AMP-activated protein kinase controls metabolism and heat production during embryonic development in birds

The transition from ectothermy to endothermy during ontogenetic development in endotherms is facilitated by an increase in mitochondrial density and enzyme activities in oxidative pathways, which leads to an increase in oxidative metabolic capacity (Seebacher et al., 2006; Walter and Seebacher, 2007). Increased mitochondrial metabolic flux is at least partly driven by increasing cellular ATP demands caused by elevated Na⁺/K⁺-ATPase activity (Walter and Seebacher, 2009; Wu et al., 2004). At the same time mitochondria become more uncoupled (Walter and Seebacher, 2009). Together, these processes can increase metabolic heat production but reduce the efficiency of ATP synthesis (Brand et al., 1991; Else and Hulbert, 1987; Hulbert and Else, 1990). Therefore, the development of endothermy exposes the growing organism to an energy allocation challenge, between growth and heat production, which requires a regulatory mechanism to match ATP production with ATP usage. Understanding these as yet unknown regulatory mechanisms will significantly advance understanding of energetic allocation during endothermic development by providing the biological mechanisms underlying theoretical models of ontogenetic energy budgets (Hou et al., 2008; Kooijman et al., 2008). We propose that metabolic co-ordination during development is achieved by AMP-activated protein kinase (AMPK).

AMPK is one of the major cellular energy sensors (Carling et al., 2008) and an intracellular low-energy warning system (Hardie and Carling, 1997; Hegarty et al., 2009). AMPK is a serine/threonine kinase with heteromeric structure, composed of a catalytic alpha and regulatory beta and gamma subunits (Hardie and Carling, 1997). In situations of high ATP use, AMP levels increase and activate AMPK (Kahn et al., 2005) by promoting net phosphorylation of the Thr172 residue on the α subunit, and also by allosteric activation of phosphorylated AMPK (Sanders et al., 2007; Winder and Thomson, 2007). Additionally, AMPK phosphorylation might be regulated by hormones (for example, by thyroid hormones) to control metabolic pathways and mitochondrial biogenesis (Irrcher et al., 2003). Once activated, pAMPK can phosphorylate a variety of downstream targets, which can lead to an increase in the rate of fatty acid oxidation and ATP generation, and to decreased rates of energy utilization (Hardie and Carling, 1997).

As an immediate action to restore cellular energy charge, AMPK enhances the availability of carbohydrates and fats as fuels for mitochondrial oxidation to produce ATP. As one of its main targets, pAMPK phosphorylates acetyl CoA carboxylase (ACC), thereby blocking fat storage and promoting fatty acid oxidation in mitochondria (Winder and Thomson, 2007). In addition to this acute (1–2 h) regulation, AMPK is involved in long-term (over several days) regulation of ATP production. Thus, AMPK controls mitochondrial biogenesis by increasing the abundance of the transcriptional co-activator PGC-1α (Irrcher et al., 2003; Reznick and Shulman, 2006), and by activating nuclear respiratory factor-1 (NRF-1) (Bergeron et al., 2001; Zong et al., 2002). PGC-1α is a prominent metabolic regulator in mammals (Finck and Kelly, 2006; Puigserver and Spiegelman, 2003), and it has recently been shown to play a role in the metabolic regulation of birds (Walter and Seebacher, 2007). Hence, because of its potential to modulate mitochondrial capacity and thereby heat production and ATP availability, AMPK possesses the right characteristics to act as a metabolic regulator during the embryonic development of endothermy.

The aim of this study was to determine whether AMPK controls metabolic capacity and heat production during ontogenetic development of birds. We tested the hypotheses that chronic AMPK activation boosts endothermic heat production and increases the oxidative capacity of mitochondria, and that thyroid hormones coordinate these events via their regulatory effect on AMPK activation.

All procedures were approved by the University of Sydney Animal Ethics Committee (Approval Number: L04/11-2008/1/4922). We used chicken embryos as model system because of the well established transition from ectothermy to endothermy; during development embryos are ectothermic and cannot regulate their body temperature metabolically, but chicks become endothermic by eight days after hatching (Decuypere, 1984; Tzschentke and Nichelmann, 1999). Fertilized chicken eggs (Gallus gallus) were obtained from a local supplier (Cordina Chicken Farms, Sydney, Australia). Eggs were incubated at a constant temperature (38°C) and relative humidity (60–67%; Octagon 40 Incubator, Brinsea Products, Sandford, North Somerset, UK). Internal piping occurred on day 19/20 and hatching on day 20/21 of incubation. After hatching the hatchlings were transferred to an enclosure in a temperature-controlled room (23.0±2.0°C), and were provided with heating lamps, and with water and commercial chicken starter feed ad libitum. In all experiments, embryos were staged using the Hamburger and Hamilton (HH) system (Hamburger and Hamilton, 1951) and were weighed to confirm that treatment groups were equal in development (Walter and Seebacher, 2009).

Tissue samples (from N=6–8 birds per time point) were collected three days before hatching (referred to as –3), one day before hatching (–1), one day after hatching (+1) and eight days after hatching (+8). Animals were euthanized by cervical dislocation. Breast muscle (musculus pectoralis) and liver were collected, frozen in liquid nitrogen for enzyme and western blot analyses, or harvested in RNAlater (Ambion, Austin, TX, USA) for mRNA analyses.

AMPK phosphorylation has previously been pharmacologically induced via AICAR administration in mammals (5-aminoimidazole-4-carboxamide-1-β-D-ribofuranoside; Toronto Research Chemicals, Toronto, ON, Canada). However, to the best of our knowledge AICAR has not been tested in birds before. Therefore, we tested two AICAR concentrations, which are commonly used in rodent models (Halseth et al., 2002; Winder et al., 2000), to determine whether we could activate AMPK in birds during development. On day 17 of incubation, we injected embryos with 0.5 mg g^–1 bodyweight AICAR (0.5 mg AICAR group; N=8), 1 mg g^–1 bodyweight AICAR (1 mg AICAR group; N=8) or a physiological saline solution (control group; N=8). A hole was made in the eggshell with a needle and all injections were given in the yolk. After the procedure the eggshell was sealed with Parafilm (Parafilm M, Livingston, Rosebery, Australia). Tissue samples were collected one hour after treatment.

For chronic AMPK activation, starting on day 16 of incubation, chicken embryos were injected in the yolk daily [three injections] with 1 mg g^–1 bodyweight of AICAR dissolved in physiological saline (AICAR group; N=16) or with a physiological saline solution (control group; N=16). To avoid an acute effect of AICAR, animals were not injected with AICAR or with the control solution on the day of sample collection. Tissue samples were collected one day before hatching (–1; control group, N=8; AICAR group, N=16) and one day after hatching (+1; control group, N=8; AICAR group, N=16). Oxygen consumption (please see below) of all animals was measured immediately before samples were collected on days –1 and +1. Plasma glucose levels were determined using a glucose analyser (TrueTrack, DiaCare, Diabetes Association of Australia, Willoughby, NSW, Australia) at the time of sample collection.

To induce hypothyroidism, embryos were injected on day 14 of incubation with 1.5 mg methimazole (Walter and Seebacher, 2009) dissolved in physiological saline (hypothyroid group; N=12) or with a physiological saline solution alone (control group; N=12). We established previously that this methimazole treatment significantly reduced thyroid hormone concentrations (Walter and Seebacher, 2009). Tissue samples were collected one day before hatching (–1).

Oxygen consumption of individual hatchlings was measured in an open respirometry system located in a temperature-controlled room. Birds were placed in a gas-tight plastic container (800 ml) fitted with an inlet and outlet for air. Incoming airflow was supplied by a pump (Applied Electrochemistry, Pittsburgh, PE, USA) with a constant flow of 210 ml min^–1 and passed through a spirometer (MK2; AD Instruments, Bella Vista, Australia). The outflowing gases passed through a drying column (MLA0343; AD Instruments, Bella Vista, Australia), and were monitored by a calibrated O₂ gas analyzer (ML205 Gas Analyser; AD Instruments). Data were collected with a computerized recording system (Powerlab 4S, AD Instruments).

Resting oxygen consumption was recorded continuously for 1.5–2 h at thermoneutral ambient temperature (35°C) (Bech et al., 1984; Klaassen et al., 1987). Resting oxygen consumption (RMR; ml h^–1) was defined as the lowest rate of oxygen consumption maintained over 5 min and calculated as the mean of all data points collected over that period.

RNA was extracted from 40–100 mg of liver and muscle samples using TRIreagent (Molecular Research Centre, Cincinnati, OH, USA), following the manufacturer's instructions. RNA quality and concentration were verified using a Bioanalyzer (Agilent Biotechnologies, Palo Alto, CA, USA). One microgram of total RNA was treated with DNase I (Sigma, Castle Hill, NSW, Australia) and reverse-transcribed using RNase H-MMLV reverse transcriptase (Bioscript, Bioline, London, UK) and random hexamer primers (Bioline, London, UK).

Quantitative RT–PCR was performed on an Applied Biosystems 7500 qRT-PCR machine (Applied Biosystems, Scoresby, VIC, Australia) according to published protocols (Seebacher et al., 2006). Primer and dual label probes were designed from sequences obtained from GenBank for AMPK subunits α1, α2, β2, γ1 and γ3, nuclear respiratory factor-1 (NRF1) and peroxisome proliferator activated receptor γ coactivator-1 alpha (PGC-1α). RNA polymerase II polypeptide E (POLR2E) was used as a housekeeping gene (Table 1). We measured the mRNA levels of AMPK subunits α1, β2 and γ1 in liver and of AMPK subunits α2, β2 and γ3 in skeletal muscle, because these genes are expressed predominantly in those tissues, respectively (Proszkowiec-Weglarz and Richards, 2007; Proszkowiec-Weglarz and Richards, 2009). Real-time PCR reactions with dual-labelled probes contained 1 × Immomix (Bioline, London, UK), 4.5 mmol l^–1 MgCl₂, 50–900 nmol l^–1 primer and probe, 1 × ROX reference dye (Invitrogen, USA), and ∼50 ng cDNA. The cycle consisted of 95°C for 7 min, 40 cycles of 95°C for 20 s, 60°C for 1 min. Real-time PCR reactions with Sybr green contained 1 × SensiMixPlus SYBR (Quantace, London, UK), 4.5 mmol l^–1 MgCl₂, 50–900 nmol l^–1 primer and ∼50 ng cDNA. The cycle consisted of 95°C for 7 min, 40 cycles of 95°C for 20 s, 60°C for 1 min. Dissociation curve analysis was performed after the amplification step to verify the presence of only a single PCR product. Relative mRNA concentration of the eight target genes at each stage of development and of AICAR treatments was calculated according to Pfaffl (Pfaffl, 2001), and normalized with POLR2E. As controls, we used adult chicken samples (N=6) to assess changes in mRNA concentration during development, and the grand mean of all samples for each tissue for acute and chronic AMPK activation.

Samples of liver and muscle were homogenized in ice-cold lysis buffer containing 50 mmol l^–1 Hepes, 10% v/v glycerol, 1 mmol l^–1 EDTA, 50 mmol l^–1 sodium fluoride, 5 mmol l^–1 sodium pyrophosphate, 1 mmol l^–1 dithiothreitol, 100 μM PMSF and 1 × complete protease inhibitors (Roche, Castle Hill, Australia) and were left for 1 h at 4°C with gentle mixing. Lysates were cleared by centrifugation (13,000 g, 10 min) and protein content determined with a Bradford assay (BioRad, Hercules, CA, USA). Lysates were mixed with loading buffer containing a final concentration of 20 mmol l^–1 DTT and 2% (w/v) SDS and left to denature at room temperature for 1 h. Aliquots containing 20–40 μg of protein were separated by SDS-PAGE using 10% acrylamide gels and then transferred to PVDF membranes. After blocking (5% skim milk in TBS-0.05% Tween, 60 min at room temperature), membranes were incubated overnight at 4°C with primary antibodies (1:1000 dilution) raised against total or phosphorylated (Thr 172) AMPK α subunit, total or phosphorylated (Ser 79) ACC (all from Cell Signalling Technology, Beverly, MA, USA) or pan-14-3-3 (Santa Cruz Biotechnology, Santa Cruz, CA, USA). Following incubation with the appropriate HRP-conjugated secondary antibodies (1:10,000 dilution), protein bands were visualized using either Western Lightning (Perkin Elmer, Glen Waverley, VIC, Australia) or SuperSignal West Dura (Pierce, Rockford, IL, USA) enhanced chemiluminescent substrates and exposed to film (FujiFilm, Super RX). Signal intensities were determined by densitometric analysis using Odyssey IR imaging system software (LI-COR Biosciences, Lincoln, NE, USA) and normalized for 14-3-3. ACC and pACC were measured as one of the principal targets of AMPK in liver only. Due to antibody limitation, we could not detect total AMPK protein at –3 d in liver. pAMPK protein concentration was measured as a proxy for AMPK activity (Sanders et al., 2007).

We assayed activities of cytochrome c oxidase (COX) and citrate synthase (CS) as indicators of mitochondrial oxidative capacity and abundance, respectively (Hoppeler, 1986). Liver and muscle tissue samples (approximately 50 mg) were homogenized in nine volumes of 50 mmol l^–1 imidazole/HCl, 2 mmol l^–1 MgCl₂, 5 mmol l^–1 EDTA, 1 mmol l^–1 reduced glutathione and 0.1% Triton X-100. All assays were conducted at 38°C, according to published protocols (Seebacher et al., 2003).

Data are presented as means ± s.e.m. mRNA and protein concentration data for different stages of development were analyzed by one-way ANOVA followed by LSD pairwise comparison. We compared acute AICAR, chronic AICAR, and hypothyroid treatments with their respective control treatments by two-tailed t-tests for independent samples, except that we used a one-tailed t-test when we explicitly tested for an increase in PGC-1α mRNA concentration and in heat production with AICAR treatment. All data were tested for homogeneity of variances using Levene's test. Data were ln transformed when Levene's test was significant. In cases where transformed data also failed Levene's test, we used Kruskal–Wallis one-way ANOVA on ranks. The significance was considered as P<0.05.

In liver, the mRNA concentrations of the AMPKα1 and β2 subunits were significantly elevated before hatching (–3) compared with at later developmental stages (α1, F_3,20=3.911, P<0.03; Fig. 1A; β2, F_3,20=7.138, P<0.003; Fig. 1B). By contrast, AMPK γ1 mRNA levels increased significantly after hatching (+1) in liver (F_3,20=8.649, P<0.001; Fig. 1C). Phosphorylation of AMPK was elevated after hatching (+8) compared with at earlier developmental stages (H_3,28=8.703, P<0.05; Fig. 1D), paralleling an increase in total AMPK protein content across this developmental period (F_3,28=3.068, P<0.05; Fig. 1E). Phosphorylated and total ACC protein content was also elevated after hatching (+8) compared with at earlier developmental stages (pACC, H_3,28=25.61, P<0.0001; Fig. 1F; total ACC, H_3,28=20.91, P<0.0002; Fig. 1G).

In skeletal muscle, the mRNA concentration of the AMPK α2 subunit remained constant (F_3,20=0.28, P>0.8; Fig. 2A). However, AMPK β2 and γ3 mRNA levels increased significantly after hatching (β2, F_3,20=9.43, P<0.001; Fig. 2B; γ3, F_3,20=41.66, P<0.0001; Fig. 2C). There was no significant change in pAMPK or total AMPK concentration during development (pAMPK, F_3,28=2.446, P>0.08; Fig. 2D; total AMPK, F_3,28=0.849, P>0.4; Fig. 2E).

In liver, a single dose of 1 mg g^–1 AICAR significantly increased AMPK phosphorylation compared with control and 0.5 mg g^–1 AICAR groups (F_2,23=18.575, P<0.001; Fig. 3A). PGC-1α and NRF-1 mRNA levels remained unchanged after AICAR treatment (1 mg g^–1) compared with the control group (PGC-1α, t₁₀=–1.152, P>0.1; NRF-1, t_5.882=0.831, P>0.2; data not shown).

In skeletal muscle, neither concentration of AICAR changed AMPK phosphorylation. Also, total AMPK concentration remained constant (pAMPK, F_2,23=0.816, P>0.4; Fig. 3B; total AMPK, F_2,23=2.352, P>0.1; Fig. 3C). As in liver, the mRNA concentration of PGC-1α and NRF-1 remained unchanged after AICAR treatment (1 mg g^–1) compared with the control group (PGC-1α, t₁₀=–0.869, P>0.2; NRF-1, t₁₀=0.131, P>0.4; data not shown).

The body mass of embryos one day before hatching (day –1; AICAR, 48.53±1.05 g; control, 51.42±2.52 g) and one day after hatching (day +1; AICAR, 49.16±1.33 g; control, 47.36±0.77 g) was not significantly different between AICAR and control groups (embryos, t_9.410=1.057, P>0.1; hatchlings, t_12.817=–1.179, P>0.1). AICAR treatment caused a significant decrease in plasma glucose in embryos (t₁₅=2.633, P<0.02; Fig. 4) and in hatchlings (t₁₆=3.333, P<0.005; Fig. 4).

In liver, chronic AICAR treatment increased pAMPK compared with the control, to a level that only reached statistical significance in hatchlings (–1, t₁₄=1.6, P>0.1; Fig. 5A; +1, t₁₄=3.42, P<0.005; Fig. 5C). Total AMPK concentration did not change with treatment either before (t₁₄=1.32, P>0.1; Fig. 5B) or after (t₁₄=0.49, P>0.1; Fig. 5D) hatching. In skeletal muscle, chronic AICAR treatment significantly increased AMPK phosphorylation before (t₁₄=2.66, P<0.03; Fig. 5E) and after (t₁₄=2.78, P<0.02; Fig. 5G) hatching, with total AMPK concentration remaining constant (–1d, t₁₄=–0.91, P>0.1; Fig. 5F; +1d, t₁₄=–0.72, P>0.1; Fig. 5H).

Chronic AMPK activation significantly increased heat production before (t₁₅=–3.888, P<0.002; Fig. 6A) and after (t₁₀=–2.266, P<0.05; Fig. 6B) hatching compared with the control groups. In liver, chronic AICAR treatment had no effect on citrate synthase (CS) activity before hatching (t₁₄=–0.582, P>0.1; Fig. 6C), but significantly increased CS activity after hatching compared with the control group (t₁₄=2.772, P<0.02; Fig. 6C). In skeletal muscle, chronic AMPK activation decreased CS activity before hatching (t₁₄=–2.176, P<0.05; Fig. 6D), but CS activity increased after hatching in the AICAR treatment group compared with the control group (t_9.675=5.304, P<0.001; Fig. 6D). Cytochrome-c oxidase (COX) activity remained unchanged after AICAR treatment compared with the control group before and after hatching in liver (before, t₁₄=–1.698, P>0.1; after, t₁₄=0.736, P>0.1; data not shown) and in muscle (before, t_8.216=1.436, P>0.1; after, t₁₄=0.012, P>0.1; data not shown).

Chronic AMPK activation had no effect on PGC-1α mRNA concentrations before hatching in liver (t₁₄=0.777, P>0.1; Fig. 7A) or in muscle (t₁₄=–0.976, P>0.1; Fig. 7B). By contrast, after hatching PGC-1α mRNA concentration was significantly elevated in the AICAR group compared with the control group in muscle (t₁₄=2.536, P<0.01; Fig. 7B) and in liver (t₁₄=1.78, P<0.05; Fig. 7A). NRF-1 mRNA levels did not change with AICAR treatment in liver or muscle before (liver, t₁₄=–0.797, P>0.1; Fig. 7C; muscle, t₁₄=0.148, P>0.1; Fig. 7D) or after (liver, t_9.69=1.19, P>0.1; Fig. 7C; muscle, t₁₄=–0.88, P>0.1; Fig. 7D) hatching.

In liver, hypothyroidism significantly reduced AMPK phosphorylation (t₁₄=3.090, P<0.009; Fig. 8A), despite an increase in total AMPK concentration in the methimazole treatment group compared with the control group (t₁₄=–2.990, P<0.02; Fig. 8B). In skeletal muscle, there was no significant difference between treatment groups in AMPK phosphorylation and total AMPK protein content (pAMPK, t₁₄=–0.816, P>0.4; Fig. 8C; total AMPK, t₁₄=–0.0138, P>0.9; Fig. 8D).

The development of endothermy during ontogenesis is accompanied by increasing ATP demands but reduced ATP synthesis efficiency (Walter and Seebacher, 2009). We show that AMPK plays a role in this process, and that increased phosphorylation of AMPK is associated with increased PGC-1α concentrations and increased mitochondrial metabolic capacities. The increased mitochondrial abundance that is known to accompany the ontogenetic transition from ectothermy to endothermy can therefore be at least partly explained by the increase in AMPK phosphorylation and subsequent increase in PGC-1α gene expression, a known inducer of mitochondrial biogenesis. These AMPK-mediated changes can contribute to the increased blood glucose utilization and heat production shown here. In addition, AMPK activity in liver is induced by thyroid hormone.

Both liver and muscle are important sites for metabolic heat production in endotherms (Rolfe and Brand, 1996; Rolfe et al., 1999; Ueda et al., 2005). Additionally, both have a high demand for ATP to sustain numerous housekeeping functions, such as gluconeogenesis in liver and contractile function in muscle. The increase in AMPK phosphorylation with development reflects the increasing demand for greater metabolic capacity to sustain both ATP-consuming functions and growth on the one hand, and heat production on the other. In skeletal muscle, the increase in AMPK phosphorylation after hatching paralleled that of liver, but the increase was not significant (P=0.08). This non-significant result is due to the high variability among samples, which might reflect differences in the timing of the increase in muscle metabolic capacities between hatchlings. As a result of this variability, our sample size was too small to detect a statistical difference at P<0.05. The mRNA concentrations of all subunits changed during development, except for AMPKα2 in muscle. However, there is no direct correspondence between mRNA and protein concentrations, which indicates that the assembly of the quaternary structure is not regulated by the mRNA concentration of any one subunit.

Mitochondrial proliferation is one of the key elements in the transition from ectothermy to endothermy, which is tightly linked to metabolic capacity (Seebacher et al., 2006). We show that, during the ontogenetic transition from ectothermy to endothermy in birds, activation of AMPK induces an increase in PGC-1α mRNA concentration that is paralleled by an increase in mitochondrial content (CS activity) and heat production. These findings suggest that the previously observed increase in PGC-1α and its target PPARγ in liver and skeletal muscle during the embryonic development of birds (Walter and Seebacher, 2007) is mediated by AMPK, and that AMPK plays a leading role in mitochondrial biogenesis by regulating PGC-1α expression. AMPK regulates PGC-1α expression either directly by phosphorylating PGC-1α (Jager et al., 2007), or indirectly by activating SIRT1. SIRT1 is a NAD1-dependent type III deacetylase (Canto et al., 2009; Nemoto et al., 2005), which stimulates the PGC-1α-dependent induction of the PGC-1α promoter (note that PGC-1α can regulate its own promoter). PGC-1α interacts with a suite of transcription factors, such as NRF-1 and NRF-2, to control mitochondrial biogenesis (Finck and Kelly, 2006; Scarpulla, 2008). NRF-1 mRNA was not affected by AICAR treatment, which indicates that AMPK does not influence NRF-1 gene expression. The significant decrease in muscle CS activity of control animals after hatching is likely to reflect hypertrophy and the consequent decrease in mitochondrial density per unit tissue of muscle cells that is associated with the onset of locomotion. The decrease in CS activity disappears four days after hatching (Walter and Seebacher, 2007), indicating that mitochondrial proliferation follows hypertrophy.

There are strong similarities between the action of AMPK during bird development and its role in metabolic regulation in response to exercise and diet in mammals (Park et al., 2002; Viollet et al., 2009b). Hence, in mammals AMPK is known to promote mitochondrial biogenesis by interacting with PGC-1α, NRF-1, NRF-2 and mitochondrial transcription factor A in response to exercise or metabolic stress (Viollet et al., 2009a). In the liver of mammals, AMPK regulates cellular energy homeostasis by controlling mitochondrial biogenesis (Guigas et al., 2007). Hepatocytes from AMPKα1α2_LS^–/– mice, which lack both α1 and α2 AMPK catalytic subunits, show a decreased oxidative capacity that is linked to a decreased cellular density of mitochondria as a result of decreased expression of PGC-1α, COX subunit I (COXI) and Fo/F1 ATPase (Guigas et al., 2007). Similarly, non-exercised red and white gastrocnemius muscles of AMPKα2^–/– mice show a 20% reduction in mitochondrial markers such as COXI protein content and CS activity (Jorgensen et al., 2007). Interestingly, a double knockout of catalytic AMPK subunits (AMPKα1 and AMPKα2) in embryonic mice is lethal at ∼10.5 days post-conception (Viollet et al., 2009a), which demonstrates the importance of AMPK in controlling energy metabolism during development. Additionally, AMPK has been suggested to mediate normal metabolic development of human neonates, and in particular the switch from glycolytic to oxidative metabolism, by regulating glucose and fatty acid metabolism (Brauner et al., 2006). These data together with our findings show that there are similarities in the pathways that regulate the ontogenetic transition from ectothermy to endothermy in birds and mammals. However, unlike in mammals, NRF-1 in birds does not respond to AICAR treatment and it is therefore unlikely to be involved in the AMPK-mediated metabolic responses.

In addition to its function in modulating mitochondrial biogenesis, AMPK controls the coordination of lipid metabolism by balancing fatty acid oxidation with long-chain fatty acid synthesis (Viollet et al., 2009b). In chicken, lipogenesis occurs only after hatching, because the embryo directly derives fatty acids from yolk lipids (Ryder, 1970). The increase in ACC indicates increased lipogenesis in the liver (Carling et al., 1987) of hatchlings that is inhibited by increasing AMPK phosphorylation, which thereby balances fat storage with ATP and heat production in mitochondria.

The decrease in blood glucose levels after AICAR treatment shows, firstly, an involvement of AMPK on whole-body glucose homeostasis during development and, secondly, the increasing energy demands caused among others by elevated mitochondrial biogenesis. Also, AICAR might possibly affect other AMP-sensitive enzymes, such as fructose-1, 6-bisphosphatase, that influence blood glucose levels independently of AMPK (Vincent et al., 1991). However, in mammals AMPK regulates glucose uptake. For example, in the liver of AMPKα1α2_LS^–/– mice, up to 40% of AICAR-induced hypoglycemia is due to the activation of AMPK (Viollet et al., 2009b). Hepatic glucose production is the primary mechanism for regulating glucose flux at rest and, in mammals, liver AMPK is important in maintaining whole-body glucose homeostasis by inhibiting hepatic glucose production if energy demands increase (Foretz et al., 2005; Viollet et al., 2009a). If circulating glucose levels do not meet metabolic energy demand, the liver releases glucose by glycogenolysis, and in the longer term by gluconeogenesis (Hegarty et al., 2009). Furthermore, because the growing embryo has only a limited energy supply (the yolk), decreasing blood glucose levels cannot be replenished. AMPK plays a role in gluconeogenesis because of its capacity to control the expression of some key gluconeogenic genes (e.g. PEPCK and G6Pase) by phoshorylating the CREB co-activator TORC2 (Koo et al., 2005; Viollet et al., 2009b). Similarly, in muscle AMPK stimulates beta oxidation and glucose utilization, which leads to an increase in metabolic capacity. Furthermore, AMPK regulates skeletal muscle capillarization and VEGF expression (Zwetsloot et al., 2008), which assures sufficient vascularization to provide substrates to sustain increased metabolic activity (Adair, 2005).

Thyroid hormones are known to play an important role in regulating the mitochondrial metabolism and internal heat production of endotherms (Lowell and Spiegelman, 2000; Silva, 1995). This effect is at least partly due to increased fatty acid oxidation via the AMPK-mediated inhibition of ACC (de Lange et al., 2008; Lombardi et al., 2009). Here we show that, in addition to blocking ACC, thyroid hormone also alters the phosphorylation state of AMPK and thereby causes an increase in PGC-1α gene expression during development. We have previously shown that T3 regulates mitochondrial function and capacity via PGC-1α during the development of endothermy (Walter and Seebacher, 2009). Those results can now be explained by a direct genomic effect of thyroid hormone on PGC-1α gene expression and by its effect via AMPK phosphorylation.

In conclusion, even though modern endotherms, mammals and birds, are phylogenetically distinct, AMPK signalling appears to be a common and conserved molecular mechanism that controls metabolic heat production. Our data suggest that, as in mammals, the ontogenetic transition from ectothermy to endothermy in birds is controlled by thyroid hormones that stimulate AMPK to induce PGC-1α expression. Consequently, as in mammals, this kinase plays a role in regulating mitochondrial biogenesis. Thus, increases in mitochondrial content lead towards an enhanced ATP production to support the growing organism, as well as to increased internal heat production. It will be interesting for future research to investigate the importance of AMPK in controlling energy demands, particularly in regulating Na⁺/K⁺-ATPase activity, that might drive increased metabolic flux through mitochondria (Walter and Seebacher, 2009). In mammalian lung epithelial cells, AMPK affects Na⁺/K⁺-ATPase activity if pharmacologically activated with metformin or AICAR under hypoxic conditions (Gusarova et al., 2009; Woollhead et al., 2007). If this interaction were generally true, AMPK would affect and control the main components of endothermy.

 
• ACC

  acetyl CoA carboxylase

 
• AICAR

  5-aminoimidazole-4-carboxamide-1-β-D-ribofuranoside

 
• AMPK

  AMP-activated protein kinase

 
• COX

  cytochrome c oxidase

 
• CS

  citrate synthase