Pharmacy Department, Kings' College London, FranklinWilkins Building, 150 Stamford Street, London SE1 8WA, UK
Received 23 October 2000; in revised form 2 June 2000; accepted 18 June 2000
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ABSTRACT |
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INTRODUCTION |
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The kidney is one of the most important organs involved in taurine regulation and there is considerable evidence to demonstrate that the kidney (Chesney, 1985) regulates the whole body homeostatis of taurine. In the kidney, there is a high affinity, low capacity, Na+-dependent, ß-amino acid specific transport system, which is responsible for the reabsorption of taurine across the renal tubular brush border membrane (Goldman and Scriver, 1967
; Rozen et al., 1979
). In general, the transport systems for taurine are specific for ß-amino acids and are both energy- and sodium-dependent. Using structural analogues of taurine, which compete for the specific ß-amino acid uptake sites into cells, it is possible to deplete the body pool of taurine. ß-Alanine is commonly used for this purpose in taurine research, as it utilizes the same ß-amino acid uptake system in the kidney (Goldman and Scriver, 1967
), and so it competitively inhibits taurine reuptake into the proximal tubular cells. The result is a loss of taurine from the body, and thus depletion of taurine in tissues such as the liver, as taurine does not accumulate in cells.
The aim of the present study was to show whether the co-administration of ß-alanine with alcohol to rats (to reduce hepatic taurine levels) would alter the pathological and biochemical lesions induced by alcohol, for example, hepatic steatosis and lipid peroxidation.
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MATERIALS AND METHODS |
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Animals
Female SpragueDawley rats (125150 g) were obtained from Charles River (UK) and acclimatized for at least 7 days after delivery. Animals were housed in communal cages, fed a rat and mouse maintenance cube diet (691 diet, Quest Nutrition Ltd., Wingham, Kent, UK) and water ad libitum. During experiments, animals were housed in individual metabolism cages designed to separate and collect faeces and urine (Techmate Ltd., Milton Keynes, Bucks, UK) and given a powdered diet (691 diet, Quest Nutritional Ltd., Wingham, Kent, UK) and water ad libitum prior to introduction of the liquid diet. Lighting was controlled to give a regular 12 h light12 h dark cycle (08:00 on20:00 off); room temperature was maintained at 21 ± 1°C. Urine samples (24 h) were collected over ice and diluted to 25 ml with UHQ water, centrifuged (2000 rpm, 10 min, 4°C) to remove hair and food debris and stored (80°C) in aliquots for later analysis. The body weight and general condition of the animals were monitored twice weekly and liquid diet intake was determined daily. Animals were used under the British Home Office regulations.
Liquid diet technique of ethanol administration
Due to the tendency of animals given alcohol to reduce their solid food consumption, animals were given a liquid diet. Chronic ethanol feeding was achieved by incorporation of ethanol in a nutritionally adequate totally liquid diet obtained from Dyets Inc. (Pennsylvania, USA). The liquid diet provided 1 kcal/ml of which 35% of total calories were derived from fat, 47% from carbohydrates and 18% from protein. Ethanol-treated animals were given diet, where maltose dextrin was isocalorically replaced by ethanol. The alcohol provided 36% of the calories (Lieber and DeCarli, 1989). Animals were started on the diet at a body weight of 125150 g and ethanol was introduced progressively with 30 g/l of the liquid diet for 2 days, 40 g/l for the subsequent 2 days followed by the final formula containing 50 g/l (Lieber and DeCarli 1989
; Kerai et al., 1998
, 1999
).
Preparation of diet. As vitamin A degrades when mixed in with other dry ingredients, vitamins and minerals were incorporated into the diet at the time of preparation of the liquid diet. The liquid diet was prepared in cold tap water using a kitchen blender. The diet was kept refrigerated in the dark and used within 1 week of preparation.
Pair-feeding.
The alcohol-fed animals were allowed liquid diet consumption ad libitum and their daily intake was monitored. The control animals were then given the same amount of liquid control diet during the following 24-h feeding period. This pair-feeding process was repeated every 24 h. The technique of daily pair-feeding was adopted to assure a strict caloric intake in both ethanol-treated animals and their individual pair-fed controls (Lieber and DeCarli, 1989).
Study design
Rats were either provided with 3% ß-alanine in the drinking water for 2 days (n = 12) or given tap water (n = 12). Rats (n = 6) were then treated with alcohol which was administered in the liquid diet for 28 days. Pair-fed control rats (n = 6) were also provided with the same liquid diet but without alcohol. A second group of animals (n = 6) received alcohol administered in the liquid diet which also contained 3% ß-alanine. Pair-fed control rats (n = 6) were given the same liquid diet containing 3% ß-alanine only. After 28 days of treatment, animals were killed and blood and tissue removed for analysis and microsomes prepared from the liver.
Post-mortem procedure
Animals were exsanguinated from the abdominal aorta under anaesthesia (Hypnorm : Hypnovel : water, 1:1:2, 3.33ml/kg, i.p.) and blood samples were collected into Microtainers (Becton Dickinson & Co., Rutherford, NJ, USA) for the separation of serum. After standing at room temperature for at least 45 min, the Microtainers were centrifuged (13 000 rpm, 45 s, MSE minifuge) and stored at 80°C. Serum was analysed for enzymes and biochemical parameters using appropriate kits (Boehringer Mannheim GmbH Diagnostica, Germany) with a centrifugal IL Monarch 2000 (Instrumentation Laboratory, UK, Ltd). The liver was removed, weighed, and approximately 200 mg taken from the right lobe and immediately homogenized in trichloroacetic acid (TCA, 10% w/v, 4 ml, 4°C), frozen in liquid nitrogen and stored at 80°C for subsequent analysis of ATP. Approximately 200 mg of liver were also taken from the right lobe and immediately homogenized in sulphosalicyclic acid (0.2 M, 2 ml, 4°C), frozen in liquid nitrogen and stored at 80°C for subsequent analysis of taurine, TNPSH, and oxidized glutathione (GSSG).
Biochemical determinations
Taurine.
A high-performance liquid chromatographic method with fluorimetric detection was used for the determination of taurine in urine, serum, and liver tissues essentially by the method of Waterfield (1994). Taurine was derivatized with o-phthalaldehyde/2-mercaptoethanol prior to injection onto a C18 column. Isocratic elution of the adduct was carried out using NaH2PO4 (0.05 M, pH 5.4) in methanol and water (43:57 v/v). Homoserine was used as an internal standard to facilitate the standardization and quantification of samples. Analysis was completed in 6 min with homoserine and taurine eluting after 3 and 4 min respectively.
Triglycerides. The hepatic content of triglyceride was determined by a modified method of Butler et al. (1962). Briefly, phospholipids were separated from triglycerides by adsorption on a synthetic Zeolite. The triglycerides were then extracted into chloroform, hydrolysed, and measured as esterified glycerol with non-esterified samples used as individual blanks.
Lipid peroxidation. Lipid peroxidation, measured as malondialdehyde production in liver samples, was determined by the method of Sawicki et al. (1963) employing malondialdehyde as standard.
ATP.
ATP content of liver samples was determined by luciferase-linked bioluminescence in TCA extracts of liver samples using a firefly lantern extract (Jenner and Timbrell, 1994).
TNPSH.
Liver TNPSH were determined by the method of Ellman (1959) as a measure of reduced liver glutathione, which constitutes most (>95%) of the liver TNPSH (De Master and Redfern, 1987).
GSSG. GSSG was determined by the method of Griffith (1980) using 2-vinylpyridine to mask GSH.
Microsomal analysis. Microsomes were prepared from livers, as described by Lake (1987). Total cytochrome P-450 content of liver samples was determined by the method of Omura and Sato (1964). 4-Nitrophenol hydroxylase activity was determined by the modified method of Prough et al. (1978). 4-Nitrophenol is a substrate for the ethanol-inducible CYP2E1. The method relies on the formation of p-nitrocatechol, which can be detected spectrophotometrically after total ionization under alkaline conditions. The protein content of microsomes was determined by the method of Lowry et al. (1951) using bovine serum albumin (BSA) as standard.
Homocysteine and cysteine. A high-performance liquid chromatographic method with fluorimetric detection was used for determination of total homocysteine and cysteine, (oxidized and reduced) in urine and serum by the procedure of Fortin and Genest (1995). Homocysteine and cysteine were reduced by 10% tri-n-butylphosphine in dimethylformamide, then derivatized with SBD-F (ammonium-7-fluorobenzo- 2-oxa-1,3-diazole-4-sulphonate) at 60°C for 1 h (stable for 1 week at 4°C), prior to injection onto a C18 column. Isocratic elution of the adduct was carried out using sodium acetate (0.1 M), acetic acid (0.1 M) and 2% methanol, pH 4.0. N-Acetylcysteine was used as an internal standard to facilitate the standardization and quantification of samples. Analysis was completed in 14 min with cysteine, homocysteine, and N-acetylcysteine eluting after 2.5, 3.5, and 6 min, respectively.
Methionine synthase. Methionine synthase was measured in the liver cytosol as described by Nicolaou et al. (1997). Assay mixtures (total volume 300 µl) contained 50 mM potassium phosphate buffer pH 7.2, 400 µM (dl)-homocysteine, 35 µM SAM, 236 µM MTHF (2658 dpm/nmol), 60 µM hydroxycobalamin, 25 mM DTT (dl-dithiothreitol) and the enzyme source. Incubations were performed in light-protected stoppered serum vials under nitrogen. Reaction mixtures were preincubated for 5 min (37°C), prior to the initiation of the reaction by the addition of homocysteine through a syringe. Incubations were performed for 45 min at 37°C. The enzyme reaction was terminated by the addition of ice-cold water (400 µl) and solutions immediately passed through a 0.5 x 5.0 cm column of BioRad AG1-X8 resin. [14C]Methionine was eluted with 2 ml of UHQ water, collected and quantified by scintillation spectrometry. Protein concentrations were determined with the BioRad protein assay based on the method of Bradford (1976) with BSA as standard.
Urinary protein. Urinary protein was measured by the Coomassie Plus Protein assay kit supplied by Pierce and Warriner (Chester, UK).
Histology
Tissues were fixed in 10.5% (v/v) phosphate-buffered formalin (pH 7.2) and embedded in paraffin wax. Sections (4 µm) were cut and stained with Mayer's haematoxylin and eosin. Frozen liver sections from fixed tissues were cut (10 µm) and stained for lipid with Oil Red O in triethylphosphate with Mayer's haematoxylin as counter stain.
Statistical analysis
Statistical evaluation of data was performed by Duncan's multiple range test to make comparisons between groups. Values quoted are means ± SEM of six animals. The level of significance (P) was set at <0.05.
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RESULTS |
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Post mortem
Both total and relative liver weights (Fig. 2a) and total kidney weights (Fig. 2b
) were significantly raised in the alcohol- and alcohol plus ß-alanine-treated animals, compared to the pair-fed controls. There was no significant difference in liver and kidney weights between the alcohol- and alcohol-plus ß-alanine-treated animals.
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DISCUSSION |
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Although hepatic triglyceride levels were found to be slightly but significantly lower in the alcohol plus ß-alanine-treated animals, compared to alcohol treatment alone, the hepatic steatosis seemed greater in the alcohol plus ß-alanine-treated animals when assessed histologically. Ethanol decreases triglyceride release into the blood (Dajani and Konyoumjian, 1967; Koga and Hirayama, 1968
; Madsen, 1969
), which may be one of the mechanisms underlying ethanol-induced hepatic steatosis. Although alcohol and alcohol plus ß-alanine treatment raised serum triglyceride levels, the difference between the two groups was not significant. Thus, unlike taurine (Kerai et al., 1999
), ß-alanine does not affect triglyceride transport from the liver.
Chronic ethanol feeding results in proliferation of the membranes of the smooth endoplasmic reticulum. As many of the enzymes involved in the synthesis of triglycerides and phospholipids are bound to the membranes of the endoplasmic reticulum, the lipid-synthesizing capacity of the liver could be enhanced. As ethanol is also metabolized by the microsomal enzyme system, it has the potential to interfere with lipid metabolism. Microsomal ethanol oxidation may interfere with lipid metabolism by generating oxygen radicals such as O2 and HO, which initiate a cascade of lipid peroxidation and damage cell membranes (Polavarapu et al., 1998). In this study, both alcohol and alcohol plus ß-alanine treatment significantly increased hepatic lipid peroxidation, but there was no significant difference between the treatment groups, suggesting that taurine depletion did not affect lipid peroxidation. Previous studies, however, have shown that taurine treatment decreases the extent of lipid peroxidation (Kerai et al., 1998
, 1999
).
Alcohol and alcohol plus ß-alanine treatment raised hepatic GSH levels and reduced GSSG levels after 28 days of ethanol administration. Fernández-Checa et al. (1993) have shown that GSH depletion precedes steatosis and lipid peroxidation, and have suggested that the depletion of GSH could be a contributing factor in the development of alcoholic liver disease. It is possible that GSH depletion occurred earlier in this study and that by 28 days, GSH levels were raised as a result of: (1) rebound synthesis in GSH; (2) conversion of homocysteine to GSH; or (3) mild cholestasis (Dahm et al., 1991; Seabra and Timbrell, 1997
). As with previous studies, there was no effect of alcohol on hepatic ATP levels. However, ß-alanine treatment alone raised ATP levels, but the reason for this is unclear.
Changes in methionine metabolism or methylation in the liver may have an important role in alcohol toxicity (Fig. 10). Methionine is converted to SAM, which is important in maintaining the integrity of the liver. SAM is important in the conversion of phosphatidylethanolamine to phosphatidylcholine. Phosphatidylcholine has been shown to be an important constituent of lipoproteins that are involved in the transport of fat from the liver. This would prevent accumulation of fat in the liver and subsequent liver injury (Gigliozzi et al., 1998
). The conversion of homocysteine to methionine has been considered to be an essential reaction for conserving methionine, detoxifying homocysteine and for the production of SAM (Lucock et al., 1996
). The initial decrease in urinary homocysteine in the control animals and ß-alanine-treated groups can be explained by the change from the powdered diet (4.7 g/l methionine), compared to the liquid diet (0.3 g/l methionine). Previous studies have shown that taurine does not protect against the alcohol-induced increase in urinary homocysteine levels and the inhibition of hepatic methionine synthase activity (Kerai et al., 1998
, 1999
). Although ethanol itself does not inhibit methionine synthase, acetaldehyde is known to inhibit highly purified methionine synthase in vitro (Kenyon et al., 1998
). However, in the present study, ß-alanine- and especially alcohol plus ß-alanine-treated animals excreted significantly greater amounts of homocysteine and cysteine into the urine.
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Consistent with this is the lack of effect of ß-alanine on hepatic lipid peroxidation. The data on the effect of ß-alanine on fatty liver and triglyceride accumulation is difficult to interpret and reconcile. Biochemically, there was a decrease in triglycerides in the liver of alcohol plus ß-alanine-treated animals, whereas histologically there seemed to be an increase. This could reflect the nature of the lipids which accumulate and are detected by Oil Red O staining, in contrast to the triglycerides measured biochemically. Thus, there could be an overall increase in liver fats but a decrease in trigylcerides caused by ß-alanine. Clearly, however, unlike taurine, ß-alanine treatment did not decrease cytochrome P-450 levels or hepatic lipid peroxidation. Also in the present study, serum bile acids and ALP were raised in alcohol-treated animals and significantly more so in the alcohol plus ß-alanine-treated animals, suggesting that cholestasis may have occurred. However, with lower levels of taurine, less taurocholate may be formed. Thus, the accumulation of toxic bile acids in the alcohol plus ß-alanine-treated animals could result in greater hepatotoxicity in these animals.
The data indicate that treatment with ß-alanine has a number of effects. These may be due to the depletion of taurine or to the direct effects of ß-alanine itself, or to a combination of these factors. This study does not distinguish between these effects. As ß-alanine is an analogue of taurine, it is quite possible that some effects may be common. For example the decrease in the level of triglycerides and the rise in the level of bile acids and ALP seen in the serum were also observed in rats treated with alcohol and taurine. Thus, in this study the depletion of hepatic and serum taurine levels by treatment with ß-alanine significantly increased the hepatotoxicity of ethanol as determined histologically and by some biochemical measurements. The present results support previous findings (Kerai et al., 1998, 1999
) that taurine has a protective role in the liver against ethanol-induced hepatic steatosis. However it should be noted that the rat is a good synthesizer of taurine, unlike humans, who rely more heavily on dietary sources. Therefore changes brought about by the depletion of taurine with ß-alanine treatment would have more impact in humans than in the rat.
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ACKNOWLEDGEMENTS |
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FOOTNOTES |
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