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Molecular Cancer Research 4:747-758 (2006)
© 2006 American Association for Cancer Research

DNA Damage and Cellular Stress Responses

Reactive Oxygen Species Mediated Sustained Activation of Protein Kinase C {alpha} and Extracellular Signal-Regulated Kinase for Migration of Human Hepatoma Cell Hepg2

Wen-Sheng Wu1, Rong Kung Tsai2,3, Chung Hsing Chang2,4, Sindy Wang1, Jia-Ru Wu1 and Yu-Xun Chang1

1 Department of Medical Technology and 2 Graduate Institute of Medicine, Tzu Chi University; and Departments of 3 Ophthalmology and 4 Dermatology, Tzu Chi General Hospital, Hualien, Taiwan

Requests for reprints: Wu Wen-Sheng, Department of Medical Technology, Tzu Chi University, No. 701, Chung Yang Road, Section 3, Hualien 970, Taiwan. Phone: 3-8567285 ext 7512; Fax: 3-8571917. E-mail: wuws{at}mail.tcu.edu.tw


   Abstract
Top
 Notes
 Abstract
Introduction
 Result
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
The tumor promoter 12-O-tetradecanoylphorbol-13-acetate (TPA) can trigger growth inhibition, epithelial-mesenchymal transition (EMT)–like cell scattering, and migration of hepatoma cells HepG2 in a protein kinase C-{alpha} (PKC-{alpha})–dependent manner. Saikosaponin a, an ingredient of antitumorigenic Chinese herb Sho-Saiko-to, inhibited cell growth but did not induce EMT-like cell scattering and cell migration of HepG2. Saikosaponin a and TPA induced transient (for 30 minutes) and sustained (until 6 hours) phosphorylation of extracellular signal-regulated kinase (ERK), respectively. Generation of the reactive oxygen species (ROS) was induced by TPA, but not saikosaponin a, for 3 hours. As expected, scavengers of ROS, such as superoxide dismutase, catalase, and mannitol, and the thiol-containing antioxidant N-acetylcystein dramatically suppressed the TPA-triggered cell migration but not growth inhibition of HepG2. The generation of ROS induced by TPA was PKC, but not ERK, dependent. On the other hand, scavengers of ROS and N-acetylcystein also prevented PKC activation and ERK phosphorylation induced by TPA. On the transcriptional level, TPA can induce gene expression of integrins {alpha}5, {alpha}6, and ß1 and reduce gene expression of E-cahedrin in a PKC- and ROS-dependent manner. In conclusion, ROS play a central role in mediating TPA-triggered sustained PKC and ERK signaling for regulation of gene expression of integrins and E-cahedrin that are responsible for EMT and migration of HepG2. (Mol Cancer Res 2006;4(10):747–58)


   Introduction
Top
 Notes
 Abstract
 Introduction
Result
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
Metastasis is one of the most complicated and major pathologic processes responsible for poor prognosis of cancer patients. Studies of the molecular mechanisms for these processes are important for developing more effective anti-metastatic strategies. The migration of tumor cells is a prerequisite for tumor cell invasion and metastasis. Before migration, the cells often exhibit dramatic morphologic changes [i.e., epithelial–mesenchymal transition (EMT)]. EMT is also an important process during embryonic development (1). Cells that undergo EMT often lose epithelial adhesion and cytoskeleton components concurrent with acquiring expression of mesenchymal components and migratory phenotype. Mounting evidence suggests that EMT is an important event in the progression of many carcinomas (2). In spite of much research, the signaling mechanisms for triggering EMT and migration are not fully understood.

Two of the candidate signaling molecules involved in EMT and cell migration are protein kinase C (PKC) and mitogen-activated protein kinases. The PKC- and mitogen-activated protein kinase–mediated signal pathways coordinate complex physiologic and pathologic events, including cell cycle control, differentiation, neoangiogenesis, and metastasis (3-5). Cytokines, such as transforming growth factor-ß, hepatocyte growth factor, and fibroblast growth factor, may stimulate tumor invasion and metastasis via PKC and mitogen-activated protein kinases (4, 5).

PKC was initially recognized as a Ca2+, phospholipid-dependent serine-threonine kinase (6). To date, >12 PKC isozymes with different cofactors requirements have been found, including the conventional PKCs (cPKC-{alpha}, cPKC-ßI, cPKC-ßII, and cPKC-{gamma}), novel PKCs (nPKC-{delta}, nPKC-{varepsilon}, and nPKC-{eta}), and atypical PKCs (aPKC{zeta}; ref. 7). Various PKC isozymes have been shown to mediate cell migration in many distinct in vitro and in vivo systems (8, 9).

The mitogen-activated protein kinase signaling cascade, including extracellular signal-regulated kinase (ERK), c-Jun NH2-terminal kinase, and P38, has also been implicated in the migration of numerous cell types (10). Specific inhibitors for ERK signaling may inhibit the migration of cells in response to extracellular matrix and growth factors, such as fibroblast growth factor and epidermal growth factor (11-13). Moreover, inhibition of ERK by an antisense strategy was shown to prevent cell migration (14).

Many genes with different functions are known to be associated with migration, among which the cell-matrix interaction molecule integrin is the most important (15). Dramatic alterations in gene expression of various integrin subtypes are frequently observed during cell migration (16). Importantly, PKC may influence the effect of integrin by regulating either its gene expression or affinity toward the extracellular matrix (17). Moreover, the integrin-mediated, outside-in signaling also coordinates with several key signal pathways to regulate cell migration (18). Transcriptional factors, such as Snail, and cell-cell adhesion molecules, such as E-cahedrin, are involved in EMT (2). Up-regulation of Snail and down-regulation of E-cahedrin are frequently associated with EMT (2) and known to be characteristics of most invasive tumors (19).

Previously, we showed that the tumor promoter 12-O-tetradecanoyl phorbol-13-acetate (TPA) can activate the PKC{alpha}-mitogen-activated protein/ERK kinase (MEK)-ERK signaling cascade for induction of gene expression of the cyclin-dependent kinase inhibitors p15INK4b and p16INK4b that ultimately leads to growth suppression of human hepatoma cell HepG2 (20, 21). However, in addition to suppression of cell growth of HepG2, TPA may trigger cell scattering accompanied with the cell's turning from an epithelial to a fibroblast-like appearance. This dramatic phenotypic change is reminiscent of EMT. Thus, it is tempting to investigate whether TPA can trigger migration of HepG2 as well and whether it is mediated through the same signaling pathway as that for growth inhibition (i.e., the PKC{alpha}-MEK-ERK cascade).

In addition to MEK and ERK, one of the key candidate messengers involved in PKC-mediated signaling is the reactive oxygen species (ROS). ROS has been reported to mediate many cellular effects, including migration (22, 23). ROS-triggered signaling can be generated by TPA in a PKC-dependent manner (24) and may crosstalk with PKC-mediated pathway (25). In this study, we found that ROS plays a central role in sustained PKC-ERK signaling for TPA-induced EMT-like cell scattering and migration of hepatoma cells.


   Result
Top
 Notes
 Abstract
 Introduction
 Result
Discussion
 Materials and Methods
 Acknowledgements
 References
 
TPA-Triggered EMT and Migration of Hepatoma Cells Was PKC-{alpha} Dependent
In addition to inhibiting cell growth, TPA may trigger cell scattering of HepG2 (i.e., from a piled up epithelial appearance into flattened, fibroblastic appearance like EMT). This can be prevented effectively by cotreatment with the PKC inhibitor bisindolylmaleimides or the MEK inhibitor E-{alpha}-(amino-((4-aminophenyl)thio) methylene)-2-(trifluoromethyl) benzeneacetonitrile (Fig. 1A ). Furthermore, we found that 50 nmol/L TPA induced cell migration of HepG2 and another hepatoma cell (Huh7) within 24 hours in the Transwell migration assay system (Fig. 2A ). It seemed that Huh7 exhibited higher migratory activity than HepG2, probably because Huh7 itself acquired a fibroblastic-like morphology (data not shown), which benefits cell migration. Quantitative analysis for the migrated cell showed that TPA increased the migration of HepG2 by 10-fold at 24 hours, which can be suppressed by 90% by cotreatment with 5 µmol/L bisindolylmaleimides. (Fig. 3A ). This result indicated that TPA can trigger migration of HepG2 in a PKC-dependent manner.


Figure 1
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  		FIGURE 1. Effects of various inhibitors on TPA-induced, EMT-like morphologic change, and growth inhibition of HepG2. HepG2 cells were untreated (con) or treated with 50 nmol/L TPA coupled with 5 µmol/L bisindolylmaleimides (BIS), 20 mmol/L N-acetylcystein (NAC), 6 µmol/L MEK inhibitor (MEK in), or none (A and B), or 15 µg/mL saikosaponin a (B) for 48 hours. The cells were photographed under x100 magnification (A) or counted by hemocytometer (B). Cell numbers of the untreated cells were taken as 100%. A. Representative of at least four experiments. B. Obtained from average of at least six experiments. The growth inhibitory effect of TPA and saikosaponin a (compared with untreated cells) was evaluated by paired t test (P < 0.0001). The preventive effect of bisindolylmaleimides, N-acetylcystein, and MEK on TPA-induced growth inhibition was evaluated by ANOVA followed by Dunnet's test using the TPA-treated group (black column) as control. *, P < 0.05; **, P < 0.005; #, P > 0.5.

 

Figure 2
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  		FIGURE 2. TPA-induced migration of HepG2 and Huh7 and effects of antisense PKC oligodeoxynucleotides on TPA-induced migration of HepG2. A. HepG2 and Huh7 cells were untreated (control) or treated with TPA for 24 hours. B. HepG2 cells were untreated (control) or transfected with 4 µmol/L antisense PKC-{alpha} (AS PKC{alpha}) or sense PKC-{alpha} (S PKC{alpha}) for 24 hours followed by TPA treatment for 24 hours. Cell migration assays were done. The migrated cells were pictured under x40 magnification. The results shown in (A) and (B) were replicated at least twice.

 

Figure 3
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  		FIGURE 3. Quantitative determination of the effects of various inhibitors and antisense PKC oligodeoxynucleotides on TPA-induced migration of hepatoma cells. A. HepG2 cells were untreated, treated with 15 µg/mL saikosaponin a, or 50 nmol/L TPA coupled with 5 µmol/L bisindolylmaleimides, 10 or 20 mmol/L N-acetylcystein, or none for 24 hours. B. HepG2 cells were untreated, or treated with 50 nmol/L TPA coupled with 200 units/mL superoxide dismutase (SOD), 100 units/mL catalase, 20 mmol/L mannitol, or none for 24 hours. C. HepG2 cells were untreated or transfected with 4 µmol/L antisense PKC-{alpha}, PKC-ßII, or PKC-{varepsilon} for 24 hours followed by TPA treatment for 24 hours. Relative quantitation of the migrated cells was done. The migration of TPA-treated samples was taken as 100% in each experiment. Each result was obtained from average of at least three separate experiments. The migratory effect induced by TPA was analyzed by t test (P < 0.0001). The preventive effect of various inhibitor on TPA-induced migration was evaluated by ANOVA followed by Dunnet's test using the TPA-treated group (black column) as control. *, P < 0.05; **, P < 0.005; #, P > 0.5.

 
To further identify the PKC isotype responsible for this effect, antisense oligodeoxynucleotides of PKC isozymes, including PKC-{alpha}, PCK-ßII, and PCK-{varepsilon}, used in our previous study were employed. We have shown that PKC-{alpha} but not PKC-ßII or PKC-{varepsilon} can be activated by TPA, and that antisense PKC-{alpha} oligodeoxynucleotide can efficiently block translation of PKC-{alpha} protein in HepG2 (21). Transfection of 4 µmol/L antisense PKC-{alpha} oligodeoxynucleotide but not sense PKC-{alpha} oligodeoxynucleotide for 24 hours dramatically blocked the TPA-induced, EMT-like cell scattering (data not shown) and migration of HepG2 (Fig. 2B). The suppressive effect of antisense PKC-{alpha} oligodeoxynucleotide on TPA-induced HepG2 cell migration was compared with that of antisense PKC-ßII and PKC-{varepsilon} oligodeoxynucleotides by quantitative migration assay. Although 76% of TPA-induced HepG2 cell migration was abolished in cells transfected with 4 µmol/L antisense PKC-{alpha} oligodeoxynucleotide, only 13% and 10% of that were prevented in cell transfected with 4 µmol/L antisense PKC-ßII and PKC-{varepsilon} oligodeoxynucleotide, respectively (Fig. 3C). Thus, PKC-{alpha}, but not the other PKC isozymes, was responsible for TPA-triggered EMT and migration of HepG2.

Sustained but not Transient ERK Activation Was Associated with TPA-Triggered EMT and Migration
Because PKC-{alpha} is also required for mediating TPA-induced growth inhibition of HepG2 (21), we further investigated whether the signaling mechanisms downstream of PKC-{alpha} are the same for TPA-induced growth inhibition, EMT, and migration. It seems that saikosaponin a, an active ingredient of the antitumorigenic Chinese herb Sho-Saiko-to (TJ-9), is suitable for addressing this issue. We previously showed that saikosaponin a can suppress cell growth of HepG2 (20, 26). As shown in Fig. 1B, treatment of the cells with 15 µg/mL saikosaponin a for 48 hours may reduce proliferation of HepG2 by 50%. After treatment with saikosaponin a, the cells turned into a flattened monolayer growth (obviously as a result of the decrease of cell number) at 24 hours; however, cell scattering and the fibroblastic appearance that were typically induced by TPA were not observed (Fig. 4A ). Consistently, saikosaponin a did not enhance cell migration of HepG2 as TPA did (Fig. 4B and Fig. 3A). One of the saikosaponin a–induced molecular effects that can be easily distinguished from that induced by TPA is the duration of ERK activation. As shown in Fig. 5A , phosphorylation of ERK was induced by 50 nmol/L TPA at 1 hour (by about 3.6-fold) and can be sustained until 6 hours (2.8-fold), whereas induction of ERK phosphorylation was observed in saikosaponin a–treated cells at 30 minutes (by 2.5-fold) and return near to basal level within 1 to 4 hours. Thus, a rapid and transient activation of ERK may be required for TPA-induced growth inhibition (as in the case of saikosaponin a), whereas sustained activation of ERK is required for TPA-triggered EMT-like scattering and migration.


Figure 4
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  		FIGURE 4. Differential cellular effects of TPA and saikosaponin a on HepG2. HepG2 cells were treated with 15 µg/mL saikosaponin a or 50 nmol/L TPA for 24 hours. The cells were photographed (x100; A) or assayed for migration (x40; B). The result was replicated at least five times.

 

Figure 5
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  		FIGURE 5. Time course for saikosaponin a or TPA-induced ERK phosphorylation and effects of various inhibitors on TPA-induced ERK phosphorylation in HepG2 cells. HepG2 cells were untreated (con), treated with 15 µg/mL saikosaponin a for 0, 1, 2, and 4 hours or 50 nmol/L TPA for 1, 2, 4, 6, and 8 hours (A). HepG2 cells were untreated (con; B, C, and D); treated with 50 nmol/L TPA coupled with 5 µmol/L bisindolylmaleimides, 10 or 20 mmol/L N-acetylcystein, and 0.2% DMSO or none for 30 minutes (B); 50 nmol/L TPA coupled with 10 to 50 nmol/L DPI, 0.5 mmol/L dithiotheritol, 0.5 mmol/L Trolox, 10 to 20 mmol/L mannitol, 200 units/mL superoxide dismutase, or none for 30 minutes (C); and 50 nmol/L TPA coupled with 100 units/mL catalase, 200 units/mL SOD, or none for 30 minutes (D). Western blot of phosphorylated ERK (pERK) was done. Western blot of ERK was used for normalization of protein. The ratio of the intensity of phosphorylated ERK versus ERK for each sample was indicated below each lane. The ratio of the control samples were taken as 1.0. A and B. Representative of three experiments with coefficient of variance of 8.0% to 15%.

 
ROS Is the Key Amplifier for Sustained PKC-ERK Signaling Required for TPA-Induced EMT and Migration
To further delineate the downstream signaling for TPA-induced EMT-like scattering and cell migration in HepG2, the role of ROS was investigated. ROS-mediated signaling was not only associated with cell migration and tumor progression (27) but also implicated in sustained ERK activation for some specific cellular effects (28, 29). Several ROS scavengers, such as mannitol (a hydroxyl radical scavenger), superoxide dismutase (catalyzing the degradation of O2–), and catalase (catalyzing auto-oxidoreduction of H2O2), and the thiol (-SH)–containing antioxidant, such as N-acetylcystein, were employed to investigate whether the TPA-induced molecular and cellular effects are mediated by ROS. Cotreatment of the cells with 20 mmol/L N-acetylcystein, efficiently suppressed the TPA-induced EMT-like cell scattering (Fig. 1A). Furthermore, cotreatment of the cells with 10 to 20 mmol/L N-acetylcystein dose-dependently prevented the TPA-triggered cell migration by 70% to 85% (Fig. 3A). In contrast, growth inhibition of HepG2 induced by 50 nmol/L TPA was not significantly prevented by 20 mmol/L N-acetylcystein, whereas it was prevented by 5 µmol/L bisindolylmaleimides and 6 µmol/L MEK inhibitor by 95% and 90%, respectively (Fig. 1B). Similarly, pretreatment of 10 mmol/L mannitol, 200 units/mL superoxide dismutase, and 100 units/mL catalase also prevented TPA-induced cell migration by 78%, 90%, and 80%, respectively (Fig. 3B), but did not prevent TPA-induced cell growth inhibition (data not shown). Taken together, the results suggested that ROS was specifically required for TPA-triggered EMT and cell migration but was not important for growth inhibition of HepG2. On the other hand, ROS generation was detected by in vivo assay using cell-permeable 2',7'-dichlorofluorescin diacetate as a ROS probe. As shown in Fig. 6A , whereas ROS generation cannot be induced by 15 µg/mL saikosaponin a at 0.5 and 1 hour, it can be induced by 50 nmol/L TPA treatment for 30 minutes to 3 hours in HepG2 cells. The TPA-induced ROS generation within this period can be prevented dramatically by 5 µmol/L bisindolylmaleimides but not 6 µmol/L MEK inhibitor. In the cell labeled with the nonpermeable, oxidation-insensitive analogue of 2',7'-dichlorofluorescin diacetate (2',7'-dichlorofluorescin) no fluorescence was detected in all the treatment groups (data not shown). For quantitative purposes, ROS assays were also done by directly measuring fluorescence of oxidized 2',7'-dichlorofluorescin in the cell lysate. As shown in Fig. 6B, a substantial basal level of ROS was detected in control cells, and a 2.5-fold increase of ROS was observed in TPA-treated cells. Cotreatment with 5 µmol/L bisindolylmaleimides or 20 mmol/L N-acetylcystein prevented the TPA-induced ROS production by 90% and 80%, respectively, whereas only 6.0% was prevented by 6 µmol/L MEK inhibitor. These results indicated that the TPA-induced activation of PKC, but not ERK, was required for ROS generation. We further investigated whether ROS are required for ERK activation. Western blot of phosphorylated ERK (Fig. 5B) showed that 5 µmol/L bisindolylmaleimides reduced the TPA-induced ERK phosphorylation at 30 minutes by about 65%, as was shown in our previous report (21). Importantly, the TPA-induced ERK phosphorylation was prevented by N-acetylcystein (20 mmol/L) by about 40% (Fig. 5B). Furthermore, 20 mmol/L mannitol, 200 units/mL superoxide dismutase, 0.5 mmol/L dithiotheritol (which is another thiol-containing antioxidant), and 0.5 mmol/L Trolox (which is a hydrophilic analogue of the antioxidant {alpha}-tocopherol) suppressed the TPA-induced ERK phosphorylation by about 49%, 52%, 37%, and 43%, respectively (Fig. 5C). In addition, 100 units/mL catalase also suppressed ERK phosphorylation induced by TPA by about 31% (Fig. 5D). Taken together, these results indicated that both PKC and ROS were required for ERK activation, and it seems that ROS is downstream of PKC and upstream of ERK in the TPA-triggered signal pathway. Although this is in accord with several previous reports showing that ROS are the signaling mediator downstream of PKC (30), there are reports suggesting that PKC may also be activated by ROS to mediate many cellular effects (25). Interestingly, ROS may amplify PKC signaling such as observed in highly glucose-treated human peritoneal mesothelial cells (31). Thus, we sought to clarify whether ROS may activate PKC to sustain the PKC-MEK-ERK pathway. As the nonradioactive PKC assay shown in Fig. 7A , the induction of PKC activity gradually increased over 0.5 to 3 hours followed by a decline at 5 hours. Cotreatment with 20 mmol/L N-acetylcystein suppressed the TPA-induced PKC activity at 30 minutes and 1 and 3 hours by 85% to 90%. Moreover, PKC activity induced by TPA at 1 hour was also dramatically suppressed by mannitol (20 mmol/L), catalase (100 units/mL), and superoxide dismutase (200 units/mL; Fig. 7B). These results indicated that ROS were indeed required for TPA-induced PKC activation. Thus, it seems that ROS is not only downstream but also upstream of PKC, serving as the signaling amplifier for sustained PKC-ERK cascade that mediate EMT-like cell scattering and cell migration induced by TPA in HepG2. ROS was previously reported to be generated by cytosolic oxidase, such as NADPH oxidase, using molecular oxygen as a substrate; therefore, we suggested that activation of NADPH oxidase may be required for the TPA-triggered signaling in hepatoma cells. As was expected, diphenyleneiodonium chloride (10 and 50 nmol/L), an inhibitor of NADPH oxidase, can not only significantly attenuate the TPA-induced ERK phosphorylation by about 37% (Fig. 5C) but also dramatically suppress the TPA-induced cell migration of HepG2 and Huh7 (Fig. 8 ).


Figure 6
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  		FIGURE 6. Time course for TPA and saikosaponin a–induced ROS generation and quantitative analysis for effects of various inhibitors on TPA-induced ROS generation. A. HepG2 cells were untreated (control), treated with 15 µg/mL saikosaponin a for 0.5 and 1 hour (left) or treated with 50 nmol/L TPA coupled with 5 µmol/L bisindolylmaleimides, 20 mmol/L N-acetylcystein, 6 µmol/L MEK inhibitor, or none for 0.5, 1.0, and 3.0 hours (right). In vivo ROS generation was done. B. HepG2 cells were untreated or treated with 50 nmol/L TPA coupled with 5 µmol/L bisindolylmaleimides, 20 mmol/L N-acetylcystein, or 6 µmol/L MEK inhibitor, or none for 30 minutes. ROS generation was assayed by measuring the amount of fluorescent oxidized 2',7'-dichlorofluorescin (DCF) product in each cell lysate normalized with protein. Relative fluorescence (RFU) for each sample was determined, with the TPA-treated sample taken as 100%. The result in (A) and (B) was replicated three times. The significant increase of relative fluorescence in the TPA-treated sample was analyzed by t test (P < 0.0001). The preventive effect of various inhibitor on TPA-induced ROS generation was evaluated by ANOVA followed by Dunnet's test using the TPA-treated group (black column) as control. *, P < 0.05; **, P < 0.005; #, P > 0.5.

 

Figure 7
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  		FIGURE 7. Time course of PKC activity induced by TPA in HepG2 cell and the prevention of TPA-induced PKC activity by N-acetylcystein. HepG2 cells were untreated (control; A and B), treated with 50 nmol/L TPA for 30 minutes and 1, 3, and 5 hours or 50 nmol/L TPA coupled with 20 mmol/L N-acetylcystein for 30 minutes and 1 and 3 hours (A) and 50 nmol/L TPA coupled with 0.5 mmol/L DTT, 20 mmol/L mannitol 100 units/mL catalase, 200 units/mL SOD, and none for 1 hour (B). Nonradioactive PKC assay was done as described in Materials and Methods. Representative of three separate experiments. u-PKC, unphosphorylated PKC; p-PKC, phosphorylated PKC.

 

Figure 8
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  		FIGURE 8. Prevention of TPA-induced migration of HepG2 and Huh7 by diphenyleneiodonium chloride. HepG2 and Huh7 cells were untreated (control), treated with 50 nmol/L TPA or TPA plus 50 nmol/L DPI for 24 hours. Cell migration assay was performed as described in Fig. 2. Representative of three separate experiments.

 
Prevention of EMT by Bisindolylmaleimides and N-Acetylcystein at Time of Post-TPA Treatment
To further examine the importance of sustained ROS{leftrightarrow}PKC signaling for TPA-induced EMT-like cell scattering and cell migration, we also investigated whether bisindolylmaleimides and N-acetylcystein could prevent these cellular effects even after the cells were treated with TPA. As shown in the previous section, TPA-induced EMT-like cell scattering can be prevented by cotreatment with 5 µmol/L bisindolylmaleimides or 20 mmol/L N-acetylcystein (Fig. 1A). When each inhibitor was added at 30 minutes and 1 and 3 hours after TPA treatment, their preventive effects on the TPA-induced EMT and migration were the same as when they were cotreated with TPA (Fig. 9A and B ). In contrast, the preventive effect of bisindolylmaleimides on TPA-induced growth inhibition of HepG2 decreased by 20%, 90%, and 95% when bisindolylmaleimides was added 30 minutes and 1 and 3 hours after TPA treatment, respectively, compared with the cells cotreated simultaneously with bisindolylmaleimides and TPA (Fig. 9C). Once again, these results can be explained simply by that an early (30 minutes) and transient ERK activation was responsible for TPA-induced growth inhibition, whereas the ROS{leftrightarrow}PKC->ERK signaling described herein must be maintained at least until 3 hours for the cell to enter EMT and migration.


Figure 9
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  		FIGURE 9. Prevention of TPA-induced EMT and migration but not growth inhibition of HepG2 by bisindolylmaleimides and N-acetylcystein at time of post-TPA treatment. HepG2 cells were untreated (control) or treated with 50 nmol/L TPA coupled with 5 µmol/L bisindolylmaleimides or 20 mmol/L N-acetylcystein added at 0, 0.5, 1, and 3 hours after TPA treatment. Cells were photographed after 48 hours under x100 magnification (A), or quantitative analysis of cell migration was done after 24 hours as described in Fig. 5B. C. HepG2 cells were untreated or treated with 50 nmol/L TPA coupled with 5 µmol/L bisindolylmaleimides added at 0, 0.5, 1, and 3 hours after TPA treatment. Cell number was determined after 48 hours as described in Fig. 1A, with the number of control cell taken as 100%. A. Representative of at least five experiments. B and C. Average obtained from three experiments. The migratory and growth inhibitory effects induced by TPA were analyzed by t test (P < 0.0001). The effect of various inhibitor on TPA-induced migration and growth inhibition at various time point was evaluated by ANOVA followed by Dunnet's test using the TPA-treated group (black column) as control. *, P < 0.05; **, P < 0.005; #, P > 0.5.

 
Regulation of Integrin and E-Cahedrin Gene Expression via PKC{leftrightarrow}ROS Pathway
Previous reports showed that TPA may regulate gene expressions of integrins (32, 33), the important cell-matrix adhesion molecules associated with cell migration. Semiquantitative reverse transcription-PCR analysis (Fig. 10A ) showed that gene expressions of the integrin subunits {alpha}5 and ß1 were induced by 50 nmol/L TPA by ~6.0- and 3.0-fold, respectively, at 12 hours in both HepG2 and Huh7. Cotreatment with 5 µmol/L bisindolylmaleimides or 20 mmol/L N-acetylcystein blocked the TPA-induced gene expression of integrin {alpha}5 and ß1 in both cells by 85% to 95%. Surprisingly, integrin {alpha}6 was induced by TPA in Huh7, but not HepG2, by ~5.0-fold, an effect that can also be prevented by bisindolylmaleimides and N-acetylcystein. Interestingly, the TPA-induced cell migration of both HepG2 and Huh7 can be suppressed by blocking antibody for integrin ß1 (Fig. 11 ), by 80% to 90% in both cells estimated by quantitative migration (data not shown). These results further implied that the integrin-mediated pathway was involved in TPA-induced migration of hepatoma cells. Whether TPA regulates expression of EMT-related genes was also investigated. As shown by reverse transcription-PCR in Fig. 10B, E-cahedrin was significantly suppressed by TPA by ~65% at 12 hours, which was also prevented by 5 µmol/L bisindolylmaleimides and 20 mmol/L N-acetylcystein, by ~85% and 60%, respectively. Consistently, significant induction of Snail, a known transcriptional repressor of E-cahedrin (2), was observed after TPA treatment for 2 hours. Maximal induction of Snail by TPA was observed at 3 hours and returned to basal level at 24 hours (Fig. 10C). Taken together, these results further suggested the role of ROS{leftrightarrow}PKC signaling cascade in mediating the expression of the genes involved in EMT and cell migration.


Figure 10
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  		FIGURE 10. Prevention of TPA-regulated gene expression of integrins {alpha}5, {alpha}6, and ß1 and E-cahedrin by bisindolylmaleimides and N-acetylcystein and induction of Snail gene expression by TPA in hepatoma cells. HepG2 and Huh7 cells (A) and HepG2 cells (B) were untreated (control) or treated with 50 nmol/L TPA coupled with 5 µmol/L bisindolylmaleimides or 20 mmol/L N-acetylcystein for 12 hours. C. HepG2 cells were treated with 50 nmol/L TPA for 0.5, 1, 2, 3, and 12 hours. Reverse transcription-PCR of integrins {alpha}5, {alpha}6, and ß1 (A); E-cahedrin (B); Snail (C); and glyceraldehyde-3-phosphate dehydrogenase (GAPDH; internal control) were done. A to C. Reproducible three times.

 

Figure 11
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  		FIGURE 11. Prevention of TPA-induced HepG2 cell migration by integrin ß1 antibody. HepG2 cells were untreated (control) or treated with 50 nmol/L TPA coupled with none or 10 µg/mL integrin ß1 antibody. Cell migration assay was done after 24 hours as described in Fig. 2. Representative of at least three experiments.

 

   Discussion
Top
 Notes
 Abstract
 Introduction
 Result
 Discussion
Materials and Methods
 Acknowledgements
 References
 
Significance of PKC-{alpha} in Mediating EMT-Like Cell Scattering, Cell Migration, and Growth Suppression of Hepatoma Cell
Although the role of PKC in regulating the migratory activity of tumor cells, such as bladder carcinoma, breast cancer cells, and hepatoma cells, induced by several cytokines has been well established (8, 9, 34-37), which PKC isozymes are responsible for tumor cell migration are not fully explored. Here, we showed that PKC-{alpha} was required for migration of hepatoma cell HepG2. The most intriguing issue is that this same PKC isozyme was also responsible for TPA-induced growth arrest (21). This revealed that the two different cellular processes can be mediated by the same upstream signaling molecule. There might be some mechanistic link between cell cycle arrest and migration/invasion of tumor cells. For example, metastasis of tumor cell induced by cytokines such as transforming growth factor-ß is often accompanied by growth inhibition (38, 39). Interestingly, a lot of studies showed that PKC might crosstalk with or even mediate the effect of transforming growth factor-ß signaling (40, 41). In addition, a recent report showed that G1-S arrest by overexpression of the cyclin-dependent kinase inhibitor p16INK4a confers anoikis resistance in MCF-10A mammary epithelial cells, which might contribute to early stages of metastasis (42). Because we also found that p16INK4a was induced by TPA (20, 26), the role of p16INK4a in TPA-induced cell cycle arrest and migration is worthy of further investigation.

Role of Sustained versus Transient ERK Activation in TPA-Induced Cell Migration
Although a previous report suggested that early, transient ERK activation was required for signaling chemotaxis for PC12 pheochromocytoma cells (43), mounting evidence shows that sustained ERK activation was associated with cell migration (44, 45). In the present study, the possibility that sustained ERK activation is required for TPA-induced cell migration was validated in two ways. First, as shown in Fig. 5A, ERK activation was maintained for 6 hours in TPA-treated HepG2, in which both growth inhibition and cell migration were observed. On the other hand, ERK activation was induced transiently (~30 minutes) in saikosaponin a–treated HepG2 cells, in which only growth inhibition was observed. Second, we found that bisindolylmaleimides and N-acetylcystein may prevent EMT and migration of HepG2 even at 30 minutes to 3 hours post-TPA treatment (Fig. 9A and B). This result strongly suggested that long-term ROS{leftrightarrow}PKC->ERK signaling is required for the cells to prepare for EMT and migration. Such long-term signaling might be beneficial for triggering gene regulation relevant to both cellular processes, as were reported previously. For example, L1-triggered sustained activation of the ERK was required for the expression of motility- and invasion-associated gene products, including the integrin ß3 subunit and small GTPases (44). Furthermore, a previous report showed that sustained Raf-MEK-ERK activated by oncogenic Ras was required for induction of ß3 integrin transcription, whereas transient activation of Raf-MEK-ERK signaling by growth factors and mitogens had no such effect (46). It is also likely that a more extended period of time is required for induction of the transcriptional factors associated with EMT and migration than that associated with growth inhibition. For example, in Fig. 10C, we found that it took at least 3 hours for TPA to induce maximal gene expression of Snail, which is the key transcriptional suppressor of E-cahedrin (2). In contrast, as shown in our previous report (20), it took only 30 minutes for TPA to induce activator protein, which has been reported to be associated with p15INK4b/p16INK4a expression and growth inhibition (47, 48). Alternatively, sustained ERK activation may be required for induction of multiple transcriptional systems for regulation of genes relevant to EMT and cell migration. For example, Est-1, one of the important transcriptional regulators associated with tumor cell invasion, must cooperate with other transcriptional factors to mediate gene expression regulated by TPA (49, 50).

ROS Trigger the Sustained PKC-ERK Signaling for Cell Migration
ROS production has been shown to be coupled with the sustained activation of the ERK signaling pathway for a variety of cellular effects, including apoptosis (51) and phagocytosis (52). Furthermore, a recent report showed that long-term oxidative stress may induce invasive potential of mammary epithelial cells (53). In addition, activations of PKC, ROS, and ERK were shown to be responsible for oleic acid– and angiotensin II–induced vascular smooth muscle cell migration (23). In our study, several lines of evidences strengthened the specific, essential role of ROS in triggering EMT and cell migration. First, ROS may be induced by TPA for at least 3 hours. Furthermore, the ROS scavengers superoxide dismutase, catalase, and mannitol and the thiol-containing antioxidant N-acetylcystein may prevent TPA-induced cell migration (Fig. 3A and B). Second, ROS generation was not only dependent on but also required for PKC activation (Figs. 6 and 7), implying its role for amplification of PKC signal. Third, we have shown that PKC-{alpha} and ROS were required for TPA-induced ERK phosphorylation (Fig. 5B; ref. 21), and that overexpression of active PKC-{alpha} was sufficient for triggering ERK phosphorylation in HepG2 cell (54). Taken together, it is very probable that ROS play essential role for sustained PKC-ERK activation induced by TPA for EMT and migration of HepG2. Whether ROS is also involved in mediating migration and invasion of hepatoma cell in vivo is worthy of further investigation.

Mechanisms for TPA-Induced ROS Generation
How ROS is generated by TPA is another important issue. ROS was previously reported to be generated by cytosolic oxidase using molecular oxygen as a substrate, but lately, the mitochondria were also suggested to be a source of this oxidant (55, 56). There is evidence showing that PKC-dependent activation of NADPH oxidase was required for the generation of ROS (57). We have shown that TPA-induced ROS generation was PKC dependent (Fig. 6), and diphenyleneiodonium chloride, an inhibitor of NADPH oxidase, can significantly prevent TPA-induced ERK phosphorylation and migration of HepG2 and Huh7 (Fig. 5B and Fig. 8), suggesting that TPA-induced ROS generation is dependent on NADPH oxidase. It is possible that the TPA-induced ROS generation also be derived from the integrin-mediated pathway. Integrin engagement is associated with cell adhesion and migration (58, 59), and intracellular ROS may be generated via mitochondria after integrin engagement (56). Our results showed that several integrin subunits are induced by TPA, among which integrin ß1 is known to be closely associated with cell migration (60). Interestingly, TPA-induced cell migration can be blocked efficiently by integrin ß1 antibody in both HepG2 and Huh7 (Fig. 11). This result implies that integrin-mediated signaling may crosstalk with the PKC-ERK pathway to mediate TPA-induced EMT and migration of hepatoma cells, probably via ROS generation.

In conclusion, we delineated the signal mechanism for TPA-induced EMT and migration compared with TPA and saikosaponin a–induced growth inhibition in HepG2 as outlined in Fig. 12 .


Figure 12
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  		FIGURE 12. Summary for signaling mechanisms for TPA and saikosaponin a–triggered EMT, migration, and growth inhibition of HepG2. TPA-induced EMT/migration of HepG2 was mediated by sustained ROS{leftrightarrow}PKC{alpha}->ERK for induction of Snail and suppression of E-cahedrin gene expression that were associated with EMT. There might be some other transcriptional factor, such as Est-1, for induction of integrin and other migration-related gene expression. The source of ROS generation was not certain. On the other hand, TPA and saikosaponin a may trigger transient ERK activation and activator protein-1 induction followed by p15INK4b/p16INK4a expression and growth inhibition.

 

   Materials and Methods
Top
 Notes
 Abstract
 Introduction
 Result
 Discussion
 Materials and Methods
Acknowledgements
 References
 
Cell Culture and Chemicals
The cultured condition and methods for cell number determination of hepatoma cell HepG2 and Huh7 were the same as described in our previous report (20). The morphology of the cells was observed for EMT-like cell scattering using an inverted microscope (Axiovert 25; Carl Zeiss, Oberkochen, Germany) under x100 magnification.

TPA, 2',7'-dichlorofluorescin diacetate, 2',7'-dichlorofluorescin, N-acetylcystein, mannitol, catalase, superoxide dismutase, dithiotheritol, and diphenyleneiodonium chloride were purchased from Sigma (St. Louis, MO). Bisindolylmaleimides, Trolox, and specific MEK1/2 inhibitor [E-{alpha}-(amino-((4-aminophenyl)thio) methylene)-2-(trifluoromethyl) benzeneacetonitrile] were from Calbiochem (La Jolla, CA). Saikosaponin a was from the Pharmaceutical Industry Technology and Development Center (Taipei, Taiwan).

Evaluation of TPA-Induced EMT of HepG2
After TPA treatment, HepG2 turned from piled-up epithelial into a flattened, fibroblastic appearance, a morphologic change like that of EMT. For quantitating the cells that exhibited EMT-like morphologic change in Figs. 1A, 4, and 9A, the number of the fibroblast-like cells, with clear cell-cell boundary, were counted under a x100 magnification field of phase-contrast microscope. Eight fields per well (of 24-well plate) were scored and averaged (data not shown). Each experiment was repeated and validated by two other investigators who did not know the treatment groups before microscopic observation.

Migration Assay
Migration assays were done using a 24-well Transwell migration insert (Nalge Nunc International, Rochester, NY). Cells were seeded at 5 x 104 per upper chamber in complete medium with or without addition of TPA or TPA coupled with various inhibitors. Cells that had migrated to the underside of the insert membrane were stained with 0.3% crystal violet. The cells in the upper side of the insert membrane were rubbed with a cotton swab. For quantitation, the migrated cells on the underside were counted based on the number of the cells under a x100 magnification field. Eight fields per insert were scored and averaged.

Transfection of PKC Oligodeoxynucleotide
The sense PKC-{alpha} and antisense PKC-{alpha}, PKC-ßII, and PKC-{varepsilon} oligodeoxynucleotides with phosphorothioate linkage throughout the entire oligodeoxynucleotide molecule were the same as used in previous reports (61-63): antisense PKC-{alpha}, 5'-CATGGTYCCCCCCAACCACC-3' (Y = T or C; antisense sequence against 20 nucleotides upstream of the AUG codon); sense PKC-{alpha}, 5'-GGTGGTTGGGGGGRACCATG-3' (R = A or G; ref. 61); antisense PKC-ßII, 5'-CGCAGCCGGGTCAGCATC-3' (62); antisense PKC-{varepsilon}, 5'-GCCATTGAACACTACCAT-3' (63). The PKCs (oligodeoxynucleotides) were transfected into HepG2 cells (at final concentration of 4 µmol/L oligodeoxynucleotide) for 24 to 36 hours using the effectene transfection kit (Life Technologies, Grand Island, NY) according to the manufacturer's instructions.

Reverse Transcription-PCR
Reverse transcription-PCR for semiquantitation of gene expressions of integrins {alpha}5, {alpha}6, and ß1; E-cahedrin; and Snail was done as described in our previous report (20). Briefly, single-stranded cDNAs were synthesized by avian myeloblastosis virus transcriptase with oligo-dT primer (Life Technologies) followed by PCR. Glyceraldehyde-3-phosphate dehydrogenase was amplified for 20 cycles to normalize the amount of cDNA in each sample. The primers used were as follows: integrin {alpha}5: forward, 5'-GTTCCAAGAGCAGCAAGGAC-3' and reverse, 5'-GGGTTGCAAGCCTGTTGTAT-3' (1,002 bp); integrin {alpha}6: forward, 5'-GGCGGTGTTATGTCCTGAGT-3' and reverse, 5'-AGGGAACCAACAGCAACATC-3' (910 bp); integrin ß1: forward, 5'-CAGACATTTACATTAAAATTCAAG-3' and reverse, 5'-ACAGATGTACTGAAGAATAACCTC-3' (1,002 bp); E-cahedrin: forward, 5'-CCATGGATAACCAGAATA-3' and reverse, 5'-GGATCCTTAATTGACCTCAGAAGATGCACTA TCTAA-3' (650 bp); Snail: forward, 5'-AAGCTTCCATGGCGCGCTTCTTTCCTCGTCAGGAAGCCC-3' and reverse, 5'-GGATCCTCAG C GGGGACATCCTGAGCAGCCGGACTCTTG-3' (598 bp); glyceraldehyde-3-phosphate dehydrogenase: forward, 5'-CGGAGTCAACGGATTTGGTCGTAT-3' and reverse, 5'-AGCCTTCTCCATGGTGGTGAAGAC-3' (306 bp).

PCR Condition
After initial denaturation at 95°C for 5 minutes, amplification was done by 35 cycles of denaturation at 94°C for 40 seconds, annealing at 55°C for 1 minute, and extension at 72°C for 40 seconds. For quantitation, the gels were scanned, and the intensity of each band was estimated with gel digitizing software Quantity One (Bio-Rad, Hercules, CA).

Western Blotting
Western blotting was done as described in our previous report (20). Briefly, the cells were lysed in buffer containing 50 mmol/L Tris at pH 7.4, 50 mmol/L NaCl, 0.1% Triton X-100, 0.1% SDS, 0.3 mmol/L sodium orthovanadate, 50 mmol/L NaF, 1 mmol/L dithiotheritol, 10 µg/mL leupeptin, and 5 µg/mL aprotinin. Proteins were separated and transferred to polyvinylidene difluoride membrane. Membranes were blocked in 20 mmol/L Tris (pH 7.6) and 250 mmol/L NaCl containing 5% dry milk and probed with antibodies against phosphorylated ERK1/2 and ERK1/2 (Santa Cruz Biotechnology, Santa Cruz, CA). After incubation with alkaline phosphatase–conjugated goat anti-mouse or goat anti-rabbit 2nd antibody, proteins were visualized with nitroblue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate for color development. For quantitation, the blots were scanned, and the intensity of each band was estimated with gel digitizing software Quantity One (Bio-Rad).

Assay for ROS Generation
Intracellular ROS production was monitored by oxidation of permeable dye 2',7'-dichlorodihydrofluorescein diacetate, which reacts with ROS to form the fluorescent product 2',7'-dichlorodihydrofluorescein. The cells were incubated with 20 µmol/L 2',7'-dichlorofluorescin diacetate for 1 hour and washed twice with PBS, and the fluorescent cell was observed and photographed under a fluorescent microscope Olympus IX71 (Olympus Corp., Tokyo, Japan). The nonpermeable, oxidation-insensitive analogue of 2',7'-dichlorofluorescin diacetate (2',7'-dichlorofluorescin) was also used for the negative control. Alternatively, the relative intensity of fluorescence in the cell lysate was determined by a fluorescence spectrophotometer (Fluoroskan Ascent FL, Labsystems Corp., Waltham, MA) at excitation and emission wavelength of 485 and 510 nm, respectively. Data were normalized on the amount of total protein in each sample.

Fractionation of Cellular Extract
Briefly, the cells were suspended in hypotonic buffer [10 mmol/L Tris (pH 7.4), 50 mmol/L NaCl, 0.3 mmol/L sodium orthovanadate, 50 mmol/L NaF, 1 mmol/L dithiotheritol, 10 µg/mL leupeptin, and 5 µg/mL aprotinin] and incubated for 30 minutes at 4°C. The cell suspensions were homogenized and centrifuged at 2,500 rpm for 3 minutes. The supernatants were then subjected to ultracentrifugation at 25,000 rpm for 1 hour. The resulting supernatants were obtained as cytosolic fractions. The pellets were dissolved in lysis buffer containing 50 mmol/L Tris (pH 7.4), 50 mmol/L NaCl, 1% Triton X-100, 0.3 mmol/L sodium orthovanadate, 50 mmol/L NaF, 1 mmol/L dithiotheritol, 10 µg/mL leupeptin, and 5 µg/mL aprotinin. After centrifugation at 15,000 rpm for 30 minutes, the supernatants were obtained as particulate fractions.

PKC Assay
Nonradioactive PKC assays were done with PepTag PKC assay kit (Promega, Madison, WI) according to manufacturer's instructions. Briefly, 20 µg protein in particulate fraction of the cellular extract was incubated with PepTag PKC substrate and lipid activator at 30°C for 30 minutes followed by inactivation at 95°C for 5 minutes. After electrophoresis in 0.8% agarose gel for 15 minutes, the phosphorylated PKC and unphosphorylated PKC substrate migrating toward the anode and cathode, respectively, were detected under UV light.

Statistics
The quantitative assessment for the effect of TPA on cell growth (Fig. 1B), migration (Fig. 3), and EMT (Figs. 1A, 4, and 9A) was simply validated by paired t test between control and TPA-treated groups. The preventive effects of various inhibitors on TPA-induced cellular effects were evaluated by one-way ANOVA followed by Dunnet's post hoc comparisons using the TPA-treated group as control. All the statistics was done with WINKS 4.70 statistic software. Significance was inferred when P < 0.05.


   Acknowledgements
Top
 Notes
 Abstract
 Introduction
 Result
 Discussion
 Materials and Methods
 Acknowledgements
References
 
We thank the National Science Council in Taiwan for financial support and C-Z. Chung BS for technical assistance.


   Notes
Top
 Notes
Abstract
 Introduction
 Result
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 4/ 7/06; revised 6/17/06; accepted 7/20/06.


   References
Top
 Notes
 Abstract
 Introduction
 Result
 Discussion
 Materials and Methods
 Acknowledgements
 References
 

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