Insulin secretion and carbohydrate metabolism in the dystrophic mouse

European Journal of Pharmacology, 53 (1979) 365--373

365

© Elsevier/North-Holland Biomedical Press

INSULIN SECRETION AND CARBOHYDRATE METABOLISM IN THE DYSTROPHIC MOUSE INGMAR LUNDQUIST and JOHN B. HARRIS *

Department of Pharmacology, University of Lund, S61vegatan 10, S-223 62 Lund, Sweden Received 6 September 1977, revised MS received 11 September 1978, accepted 6 November 1978

I. LUNDQUIST and J.B. HARRIS, Insulin secretion and carbohydrate metabolism in the dystrophic mouse, European J. Pharmacol. 53 (1979) 365--373. Dystrophic mice were investigated with regard to their regulation of blood glucose and insulin secretion in vivo. The following were also measured: tissue glycogen levels, activity of the glycogenolytic hydrolase, acid amyloglucosidase, and in vitro glucose utilization by liver, muscle and adipose tissue. Basal levels of blood glucose and plasma insulin of dystrophic mice were essentially within the same range as in the clinically unaffected littermate controls. Dystrophic mice had a decreased tolerance to glucose and glibenelamide; the secretion of insulin in response to these secretagogues was moderately reduced. Insulin release following ~-adrenergic stimulation, however, was increased in the dystrophic mice. Glycogen levels and acid amyloglucosidase activity were increased in dystrophic muscles but were normal in liver. Acid amyloglucosidase activity in pancreatic islets was lower in the dystrophic mouse. Glucose utilization in vitro appeared normal in liver tissue from dystrophic mice; in dystrophic muscle there was a threefold increase in 14CO2-production with no concomitant increase in either glycogen or 14C-incorporation into glycogen. 14CO2 production and 14C-incorporation into lipid and glycogen were increased in dystrophic adipose tissue. We suggest that the decreased glucose tolerance, and the reduced insulin response to glucose in the dystrophic mouse are compensated by an increased glucose utilization in muscle and adipose tissue and an increased ~-adrenergic-mediated secretion of insulin. Dystrophic mice Tissue glycogen

Blood glucose

Plasma insulin

1. Introduction A variety of diseases of skeletal muscle and the neuromuscular junction are reportedly associated with abnormalities in glucose metabolism (Collis and Engel, 1968). In particular, patients with myotonic dystrophy have been observed to have a high incidence of diabetes mellitus and to exhibit abnormalities of insulin secretion (BjSrntorp et al., 1973; Walsh et al., 1970). The dystrophic mouse of the 129/ReJ strain suffers an inherited form of muscular dystrophy (Michelson et al., 1955; * Present address: Muscular Dystrophy Group Laboratories, Regional Neurological Centre, Newcastle General Hospital, Newcastle upon Tyne, NE4 6BE, England.

Glucose utilization

Acid amyloglucosidase

Meier and Hoag, 1966) and so may be considered a useful model for the study of both glucose metabolism in diseased skeletal muscle and general changes in carbohydrate metabolism which might be associated with the disease. The aim of the present investigation was to study carbohydrate metabolism and regulation in the dystrophic mouse with special regard to glucose tolerance, insulin secretion and glycogen content in vivo, and the glucose utilization in vitro of skeletal muscle, liver and adipose tissue. Additionally the activity of the glucose-producing glycogenolyric hydrolase, acid amyloglucosidase, was assayed in liver and skeletal muscle tissue, and in isolated pancreatic islets.

366

2. Materials and methods 2.1. Animals

Female 129/ReJ dystrophic mice with apparently unaffected littermate controls were obtained from the Jackson Laboratory, Bar Harbor, Maine, U.S.A., at the age of about 3 months. The dystrophic animals are homozygous for the dystrophic gene (i.e. dydy); the littermate controls may be either homozygous (i.e. DyDy) or heterozygous (i.e. Dydy). Each dystrophic animal was housed, with its littermate control and kept on a standard pellet diet (Astra-Ewos, SSdert~ilje, Sweden) supplemented with wheat germ and dried milk. They had free access to food and tap water before and throughout all experiments. 2.2. Experiments in vivo

The insulin secretagogues used were dissolved in 154 mmol/1 NaC1 and injected into a tail vein of the mouse in a volume of about 0.2 ml. Blood sampling in unanaesthetized mice was performed by orbital puncture using commercial constriction pipettes as described previously (Rerup and Lundquist, 1966}. Blood glucose levels were determined enzymatically {Marks, 1959). The plasma immunoreactive insulin levels measured in mice following injection of the different insulin secretagogues were always the peak levels of the first phase of insulin secretion. Repeated experiments in this laboratory have shown that maximum concentrations of immunoreactive insulin in mouse plasma following a rapid intravenous bolus of D-glucose or the sulphonylurea derivative glibenclamide were achieved after about 2 min, whereas peak levels after the ~-adrenergic agonist L-isopropyl-noradrenaline (L-IPNA) or corticotrophin (ACTH) were obtained after 5--6 and 8--10 min respectively. The concentrations of insulin in plasma were determined by the method of Heding (1966) using 12SI-labelled pig insulin and guinea pig antipig ins[tlin

I. LUNDQUIST, J.B. HARRIS

(kindly provided by Dr. L. Heding, Novo Research Institute, Copenhagen, Denmark). The direct assay of tissue glycogen levels representative of the in vivo situation was performed according to Rerup and Lundquist {1967). The muscle specimen for this purpose consisted of the hind leg muscles after removal of the gastrocnemius for in vitro incubation and enzyme assay (see below). L-IPNA was generously supplied by H~issle AB, GSteborg, Sweden, and glibenclamide by Boehringer GmbH, Mannheim, FRG. Synthetic tetraicosapeptide corticotrophin (Synacthen®) was obtained from CIBA Ltd., Basel, Switzerland. 2.3. Experiments in vitro

The animals were sacrificed by cervical dislocation and samples of liver, muscle (gastrocnemius) and adipose tissue (parametrial fat pads) were immediately removed, weighed and rinsed (except adipose tissue). The tissue samples were incubated for 1 h at 37°C in Krebs-Henseleit solution, pH 7.4, containing 2 mg gelatine (U.S.P.), 2 mg D-glucose and 0.1pCi (liver 0.3 pCi) D-(U-14C)-glucose per ml incubation medium as previously described (Lundquist, 1972a). The amount of labelled carbon dioxide was determined according to Lyngs~e (1961), 14C-lipid according to Fain, et al. (1963) and 14C-glycogen from adipose tissue according to Leonards and Landau (1960}. 14C-Glycogen and total glycogen from liver and muscle was isolated and determined as described previously (Lundquist, 1972a). Labelled material was counted in a Packard Tri-Carb liquid scintillation spectrometer or in a flow-counter (Nuclear-Chicago) as described previously (Lundquist, 1972a). D-(U-14C)-Glucose (spec. act. 3 mCi/mmol) was obtained from the Radiochemical Centre, Amersham, England. All other chemicals were obtained from British Drug Houses Ltd., Poole, England. 2.4. Enzyme assay

The preparation of isolated pancreatic islets and the determination of acid amylogluco-

CARBOHYDRATE REGULATION IN THE DYSTROPHIC MOUSE

sidase activity in islet tissue using glycogen as substrate was performed as described previously (Lundquist, 1971b). Acid amyloglucosidase activity in homogenates of liver and muscle (gastrocnemius) was determined essentiaUy as previously reported (Lundquist, 1972c) with some modifications as follows. The reaction mixture for assay of the liver enzyme contained 1% (w/v) liver tissue in 0.25 mol/1 mannitol, 2 mmol/1 EDTA, 2.5 mmol/1 Tris, 1% (w/v) glycogen, 0.5% (v/v) Triton X-100 and 50 mmol/1 maleate buffer, pH 5.0. The assay mixture for the muscle enzyme contained 2.5% (w/v) muscle tissue in 0.25 mol/1 mannitol, 2.5 mmol/1 EDTA, 2.5 mmol/1 Tris, 1% glycogen, 0.5% Triton X-100 and 50 mmol/1 maleate buffer, pH 6.0. The reaction was carried out at 37°C for 90-300 min in triplicate and the liberated glucose was determined with a specific glucose oxidase reagent (Dahlqvist, 1961). Protein was determined by means of the Lowry procedure (Lowry et al., 1951) using crystalline bovine albumin as standard.

2.5. Statistical analysis of results The results are generally quoted as mean ± standard error of the mean (S.E.M.). Students t-test was used to compare the means and probability levels are quoted; where P < 0.05 the difference is considered significant.

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TABLE 1 Basal blood glucose levels in dystrophic mice and their littermate controls during the experimental period; 7--9 mice in each group (2 dystrophic mice died before the age of 33 weeks). P = probability level of random difference; * P < 0.005; ** P < 0.05. Age (weeks)

22 24 26 28 30 33

Blood glucose mg/100 ml (mean ± S.E.M.) Control

Dystrophic

65.3±3.3 68.6±2.4 57.7±1.6 66.6±1.5 64.8±4.4 49.8±4.2

67.6±3.6 64.8±1.7 58.6±3.2 57.5±1.9" 56.6±2.2 38.1±3.0"*

months old. The blood glucose concentrations were of the same magnitude in both groups during the whole experimental period with a slight but sometimes statistically significant

CONTROL O~mlll OYSTROPHIC

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12.5. 3.1. Blood glucose levels and glucose tolerance It was reasoned that any possible abnormalities in the carbohydrate metabolism of the dystrophic mice might increase with advancing age of the animals, and so no experiments were performed until they had reached the age of 22 weeks. At that time the colony consisted of 9 dystrophic animals with their 9 littermates. The basal blood glucose levels were then recorded every second week until the animals were sacrificed when about 8

MINUTES

Fig. 1. Glucose elimination rate (k) in dystrophic mice and their Uttermate controls at the age of about 7 months after an intravenous glucose load of 8.3 mmol/kg. The k values (means + S.E.M.) were calculated from the regression lines of the blood glucose levels obtained after subtracting the basal glucose values; 6 mice in each group. Probability level of random difference is marked by: * * * P < 0.001. Ordinate: blood glucose (log scale) normalised to 100% at rain 3. Abscissa: time (min).

368

I. L U N D Q U I S T , J.B. H A R R I S

decrease in the dystrophic animals after the age of about 7 months (table 1). There was, however, a clear difference in glucose tolerance at this age (fig. 1). Serial blood sampling was performed after a rapid intravenous injection of glucose at a dose of 8.3 mmol/kg. Fig. 1 compares the glucose elimination rate (k-value) in the dystrophic mice and their littermates, and demonstrates a significant impairment of glucose tolerance in the dystrophic animals. The k value was calculated from the regression line of the blood glucose concentrations observed between 15 and 60 min, after subtraction of the basal glucose values. Best-fit regressions were calculated by the method of least squares. The half-life of glucose was calculated to be 20.9 -+ 2.0 min in the control group compared with 43.6 + 6.6 min in the dystrophic group (mean + S.E.M.) (P < 0.001). Note that the blood glucose values recorded in the dystrophic animals do not exclude the possibility of two glucose pools. One week later an experiment was performed to study the blood glucose pattern in the dystrophic mice and their littermates following an intravenous injection of the potent sulphonylurea derivative, glibenclamide, 0.5 pmol/kg. The decrease in blood glucose after glibenclamide was much more pronounced in the controls than in the dystrophic +2 0 .

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animals (fig. 2), which further suggested an abnormality of glucose handling in the dystrophic mice. 3.2. Basal and stimulated insulin secretion

A series of experiments was designed to study basal plasma insulin levels and the secretion of insulin in response to different secretagogues in the dystrophic mice and their littermates. From the age of 5.5 months to the age of about 8 months, insulin secretion was studied either in the basal state or after the intravenous injection of glucose, glibenclamide or L-IPNA on two different occasions for each secretagogue. An interval of about one week was allowed between each test. In addition at the age of 8 months one experiment was performed with ACTH as the secretagogue. Fig. 3 shows the basal plasma insulin levels and insulin secretion in response to a near-maximal dose of glucose and a maximal dose of glibenclamide. The basal plasma insulin levels were of similar magnitude in dystrophic animals and in controls, whereas the insulin response to both glucose and glibenclamide was slightly b u t significantly lower in the dystrophic animals. The secretion

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Fig. 2. Change in blood glucose level following the intravenous injection of glibenclamide ( 0 . 5 ~ m o l / kg) in dystrophic mice and their littermate controls; 6 mice in each group. Probability level of random difference is m a r k e d by: * P < 0.05; * * P < 0.01; *** P < 0.001. Ordinate: b l o o d glucose change (rag/ 100 ml). Abscissa: time (rain).

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Fig. 3. Immunoreactive plasma insulin levels (ordinate, pU/m]) in the basal state and peak levels f o l l o w i n g the intravenous injection o f a near-maximal dose of glucose (8.3 mmol/kg) or a maximal (Jose of glibenclamide (0.5 pmol/kg) in dystrophic mice and their littermate controls. Bars indicate standard errors of the mean. P = probability level of random difference. NS = n o t significant. N = n u m b e r of observations.

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TABLE 2 Concentration of liver and muscle glycogen (mg/g tissue wet weight) in dystrophic mice and their littermate controls. The tissue specimens were assayed for glycogen immediately after removal from the animal; 6 mice in each group. Results are mean +- S.E.M. * P < 0.05. Tissue

J. Liver Muscle

<0005 17

NS 16

9

7

Fig. 4. Plasma immunoreactive insulin (ordinate, peak levels, pU/ml) following the intravenous injection of a maximal dose of L-IPNA (1.37 pmol/kg) or ACTH (0.04 pmol/kg) in dystrophic mice and their littermate controls. Bars indicate S.E.M. P = probability level of random difference. NS -- not significant. N = number of observations. o f insulin in r e s p o n s e t o a m a x i m a l d o s e o f t h e /~-adrenergic agonist L - I P N A , h o w e v e r , was m u c h increased in t h e d y s t r o p h i c m i c e as c o m p a r e d w i t h t h e i r c o n t r o l s (fig. 4). N o significant d i f f e r e n c e was o b s e r v e d a f t e r t h e i n j e c t i o n o f A C T H (fig. 4). T h e initial b l o o d glucose levels in t h e d y s t r o p h i c animals did n o t usually d i f f e r f r o m t h a t in t h e c o n t r o l s b e f o r e t h e i n j e c t i o n o f secretagogues. H o w ever in t h e A C T H e x p e r i m e n t t h e b l o o d glucose level in t h e d y s t r o p h i c a n i m a l s (38.1 + 3.0 m g / 1 0 0 ml) was significantly l o w e r t h a n in t h e c o n t r o l s ( 4 9 . 8 + 4.2 m g / 1 0 0 ml).

3.3. Glycogen concentration and acid amyloglucosidase activity G l y c o g e n levels in liver a n d m u s c l e tissue w e r e d e t e r m i n e d in d y s t r o p h i c m i c e a n d t h e i r l i t t e r m a t e s at t h e age o f a b o u t 8 m o n t h s . T h e results (table 2) s h o w t h a t t h e c o n c e n t r a t i o n o f g l y c o g e n in t h e m u s c l e was n e a r l y t w i c e as

369

Glycogen mg/g wet weight Control

Dystrophic

65.8 + 7.4 0.6 + 0.1

62.1 + 5.8 1.2 + 0.2 *

high in t h e d y s t r o p h i c animals as in t h e controls, w h e r e a s n o s u c h d i f f e r e n c e was seen in liver tissue. T h e d y s t r o p h i c m i c e w e r e also f o u n d t o h a v e a significantly e n h a n c e d activity o f t h e glucose-producing glycogenolytic hydrolase, acid a m y l o g l y c o s i d a s e , in t h e i r m u s c l e tissue as c o m p a r e d w i t h t h e l i t t e r m a t e c o n t r o l s (table 3). T h e acid a m y l o g l y c o s i d a s e activity r e c o r d e d in i s o l a t e d p a n c r e a t i c islets, h o w e v e r , was f o u n d t o b e significantly l o w e r in t h e d y s t r o p h i c g r o u p . N o d i f f e r e n c e was observed in liver tissue (table 3).

TABLE 3 Acid amyloglucosidase activity in liver, muscle (gastrocnemius) and isolated pancreatic islets of dystrophic mice and their littermate controls. Results are expressed as ~tmol glucose liberated from glycogen/g tissue protein/rain (mean + S.E.M.). Figures in parentheses denote number of animals (liver, muscle) or number of individual determinations from 2 pools of islets comprising 3 mice each. * P < 0.05; ** P < 0.02. Tissue

Liver Muscle Pancreatic islets

Acid amyloglucosidase activity (pmol glucose liberated/g protein/min) Control

Dystrophic

3.06 + 0.13 (6) 1.02 + 0.14 (6) 3.86 +_0.68 (17)

3.10 + 0.13 (6) 1.48 + 0.14 ( 6 ) * 1.93 + 0.32 (14) **

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I. L U N D Q U I S T , J.B. H A R R I S

3.4. Glucose utilization in vitro by liver, muscle and adipose tissue The utilization of glucose by isolated specimens of liver, muscle and adipose tissue from dystrophic mice and their littermates at the age of about 8 months is summarized in table 4. There were no differences in any of the parameters for liver tissue from normal or from dystrophic mice. In muscle tissue, however, the 14CO2-production from D-(U-14C) glucose was three times greater in the dystrophic muscles than in the controls. There was no concomitant increase in 14C-incorporation into glycogen and there was indeed no increase in total glycogen. In fact, the glycogen concentration in the muscle from the normal animals was now significantly greater than the concentration in the dystrophic muscles (mean increase + S.E.M. = +92.1 -+ 35.3%; paired t-test, P ~ 0.05; cf. table 2). In the adipose tissue of dystrophic mice there was a significant increase in ~4CO2-production and ~4C-incorporation into lipid from labelled glucose. Also 14C-incorporation into glycogen was enhanced (table 4). Thus all parameters recorded for glucose utilization in adipose tissue were increased in the dystrophic mouse.

4. Discussion Neither previous reports on blood glucose homeostasis in murine dystrophy (Baker and Huebotter, 1964; Baker et al., 1958) nor the data presented above showing unchanged or decreased basal blood glucose levels with advancing age in the dystrophic animals suggest that hereditary m u r i n e ' d y s t r o p h y is associated with any abnormality of carbohydrate metabolism that could eventually lead to the development of diabetes mellitus. The life-span of dystrophic mice is known to be relatively short (Meier and Hoag, 1966) and, as the present study was performed when the animals were of an unusually old age, the characteristic symptoms and signs of diabetes would have been expected to appear in a substantial number of the mice. Our experiments nevertheless revealed several abnormalities of carbohydrate metabolism and its regulation in the dystrophic mice as compared with their littermate controls. For example, it was shown that the dystrophic mouse could maintain normal basal blood glucose levels in spite of an obvious impairment in both glucose tolerance and glibenclamide tolerance. Further, it was ob-

TABLE 4 Glucose u t i l i z a t i o n a n d glycogen c o n c e n t r a t i o n in liver, muscle ( g a s t r o c n e m i u s ) and a d i p o s e tissue ( p a r a m e t r i a l ) f r o m d y s t r o p h i c mice a n d t h e i r l i t t e r m a t e c o n t r o l s a f t e r i n c u b a t i o n in vitro. Glucose u t i l i z a t i o n was m e a s u r e d as D - ( u A a c ) glucose c o n v e r s i o n to 14CO2, 14C-glycogen, a n d 14C-lipid. Results are expressed as c p m × 1 0 - 3 / g tissue w e t w e i g h t / h ( m e a n + S.E.M.). T o t a l glycogen c o n c e n t r a t i o n s are expressed as m g glycogen/g tissue wet w e i g h t ( m e a n -+ S.E.M.). 6 tissue s p e c i m e n s in each g r o u p (6 mice). * P < 0 . 0 2 ; ** P < 0.001. Control 1.7 + 3.8 + 26.5 +

Dystrophic

Liver Tissue

14CO2 14C-Glycogen .Glycogen

0.2 0.7 8.8

1.9 + 3.6 + 21.3 +

0.2 0.7 5.8

Muscle tissue

14CO2 14C-Glycogen Glycogen

2.5 +_ 1.7 202 + 12 1.7 _+ 0.1

7.9 -+ 0.9 ** 197 + 4 5 0.3 1.1 +

Adipose tissue

14CO2 14C-Glycogen 14C-Lipid

11.1 + 2.3 15.9 + 5.7 279 _+ 67

64.9 -+ 7.3 ** 113 + 30 * 1696 -+ 227 **

CARBOHYDRATE REGULATIONIN THE DYSTROPHIC MOUSE served that the secretion of insulin in response to both glucose and glibenclamide was decreased in dystrophic animals compared with the controls. It would seem then, that the decreased tolerance to glucose and glibenclamide could at least in part, be explained by the impaired insulin response to these secretagogues. In this context, it is of interest that the activity of acid amyloglucosidase in the pancreatic islets of the dystrophic mouse was reduced, since there is good evidence for a direct relationship between the level of the enzyme and the ability of glucose and glibenclamide to stimulate insulin secretion (Lundquist, 1974, 1975; Lundquist and LSvdahl, 1975, 1977). The decreased functional muscle mass in the dystrophic mice might restrict the capacity of the muscle to utilize glucose and thus also contribute to the reduced glucose tolerance. This latter possibility is consistent with current concepts regarding the decreased glucose tolerance seen in various muscle wasting diseases in man (BjSrntorp et al., 1973; Collis and Engel, 1968). The dystrophic mouse appears to have developed novel compensatory mechanisms for counteracting the abnormalities in insulin secretion and glucose handling. Glucose utilization in both the dystrophic muscle tissue and the adipose tissue of these animals was much greater than that of the controls. The pattern of glucose utilization, however, was different in the two tissues. In dystrophic muscle, the increased metabolism of glucose seemed to be related to the oxidation of 14C-glucose to 14CO2, the incorporation of ~4C into glycogen being similar to that in the control muscles. In fact, although the total glycogen concentration in the dystrophic muscles was higher than in the control muscle in vivo (table 2), the situation tended to reverse after incubation in vitro for 1 h (table 4). This observation might suggest that the dystrophic muscles contain a glycogen pool which is metabolically inaccessible and probably different from the normal metabolic pool. The reason for making tl~is suggestion is that after in vitro incubation, the glycogen

371

concentration in the dystrophic muscle fell. This would have resulted in a relative dilution of the glucose-6-P pool, which in turn would lead to a relative decrease of 14CO2 production in the dystrophic muscle, and not to a threefold increase as was the case (table 4). Furthermore, glucose carbon incorporation into glycogen was similar in both normal and dystrophic muscles. Whether the relative change in glycogen content in the dystrophic muscle was in some way associated with the increased activity in the muscle tissue of the glycogenolytic hydrolase, acid amyloglucosidase (table 3), remains to be elucidated. However since the activity of this enzyme is known to be largely associated with the lysosomal apparatus in several mammalian tissues (Lundquist, 1971a; Smith et al., 1968) it is conceivable that the hypothetical, less accessible pool of glycogen is membrane bound or associated with the lysosomal apparatus in some other way. Since enzymes of the citric acid cycle do not show any appreciable difference in activity between normal and dystrophic muscle (Jato-Rodriguez et al., 1972) the increased 14CO2 production in dystrophic muscle reported here (table 4) might be compatible with an enhanced activity of the pentose shunt. This possibility is strengthened by the earlier observations (McCaman, 1963; Stamp and Lesker, 1967) that the activity of hexokinase, glucose 6-Pdehydrogenase and 6-P-gluconic dehydrogenase was increased in dystrophic mouse skeletal muscle. The increased glucose utilization in adipose tissue of the dystrophic mice apparently involved both 14CO2-production 14C-incorporation into lipid and to a less significant extent, 14C-incorporation into glycogen. Although the adipose tissue mass is small in the dystrophic mice, it might still make a significant contribution to the maintenance of a normal glucose homeostasis. The mechanisms responsible for this increased glucose oxidation in adipose tissue are still unclear. Our data do not support any major role for insulin in this context since basal insulin levels were similar to

372 the control levels, and glucose-induced insulin secretion was decreased. T he stimulation of fl-adrenergic receptors pr om ot e s glucose utilization in parametrial adipose tissue of the mouse (Lundquist, 1972a, 1972b). It is known t h a t th er e are increased catecholamine levels in the skeletal muscles, adrenal glands and urine of the dystrophic mouse (G o rd o n and Dowben, 1966; Kabara et al., 1976). High catecholamine levels m ay be related to th e increased e x t e n t of glucose utilization in adipose tissue. Adipose tissue in the dystrophic mouse may also have an enhanced sensitivity to fl-adrenergic stimulation, as is clearly the case for fl-adrenergic stimulated insulin release. It has previously been shown t h a t the adrenergic system has a dual action on the insulin secreting mechanisms in t he mouse (Lundqulst, 1971a, 1972a, 1972b). The p r o m o t i o n o f insulin release is mediated through fl-adrenergic receptors and the inhibition o f insulin release is mediated through a-adrenergic receptors (Lundquist, 1971a; Porte and Robertson, 1973; Woods and Porte, 1974). An enhanced capability of the fl-adrenergic system o f the pancreatic B-cells to secrete insulin would be compatible with previous reports o f elevated catecholamines and increased activity of the adenylate cyclase-cyclic AMP system in ot he r tissues of hereditary dystrophic animals ( G or don and Dowben, 1966; L i n e t al., 1976; Rodan et al., 1974). Insulin release in response to/3-adrenergic stimulation is i n d e p e n d e n t o f amyloglucosidase activity (Lundquist, 1975; Lundquist and LSvdahl, 1975). Thus the enhanced secretion of insulin in response to fl-adrenergic stimulation in the dystrophic mouse (fig. 4) might be a c o m p e n s a t o r y mechanism for the reduced sensitivity to glucose; this would further support our previous suggestion (Lundquist, 1975; Lundquist and LSvdahl, 1975, 1977) th at the insulin secretory process following ~-adrenergic stimulation is at least in part qualitatively distinct from t he insulin release induced by glucose.

I. LUNDQUIST, J.B. HARRIS Acknowledgements The expert technical assistance of Mrs. Lena Kvist is gratefully acknowledged. This work was supported by the Swedish Medical Research Council (Project No. 14X-4286; 04P-4289), the Swedish Diabetes Association and the Medical Faculty, University of Lund, Sweden. J.B.H. was supported by a WellcomeSwedish Travelling Fellowship during the early stages of this project.

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