INTRODUCTION

Protein kinase C (PKC)¹ is a multigene family of phospholipid-dependent, serine-threonine kinases that plays a central role in cell growth and differentiation. Molecular cloning studies have identified 10 isozymes encoded by 9 different mRNAs (1, 2). Based on sequence homology and biochemical properties, the PKC gene family has been divided into three groups: "conventional" PKCs (, I, II, and ) regulated by calcium and diacylglycerols or phorbol esters; "novel" PKCs (, , , and ), which are calcium-independent but diacylglycerol- and phorbol ester-sensitive; and "atypical" PKCs (, and /), which are insensitive to calcium, diacylglycerol, and PMA. In addition, two related phospholipid-dependent kinases, PKCµ and protein kinase D, share sequence homology in their regulatory domains to novel PKCs and may constitute a new subgroup (3, 4).

Several studies with tumor-promoting phorbol esters suggest that PKC modulates neural differentiation. Phorbol esters induce neural tissue from ectoderm in Xenopus embryos (5) and elicit neurite outgrowth from chick sensory ganglia (6, 7), chick ciliary ganglion neurons (8), several human neuroblastoma cell lines (9, 10), and rat PC12 cells (11, 12). Studies using purified isozymes, kinase-defective mutants, and transgenic or mutant cell lines have implicated PKC, -, -, -, and - in the differentiation of nonneural cells (13-17). Overexpression of PKC or - in Xenopus embryos enhances neural induction (18), but little else is known about the identity of specific PKC isozymes that regulate neural differentiation.

Recent evidence suggests that PKC plays a role in neural differentiation and plasticity. PKC is expressed predominantly in the nervous system and is particularly abundant in the hippocampus, olfactory tubercle, and layers I and II of cerebral cortex (19). Within immunoreactive neurons, it is localized to the Golgi apparatus and to axons and presynaptic nerve terminals (19). PKC is activated by growth factors that stimulate neural differentiation such as insulin (20) and NGF (21). In addition, in developing chick brain, it is the major isozyme found in nondividing, differentiating neurons (22).

Further evidence for involvement of PKC in neural differentiation has come from studies with PC12 cells. PC12 cells are derived from neural crest and, when treated with NGF or fibroblast growth factors, undergo dramatic biochemical and morphological differentiation, developing several characteristics of mature sympathetic neurons (23). PKC-activating phorbol esters enhance NGF-induced activation of ERK1 and ERK2 mitogen-activated protein (MAP) kinases and neurite outgrowth in PC12 cells, suggesting that PKC modulates responses to NGF (11, 12, 24). Studies with ethanol-treated PC12 cells helped direct us toward the PKC isozyme responsible for this effect. Like phorbol esters, ethanol increases NGF-induced MAP kinase activation and neurite outgrowth through a PKC-dependent mechanism (11, 24). Ethanol promotes PKC-mediated phosphorylation in PC12 cells by increasing levels of messenger RNA and protein for two PKC isozymes, PKC and PKC (25, 26). Recently, we found that overexpression of PKC, but not of PKC, enhances NGF-induced MAP kinase activation and neurite outgrowth (27). These findings establish PKC as a positive modulator of neurite growth. They also suggest that PKC mediates the neurite-promoting effect of ethanol and phorbol esters in PC12 cells. However, proof of this hypothesis requires studies with PKC isozyme-specific inhibitors or cells lacking specific PKC isozymes.

In the current study, we used specific inhibitors of PKC and PKC to investigate whether PKC mediates enhancement of neurite outgrowth by phorbol esters and ethanol. To achieve this goal we used dominant-negative inhibitors based on the amino acid sequences for PKC and PKC. This approach is based on the observation that upon activation, PKC isozymes translocate to specific intracellular sites where they appear to bind anchoring proteins, termed RACKs (receptors for activated C-kinase) (28). One such protein that has been cloned is RACK1, which interacts with the C2 domain of conventional PKCs (29). The sites of interaction between RACK1 and the C2 domain of PKC have been mapped, and short peptides derived from these domains inhibit translocation and activation of PKC in cardiac myocytes and Xenopus oocytes (29-31). Homology has been noted between the unique first variable region of PKC (V1) and the C2 domain of conventional PKCs (32), suggesting that, similar to the C2 domain of conventional PKCs, V1 may contain a binding site for an PKC-specific RACK. If that is the case, then expression of an V1 fragment should inhibit PKC translocation and function. Indeed, recent work has shown that an V1 fragment and a peptide corresponding to amino acids 14-21 in this region prevent phorbol ester-induced PKC translocation and inhibition of contraction in cultured cardiac myocytes (33).

In this paper, we describe studies with PC12 cell lines that stably express the fragments V1 or V1, which are derived from the first variable domains of PKC or PKC, respectively. We found that each fragment selectively inhibited phorbol ester-induced translocation of its corresponding isozyme, indicating that these fragments can function as isozyme-selective translocation inhibitors. NGF-induced MAP kinase phosphorylation and neurite outgrowth were not enhanced by phorbol esters or ethanol in cells expressing V1, but they were increased by these agents in cells expressing V1 and in cells transfected with empty vector. These results demonstrate that PKC specifically mediates enhancement of MAP kinase activation and neurite growth by phorbol esters and ethanol in PC12 cells.

EXPERIMENTAL PROCEDURES

NGF (2.5 S) was purchased from Collaborative Research (Bedford, MA). Geneticin (G418), laminin, and poly-L-ornithine (30-70 kDa) were purchased from Sigma. Phorbol 12-myristate 13-acetate (PMA) was from LC Laboratories (Woburn, MA). Antibodies were purchased from the following sources: peroxidase-conjugated goat anti-rabbit IgG from Boehringer Mannheim, fluorescein-conjugated goat anti-rabbit IgG from Cappel (Durham, NC), anti-phospho-MAP kinase antibody from New England Biolabs (Beverly, MA), and anti-Flag M2 antibody from Eastman Kodak Co.

PC12 cells, originally obtained from Dr. John A. Wagner (Cornell University, New York, NY), were cultured in plastic tissue culture flasks at 37 °C in Dulbecco's modified Eagle's medium containing 10% heat-inactivated horse serum, 5% fetal calf serum, 2 mM glutamine, 50 units/ml penicillin, and 50 µg/ml streptomycin, in a humidified atmosphere of 90% air and 10% CO₂. For studies of neurite outgrowth, cells were plated at a density of 30-40 × 10³ cells/well on 24-well plastic culture plates pretreated for 1 h with poly-L-ornithine (100 µg/ml in 15 mM sodium borate, pH 8.4). In some experiments, cells were plated at a density of 20 × 10³ cells/well in 8-well glass chamber slides (Nunc, Naperville, IL) treated first with poly-L-ornithine and then laminin (30 µg/ml) overnight. Cells were cultured in medium containing 50 ng/ml of NGF for 4 days, and neurites were measured as described previously (27). Ethanol-treated cultures were wrapped in Parafilm to prevent evaporation of ethanol, as in prior studies (24, 25).

To detect PKC immunoreactivity, cells plated on glass chamber slides were incubated in PBS (137 mM NaCl, 2.7 mM KCl, 1.47 mM KH₂PO₄ 8 mM Na₂HPO₄, 0.5 mM MgCl₂, 0.9 mM CaCl₂, pH 7.2) containing 2% paraformaldehyde for 30 min and 4% paraformaldehyde for 30 min at 4 °C. Cells were washed three times in PBS and incubated for 2 h in PBS containing 1% normal goat serum and 0.1% Triton X-100. Cells were incubated 48 h at 4 °C in PBS containing 2 mg/ml of bovine serum albumin, 0.1% Triton X-100, and 2 µg/ml of rabbit anti-PKC antibody (34) provided by Dr. Susan C. Kiley (W. Alton Jones Cell Science Center, Lake Placid, NY). Cells were washed three times in PBS, and immunoreactivity was detected using fluorescein-conjugated goat anti-rabbit IgG as described previously (27). Images were detected using a liquid-cooled CCD camera (Photometrics Ltd., Tucson, AZ) fitted with a Thompson 7883 chip (384 × 576 pixels) attached to an Olympus IMT-2 inverted microscope equipped with a × 60 1.3 numerical aperture objective. Exposure times were 0.5 s, and images were stored on an Apple Macintosh Quadra 950 computer. Fluorescence intensity in growth cones and cytoplasm was measured using the program BDS Image (Oncor Imaging Systems, Gaithersburg, MD).

To detect PKC immunoreactivity, cells were incubated with rabbit polyclonal antibody against PKC (0.2 µg/ml) from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA) as described previously (27). Immunofluorescence in 0.5-µm optical sections was detected with a Bio-Rad MRC 1024 confocal laser-scanning microscope equipped with a Nikon × 60, 1.4 numerical aperture oil immersion objective. Nuclei were stained by incubating cells in PBS containing 2 mg/ml of bovine serum albumin, 0.1% Triton X-100, and 0.2 µM TOTO-3 (Molecular Probes, Eugene, OR) for 2 h at 25 °C. After three washes in PBS, slides were dried, and coverslips were mounted with Vectashield (Vector, Burlingame, CA). TOTO-3 immunofluorescence was detected by confocal laser-scanning microscopy. Images were analyzed in BDS Image to measure the area of each nucleus at its widest diameter.

The plasmid pDM27 (33), containing a Flag-epitope tag followed by the sequence encoding amino acids 2-144 of PKC was used to amplify a 480-base pair fragment containing a NotI site at the 5-end and a XbaI site at the 3-end. This amplified fragment, containing an ATG start codon, the Flag epitope sequence, and the PKC sequence, was subcloned into the NotI and XbaI sites of pRc/RSV (Invitrogen, San Diego, CA) to generate the plasmid pRV-1. Another plasmid, pDM68 (33), containing the Flag epitope followed by the sequence encoding amino acids 2-144 of PKC was used to amplify and subclone a homologous Flag-tagged PKC sequence into pRc/RSV to generate the plasmid pRV-1. PC12 cells (10⁷) were suspended in 0.5 ml of Ca²⁺- and Mg²⁺-free PBS containing 137 mM NaCl, 2.7 mM KCl, 1.47 mM KH₂PO₄, 8 mM Na₂HPO₄, pH 7.2, and were electroporated with 80 µg of pRV-1, pRV-1, or pRc/RSV as described previously (27). Geneticin was initially added at 400 µg/ml to select clones and later was added at 200 µg/ml to maintain cultures. For each vector, 46 clones were selected, expanded, and then examined for expression of Flag-tagged PKC fragment mRNA using RT-PCR and for Flag immunoreactivity by Western analysis.

Poly(A) mRNA was isolated from 10⁶ cells using a Micro-Fast Track mRNA isolation kit (Invitrogen, San Diego, CA). Reverse transcription was carried out with 100 ng of mRNA using a Stratagene (La Jolla, CA) RT-PCR kit according to the manufacturer's protocol. Amplification of cDNA was achieved using the forward primer (5-ACACTGGCGGCCGCATGGACTACAAGGACGACGAT-3) and the reverse primers 5-AGCGAGCTCTAGATCGTTCTTCATTGTCTTTA-3 for pRV-1, and 5-ACAGACCTCTAGAGCGGTTCATAGTTGGGAA-3 for pRV-1. These primers were designed to specifically amplify the Flag-V1 sequences and not the V1 sequences of endogenous PKC and PKC in cells. Samples were heated to 94 °C, and amplification was started by the addition of Taq DNA polymerase. The amplification cycle was as follows: annealing for 45 s at 48 °C; elongation for 2 min at 72 °C, and denaturation for 1 min at 94 °C. Amplification was repeated for 30 cycles.

To detect expression of Flag-tagged peptides, cells were cultured on poly-L-ornithine-coated, 100-mm tissue culture dishes at a density of 6 × 10⁶ cells/dish. Medium was removed, and cells were rinsed twice at 4 °C with buffer A (120 mM NaCl, 5 mM KCl, 1.4 mM CaCl₂, 0.8 mM MgSO₄, 1 mM NaH₂PO₄, 10 mM glucose, 25 mM HEPES, pH 7.4). Cells were scraped into 1 ml of buffer A containing 40 µg/ml leupeptin, 40 µg/ml soybean trypsin inhibitor, and 1 mM phenylmethylsulfonyl fluoride and then were frozen on dry ice. Concentrated 5 × sample buffer was added to 400-µl frozen samples to yield a final solution containing 62.5 mM Tris-HCl (pH 6.8), 2% SDS, 10% glycerol, 5% 2-mercaptoethanol, and 12.5 mg/ml bromphenol blue. Samples were heated at 90 °C for 10 min, passed five times through a 26-gauge needle, and centrifuged at 10,000 × g for 10 min. Samples (80 µg/lane) were separated by SDS-polyacrylamide gel electrophoresis using 14% gels. Proteins were electrophoretically transferred for 2 h at 4 °C to Hybond-C extra membranes (Amersham Corp.). Membranes were blocked for 1 h with 3% nonfat dry milk dissolved in Tris-buffered saline (TBS; 20 mM Tris HCl, pH 7.4, 137 mM NaCl). Blots were then incubated with anti-Flag M2 antibody (10 µg/ml) for 2 h at 25 °C. Blots were washed three times with TBS containing 0.05% Tween-20 (TBS-T) for two minutes and then were incubated with goat anti-mouse IgG-peroxidase-conjugated antibody (1:1000 dilution) in blocking solution overnight at 25 °C. Blots were washed three times for 15 min in TBS-T and once with TBS. Immunoreactive bands were detected with the ECL kit from Amersham.

Activation of ERK1 and ERK2 MAP kinases was assayed with a phospho-specific 42/44-kDa MAP kinase rabbit polyclonal antibody raised against a phosphotyrosine peptide corresponding to residues 196-209 of human ERK1. The antibody detects phosphorylation of ERK1 at tyrosine 204, which is required for ERK1 activation (35). The antibody also detects phosphorylation of the corresponding activating tyrosine of ERK2. Blots from 11% polyacrylamide gels were washed for 5 min with 25 ml of buffer B containing 58 mM Na₂HPO₄, 17 mM NaH₂PO₄, and 68 mM NaCl, pH 7.4. Membranes were blocked in buffer B containing 0.1% Tween 20 and 5% milk (blocking buffer) for 1 h. Blots were incubated overnight at 4 °C with 1 µg/ml of anti-phospho-MAP kinase antibody in buffer B containing 0.05% Tween 20 and 5% bovine serum albumin (incubation buffer). They were then washed three times for 5 min with 15 ml of blocking buffer and incubated with goat anti-rabbit alkaline phosphatase-conjugated antibody (1:2000 dilution) in incubation buffer for 1 h at 25 °C. Blots were finally washed three times with 15 ml of blocking buffer, and immunoreactive bands were detected using the Western-Light chemiluminescent detection system from Tropix, Inc. (Bedford, MA).

Cells (3-4 × 10⁶) were plated on 100-mm plastic tissue culture plates. After 48 h, cultures were rinsed at 37 °C twice with 10 ml of medium and incubated with or without 30 nM PMA for 2 min. Cells were rapidly rinsed twice at 4 °C with Ca²⁺- and Mg²⁺-free PBS and then were scraped into 1 ml of ice-cold buffer C containing 20 mM Tris HCl, pH 7.5, 2 mM EDTA, 10 mM EGTA, 40 µg/ml leupeptin, 40 µg/ml aprotinin, 20 µg/ml soybean trypsin inhibitor, and 1 mM phenylmethylsulfonyl fluoride. Cells were homogenized at 4 °C with 10 strokes of a Teflon-glass homogenizer. Sucrose was added to a final concentration of 250 mM, and the sample was homogenized with 10 additional strokes. An aliquot of 800 µl was centrifuged at 150,000 × g for 1 h, and the supernatant was frozen on dry ice. The pellet was dispersed in 800 µl of buffer C by sonication in a Branson Sonifier 450 for 2 s at a setting of six. Samples of supernatant and pellet suspension derived from 100 µg of crude homogenate were separated by SDS-polyacrylamide gel electrophoresis using 10% gels and analyzed for PKC and PKC immunoreactivity by Western analysis as described previously (27).

Protein concentrations were measured by the Bradford method (36) using bovine IgG standards. Results are expressed as mean ± S.E. values, and differences between means were analyzed by ANOVA. Where p < 0.05, the significance of differences between means was evaluated by the Scheffe F-test or the Newman Keuls test.

RESULTS

Stably transfected PC12 clones were tested for expression of the first variable domain of PKC or PKC by RT-PCR (Fig. 1A) and Western analysis (Fig. 1B). Two clones, V11 and V12, expressing the V1 region of PKC (V1), and two clones, V11 and V12, expressing the V1 region of PKC (V1), were expanded for further studies.

RT-PCR and Western analysis of transfected PC12 cells demonstrating expression of V1 and V1 fragments.

To determine whether the V1 and V1 fragments expressed by our PC12 clones act as PKC isozyme-selective inhibitors, we measured PMA-induced translocation of PKC and PKC to the particulate fraction of these cells. We predicted that if V1 contains a binding site for a PKC-specific RACK, it should inhibit translocation of PKC. Likewise, if V1 interacts with an PKC-specific RACK, then expression of V1 should block phorbol ester-mediated translocation of PKC. In the parent PC12 cell line and in cells transfected with the pRc/RSV vector alone (clone C1), 30 nM PMA stimulated translocation of PKC and PKC to the particulate fraction (Fig. 2). In V11 and V12 cells, which express the V1 peptide, 30 nM PMA stimulated translocation of PKC, but translocation of PKC was reduced in these clones (Fig. 2, A and B). In V11 and V12 cells, 30 nM PMA stimulated translocation of PKC, but translocation of PKC was reduced compared with control cells (Fig. 2, A and C). These results demonstrate that V1 and V1 fragments selectively inhibit PMA-induced translocation of their corresponding PKC isozymes.

The V1 and V1 fragments selectively inhibit PMA-induced translocation of their parent PKC isozymes in transfected cells.

We attempted to examine cells by indirect immunofluorescence with anti-Flag antibody to determine the subcellular localization of expressed V1 and V1 but were unsuccessful because of high background staining. We next examined the subcellular localization of endogenous PKC and PKC to determine whether their localization was altered by expression of V1 or V1. In undifferentiated cells, PKC and PKC immunoreactivity was observed throughout the cytoplasm, as previously reported (27) and was not altered in clones expressing V1 or V1 (data not shown). In PC12 and C1 cells treated with 50 ng/ml of NGF for 4 days, most PKC immunoreactivity was observed in the cytosol, asymmetrically next to the nucleus (Fig. 3). Less intense staining was seen in a thin perinuclear band and within nuclei. In the majority of V11 and V12 cells expressing V1, most PKC immunoreactivity appeared in the perinuclear region (Fig. 3). In addition, nuclei appeared larger in these cells. This was analyzed further using confocal images of TOTO-3-stained nuclei at their widest diameter. Nuclei were of similar size in PC12 (82.8 ± 1.5 µm²; n = 164) and C1 (88.3 ± 2.1 µm²; n = 145) cells but were significantly larger in V11 (106.4 ± 3.8 µm²; n = 88) and V12 (100.6 ± 2.8 µm²; n = 172) cells (p < 0.05 compared with PC12 and C1 nuclei; ANOVA and Scheffe-F test).

Indirect immunofluorescence of PKC in V1-expressing cells.

V11

V12

Following treatment with NGF, PKC immunoreactivity was observed in growth cones, neurite shafts, and the cytoplasm of the cell soma in PC12 and C1 cells (Fig. 4), as noted previously (27). In V11 and V12 cells expressing V1, PKC immunoreactivity was reduced in growth cones and neurite shafts (Fig. 4). This was examined further by calculating the ratio of mean fluorescence intensity of each growth cone and of the cytoplasm at the base of its neurite shaft. In PC12 (0.73 ± 0.02; n = 71) and C1 (0.76 ± 0.02; n = 55) cells, this ratio was significantly greater (p < 0.05; ANOVA and Scheffe F-test) than ratios measured in V11 (0.49 ± 0.01; n = 83) and V12 (0.50 ± 0.01; n = 71) cells. These results demonstrate that V1 and V1 fragments alter the localization of their corresponding PKC isozymes in cells undergoing NGF-induced differentiation. In addition, expression of V1 is associated with an increase in nuclear size.

Indirect immunofluorescence of PKC in V1-expressing cells.

V11

V12

Growth and Neurite Formation in Cell Lines Expressing V1 or V1

The growth rates of V1- and V1-expressing clones, parent PC12 cells, and C1 cells were similar before NGF treatment (data not shown). After culture in poly-L-ornithine-treated culture dishes with 50 ng/ml NGF for 4 days, the number of neurite-bearing cells and the length of neurites was similar in all cell lines (Tables I and II). However, in NGF-treated V11 and V12 cells, neither ethanol nor PMA (10 nM) increased neurite length or the percentage of cells that expressed neurites. In contrast, ethanol and PMA increased neurite length and the percentage of neurite-bearing cells in NGF-treated PC12, C1, V11, and V12 cultures. When cells were cultured on glass slides treated with poly-L-ornithine and coated with laminin, the results were qualitatively similar but more dramatic because NGF-induced neurite outgrowth was especially robust following treatment with ethanol or PMA in all but V11 and V12 cultures (Fig. 5, A and B). Therefore, expression of the V1 fragment, but not of the V1 fragment, appears to prevent enhancement of neurite extension by PMA or ethanol in NGF-treated cells.

Table I. Percentage of cells expressing neurites after treatment with NGF plus PMA or ethanol Cells were cultured on polyornithine-coated tissue culture dishes with or without 100 mM ethanol for 6 days and then with 50 ng/ml NGF in the continued presence or absence of ethanol for another 4 days. Additional cells were cultured with NGF and 10 nM PMA for 4 days without ethanol. The percentage of cells with neurites was calculated from 80-120 cells/condition in 3-5 experiments. Cell line NGF NGF + PMA NGF + ethanol PC12 38  ± 1 57  ± 1a 60  ± 2a C1 38  ± 1 56  ± 2a 60  ± 2a V11 36  ± 1 56  ± 2a 62  ± 2a V12 40  ± 1 59  ± 1a 62  ± 1a V11 32  ± 2 30  ± 1 33  ± 3 V12 40  ± 1 40  ± 1 43  ± 1 a Significantly different compared with treatment with NGF alone (ANOVA and Scheffe F-test).

Table II. Neurite length after treatment with NGF plus PMA or ethanol Cells were cultured on polyornithine-coated tissue culture dishes with or without 100 mM ethanol for 6 days and then with 50 ng/ml NGF in the continued presence or absence of ethanol for another 4 days. Additional cells were cultured with NGF and 10 nM PMA for 4 days without ethanol. Neurite length is expressed in µm and was determined by measuring 100 neurites/condition in a representative experiment repeated three times. Cell line NGF NGF + PMA NGF + ethanol PC12 24.7  ± 1.5 47.6  ± 2.9a 46.4  ± 2.8a C1 23.6  ± 1.8 42.6  ± 2.4a 43.8  ± 3.2a V11 24.2  ± 1.6 39.3  ± 2.4a 46.5  ± 2.4a V12 24.2  ± 1.6 44.8  ± 2.1a 46.8  ± 2.9a V11 21.7  ± 1.4 27.4  ± 1.7 22.5  ± 1.4 V12 21.1  ± 1.7 29.0  ± 1.9 27.9  ± 1.9 a Significantly different compared with cells treated with NGF alone (ANOVA and Scheffe F-test).

Morphology of wild type and V1 and V1 transfected PC12 cells after NGF treatment.

V11

V12

V11

V12

NGF and basic fibroblast growth factor stimulate sustained activation of ERK1 and ERK2 MAP kinases in PC12 cells, which is important for their neuronal differentiation (37). Previous work has shown that treatment with either PMA or ethanol increases NGF-induced phosphorylation and activation of ERK1 and ERK2 (24). Since overexpression of PKC also enhances NGF-induced MAP kinase phosphorylation (27), we examined whether expression of V1 would prevent enhancement of MAP kinase activation by PMA or ethanol.

ERK1 and ERK2 are activated by dual phosphorylation on neighboring tyrosine and threonine residues (35). We measured activation of ERK1 and ERK2 by Western analysis using an anti-phospho-MAP kinase antibody that specifically detects phosphorylation of the activating tyrosine of each enzyme. As described previously in PC12 cells (24), NGF stimulated phosphorylation of ERK1 and ERK2 in C1 cells with a biphasic time course (Fig. 6A). Phosphorylation was maximal after 5-10 min (peak phase) and then declined to a lower level (plateau phase) that was maintained for at least 2 h. A similar pattern of phosphorylation was observed in V11, V12, V11, and V12 cells (Fig. 6). As previously observed in PC12 cells (24), co-treatment with 10 nM PMA or pretreatment with 100 mM ethanol for 6 days increased NGF-induced ERK phosphorylation. This was particularly evident during the plateau phase of ERK phosphorylation, which was elevated to levels achieved during the peak phase in C1 and V12 cells (Fig. 6A). A similar increase in phosphorylation was also observed in V11 cells (Fig. 6, B and C). In contrast, the plateau phase of ERK phosphorylation was not increased in V11 or V12 cells treated with ethanol or PMA (Fig. 6, A-C). Therefore, expression of the V1 fragment specifically inhibits PMA- or ethanol-induced ERK phosphorylation in NGF-treated cells.

The V1 fragment prevents enhancement of NGF-stimulated MAP kinase phosphorylation by PMA or ethanol.

V11

V11

DISCUSSION

The current results identify PKC as the PKC isozyme responsible for enhancement of NGF responses by phorbol esters and ethanol in PC12 cells. We previously found that phorbol esters enhance NGF-induced neurite outgrowth and MAP kinase activation in PC12 cells (11, 24). Ethanol also increases these responses to NGF by a PKC-dependent mechanism (11, 24). Recently, we found that overexpression of PKC also enhances responses to NGF (27), suggesting that PKC mediates the neurite-promoting effect of PMA and ethanol. In this paper, we investigated this issue directly, by creating PC12 cell lines that express peptides encoding V1 domains of PKC or PKC, which act as isozyme-selective translocation inhibitors (33). We found that expression of the V1 fragment selectively inhibited PMA-induced translocation of PKC, whereas expression of the V1 fragment specifically inhibited translocation of PKC. Cells expressing these peptides showed no alterations in cell growth. However, expression of V1 prevented enhancement of NGF-induced MAP kinase activation and neurite growth by PMA or ethanol. In contrast, expression of V1 did not alter enhancement of NGF responses by these agents. These findings indicate that PKC mediates enhancement of neurite outgrowth and MAP kinase activation by PMA or ethanol in NGF-treated PC12 cells.

Our data are consistent with a recent study (33) in which a V1 fragment, an V1 fragment, and an V1-derived peptide were introduced into cardiac myocytes by transient permeabilization. These fragments selectively inhibited PMA-induced translocation of their corresponding PKC isozyme and not translocation of other isozymes concomitantly activated in these cells. Furthermore, the V1 fragment and the short peptide derived from it inhibited phorbol ester- or hormone-induced regulation of contraction rate, whereas the V1 fragment or translocation inhibitors of PKC did not. Together with the data presented here, these studies reinforce the concept that translocation of PKC is required for its function (28) and indicate that isozyme-selective inhibitors of PKC translocation can be used to determine the function of individual isozymes in a variety of cells.

The ability of the PKC and - fragments to act as isozyme-selective translocation inhibitors is also consistent with the hypothesis that each contains a binding site for a corresponding, isozyme-specific RACK (28). This hypothesis is further supported by the finding of structural homology between V1 (32) and the C2 domain of conventional PKCs, which contains a RACK1 binding site (29). Indeed, an PKC-specific RACK, RACK2, that has recently been cloned, binds the V1 fragment in vitro.²

Prominent PKC immunoreactivity was found in neurites and growth cones of NGF-treated parent PC12 and C1 cells. This was reduced in processes of cells expressing V1, suggesting that V1 displaces endogenous PKC from binding sites in neurites and growth cones. Localization of PKC to growth cones is consistent with a role for this isozyme in regulating neurite outgrowth (38). Immunoprecipitation-kinase assays indicate that NGF activates PKC in PC12 cells (21), suggesting that NGF-induced localization of PKC to neurites and growth cones may involve activation of this isozyme. This suggests that an PKC-specific RACK may reside in growth cones and neurites.

In PC12 and C1 control cells, PKC immunoreactivity was most prominent asymmetrically next to the nucleus. Expression of V1 was associated with redistribution of PKC immunoreactivity to the perinuclear region and with an increase in nuclear size. It is difficult to speculate on the physiologic significance of these changes, since the function of PKC in these cells is not yet known. However, the findings clearly indicate that V1 alters the localization of PKC and produces a unique change in cell morphology.

NGF-induced activation of ERK1 and ERK2 involves phosphorylation and binding of phospholipase C and the adapter protein Shc to the NGF receptor tyrosine kinase TrkA (39). Within minutes of NGF binding, phosphorylated Shc also forms a complex with another adapter protein, Grb2, and with the guanine nucleotide exchange factor mSOS (40), leading to activation of Ras (41) and sequential activation of B-Raf (42), MAP kinase kinase-1 (43, 44), and ERK1 and ERK2 (45). This pathway appears essential for NGF-induced neurite outgrowth, since dominant negative inhibitors of Ras (46) or MAP kinase kinase-1 (47) block NGF-induced neurite outgrowth in PC12 cells. PKC could enhance ERK activation by increasing the activity of members of this pathway. For example, PKC appears to phosphorylate and activate Raf-1 (48), suggesting that other PKC isozymes, such as PKC, might also modulate Raf kinases. In addition, NGF stimulates binding of Shc to F-actin in PC12 cells (49). NGF also activates PKC (21), and upon activation, PKC binds F-actin in nerve terminals (50). This raises the intriguing possibility that Shc or proteins complexed with Shc interact with PKC when both Shc and PKC are anchored to actin. We do not yet know if the V1 fragment prevents binding of PKC to actin, since actin binds to PKC at a site between the first and second cysteine-rich regions of the C1 domain, which is outside of the V1 domain (50). Whether binding of PKC to actin is important for regulation of MAP kinases and neurite outgrowth also remains to be determined.

The major effect of phorbol esters or overexpressed PKC is to increase the late plateau phase of MAP kinase activation rather than the initial peak phase (24, 27). These kinetics suggest that PKC may act by inhibiting dephosphorylation of ERK1 and ERK2 rather than by promoting ERK activation. MAP kinase phosphatase (MKP)-1, MKP-2, and hVH3 are dual specificity protein phosphatases that are expressed in the brain and dephosphorylate and inactivate ERKs (51, 52). MKP-1 and MKP-2 mRNAs are constitutively expressed at low levels in PC12 cells and are increased by NGF (51). Inhibition of MKP-1 expression in PC12 cells does not accelerate the early phase of MAP kinase inactivation following stimulation with growth factors (53). Instead, protein phosphatase 2A and an unidentified protein-tyrosine phosphatase appear to mediate the rapid phase of inactivation in these cells (54). The role of dual specificity phosphatases in regulating the later plateau phase of NGF-induced MAP kinase activation is not known, but NGF induction of mRNA for MKP-1 and MKP-2 peaks at 1-2 h (51), suggesting that they may regulate this phase. If this is the case, then inhibition of these phosphatases by PKC may account for enhanced activation of ERK1 and ERK2.

ERK activity is also regulated by negative feedback inhibition. Phosphorylation of mSOS by MAP kinase promotes dissociation of mSOS from tyrosine-phosphorylated Shc and epidermal growth factor receptors (55, 56). In addition, MAP kinase kinase-dependent phosphorylation of mSOS has been reported to cause dissociation of mSOS-Grb2 complexes, interrupting mSOS activation of Ras (57). Moreover, ERK1 phosphorylates MAP kinase kinase, and this phosphorylation appears to reduce MAP kinase kinase activity (58). Therefore, inhibition of mSOS or MAP kinase kinase retrophosphorylation may be mechanisms by which PKC could enhance MAP kinase activation.

Together with studies in primary neurons (20, 22, 59), our findings suggest that PKC modulates neural differentiation. This may be a mechanism for enhancement of neurite growth by neurotransmitters that activate receptors coupled to phospholipase C and could be important for activity-dependent remodeling of synapses during normal development (60). Our studies also suggest that excessive activation of PKC may contribute to abnormal neurite growth observed in certain disease states. Chronic abuse of ethanol can damage the nervous system by disrupting the growth and remodeling of dendrites and axons. In certain brain regions, ethanol increases the growth of neural processes and terminals (61-65). This is particularly striking in the hippocampus, where expression of PKC is high (19). There prenatal exposure to ethanol causes marked overgrowth of dentate granule cell axons (mossy fibers) into the stratum pyramidale and stratum oriens of CA3, which may contribute to cognitive dysfunction (65). In addition, abnormal growth of mossy fibers into the supragranular layer of the dentate gyrus is found in some humans with temporal lobe epilepsy and in animals following stimuli that induce epilepsy (66). Future studies with transgenic and PKC mutant mice or with V1-derived inhibitory peptides may allow us to examine the role of PKC in normal development, alcohol-related neurologic disorders, and epileptogenesis.