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Research Communications |
a Departments of Biochemistry and Molecular Biology, College of Medicine, University of South FloridaUSA
b The James A. Haley Veterans Hospital, Tampa, Florida 33612, USA
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: A10 cells • human AoSMC • acute hyperglycemia • posttranscriptional regulation • glucose
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, ßI, ßII, and 
. Novel protein kinase C (Ca2+ independent) and atypical protein kinase C include PKCs 
, 
, 
, 
, 
/
, 
, and µ (2, 9, 10). PKC isozymes show distinct tissue, cellular, and subcellular distribution although the same cell type may express more than one isozyme (8, 11, 12). PKCßI and PKCßII are derived from the alternative splicing of the 3' exon of a single PKCß gene (13). Both PKCßI and PKCßII are present in VSMC. In quiescent cells, PKCßII is the predominant isoform detected on Western blots. Smooth muscle cells in the arterial wall normally exist in a quiescent state until recruited to proliferate. Complications involved in cardiovascular tissue injury recovery are aggravated by episodes of acute hyperglycemia. The response to injury theory proposed by Ross (14) suggests that injury to the epithelium leads to the production of growth factors and cytokines, which stimulate smooth muscle cell movement into and proliferation within the arterial intima. Diabetic conditions, especially acute hyperglycemia, may provoke the excessive formation of atherosclerotic lesions, which may ultimately lead to the formation of the fibrous plaques (1). This study elucidates molecular events occurring after initiation of the cell cycle in quiescent vascular smooth muscle cells after exposure to acute hyperglycemic conditions. In previous studies, only PKC activity and protein levels were examined after chronic glucose exposure; no experiments were performed to determine the effects of acute exposure to high glucose on PKCß gene processing in proliferating cells from the macrovasculature. Prolonged exposure to high glucose leads to glucose-induced PKC activation in rat nerves, adipocytes, and soleus muscles, followed by the down-regulation of PKC activity (15–17). PKCß2 levels were elevated in vascular aortic cells from diabetic rats exposed to high glucose levels for 2 to 4 months and in VSMC cultured under chronic high glucose conditions for 5 to 10 days (5, 18, 19). These studies demonstrated an increase in membrane PKCßII activity as a result of increased translocation from the cytosol in cells from vascular tissue. However, the conditions of acute hyperglycemia used here to study PKCß gene expression in vascular smooth muscle cells more closely mimicked the `response to injury model' proposed by Ross (14).
Our previous studies indicated that PKCßI and PKCßII regulate the vascular smooth muscle cell cycle. In A10 cells (a clonal cell line of VSMC) overexpressing PKCßII, DNA synthesis was attenuated (20). This suggested that PKCßII may function as a cell cycle mediator during G1/S phase transition in VSMC. We found that PKCßII protein expression was decreased and the percentage of A10 cells entering the S phase was increased in VSMC in the presence of acute high extracellular glucose concentrations (32). On the basis of these findings, we hypothesized that the hyperglycemia-induced proliferation of VSMC may be related to glucose regulation of PKCß gene expression. In this study, we investigated molecular mechanisms through which high glucose could regulate PKCß gene expression in proliferating VSMC. Hyperglycemic conditions down-regulated PKCßII gene expression as reflected by protein and mRNA levels, whereas PKCßI levels remained unchanged between high and normal glucose conditions. Glucose induced posttranscriptional destabilization of PKCßII messenger RNA through cytosolic nuclease activity. The specificity of glucose-induced destabilization was confirmed via reverse transcriptase-polymerase chain reaction (RT-PCR), for PKCßII mRNA while not significantly altering PKCßI mRNA. Primary cultures from human aortic muscle cells demonstrated the same phenomenon.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Primary cultured human aortic smooth muscle cells (AoSMC) (Clonetics, San Diego, Calif.) were grown in SmGM (Clonetics) medium containing 5.5 mM glucose, 5% FBS, 10 ng/ml human recombinant epidermal growth factor, 390 ng/ml dexamethasone, 50 µg/ml gentamicin, and 50 ng/ml amphotericin-B (according to recommendation by Clonetics) at 37°C in a humidified 5% CO2, 95% air atmosphere. Cells were grown to 80% confluency and medium was changed every 5 days. Cell synchronization was achieved by serum deprivation, culturing the aortic smooth muscle cells in SmGM with 5.5 mM glucose (basal medium) containing 0.5% FBS for 72 h (22). Cells were reinitiated to proliferate by changing the medium to SmGM with 5.5 mM glucose containing 5% FBS.
Isolation of RNA and Northern blot analysis
Total cellular RNA was isolated from 100 mm plates using RNAzol B (Tel-Test, Inc., Tex.). RNA samples (10 µg) were prepared in formamide, formaldehyde, and 1x MOPS, and fractionated on 1.2% agarose-formaldehyde gels. Ethidium bromide (0.5 µg/ml) was added to the loading buffer. After fractionation, the integrity and loading of 18S and 28S RNA were assessed under UV light as described (17, 23, 24). The size-fractionated RNA was then capillary transferred to Hybond membranes (Amersham, Arlington Heights, Ill.), and the uniform transfer of RNA was assessed by visualizing 18S and 28S RNA on the membrane. RNA was then cross-linked to membranes by baking at 80°C in a vacuum oven for 2 h. Membranes were hybridized overnight at 42°C with 2 x 107 cpm of the full-length PKCß cDNA probe (labeled with [32P] dCTP by nick translation, as described) (25) per ml of hybridization buffer. Membranes were washed with high stringency conditions, and quantitated using the Molecular Dynamics Phosphoimaging System. Where mentioned, membranes were exposed to X-ray film and the autoradiograms were quantitated densitometrically. For RNA stability assessment, A10 cells were pretreated with 5 µg/ml actinomycin D (Sigma) for 30 min, then incubated with medium containing 5.5 mM or 25 mM glucose. Each Northern blot analysis was repeated at least three times to confirm reproducibility.
Western blot analysis
Synchronized A10 cells were incubated with 5.5 mM (normal or low) or 25 mM (high) glucose for 6, 10, 14, or 18 h. Cell lysates were analyzed as described previously (25). PKCßI and PKCßII antibodies were obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, Calif.). After incubation with primary antibody, blots were developed with the enhanced chemiluminescence (ECL) kit (Amersham Corp.).
Calcium phosphate transfections
For promoter studies, A10 cells were cultured in 6-well plates and synchronized in DMEM with 5.5 mM glucose containing 0.5% FBS for 48 h. To reinitiate cell proliferation, fresh medium (DMEM with 5.5 mM glucose containing 10% fetal bovine serum) was substituted on A10 cells. PKCß-chloramphenicol acetyltransferase (CAT) expression plasmids containing varying 5' upstream sequences of PKCß with a constant 3' end at position +179 base pairs (bp) inserted upstream of CAT (chloramphenicol acetyltransferase) structural gene in the promoterless CAT plasmid were obtained from S. Ohno (26). Each construct (3.5 µg) ligated to CAT as reporter gene (27) was transfected into A10 cells using calcium phosphate–DNA precipitates (28). The pSV-ß-galactosidase vector (2 µg) was cotransfected for normalization of transfection efficiency (29). The pSVACAT plasmid containing neither promoter nor enhancer activity was used as negative control. pCAT plasmid containing the SV40 promoter and enhancer sequences was used as the positive control. Twenty-four hours later, calcium phosphate–DNA precipitate was washed off with 1x Dulbecco's phosphate-buffered saline (DPBS) and replaced with fresh medium containing either 5.5 or 25 mM glucose, or 5.5 mM glucose + 100 nM 12-O-tetradecanoylphorbol-13-acetate (TPA). Cells were extracted after 6 h or periodically over 24 h (for simultaneous cell cycle studies), using reporter lysis buffer (Promega cat # E1000), and assayed for CAT activity and ß-galactosidase activity (30). A negative control extract, prepared from mock transfected A10 cells, was assayed for the presence of endogenous ß-galactosidase activity. CAT activity was normalized to ß-galactosidase activity.
RNA stability assay
To prepare postnuclear extracts (31), synchronized A10 cells reinitiated to proliferate with DMEM + 10% FBS were treated with either 5.5 or 25 mM glucose for 2 h to 24 h, then washed and collected in ice-cold 1x DPBS. The cellular pellet was resuspended in extract buffer containing 0.01 M Tris-HCl, 0.15 M NaCl, 0.5% Nonidet P-40, leupeptin (10 µg/ml), and aprotinin (10 µg/ml), incubated on ice for 10 min and carefully layered over extract buffer containing 24% sucrose. Samples were centrifuged over a sucrose gradient at 10,000 x g for 20 min at 4°C. Postnuclear extract was separated, stored on ice, and aliquots taken for protein concentration using Bio-Rad protein assay. Total RNA (10 µg) from A10 cells was incubated with 2.7% (vol/vol) of the postnuclear extracts for 30 min at 4°C in 50 µl (final volume) of extract buffer. When specifically noted, 50 mM EDTA was added to the reactions as a control to inhibit nuclease activity. The reactions were terminated by phenol-chloroform extraction, supplemented with 10 µg yeast tRNA as carrier, and precipitated with ethanol. RNA was fractionated on 1.2% formaldehyde-agarose gels and Northern blot analysis was carried out with full-length PKCß cDNA probe. The assay and analyses was repeated at least three times with separate postnuclear extracts to assess and confirm reproducibility.
Reverse transcriptase-polymerase chain reaction
Total RNA was isolated from control (5.5 mM glucose) or glucose-treated (25 mM glucose) A10 cells and 2 µg was used to synthesize first strand cDNA using an Oligo(dT) primer and Superscript II reverse transcriptase (Life Technologies preamplification kit). The upstream sense primer corresponded to the C4 kinase domain common to both PKCßI and PKCßII (5' CGTATATGCGGCCGCGTTGTGGGCCTGAAGGGG 3'), and the downstream antisense primer was specific for PKCßI (5' GCATTCTAGTCGACAAGAGTTTGTCAGTGGGAG 3') (25). These primers detect inclusion of the PKCßII exon in the mature mRNA as well as PKCßI mRNA. Sense and antisense primers for ß-actin (#5402–3) were obtained from Clontech (Palo Alto, Calif.). PCR was performed using ampliTaq Gold DNA polymerase from Perkin-Elmer (#N808–0240) on 10% of the reverse transcriptase reaction product. After 35 cycles of amplification in a Biometra Trioblock thermocycler (PKCßI and ßII: 95°C, 30 s; 64°C, 2 min for 35 cycles; and for ß-actin: 94°C, 1 min; 58°C, 1 min; and 72°C, 3 min for 35 cycles), 25% of the PCR reaction was resolved on a 1.2% agarose gel. Bands were observed under UV light and photographed.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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PKCßII protein levels decreased when A10 cells were exposed to 25 mM glucose
To better understand the biochemical mechanism underlying the acute effects of high glucose exposure on PKCß expression in aortic vascular smooth muscle cells, protein levels of PKCßI and ßII were determined in synchronized A10 cells. Cell growth synchronization was achieved by serum deprivation for 48 h, as described in Materials and Methods; after synchronization, medium on A10 cells was replaced with DMEM containing 10% FBS to reinitiate cell proliferation and incubated with high (25 mM) or normal (5.5 mM) glucose. Over a 24 h period, protein was extracted at various times from cells reinitiated by addition of serum to proliferate. As cells progressed through the cell cycle, the expression of PKCßII appeared to be cell cycle associated, with highest levels of protein expression observed at 6 and 18 h and lowest levels occurring at 14 h. PKCßII protein levels in cells treated with high glucose dramatically decreased (by 55%) within the first 6 h and remained low after 14 h of continuous high glucose treatment, beginning to return at 18 h (
Fig. 1b).
No significant changes in PKCßI protein levels were detected between high and normal glucose treatments ( Fig. 1a). To check the specificity of high glucose to down-regulate PKCßII, Western blot analysis was repeated using primary antibodies to PKC and PKC. Again, no significant changes in PKC or protein levels were detected between high and normal glucose treatments (data not shown). Thus, the down-regulation by acute glucose appeared specific for the PKCßII isoform.
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Hyperglycemic conditions markedly reduce PKCß(I+II) mRNA in A10 cells
To determine the acute effects of hyperglycemia on PKCß gene expression, steady-state levels of PKCß mRNA were examined in synchronized A10 cells (by serum deprivation for 48 h as described in Materials and Methods) that were reinitiated by changing the medium to DMEM containing 10% FBS and incubated overnight with either high (25 mM) or normal (5.5 mM) glucose. Cells incubated with 25 mM mannitol were used as osmotic controls. Total RNA was extracted and the mRNA transcript levels were detected using a full-length PKCß cDNA probe that would cross-hybridize with both PKCßI and PKCßII mRNAs. As shown in Fig. 2,
a 60–75% decrease in PKCß(I+II) mRNA levels was observed in cells treated overnight with high glucose (25 mM). The mRNA levels in the control cells (5.5 mM glucose) and osmotic control cells (mannitol-treated) remained unchanged. This suggested that high glucose down-regulated PKCß(I+II) gene expression. Since protein analysis after high glucose exposure indicated a decrease only in PKCßII levels, it was assumed that the Northern blot analysis reflected a down-regulation of PKCßII mRNA whereas PKCßI mRNA remained constant. Since PKCßII is the product of PKCß mRNA alternatively splicing, the possibility existed that transcriptional and posttranscriptional controls were altered by high glucose.
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Glucose quenches PKCß promoter activity
Our studies indicated that PKCßII levels `cycled' with synchronized A10 cells during a 24 h period (32). We chose to systematically study the effect of high glucose on PKCß promoter activity in synchronized A10 cells. Glucose may be exerting its effect on PKCß gene expression through a response element located in the common promoter upstream of the transcription start site, as reported with L-PK and S14 gene expression in hepatocytes (20, 33–36). Since PKCß is a low copy gene, nuclear run-on assays were not a practical approach to examine transcriptional regulation. Hence, the PKCß-CAT expression plasmids cloned by S. Ohno (Yokohama College of Medicine) (26) were used to study transcriptional regulation of PKCß gene expression by glucose. PKCß promoter constructs with successive deletions of the 5' region cloned in front of a promoterless CAT as the reporter gene were transiently transfected into A10 cells ( Fig. 3a).
pSV-ß-Galactosidase was cotransfected with the CAT constructs to normalize the efficiency of transfection. Simultaneous positive and negative controls were performed as described in Materials and Methods. Transcriptional repression of PKCß promoter by transcription factors in response to high glucose was anticipated. However, in A10 cells incubated with high glucose (25 mM) for 6 h, a moderate decrease or quenching in the PKCß promoter activity of constructs A (-4.1 kb to +179CAT), C (-674 to +179CAT), and D (-411 to +179CAT) was observed ( Fig. 3b). Construct E (-234 to +179CAT) showed no quenching with glucose indicating that a putative carbohydrate response element had been deleted. Construct E contains the basal response elements required for promoter activity and was not affected by glucose concentrations. Thus, construct D contained the elements involved in maximum quenching of promoter activity, presumably within a 177 bp region which was deleted in construct E. In parallel experiments, a control with 100 nM TPA increased the promoter activity by 10-fold in all constructs. This confirmed the response of PKCß basal promoter element with phorbol esters (37).
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In the case of transcriptional repression, DNA binding is inhibited by repressors blocking the binding site for a factor or forming a non-DNA binding protein–protein complex resulting in inhibition of transcription. The reduction in promoter activity observed here did not account for the dramatic down-regulating effect of glucose on total PKCß mRNA. If glucose effects were mediated exclusively at the transcriptional level, total repression would have occurred. The reduction in promoter activity observed here is most likely explained as a quenching effect that involves interfering with transcriptional activation by a DNA-bound factor (38). An example of transcriptional quenching is illustrated by the yeast factor GAL80, which inhibits gene activation by the positively acting GAL4 protein in the regulation of galactose in yeast (44). Since the extent of quenching of the PKCß promoter by glucose varied depending on the time points examined, the results suggested that as cells progress through the cell cycle, high glucose regulation of PKCß gene expression may be cell cycle mediated. Simultaneous cell cycle studies indicated an increase in the percentage of cells going into S phase in high glucose, implying that quenching of PKCß transcription may be related to cell cycle progression (N. A. Patel, M. Yamamoto, and D. R. Cooper, unpublished observations).
PKCß message is destabilized by glucose
Since the quenching of PKCß promoter activity by acute high glucose did not account for the 60–75% reduction in PKCß mRNA levels as observed by Northern blot analysis (
Fig. 2), the major effect of glucose may be exerted posttranscriptionally. However, high glucose also diminished PKCßII protein levels, suggesting that high glucose could affect the stability of the PKCßII transcript at the posttranscriptional level. Northern blot analysis of PKCß(I+II) mRNA level (see Materials and Methods) was performed on total RNA from synchronized A10 cells. Upon reinitiation of proliferation by DMEM + 10% FBS, A10 cells were pretreated with actinomycin D for 30 min, followed by incubation with normal or high glucose. The mRNA levels were detected using the full-length PKCßII cDNA probe, which recognizes both PKCßI and PKCßII mRNA. A10 cells exposed to high glucose showed a decrease in PKCß(I+II) mRNA levels within 2 h of treatment ( Fig. 4);
levels decreased further to 60% of control by 6 h. Thus, the results suggested that posttranscriptional destabilization of PKCßII mRNA by glucose was also occurring.
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In vitro assay for RNA destabilization
To assess the involvement of a nuclease activity in the destabilization of PKCß mRNA by glucose, an in vitro RNA stability assay was performed (31). A10 cells incubated with 5.5 or 25 mM glucose for 2, 6, 18, and 24 h were used to prepare control postnuclear extracts (control PNE) or glucose-treated postnuclear extracts (glucose PNE). Total RNA isolated from untreated A10 cells was incubated with 2.7% (vol/vol) postnuclear extracts or glucose-treated postnuclear extracts from A10 cells in 50 µl total volume of extract buffer. RNA incubated without postnuclear extracts was used as a positive control. A reaction with no RNA served as the negative control. Since involvement of divalent cations is a feature of some ribonucleases that regulate mammalian RNA turnover, EDTA was added to postnuclear extracts to distinguish nuclease activity from cyclizing RNases and acid lysosomal RNase. Northern blot analysis was then performed to detect nondegraded PKCß(I+II) mRNA. Incubation of RNA with postnuclear extracts from synchronized A10 cells treated with 25 mM glucose for 2 and 6 h showed 45% and 65% decreases, respectively, in PKCß(I+II) RNA levels whereas incubation for 18 and 24 h showed 8% and 12% decreases, respectively, in PKCß(I+II) RNA levels ( Fig. 5a).
No degradation of RNA was observed in the presence of EDTA either in the control or high glucose-treated postnuclear extracts ( Fig. 5b). These results indicate the activation of a divalent cation-dependent (31, 39) nuclease activity by high glucose. The destabilization of PKCß(I+II) mRNA by glucose accounted for a significant down-regulation of PKCßII expression, although these observations suggested that destabilization of PKCßII mRNA occurred as an early event in the regulation of mRNA processing by high glucose.
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Specificity of mRNA destabilization by glucose
Since a full-length PKCß cDNA probe was used for analysis of PKCß(I+II) RNA levels by the Northern blot analysis, RT-PCR was performed on total RNA from A10 cells to determine the specificity of high glucose to down-regulate PKCßII mRNA rather than PKCßI mRNA. It had been demonstrated earlier that mature PKCßII mRNA results from differential processing leading to in PKCßII exon inclusion into PKCßI mRNA transcript (25, 40). Thus, the PKCßII transcript contains the PKCßI exon with the poly A tail and a common 3' untranslated region (UTR) ( Fig. 6a).
Inclusion of the alternatively spliced exon encodes a stop codon such that the PKCßI exon facilitates poly-adenylation but the exon is not translated, thereby generating PKCßII protein. Primers were designed for the upstream common kinase region (C4) (-sense primer) and the PKCßI exon (-antisense primer) for simultaneous amplification of both PKCßI and -ßII messages by RT-PCR. In control A10 cells (5.5 mM glucose), both PKCßI and PKCßII PCR products were detected. Although this assay is semiquantitative, the ratio of PKCßII to PKCßI mRNA was about 2 to 1. In glucose-treated (25 mM) A10 cells, PKCßII PCR product decreased >90%, whereas PKCßI mRNA level did not show any significant change. ß-Actin levels remained constant in both control and glucose-treated A10 cells ( Fig. 6b). Thus, PKCßII mRNA was specifically destabilized in response to 25 mM glucose.
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To extend the physiological significance of the down-regulation of PKCßII by high glucose, we repeated the RT-PCR described above using primary cultured aortic smooth muscle cells from humans (AoSMC CC2571, Clonetics Normal Human Cell System). In high glucose medium (25 mM glucose), AoSMC showed a >95% decrease in PKCßII mRNA whereas PKCßI mRNA was not significantly affected. ß-Actin mRNA levels remained the same in both control (5.5 mM glucose) and glucose-treated (25 mM glucose) AoSMC ( Fig. 6c). Since cells from human primary culture responded in the same manner, the A10 cells proved to be a reliable model for studying the effects of high glucose on PKCß gene expression in vascular smooth muscle cells.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Glucose regulation of PKCß gene expression in VSMC may occur at transcriptional, posttranscriptional, translational, or posttranslational levels. The potential sites of regulation included repression of transcription, improper nuclear export of PKCßII mRNA, posttranscriptional destabilization, decreased translation, or multiple effects culminating in decreased PKCßII protein levels. PKCßII protein levels declined sharply after high glucose exposure whereas PKCßI protein levels did not change significantly between high and normal glucose conditions. Since protein levels often reflect the amount of mRNA transcribed, steady-state levels of PKCß mRNA were determined. It followed that the decrease in PKCß(I+II) mRNA levels observed in A10 cells exposed to high glucose was likely due to down-regulation of PKCß gene expression.
Since PKCßI and PKCßII share a common DNA promoter, transcriptional processing of PKCß in VSMC was examined to determine whether high glucose down-regulated PKCß gene expression by repressing promoter activity. Hyperglycemic conditions, however, only quenched activities of PKCß promoter constructs rather than repress promoter activity, and accounted for only a fraction of the effect of high glucose on PKCßII protein levels.
It is possible that glucose may be exerting its quenching effect through a response element located upstream of the transcription start site. A carbohydrate response element (ChoRE) that mediates its action via glucose had been reported (35, 41). The consensus sequence, 5' CACGTG 3', was described in the promoter region of a number of glucose responsive genes including L-PK (34) and S-14 in hepatocytes (20, 33, 36). Multiple copies of CACGTG motif are found in the region upstream of the transcription initiation site of the PKCß promoter region. It was likely that glucose may play a role as a quencher of positively acting regulatory factors by binding to or otherwise activating a ChoRE. Construct D (-411 to +179CAT) contains a ChoRE motif in proximity to the CAAT box and AP2 binding site (TATA box and AP1 binding site have been deleted). The ability of a bound trans-acting factor to stimulate transcription via its activation domain may be inhibited by quenching of the activation domain by glucose, which could either bind to the trans-acting factor without directly binding to DNA or bind to a site adjacent to the trans-acting factor (38).
We further investigated whether high glucose affected the posttranscriptional processing of PKCß transcripts since quenching of PKCß promoter activity did not fully explain the reduction in PKCß mRNA levels. A10 cells exposed to high glucose showed a decrease in PKCß(I+II) mRNA stability within 2 h in the presence of actinomycin D, indicating an increase in message degradation was occurring. The induction or activation by glucose of a nuclease activity involved in the destabilization of PKCßII mRNA was demonstrated by an RNA stability assay. This divalent cation-dependent nuclease activity was distinct from cyclizing RNases and acid lysosomal Rnase, since its activity was inhibited by EDTA.
To confirm that glucose specifically destabilized the PKCßII mRNA isoform, RT-PCR was performed using primers that amplified PKCßI and PKCßII mRNA simultaneously. High glucose destabilized PKCßII mRNA whereas PKCßI mRNA was not affected. Since the mature PKCßII mRNA is generated by exon inclusion (25, 40) and both PKCßI and PKCßII transcripts contain the PKCßI exon and share a common 3' UTR ( Fig. 6a), the destabilizing elements appear to be present within the PKCßII exon or are introduced around the splice sites between the exons. This is a novel observation since the destabilization elements regulated by glucose occur within the coding area of PKCßII exon rather than in the 3' UTR.
Our studies were extended to primary human AoSMC cultures where we demonstrated that glucose also induced PKCßII destabilization. The combined effects of high glucose on transcriptional quenching of the PKCß promoter and specific destabilization of the PKCßII transcript account for the reduction of PKCß mRNA levels observed under steady-state conditions in rat and human proliferating aortic smooth muscle cells. The human primary cultures further underscore the physiological relevance of acute high glucose effects on vascular smooth muscle cell proliferation. In other studies, we used CGP53353 a PKCßII-specific inhibitor to demonstrate that CGP53353 inhibits glucose-stimulated rat VSMC proliferation (M. P. Yamamoto, N. A. Patel, and D. R. Cooper, unpublished observations).
Earlier studies showed that prolonged exposure to high glucose for 2–4 months might lead to an elevation of PKCß levels in vascular tissues (4, 5, 18, 19, 42). Using an inhibitor for PKCß on diabetic rats, Inoguchi et al. (19) suggested that the activation of PKCßII might be a key step through which glucose triggers diabetic complications in microvascular cells. It might seem that certain discrepancies exist between these observations and our present experiments. However, these investigators carried out their experiments using in vivo diabetic rat models subjected to chronic glucose exposure for 2 to 4 months (5, 19). PKC activity was examined in nonsynchronized VSMC cultured under chronic high glucose conditions for 5 to 10 days (5, 18, 19, 42). Here, we studied the effects of high glucose on PKCßI and PKCßII mRNA on synchronized VSMC during the first cell cycle postsynchronization and demonstrated that glucose-induced destabilization of PKCßII mRNA occurs during the course of the cell cycle. Also, instead of PKCßII, the PKCß2 nomenclature has been used on occasions that would imply that PKCßI was the isoform increased by hyperglycemia (40, 43). Moreover, King and co-workers (5, 18, 19) reported an increase in the PKC activity as reflected by increases in the membrane PKC fraction under chronic glucose conditions. In none of these studies were PKCß mRNA levels examined with acute glucose exposure. Our present studies examined the regulation exerted by acute glucose exposure on the PKCß gene and have shown that glucose acted both transcriptionally and posttranscriptionally, with the latter effect accounting for the majority of the down-regulation of PKCßII mRNA.
We have elucidated several molecular mechanisms involved in regulating PKCß gene expression in VSMC. We suggested that acute high glucose could exert important regulatory effects through a putative carbohydrate response element located upstream of the PKCß transcription start site and through glucose-induced posttranscriptional destabilization of PKCßII message via a nuclease activity present in the cytosol. In this case, the rate of mRNA degradation plays an important role in establishing levels of gene expression. Although hormones, growth factors, and ions are known to induce changes in mRNA stability (for review, see ref 39), here we determined that PKCßII mRNA levels are regulated primarily by mRNA destabilization in response to glucose, a cellular nutrient. This multilevel regulation of gene expression by glucose suggests that PKCßII may play a pivotal role in vascular smooth muscle cell function.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Abbreviations: AoSMC, aortic smooth muscle cells; bp, base pairs; CAT, chloramphenicol acetyltransferase; ChoRE, carbohydrate response element; DMEM, Dulbecco's modified Eagle's medium; DPBS, Dulbecco's phosphate-buffered saline; FBS, fetal bovine serum; kb, kilobase(s); PKC, protein kinase C; PNE, postnuclear extract; RT-PCR, reverse transcriptase-polymerase chain reaction; TPA, 12-O-tetradecanoylphorbol-13-acetate; UTR, untranslated region; VSMC, vascular smooth mucle cells.
Received for publication July 8, 1998.
Revision received August 25, 1998.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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