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Endocrinology Vol. 138, No. 11 4721-4731
Copyright © 1997 by The Endocrine Society

ARTICLES

Evidence for Involvement of Protein Kinase C (PKC)-{zeta} and Noninvolvement of Diacylglycerol-Sensitive PKCs in Insulin-Stimulated Glucose Transport in L6 Myotubes1

Gautam Bandyopadhyay, Mary L. Standaert, Lamar Galloway, Jorge Moscat and Robert V. Farese

J. A. Haley Veterans Hospital and the Departments of Internal Medicine and Biochemistry/Molecular Biology (G.B., M.L.S., L.G., R.V.F.), University of South Florida College of Medicine, Tampa, Florida 33612; and Centro de Biologia Molecular "Severo Ochoa" (J.M.), Universidad Autonoma, Canto Blanco, 28049 Madrid, Spain

Address all correspondence and requests for reprints to: Robert V. Farese, M.D., Research Service (VAR 151), J. A. Haley Veterans Hospital, 13000 Bruce Downs Boulevard, Tampa, Florida 33612.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
We examined the question of whether insulin activates protein kinase C (PKC)-{zeta} in L6 myotubes, and the dependence of this activation on phosphatidylinositol (PI) 3-kinase. We also evaluated a number of issues that are relevant to the question of whether diacylglycerol (DAG)-dependent PKCs or DAG-insensitive PKCs, such as PKC-{zeta}, are more likely to play a role in insulin-stimulated glucose transport in L6 myotubes and other insulin-sensitive cell types. We found that insulin increased the enzyme activity of immunoprecipitable PKC-{zeta} in L6 myotubes, and this effect was blocked by PI 3-kinase inhibitors, wortmannin and LY294002; this suggested that PKC-{zeta} operates downstream of PI 3-kinase during insulin action. We also found that treatment of L6 myotubes with 5 µM tetradecanoyl phorbol-13-acetate (TPA) for 24 h led to 80–100% losses of all DAG-dependent PKCs ({alpha}, ß1, ß2, {delta}, {epsilon}) and TPA-stimulated glucose transport (2-deoxyglucose uptake); in contrast, there was full retention of PKC-{zeta}, as well as insulin-stimulated glucose transport and translocation of GLUT4 and GLUT1 to the plasma membrane. Unlike what has been reported in BC3H-1 myocytes, TPA treatment did not elicit increases in PKCß2 messenger RNA or protein in L6 myotubes, and selective retention of this PKC isoform could not explain the retention of insulin effects on glucose transport after prolonged TPA treatment. Of further interest, TPA acutely activated membrane-associated PI 3-kinase in L6 myotubes, and acute effects of TPA on glucose transport were inhibited, not only by the PKC inhibitor, LY379196, but also by both wortmannin and LY294002; this suggested that DAG-sensitive PKCs activate glucose transport through cross-talk with phosphatidylinositol (PI) 3-kinase, rather than directly through PKC. Also, the cell-permeable, myristoylated PKC-{zeta} pseudosubstrate inhibited insulin-stimulated glucose transport both in non-down-regulated and PKC-depleted (TPA-treated) L6 myotubes; thus, the PKC-{zeta} pseudosubstrate appeared to inhibit a protein kinase that is required for insulin-stimulated glucose transport but is distinct from DAG-sensitive PKCs. In keeping with the latter dissociation of DAG-sensitive PKCs and insulin-stimulated glucose transport, LY379196, which inhibits PKC-ß (preferentially) and other DAG-sensitive PKCs at relatively low concentrations, inhibited insulin-stimulated glucose transport only at much higher concentrations, not only in L6 myotubes, but also in rat adipocytes, BC3H-1 myocytes, 3T3/L1 adipocytes and rat soleus muscles. Finally, stable and transient expression of a kinase-inactive PKC-{zeta} inhibited basal and insulin-stimulated glucose transport in L6 myotubes. Collectively, our findings suggest that, whereas PKC-{zeta} is a reasonable candidate to participate in insulin stimulation of glucose transport, DAG-sensitive PKCs are unlikely participants.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
L6 MYOTUBES are particularly important in studies of insulin action, as these cultured cells serve as a useful model of skeletal muscle, the most important tissue for insulin-mediated glucose disposal in vivo. With respect to insulin signaling systems that are operative in L6 myotubes, insulin has been reported to provoke increases in diacylglycerol (DAG) (1) and activate DAG-sensitive protein kinase Cs (PKCs) (1, 2) in L6 myotubes. A role for these PKCs in insulin-stimulated glucose transport, however, was initially discounted, because phorbol ester-induced down-regulation of DAG-sensitive, total PKC enzyme activity does not substantially inhibit insulin effects on glucose transport in L6 myotubes (3). Nevertheless, both phorbol esters as DAG analogs (3) and DAG itself (4) provoke insulin-like effects on glucose transport, and PKC inhibitors, at least in relatively high concentrations, block insulin effects on glucose transport (1) in L6 myotubes. Furthermore, it has recently been suggested (5) that PKC-ß2 is largely or totally responsible for the effect of insulin on glucose transport in L6 myotubes because this effect is inhibited by (a) transient expression of a C-terminal deleted (truncated) form of PKC-ß2, and (b) a semiselective PKC-ß2 inhibitor, CG53353. Obviously, these seemingly contradictory findings in the L6 system need to be resolved.

Several other recent findings are germane to the conundrum described above. On the one hand, in support of the argument that DAG-sensitive PKCs are required for insulin action, phorbol ester-induced down-regulation of PKC has been reported to be attended by an alteration in PKC-ß messenger RNA (mRNA) splicing, thus causing a decrease in PKC-ß1 and a paradoxical increase in PKC-ß2 in BC3H-1 myocytes; moreover, this mechanism has been suggested to account for retention of glucose transport effects of insulin in this experimental paradigm (6). On the other hand, several other recent findings militate against the involvement of DAG-sensitive PKCs, and speak for the involvement of the DAG-insensitive PKC, viz., PKC-{zeta}, in insulin-stimulated glucose transport. First, phorbol esters have been found to activate a membrane-associated form of phosphatidylinositol (PI) 3-kinase in rat adipocytes (7) and 3T3/L1 cells (8), and the relatively small effects of phorbol esters on glucose transport in 3T3/L1 cells [although not in rat adipocytes (7)] are blocked by the PI 3-kinase inhibitor, wortmannin (8): thus, DAG-sensitive PKCs do not directly activate glucose transport and are unlikely to serve as downstream effectors for PI 3-kinase-dependent increases in glucose transport, at least in some cell types. Second, we have recently found that insulin provokes an increase in the enzymatic activity of immunoprecipitable PKC-{zeta} in 3T3/L1 cells (9). Third, in transfection studies in 3T3/L1 fibroblasts and adipocytes, we have found that: 1) stable expression of wild-type PKC-{zeta}, but not PKC-{alpha} or PKC-ß2, provokes increases in basal and insulin-stimulated glucose transport; and 2) stable expression of a kinase-inactive mutant form of PKC-{zeta} inhibits basal and insulin-stimulated glucose transport (9).

Presently, we have attempted to gain further insight into the question of whether DAG-sensitive or DAG-insensitive PKCs are more likely to play a role in insulin-stimulated glucose transport in L6 myotubes. To this end, we have studied in L6 myotubes: 1) the activation of immunoprecipitable PKC-{zeta} by insulin and its dependence on PI 3-kinase; 2) the completeness of depletion of specific PKC isoforms during prolonged, high-dosage phorbol ester treatment; 3) effects of phorbol ester on PKC-ß1 and PKC-ß2 mRNA; 4) effects of phorbol esters on PI 3-kinase; 5) effects of inhibitors of PI 3-kinase on acute, phorbol ester-induced increases in glucose transport; and 6) effects of stable and transient transfection of wild-type and kinase-inactive PKC-{zeta} on glucose transport. We also studied the effects of selective inhibitors of PKC-ß and other DAG-sensitive PKCs on insulin-stimulation of glucose transport both in L6 myotubes, and several other commonly used insulin-sensitive cell types, i.e. rat adipocytes, 3T3/L1 adipocytes, BC3H-1 myocytes, and rat soleus muscles.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
L6 cells (a gift of Dr. Amira Klip) were cultured to confluence and differentiated to myotubes as described (1). Myotubes were equilibrated at 37 C in Krebs Ringer phosphate (KRP) buffer containing 0.1% BSA. Vehicle, insulin (Elanco, Indianapolis, IN), tetradecanoyl phorbol-13-acetate (TPA) (Sigma Chemical Co., St. Louis, MO), myristoylated PKC-{zeta} pseudosubstrate (Quality Controlled Biochemicals, Inc., Hopkinton, MA), wortmannin (Sigma), LY294002 (BioMol Research Laboratories, Inc., Plymouth Meeting, PA), LY379196 (a selective PKC-ß inhibitor, kindly provided by Drs. Michael Jirousek and Kirk Ways, Eli Lilly Co.), CG53353 (a selective PKC-ß2 inhibitor, kindly provided by Drs. Anna Suter and Doriano Fabbro, Ciba Geigy Corp.), and other substances were added to the medium, and incubations were conducted as described in the text.

PKC isoforms in total cell lysates were immunoblotted as described (1, 10, 11, 12), except that quantitation of chemiluminescence was accomplished with a BioRad 32P/Chemiluminescence Molecular Analyst Imaging System, which provides linear responses for both 32P and chemiluminescence. Polyclonal antisera for PKC-{alpha}, {delta}, {epsilon} and {zeta} were obtained from Life Technologies (Grand Island, NY). Polyclonal antisera for PKC-ß1 and PKC-ß2 were obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Antisera specificities were confirmed by 1) immunoabsorption (i.e. loss of immunoreactivity) with the immunizing peptide and 2) by blotting against recombinant PKCs obtained from Sf9 insect cells infected with baculovirus containing isoform-specific PKC complementary DNA (cDNA), and/or NIH3T3, 3T3/L1 or L6 cells transfected with cDNAs encoding respective PKCs (see Refs. 9, 10, 12, 13, and Santa Cruz catalog). We presently could not detect PKC-{theta} in L6 myotubes using methods successfully applied to mature rat skeletal muscle (12). GLUT4 and GLUT1 glucose transporter levels were measured by Western analysis using antisera obtained from Charles River.

PKC-{zeta} enzyme activity was measured in specific immunoprecipitates as described (9). Cells were lysed by sonication in Buffer I containing 20 mM Tris-HCl (pH 7.5), 0.25 M sucrose, 1.2 mM EGTA, 20 mM ß-mercaptoethanol, 1 mM phenylmethylsulfonylfluoride (PMSF), 20 µg/ml leupeptin, 20 µg/ml aprotinin, 1 mM Na pyrophosphate, 1 mM NaF, 1% Triton X-100, 0.5% Nonidet and 150 mM NaCl. Lysates were first cleared with rabbit preimmune serum, followed by treatment with second antibody (goat antirabbit IGG antiserum) and protein AG-sepharose beads. PKC-{zeta} was then immunoprecipitated by overnight treatment at 4 C with anti-PKC-{zeta} antiserum obtained from either Life Technologies or Santa Cruz Biotechnology, followed by addition of second antibody. Precipitates were collected on protein AG-sepharose beads, washed and suspended in Buffer II containing 50 mM Tris/HCl (pH, 7.5), 5 mM MgCl2, 100 µM Na3VO4, 100 µM Na3P2O7, 1 mM NaF and 100 µM PMSF. The suspension was then incubated for 8 min at 30 C in 100 µl Buffer II containing 3–5 µCi [{gamma}-32P]ATP (New England Nuclear, Boston, MA), 50 µM ATP, 4 µg phosphatidylserine (PS), and 40 µM[159Ser]-PKC-{epsilon}(AA153–164)-NH2 (Upstate Biotechnology, Inc., Lake Placid, NY) [this PKC-{epsilon} pseudosubstrate analog is an excellent substrate for PKC-{zeta} (14) and we have found that this substrate is phosphorylated very well by PKC-{zeta} and PKC-{epsilon} (both set at 100%), and poorly by PKC-{alpha} (35%), PKC-ß1 (24%), PKC-ß2 (10%) and PKC-{delta} (5%) using PKCs purified from Sf9 insect cells infected with cDNAs encoding these PKCs (kindly supplied by Dr. Larry Ballas, Sphinx Division, Eli Lilly Co.). PKC-{zeta} pseudosubstrate (as a relatively specific PKC-{zeta} inhibitor) was used in the assay to determine blank values, i.e. cpm incorporated independently of PKC-{zeta} (~15% of total cpm), which were subtracted from total cpm. After incubation, an aliquot of the reaction mixture was spotted on p81 paper, washed in 5% acetic acid, and counted. As reported elsewhere (9), PKC-{zeta} immunoprecipitates contained no detectable PKC-{alpha}, ß, {delta} or {epsilon}; additionally, recovery of PKC-{zeta} was approximately 50% and was not influenced by insulin treatment or improved by adding a 2-fold excess of antibody.

PKC-ß1 mRNA and PKC-ß2 mRNA were measured by ribonuclease protection assay (RPA) as described (9, 13) (original cDNAs were kindly provided by Dr. Yoshitaka Ono, Kobe, Japan). In these RPAs, the protected RNA fragments spanned the splice sites and extended well into both adjacent exons of fully processed PKC-ß1 mRNA (base 1792 to 2058) and PKC-ß2 mRNA (base 1792 to 1962); accordingly PKC-ß1 and PKC-ß2 mRNAs cannot be confused with prespliced (smaller) precursors (note - the splice site is at base 1865 of the coding region of rat PKC-ß). The validity of this method has been confirmed by observing specific increases in PKC-ß2 (but not PKC-ß1) mRNA in 3T3/L1 cells transfected with cDNA encoding PKC-ß2 mRNA (9). Quantitation was accomplished with the Biorad Phosphorimager Molecular Analyst Imaging System.

Wild-type PKC-{zeta} and PKC-ß2 and kinase-inactive PKC-{zeta} (i.e. point-mutated in the ATP-binding site; see Ref.15), each contained in a pCDNA3 eukaryotic expression vector (Invitrogen, Carlsbad, CA), were transfected into L6 cells using Lipofectamine according to instructions of the manufacturer (Life Technologies). In stable transfection experiments, colonies were selected by resistance to G418 (500 µg/ml), cultured to confluence in 24-well plates (for glucose transport studies) or 100-mm plates (for immunoblot studies), and then differentiated to myotubes as described above. Growth rates and differentiation did not appear to be influenced by the transfection of vectors or vectors containing inserted PKC-{zeta} cDNAs. In transient transfection experiments, fully differentiated L6 myotubes were transfected directly in 24-well plates and then allowed to incubate for 2 days in fresh medium before experimental use. Transfection with ß-galactosidase (ß-gal)-containing constructs (also in pCDNA3) were used to judge transient transfection rates, which were at least approximately 25%.

[3H]2-Deoxyglucose (2-DOG) uptake in L6 myotubes was measured as described (1). In these assays, cells were incubated in glucose-free KRP with or without inhibitors as described in the figure legends and then stimulated for 30 min with or without agonist (insulin or TPA), followed by determination of uptake during a 5-min incubation in medium containing 0.1 mM [3H]2-DOG (New England Nuclear). In a more limited number of experiments, 2-DOG uptake was also measured in rat adipocytes, 3T3/L1 adipocytes, BC3H-1 myocytes and rat soleus muscles, as described previously (7, 9, 16, 17, respectively).

Membrane-associated PI 3-kinase activity was measured as described previously (7). In brief, total membranes were obtained by centrifugation of postnuclear homogenates at 100,000 x g for 60 min, and 100 µg membrane protein were incubated for 10 min with 50 µM [{gamma}32P]ATP, 10 mM MgCl2, 20 µg PI, and other components of the assay system in a final volume of 75 µl. After incubation, PI-3-PO4 was extracted, resolved by TLC, and counted in the Biorad Phosphorimager Molecular Analyst Imaging System.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effects of insulin and PI 3-kinase inhibitors on PKC-{zeta} activation in L6 myotubes
Although total PKC enzyme activity has been reported to be activated by insulin in L6 myotubes (1), there is no direct information on whether or not PKC-{zeta} is specifically activated in these cells. Presently, we found that 20 nM insulin provoked, on the average, 80% increases in the enzyme activity of immunoprecipitable PKC-{zeta} in L6 myotubes (Fig. 1Go). Moreover, PI 3-kinase appeared to be required for the activation of PKC-{zeta}, as this effect of insulin was not observed in myotubes incubated in the presence of either 100 nM wortmannin or 100 µM LY294002 (Fig. 1Go).



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  		Figure 1. Effects of insulin, wortmannin, and LY294002 on immunoprecipitable PKC-{zeta} enzyme activity in L6 myotubes. Cells were treated for 15 min with or without 100 nM wortmannin or 100 µM LY294002, and then for 10 min with or without 20 nM insulin. Values are mean ± SE of (n) comparisons of control and insulin-treated samples in multiple experiments with the controls arbitrarily set at 100%. P was determined by paired t test, insulin vs. control.

 
Depletion of PKC isoforms during TPA-induced PKC down-regulation in L6 myotubes
As shown in Fig. 2Go, treatment of L6 myotubes with 5 µM TPA for 24 h was attended by large (80–100%) losses of immunoreactive PKC-{alpha}, ß1, ß2, {delta} and {epsilon}; concomitantly, acute effects of TPA on glucose transport were completely lost in down-regulated myotubes. The DAG-insensitive PKC-{zeta}, on the other hand, was fully retained in TPA-treated myotubes, and this correlated well with full retention of insulin-stimulated (a) 2-DOG uptake (Fig. 2Go) and (b) translocation of both GLUT4 and GLUT1 to the plasma membrane (Fig. 3Go). Unlike what was reported in studies of BC3H-1 myocytes (see Refs. 6 and 18), neither PKC-ß2 protein (Fig. 2Go) nor PKC-ß2 mRNA (Fig. 4Go) was induced by 24-h TPA treatment in L6 myotubes. (It may be noted that acute TPA treatment, like insulin, provoked increases in plasma membrane levels of both GLUT4 and GLUT1-see Fig. 3Go).



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  		Figure 2. Effects of prolonged phorbol ester (TPA) treatment on levels of PKC isoforms, and insulin-stimulated and TPA-stimulated 2-deoxyglucose (2-DOG) uptake in L6 myotubes. Cells were treated with 5 µM TPA for 24 h as indicated and then thoroughly rinsed and placed into glucose-free KRP medium for measurement of acute 2-DOG uptake ( ± 200 nM insulin or 500 nM TPA) and immunoreactive PKC. A, Representative immunoblots show levels of indicated PKCs in untreated control cells and cells down-regulated for 24 h with 5 µM TPA; bargrams depict mean changes of four closely agreeing determinations, plotted as a function of the control set at 100%. B, Bargrams depict relative changes in insulin- and TPA-stimulated 2-DOG uptake in non-down-regulated (NON-DR) and TPA-down-regulated (DR) cells. Rates of [3H]2-DOG uptake in control and insulin-treated cells were not influenced significantly by 24-h TPA treatment: i.e. results in non-down-regulated vs. TPA-down-regulated control cells were 5206 ± 281 vs. 4946 ± 1019 cpm/well·5 min, respectively (mean ± SE, n = 3); results in non-down-regulated vs. TPA-down-regulated insulin-treated cells were 20938 ± 1398 and 20669 ± 1228 cpm/well/5 min, respectively (mean ± SE, n = 3). Similar results were obtained in at least four separate experiments (also see Ref. 3 for retention of glucose transport responses to insulin and decreases in total PKC enzyme activity in down-regulated cells). [It may be noted that insulin effects on glucose transport were greater (4-fold) in these experiments and exceeded the effects of TPA: this may reflect the fact that 200 nM insulin was used herein, whereas 20 nM insulin was used in most other studies.]

 


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  		Figure 3. Acute effects of insulin and/or TPA on plasma membrane levels of GLUT4 and GLUT1 glucose transporters in non-down-regulated and PKC-down-regulated L6 myotubes. As indicated, cells were treated for 24 h with or without 5 µM TPA, before acute treatments, as in Fig. 2Go. GLUT4 and GLUT1 levels in the plasma membrane were assessed by immunoblotting, measured after acute treatment for 30 min with or without 100 nM insulin or 500 nM TPA, as indicated. The results of a representative experiment are shown in the immunoblots: the bargrams depict mean values of two experiments. Although not shown, there were reciprocal changes in microsomal levels of GLUT4 and GLUT1.

 


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  		Figure 4. Effects of prolonged TPA treatment on levels of PKC-ß1 mRNA and PKC-ß2 mRNA in L6 myotubes. Cells were treated with 5 µM TPA for 24 h as in Fig. 2Go, and 10Go µg of total RNA was subjected to RPA analysis as described in Materials and Methods. Shown here are: 1) representative autoradiograms of protected mRNA fragments [migrating as expected (see Ref. 13) just slightly below the level of the intact probe]; and 2) mean ± SE values of four to five determinations with the mean control value set at 100%.

 
Effects of PI 3-kinase inhibitors on TPA-and insulin-stimulated glucose transport in L6 myotubes
Both wortmannin (Fig. 5Go) and LY294002 (Fig. 6Go) inhibited TPA-, as well as insulin-stimulated, 2-DOG uptake in L6 myotubes: maximal inhibition was observed at 50 nM wortmannin and 50 µM LY294002, concentrations that are well within the range of those reported to inhibit PI 3-kinase (19, 20). These findings suggested that: 1) phorbol esters activate glucose transport by cross-talking of PKC with PI 3-kinase; and 2) DAG-sensitive PKCs cannot be viewed simply as serving as downstream effectors for PI 3-kinase-dependent activation of glucose transport in L6 myotubes. It may also be noted that basal 2-DOG uptake was mildly inhibited by wortmannin and LY294002, suggesting the presence of significant (although variable; see below) basal PI 3-kinase activity in control myotubes. (This was also noted previously; see Ref.19).



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  		Figure 5. Dose-related effects of wortmannin on TPA-stimulated and insulin-stimulated [3H]2-deoxyglucose uptake in L6 myotubes. Cells were incubated for 15 min with inhibitor before addition of 500 nM TPA or 20 nM insulin. Values are mean ± SE of three determinations.

 


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  		Figure 6. Dose-related effects of LY294002 on TPA-stimulated and insulin-stimulated [3H]2-deoxyglucose uptake in L6 myotubes. Cells were incubated for 15 min with inhibitor before addition of 500 nM TPA or 20 nM insulin. Values are mean ± SE of three determinations.

 
Effects of TPA on membrane-associated PI 3-kinase in L6 myotubes
As shown in Fig. 7Go, both insulin and TPA provoked increases in membrane-associated PI 3-kinase activity. This finding is similar to those observed in rat adipocytes (7) and 3T3/L1 adipocytes (8), and, moreover, provides an explanation for the inhibitory effects of wortmannin and LY294002 on acute effects of TPA on glucose transport in L6 myotubes.



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  		Figure 7. Effects of TPA and insulin on membrane-associated PI 3-kinase activity. Myotubes were treated for 10 min with vehicle (control), 100 nM insulin, or 500 nM TPA. Membranes were then isolated and assayed for PI 3-kinase activity. A representative autoradiogram is shown here. Bargrams indicate relative values (mean ± SE) for five determinations. *, P < 0.025 (paired t test).

 
Effects of myristoylated PKC-{zeta} pseudosubstrate on insulin and TPA-stimulated glucose transport in L6 myotubes
The findings described above were compatible with the possibility that a DAG-insensitive PKC, such as PKC-{zeta}, and/or another protein kinase, distinct from DAG-sensitive PKC family members, may operate downstream of PI 3-kinase and participate in both insulin-stimulated and TPA-stimulated glucose transport. Accordingly, it was of interest to find that the cell-permeable, myristoylated PKC-{zeta} pseudosubstrate (myr-SIRRGARRWRKL-NH2) inhibited insulin-stimulated, and to a variable extent, basal 2-DOG uptake, both in non-down-regulated L6 myotubes and in L6 myotubes down-regulated by 24-h treatment with 5 µM TPA (Figs. 8Go and 9Go). Inhibition of insulin effects was complete at 10–30 µM PKC-{zeta} pseudosubstrate, concentrations of myristoylated pseudosubstrate that are well within the range of those reported to inhibit PKC in other cellular systems (see Ref.21). The partial, but variable, inhibition of basal 2-DOG uptake by the PKC-{zeta} pseudosubstrate, like that occurring with wortmannin and LY294002, probably reflects significant activation PI 3-kinase and a distal protein kinase(s) in control L6 myotubes. Also, it may be noted that, despite significant differences in amino acid sequences of various PKC pseudosubstrates (see Ref.22), the isoform specificity of the PKC-{zeta} pseudosubstrate is presently uncertain, and it is possible, if not likely, that it may also inhibit PKCs other than PKC-{zeta}: nevertheless, the inhibitory effects of the PKC-{zeta} pseudosubstrate in L6 myotubes in which all DAG-sensitive PKCs were effectively depleted by 24-h TPA treatment, suggested that a TPA-resistant PKC, possibly PKC-{zeta}, may be required for insulin-stimulated glucose transport.



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  		Figure 8. Effects of myristoylated PKC-{zeta} pseudosubstrate on insulin-stimulated [3H]2-deoxyglucose Uptake in non-down-regulated and TPA-down-regulated L6 myotubes. As indicated, cells were incubated for 24 h without (non-down-regulated) or with (down-regulated) 5 µM TPA to deplete DAG-sensitive PKCs. After rinsing and equilibrating in glucose-free KRP medium, the cells were incubated with indicated doses of myristoylated (Myr) pseudosubstrate for 90 min to allow for sufficiently complete uptake and then treated with or without 20 nM insulin for 30 min before measurement of 2-DOG uptake. Values are mean ± SE of four determinations.

 


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  		Figure 9. Effects of myristoylated PKC-{zeta} pseudosubstrate on TPA- and insulin-stimulated 2-DOG uptake in L6 myotubes. Cells were incubated for 90 min with or without indicated concentrations of myristoylated PKC-{zeta} pseudosubstrate (myr-SIRRGARRWRKL-NH2, solid lines) or a similarly sized control peptide (myr-ITRARRAPSVAN-NH2, broken lines) and then treated with or without 500 nM TPA or 20 nM insulin (as indicated) for 30 min before measurement of 2-DOG uptake. Values are mean ± SE of three to six determinations.

 
In addition to inhibiting insulin effects on glucose transport, the myristoylated PKC-{zeta} pseudosubstrate inhibited TPA-stimulated 2-DOG uptake in L6 myotubes (Fig. 9Go). This inhibition, however, appeared to be slightly less complete than that observed with insulin, suggesting that a small part of the TPA effect may be mediated by a protein kinase or other factor that is relatively resistant to the PKC-{zeta} pseudosubstrate. It may also be noted that a comparably sized, myristoylated peptide containing the amino acid sequence, myr-ITRARRAPSVAN-NH2, did not inhibit insulin-stimulated 2-DOG uptake (Fig. 9Go), suggesting at least some degree of specificity of the sequence of the PKC-{zeta} pseudosubstrate.

Effects of expression of transfected PKC-{zeta} on insulin-stimulated glucose transport in L6 myotubes
Stable expression of wild-type PKC-{zeta} and PKC-ß2 provoked 2-fold increases in immunoreactive levels of these PKCs but failed to affect insulin effects on glucose transport (data not shown); it may therefore be surmised that these PKC isoforms, at least in their inactive forms, are present in amounts that are not rate-limiting for glucose transport. On the other hand, stable expression of kinase-inactive PKC-{zeta} provoked a 50% decrease in basal and insulin-stimulated 2-DOG uptake (Fig. 10Go). Concomitantly, immunoreactive PKC-{zeta} was increased approximately 2-fold [i.e. an increase of 127 ± 11% (mean ± SE; n = 7)] above control] in cells stably transfected with kinase-inactive PKC-{zeta} (Fig. 10Go): this correlated well with the observed 50% inhibition of glucose transport, if it is assumed that transfected, catalytically inactive PKC-{zeta} competes equally with native, catalytically active PKC-{zeta} for either activating substances and/or substrates. Along these lines, it may be noted that in previous studies of 3T3/L1 adipocytes(9), we observed that, as in the present study, stable transfection of the same kinase-inactive PKC-{zeta} led to a 2-fold increase in immunoreactive PKC-{zeta}, with no change in total cellular PKC-{zeta} enzyme activity (9) (this is the expected result, if the transfected PKC-{zeta} is catalytically inactive, but does not inhibit endogenous PKC-{zeta} enzyme): this suggested that the specific enzyme activity of total cellular PKC-{zeta} was decreased by 50% in these transfected 3T3/L1 cells. We presume that the same situation prevailed in L6 myotubes stably transfected with kinase-inactive PKC-{zeta}.



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  		Figure 10. Effects of stable (upper panel) and transient (lower panel) expression of kinase-inactive (DN) PKC-{zeta} on insulin-stimulated [3H]2-deoxyglucose uptake in L6 myotubes. Values are mean ± SE of (n) clones, each assayed in triplicate at each of the designated insulin concentrations. Insets show representative blots depicting increases in immunoreactive PKC-{zeta} in cells transfected with kinase-inactive (DN) PKC-{zeta} (also see text for quantitation of mean increases in multiple samples). *, P < 0.05, t test.

 
It may be noted that inhibition of basal glucose transport by kinase-inactive PKC-{zeta} was not unexpected because basal transport was also frequently inhibited (albeit to a variable extent) by wortmannin, LY294002 and myristoylated PKC-{zeta} pseudosubstrate (this inhibition of basal activity probably reflects variable stimulatory effects of residual serum or endogenous paracrine or autocrine activators that may have survived the washings and preincubations, or continue to be released from the myocytes, thus raising the basal transport level). It may also be noted that the inhibition of glucose transport by kinase-inactive PKC-{zeta} could not be explained by changes in levels of GLUT4 and/or GLUT1 glucose transporters, as these were increased by 80 ± 17 (n = 5; P < 0.01) and 88 ± 29 (n = 6; P < 0.05) percent, respectively. Interestingly, increases in Glut4 and/or Glut1 glucose transporters were also observed in 3T3/L1 cells stably transfected with kinase-inactive PKC-{zeta} (9); it is therefore possible that decreases in basal and insulin-stimulated glucose transport may have resulted in compensatory increases in Glut4 and/or Glut1 glucose transporters. Whatever the explanation, the decrease in basal and insulin-stimulated glucose transport in these transfected cells, in the face of increases in levels of Glut4 and Glut1, suggested that 1) the levels of these glucose transporters were not rate-limiting for basal glucose transport; and 2) the inhibitory effects of kinase-inactive PKC-{zeta} may have been partly offset by increases in Glut4 and Glut1 glucose transporters.

Similar to changes observed in stable transfectants, transient transfection of kinase-inactive PKC-{zeta} led to a modest (~40%) but significant inhibition of insulin-stimulated glucose transport (Fig. 10Go). This inhibition was associated with approximately 2-fold increases in immunoreactive PKC-{zeta}, which, based upon transfection rates with ß-gal, appeared to reside in at least 25% of the cells. Inasmuch as the latter may have been underestimated by microscopic analysis, it may be surmised that there were approximately 4- to 8-fold increases in PKC-{zeta} in these transiently transfected cells, and this may explain the apparently greater relative (perhaps full) inhibitory effect of kinase-inactive PKC-{zeta} in transient transfection experiments.

Effects of selective PKC-ß inhibitors (LY379196 and CG53353) on insulin-stimulated glucose transport in L6 myotubes and other cell types
As our results suggested that PKC-ß and other DAG-sensitive PKCs are not likely to be important for insulin effects on glucose transport, it was of interest to use a PKC inhibitor that provided a wide difference between inhibitory effects on PKC-ß and other PKCs. For this purpose, we used LY379196, which (as determined with recombinant, baculovirus-derived PKCs, purified from Sf9 insect cells by Dr. Michael Jirousek, Eli Lilly Co.) provides a difference of nearly three orders of magnitude between the inhibition of PKC-ß1 (IC50, 50 nM) and PKC-ß2 (IC50, 30 nM), on the one hand, and PKC-{zeta} [IC50, 48 µM], on the other hand. [Note: IC50s for PKC-{alpha}, PKC-{gamma}, PKC-{delta}, PKC-{eta} and PKC-{epsilon} = 0.6, 0.6, 0.7, 0.3, and 5 µM, respectively; also note that these IC50s are similar in relative distribution to those of another, structurally similar, selective, PKC-ß inhibitor, LY333531; see Ref. 23]. As shown in Fig. 11Go, LY379196, in concentrations up to 10 µM, had no effect on control or insulin-stimulated 2-DOG uptake, and caused inhibition only at higher concentrations in both L6 myotubes and rat adipocytes. In contrast, LY379196, in relatively low concentrations, fully inhibited TPA-stimulated DOG uptake in L6 myotubes (Fig. 11Go); this confirmed the fact that this inhibitor readily entered these cells and effectively inhibited DAG-sensitive PKCs in these experiments. Similarly, L379196, in concentrations up to 10 µM, failed to inhibit 2-DOG uptake in 3T3/L1 adipocytes, BC3H-1 myocytes and rat soleus muscles (Fig. 12Go). Goe 6796, a selective inhibitor of PKC-{alpha}, ß and {gamma} (24), also failed to inhibit 2-DOG uptake in these cells (data not shown). These findings provided further evidence suggesting that PKC-ß1, PKC-ß2 and probably most other DAG-sensitive PKCs, are not likely to contribute importantly to insulin effects on glucose transport.



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  		Figure 11. Effects of LY379196 on insulin-stimulated and TPA-stimulated [3H]2-deoxyglucose uptake in L6 myotubes and rat adipocytes. Cells were equilibrated for 45–60 min in glucose-free KRP medium with indicated concentrations of LY379196 to allow for sufficiently complete uptake and then treated with or without 20 nM insulin or 500 nM TPA (myotubes, panels A and C) or 10 nM insulin (adipocytes, panel B) or increasing concentrations of insulin (adipocytes, panel D) for 30 min before measurement of 2-DOG uptake over 5 min (myotubes) or 1 min (adipocytes). Values are mean ± SE of either (n) or four determinations.

 


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  		Figure 12. Effects of LY379196 on insulin-stimulated [3H]2-deozyglucose uptake in 3T3/L1 adipocytes, rat soleus muscles, and BC3H-1 myocytes. Incubations were conducted as in Fig. 11Go, with additions as indicated. Values are mean ± SE of four determinations.

 
In addition to LY379197, we used CG53353, a somewhat less selective inhibitor of PKC-ß2 [as reported (5), the IC50 for PKC-ß2 is 1 µM and other PKCs are inhibited at slightly higher concentrations of CG53353]. As shown in Fig. 13Go, this inhibitor elicted similar dose-dependent inhibition curves for insulin-stimulated 2-DOG uptake in non-down-regulated and TPA-down-regulated L6 myotubes. Inasmuch as PKC-ß2 and other DAG-sensitive PKCs were markedly depleted in TPA-treated cells, it may be surmised that the inhibition of insulin-stimulated glucose transport by CG53353 cannot be attributed to the inhibition of PKC-ß2.



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  		Figure 13. Effects of CG53353 on insulin-stimulated [3H]2-deoxyglucose uptake in non-down-regulated and TPA-down-regulated L6 Myotubes. Where indicated, cells were incubated for 24 h without (open symbols) or with (closed symbols) 5 µM TPA to deplete DAG-sensitive PKCs. Cells were then washed and equilibrated for 45 min in glucose-free KRP medium with indicated concentrations of CG53353 to allow for sufficiently complete uptake, and then treated with (INS; circles) or without (CON; squares) 20 nM insulin for 30 min before measurement of 2-DOG uptake. Values are mean ± SE of four determinations.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
There appears to be a general consensus that IRS-1 and PI 3-kinase play pivotal roles in the regulation of glucose transport by insulin. Factors that operate downstream of PI 3-kinase, however, are less certain, but it nevertheless appears likely that a protein kinase (see Refs. 25 and 26) is required, as the PKC inhibitor, RO 31–8220, inhibits effects of insulin on glucose transport, without inhibiting the activation of PI 3-kinase (26). Although DAG-sensitive PKCs, e.g. {alpha} and ß, may operate downstream of PI 3-kinase (see 26), the full retention of insulin effects on glucose transport and translocation of both GLUT4 and GLUT1 to the plasma membrane in L6 myotubes (ref 3, and present findings) and certain other cells, e.g. 3T3/L1 cells (27), after prolonged phorbol ester treatment and marked depletion of DAG-sensitive PKCs, argues against their involvement in glucose transport. Indeed, as shown presently in L6 myotubes, TPA provoked marked losses of all detectable DAG-sensitive PKCs ({alpha}, ß1, ß2, {delta}, {epsilon}) without altering insulin effects on glucose transport. Moreover, contrary to findings in BC3H-1 myocytes (6, 18), TPA did not induce increases in PKC-ß2 mRNA or protein in L6 myotubes; it therefore seems clear that retention of the PKC-ß2 isoform is unlikely to explain the retention of insulin effects on glucose transport in TPA-down-regulated myotubes.

As stated above, from the present and previous (3) findings in down-regulated cells, it seems most unlikely that DAG-sensitive PKCs serve as required kinases that operate downstream of PI 3-kinase in insulin-stimulated glucose transport. As may be surmised, this conclusion differs from that of another study in which transient expression of a nine amino acid, but not a sixteen amino acid, C-terminal, truncated form of PKC-ß2, led to apparently complete inhibition of insulin-stimulated glucose transport in L6 myotubes (5). However, it is presently not clear why the sixteen amino acid-truncated form of PKC ß2 failed to inhibit insulin effects on glucose transport, as this form was enzymatically even less active than the nine amino acid-truncated form. This discrepancy was explained by postulating that the smaller truncated form of PKC-ß2 could function as a dominant-negative inhibitor of intact endogenous PKC-ß2 by virtue of an amino acid sequence that was present in the nine amino-acid form and was required for recognition of substrate or an intracellular receptor (5). This explanation is plausible but speculative. In contrast to findings of Chalfant et al. (5), our findings with PKC inhibitors and PKC depletion seem to be less subject to speculative interpretation, and, in our view, strongly suggest that PKC-ß2 is not required for insulin stimulation of glucose transport. Similarly, our finding that acute effects of TPA on glucose transport were blocked by wortmannin and LY294002 leaves little doubt that DAG-sensitive PKCs, including PKC-ß2, cannot serve to directly activate the glucose transport process during TPA treatment in L6 myotubes. Nevertheless, we are currently examining the role of PKC-ß in studies of muscles of mice in which PKC-ß has been knocked out by homologous recombination methodology.

Further support for our contention that a DAG-insensitive, rather than a DAG-sensitive, PKC is more likely to be involved in insulin stimulation of glucose transport derives from two other lines of evidence, i.e. 1) the PKC-{zeta} pseudosubstrate and CG53353, a general PKC inhibitor (5), inhibited insulin-stimulated 2-DOG uptake in cells largely depleted of DAG-sensitive PKCs (i.e. after 24-h TPA treatment); and 2) inhibitory effects of a highly selective PKC-ß inhibitor, LY379196, on insulin-stimulated 2-DOG uptake were observed only at concentrations that were far in excess of those required to inhibit PKC-ß and probably most other DAG-sensitive PKCs. Obviously, the higher concentrations of LY379196 that inhibited 2-DOG uptake may have inhibited PKC-{zeta}, other atypical PKCs, e.g. {lambda}, and/or other related kinases. Along the latter lines, it may be noted that we have previously reported that, at concentrations of 5 µM or greater, R0 31–8220, a relatively nonspecific PKC inhibitor, blocks insulin-stimulated glucose transport in L6 myotubes (1). Of further interest, we have found that comparably high concentrations of RO 31–8220 inhibit immunoprecipitated PKC-{zeta} (our unpublished data), whereas considerably lesser concentrations of RO 31–8220 have been reported to inhibit PKC-{alpha} and PKC-ß (see Ref.28).

Presently, we were able to obtain evidence that insulin activates PKC-{zeta} by a PI 3-kinase dependent mechanism in L6 myotubes. This apparent dependency on PI 3-kinase may at least in part reflect the fact that PKC-{zeta} has been found to be directly activated in vitro by polyphosphoinositides (see Refs. 29 and 30) that are generated through PI 3-kinase action. In further support of this possibility, we have recently found in studies of rat adipocytes (our unpublished data) that PI-3,4-(P04)2 and PI-3,4,5-(P04)3 activate control, but not insulin-stimulated, PKC-{zeta} immunoprecipitates, thereby diminishing or abolishing the difference in activity between these immunoprecipitates. With respect to activating mechanisms, we have also recently found that insulin and the above-stated polyphosphoinositides stimulate the phosphorylation, as well as the enzymatic activation, of PKC-{zeta} in rat adipocytes; however, it is presently not clear if this phosphorylation contributes to the enzymatic activation of PKC-{zeta}. Further studies will be required to see if the activation of PI 3-kinase is sufficient to account for insulin-induced activation of PKC-{zeta}.

Similar to findings in rat adipocytes (7) and 3T3/L1 adipocytes (8), we found that TPA activated a membrane form of PI 3-kinase in L6 myotubes. However, it should be noted that, unlike insulin, TPA does not activate PI 3-kinase through IRS-1, at least in rat adipocytes (see Ref.7). Needless to say, it is presently unclear as to how TPA activates PI 3-kinase, and what specific PI 3-kinase isoform is activated by TPA. Nevertheless, our findings with both wortmannin and LY294002 strongly suggested that PI 3-kinase activation is required for glucose transport effects of TPA in L6 myotubes. This somewhat surprising finding provides clear evidence that the simple demonstration of TPA effects on glucose transport does not necessarily imply that DAG-sensitive PKCs participate directly in activating this and other processes. As alluded to above, a more likely scenario, at least in L6 myotubes, is that TPA activates PI 3-kinase through DAG-dependent PKCs, and PI 3-kinase, in turn, apparently through factors that are clearly distinct from DAG-sensitive PKCs, is largely responsible for activating glucose transport. Whether PKC-{zeta} operates downstream of PI 3-kinase during TPA action is presently uncertain.

Finally, we recognize the fact that our findings with stable and transient expression of kinase-inactive PKC-{zeta} can only be taken as suggestive evidence that PKC-{zeta} acts as a downstream effector for PI 3-kinase in the regulation of glucose transport. Nevertheless, in view of the inhibitory effects of the PKC-{zeta} pseudosubstrate, at least one PKC, or a closely related protein kinase, appears to be required for insulin-stimulated glucose transport. Moreover, because DAG-sensitive PKCs are unlikely candidates, it is reasonable to suggest that one or more DAG-insensitive PKCs, e.g. PKC-{zeta} and/or other atypical PKCs or related protein kinases, may be involved in the activation of glucose transport in L6 myotubes by insulin. Clearly, more work will be needed to test the involvement of PKC-{zeta} and others DAG-insensitive protein kinases in insulin-stimulated glucose transport.


   Footnotes
 
1 This work was supported by funds from the Department of Veterans Affairs Merit Review Program and National Institutes of Health Grant DK-38079. Back

Received February 20, 1997.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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