* Central Product Safety Division, Miami Valley Laboratories, Procter and Gamble Co., Cincinnati, Ohio 45253; and
Department of Nutrition, University of North Carolina, Chapel Hill, North Carolina 27599
Received October 12, 2001; accepted December 18, 2001
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ABSTRACT |
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Key Words: choline; diethanolamine; phosphocholine; S-adenosylmethionine; betaine; ethanol; hepatocarcinogenesis.
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INTRODUCTION |
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DEA is structurally similar to ethanolamine and choline, important endogenous precursors in the synthesis of phospholipids that are essential for normal membrane structure and function (Pelech and Vance, 1984; Zeisel and Blusztajn, 1994
). Other choline metabolites include acetylcholine, a vital neurotransmitter, platelet-activating factor, a phospholipid that mediates many processes of inflammation and allergy, and sphingomyelin, an important precursor involved in cell signaling and differentiation (Zeisel and Blusztajn, 1994
). Choline is also oxidized to betaine, an essential methyl group donor (Finkelstein et al., 1982
).
The pathways of choline utilization and 1-carbon metabolism are shown in Figure 1. Importantly, choline utilization intermingles with the metabolic pathways that serve to protect the capacity to donate methyl groups. The interaction between these pathways is demonstrated by the fact that dietary choline deficiency in rats not only depletes hepatic choline and choline metabolites, but decreases S-adenosylmethionine (SAM) concentrations as well (Pomfret et al., 1990
; Shivapurkar and Poirier, 1983
; Zeisel et al., 1989
). The liver is distinguished from most other tissues in that it has an active alternate pathway to synthesize PC by sequential methylation of phosphatidylethanolamine (PE) by the action of PE N-methyltransferase (Vance et al., 1997
). This alternate biosynthetic process requires 3 moles of SAM, representing another example of the interdependence between the choline and the methylation pathways. Furthermore, the methylation of PE to PC is thought to be the only pathway for de novo choline biosynthesis in mammals (Zeisel and Blusztajn, 1994
).
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Dietary choline deprivation is hepatocarcinogenic in rodents (Ghoshal and Farber, 1984; Mikol et al., 1983
; Newberne and Rogers, 1986
; Newberne et al., 1982
; Zeisel, 1996
) and promotes liver tumor formation following initiation by a chemical carcinogen (Lombardi and Shinozuka, 1979
; Sells et al., 1979
; Yokoyama et al., 1985
). In contrast, dietary supplementation of choline with or without methionine reduces liver tumor incidence in carcinogen-treated mice (Fullerton et al., 1990
; Newberne et al., 1990
). The mechanisms by which choline deficiency is thought to be carcinogenic include enhanced cell proliferation, altered methylation status, and altered signal transduction (Rogers, 1995
; Zeisel, 1996
; Zeisel and Blustjazn, 1994). DEA caused morphological transformation in the Syrian hamster embryo (SHE) cell transformation assay, supporting its carcinogenic activity. However, this response was prevented when SHE cells were cultured in a medium containing excess choline (Lehman-McKeeman and Gamsky, 2000
), supporting a role for altered choline homeostasis as a mode of action for the carcinogenic effects of DEA.
Although the precise mechanisms underlying the hepatocarcinogenic action of dietary choline deficiency are not fully established, it is clear that choline deficiency is a mode of action by which liver tumors develop in rodents. Given that DEA disrupts choline homeostasis in cultured cells, the purpose of the present work was to test the hypothesis that DEA treatment could produce biochemical changes consistent with choline deficiency in mice. To this end, the biochemical changes observed in mice made choline deficient by dietary choline deprivation were compared to the effects observed in DEA-treated mice. Experiments were also conducted to define the dose-response relationship, mouse strain dependence, and possible influence of an ethanol vehicle on these biochemical effects.
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MATERIALS AND METHODS |
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Chemicals.
DEA (99% purity) was obtained from Aldrich Chemical Co. (Milwaukee, WI). Absolute ethanol (Aaper Alcohol and Chemical Co., Shelbyville, KY) was used to prepare 95% ethanol used as the vehicle in these studies. Betaine (HCl salt) was obtained from Aldrich, SAM and SAH were obtained from Sigma Chemical Co. (St. Louis, MO), and all standards for choline analyses were as described by Pomfret et al. (1989).
Dietary choline deficiency.
Male B6C3F1 mice (approximately 6-weeks-old at study initiation) were allowed ad libitum exposure to a control or choline-devoid diet (Dyets, Bethlehem, PA) for a period of 2 weeks (n = 8/group). The control diet contained 0.25% choline (as choline bitartrate) and the methionine content in the control and choline-devoid diets was 0.6%. At the end of the 2-week exposure period, livers were rapidly removed under anesthesia, immediately flash-frozen in liquid N2 and stored at 80°C pending analysis of choline metabolites. In a separate group of mice (n = 4/group), blood was collected for clinical chemistry analyses, and livers were weighed and fixed in 10% neutral buffered formalin for histopathological assessment.
DEA dose-response study.
Male B6C3F1 mice (approximately 6-weeks-old at study initiation) were housed individually and dosed dermally with DEA dissolved in 95% ethanol at 0, 10, 20, 40, 80, or 160 mg/kg (n = 6/group) for a period of 4 weeks (5 days/week). These dosages were selected to include those shown to be carcinogenic (40, 80, and 160 mg/kg/day; NTP, 1999). There were 2 control groups for this study including untreated and ethanol-treated mice. A separate group of mice was dosed with the ethanol vehicle or 160 mg DEA/kg/day for 4 weeks (5 days/week) and then allowed a 2-week recovery prior to necropsy. For dosing, the backs of all mice were shaved, and the dosing solutions (1.8 ml/kg) were applied to a region approximately 2 cm2 (from the mid-back to the interscapular region) using a pipette. All mice, including the untreated controls, were shaved on an as-needed basis during the study period. At the end of the dosing or recovery period, livers were harvested as described above and analyzed for choline metabolites.
To evaluate potential strain differences in the response to DEA, male C57BL/6 mice, also about 6-weeks-old at study initiation, were dosed with 0 or 160 mg DEA/kg for a similar 4-week period. The untreated and ethanol-treated control groups were also used in this study.
Clinical and histopathological evaluations.
Formalin-fixed livers were processed using standard conditions and stained with hematoxylin and eosin. Histopathological evaluations were made without knowledge of treatment. Clinical analyses were conducted on freshly isolated serum with a Hitachi 717 chemistry analyzer (Boehringer-Mannheim Corp., Indianapolis, IN).
Analysis of hepatic choline metabolites.
Phospholipids and aqueous choline metabolites were analyzed according to the method outlined by Pomfret et al. (1989). Briefly, frozen liver samples were pulverized under liquid N2 after which lipids were extracted according to the method of Bligh and Dyer (1959). The organic phase (for PC) was separated by thin layer chromatography (TLC), whereas the aqueous phase was injected onto a silica HPLC column, and the appropriate internal standards labeled with stable isotopes were added to calculate and correct for recovery. Following the TLC analyses, the segments representing PC were visualized, scraped, and hydrolyzed in 6 M methanolic-HCl to liberate free choline. From the HPLC analyses, the peaks representing PCho, GPC, and choline were collected, dried, and hydrolyzed to liberate choline. In all cases, choline was converted to its propionyl ester and demethylated with sodium benzenethiolate. This volatile derivative was isolated by gas chromatography and quantified with a mass selective detector.
SAM and SAH concentrations were determined by a high-performance liquid chromatographic (HPLC) method based on a modification of Wagner et al. (1984). Briefly, frozen liver samples were homogenized in 2.5% perchloric acid containing 10 mM Na metabisulfite and 1 mM EDTA. After centrifugation, protein-free supernatants were injected onto a Waters Symmetry column (2.1 x 100 cm) and SAM and SAH were separated with an acetonitrile gradient in 100 mM phosphate buffer (pH 3.0), acetonitrile, and 8 mM octanesulfonic acid. Buffers A and B contained 5 and 50% acetonitrile, respectively, and a linear gradient system of 0 to 70% buffer B over a 20-min interval was used. SAH and SAM, detected at 254 nm, eluted at approximately 10.4 and 12.7 min, respectively. The detection limit for both analytes was 0.1 nmol/g liver.
For betaine analysis, frozen liver samples were homogenized in 4 volumes of 15% trichloracetic acid, after which the protein-free supernatants were subjected to exhaustive extraction and periodide complexation as described by Barak and Tuma (1979). Betaine periodide was precipitated, resuspended in dichloroethane and analyzed spectrophotometrically at 365 nm. The detection limit for this assay was 0.3 µmol betaine/g liver.
Statistical analysis.
Data were analyzed for statistical significance by Student's t-test or ANOVA followed by a multiple comparison test (Fischer's PLSD) using StatView 4.1 (Abacus Concepts, Berkeley, CA). For statistical comparisons, a 5% significance level (p < 0.05) was used.
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RESULTS |
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One way in which B6C3F1 mice are distinguished from the maternal C57BL/6 strain is the ability to maintain nascent methylation status, a characteristic which is dependent on hepatic SAM levels (Counts et al., 1996). Figure 3
illustrates that when dosed with DEA at 160 mg/kg/day, C57BL/6 mice were sensitive to changes in PCho levels, but unlike the B6C3F1 mice, no change in hepatic SAM levels were noted in C57BL/6 mice.
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DISCUSSION |
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The pattern of changes observed in choline metabolites after DEA treatment was very similar to those observed in choline deficient mice. Specifically, hepatic PCho levels decreased, reaching a maximum 50% reduction within 4 weeks of dermal dosing. Choline metabolites and SAM levels also decreased. Therefore, these data indicate that DEA treatment produces biochemical changes in liver that are consistent with choline deficiency. Furthermore, the effects of DEA were dosage-dependent. All carcinogenic dosages of DEA (40, 80, and 160 mg/kg/day) altered choline homeostasis, supporting the hypothesis that choline deficiency is the mode of action by which DEA is hepatocarcinogenic in mice. Importantly, these changes occurred in mice consuming a choline-replete diet. Previous studies have demonstrated that DEA selectively inhibits the uptake of choline into cells and competes for the normal utilization of choline in the CDP-choline pathway (Lehman-McKeeman and Gamsky, 1999, 2000
). Thus, it is likely that the actions of DEA limit intracellular choline availability in a manner that leads to a choline-deficient condition even when choline is available in the diet. The effects of DEA on choline uptake and utilization have also been shown to be competitive and reversible, as choline supplementation can inhibit the effects of DEA in cultured cells (Lehman-McKeeman and Gamsky, 2000
). Similarly, the ability of DEA to alter choline homeostasis in vivo reversed when treatment was stopped for 2 weeks prior to necropsy, demonstrating that DEA must be continuously present in order to exert these effects.
The histopathological hallmark of choline deficiency in rodents is fatty liver. This feature, which results from the inability of the liver to synthesize adequate PC for the formation of very low density lipoproteins required for triglyceride secretion (Yao and Vance, 1988, 1990
), has been documented repeatedly for rats made choline deficient by dietary deprivation. The possibility that DEA could produce choline deficiency has been discounted by some groups due to the lack of fatty liver (NTP, 1999
). However, the present results demonstrate that no fatty change was observed in mice after 2 weeks of dietary choline deficiency. It is generally recognized that the mouse model of chronic choline deficiency differs from rats because most mouse strains do not develop liver cirrhosis and hemorrhagic necrosis that is typical of chronic choline deficiency in rats (De Carmago et al., 1985). Moreover, 2 other experimental conditions can contribute to the development and severity of fatty liver in response to choline deficiency. First, sensitivity to and severity of choline deficiency is age-dependent. Weanling animals are highly susceptible to choline deficiency, and most studies conducted to date have initiated choline deficiency in animals ranging from 34 weeks of age (Chandar and Lombardi, 1988
; da Costa et al., 1995
; Fullerton et al., 1990
). Susceptibility to choline deficiency declines rapidly, and animals that are 1012 weeks of age are unlikely to develop features of fully expressed choline deficiency (Rogers et al., 1987
). Secondly, dietary fat composition also contributes to the development of fatty liver in choline deficiency. Specifically, the fat composition of choline deficient diets is often augmented to at least 20% of the diet, whereas standard laboratory chows contain about 5% fat (Ghoshal and Farber, 1984
; Rogers et al., 1987
). Additionally, variations in fatty liver development are observed depending on whether the fat is derived from animal or plant origins (De Camargo et al., 1985
; Rogers et al., 1987
).
In the present studies, mice were approximately 6 weeks of age at the initiation of choline deficiency or DEA treatment. This age was chosen intentionally to mimic the conditions of the chronic bioassays conducted with DEA. Furthermore, the synthetic diet contained less than 10% fat and the laboratory chow used in the DEA experiments contained only 5% fat that is derived from soybean oil. Consequently, experimental conditions, namely species used, animal age, and dietary composition may have decreased the likelihood of fatty liver development, but do not detract from the biochemical evidence that DEA causes hepatic choline deficiency.
Chronic ethanol ingestion has been shown to increase hepatic choline requirements (Barak et al., 1973), and the major manifestation of this action is a reduction in hepatic betaine levels (Barak et al., 1984
; Chern et al., 2000
). As illustrated in Figure 1
, betaine, the oxidation product of choline, is an important 1-carbon donor utilized in the maintenance of cellular methylation status. In the conduct of the chronic bioassays for DEA, ethanol was used as the vehicle for dermal application of DEA, with the daily dermal dosage reaching nearly 1.5 g ethanol/kg/day (NTP, 1999
). The results presented herein indicate that the internal dosage of ethanol from dermal dosing was sufficient to reduce hepatic betaine levels. DEA treatment had no further effect on hepatic betaine levels. Given the substantial interplay between choline and methyl metabolism, it is highly likely that the hepatic effects of DEA, including its carcinogenic action, may be exacerbated by coadministration of ethanol.
In addition to providing a mode of action by which DEA is carcinogenic, the present data have important implications for human cancer risk assessment. First, from a qualitative perspective the sensitivity of the B6C3F1 mouse strain to choline deficiency and hepatocarcinogenesis must be considered. In the present study, choline deficiency, as evidenced by changes in PCho concentrations, was produced in both B6C3F1 and C57BL/6 mice. However, unlike the B6C3F1 mouse, DEA did not alter SAM concentrations in the C57BL/6 strain, which is considered resistant to liver tumor development (Counts et al., 1996; Goodman et al., 1991
). Although we have not conducted chronic studies to evaluate the carcinogenic potential of DEA in C57BL/6 mice, the results denote important strain differences in potential susceptibility that should be considered when evaluating human risk.
From a qualitative and quantitative perspective, there are marked species differences in susceptibility to choline deficiency, with rats and mice being far more susceptible than other species including humans (Zeisel and Blusztajn, 1994). In fact, rats and mice have a higher dietary choline requirement than humans in large part because rodents oxidize choline more rapidly than humans (Sidransky and Farber, 1960
). In the carcinogenicity studies, DEA was only carcinogenic in mice, not in rats. The lack of carcinogenicity in rats, a species that is highly susceptible to choline deficiency, should be an important consideration in the overall evaluation of human cancer risk. Disposition data indicate that DEA is less readily absorbed across rat skin than mouse skin, and the resulting blood and tissue concentrations of DEA are at least 3-times lower in rats than in mice at similar dosages (Mathews et al., 1997
). The present work has determined the NOEL for DEA-induced choline deficiency in mice (based on PCho concentrations) to be 10 mg/kg/day. Thus, there is a critical concentration of DEA that must be reached in order to affect choline homeostasis. In rats, the lack of a carcinogenic response suggests that it is unlikely that exposure to DEA reached this critical concentration or that rats are not as susceptible as mice to the effects of DEA on hepatic choline metabolism.
In summary, the results of the present study provide evidence that 4 weeks of DEA treatment leads to a biochemical condition of hepatic choline deficiency in mice. The changes, which provide strong evidence for a nongenotoxic mode of action, were dose-dependent, strain-dependent, and reversible. Moreover, there was a consistent dose concordance between the biochemical and hepatocarcinogenic effects of DEA. Mice, especially the B6C3F1 strain, are particularly susceptible to altered choline and methyl group homeostasis after DEA treatment, but a clear NOEL for DEA-induced changes in hepatic choline concentrations was established. The results also demonstrate that dermally applied ethanol decreased hepatic betaine concentrations and may confound or exacerbate the hepatic effects of DEA. Overall, the results suggest that the hepatocarcinogenic effects of DEA in mice are not predictive of similar susceptibility in other laboratory animals or humans.
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ACKNOWLEDGMENTS |
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NOTES |
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REFERENCES |
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Barak, A. J., and Tuma, D. J. (1979). A simplified procedure for the determination of betaine in liver. Lipids 14, 860863.[ISI][Medline]
Barak, A. J., Tuma, D. J., and Sorrell, M. F. (1973). Relationship of ethanol to choline metabolism in the liver: A review. Amer. J. Clin. Nutr. 26, 12341241.[ISI][Medline]
Barbee, S. J., and Hartung, R. (1979). The effect of diethanolamine on hepatic and renal phospholipid metabolism in the rat. Toxicol. Appl. Pharmacol. 47, 421430.[ISI][Medline]
Bligh, E. G., and Dyer, W. J. (1959). A rapid method or total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911917.[ISI]
Chandar, N., and Lombardi, B. (1988). Liver cell proliferation and incidence of hepatocellular carcinoma in rats fed consecutively a choline-devoid and a choline-supplemented diet. Carcinogenesis 9, 259263.[Abstract]
Chern, M. K., Gage, D. A., and Pietruszko, R. (2000). Betaine aldehyde, betaine and choline levels in rat livers during ethanol metabolism. Biochem. Pharmacol. 60, 16291637.[ISI][Medline]
Christman, J. K. (1995). Lipotrope deficiency and persistent changes in DNA methylation. Lipotrope deficiency and DNA methylation. In Diet and Cancer (American Institute of Cancer Research, Ed.), pp. 97106. Plenum Press, New York.
Counts, J. L., Sarmiento, J. I., Harbison, M. L., Downing, J. C., McClain, R. M., and Goodman, J. L. (1996). Cell proliferation and global methylation status changes in mouse liver after phenobarbital and/or choline-devoid, methionine-deficient diet administration. Carcinogenesis 17, 12511257.[Abstract]
Cui, A., and Vance, D. E. (1996). Expression of phosphatidylethanolamine N-methyltransferase-2 is markedly enhanced in long term choline-deficient rats. J. Biol. Chem. 271, 28392843.
Da Costa, K.-A., Garner, S. C., Chang, J., and Zeisel, S. H. (1995). Effects of prolonged (1 year) choline deficiency and subsequent re-feeding of choline on 1,2-sn-diradylglycerol, fatty acids and protein kinase C in rat liver. Carcinogenesis 16, 327334.[Abstract]
De Camargo, J. L. V., Punyarit, P., and Newberne, P. M. (1985). Early stages of nodular transformation of the B6C3F1 mouse liver induced by choline deficiency. Toxicol. Pathol. 13, 1017.[Medline]
Finkelstein, J. D., Martin, J. J., Harris, B. J., and Kyle, W. E. (1982). Regulation of betaine content of rat liver. Arch. Biochem. Biophys. 218, 169173.[ISI][Medline]
Fullerton, F. R., Hoover, K., Mikol, Y. B., Creasia, D. A., and Poirier, L. A. (1990). The inhibition by methionine and choline of liver carcinoma formation in male C3H mice dosed with diethylnitrosamine and fed phenobarbital. Carcinogenesis 11, 13011305.[Abstract]
Ghoshal, A. K., and Farber, E. (1984). The induction of liver cancer by dietary deficiency of choline and methionine without added carcinogens. Carcinogenesis 5, 13671370.[Abstract]
Goodman, J. I., Ward, J. M., Popp, J. A., Klaunig, K. E., and Fox, T. R. (1991). Mouse liver carcinogenesis: Mechanisms and relevance. Fundam. Appl. Toxicol. 17, 651665.[ISI][Medline]
Knaak, J. B., Leung, H.-W., Stott, W. T., Busch, J., and Bilsky, J. (1997). Toxicology of mono-, di- and triethanolamine. Rev. Environ. Contam. Toxicol. 149, 186.[Medline]
Lehman-McKeeman, L. D., and Gamsky. E. A. (1999). Diethanolamine inhibits choline uptake and phosphatidylcholine synthesis in Chinese hamster ovary cells. Biochem. Biophys. Res. Commun. 262, 600604.[ISI][Medline]
Lehman-McKeeman, L. D., and Gamsky. E. A. (2000). Choline supplementation inhibits diethanolamine-induced morphological transformation in Syrian hamster embryo cells: Evidence for a carcinogenic mechanism. Toxicol. Sci. 55, 303310.
Lombardi, B., and Shinozuka, H. (1979). Enhancement of 2-acetylaminofluorene liver carcinogenesis in rats fed a choline-devoid diet. Int. J. Cancer 23, 565570.[ISI][Medline]
Mathews, J. M., Garner, C. E., Black, S. L., and Matthews, H. B. (1997). Diethanolamine absorption, metabolism and disposition in rat and mouse following oral, intravenous and dermal administration. Xenobiotica 27, 733746.[ISI][Medline]
Mathews, J. M., Garner, C. E., and Matthews, H. B. (1995). Metabolism, bioaccumulation and incorporation of diethanolamine into phospholipids. Chem. Res. Toxicol. 8, 625633.[ISI][Medline]
Mikol, Y. B., Hoover, K. L., Creasia, D., and Poirier, L. A. (1983). Hepatocarcinogenesis in rats fed methyl-deficient, amino acid-defined diets. Carcinogenesis 4, 16191629.[ISI][Medline]
NTP (1999). Toxicology and carcinogenesis studies of diethanolamine in F344/N and B6C3F1 mice (Dermal Studies). NTP TR 478. National Toxicology Program, U. S. Department of Health and Human Services, Public Health Service, National Institutes of Health.
Newberne, P. M., de Camargo, J. L. V., and Clark, A. J. (1982). Choline deficiency, partial hepatectomy, and liver tumors in rats and mice. Toxicol. Pathol. 10, 95106.
Newberne, P. M., and Rogers, A. E. (1986). Labile methyl groups and the promotion of cancer. Annu. Rev. Nutr. 6, 407432.[ISI][Medline]
Newberne, P. M., Suphiphat, V., Lockniskar, M., and de Carmargo, J. L. V. (1990). Inhibition of hepatocarcinogenesis in mice by dietary methyl donors methionine and choline. Nutr. Cancer 14, 175181.[ISI][Medline]
Pelech, S. L., and Vance, D. E. (1984). Regulation of phosphatidylcholine biosynthesis. Biochim. Biophsys. Acta 779, 217251.
Pomfret, E. A., daCosta, K.-A., Schurman, L. L., and Zeisel, S. H. (1989). Measurement of choline and choline metabolite concentrations using high-performance liquid chromatography and gas chromatography-mass spectrometry. Anal. Biochem. 180, 8590.[ISI][Medline]
Pomfret, E. A., daCosta, K.-A., and Zeisel, S. H. (1990). Effects of choline deficiency and methotrexate upon rat liver. J. Nutr. Biochem. 1, 533541.[ISI]
Rogers, A. E. (1995). Methyl donors in the diet and response to chemical carcinogens. Am. J. Clin. Nutr. 61(Suppl. 3) 659S665S.[Abstract]
Rogers, A. E., Nields, H. M., and Newberne, P. M. (1987). Nutritional and dietary influences on liver tumorigenesis in mice and rats. Arch. Toxicol. 10(Suppl.), 231243.
Schneider, W. J., and Vance, D. E. (1978). Effect of choline deficiency on the enzymes that synthesize phosphatidylcholine and phosphatidylethanolamine in rat liver. Eur. J. Biochem. 85, 181187.[Abstract]
Sells, M. A., Katyal, S. L., Sell, S., Shinozuka, H., and Lombardi, B. (1979). Induction of foci of altered, -glutamyltranspeptidase-positive hepatocytes in carcinogen-treated rats fed a choline-deficient diet. Br. J. Cancer 40, 274283.[ISI][Medline]
Shivapurkar, N., and Poirier, L. A. (1983). Tissue levels of S-adenosylmethionine and S-adenosylhomocysteine in rats fed methyl-deficient, amino acid-defined diets for one to five weeks. Carcinogenesis 4, 10511057.[ISI][Medline]
Sidransky, H., and Farber, E. (1960). Liver choline oxidase activity in man and in several species of animals. Arch. Biochem. Biophys. 87, 129133.[ISI]
Vance, D. E., Walkey, C. J., and Cui, Z. (1997). Phosphatidylethanolamine N-methyltransferase from liver. Biochim. Biophys. Acta 1348, 142150.[ISI][Medline]
Wagner, J., Danzin, C., Huot-Olivier, S., Claverie, N., and Palfreyman, M. G. (1984). High-performance liquid chromatographic analysis of S-adenosylmethionine and its metabolites in rat tissues: Interrelationship with changes in biogenic catechol levels following treatment with L-dopa. J. Chromatogr. 290, 247262.[Medline]
Wainfan, E., and Poirer, L. A. (1992). Methyl groups in carcinogenesis: Effects on DNA methylation and gene expression. Cancer Res. 52(Suppl. 7), 2071s2077s.[Abstract]
Yao, Z. M., and Vance, D. E. (1988). The active synthesis of phosphatidylcholine is required for very low density lipoprotein secretion from rat hepatocytes. J. Biol. Chem. 263, 29983004.
Yao, Z. M., and Vance, D. E. (1990). Reduction in VLDL, but not HDL, in plasma of rats deficient in choline. Biochem. Cell Biol. 68, 552558.[ISI][Medline]
Yokoyama, S., Sells, M. A., Reddy, T. B., and Lombardi, B. (1985). Hepatocarcinogenic and promoting action of a choline-devoid diet in the rat. Cancer Res. 45, 28342842.[Abstract]
Zeisel, S. H. (1996). Choline. A nutrient that is involved in the regulation of cell proliferation, cell death and cell transformation. In Dietary Fats, Lipids, Hormones, and Tumorigenesis (Heber and Kritchevsky, Eds.), pp. 131141. Plenum Press, New York.
Zeisel, S. H., and Blusztajn, J. K. (1994). Choline and human nutrition. Annu. Rev. Nutr. 14, 269296.[ISI][Medline]
Zeisel, S. H., Zola, T., daCosta, K.-A., and Pomfret, E. A. (1989). Effect of choline deficiency on S-adenosylmethionine and methionine concentrations in rat liver. Biochem. J. 259, 725729.[ISI][Medline]