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Department of Internal Medicine, College of Medicine, KonKuk University, Chung Ju 380-704, South Korea
1 Department of Food and Nutrition Hoseo University, 29-1 Sechul-Ri, BaeBang-Myun, Asan-Si, Chungnam-Do 336-795, South Korea
(Requests for offprints should be addressed to S Park; Email: smpark{at}office.hoseo.ac.kr)
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Abstract |
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Introduction |
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In contrast, exercise is well known to lessen hyperglycemia and insulin resistance by improving glucose utilization in muscles and decreasing fat deposits in the body (Sato et al. 2003, Hawley 2004). Few studies have been performed to verify whether exercise affects ß-cell function and mass in diabetic rats (Shima et al. 1997). The effect of exercise and DEX on ß-cell function and mass and their mechanism have not been revealed, even though both are involved in glucocorticoid metabolism. Therefore, we studied whether (1) DEX and exercise affect islet function and mass along with insulin resistance in pancreatectomized (Px) and sham-operated rats, and (2) whether the insulin/IGF-I signaling cascade in ß-cells is associated with their modulation.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Male Sprague–Dawley rats weighing 248 ± 14 g had 90% of their pancreas removed using the Hosokawa technique (Hosokawa et al. 1996) or received a sham pancreatectomy operation. After a 90% pancreatectomy (Px), the pancreas which remained was within 2 mm of the common bile duct and extended from the duct to the first part of the duodenum. Px rats with random fed serum glucose levels less than 9.4 mM were excluded after 2 weeks from surgery, and the Px rats included in the experiments showed characteristics of mild diabetes mellitus. A sham pancreatectomy was performed by disengaging the pancreas from the mesentery and gently rubbing it between the fingers. The sham-operated rats did not have any symptoms of diabetes. All experimental animals were allowed free access to standard laboratory food (Sam Yang Co., Kangwon-Do, Korea) and water. They were housed individually in stainless-steel cages in a controlled environment (23 ° C and a 12 h light:12 h darkness cycle). All surgical and experimental procedures were performed according to the guidelines of the Animal Care and Use Review Committee at Hoseo University, Korea. Overnight fasted serum-glucose levels, food intake, and body weight were measured weekly every Tuesday at 1000 h.
Px and sham rats were divided into two groups; half of the rats ran on an uphill treadmill at 20 m/min for 30 min four times a week during the experimental periods, and were designated as the exercise group, while the rest did not exercise and were designated as such. Exercised and unexercised rats were further divided into three groups respectively; each group received a daily oral administration of 0.1 mg cellulose/kg body weight (bw) (control), 0.1 mg DEX (Yuhan Co., Seoul, Korea)/kg bw (HDEX) or 0.01 mg DEX/kg bw (a low dosage treatment of DEX (LDEX)) for 8 weeks.
Insulin secretion and insulin resistance
After 7 weeks of treatment, catheters were surgically implanted into the right carotid artery and left jugular vein of rats anesthetized with i.p. injections of ketamine and xylazine (100 mg and 10 mg/kg bw respectively). After 5–6 days of implantation, a hyperglycemic clamp was performed in conscious and overnight fasted rats to determine insulin-secretion capacity (Dobbins et al. 2002). An i.v. bolus of 25% glucose over 5 min was given to instantaneously raise blood glucose to 12 mM. Subsequently, a continuous and variable glucose infusion was applied to hold the rate at a constant 12 mM during the time from 60 to 120 min. Serum glucose and insulin levels from artery blood were measured at 0, 2, 5, 10, 60, 90, and 120 min.
Two days after use of the hyperglycemic clamp, a euglycemic hyperinsulinemic clamp (Choi & Park 2002) was used under the same conditions as the hyperglycemic clamp to determine insulin resistance. Insulin-stimulated whole body glucose flux was estimated using a prime continuous infusion of [3-3H]glucose (10 µCi bolus, 0.1 µCi/min, NEN Life Science, Boston, MA, USA) throughout the clamps. Regular human insulin (Humulin, Eli Lilly and Co.) was continuously infused at a rate of 20 pmol/kg per min in order to raise the plasma insulin concentration to approximately 1100 pM. Blood samples from arteries were collected at 10-min intervals for glucose estimation and 25% glucose was infused as needed at variable rates to clamp glucose levels at approximately 6 mM. To determine plasma [3-3H]glucose concentrations, the plasma was deproteinized with ZnSO4 and Ba(OH)2, dried to remove 3H2O, resuspended in water and disintegrations per minute (d.p.m.) of 3H were recorded. The plasma concentration of 3H2O was determined by the difference between 3H counts with and without drying. Rates of whole body glucose uptake and basal glucose turnover were determined as the ratio of the [3H]glucose infusion rate to the specific activity of plasma glucose (d.p.m./µmol) during the final 30 min of the respective experiments. Hepatic glucose production during clamps was determined by subtracting the glucose infusion rate from the whole body glucose uptake. The rats were anesthetized with sodium pentobarbital (35 mg/kg bw) (Nembutal, Abbott Laboratories) and were killed by decapitation at the end of the clamp. Tissues were rapidly dissected, weighed and frozen in liquid nitrogen, and stored at – 70 ° C until further analysis could be performed.
Serum-glucose levels were analyzed with a Glucose Analyzer II (Beckman, Palo Alto, CA, USA). Serum-insulin levels were measured by RIA (Linco Research, St Charles, MO, USA). Advanced glycated endproducts (AGE) of s.c. tissues were measured using fluorescence methods (Oddeti et al. 1990). Briefly, s.c. tissues were homogenized in PBS and the lysates centrifuged at 10 000 g for 30 min at 4 ° C. The pellet was defatted with chloroform and methanol (2:1, v:v) and digested with collagenase type 7 and proteinase K in PBS for 48 h, followed by overnight incubation with an equal amount of 0.2 M NaOH at 4 ° C. After centrifugation, half of the supernatant was used for fluorescence determination at excitation 370 nm, emission 440 nm to measure general AGE-associated fluorescence. The rest was used for assaying the content levels of hydroxyproline, which was determined by colorimetric measurement.
In order to determine the glycogen content in the liver, its lysates were centrifuged at 100 g for 10 min and the supernatants deproteinized with 1.5 M perchloric acid. The glycogen content was calculated from glucose concentrations derived from glycogen hydrolyzed by 
-amyloglucosidase in an acid buffer (Frontoni et al. 1991). Insulin content in the pancreas was measured by acid-ethanol methods (Hennige et al. 2003). The pancreas was homogenized with acid-alcohol, stored overnight at 4 ° C and followed by centrifugation at 700 g for 30 min. The supernatant was removed and stored at – 20 ° C pending measurement of insulin content by RIA kit (Linco Research).
Immunohistochemistry and islet morphometry
Five to six rats from each group were treated with 5-bromo-2-deoxyuridine (BrdU; Roche Molecular Biochemicals; 100 µg/kg bw) at the end of the 8-week experimental period. Six hours postinjection, pancreas samples were prepared and analyzed as described above (Hennige et al. 2003). The pancreas was dissected, fixed in a 4% paraformaldehyde solution (pH 7.2) overnight at room temperature and embedded in paraffin blocks. Serial 5 µm paraffin-embedded tissue sections were mounted on slides. To prevent the selection of sections with similar areas, after rehydration, every sixth or seventh section was selected to determine ß-cell area, BrdU incorporation, and apoptosis. The randomly chosen sections were immunostained as described above (Hennige et al. 2003).
Endocrine ß-cells were identified by applying a guinea pig anti-insulin antibody in paraffin-embedded pancreatic sections. ß-Cell proliferation was examined by the incorporation of BrdU in ß-cells from rats injected with BrdU. This incorporation was determined by performing a double-label immunohistochemistry with anti-insulin (Zymed Laboratories, South San Francisco, CA, USA) and anti-BrdU antibodies (Roche Molecular Biochemicals) on rehydrated paraffin sections. Apoptosis of ß-cells was measured by terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling kit (Roche) in paraffin sections of the pancreas and counterstained with hematoxylin and eosin to visualize islets (Li et al. 2003).
Pancreatic ß-cell area was measured by acquiring random (selecting every other islet in the window), non-overlapping images from two sets of eight to ten distal images of insulin-stained pancreatic sections at a magnification of 10 x with a Zeiss Axiovert microscope (Carl Zeiss Microimaging, Thorn-wood, NY, USA). Results of ß-cell quantification are expressed as the percentage of the total surveyed area containing insulin-positive cells, measured by IP Lab Spectrum software (Scanalytics, Inc., Fairfax, VA, USA). Pancreatic ß-cell mass was calculated by multiplying the percentage of insulin-positive area by the weight of the corresponding pancreatic portion. The individual ß-cell size was determined as the insulin-positive area divided by the number of nuclei counted in the corresponding insulin-positive structures in immunofluoresence staining, which were chosen at random and corresponded to 125–150 nuclei per sample. Bigger size of individual ß-cell indicates the induction of ß-cell hypertrophy. The number of small ß-cell clusters was determined as the number of measurements in an arbitrarily set area of < 250 µm2 (islets containing less than five nuclei were excluded) and expressed as the percentage of the total number of measurements in the section (Rooman et al. 2002).
To immunostain duct cells, mouse monoclonal anti-cytokeratin-19 antibodies from Zymed were used. ß-Cell proliferation was expressed in the number of BrdU+ß-cells per square millimeter pancreas and was calculated as the total BrdU+ nuclei in ß-cell nuclei per pancreas section, two sections per animal and five to six animals per group. Apoptosis of ß-cells was determined by counting the total number of apoptotic bodies in a ß-cell nucleus, and was calculated in the same manner as that of ß-cell proliferation.
Islet isolation
Pancreatic islets from nine to ten rats of each group were isolated by collogenase digestion at the end of an 8-week treatment of DEX and/or exercise (Hennige et al. 2003). Through the pancreatic duct, 3 ml 1.0 mg/ml collagenase (Sigma) in Dulbecco’s modified Eagle’s medium (DMEM)–high glucose were injected into the pancreas of rats anesthetized with sodium pentobarbital. The pancreas was immediately removed and incubated at 37 ° C for 15 min. The digested pancreas was washed with DMEM–high glucose four times on ice and islets were isolated with a separation medium consisting of Ficoll reagent (Sigma). The islets washed with cold DMEM–high glucose were pooled from two to three rats from each group. Prior to lysing islets, they were administered with 10 nM IGF-I for 10 min to determine insulin/IGF-I signaling cascade.
Immunoblot analysis
Islets isolated from rats treated with DEX and/or exercise, as mentioned above, were lysed with a 20 mM Tris buffer (pH 7.4) containing 2 mM EDTA, 137 mM NaCl, 1% NP40, 10% glycerol and 12 mM -glycerol phosphate and protease inhibitors. After 30 min on ice, the lysates were centrifuged for 10 min at 11 000 g at 4 ° C. Lysates with equivalent amounts of protein (400 µg) were immunoprecipitated with specific antibodies (anti-IRS1 and IRS2 antibodies) or resolved directly by SDS-PAGE. Lysates with equal amounts of protein (30 µg) were used for immunoblotting with specific antibodies against IRS1 (UBI, Waltham, MA, USA), IRS2 (UBI), protein kinase B (PKB or Akt, Cell Signaling Technology, Beverly, MA, USA), phosphorylated PKBSer473 (cell signaling), pancreatic homeodomain protein (PDX-1; Santa Cruz Biotechnology, Santa Cruz, CA, USA), and ß-actin (Santa Cruz Biotechnology) as described above (Giraud et al. 2004). The intensity of protein expression was determined using Imagequant TL (Amersham Biosciences). These experiments were repeated four times for each group.
Statistical analysis
All results are expressed as means ± S.D. Statistical analysis was performed using the SAS statistical analysis program (Committee of SAS Institute 1985). One-way ANOVA was carried out to determine DEX effect in cell culture studies. In animal studies, the two main effects of DEX and exercise were determined by two-way ANOVA, since there was no significant interaction between exercise and DEX. Significant differences in the main effects among groups were identified by Tukey’s tests. Differences with P < 0.05 were considered statistically significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Px rats treated with HDEX, not LDEX, exhibited hyperglycemia and hyperinsulinemia during an overnight fast and increased AGE levels at the end of the experimental periods. Exercise improved glucose homeostasis in HDEX-treated Px rats, which led them to exhibit near normoglycemia and decreased AGE levels similar to the control group (Table 1
). DEX increased serum-insulin levels in a dose-dependent manner in Px rats, compared to the control, and exercise did not reverse this effect. Sham rats maintained normoglycemia despite DEX treatments. Even sham rats with HDEX treatment displayed normoglycemia with hyperinsulinemia, while exercise corrected the hyperinsulinemia induced by HDEX without changing serum-glucose levels (no data shown).
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). Px rats, administered with HDEX, displayed decreased glucose disposal rates in comparison with the control rats, while exercise improved the rates in DEX-treated diabetic rats (P < 0.05, Fig. 1A). However, LDEX did not decrease glucose disposal rates comparable with the control (Fig. 1A).
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). Under clamped conditions of 1100 pM serum-insulin levels, HDEX increased hepatic glucose output by blocking insulin action (Fig. 1B). Exercise reversed elevated hepatic glucose output in the basal and clamped states of HDEX-treated rats. Exercise significantly suppressed glucose output in the livers of LDEX- and placebo-treated Px rats (Fig. 1B). Hepatic glycogen storage appeared to be lower in HDEX-treated Px rats, but it was not significantly different and did not change over time (Fig. 1C P, < 0.05). In contrast with diabetic rats, DEX and exercise did not change hepatic glucose output in the basal state of sham rats, consistent with fasted serum-glucose levels (Fig. 1D). However, in sham rats during clamped states, HDEX did not suppress hepatic glucose output as much as in the control group, while exercise reversed the suppression, returning output to levels of the control (Fig. 1D). Exercise and DEX modulate ß-cell function via alteration of the glucose sensing mechanism
During hyperglycemic clamp, the glucose infusion rates required to elevate basal serum-glucose levels by 6 mM were lower in DEX-treated Px rats in a dose-dependent manner (Fig. 2A). In diabetic rats, exercise returned them to the levels exhibited in the control group. However, decreased glucose infusion rates were not observed in HDEX-treated sham rats, while exercise elevated them but not as much as in Px rats (Fig. 2A).
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). Serum-insulin levels at a basal state were elevated by DEX in a dose-dependent manner in Px and sham rats, while exercise reversed the elevation caused by DEX (Fig. 2B). Insulin secretion at 2 min was lower in HDEX-treated Px and sham rats, but it increased at 5 min, compared with the control group (Fig. 2C). Sham rats treated with HDEX exhibited sufficiently elevated second-phase insulin secretion during 60, 90, and 120 min in order to compensate for impaired first-phase insulin secretion (Fig. 2D). However, HDEX treatment in Px rats did not increase second-phase insulin secretion as in sham rats. The area under the curve of serum-insulin levels tended to be elevated in HDEX-treated sham rats during hyperglycemic clamp, but it appeared to be lower in Px rats which had undergone HDEX treatment (P = 0.07; Fig. 2E). Remaining insulin in the pancreas was lowered by HDEX in Px rats, but the reduction was recovered by exercise (Fig. 2F). However, consistent with second-phase insulin secretion, HDEX did not decrease the remaining insulin contents in the pancreas of sham rats, while exercise increased them.
The expression of glucose transporter 2 (GLUT2) and glucokinase was analyzed under our different treatment paradigms to assess their roles in the impairment of insulin secretion observed in Fig. 2. GLUT2 expression remained unchanged under all conditions (Fig. 3). Glucokinase expression in islets was increased in exercised rats, while its expression was lowered by DEX when compared with the control (Fig. 3).
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After 90% pancreatectomy, the pancreas, including islets in Px rats, was regenerated up to 50–60% of the size of sham rats for 8 weeks, which was the same among all groups of Px rats (no data shown). Regeneration of the pancreas was evident due to an increase in pancreas weight. However, exercise and DEX increased absolute ß-cell mass, which was calculated by multiplying pancreas weight by ß-cell area (percentage of total pancreas area), per section in both Px and sham rats. Although the percentage of the ß-cell area in the pancreas was greater in Px rats than in sham rats, absolute ß-cell mass was lower in Px rats due to a smaller pancreas (Tables 2 and 3). HDEX treatment enlarged ß-cell area and mass in sham rats in higher proportions than in Px rats, suggesting that Px rats expand ß-cell mass at insufficient rates.
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and 3). Even with HDEX treatment, the number of small ß-cell clusters was significantly lower in sham rats when compared with Px rats. Figure 4 exhibits an example of small ß-cell clusters in Px rats. Small ß-cell clusters were checked at random to ensure they were not peripheral parts of larger islets in the series of sections. Most small ß-cell clusters were located close to the ductal complexes, and some of them were within the complex (Fig. 4). An increase in the number of small ß-cell clusters was an indication of increased neogenesis in DEX-treated Px rats, yet exercise did not affect their production in Px and sham rats.
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and 3). DEX had a greater impact on ß-cell proliferation and apoptosis in Px rats than in sham rats. Exercise increased ß-cell mass by increasing the number of ß-cells via elevating ß-cell proliferation and decreasing apoptosis. By contrast, HDEX decreased ß-cell proliferation and increased apoptosis. However, DEX expanded the mass by ß-cell hypertrophy, determined by individual ß-cell size in sham and Px rats, possibly due to increased insulin resistance (Tables 2 and 3). The individual size of a ß-cell is represented by the cellular area with one nucleus. Exercise suppressed ß-cell hypertrophy, which had been induced by DEX. LDEX plus exercise exhibited the biggest additive increase in ß-cell mass via increased neogenesis and proliferation and decreased apoptosis in Px and sham rats (Tables 2 and 3). Exercise and DEX modulate insulin/IGF-I signaling cascade in islets of Px rats via the induction of IRS2 expression
Islets isolated from DEX-administered rats exhibited decreased IRS2 expression (Fig. 5). The effect of DEX on IRS2 expression was reversed by exercise; IRS2 content was elevated in the islets from exercised rats and tyrosine phosphorylation was stimulated in parallel with IRS2 expression levels when isolated islets were administered with 10 nM IGF-I for 10 min. According to the tyrosine phosphorylation of IRS2, the insulin/IGF-I signaling cascade in the islets was positively modulated. Along with the potentiation of the insulin/IGF-I signaling cascade, PDX-1 expression was elevated in islets from exercised rats and lowered in those from HDEX-treated rats. Exercise overcame the attenuation of the insulin/IGF-I signaling cascade resulting from DEX administration in the combination of treatments.
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Discussion |
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Progression into diabetes can be viewed as having definable stages, characterized by changes in serum-glucose levels and ß-cell function. In addition, each stage is marked by important changes in ß-cell mass. Until ß-cell mass increases to compensate for insulin resistance, blood glucose levels remain within a normal range (Weir & Bonner-Weir 2004). Thus, in an insulin-resistant state, ß-cell mass needs to expand abruptly. ß-cell mass is normally tightly regulated through a balance of ß-cell birth (ß-cell replication and islet neogenesis from precursor cells) and ß-cell death through apoptosis (Taylor 1999). Since insulin resistance disrupts this regulation and impairs ß-cell growth and survival, hypertrophy (an enlargement of individual islet size) plays a predominant role in increasing ß-cell mass during insulin-resistant states. This expansion due to hypertrophy could not be sustained for long periods and fell into decompensation, yet it temporarily prevented a sudden progression of diabetes during an insulin-resistant state. In contrast, exercise blocked any hypertrophy in HDEX-treated rats and expanded ß-cell mass by hyperplasia of ß-cells, resulting in longer compensation. Thus, consistent with Taylor’s results, our data revealed that DEX and exercise expanded ß-cell mass via two independent mechanisms, contributing to the modulation of ß-cell function in order to compensate for insulin resistance.
Consistent with our results, Px rats exhibited abnormalities in ß-cell function and mass, even though ß-cell area and small ß-cell clusters were increased (Laybutt et al. 2003). The mechanism that impairs ß-cell function and mass due to insulin resistance has not yet been clarified. An increase in non-esterified fatty acids (NEFA) during an insulin-resistant state stimulated insulin secretion to a moderate extent. However, long-term treatment with NEFA inhibited glucose-induced insulin secretion. Thus, elevated NEFA in an insulin-resistant state contributed to the impairment of ß-cell function (Grill & Qvigstad 2000). Human and animal studies have shown that DEX elevates serum NEFA via increased lipolysis by impairing the anti-lipolytic action of insulin (Mokuda & Sakamoto 1999, Willi et al. 2002). In our study, LDEX-treated rats exhibited higher body weight with increased food consumption compared with the control. In contrast, HDEX treatment decreased weight gain despite increased food intake per 100 g bw compared with the control, possibly due to elevated energy expenditure through peripheral catabolism. We did not measure serum NEFA levels, but we expected that HDEX increased serum NEFA levels, which would participate in elevated insulin resistance and impaired ß-cell function.
Another possible mechanism to reduce ß-cell mass is decreased expression of transcription factors, which are important for islet development, such as PDX-1, Neuro D and hepatic nuclear factor-1a (HNF-1a) (Laybutt et al. 2003). Several studies have demonstrated that mRNA levels of these transcription factors were reduced by 50–60% in Px rats with 120 mg/dl glucose levels after 4 weeks from pancreatectomy (Jonsson et al. 1994, Stoffers et al. 1997). The mRNA expression levels of GLUT2 and glucokinase associated with glucose sensing in ß-cells were downregulated in Px rats with hyperglycemia. During insulin-resistant states, PDX-1 expression in precursor cells, such as ductal and acinar cells, may be elevated during differentiation into ß-cells, and the expression may be reduced when differentiation is completed. Thus, an increase in small clusters transiently prevents induction of hyperglycemia in order to compensate for insulin resistance.
In our study, exercise induced IRS2 expression in islets leading to an enhanced insulin/IGF-I signaling cascade, which, in turn, possibly led to ß-cell mass expansion and improved function. Several studies have demonstrated that IRS2 induction in ß-cells contributed to increasing their function and mass via enhancing insulin/IGF-I signaling (Hennige et al. 2003, Park et al. 2006). However, due to an attenuated insulin/IGF-I signaling cascade, DEX eventually exacerbated ß-cell function, even though hypertrophy could be temporarily sustained to compensate for peripheral insulin resistance. By contrast, exercise plays a crucial role in diabetic treatment, not only by lessening insulin resistance, but also through potentiating ß-cell function and mass via enhancement of the insulin/IGF-I signaling cascade. Enhanced signaling promoted the expansion of ß-cell mass by increasing hyperplasia via increased ß-cell proliferation and decreased apoptosis (Hennige et al. 2003, Hashimoto et al. 2005). Thus, hyperplasia is necessary in order to strengthen ß-cell function to compensate for insulin resistance.
In addition to proliferation, ß-cell neogenesis from precursor cells, such as ductal cells and acinar cells is known to contribute to ß-cell regeneration after 90% partial pancreatectomy and administration of a high dosage of streptozotocin (Rooman et al. 2002, Li et al. 2003, Pospisilik et al. 2003). The present study revealed that DEX increased ß-cell mass not through ß-cell proliferation, but via neogenesis in parallel with elevated insulin resistance. ß-Cell neogenesis was represented by an increased frequency of small ß-cell clusters, and it appeared to be crucial for ß-cell mass expansion during times of increased insulin resistance in which ß-cell proliferation was decreased and apoptosis was increased. Insulin resistance elevated PDX-1 expression in precursor cells of ß-cells during differentiation and in early stages of islet development (Li et al. 2003, Kulkarni et al. 2004, Jetton et al. 2005). However, the elevation of PDX-1 expression can only be maintained in ß-cells if the insulin/IGF-I signaling cascade is activated. Small ß-cell clusters generated from neogenesis required an enhanced insulin/IGF-I signaling cascade for growth and survival. However, DEX attenuated insulin/IGF-I signaling in ß-cells causing small ß-cell clusters to slowly grow into bigger islets. In contrast, the number of small clusters was fewer in exercised rats than in rats treated with DEX in this study, which can be explained as follows: exercise did not stimulate neogenesis as much as DEX, and/or it made newly created small ß-cell clusters grow into bigger islets.
In conclusion, DEX reduces ß-cell numbers but still increases ß-cell mass via hypertrophy of individual ß-cells and neogenesis, which play an important role in temporarily preventing diabetes development and progression when peripheral insulin resistance is induced. However, hypertrophy is insufficient to compensate for glucose dysregulation in Px rats due to inherent low insulin storage and abnormal ß-cell function. In contrast, exercise ameliorates glucose homeostasis by improving ß-cell function and mass as well as reducing insulin resistance. As with DEX treatment, exercise expands ß-cell mass but through a different pathway. Exercise induces hyperplasia by stimulating ß-cell proliferation and suppressing apoptosis via activation of the insulin/IGF-I signaling cascade. Thus, exercise plays an important role in preventing diabetic development and progression during insulin-resistant states.
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Acknowledgements |
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Received in final form 15 March 2006
Accepted 18 March 2006
Made available online as an Accepted Preprint 10 May 2006
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Significant EX effect in two-way ANOVA at P < 0.05. Columns with different superscript letters (a, b, c) were significantly different in Tukey’s test at P < 0.05. HDEX, high dosage DEX treatment; LDEX, low dosage DEX treatment.
