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Departments of Oncology (Y.H.L., T.L., M.H., R.P.-S.) and Molecular Genetics (Z.Y., T.K.W.), Albert Einstein College of Medicine, Bronx, New York
Received February 6, 2007; accepted April 23, 2007
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; Chellappan et al., 1998). Several lines of evidence have demonstrated that growth factors trigger cascades of intracellular signals that lead to activation of nuclear transcriptional factors that activate cyclin/CDK complexes; active cells pass the G1 checkpoint and embark on DNA replication in S phase (Matsushime et al., 1991). G1/S transition is positively controlled by two families of CDKs, including the complexes of CDK2/cyclin E and cyclin A or the complexes of CDK4/cyclin D or CDK6/cyclin D. In contrast, the complexes of CDK inhibitors can be subdivided into two families, including the INK4 family, consisting of p16INK4a, p15INK4b, p18INK4c, and p19INK4d and KIP/CIP families consisting of p21WAF1/CIP, p27KIP1, and p57KIP2 (Sherr and Roberts, 1999). Other critical events for the G1/S transition are Rb protein phosphorylation and the release of its regulatory E2F proteins, which translocate to the nucleus and induce transcription of target genes that are required for cell proliferation (Weinberg, 1995; Dyson, 1998).
The epidermal growth factor receptor (EGFR) is a tyrosine kinase receptor of the ErbB family. Upon ligand binding, EGFR may either homodimerize or heterodimerize, resulting in transautophosphorylation (Yarden and Sliwkowski, 2001). The tyrosine-phosphorylated EGFR then served as a docking molecule to initiate the activation of downstream pathways, including the activation of PI3/AKT (promoting cell survival) and/or the activation of Raf/Ras/mitogen-activated protein kinase cascades (associated with cell proliferation) (Salomon et al., 1995). Moreover, EGFR and its family are implicated in the regulation of cell growth, transformation, and apoptosis (Klapper et al., 2000). Many tumor cells, especially epithelial cell-derived tumors, express elevated levels of EGFR or express mutant versions of ErbB family members. A number of reports have demonstrated high expression of EGFR in NSCLC cells (Haeder et al., 1988; Scagliotti et al., 2004). Because increased EGFR expression is known to correlate with poor clinical outcome in patients with NSCLC, the EGFR has been considered a potential therapeutic target. In recent years, several compounds have been developed that directly target the EGFR signaling pathway and have significant anticancer activity (Noonberg and Benz, 2000; Herbst and Bunn, 2003). Erlotinib (Tarceva; OSI-774; OSI Pharmaceuticals, Melville, NY) is an orally bioavailable quinazoline derivative that selectively inhibits the EGFR tyrosine kinase by competitively inhibiting the intracellular ATP binding domain and blocking signal transduction pathways implicated in cell proliferation and survival of cancers (Moyer et al., 1997; Pollack et al., 1999). Preclinical studies demonstrated erlotinib's potent activity against tumor cell growth accompanied by suppression of EGFR activation. Erlotinib as a single agent has demonstrated significant clinical activity even in patients with NSCLC treated previously and has improved patient survival in a randomized, placebo-controlled trial (Shepherd et al., 2005), and recently it has been approved by the U.S. Food and Drug Administration as the second/third line for treatment of patients with NSCLC. Although erlotinib has marked antitumor activity in both in vitro and in vivo systems, the mechanisms of its antitumor effects remain to be elucidated. In this work, we used the erlotinib-sensitive human H322 NSCLC cell line as model to examine the effects of erlotinib on cell proliferation and cell cycle machinery. Our results demonstrate that erlotinib treatment causes cells to accumulate at G1/S phase accompanied by a decline in the expression of G1-related regulators, remarkable suppression of CDK2 and CDK4 activities, and induction of CDK inhibitor p27KIP1. In addition, we found that erlotinib treatment resulted in Rb hypophosphorylation. Moreover, we found that erlotinib induces p27KIP1 accumulation via promotion of p27KIP1 transcription and protein stabilization. Erlotinib treatment resulted in p27KIP1 translocation to the nucleus. Knockdown of p27KIP1 expression with p27 siRNA caused abrogation of G1 phase arrest and cell growth inhibition by erlotinib. It is noteworthy that we observed a direct relationship between G1 phase arrest and cell sensitivity to erlotinib in several human NSCLC cell lines. The results provide insights into the cell cycle effects of erlotinib and may be used as potential surrogate endpoints of drug action in clinical studies.
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Materials and Methods |
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Cell Lines and Cell Culture. Human non-small-cell lung cancer cell lines (H322, H358, A431, H460, A549, H596, and H1299), and human skin epidermoid carcinoma A431 cells were purchased from American Type Culture Collection (Manassas, VA). Human head and neck carcinoma HN5 cell line was a generous gift from OSI Pharmaceuticals (Farmingdale, NY). All cell lines were grown in RPMI 1640 medium with 10% fetal bovine serum in a humidified air atmosphere with 5% CO2.
Cell Growth Assay. Exponentially growing cells (2 x 104 cells/well) were plated on a 96-well plate overnight. After cell attachment, cells were exposed to various concentrations of erlotinib at 37°C for 72 h. After exposure, cell survival fractions were assessed by viable cell count with trypan blue exclusion or by colorimetric assay based on the reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium (MTT).
Cell Cycle Assay. H322 cells were exposed to various concentration of erlotinib for 24 h or to 2 µM erlotinib for the indicated times. Cells were washed twice with cold PBS solution and harvested by trypsinization. After fixing with cold 75% ethanol overnight, cells were stained with 1 µg/ml propidium iodide and exposed to 5 µg/ml RNase I at room temperature for 3 h. The cell cycle distribution was assessed by FACS flow cytometer analysis (BD Biosciences, San Jose, CA). For determination of BrdU incorporation into DNA, cells were treated with 2 µM erlotinib for the indicated time, and then 10 µM BrdU was added into cell culture. After incubation for 1 h, the incorporated BrdU was detected with an FITC-BrdU assay kit according to the manufacturer's instruction (Calbiochem, Cambridge, MA).
Immunoblot Analysis. Cells were lysed with lysis buffer containing 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM NaF, 1 mM phenylmethylsulfonyl fluoride, 1 mM dithiothreitol, 20 µg/ml leupeptin, 20 µg/ml aprotinin, 0.1% Triton X-100, and 1% SDS at 0 to 4°C for 15 min. Equal amounts of lysates (50 µg of protein) were subjected to electrophoresis on either 7 or 12% SDS-PAGE. After electrophoresis, protein blots were transferred to a nitrocellulose membrane and probed with the corresponding primary antibodies. The detected protein signals were visualized by an enhanced chemiluminescence reaction system (GE Healthcare, Chalfont St. Giles, Buckinghamshire, UK).
CDK Kinase Assay. Cells were exposed to 2 µM erlotinib for the indicated times and harvested by trypsinization. Cells were suspended in a lysis buffer on an ice-bath for 10 min. After centrifugation at 15,000g at 4°C for 10 min, the supernatant was collected for immunoprecipitation. Equal amounts of supernatant (500 µg of protein) were incubated with 2 µg of anti-CDK2 or anti-CDK4 antibodies and 25 µl of protein A/G conjugated agarose beads at 0 to 4°C overnight. After washing three times with lysis buffer, immunoprecipitates were incubated at 30°C for 15 min in 30 µl of reaction mixture containing 20 mM HEPES, pH 7.4, 10 mM p-nitrophenyl phenylphosphonate, 20 mM MgCl2, 1 mM EDTA, 1 mM Na2VO4,1 µM ATP, 1 µCi of [
-32P]ATP (Amersham), and 5 µg of histone H1 (Sigma) as a substrate for CDK2 assay or 5 µg of Rb fusion protein (Cell Signaling) as a substrate for CDK4 assay. The reaction was terminated by the addition of 2x SDS-PAGE sample buffer. After boiling for 5 min, the supernatants were collected by centrifugation at 15,000g for 5 min and then subjected to 12% SDS-PAGE. Activities of CDK2 and CDK4 were determined by autoradiography of the dried gels.
Subcellular Fractionation. Cells were treated with 2 µM erlotinib or with the same volume of medium containing 0.1% DMSO as a control for 24 h, washed twice with ice-cold PBS solution, and harvested by trypsinization. Cells were suspended in an ice-cold nuclei isolation buffer containing 10 mM HEPES, pH 7.5, 1.5 mM MgCl2, 10 mM KCl, 0.5 mM dithiothreitol, and 1% Triton X-100 and incubated on an ice-bath for 5 min. After centrifugation at 800g at 4°C for 5 min, the supernatant was collected as a cytosolic fraction. The pellets were resuspended in lysis buffer containing 1% SDS, and after incubation on an ice-bath for 5 min, the lysate was centrifuged, and supernatant was collected as a nuclear fraction. After determination of protein concentration with a Bio-Rad DC protein assay kit (Bio-Rad, Hercules, CA), equal amounts (50 µg of protein) of cytosolic and nuclear fractions were subjected to 15% SDS-PAGE, and p27KIP1 was detected by immunoblot analysis as described above.
Real-Time RT-PCR Analysis. Total RNA was isolated from H322 cells after treatment with 2 µM erlotinib for the indicated times by phenol/chloroform extraction, and cDNA was produced with SuperScript II reverse transcription (Invitrogen, Carlsbad, CA). The standard real-time RT-PCR was performed using the following primers: p27 primers: forward, 5'-CTGCCCTCCCCAGTCTCTCT-3', and reverse, 5'-CAAGCACCTCGGATTTT-3'; and β-actin primers: forward, 5'-GATGAGATTGGCATGGCTTT-3', and reverse, 5'-CACCTTCACCGTTCCAGTTT-3'. All assays were performed using duplicate samples of reverse transcriptase product. The mRNA expression of p27 was normalized using the dCt = [Ct(p27) - Ct(β-actin)] method (Livak and Schmittgen, 2001). The increased folds of p27 mRNA were calculated as relative to p27 mRNA level at time 0.
Luciferase Activity Assay. p27 promoter containing luciferase reporter construct and cDNA empty vector was a gift from Dr. T. Sakai (Department of Molecular-Targeting Cancer Prevention, Kyoto Prefectural University of Medicine, Kyoto, Japan) (Inoue et al., 1999). H322 cells were transiently transfected with p27 luciferase reporter cDNA or with cDNA empty vector by a Lipofectamine kit (Invitrogen) according to the manufacturer's instructions. After transfection, cells were treated with 2 µM erlotinib or with the same volume of medium contained 0.1% DMSO as a control for the indicated times and then harvested in 1x lysis buffer. Luciferase activity was measured by the luciferase assay system kit (Promega). For normalization of transfection efficiency, 2 µg of Renilla reniformis (sea pansy) luciferase expression plasmid (pRL-TK vector; Promega) was included in the transfection.
p27 siRNA Transfection. p27 siRNA and nonspecific siRNA were purchased from Dharmacon (Lafayette, CO), and transfections of p27 siRNA and nonspecific siRNA were performed with the use of Oligofectamine (Invitrogen) according to the manufacturer's instruction. After p27 siRNA transfection, cells were exposed to 2 µM erlotinib or the same volume of medium as a control at 37°C for 24 h. Cells were washed twice with cold PBS solution, and cell pellets were divided into two aliquots. One was for cell cycle analysis by FACS analysis as described above. The other was prepared for the determination of p27 expression by immunoblot analysis. For determination of cell growth, cells were plated on a 12-well plate and transfected with p27 siRNA or without siRNA as control. p27 siRNA-transfected and -untransfected H322 cells were exposed to 2 µM erlotinib or to the same volume of medium as a control for the indicated times. At the specified time point, cells were harvested by trypsinization, and the viable cell numbers were assessed by trypan blue exclusion.
Immunofluorescence Staining. Cells were plated on a glass cover and treated with 2 µM erlotinib or with the same volume of medium contained 0.1% DMSO as a control for 24 h. After treatment, cells were washed twice with cold PBS solution, fixed with 4% paraformaldehyde in PBS solution at room temperature for 15 min, and then treated with 1% Nonidet P-40 in PBS solution for 30 min. After blocking with 5% bovine serum albumin in PBS solution for 30 min, cells were incubated with anti-p27KIP1 antibodies (1:500) at room temperature for 1 h. After washing three times with PBS solution, cells were incubated with fluorescence FITC-conjugated secondary antibodies (1:1000) and 100 ng/ml DAPI for 30 min in a dark room. The immunofluorescence signals were visualized with a Nikon Eclipse E400 fluorescence microscope (Nikon, Tokyo, Japan).
Data Analysis. Data are presented as mean ± S.D. of three independent experiments. The comparisons were made with a t test, and the difference was considered statistically significant if the p value was <0.05.
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; Pollack et al., 1999), we sought to extend these findings to H322 cells. An accumulation of cells at G1 phase and reduction of S and G2/M phase cells occurred in H322 cells after treatment with different concentrations of erlotinib for 24 h or with 2 µM erlotinib for the indicated times. Erlotinib exposures from 2 to 10 µM resulted in a range of 
80% of cells at G1 phase compared with 52% of control cells at G1 phase (Fig. 1B). The time course studies demonstrate that erlotinib-induced G1 phase arrest (70% of cells) at 12 h, reached a maximum (81%) at 24 h, and remained high over experimental periods (Fig. 1C). Furthermore, we used BrdU incorporation into DNA to determine the effect of erlotinib on cell-cycle progression from G1 to S phase transition. As shown in Fig. 1D, the numbers of BrdU-positive cells standing for the cell cycle at S phase were dramatically reduced by 7% after 24-h exposure to 2 µM erlotinib compared with 38% of BrdU-incorporated cells at time 0 and decreased to complete abolishment (2%) after 48-h exposure. The results show that erlotinib induces cell growth inhibition accompanied by a strong blockade of cell-cycle progression from G1 to S phase.
Effects of Erlotinib on the Expression of G1/S-Related Cell Cycle Regulators, CDK Kinase Activity, and Rb Phosphorylation. Next, we investigated the effect of erlotinib on intracellular expression of cyclins A, E, D1, D2, and CDK2 and CDK4 by immunoblot analysis. With H322 cells exposed to 2 µM erlotinib for the indicated periods, reduction of intracellular levels of cyclin E and CDK2 occurred at 12 h, and reduction of cyclin A level started at 24 h; the extents of reduction of these regulators were gradually increased thereafter. In contrast, the levels of CDK inhibitor p27KIP1 were significantly induced in a time-dependent manner; i.e., the endogenous amounts of p27KIP1 were barely detected at time 0 to 8 h but were clearly induced 12 h after treatment and increased thereafter. The level of p21WAF1/CIP was barely detectable in H322 cells over time (data not shown), and the level of p16IKK4a was unchanged over experiment periods (Fig. 2A). Furthermore, we found that erlotinib treatment resulted in a time-dependent suppression of CDK2 activity as measured by the use of histone H1 as a substrate and reduction of CDK4 activity as assessed using Rb fusing protein as a substrate (Fig. 2B). The active, phosphorylated Rb is believed to play a critical role in the regulation of cell cycle progression at the G1/S phase transition (Berthet et al., 2006). We therefore examined whether erlotinib-induced G1 phase arrest could be involved in the disruption of Rb phosphorylation. The results shown in Fig. 2C demonstrate that erlotinib treatment led to down-regulation of total Rb protein levels and decrease in Rb protein phosphorylation as detected by slow migration of phosphorylated Rb bands in a time-dependent manner; i.e., the reduction of total Rb protein level and its phosphorylation were seen at 12 h after drug treatment and increased over experimental times. In addition, we compared the inhibitory effect of erlotinib on Rb phosphorylation at different sites probed by immunoblots using the corresponding antibodies and found that Rb phosphorylation at Ser780 and Ser795 were notably inhibited after 12 h of erlotinib exposure with similar patterns of inhibition of total Rb phosphorylation; however, p-Ser795 seemed more susceptible to erlotinib than p-Ser780. For example, Rb p-Ser795 was fully abolished but was only 60% reduced at Rb p-Ser780 after 24 h of erlotinib exposure.
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Erlotinib Induces the Promotion of p27KIP1 Gene Transcription. To further understand the molecular mechanisms of erlotinib action on G1/S phase arrest and the induction of p27KIP1, we examined the effect of erlotinib on p27KIP1 expression at the protein level by immunoblots and at the transcriptional level by real-time RT-PCR analysis in H322 cells after treatment with 2 µM erlotinib for the indicated times. The results as shown in Fig. 3A demonstrate that erlotinib treatment results in the induction of p27KIP1 at both protein and mRNA levels in a time-dependent manner. The time course study indicates that the elevation of p27KIP1 protein amount coincides with an increase in p27KIP1 mRNA levels. Real-time RT-PCR results show that erlotinib treatment results in approximately 2.8- and 4.6-fold increase in p27KIP1 expression at 24 and 48 h, respectively. Next, we explored whether this effect could be caused by an activation of the p27KIP1 transcriptional promoter. H322 cells were transiently transfected with a cDNA construct containing luciferase reporter, controlled by promoter regions of human p27KIP1 (p27 Luc, from -3568 to -549) or with an empty cDNA vector as a control. As shown in Fig. 3B, erlotinib treatment led to a notable and time-dependent activation of p27KIP1 promoter as measured by luciferase activity. All results suggest that the induction of p27KIP1 by erlotinib may be due, at least in part, to the activation of p27KIP1 at the transcriptional level.
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Erlotinib Induces p27KIP1 Protein Stabilization. Aside from induction of p27KIP1 transcription, the increase in the intracellular amount of p27KIP1 could be caused by a reduction of p27KIP1 protein degradation via suppression of its phosphorylation at Thr187 or interaction with the SKP2-mediated ubiquitin/proteasome pathway (Tsvetkov et al., 1999). To test this possibility, we examined p27KIP1 phosphorylation at Thr187 and the expression of SKP2 in H322 cells after treatment with 2 µM erlotinib for the indicated times. The results shown in Fig. 4A demonstrate that erlotinib treatment caused a time-dependent reduction of p27KIP1 phosphorylation at Thr187 and decrease in SKP2 expression. It is interesting that the time point at which the reduction of p27KIP1 p-Thr187 and SKP2 occurred (12 h after erlotinib treatment) was tightly consistent with the accumulation of p27KIP1. To further test the hypothesis that the increased level of p27KIP1 protein in erlotinib-treated cells is caused by stabilization of p27KIP1 protein, we determined the effect of erlotinib on p27KIP1 stability in a pulse-chase experiment. The results as shown in Fig. 4B indicate that p27KIP1 protein was rapidly degraded with calculated half-life time (t
) of 1.8 h in control cells, whereas the p27KIP1 protein was stable with t of 6 h in erlotinib-treated cells. The accumulation of p27KIP1 protein in cells treated with erlotinib may be caused by the activation of p27KIP1 transcriptional level and/or increase in p27KIP1 stabilization.
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). Here, we sought to determine whether erlotinib-induced cell G1/S arrest could be involved in the alteration of p27KIP1 subcellular localization. First, we performed immunofluorescence staining experiments to observe the localization of p27KIP1 in H322 cells after 24 h of exposure to 2 µM erlotinib or with the same volume of medium contained 0.1% DMSO as a control. Figure 5A shows a representative example of immunofluorescence staining, which shows that erlotinib treatment leads to p27KIP1 localization in the nucleus (48 ± 10% of cells) compared with that in control cells (only 9 ± 7% of cells with p27KIP1 staining in the nucleus). Furthermore, we performed cell fractionation to determine the levels of p27KIP1 in cytosolic and nuclear fractions by immunoblot analysis. We consistently found that p27KIP1 was detected predominantly in the nuclear fraction and to a lesser extent in cytosolic fraction in erlotinib-treated cells, whereas low levels of p27KIP1 protein were detected in both cytosolic and nuclear fractions of untreated cells (Fig. 5B). These results suggest that the accumulation of p27KIP1 protein in the nucleus may be one event in the response to erlotinib-induced G1/S phase arrest and cell growth inhibition.
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Relationship between G1 Phase Arrest and Cell Sensitivity to Erlotinib. G1/S phase arrest may be a major contributor to erlotinib-induced cell growth inhibition in H322 cells. Thus, we needed to determine whether there was a relationship between cell cycle response and cell sensitivity to erlotinib. We chose two sensitive NSCLC cell lines (H322, H358), human skin epidermoid carcinoma A431 cells, and human head and neck carcinoma HN5 cells, which are known to be sensitive to erlotinib (Moyer et al., 1997; Pollack et al., 1999), and four resistant NSCLC cell lines (H460, A549, H596, and H1299 cells) as models and exposed them to erlotinib at 2 µM, a concentration that is close to clinically effective doses (Hidalgo et al., 2001). After 72 h of exposure, cell survival was assessed by MTT assay. The results shown in Fig. 7, A and B, indicate that 2 µM erlotinib causes >50% cell growth inhibition (P < 0.01) in all tested sensitive cell lines but had less effect in all tested resistant cell lines. A 24-h exposure to 2 µM erlotinib consistently resulted in significant increase in G1 phase cell accumulation in sensitive cell lines tested, but lesser extents of G1 cell accumulation was observed in resistant cell lines (Fig. 7, C and D). Moreover, immunoblot analysis revealed that whereas erlotinib treatment led to suppression of p-Rb, decrease in cyclin A expression, and induction of p27KIP1 protein in all tested sensitive cells, no noticeable effect was observed in resistant cell lines (Fig. 7, E and F). These results suggest that G1/S phase arrest, the up-regulation of p27KIP1, and the alteration in the expression of cell cycle regulators may be at least partly associated with NSCLC cell response to erlotinib.
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). In this work, we demonstrate that erlotinib strongly inhibits H322 cell growth and proliferation and blocks cell cycle progression at G1/S phase transition. To better understand the molecular mechanisms underlying erlotinib-induced cell growth inhibition and cell cycle arrest, we determined the effect of erlotinib on the expression of key regulatory proteins that are required for transition past the G1/S phase restriction point of cell cycle progression. Indeed, we found that erlotinib induced a time-dependent decrease in cyclin A and cyclin E expression and inhibition of CDK2 activity. Several reports have demonstrated the G1-phase arrest induced by inhibition of mitogen-activated signal pathways is accompanied by down-regulation of D-types of cyclins and by suppression of CDK4 activity (Baldin et al., 1993). In our work, we found that erlotinib treatment significantly suppressed CDK4 activity but did not markedly affect the levels of cyclin D1 and CDK4 in H322 cells. To certify our conclusions, we used two different epitope-mapping anti-cyclin D1 antibodies to determine cyclin D1 levels in lysates from erlotinib treatment and obtained similar results, indicating that the results should be believable. In addition, our preliminary results showed that erlotinib induced G1 phase arrest but did not markedly change cyclin D1 expression in H358 cells (data not shown). The reasons for cyclin D level being stable in erlotinib-induced G1 cells remain to be understood. One possibility may be that H322 cells have too high endogenous cyclin D1 to change its levels after erlotinib treatment. In addition, we suggest that erlotinib-induced suppression of CDK4 activity may be mediated via unidentified factors that could be required for the activation of CDK4. Recent reports have demonstrated that inhibition of cyclin A/CDK2 or cyclin E/CDK2 is implicated in the suppression of CDK4 activity (Koff et al., 1992). In this work, we demonstrated that erlotinib treatment causes a time-dependent decrease in the levels of total Rb protein and inhibition of Rb phosphorylation. In addition, erlotinib inhibits Rb phosphorylation at different serine residues, particularly at Ser795, which was more susceptible than that at Ser780, suggesting that erlotinib could act on Rb phosphorylation at specific sites, by which it may specifically affect Rb protein binding to a particular partner. In addition, recent reports have shown that the ERK1/2 pathway may be involved in EGF-induced rapid Rb protein phosphorylation at Ser780 and Ser795 (Guo et al., 2005). We examined whether erlotinib-induced inhibition of Rb phosphorylation at Ser795 or Ser780 could be associated with inhibition of ERK1/2 and/or PI3/AKT related pathways. Our preliminary data show that only erlotinib and U0126, an ERK1/2 inhibitor, but not LY294002, a PI3/AKT inhibitor, caused the suppression of Rb phosphorylation at Ser795 and Ser780, suggesting that erlotinib-induced Rb hypophosphorylation may be at least in part associated with the inhibition of ERK1/2-related pathways (data not shown).
It has been well-established that cyclin-CDK inhibitors CIP/KIP p21, p27, and INK4 p15, p16, and p18 as the negative controllers play important roles in the regulation of cell cycle progression. Recent studies have shown that p21WAF/CIP1 and p27KIP1 are necessary for the assembly of the cyclin A/CDK4 or CDK6 and the formation of cyclin A/CDK2 or cyclin E/CDK2 complexes. Thus, an increase in the expression of p21WAF/CIP1 and p27KIP1 could facilitate assembly of the complexes and result in suppression of the activity of cyclin/CDKs and thereby delay cell cycle progression (Slingerland and Pagano, 2000). In this work, we determined the effect of erlotinib on the expression of CDK inhibitors and found that erlotinib did not affect the expression of p16INN4a over experiment times. The endogenous levels of p21WAF/CIP1 in H322 cells were barely detectable, and erlotinib did not induce p21WAF/CIP1 expression. However, we found that p27KIP1 levels were markedly induced by erlotinib in a time-dependent manner. The increase in p27KIP1 levels tightly correlated with G1/S phase arrest. It has been known that expression of p27KIP1 is regulated at transcriptional and post-transcriptional levels in different types of cells (Besson et al., 2006). We found that the erlotinib-induced increase in p27KIP1 level was mediated through up-regulation of gene transcriptional activity and via inhibition of protein degradation. The results from quantitative real-time RT-PCR showed that an increase in p27 mRNA expression was tightly correlation with an increase in p27KIP1 protein in cells treated with erlotinib. We also provide evidence that the region from -3568 to -549 of the p27KIP1 promoter plays an important role in erlotinib-induced p27KIP1 gene transcription. Several reports have shown that the transcriptional regulation of p27KIP1 promoter activity seems to be complex and consists of both positive and negative regulatory elements. Multiple transcriptional factor binding sites within the p27KIP1 promoter have been characterized, including forkhead transcription factor (Dijkers et al., 2000), SP1 (Fischer et al., 2005), and E2F (Wang et al., 2005). Although our results showed that erlotinib caused the up-regulation of p27KIP1 expression, the mechanism by which erlotinib-induced promotion of p27KIP1 gene transcription remains to be further elucidated. Besides activation of gene expression, the increase in p27KIP1 level may depend on the post-translational regulation by the prevention of proteolytic degradation. Several studies have demonstrated that the ubiquitin-proteasome proteolysis system is a major pathway for the regulation of p27KIP1 levels (Boehm et al., 2002). It has been known that phosphorylation of p27KIP1 at Thr187 by CDK2 prepared p27KIP1 protein for binding to ubiquitin ligase SCF-SKP2 that leads to 26S proteasome degradation (Montagnoli et al., 1999). In our study, erlotinib treatment consistently resulted in a time-dependent reduction of p27KIP1 phosphorylation at Thr187 and decrease in SKP2 levels. Overall, our findings suggest that the G1/S phase arrest by erlotinib was at least in part associated with the accumulation of p27KIP1, which was regulated by the promotion of gene expression and decrease in protein degradation. Recent studies have shown that localization of p27KIP1 protein in the nucleus is required to inhibit CDK activation by CDK-activating kinase (Yaroslavskiy et al., 1999). In addition, p27KIP1 localization is essential for controlling the cell cycle progression and cell proliferation (Jiang et al., 2000). In this work, we have investigated the effect of erlotinib on p27KIP1 subcellular localization and found that the p27KIP1 protein was significantly accumulated in the nucleus in erlotinib-treated cells compared with its predominant localization in the cytoplasm in control cells, suggesting that the alteration of p27KIP1 localization may be involved in the cell cycle arrest at G1/S phase and inhibition of cell proliferation. Several reports have shown that phosphorylation of p27KIP1 is an important determinant of its subcellular localization. It has been evident that phosphorylation of p27KIP1 at Thr157 mediated by PKB/AKT results in retention of p27KIP1 in the cytoplasm and prevention of G1 arrest (Shin et al., 2002). Ser10 is another phosphorylation site of p27KIP1 for the nuclear export of the protein mediated by exportin (Viglietto et al., 2002). Recent studies have consistently showed that the inhibition of p27KIP1 phosphorylation at Thr157 by LY294002 causes p27KIP1 accumulation in the nucleus and cell growth inhibition (Shin et al., 2005). In addition, it has been reported that inhibition of cyclin E/CDK2 results in the retention of p27KIP1 in nucleus (Ishida et al., 2002). Therefore, we suggest that erlotinib-induced p27KIP1 accumulation in the nucleus may be caused by the suppression of activity of the EGFR-PKB/AKT axis and/or by the reduction of cyclin E/CDK2 as described above. It is interesting that knockdown of p27KIP1 expression by siRNA resulted in the attenuation of G1 phase arrest and significantly overrode the inhibition of cell growth in erlotinib-treated cells, indicating that the induction of p27KIP1 is essential for erlotinib-induced G1 arrest and cell growth inhibition. These results are consistent with studies by Le et al. (2003), who showed that reduction of p27KIP1 using p27 siRNA blocked anti-HER2 antibody-induced p27KIP1 up-regulation and G1 arrest in breast cancer cells. Finally, our results demonstrate a direct relationship between G1/S phase and cell sensitivity to erlotinib. These results are consistent with other reports, which showed that inhibition of erbB-2 pathways by AG1478 caused G1 phase arrest with accumulation of p27KIP1 and a decrease in the cyclin D1 in the human breast MCF-7/ErbB2-overexpressing cell line (Lenferink et al., 2001). Nahta et al. (2004) showed that down-regulation of p27KIP1 in breast cancer cell lines was associated with both an increase in cell cycle S-phase fraction and with cell resistance to trastuzumab, an anti-erbB-2/Neu monoclonal antibody. We found consistently that induction of p27KIP1 was necessary in response to erlotinib-induced cell growth inhibition and G1/S arrest, suggesting the induction and accumulation of p27KIP1 may be one of the important determinants in response to erlotinib-induced cell cycle blockade and cell growth inhibition.
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Footnotes |
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ABBREVIATIONS: CDK, cyclin-dependent kinase; BrdU, bromodeoxyuridine; CIP, cyclin inhibitory protein; KIP, kinase inhibitory protein; EGFR, epidermal growth factor receptor; FACS, fluorescence-activated cell sorting; NSCLC, non-small-cell lung cancer; PAGE, polyacrylamide gel electrophoresis; Rb, retinoblastoma; RT-PCR, reverse transcription-polymerase chain reaction; siRNA, small interfering RNA; PI3, phosphatidylinositol 3; DMSO, dimethyl sulfoxide; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium; PBS, phosphate-buffered saline; FITC, fluorescein isothiocyanate; DAPI, 4,6-diamidino-2-phenylindole; PKB, protein kinase B; ERK, extracellular signal-regulated kinase; LY294002, 2-(4-morpholinyl)-8-phenyl-1(4H)-benzopyran-4-one hydrochloride; U0126, 1,4-diamino-2,3-dicyano-1,4-bis(o-aminophenylmercapto)butadiene; AG1478, 4-(3'-chloroanilino)-6,7-dimethoxy-quinazoline.
Address correspondence to: Dr. Roman Perez-Soler, Department of Oncology, Montefiore Medical Center, 111 East 210th Street, Hofheimer 100, Bronx, New York 10467. E-mail: rperezso{at}montefiore.org
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