The consumption of soft drinks, which contain high concentrations of fructose, has markedly increased during the last three decades. This has paralleled the increased prevalence of obesity and insulin resistance that are associated with the development of type 2 diabetes and cardiovascular disease [
The development of diabetic complications, such as cardiovascular disease, is a major cause of mortality among older diabetic patients. As first reported by Rubler et al. [
Hydrogen sulfide (H2S) was recognized as the third gasotransmitter to be identified after nitric oxide and carbon monoxide and is synthesized endogenously from L-cysteine primarily via the action of two enzymes, cystathionine-
Therefore, the aims of this study were to investigate any functional and structural changes in ageing diabetic mouse hearts that resulted from long-term (15 months) feeding of a high-fructose diet and to examine for any changes in the levels of endogenous H2S and expression of the three H2S-producing enzymes involved in the pathogenesis of DCM in these mice.
Male C57BL/6J mice (8-week-old) from the Department of Laboratory Animal Science, Fudan University were housed at constant temperature (
For a glucose tolerance test, glucose levels were measured using glucose strips (Onetouch; Johnson) in blood obtained from the tail vein immediately prior to and at 15, 30, 60, 90, and 120 min after an intraperitoneal (IP) injection of a 25% glucose solution (2 g/kg body weight) into mice that were fasted for 16 h. Insulin sensitivity was tested by IP injection of 0.5 units/kg body weight of recombinant human insulin (Humulin 70/30, Eli Lilly and Company) and plasma glucose measurements were in tail vein blood obtained at 0, 15, 30, 60, 90, and 120 min after this injection in mice that were fasted for 4 h. Areas under the curve (AUC) were determined using the trapezoidal rule.
To record 24 h water and food intakes and collect 24 h urine samples, individual mouse was placed in a metabolic cage (Tecniplast, Italy). Starting 3 days before the collection period, mice were acclimatized to this new environment for 6 h each day.
To test left ventricular function, mouse two-dimensional echocardiography was performed using a Vevo770 ultrasound device (VisualSonics Inc.), as previously described [
At the end of the experimental period, mice were fasted for 12 h and then euthanized with 6% chloral hydrate. Blood samples were collected, and their glucose levels were monitored using blood glucose strips (Onetouch; Johnson). Then, plasma was prepared by centrifuging the blood samples at 3000 rpm for 15 min. The plasma levels of triglycerides (TG), total cholesterol (CHO), low density lipoprotein cholesterol (LDL-C), high density lipoprotein cholesterol (HDL-C), blood urea nitrogen (BUN), and creatinine (Cr) were determined by automatic biochemical analyzer (Cobas 6000, Roche). The BUN/Cr index was calculated.
A heart was surgically removed to determine the heart to body weight ratio (HW/BW × 100%). For histological analysis, the ventricles were excised, fixed in 10% formalin for 48 h before dehydration using a graded ethanol series, embedded in paraffin, sectioned at 4
Paraffin embedded myocardial tissues were subjected to immunofluorescence for the detection of CBS (Santa Cruz Biotechnology Company) and CD31 (Abcam Company) which were incubated with the antibodies at a dilution of 1 : 100, overnight at 4°C. After washing, the sections were incubated with Alexa Fluor 488 and Alexa Fluor 594 (Life Technologies) secondary antibody at 37°C for 1 h in the dark. Then, sections were incubated with 4′,6-diamidino-2-phenylindole (DAPI) at room temperature for 5 min to stain nuclei. Fluorescent signals were observed under a fluorescence microscope (Olympus).
Frozen left ventricle tissues were lysed with ice-cold RIPA buffer. Proteins were extracted and quantified using BCA reagent (Shen Neng Bo Cai Corp.). Protein samples were separated on 10% SDS-PAGE gels and transferred to polyvinylidene fluoride (PVDF) membranes (Millipore-Upstate). The membranes were blocked with 5% non-fat milk at room temperature for 1 h and then incubated with antibodies directed against CSE, CBS, 3-MST, Collagen I, Bax, Bcl-2 (Santa Cruz Biotechnology Company), and Collagen III (Abcam Company) at 4°C overnight. After washing with TBST, the membranes were incubated with horseradish peroxidase-conjugated secondary antibodies at room temperature for 1 h. Specific bands were detected with SuperSignal West Pico Chemiluminescent Substrate (Thermo Scientific-Pierce).
Primary neonatal rat cardiac ventricular myocytes (NRCMs) were collected as previously described with some modifications [
The viability of NRCMs which were cultured in 96-well plates was measured by using the Cell Counting Kit-8 (CCK-8) (Dojindo Molecular Technologies), according to the manufacturer’s instructions. The absorbance of CCK-8 was obtained with a microplate reader at 450 nm. The values were normalized to the NG group.
Cellular apoptosis was determined using the Annexin V–FITC apoptosis detection kit (Dojindo Molecular Technologies), according to the manufacturer’s instructions. NRCMs were stained with Annexin V–FITC and propidium iodide (PI) and then the percentage of cell apoptosis was then determined using flow cytometry with a BD FACSCalibur platform (BD Biosciences). The apoptotic ratio was calculated according to the percentage of Annexin V positive apoptotic cells of the total cells. Fluorescent signals were also observed under a laser confocal microscope (Zeiss).
ROS levels in NRVMs were determined by dihydroethidium (DHE, Sigma-Aldrich) fluorescence using confocal microscopy. After treatments for 72 h, cells were washed with PBS and incubated with DHE (10
H2S levels in plasma, urine, and cell culture medium were measured according to previously described methods [
Results were expressed as means ± SEM. Statistical analysis was performed using an SPSS software package, version 13.0 (SPSS, Inc., Chicago, IL, USA). The results for three or more groups were compared using one-way ANOVA followed by Student-Newman-Keuls test. Comparisons between two groups were made using Student’s
Compared to mice that were fed water only, mice that were fed a 30% fructose solution for 15 months exhibited characteristics typical of a severe diabetic phenotype, including marked obesity, hyperglycaemia, and dyslipidemia, but there were no differences in BUN and BUN/Cr index (Table
Physiological and biochemical results for control and ageing diabetic mice induced by a 30% fructose solution fed for 15 months.
Control ( |
30% Fructose ( |
|
---|---|---|
Body weights (g) | 27.90 ± 0.72 | 34.19 ± 0.70 |
Total cholesterol (mM) | 1.54 ± 0.04 | 2.00 ± 0.06 |
Triglycerides (mM) | 0.46 ± 0.04 | 0.38 ± 0.02 |
HDL cholesterol (mM) | 1.13 ± 0.06 | 1.30 ± 0.07 |
LDL cholesterol (mM) | 0.26 ± 0.02 | 0.47 ± 0.04 |
BUN (mM) | 6.68 ± 0.49 | 6.07 ± 0.34 |
Cr ( |
9.56 ± 0.5 | 7.54 ± 0.42 |
BUN/Cr index | 0.7 ± 0.04 | 0.82 ± 0.05 |
Results are means ± SEM.
Twenty-four-hour metabolic characteristics of control and ageing diabetic mice induced by a 30% fructose solution fed for 15 months.
Control ( |
30% Fructose ( |
|
---|---|---|
24 h water intake (mL) | 3.74 ± 0.31 | 6.04 ± 0.38 |
24 h food intake (g) | 0.62 ± 0.19 | 0.32 ± 0.15 |
24 h urine volume (mL) | 1.02 ± 0.16 | 2.13 ± 0.25 |
Results are means ± SEM.
Fifteen months of high-fructose feeding increases fasting blood glucose levels and reduces insulin sensitivity and glucose tolerance in mice. (a) Fasting blood glucose levels of control and high-fructose-fed mice at the beginning and after 15 months of the experimental period. (b) Representative glucose tolerance test curves for control and high-fructose-fed mice after 15 months. (c) Area under the curve (AUC) of glucose tolerance test results was determined for each animal using the trapezoidal rule. (d) Representative insulin tolerance test curves for control and high-fructose-fed mice after 15 months. (e) Area under the curve (AUC) of insulin tolerance test results was determined for each animal using the trapezoidal rule. Results are means ± SEM.
Regarding glucose tolerance and insulin tolerance tests, as expected, mice that were fed the fructose solution developed both impaired glucose tolerance and insulin resistance (Figures
To assess the effects of the long-term high-fructose diet on cardiac function, we used echocardiography to measure cardiac physiological parameters. The representative M-mode images were showed in Figure
Fifteen months of high-fructose feeding induces cardiac dysfunction. (a) Representative M-mode images. (b–e) Echocardiographic parameter analysis. LVEF, left ventricular ejection fraction; LVFS, left ventricular fractional shortening; LVIDs, left ventricular internal dimension systole; LVIDd, left ventricular internal dimension diastole; LVESV, left ventricular end-systolic volume; LVEDV left ventricular end-diastolic volume. Results are means ± SEM. A
After 15 months of high-fructose feeding, increased HW/BW ratio (Figure
Fifteen months of high-fructose feeding induces cardiac remodelling and apoptosis. (a) Heart to body weight ratio (HW/BW × 100%). (b) Representative HE-stained left ventricular sections (scale bar = 250
In the long-term high-fructose-induced diabetic mice, H2S levels in both plasma and urine were significantly lower than those in control mice (Figures
Fifteen months of high-fructose feeding results in reduced H2S levels in plasma, urine, and heart tissues. (a) H2S levels in plasma, (b) H2S levels in urine, and (c) H2S levels in heart tissues. Results are means ± SEM. A
Because the H2S levels were low in the diabetic heart and H2S production depends on CBS, CSE, and 3-MST enzymes, we determined the expression levels of these three enzymes in heart samples. Western blot analysis revealed bands of 61 kDa, 45 kDa, and 33 kDa, which corresponded to CBS, CSE, and 3-MST, respectively. CSE and 3-MST protein expression levels were significantly reduced in the high-fructose-induced diabetic mice as compared with those of controls, whereas CBS protein expression levels were significantly increased in the heart tissues of diabetic mice (Figure
Fifteen months of high-fructose feeding alters CBS, CSE, and 3-MST protein expression. (a) Representative Western blots and quantitative analysis for CBS, CSE, and 3-MST expression in the myocardium. GAPDH was used as the internal control. (b) Representative double-staining immunofluorescence showing the distribution of CBS (red) in the cardiomyocytes and vessels (labelled by CD31, green) from control or ageing diabetic mice (scale bar = 250
To confirm whether endogenous H2S is involved in the diabetic myocardial injury, NRCMs were incubated in normal glucose (5.5 mmol/L) and high glucose (33 mmol/L) for 72 h to mimic the hyperglycemia in DCM
Reduction of endogenous H2S involves in high glucose-induced myocardial injury. (a) Neonatal rat cardiac ventricular myocytes (NRCMs) viability measured by CCK-8 assay at the end of the treatment for 72 h. (b) Representative images of cardiomyocyte apoptosis detected by a laser confocal microscope at the end of the treatment for 72 h. (c) Quantitative analysis for cardiomyocyte apoptosis determined by flow cytometry. (d) Representative images of ROS levels in NRCMs detected by a laser confocal microscope at the end of the treatment for 72 h. (e) Quantitative analysis for ROS levels in NRCMs. (f) H2S levels in NRCMs at the end of the treatment for 72 h. NG group, normal glucose (5.5 mmol/L); HG group, high glucose (33 mmol/L); Osmotic pressure control group, normal glucose (5.5 mmol/L) + L-glucose (27.5 mmol/L). Results are means ± SEM. A
To determine whether exogenous H2S attenuated high glucose-induced myocardial injury, different concentrations (10, 50, and 100
Exogenous H2S attenuates high glucose-induced myocardial injury. (a) Exogenous NaHS caused transient increase of H2S level in cell culture media within 6 hours.
In this study, we established an ageing diabetic mouse model by feeding mice water with 30% fructose for up to 15 months to investigate any effects of long-term high-fructose feeding on the mouse cardiovascular system. This resulted in two important findings: (1) long-term high-fructose consumption was associated with diabetic cardiomyopathy (DCM) and (2) H2S levels were reduced in the ageing diabetic heart because of alterations in the three H2S-producing enzymes.
Despite recent advances in care and management, diabetes and its associated complications continue to be a major global public health problem, which is gradually worsening, particularly in the developing nations. Although genetic predisposition is an important aetiology of this disease, environmental factors, such as diet and physical activity, are also involved. In particular, long-term consumption of overly nutritious diets that are enriched in fructose and fats can cause initiation of obesity and insulin resistance, which result in development of type 2 diabetes and its associated complications [
Although increased coronary atherosclerosis is the major cause of death among diabetic patients, particularly elderly patients, there is an increased risk for the development of heart failure that is independent of coronary artery disease and hypertension. This adverse situation is referred to DCM, which is characterized by cardiac remodelling, fibrosis, progressive cardiac dysfunction and independent of coronary artery disease [
Animal models in DCM research are quite common. However, the drawback of these models is that they only mimic a short term for DCM but do not mimic it long term. Thus, to better understand the pathogenesis of DCM, a long-term rodent model mimicking as best as possible human DCM would be of great help. In this study, ageing diabetic mice were induced by feeding with a 30% fructose water solution for 15 months (at the end, mice were 17 months old), and these mice were overweight, hyperglycaemic, insulin resistant, and dyslipidemic by the end of the study.
This long-term fructose feeding also caused morphological changes in mouse heart tissue, increased interstitial collagen deposition and expression and increased heart/body weight ratios, indicative of cardiac hypertrophy and fibrosis. The increased Bax/Bcl-2 ratio also indicated cardiomyocyte apoptosis in these mice. M-mode echocardiography confirmed that LVEF and LVFS were significantly reduced along with an increased LV volume, which suggested hyperglycaemia-induced cardiac dilation and dysfunction. In contrast, mice that were fed tap water only for the same period remained healthy. This was consistent with the results of previous reports that showed the developmental stages of cardiomyopathy in db/db diabetic mice [
A number of mechanisms have been proposed to contribute to the development of diabetic cardiomyopathy, including increased oxidative stress [
CSE, a key enzyme involved in H2S production in the cardiovascular system, was downregulated, which might have contributed to the reduced H2S levels. These findings were consistent with those in previous studies. Zhang et al. reported that glucose induced SP1 phosphorylation via p38 MAPK activation, which resulted in decreased CSE promoter activity and the subsequent downregulation of the expression of CSE gene [
As discussed above, there are conflicting reports regarding the regulation of H2S-producing enzymes in diabetes. Several
These conflicting findings on the expression of H2S-producing enzymes may be due to the different responses of different organs and different cell types. This may also depend on the stage or severity of the disease. These questions as well as the actual molecular regulation of these enzymes need to be further investigated. Despite the controversy on the expression of the three H2S-producing enzymes, it appears that endogenous H2S plays an important role in the development of diabetes and its complications.
In conclusion, our results suggest that ageing diabetic mice induced by long-term high-fructose consumption developed diabetic cardiomyopathy and that H2S levels were reduced in the diabetic heart due to alterations in the expression of the three H2S-producing enzymes, which might be involved in the pathogenesis of DCM.
The authors declare that there is no conflict of interests regarding the publication of this paper.
This study was supported by the grants from the Ministry of Science and Technology (2012ZX09501001-001-002) of China, the National Natural Science Foundation of China (81230003, 31300945, and 81402917), and the Research Center on Aging and Medicine, Fudan University (13dz2260700).