To generate Ras_G12D-transformed cell populations of mouse embryonic fibroblasts (MEFs), we first infected primary MEFs from wild type (WT) and PINK1-deficient mice with a retrovirus expressing simian virus-40 large T antigen (SV40LT), which led to the emergence of continuously growing (immortalized) cells. Subsequently, immortalized MEFs were infected twice with a retrovirus expressing human K-Ras_G12D to generate Ras_G12D-transformed cells, which were used to derive single clones of Ras_G12D-transformed MEFs by limiting dilution. We selected two single clones with similar SV40LT expression levels for each genotype ( Figure S1A ). PINK1 WT clones 1 and 6 and PINK1-deficient clones 2 and 5 expressed comparable levels of Ras protein ( Figure 1A ). These four Ras_G12D-transformed clones were therefore selected for growth comparison in soft agar, which showed that the average colony area for PINK1^-/- clones was about 5-fold smaller than that of the WT control clones (18.8% of WT). No difference in soft agar growth was observed between WT and PINK1-deficient MEFs that only expressed SV40LT ( Figure 1B ). We next measured soft agar growth with WT and PINK1^-/- Ras_G12D-transformed cell populations and also generated a cell population (PINK1^-/-plus huPINK1) in which human PINK1 was re-expressed in mouse PINK1-deficient Ras_G12D-transformed cells. Characterization of protein expression showed that Ras protein levels in PINK1^-/-, WT and PINK1^-/-plus huPINK1 population cells are similar ( Figure 1C ). We also examined the rate of glucose consumption in WT and PINK1^-/- Ras_G12D-transformed cells, and the results showed that MEFs from PINK1^-/- Ras_G12D-transformed had reduced uptake of glucose ( Figure S1C ). The average area of PINK1-deficient colonies reached only 13% of that of WT colonies ( Figure 1D ). Re-expression of human PINK1 after lentiviral infection substantially rescued the growth deficit, increasing the average colony area to 74% of WT ( Figure 1D ). We did not detect PINK1 by western blot in WT Ras_G12D-transformed MEFs ( Figure 1E ), possibly because most of PINK1 is constitutively degraded in normal cells and PINK1 only becomes stabilized upon significant mitochondrial depolarization (11, 32, 33). As expected, PINK1 was absent from PINK1-deficient Ras_G12D-transformed cells that were derived from MEFs of mice with a null mutation in PARK6 (34). In contrast, both 62 kDa and 52 kDa PINK1 species were readily detected in PINK1^-/-plus huPINK1 cells ( Figure 1E ), suggesting that human PINK1 was imported to the inner mitochondrial membrane and cleaved by PARL, but that lentiviral over-expression of PINK1 may have saturated its mitochondrial import/cleavage and proteasomal degradation (32, 33).

In nude mice, we injected the same number of MEFs from these two different genotypes (PINK1^-/- Ras_G12D and PINK1^+/+ Ras_G12D). The tumor formation of PINK1^-/- cells was significantly reduced when compared with PINK1^+/+ cells in nude mice. PINK1^-/- tumors showed a trend of slower growth at each measurement, and the WT tumors were on average 4.6-fold larger (range: 3.6-5.6) than the PINK1^-/- tumors between days 11-17. On day 17, mice with WT tumors were euthanized due to excessive tumor burden, while PINK1^-/- tumors were allowed to grow longer until day 24, when they reached similar volumes as the WT tumors had reached on day 17 ( Figure 1F and Figure S1B ). In agreement with soft agar results, these experiments show that PINK1 deficiency slows the growth of Ras_G12D-induced tumors in nude mice. Overexpression of PINK1 in the PINK1^-/-plus huPINK1 population increased tumor growth by day 19 compared to PINK1^-/- Ras_G12D-transformed cells ( Figure 1G ), although the rescue effect was less pronounced than in soft agar (mean tumor volume on day 19: WT = 1413 mm³; PINK1^-/-= 700 mm³; PINK1^-/-plus huPINK1 = 911 mm³).

We further asked whether the reduced growth of PINK1-deficient tumors is due to increased cell death or decreased cell cycle progression, as both are hallmarks of cancer and involved in Ras-induced tumorigenesis. Cell death was measured with Annexin V/propidium iodide (PI) staining assays in RasG12D-transformed MEFs with different genetic backgrounds. As shown, PINK1 loss increased the percentage of necrotic but not apoptotic cells, which was fully reversed by re-expression of human PINK1 ( Figures 2A, B ). Consistent with the function of PINK1 in maintaining mitochondrial integrity and its roles in regulating cell death, PINK1^-/- cells displayed reduced membrane potential (Δψm) ( Figure S2A ). There was a positive correlation between Δψm and mean colony area in soft agar ( Figure S2B ). In addition, total ROS and mitochondrial ROS were reduced in PINK1^-/- cells. Recovery PINK1 expression at least partially restored the reduced ROS in PINK1^-/- cells ( Figures S2D, E ). Furthermore, basal mitochondrial respiration was reduced in PINK1^-/- RasG12D-transformed MEFs as revealed by Seahorse analysis ( Figure S2C ). These data suggest that mitochondrial metabolism is changed by PINK1.

Cell cycle analysis of PI-stained cells by FACS demonstrated that PINK1 deficiency led to an increase of Ras_G12D-transformed cells in the G2/M phase and decrease of cells in the G0/G1 phase. Overexpression of PINK1 had the opposite effect, reducing the percentage of cells in G2/M and increasing the proportion of cells in G0/G1 ( Figure 2C ). In line with these results, protein levels of p16 and p21 were increased in PINK1^-/- cells by Western blot analysis ( Figures 2D, E ). In addition, p53 protein expression was increased in PINK1^-/- Ras_G12D-transformed cells ( Figures 2D, E ). These data suggest that loss of PINK1 promotes cell death and cell cycle arrest.

To study the mechanisms underlying PINK1-regualted tumor growth in vivo, we examined cell proliferation rates in tumor tissues derived from WT, PINK1^-/- and PINK1^-/-plus huPINK1 cells in nude mice. Ras_G12D protein levels were comparable in the various tumor populations ( Figure 3A ). In contrast, expression of the cell proliferation marker Ki67 in PINK1^-/- tumors was reduced compared to WT, while expression of Ki67 in PINK1^-/-plus huPINK1 tumors that over-expressed huPINK1 was increased, indicating reversible inhibition of tumor cell proliferation in vivo by ablation of PINK1 ( Figure 3B ). Thus, the reduced growth of PINK1-deficient Ras_G12D-transformed cells in vitro is validated and supported by reduced proliferation of the corresponding tumor cells in nude mice.

Using the caspase 3/7 apoptosis detection kit, we also observed increased apoptosis of PINK1^-/- tumor cells in nude mice ( Figure 3C ). Expression levels of p16, p21 and p53 were significantly increased in PINK1^-/- tumor tissues, which was reversed after re-expression of human PINK1. However, unlike in MEFs, p21 was not downregulated significantly after re-expression of human PINK1 ( Figure 3D ).

We next explored possible mitochondrial mechanisms underlying the reduced growth of PINK1-deficient Ras_G12D-transformed cells. As indicated above, PINK1 deletion did not influence Ras expression levels ( Figure 1C ). Several studies have shown that PINK1 acts as a pro-fission factor (14, 35–38), consistent with mitochondrial fission preceding mitophagy (36, 39) and the enlarged mitochondrial morphology in various cells of PINK1-deficient mice (40). To analyze whether loss of PINK1 altered mitochondrial morphology, we stained mitochondria in MEFs by immunocytochemistry with an antibody against LRP130. LRP130 is a suitable marker because it is exclusively expressed in the mitochondrial matrix where it regulates the assembly and activity of cytochrome c oxidase (complex IV) (41). The mitochondrial network was highly fragmented in the majority (74.5%) of WT Ras_G12D-transformed cells ( Figure 4A ). In contrast, only 32% of PINK1^-/- Ras_G12D-transformed cells showed a fragmented mitochondrial network, while 59.4% showed an intermediate network and 8.6% showed an elongated/fused network ( Figure 4A ). Re-expression of PINK1 restored mitochondrial fragmentation to 69.6% of the cells, similar to that in WT cells ( Figure 4A ). Thus, loss of PINK1 interfered with mitochondrial dynamics in Ras_G12D-transformed MEFs, which could be reversed by re-expression of PINK1.

Because it has been shown that PINK1 promotes mitochondrial fission by phosphorylating Drp1 at Ser616 (42), we studied whether altered mitochondrial dynamics in PINK1^-/- Ras_G12D- MEFs was related to abnormal phosphorylation or expression of Drp1. To this end, the expression of Drp1 and phosphorylated Drp1 was detected by western blotting and flow cytometry. There was no significant difference in Drp1 protein expression. PINK1^-/- MEFs showed a significant decrease of phospho-Drp1 (Ser616) levels compared to WT MEFs. Re-expression of huPINK1 in PINK1^-/- cells partially rescued the decreased phospho-Drp1 (Ser616) levels ( Figures 4B, C ). Analysis of the expression of Drp1 and phosphorylated Drp1(Ser616) showed that phosphorylation of DRP1 was also decreased in PINK1^-/- tumor tissues ( Figure 4D ). It has been also shown that ERK can increase Drp1 phosphorylation (43). However, our data did not show a reliable link between ERK phosphorylation (activation) and PINK1 expression ( Figure S2G ), suggesting that decreased phosphorylation at Drp1 (Ser616) may be due to a direct effect of PINK1 loss. In contrast to the phosphorylation at Drp1(ser616), phosphorylation at Drp1(Ser637) inhibits mitochondria fission and modulate mitophagy (44). Interestingly, PINK1 deficiency did not cause alterations in Drp1(Ser637) phosphorylation ( Figure S2F ). In line with above data, although PINK1 deficiency increased p62 and LC3-1/II, the recovery of PINK1 expression had no ability to abolish it ( Figure S2H ), suggesting that the increased p62 and LC3-1/II is not intrinsic effect of PINK1. Therefore, PINK1 deficiency regulate mitochondria morphology mainly by reducing Drp1(Ser616) phosphorylation-mediated mitochondria fission.

Drp1 affects mitochondrial fission, leading to cell cycle arrest and apoptosis (45–47), which prompted us to explore the impact of PINK1 on cell growth in the presence of the mitochondrial fission (Drp1) inhibitor Mdivi-1. Mdivi-1 significantly inhibited the growth of Ras_G12D-transformed WT cells, while the inhibitory effect of Mdivi-1 was compromised in PINK1-deficient cells. Re-expression of PINK1 in PINK1^-/- Ras_G12D MEFs restored Mdivi-1-mediated growth inhibition ( Figure S3A ). Therefore, PINK1 loss inhibits tumor growth at least in part through impaired regulation of Drp1. Together with data presented above, these results demonstrate a pro-fission and pro-growth function of the PINK1-Drp1 axis in Ras_G12D-transformed MEFs.

In MEFs, Ras_G12D was overexpressed after viral transduction. To study whether PINK1 loss similarly affected the growth of authentic human Ras_G13D-transformed tumor cells in which mutated Ras is expressed at physiological levels, we introduced CRISPR-Cas9-mediated deletions into the PINK1 gene of HCT116 cells that were originally isolated from a primary human colon carcinoma. Several PINK1-deficient HCT116 clones with varying PINK1 deletions were identified by PCR with genomic DNA using primers and DNA sequencing that flanked the two CRISPR target sites ( Figures S3B, C ). We did not detect PINK1 protein expression by Western blots in WT HCT116 cells, likely due to constitutive degradation in cells without severe mitochondrial depolarization (32). However, in addition to sequencing, we confirmed that the CRISPR/Cas9-generated genomic PINK1 deletions produced null mutations at the mRNA level by PCR amplification of total cellular cDNA with two different primers pairs ( Figure S4A ). PINK1-knockout clones 45 and 51 produced the expected PCR fragments (deletions), which were unable to encode any functional PINK1 protein. In addition, clones 3, 11 and 50 yielded no bands or only very faint PCR products, suggesting that in these clones the genomic deletions destabilized the resulting PINK1 mRNAs to levels below detection. Collectively, these results show that all CRISPR/Cas9-generated PINK1 deletions produced null mutations.

HCT116 clones lacking PINK1 showed reduced growth in soft agar when compared to the control clones, which were transfected with px458 vector only ( Figure 5A ). This was not due to altered Ras expression, because PINK1-deficient and control vector-transfected HCT116 clones expressed comparable levels of Ras ( Figure 5B ). Deletion of PINK1 also did not affect the GTPase activity of Ras ( Figure S4B ). Like in MEFs, we detected Drp1 and phosphorylated Drp1(Ser616) protein in HCT116 cells by western blotting and flow cytometry and found a decrease in phosphorylated Drp1(Ser616) in PINK1 knockout HCT116 cells ( Figures 5C, D ). In tissue culture, growth of PINK1 knockout HCT116 cells was significantly slower than that of the control group ( Figure 5E ). Overall, these results show that in human tumor cells with a common Ras mutation, the loss of PINK1 also causes impaired tumor growth.

Similar to the results reported in other studies that most of the mitochondria were fragmented in cell lines containing Ras mutations (48), mitochondrial fragmentation was also observed in HCT116 vector-transfected cells. In contrast, HCT116 PINK1 knockout cells showed significantly less fragmentation with a shift to intermediate-size mitochondria. Specifically, PINK1^-/- cells contained 29.2% fragmented mitochondria compared with 77.7% in the vector group ( Figure 6 ). It has been reported previously that upon Drp1 deletion, the mitochondrial morphology in HCT116 cells was more intermediate (48–50). In our study, we inhibited Drp1 expression in HCT116 cells by small interfering RNA (siRNA), and the knockdown efficacy was determined by western blot ( Figure S4C ). Followed by confocal microscopy analysis of mitochondrial morphology. We observed a decrease in intracellular mitochondrial fragmentation after Drp1 knockdown (KD) in the control cells. However, in PINK1^-/- HCT116 cells, mitochondrial morphology did not change much and more of it was still in fusion state ( Figure 6 ). This suggests that the effect of PINK1^-/- on mitochondrial morphology is similar to that of Drp1 reduction.

It has been shown previously that deletion of Drp1 increases apoptosis (49). Analyzing apoptosis in HCT116 cells, we found that 22.51% of the PINK1^-/- cells underwent apoptosis while only 6.95% of the WT cells were apoptotic ( Figure 7A ). Thus, similar to MEFs, cell death was also increased in PINK1^-/- HCT116 cells, although in MEFs increased death was mainly due to increased necrosis rather than apoptosis. In HCT116 cells, we also used PI to study the cell cycle. In contrast to MEFs, we found no difference in the percentage of cells in the G2/M phase in HCT116 cells lacking PINK1( Figure 7B ). In agreement with this result, expression of the cell cycle-related checkpoint proteins p16 and p53 was unaltered in PINK1^-/- HCT116 cells ( Figure 7C ).

PINK1^-/- mice have been described previously (34) and were backcrossed for at least 15 generations onto the pure C57BL/6J background. Male nude mice (BALB/c-nu, 4-6 weeks old) were purchased from Beijing Vital River Laboratory Animal Technology, China. Animal experiments were performed according to the “Guide for the Care and Use of Laboratory Animals”, 8^th Edition, 2011 (The National Academic Press, Washington, and D.C.) and approved by the Institutional Animal Care and Use Committee/Animal Experimentation Ethics Committee of the Harbin Institute of Technology (HIT-IACUC).

Oligonucleotides ( Table S1 ) and antibodies ( Table S2 ) are listed in Supporting Information. We used antibodies at the manufacturer-recommended concentrations, and obtained tissue culture media, trypsin and phosphate-buffered saline (PBS) from Thermo Fisher and fetal bovine serum (FBS) from JYK Biotechnology, China.

Retroviral vectors expressing SV40LT, EGFP and human K-Ras_G12D were based on the vector SFG-MCS, a derivative of the plasmid MFG (67). To generate recombinant retroviruses, we transfected 10 μg viral vector DNA per 6-cm plate into PlatE packaging cells (68) using the calcium phosphate co-precipitation method. To produce lentiviral particles for PINK1 re-expression, we inserted the human PINK1 coding sequence into the lentiviral vector pLVSIN-CMV-Pur (Takara Biotech) and co-transfected 293T cells in 6-cm plates with 5 μg lentiviral vector plasmid, 3.33 μg HIV gag/pol expression plasmid psPAX2 (Addgene #12260) and 1.67 μg VSV-G expression plasmid pMD2.G (Addgene # 12259) (3:2:1 ratio). Viral particles in the medium were collected 24 and 48 hours after transfection, passed through a 0.45 μm filter and stored at -80°C.

We isolated mouse embryonic fibroblasts (MEFs) from E14.5 stage embryos of WT and PINK1^-/- mice as described (69). For immortalization, we infected MEFs with SFG-SV40LT retrovirus in the presence of 4 μg/ml polybrene (Sigma H9268). The immortalized phenotype was confirmed by unlimited passage ability, in contrast to MEFs infected with SFG-EGFP retrovirus that stopped dividing several passages after infection. For transformation, we infected SV40LT-immortalized MEFs twice separated by 24 hours with SFG-Ras_G12D retrovirus in presence of 4 μg/ml polybrene to yield a Ras_G12D-transformed population. Individual Ras_G12D transformed tumor clones were obtained by limiting dilution in 96-well plates. To express human PINK1 in mouse PINK1-deficient Ras_G12D-transformed cells, we infected cells with the pLVSIN-PINK1-Pur lentiviral vector and selected stably transduced cells by growth in medium containing 2 μg/ml puromycin (Sigma P8833).

HCT116 cells were obtained from American Type Culture Collection (ATTC, CCL-247). We used a modified version of the CRISPR-Cas9 plasmid pSpCas9(BB)-2A-GFP (PX458) (Addgene 48138) that was engineered to contain a puromycin expression cassette and BbsI and BsaI cloning sites for insertion of two double-stranded oligonucleotides encoding different sgRNAs. The oligonucleotides encoding sgRNAs targeting two sites in PINK1 exon 1 are shown in Table S1 . After transfecting the CRISPR/Cas9 PINK1 targeting vector into HCT116 cells, puromycin-resistant (2 μg/ml) single clones were obtained by limiting dilution. Genomic DNA from clones was PCR-amplified with primers PINK1-KO-screen-fw and PINK1-KO-screen-rev to identify clones that produced a single, shorter PCR product compared to wildtype genomic DNA ( Figure S3B ). PCR products from such clones were sequenced and the sequences aligned with the human PINK1 gene (gene ID: 65018) to determine the exact PINK1 deletions in different HCT116 PINK1-knockout clones ( Figure S3C ). The two CRISPR target sites, clone-specific deletions and PCR primers used to identify PINK1-knockout clones are indicated in Table S1 . In all clones, the deletions resulted in severely truncated proteins containing only 72 to 113 N-terminal amino acids before running into a premature stop codon, thereby lacking the entire PINK1 kinase domain. To analyze deletions in PINK1 mRNA, cDNA was synthesized from total RNA (Prime Script cDNA kit, Takara Bio) and PCR-amplified with two different primer pairs (fw-1/rev-1 and fw-1/rev2) flanking the deletions ( Figure S4A ).

Cells were grown in DMEM with 10% FBS. Before the soft agar assay, 1.5x10⁴ cells were resuspended in 0.35% low-melting point agarose (Sigma) in DMEM/10% FBS and plated into triplicate wells of a 6-well plate containing a 2 ml base agarose layer (0.7%). After 8-10 days, images of cells were taken with a Nikon inverted microscope at 200x magnification. Three random images per well were captured at different soft agar depths (9 images total), and the colony areas were measured with Image J software (imagej.nih.gov/ij). To measure the growth of HCT116 tumor clones in tissue culture, we plated 10⁴ cells per clone and well (in triplicate wells) and measured cell numbers with the sulforhodamine B assay (70) over a period of 5 days. The average OD534 from three wells was statistically compared between PINK-1 deficient and control tumor clones using 2-way ANOVA.

Cells were detached with trypsin, resuspended in DMEM/10% FBS and centrifuged for 5 min at 500 x g. The cell pellet was washed twice in PBS and cells were counted in a hemocytometer (four big squares). Cell density was adjusted to 1.25x10⁷ or 2.5x10⁷ cells/ml, and 200 μl of cell suspension (2.5x10⁶ or 5x10⁶ cells, matched for all genotypes) was injected subcutaneously into mice. Tumor growth was monitored starting at 7 days and dimensions of tumors were measured with a caliper at the days indicated in the figures. Tumor volumes (in mm³) were calculated as length x width² x 0.5 (width being the smaller dimension) and compared using two-way ANOVA. At the end of the experiment mice were euthanized by CO₂ inhalation.

Cellular RNA was isolated with Trizol reagent, and first strand cDNA synthesized with the Prime Script RT kit (Takara Inc.) from 500 ng total RNA. Two μl of the resulting cDNA (5-fold dilution) was subjected to real-time PCR using SYBR Premix Ex Taq II (Tli RNase H Plus) master mix (Takara Inc). PCR primers are listed in Table S1 . Melting curve analysis was done to confirm single PCR products. We used the 2^-ΔCt method (71) to calculate mRNA expression of each gene relative to 18S rRNA.

Cells were lysed in modified RIPA buffer (50 mM Tris-HCl pH 8.0, 1 % Triton X-100, 0.1 % SDS, 0.14 M NaCl, 1 mM EDTA, and 1 mM EGTA) containing 1% (v/v) protease inhibitor cocktail and - where appropriate - phosphatase inhibitor cocktail (Amresco). 20-30 μg total proteins in the cleared lysates (supernatants of 10 min, 12,000 x g centrifugation) were analyzed by standard Western blot procedures. Secondary antibodies were conjugated to fluorophores or HRP, and protein bands were detected with the Odyssey Infrared Imaging System (Li-COR) or the enhanced chemiluminescence imager. Protein bands were quantified using Image J software (imagej.nih.gov/ij). Three to four independent cell lysates (prepared at different passage numbers) of each genotype (MEF population) and the indicated number of WT and PINK1-deficient cell clones were analyzed and used for statistical evaluation.

To measure expression of total Drp1 and phospho-Drp1 (Ser616) by flow cytometry, 5x10⁵ cells were fixed in 4% PFA for 30 min and centrifuged at 500 x g for 5 min. The cell pellet was gently resuspended and incubated in 100 μl PBS/0.2% Triton X-100 for 10 min, washed with PBS, and the cells were incubated with primary antibody diluted in PBS/0.05% Tween-20 (PBST) for 1 hour at RT (antibody dilutions as recommended by the manufacturer). After washing twice in PBST, the cells were incubated for 30 min with 1:400-diluted Cy3-conjugated secondary antibody, followed by a final wash and re-suspension in PBST before flow cytometry. To determine background fluorescence and set the fluorescence window for positive cells (M1), control cells were incubated with secondary antibody only. N=4 independent antibody incubations/flow cytometry experiments were carried out with each genotype/cell population for statistical analysis. To measure the mitochondrial membrane potential Δψm, cells were incubated with 50 nM TMRE for 30 min at 37°C. Δψm was compared between groups of WT and PINK1^-/- cells, where the groups were comprised of individual cell clones and the cell population for each genotype (n=3-4 clones/populations per genotype, each clone and population measured in triplicate). Mitochondrial ROS were measured in WT, PINK1^-/- and PINK1^-/-plus huPINK1 (human PINK1-overexpressing) cell populations incubated for 30 min at 37°C with 2.5 μM MitoSOX Red (ThermoFisher M36008) (n=3 dye incubations/flow cytometry experiments per genotype/population). Negative control cells were incubated in buffer without TMRE or MitoSOX to determine background fluorescence and set the window for positive cells (M1). Fluorescence (FL2-H) was measured with the BD FACSCalibur using a 585/42 nm band pass emission filter, and the mean fluorescence of the cells in the positive window (M1) was compared between genotypes/cell populations.

For analysis of mitochondrial network morphology, cells (4x10⁵/35-mm plate) grown on cover glasses were washed with PBS, fixed in 4% PFA and mitochondria were stained by immunocytochemistry with anti-LRP130 antibody and Cy3-conjugated secondary IgG. LRP130 is exclusively expressed in mitochondria (41). Nuclei were visualized with DAPI. The percentage of cells with various mitochondrial morphologies was analyzed in a blinded manner using 15 random confocal images per genotype (87-243 cells; 5-13 images from 3 cover glasses), taken with 40x objective and digitally enlarged 3x for analysis using Image J software. Cells with mostly (>80-90%) small and spherical (fragmented) mitochondria were classified as having a “fragmented” mitochondrial network, cells with appreciable proportions (>40-50%) of elongated/fused mitochondria that also contained fragmented mitochondria were classified as “intermediate”, and cells with mostly (>80-90%) elongated/fused mitochondria were classified as “elongated/fused".

Cells (10⁵/cm²) were grown for 48 hours before being collected by trypsinization and resuspended in ice-cold 70% ethanol for fixation (1 hour on ice). Fixed cells were centrifuged (400 x g for 5 min), washed once with 2 ml PBS and centrifuged again. The final cell pellet was resuspended in PBS containing 1:10 diluted propidium iodide/RNase A solution (Sungene Biotech, China), and cells were incubated for 30 min at RT and protected from light. Cells were analyzed by flow cytometry (FACSCalibur, Becton Dickinson) and data files were imported into FCS Express 6.0 Plus software (De Novo software, Glendale, CA). For processing and quantification, the main cell population was gated in the FSC vs. SSC plot. Within the main population, cell debris and aggregated cells were excluded by gating on single cells in the FL2-A vs. FL2-W plot. Markers for G0/G1, S and G2/M phases were placed in the FL2-A vs. cell count plot of WT Ras_G12D-transformed cells, and the same markers were used for all other cell samples.

Tumor tissues dissected from nude mice fixed with 10% formalin, and embedded in paraffin. The tissue sample was subjected to antigen retrieval by boiling in 0.01 mol/L citrate buffer for 5 min. Slides were then incubated with anti-RasG12D or anti-Ki-67 antibody at 4°C. for the night. Slides were then washed with PBS, fixed with 10% formalin and provided for staining. Detection was carried out by the REAL EnVision detection system (Dako) with diaminobenzidine peroxidase serving as chromogen. Images were collected, and were de-interfered and then quantitatively analyzed for gray values using Image J software.

The apoptotic rates were evaluated by Caspase-Glo^® 3/7 Assay kit (Promega, G8090). Briefly, the tumor tissues were harvested in urea buffer (2M Thiourea, 4% CHAPS, 40mM Tris-Base, 40mM DTT, 2% Pharmalyte) and sonicated to crush DNA. The same amount of tumor tissues protein lysates (500μg) was transfer into the 96-well cell culture plate. Caspase 3/7 activity assay was repeated independently for three times according to its manufacturer’s instructions, which was represented as a fold-increase of fluorescence calculated by comparing to vector groups.

The small GTPase activity of Ras was measured using the Ras activation assay Biochem kit (cytoskeleton #BK008). Cells were harvested with cell lysis buffer (50 mM Tris pH 7.5, 10 mM MgCl2, 0.5 M NaCl, and 2% Igepal when reconstituted) contains 1×Protease Inhibitor Cocktail on ice. The resulting protein lystates were immediately clarified by centrifugation at 10,000 rpm, for 2 min at 4°C and protein concentrations were determined by Bradford kit. After that, 400ug of protein lysates was incubated with Raf-RBD beads (Cat. # RF02) on a rotator at 4°C for 1h. The Raf-RBD beads were pelleted by centrifugation at 3-5,000 × g for 1 min at 4°C. 90% of the supernatant was carefully removed and the beads were washed once with 500μl Wash Buffer. The Raf-RBD beads were pelleted again by centrifugation at 3,000xg for 3 min at 4°C. The supernatant was carefully removed without disturbing the beads. 20 μl of 2×Laemmli sample buffer was added into each tube and the beads were boiled for 2 min. Then resulting lysates were subjected to the following western blot analysis of Ras.

Apoptotic and necrotic cells were detected by the Annexin V-FITC apoptosis analysis kit (Tianjin Sungene Biotech, China). Different genotypes of cells density were adjusted to 5 × 10⁶ cells/ml. About 100 μl cell suspension was incubated with 2.5 μl AnnexinV/FITC for 10 min and then 2.5 μl PI for 5 min at room temperature in dark. The rate of apoptosis was measured by flow cytometry (BD Pharmingen).

Statistics was performed with Prism 9.0 software (Graph Pad). All data are presented as mean ± SD, except tumor volumes in nude mice, which are shown as mean ± SEM. Two groups were compared by unpaired t-test, and three or more groups were compared by ANOVA and Tukey’s multiple comparisons test. Two-way ANOVA with Sidak’s multiple comparisons test was used to compare tumor growth in nude mice and the growth of WT and PINK1-knockout HCT116 clones in culture. Differences were considered significant at P<0.05.