R. Williams, S.H. Snyder, Photodynamic tumor therapy: mitochondrial benzodiazepine receptors as a therapeutic target, Mol. Med. 4 (1998) 40–45. [64] A. Beurdeley-Thomas, L. Miccoli, S. Oudard, B. Dutrillaux, M.F. Poupon, The peripheral benzodiazepine receptors: a review, J. Neurooncol. 46 (2000) 45–56.

119

[65] H. Miettinen, J. Kononen, H. Haapasalo, P. Helen, P. Sallinen, T. Harjuntausta, H. Helin, H. Alho, Expression of peripheral-type benzodiazepine receptor and diazepam binding inhibitor in human astrocytomas: relationship to cell proliferation, Cancer Res. 55 (1995) 2691–2695. [66] M. Hardwick, D. Fertikh, M. Culty, H. Li, B. Vidic, V. Papadopoulos, Peripheral-type benzodiazepine receptor (PBR) in human breast cancer: correlation of breast cancer cell aggressive phenotype with PBR expression, nuclear localization, and PBRmediated cell proliferation and nuclear transport of cholesterol, Cancer Res. 59 (1999) 831–842. [67] A. Beinlich, R. Strohmeier, M. Kaufmann, H. Kuhl, Relation of cell proliferation to expression of peripheral benzodiazepine receptors in human breast cancer cell lines, Biochem. Pharmacol. 60 (2000) 397–402. [68] M. Mesenholler, E.K. Matthews, A key role for the mitochondrial benzodiazepine receptor in cellular photosensitization with deltaaminilevulinic acid, Eur. J. Pharmacol. 406 (2000) 171–180. [69] D. Kessel, M. Antolovich, K.M. Smith, The role of the peripheral benzodiazepine receptor in the apoptotic response to photodynamic therapy, Photochem. Photobiol. 74 (2001) 346–349. [70] C.H. Park, E. Carboni, P.L. Wood, K.W. Gee, Characterization of peripheral benzodiazepine type sites in a cultured murine BV-2 microglial cell line, Glia 16 (1996) 65–70. [71] A. Kalia, V. Jain, Effects of some divalent metal ions on the aging phenomenon of hematoporphyrin and photofrin II, Ind. J. Biochem. Biophys. 26 (1989) 213–218. [72] N.K. Chaudhury, V. Jain, Effects of microenvironment on aggregation and aging in homogeneous solutions of hematoporphyrin, in: Fifth Congress of the European Society for Photobiology, Marburg, Germany, 19–26 September, 1993. [73] V. Jain, Mechanisms and metabolic modulation of photosensitization, in: V. Jain, H.C. Goel (Eds.), Selected Topics in Photobiology, Indian Photobiology Society, 1992, pp. 130–147. [74] W.W. Franke, Structure, biochemistry, and functions of the nuclear envelope, Int. Rev. Cytol. Suppl. 4 (1974) 71–236. [75] J.F. Evensen, J. Moan, Photodynamic action and chromosomal damage: a comparison of hematoporphyrin derivative (HpD) and light with X-irradiation, Br. J. Cancer 45 (1982) 456–465. [76] A. Maity, W.G. McKenna, R.J. Muschel, The molecular basis for cell cycle delays following ionizing radiation: review, Radiother. Oncol. 31 (1994) 1–13. [77] J. Dahle, H.B. Steen, J. Moan, The mode of cell death induced by photodynamic treatment depends on the cell density, Photochem. Photobiol. 70 (1999) 363–367. [78] S. Fournel, L. Genestier, J.-P. Rouault, G. Lizard, M. Flacher, O. Assossou, J.P. Revillard, Apoptosis without decrease of cell DNA content, FEBS Lett. 367 (1995) 188–192. [79] S. Hara, H. Dorota Halicka, S. Bruno, J. Gong, F. Traganos, Z. Darzynkiewicz, Effect of protease inhibitors on early events of apoptosis, Exp. Cell Res. 223 (1996) 372–384. [80] S.C. Denstman, L.E. Dillehay, J.R. Williams, Enhanced susceptibility to HPD sensitized phototoxicity and correlated resistance to trypsin detachment in SV40 transformed IMR-90 cells, Photochem. Photobiol. 43 (1986) 145–147. [81] J. He, M.L. Agarwal, H.E. Larkin, L.R. Friedman, L. Xue, N.L. Oleinick, The induction of partial resistance to photodynamic therapy by the protooncogene Bcl-2, Photochem. Photobiol. 64 (1996) 845–852. [82] Q. Yu, M. He, N.H. Lee, E.T. Liu, Identification of Myc-mediated death response pathways by microarray analysis, J. Biol. Chem. 277 (2002) 13059–13066. [83] I.R. Radford, Review: initiation of ionizing radiation-induced apoptosis: DNA damage-mediated or does ceramide have a role?, Int. J. Radiat. Biol. 75 (1999) 521–528. [84] D.W. Nicholson, N.A. Thornberry, Caspases: killer proteases, Trends Biochem. Sci. 22 (1997) 299–306.

120

S. Gupta et al. / Journal of Photochemistry and Photobiology B: Biology 69 (2003) 107–120

[85] L.B. Owen-Schaub, W. Zhang, J.C. Cusack, L.S. Angelo, S.M. Santee, T. Fujiwara, J.A. Roth, A.B. Deisseroth, W.W. Zhang, E. Kruzel, R. Radinsky, Wild-type human p53 and a temperaturesensitive mutant induce Fas /APO-1 expression, Mol. Cell Biol. 15 (1995) 3032–3040. [86] M. Bennet, K. MacDonald, S.-W. Chan, J.P. Luzio, R. Simari, P. Weissberg, Cell surface trafficking of Fas: a rapid mechanism of p53-mediated apoptosis, Science 282 (1998) 290–293. [87] G.S. Wu, T.F. Burns, E.R. McDonald, W. Jiang, R. Meng, I.D. Krantz, T. Kao, D.D. Gan, J.Y. Zhou, R. Mushcel, S.R. Hamilton, N.B. Spinner, S. Markowitz, Y. Wu, W.S. Diery, Killer / DR5 is a DNA damage inducible p53 regulated death receptor gene, Nat. Genet. 17 (1997) 141–143. [88] M.C. Wei, W.X. Zong, E.H.Y. Chery, T. Lindsten, V. Panoutsakoppoulou, A.J. Ross, K.A. Roth, G.R. MacGregor, C.B. Thompson, S.J. Korsmayer, Proapoptotic Bax and Bak: a requisite gateway to mitochondrial dysfunction and death, Science 292 (2001) 727–730. [89] T. Lindsten, A.J. Ross, A. King, W.X. Zong, J.L. Rathmell, H.A. Shiels, E. Ulrich, K.G. Waymire, P. Mahar, K. Frauwirth, Y. Chen,

[90] [91]

[92]

[93]

M. Wie, V.M. Eng, D.M. Adelman, M.C. Simen, A. Ma, J.A. Golden, G. Evans, S.J. Korsmayer, G.R. MacGregor, C.B. Thompson, The combined functions of proapoptotic Bcl-2 family members bak and bax are essential for normal development of multiple tissues, Mol. Cell 6 (2000) 1389–1399. R.V. Sinov, Y. Haupt, The cellular response to p53: the decision between life and death, Oncogene 18 (1999) 6145–6157. W.-G. Zhang, X.-W. Li, L.-P. Ma, S.-W. Wang, H.-Y. Yang, Z.-Y. Zhang, Wild-type p53 protein potentiates phototoxicity of 2-BA-2DMHA in HT29 cells expressing endogenous mutant p53, Cancer Lett. 138 (1999) 189–195. H.K. Lorenzo, S.A. Susin, J. Penninger, G. Kroemer, Apoptosis inducing factor (AIF): a phylogenetically old, caspase-independent effector of cell death, Cell Death Differ. 6 (1999) 516–524. M. Wu, L.-G. Xu, X. Li, Z. Zhai, H.-B. Shu, AMID, an apoptoticinducing factor-homologous mitochondrion-associated protein, induces caspase-independent apoptosis, J. Biol. Chem. 277 (2002) 25617–25623.