Crystallization of human liver alcohol dehydrogenase

Crystallization of human liver alcohol dehydrogenase

ARCHIVES OF BIOCHEiMIS’l~RT .1ND Crystallization of Human NABEEH Department 121, 431~439 (1967) BIOPHYSICS MOURAD? of Chemistry, Received L...

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ARCHIVES

OF

BIOCHEiMIS’l~RT

.1ND

Crystallization

of Human

NABEEH Department

121, 431~439 (1967)

BIOPHYSICS

MOURAD?

of Chemistry, Received

Liver

AXI) CHARLES

Brown, C:niuersity,

February

Alcohol

Dehydrogenase’ L. WORONICK”

Providence,

27, 1967; Accepted

Rhode Island

02912

34arch 23, 1967

Alcohol dehydrogenase has been cryst,allized from human liver. The purification procedure consisted of fractionation with ammonium sulfate, treatment with ethanolchloroform, chromatography on carboxymethylcellulose and on diethylaminoethylcellulose, and crystallization from aqueous ethanol. The crystalline protein was found to be homogeneous by ultracentrifugation and by disc electrophoresis. The enzyme is a basic protein. Amino acid analysis showed that, there is an excess of basic amino acids over acidic amino acids. Under the conditions of the assay, human liver alcohol dehydrogenase is about 71C: as active as horse liver alcohol dehydrogenase.

Part’ially purified alcohol dehydrogenase from human liver was first studied by von Wartburg et al. (1, 2) and by Blair and Vallee (3). A procedure for purifying the enzyme about ZOO-fold was reported by von Wartburg et al. (1). The present paper describes a procedure for isolating and crystallizing alcohol dehydrogenase from human liver. The crystalline protein was found to be homogeneous by ultracentrifugation and by polyacrylamide gel disc electrophoresis. MATERIALS

AND

METHODS

Normal-appearing human livers were obtained aft,er autopsy at 2-16 hours after death. The livers, weighing 900-2000 gm each, were used immediately or were frozen and used later. Crystalline ammonium sulfate (special enzyme grade) was obtained from Mann Research Laboratories. CM-cellulose of 0.74 meq/gm m-as purchased from the Sigma Chemical Company. DEAE-cellulose of 0.61 meq/gm was obtained from California Corporation for Biochemical Research. The ethyl alcohol used in the assay was 1 This work was supported in part by U.S. Public Health Service grant, GM 11463. 2 Present address: Brooklyn-Cumberland Medical Center, Pathology Department, Brooklyn, N.P. 11201. Hospital, 3 Present address : Pennsylvania Ayer Clinical Laboratory, Philadelphia, Pennsylvania 19107.

obtained by fractional distillation of USP alcohol. Enzyme assay. The enzyme was assayed spectrophotometrically at 23” by measuring the change in absorption at 340 rnr when DPN was reduced by ethyl alcohol at, pH 10. The enzyme was assayed according to the met,hod developed by Dalziel (4, 5) for horse liver alcohol dehydrogenase, except that the time required to produce an absorbance change of 0.100 was measured. The enzyme concentration was calculated in terms of enzyme unit,s rather than by using t,he equation developed by Dalziel (4, 5) for the horse liver enzyme. The enzyme unit was defined as the quantity of enzyme that will reduce 1 pmole of DPN per milliliter per minute under the conditions of the assay. The specific activity was defined as the enzyme lmits per milligram of protein. Protein assay. The prot,ein was estimated by measuring the absorbance at 280 rnp. The extinction coefficient was determined by measuring the absorbance of a solution of t,he cryst,alline enzyme and performing a dry-weight determination on the same solution. The absorbancy at 280 rnp of 1.0 mg of prot,ein per milliliter of 0.1~ sodium phosphate buffer (pH 7.0) was 0.61 f 0.01 for a l-cm light path. The absorbancy reported for horse liver ADH is 0.46 (8). This difference is caused by the greater number of tyrosine and tryptophan residues in human liver AUH (Table II). Sedimentation experiments. The sedimentat.ion properties were determined with a Spinco model E ultracentrifuge at 20” and 59,780 rpm. Disc electrophoresis. Polyacrylamide gel disc electrophoresis (6) was performed with a Canalco model 1400 electrophoresis apparatus in a 4” room. 431

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AN11 WORONICK

The amount of protein applied per column was about 25 fig and the current was 3.5 mA per column at a constant voltage of 400 V. The experiments were run at two different pH values and with the electrodes reversed. Electrophoresis on cellulose acetate strips. To each strip was applied 6 ~1 of a 1% solution of the was performed with enzyme. The electrophoresis a current of 2.5 mA at 150 V for 2 hours with cooling. The experiments were run at an ionic strength of 0.1 in the following buffers: Tris-acetate, pH 8.9; glycine-NaOH, pH 10.0; glycine-NaOH, pH 10.8; sodium phosphate, pH 11.0; and sodium phosphate pH 11.4. The cellulose acetate strips were cut in half lengthwise, and the protein on one half of the strip was located by staining wit,h ponceau S (Allied Chemical Corp.). The areas corresponding to the protein spots on the second half of the strip were cut out and the protein was eluted into a cuvette containing the reaction mixture and tested for enzyme activity. At pH 11 and below, the enzyme migrated toward the cathode, whereas at, pH 11.4 t,he enzyme migrated toward the anode. It was estimated that no migrat,ion would occllr at abollt pH 11.2. Since corrections were not made for electroendosmosis, the isoelectric point of the enzyme must be below pH 11.2. On the other hand, the behavior during disc electrophoresis indicates that. t,he isoelect,ric point is above pH 9.5. Protein hydrolysis. To 1.0 ml of solution containing 3.7 mg of crystalline human liver alcohol dehydrogenase was added 1.0 ml of concentrated HCl. The tltbe was evacuated and sealed and placed in an oven at 110” for 22 hours. The amino acid analysis was performed with a Beckman model 120 C amino acid analyzer with the Beckman model 125 digital integrator. The analysis was performed by Beckman Instruments, Palo Alto, California. EXPERIMENTAL

PROCEI>URE RESULTS

AND

Crystallization of alcohol clehydrogenase from hu?laan liver. All of the following steps were conducted at 4” unless ot’herwise indicated. Step 1: Extraction procedure. For a typical fractionation, two frozen livers were thawed by being allowed to stand in a 4’ room overnight, after which t,he livers were weighed and ground separately with a meat grinder. Then 1.5 liters of distilled water was added per kilogram of liver, and the mixtures were left. at room temperature for 2 hours with occasional stirring. The mixtures were then

filtered through cheesecloth, and a sample of each extract was assayed. Step W: Ammonium sulfate fractionation. If the assays indicated a satisfactory amount of enzyme in each liver, the filtrates were combined and t’ransferred to the 4’ room, after which 22.5 gm of solid ammonium sulfate was added per liter of liver extract. The mixture was adjusted to pH 7.0-7.5 by the careful addition of concentrated ammonium hydroxide and allowed to stand for 2 hours. Then t’he mixture was centrifuged at 7000 g for 50 minutes, and the precipitate was discarded. To each liter of supernatant solut’ion, 17.5 gm of solid ammonium sulfate was added wit’h stirring. The pH was adjusted to 7.5 with concentrated ammonium hydroxide, and the solution was allowed to stand at 4’ overnight. The solution was then centrifuged at 7000 CJfor 50 minutes. The precipitat’e, which contained 70-80 % of the enzyme, was suspended in a minimum volume of 0.05 P sodium phosphate buffer at pH 7.0, and was transferred into dialysis bags. Dialysis against 4 X 4 liters of 0.05 p sodium phosphate buffer (pH 7.0) was continued unt’il the concentrat’ion of ammonium sulfate decreased to about l-2 %. Step 3: Ethanol-chlorofor~la treatment. This step is essentially the same as the procedure described by Tsuchihashi (7) in which a 2: 1 mixture of et’hanol and chloroform is used. To each liter of dialyzed protein solution, 200 ml of et.hanol-chloroform mixture was slowly added while stirring vigorously. The mixture was t,hen stirred rapidly for 10 minutes with a magnetic stirrer. To each liter of solution, 200 ml of 0.05 p sodium phosphate buffer (pH 7.0) was added, after which the mixture was centrifuged at 7000 9 for about 30 minutes. About 80% of the enzyme was recovered in the supernatant solution. The alcohol and chloroform remaining in solution were evaporated on a flash evaporat#or with a bath t’emperature of 30’. In this entire st’ep both time and temperature are important. Aft,er most of t,he organic solvents had been evaporated, the aqueous enzyme solution remaining was further evaporated at a bath t’emperature of 40-45” until the concentrated solution became somewhat viscous. Foaming during the

vacuum evaporation was prevented by the addition of a few drops of octanol. The concent’rated enzyme solution was then dialyzed against 4 liters of 0.05 P sodium phosphate buffer at, pH 7.0 for S hours. This was followed by dialysis against 0.01 p sodium phosphate buffer at pH 7.0 for 8 hours. There was a IO-15 % loss of enzyme activity during the dialysis at 0.01 ionic strength. Step 4: Ca~boxymetkylcellulose column chromatography. The dialyzed enzyme solution was applied t,o a 5 X 50 cm CM-cellulose column which had been equilibrated with 0.01 p sodium phosphate buffer at pH 7.0. The flowrate of the column was about 100 ml/hour. The column was t’hen washed with 0.01 p sodium phosphate buffer, pH 7.0. A dark-brown band which moved down the column contained no enzymic activity and was discarded. A pink-colored band containing the enzyme was eluted from the column with 0.1 /L sodium phosphate buffer, pH 7.0. The enzyme, obtained in a volume of about 100-200 ml, was precipit’ated by the addition of 45 gm of solid ammonium sulfate per 100 ml of enzyme solution. The solution was adjusted to a pH of S.0 by the dropwise addit.ion of concentrated ammonium hydroxide solution, and was allowed to stand at 4” overnight. Step 5: nietlrylanrinoethylcellulose column. clr?.ol?zato!l~aphy. The ammonium sulfate precipitate was centrifuged at 12,000 9 for 40 minutes, redissolved in a minimum quantity of 0.05 p sodium phosphate buffer (pH S.O), and transferred to a dialysis bag. The solution was dialyzed against 4 liters of 0.05 p sodium phosphate buffer (pH S.0) for S hours, followed by dialysis for S hours against 4 liters of 0.01 p glycine-KaOH buffer at pH 9.5. There was no loss of enzyme activity if the dialysis against the glycine-KaOH buffer did not exceed 8 hours. A 3 X 30 cm DEAE-cellulose column, flow rate 300 ml/hour, was equilibrated with 0.01 p glycine-NaOH buffer, pH 9.5. When the enzvme solution (volume about 30 ml) was applied to the column, all the colored material was tightly adsorbed by the column. The enzyme was eluted as a colorless solution when the column was washed with 0.01 P glycinc-?;aOH buffer, pH 9.5. Chromatog-

raphy a second time on a CAI-cellulose column improved the specific activit(y by only a small amount. At this stage the enzyme is readlr for crystallization. The partially. purified enzyme was stored as a precipitate at 4’ in the following manner. The solution containing the enzyme was brought to SO’;6 saturation with respect to ammonium sulfate by the addition of solid ammonium sulfate, after which the mixture was adjusted to pH 8.0 with concentrated ammonium hydroxide. The enzyme could be stored in this way for several months without any loss of activity. Crystallization of the emyme. Enzyme which had been stored as an ammonium sulfate precipit,ate was centrifuged at 12,000 9 for 40 minutes. The precipitate was dissolved in a minimum volume of 0.1 p sodium phosphate buffer (pH S.O), and the resulting solution was allowed to stand for 3040 minutes. At this point unidentified crystals cont’aining a small amount of enzyme activity formed and were centrifuged and discarded. The supernatant solution was dialyzed against 4 liters of 0.05 p sodium phosphate buffer (pH S.0) for 12 hours and against 1 liter of 0.02 p sodium phosphate buffer (pH 10.6) for 6 hours. At this point the protein concentration was adjusted to l-2 gm/lOO ml. To the enzyme solution at 0” was added 90 % et,hyl alcohol at -7” until the final alcohol concentrat’ion was 15 SG. The alcohol was added dropwise with rapid stirring while cooling the enzyme solut,ion in a salt-ice bath. The mixt,ure was then kept at -7” for 24 hours, after which it was centrifuged at 12,000 g for 15 minutes at, -7”. The amorphous precipitate obtained at this point usually was discarded because it contained onlv a small amount of enzyme w&h a low specific act,ivit’y. To the supernatant solution 90 % alcohol nt -7” was added with continuous stirring unt8il the alcohol concentration reached 30 Y. The resulting solution, at a final t,emperature of about - 3”, cont’ained no precipiMe, and was transferred to a freezer at’ -1s” and allowed t#o stand for 24 hours. Crystals of pure alcohol dehydrogenase usually form at this st,age. The yield of crystals was 65-90 mg per liver. If the precipitate is found to be

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FIG. 1. Photomicrograph nification

of crystals

AND

WORONICK

of human

liver

alcohol

dehydrogenase.

The mag-

is 965 X.

amorphous, it can be centrifuged at -18” and redissolved at 0” in a minimum quantity of 0.1 p sodium phosphate buffer, pH 10.6. Crystal formation usually begins within a few minutes. It was found that human liver alcohol dehydrogenase has a strong tendency to co-crystallize with other proteins. The enzyme crystallized in the shape of flat triangles, the largest of which were 30 p along the base (Fig. 1). During photography at 4’, part of the crystals dissolved because of the rise in temperature. The

highest specific activity of the crystals obtained by this method was 3.3 enzyme units/ mg of protein. No further increase in specific activity was obtained when the crystals were redissolved and recrystallized by the low-temperature method. Results of the purification are summarized in Table I. Criteria of purity. Crystalline human liver alcohol dehydrogenase was subjected to polyacrylamide gel electrophoresis at two different pH values. The results of experiments at pH 6.6 and 9.5 are shown in Fig. 2.

IlURIAN

T,IVI:It

ALCOHOL TABLE

PT:HIFIVATION Stage

I. II. III. IT’. V. VI.

OF

of purification

Extract Ammonillm sldf:tte frwiiollatiorl Ethallol-chlorof!)rrn CM-cellrdose column DEAN-cellldose colrlmn Crystallization

HUMAN

435

I)~IIYI)ItO(;E~A~:E; I

LIVER ALCOHOL I~EHYDROGESASE Total

recovery (%I

100 70 55 33 30 10-15

FIG. 2. Photograph of polyacrylamide gels after electrophoresis and staining. The gel in tube 1 was at pH B.6, and the gel in tube 2 was at pH 9.5. In each case the electrophoresis was carried out for 105 minutes. The protein was applied to the upper end of each column as orient,ed in the photograph.

As can be seen, only one protein band was observed in tube number 1 at pH 6.6. In under some tubes run simultaneously identical conditions, 4 slower moving faint

Amount of enzyme per liver (units)

2200 1500 1200 710 650 220-300

Specific iunits/mg

activity protein)

0.0033 0.017 0.050 0.50 1.5 3.3

Purification (fold)

1 5 15 150 450 1000

bands, which may have been artifacts, also appeared. When the electrodes were reversed and the experiment was run at pH 9.5, no protein bands appeared on the gel. This indicates that there was no protein with an isoelectric point below pH 9.5 present in the crystalline enzyme preparation. Ultracentrifugation at 59,780 rpm gave only a single symmetrical peak (Fig. 3). Stability of the enzyme. The enzyme is usually stable between pH 7.0 and 10.6 at an ionic strength of 0.05 or greater. The enzyme is unstable at low ionic strength and loses activity rapidly below pH 6.0 or above 11.0. It appears to be most stable at about pH 8. Enzyme solutions in the crudest stages of preparation are stable to freezing, but NO-fold purified enzyme, for example, is destroyed by freezing. Preparations which have been purified about 500-fold can be stored for months at 4’ without loss of activity as an ammonium sulfate precipitate at pH 8. The enzyme crystals, when stored at pH 10.6 in 0.02 p sodium phsophate buffer containing 30% alcohol, lose considerable activity within one month at - 18”. Cleland’s reagent (dithiothreitol) at 10d4 M did not seem to protect the enzyme when tested at various st’ages during the purificat’ion. Attempts to crysballize t’he enzyme at 4” by dialysis of highly concentrated solutions of approximately 500-fold purified enzyme against 15% alcohol in 0.1 P sodium phosphate at pH 10.6 resulted in the conversion of the entire solution into a fairly rigid, insoluble gel. The same phenomenon was observed at pH 8. Since this was observed only in highly concentrated solutions, it was thought that a possible explanation might be that the protein was polymerized by the

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intermolecular oxidation of sulfhydryl groups to disulfide bonds. However, dithiothreitol at lop4 M did not prevent gel formation. Anzim acid analysis. Table II shows the amino acid composition of human liver ADH as well as the amino acid composition reported by Theorell et al. (S) for horse liver ADH. The values for human liver ADH were calculated for a molecular weight of S7,000, as tentatively reported by von Wartburg et al. (1). Comparison of the data shows that the amino acid compositions are quite different. In a few cases, the numbers of residues are very similar. In about half the cases, t,he differences are greater than 10%. Such differences are, of course, not unexpected for proteins isolated from different sources. If we assume that most of the ammonia obtained in t’he hydrolysis of human liver ADH is produced from asparagine or

WORONICK

AMINO

TABLE II ARID COMPOSITION OF THE ALCOHOL ~EHYL)KOGEK.\SES FROM HUIM.IN .\ND 1x0~s~ LIVERS Amino acid

Moles/mole of human liver

Valine Glycine Alanine Leucine Threonine Serine Proline Isoleucine Phenylalanine Half-cysteine Met,hionine Tyrosine Aspartic acid Glutamic acid Lysine Arginine Histidine Tryptophan Ammonia

86.1 74.2 63.3 52.1c 45.8” 45.6 45.2 36.2 31.2c 16.7 11.4c 63.0 59.0 65.6 20.6 13.4 Gd 53.6

ADHa

89.4

Moles/m~~le of horse lixer

ADHh 82 83 61 54 50 53 46 49 38 -e 19 8 54 66 62 25 14 4 -f

Q Tentative values calculated for a molecular weight of 87,000 (1). b The data for horse liver ADH was taken from Theorell el al. (8). c Corrected as described by Moore and Stein (13). d Estimated from the absorption at 280 rnp and the tyrosine content, (14, 15). e Not reported. Witter (9) and Theorell (see Ref. 9) have reported 28 sulfhydryl groups, pres[rmably from cysteine residues, in horse liver ADH. 1 Not reported.

FIG. 3. Photograph of a schlieren pattern obtained during a sedimentation experiment. The photograph was taken after 2880 seconds at 59,780 rpm at 20”. A protein concentration of 1.15% in 0.28 p sodium phosphate brtffer (pH 8.0) was used. The direction of sedimentation is from right to left.

glutamine residues, or both, this would indicate that the enzyme has an appreciable excess of basic amino acid residues over acidic amino acid residues. This will explain the high isoelectric point indicated above. In comparison, the isoelectric point of horse liver ADH is pH 6.8 (5), whereas the isoelectric point of a steroid-active alcohol dehgdrogenase crystallized by Theorell et al. (8) from horse liver is about pH 10. The number of sulfhydryl groups in human liver ADH may perhaps be similar to the number in horse liver ADH. The number

of

half-cysteine

residues

in human

FIG. 4. The absorption spectrllm of human liver alcohol dehydrogenase at, 20” in 0.1 G Tris-acetate buffer, pH 9.0. The absorbancy ratio at 280/260 rnp is 1.28 at this pH. The spectrum was made with a Cary model 14 recording spectrophotometer and a l-cm light path.

liver ADH is 31, whereas the horse liver enzyme contains 28 sulfhydryl groups (9). The absorption spectrum is shown in Fig. 4. Relative activity of the enzyme. In order to obt,ain a comparison of the relabive activities

of human liver ADH and horse liver ADH under the same conditions, the purest preparation of human liver ADH was assayed according to the method of Dalziel (4, 5), in which the time for an absorbance change of 0.200 at 340 ml* is measured. Since the

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reactions for both human liver ADH and horse liver ADH are not zero order during the entire reaction periods, the specific activity of human liver ADH was calculated to be 3.5 units/mg of protein under these conditions. In contrast, Dhe specific activity of the purest preparation of crystalline horse liver ADH prepared by Dalziel (5) can be calculated to be 5.1 units/mg of protein. In more specific terms, the apparent catalytic center activity of human liver ADH under the assay conditions was 152 min-L per active site, whereas the value for horse liver ADH was 214 mine1 per active site. Thus, under conditions of the assay, human liver ADH is 68 % as active as horse liver ADH when compared on an equal weight basis, or 71% as active when compared on an equal mole basis. The latter figure assumes that the molecular weight of human liver ADH is 87,000 as tentatively reported by von Wartburg et al. (1). The difference in activity must be due to differences in the values of the kinetic parameters. von Wartburg et al. (1) have claimed that the most striking difference between human liver ADH and horse liver ADH is in their substrate specificities. This conclusion was based partly on the report of Winer (10) that methanol and 2-propanol were not oxidized at a detectable rate when tested at an alcohol concentration of 0.001 M. The literature contains contradictory reports concerning methanol as a substrate for horse liver ADH. Kini and Cooper (11) reported in 1961 that methanol at a concentration of about 0.4 M has appreciable activity with crystalline horse liver ADH, as well as with partially purified rhesus monkey liver ADH. However, Wratben and Cleland (22) have recently report’ed that methanol at a concentration of 0.1 M is not a substrate for horse liver ADH. As for 2-propanol, Witter (9) in 1960 reported an apparent Michaelis constant of crystalline horse liver ADH for this alcohol. More recent,ly, Dalziel and Dickinson (12) have reported extensive kinetic studies in which 2-propanol was used as a substrate for crystalline horse liver ADH. Since this alcohol is a poor subst’rate for horse liver ADH, a possible reason for this discrepancy is that in Winer’s (10)

experiments t’he alcohol was apparent,ly test’ed at a concentration that was too low with respect to t’he value of its Michaelis con&ant. Blair and Vallee (3), on the other hand, have reported t,hat certain halogenated alcohols are substrates for human liver ADH but are not oxidized by horse liver ADH when tested under similar conditions. Since alcohol dehydrogenases from different. sources can have quite different Michaelis constants for a particular alcohol, these results should be interpreted cautiously unless it can be shown that the alcohols were tested over a sufficiently wide range of substrate concentrations. DISCUSSION

As can be calculated from Table I, the aqueous extract from a typical human liver weighing about 1400 gm contains approximately 670 mg of alcohol dehydrogenase, or about 480 mg/kg. In horse liver, Bonnichsen (16) estimated the concentration of ADH to be about 1000 mg/kg. It would be interesting to calculate whether the amount of ADH in a human liver is sufficient to account for the observed rate of ethanol oxidation in the human body. In a 70-kg man, alcohol is removed from the blood stream at a rate of 7 gm/hour, which is equivalent to 0.15 mole/hour (17). The total amount of ADH that can be extracted from a 1400-gm human liver can catalyze the oxidation of 2200 pmoles of ethanol per minute, or 0.13 mole/hour at pH 10 and 23’. According to the data of von Wartburg et al. (I), the rate at pH 7.4 and 23’ is about 45% as great’ as the rate at pH 10. This would reduce the rate of alcohol oxidation to about 0.05s mole/ hour. On the other hand, the rate at 37” is undoubtedly higher. From the data of Dalziel (18) at pH 7.1, it can be estimated that for horse liver ADH, the rate constant for alcohol oxidation at 37” is approximately 1.4 times as great as the rate constant at 23”. If we assume that a similar change in rate occurs with human liver ADH, we can estimate that at 37” a typical human liver would be capable of oxidizing approximately 0.082 mole of ethanol per hour. This rate is about 55 % as great as the rate

at’ which alcohol is removed from the blood stream. There are several possible reasons for this discrepancy, one of which is that the temperature coefficient for the human liver enzyme may be greater than the temperat,ure coefficient, for the horse liver enzyme. A very important factor to consider is that in making the aqueous extract of the liver, it is highly probable that not all the liver cells were broken during grinding, and not all the enzyme was extracted. Also some alcohol may be oxidized by the alcohol dehydrogenase found in human kidney (2), and erythrocytes (19), and a small amount is not oxidized at all but is excreted. Another source of uncertainty is that the effective concentrations of coenzyme, substrate, and hydrogen ion in t’he ccl1 are unknown. Finally, it is entirely possible that the human body has other mechanisms for oxidizing alcohol. Other syst’ems have been found by Orme-Johnson and Ziegler (20), and by Tephley et al. (21) in other animals. We can conclude that although liver ADH can probably account for the rate of alcohol oxidation in man, we cannot exclude the possibility that other systems may contribute significantly to the overall rate t’hat is observed. ACKNOWLEDGMEETS We are deeply indebted to Dr. Jacob Dyckman of Miriam Hospital, Providence, for his cooperation and continued interest. The sedimentation experiments were performed by Mr. Malcolm Smart at Harvard TJniversity through the courtesy of Professor Paul Doty. We wish to thank Professor Joseph Hteim for helpful discussions.

1. VON W~KTBUI~G, J. P., BETHUNE, J. L., AXD VALLEE, B. L., Hiochemistry 3, 1775 (1964). 2. VON WAIV~BURG, J. P., PAPENBERG, J., .4xr1 AEBI, H., Can. J. Biochem. 43, 889 (1965). 3. BLAIR, A. H., AXI) YALLEE, B. L., Biochemistr:tg 6, 2026 (19GG). i. DALZIEL, K., dcla Chenl. Stand. 11, 397 (1957). 5. DALZIEL, K., Beta Chem. &and. 12, 459 (1958). of Disc Electro6. ORNSTEIN, L., “Theory phoresis.” Canalco, Rockville, Md. (1962). 7. TSUCHIHAHHI, M., Biochem. %. 140, 62 (1923). 8. THEORELL, H., TASIGUCIII, S., AKESON, a., AND SKUILSKY, L., Biochenr. Biophys. Res. Commun. 24, GO3 (1966). 9. WITTER, A., Ada Chem. Stand. 14, 1717 (1960). 10. WINER, A. L)., ilcla Chem. Stand. 12, 1095 (1958). 11. KINI, M. M., AND COOPER, J. It., Hiochenr. Pharmacol. 8, 207 (1961). 12. DALZIEL, K., A?;D UICKINSON,F. M., Biochem. J. 100, 34 (196G). 13. MOORE, S., AND STEIN, W. H., Methods Enzymol. 6,819 (1963). 14. BEAVEN, C:. H., AND HOLIDAY., E. I<., 3rlvan. Protein Chem. 7, 113 (1952). 15. FKOMAGEOT, C., AND SCHNEK, (;., Biochim. Biophys. 4cta 6, 113 (1961). 16. BONNICHSER, 13. K., dcta Chem. &and. 4, 715 (1950). 17. THEORELL: H., AND BONXIPIISEN, It. Ii., ;lcta Chem. ScancZ. 5, 1105 (1951). 18. I)ALZIEL: K., 11&a Chem. &and. 17, S27 (1963). 19. THEORELI,, H., personal communication. 20. ORLIE-JOHSSON,W. H., AND ZIEGLER, 11. ill., Biochem. Biophys. Res. Commvn. 21, 78 (1985). 21. TEPHLEY, T. I:., PARKS, I<. E., JR., AND MANNERING, G. J., J. Pharmucol. Expll. Therap. 143, 292 (1964). 22. WHATTEX, C. C., AND CLELAND, W. W., Biochemisfry 4, 2442 (1965).