We investigated cellular responses to chlorin-based photosensitizer DH-II-24 under darkness in human gastric adenocarcinoma AGS cells. Cells were loaded with 0.5–10 μg/mL DH-II-24 for 12 h, and intracellular reactive oxygen species (ROS) and intracellular Ca2+ levels, in situ tissue transglutaminase (tTGase) activity, cell viability, cell morphology and cell cycle were examined. DH-II-24 treatment had no effect on intracellular ROS production or cell morphology, and did not induce cell detachment at any concentrations tested. In addition, cell viability and cell cycle progression were not altered by the photosensitizer. However, DH-II-24 treatment elevated the basal level of intracellular Ca2+ in a dose-dependent manner and inhibited tTGase activity without affecting tTGase expression levels. Furthermore, DH-II-24 inhibited lysophosphatidic acid-induced activation of tTGase in a dose-dependent manner. In contrast, photodynamic therapy (PDT) with 1 μg/mL DH-II-24 significantly elevated intracellular ROS and in situ tTGase activity in parallel with a rapid and large increase in intracellular Ca2+ levels. DH-II-24-mediated PDT decreased cell viability and induced cell detachment. These results demonstrate that DH-II-24 treatment alone under darkness induced different cellular responses to DH-II-24-mediated PDT. (Cancer Sci 2011; 102: 549–556)
Photodynamic therapy (PDT) is a promising therapeutic modality for cancer treatment that involves photosensitizers, irradiation with appropriate wavelengths, and molecular oxygen.(1,2) Activated photosensitizers lead to significantly increased levels of intracellular reactive oxygen species (ROS) and intracellular Ca2+. These changes result in the killing of tumor cells either directly or by damaging the tumor-associated vasculature through apoptosis and necrosis. In addition, PDT indirectly induces a significant stimulatory effect on the immune system.(3–6)
Photodynamic therapy has advantages over other methods of cancer treatment, such as non-invasive treatment, by avoiding systemic immune suppression and selectively destroying tumors.(7–13) However, it is essential to consider the side-effects of non-irradiated photosensitizers under darkness in patients. Photofrin, the most widely used photosensitizer in clinical PDT, has several weaknesses within clinical applications, including adverse reactions involving skin damage caused by slow clearance from the circulation system and low photosensitivity.(2,3,14) To avoid adverse effects, patients should remain in a dark condition until excretion of the photosensitizer following intravenous administration. Remaining in the dark reduces quality of life.(15,16) New photosensitizers have been developed with reduced side-effects and increased effectiveness.(2,8,14,17,18) Previously, we reported a new chlorin-based photosensitizer, DH-II-24, and examined its photosensitivity in human gastric adenocarcinoma (AGS) cells, human colorectal carcinoma (HCT116) cells and xenografts.(19,20) Photo-activated DH-II-24 elevated the levels of intracellular ROS and Ca2+, indicating that DH-II-24 is effective for use in PDT.(19) Several reports focus on the PDT-induced mechanisms of cell death using photosensitizers.(19,21,22) However, the responses of cells or animals to non-irradiated photosensitizers remains to be elucidated.
Dark toxicity of photosensitizers has been studied through cell viability assays in various cell types.(23–28) Incubation with various concentrations (up to 6 μM) of the valine-derivatized porphyrin for 24 h showed little or no toxicity under darkness in human breast carcinoma (MCF-7) cells.(25) Meta-tetra(hydroxyphenyl) chlorin (mTHPC; Foscan) was applied to several cancer cell lines, including gall bladder cancer cells, bile duct cancer cells and MCF-7 cells,(23,28) and cytotoxicity was examined using methylthiazolyldiphenyl-tetrazoliumbromide (MTT) assays. In gall bladder and bile duct cancer cells, no dark toxicity was observed with 2–3 μg/mL mTHPC, although cell viability rapidly decreased in response to higher concentrations.(23) Similar results were obtained using mTHPC in MCF-7 cells.(28) However, much higher concentrations of mTHPC derivatives, such as a PEG2000 derivative of mTHPC (m-THPC MD) and a liposomal formulation of mTHPC (Foslip), were required to induce similar dark toxicity.(23,28) Dark toxicity was also investigated using other photosensitizers such as hematoporphyrin monomethyl ether,(24) 18m-ALA (a dendrimer containing 18 aminolevulinic acids)(26) and dithiaporphyrin(27) in various cancer cells. The results indicated minimal dark toxicity in concentration ranges similar to porphyrin and mTHPC. Thus, dark toxicity of various photosensitizers has been examined in cancer cells. However, the effects of photosensitizers on intracellular responses remain to be elucidated.
In the present study, we examined cellular responses to non-irradiated DH-II-24 in human gastric adenocarcinoma AGS cells, including intracellular ROS and Ca2+ accumulation, in situ tissue transglutaminase (tTGase) activity, cell viability and cell cycle progression. tTGase, a multifunctional protein, is regulated by several factors including intracellular Ca2+ and redox potential, and is involved in a wide variety of cellular processes including cell death, inflammation and wound healing.(29) It is reported that PDT with DH-II-24 elevated the level of intracellular Ca2+ and tTGase is activated by intracellular Ca2+ in AGS cells.(19,30) DH-II-24 treatment had no significant effect on intracellular ROS levels, but the basal level of intracellular Ca2+ was elevated and in situ tTGase activation was inhibited. However, PDT with DH-II-24 (1 μg/mL) treatment decreased cell viability in parallel with elevated intracellular ROS levels and in situ tTGase activity. In addition, dark toxicity was not observed at up to 10 μg/mL DH-II-24, a 10-fold greater concentration than that used for PDT. Therefore, higher-concentration DH-II-24 treatments may be more beneficial for cancer treatment by achieving more efficient PDT, while maintaining the previously known advantages of photo-activation with a long wavelength light, significant photoxicity and rapid clearance from the body.
Materials and Methods
Chemicals and reagents. H2DCFDA and Fluo-4 AM were purchased from Molecular Probes (Eugene, OR, USA). 5-(Biotinamido)pentylamine and FITC-conjugated streptavidin were obtained from Thermo Fisher Scientific (Rockford, IL, USA). Lysophosphatidic acid (LPA) was purchased from Cayman Chemical (Ann Arbor, MI, USA). Methylthiazolyldiphenyl-tetrazoliumbromide (MTT) and propidium iodide were obtained from Sigma (St. Louis, MO, USA). DH-II-24 (methyl-13b-[2-dimethylaminoethoxycarbonyl]-13b-demethoxycarbonyl-pheophorbide α) was kindly provided by Professor Chang-Hee Lee (Kangwon National University, Korea).
Cell culture and loading of DH-II-24. AGS human gastric adenocarcinoma cells (American Type Culture Collection; Manassas, VA, USA) were maintained at 37°C in RPMI 1640 medium supplemented with 10% fetal bovine serum, 100 U/mL penicillin and 100 μg/mL streptomycin in a humidified 5% CO2 atmosphere. DH-II-24 was loaded into AGS cells by incubation with various concentrations of the photosensitizer under darkness. The DH-II-24-loaded cells were subjected to various analyses, such as measurements of intracellular ROS and intracellular Ca2+ levels, and in situ tTGase activity assays.
Photodynamic therapy with an on-stage system. AGS cells were cultured on round coverslips for measurement of intracellular ROS and intracellular Ca2+ levels, and for in situ tTGase activity assays, or on 24-well culture plates for cell viability assays, and loaded with 1 μg/mL DH-II-24 for 12 h. Coverslips were mounted onto a perfusion chamber for measurement of intracellular Ca2+ levels. The perfusion chamber or culture plates were placed on the stage of a confocal microscope (FV300; Olympus, Tokyo, Japan) at a distance of 5 cm from the condenser to allow for cell irradiation. Cells were irradiated at 1.45 mW/cm2 for 60 s (87 mJ/cm2) under darkness by the built-in 100 W halogen lamp. The light was filtered with a long-pass filter (LP630; Fiber Optic Korea, Cheonan, Korea) set in the filter holder of the illumination column at the end of the optical fiber cable from a halogen lamp. The irradiated cells were incubated under darkness and subjected to measurement of intracellular ROS and Ca2+ levels, in situ tTGase activity assays and MTT assays.
Measurement of intracellular ROS levels. Intracellular ROS levels were determined according to the procedures of Kim et al.(31) (2002). Briefly, cells grown on round coverslips were incubated with 10 μM H2DCFDA in culture medium lacking phenol red for 10 min immediately prior to the determination of intracellular ROS levels by confocal microscopy (FV300; Olympus). The cells were excited by a 488-nm argon laser, and the images were filtered by a bandpass filter (BP 510–530 nm). Approximately 30 cells were randomly selected from three independent experiments, and the DCF fluorescence intensities of the DH-II-24-loaded cells were compared with those of non-loaded control cells.
Measurement of intracellular Ca2+ levels. Intracellular Ca2+ levels were monitored according to the procedures of Yoo et al.(19) (2009). Briefly, AGS cells cultured on round coverslips were loaded with various concentrations of DH-II-24 for 12 h or incubated with 10 μg/mL DH-II-24 for various times, followed by incubation with 2 μM Fluo-4 AM for 30 min. Each coverslip containing stained cells was scanned by confocal microscopy with a 488-nm excitation argon laser and a 510-nm longpass emission filter. Approximately 30 cells were randomly selected from three independent experiments and Fluo-4 fluorescence intensities of DH-II-24-loaded cells were compared with those of non-loaded control cells.
Measurement of in situ tTGase activity by confocal microscopy. In situ tTGase activity was determined by the confocal microscopic assay according to the procedures of Lee et al.(32) (2003). Briefly, AGS cells on round coverslips were loaded with various concentrations of DH-II-24 for 12 h and incubated with 1 mM 5-(biotinamido)pentylamine (a pseudosubstrate of tTGase) for 1 h at 37°C. Cells were incubated with 5 μg/mL LPA for various times, fixed with 3.7% formaldehyde in PBS for 30 min and permeabilized with 0.2% Triton X-100 in PBS for 30 min at room temperature. Following incubation with 2% BSA in TBST blocking solution (20 mM Tris, pH 7.6, 138 mM NaCl and 0.1% Triton X-100) for 30 min, cells were treated with FITC-conjugated streptavidin (1:200, v/v) in the blocking solution for 1 h at room temperature and observed by confocal microscopy. Approximately 30 cells were randomly selected from three independent experiments and fluorescence intensities were determined by processing the FITC images at a single-cell level. tTGase activity was determined by comparing the fluorescence intensities of DH-II-24-loaded cells with those of non-loaded or unstimulated control cells.
Western blot analysis. AGS cells were incubated for 30 min in lysis buffer (50 mM Tris–HCl, pH 7.5, 1% Triton X-100, 10% glycerol, 150 mM NaCl, 1 mM EDTA, 0.1 mM PMSF, 10 μg/mL aprotinin and 10 μg/mL leupeptin), scraped off and centrifuged at 18,000g for 10 min at 4°C. The resulting cell lysates were separated by SDS-PAGE and transferred to PVDF membranes. The membrane was blocked with 5% non-fat dried milk in TBST for 1 h at room temperature. Proteins were analyzed with antibodies against tTGase and β-actin.
Cell viability assay. Cell viability was determined by the MTT reduction assay.(20) Cells were incubated with various concentrations of DH-II-24 for 12 h in 24-well culture plates and washed with culture medium. Cells were incubated with 1 mg/mL MTT solution for 4 h at 37°C. The resulting insoluble purple formazan crystal produced by active mitochondrial dehydrogenases from living cells was dissolved in isopropanol, and absorbencies at 570 nm were measured with a microplate reader (Molecular devices, Sunnyvale, CA, USA). Cell viability was proportional to the A570 value and was expressed as a percentage of that for non-irradiated control cells.
Cell cycle analysis by flow cytometry. After DH-II-24 treatment, cells were washed with serum-free medium, trypsinized and fixed overnight with 70% ethanol at −20°C. Cells were stained for 30 min at room temperature with 100 μg/mL propidium iodide solution in PBS containing 0.1% Triton X-100 and 100 μg/mL DNase-free RNase A. Cells were analyzed on a FACScan with CellQuest software (Becton Dickinson, San Jose, CA, USA). At least 10 000 fluorescent events were counted per sample.
Effect of DH-II-24 treatment on the levels of intracellular ROS and Ca2+. To investigate cellular responses to the chlorin-based photosensitizer DH-II-24 in cancer cells, we examined changes of intracellular ROS levels in response to DH-II-24 treatment in human gastric adenocarcinoma AGS cells. As shown in Figure 1A, DH-II-24 did not induce any significant changes in intracellular ROS levels at any concentration tested. However, PDT using 1 μg/mL DH-II-24 elevated intracellular ROS levels approximately 2.3-fold over the non-irradiated control cells (P < 0.001; Fig. 1B). Therefore, the photosensitizer alone at concentrations up to 10 μg/mL, 10 times higher than the concentration used for PDT, does not have any significant effect on intracellular ROS.
Because PDT increases intracellular Ca2+ concentrations in cancer cells,(19,33,34) we investigated changes in intracellular Ca2+ levels with respect to DH-II-24 concentrations and treatment times without irradiation. As shown in Figure 2A, DH-II-24 increased basal levels of intracellular Ca2+ in a dose-dependent manner, with a sub-maximal elevation at 2 μg/mL. DH-II-24 treatment slowly increased intracellular Ca2+ levels over time, with a maximal elevation (approximately 2.2-fold over the control level) at 9 h. The elevated levels were maintained at 12 h (Fig. 2B).
We monitored transient changes of intracellular Ca2+ levels induced by PDT with 1 μg/mL DH-II-24, and compared those data with changes in intracellular Ca2+ levels induced by incubation with 1 μg/mL DH-II-24 only. As shown in Figure 2C, DH-II-24-mediated PDT induced a rapid and transient increase in intracellular Ca2+ levels that then slowly decreased. However, incubation with 1 μg/mL DH-II-24 induced only a slight increase in intracellular Ca2+ levels during the scanning time, which might be due to irradiation of the photosensitizer during scanning. Thus, incubation with DH-II-24 induced a dose-dependent increase of intracellular Ca2+ levels in AGS cells, whereas PDT with 1 μg/mL DH-II-24 produced a transient, but much higher increase in intracellular Ca2+ levels than 1 μg/mL DH-II-24 alone.
DH-II-24 treatment-induced changes in tTGase activity and expression. Because PDT with DH-II-24 elevated intracellular Ca2+ levels in AGS cells and because tTGase is activated by intracellular Ca2+,(19,29) we investigated whether DH-II-24 treatment could activate tTGase in AGS cells. Cells were incubated with various concentrations of DH-II-24 for 12 h, and in situ tTGase activity was determined by confocal microscopic measurement of pentylamine-modified proteins.(32) As shown in Figure 3A, DH-II-24 treatment decreased in situ tTGase activity in a dose-dependent manner, with a significant decrease at 5 μg/mL DH-II-24 (P < 0.05). However, DH-II-24 had no effect on the expression level of tTGase at any concentrations tested, demonstrating that DH-II-24 alone inhibited in situ tTGase activity with no effect on its expression (Fig. 3B). We examined the effect of DH-II-24-mediated PDT on in situ tTGase activity. As shown in Figure 3C, DH-II-24-mediated PDT elevated in situ tTGase activity approximately 2.3-fold over the control level. Thus, incubation with DH-II-24 inhibited in situ tTGase with no effect on its expression; however, in situ tTGase was activated by DH-II-24-mediated PDT in AGS cells.
DH-II-24 inhibits LPA-induced in situ activation of tTGase. To confirm our previous finding (Fig. 3), we investigated the possible inhibition of LPA-induced activation of tTGase in AGS cells by DH-II-24. LPA activates tTGase in situ in Swiss 3T3 fibroblasts.(32) We determined that LPA rapidly elevated in situ tTGase activity in a time-dependent manner in AGS cells, with significant activation at 30 min and a maximal activation (approximately 4-fold over the control level) at 120 min (Fig. 4A). The activation level rapidly decreased over 2 h. We then studied the effect of DH-II-24 on LPA-induced activation of tTGase, in which we incubated AGS cells with various concentrations of DH-II-24 and determined the in situ tTGase activity in the presence of LPA. As shown in Figure 4B, incubation with DH-II-24 inhibited LPA-induced activation of tTGase in a dose-dependent manner, with a significant inhibition at 0.5 μg/mL (P < 0.001). Activation was completely suppressed by 2 μg/mL DH-II-24. Thus, DH-II-24 functioned as an inhibitor of tTGase with no effect on its expression levels in AGS cells.
Changes in cell morphology following DH-II-24 treatment. Changes in cell morphology were examined by confocal microscopic observation of AGS cells treated with various concentrations of DH-II-24 for 12 h. As shown in Figure 5A, DH-II-24 alone did not induce changes in cell morphology or cell detachment at any concentrations examined. This suggests that 0.5–10 μg/mL DH-II-24 alone is not cytotoxic in AGS cells. Phototoxicity of DH-II-24 was evaluated by microscopic observation of changes in cell morphology and cell detachment at 6 and 12 h after PDT with 1 μg/mL DH-II-24 (Fig. 5B). Most cells were shrunken and formed vacuoles by 6 h, and the majority of cells (approximately 67%) were detached by 12 h. Thus, DH-II-24 alone with concentrations 10-times greater than that used for PDT had no effect on cell morphology and did not induce cell detachment, whereas DH-II-24-mediated PDT did induce cytotoxic effects in AGS cells.
DH-II-24 treatment did not alter cell viability. We investigated the effects of DH-II-24 on AGS cell viability by MTT-reduction assays and cell cycle analyses using flow cytometry. As shown in Figure 6A, DH-II-24 treatment without irradiation did not induce changes in cell viability at concentrations up to 10 μg/mL. Cell cycle analysis using 10 μg/mL DH-II-24 showed no significant changes in cell numbers in the G2-M and G1 phases, indicating that cell cycle progression was not affected by DH-II-24 treatment (Fig. 6B). However, PDT with 1 μg/mL DH-II-24 significantly decreased cell viability to 56% over the control level (P < 0.001; Fig. 7A). In addition, the PDT increased the cell number of sub G1, in parallel with a decrease in the percentage of G1 and G2 cells (P < 0.001; Fig. 7B), indicating apoptotic cell death induced by DH-II-24-mediated PDT. These results indicate that DH-II-24 alone with concentrations up to 10-times higher than that used for PDT had no effect on cell viability or cell cycle progression.
Although a tremendous number of photosensitizers have been reported for in vitro and in vivo PDT treatments, only a few of these photosensitizers have ideal properties for clinical trials, such as chemical purity, tumor-selective accumulation, activation at longer wavelengths of light for tissue penetration, rapid clearance from the body and high photosensitivity.(35–37) Thus, a new generation of photosensitizers are still under investigation. Photosensitizers can be structurally classified into porphyrin derivatives, chlorins, phthalocyanins and porphycenes, each reacting differently and having different photosensitivities.(38) Photosensitizers are activated by specific wavelengths of light and can increase intracellualar ROS and Ca2+ levels, important factors for PDT-induced cell death.(3–6,8,19,39,40) Thus, the determination of cell death mechanisms has been an important topic in PDT treatment. However, to improve PDT efficacy by minimizing side-effects, it is essential to understand the effects of photosensitizers accumulated in non-target tissues under darkness. Although there are several reports on dark toxicity of photosensitizers,(23–28) to our knowledge this is the first report focusing on cellular responses to photosensitizers under darkness. Furthermore, this report provides a foundation for understanding the possible side-effects of photosensitizers that remain in non-targeted tissues, including skin, after PDT treatment.
We previously reported that DH-II-24-mediated PDT increased the levels of intracellular ROS and Ca2+ in AGS cells,(19) and that TGase was activated by intracellular ROS and Ca2+ in fibroblasts.(30,32,41) Thus, we examined changes in intracellular ROS and Ca2+ levels, in situ tTGase activity and cell viability in response to DH-II-24 treatment in human gastric adenocarcinoma cells. DH-II-24 treatment without photo-activation had no effect on intracellular ROS levels and cell cycle progression, and was not cytotoxic. However, this treatment did elevate the basal level of intracellular Ca2+ and inhibited tTGase activity without altering tTGase expression levels. On the other hand, DH-II-24-mediated PDT elevated intracellular ROS levels and in situ tTGase activity in parallel with a transient but strong increase in intracellular Ca2+ resulting in decreased cell viability. The expression level of tTGase was not altered by PDT (data not shown). Thus, DH-II-24 treatment under darkness induced different cellular responses than DH-II-24-mediated PDT in AGS cells. Furthermore, DH-II-24 treatment did not induce dark toxicity at concentrations up to 10 μg/mL, 10-times greater than that used for PDT (1 μg/mL), indicating that more effective PDT can be performed using higher concentrations of DH-II-24.
It is interesting that DH-II-24 treatment inhibited tTGase activity, while DH-II-24-mediated PDT stimulated tTGase activity. It was expected that DH-II-24 would activate tTGase because TGase is activated by intracellular Ca2+.(29,30,41) However, tTGase activity was significantly inhibited by DH-II-24 treatment and tTGase stimulation by LPA was also prevented by the photosensitizer in a dose-dependent manner. In addition, we performed a tTGase activity assay in the presence of 20 μM aminoguanidine, a potent inhibitor of amineoxidase, and similar inhibitory effects of DH-II-24 on in situ tTGase activity were also observed (data not shown). DH-II-24 treatment increased the basal level of intracellular Ca2+ 9 h after incubation with 10 μg/mL of the photosensitizer with no effect on intracellular ROS levels. This indicated that elevation of basal intracellular Ca2+ levels was not enough to alter in situ tTGase activity. Thus, DH-II-24 inhibited tTGase in AGS cells through an unknown mechanism(s) independent of intracellular Ca2+ levels. It would be very useful to examine the inhibitory effects of other photosensitizers on tTGase activity and their mechanisms of tTGase inhibition.
Tissue transglutaminase is a multifunctional enzyme that is implicated in a number of cellular events including apoptosis and cell adhesion, mobility and invasion.(29) There is a report that PDT induced resistance to trypsinization and elevated in vitro tTGase activity in Met B cells;(42) however, in situ tTGase activation by PDT is not known. In this report, we demonstrated that DH-II-24-mediated PDT activated in situ tTGase and elevated the levels of intracellular ROS and Ca2+. Because tTGase is activated by intracellular ROS and Ca2+,(30,32,41) it is possible that DH-II-24-mediated PDT might activate tTGase via elevation of intracellular ROS and Ca2+ levels. Considering the role of tTGase in apoptosis,(29) it is likely that tTGase mediates cell death by PDT, although it is necessary to elucidate the mechanism(s) of tTGase activation by PDT and its role in PDT-induced cell death.
In summary, we examined cellular responses to a cholin-based photosensitizer, DH-II-24, under darkness in human gastric adenocarcinoma AGS cells. DH-II-24 without photo-activation was not cytotoxic and increased basal levels of intracellular Ca2+ and inhibited tTGase, whereas DH-II-24-mediated PDT was cytotoxic, elevated intracellular ROS levels, increased in situ tTGase activity and showed a transient peaking of intracellular Ca2+ levels.
This work was supported in part by the Korea Research Foundation through the Basic Research Program (2008-05943) and by the Ministry of Health and Welfare through the National R&D Program for Cancer Control (1020420).