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Fatty acid ethyl esters: short-term and long-term serum markers of ethanol intake.

In the context of ethanol abuse, trait markers are those that identify a genetic predisposition to ethanol abuse or to development of complications from excess ethanol intake. State markers reflect the likelihood of acute and chronic ethanol intake. Specifically, the trait markers are used to address two major questions: individual predisposition to developing alcoholism and individual susceptibility to developing alcoholic cirrhosis. The state markers are valuable in addressing three separate questions: recent intake of ethanol, evidence of chronic ethanol intake, and evidence of end organ damage as a result of ethanol intake. Clinically useful markers, particularly those detectable in blood and urine, are needed to provide answers to these five questions. Several review articles provide an overview of the state and trait markers of ethanol intake [1-3].

Trait Markers

IS THIS INDIVIDUAL PREDISPOSED TO DEVELOP ALCOHOLISM?

Several markers have been under investigation to address this question, particularly for those whose risk is higher because of a family history of alcoholism [4]. Substantial attention has been given to the presence of the A1 allele of the D2 dopamine receptor [5]. The role of the D2 dopamine receptor allele as a trait marker for development of alcoholism has been widely publicized [5-7] and seriously challenged [8-10]. Platelet monoamine oxidase B has also been proposed as a marker to identify those predisposed to alcohol abuse. It has been reported that a low monoamine oxidase B activity in platelets is a marker for predisposition to alcoholism X11-14]. Subjects with low platelet monoamine oxidase B activities typically have sensation-seeking traits and an inability to abstain from ethanol [13,14]. Use of this assay is not widespread. Finally, a low value for lymphocyte and platelet adenylyl cyclase has also been proposed as a marker to identify those predisposed to alcohol abuse [15,16]. In a recent study, no difference was found in newly identified tetranucleotide polymorphisms in the human adenylyl cyclase type 7 gene between alcoholics and nonalcoholic controls [17]. The assay for adenylyl cyclase has not been popularized as a test for assessing inheritance for alcoholism.

DOES THIS INDIVIDUAL HAVE A HIGH SUSCEPTIBILITY TO DEVELOP ALCOHOLIC CIRRHOSIS?

Even less accepted than the genetic markers for predisposition to alcoholism are the proposed markers for determining susceptibility to cirrhosis by alcohol abusers. Several HLA antigens (B8, BW40, B13, A2, DR3, and DR2) have been proposed as markers to address this question [18]. In addition, the collagen a(I)2 gene polymorphism [19], the alcohol dehydrogenase 3*1 gene [20,21], and the aldehyde dehydrogenase 2*2 allele [22] have been evaluated in preliminary studies as markers for susceptibility to cirrhosis.

State Markers

HAS THERE BEEN RECENT INTAKE OF ETHANOL?

Blood ethanol is a widely accepted marker for recent ethanol intake (within the last 4-6 h). However, the rapid elimination of ethanol from the blood nearly always makes it impossible to assess ethanol ingestion beyond the most recent 6-8 h. Alcohol consumption can lead to increased concentrations of 5-hydroxytryptophol and decreased production of 5-hydroxyindole-3-acetic acid [23,24]. Urinary 5-hydroxytryptophol/5-hydroxyindole3-acetic acid ratios increase in a dose-dependent fashion with consumption of alcohol. The ratio remains increased for 5-15 h after blood ethanol is no longer detectable [24]. Serum fatty acid ethyl ester (FAEE) has recently emerged as a potential marker to assess intake of ethanol [25,261 because it is detectable in the blood both when ethanol is present and long after ethanol has been removed from the circulation. FAEEs are described in detail below.

IS THERE EVIDENCE OF CHRONIC ETHANOL INTAKE?

Obtaining evidence for chronic ethanol intake is often problematic because individuals with chronic alcoholism may abstain long enough before presenting to a physician to have a blood ethanol of zero at the time of the visit. Acetaldehyde adducts to proteins, such as hemoglobin [27], albumin [28], and lipoproteins [29], have been proposed as markers of chronic ethanol ingestion. FAEE is also potentially useful in this setting. An additional marker, carbohydrate-deficient transferrin (CDT), has emerged to identify individuals who have been ingesting large amounts of ethanol for prolonged periods but are not acutely intoxicated at the time of evaluation [1-3, 30].

IS THERE EVIDENCE OF END ORGAN DAMAGE AS A RESULT OF CHRONIC ETHANOL INTAKE?

The most common target organs for ethanol-induced end organ damage are the liver and the pancreas. For that reason, calibrated liver and pancreatic function tests are useful in the evaluation of end organ damage from ethanol abuse. One particular liver function test, the assay for y-glutamyltransferase, has long been used as a marker of liver injury following ethanol intake [3, 31]. This serum enzyme becomes increased more readily than other liver enzymes after an episode of ethanol abuse. In addition, aspartate aminotransferase and alanine aminotransferase have been used in assessment of liver injury. However, the specificity of all of these markers for liver dysfunction as a result of ethanol abuse is low [3]. Injury to the pancreas is assessed with assays for pancreatic amylase and pancreatic lipase, but these tests are also not reflective of pancreatic dysfunction specifically as a result of ethanol abuse [3]. Independent of liver and pancreatic function, an increase of the mean corpuscular volume of red blood cells has also been used as a marker of chronic ethanol intake [3,311. An increased mean corpuscular volume reflects dysfunctional production of red blood cells. Markers of injury to other organs may be valuable in a patient with signs and symptoms related to dysfunction of organs other than the liver and the pancreas, such as serum creatine kinase isoenzyme MB for damage to the heart. As with the markers originating from the liver and the pancreas, however, these markers also cannot specifically implicate alcohol as the cause for the dysfunction.

CDT: Emerging Marker for Chronic Ethanol Intake

Serum transferrin, which has a molecular mass of 80 kDa, is synthesized in the liver. The most important known function of transferrin is to transport and deliver iron. It has two N-linked carbohydrate units that are added to the amino acid chain posttranslationally. The biological half-life of serum transferrin is 6-12 days.

Chronic alcohol intake interferes with the metabolism of several glycoconjugates, of which transferrin is one. Regular high amounts of alcohol consumption result in the appearance of isoforms of serum transferrin that are deficient in their carbohydrate moiety [30]. These isotransferrins are less negatively charged, and thus they have higher isoelectric points than normal transferrin. Because of this, these isoforms can be detected by separation methods on the basis of charge. However, immunoassay is the most commonly used methodology for determination of CDT [32-35].

Daily alcohol intake of more than 60 g of ethanol (~4.5 drinks in the US) for at least 1 week, in most cases, will result in an increased concentration of CDT in the plasma. During alcohol abstinence, the values normalize with a mean half-life for the CDT of 14-17 days [30]. The mechanism for generation of CDT in alcohol abuse may be an acetaldehyde-mediated inhibition of glycosyltransferase. However, this has not yet been established. There have been several recent studies on CDT as a marker for ethanol intake [32-35]. The results of these investigations are that CDT is not a discerning marker for detection of as much as 80 g of ethanol ingested daily for 3 weeks by healthy subjects [32], that the diagnostic detection limit of CDT as a marker for chronic ethanol intake is not sufficient to permit its use as a screening test in the general population [33], that an increased CDT cannot be regarded as a reliable indicator for chronic alcohol abuse in patients with liver disease because such patients may have an increased CDT on the basis of liver disease alone [34], and that changes in blood CDT concentrations of 20-30% may be the most sensitive indicator of a change in ethanol intake [35].

FAEE--An Overview

FAEEs are esterification products of ethanol and fatty acids. As shown in Fig. 1, ethanol can be metabolized by oxidative and nonoxidative pathways [36]. In the oxidative pathway, ethanol can be converted to acetaldehyde through the action of alcohol dehydrogenase, the microsomal ethanol-oxidizing system, or catalase. Acetaldehyde is then subsequently metabolized to acetate through the action of aldehyde dehydrogenase. In one of the nonoxidative pathways of ethanol metabolism, ethanol can be inserted as the head group of a phospholipid to form phosphatidylethanol. This transformation occurs through the action of phospholipase D on phosphatidylcholine in the presence of ethanol. The nonoxidative ethanol pathway that is the focus of this review is the pathway leading to the synthesis of FAEE. This is an enzyme-mediated esterification of fatty acid or fatty acylCoA and ethanol.

[FIGURE 1 OMITTED]

It has long been known that ethanol abuse leads to end organ damage in the liver and pancreas and, to a smaller extent, in the heart and brain. Acetaldehyde has been proposed as a mediator of this organ damage. However, acetaldehyde has been shown to be generated primarily in the liver with little or no synthesis in the pancreas [37-40]. For this reason an ethanol metabolite other than acetaldehyde has been sought to account for the toxicity of ethanol. A 1986 autopsy study involving subjects acutely intoxicated at the time of death demonstrated that the organs most frequently damaged by ethanol abuse, the pancreas and liver, have the highest concentrations of both FAEE and FAEE synthase, the enzyme responsible for FAEE synthesis [41]. An enzyme now known as FAEE synthase has been purified from several different organs [42-44], but it is not clear whether this enzyme is responsible for the bulk of FAEE synthesis. Carboxylester lipase, which has the ability to liberate fatty acids from complex lipids to which they are esterified, has FAEE synthase capability [45]. This observation has raised the possibility that hydrolysis of a fatty acid from a phospholipid or a triglyceride molecule in the presence in ethanol can lead to formation of FAEEs.

To evaluate the biochemical mechanism for FAEE synthesis, secretion, and degradation, and to evaluate the toxic effects of FAEE, it was first necessary to develop a system for solubilization of the highly nonpolar FAEE in aqueous medium. We had found in a clinical study that FAEEs appear in the serum after ethanol ingestion bound to albumin and in the core of lipoproteins with other neutral lipids [25]. With this finding in mind, a method was developed for the solubilization of FAEE in isolated LDL particles [46]. In this method, LDL are isolated, and the core lipids are removed with heptane. FAEEs, which can be synthesized from triglyceride incubated with 0.5 mol/L KOH in ethanol and subsequently purified by solid-phase extraction [47], are added to the core of the delipidated LDL particle. This results in the accumulation of FAEEs into the core of the water-soluble LDL particle.

SYNTHESIS AND SECRETION OF FAAES

We have demonstrated that a human hepatoma cell line (HepG2 cells) exposed to ethanol will synthesize and secrete FAEEs. In these studies, radiolabeled fatty acid is added to HepG2 cells for 12 h, and then the cells are exposed to ethanol for an additional 10 h. The culture medium and cell monolayer are harvested, the lipids are extracted from each, and the FAEEs are isolated from all other lipids and quantitated. We have shown that FAEE synthesis and secretion are linearly correlated to the ethanol concentration in the culture medium of HepG2 cells. We have also demonstrated that secretion is highly dependent on the presence of a carrier for FAEE in the medium. In the absence of any carrier in the medium for the FAEEs, secretion of FAEEs into the medium is very limited. FAEEs secreted into the HepG2 cell culture medium are associated with lipoproteins, most predominantly an HDL secreted by the cells. The secretion of FAEE from HepG2 cells can be interrupted by cycloheximide, brefeldin, and monensin-inhibitors of protein synthesis and various stages of vesicular transport [A. Kabakibi and M. Laposata, unpublished observations].

TOXICITY OF FAEES

There have been several reports suggesting that FAEEs are toxic metabolites of ethanol. In 1983, FAEEs in emulsions were shown to cause uncoupling of oxidative phosphorylation in mitochondria [48]. In 1986, as noted earlier, an autopsy study demonstrated the presence of FAEEs selectively in the organs damaged by ethanol abuse. However, no causal association of FAEE for toxicity was shown in this investigation [41]. In 1988, FAEEs in emulsions were found to produce changes in membrane fluidity in synaptosomal membranes [49]. In 1993, FAEEs in emulsions were found to increase rat pancreatic lysosomal fragility [50]. In none of these studies, however, were FAEEs shown to be toxic for intact cells, and there was little acceptance after these reports of the suggestion that FAEEs are cytotoxic.

For that reason, we performed a study with HepG2 cells incubated with LDL containing FAEE in the core of a human LDL particle [51]. The HepG2 cells were incubated with the FAEEs in LDL for 12 h, and tritiated thymidine was then added for 5 h. Ethyl oleate and ethyl arachidonate substantially inhibited the proliferation of HepG2 cells, while native LDL and LDL reconstituted with cholesterol esters or triglycerides had no effect (Fig. 2). These two different FAEE species were also shown to decrease the synthesis of [[sup.35]5]methionine-labeled protein by the HepG2 cells. Thus, this study demonstrated that FAEEs could be toxic for intact cells.

To determine whether FAEEs could be toxic in vivo, we performed a study in which FAEEs in reconstituted LDL particles were delivered as an intraarterial bolus followed by subsequent infusion into the circulation of rats [52]. Control animals received saline or LDL reconstituted with cholesterol esters. After periods up to 12 h, the animals were killed, and blood and pancreas were removed for analysis. The toxicity to the pancreas was determined by assessment of edema formation, measurement of trypsinogen activation peptide for pancreatic cell injury, and histologic and electron microscopic examination of the pancreas. The increase over control values in edema formation and in trypsinogen activation peptide concentrations 3 and 6 h after infusion of FAEEs were highly statistically significant (P <0.001 for all comparisons). Ultrastructurally, the cells exposed to FAEE showed dilatation of the endoplasmic reticulum and an increased number of lipid droplets and secondary lysosomes.

[FIGURE 2 OMITTED]

[FIGURE 3 OMITTED]

All of these measurements demonstrated that only lipoprotein particles containing FAEEs produced injury to the pancreas. With evidence that FAEEs can produce a toxic effect, we investigated whether orally ingested FAEEs, used clinically to supplement patients with specific fatty acids, can be toxic in vivo. Before the availability of FAEEs, fatty acids used for therapeutic purposes were provided as triglycerides. To evaluate whether FAEE supplements are associated with organ toxicity, we first evaluated the degradation of FAEE in the gastrointestinal tract and in the blood [53]. Radiolabeled FAEEs were delivered as an oil directly into the rat stomach through a gastrostomy. Blood was collected from each rat at 5, 15, 30, 60, 90, and 120 min, after which the animal was killed and the organs were harvested. The organ distribution of total radioactivity from the radiolabeled FAEEs 2 h after delivery into the stomach, for both radiolabeled ethyl oleate and radiolabeled ethyl eicosapentaenoate, indicated that the radioactivity was largely present in the gastrointestinal tract. The highest amounts were in the stomach, duodenum, jejunum, and liver. When the radioactive lipid classes were quantitated in these organs to determine the percentage of radioactivity remaining as FAEE, only a partial hydrolysis of the FAEE was found in the stomach. In the duodenum, however, there were no residual FAEEs. This suggests that lipases in the gastrointestinal tract, primarily in the duodenum, can result in hydrolysis of the FAEEs and thereby limit any toxic effect from FAEE supplements.

Because of the likely absorption into the blood of undegraded FAEEs through the stomach, we evaluated the hydrolysis of FAEEs in LDL in the vascular compartment after intraarterial injection into the rat. We demonstrated that the degradation of FAEEs in the blood is extremely rapid, with a half-life of 58 s. This provides additional evidence that FAEEs ingested as fatty acid supplements are unlikely to produce toxic effects.

FAEES AS MARKERS FOR ACUTE AND CHRONIC ETHANOL INTAKE

We have recently reported that FAEEs may be useful as markers for both acute and chronic ethanol intake [26]. We performed a study in which seven subjects were given ethanol to drink at a controlled rate over 90 min. Multiple samples were then collected from the subjects for blood ethanol and serum FAEEs for up to 24 h. The results from this study indicate that the concentration of FAEE in the blood closely parallels the concentration of blood ethanol (Fig. 3). Importantly, however, the serum FAEEs in these subjects, who all achieved blood ethanol concentrations >1.5 g/L (1500 mg/L, 32.5 mmol/L), were still detectable 24 h after ethanol ingestion (Fig. 4). Thus, this observation identifies individuals who have ingested ethanol within 24 h. Individuals who had ethanol values very slightly above baseline and <0.10 g/L ethanol (100 mg/L, 2.2 mmol/L) and would be considered negative for ethanol in our clinical laboratory were all found to be positive for FAEE. This suggests that FAEE may be a more discerning marker for ethanol intake than ethanol itself. Thus, serum FAEE may evolve into both a short-term and a long-term marker of ethanol ingestion.

[FIGURE 4 OMITTED]

[FIGURE 5 OMITTED]

[FIGURE 6 OMITTED]

Algorithm to Determine Recent Intake of Ethanol

Figure 5 shows the algorithm to assess recent intake of ethanol. This algorithm begins with a test for blood ethanol. If the answer is negative, the person evaluating the patient should assess the degree of suspicion for ethanol intake within the past 24 h. If there is no suspicion, then the evaluation can be ended. However, if there is still a suspicion of ethanol intake, an assay for serum FAEE would be valuable. A negative blood ethanol with a positive FAEE is consistent with ethanol intake 4-24 h before blood collection.

If the test for blood ethanol is positive and confirming the positive blood ethanol or assessing the timing of ethanol intake is desired, an assay for serum FAEE could be performed. If the assay for serum FAEE is positive, it can be concluded that ethanol intake has occurred 0-6 h before blood collection. If the FAEE test is negative, the ethanol and FAEE tests should be repeated because a positive blood ethanol with a negative FAEE is not known to occur.

Algorithm to Assess Chronic Ethanol Intake and End Organ Damage in the Absence of Acute Intoxication

In this algorithm (Fig. 6), the first step for a patient suspected of chronic ethanol abuse is to document a negative blood ethanol to rule out ethanol intake within the past 6 h. Assuming that the result for blood ethanol is negative, the next step is to perform an FAEE assay to assess ethanol intake within the last 24 h. If FAEEs are detected, it can be concluded that substantial ethanol intake has occurred within the last 24 h, and therefore, chronic ethanol intake should be suspected. With or without detection of serum FAEE, an assay for CDT to assess chronic alcohol intake should be performed. If the CDT assay is positive, whether or not FAEEs are present, evidence of substantial chronic ethanol intake exists, and the patient should be evaluated with clinical and laboratory evaluations of the liver and pancreas. If the CDT assay is negative and the person evaluating the patient still suspects chronic ethanol intake, the CDT assay should be repeated at a later date. If the CDT result is positive at that time, the patient should be evaluated as above for end organ damage. If the CDT assay is repeatedly negative but suspicion of ethanol abuse persists, other markers for chronic ethanol intake (many are in development) could be sought. If any are positive, this may provide evidence for chronic ethanol intake and lead to a reevaluation of the patient at a later date with FAEE and CDT assays and tests for end organ damage.

If none of the new ethanol intake markers are available and the CDT has been repeatedly negative, the next question is to ask whether FAEEs were detectable in the initial analysis. If serum FAEEs have never been detected in a patient who is CDT-negative on two occasions and has no other markers for chronic ethanol intake, there is no evidence for chronic ethanol intake. However, if FAEEs are detected with a repeatedly negative CDT, the question of whether it is the first time for FAEE detection becomes important. If the FAEEs have been detected on more than one occasion, this is strong evidence for chronic ethanol intake, even with a negative CDT test. If this is the first time for FAEE detection, it would be most prudent to repeat the evaluation from the beginning at a later date.

A clinical need for markers of ethanol intake exists. Although tests of liver function provide some evidence for excess ethanol intake, several newer markers, notably CDT and FAEE, could become widely used. Ongoing work with these new indicators of ethanol intake should provide important information regarding their clinical utility. An initial proposal for their clinical use is shown in the algorithms provided in this report (Figs. 5 and 6).

Received February 20, 1997; revised April 30, 1997; accepted May 16, 1997.

References

[1.] Rosman AS, Lieber CS. The diagnostic utility of laboratory tests in alcoholic liver disease [Review]. Clin Chem 1994;40:1641-51.

[2.] Chan AWK. Recent developments in detection and biological indicators of alcoholism. Drugs Society 1993;8:31-67.

[3.] Conigrave KM, Saunders JB, Whitfield JB. Diagnostic tests for alcohol consumption. Alcohol Alcohol 1995;30:13-26.

[4.] Schuckit MA, Smith TL. An 8-year follow-up of 450 sons of alcoholic and control subjects. Arch Gen Psychiatry 1996;53:20210.

[5.] Blum K, Noble EP, Sheridan PJ, Montgomery A, Ritchie T, Jagadeeswaran P, et al. Allelic association of human dopamine DZ receptor gene in alcoholism. JAMA 1990;263:2055-60.

[6.] Blum K, Noble EP, Sheridan PJ, Finley 0, Montgomery A, Ritchie T, et al. Association of the A1 allele of the DZ dopamine receptor gene with severe alcoholism. Alcohol 1991;8:409-16.

[7.] Parsian A, Todd RD, Devor EJ, O'Malley KL, Suarez BK, Reich T, Cloninger CR. Alcoholism and alleles of the human dopamine DZ receptor locus: studies of association and linkage. Arch Gen Psychiatry 1991;48:655-63.

[8.] Bolos AM, Dean M, Lucas-Derse S, Ramsburg M, Brown GL, Goldman D. Population and pedigree studies reveal a lack of association between the dopamine D2 receptor gene and alcoholism. JAMA 1990;264:3156-60.

[9.] Gelernter J, O'Malley S, Risch N, Kranzler HR, Krystal J, Merikangas K, et al. No association between an allele at the D2 dopamine gene (DRD2) and alcoholism. JAMA 1991;266:1801-7.

[10.] Gelernter J, Goldman D, Risch N. The A1 allele at the D2 dopamine receptor gene and alcoholism. a reappraisal. JAMA 1993;269:1673-7.

[11.] Schuckit MA, Shaskon E, Duby J. Platelet MAO activities in relatives of alcoholics and controls. Arch Gen Psychiatry 1982; 39:137-40.

[12.] Sullivan JL, Baenziger JC, Wagner DL, Rauscher FP, Numberger JI Jr, Holmes JS. Platelet MAO in subtypes of alcoholism. Biol Psychiatry 1990;27:911-22.

[13.] von Knorring L, Oreland L. Platelet MAO activity in type 1/type 2 alcoholics. Alcoholism Clin Exp Res 1996;20:224A-30A.

[14.] Buschbaum MS, Coursey RD, Murphy DL. The biochemical highrisk paradigm: behavioural and familial correlates of low monoamine oxidase activity. Science 1976;194:339-41.

[15.] Diamond IB, Wrubel B, Estrin W, Gordon A. Basal and adenosinereceptor-stimulated levels of cAMP are reduced in lymphocytes from alcoholic patients. Proc Natl Acad Sci U S A 1987;84: 1413-6.

[16.] Tabakoff B, Hoffman PL, Lee JM, Saito T, Willard B, De Leon-Jones F. Differences in platelet enzyme activity between alcoholics and nonalcoholics. N Engl J Med 1988;318:134-9.

[17.] Hellevvo K, Welborn R, Menninger JA, Tabakoff B. Human adenylyl cyclase type 7 contains polymorphic repeats in the 3' untranslated region: investigations of association with alcoholism. Am J Med Genet 1997;74:95-8.

[18.] Dover EJ, Cloninger CR. Genetics of alcoholism. Annu Rev Genet 1989;23:19-36.

[19.] Weiner FR, Eskries DS, Compton KV, Orrego H, Zem M. Haplotype analysis of a type I collagen gene and its association with alcoholic cirrhosis in man. Mol Aspects Med 1988;10:159-68.

[20.] Day CP, Bashir R, James OFW, Bassendine MF, Crabb DW, Thomasson HR, et al. Investigation of the role of polymorphisms at the alcohol and aldehyde dehydrogenase loci in genetic predisposition to alcohol-related end-organ damage. Hepatology 1991; 14:798-801.

[21.] Poupon RE, Nalpas B, Coutelle C, Fleury B, Couzigou P, Higueret D (French Group for Research on Alcohol and Liver). Polymorphism of the alcohol dehydrogenase, alcohol and aldehyde dehydroge nase activities: implications in alcoholic cirrhosis in white patients. Hepatology 1992;15:1017-22.

[22.] Enomoto N, Takase S, Takada N, Takada A. Alcoholic liver disease in heterozygotes of mutant and normal aldehyde dehydrogenase-2 genes. Hepatology 1991;13:1071-5.

[23.] Helander A, Beck 0, Jacobsson G, Lowenmo C, Wikstrom T. Time course of ethanol-induced changes in serotonin metabolism. Life Sci 1993;53:847-55.

[24.] Helander A, Beck 0, Jones AW. Laboratory testing for recent alcohol consumption: comparison of ethanol, methanol, and 5-hydroxytryptophol. Clin Chem 1996;42:618-24.

[25.] Doyle KM, Bird DA, AI-Salihi S, Hallaq Y, Cluette-Brown JE, Goss KA, Laposata M. Fatty acid ethyl esters are present in human serum after ethanol ingestion. J Lipid Res 1994;35:428-37.

[26.] Doyle KM, Cluette-Brown JE, Dube DM, Bernhardt TG, Morse CR, Laposata M. Fatty acid ethyl esters in the blood as markers for ethanol intake. JAMA 1996;276:1152-6.

[27.] Lin RC, Shahidi S, Kelly TJ, Lumeng C, Lumeng L. Measurement of hemoglobin-acetaldehyde adduct in alcoholic patients. Alcohol Clin Exp Res 1993;17:669-74.

[28.] Niemela 0. Acetaldehyde adducts of proteins: diagnostic and pathogenic implications in diseases caused by excessive alcohol consumption. Scand J Clin Lab Invest 1993;213(Suppl):45-54.

[29.] Wehr H, Rodo M, Lieber CS, Baraona E. Acetaldehyde adducts and autoantibodies against VLDL and LDL in alcoholics. J Lipid Res 1993;34:1237-44.

[30.] Stibler H. Carbohydrate-deficient transferrin in serum: a new marker of potentially harmful alcohol consumption reviewed. Clin Chem 1991;37:2029-37.

[31.] Chick J, Kreitman N, Plant M. Mean cell volume and [gamma]-glutamyltranspeptidase as markers of drinking in working men. Lancet 1981;i:1249-51.

[32.] Lesch OM, Walter H, Antal J, Heggli D-E, Kovacz A, Leitner A, et al. Carbohydrate-deficient transferrin as a marker of ethanol intake: a study with healthy subjects. Alcohol Alcohol 1996;31:265-71.

[33.] Vermes I, van den Bergh FAJTM. Clinical utility of carbohydrate deficient transferrin to detect alcohol abuse in a general population [Letter]. Clin Chem 1996;42:2048-9.

[34.] Radosavljevic M, Temsch E, Hammer J, Pfeffel F, Mayer G, Renner F, et al. Elevated levels of serum carbohydrate deficient transferrin are not specific for alcohol abuse in patients with liver disease. J Hepatol 1995;23:706-11.

[35.] Anton RF, Sillanaukee P. The use of carbohydrate deficient transferrin as an indicator of alcohol consumption during treatment and follow-up. Alcoholism Clin Exp Res 1996;20:54A-6A.

[36.] Lieber CS. Medical disorders of alcoholism. N Engl J Med 1995; 333:1058-65.

[37.] Lieber CS, DeCarli LM. Ethanol oxidation by hepatic microsomes: adaptive increase after ethanol feeding. Science 1968;162: 917-8.

[38.] Lochner A, Cowley R, Brink AJ. Effect of ethanol on metabolism and function of perfused rat heart. Am Heart J 1969;78:770-80.

[39.] Raskin NH, Sokoloff L. Enzymes catalyzing ethanol metabolism in neural and somatic tissues of the rat. J Neurochem 1972;19: 273-82.

[40.] Hamamoto T, Yamada S, Hirayama C. Nonoxidative metabolism of ethanol in the pancreas: implication in alcoholic pancreatic damage. Biochem Pharmacol 1990;39:241-5.

[41.] Laposata EA, Lange LG. Presence of nonoxidative ethanol metabolism in human organs commonly damaged by ethanol abuse. Science 1986;231:497-9.

[42.] Mogelson S, Lange LG. Nonoxidative ethanol metabolism in rat myocardium: purification to homogeneity of fatty acid ethyl ester synthase. Biochemistry 1984;23:4075-81.

[43.] Sharma R, Gupta S, Singhal SS, Ahmad H, Haque A, Awasthi YC. Independent segregation of glutathione Stransferase and fatty acid ethyl ester synthase from pancreas and other human tissues. Biochem J 1991;275:507-13.

[44.] Tsujita T, Horomichi 0. Fatty acid ethyl ester synthase in rat adipose tissue and its relationship to carboxylesterase. J Biol Chem 1992;267:23489-94.

[45.] Tsujita T, Okuda H. The synthesis of fatty acid ethyl ester by carboxylester lipase. Eur J Biochem 1994;224:57-62.

[46.] Bird DA, Szczepiorkowski ZM, Trace VC, Laposata M. Low-density lipoprotein reconstituted with fatty acid ethyl esters as a physiological vehicle for ethyl ester delivery to intact cells. Alcohol Clin Exp Res 1995;19:1265-70.

[47.] Bernhardt TG, Cannistraro PA, Bird DA, Doyle KM, Laposata M. Purification of fatty acid ethyl esters by solid-phase extraction and high-performance liquid chromatography. J Chromatogr B Biomed Appl 1996;675:189-96.

[48.] Lange LG, Sobel BE. Mitochondrial dysfunction induced by fatty acid ethyl esters, myocardial metabolites of ethanol. J Clin Invest 1983;72:724-31.

[49.] Hungund BL, Goldstein DB, Villegas F, Cooper TB. Formation of fatty acid ethyl esters during chronic ethanol treatment in mice. Biochem Pharmacol 1988;37:3001-4.

[50.] Haber PS, Wilson JS, Apte MV, Pirola RC. Fatty acid ethyl esters increase rat pancreatic lysosomal fragility. J Lab Clin Med 1993; 121:259-64.

[51.] Szczepiorkowski ZM, Dickerson GR, Laposata M. Fatty acid ethyl esters decrease human hepatoblastoma cell proliferation and protein synthesis. Gastroenterology 1995;108:515-22.

[52.] Werner J, Laposata M, Fernandez-del Castillo C, Saghir M, lozzo RV, Lewandrowski KB, Warshaw AL. Pancreatic injury in rats induced by fatty acid ethyl ester, a non-oxidative metabolite of alcohol. Gastroenterology 1997;113:286-94.

[53.] Saghir M, Werner J, Laposata M. The rapid and extensive hydrolysis of fatty acid ethyl esters, toxic nonoxidative metabolites of ethanol, in the gastrointestinal tract and circulation of rats. Am J Physiol 1997, in press.

MICHAEL LAPOSATA*

*Address for correspondence: Room 235, Gray Bldg., Massachusetts General Hospital, Fruit St., Boston, MA 02114. Fax 617-726-3256; e-mail LAPOSATAMI@A1.mgh.harvard.edu.
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Title Annotation:Beckman Conference
Author:Laposata, Michael
Publication:Clinical Chemistry
Date:Aug 1, 1997
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