Cellular Mechanism of Nutritionally Induced Insulin Resistance in Psammomys Obesus: Overexpression of Protein Kinase Cε in Skeletal Muscle Precedes the Onset of Hyperinsulinemia and Hyperglycemia

Rabbit antipeptide antibodies against PKC isoforms α, β_Ι, β_ΙΙ, δ, ε, θ, η, ι, and ζ and glutathione S-transferase (GST) were from Santa Cruz Biotechnology (Santa Cruz, CA). PKC l antibodies were from Transduction Laboratories (Lexington, KY). Phospho-Akt (Ser473) antibody and Akt antibody were from New England Biolabs (Beverly, MA). Polyclonal rabbit antisera against the IR were produced by immunization with synthetic peptides covering the last 15 COOH-terminal amino acids. Horseradish peroxidase–conjugated anti-mouse and anti-rabbit IgG were purchased from Amersham (Braunschweig, Germany).

Recombinant human insulin and [¹²⁵I]Tyr^A14-labeled human insulin were obtained from Novo Nordisk (Bagsvaerd, Denmark).

The cDNAs for the type A IR (19), PKCε (Wang et al., unpublished data, Genbank accession number AF028009), and GST (kindly provided by R.A. Bradshaw) (20) were cloned into a cytomegalovirus promotor-enhancer-driven expression vector.

Male and female Psammomys obesus from the DP and DR lines were bred at the Hebrew University-Hadassah Medical School animal farm. The animals were weaned at 3 weeks of age to an LE or HE diet (2.38 and 2.93 cal/g, respectively) manufactured by Koffolk, Petach (Tikva, Israel). Psammomys were housed in individual polypropylene cages; water and food were supplied ad libitum. All experimental procedures performed in this study were authorized by the Institutional Animal Care Committee.

The animals were killed by decapitation in the fed state in the morning. Blood was collected for determination of serum glucose and triglyceride levels by enzymatic methods (Roche, Basel, Switzerland) and serum insulin levels by radioimmunoassay, using anti-human antibodies (Medgenix, Brussels, Belgium). The liver and gastrocnemius muscles from both legs were excised, immediately frozen in liquid nitrogen, and then stored at −80°C.

Muscles and livers were extracted in ice-cold solubilization buffer using a motor-driven Potter homogenizer (final concentrations: 20 mmol/l HEPES, 8 mmol/l EDTA, 0.2 mmol/l Na₃VO₄, 10 mmol/l Na₄P₂0₇, 2.5 mmol/l phenylmethylsulfonyl fluoride, 1 mg/ml aprotinin, 2.5 mg/ml benzamidine, 2.5 μg/ml pepstatin, 2.5 μg/ml leupeptin, 160 mmol/l NaF, 2 mmol/l dichloroacetic acid, 1% Triton X-100, pH 7.4. After 20 min at 4°C, the samples were centrifuged at 20,000g for 60 min to remove insoluble material and the clarified extracts were stored at −80°C.

In 2 ml of KH buffer (0.15 mol/l KCl, 20 mmol/l HEPES, pH 7.5, 1 mmol/l ATP), 0.2 g of powdered muscle was homogenized using a Polytron. Samples were maintained on ice during subsequent steps, and all centrifugations were performed at 4°C. KCl was added to a final concentration of 0.65 mol/l, followed by centrifugation at 700g for 10 min. The supernatant S1 was saved and the pellet was reextracted in 2 ml of KH buffer and centrifuged, yielding supernatant S2. S1 and S2 were combined in a total of 5 ml of KH buffer and centrifuged at 183,000g for 60 min. The supernatant was designated as the cytosolic fraction. The pellet was resuspended in 0.2 mol/l Tris-HCl, pH 7.4, 1 mmol/l EDTA, 1 mmol/l EGTA, 255 mmol/l sucrose, layered on a 38.3% sucrose gradient and centrifuged in a swing-out rotor at 60,000g for 40 min. The interphase was collected and centrifuged at 183,000g for 90 min. The pellet was resuspended in 5 ml of lysis buffer containing 20 mmol/l Tris-acetate, pH 7.0, 0.27 mol/l sucrose, 1 mmol/l EDTA, 1 mmol/l EGTA, 1 mmol/l 4-(2-aminoethyl)benzenesulfonyl fluoride, 1 mmol/l Na₃VO₄, 100 mmol/l NaF, 1% Triton X-100, 10 mmol/l Na₄P₂O₇, 10 mmol/l C₃H₇O₆PNa₂, 1 mmol/l benzamidine, 1 mmol/l dithiothreitol, and 1 μmol/l leupeptin. The samples were centrifuged at 15,000g for 10 min to remove insoluble material and then stored at −80°C. Equal amounts (vol/vol) of cytosolic and membrane fractions were subjected to SDS-PAGE to determine the subcellular distribution of PKC isotypes.

Human embryonic kidney fibroblasts (HEK293; ATTC CRL 1573) were cultured in Dulbecco’s modified Eagle’s medium supplemented with 10% (vol/vol) fetal bovine serum (Life Technologies, Eggenstein, Germany), 10 mg/ml streptomycin, and 100 U/ml penicillin at 37°C in a 5% CO₂-enriched, humidified atmosphere.

HEK293 cells were transiently transfected using CaCl₂, as previously described by Chen and Okayama (21) and Gorman et al. (22). Twenty-four hours after transfection, the cells were starved overnight in medium containing 5 mmol/l glucose and 0.5% fetal bovine serum.

After stimulation, the cells were washed once with phosphate-buffered saline and homogenized in lysis buffer containing 20 mmol/l HEPES, pH 7.5, 150 mmol/l NaCl, 10% glycerol, 1.5 mmol/l MgCl₂, 4 mmol/l EGTA, 1 mmol/l EDTA, 1% Triton X-100, 1 mmol/l AEBSF, 30 mmol/l Na₄P₂O₄, 100 mmol/l Na₃VO₄, 10 mmol/l NaF, 5 μg/ml leupeptin, and 10 mmol/l benzamidine. After removal of cellular debris by centrifugation at 15,000g for 10 min at 4°C, the protein content in each sample was measured using Bio-Rad Protein Assay according to the manufacturer’s instructions (Bio-Rad, Richmond, CA).

Equal amounts of cell lysate were dissolved in 2× Laemmli buffer and subjected to SDS-PAGE. The proteins were transferred to nitrocellulose membranes (Schleicher & Schüll, BA85). Immunoreactive proteins were made visible using horseradish-peroxidase–coupled secondary antibodies and enhanced chemiluminescence reagents according to the manufacturer’s instructions (Amersham). The specific bands were quantitated using ImageQuant software (Molecular Dynamics, Sunnyvale, CA). For the Akt experiments, we used Fujifilm LAS1000 Luminescent Image Analyzer for detection of bands and Fujifilm Image Gauge V2.53 for quantification.

Powdered muscle tissue (100 mg) was extracted with 3 ml of chloroform/methanol (1:2, vol/vol) before addition of 0.8 ml of 1 mol/l NaCl. After monophase mixing, 1 ml of chloroform and 1 mol/l NaCl were added to break the phases. After centrifugation at 5,000g for 2 min, the chloroform phase was analyzed for DAG using the DAG kinase method (sn-1,2-DAG assay reagents system, Amersham). [³²P] phosphatidic acid was separated by thin-layer chromatography, and the respective bands were scanned on a PhosphorImager and quantitated using ImageQuant software (Molecular Dynamics).

Insulin-binding activity was measured by the method described by Klein et al. (23), with a slight modification. Microwells coated with an anti-IR antibody were provided by Dr. H.H. Klein (Medical University of Lübeck, Lübeck, Germany). Lysate (60 μl) was added to each well and incubated for 16 h at 4°C. The lysates were then removed, the wells were washed five times with wash buffer (0.05% Triton X-100, 100 mmol/l NaCl, 2.5 mmol/l KCl, 1 mmol/l CaCl₂, 0.1 mmol/l Na₃VO₄, 20 mmol/l HEPES, 10% glycerol, 0.05% NaN₃, 0.5% BSA, pH 7.4), and ¹²⁵I-insulin and unlabeled insulin (6.4 ng/ml and 121.6 ng/ml, respectively) were added. After 16 h at 4°C, the wells were washed again and bound ¹²⁵I-insulin was subsequently collected by twice adding a solution of 2% SDS at room temperature for 30 min. The radioactivity in the collected SDS solution was measured by γ-counting.

All data are presented as the mean ± SE. Group comparisons were made by unpaired Student’s t test. Correlations were assessed by Pearson’s least-squares method. P values less than 0.05 were considered significant.

The general data on animals used in this study are summarized in Table 1. There was no significant difference in glucose and insulin values of animals in stage A, whether from the DR line maintained on an HE diet or DP animals maintained on an LE diet. As expected, the transfer of DP animals to the HE diet induced hyperinsulinemia and hyperglycemia. Well defined animal groups (n = 11–14 per group) were taken for PKC assays.

We first aimed to determine the expression level of several PKC isoforms in skeletal muscle (gastrocnemius) tissues. Western blots of muscle lysates were probed with isozyme-specific antibodies for the classic PKC isoform α, β_Ι, and β_ΙΙ, the novel PKC isoforms δ, ε, θ, and η, and the atypical PKC isoform ζ, ι, and λ. An example of such a Western blot is shown for a subset of the samples in Fig. 1. The quantitation and statistical analysis of the specific bands from all samples analyzed is summarized in Table 2. Expression levels of PKC isoforms α, ε, and ζ were significantly increased in the diabetic animals (DP/C) as compared with the DR line. Interestingly, there was also a significant increase in PKCε expression level in the normoinsulinemic, normoglycemic animals (which may therefore be termed the “prediabetic stage”) of the DP line as compared with the DR line. The values were calculated from 10–11 animals per group and are expressed as means ± SE. PKC isoforms β_Ι, β_ΙΙ, θ, and η were not changed in the DP line, neither in the prediabetic nor in the diabetic state. The expression level of PKC isoforms δ, ι, and λ were undetectable by Western blotting or too low to allow quantitiation of the data.

The Western blots had shown a significant increase in the expression level of several PKC isoforms, both in the prediabetic (PKCε) and the diabetic animals (PKCα, ε, ζ). To determine whether this was associated with increased activity of these isoforms, we performed subcellular fractionation of the muscle tissue. Translocation of PKCs from the cytosolic to the particulate fraction is a prerequisite of activation. Because of the lack of sensitive isoform–specific PKC activity assays, plasma membrane (PM) association of PKC is often used as a measurement of its activity. PM and cytosolic fractions of the muscle samples were run on Western blots and probed with PKC isoform–specific antibodies. The autoradiograms were scanned and quantitated using ImageQuant (Molecular Dynamics). The percentages of PM-associated PKCα, β_Ι, β_ΙΙ, ε, η, and ζ in the three groups are shown in Table 3. Interestingly, the samples from diabetic animals revealed significant increases in the PKCα, β_Ι, and ε content of the PM fraction, indicating a chronic activation of these PKC isotypes. Smaller increases in PKCα and ε PM association in the “prediabetic” animals were not significant within the study. PKCθ revealed the highest degree of association with the PM compared with other PKC isoforms; surprisingly, however, the percentage of PM-associated PKCθ decreased in diabetic animals. Despite significant increases in the expression level of PKCζ, we did not observe increased PM association of this isotype in the diabetic animals.

Both classic and novel PKC isoforms require DAG for activation (12). To investigate whether the increased PM association of PKCα, θ, and ε could be caused by changes in DAG levels, we determined the DAG content in the muscle samples. Figure 2 shows linear regression analysis of PM association of PKC isoforms versus the DAG content in the individual samples. A significant correlation between these parameters was observed for α and ε isoforms of PKC. In contrast, we could not detect such a correlation for PKCζ. This was in agreement with the finding of unaltered PM content of PKCζ in the three different groups of animals (see Table 3). It was also expected because the activation of atypical PKC isoforms is independent of DAG (13).

Interestingly, PKCε was already overexpressed in the prediabetic stage. To evaluate whether the increased PKCε expression level correlated with any of the parameters characteristic for insulin resistance/diabetes, we performed linear regression analysis of PKCε versus the serum levels of glucose, insulin, and triglycerides, respectively, in the individual samples. As demonstrated in Fig. 3, there was no significant correlation with any of these parameters.

To determine whether the changes in PKCε expression levels were specific for skeletal muscle, we prepared liver extracts from the same animals to look at another insulin target tissue. No increase in PKCε expression was detectable in the group of prediabetic animals. The group of diabetic animals (DP/C) showed increased PKCε expression; however, this did not reach significance (DR, 7.5 ± 0.9; DP/A, 7.5 ± 0.9; DP/C, 10.1 ± 1.1, P = 0.08 vs. DR).

A previous report suggested that PKCα was capable of degrading the IR (18). We were therefore interested to see whether a similar effect could be observed with PKCε. HEK293 fibroblasts were transfected either with the IR alone or with the IR and PKCε. As a control, GST was coexpressed in all samples. Cells were stimulated for 1 or 6 h with 1 μmol/l tetradecanoylphorbol acetate (TPA) to activate PKC and were subsequently lysed. Figure 4A shows a representative Western blot of lysates probed with an antibody specifically recognizing the IR β-subunit (upper panel), PKCε (middle panel), or GST (lower panel). Clearly, the endogenous background of PKCs has no effect on the expression level of the IR. However, when PKCε is coexpressed with the IR, there is a marked downregulation of IR expression, which is further enhanced after 6 h of TPA stimulation. PKCε itself is also partly downregulated after 6 h of TPA stimulation, in agreement with previous reports. No effect on the expression level of the GST could be detected, indicating that the changes in the expression level of the IR and PKCε are specific. A quantitation and statistical analysis of six experiments shown in Fig. 4B underlines the significance of this finding (P < 0.001). Similar effects could be detected for other classic and novel PKC isoforms, but not for atypical PKC isoforms (24).

Because PKCε overexpression and activation led to decreased IR expression, we would obviously expect an effect on IR downstream signaling. To test this hypothesis, we determined the activity of PKB/Akt, which upon insulin stimulation is activated through the IR via the phosphatidylinositol 3-kinase pathway. HEK293 cells were transfected with either the IR alone or a combination of IR and PKCε and stimulated with insulin or TPA plus insulin, respectively. Cells were then lysed; proteins were separated on SDS-PAGE and subjected to Western blotting. The respective bands of the IR and Akt were detected with specific antibodies and quantified. For Akt, we determined both the active (Akt-Ser) and total amount of protein. Figure 4C shows the result of a quantification of three independent experiments performed in duplicate (n = 6). Insulin-stimulated samples from cells expressing only the IR were set to 1, and all other samples were calculated accordingly. Coexpression and activation of PKCε led to a significant reduction of IR expression level (P < 0.001) and a concurrent inhibition of Akt activation (P < 0.01).

We therefore addressed the question of whether overexpression of PKCε could have a similar effect on the IR expression level in vivo. Because Psammomys have a very low number of IRs, we used a sensitive ELISA to indirectly determine the number of IRs, by measuring binding of ¹²⁵I-insulin to the IR (23). There was a small but statistically significant reduction in ¹²⁵I-insulin binding in lysates from the DP/C animals when compared with the DR animals (DR, 199.2 ± 9.2; DP/A, 188.8 ± 21.2; DP/C, 170.8 ± 8.5, P < 0.05 vs. DR). We therefore performed linear regression analysis of the IR expression level (¹²⁵I-insulin binding) versus the PKCε expression levels. Figure 5A (2) shows that there is a significant inverse correlation between these two parameters (P < 0.05). This seemed to be specific for PKCε expression, because comparison of ¹²⁵I-insulin binding to the expression levels of the other PKC isoforms (Fig. 5A) as well as to blood glucose and plasma insulin levels (data not shown) did not reveal any significant correlation. An even stronger correlation was detected between ¹²⁵I-insulin binding and the PM-associated fraction of PKCε (Fig. 5B).

In this study, we provide evidence that several PKC isoforms (α, ε, ζ) are overexpressed in the skeletal muscle of diabetic P. obesus, taking the individuals from the DR line as a basal reference. This may indicate that this overexpression could be the primary reason for the insulin resistance in these animals. It is not a general phenomenon of all PKC isoenzymes, because, for example, PKC β_Ι, β_ΙΙ, θ, and η expression levels are unchanged. Most notable is the expression of PKCε in even the prediabetic DP/A animals, which is consistent with the possibility that it could be causally related to the development of diabetes in these animals and may be an early event or mark an inherent susceptibility to the development of the disease. Overexpression of PKCε alone seems to have no consequences for the animals as long as they are restricted in their diet. Only after the introduction of a high-calorie diet is the increased expression of PKCε also reflected in higher PM association and thus presumably increased activity of the enzyme. The finding of increased plasma membrane association and therefore probable chronic activation of PKCε in diabetic DP/C, but not prediabetic DP/A animals, suggests that factors contributed by the diabetic milieu, i.e., hyperlipidemia, hyperglycemia, and/or hyperinsulinemia, are required to activate PKCε. Therefore, in light of a multifactorial origin for diabetes, the overexpression of PKCε might explain the genetic susceptibility of the DP line of P. obesus to the de-velopment of insulin resistance, which, together with environmental influences, may lead to development of the disease.

This PKC overexpression is particularly distinctive in the skeletal muscle of these animals but may also play a role in other insulin target tissues such as liver. In fact, when we determined the expression level of PKCε in the liver samples of the animals, we detected a trend of increased PKCε expression in the diabetic group; however, a larger number of samples would be needed to determine whether this was significant. Overexpression of PKC has previously been demonstrated in the liver of humans and rats with type 2 diabetes (8). In skeletal muscle of high-fat–fed rats (9), chronic activation of PKC isoforms ε and θ was observed. This was connected with decreased expression levels of PKCθ and no changes in PKCε expression level. In this study, the rats were fed a diet containing >30% fat, representing ∼60% of their caloric intake. They were insulin-resistant, as determined during an euglycemic-hyperinsulinemic clamp, but only mildly hyperinsulinemic or hyperglycemic. It was speculated that elevated DAG levels originating from lipolysis of two- to threefold increased triglyceride content in the muscles was responsible for increased PKC activation. This is different from the Psammomys that were fed a regular diet containing only ∼4% of fat, whose insulin resistance stemmed from a caloric load inappropriate to their metabolic capacity. Interestingly, however, we detected increased DAG levels in the DP line of P. obesus. The elevated DAG levels, possibly arising from increased synthesis and breakdown of glycerolipids, correlated well with membrane-associated fractions of PKCα and ε, suggesting that they may be causally related to the activation of these PKC isoforms. Increased DAG levels in aorta, heart, and retina of hypergalactosemic and diabetic dogs and rats have been suggested to be responsible for the activation of PKCβ (25,26). PKC enzyme activity and levels of immunoreactive PKCα, β, ε, and δ were shown to be increased in the membrane fractions of Goto-Kakizaki rats (10). Furthermore, an increased activity of PKC was linked to reduced IR phosphorylation in the liver of starved rats (27). These data, however, are in contrast to another study that found a decreased expression of PKCα, β, and ε in the soleus muscle of Zucker obese (fa/fa) rats (28). In most of these studies, the elevated expression/activation of PKC seemed to be a consequence of the diabetic milieu and therefore a late event in the development of the disease, correlating with muscle triglyceride levels (9) or hyperinsulinemia (10). The Psammomys have the advantage of being extremely well characterized with respect to the development of diabetes, and relatively homogeneous groups of animals representing the different stages (A, B, C, D) of the disease (4) can be studied. We can show that there is no correlation between PKC overexpression and hyperinsulinemia or serum triglyceride levels. In this animal model, overexpression of PKCε is clearly primary to the development of a diabetic milieu. Of course, it is possible that changes in the diabetic milieu add to the overexpression/activation of PKC in DP/C. Although it has been suggested that PKC activation arises from increased glucose incorporation into DAG (25,26), neither our own nor other studies (9,10) in diabetic animals show conclusive evidence that hyperglycemia could cause the observed changes. Hyperglycemia-induced changes of PKC activity (14) may, however, be an important factor in later stages of diabetes, in which PKCβ has been shown to be responsible for late complications of the disease (29).

Several studies in different cell lines as well as in vitro studies have suggested that PKC directly interferes with insulin signaling through serine/threonine phosphorylation of either the IR itself or one of its major substrates, the insulin receptor substrate 1 (IRS-1) (14–17,30). There seems to be agreement that the ε isoform of PKC, in particular, is subject to regulation in diabetic individuals (7–10). Interestingly, an in vitro study provided evidence that PKCε may play an important role in mediating the inhibitory effect of tumor necrosis factor-α on insulin signaling (31,32), the major cause of obesity-linked insulin resistance. It is believed that serine/threonine phosphorylation of IRS-1, via a feedback loop, is able to inhibit the tyrosine kinase activity of the IR (33). In P. obesus, insulin resistance becomes considerably accentuated at stages B and C on the HE diet. This was shown to be associated with impaired IR function, which seems to be related to a decrease in the amount of phosphorylated tyrosine residues on the IR β-subunit (34). The extent of insulin-induced activation of the IRK was markedly reduced in stages B and C, using stage A as a reference. Hyperinsulinemia by itself was shown to be a detrimental factor for IR activation (34), because development of hyperinsulinemia, even without hyperglycemia, was associated with a decrease in insulin-induced IRK activation in liver and muscle. In the skeletal muscle, hyperinsulinemia was accompanied by a reduction in basal IRK activity. Diet restriction leading to the return of insulin level to normal restored the IRK activity (4,34). Therefore, the overexpression of PKC, particularly PKCε, may be an additional factor inducing the IR malfunction. In the present study, we provide evidence that, in addition to serine/threonine phosphorylation of the IR or IRS-1, a downregulation of IR numbers could be a possible mechanism for induction of insulin resistance. Skeletal muscle of diabetic Psammomys DP/C showed a reduction in ¹²⁵I-insulin binding, most probably representing a decreased number of IRs, although we cannot completely rule out changes in the affinity for insulin. In fact, there was a clear correlation between PKCε expression levels or activity and ¹²⁵I-insulin binding. This is in good agreement with previous reports showing that PKCα is capable of downregulating tyrosine kinase receptors (18) and that several PKC isoforms mediate degradation of the IR (24). The significant inverse correlation between insulin binding and PKCε expression further suggests that the PKC-induced downregulation of IR number is a causal factor in insulin resistance characterizing P. obesus. Insulin resistance is generally believed to have a major impact on skeletal muscle, liver, and adipose tissue. Surprisingly, mice with a targeted disruption of the IR gene in skeletal muscle displayed elevated levels of fat mass, serum triglycerides, and free fatty acids but never developed diabetes, suggesting that insulin sensitivity in muscle may not be the primary cause for the development of diabetes (35). However, the knockout mice reflect an isolated effect of IR expression in skeletal muscle, whereas in Psammomys, we study the progression of the disease in a much more complex situation of nutritionally induced diabetes. We describe only one aspect of several factors that may lead to insulin resistance in these animals. Other factors, such as the inhibition of IR tyrosine phosphorylation (34) and effects from other tissues, are contributing to the pathogenesis of diabetes. This is also a likely scenario in the context of a growing number of reports that suggest that type 2 diabetes is a multifactorial disease (36). In particular, this has been very elegantly demonstrated in heterozygotes combining several knockouts of genes of the insulin signaling pathway. It is also worthwhile to note that Psammomys has a very small number of IRs compared with the Albino rat (34). Therefore, even a minor change in IR number could have an influence on insulin signaling. In addition, the impact of decreased IR number could become more overt if, at the same time, IR tyrosine phosphorylation is inhibited (34).

Smaller changes in PKCε expression level and insulin binding (data not shown) were found in the liver of Psammomys. The hepatic insulin resistance is manifest mainly by nonsuppression of gluconeogenesis by insulin. This aspect of insulin resistance may not be exclusively related to the overexpression of PKCε.

In summary, we have demonstrated changes in the expression level and activity of PKC in gastrocnemius muscle of P. obesus. Our data suggest that, in particular, PKCε could be causally related to the development of insulin resistance in these animals. However, our correlative studies do not provide evidence of causality between PKCε activation and in vivo insulin resistance, although the proposed mechanism of its inhibitory action on insulin signaling through downregulation of IR numbers is consistent with such a model. Further work is required to establish a causative link between the observed changes in PKC expression/activity and the development of insulin resistance.

We thank Patricia Holst for critical reading of the manuscript; K. Seedorf for valuable discussions throughout the study; A. Ullrich for IR cDNAs; H. Mischak, A. Ullrich, and T. Biden for PKC cDNAs; and H. Klein for ELISA plates coated with IR antibodies. The Hagedorn Research Institute is an independent basic research component of Novo Nordisk A/S.