ARCHIVES

OF

BIOCHEMISTRY

Localization

AND

BIOPHYSICS

638-649

(19763

and Characteristics of Rat Liver Mitochondrial Dehyd rogenasesl CHAKWAN

Department

176,

of Pharmacology,

SEW,’

University

RICHARD

Aldehyde

A. DEITRICH3

of Colorado School of Medicine, Denver, Colorado 80220

4200

East Ninth

Avenue,

AND

V. GENE School

of Pharmacy,

University Received

ERWIN

of Colorado, March

Boulder,

Colorado

80301

12, 1976

Two NAD-dependent aldehyde dehydrogenase enzymes from rat liver mitochondria have been partially purified and characterized. One enzyme (enzyme I) has molecular weight of 320,000 and has a broad substrate specificity which includes formaldehyde; NADP is not a cofactor for this enzyme. This enzyme hasK,n values for most aldehydes in the micromolar range. The isoelectric point was found to be 6.06. A second enzyme (enzyme II) has a molecular weight of 67,000, a K,,, value for most aldehydes in the millimolar range but no activity toward formaldehyde. NADP does serve as a coenzyme, however. The isoelectric point is 6.64 for this enzyme. By utilization of the different substrate properties of these two enzymes it was possible to demonstrate a timedependent release from digitonin-treated liver mitochondria. The high K,,!, low molecular weight enzyme (enzyme II) is apparently in the intermembrane space while the low K,, high molecular weight enzyme (enzyme I) is in the mitochondrial matrix and is most likely responsible for oxidation of acetaldehyde formed from ethanol.

The role of various liver aldehyde dehydrogenase enzymes (EC 1.2.1.3) in aldehyde metabolism, especially as related to ethanol oxidation, has been a subject of increasing interest. It is generally agreed that aldehyde dehydrogenase enzymes exist in the supernatant, mitochondrial, and microsomal fractions as isolated from rat liver (l-4). The distribution of activity between these cellular components is the subject of some

debate; however, by and large the activity in the supernatant is 520% of the total, that in the microsomes between 10 and 30%, and the remainder in the mitochondria. The variation in the figures may be related to the assay, the strain of rat, the substrate, or the cofactor employed. Other evidence from liver perfusion studies (5-7) indicates that the mitochondrial fraction is primarily responsible for the oxidation of acetaldehyde produced during ethanol metabolism. This would agree with our finding (1) and that of others (3, 4) that rat liver mitochondria contain a low K,, enzyme . During ethanol oxidation large amounts of NADH are generated in the cytoplasm by the action of alcohol dehydrogenase. If the second step of ethanol metabolism, NAD-linked acetaldehyde oxidation, also

’ Supported by Grant No. AA00263. A preliminary communication has appeared: (1974) Fed. Proc. 33, 538. ’ Predoctoral fellow, Training Grant No. GM 1982. Present address: Department of Pharmacology, Stanford Medical School, Stanford, Calif. s Recipient of Career Development Award No. GM 10,475; author to whom inquiries should be addressed. 638 Copyright All rights

0 1976 by Academic Press, Inc. of reproduction in any form reserved.

MITOCHONDRIAL

ALDEHYDE

occurs in the cytoplasm, two molecules of NADH will be produced for each molecule of ethanol oxidized in that compartment. These considerations have led to the postulated necessity of shuttle mechanisms to transfer reducing equivalents from NADH to the electron transport system of the mitochondria (8). If, however, the step of aldehyde oxidation occurs inside the intramitochondrial membrane, then the shuttle systems might be less important. Other aspects of aldehyde metabolism also require a knowledge of the localization of aldehyde dehydrogenase enzymes. Thus, the presence of monoamine oxidase on the outer mitochondrial membrane catalyzes the production of aldehydes at this location (9). The subsequent oxidation of these aldehydes would be facilitated by the close proximity of an aldehyde dehydrogenase. This study was carried out to localize and characterize the mitochondrial aldehyde dehydrogenase enzymes. EXPERIMENTAL

PROCEDURE

Materials Water was deionized and double distilled in a quartz still. All reagents were of the highest quality obtainable from commercial sources. Aldehydes were distilled under nitrogen at atmospheric pressure. The concentrations of freshly prepared aldehyde solutions were determined enzymatically immediately before use (10). Sephadex G-200 (Pharmacia Fine Chemicals, Inc.) columns were prepared by suspending dry Sephadex G-200 in excess of 1 or 50 mM sodium phosphate, pH 7.4, and allowing it to swell for several days prior to packing by gravity in a refrigerated column (2.5 x 25 cm). The column was calibrated with the following proteins: bovine liver catalase CM,, 240,000), yeast alcohol dehydrogenase CM,, 150,000), soya bean trypsin inhibitor (M,, 21,500), and bovine serum albumin CM,, 70,000). The void volume of the column was determined by using blue dextran. The elution volumes (V,.) of blue dextran and bovine serum albumin were determined spectrophotometrically. The presence of catalase, alcohol dehydrogenase, and soya bean trypsin inhibitor was determined enzymatically. DEAE-cellulose (diethylaminoethyl cellulose,4 Bio-Rad Laboratories), and hydroxylapatite (Hypatite C, modified calcium phosphate, Clarkson Chem4 Abbreviations used: ethyl; D’IT, dithiothreitol; hyde dehydrogenase.

DEAE-, (in figures)

diethylaminoALDH, alde-

639

DEHYDROGENASES

ical Company, Inc.) were prepared for use according to manufacturers’ recommendations. Protein determinations were done by the biuret method (11) using bovine serum albumin as a standard or by the absorption of the protein solutions at 280 nm. Specially purified enzyme grade ammonium sulfate was obtained from Schwa&Mann Research Company. All percentage of saturation figures are those at 4°C. METHODS Subcellular localization of NAD-dependent aldehyde dehydrogenase activity in rat liver. The subcellular distribution of rat liver NAD-dependent aldehyde dehydrogenase activity was determined by using livers obtained from adult Sprague-Dawley or Long-Evans rats of both sexes. All steps were carried out at 0-4°C. Animals were deprived of food for 24 h prior to the beginning of the experiment. The livers were removed immediately following stunning and decapitation of the animals. The livers were then placed in ice-cold 0.25 M sucrose, cut into small pieces, rinsed several times to remove most of the blood, and homogenized in sufficient fluid to make a 20% homogenate. Homogenization was performed in a glass homogenizer with a tight-fitting Teflon pestle at 800 rpm for 10 complete passes of the pestle. Unless stated otherwise, centrifugation was carried out in a Servall RC-2 centrifuge at 0°C and g forces are for the middle of the tube. The homogenate was initially centrifuged at 480g for 10 min and the pellet resuspended and recentrifuged twice as before. The final pellet obtained was designated as the nuclear fraction. The combined supernatant fluid was centrifuged for 10 min at 4200g. The resulting supernatant fluid (designated as the postmitochondrial fluid) was carefully decanted, and the mitochondrial pellet was gently resuspended in a volume of medium equivalent to the wet weight of the liver and recentrifuged at 42OOg for 10 min. This washing was combined with the postmitochondrial supernatant fluid while the combined supernatant fluid was recentrifuged at 19,OOOg for 10 min. The pellet obtained was designated the lysosomal fraction. The postlysosomal supematant was combined with the washings and centrifuged at 76,000g. The pellet was designated the microsomal fraction and the supernatant designated the final supernatant. All pellets were then resuspended in 1 mM sodium phosphate buffer, pH 7.4, containing 1 mM dithiothreitol. All resuspended pellets were subject to sonic disruption in an ice-cold ultrasonic bath. A Branson Model 125 Sonifler with a a/s-in. step horn at a setting of 3 which gives 3.5 A was used. The sonic pressure was 3g as determined by the recommendation of the manufacturer. The time of sonication was 3 min in 30-s intervals interrupted by several minutes to allow cooling of the material. All

640

SIEW.

DEITRICH

live fractions (nuclear, mitochondrial, lysosomal, microsomal, and final supernatant fluid) were centrifuged for 1 h at 76,000g before being assayed for NAD-dependent aldehyde dehydrogenase, glutamate dehydrogenase (121, P-glucuronidase (131, aminopyrine demethylase (14), and alcohol dehydrogenase (15) activity. Aldehyde dehydrogenase activity was measured in glass cuvettes containing a final concentration of: 15 mM sodium pyrophosphate buffer, pH 9.6; 1 mM NAD; 3.3 rnM propionaldehyde; 0.1 ml of enzyme solution; and water to give a final volume of 3.0 ml. Formation of NADH was determined spectrophotometrically at 340 nm. Purification of rat liver mitochondrial NAD-dependent alde$yde dehydrogenase. Two NAD-dependent aldehyde dehydrogenases were partially purified from rat liver mitochondria. Rat liver obtained as previously described was homogenized in isolation medium in a glass homogenizer, and the mitochondrial fraction was obtained by differential centrifugation. The mitochondrial fraction was either washed three times with medium or after one wash the fraction was resuspended in a volume of isolation medium equivalent to the wet weight of the liver and layered over two volumes of 1.2 M sucrose and centrifuged at 10,OOOg for 45 min. The resulting mitochondrial pellet obtained from either method was suspended in the original volume of 1 mM sodium phosphate, pH 7.4, containing 1 mM dithiothreitol (DlT) and sonicated at 3.5 A for six 0.5-min intervals (a total of 3 min) in an ice-cold bath with a Branson Soniiier and then centrifuged at 144,OOOg for 1 h. The resulting clear supernatant fluid was decanted and fractionated either with solid ammonium sulfate or saturated ammonium sulfate solution at 4°C (appropriate corrections for volume changes were made throughout the fractionation procedures). The precipitated materials obtained between 0.30-0.55 saturation containing enzyme I and 0.55-0.85 saturation containing enzyme II were sedimented by centrifugation at 20,OOQg for 20 min. The precipitates were dissolved in a mixture of 1 mM sodium phosphate, pH 7.4, and 1 mM DTT and passed separately through a Sephadex G-200 column, equilibrated with the same buffer. Enzyme II, which precipitates between 0.55 and 0.85 saturated ammonium sulfate, was then placed on a DEAEcellulose column. The enzyme did not bind and was eluted with the starting buffer. The enzyme which precipitated between 0.30 and 0.55 saturated ammonium sulfate (enzyme I) was treated similarly and was bound to DEAE-cellulose and was eluted with 50 mM sodium phosphate, pH 7.4. Similar studies were carried out with sonicated mitochondrial extracts. Enzyme fractions separated in this way were applied separately to hydroxylapatite columns and eluted with phosphate buffer. Both enzymes were then further purified after the DEAE-cellulose step

AND

ERWIN

by ampholyte electrofocusing according to the method recommended by the manufacturer (LKB Produkter AB, Stockholm, Sweden). The molecular weights of the two rat liver mitochondrial aldehyde dehydrogenases were determined. The Stokes radii were determined by the method of Ackers (16) using Sephadex G-200 gel filtrations in 50 mM sodium phosphate at pH 7.4, with xanthine oxidase, catalase, bovine serum albumin, and soya bean trypsin inhibitor as standard proteins. A linear sucrose density gradient ultracentrifugation technique according to Martin and Ames (17) was used to determine the sedimentation constants using catalase and horse liver alcohol dehydrogenase as standard proteins. Submitochondrial localization of aldehyde dehydrogenases. Submitochondrial localization of NADdependent aldehyde dehydrogenases was determined by a modified method of Schnaitman et al. (18). A 1% stock solution of digitonin (Sigma) was prepared by adding hot (=8X!) 0.25 M sucrose to powdered digitonin, mixing briefly, and sonicating for 2 min. The resulting solution was cooled and remained clear for at least 60 min at 4°C. All digitonin solutions were prepared immediately prior to use. Mitochondrial suspensions containing 100 mg of protein/ml were placed in an ice bath, and an equal volume of cold digitonin solution was added with continuous stirring. The digitonin: protein ratio was 1.0 mg of digitonin per 10.0 mg of protein. The suspensions were incubated at 4”C, and at various time intervals aliquots of digitonin-treated mitochondria were removed and diluted immediately by the addition of three volumes of cold medium. The diluted suspensions were centrifuged at 95OOg for 10 min. The supernatant fluid was carefully decanted from the pellet. In some experiments, the pellet was resuspended in 1 mM sodium phosphate at pH 7.4 and was sonicated for 0.5 min. The sonicated suspension was centrifuged at 144,OOQg for 1 h, and the resulting supernatant fluid was designated as the matrix fraction. The high speed pellet was resuspended and homogenized in 1 mM sodium phosphate, pH 7.4, and designated as the inner membrane fraction. All fractions, outer membrane plus soluble intermembrane space material, matrix, and inner membrane, were sonicated for an additional 2 min prior to enzyme assays. In other experiments only the centrifugation at 9500g was carried out and the resultant supernatant fluid assayed for released enzyme. Monoamine oxidase (MAO) was selected as a mitochondrial outer membrane marker enzyme and was assayed by the method described by Deitrich and Erwin (19). Malate and glutamate dehydrogenase (MDH and GDH) were selected as the marker enzymes for the matrix fraction and were assayed by the methods of Ochoa (20) and Strecker (12). The inner membrane marker enzyme was succinic dehydrogenase, and this enzyme activity was

MITOCHONDRIAL

ALDEHYDE

Following sonication of the mitochondria, the high speed supernatant fraction was subjected to the isolation procedure as outlined in the methods section. Results of the separation are presented in Table I. Two fractions, enzyme I and enzyme II, were obtained by ammonium sulfate fractionation. These fractions were applied to a Sephadex G-200 column, separately. The elution patterns are shown in Fig. 1. Nearly superimposable elution profiles were obtained when mitochondrial supernatant was applied directly to such columns. Figure 2 illustrates the elution pattern obtained when the enzymes were applied separately to DEAE-cellulose anion-exchange columns. Similar experiments with mitochondrial supernatant containing a mixture of the enzymes yielded essentially the same picture. These data taken together allowed identification of the first peak as enzyme II, as designated by ammonium sulfate fractionation, and the activity eluting with 50 mM phosphate as enzyme I. Enzymes from this step were applied to hydroxylapatite columns as shown in Fig. 3. It is apparent that the two enzymes have different binding affinities for this material. Electrofocusing of the two enzymes from the DEAE-cellulose step was carried out as shown in Figs. 4 and 5 with pH 5-7 ampholyte. Enzymes at this stage were very unstable even in the presence of 1 mM dithiothreitol. For this reason the follow-

assayed by the method of Arrigoni and Singer (21). Adenylate kinase was used as a marker enzyme for the soluble intermembrane space fraction, assayed by the method of Schnaitman and Greenawalt (22). RESULTS

SEPARATION AND CHARACTERIZATION MITOCHONDRIAL ENZYMES

OF

Since the major aim in this study was characterization of mitochondrial enzymes, the isolation methods were designed to yield relatively pure mitochondria. This was reasonably successful as evidenced by recovery of only 2% of the total aminopyrine demethylase as a microsomal marker and no detectable alcohol dehydrogenase as a marker for the cytosolic fraction in the mitochondrial fraction. Isolated mitochondria contained 47% of the recovered aldehyde dehydrogenase activity, the microsomal fraction contained 13%, and the supernatant fluid contained 24%, the remainder being found in the nuclear and lysosomal fractions which contained 8% each. Purification

of mitochondrial

enzymes.

Here two approaches were taken. A puriflcation scheme was developed to yield partially purified enzymes. In addition, steps of the procedure were applied directly to mitochondrial supernatant in order to demonstrate clearly the presence of two enzymes by a variety of techniques and decrease the chances that any given procedure was bringing about alterations in the characteristics of the enzymes. TABLE PURIFICATION

Mitochondrial

OF NAD-DEPENDENT Fraction extracts

ALDEHYDE

I

DEHYDROGENASES Total

(sonicated)

641

DEHYDROGENASES

FROM

protein (ma)

RAT LIVER Total

activity”

MITOCHONDRIA Specititcvactivi-

808.2

35.6

44.1

(NH&SO, (0.30-0.55) saturation, enzyme I Sephadex G-200 gel filtration DEAE-cellulose ion-exchange chromatography Ampholyte electrofocusing

318.8 245.1 114.0 22.7

18.9 17.1 15.1 7.1

47.4 69.6 132.2 231.7

(NH&SO, (0.55-0.85) saturation, enzyme II Sephadex G-200 gel filtration DEAE-cellulose ion-exchange chromatography Amphol.yte electrofocusing

156.3 94.6 29.6 8.8

13.0 8.8 8.2 6.8

83.0 92.6 276.8 798.3

” Total activity is expressed as micromoles h Specific activity is expressed as nanomoles

of NADH formed per fraction per minute. of NADH formed per milligram of protein

per minute.

SIE W, DEITRICH

642

AND ERWIN

Enzyme

FRACTION

II:

NUMBER

FIG. 1. Elution of aldehyde dehydrogenase enzymes from Sephadex G-200 columns. Enzymes from the (NH&SO, fractionation step were placed on the column in separate experiments. The column was 35 x 2.5 cm, equilibrated with sodium phosphate buffer, 1 mM, pH 7.4. Fractions 38

_

30

-

34

-

32

-

30

-

25

-

0’

24

-

._ ;;

24

-

ID lL

were 3.6 ml each.

-j

1400

-I

1300

z ii I

22

-

;= P

20

-

Ex

lb 16

_-

-

so0

z, ;i

c ._ P)

14

-

-

700

;

;

0’

12

-

-

600

10

-

-

500

I

-

-

400

-

300

6

-

4

-

2

-

1s

200

,,’ I?

19

21

23

25

27

29

31

33

35

37

39

FRACTION

41

51

53

55

57

59

61

63

65

67

69

r< 0 4 z : ” : _

lo: 71

73

NUMBER

FIG. 2. Elution of aldehyde dehydrogenase enzymes from DEAE-cellulose columns. After dialysis, enzymes from (NH&SO, fractionation step were placed on 33 x 2.5cm DEAEcellulose columns. Enzyme II eluted with the starting buffer, 1 mM sodium phosphate, pH 7.4; enzyme I eluted with 50 mM of the same buffer. A similar profile was obtained when sonicated rat liver mitochondrial supernatant was applied directly to such columns.

=I

MITOCHONDRIAL

ALDEHYDE

FRACTION

643

DEHYDROGENASES

NUMBER

FIG. 3. Elution of aldehyde dehydrogenase enzymes from hydroxylapatite zymes obtained from DEAE-cellulose chromatography were placed separately hydroxylapatite columns. Enzyme I was eluted with 0.2 M sodium phosphate while enzyme II was eluted with 0.4 M of the same buffer.

ing studies were conducted with enzymes from DEAE-cellulose columns. Properties of the enzymes. The cofactor and aldehyde specificities for the two enzymes are given in Table II. The finding that only enzyme II will utilize NADP as cofactor but fails to oxidize formaldehyde is the most unique difference between the enzymes. The much higher K, values for most aldehydes is also a characteristic of enzyme II. These values for V can be roughly correlated with Taft’s u* value (23). Such a plot gives a p value of +0.47 with a correlation coefficient of 0.92 for enzyme I and a p value of +0.47 with a correlation coefficient of 0.82 for enzyme II. Inhibitor sensitivity is presented in Table III. In general, enzyme II is somewhat less sensitive to the usual aldehyde dehydrogenase inhibitors than is enzyme I but not markedly so. Molecular weights of the two enzymes were estimated as outlined in the methods

columns. Enon 30 x 2.5-cm buffer, pH 7.4,

section and found to be 320,000 for enzyme I and 67,000 for enzyme II. Heat denaturation curves for enzymes I and II carried out at 50°C in 50 mM sodium phosphate buffer, pH 7.4, at a protein concentration of 2 mg ml-‘, reveal a significant difference in heat stability between the two enzymes (Fig. 6). Submitochondrial localization zymes . Initial studies with

of the en-

digitonintreated mitochondria had indicated that there was a biphasic release of aldehyde dehydrogenase activity (24). The finding of two enzymes with markedly different substrate characteristics provided an opportunity to identify more clearly the submitochondrial compartment for these enzymes. In Fig. 7 is illustrated the early phase of release of enzyme I (low K, and formaldehyde activity) and enzyme II (high K,,

NADP)

from mitochondria

mg of digitonin

treated

per 10 mg of protein.

with

1

It is

clear that enzyme II is released at a much faster rate than enzyme I which is released

644

SEW,

DEITRICH

AND

( Enzyme1

ERWIN

) 900

1.1 I.1

*

1.8

F -

1.5

-

1.k

-

1.3

-

1.2

-

1.1

-

1.0

-

0.9

-

0.8

-

0.1

-

0.8

-

0.5

-

i

800

0 2

8.9

0I

n ! 0 t a

‘iD

0.3 0.2 0.1

4

0

O t

5

L 10

15

20

25

36

35

,O

VOLUME

,5

50

55

( ml

60

70

75

80

85

90

1

( Enzyme

1.8

65

II )

1.7 F

1.8 t

7.0

#

6.64 i

pI 6.5

-

3000

;

-

2800

2

-

2800

z

-

2400

f

-

2200 2000

:

-

1800

-

1600

c,; If3

;D

D D

1

I a

6.0

5.5

VOLUME

( ml

1

FIGS. 4 and 5. Isoelectric focusing of aldehyde dehydrogenases. Figure 4 is the pattern obtained with enzyme I which has been purified through the DEAE-cellulose step. Figure 5 is the pattern for enzyme II. Narrow range (pH 5-7) ampholyte was used for both.

&

=

MITOCHONDRIAL

ALDEHYDE TABLE

MICHAELIS

CONSTANTS

Substrate

AND

__-K,,,

MAXIMAL

(X

l@

Formaldehyde Acetaldehyde Propionaldehyde Butyraldehyde Isobutyraldehyde Valeraldehyde Heptylaldehyde Glycoaldehyde Monochloroacetaldehyde Glyoxal Crotonaldehyde Pyruvic aldehyde p-Carboxybenzaldehyde 4-Cyanobenzaldehyde Phenylacetaldehyde Cofactor NAD NADP Deamino-NAD 2-Acetylpyridine-NAD

II

VELOCITIES

Enzyme M)

645

DEHYDROGENASES

FOR VARIOUS

ALIPHATIC

I V (relative)

3.10 0.15 3.10 4.60 3.60 0.07 0.56 4.50 4.30 4.30 3.20 1.30 2.40 1.40 2.70

120 97.0 100 83 59 0.7 72 294 317 29 12 33 23 62 88

0.3

100 N.D. 30 53

326 22

ALDEHYDES”

Enzyme K,a (X

lo4 M)

15.0 4.5 7.4 3.8 0.23 0.47 1.6 2.7 19.0 19.0 32.0 0.41 0.05 0.04

6.0 170 46 14

V (relative) N.D. 67 100 61 30 78 41 30 196 45 30 62 40 132 74

II u* +0.490 0.000 -0.100 -0.115 -0.190

+ 1.050

+0.215

100 50 23 25

n Michaelis constants (K,) and maximal velocities (V) were determined by conventional LineweaverBurk plots. The rate of NADH formation was measured spectrophotometrically as described in the text. Partially purifed rat liver mitochondrial aldehyde dehydrogenase (from DEAE-cellulose step) was used. NAD, 1 mM, and various concentrations of aldehyde or 1 mM propionaldehyde and various concentrations of NAD analogs were used as substrates. Relative V values were calculated utilizing that obtained with NAD and propionaldehyde as 100. N.D., not detected. u* are Taft values.

at a rate much closer to that of glutamate dehydrogenase (and malate dehydrogenase also, not shown). Numerous studies previously showed that less than 2% of succinate dehydrogenase was released into the 9500g supernatant by even twice the digitonin concentration employed here (24). These results indicate that enzyme I is localized in the matrix of the mitochondria and enzyme II may be localized in the intermembrane space. A summary of the results is found in Table IV. DISCUSSION

The work reported in this study confirms the presence of at least two aldehyde dehydrogenase enzymes in rat liver mitochondria by a variety of techniques and, in addition, describes their properties and possible intramitochondrial localization. Two mitochondrial enzymes were reported by Tottmar et al. (4) and given the desig-

nations of enzyme I and enzyme II; separation by kinetic means was achieved. These designations of enzymes I and II fortuitously conform to our nomenclature of these enzymes as well. Similarly Koivula and Koivusalo (25) also find two mitochondrial enzymes and designate them enzymes I and II. Grunnet (3) also presented evidence suggestive of two mitochondrial enzymes but could not achieve separation of these activities. None of these groups achieved significant purification of these enzymes. In order to establish the presence of two independent enzyme activities a number of separation procedures was applied directly to the sonicated mitochondrial supernatant fraction. Thus, ammonium sulfate fractionation, Sephadex, DEAE-cellulose, and hydroxylapatite column chromatography all achieved separation of two enzyme activities. It is unlikely that each of these procedures would result in disso-

646

SIEW,

DEITRICH TABLE

EFFECT

OF INHIBITORs’

ON PARTIALLY

PURIFIED

DEHMROCENASE

Inhibitor

p-Chloromercuribenzoate

Sodium

Chloral

Disultiram

arsenite

hydrate

AND III RAT

LIVER

MITOCH~NDRIAL

ALDEHYDE

in Vitro”

Solvent

Final

Water

Water

Water

1%

ERWIN

Propylene glycol

concentration (M)

3.0 8.0 1.0 1.3

x 10-5

3.0 6.0 2.0 3.0

x IO-3 x 1O-3 x 10-Z

1.0 2.0 3.0 6.0

x 10-G

1.0

x 10-C

Inhibition Enzyme

x lo-” x IO-4 x 10-d

x 10-Z

x 10-G x 10-G

x 1Om6

3.0 x 10-G 6.0 x 10-O

I

(%a) Enyzme

17.7 86.8 88.6 96.2

0.0 72.5 76.4 95.0

24.3 30.0 35.6 48.5

21.4 26.8 32.8 37.5

49.3 54.3 55.9 64.3

24.7 27.0 22.4 53.6

42.0 84.1 67.2

20.0 35.3 53.0

II

” The rate of NADH formation was measured spectrophotometrically at 340 nm as described in the text. Partially purified enzymes from the DEAE-cellulose step were used. Enzyme and inhibitor were incubated at 25°C for 3.0 min before reaction was started by adding NAD (1 mM) and propionaldehyde (3.3 mm).

TIME

(SECONDS

Enzyme

I

Enzyme

II

)

FIG. 6. Heat denaturation curves for aldehyde dehydrogenase. Enzymes I and II as obtained from the DEAE-cellulose step were heated at 50°C in 50 mM sodium phosphate buffer, pH 7.4. Aliquots were removed, cooled, and centrifuged, and enzyme activity was determined at the indicated times. Log of the initial activity (taken as 100%) was plotted as a function of time. Protein concentration was 2.0 mgiml.

ciation of a polymeric enzyme to yield the same two enzymatic activities. Partial purification of the two enzyme activities was carried out by a combination of these techniques. Study of the characteristics of these two enzymes reveals the presence of a low K, enzyme of molecular weight of 320,000 (enzyme I) and of a high K,,, enzyme (II) with a molecular weight of

67,000. The substrate specificities of these enzymes reveal that only enzyme I will significantly catalyze the oxidation of formaldehyde. Only enzyme II will utilize NADP as a cofactor. The possibility of the presence of an aldehyde dehydrogenase enzyme in liver that would utilize NADP as cofactor was reported earlier (261, could not be demonstrated in mitochondria by

MITOCHONDRIAL

ALDEHYDE

Grunnet (31, but was found by Tottmar et al. (4). The partially purified enzyme II utilizes NADP as cofactor with a reasonable V (50% of the V with NAD), but the K,,, is very high, calling into question the possible physiological significance of such activity. This activity, however, proved useful in submitochondrial localization studies. The correlation of the reaction rates with Taft’s o* values demonstrates that these aldehyde dehydrogenases function more efficiently with aldehyde substrates possessing more strongly electronegative carbonyl carbon atoms. This feature of the aldehyde dehydrogenase mechanism

Ti:E

(min.)

FIG. 7. Release of mitochondrial enzymes during incubation with digitonin. Rat liver mitochondria were prepared as described in the text. They were incubated at 0°C with 1 mg of digitonin per 10 mg of protein (final protein concentration was 50 mg/ml) for the time indicated, diluted with three volumes of cold 0.25 M sucrose, and centrifuged at 95OOg for 10 min. The supernatant was assayed for adenylate kinase (A.K.), monoamine oxidase (MAO), glutamate dehydrogenase (G.D.), high K,,, aldehyde dehydrogenase (Ald.D.), NADP aldehyde dehydrogenase, low K,,, aldehyde dehydrogenase, and formaldehyde dehydrogenase (Formald.), as indicated. Percentage of release is calculated by taking the total present as the amount of enzyme in a sonicated preparation. Samples were taken up to 60 min, of which only the first 15 min are shown.

DEHYDROGENASES

647

seems to be common to all such enzymes so far studied (27, 28). Inhibition by sulfhydryl reagents is a common feature of aldehyde dehydrogenase enzymes, and these are no exceptions. There are minor differences in sensitivity to these reagents as well as to chloral hydrate but they are not large enough to afford a tool for differentiation of the enzyme activities in crude preparations. In results reported since completion of this work a similar picture is seen (25). The submitochondrial localization of these enzymes is of importance. Smith and Packer (29) as well as Grunnet (3) suggested that a portion of the total aldehyde dehydrogenase activity of the mitochondria must lie outside the inner membrane because there is some NAD-dependent aldehyde oxidation which is not coupled to the electron transport chain. Schnaitman and Greenawalt (22) as well as Grunnet (3) utilized digitonin at one concentration for a single time and concluded that the enzyme activity was primarily in the matrix but some possibly in the outer membrane. Smith and Packer (29) concluded that the enzyme is membrane bound. Data from Tottmar and Kiessling (personal communication), who utilized swelling in phosphate, reveal the presence of a low K, enzyme in the matrix and inner membrane, while another high K, enzyme is associated with the outer membrane and with matrix. By utilizing two propionaldehyde concentrations and formaldehyde as substrates with both NAD and NADP as cofactors it was possible to demonstrate a time-dependent release of at least two aldehyde dehydrogenase enzymes from digitonin-treated mitochondria. The high K, enzyme activity and the NADP-dependent enzyme activity appear to be similar in their rate of release from mitochondria and also similar but somewhat slower than the release of monoamine oxidase and adenylate kinase. This is presumptive evidence for the localization of the high K,, aldehyde dehydrogenase (enzyme II) in the inter-membrane space. The low K, enzyme activity (enzyme I) is released at essentially the same rate as glutamate and malate dehydrogenases.

648

SIEW,

DEITRICH

AND

TABLE SUMMARY

OF THE

PROPERTIES

AND

POSSIBLE

Molecular Substrate Cofactor Inhibitor

weight specificity specificity sensitivity

PI

DEAE-cellulose Hydroxylapatite Possible location

binding

IV

SUBMITOCHONDRIAL

ALDEHYDE

Enzyme

ERWIN

LOCATIONS

OF Two

NAD-DEPENDENT

Enzyme

II

DEHYDROGENASES

I

320,000Low K,,, (micromolar) +HCOH NAD More sensitive 6.06 Eluted at 50 mM phosphate, Eluted at 0.2 M phosphate, Matrix

This release is similar but does not appear to be identical with the release of formaldehyde-dependent activity. The partially purified enzyme I is more active with formaldehyde than propionaldehyde as a substrate so it is difficult to postulate the presence of a second enzyme in the matrix, although that is not ruled out. Perhaps digitonin has some selective effect on the activity with formaldehyde as a substrate, if a single enzyme is responsible. The absolute amount of enzyme in each of the two compartments, intermembrane space and matrix, can be estimated utilizing the 60-min point in the digitoninrelease experiments. At this time all of both enzymes has been released. The low K,, enzyme activity accounts for about 35% of the total as measured with high propionaldehyde concentrations. Comparison with the data of Smith and Packer (29) allows the calculation that 37% of the total aldehyde dehydrogenase is associated with the inner membrane; Grunnet (3) suggested that 80% of the activity ‘is in the matrix while Tottmar and Kiessling (personal communication) find that about 50% of the total enzyme activity is due to the low K, enzyme. They found that enzyme was associated primarily with the matrix plus intermembrane space and inner membrane. In a paper published since submission of this communication, Cinti et al. (30) found that a formaldehyde dehydrogenase is localized in the matrix fraction of rat liver mitochondria. They find 80% of the activity in this fraction of the mitochondria and this enzyme is dependent upon a functional electron transport system in the absence of added NAD. In a

pH 7.4 pH 7.4

67,000 High K,,, (millimolar) - HCOH NAD and NADP Less sensitive 6.64 Eluted at 1 mM phosphate, Eluted at 0.4 M phosphate, Intermembrane space

pH 7.4 pH 7.4

recent study Horton and Barrett (31) conclude that enzyme II is chiefly located in the outer membrane. Since our method of fractionation of mitochondria utilizes digitonin and their method employs phosphate swelling, it is possible that the divergence between our conclusions and those of Horton and Barrett may be due to release of enzyme II from outer membrane by digitonin. It would be our view that the low K, enzyme is confined to the matrix, and thus oxidation of acetaldehyde, when that aldehyde is generated from ethanol, will take place in the matrix. In the face of higher concentrations of acetaldehyde such as occur after certain drug treatments, or possibly in alcoholic humans (32), some oxidation may occur in the intermembrane space or cytoplasm. The oxidation of other aldehydes will depend upon the V and K,,? values for these substrates with the various enzymes. The role of high K, aldehydesdehydrogenase enzymes in normal or abnormal biochemical processes remains to be defined. REFERENCES R. A., AND SIEW, C. (1974) in Alcohol and Aldehyde Metabolizing Systems (Thurman, R. G., Yonetani, T., Williamson, J. R., and Chance, B., eds.), pp. 125-135, Academic Press, New York. 2. MARJONEN, L. (1972) Rio&em. J. 127, 633-639. 3. GRUNNET, N. (1973) Eur. J. Biochem. 35, 236243. 4. TO~TMAR, S. 0. C., PETTERSSON, H., AND KIESSLING, K-H. (1973) Biochem. J. 135, 577-586. 5. LINDROS, K. O., OSHINO, N., PARRILLA, R., AND WILLIAMSON, J. R. (1974) J. Biol. Chem. 249, 7956-7963. 1. DEITRICH,

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R., ZIMMERMAN, WILLIAMSON,

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OHKAWA, K., LINDROS, K. O., U. U. P., KOBAYASHI, K., AND J. R. (1974) J. Biol. Chem. 249,

4926-4933. 7. GRUNNET, N., QUISTORFF, B., AND THIEDEN, H. I. D. (1974): in Alcohol and Aldehyde Metabolizing Systems. (Thurman R. G., konetani, T., Williamson, J. R., and Chance, B., eds.) pp. 137-146, Academic Press, New York. 8. CEDERBAUM, A. I., LIEBER, C. S., BEATTIE, D. S., AND RUBIN, E. (1973)Arch. Biochem. Biophys. 158, 763-781. 9. DEITRICH, R. A., AND ERWIN, V. G. (1975) Fed. Proc. 34, 1962-1968. 10. DEITRICH, R. A., AND HELLERMAN, L. (1963) J. 11.

Biol. LAYNE,

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238, 1683-1689. (1975) in Methods

E. in Enzymology (Colowick, S. P., and Kaplan, N. O., eds.), Vol. 3, pp. 450-451, Academic Press, New York. 12. STRECKER, H. J. (1955) in Methods in Enzymology (Colowick, S. P., and Kaplan, N. O., eds.), Vol. 2, pp. 220-225, Academic Press, New York. 13. GIANETTO, R., AND DEDUVE, C. (1955) Biochem. J. 59, 433-438.

14. MAZEL, P. tabolism Mandel, Williams 15. LUNDQUIST,

(1971) in Fundamentals of Drug Meand Drug Disposition, (LaDu, B. N., H. G., and Way, E. L., eds.), p. 546, and Wilkins, Baltimore. F., AND WOLTHERS, H. (1958) Acta Phurmacol. Toxicol. 14, 265-289. 16. ACKERS, G. K. (1964) Biochemistry 3, 723-730. 17. MARTIN, R. B., AND AMES, B. N. (1961) J. Biol. Chem. 236, 1372-1379.

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C. A., ERWIN, V. G., AND GREENAWALT, J. W. (1967) J. Cell Biol. 32, ‘719-735. 19. DEITRICH, R. A., AND ERWIN, V. G. (1969)Anal. Biochem. 30, 395-402. 20. OCHOA, S. (1955): in Methods in Enzymology,

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(Colowick, S. P., and Kaplan, N. O., eds.), pp. 735-739, Academic Press, New York. ARRIGONI, O., AND SINGER, T. P. (1962) Nature (London) 193, 1256-1258. SCHNAITMAN, C. A., AND GREENAWALT, J. W. (1968) J. Cell Biol. 38, 158-175. TAFT, R. W., JR. (1956) in Steric Effects in Organic Chemistry (Newman, M. S., ed.) pp. 591-595, Wiley, New York. SIEW, C. (1975) Thesis, University of Colorado, Boulder, Colo.

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L., AND WEIN, J. (1962) J. Biol. Chem. 237, 560-564. 28. FELDMAN, R. I., AND WEINER, H. (1972) J. Biol. Chem. 247, 267-272. 29. SMITH, L., AND PACKER, L. (1972) Arch. Biothem. Biophys. 148, 270-276. 30. CINTI, D. L., KEYES, S. R., Lemelin, M. A., DENK, H., AND SCHENKMAN, J. B. (1976) J. Biol. Chem. 251, 1571-1577. 31. HORTON, A. A., AND BARRETT, M. C. (1975)Arch. Biochem. Biophys. 167, 426-436. 32. KORSTEN, M. A., MATSUZAKI, S., FEINMAN, L., AND LIEBER, C. S. (1975) N. Engl. J. Med. 292, 386-389.

Localization and characteristics of rat liver mitochondrial aldehyde dehydrogenases.

ARCHIVES OF BIOCHEMISTRY Localization AND BIOPHYSICS 638-649 (19763 and Characteristics of Rat Liver Mitochondrial Dehyd rogenasesl CHAKWAN D...
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