INTRODUCTION

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NONSHIVERING THERMOGENESIS (NST) is a common adaptive response to cold found in a number of mammalian species (9) and a few species of birds, including chickens, ducklings, and penguins (2, 13, 19). However, the sites and mechanisms of NST appear to differ in these classes. Brown adipose tissue (BAT) is well known to account for a large proportion of mammalian NST, whereas skeletal muscle is the main site of NST in ducklings (12). BAT thermogenesis is based on an uncoupling of the mitochondrial respiratory chain by the proton-translocator uncoupling protein (28). By contrast, avian mitochondria contain no mammalian-like uncoupling protein (29). Proposed mechanisms for avian NST include those based on fatty acid-induced loose-coupling of mitochondrial respiration (5) and also increased Ca²⁺ cycling by the sarcoplasmic reticulum (17). Although the sympathetic control of BAT NST is well documented, far less is known about the endocrine control of avian muscle NST.

On the basis of its marked thermogenic and lipolytic effects in birds (3, 4, 6, 14), glucagon appears as a potential mediator of avian NST. Moreover, plasma glucagon concentration is increased in cold-acclimated ducklings (4), and chronic administration of glucagon to ducklings kept at thermoneutrality leads to the development of NST (3). It was postulated that glucagon may trigger muscle NST by stimulating the release of fatty acids from a multilocular adipose tissue differentiated for lipolytic activity (2). Released fatty acids may then affect the respiration of muscle mitochondria, which show a higher basal metabolic rate (MR) and a higher increase in respiration due to the uncoupling effect of fatty acids after cold acclimation (5). As reflected by in vivo measurements of muscle blood flow and arteriovenous differences in oxygen content, muscle NST can be stimulated by exogenous glucagon (14). However, whether the action of glucagon is indirect through lipolysis or direct on myocytes cannot be inferred from these experiments. Besides the action of glucagon, other hormones such as catecholamines could also play a part in the stimulation of avian NST on account of some thermogenic effects of these hormones in birds, both in vivo (6, 24, 31) and in vitro (25). However, such calorigenic action is not invariably found (9) and is much lower than the calorigenic action of glucagon (6).

The use of in vitro perfused muscle preparations has provided numerous insights to the humoral control of muscle metabolism in mammals (reviewed in Ref. 26). Extended studies with perfused hindlimbs in rats have underlined the major role of catecholamines and vasoconstrictors in the modulation of muscle oxygen consumption (11). Recently, with the use of a perfused leg muscle preparation, it has been shown that catecholamines in interaction with glucagon could be potent stimulators of muscle resting oxygen uptake in chickens reared at thermoneutrality (18). It is, however, yet to be established whether this effect is potentiated in animals developing NST and is thus of adaptive interest.

The aim of this experiment was therefore to develop a perfused muscle preparation in ducklings allowing us to address the hypothesis that muscle thermogenesis is under direct hormonal control. Potential thermogenic and vascular effects of catecholamines and glucagon have been investigated in control ducklings reared at thermoneutrality (TN), and also in cold-acclimated (CA) and glucagon-treated (GT) ducklings, the latter two groups of animals showing muscle NST in vivo. Possible calorigenic effects of epinephrine were also investigated in vivo.

	MATERIALS AND METHODS

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Animals. Male Muscovy ducklings (Cairina moschata L, pedigree R31, Institut National de la Recherche Agronomique) were obtained from a commercial stock breeder (Ets Grimaud). They were fed a commercial mash (Aliment Genthon Démarrage) ad libitum and had free access to water. Ducklings were kept in a constant photoperiod (8:16-h light-dark cycle). The cold-acclimation schedule described by Barré et al. (2) was used: from the age of 1 wk, ducklings were caged in groups of six for a period of 4 wk at either 4°C ambient temperature (T_a) (CA) or 25°C T_a (TN). The glucagon treatment schedule described by Barré et al. (3) was used. From the age of 1 wk, ducklings were caged in groups of six for a period of 4 wk at 25°C T_a and received twice daily an intraperitoneal injection of glucagon (103 nmol/kg; GT).

To standardize the basal values of oxygen consumption and perfusion pressure among the three groups, animals were kept at thermoneutrality (25°C) for 2 h before the start of the surgery.

Birds were cared for under the French Code of Practice for the Care and Use of Animals for Scientific Purposes, and the experimental protocols were approved by the French Ministry of Agriculture Ethics Committee (Animals).

Muscle preparation. The duckling limb surgical preparation was derived from that described by Eldershaw et al. (18) on chicken. Ducklings were anesthetized by continuous halothane inhalation. The left lower limb was surgically isolated and perfused. The major skin vessels were ligated, and the skin covering the limb was carefully removed. A prior intracardiac injection of heparin (2,000 U/kg) prevented blood coagulation during catheterization and the subsequent replacement of blood with artificial perfusate. The popliteal fossa was incised to expose the popliteal artery and vein. The popliteal nerve was divided and the hamstring muscles were ligated and resected proximal to the fossa to give good access for cannulation of the popliteal artery and vein with a polyethylene tubing (1.2 mm ID) filled with heparinized saline. To prevent flow to other tissues, tight ligatures were placed around the ankle and thigh, just above the cannulation site. The duckling was killed with an intracardiac injection of a lethal dose of pentobarbital. The arterial catheter was immediately connected to the perfusion open circuit. The entire surgical procedure routinely lasted ~30 min. During perfusion, the animal was laid on its back and the limb was supported partially aloft. Exposed muscle was covered in plastic cling wrap. After the perfusion had started, the contralateral limb was ligated similarly and excised. The muscles were removed and weighed to allow calculation of required perfusate flow to the perfused limb. This method of estimating perfused muscle mass was validated by infusing 1% Evans blue dye with the perfusate, followed by removal and weighing of stained muscle to determine the mass of muscle perfused, generally in the range of 23-27 g.

Perfusion system. The perfusion system was similar to that described previously (18). Briefly, it consisted of a nonrecirculating constant flow system thermostated at 25°C to ensure adequate oxygenation without the use of red blood cells. A Krebs-Henseleit buffer was used as the perfusion medium, with a composition of (in mM) 120 NaCl; 4.8 KCl; 1.2 KH₂PO₄; 1.2 MgSO₄, 7H₂O; 2.5 CaCl₂; and 24 NaHCO₃; this buffer contained 8.3 mM glucose and 2% bovine serum albumin (BSA), pH 7.4. Before use, the perfusion medium was filtered (Millipore, 0.45 µm). The buffer was continuously gassed with carbogen (95% O₂-5% CO₂) both in a reservoir placed on ice and by passage through a Silastic tubing lung (5 m; 0.058 cm ID; 0.077 cm OD) before entering the limb muscles to ensure a constant high saturation of arterial O₂ and CO₂. The whole circuit, which consisted of a heat exchanger, bubble traps, and Silastic lung, was placed in a 25°C thermostated water bath. A small mixing chamber, placed on an electromagnetic stirrer, allowed the injection of hormones directly in the perfusate flow. Buffer solution was pumped from the reservoir through the perfusion system and into the arterial catheter by a Gilson peristaltic pump (Minipuls 8). An in-line pressure transducer (PDCR 75 Nortek Bio 1000) was situated immediately upstream from the arterial catheter. A mercury manometer was used to calibrate the transducer at the start and at the end of a series of experiments. The venous effluent flowed through a 1.5-ml thermostated chamber (25°C) containing a Clark-type oxygen electrode connected to a Gilson apparatus, and the effluent was discarded after measurement of its oxygen partial pressure. The venous tubing and associated electrode chamber did not exert any back pressure on the isolated muscle preparation. The electrode was calibrated before and after each experiment with gas mixture at 100% and 20.93% (atmospheric air) of oxygen. Muscle O₂ consumption (O₂) was calculated from arteriovenous difference in O₂ content and flow rate; the oxygen dissolution Bunsen coefficient of O₂ in plasma was used. Where stated, infusions of hormones into the mixing chamber were made continuously at <1% of the total flow.

Flow rate. Perfusions were conducted at flow rates of either 0.33 ml · min¹ · g¹ (n = 5-6 per group) or 0.47 ml · min¹ · g¹ (n = 5 per group). The lower flow rate was chosen to be within the range of average skeletal muscle blood flow measured in vivo, whereas the higher rate corresponded to an increase in flow equivalent to that seen in vivo after a glucagon injection to the animal (14). The chosen flow rate remained constant throughout each experiment.

Hormone infusion. The effects of infused glucagon and catecholamines [norepinephrine (NE) and epinephrine] were studied. Catecholamines were dissolved in saline (9 g/l NaCl) containing 0.1% ascorbic acid. Solutions were freshly prepared for each experiment. Glucagon solution (Novo-Nordisk) also contained 107 mg lactose/mg glucagon. In each series of experiments, infusion of vehicle alone had no effect on O₂ or perfusion pressure. Hormones were infused in the perfusate flow by a precision peristaltic pump (Gilson minipuls). Final hormonal concentrations tested were between 1 nM and 1 µM.

Muscle metabolites. Muscle metabolite concentrations were determined in limb muscle samples previously freeze clamped with tongs precooled in liquid nitrogen. Samples were obtained after 2 h of perfusion at 25°C at low (0.33 ml · min¹ · g¹) or high (0.47 ml · min¹ · g¹) flow rate with or without hormone stimulation (3-6 determinations per group). Frozen samples were stored at 80°C and powdered under liquid nitrogen. Creatine phosphate, ATP, ADP, and AMP were assayed by spectrophotometric methods. The values obtained with perfused muscles in vitro were compared with in vivo data obtained from a separate batch of ducklings (n = 4-6 ducklings per group). These birds were catheterized (jugular vein) 2 days before sampling under halothane anesthesia. At the time of sampling, ducklings were placed in individual boxes and the catheter was connected to polyethylene tubing. Animals were rested for 2 h and were then anesthetized with 1 ml of pentobarbital sodium (50 mg/kg) injected through the catheter without disturbing the animal. Immediately after anesthesia, limb muscles were exposed and freeze clamped. With this protocol, in vivo samples were obtained with minimal prior contraction of the sampled muscles.

Estimation of edema. Muscle samples were taken from the perfused limb after 2 h of perfusion (n = 3-6 values per group) or from the contralateral limb at the beginning of the experiment (denoted in vivo; n = 4-6 values per group). Samples were dried at 80°C to a constant weight. The ratio of fresh weight to dry weight was calculated from in vivo and perfused muscle samples to estimate the magnitude of edema after 2 h of perfusion with a 2% BSA buffer.

In vivo MR measurement. MR was measured in vivo by indirect calorimetry in an open circuit as described previously (2). To allow saline or epinephrine infusion, the jugular vein was cannulated under halothane anesthesia 2 days before the experiment. On the day of the experiment, the vascular catheter was connected to a continuous infusion pump (A99, Bioblock Scientific), and the bird was positioned in the thermostatic chamber set at thermoneutrality (25°C). As performed with the protocol with perfused muscle in vitro, increasing hormone doses were tested successively in vivo after a 120-min preliminary equilibration period. Each epinephrine infusion lasted 20 min and was followed by a 40-min recovery period. MR was monitored throughout. Total doses (and infusion rates) of epinephrine bitartrate salt were 2 µg/kg (100 ng · kg¹ · min¹), 20 µg/kg (1 µg · kg¹ · min¹), and 100 µg/kg (5 µg · kg¹ · min¹), thus in the range of 6-300 nmol/kg. Epinephrine solutions were freshly prepared immediately before infusion. In vivo experiments were performed with TN (n = 8) and CA (n = 5 or 6) ducklings. Epinephrine was chosen because this hormone induced a higher thermogenic effect than NE in king penguin chicks in vivo (6).

Statistics. Values are presented as means ± SE. Statistical significance of observed variations was assessed by the one-factor analysis of variance (ANOVA) for repeated measures; observed differences between means were then tested by Scheffé's F test. Differences between values from the same group were assessed by Student's paired t-tests. The three-factors ANOVA (hormone concentration × group × flow rate) was reduced to only two factors by calculating the integrated response for the entire range of concentrations tested (area under the curve); a two-way ANOVA was then used. Statistical significance was recognized at P < 0.05.

	RESULTS

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Characterization of the perfused muscle preparation. Evans blue dye (1%) was routinely infused at the end of perfusion through the arterial catheter to assess the degree of perfusion of the preparation. The resultant staining confirmed that perfusate flow was confined to the lower limb in both hormone-stimulated and nonstimulated preparations. More than 98% of the isolated muscle tissue was actually perfused in this model.

Muscle wet weight-to-dry weight ratio (Table 1) was not different among the three in vivo groups, and no significant change occurred after perfusion at a low flow rate. At the high flow rate, the ratio slightly increased, indicating some edema formation in both TN and CA groups after >2 h of perfusion at this flow rate.

Muscle phosphagen concentrations (Table 1) were no different from in vivo levels after 2 h of perfusion at the low (0.33 ml · min¹ · g¹) or high (0.47 ml · min¹ · g¹) flow rate without hormonal stimulation. Muscle energy charge of the adenylate system, defined as ([ATP] + 0.5 × [ADP])/([ATP] + [ADP] + [AMP]), where brackets indicate concentration, remained at the in vivo value regardless of the perfusion flow rates. Muscle phosphagens concentrations and energy charge were also maintained at the in vivo levels after perfusion with hormonal stimulation. A slight increase in the wet weight-to-dry weight ratio was, however, noted after very high doses of catecholamines.

Basal values of O₂ and perfusion pressure. At the low perfusion rate, basal O₂ was higher in CA (10.1 ± 0.4 µmol · g¹ · h¹, P < 0.01) than in TN (8.1 ± 0.4 µmol · g¹ · h¹) and GT (8.5 ± 0.3 µmol · g¹ · h¹) ducklings (Table 2). Yet the difference between CA and TN ducklings was not significant at the high perfusion flow rate. A significant group effect (P < 0.05) was also observed at low flow rate on basal perfusion pressure, which was higher in TN (43.8 ± 1.6 mmHg) than in CA (35.8 ± 1.1 mmHg) and GT (31.3 ± 1.3 mmHg) ducklings (P < 0.05 in each case). The difference in pressure persisted at the high perfusion flow rate between TN (53.1 ± 2.9 mmHg) and CA (41.3 ± 2.3 mmHg) ducklings (P < 0.05).

NE effects on O₂ and perfusion pressure. Increasing concentrations of NE, including 1, 10, 20, 50, and 300 nM and 1 µM, were tested. Regardless of prior treatment or flow rate, NE infusion induced a rapid increase in perfusion pressure as well as a rapid decrease in venous PO₂ (Fig. 1), indicating a rapid onset of vasoconstriction and an increase in O₂. Effects were sustained throughout the period of NE infusion and were rapidly reversed on cessation of NE infusion. Dose-curve responses for both O₂ and perfusion pressure are shown in Fig. 2. In all treatment groups, NE induced significant increases in O₂ and perfusion pressure at concentrations higher than 1 nM (P < 0.05). Effects on O₂ were higher in CA and GT ducklings than in TN controls, whereas effects on perfusion pressure were lower. The maximal NE-stimulated increase in O₂ at the low flow was 2.5 µmol · g¹ · h¹ in TN (+31% over basal), 3.6 µmol · g¹ · h¹ in CA (+36% over basal), and 3.7 µmol · g¹ · h¹ in GT ducklings (+43% over basal). At higher doses of infused NE (>300 nM), O₂ tended to decrease. The inhibitory effect of high doses of NE was, however, much more marked at the high flow rate, O₂ being significantly decreased in TN ducklings (P < 0.05), whereas it was still increased (+35%) in CA ducklings. In TN animals, the O₂ obtained with the highest dose of NE was not significantly different from the basal value without hormone stimulation. This was despite a dose-dependent continuous increase in perfusion pressure. Perfusion pressures associated with maximal O₂ effects at the low flow rate tended to be higher in TN (+330%) than in CA (+280%) and GT (+260%, P < 0.05) ducklings. Similar trends were observed at the high flow rate. There was a trend for half-maximal effective concentration values to be lower in CA and GT ducklings (3.6 ± 1.7 and 4.1 ± 1.4 nM, respectively) than in TN controls (13.5 ± 1.5 nM).

View larger version (15K): [in this window] [in a new window]   Fig. 1.   Representative tracings of venous PO2 (top) and perfusion pressure (P, bottom) obtained with perfused leg muscles of a cold-acclimated (CA) duckling. Tracings were recorded at the low flow rate (0.33 ml · min1 · g1) with a series of increasing norepinephrine concentrations, which were continuously infused for periods represented by the length of the solid bars.

View larger version (24K): [in this window] [in a new window]   Fig. 2.   Effects of norepinephrine on muscle oxygen uptake (O2) and perfusion pressure in thermoneutral (TN), CA, or glucagon-treated (GT) ducklings. Leg muscles were perfused at a low (0.33 ml · min1 · g1, left) or high (0.47 ml · min1 · g1, right) flow rate. Data are means ± SE from 5 or 6 experiments. In each group, norepinephrine induced a significant increase in both O2 and P for concentrations 10 nM (P < 0.05). When not visible, error bars are within symbols.

View larger version (30K): [in this window] [in a new window]   Fig. 3.   Integrated responses to norepinephrine (NE) on O2 and perfusion pressure in TN, CA, or GT ducklings. Values correspond to the integrated response (area under the curve) obtained with low norepinephrine concentrations (1-10 nM). Values are expressed by difference with basal values in each group (see values in Table 2). Leg muscles were perfused at a low (0.33 ml · min1 · g1, open bars) or high (0.47 ml · min1 · g1, hatched bars) flow rate. In GT ducklings, experiments were only performed at the low flow rate. Data are means ± SE of 5 or 6 ducklings. Values with different letters are significantly different (P < 0.05).

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Glucagon effects on O₂ and perfusion pressure. Increasing concentrations of glucagon (0, 1, 10, and 100 nM and 1 µM) were tested on perfused muscles of TN, CA, and GT ducklings (Fig. 4, low flow only). In TN and CA groups, experiments were conducted at the low and high (data not shown) flow rates. After glucagon infusion, a decrease in perfusion pressure was associated with an increase in venous PO₂, indicating some vasodilation and a decrease in basal O₂. These effects were usually significant for glucagon concentrations >10 nM (P < 0.05); the decreases in O₂ and perfusion pressure were then proportional to the glucagon concentration tested. The lack of direct thermogenic effect of glucagon was not due to the low perfusion temperature (25°C), because perfusion experiments performed at 40°C showed similar results (data not shown).

View larger version (17K): [in this window] [in a new window]   Fig. 4.   Effects of glucagon on O2 and perfusion pressure in TN, CA, or GT ducklings. Leg muscles were perfused at the low (0.33 ml · min1 · g1) flow rate. Number of perfusions is given in parentheses. Data are means ± SE. When not visible, error bars are within symbols. In each group, glucagon induced a significant decrease in both O2 and perfusion pressure for concentrations between 10 and 100 nM (P < 0.05).

View larger version (16K): [in this window] [in a new window]   Fig. 5.   Effects of glucagon (Gluc) and nitroprusside (NP) on norepinephrine-induced increases in O2 and perfusion pressure in TN (open bars) or CA (solid bars) ducklings. Leg muscles were perfused at the low (0.33 ml · min1 · g1) flow rate, and a dose of 100 nM norepinephrine (NE) was used for preconstriction. Data are means ± SE from 4 or 5 experiments. * P < 0.05 vs. effect in TN ducklings.

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View larger version (15K): [in this window] [in a new window]   Fig. 6.   In vivo effects of epinephrine infusion on metabolic rate in TN () or CA () ducklings at thermoneutrality (25°C). Epinephrine was infused intravenously for 20 min (horizontal bar) at either 100 ng · kg1 · min1 (A), 1 µg · kg1 · min1 (B), or 5 µg · kg1 · min1 (C). Data are means ± SE of 8 TN and 5 or 6 CA ducklings. When not visible, error bars are within symbols. * P < 0.05 vs. resting metabolic rate before epinephrine infusion.

	DISCUSSION

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The present study has investigated for the first time the endocrine control of resting muscle thermogenesis in ducklings. It has been demonstrated that perfusion flow rate and catecholamines can directly stimulate muscle thermogenesis in vitro. Furthermore, the thermogenic effect of catecholamines on perfused muscles in vitro was enhanced in CA and GT ducklings, two groups exhibiting regulatory muscle NST in vivo (2, 3, 12, 14). By contrast, glucagon had no direct thermogenic effect on perfused muscle in vitro. Finally, epinephrine at low doses exerted a small thermogenic effect in vivo in CA ducklings.

Validation of the perfused limb muscle preparation in ducklings. In the present study, the lower limb perfusion preparation originally described in the chicken (18) was successfully applied to ducklings. Perfusions were performed at 25°C to avoid the need for erythrocytes in the perfusate but to allow delivery of sufficient oxygen to the preparation, which was stable with respect to basal O₂, basal perfusion pressure, and high energy phosphate levels throughout the experimental period. Similar studies of muscle metabolism with perfused or incubated mammalian muscle preparations at 25°C gave similar results, with a correction factor, to those performed at 37°C (8, 11). The lower flow rate of 0.33 ml · min¹ · g wet wt¹ of muscle in the popliteal bed was chosen according to microsphere studies of blood flow in birds (12). At this flow rate, the venous PO₂ did not fall below 200 mmHg, and muscle phosphagen levels were maintained at their in vivo levels. The energy charge was consequently well preserved, and no effect of increased flow rate on phosphagen levels was observed. Because of the absence of edema development at this low flow rate even after long perfusions, Krebs buffer containing 2% BSA appears to be a suitable perfusate for duckling muscle at low temperatures. At the higher flow rate or after high doses of catecholamines, a slight edema was observed especially in TN ducklings, but similar degrees of edema were reported by others in perfused rat muscles (10). However, this did not cause any rise in basal perfusion pressure over time. The lower vascular resistance of CA and GT ducklings could be partly accounted for by a higher vascular density than in TN animals (15).

Basal values of O₂. The oxygen consumption of resting perfused muscles in TN duckling at 25°C (8.1 ± 0.4 µmol · g¹ · h¹) is similar to that measured in the chicken in the same conditions (7.4 ± 0.3 µmol · g¹ · h¹, Ref. 18). Limb muscles of CA ducklings showed a higher basal O₂ than those of TN ducklings (+25%), although all animals had been submitted to a 2-h fast at thermoneutrality before the surgical preparation. Similarly, the O₂ of incubated skeletal muscle slices from CA sparrows was higher than in controls (1). An increased basal O₂ was also noted in incubated muscles of CA pigs (23), a large mammal devoid of BAT. By contrast, such a difference was not observed with perfused muscles of CA and TN rats (21, 30). This higher basal O₂ of CA vs. TN ducklings could contribute to the higher resting MR measured in vivo at thermoneutrality (Ref. 2 and the present study). It may involve stimulations of several ion-transport mechanisms, such as the membrane Na⁺-K⁺-adenosinetriphosphatase (ATPase) activity, as shown in muscles of CA pigs (23), or the sarcoplasmic reticulum Ca²⁺-ATPase, as demonstrated in muscles of CA ducklings (17). Furthermore, it is in agreement with the increase in mitochondrial basal respiration observed in CA ducklings (5).

Flow-induced increase in O₂. An increase in the perfusion flow rate, within limits compatible with physiological variations, entailed an increase in O₂. It was not caused by an hypoperfusion at the low flow rate because no change in phosphagen levels was induced by the increase in flow, levels remaining similar to those measured in vivo. This phenomenon has already been observed in several perfused muscle preparations in rats and dogs (Ref. 10; review in Ref. 11). This flow-limited O₂ uptake can therefore be extended to bird muscles, as shown in ducklings (this study) and in chickens (18). The relationship between O₂ and perfusion flow could be interpreted as a physiological limitation of O₂ at rest by O₂ supply (10). Such a phenomenon could possibly account for the heterogeneity of perfusion observed in mammalian muscle at rest, possibly involving vasomotion in terminal arterioles (16). This does not mean that the muscle is hypoxic, because a marked decrease in phosphocreatine concentration should be observed under such circumstances. Heterogeneous perfusion of muscle has, however, not been reported, to our knowledge, in bird muscles. Other mechanisms could also be involved in the flow-induced O₂. Given the link between the increase in O₂ and the rise in perfusion pressure, a direct role of the smooth muscle cells of the vascular network has been suggested (11). Although the contribution of the vascular cell O₂ uptake to total O₂ remains to be fully evaluated, it can be noted that the lower flow-induced rise in basal O₂ of CA ducklings was accompanied by a lower rise in perfusion pressure. Alternatively, shear stress-dependent release of autacoids by endothelial cells might have a role in a paracrine control of skeletal muscle metabolism (11). The respective contribution, if any, of these different mechanisms remains unclear.

Glucagon effects on O₂ and perfusion pressure. On the basis that the higher muscle thermogenic response to catecholamines observed after cold acclimation was reproduced by the chronic administration of glucagon at thermoneutrality (GT ducklings), the present results support assertions that glucagon may play a major role in the long-term development of muscle NST in birds (3, 6). This is in keeping with other results obtained in vivo suggesting that glucagon could be a potential mediator of avian muscle NST (3, 6, 14). It was, however, not known whether the short-term thermogenic effect of glucagon was direct on skeletal muscle cells or mediated indirectly for instance by the release of endogenous stimulators such as fatty acids.

Perspectives