The Parkin gene (PARK2) is the causative gene of the autosomal recessive form of Parkinson’s disease (1-3). This gene encodes an E3 ubiquitin ligase that catalyzes the formation of a polyubiquitin chain on substrate proteins, which initiates protein degradation via the ubiquitin-proteasome system (4-6). PARK2 was mapped to the FRA6E region, a chromosomal fragile site frequently affected in the development of various cancers, and deletional mutations are the most common Parkin mutations (7-9). Since then, multiple lines of evidence have suggested that loss of PARK2 heterozygosity and copy number can be implicated not only in Parkinson’s disease but also in various cancers (10-12). Hence, the tumor-suppressive role of the Parkin protein (Parkin) has rapidly gained attention.
In the past two decades, studies directed at understanding the mechanisms underlying Parkin‑mediated tumor suppression revealed that the tumor-suppressive effect of Parkin largely depends on the promotion of apoptosis and/or inhibition of cell cycle progression (13, 14). Parkin triggers the poly-ubiquitination and degradation of the anti-apoptotic molecule myeloid cell leukemia-1, thereby promoting the apoptosis of human cervical cancer cells (13). Parkin also ubiquitinates phosphoglycerate dehydrogenase (PHGDH), the first rate-limiting enzyme in the serine synthesis pathway, which is critical for the excessive proliferation of cancer cells, leading to the degradation of PHGDH to prevent tumor development in human Hs578T breast cancer and H1299 lung cancer cells (15). Another study showed that Parkin binds to microtubules and facilitates the interaction between anticancer chemicals and microtubules, increasing the sensitivity of HeLa cells to anticancer chemicals (16, 17). In addition, Parkin overexpression induces caspase-dependent apoptosis and cell cycle arrest at the G1 phase (10) in non-small cell lung cancer (NSCLC) cell lines. Similarly, Parkin overexpression impedes the proliferation of colorectal cancer cells, HCT116 and DLD1, by suppressing the entry of cells into the S phase (18). The doubling time of Parkin-overexpressing cells dramatically increases by two- to threefold (18). These studies show that Parkin suppresses tumor development via various molecular pathways.
The transcription factor p53 is a crucial molecule that is highly induced by diverse stresses, including DNA damage, which causes cell cycle arrest to maintain genomic integrity during cell division (19-21). Upon DNA damage, p53 becomes activated and elicits the transcriptional regulation of several cell cycle regulatory proteins (22). Among these proteins, p21 is the most well-studied transcriptional target of p53. It contributes to cell cycle arrest (23) by acting as a CDK (Cyclin-dependent kinase) inhibitor (24). Cyclin B1 is another important molecule regulated by the p53 cascade in response to DNA damage (25). The CDK1/cyclin B complex is responsible for G2/M phase transition (26). In response to DNA damage, activated p53 induces the transcriptional downregulation of cyclin B, which consequently inhibits the activity of the CDK1/cyclin B complex and results in cell cycle arrest at the G2/M phase (27).
In the present study, we aimed to examine the tumor-suppressive role of Parkin in lung and colorectal cancer cells.
Parkin acts as a tumor suppressor in various cancer cells, including lung cancer cells (11). The large-scale datasets from cBioportal revealed that PARK2 alterations can be observed in 4.83% out of 3,622 lung adenocarcinoma cases (28, 29). However, the mechanisms underlying the tumor-suppressive role of Parkin in lung cancer cells remain to be elucidated. In the present study, we examined whether Parkin affects the viability of A549 lung cancer cells. Parkin was overexpressed in A549 lung cancer cells, and the number of viable cells was counted. Parkin overexpression reduced the number of viable cells in a dose- and time-dependent manner (Fig. 1A, B). In addition, the Parkin-induced reduction in cell viability was partly rescued when Parkin overexpression was partially reduced by the introduction of Parkin-specific small-interfering RNA (siRNA, Fig. 1C). These results imply that Parkin can suppress the viability of lung cancer cells. To elucidate the mechanisms by which Parkin reduces cell viability, we investigated whether this protein promotes the apoptosis and/or suppresses the proliferation of lung cancer cells. As shown in Fig. 1D, no significant cleavage of PARP, a hallmark of apoptosis, was observed when Parkin was overexpressed in a dose-dependent manner. However, flow cytometry results showed Parkin overexpression induced G2/M cell cycle arrest in the A549 lung cancer cells in a dose- and time-dependent manner (Fig. 1E-H). Taken together, these results suggest that Parkin reduces the viability of A549 cells by decreasing their proliferation and not by promoting their apoptosis.
G2/M cell cycle arrest results from many causes, including DNA damage (30). A recent study has shown that ionizing radiation induces the upregulation of Parkin in lung cancer cells (31). In the present study, we investigated whether Parkin expression causes DNA damage. Parkin was overexpressed in A549 cells, and a comet assay was performed. Comet tails, which consist of broken DNA strands indicating DNA damage, were clearly observed in the Parkin-overexpressing group, whereas no comet tails were observed in the control group (Fig. 2A). In addition, the phosphorylation of ATM, one of the molecules associated with DNA damage response in cells, was increased in the Parkin-overexpressing cells in a time-dependent manner (Fig. 2B). We also found that the Parkin-induced reduction in cell viability was partially recovered by ATM inhibition (Fig. 2C). p53 is involved in DNA damage response (32) and phosphorylated in response to DNA damage, regulating various downstream molecules to facilitate cell cycle arrest (33). Therefore, we explored whether Parkin expression influences p53 expression. We found that Parkin expression increased the protein expression and phosphorylation of p53 in a dose-dependent manner (Fig. 2D). Moreover, the Parkin-induced increase in the amount and phosphorylation of p53 was partly restored when Parkin overexpression was partially reduced after introducing Parkin-specific siRNA (Fig. 2E). The Parkin-induced decrease in cell viability was partly recovered when p53 was partially downregulated after introducing p53-specific siRNA, indicating that the Parkin-induced reduction in cell viability was mediated by p53 (Fig. 2F). We also found that ATM inhibition reversed Parkin-induced p53 activation in an ATM inhibitor dose-dependent manner (Fig. 2G), implying that ATM was an upstream molecule of p53. Collectively, these results suggest that Parkin induces DNA damage and ATM phosphorylation, which consequently activate p53 and decrease the viability of lung cancer cells. Previous studies have demonstrated the protective role of Parkin on DNA damage. After exposure to various DNA-damaging agents, Parkin translocates to the nucleus to promote DNA repair and reduce DNA damage in SH-SY5Y cells (34). Similarly, Parkin protects cells against reactive oxygen species-induced mitochondrial DNA damage by binding to DNA to facilitate DNA repair in SH-SY5Y cells (35). However, whether Parkin itself induces DNA damage remains unclear. In the present study, we demonstrated that Parkin induced DNA damage in the lung and colorectal cancer cells. Parkin seems to function differently depending on the proliferative capacity of cells. Parkin may decrease DNA damage in fully differentiated cells that are incapable of proliferation, such as neuronal cells. By contrast, Parkin may cause DNA damage in proliferating cells, such as lung and colorectal cells.
The DNA damage-induced activation of p53 leads to the transcriptional regulation of genes associated with the cell cycle, such as p21, a cyclin-dependent kinase inhibitor (36). In the current study, we examined whether Parkin regulates p21 expression. We found that Parkin induced the upregulation of p21 at the transcriptional and translational levels in a dose-dependent manner (Fig. 3A, B). Cyclins are important molecules involved in cell cycle regulation, and decreased cyclin A2 and/or cyclin B1 levels cause cell cycle arrest at the G2/M phase in several cancer cell models (37-39). Therefore, we examined whether Parkin regulates the expression of cyclin A2 and cyclin B1. In our A549 cell model, Parkin expression elicited the transcriptional and translational downregulation of cyclin B1 but not cyclin A2 (Fig. 3C, D). The Parkin-induced upregulation of p21 and downregulation of cyclin B1 were restored when the Parkin-induced activation of p53 was reversed after introducing p53-specific siRNA (Fig. 3E, F). This result indicates that the Parkin-induced regulation of p21 and cyclin B1 is mediated by p53. A previous report showed that Parkin overexpression induces cell cycle arrest at the G1 phase in H1299 and H460 NSCLC cell lines (10). This study suggested that the G1 arrest is mediated by the upregulation of p21 and p18, potent inhibitors of the G1/S cell cycle transition, and downregulation of cyclin D1/CDK4, an inducer of cell cycle progression from the G1 to S phase. Further studies are warranted to elucidate the mechanisms by which Parkin influences the proliferation of lung cancer cells, which may vary depending on the cell type.
Parkin overexpression impedes cell proliferation in colorectal cancer cells, and the large-scale datasets from cBioportal show PARK2 alterations in 7.83% out of 1,482 colorectal cancer cases (28, 29). To investigate whether the Parkin-associated anti-proliferative mechanism elucidated above is limited to lung cancer cells, we examined the anti-proliferative effect of Parkin in HCT-15 human colorectal cancer cells. We observed that Parkin overexpression dramatically reduced the viability of HCT-15 cells (Fig. 4A). In accordance with the results obtained in A549 lung cancer cells, Parkin expression induced G2/M cell cycle arrest in HCT-15 colorectal cancer cells (Fig. 4B). In addition, Parkin expression induced the upregulation of p53 and p21 and downregulation of cyclin B1 in colorectal cancer cells, similar to that in lung cancer cells (Fig. 4C). These results suggest that the suppressive effects of Parkin on cancer cell proliferation are not limited to lung cancer cells.
Parkin is a tumor suppressor that has been actively investigated. However, the mechanisms by which Parkin prevents tumorigenesis are not completely understood. In this study, we explored the mechanisms underlying Parkin-mediated tumor suppression in lung and colorectal cancer cells. Our findings clearly showed that 1) Parkin expression induces DNA damage, 2) Parkin-mediated DNA damage elicits the ATM-dependent activation of p53, and 3) activated p53 induces the upregulation of p21 and downregulation of cyclin B1, which consequently halt the cell cycle at the G2/M phase and inhibit cancer cell proliferation (Supplementary Fig. 1). In conclusion, we report the tumor-suppressive role of Parkin in lung and colorectal cancer cells. The findings of this study highlight the potential of Parkin as a target molecule for gene therapy to manipulate cancers derived from the loss of PARK2 heterozygosity.
siRNA transfection was performed as previously described (40). Briefly, cells were seeded in a six-well plate at a density of 2 × 105 cells/well. After 24 h, the cells were infected with the mock or Parkin virus and treated with siRNA simultaneously and then incubated for another 48 h. Prior to transfection, siRNAs (200 pmol) were mixed with liposomes in serum-free RPMI 1640 medium and then incubated for 30 min at room temperature. The mixture and the virus were added to the cells. After 4 h incubation, the cells were supplemented with fresh RPMI 1640 medium containing FBS (to set a final concentration of 10%) and then incubated at 37°C in a CO2 incubator for a given time. The siRNA sequences were as follows: siRNA specific for Parkin, sense 5’-GCA CCU GAU CGC AAC AAA UTT-3’ and antisense 5’-AUU UGU UGC GAU CAG GUG CTT-3’; p53, sense 5’-GCG UGU GGA GUA UUU GGA UTT-3’ and antisense 5’-AUC CAA AUA CUC CAC ACG CTT-3’; non-specific siRNA, sense 5’-AUG AAC GUG AAU UGC UCA ATT-3’ and antisense 5’-UUG AGC AAU UCA CGU UCA UTT-3’. Non-specific siRNA was used as a universal control.
The comet assay was performed as previously described (41). Briefly, cells were centrifuged, washed with ice-cold PBS, resuspended in a 0.5% (w/v) solution of low temperature-melting agarose in PBS at 37°C, and then layered onto Comet slides (Trevigen, Gaithersburg, USA). The agarose-cell mixture was incubated at 4°C for 30 min and then placed in a lysis buffer for 30 min at 4°C in the dark. Subsequently, the slides were immersed in an alkaline unwinding solution. Electrophoresis was performed in an alkaline buffer at 1 V/cm for 25 min at 4°C, and the gel was washed twice in distilled water. After washing, the slides were dried at 45°C for 20 min and then stained with SYBR Green. Images were captured using a laser confocal scanning microscope (LSM 710; Zeiss, Heidenheim, Germany).
The authors have no conflicting interests.