Am J Clin Nuir l992;56:467-74. Printed in USA. © 1992 American Society for Clinical Nutrition 467

Reduced tissue arachidonic acid concentration with chronicethanol feeding in miniature pigs14

Manabu T Nakamura, Anna B Tang, Jesus Villanueva, Charles H Halsted, and Stephen D Phinney

ABSTRACF The effect ofethanol feeding on the essential

fatty acid content of tissues has been contradictory. To define

the effect, we analyzed fatty acid profiles in various tissues from

five miniature pigs fed daily 105 Id basal diet/kg body wt and

146 kJ ethanol/kg body wt, and also five control pigs pairfed

the same amount of basal diet but with corn starch substituted

for ethanol. After 12 mo, biopsy samples were taken, and tissue

fatty acid profiles were analyzed. In the phospholipid fraction

from the ethanol group there was a uniform decrease in ara-

chidonic acid (AA) and an increase in oleic acid in liver, serum,

and muscle. AA was consistently decreased in the triglyceride

fractions ofliver, serum and subcutaneous adipose ofthe ethanol

group. Possible explanations for this general reduction in tissue

AA with ethanol feeding include decreased activities of �6 and

� desaturases, and a displacement of AA from lipid fractions

by other fatty acids. Am J Clin Nutr 1992;56:467-74.

KEY WORDS Pig, ethanol, arachidonic acid, fatty acid

profile, essential fatty acid metabolism

Introduction

The essentiality of arachidonic acid (AA) in animals is well

established. A deficiency of AA causes growth stunting, der-

matosis, and a variety ofmetabolic disorders in animals including

humans (1). AA is one of the major fatty acids (FAs) in phos-

pholipids (PLs) of biological membranes and is a precursor of

eicosanoids, which have a variety of regulatory roles in physi-

ological functions. AA can be synthesized from linoleic acid

(LA) in most animals through �6 desaturase, elongase, and �5

desaturase (Fig 1). It is generally assumed that humans can syn-thesize enough AA from LA to fulfill their daily requirement,

and that dietary intake of linoleic acid from vegetable sources

provides copious substrate for this process.

Many studies have reported alterations of LA and AA con-

centrations in various tissues with chronic ethanol intake both

in humans and animal models (2-22). These studies have also

suggested that the changes in essential fatty acids (EFAs) may

be related to the pathology of alcoholism with respect to fatty

liver (3, 18-20), inefficient energy utilization (8), hemostasis (6,

7, 12), or changes in blood pressure (6). However, the results of

these studies were sometimes contradictory, and there is yet nogeneral agreement on the effect of ethanol on EFA metabolism

or the role of altered EFA profile on the pathology induced by

ethanol consumption.

In human studies, Holman and Johnson (2) observed de-

creased AA in serum PLs of alcoholics, whereas LA was un-

changed. Cairns and Peters (3) reported a decrease in the AA-

LA ratio in the liver PLs of human alcoholics with fatty liver.

Alling et al (4, 5) found a decrease in LA but not in AA in serum

PLs. Also, there are observations ofincreased bleeding time and

altered eicosanoid synthesis in human alcoholics (6, 7).

In animal models ofalcoholism, the most common abnormal

pattern of EFAs is a decreased AA-LA ratio in tissue PLs ofethanol-fed animals, suggesting a deficiency ofAA (8-12). How-

ever, this change was not uniformly observed (1 3-1 7). In par-

ticular, the effect of ethanol on the FA profile of liver mito-

chondria is inconsistent (8-1 1 , 1 3, 14), and the relationship be-

tween the FA composition in mitochondrial membrane and the

respiratory function of mitochondria is not conclusive (8, 13,

14). The effects of dietary supplementation with AA are also

controversial in animal studies. Goheen et al (18) first reported

that dietary AA alleviated alcoholic fatty liver and increased

weight gain in ethanol-fed rats. Karpe et al (19) and Segarnick

et al (20) showed alleviation of ethanol-induced fatty liver by

administration of AA or -y-linolenic acid, respectively. On the

other hand, Goheen et al (2 1) failed to reproduce their results,

and Stem et al (22) recently reported no beneficial effect of dietary

AA on fatty liver or weight gain.

These discrepancies in both human and animal studies maybe due to many factors such as animal species, tissues examined,diet composition, and experimental procedures for ethanol ad-

ministration (level, duration, and form). Differences in dietary

fat may be another confounding factor because the amount and

composition of dietary fat affect both EFA profile (1) and pa-

thology ofalcoholism (23-25). Another likely factor contributing

to the inconsistent results is the focus of most studies on the FA

composition ofonly one or two tissues, rather than determining

whole-body EFA status in ethanol-fed animals by measuring

From the Division ofClinical Nutrition and Metabolism, School ofMedicine, University ofCalifornia, Davis.

2 Presented in part at the annual meeting ofthe American Society for

Clinical Nutrition, Seattle, WA, May 1991.3 Supported by grants NIAAA 06938 and NIDDK 35747 and a grant

from the Alcoholic Beverage Medical Research Foundation.4Address reprint request to SD Phinney, Division of Clinical Nutrition

and Metabolism, School of Medicine, TB-156, University of California,Davis, CA 95616.

Received October 1, 1991.Accepted for publication March 12, 1992.

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468 NAKAMURA ET AL

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FIG 1. Anabolic pathway of long-chain fatty acids. The arrows withtail show conversion between fatty acids. The arrow heads show actingpoints of enzymes listed in the leftmost column. EPA, eicosapentaenoicacid; DHA, docosahexaenoic acid.

major pools like muscle for PLs and subcutaneous adipose for

triglycerides (TGs).

The objectives of the present study were 1) to define the effect

of chronic ethanol feeding on EFA status in a pig model and 2)

to assess possible mechanisms of this ethanol effect. To achieve

the objectives we conducted a comprehensive study examining

the FA profile in four major lipid fractions in a number of tissues

with control of dietary composition, intake, and weight gain.

Materials and methods

Animals and diets

Details of animals and diets have been reported else-

where (26). Briefly, ten Hanford miniature pigs (University of

California, Davis, CA), both barrow and female, were paired by

weight and sex for control and ethanol diets. At the start of the

experiment they were 8 mo of age with mean weights of 46.5

± 2.6 and 48.0 ± 3.6 kg(I ± SD) for control and alcohol groups,

respectively. The dietary ethanol amount was gradually in-

creased, reaching 60% of total metabolizable energy 4 mo after

beginning the experiment. For 12 mo after reaching 60% of total

metabolizable energy, the maximum amount, pigs in the ethanol

group were fed a basal diet (8.3 g/kg body wt) and ethanol (5 g/

kg body wt). The basal diet was made of a pelleted commercial

diet (Lab Mini-Pig Chow Starter 5080; Purina Mills, Richmond,

IN), corn oil, and fiber (Solka Hoc; James River Corp,

Hackensack, NJ), supplying 1.5 g protein, 0.6 g fat, 3.9 g car-

bohydrate, and 105 U metabolizable energy/kg body wt. The

dietary fatty acids were primarily LA and contained only trace

amounts (< 0.07% of total dietary fat) of AA and eicosapenta-

enoic acid. Vitamins and minerals supplied by the basal diet

met The National Research Council’s (NRC’s) requirement for

swine (27). The control pigs were fed the same amount of the

basal diet as that of the paired ethanol-fed pigs but ethanol was

replaced with corn starch; that is, both groups were given equal

nutrients per body weight except for cornstarch and ethanol.

The amount of corn starch was adjusted to half the metabolizable

energy of ethanol, so that weight gain of the control pigs was

comparable with that of the ethanol-fed pigs. The diet was divided

into three portions and fed at 0800, 1200, and 1700. The animals

in both groups routinely ate the given meal by the time of next

feeding.

Blood samples were taken every other month from the vena

cava. Twelve months after beginning the maximal ethanol feed-

ing, the pigs were fasted overnight; pretreated with ketamine (20

mg/kg), acepromazine (0.2 mg/kg), and atropine sulfate (0.04

mgjkg); and anesthetized with halothane. Biopsy samples of liver,

omentum adipose, abdominal muscle, and subcutaneous adipose

were taken for lipid analysis. The samples were washed with ice-

cold saline, blotted, and stored at -70 #{176}Cuntil analysis. This

protocol was approved by the Animal Use and Care Adminis-

trative Advisory Committee at the University of California at

Davis.

Lipid extraction and separation

Tissue lipids were extracted by the method of Folch et al (28).

Tissues (� 10 mg for adipose and �250 mg for liver and muscle)

were weighed and homogenized in 5 mL chloroform:methanol

(2: 1 by volume) with a Ten Broek (Corning Glass Works,

Horsehead, NY) tissue grinder under ice-cold conditions. Before

extraction 100 � each ofdiheptadecanoyl phosphatidyl choline

(Sigma Chemical, St Louis), heptadecanoic acid, triheptadeca-

noin, and cholesteryl heptadecanoate (Nu Chek Prep, Elysian,

MN) were added as internal standards. All subsequent steps were

performed under nitrogen to prevent peroxidation of polyun-

saturated fatty acids. After the addition of 3 mL water, the ho-

mogenate was vortexed and centrifuged at 1000 X g for 3 mm

at room temperature. Lipids were recovered in the chloroform

layer. For serum lipids, 2 mL serum was vortexed with 6 mL

chloroform:methanol; the mixture was then centrifuged and the

aqueous layer was removed. The chloroform layer was processed

through Whatman #1 filter paper to remove the proteins. The

chloroform extract was blown down to dryness with nitrogen,

resolubilized with a minimum amount (� 160 �tl) of chloroform,

and applied on a prerun silica gel H thin-layer chromatography

plate. The plate was developed with petroleum ether:diethyl

ether:acetic acid (80:20: 1 by volume), and the lipids were sap-

arated into four fractions: PLs, free FAs, TGs, and cholesteryl

esters (CEs). The plate was sprayed with 0. 1% dichlorofluorescein

in ethanol and the lipids were identified under ultraviolet light.

Fatty acid analysis

This procedure was based on the method by Holman et al

(29). The silica gel containing each lipid fraction was scraped

into a tube, and 3 ml ofa methylating reagent (5 wt:vol % acetyl

chloride in methanol) was added. The lipids were incubated at

75 #{176}Cfor 45 mm for free FAs and TGs and for 90 mm for PLs

and CEs. The fatty acid methyl esters were extracted by petro-

leum ether after 2 ml each of water and petroleum ether were

added. The petroleum ether extract was evaporated to dryness

with nitrogen, resolubilized with heptane, and quantitated by

gas chromatography. A Hewlett-Packard (Sunnyvale, CA) 5890

gas chromatograph with a 50 m X 0.25 mm fused silica capillary

column, Bonded 007 FFAP (Quadrex, New Haven, CT), was

programmed for a temperature increase of 2 #{176}C/min from I 90

to 220 #{176}Cand then 0.4 #{176}C/minto 232 #{176}C.Helium was used as

carrier gas with a flow rate of 1 .4 mL/min. The split ratio was

1:65. The injector and detector temperatures were 250 and 270

#{176}C,respectively. The peaks were identified by comparison with

authentic standards (Nu Chek Prep, Supelco, Bellefonte, PA).

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REDUCED ARACHIDONIC ACID BY ETHANOL 469

The peak areas were calculated with a Hewlett-Packard 3396 A

integrator, and each fatty acid was quantitated by comparison

with the internal standard, 17:0. As many as 40 FA methyl esters

from 12:0 to 22:6w3 were identified and quantitated with this

procedure.

Blood analysis

Serum enzymes were assayed with a DACOS automated an-

alyzer (Coulter Electronics, Hialeah, FL). Blood ethanol was

analyzed enzymatically with a commercial kit (Sigma Diagnos-

tics, St Louis).

Histology

Liver-biopsy samples fixed with formalin were examined after

hematoxylin/eosin or Masson’s trichrome staining. Histochem-

ical assay of succinate dehydrogenase activity was performed

with frozen liver sample by the method of French (30).

Statistical analysis

Data were presented as I ± SE. Data ofethanol-fed and control

groups were compared by the two tailed t test, and a P value

< 0.05 was considered statistically significant.

Results

The details of animal characteristics were reported elsewhere

(26). Briefly, in the ethanol groups the mean blood ethanol con-

centration after 7 mo of ethanol feeding exceeded 43 mmol/L

after the first meal and persisted over a 12-h period with sub-

sequent feedings. Serum enzymes (alkaline phosphatase, aspar-

tate aminotransferase, alanine aminotransferase, ‘y-glutamyl

transferase, and lactate dehydrogenase) in ethanol-fed pigs tended

to be elevated throughout the experimental period, sometimes

but not always statistically significant. The histological exami-

nation of the liver showed a centrilobular shift of succinate de-

hydrogenase in the ethanol group, showing mitochondrial dam-

age (30). However, fatty liver, stainable collagen, and megami-

tochondria were not observed in the ethanol group.

Table 1 shows the FA profile and the amount of each lipid

fraction in the liver. The notation and metabolic relationships

of polyunsaturated FAs are shown in Figure 1 . Uniformly ob-

served changes in all four fractions were a decrease in AA and

22:5w3 and an increase in oleic acid and l8:3w3 in the ethanol

group, although the changes in these FAs were not statistically

significant in one of the four lipid fractions. The amounts of

TGs and free FAs in the ethanol-fed pigs were significantly higherthan those in the control pigs, suggesting impairment of lipid

metabolism by ethanol.

Table 2 shows the FA profile and the amount of each serum

lipid fraction. A significant decrease in AA and an increase in

oleic acid were observed in the serum PL, TG, and CE fractions

in the ethanol group, similar to the changes in the liver, whereas

the serum free FA fraction showed little change with ethanol

feeding. The pattern of changes in the TG fraction with ethanol

feeding was similar in the liver and serum; that is, an increase

in stearic (18:0) and oleic acids and a decrease in LA and AA

(Tables 1 and 2).

Table 3 shows the FA profile of PLs from abdominal muscle

and TGs from subcutaneous and omentum adipose. In the

ethanol group a decrease in AA was observed in the muscle PLs

and the subcutaneous adipose TGs but not in omentum adipose

TGs, which showed a remarkable resistance to the effect of

ethanol. Oleic acid was increased in muscle PLs with ethanol

feeding, but not in TGs from subcutaneous or omental adipose.

In the subcutaneous adipose there were no changes in the major

FAs between the two groups. The only significant change inunsaturated FAs with ethanol feeding was a decrease in I 8:3w6

and AA, the products of �6 and �5 desaturase, respectively.

In the PL fraction similar changes were observed among liver,serum, and muscle (Tables 1-3). The most consistent change

with ethanol feeding was an increase in oleic acid and a decrease

in AA. A nonsignificant and modest increase in LA was also

observed in each tissue. In the PLs ofabdomirial muscle (Table3), products of �6 and �5 desaturases (l8:3w6 and AA) were

significantly lower in ethanol-fed pigs than in controls with non-

significant elevation of substrates (LA and 20:3w6). The liver

PLs showed the same tendency although the values were notstatistically significant (Table 1).

Figure 2 shows the concentrations of free FAs in the liver. As

shown in the upper panel, there was a marked increase in theabsolute amount of oleic acid and LA in ethanol-fed pigs, but

there was no change in the concentration of AA. The lower

panel shows the free FAs in the desaturase-elongase pathway in

the liver. In the free FA fraction, all the precursors were signif-

icantly increased in ethanol-fed pigs but no decrease was observed

in the desaturase products.

Discussion

The major finding ofthis study is that in ethanol-fed pigs, AAwas decreased in all tissues and lipid fractions examined, in-

cluding the largest PL pool (muscle) and the largest TG pool

(subcutaneous adipose) with the exception of the TG fraction

from omentum adipose and the free FA fraction from serum.A concomitant increase of oleic acid was also observed except

for the TO fractions from subcutaneous adipose and omentum

adipose. To our knowledge this is the first comprehensive studyof the effect of ethanol feeding on the body pool of AA.

Despite the minimal liver damage in the ethanol-fed pigs (26),

the lipid analysis showed a four-fold increase in liver TO and

alteration of tissue FA profiles in the ethanol-fed pigs. This in-

dicates an impairment of lipid metabolism by ethanol before

the overt histological manifestation offatty liver. This relatively

minor liver damage compared with that in other animal models

may be due to the low amount of dietary fat in our model (9%

of total energy). Lieber and DeCarli (23) have shown that the

severity of fatty liver was correlated with the amount of dietary

fat in a rat liquid-diet model. In other rat models, a high-fat diet

exacerbated ethanol-induced fatty change, and progression tofibrosis was observed only with the high-fat diet (24, 25). Another

possible explanation would be a species difference. O’Hea and

Leveille (3 1) have reported that fatty acid synthesis by the liver

was minor compared with the synthesis by adipose tissue in pigs.

Thus pigs may be more resistant to the development of alcoholic

fatty liver than are rats and humans, which have significant ca-

pacity of FA synthesis by the liver.

The FA profiles in various tissues should be interpreted in the

context of the systemic metabolism of EFAs. Because AA wasnot appreciably supplied from the diet in our study, the liver

would be the primary endogenous source of AA via synthesis

from LA. In the fasting state, the serum FA composition is likely

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470 NAKAMURA ET AL

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REDUCED ARACHIDONIC ACID BY ETHANOL 471

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472 NAKAMURA ET AL

TABLE 3Fatty acid profile in other tissues5

Muscle phospholipidsSubcutaneous adipose

triglycerides Omentum adi pose triglycerides

Control Alcohol Control Alcohol Control Alcohol

Fatty acid (% by wt)14:0 0.21 ± 0.03 0.17 ± 0.03 1.57 ± 0.1 1 1.22 ± 0.09t 1.51 ± 0.26 1.48 ± 0.1116:0 isomersf 8.80 ± 0.42 7.47 ± 0.55 0.09 ± 0.02 0.10 ± 0.02 0.01 ± 0.01 0.01 ± 0.0116:0 18.84 ± 1.12 16.55 ± 0.79 23.08 ± 1.18 20.37 ± 1.07 25.95 ± 1.41 26.36 ± 1.40

16:lw7 0.48 ± 0.05 0.54 ± 0.09 2.86 ± 0.14 2.75 ± 0.18 2.13 ± 0.31 2.26 ± 0.2018:0 isomerst 7.59 ± 0.67 8.01 ± 0.40 0.25 ± 0.03 0.39 ± 0.02� 0.15 ± 0.01 0.21 ± 0.Olfl18:0 7.30 ± 0.33 8.36 ± 0.33 8.43 ± 0.83 6.95 ± 0.49 16.36 ± 1.65 17.42 ± 1.08l8:lw9 5.71 ± 0.29 7.27 ± 0.60t 39.00 ± 2.12 42.55 ± 2.01 32.70 ± 4.16 32.47 ± 1.32l8:1w7 2.35 ± 0.22 2.36 ± 0.15 3.25 ± 0.20 3.26 ± 0.32 2.18 ± 0.40 2.29 ± 0.26l8:2w6 26.20 ± 096 2950 ± 2.21 16.40 ± 1.17 17.35 ± 1.1 1 15.45 ± 4.87 14.05 ± 2.79l8:3w6 0.14 ± 0.01 0.10 ± 0.Olt 0.03 ± 0.00 0.02 ± 0.00� 0.04 ± 0.03 0.04 ± 0.04l8:3w3 0.18 ± 0.02 0.26 ± 0.03 0.48 ± 0.05 0.53 ± 0.04 0.49 ± 0.1 1 0.51 ± 0.0920:0 0.12 ± 0.01 0.18 ± 0.04 0.19 ± 0.02 0.14 ± 0.01 0.29 ± 0.05 0.23 ± 0.0520:1w9 0.22 ± 0.02 0.30 ± 0.04 0.94 ± 0.1 1 0.98 ± 0.12 0.79 ± 0.12 0.75 ± 0.0820:2w9 0.00 ± 0.00 0.00 ± 0.00 0.07 ± 0.00 0.06 ± 0.0 1 0.03 ± 0.03 0.05 ± 0.0120:2w6 0.54 ± 0.09 0.64 ± 0.08 0.8 1 ± 0.09 0.85 ± 0.08 0.58 ± 0. 17 0.56 ± 0.0920:3w9 0.05 ± 0.00 0.08 ± 0.03 0.03 ± 0.01 0.04 ± 0.01 0.02 ± 0.02 0.02 ± 0.0220:3w6 0.64 ± 0.02 0.86 ± 0.10 0.13 ± 0.01 0.10 ± 0.01 0.10 ± 0.02 0.1 1 ± 0.0120:4w6 12.22 ± 0.42 9.36 ± 0.8It 0.24 ± 0.02 0.15 ± 0.0l� 0.29 ± 0.07 0.28 ± 0.0420:3w3 0.07 ± 0.01 0.10 ± 0.03 0.15 ± 0.01 0.15 ± 0.01 0.07 ± 0.01 0.09 ± 0.0120:5w3 0.21 ± 0.01 0.24 ± 0.04 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.0122:0 0. 18 ± 0.02 0.20 ± 0.03 0.00 ± 0.00 0.00 ± 0.00 0.02 ± 0.04 0.01 ± 0.0122:4w6 1.32 ± 0.16 1.25 ± 0.17 0.32 ± 0.10 0.28 ± 0.09 0.14 ± 0.04 0.12 ± 0.0222:5w6 0.78 ± 0.21 0.71 ± 0.20 0.00 ± 0.00 0.00 ± 0.00 0.00 ± 0.00 0.03 ± 0.0722:5w3 0.68 ± 0i5 0.58 ± 0.16 0.00 ± 0.00 0.00 ± 0.00 0.06 ± 0.04 0.07 ± 0.0122:6w3 0.93 ± 0.20 0.89 ± 0.29 0.00 ± 0.00 0.00 ± 0.00 0.01 ± 0.01 0.00 ± 0.00Others 4.22 ± 0.45 3.99 ± 0.46 1.69 ± 0.22 1.78 ± 0.14 0.65 ± 0.09 0.58 ± 0.06

S � � SE.t�II Significantly different from control value (t test): tP < 0.05, §P < 0.01, lIP < 0.00 1.

t May contain ether lipids.

to reflect PLs and TGs secreted from the liver. We observed auniform increase in oleic acid and LA and a decrease in AA in

the PLs from liver, serum, and muscle (the largest PL pool in

the body) of ethanol-fed pigs (Tables 1-3). This consistent de-crease in AA in all three tissues suggests that the output of AAfrom the liver was decreased in ethanol-fed pigs. In the TO frac-tions, the ethanol effect on the FA profile was similar in the liverand serum. In the subcutaneous adipose, the largest TO pool in

the body, the only difference in unsaturated FAs betweenethanol-fed and control groups was a significant reduction of18:3w3 and AA (Table 3), possibly because of the dependence

of adipose tissue on the liver for these FAs despite its ability tosynthesize saturated and monounsaturated FAs and to take upLA from the diet. Therefore, the consistent decrease of AA in

the TO fraction ofthese three tissues is a further indication thatthe output of AA from the liver was decreased in ethanol-fedpigs. The FA profile of the omentum adipose TO showed the

least difference between the two groups in this study (Table 3),suggesting that lipid metabolism in omentum adipose is different

from that in subcutaneous adipose tissue.

Our study showed that ethanol administration was associatedwith uniform reductions in AA (the major product of the de-

saturase-elongase pathway) with no decrease in LA (the sub-strate), with the only exception being the liver and serum TO

fractions. Also, �,3 products ofthe pathway (22:5w3 and 22:6w3)

were decreased with a concomitant increase in the substrate ( I 8:

3o�3) in the liver PL fraction of the ethanol-fed pigs. These

changes could be caused by reduced activities of �6 and i�5

desaturases in the ethanol group. Although we did not measuredesaturase enzyme activities in this study, studies with rats havereported a decrease in �6 and �5 desaturase activities with

ethanol ingestion (32-35).

Whereas the changes in FA profile in PLs and TOs could be

the result of decreased activity of z�6 and �.5 desaturases in

ethanol-fed pigs, another possibility is changes in hepatic acyl

CoA pool, the substrate pool for PL and TO synthesis. Becausethere is fairly rapid recycling of FAs in the PL pool via free FAsand FA CoA pools in the liver, free FAs may better represent

the true substrate (acyl CoA) pool than the FA profile of the

PLs and TOs (36). In the free FA profile in the liver from our

study, the relative amount of AA and some other desaturase-

elongase products were decreased in ethanol-fed pigs (Table 1),

but not the absolute amount (Fig 2). Their relative reductionswere due to large increases in other FAs (mainly oleic and linoleic

acids), which could be due to impaired oxidation of these FAs

by ethanol feeding (37). Also, the increased stearic and oleic

acids in liver and serum TOs in ethanol-fed pigs (Table 1 and

2) suggests an elevated de novo synthesis ofthese FAs by ethanol

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REDUCED ARACHIDONIC ACID BY ETHANOL 473

1. Holman RT. Essential fatty acid deficiency. Prog Chem Fats Lipids

l976;9:275-348.

1400

1200

1000

80� ..

600

:� �xi

FIG 2. Amount of free fatty acids in the liver (�zg fatty acids/g wetliver). Top: five major fatty acids. 0, control group; 0, ethanol-fed group(n = 5 for each group). I ± SE. Under “others” are grouped all fattyacids except for the five fatty acids shown individually. Among fatty

acids grouped in “others”, 16:lw7 and 18:1w7 were responsible for the

significant increase in ethanol-fed pigs. Bottom: free fatty acids in thedesaturase-elongase pathway in the liver. The horizontal axis representsthe sequential steps of �6 desaturase, elongase, and �5 desaturase. Thevertical axis shows the relative difference of fatty acid concentration in

ethanol-fed pigs compared with the control pigs. For both panels, the

asterisk shows statistical significance between control and ethanol-fed

groups by t test: �, P < 0.05; �, P < 0.01.

feeding. Thus, the change of FA profile in PLs and TOs may bedue to the relative increase in other FAs (stearic, oleic, and lin-

oleic acids) over desaturase products, because these FAs alsocompete with their products for the incorporation into PLs (38)

and TGs (39). However, this argument must be viewed with

caution because the free FA profile ofthe liver might have been

distorted by post mortem hydrolysis of PLs, because the liver

sample was not freeze-clamped. Also, the possibility that thechanges in FA profile in PLs may be due to the change in the

proportion of PL subspecies or subcellular organelles cannot be

ruled out, because we did not analyze PL subspecies or organelles

separately in this study. This, however, would not be the case

in the TO fraction because there are no subfractions in TO.

Increased peroxidation could be another possible mechanismfor the decreased AA in serum and tissues. Although the effect

of ethanol intake on lipid peroxidation has been controversial

(40-43), it has been reported that ethanol consumption decreased

the glutathione concentration (37), and increased both free rad-

ical formation (41, 44) and iron mobilization (42). All of these

changes could increase AA peroxidation. The status of the freeradical defense system and lipid peroxidation in our study wasreported elsewhere (26). Briefly, no clear reduction was observedin the free radical scavenging system, and thiobarbituric acid-

reactive substances (TBARS) were decreased in the liver ho-mogenate of the ethanol group. However, this decrease inTBARS was correlated with the peroxidizability index (P1) of

the liver PL (r = 0.82, P < 0.01). This fairly high correlation

could be due to either increased peroxidation by ethanol or de-

creased highly unsaturated FAs in the ethanol group caused by

other mechanisms than peroxidation. The result presented in

this paper favors the latter possibility because the peroxidativedegradation of highly susceptible FAs does not explain the se-

lective reduction in polyunsaturated FAs without depletion of

precursors in the ethanol-fed animals seen in Tables 1-3.

Foudin et al (16) also reported the effects of ethanol on the

FA profile of serum lipids in miniature pigs. Their observationsof increased oleic and stearic acids in serum TGs and increased

oleic acid in phosphatidyl choline are similar to our results, sug-

gesting increased endogenous FA synthesis from ethanol. How-

ever, their reports of decreased LA and increased AA in serum

phosphatidyl choline with ethanol feeding are opposite our find-

ings. Although direct comparison of the two studies is not pos-

sible because of the differences in experimental design (such as

duration and diet), note that the discrepancy between the two

studies is not due to an absolute difference in FA profile of

ethanol-fed animals. Rather, it appears to be due to differences

between the groups of control animals (their controls showed

much higher LA and 20:3 and much lower AA than ours). This

large difference between controls must be explained by factorsother than an ethanol effect. They did not state whether the

samples were taken in the fed or fasting state. If the animalswere not fasted before sample collection, changes in meal patternby ethanol feeding could also affect the serum fatty acid profiles.

The high LA content in both serum lOs and phosphatidyl cho-line in their control group may have been a reflection of the

composition of the soybean oil they used in their diet.

Although decreased tissue AA observed in our study couldhave pathophysiological effects by affecting the function of tissue

membranes and the production of eicosanoids in chronic al-coholism, we cannot extrapolate the results of our pig model

directly to humans. However, possible implications ofthis study

for human alcoholism are as follows. A nutritional need for

specific fatty acids (such as AA) may be induced by alcoholism

even before the progression to liver damage. On the other hand,

because a high AA concentration in platelets promotes platelet

aggregation in humans (45), the decrease in AA with ethanol

intake may be related to a longer bleeding time in alcoholics (7)

and may explain the lowered risk of coronary heart disease in

the population of moderate alcohol consumption (46).

In conclusion, this study demonstrated that AA was decreasedwith a concomitant increase of oleic acid in the tissues repre-

senting the major body pools in the ethanol-fed pigs. The possible

mechanism is a decrease in desaturase activity, and/or compet-

itive displacement ofAA from lipid fractions by other FAs, pri-marily oleic and linoleic acids. 13

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474 NAKAMURA ET AL

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