The human renal carcinoma cell line, OS-RC-2, was obtained from the Type Culture Collection of the Chinese Academy of Sciences (Shanghai, China). The cells were cultured under 5% CO₂ at 37°C in RPMI-1640 medium with 5% FBS and 1% penicillin/streptomycin (all from HyClone, Auckland, New Zealand).

Target gene accession numbers NM_033437 and NM_003109 were obtained from GenBank. According to the principles of Ambion for the design of siRNAs, the sequences of 21 nucleotides beginning with AA and containing 30–50% GC were selected and used as target sites. The siRNA used were as follows: PDE5A siRNA, 5′-GGAAGAAACAAG AGAGCUAdTdT-3′ (sense) and 5′-UAGCUCUCUUGUUUCU UCCdTdT-3′ (antisense); Sp1 siRNA, 5′-CAUCCAAGGCUGU GGGAAAdTdT-3′ (sense) and 5′-UUUCCCACAGCCUU GGAUG dTdT-3′ (antisense). The control and negative control sequences were provide by the company, and a report was sent to ensure the effects. No homology sequence was found by Blast analysis. The siRNAs were purchased from Wuhan Genesil Biotechnology Co., Ltd. (Wuhan, China), and 10 nmol/l of each siRNA were transfected into the OS-RC-2 cells, using HiPerFect transfection reagent (Qiagen, Valencia, CA, USA).

The activity of caspase-3 was determined using the caspase-3 activity kit (Beyotime Institute of Biotechnology). Cell lysates were prepared after their respective treatment. Subsequently, the OS-RC-2 cells were scraped, centrifuged, resuspended and lysed in lysis buffer (RIPA buffer and protease inhibitor PMSF; Solarbio, Beijing, China). Assays were performed in 96-well microtiter plates by incubating 10 μl protein of cell lysate per sample in 80 μl reaction buffer (1% NP-40, 20 mM Tris-HCl, pH 7.5, 137 mM Nad and 10% glycerol) containing 10 μl/mM caspase-3 substrate (Ac-DEVD-pNA). The lysates were incubated at 37°C for 4 h. Samples were measured using a microplate reader (Bio-Rad Laboratories, Inc., Hercules, CA, USA) at an absorbance of 405 nm, according to the manufacturer’s instructions. Cell proliferation was determined by MTT assay according to the instructions provided with the MTT cell proliferation assay kit (Beyotime Institute of Biotechnology). The absorbance was measured at 570 nm on a microplate reader (Bio-Rad Laboratories, Inc.).

Total RNA was extracted using TRIzol reagent (Invitrogen, Carlsbad, CA, USA), reverse-transcribed into cDNA templates and amplified using a ReverTra Ace qPCR RT kit (Toyobo, Osaka, Japan). RT-qPCR was performed using a SYBR-Green Real-time PCR Master Mix-Plus (Toyobo). The PCR primers (Table I) were used for gene amplification. Target gene expression levels are presented as a ratio of the levels in the treated cells and the control cells using the ΔΔCt method, as previously described (13).

The OS-RC-2 cells were collected using the rubber policeman, washed in phosphate-buffered saline (PBS) and lysed in ice-cold PBS buffer [0.l M Tris-HCl, pH 8.0, 100 mM NaCl, 1 mM EDTA, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% sodium dodecylsulfate (SDS), and 0.1% protease inhibitor cocktail and 10 mM phenylmethylsulphonyl fluoride (PMSF)] for 30 min on ice. The lysates were sonicated for 2 sec to shear the DNA to reduce its viscosity, followed by centrifugation at 28,100 × g for 30 min at 4°C. The supernatant was collected, and protein concentrations were measured using the Bradford method (Bio-Rad Laboratories, Inc.). Twenty micrograms of protein determined by Bradford assay were electrophoretically separated using a 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gel and transferred onto polyvinylidene fluoride (PVDF) membranes, and then immunoblotted with the corresponding antibodies. Immuno-detection was performed with an enhanced chemiluminescence (ECL) detection kit (Beyotime Institute of Biotechnology). The protein bands were analyzed using Quantity One 4.4 software.

The wild-type and mutant constructs of PKG, including PKG Iα and PKG Iβ were designed and purchased from BioTech Co., Ltd. The PKG constructs were subcloned into the expression vector pMIG upstream of the IRES-enhanced green fluorescent protein (GFP) sequence. The cells were transfected with pMIG as previously described (14).

All data are presented as the means ± standard deviation (SD). Significant differences among the groups were determined using the unpaired Student’s t-test. A value of P<0.05 was accepted as an indication of statistical significance. All data in the figures were obtained from at least 3 independent experiments.

We detected the mRNA and protein expression of PDE5 in the OS-RC-2 cells, and then used specific siRNA to silence PDE5 expression. Following transfection with PDE5 siRNA, real-time PCR revealed that the level of PDE5 mRNA expression was significantly reduced in the transfected cells when compared with the negative control (Fig. 1A). Western blot analysis also revealed a significant decrease in the levels of the 105 kDa PDE5 in the OS-RC-2 cells transfected with PDE5 siRNA, when compared with the control (Fig. 1B and C). Thus, our data indicated that treatment with PDE5 siRNA successfully suppressed the expression of PDE5 in the OS-RC-2 cells.

PDE5 siRNA-transfected OS-RC-2 cells had much lower growth rates than the control and negative control cells. The average rate of growth inhibition was approximately 50% in the PDE5 siRNA-transfected cells. When 8-Br-cGMP, a cell membrane permeable cGMP derivative was added, a further inhibition was observed. However, the inhibitory effect in the PDE5 siRNA-transfected OS-RC-2 cells was blocked by KT5823, an inhibitor of PKG (Fig. 2A). Conversely, there was a 2-fold increase in the relative level of caspase-3 in the transfected cells compared to the control cells. Caspase-3 is an early indicator of apoptosis. Caspase-3 activity was enhanced 3-fold following treatment with 8-Br-cGMP and was inhibited following treatment with KT5823 (Fig. 2B).

To elucidate the mechanisms responsible for the reduced proliferation and increased apoptosis in the PDE5 siRNA-transfected cells, the levels of cell cycle-specific molecules, including cyclins (cyclin A, B and D1) and CDK inhibitors (CKIs, p21 and p27) were determined. Our results revealed that no changes in the levels of cyclin A and B1 were detected in the PDE5 siRNA-transfected cells; however, a 50% reduction in the levels of cyclin D1 was observed in these cells. Further study indicated that both the mRNA and protein expression of cyclin D1 was decreased in a time-dependent manner in these cells. A 2- to 3-fold increase in the expression of p21, a mitotic inhibitor, was observed in the PDE5 siRNA-transfected cells when compared to the control cells. However, there were no changes observed in p27 expression, another mitotic inhibitor involved in the cyclin D1-CDK2, CDK4 and cyclin E-CDK2 inhibition at the G1 phase (Fig. 3). Therefore, cyclin D1 and p21 may play an important role in the inhibition of proliferation and the increase in apoptosis.

To explore whether the time-dependent decrease in the expression of cyclin D1 involves the regulation of β-catenin in this study, we examined the mRNA and protein levels of β-catenin. Of note, the protein levels of β-catenin also showed a time-dependent reduction following transfection. However, no changes in the mRNA level of β-catenin were detected (Fig. 4). This may partly be explained by the changes which occur in the expression of cyclin D1, as previously described (15).

Our results also indicate that a decrease in β-catenin expression is caused by an effect at the protein level rather than at the mRNA level in the activated PKG pathway. Furthermore, a significant increase in the phosphorylation of β-catenin was detected in the PDE5 siRNA-transfected cells, which was also consistent with previous data indicating that PKG has a similar role to GSK3β and directly phosphorylates β-catenin/TCF, leading to proteasomal degradation (16). MG132, an ubiquitin-proteasome inhibitor, was used to determine whether the decrease in β-catenin expression induced by PKG activation could be blocked at the proteasomal level. Our results revealed that MG132 prevented the reduction in β-catenin expression which was caused by activated PKG, and the inhibitor alone did not alter the level of β-catenin. These results indicated that the protein was processed by the ubiquitin-proteasome system (Fig. 5A). An increased expression of cyclin D1 levels was also detected compared to the control cells, after MG132 was used to inhibit the ubiquitin-proteasome system (Fig. 6A).

A previous study indicated that JNK activation was required for the PKG-dependent regulation of TCF activity (17). Therefore, in this study, we detected the level of phosphorylation of JNK and found that there was a significant increase in the phosphorylation of JNK in the transfected cells (Fig. 6B). This result is consistent with that of a previous study showing that the activation of PKG directly phosphorylates JNK (18). In addition, there was an increase in the level of cyclin D1 and no change in the levels of p-β-catenin following treatment with an inhibitor of JNK, indicating that activated JNK may affect the downstream molecules of β-catenin/TCF (Fig. 6A).

To examine whether the activation of PKG alters the transcription of p21, we treated the OS-RC-2 cells with PDE5 siRNA or 8-Br-cGMP. Our results revealed that the mRNA levels of p21 were increased in the cells treated with PDE5 siRNA or 8-Br-cGMP; however, the increase was not time-dependent (data not shown). Previous studies have confirmed that Sp1 is required for the PKG-induced upregulation of p21, p27 and HINT1 in SW480 cells, and can be phosphorylated by activating PKG on serine residues (11,12). To further investigate the requirement of Sp1 in the increase in p21 expression induced by PKG in OS-RC-2 cells, we knocked down the expression of endogenous Sp1 by transfecting the cells with Sp1 siRNA. The level of Sp1 in the OS-RC-2 cells was markedly reduced by Sp1 siRNA. The activation of p21 protein expression was clearly observed in the control group following treatment with 8-Br-cGMP, and was inhibited by the suppression of Sp1 (Fig. 7). Thus, our data indicate that the increased expression of p21 may be regulated through Sp1 by activating PKG.

PKG has various isozymes, including PKG I and PKG II; the N terminus of PKG I is encoded by 2 alternatively spliced exons that produce the isoforms α and β (19). In this study, we investigate which type of PKG is responsible for the anticancer activities in OS-RC-2 cells. The mRNA expression of PKG I and PKG II was detected in the cells treated with PDE5 siRNA. The results revealed that the mRNA expression of PKG Iα and PKG Iβ was significantly increased. The same result was also observed in the nitric oxide (NO) donor SNAP group, which indicated that the NO-cGMP pathway was involved in the anticancer activities induced by the activation of PKG. Unexpectedly, there were no changes observed in the PKG II levels in the cells treated with PDE5 siRNA of SNAP when compared with the control (Fig. 8A). These results suggested that not PKG II, but PKG I was involved in the anticancer activities. Next, we investigated the effects of PKG Iβ overexpression on cell proliferation and apoptosis. The results revealed that the cells with an overexpression of PKG had significantly increased growth rates, whereas this increase was abolished by treatment with the mutants (Fig. 8B). Relative caspase-3 expression was significantly increased in the cells with PKG Iβ overexpression, but there was no obvious change in the cells with PKG Iα overexpression (Fig. 8C). However, the role of PKG I was according to its intact structure as no changes in cell proliferation and apoptosis were observed in the cells overexpressing mutant PKG Iα and Iβ (Fig. 8B and C). Futhermore, immunoblot analysis revealed that the protein expression of β-catenin was reduced and the protein expression of p21 was increased in the PKG Iβ group (Fig. 8D). These results suggest that PKG Iβ plays a vital role in anticancer activities in OS-RC-2 cells.