3p21.3 Tumor Suppressor Gene H37/Luca15/RBM5 Inhibits Growth of Human Lung Cancer Cells through Cell Cycle Arrest and Apoptosis

The loss of tumor suppressor gene (TSG) function is a critical step in the pathogenesis of human lung cancer. Deletion at chromosome 3p21.3 is the most frequent genetic alteration identified in lung cancer, observed in >90% of small cell lung cancers (SCLC) and in 50% to 80% of non–small cell lung cancers (NSCLC), indicating that the region contains one or more TSGs (1). Furthermore, loss of alleles at a 370-kb region of 3p21.3 is the earliest alteration found in preneoplastic lesions of the lung and sometimes in histologically normal lung epithelium of smokers (2). This timing suggests that 3p21.3 TSGs play a role in preventing lung cancer initiation. Thus, isolation and characterization of the 3p TSGs could be a first step toward developing new targeted treatments and diagnostic tools for lung cancer. Accordingly, the search for TSGs in this region has been the focus of extensive research over the past two decades, and many genes mapping to this region have recently been characterized (3–9). The two most prominent genes, already being evaluated for clinical utility, are fus-1 (phase I clinical trial; refs. 10, 11) and RASSF-1A (epigenetic biomarker; refs. 12, 13).

We previously showed that the H37 gene (also called Luca15 or RBM5; refs. 14, 15), concurrently cloned by us and others in 1996 and located in the 3p21.3 tumor suppressor region, has the characteristics of a TSG. We found reduced expression (compared with adjacent normal tissue) of the H37 transcript in 9 of 11 (82%) primary NSCLCs (16). By generating a polyclonal H37 antibody and using it for immunohistochemical analysis of primary NSCLC specimens, we further showed that 46 of the 62 (73%) cancers underexpressed H37 protein (16). Moreover, when we introduced H37 cDNA into human breast cancer cells that had 3p21-22 deletions, both anchorage-dependent and anchorage-independent growth was reduced (16). We found that subsequent transfection of H37 cDNA into one of the human lung cancer cell lines with a homozygous deletion in the TSG region (NCI-H740) resulted in a very low yield of H37-expressing clones, suggesting that H37 has antiproliferative properties (16). We also found that H37 suppressed anchorage-dependent and anchorage-independent growth in A9 mouse fibrosarcoma cells and inhibited tumor formation in nude mice (16).

H37's involvement in malignancy has been explored by other investigators as well, with results consistent to our findings. For example, H37 was identified as one of the serum antigens reacting with autologous antibody in renal cancer patients (17). H37/Luca15 is down-regulated in human schwannomas (18) as well as in ras-transformed Rat-1 rat embryonic fibroblastic cells (19). Ectopic expression of the H37 gene also suppresses growth of HT 1080 human fibrosarcoma cells (19). Several alternatively spliced transcripts from the H37 locus, and an antisense fragment, seem to function as apoptotic regulators in hematopoetic cells (20–24). In particular, growth inhibition by the full-length H37 correlated with higher apoptotic rates and G₁ cell cycle arrest in CEM-C7 human T cells (24). Most significantly, the H37/RBM5 gene was included in the 17 common gene signature associated with metastasis (one of the nine genes down-regulated in metastases) identified in multiple solid tumor types (25). Solid tumors carrying this gene expression signature had high rates of metastasis and poor clinical outcome (25). The fact that H37 shows tumor inhibitory properties in multiple different tissues is consistent with the finding that 3p21 is deleted in multiple other tumor types (although not as frequent as in the lung cancer), suggesting that this 3p21.3 TSG may be a general TSG (1).

Taken together, these data suggest that the H37 gene plays a significant role in tumor formation. However, tumor suppression by H37 has never been shown, systematically, in the tissue where H37 TSG would presumably be the most important (the lung). Therefore, there remains the need to develop an appropriate lung cancer model to thoroughly test H37 tumor suppression and investigate the physiologic function(s) of H37 and its molecular mechanism of tumor suppression in the most relevant/appropriate tissue type. To this end, we have developed a lung cancer cell line model in which H37 is ectopically expressed. By using this cell system, we have confirmed tumor suppressive activity of H37 in the lung and begun to characterize the molecular mechanism/biochemical pathways for H37's tumor suppression.

Cell culture. A549 cells were purchased from the American Tissue Type Collection (Manassas, VA). Cells were grown in RPMI 1640 supplemented with 10% fetal bovine serum, 2 mmol/L glutamine, and 1% penicillin G/streptomycin/fungizone solution.

Plasmid transfection and generation of stable H37 transfectants. The full-length H37 cDNA was subcloned in the pBK-CMV eukaryotic expression vector. The plasmid was transfected into cells by LipofectAMINE (Invitrogen, Carlsbad, CA) according to the manufacturer's protocol. After selection in G418 (700 μg/mL) for 2 weeks, cells were plated into 10-cm dishes at limiting cell dilutions. Single cells that grew into individual colonies were removed from the dish by cell scraping and transferred into a six-well plate. Each cell line was then expanded and tested for H37 expression. The transfected cells were strictly maintained in the same concentrations of G418 to avoid reversion of the cells.

Cell counts and soft agar assay. Cells were plated in triplicate in six-well plates (5,000 per well). After plating (day 0), seeding density was measured at day 1, and cells were counted at days 3, 5, and 7 by Coulter cell counter. For soft agar assay, cells were plated in 0.5% (bottom) and 0.3% (top) soft agar layers, in triplicate, in 35-mm dishes (20,000 per dish) and were incubated for up to 2 weeks before counting and photography. Colonies with >20 cells were counted.

Cell cycle analysis. Cells (1 × 10⁶) were harvested and washed in cold PBS followed by fixation in 70% alcohol for 30 minutes on ice. After washing in cold PBS three times, cells were resuspended in 0.8 mL of PBS solution with 40 μg of propidium iodide and 0.1 μg of RNase A for 30 minutes at 37°C. Samples were analyzed for DNA content by FACSCalibur (Becton Dickinson, San Jose, CA) using blue 488-nm laser for excitation and fl2 detector 585 ± 21 nm for emission.

Hoechst staining for apoptotic cell nuclei. Cells grown in four-well chamber slides (8,000 per chamber) were fixed in Carnoy's Fixative (one part glacial acetic acid to three parts absolute methanol; MP Biochemicals, Irvine, CA) and subsequently stained with Hoechst 33258 (0.05 μg/mL) in HBSS (MP Biochemicals). The cells were observed under a fluorescence microscope, and blue cells with apoptotic nuclear characteristics, such as nuclear condensation and fragmentation, were scored as apoptotic. Results are expressed as the percentage of the number of apoptotic cells compared with the total number of cells in 10 random 0.159-mm² fields.

Terminal deoxynucleotidyltransferase–mediated nick end labeling staining. The assay was done using the In situ Cell Death Detection kit, Fluorescein (Roche Applied Science, Indianapolis, IN), according to the manufacturer's instructions for adherent cell preparations. A terminal deoxynucleotidyltransferase–mediated nick end labeling (TUNEL) reaction mixture without terminal transferase served as a negative control. For a positive control, permeabilized cells were incubated with DNase I for 10 minutes at 15°C to 25°C to induce DNA strand breaks before labeling procedures. Cells plated onto microscope slides were air-dried and fixed with 4% paraformaldehyde (pH 7.4) for 1 hour, then incubated in 0.1% Triton X-100 with 0.1% sodium citrate on ice for 2 minutes. Fifty microliters of the TUNEL reaction mixture containing fluorescein-dUTP and terminal deoxynucleotidyl transferase were added to the cells with subsequent incubation in a humidified chamber for 60 minutes at 37°C in the dark. Apoptotic cells exhibiting a strong nuclear green fluorescence were detected using a standard fluorescein filter (520 nm). To quantify the apoptotic events, the number of cells undergoing apoptosis was counted in ten 0.159-mm² fields and expressed as a percentage of the total number of tumor cells in the same field.

Western blot analysis. Western blot analysis was done according to the standard protocol (26). Antibodies for the cell cycle–related proteins were purchased from Santa Cruz Biotechnology (Santa Cruz, CA), and all the antibodies for apoptotic pathways were purchased from BD Biosciences (San Jose, CA).

Bax knockdown by small interfering RNA. The SMARTpool small interfering RNA (siRNA; Dharmacon, Lafayette, CO) for Bax was used, which contains a mixture of four siRNA duplexes selected by the SMARTselection algorithm by the manufacturer. The siRNA transfection was done using the DharmaFECT1 transfection reagent (Dharmacon) according to the manufacturer's instructions. Briefly, A549 cells were seeded in 24-well plates (1.25 × 10⁴ in 0.5 mL per well) in antibiotic-free growth media 1 day before the transfection. For transfection, the growth media was replaced with the 500 μL of the transfection mixture (per well) containing 50 pmol siRNA and 2 μL DharmaFECT1 in the Opti-MEM I–reduced serum medium (Invitrogen). The transfection medium was replaced with the complete growth medium 24 hours after the transfection. The cells were harvested 96 hours after the transfection for the Bax expression analysis by Western blot and cell counts and TUNEL assays. Appropriate controls (all from Dharmacon) were used as follows: siGlow and siTox (positive transfection controls), siGAPDH (positive knockdown control), and siNon-Targeting Pool (negative control).

Mitochondrial membrane potential breakdown assay. Cells were seeded in four-well chamber slides 24 hours before staining. Cells were stained with the JC-1 reagent (Stratagene, La Jolla, CA) and incubated in the tissue culture incubator for 15 minutes according to the manufacturer's protocol. JC-1 aggregates in the mitochondria (only in live cells) were detected as the red color by a fluorescence microscope at excitation/emission = 540/570 nm; JC-1 monomers in the cytosol (in both live and apoptotic cells) were detected as green at excitation/emission = 490/520 nm.

Cytochrome c release assay. Cell fractionation was done by using the cytochrome c release apoptosis assay kit (Calbiochem, San Diego, CA) according to the manufacturer's protocol. Briefly, the cell pellet (5 × 10⁷ cells) was resuspended in 1 mL cytosol extraction buffer mix containing DTT and protease inhibitors, incubated on ice for 10 minutes, and disrupted with 30 to 50 passes of a Dounce homogenizer. The homogenate was centrifuged at 700 × g to separate the pellet containing the nuclei, unbroken cells, and large membrane fragments from the supernatant containing cytosolic fraction. The pellet was resuspended in 0.1 mL mitochondrial extraction buffer mix to obtain mitochondrial fraction; 10 μg of each cytosolic and mitochondrial fraction was subjected to 12% SDS-PAGE electrophoresis and Western blot analysis with the monoclonal cytochrome c antibody.

Caspase-9 and caspase-3 activity assays. Caspase-9 activity was analyzed using a fluorometric assay kit (R&D Systems, Minneapolis, MN) according to the manufacturer's instructions. Briefly, the cell pellet (1 × 10⁶ cells) was lysed in the lysis buffer on ice. After protein standard assay, 50 μL of cell lysate containing 125 μg of protein and the reaction buffer was pipetted into a 385-well plate. Subsequently, 5 μL of caspase-9 fluorogenic substrate [LEHD peptide conjugated to 7-amino-4-trifluoromethyl coumarin (AFC)] was added to each reaction well. After 1 hour of incubation at 37°C, the plate was read in a fluorescence microplate reader (Flexstation II, Molecular Devices, Sunnyvale, CA) at 400 and 505 nm excitation and emission wavelengths, respectively.

Caspase-3 activity was analyzed using a fluorometric immunosorbent enzyme assay kit (Roche Applied Science) according to the manufacturer's instructions. Briefly, caspase-3 from cellular lysates (2 × 10⁶ cells) was captured by a monoclonal anti-caspase-3 antibody precoated to the microtiter plate. Following the washing step, added substrate (Ac-DEVD-AFC) was cleaved proportionally to the amount of activated caspase-3. Generated free fluorescent AFC was determined fluorometrically at 505 nm. Results are reported as formed AFC (μmol/L) as determined against a standard curve.

Screening of multiple adherent lung cancer cells for the H37 protein expression and the H37 gene transfection. Prior experiments designed to ectopically express H37 in lung cancer cells, which were deleted at 3p21.3, were limited due to the cell line availability (16). The NCI-H740 cells would have been an ideal model system to test because both copies of 3p21.3 are deleted. However, they grow in suspension and are difficult to maintain in culture. Thus, we were only able to show H37's growth inhibition to a limited extent (16) because we could not grow the cells on soft agar for anchorage-independent growth assay nor generate stable transfectant cell clones to further characterize biochemical pathways of H37 tumor suppression. As an alternative strategy, we screened multiple adherent lung cancer cell lines to compare H37 protein expression levels. We found that among seven commonly used adherent lung cancer cell lines, A549 cells have the lowest H37 protein expression by Western blot analysis (Fig. 1A). These cells were used as parental cells for H37 transfection using pBK-CMV plasmid expression vector. Both transiently (3 days after transfection) and throughout the 2-week selection in neomycin, the H37 transfectants showed consistent growth retardation compared with the vector controls (Fig. 1B). The A549/H37 cells grew to only 20% of the control cells at the end of the 2-week selection period (data not shown). Morphologically, examination of the control cells revealed organized, tightly packed, uniform-shaped cancer cell colonies with a “cobblestone” appearance, whereas the H37 cells displayed an irregular, disorganized array of colonies consisting of flattened giant cells with cytoplasmic extensions (Fig. 1B). This disordered cell feature has been reported to be related to TSG transfections with other genes, including p53 (27).

Generation of stable clones and cell proliferation assays. To generate stable transfectants with defined levels of H37 expression, numerous single transfected cells were expanded to cell clones. As expected, these cells were difficult to propagate due to the growth inhibition induced by H37. A total of 28 H37 clones and 12 control clones were generated, and H37 expression levels in each clone were analyzed by Western blot analysis. H37 expression levels from two representative H37 clones (hereafter called H1 and H2) and two control clones (hereafter called C1 and C2) are shown in Fig. 2A. To measure proliferation rates, growth curves were generated for each clone over a 1-week period. By day 7, growth of H1 and H2 cells was reduced by 78% and 72%, respectively, compared with the average control level. In addition, significantly reduced numbers (as well as size) of the soft agar colonies resulted from H37 transfection in an anchorage-independent growth assay (Fig. 2C). The numbers of H1 and H2 colonies were reduced by 69% and 76%, respectively, compared with the average control levels (Fig. 2C). Subsequently, growth inhibition of the H37-overexpressing A549 cells were further evaluated by s.c. injection into nude mice. In contrast to the control cells that formed relatively large tumors within a short period of time, animals injected with H37.1 and H37.2 cells exhibited almost complete inhibition of tumor formation (Fig. 2D).

Analysis of cell cycle alterations induced by H37. We next explored the mechanisms underlying growth inhibition by H37. The lack of growth in H37 transfectants may result from cell cycle effects, cell death, or a combination of the two processes. The cell cycle distribution of cells transfected with control vectors or H37 was determined by fluorescence-activated cell sorting analysis of propidium iodide–stained cells, a measure of DNA content. As shown in Fig. 3A, H37 clones showed a higher proportion of cells in G₀-G₁ phase (69.44% and 61.71% for H1 and H2, respectively), compared with control cells (50.73% and 49.67% for C1 and C2, respectively), and a concomitant decrease in the proportion of cells in S phase (22.32% and 25.28% for H1 and H2, respectively) relative to that observed in controls (39.11% and 37.51% for C1 and C2, respectively).

Analysis of apoptosis induced by H37. To determine the contribution of cell death to H37-induced reductions in cell growth, we employed two different methods to detect apoptotic cells: Hoechst staining and TUNEL labeling. Hoechst staining, by fluorescence detection, enables more direct visualization of apoptotic cells, which undergo compaction and fragmentation of the nuclear chromatin, shrinkage of the cytoplasm, and loss of membrane asymmetry. As shown in Fig. 3B, condensed and fragmented nuclei were more frequently observed in H37-transfected cells and were only rarely seen in the control cells (∼3.5-fold increase in the H37 cells compared with the control). These findings were confirmed by TUNEL assay, in which labeled uridine bases are attached to the DNA nicks characteristic of apoptotic cells showing green fluorescence. As shown in Fig. 3C, approximately three times more of the H37 cells were apoptotic compared with the control cells. In addition, most of the cells counted apoptotic showed stronger intensity of green in the H37 cells.

Analysis of cell cycle– and apoptosis-regulated proteins in association with H37 overexpression. To investigate potential molecular mechanisms for H37-induced cell cycle alterations and apoptosis, we examined the effects of H37 on cell cycle and apoptosis regulatory proteins. Whole-cell protein extracts prepared from either the H37 and/or control A549 cells were analyzed for protein levels by Western blotting. Although a spectrum of cell cycle regulatory proteins was tested, including p53, pRB, cyclins A, B, C, D1, D2, D3, and E, p21, p27, cyclin-dependent kinase 2 (cdk2), cdk4, and cdk7/CAK, we detected noticeable changes in only two proteins, cyclin A and hyperphosphorylated RB, both of which were down-regulated in H37 cells (Fig. 3D). There were no marked changes in expression level in any of the other proteins tested (data not shown). Both cyclin A and hyperphosphorylated RB function in progressing cells from G₁ to S phase, and a decrease expression of these proteins is consistent with the G₁ arrest induced by H37. Similarly, of a large number of apoptosis regulatory proteins tested (p53, Bad, Bax, Bcl-2, Bcl-x, caspase-2, caspase-3, caspase-7, DFF45, FADD, Fas, PARP, Apaf-1, and CAS), we observed a significant alteration in only one protein, the proapoptotic protein Bax. Its expression was increased significantly in the H37 cells (Fig. 3D). Bax, downstream of p53, promotes apoptosis by triggering caspases cascades in apoptotic pathways, and its increased expression in H37 cells is consistent with increased apoptotic rates in H37 cells.

H37 transfection into the p53-null lung cancer cells. Our observation that H37-induced tumor suppression is not associated with a change in p53 protein expression in A549 cells prompted us to test a hypothesis that H37 may suppress tumor growth independently of p53. To this end, we transfected H37 into the H1299 human NSCLCs (p53 deleted) in parallel with A549 and H460 cells (both p53 wild type). Despite lack of p53, H1299 cells, upon H37 transfection, still manifested reduced cell proliferation and increased apoptosis (Fig. 4A). These changes were comparable with the levels seen in the A549/H37 and H460/H37 transfectants, although we could not detect any noticeable change in Bax protein expression in the H1299/H37 cells in contrast to the A549/H37 and the H460/H37 cells (Fig. 4A). Bax may not be active/functional in H1299 cells due to lack of its immediate upstream activator, p53, and/or other potential biochemical alterations associated with p53's absence; therefore, H37 may initiate mitochondrial apoptotic pathway through molecules further downstream of Bax in this cell line. The possibility still exists that H37- induced change in p53 protein expression level may be too subtle to be detected by Western blot, or p53 is activated by post-translational modifications and/or by any other alternative mechanisms in the p53 wild-type A549/H37 and H460/H37 cells. Nonetheless, our preliminary data showing inhibition of growth by H37 in the p53- null lung cancer cells may suggest existence of a novel, H37-mediated apoptotic pathways operating independent of the p53 regulations. It is intriguing to note that the p53-null H1299 has the highest H37 expression level (Fig. 1A), and investigation of multiple other p53-null lung cancer cells is warranted.

Analysis of Bax's biological involvement in H37-mediated apoptosis using Bax siRNA. To further confirm that Bax plays a direct functional role downstream of the H37-induced apoptotic pathway, we employed siRNA knockdown to reverse Bax expression comparable with the level seen in the A549/control cells (Fig. 4B). Bax knockdown in A549/H37 cells showed 280% increase in cell proliferation and 79% decrease in apoptotic rate compared with the negative control siRNA transfection (Fig. 4B). These changes are in contrast to the only 68% increase and 29% decrease, respectively, observed in the A549/control cells (Fig. 4B). The reverted growth properties in the H37 cells upon knockdown of Bax expression (which was initially induced by H37) suggests Bax functions biologically in the downstream of H37 in A549 cells.

H37-induced mitochondrial apoptotic pathways. To further characterize the molecular mechanisms of the H37/Bax–mediated apoptotic pathways, we investigated the influence of mitochondrial apoptosis signaling. Fluorescence microscopy following JC-1 staining revealed breakdown of mitochondrial membrane potential by H37. As shown in Fig. 5A, decrease in mitochondrial red staining in the H37 cells (due to loss of JC-1 aggregates in the mitochondria) indicated reduced mitochondrial membrane potential, and the number of the cells with intact mitochondrial membrane potential was shown to be decreased in H1 and H2 by 47% and 43%, respectively, over the average control level. Thus, the contents of the inter membrane space, including cytochrome c, were released into the cytosol in the H37 cells (Fig. 5B). Subsequently, cytochrome c accumulated in the cytosol-triggered caspase-9 activity, 96% and 83% increase in H1 and H2, respectively, over the average control level (Fig. 5C). Caspase-9, in turn, activated its downstream caspase-3, 78.8% and 88.2% increase in H1 and H2, respectively, over the average control level (Fig. 5D). The activated caspase-3 is responsible for the cytologic changes characteristic of apoptosis, including DNA fragmentation. The current data suggest that although H37's involvement in the death receptor–mediated apoptotic pathway remains to be in depth investigated, H37-mediated growth suppression, at least in part, employs regulation of mitochondrial apoptotic pathways.

The earliest and most common genetic change observed in lungs of smokers maps to the short arm of chromosome 3, specifically, 3p21.3 (1). This is the region where deletions are observed most frequently in lung cancer, and occasionally, the whole band is homozygously deleted. These observations suggest that TSGs reside at 3p21.3. The fact that specific alterations at this location can be detected in preneoplastic or preinvasive lesions also suggests that one or more 3p TSGs may function as “gatekeepers” for initiation of carcinogenesis and hence hold particular promise as biomarkers as well as potential therapeutic targets. Such biomarkers could be used to (a) identify individuals at high risk for lung cancer, (b) diagnose early lung cancer, and (c) assess the efficacy of lung cancer therapies and chemopreventive agents. In addition, 3p-linked biomarkers could have prognostic value.

H37/Luca15/RBM5 is among the 35 genes located within the 370-kb minimal area of deletions at 3p21.3 (14). The current report represents the first characterization of H37 tumor suppressive activity in lung cancer cells. The data showed that H37 inhibition of human lung cancer cell growth involves both interference with cell cycle progression and induction of apoptosis. Among the various cell cycle–regulated proteins tested, we have identified decreased cyclin A and phosphorylated RB to be associated with H37 overexpression. In G₁ phase, the cyclin D–dependent kinases initiate Rb phosphorylation, activating cyclin E for S-phase entry (28). Cyclin E/cdk2 then completes RB phosphorylation at the G₁-S transition, after which cyclin E is degraded and replaced by cyclin A during the S phase (28). The data showing H37-induced decreases in cyclin A and phospho-RB protein expression are consistent with the decreased proportion of S-phase cells and increased proportion of G₁ phase cells observed in these studies. The precise mechanism by which these events are achieved remains to be clarified, and studies are in progress to elucidate the potential signaling pathway(s) involved in H37 growth inhibition by G₁ phase cell cycle arrest.

Apoptosis is a complex, multistage process involving many genes. Among the wide spectrum of apoptosis regulatory proteins tested in our studies, the proapoptotic protein Bax was found to be overexpressed in H37-overexpressing lung cancer cells, with concomitant triggering of the mitochondrial apoptotic pathways, including caspase-9 and caspase-3 activations. A major regulation of the apoptotic death signal resides with the bcl-2/bax genes, the former an apoptosis-suppressing and the latter an apoptosis-promoting protein (29). Bax and Bcl-2 can form heterodimers, and overexpression of one antagonizes the other's effect; p53 functions, in part, by regulating the ratio of Bax/Bcl-2. In our studies, bcl-2 expression was not changed in the H37 cells. Thus, H37 expression may result in increased bax protein expression, leading to an increased apoptotic rate.

In an attempt to understand further the cellular/molecular functional mechanism of the H37-mediated tumor suppression, we have recently done computational algorithms to search for the transcription factor binding sites commonly present in the H37 gene promoter across species. Among those predicted to be true transcription factor binding sites was for the ZNF143/STAF protein, a transcription factor for eight different vertebrate small nRNA genes that function in RNA processing, such as splicing and polyadenylation (30, 31). This finding, although it needs to be validated by actual experiments, such as a gel-shift assay, is in a good correlation with the previous observations that the H37 protein contains RNA-binding motifs (i.e., RBM5 is another name for H37/Luca15; refs. 15, 32), and H37 shows in vitro RNA-binding activity (33). Taken together, our current hypothesis is that H37 may function in some aspects of post-transcriptional regulations of certain important proteins involved in cell cycle and/or apoptosis (i.e., Bax, cyclin A, RB, etc.). Based on this idea and in summary of our current findings, here, we propose a putative model for the molecular/cellular mechanism of the H37-mediated tumor suppression (Fig. 6). In this model, the RNA-binding protein H37 may directly interact with mRNA transcripts of certain important regulatory proteins involved in tumor formation, either stabilizing tumor-suppressing genes (i.e., Bax) and/or destabilizing tumor-promoting genes (i.e., cyclin A; Fig. 6). The RNA-binding proteins control mRNA stability and translation of the specific target mRNAs through a complex network of RNA/protein interactions. HuR, mdm-2, and VHL are only the few examples of an ever-increasing number of RNA-binding proteins carrying out important functions in cancer (34). Identification of the specific mRNA targets of H37 and/or its RNA-protein complex warrants future investigations.

The A549/H37 versus control cell pair system may be further used to examine H37's role in various physiologic pathways (e.g., transformation prevention, growth signal transduction, differentiation, genomic stability, and DNA damage response). This system may also be used for identifying downstream target molecules involved in the H37 growth suppression pathway. The genes differentially expressed in association with H37 growth inhibition are being studied by microarray analyses comparing the two cell populations differing only in H37 expression level, and we hope that these data will yield better insights into the cellular function of H37. The isogenic cell paired system may also be used to identify H37 protein partners or to validate the found partners by the yeast two-hybrid screening, which is currently under way. However, the most rigorous test of validating H37 as a bona fide tumor suppressor could come from generation of H37 knockout mice, and these studies are currently in progress.

In sum, the current studies provide a foundation to further explore the role of H37 in lung cancer pathogenesis and suggest a strategy for the development of rationally designed therapeutics using H37 as a target.

Grant support: Developmental Research Program Award from the University of California at Los Angeles Lung Cancer Specialized Programs of Research Excellence grant P50 CA90388 (J.J. Oh), American Lung Association of California Research Grant (J.J. Oh), and the Wolfen Family Lung Cancer Clinical/Translational Research Program at University of California at Los Angeles Jonsson Comprehensive Cancer Center.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Dr. David Reese for helpful discussion of this manuscript and Dr. Robert Damoiseaux at University of California at Los Angeles Molecular Shared Screening Resources for his help with caspase-9 activity assay.