Hydroxycamptothecin Induces Apoptosis of Human Tenon's Capsule Fibroblasts by Activating the PERK Signaling Pathway

Abstract

Purpose.: Hydroxycamptothecin (HCPT) has been proven to induce apoptosis in fibroblasts. In this study, we investigated whether the PRKR-like ER kinase (PERK) pathway is implicated in apoptotic signaling of human Tenon's capsule fibroblasts (HTCFs) by HCPT.

Methods.: Normal and PERK-knockdown HTCFs were used in this study. Apoptosis was determined by the cell viability assay, Annexin V/propidium iodide (PI) dual-staining, cell cycle analysis in HTCFs treated with HCPT in various doses and for various durations. Endoplasmic reticulum (ER) stress markers and sensor proteins were detected by Western blot analysis. Mitochondrial dysfunction was measured by detecting the mitochondrial membrane potential (ΔΨm) and measuring the expression of cytochrome c (cyt c).

Results.: HCPT induced apoptosis in the HTCFs, which was characterized as decreased cell viability and sub-S fraction of the cell cycle and increased apoptosis rate by Annexin V/PI dual-staining. The activity levels of caspase-3 and caspase-9 were significantly increased and were accompanied by cytosolic release of cyt c and decreased ΔΨm in response to HCPT. Treatment with HCPT increased the expression of glucose-regulated protein 78 (GRP78), phospho-PERK, activating transcription factor 6 (ATF6), phosphoinositol-requiring kinase 1 (IRE1), C/EBP homologous protein (CHOP), Bax, and phospho-c-Jun N-terminal kinase (JNK) and decreased the expression of Bcl-2. Knockdown of PERK attenuates HCPT-induced apoptosis in HTCFs, dependent upon both ER stress and the mitochondrial apoptotic pathway.

Conclusions.: This study suggests that the ER stress response and mitochondrial dysfunction are involved in apoptosis induced by HCPT in HTCFs, which might be mediated by PERK; thus, this study offers new insight into preventing postoperative scarring via treatment with HCPT.

Introduction

Filtering surgery is a major method for managing glaucoma, and advanced trabeculectomy requires a treatment modality that would minimize fibrosis and yield better outcomes from glaucoma filtration surgery (GFS). ¹ Scarring in the surgical area is the major cause of the failure of trabeculectomy and other glaucoma surgeries. Therefore, the single intraoperative application of antimetabolite agents, including mitomycin-C (MMC) and 5-fluorouracil (5-FU), are widely used to inhibit the formation of subconjunctival scars by preventing fibroblast proliferation. ^2–4 However, treatment with MMC and 5-FU could induce serious damage to normal tissues, including wound leaks, hypotony, macular degeneration, corneal epithelial damage, endophthalmitis, and hyphema. ^5,6  

Hydroxycamptothecin (HCPT) is one of a series of camptothecin derivatives with fewer side effects and is widely used to treat various cancers. ^7,8 Unlike MMC and 5-Fu, HCPT is a cell cycle–specific agent that mainly arrests cells during DNA synthesis (S phase) and inhibits the proliferation of cells. ⁹ Moreover, HCPT reduces epidural fibrosis by preventing the proliferation of fibroblasts in scar tissue ^10,11 and induces apoptosis of fibroblasts by downregulating their metabolism. ¹² Caspase-3 is one of the most important caspases for the execution of apoptosis in various cell types and is activated by initiator caspases, such as caspases 8 and 9, in response to proapoptotic signals. ¹³ Recently, it was reported that HCPT may have beneficial roles in filtering surgery by triggering apoptosis in human Tenon's capsule fibroblasts (HTCFs). ¹⁴ However, the possible mechanisms that are involved require further investigation. 

Both mitochondrial dysfunction and endoplasmic reticulum (ER) stress–mediated cell death pathways are involved in toxic chemical (arsenic)-induced apoptosis in mammalian cells by increasing the Bax/Bcl-2 ratio. ¹⁵ The Bcl-2 family plays a central role in the apoptosis pathway by regulating the release of cytochrome c (cyt c) and the activation of caspases. ^16,17 Moreover, the Bcl-2 family is downstream of C/EBP homologous protein (CHOP) and is considered to be a critical mediator of apoptosis in the ER stress–mediated cell death pathways. 

The ER is responsible for the synthesis, folding, assembly, modification, and transport of nascent proteins. Aggregation of unfolded/misfolded proteins activates the unfolded protein response (UPR) ^18,19 or ER stress, with three downstream signals, including PRKR-like ER kinase (PERK), inositol-requiring kinase 1 (IRE1), and activating transcription factor 6 (ATF6). Activation of the UPR induces an adaptive response to restore normal ER function, and prolonged stress or a failed adaptive response leads to an accumulation of glucose-regulated protein 78 (GRP78/BiP), and activation of IRE1, ATF6, and PERK. PERK phosphorylates eukaryotic initiation factor-2 (eIF2α) and induces activating transcription factor 4 (ATF4) and CHOP expression, downregulates Bcl-2 expression, and upregulates Bax. ^20–22 CHOP expression is a consequence of PERK phosphorylation; therefore, we hypothesized that HCPT activates the apoptotic pathway in HTCFs through the activation of PERK. The purpose of this study was to investigate the effects of PERK on the apoptotic pathway in human HTCFs. 

Materials and Methods

Cell Culture

A primary cell line of HTCFs was established from subconjunctival Tenon capsules isolated from patients during pterygium surgery. All of the patients gave their informed consent before surgery. This study was approved by the Nanjing Medical University Medicine ethics committee in accordance with the provisions of the Declaration of Helsinki. The HTCFs were cultured at 37°C with 5% CO₂ in Dulbecco's modified Eagle's medium (DMEM; Gibco, Grand Island, NY) with L-glutamine (2 mM), 10% fetal bovine serum (FBS; Gibco), penicillin (100 IU/mL), and streptomycin (100 μg/mL) (PS; Thermo, Rockford, IL). Cells in the exponential growth phase were used between passages 4 and 7 in all of the experiments. 

HCPT Treatment

HTCF monolayers that had been seeded into 24-well plates, 96-well plates, or 10-cm dishes overnight were washed with PBS (pH 7.4) and treated with HCPT (Santa Cruz Biotechnology, Santa Cruz, CA) in various concentrations and for various durations. The controls were treated with 5-minute applications of PBS only. After treatment, the cells were immediately washed three times with PBS and were maintained in the growth medium for subsequent experiments. 

Cell Viability

Cell viability was measured using a Cell Counting Kit-8 (CCK-8; Dojindo, Tokyo, Japan). The HTCFs were cultured in triplicate in 96-well plates and were treated with 0.06, 0.25, 1, and 4 mg/L HCPT. The HTCFs were maintained in DMEM for 12, 24, or 48 hours, as described in our previous report, ¹⁴ and were further incubated with 10 ul WST-8 (2-[2-methoxy-4-nitrophenyl]-3-[4-nitrophenyl]-5-[2,4-disulfonyl]-2H-tetrazolium; Dojindo) for 1 hour at 37°C. Cells that stained positively with WST-8 were considered viable cells and were expressed as a percentage compared with the control cells. 

Flow Cytometric Analysis of Apoptosis

Annexin V/PI dual-staining and cell cycle analysis was used to detect cell apoptosis. The HTCFs were plated in 60-mm wells (2 mL, 1 × 10⁶ cells/well) and incubated for 24 hours at 37°C. After treatment with 1 mg/L HCPT for 24 hours, the detached and adherent cells were collected and washed twice with ice cold PBS buffer. Cells were then resuspended in binding buffer at 1 × 10⁶ cells/well and incubated with Annexin V-FITC and PI (BD Biosciences, Singapore) for double-staining according to the manufacturer's protocol. Prior to analysis, the mixture was incubated in the dark for 15 minutes at room temperature. For cell cycle analysis, the collected cells were fixed at −20°C in ice cold 70% ethanol overnight. After two washes with PBS, the cells were stained with PI, and the data were collected and analyzed using a Beckman Coulter FC500 Flow Cytometry System with CXP Software (Beckman Coulter, Fullerton, CA). 

Mitochondrial Membrane Potential Transition (ΔΨm)

HTCF mitochondria were incubated with 0.2 μg/mL JC-1 (5,5,6,6-tetrachloro-1,1-3,3′-tetra ethyl benzimidazolocarbocyanine iodide) dye (BD Biosciences) at 37°C for 10 minutes. After incubation, the cells were washed three times with PBS, collected, and analyzed by flow cytometry. 

Western Blot Analysis

HTCFs were lysed in 50 mM Tris (pH 8.0), 5 mM EDTA, 150 mM NaCl, 0.5% Nonidet P-40, and 1 mM phenylmethylsulfonyl fluoride (PMSF). The protein concentration was determined by the BCA Protein Assay Kit (Thermo). Equal amounts (25 μg/lane) of total proteins were subjected to electrophoresis on a 10% SDS-PAGE. Following electrophoresis, the proteins were electrotransferred onto polyvinylidene difluoride membranes (Millipore, Bedford, MA). The membranes were then blocked with 5% skim milk in tris-buffered saline and Tween 20 (TBST) at room temperature for 2 hours and subsequently incubated with primary antibodies against GRP78, CHOP, cleaved caspase-3, PERK, phospho-PERK, Bax, Bcl-2, cyt c (Cell Signaling Technology, Danvers, MA), phospho-JNK, JNK1, β-Actin (Santa Cruz Biotechnology), ATF6, phospho-IRE1α, and IRE1α (Abcam, Hong Kong, China) at 4°C overnight, with dilutions ranging from 1:500 to 1:1000. The membranes were washed three times in TBST and incubated with horseradish peroxidase-conjugated goat anti-mouse or anti-rabbit IgG (diluted 1:5000) for 1 hour. The immune complexes were visualized by fluorography enhanced by the electrochemiluminescence system (Millipore). 

Cytochrome c Release From Mitochondria

To isolate the mitochondrial and cytosolic fractions, the cells were lysed in buffer containing 10 mM Tris–HCl (pH 7.4), 10 mM NaCl, and 12.5 mM EDTA (Thermo), and the cell extract was centrifuged at 600g for 5 minutes to pellet the nuclei. The supernatant was centrifuged at 16,000g for 20 minutes to pellet the mitochondrial fraction. The supernatant contained the cytosolic fraction. The expression of cyt c in the mitochondrial and cytosolic fractions was detected by Western blot analysis. 

Quantitative Real-Time PCR Analysis

HTCFs were harvested at various time points after treatment with HCPT. Subsequently, the total RNA for all procedures was extracted using TRIzol reagent (Invitrogen, Grand Island, NY), according to the manufacturer's recommendations. First-strand cDNA was synthesized using random primers (Invitrogen) (Table). Real-time quantitative RT-PCR reactions were performed using an ABI Step One (Applied Biosystems, Grand Island, NY) with the following cycle conditions: 95°C for 1 minute, followed by 40 cycles of denaturation at 95°C for 15 seconds, and annealing and extension at 60°C for 1 minutes using the SYBR Green PCR Master Mix (Applied Biosystems). The relative mRNA levels of all of the genes were normalized to the levels of glyceraldehyde-3-phosphate dehydrogenase (GAPDH). 

Table

 

Sequence of Primer

Primer	Sequence
CHOP	Forward: 5′-ACCAAGGGAGAACCAGGAAACG-3′
	Reverse: 5′-TCACCATTCGGTCAATCAGAGC-3′
GRP78	Forward: 5′-CGGGCAAAGATGTCAGGAAAG-3′
	Reverse: 5′-TTCTGGACGGGCTTCATAGTAGAC-3′
GAPDH	Forward: 5′-GGGCTCTCCAGAACATCATCC-3′
	Reverse: 5′-GTCCACCACTGACACGTTGG-3′

Generation of Stable PERK-Silenced HTCF Cell Lines

Stable HTCFs with reduced levels of PERK were produced by targeting PERK mRNA with shorthairpinRNA (shRNA) that was generated using the lentiviral expression vector pLKO.1 (Shanghai Genechem Co. Ltd., Shanghai, China) at a multiplicity of infection (MOI) of 20 and puromycin selection (3 μg/mL for 7 days) using polybrene (2 μg/mL) to increase the transduction efficiency for subsequent experiments. 

Statistical Analysis

The data are expressed as the mean ± SD of triplicate experiments. Statistical differences between the treatment groups were analyzed by one-way ANOVA followed by the Dunnett test or Student's t-test using Statistical Package for the Social Sciences (SPSS) 13.0 software (Statistical Product and Service Solutions, Chicago, IL). Statistical significance was set at P less than 0.05. 

Results

Apoptotic Effect of HCPT on HTCFs

Exposing HTCFs to HCPT led to time- and dose-dependent increases in the number of dead cells, as assessed by the CCK-8 assay. Compared with the controls, exposure to HCPT for 24 hours at concentrations of 0.06, 0.25, 1, and 4 mg/L led to survival rates of 85.8%, 74.6%, 72.1%, and 60.0% of the cells, respectively (Fig. 1A). Time-dependent kinetics of cell viability upon HCPT treatment demonstrated that 12 hour of exposure to 1 mg/L did not cause significant cytotoxicity, but the effects of exposure became significant after 24 and 48 hours in the HTCFs (Fig. 1B). Consistent with the findings of the CCK-8 assay, Annexin V/PI dual-staining demonstrated significantly increased apoptosis (from 2.88%–15.98%) in cells treated with 1 mg/L HCPT for 24 hours (Figs. 1C, 1D). The cell cycle was analyzed by flow cytometry in HTCFs stained with PI (Figs. 1E, 1F). In contrast with the control culture, exposure to HPCT (1 mg/L) for 24 hours increased the S fraction of the HTCFs (from 5.09%–13.6%). These data indicated that HCPT treatment decreased the viability of the HTCFs via apoptosis. 

Figure 1

 

HCPT caused HTCF apoptosis. (A, B) Dose- (left) and time-dependent (right) effects on HTCF viability were determined by CCK-8 assay at 24 hours with various doses or up to 48 hours at the 1 mg/L concentration. (C, D) Apoptotic cells were stained by Annexin V/PI dual-staining and (E, F) cell cycle analysis was performed by flow cytometry after staining with PI in cells treated with 1 mg/L HCPT for 24 hours. The data are presented as the mean ± SD of three independent experiments (*P < 0.05, **P < 0.01).

Activation of Caspase-3, Caspase-9, and Phospho-JNK With Mitochondrial Dysfunction in HCPT-Treated HTCFs

To determine the signaling pathway in apoptosis of HTCFs, cells were maintained in the presence of HCPT for 24 hours at concentrations of 0.06, 0.25, 1, and 4 mg/L. The lysate was used to measure cleaved caspase-3, levels of upstream caspase-9, and phospho-JNK by Western blot analysis (Figs. 2A, 2B). Activation of caspase-3 began at 1 and attained its peak at 4 mg/L, and active caspase-9 showed an increase significantly at 4 mg/L. The expression of phospho-JNK began to be detectable when cells were treated with HCPT at 1 mg/L HCPT (P < 0.01). To test whether the activation of caspase-3 resulted from mitochondrial dysfunction, ΔΨm, and the cytosolic release of cyt c was also measured. 

Figure 2

 

HCPT induced mitochondrial dysfunction in the HTCFs. (A, B) Expression levels of caspase-3, caspase-9, JNK, and phospho-JNK were determined by Western blot analysis in cells after treatment with 0.06, 0.25, 1, and 4 mg/L HCPT for 24 hours. β-actin was selected as the loading control. (C, D) Mitochondrial ΔΨm was determined by JC-1 staining and was detected by flow cytometry after treatment with 1 mg/L HCPT for 24 hours. (E, F) Cytosolic release of cytochrome c was measured by Western blot analysis in the cytosolic and mitochondrial fractions after treatment with 0.06, 0.25, 1, and 4 mg/L HCPT for 24 hours. The mitochondrial fraction was verified using an anti–voltage-dependent anion channel (VDAC) antibody. The data are presented as the mean ± SD of three independent experiments (*P < 0.05, **P < 0.01).

We determined the ΔΨm of the HTCFs by JC-1 staining after treatment with 1 mg/L HCPT for 24 hours (Figs. 2C, 2D). At a high mitochondrial membrane potential, the mitochondrial JC-1-aggregates would be detected under red wave length (FL2-H). When mitochondrial membrane potential is low, the cytoplasmic JC-1 monomer will be detected under green fluorescence channel (FL1-H). The ratio of Y Mean/X Mean was 4.99 (Y Mean = 1926.45, X Mean = 385.97) in control group, while 1.18 (Y Mean = 1026.18, X Mean = 872.05) in 1 mg/L group (Fig. 2C). ΔΨm was decreased significantly with 1 mg/L HCPT for 24 hours (Fig. 2D). After fractionation of the cell lysate into mitochondrial and cytosolic parts, two fractions were separately used to measure the expression of cyt c by Western blot analysis. Cyt c in the cytosolic fraction was markedly increased at 1 mg/L and 4 mg/L. Mitochondrial cyt c was decreased significantly at 4 mg/L (Figs. 2E, 2F). These data indicated that HCPT activated the caspase-9, caspase-3, and JNK in the apoptosis signaling of HTCFs, decreased the ΔΨm of HTCFs, and eventually induced mitochondrial apoptosis. 

ER-Stress Induced by HCPT

We determined whether ER stress is induced by HCPT by testing the expression of GRP78 and CHOP, all of which are markers of ER stress. As detected by quantitative real-time PCR, the mRNAs level of CHOP increased at 1 and 4 mg/L after treatment with HCPT. And the mRNAs level of GRP78 increased at 0.25 mg/L and reached a maximum at 4 mg/L (Fig. 3A). Furthermore, HCPT treatment markedly decreased antiapoptogenic Bcl-2 expression, whereas the drug increased apoptogenic Bax and CHOP expression in a dose-dependent manner. The ration of Bax/Bcl-2 was increased significantly at 1 and 4 mg/L (Figs. 3B, 3C). After treatment with 1 mg/L HCPT, the GRP78 expression was markedly increased at 4 hours after treatment, and sustained until 12 hours. Treatment of the cells with 1 mg/L HCPT markedly induced the activation of IRE-1 and PERK, but decreased the expression of ATF-6 (Figs. 3D, 3E). These data indicated that HCPT induced ER stress, which eventually increased the Bax/Bcl-2 ratio in the apoptosis signaling of the HTCFs. 

Figure 3

 

HCPT induced ER stress in the HTCFs. (A) The mRNA expression levels of the ER stress markers CHOP (left) and GRP78 (right) were analyzed by real-time quantitative PCR detecting system in cells after treatment with 0.06, 0.25, 1, and 4 mg/L HCPT for 24 hours. Relative mRNA levels of all of the genes were normalized to the levels of GAPDH. (B, C) The expression of CHOP and the downstream Bcl-2 and Bax were determined by Western blot analysis in cells after treatment with 0.06, 0.25, 1, and 4 mg/L HCPT for 24 hours. β-actin was used as the loading control. (D, E) The expression levels of ER stress markers, including GRP78, phospho-PERK, PERK, ATF6 p90, phospho-IRE1, and T-IRE1, were determined by Western blot analysis in cells after treatment with 1 mg/L HCTP for 2, 4, 8, 12, and 24 hours. β-actin was used as the loading control. The data are presented as the mean ± SD of three independent experiments (*P < 0.05, **P < 0.01).

Suppression of HCPT Induced Apoptosis in HTCFs Via Knockdown of PERK

PERK has been implicated in the expression of CHOP, Bal-2, and Bax during ER stress in cells. Therefore, we pretreated HTCFs with lentiviral vectors bearing the PERK target gene (shRNA). 

After silencing the expression of PERK (Fig. 4C), both of PERK^−/− and PERK^+/+ HTCFs were treated with 1 mg/L HCPT for 24 hours. Flow cytometric analysis of apoptosis demonstrated that the apoptotic rate was 11.15% and 6.5% in PERK^+/+ group and PERK^−/− group, respectively. Thus, knockdown of PERK inhibited apoptosis significantly in cells treated with 1 mg/L HCPT for 24 hours (Figs. 4A, 4B). Meanwhile, the expressions of the ER stress markers (GRP78, CHOP) and the ratio of downstream Bax/Bcl-2 were decreased significantly in PERK^−/− group (Figs. 4D, 4E; *P < 0.05, **P < 0.01). 

Figure 4

 

PERK knockdown attenuated the HCPT-induced ER stress and apoptosis in the HTCFs. (A, B) Apoptosis in normal and PERK-knockdown HTCFs was measured by Annexin V/PI dual staining and analyzed by flow cytometry after treatment with 1 mg/L HCPT for 24 hours. (C) HTCFs were transfected with a PERK-mediated lentivirus. Silenced PERK expression was verified by Western blot analysis. (D, E) The expression levels of the ER stress markers GRP78 and CHOP and the downstream Bcl-2 and Bax were determined by Western blot analysis in PERK-knockdown and control HTCFs after treatment with 1 mg/L HCPT for 24 hours. Significant differences were detected PERK knockdown groups and control groups. β-actin was used as the loading control. The data are presented as the mean ± SD of three independent experiments (*P < 0.05, **P < 0.01).

Mitochondrial Dysfunction and Mitochondrial Apoptosis in HCPT-Treated HTCFs is PERK-Dependent

After application of 1 mg/L HCPT for 24 hours, the knockdown of PERK decreased the loss of ΔΨm. The ratio of Y Mean/X Mean was 0.93 (Y Mean = 693.40, X Mean = 748.91) in PERK^+/+ group, while 1.91 (Y Mean = 987.21, X Mean = 516.46) in PERK^−/− group. The difference was significant (Fig. 5A). PERK knockdown attenuated the mitochondrial dysfunction (Figs. 5B, 5C). The release of cyt c from mitochondria was enhanced as the level of cyt c in cytosol was markedly increased, whereas the level of in mitochondrial cyt c was decreased significantly (*P < 0.05, **P < 0.01). Similarly, HCPT-induced expression of activated caspase-3, caspase-9, and p-JNK (the downstream mitochondrial apoptosis markers) were also attenuated by PERK knockdown in the HTCFs (Figs. 5D, 5E). ²³  

Figure 5

 

PERK knockdown attenuated HCPT-induced mitochondrial dysfunction in HTCFs. (A) Mitochondrial ΔΨm in normal and PERK-knockdown HTCFs was determined by JC-1 staining and was detected with flow cytometry after treatment with 1 mg/L HCPT for 24 hours. (B, C) Cytosolic release of cyt c in normal and PERK-knockdown HTCFs was measured by Western blot analysis in the cytosolic and mitochondrial fractions after treatment with 1 mg/L HCPT for 24 hours. The mitochondrial fraction was verified via an anti-VDAC antibody. (D, E) Expression levels of cleaved caspase-3, caspase-9, phospho-JNK, and JNK were determined by Western blot analysis in control and PERK-knockdown HTCFs after treatment with 1 mg/L HCPT for 24 hours. β-actin was used as the loading control. The data are presented as the mean ± SD of three independent experiments (*P < 0.05, **P < 0.01).

Discussion

Camptothecin (CPT) is a natural alkaloid that was isolated in the 1960s from extracts of Camptotheca acuminate, a native plant in China ^24–26 ; The drug's analogue, 10 HCPT, exhibits strong antitumor activity by inducing apoptosis in cancer cells with less toxicity both in experimental animals and human clinical evaluations. ^8,27–29 CPT has been proven to effectively inhibit scar formation in the filtering bleb, and complications in CPT-treated subjects are much less common than in 5-Fu–treated subjects. ³⁰ Recently, HCPT has been used to reduce epidural fibrosis by preventing proliferation and inducing apoptosis in fibroblasts, thus, exhibiting a potentially beneficial effect in filtering surgery. ^11,14,30  

In this study, in vitro experiments using fibroblasts were performed to clarify the precise and detailed mechanism of action of HCPT as an antifibrosis agent. HCPT was found to inhibit the proliferation of fibroblasts in a dose- and time-dependent manner. After treatment with 1 mg/L HCPT for 24 hours, flow cytometry analysis revealed that the number of apoptotic cells increased from 2.88% to 15.98%, and cell growth was arrested in the S phase. These data suggest that the HCPT-induced inhibition of proliferation in fibroblasts is a consequence of S phase arrest and apoptosis, consistent with previously published data from tumor cells. ⁹  

Apoptosis is an important biologic process in many systems and can be triggered by a variety of stimuli. Apoptosis generally occurs via intrinsic and extrinsic pathways. The extrinsic pathway, also known as the death receptor pathway, involves the initiation of apoptosis through ligation of cell–surface death receptors (such as the tumor necrosis factor receptor [TNFR] family) located on the plasma membrane. ³¹ In contrast, the intrinsic pathway is triggered by proapoptotic signals that result in the disruption of mitochondrial functions, leading to the production of excessive reactive oxygen species (ROS), ³² disruption of ΔΨm, and release of cyt c, and apoptosis-inducing factor (AIF); those events subsequently activate the caspase cascade and apoptosis. ^33–36 Our results demonstrated that HCPT treatment efficiently caused mitochondrial dysfunction in fibroblasts, as evidenced by decreased ΔΨm and release of cyt c from the mitochondria into the cytosol. Western blot analysis assay of apoptotic-related proteins (including cleaved caspase-3, caspase-9, and phospho-JNK) further verified that the antiproliferation effects of HCPT on fibroblasts were mediated via the mitochondria-mediated apoptosis pathway, as confirmed by our earlier studies. ¹⁴  

Bcl-2 family members have important function in regulating of almost all known pathways of apoptosis. ³⁷ Bcl-2 and Bax are the members of Bcl-2 subfamilies, which act as antiapoptotic protein and proapoptotic protein, respectively. ^38,39 Bcl-2 proteins, which are antiapoptotic proteins, could protect the mitochondria function during apoptotic process, whereas the proapoptotic family members, such as Bax, appear to disrupt mitochondria homeostasis. ^40,41  

Bcl-2 is also regulated by the ER and nuclear membranes. ^42,43 The ER stress could activate UPR, upregulate CHOP expression, downregulate Bcl-2 expression, and upregulate Bax expression. An increase in the Bax/Bcl-2 ratio contributes to the promotion of apoptosis. ^44,45 In our study, HCPT elicited a time- and dose-dependent activation of UPR, accompanied by increased CHOP expression and increased Bax/Bcl-2 ratios in fibroblasts, strongly suggesting that HCPT induces apoptosis in fibroblasts by activation of ER stress. 

PERK is key regulator of CHOP transcription. Activation of PERK leads to eIF2α phosphorylation and ATF4 synthesis, which, together with C/EBP-β, binds to the composite site and transactivates the CHOP promoter. ^46–48 In this study, we found that the activation of PERK occurred during the HCPT-mediated upregulation of CHOP in the fibroblasts. The depletion of PERK led to decreases in the expression of CHOP, the Bax/Bcl-2 ratio, and the apoptosis rate in HCPT-treated fibroblasts, indicating the significance of this pathway. 

ER and mitochondria are mainly connected in the mitochondria-associated ER membranes (MAMs), and PERK is a component of the MAMs. ^49,50 Under oxidative stress, leakage of Ca²⁺ from the ER lumen increases, causing rapid depletion of the ER Ca²⁺ stores and ultimately leading to mitochondrial dysfunction and apoptotic cell death. ⁵¹ PERK knockdown cells display disturbed ER morphology and Ca²⁺ signaling and significantly weakened ER–mitochondria contact sites. ²³ In our case, the depletion of PERK also led to increased ΔΨm, decreased release of cyt c from the mitochondria into the cytosol, and attenuated phosph-JNK and caspase signaling (including caspase-3 and caspase-9) in HCPT-treated fibroblasts. These results indicate that PERK also plays a key role in the mitochondrial pathway of HCPT-induced apoptosis in fibroblasts. There is compelling evidence that an increase in the Bax/Bcl-2 ratio can disrupt the mitochondrial membrane potential. Combined with our results, this phenomenon strongly suggests that HCPT induces apoptosis of fibroblasts via the mitochondrial and ER stress–signaling pathway. PERK is crucial in these apoptotic signaling pathways. ⁵² The reason for the activation of PERK and ER stress that is induced by HCPT in fibroblasts still needs to be explored. 

In our previous study, we demonstrated that activation of caspase-3 resulted from activation of intrinsic caspase-9 with mitochondrial dysfunction in HCTFs exposed to HCPT. ¹⁴ The results of this study demonstrated that HCPT caused activation of mitochondrial dysfunction and ER stress–mediated cell apoptosis pathways in fibroblasts. Moreover, depletion of PERK effectively attenuated the ER stress and reversed the mitochondrial dysfunction and caspase activity in the HCPT-treated fibroblasts. Based on these results, we believe that PERK activation plays a crucial role in HCPT-induced fibroblasts apoptosis. To the best of our knowledge, no literature exists that investigates the role of PERK activation in HCPT-induced fibroblast death. 

In conclusion, our current study suggests that the ER stress response and mitochondrial dysfunction is involved in HCPT-induced apoptosis in fibroblasts, which may be mediated by PERK; thus, this investigation offers new insight into preventing postoperative scarring via treatment with HCPT. 

Acknowledgments

Supported by a grant from the National Natural Science Foundation of China (no. 81271001). 

Disclosure: X. Yin, None; H. Sun, None; D. Yu, None; Y. Liang, None; Z. Yuan, None; Y. Ge, None 

Footnotes

 XY and HS contributed equally to the work presented here and should therefore be regarded as equivalent authors.