Abstract
Objectives
The PLCG2 (PLCγ2) gene is a member of PLC gene family encoding transmembrane signalling enzymes involved in various biological processes including cell proliferation and apoptosis. Our earlier study indicated that PLCγ2 may be involved in the termination of regeneration of the liver which is mainly composed of hepatocytes, but its exact biological function and molecular mechanism in liver regeneration termination remains unclear. This study aims to examine the role of PLCγ2 in the growth of hepatocytes.
Materials and methods
A recombinant adenovirus expressing PLCγ2 was used to infect primary rat hepatocytes. PLCγ2 mRNA and protein levels were detected by qRT‐PCR and Western blot. The subcellular location of PLCγ2 protein was tested by an immunofluorescence assay. The proliferation of hepatocytes was measured by MTT assay. The cell cycle and apoptosis were analysed by flow cytometry. Caspase‐3, ‐8 and ‐9 activities were measured by a spectrophotometry method. Phosphorylation levels of PKCD, JNK and p38 in the infected cells were detected by Western blot. The possible mechanism underlying the role of PLCγ2 in hepatocyte growth was also explored by adding a signalling pathway inhibitor.
Results
Hepatocyte proliferation was dramatically reduced, while cell apoptosis was remarkably increased. The results demonstrated that PLCγ2 increased the phosphorylation of PKCD, p38 and JNK in rat hepatocytes. After PKCD activity was inhibited by the inhibitor Go 6983, the levels of both p‐p38 and p‐JNK MAPKs significantly decreased, and PLCγ2‐induced cell proliferation inhibition and cell apoptosis were obviously reversed.
Conclusions
This study showed that PLCγ2 regulates hepatocyte growth through PKCD‐dependently activating p38 MAPK and JNK MAPK pathways; this result was experimentally based on the further exploration of the effect of PLCγ2 on hepatocyte growth in vivo.
1. INTRODUCTION
The recovery of liver volume after resection is probably regulated by the functional needs of the organism, when liver regeneration (LR) process enters into the terminal phase during which the normal liver‐to‐bodyweight ratio has been restored. A large body of research has been done to investigate the mechanisms controlling the early phase of LR.1, 2, 3 However, the mechanisms controlling the terminating phase are not explored to the same extent,4, 5 and until now, the limited information has been available about the inhibitory factors and stop signals.
Some cytokines, such as transforming growth factor β (TGFβ) and activins, are currently accepted as important factors resulting in the termination of the regenerative process through negatively regulating hepatocyte growth.6 For example, it is well‐known that TGFβ inhibits hepatocyte proliferation both in vitro and in vivo.7 Extracellular matrix (ECM) components were considered to be essential for the proper termination of LR.8 PPARγ would inhibit hepatocyte proliferation once the cell number is sufficient.9 In addition, the study from Oakley et al10suggested that neurotrophins (NTs, consisting of NGF, BDNF, NT3 and NT4/5) may play a role in hepatic regeneration because of their elevated expressions in both mRNA and protein levels in association with hepatocyte proliferation after partial hepatectomy (PH) in mice. Moreover, Asai and his coworkers demonstrated that NGF produced by regenerating hepatocytes could induce apoptotic death of activated HSCs via the receptor p75NTR, thus leading to the termination of LR.11 In earlier study, we thoroughly analysed the changes in genome‐scale gene expression during LR in a rat 2/3 PH model and found that PLCG2 gene was significantly upregulated at the termination phase of LR (Table S1). And PLCG2 was referred as an important gene for the termination of LR according to the result of systems biology analysis.
PLCG2 encodes phospholipase C gamma 2 (PLCγ2), one member of the PLCγ subfamily including PLCγ2 itself and PLCγ1,12 and is responsible for the hydrolysation of phosphatidylinositol to generate diacylglycerol (DAG) and inositol 1,4,5‐trisphosphate (IP3) upon activation. IP3 mediates Ca2+ signalling, while DAG activates protein kinase C (PKC)‐mediated signalling,13 which is further involved in various cellular processes such as proliferation, differentiation and immunity.14 Evidences has indicated that PLCγ2 plays a key role in the development of the immune system. For instance, Ichise et al15found that the PLCγ2‐lacking mutant mice exhibited lymphatic vessel abnormalities, resulting from malfunctions in the lymphatic circulation and misconnections between blood and lymphatic vessels. Additionally, some studies showed the positive effect of PLCγ2 on cell apoptosis. For example, the findings of study conducted by Tomlinson et al. indicated the absolute necessity of PLCγ2 in Bruton's tyrosine kinase (BTK): ER‐induced apoptosis response.16 Zhang et al17 also reported the important role of the translocation and upregulation of PLCγ2 in TPA‐induced apoptosis in gastric cancer cells. Nevertheless, the function of PLCγ2 on hepaotcyte growth and the underlying mechanisms remain unclear.
PKC is a family of enzymes which are subgrouped into three classes, including the classical, novel and atypical. The activation of classical (α, β and γ) and novel PKCs (δ, ε, θ and η) mainly rely on phospholipids, whereas atypical PKCs (ζ, ι and λ) do not require these lipids.18 It is noteworthy that different PKC isoforms somewhat differ in their biological functions. For example, many studies have suggested the involvement of PKCD in the activation of both p38 MAPK in heart failure19 and JNK MAPK pathway in mitochondria‐dependent hepatic cell apoptosis.20 p38 and JNK MAPK signalling pathways are two important intracellular signalling pathways which have been proved to be tightly associated with tumour cell apoptosis.21, 22 On the whole, as mentioned above, PLCγ2 induces cell apoptosis or growth inhibition in some cancers, such as gastric cancer and lymphoma.17, 23 However, whether PLCγ2 is involved in hepatocyte apoptosis via PKCD‐dependent p38 or JNK MAPK signalling pathways remains to be elucidated.
Based on the abovementioned studies, the present study aimed to discuss the effects of PLCγ2 gene overexpression on hepatocytes and to investigate the possible molecular mechanisms underlying these effects. Our study found that PLCγ2 inhibited proliferation and promoted apoptosis in rat hepatocytes in vitro and that the activation of PKCD/p38 and PKCD/JNK signalling might be involved in these processes. These results offer a new insight for comprehensively understanding the mechanism of LR termination.
2. MATERIALS AND METHODS
2.1. Materials
An empty adenovirus expressing green fluorescent protein (Ad‐GFP) and a recombinant adenovirus expressing PLCγ2 (Ad‐PLCγ2) with GFP were constructed in our earlier study. Collagenase IV and MTT were obtained from Sigma (San Francisco, USA). 0.25% trypsin was obtained from Gibco (Grand Island, USA). DNaseIwas obtained from Roche (Indianapolis, USA). Hepatocyte growth factor (HGF) was obtained from PeproTech (Rocky Hill, USA). The TRIzol reagent was purchased from Aidlab (Beijing, China). The semi‐quantitative RT‐PCR reagents were purchased from Genecopoeia (Rockville, USA). The RT‐qPCR reagents were purchased from VAZYME (Nanjing, China). The PKCD inhibitor Go 6983 was purchased from Selleck (Houston, UK). Rabbit anti‐PLCγ2, anti‐p38 and anti‐p‐p38 antibodies were purchased from Abcam (Cambridge, UK). Rabbit anti‐PKCD, anti‐p‐PKCD, anti‐JNK and anti‐p‐JNK antibodies were purchased from CST (Beverly, USA). The mouse anti‐β‐actin antibody and the secondary antibodies (an HRP‐conjugated goat anti‐mouse IgG antibody, an anti‐rabbit IgG antibody, a Cy3‐conjugated goat anti‐mouse IgG antibody, an anti‐rabbit IgG antibody) were purchased from BOSTER (Wuhan, Hubei, China). Hoechst 33258, Caspase activity detection kit and Western blot detection reagents were purchased from the Beyotime Institute of Biotechnology (Jiangsu, China).
2.2. Methods
2.2.1. Isolation and primary culture of hepatocytes from rat liver
The hepatocytes were isolated and purified from Sprague‐Dawly (SD) rats by two‐step collagenase perfusion, filtration and centrifugation (50 × g for 5 minutes). The cell viability was between 96% and 98%, as determined by trypan blue staining. The isolated hepatocytes were cultured in DMEM supplemented with 10% FBS and 5 ng/mL HGF. The cell concentration was adjusted to approximately 1.5 × 106 cells/mL, and the cell suspension was plated in a 24‐well plate in a humidified atmosphere with 5% CO2 at 37°C. After 4 hours, the cultures were washed three times and incubated for an additional 24 hours in the culture medium as mentioned above. The medium was replaced with fresh culture medium every 2 days for 6‐7 days. Morphological changes in hepatocytes of rats were observed under an inverted microscope.
2.2.2. Adenovirus transfection and grouping
After 1 week of primary culturing, the rat hepatocytes were seeded into a 6‐well plate (1 × 106 cells/well) and cultured overnight. The rat hepatocytes cultured in a six‐well plate were infected with the recombinant adenovirus expressing PLCγ2 (Ad‐PLCγ2) at a multiplicity of infection of 100. The cells were divided into three groups as follows: the experimental group, in which the cells were infected with Ad‐PLCγ2; the Ad‐dull group, in which the cells were infected with empty adenovirus Ad‐GFP; and the blank control group, in which the cells were not infected with any adenovirus. After infection for 6 hours, the medium was replaced with fresh medium followed by cell culturing for an additional 24 hours. The cells were observed under a fluorescence microscope to assess the infection efficiency.
2.2.3. RNA extraction, semi‐quantitative RT‐PCR and RT‐qPCR analysis
Total RNA was extracted from rat hepatocytes using the TRIzol reagent according to the manufacturer's instructions. First‐strand DNA was synthesized using the HiScript Reverse Transcriptase. Semi‐quantitative RT‐PCR measuring gene expression level was performed according to the following condition: 30 cycles at 94°C × 30 seconds, 56°C × 30 seconds, 72°C × 25 seconds; 72°C × 4 minutes for 1 cycle and 4°C × 4 minutes for 1 cycle. PCR products were separated by electrophoresis on a 1.5% agarose gel. RT‐qPCR was run in the ViiA™ 7 Real‐Time PCR System (Applied Biosystems, Carlsbad, USA) using SYBR Green/Flourescein qPCR Master Mix (VAZVME) according to the following protocol: 50°C for 2 minutes; 95°C for 10 minutes; then, 40 cycles of 95°C for 30 seconds and 60°C for 30 seconds. β‐Actin was used as an internal control. The primers were designed according to gene sequence and were synthesized by Beijing Qing Ke Biotech Corp (Table 1). The data were analysed according to the 2−ΔΔCt method. The mRNA levels were normalized to β‐Actin. Three independent experiments were performed for each group.
Table 1.
Primer sequences for semi‐quantitative RT‐PCR and real‐time qPCR
| Genes | Primer sequence (5′→3′) |
|---|---|
| PLCγ2 | FP: CCGACTCTTACGCCATCA |
| RP: GGGTAGCGAAGCCTCATC | |
| β‐Actin | FP: CACGATGGAGGGGCCGGACTCATC |
| RP: TAAAGACCTCTATGCCAACACAGT |
2.2.4. Western blot assay
Briefly, primary rat hepatocytes with different treatments were lysed with ice‐cooled RIPA buffer containing PMSF (100 mmol/L) for 30 minutes and centrifuged (10000 × g) for 5 minutes at 4°C, and finally the supernatants were collected. The concentration of total protein was determined by the BCA assay. An equal amount of the cell extracts was separated on SDS‐PAGE gels (5% concentration gel, 10% separation gel) and then transferred onto PVDF membranes. After blocking with 5% non‐fat dried milk in TBST for 2 hours at room temperature, the membranes were probed with the rabbit anti‐PLCγ2 (1:200 dilution) primary antibody and incubated at 4°C overnight. HRP‐conjugated secondary antibodies were added at a dilution of 1:50000 followed by an incubation at 37°C for 2 hours. β‐Actin served as an internal reference. The blots were developed with ECL (Thermo, Waltham, USA) and were exposed to X‐ray film. Three individual experiments were performed for each group.
2.2.5. Immunofluorescent staining
First, the cells were fixed with 4% paraformaldehyde for 15 minutes to enhance membrane permeability with 0.3% Triton X‐100. After blocking with normal goat serum (Beyotime, China) for 30 minutes, the cells were incubated with a rabbit anti‐PLCγ2 primary antibody overnight at 4°C. After washing with PBS buffer, the cells were incubated with a Cy3‐conjugated goat anti‐rabbit IgG secondary antibody (1:100 dilution) for 1 hour at room temperature. The cells were then stained with DAPI to visualize the nuclei. The fluorescence was observed under a fluorescence microscope (BX53, Olympus Corporation, Tokyo, Japan).
2.2.6. Cell viability assay
Cell proliferation was analysed by the 3‐(4,5)‐dimethylthiazol(‐z‐yl)‐3,5‐diphen ‐yltetra‐zolium bromide (MTT) assay. Briefly, the adenoviruses‐infected cells or the normal cells were seeded in a 96‐well plate with 2 × 104 cells/well in DMEM supplemented with 10% FBS and 5 ng/mL HGF. At the indicated time‐points including 24, 48 and 72 hours, 20 μL of MTT reagent (5 mg/mL) was added to each well, and the mixture was cultured for another 4 hours. The MTT solution was removed and 150 μL of DMSO was added to dissolve the formazan crystals, followed by the measurement at 568 nm using a microtiter plate reader (Thermo, USA). The percentage of cell viability was calculated using the following formula: the absorbance of test wells/the absorbance of control wells × 100%.
2.2.7. Cell cycle and apoptosis analysis
Cell cycle and apoptosis were evaluated using flow cytometry (FCM) method. Briefly, the cells were seeded in a 6‐well plate at a density of 2 × 105 cells/well and treated with the relevant adenovirus for 24 hours. The cells from each group were digested with 0.25% trypsin and then harvested by centrifugation. After two washes in ice‐cold PBS and resuspending, an aliquot of the samples was fixed with pre‐cold 70% ethanol for 4 hours and then treated with RNase A for 30 minutes at 37°C, followed by staining with 50 μg/mL PI for 30 minutes at 4°C in darkness. The remaining samples were added into 5 μL 7‐AAD for 15 minutes in darkness at room temperature, followed by the addition of 1 μL of Annexin V‐APC. Finally, a FACSCalibur flow cytometer (BD Biosciences, San Jose, USA) was used to assay the apoptosis rate and cell cycle. Three independent experiments were performed for each group.
2.2.8. Caspase‐3,‐8 and ‐9 activity assay
Caspase‐3, ‐8, and ‐9 activities were determined by measuring the absorbance at 405 nm after the cleavage of the synthetic substrate Ac‐DEVD‐pNA. In detail, the cells were collected and lysed on ice for 30 minutes in cell lysis buffer. A total of 50 μL of lysates was reacted with 10 μL of Ac‐DEVD‐pNA (2 mmol/L) and 40 μL of reaction buffer. The mixtures were maintained for 1 hour at 37°C and subsequently analysed by a microtiter plate reader (Thermo, USA). The enzyme activity was determined from a standard curve prepared using ρNA. The relative level of pNA was normalized against the protein concentration for each extract.
2.2.9. PKCD/p38 and PKCD/JNK pathway activity assay
After 48 hours of infection, the PKCD/p38 and PKCD/JNK pathway activities were evaluated through detecting expression levels of total PKCD, p38, JNK1/2, and p‐PKCD, p‐p38, p‐JNK1/2 in the cells from each group using Western blot method. The detailed procedure was the same as described above. The dilutions of the primary antibodies rabbit anti‐PKCD, p38, JNK1/2, p‐PKCD, p‐p38 and p‐JNK1/2 were 1:1000, 1:1000, 1:1000, 1:1000, 1:1000 and 1:3000, respectively. The secondary antibody IgG coupled to HRP was diluted to 1:50000. Three separate experiments were performed for each group.
2.2.10. Inhibition of a PKCD inhibitor on p38 and JNK activities and its effect on PLCγ2‐regulated growth of hepatocytes
The cells were pre‐treated with 10 μM Go 6983(PKCD inhibitor)for 1 hour, followed by treatment with Ad‐dull or Ad‐PLCγ2 for the indicated time points. After infection of 48 hours, the phosphorylation of PKCD, p38 and JNK was detected by Western blot as described above. In addition, the cell viability was assayed by the abovementioned MTT method. Cell proliferation was assessed by the FCM and BrdU incorporation tests. Cell apoptosis was evaluated by FCM and Hoechst 33258 staining. The procedure for FCM was the same as above, and that of Brdu incorporation and Hoechst staining were as follows:
BrdU incorporation
Dividing cells were detected using a BrdU incorporation experiment. After 24 hours of adenovirus infection, rat hepatocytes continued to grow in the medium with 10 μmol/mL BrdU solution. After culturing for another 24 hours, the cells were fixed with 4% paraformaldehyde for 15 minutes at temperature, and incubated with 0.5% Triton X‐100 for 20 minutes to enhance the membrane permeability. After treatment with 2N HCl and 0.1 mol/L boracic acid buffer, the cells were incubated with primary antibody BrdU (1:100 dilution) overnight at 4°C, allowing BrdU to incorporate into DNA, followed by an incubation with a Cy3‐labeled IgG secondary antibody for 1 hour at 25°C and then staining with DAPI for visualizing the nuclei. Finally, fluorescent photos were observed and captured under a fluorescence microscope (BX53, Olympus, Tokyo, Japan).
Hoechst 33258 staining assay
The Hoechst staining assay was performed according to the manufacturer's instructions using Hoechst 33258. The cells seeded in a 12‐well plate with prepared cell sheets were treated with adenovirus for 48 hours and washed three times in PBS. The cells were fixed with 4% paraformaldehyde for 15 minutes and washed twice in PBS. A sufficient Hoechst 33258 staining solution (10 μg/mL) was added onto each cell climbing sheet, and the cells were incubated for 10 minutes at room temperature in darkness. The cells were washed three times in PBS and viewed under a fluorescence microscope (BX53, Olympus, Tokyo, Japan).
2.2.11. Statistical analysis
The data are expressed as the means ± standard deviation (SD). A one‐way analysis of variance (ANOVA) was used to analyse the differences between groups, and the LSD method of multiple comparisons was used when the probability for ANOVA was statistically significant using SPSS 17.0. Statistical significance was set at P < .05.
3. RESULTS
3.1. PLCγ2 is successfully overexpressed in rat hepatocytes
To identify the role of PLCγ2 in hepatocytes, the recombinant adenovirus Ad‐PLCγ2 was used to infect primary rat hepatocytes. In the present study, the infection efficiency of Ad‐PLCγ2 in rat hepatocytes was >90% under fluorescence microscopy (Figure 1A). In addition, semi‐quantitative RT‐PCR (Figure 1B), RT‐qPCR (Figure 1C) and western blot assays (Figure 1D) were performed to measure the expression level of PLCγ2 after 24 hours of infection. An obvious increase in PLCγ2 expression was observed in the Ad‐PLCγ2 group compared with the control and Ad‐dull groups (P < .01 and P < .05) in the hepatocytes. Additionally, the result of immunofluorescence assay demonstrated that PLCγ2 was uniformly localized in the cytoplasm of rat hepatocytes, and the fluorescence intensity in Ad‐PLCγ2 group was significantly higher than the blank control and Ad‐dull groups. These data indicated that PLCγ2 was successfully overexpressed in hepatocytes.
Figure 1.
PLCγ2 was overexpressed in rat hepatocytes. (A) Constructed adenovirus vector Ad‐PLCγ2 was used to infect rat hepatocytes. The infection efficiency of Ad‐PLCγ2 (MOI = 100) in the hepatocytes was >80% under fluorescence microscopy. (B) RT‐pCR, (C) RT‐qPCR and (D) Western blot assays were used to measure the expression of PLCγ2, which showed an obvious increase in PLCγ2 expression in the Ad‐PLCγ2 group compared with the Ad‐dull and blank groups (**P < .01 and * P < .05 vs the Ad‐dull and blank control groups). No difference was found between the Ad‐dull and blank control groups (P > .05). (E) Immunofluorescence assay was carried out to observe the distribution and expression intensity of PLCγ2 in rat hepatocytes. PLCγ2 was distributed throughout the cytoplasm and its intensity was obviously increased in the Ad‐PLCγ2 group compared with the Ad‐dull and control groups
3.2. PLCγ2 inhibits cell viability of rat hepatocytes
To study the effect of PLCγ2 on cell growth, the rat hepatocytes were treated with Ad‐PLCγ2 for 24, 48 and 72 hours, and cell viability was assessed by MTT method. As shown in Figure 2A, the viability of Ad‐PLCγ2‐treated hepatocytes was significantly inhibited at 48 hours (P < .01) when compared with the Ad‐dull and blank control groups. And there was no significantly difference (P > .05) in the cell viability between the Ad‐dull group and the control group. Therefore, we selected 48 hours as the time‐point for the remaining experiments.
Figure 2.
PLCγ2 inhibits the proliferation of rat hepatocytes. A, Rat hepatocytes were infected with Ad‐PLCγ2 for 24, 48 and 72 hours. At the end of the indicated time, cell viability was determined by MTT. B, Rat hepatocytes were infected with Ad‐PLCγ2 for 48 hours. The distribution of the cell cycle was analysed by flow cytometry. The assay was performed in triplicate. **P < .01 and *P < .05 vs the Ad‐dull and blank control groups
3.3. PLCγ2 induces cell cycle arrest in rat hepatocytes
Regulation of cell cycle progress directly influences and determines the process of cell proliferation. FCM analysis was used to detect the cell cycle distribution after PLCγ2 was overexpressed (Figure 2B). The result showed that PLCγ2 expression decreased the percentage of rat hepatocytes in the S phase from (16.10 ± 0.94)% to (11.23 ± 0.89)% (P < .05) compared to the control group, and increased the percentage of G1 phase from (76.64 ± 0.32)% to (83.02 ± 0.45)% (P < .05) compared to the control group.
3.4. PLCγ2 stimulates apoptosis of rat hepatocytes in vitro
To evaluate cell apoptosis in rat hepatocytes after treatment with Ad‐PLCγ2, FCM assay and caspases activity detection were performed. In Figure 3A, it was shown that, after treatment with Ad‐PLCγ2 for 48 hours, the apoptotic rate of rat hepatocytes was significantly increased from (8.47 ± 0.41)% to (14.95 ± 0.60)% (P < .01) compared to the Ad‐dull group, and the Ad‐dull group showed almost no change in the apoptotic rate when compared with the control group (P > .05). Furthermore, caspase‐3, ‐8 and ‐9 activities were examined to confirm whether PLCγ2 increased the activities of these caspases using Ac‐DEVD‐ρNA as a substrate, and the data showed a significant increase in caspase‐3, ‐8 and ‐9 activities in rat hepatocytes after Ad‐PLCγ2 infection (P < .05, Figure 3B).
Figure 3.
PLCγ2 induces apoptosis in rat hepatocytes in vitro. A, Rat hepatocytes were infected with Ad‐PLCγ2 for 48 hours and were then analysed for apoptosis by flow cytometry. The percentage of apoptotic cells in rat hepatocytes for each group are quantified in the right panel (**P < .01 vs Ad‐dull group). B, PLCγ2 expression stimulated the activities of caspase‐3, ‐8 and ‐9 in rat hepatocytes. The cells were treated with adenovirus for 48 hours and the activities of caspase‐3, ‐8 and ‐9 were determined by a spectrophotometry method as described in the Materials and Methods. **P < .01 vs Ad‐dull group
3.5. PLCγ2 induces the activation of PKCD‐dependent MAPK signalling pathways in rat hepatocytes
It was previously reported that the activation of p38 and JNK MAPK signalling pathways was associated with cell apoptosis. To determine whether PLCγ2 expression is involved in the activation of PKCD‐dependent MAPK signalling pathway in rat hepatocytes, this study detected the phosphorylation of PKCD and MAPKs in the cell lysates of rat hepatocytes from different experimental groups by Western blot analysis. The result showed that PLCγ2 overexpression caused the enhancement of the phosphorylation levels of PKCD, p38 and JNK in rat hepatocytes (P < .01, P < .05 and P < .01, Figure 4A, B and C, respectively), preliminarily indicating that PLCγ2 promoted the activity of PKCD‐dependent p38 and JNK MAPK signalling pathways in rat hepatocytes.
Figure 4.
PLCγ2 activates the PKCD‐dependent p38/MAPK and JNK/MAPK signalling pathways in rat hepatocytes. (A) The phosphorylation levels of PKCD, (B) The phosphorylation levels of p38 and (C) JNK in Ad‐PLCγ2‐infected hepatocytes were detected by Western blot analysis. The densitometric quantification data are shown in the right panel. **P < .01 and *P < .05 vs the Ad‐dull and blank control groups
3.6. Impact of a PKCD inhibitor on the activities of p38 MAPK and JNK MAPK
To investigate whether PKCD mediates the activation of p38 and JNK MAPK signalling pathways in PLCγ2‐induced apoptosis of rat hepatocytes, the PKCD inhibitor Go 6983 was used to pre‐treat primary rat hepatocytes. Western blot analysis showed that the phosphorylation level of PKCD was significantly decreased (P < .05) in the Go 6983‐pre‐treated cells infected with Ad‐PLCγ2, when compared with the Ad‐PLCγ2 group (Figure 5A), suggesting the effective inhibition of Go 6983 against PKCD activity. Consistently, the PLCγ2‐induced phosphorylation of p38 and JNK1/2 was also blocked by the PKCD inhibitor Go 6983 (P < .05, Figure 5B).
Figure 5.
Go 6983 suppresses the activation of p38 and JNK MAPKs in the PLC γ2‐induced apoptosis of rat hepatocytes. A, The effect of Go 6983 on PKCD activity was detected by Western blot. The phosphorylation of PKCD was obviously inhibited by Go 6983; *P < .05 vs. the Ad‐PLC γ2 group. B, The effects of Go 6983 on the activation of p38 and JNK were tested by Western blot. The phosphorylation of p38 and JNK was suppressed by Go 6983; **P < .01 vs Ad‐PLC γ2 group. The densitometric quantification data are shown in the right panel
3.7. Impact of the inhibition of PKCD/p38 and PKCD/JNK signalling pathways on PLCγ2induced apoptosis of hepatocytes
To investigate the impact of PLCγ2 on hepatocyte growth when inhibiting the activation of the PKCD/p38 and PKCD/JNK pathways by the inhibitor Go 6983, firstly, we used the MTT method to measure the cell viability of Go 6983‐pre‐treated hepatocytes after 24, 48, 72 hours of infection with Ad‐PLCγ2. As shown in Table 2, Go 6983 significantly suppressed the PLCγ2‐induced anti‐proliferative effect at 48 and 72 hours (P < .01). Thus, we selected 48 hours as the time‐point for the following experiments.
Table 2.
The effect of PLCγ2 on the cell viability of hepatocytes with or without Go 6983 pretreatment
| Group | Infection time points (hours) | ||
|---|---|---|---|
| 24 | 48 | 72 | |
| Control | 1.00 ± 0.00A | 1.00 ± 0.03A | 1.000 ± 0.04A |
| Ad‐dull | 0.99 ± 0.03A | 0.99 ± 0.02A | 0.98 ± 0.04A |
| Ad‐PLCγ2 | 0.92 ± 0.03A | 0.88 ± 0.018B | 0.83 ± 0.02B |
| Ad‐PLCγ2 + Go 6983 | 0.97 ± 0.02A | 0.95 ± 0.01AC | 0.91 ± 0.02C |
Same capital letters within the same column denote no significant difference (P > .05); the different capital letters denote significant difference (P < .05).
The cell apoptotic death of rat hepatocytes was visualized in the presence and absence of Go 6983 through Hoechst 33258 staining (Figure 6A). The cells were examined under a fluorescence microscope (200×). Compared with the blank control and Ad‐dull groups, a significantly higher percentage of the cells in the Ad‐PLCγ2 group showed the apoptotic morphology with irregular and condensed nuclei. However, after pre‐treatment with the PKC inhibitor Go 6983, the percentage of apoptotic cells was significantly decreased as observed by their normal morphology. In addition, the FCM analysis showed that the percentage of apoptotic cells was (22.93 ± 1.58)% in the Ad‐PLCγ2 group, which was significantly higher (P < .01) than that in the control group, while the apoptotic rate of the cells in the Go 6983‐pre‐treated group markedly fell to (9.93 ± 1.20)% (P < .01 vs. Ad‐PLCγ2 group; Table 3, Figure 6B).
Figure 6.
Go 6983 suppresses PLCγ2‐induced apoptosis in rat hepatocytes. A, At the end of the 48‐h incubation, the hepatocytes were stained with Hoechst 33258 and the morphological changes were observed by fluorescence microscopy. B, Cell apoptosis was assayed by flow cytometry. The cells in the presence or absence of Go 6983 were infected by Ad‐PLCγ2 for 48 hours and were then analysed for the apoptosis index by flow cytometry. C, DNA synthesis was assayed by BrdU incorporation method. The hepatocytes pretreated with or without Go 6983 were infected with Ad‐PLCγ2 for 48 hours and DNA synthesis in the cells was visualized by BrdU incorporation. D, Cell cycle was assayed by flow cytometry. The distribution of the cell cycle was analysed by FCM
Table 3.
The impact of PLCγ2 on the apoptotic rate and cell cycle distribution of rat hepatocytes with or without Go 6983 pretreatment
| Group | Apoptotic rate (%) | Cell cycle (%) | ||
|---|---|---|---|---|
| G1 | S | G2/M | ||
| Blank control | 9.73 ± 0.32A | 66.51 ± 0.31A | 16.34 ± 0.23A | 17.15 ± 0.53A |
| Ad‐dull | 9.93 ± 1.20A | 67.26 ± 0.36A | 16.05 ± 0.56A | 16.70 ± 0.43A |
| Ad‐PLCγ2 | 22.93 ± 1.58B | 75.65 ± 0.23B | 9.53 ± 0.51B | 14.81 ± 0.72B |
| Ad‐PLCγ2 + Go 6983 | 9.93 ± 1.20AC | 70.97 ± 0.63AC | 11.76 ± 0.57AC | 17.27 ± 0.21AC |
Same capital letters within the same column denote no significant difference (P > .05); the different capital letters denote significant difference (P < .05).
Cell cycle progression in the PLCγ2‐infected hepatocytes pre‐treated with or without Go 6983 was investigated using BrdU incorporation assay (Figure 6C). The results showed that the proportion of the cells in the DNA synthesis phase was significantly reduced in the Ad‐PLCγ2 group compared to the blank control and Ad‐dull groups. However, the proportion of cells in the DNA synthesis phase was obviously recovered after pre‐treating with Go 6983, as observed under a fluorescence microscope. Additionally, FCM analysis result showed that Go 6983 pre‐treatment significantly increased the S‐phase cell fraction and reduced the G1‐phase cell fraction (P < .01 vs. the Ad‐PLCγ2 group; Table 3, Figure 6D). These results demonstrated that Go 6983 significantly alleviated the PLCγ2‐induced anti‐proliferative effect.
4. DISCUSSION
The liver is highly susceptible to damage by physical and chemical factors. In essence, liver regeneration process is a protective mechanism by which liver tissue recovers from physical or chemical injury.24 whereas the precise molecular mechanism of liver regeneration has not yet been clarified. In the earlier study, we found the significant upregulation of PLCγ2 gene at the termination phase of rat LR through a high‐throughput gene profiling analysis (Table S1). Additionally, a recent study conducted by our lab showed that the knockdown of an endogenous PLCγ2 gene resulted in the enhanced proliferation and the decreased apoptosis in rat hepatocytes in vitro (Figure S1). These data obtained from just only RNA interference technology suggest that PLCγ2 plays an important role in the end of LR via regulating hepatocyte growth. To further validate the role of PLCγ2 in hepatocytes, this study upregulated the expression of endogenous PLCγ2 gene, and investigated the anti‐proliferative role and the underlying mechanism of this gene in hepatocyte growth. As PLCγ2 appears to be critical for controlling hepatocyte growth and is potentially associated with the development of liver cancer, our research supplies a valuable reference for the study of PLCγ2 in liver tumourigenesis.
PLCγ2 is one member of the PLCγ subfamily that cleaves PIP2 to generate the secondary messenger DAG, subsequently triggering PKC activation.25 The PLCγ1 isoform, another member of the PLCγ subfamily, is ubiquitously expressed and is essential for embryonic development, while PLCγ2 is mainly located in the immune system and is critical in the development and maturation of haematopoietic lineage cells.26 Evidences has proved that the absence of PLCγ2 gene could cause the defects in the differentiation of haematopoietic lineage.27, 28 Additionally, the findings from some studies have suggested that PLCγ2 could induce apoptosis in some cancer cells, such as gastric cancer cells and the DT40 cell line.17, 23 Similarly, our recent study found that silencing PLCγ2 could significantly increase the cell viability of rat hepatocytes (Figure S2). Nonetheless, the biological basis for its effect and the exact role of this gene in hepatocyte growth has not been clarified.
In the present study, we first measured the expression of PLCγ2 in rat hepatocytes infected with recombinant adenovirus expressing PLCγ2 through semi‐quantitative RT‐PCR, RT‐qPCR and Western blot analysis, and found an obvious increase in the expression level of PLCγ2 when compared with the empty vector and the control groups. Moreover, an immunofluorescence assay showed that the fluorescent intensity of PLCγ2 in rat hepatocytes from the Ad‐PLCγ2 group was significantly higher than that in the Ad‐dull and the control groups, suggesting the successful overexpression of this gene in hepatocytes. Then, we investigated the effect of PLCγ2 on rat hepatocytes, and the result showed that PLCγ2 overexpression had a significant inhibitory effect on cell viability, which was consistent with the result of our earlier study. A possible explanation for the growth inhibition is cell cycle arrest or apoptosis induction. Interestingly, the results of our FCM analysis demonstrated that PLCγ2 overexpression caused the cell cycle arrest in G1 phase, leading to a decrease in proliferation ability. It is well known that G1 phase is critical for the preparation for DNA replication into S phase; therefore, the abnormity of this phase would irreversibly impact DNA replication in the cells and finally suppress the cell reproduction.29 At the same time, this study also showed the apoptosis‐inducing effect of PLCγ2 in rat hepatocytes. Flow cytometry analysis of apoptosis indicated a significantly increased apoptotic rate in the cells from the Ad‐PLCγ2 group compared to that in the Ad‐dull and the control groups. Caspases‐8 and ‐9 are the most upstream components in the caspase cascade. The former is activated by cell membrane death receptors (DR) to trigger the DR‐dependent extrinsic apoptotic pathway, whereas the latter is activated by cytochrome C released from the mitochondria to induce the mitochondria‐dependent intrinsic apoptotic pathway.30 Caspase‐3 is a downstream effector of the two apoptotic pathways and cleaves various cytoplasmic and nuclear substrates including DNA fragmentation factor and DNA‐dependent protein kinase,31 eventually leading to cell apoptosis once proteolytical activation occurs.32 Our results detected a marked enhancement of caspase‐3, ‐8 and ‐9 activities in the Ad‐PLCγ2 group compared with that in the Ad‐dull and the control groups, thus reconfirming that PLCγ2 may enhance apoptosis in rat hepatocytes.
MAPK is composed of a family of serine/threonine kinases that mediate various signalling pathways.33, 34 Amongst the MAPK family, p38 and JNK are considered as two important pathways involved in the induction of cell apoptotic death.35, 36 PKC is a family of enzymes that consist of at least 12 members and is involved in the phosphorylation of other proteins. The different PKC isoforms play specific roles in signal transduction. Evidence has proven that, among the PKC family members, PKCD participates in the activation of p38 and JNK.19, 20 In this current study, Western blot analysis revealed that when the phosphorylation level of PKCD was elevated due to the overexpression of PLCγ2, the levels of both p‐p38 and p‐JNK also significantly increased in rat hepatocytes in vitro. Furthermore, the inhibition of p38 and JNK MAPKs activities by the PKCδ inhibitor Go 6983 strongly suggested that PLCγ2 could induce the activation of p38 and JNK through phosphorylating PKCD.
Our further studies showed that after the suppression of PKCD activity by its inhibitor, the degree of apoptosis in rat hepatocytes was apparently improved in the Go 6983‐pre‐treated cells infected with Ad‐PLCγ2. In particular, MTT result showed that cell viability in hepatocytes was significantly recovered after Go 6983 pre‐treatment; BrdU incorporation and Hoechst staining assays indicated that compared with Ad‐PLCγ2 group, the proportion of DNA synthesis phase cells was obviously increased, and that of the apoptotic cells was significantly decreased, as observed by their normal morphology in the Go 6983‐pre‐treated group; FCM analysis also showed that there was a significant increase in the S‐phase cell fraction, while there was a marked decrease in the G1‐phase cell number and apoptotic rate after pre‐treatment with Go 6983. The above results from this present study reconfirm that PLCγ2‐induced cell proliferation inhibition and apoptosis were reversed by Go 6983, indicating that PLCγ2 is involved in hepatocyte apoptosis by activating the PKCD‐dependent p38 and JNK MAPK signalling pathways. These results reinforce the conclusion drawn from our previous findings obtained by RNA interference technology.
As far as we know, in this study we first confirmed that PLCγ2 expression can inhibit hepatocyte proliferation while promoting apoptosis in rat hepatocytes in vitro. We also demonstrated that the PLCγ2‐induced activation of the PKCD/p38 and PKCD/JNK MAPK signalling pathways may be involved in these biological processes. These results offer an experimental basis for further exploring the effects of PLCγ2 on hepatocytes in vivo. Moreover, our results indicate that PLCγ2 overexpression may prove to be a valuable tool for controlling the liver tumour growth.
CONFLICT OF INTEREST
The authors have declared that no competing interests exist.
Supporting information
ACKNOWLEDGEMENTS
This work was financially supported by the National Natural Science Foundation of China (No. 31401209), Ministry of Science and Technology of the People's Republic of China (MOST). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Chen X, Lv Q, Ma J, Liu Y. PLCγ2 promotes apoptosis while inhibits proliferation in rat hepatocytes through PKCD/JNK MAPK and PKCD/p38 MAPK signalling. Cell Prolif. 2018;51:e12437 10.1111/cpr.12437
REFERENCES
- 1. Arai M, Yokosuka O, Chiba T, et al. Gene expression profiling reveals the mechanism and pathophysiology of mouse liver regeneration. J Biol Chem. 2003;278:29813‐29818. [DOI] [PubMed] [Google Scholar]
- 2. Fukuhara Y, Hirasawa A, Li XK, et al. Gene expression profile in the regenerating rat liver after partial hepatectomy. J Hepatol. 2003;38:784‐792. [DOI] [PubMed] [Google Scholar]
- 3. Locker J, Tian JM, Carver R, et al. A common set of immediate‐early response genes in liver regeneration and hyperplasia. Hepatology. 2003;38:314‐325. [DOI] [PubMed] [Google Scholar]
- 4. Taub R. Liver regeneration: from myth to mechanism. Nat Rev Mol Cell Biol. 2004;5:836‐847. [DOI] [PubMed] [Google Scholar]
- 5. Fujiyoshi M, Ozaki M. Molecular mechanisms of liver regeneration and protection for treatment of liver dysfunction and diseases. J Hepatobiliary Pancreat Sci. 2011;18:13‐22. [DOI] [PubMed] [Google Scholar]
- 6. Gilgenkrantz H, de Collin l'Hortet A. New insights into liver regeneration. Clin Res Hepatol Gastroenterol. 2011; 35:. [DOI] [PubMed] [Google Scholar]
- 7. Nguyen LN, Furuya MH, Wolfraim LA, et al. Transforming growth factor‐beta differentially regulates oval cell and hepatocyte proliferation. Hepatology. 2007;45:31‐41. [DOI] [PubMed] [Google Scholar]
- 8. Apte U, Gkretsi V, Bowen WC, et al. Enhanced liver regeneration following changes induced by hepatocyte‐specific genetic ablation of integrin‐linked kinase. Hepatology. 2009;50:844‐851. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 9. Yamamoto Y, Ono T, Dhar DK, et al. Role of peroxisome proliferator‐activated receptor‐gamma (PPARgamma) during liver regeneration in rats. J Gastroenterol Hepatol. 2008;23:930‐937. [DOI] [PubMed] [Google Scholar]
- 10. Oakley F, Trim N, Constandinou CM, et al. Hepatocytes express nerve growth factor during liver injury: evidence for paracrine regulation of hepatic stellate cell apoptosis. Am J Pathol. 2003;163:1849‐1858. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 11. Asai K, Tamakawa S, Yamamoto M, et al. Activated hepatic stellate cells overexpress p75NTR after partial hepatectomy and undergo apoptosis on nerve growth factor stimulation. Liver Int. 2006;26:595‐603. [DOI] [PubMed] [Google Scholar]
- 12. Mizuguchi M, Yamada M, Kim SU, Rhee SG. Phospholipase C isozymes in neurons and glial cells in culture: an immunocytochemical and immunochemical study. Brain Res. 1991;548:35‐40. [DOI] [PubMed] [Google Scholar]
- 13. Rohacs T. Phosphoinositide regulation of non‐canonical transient receptor potential channels. Cell Calcium. 2009;45:554‐565. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 14. Kim D, Jun KS, Lee SB, et al. Phospholipase C isozymes selectively couple to specific neurotransmitter receptors. Nature. 1997;389:290‐293. [DOI] [PubMed] [Google Scholar]
- 15. Ichise H, Ichise T, Ohtani O, Yoshida N. Phospholipase C gamma2 is necessary for separation of blood and lymphatic vasculature in mice. Development. 2009;136:191‐195. [DOI] [PubMed] [Google Scholar]
- 16. Tomlinson MG, Woods DB, McMahon M, et al. A conditional form of Bruton's tyrosine kinase is sufficient to activate multiple downstream signaling pathways via PLC Gamma 2 in B cells. BMC Immunol. 2001;2:4. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17. Zhang B, Wu Q, Ye XF, Liu S, Lin XF, Chen MC. Roles of PLC‐gamma2 and PKCalpha in TPA‐induced apoptosis of gastric cancer cells. World J Gastroenterol. 2003;9:2413‐2418. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18. Grunicke HH, Spitaler M, Mwanjewe J, Schwaiger W, Jenny M, Ueberall F. Regulation of cell survival by atypical protein kinase C isozymes. Adv Enzyme Regul. 2003;43:213‐228. [DOI] [PubMed] [Google Scholar]
- 19. Chen C, Du J, Feng W, et al. β‐Adrenergic receptors stimulate interleukin‐6 production through Epac‐dependent activation of PKCδ/p38 MAPK signalling in neonatal mouse cardiac fibroblasts. Br J Pharmacol. 2012;166:676‐688. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20. Das J, Ghosh J, Manna P, Sil PC. Protective role of taurine against arsenic‐induced mitochondria‐dependent hepatic apoptosis via the inhibition of PKCdelta‐JNK pathway. PLoS ONE. 2010;5:e12602. [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
- 21. Kuo WH, Chen JH, Lin HH, Chen BC, Hsu JD, Wang CJ. Induction of apoptosis in the lung tissue from rats exposed to cigarette smoke involves p38/JNK MAPK pathway. Chem Biol Interact. 2005;155:31‐42. [DOI] [PubMed] [Google Scholar]
- 22. Mansouri A, Ridgway LD, Korapati AL, et al. Sustained activation of JNK/p38 MAPK pathways in response to cisplatin leads to Fas ligand induction and cell death in ovarian carcinoma cells. J Biol Chem. 2003; 278:19245‐19256. [DOI] [PubMed] [Google Scholar]
- 23. Takata M, Homma Y, Kurosaki T. Requirement of phospholipase C‐gamma 2 activation in surface immunoglobulin M‐induced B cell apoptosis. J Exp Med. 1995;182:907‐914. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 24. Lai SS, Zhao DD, Cao P, et al. PP2Acα positively regulates the termination of liver regeneration in mice through the AKT/GSK3β/Cyclin D1 pathway. J Hepatol. 2016;64:352‐360. [DOI] [PubMed] [Google Scholar]
- 25. Ibarguren M, López DJ, Escribá PV. The effect of natural and synthetic fatty acids on membrane structure, microdomain organization, cellular functions and human health. Biochim Biophys Acta. 2014;1838:1518‐1528. [DOI] [PubMed] [Google Scholar]
- 26. Wang D, Feng J, Wen R, et al. Phospholipase Cgamma2 is essential in the functions of B cell and several Fc receptors. Immunity. 2000;13:25‐35. [DOI] [PubMed] [Google Scholar]
- 27. Graham DB, Robertson CM, Bautista J, et al. Neutrophil‐mediated oxidative burst and host defense are controlled by a Vav‐PLCc2 signaling axis in mice. J Clin Invest. 2007;117:3445‐3452. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 28. Jakus Z, Simon E, Frommhold D, Sperandio M, Mo′csai A. Critical role of phospholipase Cc2 in integrin and Fc receptor‐mediated neutrophil functions and the effector phase of autoimmune arthritis. J Exp Med. 2009;206:577‐593. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 29. Nojima H. G1 and S‐phase checkpoints, chromosome instability, and cancer. Methods Mol Biol. 2004;280:3‐49. [DOI] [PubMed] [Google Scholar]
- 30. Lee KW, Chung KS, Seo JH, et al. Sulfuretin from heartwood of Rhus verniciflua triggers apoptosis through activation of Fas, Caspase‐8, and the mitochondrial death pathway in HL‐60 human leukemia cells. J Cell Biochem. 2012;113:2835‐2844. [DOI] [PubMed] [Google Scholar]
- 31. Casciola‐Rosen LA, Anhalt GJ, Rosen A. DNA‐dependent protein kinase is one of a subset of autoantigens specifically cleaved early during apoptosis. J Exp Med. 1995;182:1625‐1634. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 32. Derfuss T, Fickenscher H, Kraft MS, et al. Antiapoptotic activity of the herpesvirus saimiri‐encoded Bcl‐2 homolog: stabilization of mitochondria and inhibition of caspase‐3‐like activity. J Virol. 1998;72:5897‐5904. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 33. Shearn CT, Reigan P, Petersen DR. Inhibition of hydrogen peroxide signaling by 4‐hydroxynonenal due to differential regulation of Akt1 and Akt2 contributes to decreases in cell survival and proliferation in hepatocellular carcinoma cells. Free Radic Biol Med. 2012;53:1‐11. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 34. Wang Z, Zhang H, Xu X, et al. bFGF inhibits ER stress induced by ischemic oxidative injury via activation of the PI3K/Akt and ERK1/2 pathways. Toxicol Lett. 2012;212:137‐146. [DOI] [PubMed] [Google Scholar]
- 35. Wagner EF, Nebreda AR. Signal integration by JNK and p38 MAPK pathways in cancer development. Nat Rev Cancer. 2009;9:537‐549. [DOI] [PubMed] [Google Scholar]
- 36. Tiwary R, Yu W, Li J, Park SK, Sanders BG, Kline K. Role of endoplasmic reticulum stress in α‐TEA mediated TRAIL/DR5 death receptor dependent apoptosis. PLoS ONE. 2010;5:e11865. [DOI] [PMC free article] [PubMed] [Google Scholar]
Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials