Purpose: The lack of effective treatment for pancreatic cancer results in a very low survival rate. This study explores the enhancement of the therapeutic effect on human pancreatic cancer via the combination of triptolide and ionizing radiation (IR).
Experimental Design:In vitro AsPC-1 human pancreatic cancer cells were treated with triptolide alone, IR alone, or triptolide plus IR. Cell proliferation was analyzed with sulforhodamine B (SRB) method and clonogenic survival; comparison of apoptosis induced by the above treatment was analyzed by annexin V–propidium iodide (PI) staining. Furthermore, the expression of apoptotic pathway intermediates was measured by the assay of caspase activity and Western blot. Mitochondrial transmembrane potential was determined by JC-1 assay. In vivo, AsPC-1 xenografts were treated with 0.25 mg/kg triptolide, 10 Gy IR, or triptolide plus IR. The tumors were measured for volume and weight at the end of the experiment. Tumor tissues were tested for terminal nucleotidyl transferase–mediated nick end labeling (TUNEL) and immunohistochemistry.
Results: The combination of triptolide plus IR reduced cell survival to 21% and enhanced apoptosis, compared with single treatment. In vivo, tumor growth of AsPC-1 xenografts was reduced further in the group treated with triptolide plus IR compared with single treatment. TUNEL and immunohistochemistry of caspase-3 cleavage in tumor tissues indicated that the combination of triptolide plus IR resulted in significantly enhanced apoptosis compared with single treatments.
Conclusions: Triptolide in combination with ionizing radiation produced synergistic antitumor effects on pancreatic cancer both in vitro and in vivo and seems promising in the combined modality therapy of pancreatic cancer.
Pancreatic adenocarcinoma remains one of the most lethal of malignancies. The incidence of pancreatic cancer has steadily increased over the past four decades (1). Satisfactory treatment is available only for the minority of patients who present with very early-stage disease. Despite recent research and improvements in imaging, efforts to detect tumors at an earlier stage or augment standard therapy have done little to change the dismal prognosis. The 5-year survival rate is <5% (1), ranking this cancer as the fourth leading cause of cancer death (2). Importantly, at the time of diagnosis, the majority of patients (80-90%) already have locally advanced, metastatic, or inoperable tumors. Radiation therapy alone or in combination with chemotherapy has shown only modest efficacy in local control and palliation (3, 4). A new therapeutic strategy is urgently needed to control this aggressive cancer.
Triptolide, a diterpenoid triepoxide (MW, 360) derived from the herb Tripterygium wilfordii, has been used as a natural medicine in China for hundreds of years (5). Several recent papers have evaluated triptolide as an antitumor agent (6–8). Among its actions, triptolide slows proliferation of a variety of cell types. Slowly proliferating cells accumulate more genetic damage per cell cycle during a course of radiation therapy than more rapidly dividing cells. This heightened accumulation of DNA damage might overcome repopulation, the process in which cells continue to divide during a protracted course of therapy. Triptolide is also reported to be an effective inducer of apoptosis in solid cancer cells, including breast, prostate, and lung cancer (9). Although the mechanism is not well elucidated, it is suggested that triptolide might induce apoptosis by altering pathways involving p21 and p53 (10). Several studies have also shown that triptolide induces DNA damage (11, 12). In this study, we explore the effect of triptolide in combination with IR in an attempt to evaluate the underlying mechanisms by which this new therapeutic approach controls pancreatic cancer.
Materials and Methods
Cells and reagents. AsPC-1, a human pancreatic cancer cell line, was obtained from the American Type Culture Collection. The cells were maintained as monolayer cultures in DMEM supplemented with 10% fetal bovine serum, 100 μg/mL of streptomycin, and 100 units/mL of penicillin. The cells were incubated at 37°C in a humidified atmosphere of 5% CO2.
Triptolide with 99.9% purity was purchased from the Institute of Medical Research (Fuzhou, China); sulforhodamine B (SRB) was purchased from Sigma; caspase-3 assay kits were purchased from Molecular Probes; caspase-8 and caspase-9 assay kits were purchased from BioVision; JC-1, annexin V, and propidium iodide (PI) were purchased from Molecular Probes; cytochrome c, cleaved caspase-3, and poly(ADP-ribose) polymerase (PARP) polyclonal antibodies were purchased from Cell Signaling Technology, Inc.; and ApopTag in situ apoptosis detection kits were purchased from Chemicon International, Inc.
Cell viability assay. Viability of AsPC-1 after different treatments was evaluated by SRB assay (13). Briefly, AsPC-1 cells (1 × 103 cells per well) were plated in triplicate in 96-well plates overnight and changed to fresh media. The cells were treated with different concentrations (0, 12.5, 25, or 50 nmol/L) of triptolide alone or followed by 4 Gy IR at a dose rate of 280 cGy/min delivered by a Cs-137 Mark I irradiator. The control cells were treated with the same concentration of vehicle (0.01% DMSO) or mock IR. Forty-eight hours later, 50 μL of 10% trichloroacetic acid was added to fix cells at 4°C for 2 h, stained with 70 μL of 0.3% SRB for 30 min, color developed with 200 μL Tris base (10 mmol/L; pH, 10.5), and read at A490.
Clonogenic survival assay. AsPC-1 cells were treated with vehicle (DMSO) alone, triptolide alone (0, 3.125, 6.25, 12.5, or 25 nmol/L), triptolide plus IR at a dose of 0, 2, 4, 6, or 8 Gy, or IR alone and plated in 60-mm dishes at different densities based on the stringency of treatments. The number of cells was adjusted to generate 50 to 200 colonies per dish at each radiation dose. After 21 days, the colonies (containing ≥50 cells) were stained with crystal violet, and the numbers of colonies were counted with FluorChem SP (Alpha Innotech). The surviving fraction (SF) was calculated as a ratio of the number of colonies to the number of cells plated (plating efficiency) divided by the same ratio calculated for the nonirradiated group. D0 (the incremental dose required for reducing the fraction of colonies to 37%, indicative of single-event killing) was calculated using the formula of the single hit multitarget (SHMT) model [SF = 1 − (1 − e−(D/D0))n; ref. 14]. SF2 is the surviving fraction of exponentially growing cells when irradiated at the clinically relevant dose of 2 Gy.
Flow-cytometric analysis of apoptosis. Exactly 48 h after treatment with triptolide (25 nmol/L) alone, IR alone (4 Gy), or triptolide plus IR, cells were harvested, stained with annexin V for 30 min and then with PI, immediately followed by flow-cytometric analysis according to the manufacturer's instructions. The percentage of cells that were annexin V positive but PI negative was compared among the different treatment groups. For the cell cycle assay, the harvested cells were immediately fixed in 75% alcohol overnight, then treated with 1% RNase A for 30 min at room temperature, and stained with PI for 10 min; samples were then measured by flow cytometry (FACS, Becton Dickinson). The data were analyzed with CellQuest software.
JC-1 analysis for mitochondria membrane potential. Mitochondrial membrane potential was measured by flow cytometry using JC-1 staining. About 2 μg of JC-1 in 30 μL of saline was added to 100 μL of single cell suspension that had been treated with 25 nmol/L triptolide alone (48 h exposure) or 4 Gy IR alone, or both in combination (IR followed immediately by triptolide treatment). After 10 min, cells were washed twice with PBS and immediately subjected to flow-cytometric analysis. The percentage of cells in the high-red region or low-red and high-green region was measured under the different treatments.
Assays for activities of caspase-3, caspase-8, and caspase-9. The activity of caspase-3 was measured with fluorescent substrate assay according to the manufacturer's instructions (Molecular Probes). Briefly, 1 × 106 cells treated with either vehicle alone, triptolide alone (25 nmol/L), IR alone (4 Gy), or both in combination (IR followed immediately by triptolide treatment) for 24, 48, and 72 h were collected and resuspended in cold lysis buffer. About 50 μL of 2× reaction buffer was added to 50 μL of cell lysate and incubated for 2 h at 37°C with DEVD-R110, a caspase-3 substrate, which released fluorescence after its cleavage. Similarly, caspase-8 and caspase-9 activities were measured using kits purchased from BioVision following the manufacturer's protocols.
Western blot analysis. For detection of released cytochrome c, AsPC-1 cells were treated with 25 nmol/L triptolide alone, 4 Gy IR once, or both in combination. Exactly 48 h later, cells were washed with PBS and harvested in a buffer [20 mmol/L HEPES (pH, 7.5), 10 mmol/L KCl, 1.5 mmol/L MgCl2, 1 mmol/L EDTA, 1 mmol/L EGTA, 1 mmol/L DTT, 250 mmol/L sucrose, 1 mmol/L phenylmethylsulfonyl fluoride, 1 mg/mL aprotinin, and 1 mg/mL pepstatin A] for 20 min at 4°C and then homogenized using a Dounce homogenizer (40 strokes) and centrifuged at 10,000 rpm for 15 min at 4°C for the cytosol portion. Cytosol protein (50 μg) was subjected to 12% SDS-PAGE, transferred to immunoblotting membrane, stained with monoclonal antibody against cytochrome c, followed by antimouse secondary antibody conjugated to horseradish peroxidase (HRP), and then visualized with enhanced chemiluminescence (ECL; Amersham). The cleavage of caspase-3 and PARP is a characteristic marker of apoptosis. For the detection of this, AsPC-1 cells in 100-mm dishes at 80% confluence were treated with vehicle alone, 25 nmol/L triptolide alone, 4 Gy IR alone, or triptolide and IR in combination for 48 h, and then harvested with 1 mL of lysis buffer (1% Triton X-100, 0.5% Na deoxycholate, 0.5 μg/mL leupeptin, 1 mmol/L EDTA, 1 μg/mL pepstatin, and 0.5 mmol/L phenylmethylsulfonyl fluoride). The protein concentration of the lysate was determined by the bicinchoninic acid method (Pierce). About 30 μg of protein was loaded onto a 10% SDS-PAGE, electrophoresed, and transferred to a nitrocellulose membrane. The loading and transferring of equal amounts of protein were confirmed by staining the membrane with a Ponceau S solution (Sigma). The membranes were blocked with 5% fat-free milk in PBS (pH, 7.4) for 30 min and then incubated overnight with 0.2 μg/mL of anticleaved caspase-3 or PARP polyclonal antibodies separately. After washing, the membranes were incubated with HRP-labeled secondary antibodies for 1 h followed with ECL exposure. For reprobing glyceraldehyde-3-phosphate dehydrogenase (GAPDH), the blots were stripped with a buffer containing 50 mmol/L Tris-HCl (pH, 6.8), 2% SDS, and 0.1 mol/L β-mercaptoethanol.
Xenografts in nude mice. AsPC-1 cells were grown in 15-cm2 dishes to 80% confluence, harvested with 10 mmol/L EDTA, and resuspended in PBS for 107 cells/mL. A suspension of 2 × 106 cells in 0.2 mL PBS was injected s.c. into the hind leg of athymic nude mice using a 27.5-gauge needle. Tumors were allowed to grow for 7 days before treatment. Thirty-two nude mice with established tumors (all ∼100 mm3) were divided into four groups and treated with (a) vehicle (PBS) alone; (b) a single dose of 10 Gy IR; (c) 0.25 mg/kg of triptolide twice weekly for 4 weeks; or (d) triptolide plus IR (first dose of triptolide administered immediately after IR). The IR treatment of tumors grown in hind legs was carried out at a dose rate of 128 cGy/min with a Cs-137 Mark I irradiator on day 7 (initiation of treatment). Tumor size was measured thrice per week with Vernier calipers. The tumor volume was determined according to the formula: (length × width2)/2. The body weight of each mouse in each group was recorded once per week. Growth delay time (GD) was calculated as the time for treated tumors to reach ≥400 mm3 in volume minus the time for control tumors to reach ≥400 mm3 in volume. Most tumors were harvested on day 38; however, half of the animals (n = 4) from the combined treatment group had no detectable tumor on day 38, and these animals were therefore followed with continued measurements until recurrence. The enhancement factor (EF) was then determined as follows: EF = (GDIR + triptolide − GDtriptolide)/GDIR.
Terminal nucleotidyl transferase–mediated nick end labeling. Paraffin-embedded harvested tumor tissue sections were obtained by placing harvested tumors in 10% formalin with subsequent paraffin embedding after 48 h. Paraffin sections were subsequently placed on Superfrost-plus slides and were deparaffinized in xylene, rehydrated in graded ethanol, and transferred to PBS. The harvested tumor tissues were assessed for degree of apoptosis using terminal nucleotidyl transferase–mediated nick end labeling (TUNEL) assay. ApopTag in situ staining kits were used according to the manufacturer's instructions. The apoptotic cells were counted in eight independent tumors from each treatment group. The percentage of apoptosis was calculated by dividing the number of TUNEL-positive cells by the total number of AsPC-1 cells measured under the observation of light microscope.
Immunohistochemistry. Paraffin-embedded tissue sections were deparaffinized and rehydrated as above. The slides were rinsed twice with PBS, and endogenous peroxidase was blocked using 3% hydrogen peroxide in PBS for 1 h, washed section thrice with PBS, and incubated for 20 min at room temperature with a protein-blocking solution consisting of PBS (pH, 7.5) containing 5% normal horse serum. Tissue samples were incubated overnight at 4°C with a 1:50 dilution of rabbit-polyclonal anti–caspase-3 cleavage antibody. The samples were rinsed four times with PBS and incubated for 60 min at room temperature with the appropriate dilution of peroxidase-conjugated anti-rabbit immunoglobulin G. The slides were rinsed with PBS and incubated for 5 min with diaminobenzidine. The sections were then washed thrice with distilled water. A positive reaction was indicated by a brown staining. The counting method is the same as TUNEL.
Statistical analyses. All data are expressed as the mean ± SD of at least three determinations unless otherwise stated. The differences between two groups were determined by the two-sample Student's t test or one-factor ANOVA.
Triptolide enhances the effect of radiation on AsPC-1 cells in vitro. Triptolide at a dose of 50 nmol/L reduced the cell survival to 32%, whereas at a lower dose of 25 nmol/L, 52% of cells survived. Triptolide at 25 nmol/L was used to provide a window for measuring interaction with IR (Fig. 1A). As shown in Fig. 1B, with treatment of 4 Gy IR alone, more than 90% of AsPC-1 cells were viable as assessed by SRB, providing room for determination of combined effects with triptolide. Indeed, cell viability was reduced to 21% when IR at 4 Gy was combined with triptolide at 25 nmol/L, suggesting that the combination of triptolide and IR was complementary. To further delineate if this is a synergistic effect, a standard clonogenic assay was done. Cells were treated with 0, 3.125, 6.25, 12.5, or 25 nmol/L triptolide in combination with IR at a dose of 0, 2, 4, 6, or 8 Gy and cultured in 60 mm dishes for 3 weeks. The results were assessed with Chou-Talalay's synergism analysis, a widely used method to analyze and quantify the synergy in combination therapies (refs. 15–19; Fig. 1C and D). Radiation-induced clonogenic inhibition was correlated with the dose of triptolide (r = 0.988, P < 0.001). Table 1 shows the synergism analysis. At the chosen radiation and triptolide dose ranges, all combined treatments exhibited a combination index (CI) of <1 and a dose reduction index (DRI) of >1, supporting synergistic clonogenic cell killing by the combination of triptolide and IR (20, 21). The treatment of 25 nmol/L triptolide combined with IR (dose, 4-8 Gy) induced the most powerful synergistic effect (CI, 0.25; DRI, 7.78). Therefore, we chose the combination of 25 nmol/L triptolide and 4 Gy IR to investigate the interaction mechanism of triptolide and IR in vitro.
|TPL (nM) .||D0 (cGy) .||SF2 .||ER .||CI .||DRI .|
|TPL (nM) .||D0 (cGy) .||SF2 .||ER .||CI .||DRI .|
NOTE: Radiobiological parameters calculated from survival curves in (C) are based on single-hit multitarget (SHMT) model curve fit. The synergism analysis of triptolide with radiation at different doses were done with Chou-Talalay's combination index-isobologram and multiple drug-dose effect analysis method as described in Materials and Methods. A CI of <1 and DRI of >1 indicate synergy of triptolide and IR. *, P < 0.05, in comparison with control. §, P < 0.05, in comparison with triptolide alone.
Abbreviations: D0, dose required for reducing the fraction of cells to 37%, indicative of single-event killing; ER, enhancement ratio; SF2, survival fraction at 2 Gy.
Triptolide plus IR trigger apoptosis via both mitochondrial and death receptor pathways. We speculated that the underlying mechanism for the synergistic effect of the combined treatment might relate to one or both signal pathways for apoptosis. First, the mitochondrion-dependent apoptotic pathway was examined. When treated cells were stained with JC-1 (a dye indicating mitochondrial membrane depolarization, which can occur during early stages of apoptosis; refs. 22, 23), the percentage of cells in high-green and low-red regions (Fig. 2A) was increased to 76% in the combined treatment group, compared with 33% after 4 Gy IR alone or 51% after 25 nmol/L triptolide alone. Thus, mitochondrial membrane damage was increased after combined treatment. We then examined the release of cytochrome c by Western blot analysis. Although IR or triptolide alone increased the release of cytochrome c from the mitochondria, the combination further increased this release (Fig. 2B). The biofunction of released cytochrome c was shown by the increased activity of caspase-9 (Fig. 2C), a downstream precursor of cytochrome c. These data suggest that the enhanced killing effect of dual treatment is associated with the mitochondrion-dependent apoptotic pathway.
Second, the death receptor–related apoptotic pathway was explored. After combined treatment, the activity of caspase-8 (measured by enzyme substrate) was increased compared with the single treatment (Fig. 3A). For example, after 48 or 72 h of triptolide exposure, caspase-8 activity was elevated by 3.7- and 3.4-fold, respectively, and for triptolide in combination with IR, caspase-8 levels were increased to 5.8- and 5.1-fold, respectively. This increased activity occurred at 24 h after treatment, peaked at 48 h, and decreased at 72 h.
Two key apoptotic molecules in the common pathway were also studied. Here, caspase-3 activity was enhanced (Fig. 3B), peaked at 48 h, and remained elevated for 72 h after combined treatment, which was consistent with the increased cleaved form of caspase-3 as shown by Western blot (Fig. 3C). Triptolide exposure for 48 and 72 h increased caspase-3 levels by 22- and 20-fold, respectively. The combined treatment further increased caspase-3 levels by 41- and 35-fold, respectively. Again, IR alone had a minimal effect on caspase-3 levels, with elevation of only 1.2-fold at both 48 and 72 h after treatment. We used Western blot to detect caspase-3 cleavage. The data is consistent with the results of the caspase-3 activity assay. PARP, an enzyme responsible for the cleavage of DNA, was also highly activated 48 h after dual treatments compared with the single treatment, as evidenced by its cleavage 85 kDa measured by Western blot (Fig. 3D). Taken together, the synergistic effect of triptolide plus IR is likely related to the triggering of apoptosis via both mitochondrial and death receptor pathways.
Triptolide plus IR enhances apoptosis induction and G2-M accumulation. Finally, evidence for ongoing apoptosis is supported by annexin V staining, a characteristic marker for apoptosis. In the cells treated with 4 Gy IR alone, the percentage of cells stained positive with annexin V without PI was 15.1%, whereas in the cells treated with 25 nmol/L triptolide alone, the percentage was 42.1%. The combined treatment increased positive annexin V–PI staining to 70% (Fig. 4A). This shows that combined treatment augments the apoptosis induced by single treatment.
To determine the effect of triptolide on cell cycle, standard flow-cytometric analysis was done. Triptolide significantly increased cells in the G1 phase, along with a slight increase in G2-M (Fig. 4B). When combined with IR, the proportion of cells in G2-M was dramatically increased.
Combination of triptolide and IR enhances therapeutic effect in vivo. To determine whether the synergistic inhibitory effect of triptolide and IR observed in pancreatic cancer cells in vitro was translatable in vivo, AsPC-1 cells were injected into the hind legs of nude mice to establish a xenograft model. When tumors reached a minimum size of 100 mm3 (7 days after cell inoculation), tumor-bearing mice were randomly divided into four groups and treated with vehicle (PBS), triptolide alone (0.25 mg/kg, twice weekly), 10 Gy IR, or the combination (triptolide delivered immediately after IR). On day 38, mice with tumors were sacrificed, and the tumors were weighed. As shown in Fig. 5A, triptolide alone and IR alone each produced significant tumor volume regression and growth delay (P < 0.01); however, the combined treatment was even more effective, with four out of eight tumors decreasing in size from 100 mm3 to impalpable over about 3 weeks. The four animals with tumors that were undetectable at day 38 were also followed and showed a prolonged growth-delay time. Mock-treated tumors were large (≈1.7 g), and both single modality treatments reduced tumor weight to 0.2 to 0.3 g (control versus single modality, P < 0.01). Tumor mass after combined treatment for the tumors detectable at 38 days was 0.08 g (single modality versus combined P < 0.01, Fig. 5B). Tumor growth delay (TGD) was defined as the time delay for tumors to reach ≥400 mm3 in the treated group compared with control, vehicle-treated animals. As shown in Fig. 5A (inset), the TGD in vehicle-, IR-, and triptolide-treated groups was 0, 11, and 14 days, respectively. The TGD of the combination treatment group was 53 days, a significant difference compared with either single treatment. EF was 3.5.
Figure 5C showed that triptolide-treated mice had a transient decrease of body weight measured during week 3. All animals recovered with no deaths within 2 weeks. Interestingly, the combination of IR and triptolide did not reduce body weight at any time (Fig. 5C). Thus, low-dose, but effective triptolide produced low toxicity and reversible side effects.
Combination of triptolide and radiation enhances apoptosis in vivo. To determine whether apoptosis was involved in the triptolide-enhanced regression of AsPC-1 tumors, the TUNEL assay was used to quantify apoptosis in tumor sections from all groups (Fig. 6A). As shown in Fig. 6B, vehicle-treated tumors had an apoptotic index of 2.8%, the IR alone had an apoptotic index of 13.2%, the triptolide alone had an apoptotic index of 50.2%, and the combined treatment had an apoptotic index of 71.8% (P < 0.01). We confirmed that the TUNEL data were consistent with immunohistochemistry for caspase-3 cleavage (Fig. 6C). As shown in Fig. 6D, combined treatment-induced caspase-3 cleavage was 67.3% in comparison with triptolide alone (42.2%) and IR alone (7.1%; P < 0.01), indicating that the triptolide-enhanced antitumor effect was related to an induction of apoptosis, consistent with the findings in vitro.
Due to its anatomic location and physiologic function, pancreatic carcinoma is one of the most difficult cancers to treat. Only 10% of patients are eligible for surgical resection, and 90% of surgical patients experience cancer recurrence (24, 25). Gemcitabine is probably the most active chemotherapeutic agent, but it produces only modest response rates, with a median survival of 5.6 months (26). Other chemotherapeutic agents produce similarly poor response and survival rates (27, 28). Radiation is currently used primarily for palliative pain control. However, with the recent advancement of radiation delivery technologies, it is now possible to deliver highly conformal treatments to the pancreas and to metastases in the liver, providing an opportunity to intensify existing treatments. Unfortunately, even high-dose radiation when used alone is unlikely to produce a high rate of local control (1).
Based on the clinical observation that most pancreatic cancers are unresectable upon diagnosis, it is reasonable to conclude that highly focused radiation and chemotherapy are likely to remain mainstream therapy options, and it is therefore imperative to improve the efficacy and increase the potency of radiation therapy and drug treatments. In accordance with this need, we sought new agents and found that triptolide at an extremely low dose (25 nmol/L in vitro and 0.25 mg/kg in vivo) powerfully reduced growth of AsPC-1 cells and tumors. Indeed, in side-by-side comparisons, we found triptolide to have responses superior to gemcitabine (data not shown). Importantly, in our study, half of the tumors in the combined treatment group transiently disappeared, although the animals had only received a single radiation dose of 10 Gy. We speculate that regrowth might have been prevented by either a better timed radiation dose or a repeating cycle of drug plus radiation.
Regarding the toxicity at the effective drug dose (0.25 mg/kg i.v.), we noted that triptolide alone caused a transient weight loss that was most pronounced on day 20, but mice all regained their weight within 2 weeks without death. Interestingly, the mice given combined treatment did not experience weight loss. This might be due to better tumor control leading to lower deleterious effects of the growing tumor on the host. The tolerable side effects of triptolide in our study are consistent with other reports (29). In similar side-by-side experiments (data not shown), we found that mice given a dose of gemcitabine that produced a smaller benefit than that seen with triptolide also lost substantial weight. In that case, the animals did not fully recover from the weight loss. Notably, we used i.v. administration to treat mice by triptolide, similar to the administration of chemotherapy in the clinic. Triptolide is now in phase II trials for arthritis therapy, and toxicity is tolerable (30). Therefore, it is expected that triptolide could also be combined with radiation for pancreatic cancer treatment in humans with tolerable side effects.
As might be expected, the sensitivity of different cell lines to radiation or drug is variable (31–33). We therefore initially evaluated apoptosis after triptolide in three different pancreatic cancer cell lines. We found that AsPC-1 was least sensitive to triptolide or gemcitabine, compared with Panc-1 and MiaPaCa-2 (data not shown). We chose AsPC-1 for the current combination studies because it is one of the most resistant cell lines to radiation (34).
To determine which apoptotic path was triggered by the combination of triptolide and IR, molecules related to the mitochondrial and death receptor pathway were examined following single or combined treatment. The results indicated that the mitochondrial membrane was depolarized more with the combined treatment than with IR or triptolide alone. The damaged mitochondrial membrane was leaky (35), which allowed the movement of cytochrome c from mitochondria to cytoplasm, as evidenced by the increased level of translocated cytochrome c in the cytoplasm. The activated caspase-9 subsequently released by cytochrome c was evidenced by the increased activity of caspase-9. Caspase-8, a critical molecule in the death receptor pathway, was also highly activated by the combined treatment. Subsequent downstream apoptotic exertive molecules, caspase-3 and PARP, were highly activated. These data strongly suggest that the triptolide-enhanced IR tumor cell–killing effect is associated with the enhancement of the mitochondrial and death receptor pathways. It is noteworthy that only a single dose of 4 Gy IR was administered. IR damage was likely to be amplified by triptolide. The enhanced apoptosis in the presence of triptolide was observed to last for a relatively long period (at least 3 days). It is known that cross-talk exists between the mitochondrial and the death receptor pathways of apoptosis (36, 37). Because triptolide is a small hydrophobic molecule, it passes through the plasma membrane and penetrates and disturbs the mitochondria (38–41). Triptolide can also trigger the death receptor pathway. For example, triptolide can shift tumor cells from TRAIL resistant to TRAIL sensitive (42, 43). At this time, the pathway initially targeted by triptolide remains unknown.
It is well accepted that IR kills cells by damaging DNA, which results in reproductive dysfunction with or without apoptosis (44). The phase of the cell cycle is an important factor in cell sensitivity to IR, with the G1 and S phases being more radioresistant and the G2-M being the most radiosensitive. The accumulation of cells in the G2-M being the phase (62% measured at 48 h after IR in the present study) is consistent with reproductive failure and resulting tetraploid giant cells, and with synchronization of cells in the most radiosensitive portion of the cell cycle. The accumulation of cells in the tetraploid state may account for the prolonged proapoptotic state seen long after irradiation.
In conclusion, we have shown that triptolide not only exhibits a potent therapeutic effect on pancreatic cancer, but also produces a synergistic antitumor effect in combination with radiation both in vitro and in vivo. The synergy includes activation of the death receptor and mitochondrial apoptotic pathway, leading to prolonged apoptosis. Enhanced cell killing and tumor response are seen in a pancreatic cell line and tumor model that was previously shown to be one of the most resistant to both IR and standard chemotherapy. These results suggest that triptolide may be a promising candidate for combined modality therapy of pancreatic cancer.
Grant support: U.S. Army Medical Research and Materiel Command (DAMD17-00-1-0081 and DAMD17-01-1-0708) and Susan G. Komen Foundation (L. Zhang), the DAMD17-01-1-0708 (S. Yang), and the National Cancer Institute/Institute of Aging (W. Wang).
We thank Amy K. Huser for writing and editing assistance.