The present study examined whether administration in vivo of a maximally stimulating dose of growth hormone (GH) was capable of modulating selected aspects of the GH–insulin-like growth factor (IGF) system to the same extent in alcohol-fed and control animals. Rats were maintained on an alcohol-containing diet for 14 weeks, while control animals were fed isocalorically. After surgical implantation of a catheter in the carotid artery, rats were starved overnight. The next morning, rats were injected with recombinant human GH (500 μg/kg, s.c.) or an equal volume of saline at time 0 and 12 h. Blood samples were collected prior to GH and at 6, 12 and 24 h thereafter, tissues were collected at the end of the study. Time-matched control and alcohol-fed rats not receiving GH were also included. Although the plasma concentrations of both total and free IGF-I were decreased 30–40% in alcohol-fed rats, the ability of GH to elevate circulating IGF-I was not diminished. GH was equally effective at increasing IGF-I peptide levels in both liver and skeletal muscle. GH also produced comparable increases in IGF-I mRNA in muscle in both groups. Hepatic GH receptor (GHR) peptide levels were not significantly altered by either alcohol or GH. Alcohol feeding decreased plasma levels of IGF binding protein (IGFBP)-3 and increased IGFBP-1, and GH did not significantly alter this profile. Hepatic expression of suppressor of cytokine signalling (SOCS-3) mRNA was not different between the groups. However, SOCS-3 mRNA was increased by ~50% in control animals in response to GH, but remained unchanged in alcohol-fed rats. These data indicate that the decrease in hepatic IGF-I synthesis and plasma IGF-I observed in alcohol-fed rats was independent of a change in GHR levels. In contrast, the ability of a maximally stimulating dose of GH to modulate selected biological responses in vivo was not impaired by chronic alcohol consumption and was associated with a lack of a GH-induced increase in SOCS-3 mRNA. Acute alcoholism.
Growth hormone (GH) is generally considered to be an anabolic hormone. However, elevated levels of GH, resulting from endogenous overproduction or exogenous administration, can lead to a diabetic-like condition associated with severe insulin resistance ( Scanes, 1995 ). The presence of an adequate circulating concentration of GH is necessary for normal accretion of muscle protein in children and the maintenance of lean body mass in adults ( Florini et al., 1996 ). Although GH can directly influence various aspects of muscle metabolism, the majority of its effects appear to be mediated indirectly via enhanced synthesis and secretion of insulin-like growth factor (IGF)-I ( Daughaday, 1989 ). Previous studies have demonstrated that GH administration in vivo increases IGF-I mRNA expression in liver, which is primarily responsible for the increase in blood-borne IGF-I ( Schwander et al., 1983 ). GH also increases IGF-I in peripheral tissues, such as skeletal muscle ( Pell and Bates, 1992 ).
Most studies have shown that chronic alcohol consumption, in rodents or humans, decreases the circulating concentration of IGF-I ( Sonntag and Boyd, 1988 , Soszynski and Frohman, 1992 , Santolaria et al., 1995 , Lang et al., 1998 ). Because decreases in IGF-I have been correlated with concomitant reductions in muscle protein synthesis ( Lang et al., 1996 ), the reduction in IGF-I has been postulated to be responsible, at least in part, for the alcohol-induced myopathy ( Lang et al., 1998 ). The decrease in circulating IGF-I observed following chronic alcohol consumption could be caused by a decrease in plasma GH levels. In this regard, chronic alcohol feeding has been shown to inhibit spontaneous pulsatile GH secretion ( Soszynski and Frohman, 1992 , Badger et al., 1993 ). Alternatively, or in addition, alcohol might decrease IGF-I synthesis by producing a GH-resistant condition. Other catabolic conditions, including infection, thermal injury and diabetes ( Dahn et al., 1988 , Belcher and Ellis, 1990 , Day et al., 1998 ), may result in hepatic GH resistance, as evidenced by the diminished ability of maximally stimulating doses of GH to increase plasma IGF-I levels. In this regard, a recent study by Assy et al. (1997) reported that patients with liver cirrhosis had a severely blunted increase in serum IGF-I in response to GH administration, compared to age-matched healthy controls. Moreover, in vitro GH-stimulated IGF-I secretion was attenuated when liver slices were incubated with high levels (175 mM) of ethanol ( Xu et al., 1995 ). However, neither of these studies adequately mimic the pathological condition produced by chronic alcohol consumption in rodents where blood levels of IGF-I are decreased. In this model, alcohol-fed rats typically demonstrate only moderate elevations in blood alcohol (~50 mM) and mild signs of hepatic injury. Therefore, the purpose of the present study was to determine whether chronic alcohol consumption alters the ability of a maximally stimulating dose of GH administered in vivo to modulate IGF-I and other components of the GH–IGF axis.
METHODS AND MATERIALS
Specific pathogen-free, male Sprague–Dawley rats (Charles River Breeding Laboratories, Cambridge, MA, USA) were maintained on an ethanol-containing agar block diet for 14 weeks ( Bautista, 1997, 1998 , Lang et al., 1998 ). Initially, all rats were provided agar without alcohol for 2 days. Thereafter, half of the animals were fed agar containing 10% alcohol, while the others were provided agar containing an equal caloric amount of maltose–dextrin. The ethanol content of the agar was increased to 20% and finally to 30%, at 1-week intervals. Rats were maintained on the 30% ethanol–agar block for the remainder of the experimental protocol. Agar blocks containing alcohol were made fresh daily. Calculated ethanol intake during the final 10 weeks of the experimental protocol averaged 15 ± 2 g/kg/day (105 ± 14 kcal/kg/day). Control animals were provided the same amount of agar, but the alcohol content was substituted isocalorically with maltose–dextrin.
The agar block provides a source of water and/or ethanol for the animals, but has nominal nutritional value. The nutrient intake for both groups of animals was supplied by consumption of standard solid rodent chow (Harlan Teklad #8604, Madison, WI, USA). The alcohol-fed rats were permitted solid chow ad libitum. Control animals were provided the same amount of solid chow as that consumed by the alcohol-fed group. Over the final 10 weeks, alcohol-fed rats had an average total energy consumption of 261 ± 19 kcal/kg/day, which is sufficient to meet the nutritional requirements of growing rats ( National Research Council, 1995 ). This energy intake did not differ from that of control rats (254 ± 17 kcal/kg/day). The nutrient intakes (except for ethanol) of both groups were also essentially identical since the sole source of nutritional support was derived from solid chow (~24% protein, 5% fat and 71% carbohydrate). The total caloric intake and the amount of ethanol consumed by the rats used in this study were comparable to those reported for other rodent models of chronic alcohol feeding ( Lieber and DeCarli, 1989 , Badger et al., 1993 ).
The day prior to the experiment, rats were anaesthetized with an intramuscular injection of ketamine and xylazine (90 and 9 mg/kg, respectively), and sterile surgery was performed to implant a catheter in the arch of the aorta via the left carotid artery. Animals were returned to individual cages and were then starved overnight. The next morning, rats were injected subcutaneously with either recombinant human (rh) GH (500 μg/kg Serostim, Sorono Laboratories, Norwell, MA, USA) or an equal volume (250 μl) of vehicle at time 0 and 12 h. rhGH was used in the current study because of its greater availability and because preliminary studies indicated that, at the dose used, there was no significant difference in the IGF-I response in rats injected with rhGH or rat GH (data not shown). In both control and alcohol-fed rats an arterial blood sample (1 ml) was obtained prior to the first GH injection and at 6, 12 and 24 h thereafter. Blood samples of comparable volume and number were obtained from time-matched control and alcohol-fed rats injected with saline (no GH). After the final blood sample, animals were anaesthetized with pentobarbital, and liver (left lateral lobe) and skeletal muscle (gastrocnemius) rapidly excised. Tissues were rinsed thoroughly of blood, blotted dry and immediately frozen between liquid nitrogen-cooled aluminum blocks. All tissues were stored at –80°C until analysis. These tissues were used to quantify the peptide content and mRNA expression for various components of the IGF–GH axis.
Animals in this first study had an arterial catheter implanted so that serial blood samples could be obtained in order to detect any difference in the temporal progression of IGF-I in response to GH between groups. We cannot exclude the possibility that surgical implantation of the catheter, which results in a mild stress response, may have influenced GH action. However, both control and alcohol-fed rats were treated similarly. It is also noteworthy that all animals were starved during the period of time GH action was assessed. Starved rats were used for two reasons: (1) preliminary studies indicated that there was a larger and more consistent increase in the GH-induced increment in IGF-I in starved, compared to fed, rats, (2) we were particularly interested in whether prolonged alcohol ingestion, not necessarily the presence of alcohol per se in the blood, produced a sustained change in GH action. Therefore, this study did not directly assess whether GH action was altered when detectable levels of blood alcohol were present at the time of GH administration, and therefore could not exclude changes induced by alcohol withdrawal.
In order to address one of above-mentioned caveats, an additional study was performed. In this second protocol, naive (no prior surgery) overnight-starved rats were injected intraperitoneally with either 75 mmol/kg of ethanol or an equal volume (4 ml/kg) of saline. Thirty minutes later, half of the rats in each group received a single subcutaneous injection of rhGH (500 μg/kg), whereas the remainder of the animals were injected with saline. Rats were anaesthetized with pento-barbital 12 h after injection of GH and blood was collected into a heparinized syringe from the abdominal aorta for the measurement of total IGF-I. This study was performed to determine whether the presence of alcohol in the circulation alters the GH-induced increase in IGF-I. This model of acute alcohol intoxication is well-characterized and was chosen because of its reported ability to impair protein synthesis in liver and skeletal muscle ( Tiernan and Ward, 1986 , Preedy et al., 1988 ). All experiments were approved by the Animal Care and Use Committee at the Pennsylvania State College of Medicine and adhere to the National Institutes of Health guidelines for the use of experimental animals.
The concentration of total IGF-I in plasma was determined using a modified acid–ethanol (0.25 M HCl:87.5% ethanol) extraction procedure with cryo-precipitation, and tissues were processed using acid homogenization and Sep-pak (C-18) extraction ( Lang et al., 1996, 1998 ). The efficacy of this method in removing IGF binding proteins has been previously reported ( Frost et al., 1996b ). The tissue eluate was evaporated and the dried sample was reconstituted with radioimmunoassay (RIA) buffer containing 0.25% bovine serum albumin (BSA) for IGF-I determination. IGF-I in plasma and tissues was determined by RIA. Recombinant human [Thr
]IGF-I was used for iodination and standards (Genetech, South San Francisco, CA, USA). The ED
for this assay is 0.03–0.08 ng/tube. IGF-I was determined on all plasma samples.
The plasma concentration of free IGF-I was determined by centrifugal ultrafiltration, as originally described by Frystyk et al. (1994). Briefly, the plasma samples were diluted 1:5 with Krebs–Ringer bicarbonate buffer (pH 7.4, with 5% BSA) and prefiltered through a 0.22 μm filter (Millex-GV, Millipore, Molsheim, France) to remove debris. The prefiltered samples were then added to Amicon YMT 30 membranes and MPS-1 supporting devices (Amicon Division, W. R. Grace, Co., Beverly, MA, USA) and centrifuged at 300 g (1500 r.p.m.) at 37°C for up to 100 min. The ultrafiltrate was collected after 40–100 min of centrifugation and used for the IGF-I RIA.
RNA extraction and Northern blotting
Total RNA was isolated from liquid nitrogen-frozen gastrocnemius and liver using TRI Reagent TR-118 as outlined by the manufacturer (Molecular Research Center, Inc., Cincinnati, OH, USA). Samples (20 μg) of total RNA were run under denaturing conditions in 1% agarose/6% formaldehyde gels. The running buffer was 1× HEPES. Northern blotting occurred via capillary transfer to Zeta-Probe GT blotting membranes (Bio-Rad Laboratories, Hercules, CA, USA). An 800 bp probe from rat IGF-I (Peter Rotwein, Portland, OR, USA), a 600 bp probe from rat growth hormone receptor (GHR, Lawrence Mathews, Ann Arbor, MI, USA), and a 681 bp from murine suppressor of cytokine signalling (SOCS-3, Douglas J. Hilton, Victoria, Australia) were labelled using a Random Primed DNA Labeling kit (Roche Molecular Biochemicals, Indianapolis, IN, USA). For normalization of RNA loading, a rat 18S oligonucleotide was radioactively end-labelled using polynucleotide kinase (Amersham Pharmacia Biotech, Piscataway, NJ, USA). Membranes were pre-hybridized and hybridized at 42°C in 50% formamide/6× SSPE/5× Denhardt's/1% sodium dodecyl sulphate (SDS)/10% dextran sulphate/herring testis DNA (100 μg/ml). All membranes were washed at room temperature twice in 2× SSC/0.1% SDS for 5 min and once in 0.1× SSC/0.1% SDS for 15 min. Additionally, membranes hybridized with rat IGF-I or GHR were washed at 65°C in 0.1× SSC/0.1% SDS for 15–30 min. All data were normalized to ribosomal 18S RNA. Finally, membranes were exposed to a phosphoimager screen and the resultant data quantified using ImageQuant software (Molecular Dynamics, Sunnyvale, CA, USA).
Alcoholism post acute withdrawal
Frozen liver was powdered in a mortar pre-cooled with liquid nitrogen and GHR peptide levels determined by Western blot analysis. Briefly, ~80 μg of liver were added to 7 vol of ice-cold homogenization buffer containing (in mM) HEPES 20 (pH 7.4), EGTA 2, NaF 50, KCl 100, EDTA 0.2, β-glycerophosphate 50, DTT 1, PMSF 0.1, benzamidine 1, and sodium vanadate 0.5. Tissue was homogenized and centrifuged at 10 000 g for 10 min at 4°C. The supernatant was then mixed with an equal volume 2× Laemmli SDS buffer. Samples were boiled for 3 min and centrifuged at 13 000 r.p.m. at room temperature. Equal amounts of protein were then electrophoresed in an 8% polyacrylamide gel. After electrophoresis, proteins in the gel were transferred to nitrocellulose. After blocking for 1 h with non-fat milk (5% w/v) in 25 mM Tris (pH 7.6) containing 0.9% (w/v) saline, membranes were washed extensively in Tris–saline containing 0.01% (w/v) Tween 20. The nitrocellulose was incubated overnight at 4°C with a polyclonal antibody (1:200 dilution) which recognizes the GHR (WR Baumbach, American Cyanamid, NJ, USA). Thereafter, membranes were washed with Tris–saline–Tween (0.1%) for 15 min. Antigen–antibody complexes were identified with goat anti-rabbit IgG tagged with horseradish peroxidase (Sigma, St Louis, MO, USA), and exposed to the enhanced chemiluminescence (ECL) detection system (Amersham, Arlington Heights, IL, USA) for 1 min and to X-ray film for various periods of time. The GHR band migrated with an apparent molecular weight of 100 kDa ( Sadeghi et al., 1990 ). Bands were scanned (Microtek ScanMaker IV) and quantified using NIH Image 1.6 software. Representative samples from all experimental groups were electrophoresed on the same gel, data are expressed as a percentage of the control value.
Plasma samples were separated on a 12.5% SDS–polyacrylamide gel electrophoresis (PAGE) gel under non-reducing conditions. Separated proteins were electro-blotted onto nitrocellulose, and blocked for 2 h at room temperature with Tris-buffered saline containing 1% non-fat dry milk. The membranes were then incubated with a 1:2000 dilution of antiserum against rat IGFBP-1 at room temperature for 2 h. Antigen–antibody complexes were identified with goat anti-rabbit IgG tagged with horseradish peroxidase and exposed to the ECL detection system for 1 min and to X-ray film for 10–30 s. Bands were quantified as described above for the GHR.
IGF binding protein (IGFBP)-3 in plasma was determined by Western ligand blot analysis, as described by Hossenlopp et al. (1986) and slightly modified by our laboratory ( Frost et al., 1996a , Lang et al., 1996, 1998 ). Samples were subjected to SDS–PAGE without reduction of disulphide bonds. The electrophoresed proteins were transferred onto nitrocellulose in Tris–methanol–glycine buffer. Nitrocellulose sheets were washed and then incubated overnight with radiolabelled IGF-I. The nitrocellulose sheets were washed extensively in Tween 20, dried, and autoradiographed with X-ray film (Kodak X-Omat AR, Eastman Kodak Co., Rochester, NY, USA) and intensifying screens (DuPont, Wilmington, DE, USA) at –70°C for 2–4 days.
Plasma glucose, alcohol, hormone and amino acid concentrations
The plasma glucose and alcohol concentrations were determined using a rapid analyser (GL5, Analox Instruments, Lunenburg, MA, USA). The plasma concentration of insulin was determined by rat-specific RIA (Linco, St Louis, MO, USA). Aspartate aminotransferase (AST, 220.127.116.11) activity in plasma was determined using a standard enzymatic assay (Sigma). Plasma was deproteinized with sulphosalicylic acid (5% w/v) and the supernatant used for amino acid analysis by ion-exchange high-power liquid chromatography (model 6300, Beckman Instruments, Fullerton, CA, USA). Absorbance was measured at 440 and 570 nm after post-column ninhydrin treatment.
An aliquot of powdered liver was extracted in cold perchloric acid (PCA), neutralized and used for determination of adenosine triphosphate (ATP), by standard fluorometric methods.
Experimental values are presented as means ± SEM. The number of rats per group is indicated in the legends to the figures and tables. For the plasma IGF-I concentration, the area under the curve was calculated using the trapezoidal solution after the mean time-matched value from either the saline-injected control or alcohol-fed animals was subtracted from the respective values after the injection of GH. Data were analysed using analysis of variance followed by the Newman– Keuls test to determine treatment effect. Statistical significance was set at P <, 0.05.
Prior to being placed on the alcohol or control diet, the body weight of all animals averaged (± SEM) 145 ± 5 g. The body weight of rats in both groups increased gradually over the 14-week protocol. However, at the time experiments were performed, body weight was 8% lower in animals fed the alcohol-containing diet (498 ± 13 g), compared to pair-fed control animals (542 ± 11 g, P <, 0.05).
Plasma concentrations of alcohol, AST, glucose, insulin and ATP
Plasma alcohol levels averaged 51 ± 22 mg/dl in alcohol-fed rats at the time of surgery. The next morning, after an 18-h fast, alcohol levels were not detectable. Plasma alcohol levels were also undetectable in control animals at all time points examined.
As illustrated in Table 1, there was no significant difference in the plasma glucose concentration between control and alcohol-fed rats, and there was no detectable influence of GH on this parameter. Similarly, plasma insulin levels were not different between the groups and not influenced by GH. These data suggest that the relatively short-term administration of GH employed in the current study does not produce an overt insulin resistance, as observed after more chronic GH administration ( Scanes, 1995 ). Furthermore, these data also suggest that a secondary change in insulin (or glucose) is an unlikely mediator of any of the GH-induced responses described below.
Alcohol-induced hepatic injury was assessed by measuring plasma transaminase activity (Table 1 ). AST levels were moderately elevated by ~90% in alcohol-fed rats, compared to control animals. A similar elevation in AST was still present following GH treatment, suggesting that GH per se did not exacerbate the mild hepatic injury observed in alcohol-fed rats.
There was no significant difference in the hepatic ATP concentration between control and alcohol-fed rats (Table 1 ). Moreover, injection of GH also did not influence ATP in this tissue in either group of animals.
Because chronic GH treatment is known to cause water retention ( Moller et al., 1991 ), the water content of liver and the gastrocnemius muscle was determined. There was no significant difference in the water content (as determined by the wet-to-dry weight ratio) for either liver or muscle between control and alcohol-fed rats (data not shown).
Plasma and tissue IGF-I
The concentration of total IGF-I in plasma was reduced 30% in alcohol-fed rats, compared to pair-fed control animals (Fig. 1, top panel). Both control and alcohol-fed rats demonstrated a significant GH-induced increase in total IGF-I at the 12- and 24-h time points, compared to either their own pre-values or compared to time-matched values from animals injected with saline. The area under the curve for total IGF-I was 29% greater in alcohol-fed rats, than in control animals (Fig. 1, bottom panel).
The circulating concentration of free IGF-I was decreased 41% in alcohol-fed rats, compared to values from time-matched control animals (Fig. 2 ). Despite the lower starting concentration, GH increased free IGF-I levels to the same extent in both control and alcohol-fed rats. The relative amount of free IGF-I, compared to total IGF-I, did not differ between alcohol-fed (9.1 ± 1.2%) and control (10.6 ± 1.1%) rats, and was not altered by GH administration (10.3 ± 1.7% and 11.0 ± 1.6%, respectively). Free IGF-I levels were only quantified in the final blood sample.
The IGF-I peptide content was determined in tissues obtained 12 h after the second injection of GH (Fig. 3 ). Hepatic IGF-I content in alcohol-fed rats was reduced by 39%, compared to control values. Injection of GH increased hepatic IGF-I in both groups to a similar extent (75–100 ng/g tissue). The IGF-I content in skeletal muscle tended to be decreased in alcohol-fed rats (18%), but this change did not reach statistical significance. Again, in response to GH administration, IGF-I peptide was increased to the same extent in both groups of animals.
Hepatic IGF-I mRNA expression was reduced 31% in alcohol-fed rats, compared to control animals (Fig. 4 ). Administration of GH did not alter hepatic IGF-I mRNA in either group at the time point examined. In contrast, the abundance of IGF-I mRNA was not decreased in muscle from alcohol-fed rats, compared to control animals. However, GH increased IGF-I mRNA abundance in skeletal muscle by ~50% in both groups of animals (Fig. 5 ).
GH receptor mRNA and peptide
Chronic alcohol consumption resulted in a 27% reduction in the expression of GHR mRNA in liver, compared to control values (Fig. 6 ). GH administration in control animals reduced hepatic GHR mRNA levels by 40%. In contrast, no GH-induced reduction in GHR mRNA was detected in alcohol-fed rats. GHR mRNA levels in muscle were not significantly altered by either alcohol feeding and/or treatment with GH (data not shown).
Hepatic GHR peptide levels are illustrated in Fig. 7 and, in general, did not show the same alcohol- or GH-induced changes as observed for GHR mRNA. There was no difference in the relative GHR peptide levels between control and alcohol-fed rats. GHR peptide levels tended to increase in both groups in response to GH administration, but this change did not achieve significance for either group.
Plasma IGFBP-3 and IGFBP-1
The plasma concentration of IGFBP-3 was reduced by 53% in alcohol-fed rats, compared to control values (Fig. 8, top panel). In contrast, circulating levels of IGFBP-1 were elevated more than 6-fold in rats consuming alcohol (Fig. 8, bottom panel). GH failed to significantly alter either IGFBP-3 or BP-1 levels in either group of animals.
SOCS-3 is one of a family of negative regulators of certain cytokines (e.g. interleukin-6) and haemopoietic growth factors, which has been demonstrated to inhibit signal transduction induced by GH ( Nicholson and Hilton, 1998 ). Figure 9 illustrates that the expression of SOCS-3 mRNA in liver was not different between control and alcohol-fed rats. In contrast, GH increased SOCS-3 mRNA in control rats by ~50%, but had no detectable effect on SOCS-3 expression in liver from alcohol-fed rats. SOCS-3 mRNA expression was not detectable in muscle from any group (data not shown).
Plasma amino acids
The prevailing concentration of amino acids is known to mediate various aspects of the IGF system ( Thissen et al., 1994 , Jousse et al., 1998 ). Therefore, we determined the effect of GH on the plasma concentrations of individual amino acids in control and alcohol-fed rats (Table 2 ). In vehicle-treated rats, there was no significant difference between the two groups for the various gluconeogenic, branched-chain or aromatic amino acids. Alcohol feeding decreased the concentration of threonine (30%) and citrulline (29%) as well as increasing the concentration of taurine (115%), 3-methylhistidine (34%) and α-amino-n-butyric acid (6-fold). However, the plasma concentration of total amino acids in vehicle-treated rats was essentially identical between both groups.
Injection of GH in control rats had minimal effects on plasma amino acids. In these rats, GH increased the concentration of glutamine (29%), arginine (53%) and 3-methylhistidine (41%). However, the increase in these amino acids was not of sufficient magnitude to significantly elevate the concentration of total amino acids in the plasma of control rats. GH treatment of alcohol-fed rats produced a relatively greater number of changes. In addition to increasing circulating levels of glutamine (43%) and arginine (74%), as in control rats, GH also increased the plasma concentration of the branched-chain amino acids valine, leucine and isoleucine (35–55%) in alcohol-fed rats. Furthermore, GH reversed or attenuated the alcohol-induced decrease in threonine and the increase in α-amino-n-butyric acid.
GH action after acute alcohol intoxication
In the second experimental study, acute alcohol intoxication was produced in naive rats (no prior surgery) and GH action assessed 12 after a single injection of GH. Figure 10 illustrates plasma IGF-I concentrations in this series of animals. Total IGF-I levels were decreased 19% in alcohol-injected rats, compared to values from control animals. The injection of GH in control rats increased IGF-I by ~270 ng/ml. A comparable GH-induced increase in IGF-I (~290 ng/ml) was observed in alcohol-injected rats. There was no significant difference in the concentration of total IGF-I in plasma between GH-treated control and alcohol-injected rats. Levels of free IGF-I were not determined in this study, because of the similar changes in total and free IGF-I determined in the first study using chronic alcohol-fed rats.
The circulating concentrations of both total and free IGF-I determined prior to GH were decreased 30–40% in alcohol-fed rats, compared to time-matched pair-fed control animals. It is noteworthy that although the alcohol-fed rats had been maintained on the alcohol-containing diet for 14 weeks, at the time the starting sample was collected rats had been fasted for ~18 h and alcohol was not detectable in blood. Hence, the decrease in plasma IGF-I produced by chronic alcohol feeding was not reversed within 18 h of ethanol withdrawal. The liver is the primary site of synthesis for blood-borne IGF-I ( Yakar et al., 1999 ). Therefore, the observed reduction in both hepatic IGF-I peptide and mRNA in alcohol-fed rats confirms previous reports ( Srivastava et al., 1995 , Lang et al., 1998 ) and is consistent with the above-mentioned decrease in circulating IGF-I. Since GH must first bind to its cognate cell membrane-bound receptor prior to initiating its biological response, we also examined the effect of alcohol on hepatic GHR levels. Although GHR mRNA expression was decreased 30% in liver from alcohol-fed rats, compared to control values, there was no significant alteration in GHR peptide levels between the two groups. Hence, the observed reduction in hepatic IGF-I synthesis appears independent of a reduction in the number of GHR and suggests a defect in post-receptor GH signalling under conditions where physiological levels of GH are present. Alternatively, an alcohol-induced decrease in GH secretion, as observed in some ( Soszynski and Frohman, 1992 , Badger et al., 1993 ) but not all ( Sonntag and Boyd, 1989 ) studies, might also contribute to the reduction in starting IGF-I levels. Based on the moderate increase in AST levels and the maintenance of ATP content, it seems unlikely that the alcohol-induced decrease in IGF-I resulted from the presence of severe hepatic injury.
Alcoholism and acute kidney failure
The presence of IGF-I mRNA has been confirmed in a large number of extra-hepatic tissues ( Lowe et al., 1988 ) and suggests that this hormone may also function in a paracrine/autocrine manner (Yaker et al., 1999). Skeletal muscle is known to be an important target tissue for IGF-I action. Elevations in IGF-I increase protein synthesis and decrease proteolysis in skeletal muscle, thereby improving nitrogen balance ( Fryburg, 1994 , Bark et al., 1998 ). In the present study, there was no difference in IGF-I peptide content or the expression of IGF-I mRNA in gastrocnemius obtained from vehicle-treated alcohol-fed and control rats, and suggests that circulating levels of GH are sufficient to maintain normal rates of IGF-I synthesis in this tissue.
The large majority of circulating IGF-I is carried by a family of IGF binding proteins which alter the bioactivity and availability of IGF-I (as reviewed in Rajaram et al., 1997). More than 80% of the IGF-I in the blood is carried in a ternary complex consisting of IGF, an acid labile subunit and IGFBP-3. A much smaller percentage of IGF-I is bound to IGFBP-1. Both of these binding proteins are believed to be secreted primarily by the liver ( Rajaram et al., 1997 ). Although the synthesis of IGFBP-3 has historically been considered to be GH-dependent, insulin and nutrient status are the predominant regulators of IGFBP-1 ( Lee et al., 1997a , Rajaram et al., 1997 ). Chronic alcohol consumption reduced the plasma concentration of IGFBP-3 by ~50% and increased IGFBP-1 levels by more than 6-fold. These data confirm previous observations by our laboratory ( Lang et al., 1998 ) and others ( Santolaria et al., 1995 ). The decrease in plasma IGFBP-3 is independent of changes in the number of GHR and may result from an enhanced proteolysis of circulating IGFBP-3, as observed in other catabolic conditions ( Frost et al., 1996a, b ). In contrast, the increased IGFBP-1 levels seen in alcohol-fed rats is most likely mediated by other factors, such as increases in pro-inflammatory cytokines, as opposed to changes in GH signalling ( Fan et al., 1995 , Li et al., 1997 ). Despite a marked alcohol-induced decrease in the IGFBP-3/IGFBP-1 ratio in blood of alcohol-fed rats, no significant change in the percentage of free IGF-I, relative to total IGF-I, was detected.
The amount of dietary protein and the prevailing concentrations of specific amino acids can dramatically alter synthesis of IGF-I and various IGFBPs ( Thissen et al., 1994 , Jousse et al., 1998 ). However, chronic alcohol ingestion produced only mild changes in a small number of individual amino acids, and did not change the total concentration of amino acids in the blood. Therefore, it appears unlikely that alcohol-induced decreases in plasma amino acids were responsible for any of the observed changes in the IGF system in our alcohol-fed rats. These data are consistent with those of a previous study which also failed to detect major changes in plasma amino acids in rats fed a nutritionally adequate liquid diet containing alcohol for 8–10 weeks ( Shoemaker and Visek, 1988 ). However, the 6-fold elevation in α-amino-n-butyrate in alcohol-fed rats is noteworthy. An alcohol-induced increase in this particular amino acid has been previously reported ( Shaw and Lieber, 1978 ) and appears to result from an enhanced hepatic production ( Shaw and Lieber, 1980 ). In contrast, the plasma concentration of α-amino-n-butyrate (and the branched-chain amino acids) is decreased in animals and humans by dietary protein deficiency ( Shaw and Lieber, 1978 ). These data further suggest that dietary protein intake was adequate in our alcohol-fed rats and that a difference in nutrient consumption was not a primary mediator of the observed changes in the IGF system.
GH-induced changes in the IGF system
Administration of a maximally stimulating dose of GH increased the concentration of total IGF-I in plasma of both control and alcohol-fed rats. The integrated area under the curve for plasma total IGF-I was used to estimate GH responsiveness because of the difference in starting IGF-I levels between groups, and clearly demonstrated that GH responsiveness was not impaired in alcohol-fed rats. Furthermore, there was no difference in the GH-induced increase in free IGF-I between the two groups. GH also did not alter the percentage of free IGF-I, relative to total IGF-I, compared to that determined in vehicle-treated rats. This finding is similar to that previously reported in humans injected with a single dose of GH ( Lee et al., 1997b ). It appears GH treatment alters plasma levels of free IGF-I only when it is administered chronically and accompanied by concomitant changes in various IGFBPs ( Kawai et al., 1999 ). Because starved rats were used in our first study, the data do not directly address whether alcohol must be present in the circulation in order for GH action to be impaired. This possibility was addressed in our second study, where the GH-induced increment in plasma IGF-I was assessed in a model of acute alcohol intoxication. Although blood alcohol levels were not determined in the study, previous reports indicate that ethanol (~65 mM) was present in the blood throughout much of the time when GH action was determined ( Tiernan and Ward, 1986 ). Despite the presence of alcohol in the blood, however, plasma IGF-I levels were similarly increased in control and alcohol-injected rats in response to maximal GH stimulation. The results of this study are consistent with the previously reported ability of alcohol to suppress GH-induced secretion of IGF-I from isolated liver slices when present in high concentrations (175 mM), but not at lower concentrations (35 mM) ( Xu et al., 1995 ). The ability of a maximally stimulating dose of GH to increase plasma IGF-I in alcohol-treated rats is similar to the response reported in patients with AIDS wasting ( McNurlan et al., 1997 ), but is in contrast to other catabolic conditions (e.g. bacterial or viral infection, burn and diabetes) associated with low prevailing levels of IGF-I, which demonstrate a dramatic decrease in GH responsiveness ( Dahn et al., 1988 , Belcher and Ellis, 1990 , Day et al., 1998 ).
We also observed comparable GH-induced increases in hepatic IGF-I peptide levels in both groups, but failed to detect an increase in hepatic IGF-I mRNA expression in either group. However, in preliminary studies, we observed an ~50% increase in hepatic IGF-I mRNA abundance 4 h after injection of GH (unpublished observations). Thus, the GH-induced increase in hepatic IGF-I mRNA may have returned to control levels at the time we sampled tissue (i.e. 12 h after the last injection of GH). In skeletal muscle, GH administration resulted in proportional increases in IGF-I mRNA expression and peptide content, the magnitude of which did not differ between control and alcohol-fed rats.
The total concentration of amino acids in the plasma did not differ between control and alcohol-fed rats injected with GH. Therefore, a decrease in amino acid availability appears to be an unlikely mediator for any GH-induced differences between control and alcohol-fed rats. However, alcohol-consuming rats did demonstrate significant increases (35–55%) in the circulating levels of leucine, isoleucine and valine. Elevation in these branched-chain amino acids is suggestive of an increase in muscle proteolysis. An increased breakdown of myofibrillar protein in alcohol-fed rats in response to GH was also suggested by the tendency for plasma 3-methylhistidine to be increased. Data from several groups provide indirect evidence that exogenous GH may increase muscle proteolysis ( Yarasheski et al., 1995 , McNurlan et al., 1997 ), although the exact mechanism is unknown. Increasing the concentration of leucine has also previously been demonstrated to decrease IGFBP-1 synthesis and secretion ( Jousse et al., 1998 ). However, the GH-induced increase in leucine observed in alcohol-fed rats was not associated with a reduction in IGFBP-1 in the present study.
The cell signalling pathway for GH includes receptor activation of the Janus kinase (JAK) family of protein tyrosine kinases, which ultimately phosphorylates and activates another family of proteins termed the signal transducers and activators of transcription (STAT) ( Yoshimura, 1998 ). The recently cloned SOCS family of proteins appears to inhibit signal transduction by IL-6 and GH, possibly by interacting with elements of the JAK/STAT signalling cascade ( Nicholson and Hilton, 1998 , Yoshimura, 1998 ). Because SOCS genes inhibit GH signalling, they may represent an important com-ponent of a classic negative feedback system. In this regard, administration of GH in vivo has been shown to increase hepatic expression of SOCS-3 ( Adams et al., 1988 ). Our data from GH-injected control animals confirm this earlier finding. However, in contrast to control animals, alcohol-fed rats demonstrated no increase in SOCS-3 mRNA at the time point examined. We speculate that the loss of SOCS-3 induction in alcohol-fed rats may represent a compensatory response which minimizes production of an inhibitory regulator, allowing for continued GH signalling.
In summary, although these data indicate that hepatic GH responsiveness was not impaired by alcohol feeding, they do not necessarily exclude the presence of GH resistance in alcohol-fed rats. Hormone ‘resistance' can be produced by either decreasing the ‘responsiveness' of the system to a maximally stimulating dose of hormone and/or by decreasing hormone ‘sensitivity' ( Kahn, 1978 ). A decrease in hormone sensitivity is best evidenced by a right-shift in the dose–response curve with a concomitant increase in the ED
. Because a complete GH dose–response curve was not generated in the present study, we cannot make definitive conclusions regarding the presence or absence of GH resistance in rats provided alcohol. Indeed, the reduction in starting IGF-I in alcohol-fed rats, despite the presence of normal GHR peptide levels, is suggestive of hepatic GH resistance under conditions where the prevailing circulating concentration of GH is relatively low and physiological.
This work was supported in part by a grant from the National Institute on Alcohol Abuse and Alcoholism AA1290. We thank the National Hormone and Pituitary Program (National Institute of Diabetes and Digestive and Kidney Diseases) for the generous gift of the IGF-I antibody and Genetech (South San Francisco, CA, USA) for the IGF-I used in these studies. We also thank Dr Douglas Hilton (The Walter and Eliza Hall Institute for Medical Research and The Cooperative Research Centre for Cellular Growth Factors, Parkville, Victoria, Australia) for the SOCS-3 cDNA, and Dr William R. Baumbach (American Cyanamid Co., Princeton, NJ, USA) for the GHR antibody.