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Article |
Essential role of protein kinase C 
in transducing a motility signal induced by superoxide and a chemotactic peptide, fMLP
Correspondence to Yoshiro Niitsu: niitsu{at}sapmed.ac.jp
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Under various pathological conditions, including infection, malignancy, and autoimmune diseases, tissues are incessantly exposed to reactive oxygen species produced by infiltrating inflammatory cells. We show augmentation of motility associated with morphological changes of human squamous carcinoma SASH1 cells, human peripheral monocytes (hPMs), and murine macrophage-like cell line J774.1 by superoxide stimulation. We also disclose that motility of hPMs and J774.1 induced by a chemotactic peptide (N-formyl-methionyl-leucyl-phenylalanine [fMLP]) was inhibited by superoxide dismutase or N-acetylcystein, indicating stimulation of motility by superoxide generated by fMLP stimulation. In these cells, protein kinase C (PKC)
was activated to phosphorylate RhoGDI-1, which liberated RhoGTPases, leading to their activation. These events were inhibited by dominant-negative PKC in SASH1 cells, myristoylated PKC peptides in hPMs and J774.1, or a specific inhibitor of RhoGTPase in SASH1, hPMs, and J774.1. These results suggest a new approach for manipulation of inflammation as well as tumor cell invasion by targeting this novel signaling pathway.
Abbreviations used in this paper: DHR, dihydrorhodamine; DN, dominant-negative; DPI, diphenylene iodonium; fMLP, N-formyl-methionyl-leucyl-phenylalanine; hPM, human peripheral monocyte; HPX, hypoxanthine; IP, immunoprecipitation; NAC, N-acetylcystein; ROS, reactive oxygen species; SOD, superoxide dismutase; XOD, xanthine oxidase.
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Introduction |
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TopAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
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Cell motility results from remodeling of acto-myosin system, which is regulated by Rho family of small GTP-binding proteins (Ridley, 2001). There is diversity in extracellular stimulants for cell migration, such as lysophophatidic acid (Moolenaar et al., 2004), platelet-derived growth factor (Chiarugi et al., 2000), and hepatocyte growth factor (Kodama et al., 2000), and many different intracellular signaling molecules that correspond to each stimulant are implicated in the activation of RhoGTPases. However, despite the fact that superoxide and chemokines are considerably important as stimulants of motility, not only from the view of tumor biology but also from the view of innate immunity, no detailed exploration on the motility relevant to these stimulants has been performed to date.
In the present study, we first reveal that, in human squamous carcinoma SASH1 cells, superoxide activates PKC, which phosphorylates RhoGDI-1, in turn liberating RhoGTPases from RhoGDI-1, leading to their activation. Then, using human peripheral monocytes (hPMs) and murine macrophage-like cell line J774.1, we examined whether the superoxide extracellularly generated by hypoxanthine/xanthine oxidase (HPX/XOD), or which they themselves produced upon treatment with a chemotactic peptide, N-formyl-methionyl-leucyl-phenylalanine (fMLP), stimulated their motility, and found that both extracellularly generated and self-produced superoxide augmented their motility. Furthermore, we confirmed that the PKC–RhoGDI-1 phosphorylation–RhoGTPases activation signaling pathway was also involved in the motility of hPMs and J774.1 stimulated with superoxide or fMLP.
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Results |
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TopAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
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Superoxide reorganizes actin structure and activates Rho family GTPases in SASH1 cells
To investigate the mechanism of enhancement of cell motility by superoxide, we first examined the morphological changes in SASH1 cells (Fig. 2 a).
The parental cells had a rather round shape with small lamellipodia. In the parental cell stimulated with superoxide, there was increment of F-actin and a more discrete formation of lamellipodia and filopodia than in that without stimulation. Cu-Zn SOD transduced clone 1 showed less actin staining with spindle-shaped morphology (dendrite-like formation). Similar morphological features were observed with NAC-treated cells. This spindle-shaped morphology was unchanged even after superoxide stimulation.
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Effects of inhibitors of Rho, Rac, and Cdc42 on the motility and morphological change relevant to superoxide in SASH1 cells
To verify the involvement of Rho, Rac, and Cdc42 in motility and morphological changes relevant to superoxide, we examined the effect of specific inhibitors of the proteins on these cellular events in SASH1 cells. Treatment with C3 substantially suppressed the motility of SASH1 cells down to the basal levels, equivalent to that of NAC-pretreated cells, irrespective of superoxide stimulation.
Transfectants of dominant-negative (DN) Cdc42 (DNCdc42) and Rac1 (DNRac1), exhibited impaired motility similar to that of C3-treated cells treated with or without superoxide (Fig. 3 a). When the morphology of SASH1 cells was examined, treatment with C3 resulted in a slight reduction of F-actin intensity (Fig. 3 d) compared with that of nontreated cells (Fig. 3 b) and showed new dendrite-like formations and multiple nuclei in a single cell caused by inhibition of cytoplasmic division. In these cells, superoxide treatment did not increase F-actin intensity, but apparently induced lamellipodia or filopodia formation (Fig. 3 e). DNRac1 transfectant was not substantially different from the parental cells without the stimulation (Fig. 3 f), whereas superoxide treatment of the cells induced F-actin increment and filopodia formation, although lamellipodia formation was not observed (Fig. 3 g). Transduction of DNCdc42 caused loss of cell polarity with relatively concentrated F-actin staining in the center of the cell (Fig. 3 h). The morphological characters of DNCdc42 became more apparent by treatment with superoxide (Fig. 3 i). These results are compatible with the previous notion that F-actin is regulated by Rho; that activation of Rac1 is associated with lamellipodia formation (Nobes and Hall, 1995), although it does not associate much with F-actin or filopodia formation; and that Cdc42 regulates cell polarity and filopodia (Etienne-Manneville, 2004).
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in SASH1 cells
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, -ß, -
, -
, -
, -µ, and -, were found to be expressed (Fig. 4 c, lane 1). As members of the Rho family of small GTPases form complexes with RhoGDI-1 in cytosol in their resting state and the first step of Rho family activation is their liberation from RhoGDI-1 (Ueda et al., 1990; Isomura et al., 1991; Chuang, et al., 1993; Hancock and Hall, 1993; Hart et al., 1998), we next identified the PKC isozyme that interacted with RhoGDI-1–RhoGTPases by pull-down assay using the anti–RhoGDI-1 antibody. As shown in Fig. 4 c, among seven isozymes expressed in SASH1 cells, only PKC was detected in the immunoprecipitates, and the association was enhanced by superoxide treatment. Furthermore, this coprecipitation of PKC with RhoGDI-1 was clearly inhibited in NAC-pretreated (Fig. 4 e) or Cu-Zn SOD transduced cells (Fig. 4 f), indicating dependency of PKC activation on superoxide stimulation. We then examined whether PKC kinase activity is stimulated in superoxide-treated cells. As shown in Fig. 4 g, phosphorylated PKC was increased in superoxide- treated cells as compared with nontreated cells revealed by in vitro phosphorylation assay. The PKC activity was suppressed by calphostin C in a dose-dependent manner as described previously (Xu and Clark, 1997; Furukawa et al., 1999; Fitzgerald et al., 2000; Liu et al., 2000). Further, PKC activation was confirmed by its translocation to the plasma membrane (Fig. 4 d). The PKC molecule consists of two functional domains, reportedly, a regulatory N-terminal domain and a catalytic C-terminal domain (Spitaler and Cantrell, 2004). We investigated which domain interacts with RhoGDI-1 in superoxide-treated SASH1 cells by pull-down assay. As shown in Fig. S3 a (available at http://www.jcb.org/cgi/conent/full/jcb.200607019/DC1), RhoGDI-1 did associate with the C-terminal catalytic domain but not with the N-terminal regulatory domain. The fact that PKC is a member of serine-threonine kinases, and RhoGDI-1's interaction with the catalytic domain of PKC, prompted us to examine the possibility of phosphorylation of RhoGDI-1 by PKC. Immunoprecipitates with anti–RhoGDI-1 antibody were found to contain threonine but not serine-phosphorylated RhoGDI-1 in both superoxide-treated and nontreated cells, though in the latter the threonine-phosphorylated RhoGDI-1 increased more than in the former (Fig. 4 h).
To further elucidate whether PKC directly phosphorylate RhoGDI-1 upon superoxide stimulation or, rather, activates a putative kinase, which in turn phosphorylates RhoGDI-1, RhoGDI-1 was purified from SASH1 cell lysate by immunoprecipitation (IP) and incubated with recombinant active PKC. As shown in Fig. 4 i, the intensity of the band representing [32P]RhoGDI-1 in the PKC-treated preparation was apparently increased compared with that in the nontreated preparation, indicating the direct action of PKC on RhoGDI-1.
On the other hand, when recombinant RhoGDI-1 was simply incubated with recombinant active PKC, phosphorylation of RhoGDI-1 was undetectable (unpublished data). Therefore, it is highly conceivable that some cofactors that coprecipitated with RhoGDI from the cell lysate were required for RhoGDI-1 phosphorylation by PKC. The small RhoGTPase family proteins are potential cofactors. Upon superoxide stimulation, PKC interacts with RhoGDI-1/small RhoGTPase family protein complex; binding of PKC to the complex stimulates the release of the small RhoGTPases from RhoGDI (Fig. 4 j). Incidentally, weak but nonnegligible phosphorylation of RhoGDI-1 observed in the PKC-nontreated preparation (Fig. 4 i, left) may be ascribed to the activity of endogenous PKC that has been coprecipitated with RhoGDI-1.
It is plausible that there is a kinase that phosphorylates RhoGDI downstream of PKC. To test this, recombinant active PKC was incubated with cell lysate to activate the putative kinase. PKC was subsequently removed by IP, and recombinant GST–RhoGDI-1 was incubated with the lysate but was not phosphorylated by the PKC-deficient cell lysate (Fig. S3 b). The results denied the possibility of the other kinase downstream of PKC.
We then extended our investigation to examine the kinetics of complex formation of RhoGDI-1 and Rho family GTPases or PKC by IP using anti–RhoGDI-1 antibody. An immunoreactive band of PKC increased in intensity from 30 s until 2 min after superoxide treatment (Fig. 4 j). However, in the same time course, RhoGDI-1 showed no appreciable changes, whereas the bands of all Rho family GTPases showed gradual decrement, indicating the dissociation of RhoGTPases from the complex and their eventual activation upon initiation of superoxide stimulation. These dissociation and association phenomena were not obvious in DNPKC transduced cells (Fig. 5 e).
DNPKC inhibits superoxide-induced morphological change, Rho family GTPase activation, and motility of SASH1 cells
To further validate the implication of PKC in morphological changes, Rho family activation, and cell motility induced by superoxide, SASH1 cells were transfected with DNPKC plasmid to suppress PKC activity. We first confirmed that the transfectants expressed the DNPKC protein (unpublished data) and that DNPKC was truly active to inhibit autophosphorylation of PKC in vitro (Fig. 5 a).
This DNPKC transfectant showed morphological features that were similar to those of the parental cells (Fig. 5 b), and these features were not apparently affected by superoxide treatment. Moreover, activation of Rho family proteins with superoxide stimulation was not seen in the DNPKC transfectants (Fig. 5 d). Furthermore, intrinsic cell motility or cell motility induced by superoxide was also impaired in the transfectant (Fig. 5 c). In the transfectant of DNPKC, the associations of RhoA, Rac1, and Cdc42 with RhoGDI were not affected (Fig. 5 e) and showed no increase of threonine phosphorylation of RhoGDI even after HPX/XOD treatment (Fig. 5 e). These results indicate that the activation of Rho family, the changes of cell morphology, and stimulation of cell motility relevant to superoxide were evoked by PKC.
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and RhoGTPases
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involvement in this signaling pathway. Among PKC, -ß, -, -, -, -µ, and -, only PKC associated with RhoGDI-1 examined by pull-down assay using RhoGDI-1 antibody (unpublished data). To confirm the role of PKC in cell motility and RhoGTPase activation, J774.1 cells were treated with calphostin C. Both cell motility (Fig. 6 f) and RhoGTPase activation (Fig. 6 g) of superoxide-treated and nontreated cells were substantially suppressed to basal levels by 500 nM calphostin C. We used myristoylated pseudosubstrate peptide of PKC (myr-PKCp) to inhibit the enzyme activity. Motility of J774.1 cells with or without superoxide stimulation was impaired by myr-PKCp compared with control peptide–treated cells (Fig. 6 h). The phenomena of association of PKC and dissociation of GTPases from the RhoGDI-1–GTPases complex upon superoxide stimulation found in SASH1 cells were also confirmed in this cell line (Fig. 6 i), and dissociation and association were inhibited by calphostin C (not depicted). As shown in Fig. S4 (available at http://www.jcb.org/cgi/content/full/jcb.200607019/DC1), Rac2 was also activated by superoxide in J774.1 cells.
Motility of human peripheral blood monocytes is induced by superoxide through PKC–RhoGDI-1–RhoGTPases signal
We further extended our study by examining human monocytes collected from the peripheral blood of a healthy volunteer. As shown in Fig. S5 a (available at http://www.jcb.org/cgi/content/full/jcb.200607019/DC1), addition of Cu-Zn SOD significantly lowered the motility of hPMs, indicating autocrine activation of motility by superoxide, which they generated (automotility). Treatment with NAC almost completely suppressed their motility, a finding consistent with that observed with J774.1. Augmentation of motility by superoxide treatment was also observed with hPMs (Fig. S5 a), as was the case with SASH1 and J774.1, and the motility was positively related with superoxide production (Fig. S5 b). Morphologically, NAC-treated hPMs showed spindle form with dendrite formation, whereas nontreated cells exhibited a rather round shape with slightly increased F-actin staining and formation of some small filopodia (Fig. S5 c). Such F-actin staining or filopodia formation of hPMs was further strengthened by treatment with superoxide (Fig. S5 c). As shown in Fig. S5 d, the activation of RhoGTPases by superoxide was also confirmed. Moreover, movement of hPMs treated with or without superoxide was strongly suppressed by C3 to the basal levels (Fig. S5 e). To investigate whether PKC is also involved in the motility of hPMs, we pulled down the isozyme with anti–RhoGDI-1 antibody in the cells treated with or without superoxide. Coprecipitation of PKC with RhoGDI-1 was clearly observed in cells treated with superoxide and seen with less intensity in those without superoxide (Fig. 7 a).
RhoGDI-1 was apparently phosphorylated, as revealed by immunoblotting using anti-phosphothreonine antibody in both superoxide-treated and nontreated hPMs, with much higher intensity in the former than the latter (Fig. 7 a). As shown in Fig. 7 b, PKC kinase activity was increased by superoxide stimulation. To examine the involvement of PKC in the motility of hPMs, the cells were treated with myr-PKCp. As shown in Fig. 7 c, motility of both superoxide-stimulated and nonstimulated cells were almost completely suppressed to basal level by myr-PKCp. The activation of RhoGTPases in hPMs was also clearly suppressed by treatment with myr-PKCp (Fig. 7 d). Association of PKC with RhoGDI-1 upon superoxide stimulation and dissociation of RhoGTPases from RhoGDI-1 were confirmed with hPMs as well (Fig. 7 e). The effect of myr-PKCp on the morphology of hPMs was further examined (Fig. 7 f). hPMs had a round shape with small filopodia and mild F-actin staining. Filopodia formation and F-actin staining were enhanced, and lamellipodia formation became evident by treatment with superoxide. In myr-PKCp–treated cells, filopodia and lamellipodia formation were hardly seen and intensity of F-actin was rather weak compared with nontreated cells. The association of PKC with and dissociation of RhoGTPases from RhoGDI-1 were impaired in myr-PKC–treated cells (unpublished data).
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p, the motility of the cells was totally suppressed to basal levels. Essentially the same results were obtained by migration assay (Fig. S1 b). Thus, it was suggested that PKC is directly activated by fMLP, which utilizes the same PKC–RhoGDI-1–GTPases signal pathway as superoxide.
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Discussion |
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Regarding the motility of cancer cells (SASH1), the autocrine mechanism was not operating, as self-produced superoxide was not detectable. However, as they respond to exogenous superoxide (HPX/XOD), it is suggested that cancer cells acquire invasiveness and metastatic ability as a consequence of enhanced motility stimulated by the superoxide generated by infiltrating inflammatory cells in a paracrine manner in vivo. The observation that NAC treatment of hPMs and J774.1 or Cu-Zn SOD transfection to SASH1 cells almost completely nullified the motility of these cells implies the essentiality of intracellular superoxide as a generator of cell motility at the basal level (intrinsic motility). This recognition of the essentiality of intracellular superoxide for cell motility is consistent with our previous observation that an inverse relationship existed between intracellular Cu-Zn SOD activity and invasiveness of tumor cells. We used a tetracycline regulation system for transient expression of Cu-Zn SOD, as it was previously reported that forced long-term expression of SOD gene brought about growth suppression of the transfectants (Liu et al., 1997). With respect to the mechanism whereby extracellular superoxide increases intracellular superoxide, penetration through cell membrane may be a possibility (Lynch and Fridovich, 1978; Gus'kova et al., 1984; Mao and Poznansky, 1992; Gomes et al., 1993), although the proof remains to be established.
To substantiate the idea that superoxide induces motility, we investigated the underlying molecular mechanism. The first issue we addressed was whether, and to what extent, the Rho family of small GTP-binding proteins was involved in the signal transduction of superoxide-induced cell movement, as these proteins are well accepted as common mediators of the cell signals for various stimuli. RhoA, Rac1, and Cdc42 were each activated by superoxide stimulation in all three cells examined. The motility of SASH1 cells was markedly suppressed by C3, DNRac1, and DNCdc42, with C3 limiting motility to basal levels in hPMs and J774.1 cells. Together, these results indicate that ROS induces cell motility via Rho family GTPase activation. Furthermore, each single GTPase may be indispensable for the cell movement induced by superoxide.
Next, we searched for the molecule that may link the superoxide signal with Rho family GTPases. It has been reported that stimulators of cell motility, such as lysophophatidic acid and insulin, activate both Rho family GTPases and PKC simultaneously (Hall, 1994; Machesky and Hall, 1996). Translocation of RhoA to membrane and RhoA-dependent phospholipase D activation induced by GTPS are reportedly blocked by calphostin C in MDCK cells (Balboa and Insel, 1995). Activation of RhoA by phosphorylating RhoGDI-1 by PKC has been observed in human umbilical venular endothelial cells stimulated with thrombin (Mehta et al., 2001). In addition to these reports, previous evidence indicating activation of PKCs by oxidative stress (Konishi et al., 1997; Klann et al., 1998) led us to hypothesize that PKC is the linking molecule. This hypothesis was supported by the fact that calphostin C suppressed motility and activation of RhoGTPases in all three types of cells. With regard to the mechanism for activation of PKC by superoxide, the involvement of PI3-kinase in linking these molecules is conceivable because PI3-kinase has been known to activate PKC via PDK-1 phosphorylation in vitro (Chou et al., 1998). However, the fact that the inhibitor of PI3-kinase, LY294002, did not inhibit superoxide-induced RhoA or Cdc42 activation in our cells (unpublished data) negated this possibility. Thus, it seems likely that superoxide activates PKC either through activation of kinases other than PI3-kinase or through direct activation of PKC by superoxide, possibly by causing conformational change (Palumbo et al., 1992). Elucidation of these putative mechanisms remains a future task.
Given that PKC is involved in signaling from superoxide to all three RhoGTPases, it is highly plausible that PKC interacts with RhoGDI-1 to activate RhoGTPases, because inactive RhoA, Rac1, and Cdc42 are commonly bound to RhoGDI-1. Therefore, using SASH1, J774.1 cells, and hPMs, we examined the direct interaction between PKCs and RhoGDI-1 by IP and found that only one specific isozyme of PKCs, PKC, was coprecipitable with RhoGDI-1. This observation, taken collectively with the facts that treatment with DNPKC or inhibitory peptide of PKC caused suppression of motility and the activation of the GTPases, strongly supports the notion that this particular PKC isozyme is a signal transducer of superoxide.
It has been shown that peroxide activates various PKC isozymes expressed in COS cells (Konishi et al., 1997). We also found that superoxide treatment of SASH1 brought about activation of all types of PKCs expressed in the cell, as determined by their translocation to the plasma membrane. In this context, the specific binding of PKC to RhoGDI-1 can be rationalized by the speculation that this isozyme has higher binding affinity than other PKC isotypes to the substrate (RhoGDI-1), and not by the specific activation of PKC by superoxide.
It is generally accepted that for activation of RhoGTPases, their release from the RhoGDI-1 molecule is required. It was recently reported that PKC, activated by thrombin, can phosphorylate RhoGDI-1, catalyzing the release of bound GTPases (Mehta et al., 2001). Our present results indicate that superoxide stimulates PKC, which in turn leads to RhoGDI-1 phosphorylation at threonine sites to liberate RhoGTPases. These results are compatible with the aforementioned study, except for the stimulant (superoxide) and signal transducer (PKC).
As inflammatory cells are usually attracted and activated by chemokines in the lesion, we further studied the role of superoxide in motility of J774.1 and hPMs induced by fMLP, a chemotactic peptide that is known to interact with 7-transmembrane G-coupled formyl-peptide receptors (Le et al., 2002). Our findings that fMLP treatment of these cells stimulated superoxide generation, which was suppressed by DPI, a specific inhibitor of NADPH oxidase, confirmed the notion that chemokines activate PKC to stimulate NADPH oxidase at plasma membrane, in turn generating superoxide (Bokoch 1995; Dang et al., 2001). On the basis of these findings, we extended our investigation to elucidate the relevance of fMLP-induced motility to superoxide generated by the cells treated with fMLP. The results that Cu-Zn SOD or DPI treatment suppressed the motility of J774.1 and hPMs support the concept of autocrine activation of chemokine-induced motility via superoxide. It was speculated that NAC should scavenge out intracellular superoxide and thereby abrogate superoxide-induced cell motility. However, when the fMLP-stimulated J774.1 cells or hPMs were pretreated with NAC, a certain magnitude of motility unexpectedly remained. The results therefore imply that fMLP interaction with chemokine receptor stimulates multiple pathways to activate cell motility. Furthermore, the fact that myr-PKCp suppressed the motility of J774.1 cells and hPMs to basal level suggests the possibility that the other signal (as well as superoxide-relevant one) is also mediated by PKC. This notion, activation of PKC through interaction of chemokine with its receptor, is veritably compatible with the recent report that atypical PKC regulates stromal cell–derived factor-1 mediated migration of human CD34+ progenitor cells (Petit et al., 2005). Hence, it may be reasonable to deduce that fMLP binds its receptors to activate PKC to generate superoxide, which in turn stimulates the motility in an autocrine manner via the PKC–RhoGDI-1–RhoGTPase pathway. On the other hand, PKC activated by fMLP simultaneously induces motility via the common PKC–RhoGDI-1–RhoGTPase pathway. Accordingly, the motility induced by chemokine is considered to be at least partly dependent on superoxide.
In conclusion, we disclosed that superoxide plays a pivotal role in the motility of SASH1, J774.1, and hPMs through a novel signaling pathway of PKC–RhoGDI-1–RhoGTPases. Thus, these results suggest a new approach for manipulation of inflammation as well as tumor cell invasion by targeting this novel signaling pathway.
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Materials and methods |
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Culture of peripheral blood monocytes
CD14+ monocytes were isolated from peripheral blood mononuclear cells of healthy volunteers obtained by the standard Ficoll-Paque method and separated by negative magnetic depletion using hapten-conjugated CD3, CD7, CD19, CD45RA, CD56, and anti-IgE antibodies (MACS; Miltenyi Biotec) and a magnetic cell separator (MACS) according to the manufacturer's instructions.
Superoxide and NAC treatment
Serum-starved cultures on 100-mm dishes were treated with 4 µg/ml HPX (Sigma-Aldrich) and 7 x 10–3 U/ml XOD (Sigma-Aldrich) for various incubation periods noted in each experiment. For NAC treatment, NAC (Sigma-Aldrich) was dissolved in the medium, pH adjusted to 7.4, and the cells were treated for 1 h.
Measurement of phagokinetic tracks
A uniform carpet of gold particles was prepared as previously described (Albrecht-Buehler, 1977). In brief, sterilized coverslips were coated with 1% bovine serum albumin and were placed in 35-mm tissue culture dishes. Gold colloidal solution (184 µM HAuCl4-4H20 and 11.6 mM Na2CO3) was boiled shortly to make gold colloidal particles, and the coverslips were coated with them. 2 x 103 serum-starved cells with various pretreatments described in each experiment were seeded, attached to the coverslips, treated with or without superoxide, and 2 h later fixed by 0.1% formaldehyde. The areas cleared of gold particles were examined under microscope and quantified by image processing (KS400; Carl Zeiss MicroImaging, Inc.).
Evaluation of intracellular ROS level
Intracellular ROS level was evaluated using DHR 123 (Invitrogen) as a fluorescent probe. After cells were treated with various conditions, as indicated, 1 µl of 43.3 mM DHR was added to the cells suspended with 1 ml HBSS for 20 min, cells were washed, the generation of rhodamine 123 was monitored on a FACScan (Becton Dickinson) with excitation at 488 nm, and the emitted fluorescence was collected at 525 nm.
F-actin staining
The cells were fixed in 3% paraformaldehyde for 20 min, permeabilized with 0.2% Triton X-100 for 5 min, and stained with rhodamine-labeled phalloidin (Invitrogen) for 40 min to visualize F-actin. Fluorescence images were obtained by a confocal laser-scanning microscope system with 40x objective lens (LSM5 PASCAL; Carl Zeiss MicroImaging, Inc.).
Preparation of recombinant Clostridium botulinum C3 exoenzyme and ADP-ribosylation assay
Preparation of recombinant C. botulinum C3 exoenzyme and ADP-ribosylation assay were performed as described previously (Morii and Narumiya, 1995). The C3-expressing plasmid pET3a C3 was provided by S. Narumiya (Kyoto University Faculty of Medicine, Kyoto, Japan).
Immunodetection of RhoGDI-1–associated proteins
Cells were lysed in lysis buffer (20 mM Tris/HCl, pH 7.5, 10 mM MgCl2, 1 mM pefabloc, 5 mM leupeptin, and 5 mM pepstatin) containing 1% NP-40 and were incubated with protein G–agarose beads (GE Healthcare) for 3 h to block the nonspecific binding. After centrifugation, the supernatant was incubated with anti–RhoGDI-1 antibody (Santa Cruz Biotechnology, Inc.) for 3 h, followed by the addition of protein G–agarose beads, and agitated overnight. The beads were collected, washed, and further analyzed by immunoblotting. For immunoblotting, each aliquot of the sample was eluted by RhoGDI-1 peptide, mixed in SDS sample buffer, heated, separated on SDS-PAGE, and electroblotted. Antibodies against RhoA (26C4), RhoGDI-1, and PKC were purchased from Santa Cruz Biotechnology, Inc. Antibodies against Rac1 and PKC, -ß, -, -, -, -
, and -µ were purchased from BD Biosciences. Antibodies against Cdc42 were purchased from Upstate Biotechnology. Antibodies against phosphothreonine and phosphoserine were purchased from Zymed Laboratories and QIAGEN.
Rho family activation assay
Activity of Rho family small GTP-binding proteins was assayed by pull-down assay using each assay kit (Upstate Biotechnology) according to the manufacturer's instruction.
PKC activity assay
PKC activity from 3 x 106 cells was assayed by assessing its phosphorylation state as described previously (Xu and Clark, 1997). In brief, PKC in cell lysates was immunoprecipitated with the anti-PKC antibody, washed, mixed in assay solution (35 mM Tris, pH 7.5, 15 mM MgCl2, 1 mM MnCl2, 0.5 mM EGTA, 0.1 mM CaCl2, 1 mM sodium orthovanadate, and 100 µM -[32P]ATP), and incubated at 30°C for 10 min. After reactions were stopped by the addition of gel loading buffer, the samples were boiled and analyzed by SDS-PAGE followed by autoradiography.
In vitro kinase assay for phosphorylation of RhoGDI
Phosphorylation of RhoGDI-1 in vitro was performed using RhoGDI-1 immunoprecipitated from SASH1 cell lysate with RhoGDI-1 antibody. Confluent cells grown in 100-mm dishes were washed with ice-cold PBS and lysed in IP buffer containing 50 mM Tris, pH 7.4, 150 mM NaCl, 0.25 mM EDTA, pH 8.0, 1% deoxycholic acid, 1% Triton X-100, 5 mM NaF, 1 mM sodium orthovanadate, 1 mM PMSF, 5 µg/ml leupeptin, 5 µg/ml aprotinin, and 1 µg/ml pepstatin A. The cells were collected and then cleared by centrifugation at 4°C at 14,000 g for 10 min. The lysate was incubated with anti–RhoGDI-1 antibody for 1 h followed by the addition of protein G–Sepharose beads overnight at 4°C. The beads were collected, washed twice with ice-cold lysate buffer, washed three times with PBS, and washed once with kinase buffer (8 mM MOPS, pH 7.4, and 0.2 mM EGTA). The immunoprecipitated RhoGDI-1 was incubated with 5 ng of recombinant active PKC in kinase assay buffer for 10 min at 30°C followed by the addition of magnesium/ATP cocktail (Upstate Biotechnology): 4 mM MgCl2 and 25 µM ATP in 1 mM MOPS, pH 7.2, 1 mM ß-glycerol phosphate, 0.2 mM EGTA, 50 µM sodium orthovanadate, 50 µM dithiothreitol, and 10 µCi -[32P]ATP (stock 1 mCi/100 µl: 3,000 Ci/mmol; PerkinElmer). The reaction was stopped by the addition of Laemmli sample buffer. The samples were electrophoresed on a 10% SDS-polyacrylamide gel, transferred to nitrocellulose membrane, and exposed to x-ray film. The blots were then subjected to Western blotting with anti–RhoGDI-1 antibody to verify that an equal amount of the protein loaded in each lane.
Subcellular fractionation
Cells were washed three times with ice-cold PBS and scraped in lysis buffer (20 mM Tris/HCl, pH 7.5, 10 mM MgCl2, 1 mM pefabloc, 5 mM leupeptin, and 5 mM pepstatin). The cells were lysed by 18 passes through a 26-gauge needle on ice. Trypan blue staining of the lysate indicated >95% disruption of plasma membrane. The subcellular fractionation was performed by the method as described by Fleming et al. (1996). The lysate was first centrifuged at 500 g for 10 min to prepare low-speed pellet, and the supernatant was recentrifuged at 120,000 g for 45 min to pellet the remainder of the particulate fraction (high-speed pellet). The low-speed pellet was further purified by sucrose gradient centrifugation to obtain plasma membrane fraction. Protein concentrations were determined using BCA protein assay (Pierce Chemical Co.) according to the manufacturer's directions. Alkaline phosphodiesterase I, cytochrome c oxidase, and lactate dehydrogenase were used as marker enzymes for plasma membrane, mitochondria, and cytosol, respectively (Storrie and Madden, 1990). DNA and RNA were used as markers for nucleus and endoplasmic reticulum, respectively, and were detected on agarose gels by ethidium bromide staining, with or without RNase treatment. After subfractionation of the cells, plasma membrane fractions were subjected for immunoblotting to measure PKC or RhoGTPase activity.
Cu-Zn SOD plasmid
The pTA-Hyg and pET2a vectors were provided by J. Yokota (National Cancer Center Research Institute, Tokyo, Japan). cDNA for the human Cu-Zn SOD expression vector was generated by PCR using the forward primer 5'-TTTCCGTTGCAGTCCTCGGAA-3' and the reverse primer 5'-CCTCAGACTACATCCAAGGGA-3' with the human cDNA library as the template. The PCR fragment was ligated into the pET2a vector to generate pET2a Cu-Zn SOD vector, which expresses Cu-Zn SOD when tetracycline is omitted from the medium. SASH1 cells were first transduced with the pTA-Hyg vector, selected by hygromycin B, transfected with the pET2a Cu-Zn SOD vector, and selected by G418.
Plasmid for DNRac1 and DNCdc42
cDNA of N17Rac1 and N17Cdc42, which are DN cDNAs of Rac1 and Cdc42, were provided by Y. Takai (Osaka University, Osaka, Japan). N17Rac1 was first ligated into the pSVneo-Myc/Sra vector and then cDNA coding for Rac1N17 tagged with Myc was digested and ligated into the pIRES/neo vector, resulting in pIRES-Rac1N17-Myc/neo. N17Cdc42 was first ligated into the pFlag-CMV vector and then cDNA coding for N17Cdc42 tagged with Flag was digested and ligated into the pIRES/hyg vector, resulting in pIRES-Cdc42N17-Flag/hyg.
Plasmid for N- and C-terminal of PKC
cDNA coding for the N- or C-terminal of human PKC was generated by PCR with the human cDNA library as the template using the First strand cDNA Synthesis kit (Boeringer). To produce plasmid pRx-bsr-PKCN that contains the N-terminal regulatory domain of human PKC (nucleotides 1–738), we used 5'-CCGGAATTCACCCAAGATGGAAGGGAGCGGCGGC-3' as the forward primer and 5'-CGCGGATCCTCATAGGTCAAAGTCCTGCAGCCCAAGC-3' as the reverse primer. PCR products were separated, sequenced, digested with EcoRI and NotI, and ligated into a pcDNA3.1/His (Invitrogen), and the fragment containing His6 was excised by HindIII and NotI. It was inserted into a retroviral vector, pRx-bsr, supplied by H. Hamada (Sapporo Medical University, Sapporo, Japan). We used pRx-bsr-PKCN regulatory domain expressing vector as DN (DNPKC), as this domain reportedly showed autoinhibitory activity (Jaken, 1996).
To produce plasmid PKCC that contains the C-terminal catalytic domain of human PKC (nucleotides 739–1755), cDNA was generated by PCR using the forward primer 5'-CCGGAATTCAATCAGAGTCATCGGGCGCGGGAGC-3' and the reverse primer 5'-CGGGGTACCTCACACCGACTCCTCGGTGGACAGC-3'. The PCR product was digested with EcoRI and NotI and ligated into a pcDNA3.1/V5-His (Invitrogen), and the fragment containing (His)6 was excised by EcoRI and PmeI. The excised N-terminal PKC fragment was inserted into a retroviral vector, pRx-bsr, to construct PKCC that contains the human C-terminal fragment of human PKC cDNA and (His)6. The plasmid of N- or C-terminal fragments of PKC was transfected into the ecotropic retroviral packaging cell line BOSC23 by the calcium phosphate precipitation method to obtain the retroviral supernatant. The amphotropic 
Crip packaging cells were then infected with 2 ml of filtered retroviral supernatants of each plasmid in the presence of polybrene (Sigma-Aldrich).
Measurement of superoxide production
Phenol red–free medium was used in the assay. 106 cells were stimulated for 1 min with HPX/XOD, and the cells were further incubated for 1 h in the presence of 50 µM cytochrome c. The medium was removed and placed on ice, and the absorbance at 550 nm was immediately read by spectrophotometer with H20 as a blank. Superoxide-specific reduction of cytochrome c was expressed as the difference in absorbance between cells incubated with or without SOD using an extinction coefficient of 21 mM–1 cm–1.
Peptides
Myr-PKCp, SIYRRGARRWRKLYRAN (positions 113–129, which is pseudosubstrate region of human PKC), and myr-control peptide, RLRYRNKRIWRSAYAGR (Laudanna et al., 1998), were custom made by Sawaday Technology Co. They were solubilized immediately before use at 1 mM concentration in PBS, pH 7.2, and heated at 40°C to achieve complete solubility.
Statistical analysis
Statistical analysis of data was performed by standard techniques with the aid of the Stat View computer program for the Macintosh (Abacus Concepts). The means of values were provided with associated SEM. Statistical significance was defined as P < 0.01.
Online supplemental material
Fig. S1 shows a migration assay of SASH1 cells and J774.1 cells. Fig. S2 shows translocation of Rho family small GTPases to plasma membrane in SASH1 cells treated with superoxide. Fig. S3 shows the interaction of PKC catalytic domain with RhoGDI and that a SASH1 cell lysate without PKC did not phosphorylate recombinant RhoGDI-1. Fig. S4 shows activation of GTP-Rac2 by superoxide in J774.1 cells. Fig. S5 shows that superoxide stimulates RhoGTPases activity and motility of hPMs. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200607019/DC1.
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Acknowledgments |
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Submitted: 5 July 2006
Accepted: 21 February 2007
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