* Laboratory of Food Science and Nutrition, Department of Life Style Studies, School of Human Cultures, University of Shiga Prefecture, Hikone, Japan; and Division of Food Science and Biotechnology, Graduate School of Agriculture, Kyoto University, Uji, Japan
Received March 18, 2004; accepted June 20, 2004
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ABSTRACT |
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Key Words: phthalate ester; endocrine disrupter; tryptophan metabolism; quinolinate; metabolic disrupter.
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INTRODUCTION |
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The tryptophanNAD pathway consists of the kynurenine pathway and the NAD pathway. The kynurenine pathway is the main route of tryptophan metabolism (Fig. 1). This pathway is initiated by the oxidation of tryptophan by tryptophan oxygenase (TDO) in the liver or by indoleamine dioxygenase (IDO) in other tissues including the brain. The metabolite at a branching point in the tryptophanNAD pathway is -amino-ß-carboxymuconate-
-semialdehyde (ACMS), which is converted by ACMS decarboxylase (ACMSD, EC4.1.1.45) to
-aminomuconate-
-semialdehyde (AMS). AMS eventually leads to acetyl-CoA through the glutarate pathway, or otherwise non-enzymatic cyclization of ACMS results in the formation of quinolinate (QA), from which NAD is synthesized through the NAD pathway. Thus, ACMSD activity plays a critical role in the tryptophanNAD pathway. In mammals, NAD is also synthesized from niacin (nicotinate (NiA) and nicotinamide (Nam)) that can be obtained primarily from dietary sources.
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The structural similarity of phthalates with tryptophan metabolites prompted us to examine the effects of phthalate esters on the pathway of tryptophan metabolism. NAD can be supplied from tryptophan in the dietary protein. Therefore, administration of a niacin-deficient diet containing phthalate esters to rats and measurement of the tryptophan metabolites excreted in the urine make it possible to estimate phthalate esterinduced changes in tryptophan metabolism (Fukuwatari et al., 2002a, 2002b
; Shibata et al., 2001
). We previously reported that di-n-butyly phthalate (DBP) (Shibata et al., 2001
) and DEHP (Fukuwatari et al., 2002a
, 2002b
) stimulated conversion of tryptophan to NAD. In this article, we show that phthalate esters elevate QA and its downstream metabolites in the urine, whereas excretion of 3-hydroxyanthranilate (3-HA) remains unchanged. Of the phthalate esters tested, DEHP and its primary metabolite, mono(2-ethylhexyl)phthalate (MEHP), were the most potent disrupters of tryptophan metabolism. We also present results showing that direct inhibition of ACMSD by phthalate esters is primarily responsible for the phthalate esterinduced change in tryptophan metabolism.
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MATERIALS AND METHODS |
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Animals and diets. The care and treatment of the experimental animals conformed to The University of Shiga Prefecture guidelines for the ethical treatment of laboratory animals. Rats and mice were obtained from Clea Japan (Tokyo), and housed in a room maintained at 22 ± 1°C with 60% humidity and a 12 h light/12 h dark cycle (light onset at 6:00 A.M.). Mice were used for preparation of ACMSD as described later. Body weight and food intake were measured daily at 10:00 A.M., and food and water were renewed daily. Male Wistar rats at 5 weeks old were placed in individual metabolic cages (CT-10; Clea Japan) and acclimated for 1 week. They were fed the control diet containing no phthalate esters. Experiments (five animals per group) were started by using rats at 6 weeks of age. The control diet consisted of 20% casein, 0.2% L-methionine, 45.9% gelatinized cornstarch, 22.9% sucrose, 5% corn oil, 5% mineral mixture (AIN-93 mineral mixture), and 1% vitamin mixture (niacin-free AIN-93 vitamin mixture). The phthalate esters tested were DMP, DEP, DBP, DOP, DEHP, MBP, MHP, or MEHP. Rats were fed with a diet containing 2.6 mmol phthalate ester/kg diet ad libitum for 21 days, and controls were fed without phthalate ester. The weight percent of individual phthalate esters in the diet ranges from 0.05% (500 ppm) of DMP to 0.1% (1000 ppm) of DEHP depending on their molecular weight values. Urine samples on the last day (10:00 A.M.10:00 A.M.; 24-h urine) were collected in amber bottles containing 1 ml of 1 mol/l HCl, and stored at 25°C until use.
Determination of tryptophan metabolites in the urine. Tryptophan metabolites were determined by high-performance liquid chromatography (HPLC). To determine 3-HA (Shibata and Onodera, 1992), urine samples were filtered through a 0.45-µm microfilter, and 20 µl of the filtrates was injected into a STR ODS II column (4.6 x 250 mm I.D., particle size 7 µm) (Shinwa Chemical, Kyoto, Japan). The mobile phase was 50 mmol/l KH2PO4 (pH 3.0)-acetonitrile (100:10 v/v) containing 3 mg/l ethylene diamine tetraacetic acid (EDTA)-2Na, the flow rate was 1 ml/min, the column temperature was maintained at 40°C, and 3-HA was detected at +500 mV electrochemical detection (ECD).
To determine QA (Mawatari et al., 1995), urine samples were filtered through a 0.45-µm microfilter, and 20 µl of the filtrates was injected into a Unisil Q C18 column (4.6 x 250 mm I.D., particle size 5 µm) (GL Sciences, Tokyo). The mobile phase was 20 mmol/l KH2PO4, pH 3.8, containing 0.00045% tetramethylammonium hydroxide and 1.2% hydrogen peroxide, the flow rate was 0.6 ml/min, and the column temperature was maintained at 40°C. The fluorescence intensity at 380 nm was measured upon excitation at 326 nm.
Nam, 2-Py, and 4-Py in the urine samples were measured simultaneously (Shibata, 1987a). Briefly, 1 ml of urine samples was mixed with 10 µl of 1 mg/ml isonicotinamide as an internal standard, 1.2 g of potassium carbonate, and 10 ml of diethylether. The mixtures were shaken vigorously for 5 min, and centrifuged at 800 x g for 5 min. The organic layers were evaporated, and dissolved in 0.5 ml of water. Aliquots of each sample were filtered through a 0.45-µm microfilter, and 20 µl of the filtrates was injected into a CHEMCOSORB 7-ODS-L column (4.6 x 250 mm I.D., particle size 7 µm) (Chemco Scientific, Osaka, Japan). The mobile phase was 10 mmol/l KH2PO4 (pH 3.0)acetonitrile (96:4 v/v), the flow rate was 1 ml/min, the column temperature was maintained at 40°C, and the detection wavelength was 260 nm.
To determine MNA (Shibata, 1987b), urine samples (0.1 ml each) were mixed with 0.7 ml of water, 0.2 ml of 1 mmol/l isonicotinamide, 0.5 ml of 0.1 mmol/l acetophenone, and 1 ml of 6 mol/l sodium hydroxide. After the mixtures were cooled on ice for 10 min, 0.5 ml of 99% formic acid was added, followed by boiling in a water bath for 5 min. The mixtures were cooled on ice, filtered through a 0.45-µm microfilter, and 20 µl of the filtrates was injected into a Tosoh 80Ts column (4.6 x 250 mm I.D., particle size 7 µm) (Tosoh, Tokyo). The mobile phase was a mixture of 20 mmol/l KH2PO4, pH 3.0-acetonitrile (97:3 v/v) containing 1 g/l sodium hepansulfonate and 1 mmol/l EDTA-2Na, the flow rate was 1 ml/min, and the column temperature was maintained at 40°C. The fluorescence intensity at 440 nm was measured upon excitation at 382 nm.
Enzymes and assays. Because the dietary protein has been shown to induce ACMSD in the rat liver (Fukuoka et al., 1998), male Wistar rats (10 weeks old) were fed a high-protein diet (40% casein) for 4 weeks. Male ICR mice (9 weeks old) were fed the control diet (20% casein) for 1 week. Animals were sacrificed by decapitation, and the liver and kidneys were removed from rats and mice, respectively. The organs were immediately homogenized with a polytetrafluoroethylene (PTFE)-glass homogenizer in 5 volumes of cold 50 mmol/l potassium phosphate buffer, pH 7.0. The homogenate was centrifuged at 55,000 x g for 20 min, and the supernatant was used as an enzyme source. Four or five animals per group were used and the enzyme activities were assayed with the supernatant prepared from each organ.
Human ACMSD (Fukuoka et al., 2002) or human quinolinate phosphoribosyltransferase (QPRT, EC 2. 4. 2. 19) (Fukuoka et al., 1998
) transiently expressed in COS-7 cells was prepared from cells cultured for 72 h after transfection. Cells were harvested and lysed with 50 mmol/l Tris-HCl buffer, pH 7.6, containing 137 mmol/l sodium chloride, 1% Triton X-100, 5 mmol/l EDTA, 100 µmol/l leupeptin, and 20 µg/ml FOY-305. The homogenates were centrifuged at 100,000 x g for 15 min, and the supernatants were used for assaying enzyme activity.
The activity of ACMSD was measured as described (Ichiyama et al., 1965). The reaction mixture containing 10 µl of 3.3 mmol/l 3-HA (in 50 mmol/l Tris-acetate buffer, pH 8.0); 0.5 ml of 0.2 mol/l Tris-acetate buffer, pH 8.0; and 0.8 ml of water was incubated in a cuvette for 5 min at 25°C. ACMS was produced by the addition of an excess quantity of the purified 3-HA oxygenase (50 µl containing 0.4 mg protein). After the formation of ACMS was complete, as judged by its absorbance at 360 nm, 0.1 ml of the ACMSD preparation was added. The decrease in absorbance at 360 nm was followed for 5 min against a control incubation that contained all the ingredients except 3-HA. When the effects of phthalate monoesters were examined, 50 µl of the esters dissolved in ethanol was added before the addition of the enzyme. The control incubation contained 50 µl of ethanol. The effects of phthalate diesters could not be tested because of their low solubility in the enzyme assay mixture.
QPRT was assayed as described (Shibata et al., 2000). The incubation medium contained 50 µl of 500 mmol/l potassium phosphate buffer, pH 7.0, 50 µl of 10 mmol/l QA, 50 µl of 10 mmol/l phosphoribosylpyrophosphate, 10 µl of 100 mmol/l MgCl2, 20 µl of phthalate monoester dissolved in ethanol, 270 µl of water, and 50 µl of the enzyme preparation. The control incubation contained 20 µl of ethanol. The reaction was started by addition of the enzyme, and the incubation was carried out at 37°C for 1 h. The reaction tube was placed in a boiling water bath for 5 min to stop the reaction, cooled on ice for 5 min, and centrifuged at 10,000 x g for 5 min. The supernatant was filtered through a 0.45-µm microfilter, and 20 µl of the filtrate was injected into a HPLC column, Tosoh 80Ts (4.6 x 250 mm I.D., particle size 7 µm) (Tosoh, Tokyo). The mobile phase was 10 mmol/l potassium phosphate buffer, pH 7.8, containing 1.48 g/l tetra-n-butylammonium bromide-acetonitrile (90:10 v/v), the flow rate was 1.0 ml/min, and the column temperature was maintained at 40°C. The product was detected at 265 nm.
Statistical analysis. The values are expressed as the mean ± SEM. The statistical significance was determined by ANOVA followed by Tukey's multiple comparison test.
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RESULTS |
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DEHP causes hepatomegaly in rodents by proliferating peroxisome (Elcombe and Mitchell, 1986; Ward et al., 1986
). However, the liver weights of phthalate esterfed groups measured at the end point of experiments (9 weeks of age) did not differ from those of the control groups, indicating that DHEP at the dose level given in this experiment does not cause significant peroxisome proliferation.
Effects of Phthalate Diesters on the Urinary Excretion of the Tryptophan Metabolites
To assess the effects of various phthalate diesters on the tryptophanNAD pathway, rats at 6 weeks of age were fed with a diet containing DMP, DEP, DBP, DOP, or DEHP for 21 days, and the urinary contents of tryptophan metabolites such as 3-HA, QA, Nam, MNA, 2-Py, and 4-Py were measured. The sum of Nam, MNA, 2-Py, and 4-Py was expressed as Nam metabolites. As shown in Figure 2A, the urinary excretion of 3-HA was not changed by any of the phthalate diesters used. In contrast, QA (Fig. 2B) and its downstream metabolites (Nam metabolites in Fig. 2C) were markedly elevated by DEHP. Both DBP and DOP also increased the urinary excretion of QA but to a lesser extent; DMP and DEP, however, had no effect (Fig. 2B). DME, DEP, DBP, and DOP did not affect the excretion of Nam metabolites (Fig. 2C). Thus the length and structure of side chains in the esters appear to be crucial for the urinary excretion of tryptophan metabolites. DEHP that has long and branched side chains was the most powerful disruptor of tryptophan metabolism.
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DISCUSSION |
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Isenberg et al. (2000) explored the metabolism of DEHP given orally to rats. MEHP was the most prominent hepatic metabolite of DEHP, and elevation of the hepatic MEHP concentration was time-dependent and dose-dependent, whereas the levels of DEHP and phthalate were minimal and did not correlate with the dose of DEHP or the time after its administration (Isenberg et al., 2000
). Oral administration of phthalate and 2-ethylhexanol, hydrolysis products of DEHP, did not affect the conversion rate of tryptophan to NAD (Fukuwatari et al., 2002b
). In agreement with the proposal that phthalate monoesters are produced from the diesters in the intestine before absorption (Lake et al., 1977
), these results indicate that MEHP is mainly responsible for the perturbation of tryptophan metabolism.
According to Isenberg et al. (2000), hepatic MEHP concentration in rats fed 1000 ppm DEHP for 2 weeks, conditions similar to those used in the present experiments, was 9 µmol/g tissue. In vitro inhibition of ACMSD by MEHP was apparent (33%) at 0.3 mmol/l and greater than 90% at 3 mmol/l. These results suggest that the liver in rats fed 1000 ppm DEHP accumulates MEHP at the concentration sufficient to exhibit its inhibitory effect on ACMSD, although all of the MEHP molecules in the liver may not necessarily be available for this inhibition.
Previously we showed that ACMSD activity in the liver extracts from rats fed DEHP was similar to that of control animals (Fukuwatari et al., 2002b). This result is not contradictory to our present finding that ACMSD is inhibited in vitro by MEHP. The inhibition is reversible, and therefore, even if MEHP is accumulated in the liver of rats fed DEHP at concentrations sufficient to block ACMSD, MEHP would be washed out during the preparation of the enzyme; the resulting enzyme preparations would contain MEHP at levels that show little inhibition of ACMSD. Taken together, we conclude that phthalate esters perturb tryptophan metabolism through direct inhibition of ACMSD but not by reducing the ACMSD protein level. The mechanism by which ACMSD is inhibited remains to be examined. The very labile nature of ACMS, the substrate of ACMSD, hampers kinetic studies of this enzyme.
Quinolinate is a potential endogenous toxin; it is neurotoxic and has been suspected of being involved in the development of a number of brain diseases (see review by Stone and Darlington, 2002). Although it is believed that the liver is a major site of tryptophan metabolism, expression of ACMSD, as well as its mRNA in the brain and kidney (Fukuoka et al., 2002
), suggests that the phthalate esterinduced metabolic alteration occurs in these organs. However, to date, there are no reports that show neurotoxicity of phthalate esters. When DEHP (0200 mg/kg/day) was given to rats via oral gavage, few adverse effects on neurobehavioral evaluations were found (Moser et al., 2003
). DEHP given to mice through the diet to provide levels of 0.010.09% did not show detrimental effects on neurobehavioral parameters (Tanaka, 2002
). Examination of tissue distribution by the use of the radioactive DEHP did not show significant accumulation of the radioactivity in the brain of rats (Tanaka et al., 1975
) and mice during the pre-weaning period (Eriksson and Darnerud, 1985
). Quinolinate also can be a uremic toxin responsible for anemia with renal failure by reducing the renal production of erythropoietin, a growth factor essential for erythrocyte formation (Pawlak et al., 2003
). When DEHP at a high dose (12,500 ppm in the diet) was given to rats, the erythrocyte count, hemoglobin, and hematocrit values were significantly lower than controls, but these effects were not found with a lower dose (2500 ppm; David et al., 2000
). Although no data indicating that phthalate esters exhibit adverse effects through accumulation of QA are available, effects of the concurrent intake of a tryptophan-rich diet and phthalate esters are worthy of further investigation. Such a series of misfortunes may contribute to triggering and/or exacerbating various diseases.
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ACKNOWLEDGMENTS |
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NOTES |
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1 To whom correspondence should be addressed at Laboratories of Food Science and Nutrition, Department of Life Style Studies, School of Human Cultures, University of Shiga Prefecture, Hikone, Shiga 522-8533, Japan. E-mail: kshibata{at}shc.usp.ac.jp.
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