NSCLC patients with unresectable stage III disease or medically inoperable disease account for about 40% of all patients diagnosed with NSCLC. Currently, radiation therapy plays an important role in achieving local control of the tumor and in the relief of symptoms resulting from metastatic disease. However, complete response rates are low and long-term survival remains poor. In addition, therapeutic efficacy is limited when cancer cells develop resistance to radiation. This underscores an urgent need for the development of more effective and less toxic treatments.
Matrix metalloproteinases (MMPs) comprise a large family of related proteolytic enzymes that includes collagenases, gelatinases, stromelysins, elastases and membrane-type (MT-MMPs).1 The expression and activity of MMPs increase in almost all human cancer types and are associated with advanced tumor stage and poor survival.2 MMP-2 belongs to the gelatinase subfamily and is believed to play an important role in extracellular matrix (ECM) turnover. MMP-2 can degrade denatured interstitial collagens I and III as well as native collagen IV, which is an important component of the basement membrane.
Previous reports have described the increased expression of MMP-2 after irradiation. Ionizing radiation increased MMP-2 activity in rat astrocytes,3 rat mesangial cells,4 rat kidney tubule epithelial cells,5 human bronchial epithelial cells and A549 lung cancer cells.6 In addition, irradiation enhanced secretion of MMP-2 in subcutaneous Lewis lung carcinoma (LLC-LM) tumors7 and pancreatic cancer cells.8 Analogous to its surgical removal, radiation therapy of a primary LLC-LM tumor is followed by the explosive growth of previously dormant metastases.9 The inflammatory process of radiation-induced lung injury includes infiltration by various inflammatory cells known to release high amounts of MMP-2.10 These studies suggest that a subset of patients may be at increased risk of metastatic growth after the surgical resection or radiation treatment of their primary tumors because of increased MMP-2 expression. These patients could benefit from treatment that includes the addition of MMP-2 inhibitors to radiotherapy.
Ionizing radiation leads to direct apoptotic events in the tumor cell population and/or induces arrest at cellular interphase checkpoints. This arrest allows the cell to repair DNA strand breaks before continuing the cell cycle or induces apoptosis if DNA repair is not possible.11 Our study was designed to examine the effects of the combining MMP-2 inhibition and radiation in lung cancer cells in vitro and in vivo. Our results demonstrate that MMP-2 inhibition enhances the therapeutic response of lung cancer cells to irradiation. We also show MMP-2 inhibition abrogates FoxM1 transcriptional pathways and its downstream signaling effector checkpoint kinase (Chk2)-mediated DNA repair response in irradiated lung cancer cells.
The following antibodies and reagents were used in our study: anti-Cyclin-B1 (Biomeda, Foster City, CA); anti-phospho-p53 (Ser-15) and anti-phospho-Cdc-25C (Ser-216) (Cell Signaling, Boston, MA); anti-FoxM1 (Affinity BioReagents, Golden, CO), anti-FoxM1 (for IP), anti-MMP-2, anti-p53, anti-p21Cip1/Waf1, anti-Cdc2, anti-phospho-Cdc2 (Thr-14/Tyr-15), anti-Chk1, anti-Chk2, anti-XRCC1 and anti-GAPDH primary antibodies, HRP-conjugated secondary antibodies, mouse IgG, p53 siRNA, FoxM1 siRNA and control siRNA (Santa Cruz Biotechnology, Santa Cruz, CA); anti-phospho-γ-H2AX (Ser-139), anti-phospho-Ser/Thr-Proline-MPM-2 antibodies and human recombinant MMP-2 protein (Calbiochem, San Diego, CA), ECL reagent (Amersham Pharmacia, Piscataway, NJ), Vectashield mounting medium with DAPI (Vector Laboratories, Burlingame, CA), DAB peroxidase substrate (Sigma, St. Louis, MO), TUNEL Detection kit (Roche Molecular Biochemicals, Indianapolis, IN), Annexin-V-FITC Apoptosis Detection Kit (BioVision Mountain View, CA); MTT cell growth assay kit (Millipore Corporation, Billerica, MA) and CytoTox-96 nonradioactive cytotoxicity assay kit (Promega, Madison, WI).
Cell lines and culture conditions
A549 cells were obtained from the American Type Culture Collection (ATCC, Manassas, VA) and maintained as a monolayer in RPMI 1640 medium (ATCC) supplemented with 10% fetal bovine serum (FBS; Japan), 50 U/mL penicillin and 50 μg/mL streptomycin (Life Technologies, Frederick, MD), hereafter referred to as complete medium, at 37°C in a humidified 5% CO2 atmosphere.
Adenoviral siRNA constructs, infection and radiation
Adenoviral siRNA for MMP-2 (Ad-MMP-2-Si) and scrambled vector (Ad-SV) were constructed and amplified as described previously.12 Viral titers were quantified as pfu/mL after infection of 293 cells. Titers obtained for the viruses used in this work are as follows: Ad-SV (7.6 × 1011 pfu/mL) and Ad-MMP-2-Si (5.0 ×1011 pfu/mL). The amount of infective adenoviral vector per cell in culture media was expressed as multiplicity of infection (MOI). Viral constructs were diluted in serum-free medium to the desired concentration, added to infected cells and incubated at 37°C for 1 hr. The necessary amount of complete medium was then added; cells were incubated for 24 hr and irradiated with X-ray radiation (a single dose of 10 Gy). After addition of complete medium, cells were incubated for 12 hr.
To prepare tumor-conditioned medium, A549 cells were infected with mock (PBS), 50 MOI of either Ad-SV or Ad-MMP-2-Si. After 24 hr of incubation, cells were irradiated with 10 Gy and cultured for 12 hr in serum-free DMEM/F-12 medium. The tumor-conditioned medium was collected, and equal amounts of proteins were used to determine MMP-2 activity. Gelatin zymography was performed as described previously.13
Western blot analysis
A549 cells were infected with mock, 50 MOI of Ad-SV or the indicated MOI of Ad-MMP-2-Si and irradiated as described earlier. Whole cell lysates were prepared by lysing cells in RIPA lysis buffer [50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1% IGEPAL, 1 mM EDTA, 0.25% sodium deoxycholate, 1 mM sodium fluoride, 1 mM sodium orthovenadate, 0.5 mM PMSF, 10 μg/mL aprotinin, 10 μg/mL leupeptin]. Equal amounts of protein were resolved by SDS-PAGE and transferred to a PVDF membrane as per the manufacturer's instructions. After blocking with 5% nonfat dry milk and 0.1% Tween-20 in PBS, membranes were incubated with 1:1,000 dilutions of primary antibodies. HRP-conjugated secondary antibodies were used to detect chemiluminescent signals with the ECL system. GAPDH levels served as a loading control. For detection of phospho-FoxM1, total cellular extracts were immunoprecipitated with FoxM1 antibody and immunocomplex was subjected to western blotting with antibodies for MPM-2, which recognizes the phosphorylated protein sequence phosphoserine/phosphothreonine–proline.
Total RNA was extracted using TRIZOL reagent (Life Technologies, Grand Island, NY) according to manufacturers' protocol, and RT-PCR was performed as described previously.12 PCR products were resolved on 2% agarose gels and were visualized by ethidium bromide staining. To normalize for the amount of input RNA, RT-PCR was performed with primers for the GAPDH. The specific primers used in our study were as follows: MMP-2, forward 5′-GTGCTGAAGGACACACTAAAGAAGA-3′ and reverse 5′-TTGCCATCCTTCTCAAAGTTGTAGG-3′; GAPDH, forward 5′-TGAAGGTCGGAGTCAACGGATTTGGT-3′ and reverse 5′-CATGTGGGCCATGAGGTCCACCAC-3′. To determine the quantity of PCR products on the agarose gel, images were generated by Alpha Innotech Image Acquisition and Analysis Software and processed for display.
MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay was carried out in 96-well plates using a MTT cell growth assay kit. Cells (5 × 103/well) were plated and infected with mock, 50 MOI of Ad-SV or Ad-MMP-2-Si in 16-wells/sample for 24 hr and then irradiated with 10 Gy. The proliferation rate was measured after irradiation at 12, 24, 36 and 48 hr. At the end of various treatments, 20 μL of MTT solution was added to each well and the plates were further incubated for 1 hr. Samples were read on a microplate reader using test wavelength 550 nm and reference wavelength 655 nm. The % of viability was calculated as 100 × mean absorbance of sample divided by mean absorbance of mock. Values were plotted accordingly.
Cells (1 × 104 cells/well) were plated in 96-well plates, infected and irradiated as described earlier. Forty-eight hours after irradiation, medium from each well was collected to measure the amount of released lactate dehydrogenase (LDH). Separate wells were exposed to lysis buffer (9% Triton X-100), and the medium was collected to measure the total amount of cellular LDH. The amount of cell death was calculated as a percentage of released LDH over total intracellular LDH.
Cells were infected with 50 MOI of either Ad-SV or Ad-MMP-2-Si for 24 hr and irradiated with 10 Gy. Cells were trypsinized and seeded (200 cells) in 100-mm Petri dishes. On Day 10 after irradiation, cells were fixed in methanol and stained with Giemsa and colonies (>50 cells) were counted. Survival fraction was defined as number of colonies divided by the number of plated cells.
Terminal deoxynucleotidyl transferase-mediated nick labeling (TUNEL) assay
We tested for apoptotic nuclei using the terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) enzyme reagent (Roche Molecular Biochemicals, Indianapolis, IN) as per the manufacturer's instructions. To determine the number of apoptotic cells, fluorescent-labeled nuclei were counted and imaged. The apoptotic index was calculated as follows: apoptotic index (%) = 100 × (apoptotic cells/total cells).
Propidium iodide (PI) positive cells were determined as described earlier.14 Briefly, A549 cells were infected and irradiated as described earlier. Cells were trypsinized, washed with cold PBS and incubated with 5 μL of PI in 1× binding buffer at room temperature for 5–10 min in the dark. FACS analysis was performed using a FACS Calibur Flow Cytometer (BD Biosciences, San Jose, CA) with an excitation wavelength of 488 nm and emission wavelength of 530 nm. Data acquisition and analysis were performed using CellQuest software (BD Biosciences).
Harvested A549 cells in exponential growth phase by brief exposure to trypsin-EDTA solution and cell viability was determined by trypan blue exclusion, and only cells with >95% viability were used. To determine the tumorigenic ability, tumor cells (5 × 106 cells/mouse) were injected s.c. into the flank of 4- to 6-week-old female nude mice. When mean tumor diameter reached 3–6 mm (12th day), the mice were separated into groups of 5 animals per group, and each group was injected intratumorally with PBS (mock) or 5 × 107 pfu of either Ad-SV or Ad-MMP-2-Si (10–25 μL volume) and were given 3 doses on alternate days. Between the 1st and the 2nd injections, and the 2nd and the 3rd injections, 1 group was radiated with a dose of 5 Gy, each time. Subcutaneous tumor growth was measured every 3rd day and tumor volume was calculated as π/6 × (Rmax × Rmin2). Regression of the subcutaneous tumor growth was followed for up to 4 weeks. Mice were euthanized when the tumor diameter in control mice measured between 13 and 14 mm. The subcutaneous tumors were removed and fixed in 10% phosphate-buffered formaldehyde.
The fixed tissue samples were then processed into paraffin blocks. Tissue sections (4–5 μm thick) were subjected to immunostaining for MMP-2 and FoxM1 as described earlier.14 Briefly, after deparaffinization and rehydration, tissue sections were permeabilized in 0.1% TritonX-100 in PBS. Endogenous peroxidase activity was quenched by 3% H2O2 in methanol for 20 min followed by PBS rinses. Nonspecific binding was blocked for 1 hr at RT with 1% BSA in PBS followed by incubated for 1 hr at RT with anti-MMP-2, anti-FoxM1 or anti-mouse IgG (negative control) at a concentration of 1.0 μg/mL. The slides were rinsed in PBS and incubated for 1 hr with HRP-conjugated secondary antibody (1:200). The slides were washed and stained with DAB peroxidase substrate solution followed by counterstained with hematoxylin and mounted. Mounted specimens were observed and photographed using light microscopy.
All data are presented as means ± SE of at least 3 independent experiments, each performed at least in triplicate. In the animal experiments, the mean differences in tumor volumes were compared among treatment groups using a one-way analysis of variance (ANOVA). Statistical differences are presented at probability levels of p < 0.05, < 0.01 and < 0.001.
To examine the effect of radiation on MMP-2 expression, we determined MMP-2 activity, protein and mRNA levels in A549 lung cancer cells irradiated with various doses of X-ray. Radiation-induced MMP-2 activity increased up to 10 Gy and decreased thereafter. The decreased levels of MMP-2 activity beyond the 10 Gy could be due to severe damage of DNA and other proteins. Maximal induction in MMP-2 activity was observed at 10 Gy with a ∼3-fold increase compared to the control (0 Gy). Western blot and RT-PCR analyses indicated that MMP-2 protein and mRNA levels increased up to ∼3-fold in 10 Gy-irradiated cells compared to mock (nonirradiated) controls (Fig. 1a). It has been hypothesized that MMPs may play a role in the induction of a malignant phenotype after ionizing radiation. Therefore, we first determined whether MMP-2 inhibition at 10 Gy enhances the radiosensitivity of A549 lung cancer cells. We have used an adenovirus carrying siRNA against MMP-2 (Ad-MMP-2-Si) to inhibit MMP-2 expression. Our previous studies clearly demonstrate that Ad-MMP-2-Si specifically inhibits MMP-2 expression without inducing the interferon pathway.12, 14 Figure 1b demonstrates cytotoxicity after Ad-MMP-2-Si infection at 10–50 MOI with 10 Gy radiation. Cytotoxicity derived by LDH release assay 48 hr after treatment is shown. Maximal effect was observed with the combination of 50 MOI and 10 Gy radiation (p < 0.01). The MOI of Ad-MMP-2-Si that gave the greatest radiosensitization at 10 Gy in A549 cells (50MOI) was used to determine the underlying mechanism in Ad-MMP-2-Si-mediated radiosensitivity. Figure 1c shows the cell survival rate after exposure to Ad-MMP-2-Si and irradiation alone and in combination as determined by MTT viability assay. The combination of Ad-MMP-2-Si and radiation caused a slight decrease in cell viability (∼80%) at 12 hr. By 48 hr, there was a dramatic reduction in viability to ∼20% when compared to the control population. No significant changes in cell growth were observed for the combination of Ad-SV (scrambled vector) and radiation when compared to treatment with radiation alone. We next determined the effect of Ad-MMP-2-Si on irradiation-induced MMP-2 expression. Figure 1d indicates that Ad-MMP-2-Si infection prior to radiation inhibits irradiation-enhanced MMP-2 activity, expression and mRNA levels by more than 90%.
Ad-MMP-2-Si infection inhibits colony formation and induces DNA damage and apoptosis in A549 cells
To determine whether the combination of MMP-2 inhibition and radiation has an effect on the kinetics of cell death when compared to cells treated with single agents alone, we carried out the colony-forming assay. As shown in Figure 2a, treatment of A549 cells with either Ad-MMP-2-Si or radiation decreased the colony-forming ability of cells by ∼40%. A 95% inhibition of colony-forming ability was observed in cells treated with a combination of Ad-MMP-2-Si and radiation when compared to nonirradiated cells. In contrast, the number of colonies in the Ad-SV-treated group was not affected. We next evaluated whether the combination treatment caused increased DNA damage in A549 cells. The histone variant H2AX is phosphorylated on Ser-139 (γ-H2AX) during DNA damage.15 Figure 2b shows a 2-fold increase in p-γ-H2AX in cells treated with Ad-MMP-2-Si and radiation treatments when compared to irradiated cells indicating an increase in DNA damage in the cells that received the combined treatments compared to radiation alone. We performed the TUNEL assay to determine if the cell death observed in Ad-MMP-2-Si plus radiation-treated cells was associated with apoptosis. Indeed, an increase in the number of apoptotic cells was observed with combination treatment of radiation and Ad-MMP-2-Si when compared to cells treated with radiation (Fig. 2c). Quantitative analysis indicated that the percentage of apoptotic cells was ∼12–12.5% in radiation or radiation plus Ad-SV-treated cells when compared to controls. However, there was a remarkable increase of ∼65% in TUNEL-positive cells in cells infected with Ad-MMP-2-Si prior to radiation.
Most mammalian cells exhibit transient delays in the G1 and G2 phases of the cell cycle after treatment with radiation to allow the cell to correct possible defects.16 To identify whether the growth inhibitory effect in cells that received the combined treatment of radiation and Ad-MMP-2-Si is caused by specific perturbation of cell cycle-related events, DNA contents of A549 cells were measured by flow cytometric analysis. As shown in Figure 3a, treatment of A549 cells with radiation resulted in an increase in the percentage of G2/M cells (from 13 to 50%) when compared to control cells. In cells infected with Ad-MMP-2-Si prior to radiation, the percentage of cells in G2/M phase decreased, the percentage of cells in sub-G1 increased to a maximum of 33% and the majority of cells were in the S phase at 24%.
Ad-MMP-2-Si suppresses p53 activation in response to radiation
To gain insights into the molecular mechanism(s) for Ad-MMP-2-Si and radiation-mediated attenuation of G2/M, we measured changes in the levels of proteins that are implicated in the G2/M transition. In response to genotoxic stress, the p53 protein is stabilized and activated by phosphorylation and, via its transcription activity, it induces the expression of numerous genes.17 Among them, p21Cip1/Waf1, an inhibitor of cyclin-dependent kinases, negatively regulates the G1/S and the G2/M phases of the cell cycle.18, 19 To study the effect of MMP-2 inhibition on radiation-induced p53 expression, cells were infected with Ad-MMP-2-Si before exposure to radiation and were analyzed by western blotting at 12 hr postradiation (before the onset of apoptosis). Figure 3b shows that p53 expression and phosphorylation on serine-15 residues were increased in irradiated cells. Further, Ad-MMP-2-Si infection prior to radiation strongly inhibited radiation-induced p53 phosphorylation (Fig. 3b).
MMP-2 inhibition reduces radiation-induced Chk2 expression and phopho-Cdc25C (Ser-216)
Mammalian cells respond to DNA-damaging agents by activating cell cycle checkpoints. Chk2 is required for the maintenance of DNA damage-induced G2 arrest.20 Irradiated cells exhibited a significant increase in Chk2 levels, whereas Chk1 activity displayed very small changes. Further, radiation-induced Chk2 levels were decreased in cells infected with Ad-MMP-2-Si prior to irradiation (Fig. 3c). Chk2 phosphorylates Cdc25C on serine-216,21 which creates a binding site to 14-3-3.22 The 14-3-3-bound Cdc25C phosphatase is then sequestered in the cytoplasm and prevents it from activating its nuclear substrate Cdc2 (Cdk1), a critical kinase for driving cells through G2.23 Corresponding to these studies, we show that radiation induced phosphorylation of Cdc25C on serine-216. Activation of Cdc2 kinase requires dephosphorylation at Thr-14/Tyr-15 residues.24 We therefore determined the Thr-14/Tyr-15 phosphorylation status of Cdc2 in cells treated with radiation and Ad-MMP-2-Si. Figure 3c indicates the accumulation of the phosphoryled-Cdc2 (Thr-14/Tyr-15) (negatively regulates cell cycle progression) in irradiated cells. However, MMP-2 inhibition in combination with radiation decreased radiation-induced phosphorylation of Cdc25C, which in turn results in activation of Cdc2 (dephosphorylates Cdc2) leading to cell cycle progression through G2 phase. Cyclin-B1 is the regulatory subunit of Cdc2 serine/threonine kinase, and accumulation of cyclin-B1 in the late G2 phase of the cell cycle is a prerequisite for mitotic initiation in mammalian cells.25 Others have shown that induction of cyclin-B1 after radiation abrogated the G2 delay.26 Therefore, we next determined the cyclin-B1 expression. Our studies show that cyclin-B1 expression decreased in irradiated cells, whereas cyclin-B1 levels were induced in cells infected with Ad-MMP-2-Si prior to radiation (Fig. 3c). These results suggest that MMP-2 inhibition removes the radiation-induced phosphorylation of Cdc2 at Thr-14/Tyr15, thereby activating Cdc2 and inducing cyclin-B1 and leading to cell cycle progression.
As FoxM1 transcription factor regulates the expression of cell cycle genes essential for DNA replication and mitosis during organ repair and cancer progression, we next determined the role of FoxM1. Figure 4a indicates an increase in FoxM1 expression and phosphorylation in irradiated cells compared to controls. However, Ad-MMP-2-Si infection prior to irradiation inhibited FoxM1 expression and radiation-induced phosphorylation. As FoxM1 is involved in regulating transcription of DNA repair factor XRCC1, we determined the expression of XRCC1 in the irradiated cells and those that received both Ad-MMP-2-Si and radiation. XRCC1 expression was induced in irradiated cells, but we could not detect XRCC1 in cells infected with Ad-MMP-2-Si prior to irradiation (Fig. 4a).
To evaluate the functional role of FoxM1 in radiation-induced cell cycle arrest, we downregulated the expression of endogenous FoxM1 in cells using RNA interference. A pool of siRNA oligonucleotides, directed against FoxM1, efficiently depleted cells of endogenous FoxM1. Irradiated cells transfected with FoxM1 siRNA displayed decreased levels of the XRCC1 repair gene and high levels p-γH2AX compared to cells subjected to radiation alone. The increased number of DNA breaks in FoxM1 decreased A549 cells, which was correlated with decreased phosphorylation of p53 (ser-15) and p21Cip1/Waf1 expression (Fig. 4b), thereby suggesting that FoxM1 functions upstream of p53 expression. Taken together, these results suggest that MMP-2 inhibition stops radiation-induced FoxM1 expression, which in turn inhibits cell cycle arrest and DNA repair.
To further test the hypothesis that MMP-2 inhibition leads to FoxM1 reduction, mock and Ad-MMP-2-Si-infected cells were cultured in the presence of recombinant human-MMP-2. As shown in Figure 4c, FoxM1 expression was induced by 1.61- and 2.71-fold in the presence of 50 and 100 ng/mL of MMP-2 compared to control (mock) cells. Similarly, addition of MMP-2 restored FoxM1 expression in Ad-MMP-2-Si-infected A549 cells. Our results demonstrate that MMP-2 induction contributes directly to FoxM1 expression in irradiated cells.
MMP-2 inhibition enhances radiosensitivity in vivo
On the basis of the encouraging results from the in vitro evaluation of enhancement of radiosensitivity, we evaluated the effects of MMP-2 inhibition in combination with radiation on A549 tumors propagated in the flanks of athymic nude mice. Treatment began on Day 12 when the A549 tumors had reached 3–6 mm diameter. Tumor growth was measured and the change in tumor size determined over time. As shown in Figure 5a, on Day 24 posttreatment, radiation alone (2 × 5Gy) and Ad-MMP-2-Si alone delayed A549 tumor growth more than mock (control) by ∼17% (750 ± 39.54 vs. 900 ± 53.21 mm3) and 56% (400 ± 37 vs. 900 ± 53.21 mm3), respectively. Notably, MMP-2 inhibition in combination with radiation reduced A549 tumor size completely.
One set of tumors from each treatment was excised at Day 5 posttreatment, and the tissue sections were stained to determine whether radiation induced MMP-2 and FoxM1 in vivo. Tumor sections from mice that received radiation alone showed intense staining for MMP-2 and FoxM1. However, MMP-2 expression and FoxM1 nuclear translocation were decreased significantly in tissue sections from mice treated with Ad-MMP-2-Si and radiation, thereby confirming our in vitro results (Fig. 5b; middle panel). Furthermore, we carried out the TUNEL assay to detect apoptotic cells (Fig. 5b; bottom panel). Corresponding to their growth characteristics, we found few apoptotic cells (∼40 ± 6.75 cells/field) in irradiated tumors. In contrast, we found abundant TUNEL-positive cells in the tumor sections from mice that received Ad-MMP-2-Si and radiation (210 ± 11.58 cells/field) (Fig. 5c).
The combination of radiotherapy and biological therapies, which interfere with cancer cell growth signaling, provides several potential benefits for improving therapeutic outcome. The combination may increase tumor response, protect normal tissues and exhibit nonoverlapping toxicities. Additionally, 2 partially effective therapeutic modalities may be combined without having to significantly reduce their dose levels to avoid treatment-related toxicities. In our study, our data demonstrated that radiation enhanced a dose-dependent increase in MMP-2 expression of A549 lung cancer cells. Infection with adenovirus carrying siRNA against MMP-2 (Ad-MMP-2-Si) markedly decreased MMP-2 expression and viability of irradiated lung cancer cells. We also demonstrated that MMP-2 inhibition abrogated radiation-induced cell cycle arrest and enhanced radiation-induced apoptosis.
We next sought to determine the mechanisms underlying MMP-2 inhibition-mediated, radiation-induced abrogation of G2/M arrest and apoptosis. Progress through the cell cycle is controlled by various surveillance mechanisms that block or delay transitions until each phase of the cell cycle is accurately completed. Integrity of the control pathways, which are termed “check points,” are critical for genomic integrity27 as well as for the repair and survival of cells exposed to DNA-damaging agents.24 A well-known tumor suppressor, p53 protein, senses DNA damage at several stages of the cell cycle, and accordingly determines whether the cell needs to arrest at the subsequent checkpoint to undergo DNA repair or proceed through it.28 Recent studies have shown evidences that p53-mediated pathways promote radiation-induced cell cycle arrest in G2 and that the duration of G2 arrest was longer in cell lines with wild-type p53.29–32 It has been shown that p21Cip1/Waf1 could also be involved in p53-mediated G2 arrest.33 p53-defective cells fail to maintain the DNA damage-induced arrest, and cells gradually escape the checkpoint and progress into the M phase, which is when they eventually undergo aberrant mitoses.34 Consistent with these observations, we show that MMP-2 inhibition reduces radiation-induced p53 and p21Cip1/Waf1 expression in A549 cells.
Activated p53 transcriptionally upregulates genes that are involved in DNA repair and/or cell cycle arrest.35 Checkpoint kinase-2 (Chk2) activation is most pronounced in response to DNA damage caused by DNA double-strand breaks such as those produced by radiation.36 Our studies clearly indicate that radiation-induced Chk2 is more pronounced than Chk1. Further, the radiation-induced Chk2 levels were reduced to lower than control levels in cells infected with Ad-MMP-2-Si prior to radiation. These observations are consistent with previous reports indicating that Chk2 rather than Chk1 is the primary checkpoint kinase responsive to radiation treatment.37 G2 delay after radiation is marked by delayed expression of cyclin-B1.26 Previous studies indicate that the cellular decision to enter into apoptosis is regulated, at least in part, by the abundance of cyclin-B1 protein. Antisense inhibition of cyclin-B1 accumulation prevents γ-radiation-induced apoptosis in human hematopoietic cells, and ectopic cyclin-B1 expression is sufficient to induce apoptosis.38 In human cells, an inhibition of cyclin-B1 transcription by the p53 tumor suppressor prevents G2/M transition.39 Accordingly, our data also indicate that radiation inhibited cyclin-B1 expression, whereas MMP-2 inhibition induced cyclin-B1 expression in irradiated cells. Taken together, our studies indicate that MMP-2 inhibition enhances radiosensitivity by abrogating the radiation-induced G2/M arrest.
FoxM1 signaling is involved in cell proliferation and apoptosis, which affects the development and function of many organs.40 Loss of FoxM1 expression generates mitotic spindle defects, delays cells in mitosis and induces mitotic catastrophe.41 We show the involvement of FoxM1 in the transcriptional response during DNA damage/checkpoint signaling in cells treated with Ad-MMP-2-Si and radiation. MMP-2 inhibition prior to radiation abrogated radiation-induced FoxM1 expression. We also demonstrated that siRNA silencing of FoxM1 levels in irradiated A549 cells reduced p53 and p21Cip1/Waf1 expression. The depletion of FoxM1 also increased cell sensitivity to radiation as determined by decreased expression of the repair gene XRCC1 and increase in DNA damage as determined by increased expression of phosphorylated-γ-H2AX. Interestingly, we observed that addition of recombinant MMP-2 induces FoxM1 expression in A549 cells. Previous reports indicate that FoxM1 induces MMP-2 expression and has a binding site for the MMP-2 promoter.42 Currently, we do not fully understand how MMP-2 alters FoxM1 expression, and as such, it is the focus of our ongoing investigation. Consistent with in vitro findings, A549 tumor xenografts treatment with the combination of Ad-MMP-2-Si and radiation suppressed tumor growth significantly more than either Ad-MMP-2-Si or radiation alone. Further, tumor sections from mice that received the combination treatment of Ad-MMP-2-Si and radiation showed less immunoreactivity for MMP-2 and FoxM1 when compared to nonirradiated and irradiated tumors. These results identify a novel role for FoxM1 in the transcriptional response during DNA damage/checkpoint signaling in irradiated lung cancer cells. Previous studies have shown that a cell-penetrating ARF peptide inhibitor of FoxM1 selectively induced apoptosis in human hepatocellular carcinoma cell lines and mouse models.43 Downregulation of FoxM1 by FoxM1 siRNA inhibited the expression of MMP-2, MMP-9 and VEGF and inhibited cell growth in pancreatic cancer.44 Partial deletion of mouse FoxM1-floxed targeted allele in vivo was sufficient to significantly reduce the number and size of lung adenomas induced by urethane.45 Thus, inhibition of FoxM1 along with radiation represents potential new clinical strategies and a worthwhile target for therapeutic intervention in lung cancer.
In summary, MMP-2 inhibition can overcome the G2/M arrest in irradiated cells, thereby driving cells through a lethal mitosis (Fig. 6). As such, it is potentially useful as a radiosensitizer in lung cancer therapy. These findings suggest that the concomitant use of MMP-2 inhibitor during radiotherapy could be a potential therapeutic approach to improve the efficacy of radiotherapy for lung cancer patients.
The authors thank Ms. Noorjehan Ali for technical assistance, Ms. Shellee Abraham for assistance in manuscript preparation and Ms. Diana Meister and Ms. Sushma Jasti for review of this paper. This study was supported by grant from Caterpillar, Inc., OSF Saint Francis, Inc., Peoria, IL (to J.S.R.) and American Cancer Society Grant #06-03 (S.S.L.).