1 Department of Biochemistry and Molecular Biology and 2 Department of Internal Medicine, University of South Florida, College of Medicine, and 3 Research Service, J. A. Haley Veterans Hospital, Tampa, Florida 33612
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ABSTRACT |
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Accelerated vascular smooth muscle cell
(VSMC) proliferation contributes to the formation of atherosclerotic
lesions. To investigate protein kinase C (PKC)-II functions with
regard to glucose-induced VSMC proliferation, human VSMC from aorta
(AoSMC), a clonal VSMC line of rat aorta (A10), and A10 cells
overexpressing PKC-
I (
I-A10) and PKC-
II (
II-A10) were
studied with the use of three techniques to evaluate glucose effects on
aspects affecting proliferation. High glucose (25 mM) increased DNA
synthesis and accelerated cell proliferation compared with normal
glucose (5.5 mM) in AoSMC and A10 cells, but not in
I-A10 and
II-A10 cells. The PKC-
II specific inhibitor CGP-53353 inhibited
glucose-induced cell proliferation and DNA synthesis in AoSMC and A10
cells. In flow cytometry analysis, high glucose increased the
percentage of A10 cells at 12 h after cell cycle initiation but
did not increase the percentage of
I-A10 or
II-A10 cells entering
S phase. PKC-
II protein levels decreased before the peak of DNA
synthesis, and high glucose further decreased PKC-
II mRNA and
protein levels in AoSMC and A10 cells. These results suggest that high
glucose downregulates endogenous PKC-
II, which then alters the
normal inhibitory role of PKC-
II in cell cycle progression,
resulting in the stimulation of VSMC proliferation through acceleration of the cell cycle.
atherosclerosis; cell cycle; diabetes mellitus; protein kinase C inhibitor; thymidine
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INTRODUCTION |
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THERE IS AN APPARENT INCREASE in the prevalence of atherosclerosis in diabetic patients (31). Coronary atherosclerotic disease is a primary cause of death in diabetic patients (7), and diabetes mellitus is a well-established risk factor for peripheral vessel disease caused by atherosclerosis (20). Hyperglycemia is probably an important etiologic factor in the development of macrovascular complications in diabetic patients (37, 39). Although the development of atherosclerotic lesions in diabetes mellitus is a complex and dynamic interaction of many factors, the mechanism by which hyperglycemia might cause vascular complications remains unknown.
It is widely recognized that vascular smooth muscle cell (VSMC) proliferation is a key event in the formation of atherosclerotic lesions (29). Abnormal proliferation of VSMC with subsequent formation of intimal thickening is thought to play a role in the development of lesions (34). Prolonged high glucose exposure alters calcium channels (30), growth rates (23, 9), and protein kinase C (PKC) levels (8, 11, 19, 38, 41) in VSMC. High glucose activates PKC in several cell types including VSMC (8, 11, 19, 23, 38, 41).
PKC is a complex family of at least 11 isozymes classified into 3 groups: classic PKCs (PKC-, -
I, -
II, and
), which require diacylglycerol (DAG), phospholipids, and Ca2+ for full
activity; novel PKCs (PKC-
, -
, -
, and -
), which are
phospholipid and DAG dependent but Ca2+ independent; and
atypical PKCs [PKC-
, -(
)
, and -µ], which require only
phospholipids (12, 25, 35). In
VSMC, PKC modulates contraction (28), signal transduction
(4, 18, 40), growth rates
(23), and DNA synthesis or cell cycle progression
(1, 13, 14, 15,
24, 32, 33, 36).
Although numerous studies have been performed, little is known about
isozyme-specific actions of PKC on VSMC proliferation.
We have shown that PKC-I has a stimulatory effect and PKC-
II an
inhibitory effect on the cell cycle progression in VSMC, because stable
overexpression of PKC-
I (
I-A10) and PKC-
II (
II-A10) in A10
cells, a clonal VSMC line derived from rat aorta, altered cell cycle
regulation (42). In the present study, we investigated acute effects of extracellular high glucose on AoSMC (primary VSMC
cultures of human thoracic aorta) as well as A10,
I-A10, and
II-A10 cell cycle progression.
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METHODS |
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Materials.
A10 (ATCC CRL 1476) were obtained from American Type Culture
Collection, and AoSMC were obtained from Clonetics (San Diego, CA).
Dulbecco's modified Eagle's medium (DMEM), 0.04% trypsin-EDTA, fetal
bovine serum (FBS), Dulbecco's phosphate-buffered saline (DPBS),
penicillin G, and streptomycin sulfate were purchased from GIBCO BRL
(Gaithersburg, MD). Smooth muscle growth medium (SMGM), human
recombinant epidermal growth factor (hEGF), human recombinant
fibroblast growth factor (hFGF), dexamethasone, gentamicin, and
amphotericin-B were purchased from Clonetics. Culture materials including 75- and 25-cm2 flasks, 6-, 12-, 24-, and
96-well plates, and 100-mm culture dishes were purchased from
Costar (Cambridge, MA). EDTA and neomycin derivative G418
(Geneticin) were purchased from GIBCO (Grand Island, NY).
D-Glucose, mannitol, propidium iodide (PI),
phenylmethylsulfonyl fluoride (PMSF), leupeptin, aprotinin, and
nonylphenoxypolyethoxy ethanol (NP-40) were purchased from Sigma (St.
Louis, MO). The cell proliferation assay kit was purchased from Promega
(Madison, WI). [3H]thymidine (dThd; 1 mCi) was purchased
from Du Pont NEN (Boston, MA). The protein assay kit with bovine serum
albumin (BSA) for a standard and goat anti-rabbit -globulin coupled
to horseradish peroxidase (HRP) were purchased from Bio-Rad (Hercules,
CA). Specific antibodies for PKC-
I and -
II were purchased from
Santa Cruz (Santa Cruz, CA). The enhanced chemiluminescence (ECL)
system was purchased from Amersham (Arlington Heights, IL). Specific PKC inhibitors, CGP-53353 and CGP-41251, were kindly provided by Dr. D. Fabbro (Novartis Pharma, Basel, Switzerland). CGP-53353 inhibits
PKC-
II with an IC50 of 0.41 mM and PKC-
I with an
IC50 of 3.8 mM. CGP-41251 is a staurosporine derivative
that inhibits cPKC (IC50 = 0.05 µM) (7a).
Cell culture. The clonal VSMC line A10 derived from fetal rat aorta has been characterized as a model for the investigation of various drug and hormone effects on biochemical changes (16). Cells were grown in DMEM with 5.5 mM glucose containing 10% FBS, 50 U/ml penicillin G, and 50 µg/ml streptomycin sulfate at 37°C in a humidified, 5% CO2-95% air atmosphere in either 25-cm2 flasks (for cell cycle analysis), 6-well plates (for Western blot analysis), 24-well plates (for [3H]dThd incorporation study), or 96-well plates (for cell proliferation assay). Early passages, less than five following plating from frozen stocks, of cells were used here. To evaluate A10 as a model for examining glucose effects on cell cycle progression, we used primary cultures of AoSMC. These cells were grown in SMGM containing 5.5 mM glucose, 5% FBS, 10 ng/ml hEGF, 2 ng/ml hFGF, 390 ng/ml dexamethasone, 50 mg/ml gentamicin, and 50 ng/ml amphotericin-B at 37°C in a humidified, 5% CO2-95% air atmosphere according to the company recommendations. Cells were grown to >90% confluency, and media were changed every 5 days during growth.
Overexpression of PKC-I and -
II isoforms in A10 cells.
The rat cDNA encoding PKC-
I in the expression vector pMV7
(10) and PKC-
II in pMTH (22) were used to
transfect A10 using calcium phosphate coprecipitation. The PKC
expression vectors were kind gifts from Dr. Harald Mischak (GSF,
Münich, Germany) (22). A10 cells maintained in DMEM
containing 10% FBS with 5.5 mM glucose were trypsinized with 0.1%
trypsin-0.4% EDTA, seeded on 100-mm culture dishes in the same medium,
and grown to confluency. Cells were then incubated for 4 h with
calcium phosphate-DNA coprecipitates containing 15 µg of the
expression vector or the empty vector as a control. After 18 h,
cells were switched to fresh medium. Within 48 h, cells were
trypsinized and replaced in DMEM supplemented with 10% FBS with 600 µg/ml of the Geneticin to select for cells with neomycin resistance
as described previously (42). After 10-16 days in
selection medium, cells were examined for the presence of PKC-
I and
-
II isoform overexpression by Western blot analysis. Overexpression
was defined for these cells as a four- to fivefold increase in PKC
content measured by Western blot analysis compared with A10 or empty
vector-transfected control cells as described previously
(42).
Cell proliferation assayed by MTT method.
The effects of glucose on cell proliferation were examined in AoSMC,
A10, I-A10, and
II-A10 cells. A10,
I-A10, and
II-A10 cells
in secondary cultures were trypsinized and seeded in 96-well plates at
~1 × 103 cells/well in 100 µl of DMEM containing
10% FBS with various concentrations (5.5-40 mM) of
D-glucose and 5.5 mM D-glucose plus 34.5 mM
mannitol as an osmotic control. To determine the contribution of
PKC-
II function in glucose-induced cell proliferation, cells were
also treated with CGP-53353, a PKC-
II-specific inhibitor (27), and CGP-41251, a classic PKC inhibitor (7a). Cell
proliferation assay was performed using the cell proliferation assay
kit as follows. After incubation at 37°C for 48-96 h, 15 µl
of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT)
were added (final concentration 0.5 µg/ml). After 4 h
incubation, 100 µl of 20% sodium dodecyl sulfate (SDS) were added to
dissolve the formazan crystals formed. Optical density was measured
with an ELISA plate reader using test and reference wavelengths of 570 and 630 nm, respectively. AoSMC were also examined under the same
conditions but with SMGM to determine whether glucose effects were
similar in AoSMC. For the A10 and AoSMC, we established a linear
relationship between the MTT reduction assay and viable cell number.
Expression of the empty vector in A10 cells had no effect on the
response to glucose reported here.
Measurement of [3H]dThd incorporation into DNA.
The effect of glucose on DNA synthesis in A10, I-A10, and
II-A10
cells was determined using [3H]dThd incorporation in 5.5 mM (normal) and 25 mM (high) glucose as described previously
(42). Cells were seeded on 24-well plates, and
G0/early G1 synchronization was achieved by
serum deprivation. Media were switched to DMEM containing 10% FBS with
normal or high glucose for 48 h. Every 6 h, cells were
incubated with 1 ml of the same medium containing 2.0 µCi/ml
[3H]dThd for 1 h at 37°C. Cells were rinsed with
ice-cold DPBS buffer, and acid-soluble radioactivity was removed by
treatment with 10% trichloroacetic acid. Cells were then dissolved
with 1 N NaOH and neutralized with 1 N HCl. The [3H]
radioactivity in acid-soluble pools was determined, and DNA synthesis
was estimated [counts per minute (cpm)/well]. Expression of the empty
vector had no effect on DNA synthesis.
Cell cycle analysis using flow cytometry.
The effects of glucose on cell cycle progression were analyzed in A10,
I-A10, and
II-A10 cells using flow cytometry as described previously (42). Cells were seeded, and
G0/early G1 synchronization was achieved by
serum deprivation. Media were switched to DMEM containing 10% FBS with
5.5 mM (normal) or 25 mM (high) glucose to initiate the cell cycle.
Cells were sampled at the indicated time after initiation of the cell
cycle. Cells were washed with ice-cold DPBS and fixed with iced
ethanol. Cells were then removed from ethanol and stained with 1 ml of
DPBS containing 50 µg/ml PI. Analysis of the stained nuclei was
performed by using a fluorescence-activated cell analyzer with cell
cycle-analyzing computer software (FACStarPlus, Becton Dickinson, San
Jose, CA) in 1×106 total cells. The expression of empty
vector had no effect on cell cycle progression reported here.
Western blot analysis.
The changes in immunoreactive PKC-I and -
II after 5.5 mM (normal)
and 25 mM (high) glucose exposures were measured in AoSMC, A10,
I-A10, and
II-A10 cells by Western blot analysis using specific
PKC-
I and -
II antibodies. Cells in secondary cultures maintained
in DMEM containing 10% FBS or SMGM containing 5% FBS with 5.5 mM
glucose were trypsinized and seeded on 6-well plates at a density of
1×106 cells/well in 3 ml of the same medium. When cells
were grown to 80% confluency, G0/early G1
synchronization was achieved by serum deprivation for 72 h. The
cell cycle was then initiated, and cells were incubated for the
indicated times at 37°C with 5.5 mM (normal) or 25 mM (high) glucose.
Subsequently, cells were washed with ice-cold DPBS, scraped from dishes
with a rubber policeman, and centrifuged at 2,000 rpm for 5 min, and
the cell pellet was lysed in a buffer containing 20 mM Tris · HCl (pH 7.5), 145 mM NaCl, 10% (wt/vol) glycerol, 5 mM EDTA, 0.2 mM
Na2VO4, 0.1 mM PMSF, 10 µg/ml leupeptin, 10 µg/ml aprotinin, and 1% (wt/vol) NP-40. The suspension was sonicated
for 5 s and centrifuged at 2,000 rpm for 5 min, and the resultant
supernatant was used as the whole cell lysate fraction. The protein
concentration was determined by the method of Bradford (2)
with BSA as a standard. All of the procedures were performed at 4°C.
The samples were dissolved in Laemmli's sample buffer containing 1%
SDS (17) and resolved by SDS-PAGE on 4.5% (wt/vol)
stacking gels and 10% (wt/vol) resolving gels in a Mini-Protein II
dual-slab cell (Bio-Rad) and electrophoretically transferred to
nitrocellulose membranes (Bio-Rad). Membranes were equilibrated in
Tris-buffered saline (TBS; 20 mM Tris · HCl and 50 mM NaCl, pH
7.40), and nonspecific binding sites were blocked by 10% dried milk in
TBS containing 0.1% Tween 20 (TBS-T) at room temperature for 1-2
h. Membranes were then incubated in TBS containing antibodies to
PKC-
I and -
II (Santa Cruz) at room temperature for 3 h.
After being washed with TBS-T, membranes were incubated in TBS-T
containing a goat anti-rabbit
-globulin coupled to HRP (Bio-Rad) for
30 min. The blots were developed with the ECL system (Amersham) and
visualized by exposure to Hybond-C extra ECL film (Amersham). The
developed films were analyzed by a densitometric scanner linked to a
Macintosh computer (Apple, Cupertino, CA). Less than five consecutive
passages of A10 cells were used for determination of PKC-
II levels,
because it has been reported that PKC isozyme expression varies with
multiple passages of VSMC (8). Expression of empty vector
had no effect on the PKC-
II expression reported here, as described
previously (42).
Data analysis. Experiments were repeated at least three times to ensure reproducible results. Data are expressed as means ± SE of the number of observations. The statistical significance was assessed by one-way analysis of variance (ANOVA) or Student's t-test. P < 0.05 was considered statistically significant. Significance was determined after three or more separate experiments.
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RESULTS |
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A10 cells as a model for human VSMC to demonstrate high glucose
effects on cell proliferation and DNA synthesis.
To establish A10 cells as an appropriate model to demonstrate acute
effects of high glucose on VSMC proliferation, we examined cell
proliferation in both human AoSMC and rat A10 cells using a dye-based
assay that measured relative cell number. AoSMC proliferation increased
up to 182.6 ± 17.3% of control (P < 0.01) as
media glucose concentrations increased; A10 cell proliferation also
increased up to 151.8 ± 13.0% of control (P < 0.01), similar to AoSMC. Mannitol, an osmotic control, had no
stimulatory effect on AoSMC and A10 cell proliferation (Fig.
1A). dThd incorporation was
also assessed, because its incorporation directly reflects DNA
synthesis. After serum was added to reinitiate the cell cycle in
synchronized cell cultures, DNA synthesis was calculated as
[3H]dThd activity (cpm/well). In AoSMC,
[3H]dThd incorporation increased following serum addition
to a maximum level of 729.8 ± 77.1% of control by 30 h with
normal (5.5 mM) glucose. With high (25 mM) glucose,
[3H]dThd incorporation increased to a maximum level of
985.0 ± 14.0% of control after 24 h, which demonstrated
that high glucose stimulated and accelerated DNA synthesis in AoSMC . In A10 cells, DNA synthesis increased to 704.3 ± 78.0% of
control by 24 h with normal glucose. With high glucose, DNA
synthesis increased to 927.1 ± 71.0% of control 12 h after
reinitiation of the cell cycle (Fig. 1B). Although the peak
of [3H]dThd incorporation into DNA occurred at slightly
different times in A10 and AoSMC, high glucose increased and
accelerated DNA synthesis in both A10 and AoSMC. From these results,
A10 cells were determined to be an analogous model for elucidating
glucose-induced cell proliferation mechanisms in VSMC.
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PKC-II was involved in high glucose-induced A10 cell
proliferation.
Cell proliferation as determined by cell number was significantly
increased in A10 and AoSMC as media glucose concentrations increased
(Fig. 1A). Glucose-induced cell proliferation in A10 cells
was blocked by CGP-53353, a PKC-
II-specific inhibitor, at 1 µM
(Fig. 2A). High glucose
concentrations did not significantly alter
I-A10 and
II-A10 cell
proliferation rates, although PKC-
I overexpression increased and
PKC-
II overexpression decreased cell proliferation in normal glucose
as described previously (42). Stable overexpression of
PKC-
I and -
II blocked the effect of glucose concentrations on
cell proliferation. Although CGP-53353 did not have a significant block
on
I-A10 cell proliferation (Fig. 2B), CGP-53353
increased
II-A10 cell proliferation slightly (Fig. 2C) as
anticipated, demonstrating specific inhibition for PKC-
II by
CGP-53353. Longer treatment periods (1 wk) with CGP-53353 increased
II-A10 cell proliferation more than that shown in Fig. 2, but cell
viability of
I-A10 cells was decreased 72-96 h postincubation (data not shown). CGP-41251, a cPKC inhibitor, inhibited A10,
I-A10,
and
II-A10 cell proliferation by 60-80% without regard for
glucose concentrations (data not shown). Mannitol had no stimulatory effect on A10,
I-A10, or
II-A10 cell proliferation.
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Glucose-stimulated [3H]dThd incorporation was blocked
by CGP-53353 in A10 but not in I-A10 and
II-A10 cells.
DNA synthesis was examined by dThd incorporation following serum
addition to reinitiate the cell cycle in synchronized cells. DNA
synthesis occurred between 6 and 30 h after cell cycle initiation in A10 cells (Fig. 3A).
[3H]dThd incorporation was observed only at the 6-h time
point in
I-A10 cells (Fig. 3B), but incorporation
occurred between 12 and 42 h in
II-A10 cells with normal
glucose levels (Fig. 3C). This alteration in DNA synthesis
induced by PKC-
I and -
II overexpression supports our previous
demonstration that PKC-
I overexpression accelerated and PKC-
II
overexpression delayed the peak of [3H]dThd incorporation
(42). In A10 cells, high glucose significantly increased
the incorporation of [3H]dThd from 48.6 ± 8.2 ×10
4 cpm/well at 24 h to 65.0 ± 4.9 ×10
4 cpm/well at 6 h after cell cycle initiation
(P < 0.05) that was inhibited by CGP-53353 in A10
cells (Fig. 3A). However, no significant effects of high
glucose were found in
I-A10 and
II-A10 cells. Stable
overexpression of PKC-
I or -
II made the cells resistant to high
glucose effects, and CGP-53353 had no effect on DNA synthesis in
I-A10 cells but increased DNA synthesis slightly in
II-A10 cells
(Fig. 3, B and C).
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High glucose increased the percentage of A10 cells, but not
I-A10 and
II-A10 cells, that entered S phase.
To further characterize the effects of glucose on the cell cycle,
flow cytometry was performed. In contrast to [3H]dThd
incorporation, flow cytometry reveals the percentage of cells that have
completed DNA synthesis and the percentage of cells undergoing
G2/M phase. Flow cytometric analysis demonstrated that the
duration of S phase was ~8 h (from 8 to 16 h), and the peak of S
phase occurred 12 h after cell cycle initiation in A10 cells (Fig.
4A). In A10 cells, high
glucose exposure increased the percentage of S-phase cells at 12 h
compared with normal glucose exposure (Fig. 4A), and
CGP-53353 blocked the glucose-induced increase of S-phase cell
percentage (data not shown). However, no significant effect of high
glucose on phase distribution was noted in
I-A10 and
II-A10 cells
(Fig. 4, B and C) even though overexpression of
PKC-
I and -
II altered the duration of S phase as described
previously (42). PKC-
I overexpression accelerated and
shortened S phase, and PKC-
II overexpression slowed and prolonged S
phase. The flow cytometry data supported the results of
[3H]dThd incorporation into DNA. In this case, however,
cells that had completed DNA synthesis were sorted later than those
reflecting active DNA synthesis.
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High glucose did not affect PKC-I expression in AoSMC, A10, or
I-A10 cells.
During cell cycle progression in AoSMC, A10, and
I-A10 cells,
contents of PKC-
I were analyzed by Western blot (Fig.
5). Levels of PKC-
I did not change
significantly (from 75 ± 5.5% to 120 ± 6.5% of control)
compared with the control levels during normal and high glucose
exposure in AoSMC, A10, and
I-A10 cells. The overexpression of
PKC-
I is not evident in this blot because exposure time with the
film was shortened to stay in the linear range of the ECL reagent.
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High glucose exposure-induced downregulation of PKC-II in AoSMC
and A10 but not in
II-A10 cells.
During cell cycle progression in AoSMC, A10, and
II-A10 cells,
PKC-
II immunoreactivities were demonstrated by Western blot analysis
(Fig. 6). Peaks of DNA synthesis as
measured by [3H]dThd incorporation correlated to changes
in PKC-
II levels following normal and high glucose exposure as
follows. With normal glucose, PKC-
II levels in AoSMC decreased to
68.0 ± 5.1% of control before the peak of DNA synthesis, and
PKC-
II levels in A10 cells decreased to 49.0 ± 3.5% of
control before the peak of DNA synthesis, similarly. With high glucose,
PKC-
II levels in AoSMC decreased to 25.0 ± 1.9% of control
before the peak of DNA synthesis, and PKC-
II levels in A10 cells
decreased to 20.0 ± 1.8% of control. Decreases in PKC-
II
levels were observed at ~10-12 h before DNA synthesis in both
AoSMC and A10 cells and with high glucose exposure. Greater decreases
in PKC-
II levels occurred for longer periods of time in high glucose
than with normal glucose exposure. The downregulation of PKC-
II in
A10 cells presumably reflects the glucose-induced activation and
turnover of enzyme as well as the destabilization of PKC-
II mRNA
(26). In
II-A10 cells, PKC-
II levels were sustained
in normal glucose (74.0 ± 4.3-85 ± 7.3% of control) and in high glucose (105 ± 7.3-120 ± 10.3% of
control) during the cell cycle and increased somewhat with high glucose
compared with normal glucose. The development of the blots was
optimized to detect relative changes in PKC-
II levels with time, and
overexpression was not pronounced. We previously demonstrated PKC-
I
plus -
II overexpression (42). The sustained increase of
PKC-
II in
II-A10 cells probably reflects the high level of
heterogeneous promoter-driven PKC-
II overexpression that is
resistant to glucose regulation at transcriptional and
posttranscriptional levels (26). Vector controls were
identical to wild-type A10 cells.
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DISCUSSION |
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In this report, we used three methods to evaluate cell
proliferation in response to normal and high glucose levels. Each
method reveals a different aspect of cell proliferation. The uptake of [3H]thymidine reflects active DNA synthesis at specific
times. The G1/S flow cytometry data reflect the total
percentage of cells relative to DNA content. The MTT dye method
reflects cell number. When the methods are examined together, they
present a cohesive description of glucose effects in vascular smooth
muscle cells. This study showed that human and rat VSMC proliferation
was increased by high (25 mM) glucose exposure compared with normal
(5.5 mM) glucose, and proliferation was dependent on increasing glucose concentrations. High glucose-induced stimulatory effects were not due
to changes in osmolarity. The high glucose effect on cell proliferation
is through increased DNA synthesis, resulting in acceleration of the
cell cycle. High glucose increased and accelerated DNA synthesis and
the entry of cells into S phase and accelerated G1/M phase
as shown in Fig. 4. The agreement of glucose effects in both AoSMC and
A10 cells suggests that A10 cells are a suitable model for further
examining glucose-stimulated cell proliferation mechanisms in VSMC. The
stimulatory effects of high glucose on cell proliferation as shown by
all three methods did not occur in I-A10 and
II-A10 cells, where
PKC-
I and -
II expression was controlled by a promoter-driven
mechanism introduced by the expression vector. The report that high
glucose stimulates the progression of cells from G1 to S/M
phases in rat primary cultured VSMC and A10 cells (9) is
in keeping with our results, because glucose increased the percentage
of S phase cells as shown by flow cytometry.
This study provides evidence that high glucose decreased PKC-II
levels during early S phase in AoSMC and A10 cells as determined by
[3H]dThd incorporation and Western blot analysis.
Although PKC-
II levels were decreased somewhat in early S phase with
normal glucose, decreases in PKC-
II levels were more prominent and
were accompanied by acceleration of DNA synthesis with high glucose
exposure. However, in
II-A10 cells, PKC-
II levels were sustained,
suggesting that PKC-
II overexpression made cells resistant to
glucose-induced changes before S phase. Significant changes in PKC-
I
levels were not observed before S phase in AoSMC, A10, or
I-A10 cells.
PKC was activated by high glucose in VSMC (23). Several
studies suggested that prolonged high glucose exposure leads to the
downregulation of PKC activity following the glucose-induced PKC
activation in rat nerves (6) and soleus muscles
(5). We previously demonstrated that overnight glucose
exposure downregulated PKC- (
I +
II) levels in A10 cells,
although no distinction between PKC-
I or -
II was made
(4). In this study, the specific downregulation of only
PKC-
II by acute high glucose exposure during early S phase was
demonstrated. This decrease might result from PKC-
II activation by
acute high glucose exposure; however, regulation of PKC-
II gene
expression may also be occurring (26).
Other studies suggested that prolonged or chronic high glucose exposure
leads to a sustained elevation of PKC levels in vascular tissues
(8, 11, 19, 38,
41). These findings may seem to be at odds with the
present study; however, several differences between our experiments and
those studies exist. First, we examined PKC-I and -
II
immunoreactivities, whereas previous studies demonstrated total PKC
activity. Second, we synchronized proliferating cells before high
glucose exposure to demonstrate glucose effects during the first cell
cycle following synchronization. The period from 0 to 18 h and
proceeding to 48 h after cell cycle initiation was observed for
acute glucose effects on PKC-
I and -
II expression in the first
cell cycle postsynchronization. Previous studies examined PKC levels
after chronic high glucose exposure (7-14 days) of nonsynchronized
(quiescent) cell cultures in vitro (8, 11,
19, 38, 41) or diabetic animals
in vivo (11, 19). Finally, we demonstrated
PKC-
I and -
II immunoreactivity in whole cell lysates to show
changes of total PKC-
I and -
II expression during cell cycle
progression, whereas previous studies demonstrated subcellular
glucose-induced PKC activity translocation (8, 11, 19, 38, 41) or
PKC immunoreactivity (8, 11) in cytosol and
membrane fractions. PKC activation occurs unavoidably with various
homogenization techniques used in PKC assay studies, and
chromatographic purification of PKC activity may not actually correlate
with PKC protein levels (3). Thus PKC-
II downregulation induced by acute high glucose exposure preceding early S phase as
demonstrated here is a different phenomenon from chronic high glucose-induced effects previously reported (8,
11, 19, 38, 41).
Although previous studies demonstrated involvement of PKC in VSMC
proliferation using PKC inhibitors, phorbol esters, and various growth
factors (1, 13-15, 18,
24, 32, 33, 36, 40), little is known about the roles of specific PKC
isoforms. In this study, we extended our previous report, which
demonstrated that PKC-I and -
II might have regulatory roles in
the G1/S transition of A10 cell cycle progression
(16) to primary cultures of human VSMC. In AoSMC, like in
A10 cells, decreases in PKC-
II levels were significantly sustained
before early S phase in high glucose compared with normal glucose
conditions. Some quenching of PKC-
II expression occurred normally in
G1/S or early S phase during progression of the cell cycle.
PKC-
II levels were unaltered in
II-A10 cells overexpressing the
isoform in both high and normal glucose conditions. This suggests that
the vector-driven overexpression of PKC-
II overcomes glucose effects
on gene expression during the cell cycle. This is anticipated because
the PKC-
II cDNA encodes only translated regions of PKC-
II, and
cis-elements regulating transcriptional and
posttranscriptional processing of mRNA are not present. PKC-
II overexpression blocked glucose-stimulatory effects on proliferation as
predicted when PKC-
II levels could no longer be downregulated.
Cell cycle progression may be regulated by several PKC isoforms in a
multilevel fashion, because proliferative and antiproliferative actions
of PKC on VSMC proliferation were described previously (1,
9, 13, 14, 16,
18, 24, 32, 33,
36, 40). It is not possible to explain the
changes in the entire cell cycle time course only by alterations in
PKC-II levels. However, PKC-
II may operate as a negative
gate keeper in serum-stimulated VSMC proliferation even in the absence
of exogenous PKC activators, because downregulation of PKC-
II was
associated with accelerated DNA synthesis and increased entry of cells
into S phase.
The glucose-induced subcellular translocation of PKC-II in VSMC was
previously demonstrated (8, 11), but the
underlying mechanisms responsible for the specific regulation of
PKC-
II by elevated glucose levels in VSMC remain unclear. To our
knowledge, this is the first demonstration of a link between
glucose-stimulated PKC-
II isozyme downregulation and the
glucose-induced acceleration of cell cycle progression in VSMC.
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ACKNOWLEDGEMENTS |
---|
We thank Dr. Harald Mischak, Institut für Klinische Molekular Biologie und Tumorgenetik, GSF (Münich, Germany) for providing the PKC expression vectors. We thank Bobbye Hill, Moffit Cancer Center, and Jian Zhang, Internal Medicine, University of South Florida Medical School, for helpful advice in establishing the cell cycle analysis of VSMC, and Todd Stadeford for assisting with Western blots.
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FOOTNOTES |
---|
This study was supported by American Heart Association, Florida Affiliate Grant-in-Aid 9810139 (to D. R. Cooper), by funds from the Medical Research Service of The Veterans Administration (to D. R. Cooper), and by National Science Foundation Grant MCB-9808128 (D. R. Cooper). M. Yamamoto was a recipient of an Uehara Memorial Foundation Research Fellowship.
Address for reprint requests and other correspondence: D. R. Cooper, J. A. Haley Veterans Hospital, VAR 151, 13000 Bruce B. Downs Blvd., Tampa, FL 33612 (E-mail: dcooper{at}com1.med.usf.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Received 30 July 1999; accepted in final form 16 March 2000.
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