Human Molecular Genetics, 2001, Vol. 10, No. 2 145-152
© 2001 Oxford University Press
Abnormalities in the functioning of adipocytes from R6/2 mice that are transgenic for the Huntington’s disease mutation
Departments of 1Molecular Sciences and 2Anatomy and Neurobiology, College of Medicine, University of Tennessee Health Science Center, Memphis, TN 38163, USA
Received 18 September 2000; Revised and Accepted 7 November 2000.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
In an effort to characterize the basis of abnormalities in body weight regulation (i.e. wasting) in Huntington’s disease (HD), we examined adipocytes in a transgenic model of HD, the R6/2 mouse. These mice typically show severe wasting beginning at
12 weeks of age and die between 12 and 15 weeks. Despite an overall growth retardation compared with wild-type littermates, we observed an enhanced accumulation of body fat at 8–9 weeks of age in R6/2 mice fed laboratory chow or a synthetic high fat, high sugar diet. The obesity was not accompanied by symptoms associated with diabetes, as there were no abnormalities in serum glucose, serum insulin or the ability of insulin to stimulate glucose metabolism in epididymal adipose tissue. As expected, the obesity in the high fat, high sugar-fed R6/2 mice was accompanied by increased serum leptin. The ability of insulin to stimulate leptin release from isolated epididymal adipose tissue was also enhanced in R6/2 mice. In contrast, the ability of isoproterenol to inhibit leptin release was reduced in adipose tissue from R6/2 mice, as was the lipolytic effect of isoproterenol. These data suggest that the obesity observed at 8–9 weeks in R6/2 mice may stem from a defect in fat breakdown by adipocytes. |
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
In addition to neurological impairment, cachexia (weight loss) is a prominent characteristic of late-stage Huntington’s disease (HD) (1,2). This cachexia is characterized by below-normal body weight even in the face of high caloric intake. Although the neurological impairment appears attributable to the well documented cortical and striatal neuron loss that occurs in HD (3–8), the basis of the wasting is unclear. At one time it was thought that the hyperkinesia in early-stage HD might account for the weight loss, due to the presumed increased metabolic need accompanying increased activity associated with hyperkinesia. This explanation, however, does not adequately account for the weight loss seen in HD, since such weight loss is seen also in HD patients who are rigid rather than choreic in their movement disorder (1,2).
Although the gene defect in HD has been known for several years, the relationship of the weight loss to the central defect in HD is unknown (9–11). It may be that the gene mutation in HD causes parallel disturbances in the cortex and striatum and in the physiological mechanisms regulating food intake (12). Under these circumstances, the weight loss might contribute to the overall health decline of the HD patients. On the other hand, some authors have suggested that the brain degeneration in HD may stem from metabolic irregularities that are pathogenic because they promote excitotoxicity via bioenergetic failure in neurons (6,13). Under these circumstances, the weight loss could be a manifestation of an underlying metabolic defect that also causes or contributes to the brain neurodegeneration. For these reasons, it is important to more precisely characterize the nature and basis of any disturbances that might contribute to the defects in body weight regulation in HD victims.
The R6/2 transgenic mouse, which bears a transgene coding for exon 1 of the human HD gene, with 141–157 CAG repeats under the control of the human HD gene promoter, shows some of the characteristic features of HD, including movement abnormalities and premature death (14,15). Of present note, these mice show weight loss beginning at 12–13 weeks of age, with death following within a week or two. Because of the resemblance of this wasting phenotype to that in HD, we chose to explore the possible basis of the wasting observed in R6/2 mice. Fat metabolism and leptin release by adipocytes play a major role in the short- and long-term regulation of feeding and body weight and are under the homeostatic control of various humoral and neurally derived factors such as insulin, corticosteroids and noradrenalin (16–18). We therefore examined the influence of these factors on adipocyte response in R6/2 mice, using 8- to 10-week-old mice to avoid confounding effects of the possible multi-system failure that leads to their demise, typically between 12 and 15 weeks of age (14,19). We found that R6/2 mice were slightly obese, and this tendency was exacerbated by a high fat, high sugar diet. Our data suggest that obesity at 8–10 weeks may be attributable to defective fat breakdown by adipocytes.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
In vitro studies of adipose tissue from mice fed laboratory chow
General features of mice.
We initially attempted to compare the metabolism of adipose tissue from male R6/2 mice, fed laboratory chow, at 8–10 weeks of age with male littermate controls also fed laboratory chow. This approach encountered the difficulty that the controls had <0.1 g of epididymal adipose tissue per mouse, whereas the R6/2 mice, despite their lesser body weight and smaller size compared with their age-matched wild-type littermates (Table 1), had
0.4 g of epididymal fat. Since there was not enough adipose tissue in the age-matched wild-type littermates to examine the adipocyte metabolism in vitro, we turned to the use of older male controls, so that we could have the same amount of adipose tissue as in 8- to 10-week-old R6/2 mice (Table 1). Furthermore, using this type of control ruled out the possibility that any effects in the R6/2 mice were a consequence of the greater obesity in these mice. Although the 16- to 32-week-old control mice matched the 8- to 10-week-old R6/2 mice in epididymal fat, the control mice were, nonetheless, larger and heavier than the R6/2 mice (Table 1). Thus, the R6/2 mice possessed a higher percentage of body fat than these older control mice (Table 1). Consistent with our effort to fat-match the R6/2 mice with wild-type controls, the serum leptin values for the male R6/2 mice fed laboratory chow were not significantly different than those in the fat-matched wild-type controls (Table 1).
|
Leptin release evoked by dexamethasone from in vitro adipocytes.
We first examined the release of leptin from adipose tissue in vitro from control mice over a 24 h incubation in the presence of 25 or 200 nM dexamethasone (Fig. 1). We used dexamethasone since it is a synthetic glucocorticoid stimulator of leptin release and it allows the initial rate of leptin release to be maintained over a 24 h incubation (20–23). Leptin release from control adipocytes was found to be enhanced by dexamethasone, particularly by the 200 nM concentration (Fig. 1). The enhanced release of leptin seen with a dexamethasone concentration of 200 nM compared with the 25 nM concentration was primarily seen during the last 18 h of the incubation. The effects of 25 or 200 nM dexamethasone on the release of leptin by adipose tissue from R6/2 mice fed the laboratory chow was not significantly different from the effects in the wild-type cells (Table 1). All subsequent in vitro studies examining the effects of additional agents (i.e. insulin or isoproterenol) on adipocyte behavior were done in the presence of 200 nM dexamethasone to ensure a high baseline level of leptin release for assessing the effects of other drugs on leptin release.
|
Leptin release evoked by insulin or isoproterenol.
There were statistically significant differences between control and R6/2 mice in their response to both insulin and isoproterenol, a specific ß1 adrenergic agonist at low concentrations (24). The release of leptin evoked by 10 nM insulin was considerably greater in adipose tissue from 8- to 10-week-old R6/2 mice fed laboratory chow than in tissue from 16- to 32-week-old control mice fed the same diet (Table 2).For controls, the increase with insulin was 146% of baseline release and for the R6/2 adipose tissue the increase with insulin was 284% of baseline release. Furthermore, when adipose tissue was incubated with 10 nM isoproterenol in the presence of 200 nM dexamethasone, the inhibition of leptin release caused by isoproterenol was significantly less for R6/2 adipocytes than for the adipocytes from fat-matched control mice. For the controls, leptin release in the presence of dexamethasone and isoproterenol was 23% of that in the presence of dexamethasone alone, whereas for the R6/2 adipocytes the leptin release in the presence of dexamethasone and isoproterenol was 70% of that in the presence of dexamethasone alone (Table 2). As a consequence, adipocyte leptin release in the combined presence of dexamethasone and isoproterenol was much greater for the R6/2 than for fat-matched control mice.
|
Lipolysis evoked by isoproterenol.
Fat breakdown in the presence of isoproterenol was defective in the R6/2 adipose tissue from animals on a regular laboratory chow diet. The defect in fat breakdown by adipose tissue from R6/2 mice fed laboratory chow was evident as a significantly lesser lipolytic response (at least at 6 h) to the addition of 10 nM isoproterenol (in the presence of dexamethasone) than by adipose tissue from fat-matched animals on the same diet (Table 2). In addition, isoproterenol added in the combined presence of dexamethasone and insulin evoked a lipolytic response from the fat-matched wild-type adipose tissue, but failed to elicit such a response in adipose tissue from R6/2 mice (Fig. 2).
|
In vitro studies of adipose tissue from mice fed high fat, high sugar diet
General features of mice.
We next examined the effect of feeding male R6/2 mice a high fat, high sugar diet. A marked increase in epididymal adipose tissue weight was found in 8- to 10-week-old R6/2 mice after 3–4 weeks on the high fat, high sugar diet (Table 1). There was a 112% increase in epididymal adipose tissue content of male R6/2 mice compared with wild-type littermate control mice fed the same diet (Table 1). The doubling of epididymal fat content is readily evident in Figure 3B, which shows the appearance of the abdominal fat in an R6/2 mouse. These data indicate that the R6/2 mice develop obesity to a far greater extent than age-matched control mice of the same genetic background when fed a high fat, high calorie diet for only 3–4 weeks. Although fatter, the R6/2 mice were, nonetheless, shorter in length than littermate controls (Fig. 3A). Note that in separate experiments on R6/2 mice fed this enriched diet, we sought to determine whether the diet would extend the lifespan of these animals. We found that at 12–13 weeks of age, R6/2 mice that had been fed this enriched diet beginning at a mean age of 44.2 days (n = 6), began to show the typical wasting and morbidity that R6/2 mice begin to show at this age (14). They were sacrificed before they succumbed to the morbidity, at a mean age of 89 days. A cohort of R6/2 mice (n = 10) raised on laboratory chow and born at the same general time as these enriched-diet R6/2 mice showed similar morbidity when sacrificed at a comparable age (89 days). Thus, the high fat, high sugar diet did not extend R6/2 lifespan and the excess fat was lost by 89 days.
|
The serum leptin values were elevated 2-fold in the male R6/2 mice fed the high fat, high sugar diet compared with serum leptin levels in the age-matched littermates fed the same enriched diet (Table 1). The enhanced levels of serum leptin might, however, be expected, in light of the greater obesity of R6/2 mice and the findings that serum leptin levels tend to correlate with degree of obesity (16,18,25).
We also examined the effect of the high fat, high sugar diet in female R6/2 mice. The serum leptin levels were 20 ± 10 ng/ml (mean and range of values for two 9- to 10-week-old R6/2 mice) after 4 weeks on the high fat, high sugar diet, compared with 4 ng/ml in an age-matched wild-type littermate female mouse on the same diet. The amount of parametrial adipose tissue in the two female R6/2 mice on the enriched diet was 1.8 ± 0.6 g/mouse, compared with 0.5 g in the age-matched wild-type littermate female mouse on the same diet. Thus, female R6/2 mice also more readily became obese than age-matched littermate wild-type female mice.
Leptin release evoked by dexamethasone from in vitro adipose tissue.
The total release of leptin by epididymal fat pads from enriched diet-fed R6/2 mice in the presence of 200 nM dexamethasone over a 24 h incubation was significantly greater than that for the fat pads from wild-type control mice fed the same diet (Table 1). The elevated level of leptin release by the incubated fat pads and the elevated serum leptin in the R6/2 mice indicate that the obesity of these mice is not caused by a lack of leptin release, as is obesity in ob/ob mice (16).
Epididymal adipocyte diameters
We found that the average cross-sectional area of individual fat cells in 3-month-old wild-type mice fed laboratory chow (n = 3; mean age at death, 91 days) was 1070 ± 77 µm2. In laboratory chow-fed R6/2 mice (n = 3; mean age at death, 81 days), the mean individual fat cell area was 3218 ± 161 µm2, which is about three times that in the wild-type control mice fed the same diet. In high fat, high sugar-fed R6/2 mice that had been placed on the enriched diet beginning at a mean age of 54 days (n = 3; mean age at death, 89 days), the mean area of individual fat cells was 5767 ± 372 µm2.
Evidence against diabetes in R6/2 mice
While our studies were in progress, it was reported that R6/2 mice at 7 weeks of age had blood glucose values of 226 mg/dl, in contrast to age-matched controls with blood glucose values of 164 mg/dl (26). However, in male R6/2 mice from our colony, the fed blood glucose value at 5 weeks of age was 133 ± 6 mg/dl (mean ± SEM; n = 7). Even after 2–3 weeks on the high fat, high sugar diet, the blood glucose values in these same mice was only 150 ± 10 mg/dl. These values were comparable to those of littermate controls fed ad libitum on the high fat, high sugar diet, whose mean blood glucose level at 5 weeks of age was 142 ± 7 and whose mean blood glucose level after 2–3 subsequent weeks was 152 ± 9 mg/dl (mean ± SEM of four mice). In two female R6/2 mice on the high fat, high sugar diet for 4 weeks, the blood glucose value was 114 ± 28 mg/dl (mean ± SEM) at 8–10 weeks of age, whereas in four age-matched female control mice the blood glucose value was 124 ± 8 mg/dl (mean ± SEM).
Additionally, we found that in adipose tissue from R6/2 mice fed the laboratory chow diet, the total conversion of glucose to lactate, which is the major metabolic product of glucose metabolism in fat from obese rodents (27), was no different than in wild-type mice fed the same diet (Table 2). Furthermore, the stimulation of glucose conversion to lactate in response to 10 nM insulin was not significantly different for the adipocytes of R6/2 and wild-type mice on the laboratory chow diet (Table 2). Thus, glucose metabolism appeared normal in the R6/2 adipocytes from the laboratory chow-fed mice. Similarly, in adipose tissue from R6/2 mice fed the high fat, high sugar diet, the total conversion of glucose to lactate was no different per fat pad than in wild-type mice fed the same diet (Table 3). Furthermore, the stimulation of glucose conversion to lactate in response to 10 nM insulin was not significantly different for adipose tissue from R6/2 mice fed the high fat, high sugar diet than for wild-type mice fed the same diet (Table 3). Thus, glucose metabolism by adipose tissue appeared normal even in R6/2 mice fed the enriched diet.
|
It is clear that our R6/2 mice had no abnormalities in the regulation of blood glucose or glucose metabolism by adipocytes, even after 2–3 weeks on the high fat diet. Our results are comparable to those of Carter et al. (19) in this regard. We also measured the serum insulin in the mice fed the high fat, high sugar diet for 3–4 weeks, whose adipose tissue was utilized for the studies shown in Table 1. The mean serum insulin in the control mice was 1.9 ng/ml and the range of values was 0.7–4.0 ng/ml. The mean serum insulin value for five of the six R6/2 mice was 1.7 ng/ml and the range was 0.4–4.2 ng/ml, whereas the sixth R6/2 mouse had a serum insulin value of 13.2 ng/ml. Thus, this one mouse may have had some degree of insulin resistance, which was compensated for by an increase in serum insulin.
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
The present results suggest that among the changes seen in the HD-like R6/2 mice is a greater propensity for fat accumulation. The deposition of fat in R6/2 mice is especially pronounced in mice fed a synthetic diet high in fat and sugar. The enhanced deposition of fat seen in mice fed the high fat, high sugar diet does not, however, prevent the subsequent cachexia and morbidity seen at 12–15 weeks in our colony, the typical age of death for R6/2 mice (14). Accompanying the increase in fat stores in the R6/2 mice is an increase in leptin release and serum leptin. Thus, although obesity in some mutant mouse strains (such as ob/ob) stems from the failure to release leptin from adipocytes (19), the obesity in R6/2 mice does not appear to be related to any deficiency in leptin release by adipose tissue. Rather, high leptin release is observed from R6/2 adipocytes, as occurs for normal adipocytes rich in fat stores as part of their role in the lipostatic regulation of body weight (18,19).
Surprisingly, the elevated serum leptin in the high fat, high sugar-fed R6/2 mice does not curtail the fat accumulation. In this regard, the R6/2 mice resemble db/db mice (16) and possibly some types of obese human (28). The db/db mice become obese despite high serum leptin levels, because of an inactivating mutation in the leptin receptor gene (16). Whereas we cannot rule out the possibility that the R6/2 transgene affects leptin receptor levels and, thereby, promotes fat accumulation in these mice, our findings suggest that the fat accumulation, at least in part, stems from a defect in lipolysis by adipocytes. Our histological measurements show that adipocyte size is increased in R6/2 mice, especially those fed the enriched diet. This further confirms the increased fat accumulation in R6/2 mice and is consistent with enhanced fat deposition and/or defective fat breakdown in R6/2 adipocytes, which would thereby favor fat accumulation. Whether elevated serum leptin levels in R6/2 mice, in fact, curtail appetite and thereby diminish growth (thus accounting for the smaller lengths of R6/2 mice) is uncertain and requires monitoring of food intake in young R6/2 mice (<10 weeks of age). It may also be the case that the R6/2 transgene affects growth by acting on hypothalamic and/or hypophyseal mechanisms of feeding or growth regulation (17).
We found that ß-adrenergic agonists are less effective than normal in stimulating lipolysis and decreasing leptin release by adipocytes in R6/2 mice. The basis of the decreased response to the ß-adrenergic agonist isoproterenol in R6/2 adipocytes is uncertain. The change could stem from a reduction in the number of ß1 catecholamine receptors on these cells, or it could stem from changes in postreceptoral mechanisms. In any case, these data again suggest that defective lipolysis by adipocytes and impaired regulatory control of lipolysis are likely contributors to the obesity of the R6/2 mice.
The leptin response and the lipolytic response of the R6/2 adipocytes to isoproterenol is of potential interest to the end-stage cachexia shown by R6/2 mice. Adrenergic activation of adipocytes plays a role in the adaptive response to starvation and such activation promotes feeding by decreasing leptin release and promotes energy mobilization by increasing breakdown of stored fat (18,29). We have shown that both adrenergic inhibition of leptin release and adrenergic stimulation of energy mobilization from stored fat are defective in R6/2 mice. Thus, once R6/2 mice begin to show the motor disturbances that may make feeding more difficult (14), the defect in the adipocyte response to starvation in R6/2 mice, particularly the failure to shut down leptin release, may hasten their wasting and decline. The end-stage cachexia in R6/2 mice could also, however, stem from or receive contributions from currently undiscovered disturbances in feeding regulatory mechanisms distinct from adipocyte behavior. For example, dysfunction could occur in hypothalamic regions involved in appetite regulation (17).
Diabetes and R6/2 mice
The R6/2 mice in our study developed obesity without diabetes, as evidenced by their failure to show hyperglycemia. Of particular note, in the R6/2 mice fed the high fat, high sugar diet there were no signs of diabetes, since the blood glucose and insulin values were comparable with those of controls and the ability of insulin to stimulate glucose conversion to lactate by adipose tissue was normal. Our findings are similar to those of Carter et al. (19), who found elevated blood glucose values in only two of eight R6/2 mice at 13 weeks. In contrast, Jenkins et al. (30) found that in mice deprived of food for 7 h, the blood glucose values for 9 of 11 R6/2 mice were >200 mg/dl. Hurlbert et al. (26) also reported a marked hyperglycemia and loss of pancreatic insulin in R6/2 mice (26). The prior studies reporting diabetes in R6/2 mice have used shipped R6/2 mice and non-littermate controls (26,30). In contrast, our study and that of Carter et al. (19) used colony-bred R6/2 mice and littermate controls. These differences may have contributed to the disparity. It is also possible that our R6/2 mice and theirs differed in some unknown gene or genes that favored development of diabetes in their R6/2 mice. In any case, our findings and those of Carter et al. (19) indicate that the death of R6/2 mice by 13–15 weeks of age is not necessarily a complication of diabetes engendered by the R6/2 transgene.
Implications for humans and HD
The basis of the cachexia in late-stage HD is uncertain (1,2). It is clear that the hyperkinesia associated with HD does not account for the cachexia, since such weight loss is seen even in HD patients who are rigid, rather than choreic, in their movement disorder (1,2). It may be that the gene mutation in HD causes a disturbance in one or more of the diverse central or peripheral physiological mechanisms regulating food intake (12). For this reason, our current findings in R6/2 mice are of interest for the light they shed on possible defects in body weight regulation in HD victims. The R6/2 transgenic mouse bears a transgene coding for exon 1 of the human HD gene with 141–157 CAG repeats under the control of the human HD gene promoter. The mouse resembles HD victims in some respects, including that R6/2 mice show a progressive neurological decline culminating in premature death, that neurons in the R6/2 forebrain contain ubiquitinated nuclear accumulations of the mutant huntingtin protein fragment and that R6/2 mice show wasting in the period when the movement disorder is manifest (14,15,31,32). Such similarities encourage the belief that findings from R6/2 mice may have pertinence to HD pathogenic mechanisms. Nonetheless, the prominent differences between the R6/2 mice and HD victims, which include the much longer CAG repeat size in the mice than in the human disease, the extreme rapidity with which death occurs in the mice and the absence of striatal pathology in the mice (14), suggest the use of caution in generalizing from R6/2 mice to human HD mechanisms.
With such pertinence as well as possible caveats in mind, our current findings are of interest for the weight loss in HD. We proposed above that the tendency of R6/2 adipocytes to release abnormally high levels of leptin and respond inadequately to starvation signals could contribute to wasting in R6/2 mice. If similar adipocyte defects occur in human HD victims, they might contribute to the cachexia characteristic of this disease. Our finding of slightly enhanced fat accumulation in young R6/2 mice seems surprising in light of the wasting reported to be characteristic of HD. Nonetheless, it may be the case that a similar phenomenon to that observed in young R6/2 mice occurs early in HD. Finally, it is uncertain whether the defective mobilization of energy from fat stores occurs only in adipocytes. If the defect is systemic, then a similar defect may also be present in HD victims, alone or in combination with other metabolic defects, and may possibly contribute to the cerebral hypometabolism observed in HD victims (33–36), which some have suggested could contribute to the neurodegeneration in this disorder (13,36,37). Even if adipocyte defects occur in HD victims and contribute to their defective weight regulation, the possibility remains that the HD mutation affects other regulatory systems, including those in the hypothalamus involved in feeding and weight regulation (12,17). Pratley et al. (38) have recently shown that there is no intrinsic metabolic defect that contributes to cachexia in patients with HD. Nonetheless, our present findings for R6/2 mice suggest that studies of HD adipocytes might be of interest for the information they provide on the basis of the weight regulation defects in HD.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
Subjects
Heterozygous R6/2 breeder mice for establishment of a colony were originally obtained from the Jackson Laboratory, where they are maintained on a (CBA x C57B6) F1 hybrid background (B6CBAF1). Our colony of R6/2 mice at the University of Tennessee in Memphis is maintained on the same hybrid background. R6/2 mice were produced by the mating of B6CBAF1 females to heterozygous R6/2 males or by the mating of B6CBAF1 males to females that carried transplanted R6/2 ovaries. For all experiments reported here, the R6/2 mice used were heterozygous for the TgN(HDexon1)62Gpb transgene. Genotype was confirmed by PCR, based on a modification of the procedure described in Mangiarini et al. (14). Genomic DNA extracted from tail biopsies was used to identify mice bearing the R6/2 transgene. The primers for the detection of the R6/2 transgene were 5'-CGGCTGAGGCAGCAGCGGCTGT-3' and 5'-GCAGCAGCAGCAGCAACAGCCGCCACCGCC-3'. The amplified huntingtin DNA fragment has a size of 120 bp. One microliter of DNA template (275 ng/µl) was used and the PCR was run in thin-walled PCR tubes. The PCR reaction solution contained the following: 5.0 µl of 10x PCR buffer, 3.0 µl of 25 mM MgCl2, 1.0 µl of 10 mM dNTPs, 5.0 µl of 20 µM of each two primers, 6.9 µl of DNA dye, 18.9 µl of H2O, 5.0 µl of DMSO and 0.2 µl of Taq polymerase. Contamination from extraneous DNA was checked by replacing the cellular template with water. Amplification was performed on a thermal cycler (MJ Research), typically under the following cycle conditions: denaturation at 94°C for 30 s, annealing at 65°C for 30 s and extension at 72°C for 90 s for a total of 30 cycles. Following PCR amplification, aliquots of reaction product were analyzed by electrophoresis on ethidium bromide impregnated 1.5% agarose gels.
Experimental groups for in vitro studies of adipocytes
Adipose tissue from two separate groups of R6/2 mice was studied in vitro in comparison with adipose tissue from comparable groups of wild-type mice. In one R6/2 group, the behavior of adipose tissue from 8- to 10-week-old R6/2 mice on a normal laboratory chow diet was examined and compared with that of adipose tissue from a group of wild-type mice fed the same diet. Adipose tissue from the epididymal fat pads was utilized for these studies. We used 16- to 32-week-old male B6CBAF1 mice as the wild-type control group for this line of study because 8- to 10-week-old wild-type littermates of the R6/2 mice possessed extremely meager epididymal fat pads which yielded insufficient fat for the in vitro studies (Table 1). In contrast, 16- to 32-week-old male B6CBAF1 mice possessed epididymal fat pads equal in size to those of the 8- to 10-week-old R6/2 mice (Table 1). This allowed us to compare the behavior of adipose tissue in R6/2 mice with that in wild-type mice, without any confounding effects of differences in the average amount of lipid stored in individual fat cells.
In the second set of in vitro studies, the behavior of adipose tissue from a group of 8- to 10-week-old R6/2 mice fed a high fat, high sugar diet was examined and compared with that of adipose tissue from a group of age-matched wild-type littermates fed the same diet. The pelleted high fat, high sugar diet contained 27% casein, 20% Crisco, 46% sucrose, 2% Ralston-Purina (RP) vitamin mix and 5% RP mineral mix number 10 (Purina Mills) and was fed to mice for 3–4 weeks prior to harvesting of the epididymal fat pads at 8–10 weeks. Because of the high fat, high sugar diet, the epididymal fat pads in the wild-type mass were ample enough to provide sufficient adipose tissue for study.
In vitro studies with adipose tissue
The epididymal adipose tissue was removed from each mouse and weighed prior to being cut with scissors into small pieces (5–10 mg each). In the studies using mice fed laboratory chow, 18 control and 18 R6/2 mice were examined in six experiments. In each experiment, the adipose tissue was pooled from sets of three R6/2 mice or three control mice (so as to provide adequate tissue per assay). A total of 50–150 mg of pooled adipose tissue from the three R6/2 or control mice was then placed in a given 50 ml plastic tube containing 5 ml of incubation buffer. In the studies using mice fed the high fat, high sugar diet, six R6/2 mice and six control mice were examined. In these studies, each of the six experiments utilized cut tissue from either an individual R6/2 or an individual wild-type mouse. The amount of tissue per flask was 130 mg in the studies using tissue from the R6/2 mice on the high fat, high sugar diet, whereas the amount of tissue per flask averaged 60 mg for the control mice fed the same diet.
The buffer for incubation of adipose tissue was Dulbecco’s modified Eagle’s medium/Ham’s F12 (1:1; Sigma) containing 17.5 mM glucose, 121 mM NaCl, 4 mM KCl, 1 mM CaCl2, 25 mM HEPES, 2.4 mM sodium bicarbonate, 40 mg/ml bovine serum albumin (Bovuminar; lot L59410; Intergen), 5 µg/ml ethanolamine, 0.2 ng/ml sodium selenite, 90 µg/ml penicillin G, 150 µg/ml streptomycin sulfate, 50 µg/ml gentamicin, 55 µM ascorbic acid, 1 µg/ml leupeptin and 1 µg/ml aprotinin. The pH of the buffer was adjusted to 7.4 and then filtered through a 0.2 µm filter. Pharmacological agents were added at the start of the incubation to examine the behavior of the fat cells in response to various regulatory signals. These included the steroid dexamethasone (which normally exerts an anti-lipolytic effect and increases leptin release) (20,21,23,39), insulin (which normally increases leptin release and increases fat deposition) (16,18,39,40) and isoproterenol (which normally mobilizes a compensatory response of fat cells to starvation conditions by exerting a lipolytic effect while decreasing leptin release) (18,29). The incubations were carried out under sterile conditions with the tubes maintained in an upright position in a gyratory water bath shaker at 37°C for 24 h at 100 r.p.m. Aliquots of the medium were taken at 24 h and stored at –20°C for measurement of lactate, leptin or glycerol. The leptin content of 20–50 µl aliquots of the incubation medium was determined using radioimmunoassay kits from Linco Research. Lipolysis was based on analysis of glycerol release into the medium and determined on 20–50 µl aliquots by the procedure of Boobis and Maughan (41). Lactate was determined by the same procedure as for glycerol using lactate dehydrogenase. Statistical comparisons were made by applying Student’s t-test to the paired differences.
In vivo assays
Blood glucose levels were taken using the One-Touch system (LifeScan). All blood samples for glucose measurement were taken early in the morning by a single cut of the mouse tail from mice fed ad libitum. Serum leptin and insulin levels were determined from trunk blood taken at the time of sacrifice. Radioimmunoassay for plasma leptin determination was carried out as noted above and insulin determination was also carried out using radioimmunoassay kits from Linco Research.
Histological studies of adipocyte diameters
In an additional set of studies, we used histological methods to measure the diameter of adipocytes in the epididymal fat pads of R6/2 mice fed laboratory chow, wild-type mice fed laboratory chow or R6/2 mice fed the high fat, high calorie diet. The goal of these studies was to determine whether the greater weight of the fat pads in the R6/2 mice stemmed from increased lipid storage in individual adipocytes, as reflected by an increase in their cross-sectional area. Under deep Avertin anesthesia, mice to be used for histological analysis were perfused transcardially with phosphate-buffered saline (0.1 M sodium phosphate buffer at pH 7.4 with 0.9% NaCl) followed by 4% paraformaldehyde, 0.1 M lysine and 0.1 M sodium periodate in 0.1 M PBS. The epididymal fat pads were removed and stored in a 20% sucrose solution at 4°C and later sectioned at 20 µm on a cryostat, with the sections mounted on Superfrost+ glass slides. Sections were stained with hematoxylin and eosin to reveal cellular boundaries. Cell diameters were measured using the program NIH Image (version 1.57) from captured images.
|
ACKNOWLEDGEMENTS |
|---|
The authors wish to thank Lisa Pouncey and Lydia Hu for their skilled technical assistance. This work was supported by the Harriett S. Van Vleet Chair of Excellence in Biochemistry (J.N.F.), the Hereditary Disease Foundation (D.G. and A.R.) and NS-28721 (A.R.).
|
FOOTNOTES |
|---|
+ To whom correspondence should be addressed. Tel: +1 901 448 4343; Fax: +1 901 448 7360; Email: jfain@utmem.edu
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
1 Bruyn, G.W. and Went, L.N. (1986) Huntington’s chorea. In Vinken, P.J., Bruyn, G.W. and Klawans, H.L. (eds), Handbook of Clinical Neurology 2. Elsevier, Amsterdam, The Netherlands, Vol. V, pp. 267–313.
2 Greenamyre, J.T. and Shoulson, I. (1994) Huntington’s disease. In Calne, D.B. (ed.), Neurodegenerative Diseases. W.B. Saunders, Philadelphia, PA, pp. 685–704.
3 Kowall, N.W., Ferrante, R.J. and Martin, J.B. (1987) Pattern of cell loss in Huntington’s disease. Trends Neurosci., 10, 24–29.
4 De La Monte, S.M., Vonsattel, J.P. and Richardson Jr, E.P. (1988) Morphometric demonstrations of atrophic changes in the cerebral cortex, white matter, and neostriatum in Huntington’s disease. J. Neuropathol. Exp. Neurol., 44, 516–525.
5 Reiner, A., Albin, R.L., Anderson, K.D., D’Amato, C.J., Penney, J.B. and Young, A.B. (1988) Differential loss of striatal projection neurons in Huntington’s disease. Proc. Natl Acad. Sci. USA, 85, 5733–5737.
6 Albin, R.L., Reiner, A., Anderson, K.D., Penney, J.B. and Young, A.B. (1990) Striatal and nigral neuron subpopulations in rigid Huntington’s disease: implications for the functional anatomy of chorea and rigidityakinesia. Ann. Neurol., 27, 357–365.[ISI][Medline]
7 Hedreen, J.C., Peyser, C.E., Folstein, S.E. and Ross, C.A. (1991) Neuronal loss in layers V and VI of cerebral cortex in Huntington’s disease. Neurosci. Lett., 133, 257–261.[ISI][Medline]
8 Vonsattel, J.P.G. and DiFiglia, M. (1998) Huntington’s disease. J. Neuropath. Exp. Neurol., 57, 369–384.[ISI][Medline]
9 Huntington’s Disease Collaborative Research Group (1993) A novel gene containing a trinucleotide repeat that is expanded and unstable on HD chromosome. Cell, 72, 971–983.[ISI][Medline]
10 Albin, R.L. and Tagle, D.A. (1995) Genetics and molecular biology of Huntington’s disease. Trends Neurosci., 18, 11–14.[ISI][Medline]
11 Sharp, A.H. and Ross, C.A. (1996) Neurobiology of Huntington’s disease. Neurobiol. Dis., 3, 3–15.[ISI][Medline]
12 Kremer, H.P. and Roos, R.A. (1992) Weight loss in Huntington’s disease. Arch. Neurol., 49, 349.
13 Beal, M.F. (1992) Does impairment of energy metabolism result in excitotoxic neuronal death in neurodegenerative illnesses? Ann. Neurol., 31, 119–130.[ISI][Medline]
14 Mangiarini, L., Sathasivam, K., Seller, M., Cozens, B., Harper, A., Hetherington, C., Lawton, M., Trottier, Y., Lebrach, H., Davies, S.W. and Bates, G.P. (1996) Exon 1 of the HD gene with an expanded CAG repeat is sufficient to cause a progressive neurological phenotype in transgenic mice. Cell, 87, 493–506.[ISI][Medline]
15 Sathasivam, K., Bobbs, C., Mangiarini, L., Mahal, A., Turmaine, M., Doherty, P., Davies, S.W and Bates, G.P. (1999) Transgenic models of Huntington’s disease. Philos. Trans. R. Soc. Lond. B Biol. Sci., 354, 963–969.[ISI][Medline]
16 Friedman, J.M. and Halaas, J.L. (1998) Leptin and the regulation of body weight in mammals. Nature, 395, 763–770.[Medline]
17 Elmquist, J.K., Elias, C.F. and Saper, C.B. (1999) From lesions to leptin: Hypothalamic control of food intake and body weight. Neuron, 22, 221–232.[ISI][Medline]
18 Himms-Hagen, J. (1999) Physiological roles of the leptin endocrine system: differences between mice and humans. Crit. Rev. Clin. Lab. Sci., 36, 575–655.[ISI][Medline]
19 Carter, R.J., Lione, L.A., Humby, T., Mangiarini, L., Mahal, A., Bates, G.P., Dunnett, S.B. and Morton, A.J. (1999) Characterization of progressive motor deficits in mice transgenic for the human Huntington’s disease mutation. J. Neurosci., 19, 3248–3257.
20 Slieker, L.J., Sloop, K.W., Surface, P.L., Kriauciunas, LaQuier, F., Manetta, J., Bue-Valleskey, J., Stephens, T.W. (1996) Regulation of expression of ob mRNA and protein by glucocorticoids and cAMP. J. Biol. Chem., 271, 5301–5304.
21 Murakami, T., Iida, M. and Shima, K. (1995) Dexamethasone regulates obese expression in isolated rat adipocytes. Biochem. Biophys. Res. Commun., 214, 1260–1267.[ISI][Medline]
22 Fain, J.N. and Bahouth, S.W. (1998) Effect of tri-iodothyronine on leptin release and leptin mRNA accumulation in rat adipose tissue. Biochem. J., 332, 361–366.
23 De Vos, P., Lefebvre, A.M., Shrivo, I., Fruchart, J.C. and Auwerx, J. (1998) Glucocorticoids induce the expression of the leptin gene through a non-classical mechanism of transcriptional activation. Eur. J. Biochem., 253, 619–626.[ISI][Medline]
24 Van Liefde, I., Van Witzenburg, A. and Vauquelin, G. (1992) Multiple beta adrenergic receptor subclasses mediate the L-isoproterenol-induced lipolytic response in rat adipocytes. J. Pharmacol. Exp. Ther., 262, 552–558.
25 Caro, J.F., Sinha, M.K., Kolaczynski, J.W., Zhang, P.L. and Considine, R.V. (1996) Leptin: the tale of an obesity gene. Diabetes, 45, 1455–1461.[ISI][Medline]
26 Hurlbert, M.S., Zhou, W., Wasmeier, C., Kaddis, F.G., Hutton, J.C. and Freed, C.R. (1999) Mice transgenic for expanded CAG repeat in the Huntington’s Disease gene develop diabetes. Diabetes, 45, 649–651.
27 DiGorolamo, M., Newby, D.F. and Lovejoy, J. (1992) Lactate production in adipose tissue: a regulated function with extra-adipose implications. FASEB J., 6, 2405–2412.[Abstract]
28 Fox, C.S., Esparza, J., Nicolson, M., Bennett, P.H., Schultz, L.O., Valencia, M.E. and Ravussin, E. (1998) Is a low leptin concentration, a low resting metabolic rate, or both the expression of the ‘thrifty genotype’? Results from Mexican Pima Indians. Am. J. Clin. Nutr., 68, 1053–1057.[Abstract]
29 Trayhurn, P., Hoggard, N., Mercer, J.G. and Rayner, D.V. (1999) Leptin: fundamental aspects. Int. J. Obes. Relat. Metab. Disord., 23, 22–28.
30 Jenkins, B.G., Klivenyi, P., Kustermann, E., Andreassen, O.A., Ferrante, R.J., Rosen, B.R. and Beal, M.F. (2000) Nonlinear decrease over time in N-acetyl aspartate levels in absence of neuronal loss and increases in glutamine and glucose in transgenic Huntington’s disease mice. J. Neurochem., 74, 2108–2119.[ISI][Medline]
31 Davies, S.W., Turmaine, M., Cozens, B.A., DiFiglia, M., Sharp, A.H., Ross, C.A., Scherzinger, E., Wanker, E.E., Mangiarini, L. and Bates, G.P. (1997) Formation of neuronal intranuclear inclusions underlies the neurological dysfunction in mice transgenic for the HD mutation. Cell, 90, 537–548.[ISI][Medline]
32 DiFiglia, M., Sapp, E., Chase, K.O., Davies, S.W., Bates, G.P., Vonsattel, J.P. and Aronin, J.P. (1997) Aggregation of huntington in neuronal intranuclear inclusions and dystrophic neurites in brain. Science, 277, 1990–1993.
33 Grafton, S.T., Mazziotta, J.C., Pahl, J.J., St George-Hyslop, P., Haines, J.L., Gusella, J., Hoffman, J.M., Baxter, L.R. and Phelps, M.E. (1992) Serial changes of cerebral glucose metabolism and caudate size in persons at risk for Huntington’s disease. Ann. Neurol., 28, 614–621.
34 Kuwert, T., Lange, H.W., Boeker, H., Titz, H., Herzog, H., Aulich, A., Wang, B.C, Nayak, U. and Feinendegen, L.E. (1993) Striatal glucose consumption in chorea-free subjects at risk of Huntington’s disease. J. Neurol., 241, 31–36.[ISI][Medline]
35 Sax, D.S., Powsner, R., Kimn, A., Tilak, S., Bhatia, R., Cupples, L.A. and Myers, R.H. (1996) Evidence of cortical metabolic dysfunction in early Huntington’s disease by single-photon-emission computed tomography. Mov. Disord., 11, 671–677.[ISI][Medline]
36 Browne, S.E., Bowling, A.C., MacGarvey, U., Baik, M.J., Berger, S.C., Muquit, M.M.K., Bird, E.D. and Beal, M.F. (1997) Oxidative damage and metabolic dysfunction in Huntington’s disease: selective vulnerability of the basal ganglia. Ann. Neurol., 41, 646–653.[ISI][Medline]
37 Albin, R.L. and Greenamyre, J.T. (1992) Alternative excitotoxic hypotheses. Neurology, 42, 733–738.
38 Pratley, R.E., Salbe, A.D., Ravussin, E. and Caviness, J.N. (2000) Higher sedentary energy expenditure in patients with Huntington’s disease. Ann. Neurol., 47, 64–70.[ISI][Medline]
39 Fain, J.N. and Bahouth, S.W. (2000) Regulation of leptin release by mammalian adipose tissue. Biochem. Biophys. Res. Commun., 274, 571–575.[ISI][Medline]
40 Woods, S.C., Seeley, R.J., Porte Jr, D. and Schwartz, M.W. (1998) Signals that regulate food intake and energy homeostasis. Science, 280, 1378–1382.
41 Boobis, L.H. and Maughan, R.L. (1983) A simple one-step enzymatic fluorometric method for the determination of glycerol in 20 µl of plasma. Clin. Chim. Acta, 132, 173–179.[ISI][Medline]
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  A. Fergani, H. Oudart, J.-L. Gonzalez De Aguilar, B. Fricker, F. Rene, J.-F. Hocquette, V. Meininger, L. Dupuis, and J.-P. Loeffler Increased peripheral lipid clearance in an animal model of amyotrophic lateral sclerosis J. Lipid Res., July 1, 2007; 48(7): 1571 - 1580. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Bjorkqvist, A. Petersen, K. Bacos, J. Isaacs, P. Norlen, J. Gil, N. Popovic, F. Sundler, G. P. Bates, S. J. Tabrizi, et al. Progressive alterations in the hypothalamic-pituitary-adrenal axis in the R6/2 transgenic mouse model of Huntington's disease Hum. Mol. Genet., May 15, 2006; 15(10): 1713 - 1721. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. M. Van Raamsdonk, W. T. Gibson, J. Pearson, Z. Murphy, G. Lu, B. R. Leavitt, and M. R. Hayden Body weight is modulated by levels of full-length Huntingtin Hum. Mol. Genet., May 1, 2006; 15(9): 1513 - 1523. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. Luthi-Carter, S. A. Hanson, A. D. Strand, D. A. Bergstrom, W. Chun, N. L. Peters, A. M. Woods, E. Y. Chan, C. Kooperberg, D. Krainc, et al. Dysregulation of gene expression in the R6/2 model of polyglutamine disease: parallel changes in muscle and brain Hum. Mol. Genet., August 15, 2002; 11(17): 1911 - 1926. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Sipione, D. Rigamonti, M. Valenza, C. Zuccato, L. Conti, J. Pritchard, C. Kooperberg, J. M. Olson, and E. Cattaneo Early transcriptional profiles in huntingtin-inducible striatal cells by microarray analyses Hum. Mol. Genet., August 15, 2002; 11(17): 1953 - 1965. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. P. Mattson, S. L. Chan, and W. Duan Modification of Brain Aging and Neurodegenerative Disorders by Genes, Diet, and Behavior Physiol Rev, July 1, 2002; 82(3): 637 - 672. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||