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1985 Amino acid metabolism in ribbed mussel gill tisue during hypersmotic stress: role of transaminases and pyruvate dehydrogenase Kennedy T. Paynter Jr. Iowa State University

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Paynter, Kennedy T., Jr.

AMINO ACID METABOLISM IN RIBBED MUSSEL GILL TISSUE DURING HYPEROSMOTIC STRESS: ROLE OF TRANSAMINASES AND PYRUVATE DEHYDROGENASE

Iowa State University PH.D. 1985

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I nternstionâ! 300 N. Zeeb Road, Ann Arbor, Ml 48106

Amino acid metabolism in ribbed mussel gill tissue

during hyperosmotic stress:

Role of transaminases and pyruvate dehydrogenase

by

Kennedy T. Paynter, Jr.

A Dissertation Submitted to the

Graduate Faculty in Partial Fulfillment of the

Requirements for the Degree of

DOCTOR OF PHILOSOPHY

Major: Zoology

Approved: Members of the committee:

Signature was redacted for privacy. Signature was redacted for privacy. in C^i^e'of Major Work

Signature was redacted for privacy.

For the Major Department

Signature was redacted for privacy.

For the Graduate College

Iowa State University Ames, Iowa

1985 ii

TABLE OF CONTENTS

Page

GENERAL INTRODUCTION 1

PART I. PARTIAL PURIFICATION AND CHARACTERIZATION 5 OF ASPARTATE AMINOTRANSFERASE FROM RIBBED MUSSEL GILL TISSUE

INTRODUCTION 6

MATERIALS AND METHODS 9

RESULTS 14

DISCUSSION 26

PART II. CELLULAR LOCATION AND CHARACTERIZATION 30 OF ALANINE AMINOTRANSFERASE FROM RIBBED MUSSEL GILL TISSUE

INTRODUCTION 31

MATERIALS AND METHODS 34

RESULTS 37

DISCUSSION 49

PART III. SUBCELLULAR DISTRIBUTION OF AMINOTRANS- 55 FERASES, AND PYRUVATE BRANCH POINT ENZYMES IN GILL TISSUE FROM FOUR BIVALVES

INTRODUCTION 56

MATERIALS AND METHODS 58

RESULTS 61

DISCUSSION 64

PART IV. PYRUVATE DEHYDROGENASE FROM RIBBED 67 MUSSEL GILL TISSUE

INTRODUCTION 68

MATERIALS AND METHODS 70

RESULTS AND DISCUSSION 73 iii

Page

GENERAL DISCUSSION 84

ACKNOWLEDGEMENTS 87

LITERATURE CITED 88 1

GENERAL INTRODUCTION

Many marine bivalves are capable of withstanding a wide range of salinities and long periods of anoxia. The ribbed mussel, Modiolus demissus. a euryhaline bivalve found in salt marshes along the

Atlantic and Pacific coast of North America, and the blue mussel,

Mvtilus edulis. have been the two most commonly used animals to study cellular adaptations to osmotic and anaerobic stress in bivalves.

Tissues of the ribbed mussel accumulate intracellular free amino acids in response to hyperosmotic stress (Baginski and Pierce, 1975, 1977,

1978; Bishop, 1976, Bishop et al., 1981; Pierce, 1982; Strange and

Crowe, 1979a,b). Alanine accumulates to high (0.1-0.2 M) levels within a few hours after a hyperosmotic shift. Glycine levels rise more slowly, while proline and taurine accumulate after extended

<10-20 h) shifts. Aspartate levels, on the other hand, fall quickly with hyperosmotic shock. These amino acid shifts are similar to those- which occur with hypoxic or anoxic stress in Modiolus (Baginski and

Pierce, 1975, 1977) and other bivalves (Collicutt and Hochaclika, 1977; deZwaan et al., 1983a,b). Under hypoxic conditions, succinate accu­

mulates quickly and with prolonged anoxia, propionate accumulates (Ho

and Zubkoff, 1982, 1983). In addition, the cellular pH falls during

anoxia (Wijsman, 1975; Barrow et al., 1980; Ellington, 1983). 14 Studies with C-aspartate have shown that at least part of the

accumulated succinate is derived from aspartate (Collicutt and

Hochachka, 1977; deZwaan, 1977). Other studies using hadacidin (a

purine nucleotide cycle inhibitor) have demonstrated that the purine 2

nucleotide cycle plays a negligible role in the catabolism of aspar­ tate in mussels (Bishop et al., 1981; deZwaan et al., 1981). The transaminase inhibitor, aminooxyacetic acid, completely blocked the

catabolism of aspartate and the accumulation of alanine, indicating that the metabolism of these amino acids during hyperosmotic or anaerobic stress is via a transaminase-linked pathway (Greenwalt and

Bishop, 1980; Bishop et al., 1981; deZwaan et al., 1981).

Many alternate metabolic pathways have been proposed to account

for accumulated end products and energy production in invertebrates

exposed to anoxia (Hochachka, 1981; Ho and Zubkoff, 1982; Kluytmans et

al., 1975; Rew and Saz, 1974; deZwaan, 1977; deZwaan et al., 1981;

Saz, 1981). Central to these metabolic hypotheses are several consid­

erations. First, much attention has been given to the so-called

"redox balance" or NAD/NADH ratio which is believed to change sig­

nificantly during anaerobiosis in mussels (although no experimental

evidence has been published to verify this assumption). Second, amino

group balance must be maintained. Third, subcellular compartmentation

of these metabolites must be considered and experimentally substan­

tiated, especially with regard to enzyme activités and metabolite flux

between compartments. Many groups (see Hochachka, 1981) assume cer­

tain enzyme properties or locations based on mammalian or other animal

literature.

The control point for most of these alternate anaerobic pathways

has been proposed by a number of laboratories to be at the pyruvate

kinase reaction, regulating the "PEP branchpoint" (deZwaan, 1977; 3

Hochachka, 1981; Holwerda et al., 1981, 1984; deZwaan and Dando,

1984). Pyruvate kinase (PK) has been shown to be a covalently mod­ ified (phosphorylated) enzyme with different properties between phospho- and dephospho- forms (see Munday et al., 1980; Holwerda et al., 1984), and is inhibited allosterically by alanine. According to this hypothesis, the basic mechanism for the switch to anaerobic metabolism in bivalves was the inhibition of PK by covalent modifi­ cation or allosterically by alanine as that amino acid began to accumulate. Additionally, as the pH decreased during hypoxia, phosphoenolpyruvate carboxykinase (PEPCK) would be activated and glycolytically derived carbon skeletons would be shunted to oxalo- acetate to produce malate, not pyruvate, which would be transported into the mitochondria for metabolism to succinate (see deZwaan, 1977; deZwaan and Dando, 1984). While this pathway accounts for many aspects of the anaerobic metabolism, it ignores the role of the mito­ chondria. This is not surprising since virtually no data are available on the molluscan pyruvate dehydrogenase complex, which would control the pyruvate metabolism in the'mitochondria.

The above pathway has received general acceptance, but it leaves several unanswered questions. First, how does the control of aspar­ tate metabolism fit in? Second, what controls the initial accumula­ tion of alanine? It was generally assumed by many investigators that aspartate was transdeaminated to oxaloacetate, which was reduced to

malate via malate dehydrogenase, and that the amino group from aspar­ tate was used to transaminate pyruvate to form alanine. 4

Considering these assumptions, and the lack of an "immediate" or ultimate control point in the models, the aspartate and alanine amino­ transferases were determined to be critical "gaps" of knowledge. A characterization of these enzymes from the ribbed mussel would shed some light on the controls of aspartate catabolism and alanine accumulation. Part I of this work will focus on the kinetic proper­ ties of the aspartate aminotransferase activities from the cytosolic and mitochondrial fractions of ribbed mussel gill tissue. Part II will deal with the cellular location and properties of the alanine aminotransferase from this tissue. Part III is a short presentation of some comparative data dealing with enzyme levels and identities within mitochondrial or cytosolic fractions. Lastly, Part IV will describe the preparation, and partial characterization of the pyruvate dehydrogenase complex from mitochondria isolated from the gill tissue. 5

PART I. PARTIAL PURIFICATION AND CHARACTERIZATION OF ASPARTATE AMINOTRANSFERASE FROM RIBBED MUSSEL GILL TISSUE

The cytosolic (cAAT) and mitochondrial (niAAT) aspartate amino­ transferases were partially purified from gill tissue of the ribbed mussel, Modiolus demissus, and individually characterized with respect to heat stability, electrophoretic mobility, pH optimum, for substrates, rates in the forward vs. reverse direction, and reactivity with aminooxyacetic acid (ADA). Isozyme analysis for this population of ribbed mussels indicated three alleles for the cAAT and a single allele for the mAAT. The mAAT was a single isozymic form with a broad pH optimum and relatively low K^s for aspartate, oxaloacetate, and a-ketoglutarate and high for glutamate. The cAAT used for the kinetic studies was a single (homozygous) isozymic form, showed great variation in and V with pH, and high K s for the m max m amino acid substrates and low K s for the keto acid substrates. m Both enzymes were inhibited to the same degree by AOA. It is

hypothesized that the cAAT is involved more with aspartate synthesis

and the mAAT with aspartate catabolism. 6

INTRODUCTION

Aspartate aminotransferase (AAT) (L-aspartate: 2-oxoglutarate aminotransferase, E.G. 2.6.1.1) has been found in tissues of most organisms. The active form of the enzyme is a dimer with identical or nearly identical subunits in all tissues investigated (Fleisher et al., 1960; Odense et al., 1966; Magee and Phillips, 1971; Orlacchio et al., 1979; Porter el al., 1981a,b; Doonan et al., 1981). The two basic types, mitochondrial AAT (mAAT) and cytosolic AAT (cAAT), often show electrophoretic polymorphism of probable genetic origin plus differ­

ences in biosynthetic and assembly patterns, molecular weights, kine­ tic properties and reactivity with inhibitors (Wada and Merino, 1964;

Martinez-Carrion and Tiemeier, 1967; Michuda and Martinez-Carrion,

1969a,b; Davidson et al., 1970; Turner, 1973; Smith and Freedland,

1981; Ueno et al., 1982; Behra et al., 1981, 1982).

AAT has been found in tissues of all molluscs investigated

(Bishop et al., 1983; Campbell and Bishop, 1970). The levels of

activity in molluscan tissues vary from the very high in muscle of

some cephalopods to the very low in the tissues of some freshwater

bivalves and gastropods. Multiple molecular forms of the AAT activity

have been detected in some marine bivalves. In blue mussels (Mvtilus

edulis), Johnson and Utter (1973) report thirteen phenotypes with

evidence of a six allelic system in whole body homogenates. Johnson

et al., 1972 reported six phenotypes reflecting multiple alleles in

oyster (Ostrea lurida) adductor muscle homogenates. Johnson made no

attempt to determine which of the AAT isozymes were mAAT or cAAT. 7

Although AAT activity has been detected in both the cytosolic and mitochondrial compartments of tissues from sea mussels (Addink and

Veenhof, 1975); pulmonate snails (Sollock et al., 1979); and oysters

(Chambers et al., 1975), comparisons of the properties of cAAT and mAAT have not been made.

In other studies with a number of marine bivalves, Hammen (1968) found that the smaller animals had higher specific activities in their tissues than the larger animals. Although DuPaul and Webb (1974) found little difference in relative AAT activity in the gill tissue of

bivalves adapted to different salinities, Wickes and Morgan (1976)

reported that tissue AAT activities were higher in oysters collected

at high salinités than in oysters collected at low salinities. Read

(1962) found differences in the thermal stability of AAT in whole body

horaogenates of a number of marine bivalves, including Modiolus

demissus and Mvtilus edulis. Although these data are not conclusive,

they seem to indicate some differences in AAT properties among tissues

and compartments within tissues which may be of physiological signif­

icance.

The changes in aspartate and other amino acid levels in heart and

gill tissue subjected to osmotic stress are blocked by addition of

aminotransferase inhibitor aminooxyacetic acid (AOA) (Greenwalt and

Bishop, 1980; Bishop et al., 1981). The catabolism of aspartate to

CO^ by ribbed mussel gill tissue is inhibited 80-90% by AOA, indi­

cating that the major route of aspartate catabolism is via a trans­

aminase linked pathway rather than via the purine nucleotide cycle 8

(Bishop et al., 1981). Although Greenwalt and Bishop (1980) demon­ strated that AOA inhibited the total AAT and other transaminase activities in tissue homogenates, there was no attempt to define the sensitivity of the various isozymic forms of the transaminases to AOA.

The mAAT and cAAT activities from gill tissue of demissus have

been partially purified and partially characterized to assess possible

physiological roles of the two enzymes in regulating amino acid levels

in the tissues of this estuarine bivalve. 9

MATERIALS AND METHODS

Ribbed mussels (Modiolus demissus), purchased from Northeast

Environmental Laboratories (Monument Beach, Mass.), were kept in artificial sea water (Jungle Laboratories Inc., Sanford, Fla.) and maintained as described by Greenwalt and Bishop (1980). Except where noted, all reagents were purchased from Sigma Chemical Co., St. Louis,

MO. Ammonium sulfate (Enzyme grade) was obtained from Schwartz-Mann

(Orangeburg, N.Y.).

Enzyme assay

During purification and standard assay, the AAT activity was determined spectrophotometrically by measuring oxaloacetate production as NADH oxidation (340nm) in a reaction mixture containg 20 mM aspar­ tate, 10 mM a-ketoglutarate, 70 yM NADH, 4 units malate dehydrogenase,

100 mM Tris-HCl (pH 8.3), and enzyme in 2 ml. Incubations were at room temperature (23°C). Modifications of this assay mixture for kinetic measurements are described below. One unit of activity equals one ymole of product produced per min under the conditions specified. Kinetic constants were determined using computer analysis of initial rates (Cleland, 1979). This method utilizes a least squares analysis to determine the best linear fit.

Isozyme distribution

Isozyme distribution and purity of electromorphic types was

determined using starch-gel electrophoresis (Odense et al., 1966).

A liter of stock electrophoresis buffer contained 15.1g Tris (Base), 10

1.5g EDTA, 1.15g boric acid, and IM acetic acid to pH 5.5. Samples of whole tissue were homogenized in an equal volume of electrophoresis buffer using a Polytron (Brinkman Inst., Westbury, N.Y.) tissue homo- genizer with the small probe. Thirteen percent (w/v) horizontal starch gels (12 cm x 20 cm x 0.8 cm) were electrophoresed at approxi­ mately 35 ma at 4°C. Progress was marked with Bromphenol blue.

Gels were sliced and stained at 28°C in 200 mg L-aspartate, 100 mg a-ketoglutarate, and 150 mg Fast Blue BB in 100 ml of 50 mM

Tris/HCl pH 8.3 (Harris and Hopkinson, 1976).

Enzyme preparation

A summary of results from the enzyme preparation is found in

Table 1. Gills were removed from the mussels and collected in a chilled, tared beaker until approximately 75 g of tissue had been accumulated. The tissue was rinsed in sea water, then homogenized in

750 ml of isolation buffer (0.5 m sucrose, 0.15 M KCl and 20 mM HEPES at pH 7.5) with an "Ultra-Turrax" homogenizer (Tekmar Co., Cincinnati,

OH) using three ten-second bursts at a speed setting of "40". The homogenate was filtered through Miracloth (Calbiochem) to remove large particulate debris. This filtrate was centrifuged at 1500 x g for 8

minutes, the supernatant fluid collected and then centrifuged at 9000

X g for 15 minutes. The higher speed centrifugation yielded a pel­

let containing mitochondria which was resuspended in 20 mM potassium

phosphate buffer pK 6.8, and held at -20°C until needed. The super­

natant fluid from the higher speed centrifugation was saved for

purification of the cAAT activity. 11

For the preparation of the cAAT (see Table 1), finely ground ammonium sulfate (35g/100ml) was added slowly to the supernatant fluid from the mitochondria separation step with constant stirring at ice water temperature. After 30 minutes, the precipitate was removed by centrifugation at 15,000 x g for 30 minutes. An additional 17.5 g ammonium sulfate was added slowly with constant stirring to each 100

ml of the supernatant fluid. After 30 minutes, the precipitate, containing most of the AAT activity, was collected by centrifugation and dissolved in a minimal amount (50 ml) of 20 raM potassium phosphate

(pH 6.8). Potassium a-ketoglutarate (0.2M) was added to bring the

dissolved enzyme solution to 5 mM a-ketoglutarate. This

preparation was then placed in a 80°C water bath and heated to

70°C with constant swirling, then held at 70°C for 10 minutes.

After cooling on ice to ice water temperature, the preparation was

centrifuged at 15,000 x 3 for 30 minutes. The supernatant was

decanted into washed dialysis tubing and dialyzed for 12 hrs at 4°C

against four liters of 10 mM potassium phosphate (pH 6.8). Hydroxyl-

apatite (Bio-Rad Laboratories, Richmond, CA) was washed in degassed 10

mM potassium phosphate (pH 6.8), the fine particles removed and the

slurry poured into a column to give a 2 x 10 cm bed. After equili­

bration of this column with degassed dialysate (see above), the

preparation from within the dialysis bag was added into the column and

the column was washed with 150 ml of degassed dialysate. The AAT

activity was eluted as a single peak by washing the column with 100 ml

of 150 mM potassium phosphate buffer (pH 6.8). Fractions (1 ml) were 12

collected and the active fractions pooled (5 ml) to constitute the final preparation of the cAAT activity {Table lA).

Table 1. Partial purification of AAT activities from gill tissue of ribbed mussels^

A. Cytosolic AAT Activity

Fraction Volume Total Total Specific Protein Activity Activity (ml) (mg) (units) (units/mg)

Crude Extract 600 2328 54.0 0.023 (NH.).SO. 45 212.4 15.9 0.075 Heat Extract 43 80.8 14.6 0.181 Dialysis 69 41.4 16.4 0.396 HAP 4 3.8 6.9 2.167

B. Mitochondrial AAT activity

Fraction Volume Total Total Specific Protein Activity Activity (ml) (mg) (units) (units/mg)

Crude Extract 17 31.96 14.2 0.44 Heat Extract 17 17.68 10.9 0.62 («"4)2^4 2.5 0.70 9.6 13.7

^Activities were assayed in the oxaloacetate forming direc­ tion using the standard procedure described in Materials and Methods.

The mAAT activity was prepared (Table IB) from mitochondria lysed

by several freeze/thaw cycle treatments of the mitochondria. After

final thawing, the suspension was homogenized vigorously, then centri-

fuged at 15,000 x g for 15 minutes to remove debris. The superna­

tant fluid from the lysate was mixed with enough 0.2 M potassium 13

a-ketoglutarate to bring the solution to 5 mM, then heated quickly to 55°C in a 75°C water bath. The resulting precipitate was removed by centrifugation at 15,000 x g for 30 minutes. Ammonium sulfate (45g/100 ml) was added slowly to the supernatant fluid with stirring, and the precipitate removed by centrifugation was discarded.

Additional ammonium sulfate (llg/100 ml) was slowly stirred into this supernatant fluid and the fine precipitate collected by centrifugation at 15,000 X 3 for one hour. The fine, sometimes clear, pellet was dissolved in one ml of 20 mM potassium phosphate pH 6.8, and used for subsequent experiments.

Protein was measured by the Lowry method as modified by Miller

(1959). 14

RESULTS

Electrophoresis of cell free gill extracts from fifty individual^ animals indicated that there were two major AAT isozymic families

(Figure 1). One group of isozymes migrated toward the anode and the other migrated toward the cathode at both pH 9 (Figure la) and pH 6.5

(Figure lb). At pH 6.5, two of the fifty animals had three banded

electrophoretic patterns with respect to the anodally migrating AAT

(Figure a,b), typical of heterozygotes for two different isozyme

alleles. All of the other preparations showed the tissue to be single

banded (homozygous) for the intermediate anodally migrating form

(Figures la,b). The purified cAAT (Table lA) migrated anodally as the

single intermediate (homozygous) for the intermediate anodally mi­

grating form (Figures la,b). The purified cAAT (Table lA) migrated

anodally as the single intermediate (homozygous) allozyme (Figure Ic).

The purified mAAT (Table IB) migrated cathodally as a single zone of

activity (Figure Ic).

The levels of the mAAT and cAAT activities in four tissues are

listed in Table 2. The mantle tissue appeared to have the lowest and

the adductor muscle the highest total AAT activity. The gill and

hepatopancreas tissues had about equal amounts of mAAT and cAAT activ­

ity, whereas most the AAT activity in the adductor muscle was cyto-

solic, possibly reflecting a small number of mitochondria and the

basic glycolytic nature of adductor muscle tissue (Addink and Veenhof,

1975; deZwaan, 1977; Zurburg and Kluytmans, 1980). An electrophoretic

evaluation of the AAT isozyme distribution in four tissues (gill, -f" +

o

b c

Figure 1. Distribution of AAT isozymes using starch gel electrophoresis. Whole gill homogenates were applied to wells and electrophoresed at pH 9.0 (a) and pH 6.5 (b). Purified preparations (c) (Table 1) were applied separately (lanes 2 and 3) and mixed (lane 1). Procedures are described in Materials and Methods. Each lane represents a different animal and lanes 9 and 10 in (a) represent hétérozygotes for AAT. The same animals are repeated in (b) 16

mantle, hepatopancreas, and adductor muscle) from fifty individual mussels indicated that all four tissues had the same isozyme pattern and the same isozymes (data not shown).

Table 2. Distribution of AAT activities in different organs and tissue compartments of ribbed mussel tissues

Activity (munits/g tissue wet wt)

AAT Isozyme Gill Mantle Hepatopancreas Adductor

Mitochondrial 213.6 27.2 261.6 90.8

Cytosolic 200.0 58.8 281.1 1084.8

Because the major isozymes of the cAAT and mAAT activities were similar in all tissues, a series of experiments were performed to evaluate the similarities and differences between the cAAT and mAAT activities in ribbed mussel gill tissue. Partially purified AAT preparations of the homozygous cytosolic and mitochondrial isozymes

(see Materials and Methods; Table 1A,B; Figure Ic) were used to avoid

possible confounding of results by the genetic variation.

Both the mAAT and cAAT activities were relatively heat stable and

both were protected from heat inactivation by the presence of a-keto-

glutarate in the incubation solution (Figure 2a,b). Of the two, the

mitochondrial enzyme (Figure 2a,b) was less heat stable than the cyto­

solic activity (Figure 2a,b).

In preliminary experiments prior to evaluation of substrate

binding, the pH for optimal activity of both activities were found to 17

ISO-

100- Activity

50-

100 TCC)

150-

100- Activity

50-

100

Figure 2. Thermal stability of cAAT (C,0) and mAAT (M,+) isolated from ribbed mussel gill tissue. The heat lability of both isozymes was tested in the absence (A) and presence (B) of stabilizer (a-ketoglutarate). Approximately 30 ul of enzyme was incubated in a waterbath for 2 min at each temperature; 20 ul was then added to a 2-ml reaction cocktail at 23°C and assayed as described in Materials and Methods (oxaloacetate formation) 18

°/o Activity

Figure 3. Relative velocities of mAAT (M) and cAAT (C) at different pH values. Activity was assayed in the oxaloacetate forming direction using standard assay substrate concen­ trations (see Materials and Methods) except that 100 mM potassium phosphate buffer was used instead of Tris/HCl 19

be about pH 8.3 (Figure 3) using the substrate concentrations of the standard assay mixture. Under these conditions, both activities showed a decline in activity above pH 8.5 but very different activ­ ities below pH 7.5. The mAAT was apparently fully active at pH 6.5 with 50% activity at about pH 5.5, whereas the cAAT showed 50% activ­ ity at pH 7 and no activity at pH 5.5.

Substrate binding and relative rates of the AAT activities in both the aspartate synthesizing direction and glutamate synthesizing direction indicated that there were important differences between the cAAT and mAAT activities that may have physiological significance.

Table 3. Apparent kinetic constants of mitochondrial and cytosolic AAT activities in ribbed mussel gill tissue®

Apparent Km for substrates Ratio of Maximal Velocity

AAT Asp Synthesis Isozyme ««.kg «oaa ^glu ^lu Synthesis

Cytoso J. 1 c 5*22 0.04 0,02 5.00 3*2

Mitochondrial 0.59 0.58 0.05 19.80 4.6

Values were determined by the least squares method of Cleland (1979). Standard error for the measurements was 8% in the glutamate forming direction and 20.5% in the aspartate forming direction. Reaction rates in the aspartate forming direction were estimated by measuring the decrease in absorbancy (@260 nm) produced by the decline in oxaloacetate concentration (Velick and Vavra, 1962) using the appropriate 0.2, 0.4, and 1.0 cm light path cuvettes. Fixed concen­ trations of the alternate substrates were 20mM aspartate, lOmM a-ketoglutarate, lOmM glutamate with the cAAT and 50raM glutamate with the mAAT and 4raM oxaloacetate at pH 8.3 in 0.1 M Tris-HCl. 20

0.2-

% V

0.2

0.1

0

Figure 4, Double reciprocal plots of initial velocities of cMT (C) and mAAT (M) at different aspartate (A) and different a-ketoglutarate (B) concentrations. Assays were performed as described in Materials and Methods, keeping the levels of one substrate at or near saturating concen­ tration while the other substrate concentration was manipulated 21

max

pH

Figure 5. Variation in relative maximal velocities of cytosolic (C) and mitochondrial (M) AAT at different pH values. All substrates and 1 M Tris were prepared at the proper pH by titration with 6 M HCl. The mitochondrial AAT (0) and cytosolic (®) AAT were assayed as described in Materials and Methods 22

A B 200 2-Oxoglutarate

• 100

50

1

.5f Km (mM) 10 Km (mM) .1

.05

.02 -1 i_ o 7 10 8 9 10 pH pH

Figure 6. Variation in apparent K for a-ketoglutarate (A) and aspartate (B) for cytosolic (C) and mitochondrial (M) AAT at different pH values. All substrates and 1 M Tris were prepared by titration with 6 M HCl or 4 M KOH. The mitochondrial (0) and cytosolic (©) AAT activities were assayed as described in Materials and Methods 23

The rate of both the cAAT and mAAT activities increased in a direct linear relationship with time and enzyme concentration at all of the various pH values tested using near saturating substrate concentra­ tions. Figure 4a,b shows that there are marked differences between the cytosolic and the mitochondrial enzyme with respect to the apparent for aspartate and or a-ketoglutarate at near saturating concentrations of a-ketoglutarate and aspartate, respectively.

Experiments similar to those in Figure 4a,b were performed for all substrates and data for the specific apparent values at pH 8.3 are summarized in Table 3. Both the cytosolic and mitochondrial AAT activities showed "ping-pong" kinetic patterns (Cleland, 1979) similar to AATs in other tissues (Michuda and Martinez-Carrion, 1969a,b).

With the exception of L-glutamate, the values for all substrates with the mAAT were low whereas the K^s for aspartate and glutamate with the cAAT were relatively high.

The differences in observed velocity between the gill cAAT and mAAT at different pH values (Figure 3) prompted experiments to eval­ uate the change in and relative maximal velocity at a low phys­

iological pH value (pH 6.5), at a near optimal pH value (pH 8.3) and at a high pH value (pH 9.5). With the cAAT, the maximal velocity

increased with decreasing pH while the maximal velocity of the mAAT

increased slightly with increasing pH (Figure 5). With the cAAT, the

for aspartate increased from about 1 mM at pH 9.5 to over 150 mM

at pH 6.4 (Figure 6b). The for a-ketoglutarate increased

from about 0.03 mM at pH 6.4 to over 1 mM as the pH increased at 9.5 24

(Figure 6a). On the other hand, with the mAAT, the K^s for aspartate and a-ketoglutarate were similar at the different pH values (Figure 6a,b). These kinetic data (Table 3) explain the apparent narrow pH range for optimal activity with the cAAT and the broad apparent optimal range for the mAAT activity at the fixed substrate concentrations used in the standard assay mixture (Figure 3).

Previous studies on physiological processes regulating the intracellular free amino acid levels have employed the pyridoxal- phosphate antagonist, aminooxyacetic acid, as a tool in determining probable origins and fates of these amino acids (Greenwalt and Bishop,

1980; Bishop et al., 1981). In these studies (Greenwalt and Bishop,

1980) the apparent inhibition of the AAT activity in the tissues of the ribbed mussel was evaluated using tissue homogenates rather than purified and separated activities. Figure 7 shows that the mAAT and cAAT activities were inhibited to the same degree by AOA with 50%

inhibition of the observed activity (I^g) at 2-3 x 10~^M AOA.

This Igg concentration for AOA is essentially the same as that

reported previously for the mixed enzyme activities in tissue

homogenates (Greenwalt and Bishop, 1980). Beta-chloro-L-alanine and

vinylglycine did not inhibit either the mAAT or the cAAT activities at

concentrations up to 10 mM. 25

100

Activity

log (AO A)

Figure 7. Inhibition of cytosolic and mitochondrial AAT activities by aminooxyacetic acid. Aminooxyacetic acid was prepared in distilled water and added to aliquots of enzyme prep­ aration to give the proper concentration of inhibitor. The enzyme was incubated with inhibitor for 2 min at room temperature and then assayed. Both cytosolic (0) and mitochondrial (+) enzymes showed the same kinetics of inhibition 26

DISCUSSION

The cAAT and mAAT differed from each other in terms of the K„ m values for the amino acids and a-ketoglutarate ; the for OAA was low with both enzymes (Table 3). These values at pH 8.3 are similar to those reported for the cAAT and mAAT from pig heart (Velick and Vavra, 1962; Wada and Morino, 1964; Martinez-Carrion and Tiemeier,

1967; Michuda and Martinez-Carrion, 1969a,b). The ratios of rates

(aspartate forming:glutamate forming) listed in Table 3 for both gill isozymes are similar to the ratio of 4.33 reported for the pig heart

AATs (Velick and Vavra, 1962; Michuda and Martinez-Carrion, 1969b).

Awapara and Campbell (1964) report activity ratios between 1.7 and 3.0 for total AAT activity in homogenates of tissues of two bivalves and a gastropod under standard assay conditions. The relative calculated Keq values for the gill enzymes were 6.68 for the cAAT and 1.59 for the mAAT. Considering the error in the measurements, these Keq values are reasonably close to the values reported by Velick and Vavra (1962).

The mAAT and cAAT differed from each other in terms of relative maximal activity and K for the substrates at the different pH m values tested (Figures 3, 5, 6a,b). The pH profile obtained for the gill mAAT activity is very similar to the profile obtained for the

mAAT and cAAT from pig heart (Velick and Vavra, ,1962; Wada and Morino,

1964). The high (150 mM) for aspartate at pH 6.4 (Figure 6b) and the increase in relative V^^ with decreasing pH seen with the gill

cAAT (Figure 5) is unlike any other AAT. The relative activity pro­

file at the various pHs for the cAAT at fixed substrate concentra- 27

tions {Figure 3) is similar to the profile reported for the cytosolic form of the alanine aminotransferase from rat liver (Swick et al.,

1965).

The lack of difference between the gill cAAT and mAAT with regard to inhibition by AOA (Figure 1) means that experiments reported pre­ viously (Bishop et al., 1981; Greenwalt and Bishop, 1980) on the effect of AOA on changes in aspartate and other amino acid levels during osmotic stress are not compromised with respect to the AAT activity. Although deZwaan et al. (1982) find that high AOA concen­ trations block the changes in aspartate and alanine levels in sea mussel adductor muscle subjected to anaerobic stress, there has been no testing of AOA sensitivity or other characterization of the sea mussel AAT activities.

The lack of inhibition by 10 mM B-chloro-L-alanine is not surprising in that incubations with high inhibitor concentrations

(40-60 mM) for long periods (6-20 min) are required for significant inactivation of the pig heart AAT (Morino and Okamoto, 1973; Silverman and Abeles, 1976; Ueno et al., 1982). In other experiments (data not shown), the alanine aminotransferase and the serine hydroxymethyl- transferase are strongly inhibited by 0.1-1.0 mM

B-chloro-L-alanine.

Although differences in tissue aspartate levels have been ob­

served in a number of bivalves (Bishop et al., 1983; deZwaan, 1977)

under a number of environmental stresses, we cannot at this time

attribute changes in aspartate levels to differences in the relative 28

tissue AAT levels because of the paucity of information on AATs in the other bivalves. However, with anaerobic stress, tissue pH and aspar­ tate levels fall while tissue alanine, opine, succinate, and propion­ ate levels rise (Barrow et al., 1980; Collicutt and Hochachka, 1977;

Ellington, 1981; Kluytmans et al., 1977, 1978; Wijsman, 1975; Zurburg and Ebberink, 1981; Zurburg and Kluytmans, 1980; deZwaan et al.,

1982). Since no "cytosolic succinate dehydrogenase" has been found in

molluscs (deZwaan, 1977), the increase in succinate and propionate

levels during anaerobiosis has been attributed to a shuttling of

carbon from glycolysis and aspartate catabolism into the mitochondria

for conversion to these products (deZwaan et al., 1982). The AATs in these other bivalves are similar to the AATs in ribbed mussels, and given the kinetic behavior of the cAAT with pH change (Figure 5,

6a,b), one would predict that the cAAT with a high for aspartate

would be mainly involved in aspartate synthesis and would be very much

less active with the low tissue pH associated with the anaerobic

state. On the other hand, the mAAT with a low K for aspartate and m high for glutamate probably functions in aspartate catabolism and

would be fully active even with the low tissue pH of anaerobiosis.

Therefore, one would predict that aspartate levels fall during

anaerobiosis because the more anabolic cAAT is less active while the

more catabolic mAAT retains full activity. If this model were to

operate as supposed, then one would predict that the alanine

aminotransferase activity should be high in these mitochondria.

Studies have shown that the alanine aminotransferase of ribbed mussel 29

gill tissue is mainly mitochondrial (Part II).

This model for aspartate metabolism requires a shuttling of aspartate between the cytosol and mitochondria. Shuttles for inter­ mediate metabolites between the cytosolic and mitochondrial compart­

ments in molluscan tissues are not well understood. It would appear that a malate or dicarboxylic acid shuttle may function in some

instances in blue mussel tissues (deZwaan et al., 1981) and the

aspartate-glutamate, or aspartate-succinate or malate shuttle systems

have been suggested for bivalve tissues on the basis of indirect

measurements (deZwaan, 1977; Hochachka, 1981). The exchange system

would be similar to that suggested for the fish liver mitochondria-

cytosolic system (Casey et al., 1983; Van Waarde et al., 1982).

A similar system may apply during hyperosmotic stress under

aerobic conditions and account for the initial decline in aspartate

levels and some of the increase in alanine levels (Baginski and

Pierce, 1975, 1977, 1978; Bishop et al., 1981; DuPaul and Webb, 1971;

Greenwalt and Bishop, 1980; Livingstone et al., 1979; Shumway et al.,

1977). The differences in AAT levels and distributions of the cAAT vs

mAAT activities in the various tissues (Table 2) may account for some

of the tissue differences in aspartate, alanine and organic acid me­

tabolism in mussels subjected to anaerobic stress (Zurburg and

Ebberink,' 1981) and osmotic stress (Baginski and Pierce, 1977). 30

PART II. SUBCELLULAR LOCATION AND CHARACTERIZATION OF ALANINE AMINOTRANSFERASE FROM RIBBED MUSSEL GILL TISSUE

Differential centrifugation of ribbed mussel gill tissue homo- genates and extraction of the mitochondrial fraction demonstrated that

most (72%) alanine aminotransferase (AlAT) activity was mitochondrial.

Subsequent characterization of the cytosolic activity demonstrated

properties identical to those demonstrated by the mitochondrial en­ zyme. Both enzyme fractions showed little variation in with

pH, had low K^s for keto acid substrates, and were inhibited by aminooxyacetic acid (AOA), L-cycloserine, and 8-chloro-L-alanine.

It appears that the AlAT in ribbed mussel gill tissue is strictly

mitochondrial and that alanine production during hypoxia or

hyperosmotic stress must be mitochondrial. 31

INTRODUCTION

Amino acids, particularly alanine and glycine, comprise a sub­ stantial portion of the osmotically active constituents within the

cells of osmoconforming euryhaline bivalves (Lange, 1972; Bishop

1976). In the last few years, the processes regulating the cellular

concentrations of these amino acids have focused on aspects of the

membrane permeability that result in cellular retention of these amino

acids (see Pierce, 1982) and on the metabolic processes that determine

which amino acids accumulate (see Bishop et al., 1983). Early studies

by Read (1962) on the aminotransferases in tissues of some marine

bivalves have been followed by a number of studies suggesting some

relationship between the accumulation of some of the end-products of

anaerobic metabolism (alanine) and the overall regulation of metab­

olite flow during the reduction in metabolic rate that occurs with

hyperosmotic stress (Baginski and Pierce, 1975, 1977, 1978; Bishop et

al., 1981; Henry and Mangum, 1981a,b; Henry et al., 1981). The spe­

cific requirement for active transaminase activities in the alanine

and proline accumulation process was demonstrated in a series of

experiments employing transaminase inhibitors with ribbed mussel

tissues subjected to hyperosmotic stress (Greenwalt and Bishop, 1980;

Bishop et al., 1981).

The specific properties of the transaminase activities in mollus-

can tissues have received very little attention. All tissues of all

molluscs assayed have both the alanine aminotransferase (AlAT) and

aspartate aminotransferase (AAT) activities (Bishop et al., 1983). 32

Other studies (Part I) indicate that ribbed mussel tissues have both cytosolic and mitochondrial AAT isozymes and that the activities of these two isozymes differ considerably in terms of kinetic properties, heat stability, and electrophoretic migration. The cytosolic AAT had unusually high K^s for the amino acid substrates, particularly for aspartate at low pHs (150 mM at pH 6.5). The mitochondrial AAT showed reasonably low K^s for all substrates except glutamate throughout the pH range (pH 6.5-9.5); these values were in the range generally reported for AAT activities from mammalian tissues.

In light of the results with the AAT activities, it was important to determine the cellular distribution of the AlAT activities in ribbed mussel tissues and to compare similarities or differences among

isozymes that might be important in the regulation of amino acid accumulation during hyperosmotic stress. Isoenzymes of AlAT have been

detected in the cytosol and mitochondria of most mammalian tissues

(Hopper and Segal, 1962; Swick et al., 1965; DeRosa and Swick, 1975;

Ruscak et al., 1982). Recently, Burton and Feldman (1983) have shown

that marine copepods (Tiqriopus califomicus) homozygous for the AlAT-

allozyrae with the highest specific activity, accumulate alanine more

rapidly during hyperosmotic stress than either the heterozygotes or

those homozygous for the AlAT allozyme with the lower specific

activity.

Although AlAT activities have been found in both the cytosol and

mitochondria of tissue homogenates of some pulmonate snails by Sollock

et al. (1979) and in whole body homogenates of oysters (Crassostrea 33

virqinica) by Chambers et al. (1975), there have been no studies on the properties of the AlAT activities and whether or not the cytosolic and mitochondrial activities are associated with different isozymes.

In oysters, Chambers et al. (1975) found that the ratio of the specif­

ic activity of the mitochondrial to cytosolic AlAT activities varied

with the season from a low from November through July (0.49-0.59) to a

high in September and October (2.9). Although the reasons for these

activity variations are unclear, they may be related to yearly salin­

ity shifts, feeding, or possible reproductive cycles. For instance,

Wickes and Morgan (1976) have reported higher tissue AAT activities in

oysters collected at high salinity compared to low salinity environ­

ments. On the other hand, in a controlled study, DuPaul and Webb

(1974) found no tissue activity differences for AAT and AlAT in a

number of bivalves subjected to short term hyperosmotic stress. As an

additional complicating factor, Hammen (1968) has shown that in a

number of marine bivalves, the tissue activities of the transaminases

vary with the size of the animal.

This part reports the partial purification and some properties of

gill AlAT activities. In this gill tissue most of the AlAT is bound

within the mitochondrial compartment. 34

MATERIALS AND METHODS

Ribbed mussels (Modiolus demissus), purchased from Northeast

Environmental Laboratories (Monument Beach, Mass.), were kept in artifical sea water (Jungle Laboratories Inc., Sanford, Fla.) and maintained as described by Greenwalt and Bishop (1980). Except where noted, all reagents and coupling enzymes were purchased from Sigma

Chemical Co., St. Louis, MO. CTAB (hexadecyltrimethylammonium bromide) and ammonium sulfate (enzyme grade) were obtained from

Eastman Organic Chemicals, (Rochester, N.Y.) and Schwartz-Mann

(Orangeburg, N.Y.), respectively. B-chloro-L-alanine and vinylglycine were purchased from Calbiochem (La Jolla, CA)•

The digitonin used in these experiments was recrystallized ac­ cording to Kun et al. (1979). Digitonin (Sigma grade) was dissolved in absolute ethanol (1 g/25 ml) at 75°C then precipitated by chilling the solution on ice for 20 minutes and collected by centrifugation at

4°C. This procedure was repeated and the precipitate dried in a desiccator.

Enzyme assay

During purification and standard assay, the AlAT activity was determined spectrophotometrically by measuring pyruvate production as

NADH oxidation (340 nm) using a Beckman 3600 recording spectropho­ tometer in a reaction mixture containing 20 mM alanine, 10 mM

a-keto- glutarate, 70 wM NADH, 5 units of lactic dehydrogenase

(LDH), 50 mM Tris/HCl (pH 8.3), and enzyme in 2 ml. The reverse 35

(alanine forming) direction was measured by coupling the transaminase

reaction to a-ketoglutarate dehydrogenase and measuring the

reduction of NAD (340 nm) in a reaction mixture containing 20 mM

glutamate, O.SmM pyruvate, 70 jiM NAD, 0.5 mM CcA, 0.5 units

a-ketoglutarate dehydrogenase, 50 mM Tris/HCl, and enzyme in 1 ml

at pH 8.3. Control assays without gill extract demonstrated the

absence (< 1%) of any contaminating transaminase activity in either

the LDH or thé a-ketoglutarate dehydrogenase. Glutamate

dehydrogenase (GDH) was assayed in the glutamate forming direction by

the procedure of Reiss et al. (1977) with 1 mM ADP. Incubations were

at room temperature (23°C). One unit of activity was equal to one

umole of product produced per minute under the conditions

specified. Procedural modifications for kinetic experiments are

described below. Kinetic constants were determined using computer

assisted analysis of initial rates (Cleland, 1979) using least squares

analyses for the best linear fit.

Isozyme determination

Isozyme distribution and purity of electromorphs was determined

using starch gel electrophoresis. One liter of electrode buffer

contained 15.5 g Tris (Base), 1.5 g EDTA, 1.15 g boric acid, and

glacial acetic acid to pH 7.0. Electrode buffer was diluted 1:10 to

make the gel buffer. To examine tissue distribution, samples of whole

tissues were homogenized in an equal volume of gel buffer using a

Polytron (Brinkman Inst., Westbury, N.Y.) tissue homogenizer with a

small probe. The suspension was applied to individual slots in 36

thirteen percent (w/v) horizontal starch gels (12 cm x 20 cm x 0.8 cm) and electrophoresis performed at approximately 35 ma at 4°c.

Progress was marked with bromphenol blue. Gels were sliced and stained according to the alanine aminotransferase detection procedure of Harris and Hopkinson (1976) using a filter paper overlay and a hand-held ultra-violet light to scan, for NADH oxidized (NAD).

Cytosolic and mitochondrial fractions of gill homogenates were prepared using the previously described differential centrifugation procedure (Part I). Protein was determined by a modified Lowry procedure (Miller, 1959). 37

RESULTS

Isozyme distribution

Starch gel electrophoresis of samples from cytosol and unbroken mitochondria [suspended in 10 mM phosphate (pH 6.8)] indicated elec- tromorphs in both the cytosolic and mitochondrial compartments (Figure

1, lanes a,b). However, it appeared that more than half of the raAlAT activity remained at the origin in the wells (Figure 1, lanes a). It appeared that a major fraction of the mAlAT was not released from the mitochondria by suspension of the mitochondria in dilute buffer (10 mM). However the mAlAT activity that was released by this treatment had the same electrophoretic mobility as the major cytosolic AlAT suggesting that almost all of the gill tissues AlAT activity was localized within the mitochondria and that this "cytosolic" activity

was released during preparation of the mitochondria. With prolonged

staining of the electrophoretic gels (12 hrs.), a second faintly

staining, anodally migrating "AlAT electromorph" was detected in the

cytosolic fraction.

It was now critical to extract the remaining AlAT activity from

the gill mitochondrial and compare the properties of the mAlAT to

those of the "cytosolic AlAT". Freezing and thawing of the mito­

chondria as described previously for the release of the mitochondrial

aspartate aminotransferase (Part I) liberated very little additional

soluble mAlAT activity from the mitochondria. Treatment of 1 ml of

suspended mitochondria (3.8 rag protein/ml buffer) with 9 ml of 0.1%

CTAB released little or no itiAlAT and apparently inactivated the AlAT. 38

+ +

abc d e : B 1 1 B B B >

Figure 1. Distribution of AlAT gill tissue isozymes using starch gel electrophoresis. Four lanes of identical samples were run for each preparation. Mitochondrial (lanes a) and cyto- solic (lanes b) fractions from gill tissue were applied to wells and electrophoresed at pH 7.0. Mitochondrial digi- tonin extract was applied in lanes c. Cytosolic salt fraction 1 was applied in lanes e. See text and Table 1 for definition of procedures of electrophoresis and frac­ tions assayed 39

Treatment with 1 ml of a 0.1% CTAB solution released about 50% of the mAlAT activity with a loss of 50% of total activity. Treatment of these mitochondrial suspensions with 1%, 0.5% and 0.25% Triton X-114 at 23°C for 30 min did not cause a release of additional mAlAT activity and caused loss of total activity. Treatment of the mito­ chondrial with digitonin released most of the mAlAT activity and did not appear to inhibit the AlAT activity (Figure 1, lanes c).

A comparison of the release of AlAT and GDH from these gill mitochondria in a series of digitonin treatment experiments was made in order to determine the relative efficiency of extraction and the relative tightness of binding within the mitochondria. Results of the digitonin extraction are in Table 1 (extraction procedure outlined in footnote). Two successive digitonin extractions liberated 94% of the total mitochondrial AlAT and 86% of the total mitochondrial GDH.

Total tissue activities were 772 mUnits/gm wet wt. (AlAT) and 228 raUnits/gm wet wt. (GDH). The major cAlAT activity released during the

initial tissue homogenization and mitochondrial preparation procedure

had the same electrophoretic migration as the AlAT released from the

mitochondria with digitonin treatment; this activity was termed the

cytosolic mitochondrial AlAT or cmAlAT. In a series of experiments

with gill tissue from 30 individual animals, there was no variation in

this single mAlAT-cmAlAT isozyme pattern. 40

Partial purification of the gill AlAT activities

During typical mAlAT purification, mitochondria from approxi­ mately 20 g gill tissue were resuspended in 20 ml of 2.5 mg/ml newly recrystallized digitonin suspension in 10 mM potassium phosphate

Table 1. Intracellular distribution of alanine aminotransferase and glutamate dehydrogenase in ribbed mussel gill tissues^

Gill Tissue Fraction % Activity in Fraction Mitochondrial Protein AlAT GDH (mg/ml)

/

Cytosol (Total) 27 8.3 -

Salt Fraction 1 17 8.0 -

Salt Fraction 2 10 0.3 -

Mitochondria Total Extracted 73 91.7

(unextracted apparent) (32) (43.5) (2.5)

Extraction 1 32 43.5 1.4

Elxtraction 2 37 37.4 0.7

Extraction 3 4 11.1 0.3

Mitochondrial and cytosolic fractions were prepared as described in the text. The mitochondria were suspended in 20 ml of 10 mM potassium phosphate buffer and assayed (unextracted mitochon­ dria). This suspension was then treated with 25 rag digitonin (.5 mg digitonin/ mg protein) with constant stirring for 10 min on ice. The suspension was centrifuged (15,000 x g? 20 min) and the supernatant fluid assayed as Extraction 1. The pellet was resuspended in 20 ml of a digitonin solution (2.5 mg/ml at Img digitonin/mg protein), stirred for 30 min on ice, centrifuged and the supernatant assayed as Extraction 2. The residue was again resuspended in the same digi­ tonin solution (2.5mg/ ml) then stirred on ice for 30 min, centri­ fuged and the supernatant fluid assayed as Extraction 3. Assays are described in Materials and Methods. 41

buffer (pH 6.8) and stirred for at least 30 minutes at 0°C in a beaker with a magnetic stirring bar. The resulting suspension was centrifuged (10,000 g for 20 minutes) and the supernatant fluid containing most of the mAlAT activity (174 mUnits/mg protein) was dialyzed against 10 mM potassium phosphate buffer (pH 5.8) overnight at 4°C. Hydroxyl-apatite (HAP) (Bio-Rad Laboratories, Richmond, CA) was washed in degassed 10 mM potassium phosphate (pH 6.8), the fine particles were removed, and the slurry was poured into a column to give a 2 X 10 cm bed. After equilibration of this column with degassed dialysate buffer (see above), the preparation from the dialysis bag was added onto the column and the column was washed with

150 ml of degassed dialysate. Most of the protein was eluted from the column by washing with 150 ml of 50 mM potassium phosphate (pH 6.8); the mAlAT activity remained on the column. The mAlAT activity was then eluted as a single peak of activity by washing the column with

100 ml of 200 mM potassium phosphate (pH 6.8). The fractions (1 ml)

with the highest specific AlAT activity (520 mUnits/mg protein)

eluting behind the buffer front were pooled. These pooled fractions

constituted the final partially purified preparation of mAlAT activity

used for the kinetic experiments.

Approximately 20% of the total GDH activity extracted from the

mitochondria by the digitonin treatment was eluted from the HAP column

with the mAlAT activity (200 mM buffer wash). There was no (<1%) NAD

reduction associated with the mAlAT activity in the presence of only

glutamate, NAD, and enzyme (homogenate or enzyme preparation). GDH 42

-sr 100 +

4-" •> S 50

fC)

Figure 2. Thermal stability of AlAT found in the cytosol fraction (0) and extracted from the mitochondria (+)» Approxi­ mately 150 ul of enzyme was incubated in a water bath for 2 minutes at each temperature. One hundred microliters was added to a 1.9 ml reaction cocktail at 23°C and assayed as described in Materials and Methods for pyruvate formation 43

100 >1 4-' >

0 6 10 pH

Figure 3. Relative velocity of mAlAT at different pH values. Activity was assayed in the pyruvate-forming direction using the standard assay substrate concentrations (see Materials and Methods) with 50 mM potassium phosphate instead of Tris 44

activity was dependent on the presence of ADP in the reaction cocktail

in that there was a 5-fold loss of activity without 1 mM ADP.

The AlAT activity in the cytosolic fraction (Table 1) was par­

tially purified. Addition of 42 mg/100 ml anunonium sulfate to the

cytosolic fraction precipitated most (75%) of the AlAT (cytosolic salt

fraction 1). Addition of an additional 14 gm/100 ml ammonium sulfate

precipitated cytosolic salt fraction 2 of the AlAT (5% of the total

tissue activity). With starch gel electrophoresis, the AlAT in

cytosolic salt fraction 1 migrated as the mAlAT (cmAlAT) and the AlAT

in cytosolic salt fraction 2 contained approximately equal amounts of

the cmAlAT and the anodally migrating, slow-staining AlAT activity

described previously (Fig. ld,e). The cytosolic salt fraction 1 was

dialyzed and chromatographed on hydroxyl-apatite using the procedure

described for the mAlAT activity. This cmAlAT activity eluted with

the 200 mM phosphate (pH 6.8) buffer wash in a manner identical to the

mAlAT activity and had the same electrophoretic mobility as the mAlAT.

Properties of the mAlAT

The partially purified AlAT activity was characterized with res­

pect to substrate binding, heat stability, and reactivity with inhi­

bitors.

The heat stabilities of the cmAlAT and mAlAT were identical (Fig.

2). The activity in both fractions was lost at temperatures above

50°C. This heat stability was slightly lower than the heat stability

reported by Bulos and Handler ('65) for the beef heart AlAT activity.

The gill AlAT was much more heat sensitive than either the gill mAAT 45

or cAAT (see Part I) and apparently not as labile as the mammalian mitochondrial enzyme (Swick et al., 1965).

Kinetic analysis of the partially purifed mAlAT showed optimal activity over a broad pH range from pH 5.6 to pH 9.5 (Fig. 3). Sub­ strate binding constants (apparent K^) were determined in the for­ ward (pyruvate forming) and reverse (alanine forming) directions (data not shown). The apparent values and relative maximal vélocités did not change markedly between pH 6.4 and pH 9.5 (Table 2). The

K^s for substrate binding and ratios for rates of the forward vs the

reverse direction at pH 8.3 with the cmAlAT were essentially identical to these values with the mAlAT (Table 2).

Table 2. Apparent Km values for gill tissue mitochondrial (mAlAT) and mitochondrial-like cytosolic (cmAlAT) alanine amino- tranferases^

Apparent Km (mM)

Ratio Form pH Ala a-kg Pyr Glu F/R

mAlAT 6.4 4.2 0.12 0.01 3.7 5.47

mAlAT 8.3 4.3 0.11 0.05 5.8 5.61

mAlAT 9.5 2.3 0.12 0.13 2.7 3.45

cmAlAT 8.3 5.5 0.08 0.07 5.0 5.51

kinetic constants were generated by computer analysis (Cleland, 1979). F/R represents the maximal velocities of the forward (F) or pyruvate forming direction to the reverse or alanine forming direction. Assays are described in Materials and Methods. Alternate substrate concentrations for the reactions were: 20 mM alanine and 10 mM a-ketoglutarate for the pyruvate forming reaction, and 20 mM glutamate and 5mM pyruvate for the alanine forming reaction. Standard error for all measurements averaged 11%. 46

AOA inhibited the tnAlAT with an of approximately 7 x 10~® M and L-cycloserine inhibited the mAlAT activity with an I^q of 0.18 mM

(Fig. 4). These values were in general agreement with those reported by

Greenwalt and Bishop (1980) for the AlAT in ribbed mussel tissue homogenates.

L-serine-O-sulfate (10 mM) and vinylglycine (10 mM) did not inhibit enzyme activity under the conditions described in Figure 4.

Beta-chloro-L-alanine (g-CA) inhibited the mAlAT in a time- concentration-dependent fashion (Fig 5). The apparent (6-CA) generated from the replot in Figure 5 was 25 uM, which was in general agreement with the value of 75 yM found for the pig heart

AlAT (Golichowski and Jenkins, 1978). Using the standard assay

(pyruvate formation), there were no apparent inhibitory or stimulatory effects on the observed velocity by NaCl, KCl, glycine, or succinate

(up to 250 mM); taurine was a poor noncompetitive inhibitor (K^ 290

mM; data not shown). These results agree with those reported by

Gilles (1969) on the effects of various salts on aminotransferases in

some crustacean tissues. 47

M•> 50

Figure 4. Inhibition of the gill tissue mAlAT and cmAlAT activities by aminooxyacetic acid (-o-o-) and L-cycloserine (-0-D-) • Enzyme (125 jil) was incubated for 2 minutes at 23 with 125 wl of inhibitor. Two hundred microliters of the enzyme-inhibitor mixture were then added to a 1.8 ml reaction cocktail and assayed as described in Materials and Methods. 48

BMI VVSQDI 10

v\o5 Vfelocity Ol

It 0.6 1 l - CO ^ y/ V 0 1 2 a TlmeûTiin) o

0.2 -

.1 y 1 1 - -J

p-Cl-AlaOnM)

Figure 5. Inhibition of the gill tissue raAlAT activity by 6-chloro-alanine (B-Cl-Ala) and assayed (lOOul) after 1 minute and 2 minutes (see inset). Initial rates of inhibition (I-r,) were determined from the change in activity during ène' first minute of incubation (see inset). The I^^ values were replotted as a function of g-Cl-Ala concentration 49

DISCUSSION

The results indicate that most, if not all, of the alanine aminotransferase in ribbed mussel gill tissue is located in the mitochondria. Although the AlAT activity is found in both the cytosolic and mitochondrial fractions separated from homogenates by differential centrifugation, both the major "cytosolic" form and the mitochondrial form show the same electrophoretic, kinetic and heat stability properties, suggesting that this "cytosolic AlAT" is iden­ tical to the mitochondrial AlAT and may be released from the mito­ chondria due to mitochondrial breakage during tissue preparation. In this regard, more than half of the mAlAT activity and most of the

mitochondrial glutamate dehydrogenase activity are cryptic or tightly

bound within the mitochondria and only released with successive digi- tonin (detergent) extractions. Although preliminary studies with sea

mussel tissues (Addink and Veenhof, 1975) indicated that release of

the mAlAT required digitonin treatment of these mitochondria, these

authors make no comment on a possible cytosolic or "soluble" AlAT

activity.

The same homogenization-differential centrifugation procedure was

used as in Part I to separate mitochondrial and cytosolic aspartate

aminotrasferase (AAT) activities. Very little mitochondrial breakage

as evidenced by cross AAT isozyme contamination in the cytosolic and

mitochondrial fractions was apparent, even though the mAAT is appar­

ently not membrane bound and is easily released by a freeze/thaw

treatment. The release of a small amount of otherwise tightly bound 50

mAlAT and GDH during the homogenization-centrifugation procedure may indicate a binding mode within the mitochondria that differs consid­ erably from the mAAT activity. At this point, there is no reason to postulate that in vivo some of the gill tissue mAlAT and GDH are cytosolic as has been suggested for the sea mussel GDH (deZwaan,

1977).

In other studies by Chambers et al. (1975) with oyster tissue and by Sollock et al. (1979) with hepatopancreas tissue from a number of land snails, both groups report both cytosolic and mitochondrial AlAT activities in most species but neither used digitonin extraction treatment with isolated mitochondria and neither determined the

"isozyme" distribution. Sollock et al. (1975) subjected the snail

mitochondria to sonic disruption rather than detergent extraction

before assay. The presence of isozymic AlAT forms in homogenates of

hepatopancreas tissue from two fresh water bivalves, Anodonta

couperiana and Popenaias bucklevi is suggested from the ammonium

sulfate precipitation experiments of Falany and Friedl (1981). In

contrast to the results presented here for mussels, preliminary

results with oyster (Crassostrea virqinica) tissues (Burcham et al.,

1984; Ellis et al., 1984) indicate separate AlAT isozymes in the

cytosolic and mitochondrial compartments. In conclusion, it would

appear that a combination of isozyme analysis and differential

isolation of these cellular compartments followed by specific

extraction to solubilize "cryptic activities" may be required to

resolve differences in the reported relative amounts of the AlAT 51

activities in the tissues of molluscan species (Bishop et al., 1983).

The small amount (5% of total AlAT) of anodal migrating, faintly staining cytqsolic AlAT could be concentrated in the 60-80% ammonium sulfate fraction (Table 1) and resolved with a greater degree of con­ fidence as an AlAT activity (Figure 1, lanes e). Attempts to purify this activity using the hydroxyl-apatite column have been unsuccess­ ful. Preliminary results on the AlAT isozyme distribution in the other tissues of the ribbed mussel indicate that all had the same mAlAT isozyme as the gill tissue but it is not at this time certain that all of this activity is located within the mitochondria of these tissues. Most mammalian tissues show separate AlAT isozymes for both the mitochondrial and cytosolic compartments (Hopper and Segal, 1962;

Swick et al., 1965; DeRosa and Swick, 1975; Ruscak et al., 1982). On the other hand chicken heart, liver and kidney apparently lack the cAlAT (DeRosa and Swick, 1975). Burton and Feldman (1982, 1983) report fast and slow migrating "soluble" AlAT alleles in hétérozy­ gotes and fast or slow migrating AlAT alleles in respective homozy­ gotes of marine copepod (T^ califomicus) populations. The method used for preparation of the copepod homogenates involves a buffered sucrose solution of 600-650 mosmoles (Burton and Feldman, 1981) which may not break the mitochondria. These results suggest that these

copepod AlAT alleles may be cytosolic rather than mitochondrial; there

is no mention of possible mAlAT positive staining material remaining

in the wells at the origin after electrophoresis.

For the most part, the apparent K^s for the gill AlAT are 52

somewhat lower than those reported for rat and beef heart and liver cAlATs (Hopper and Segal, 1962; Swick et al., 1965; Bulos and Handler,

1965; DeRosa and Swick, 1975). The variation in activity with pH

(Figure 3) is very similar to that found by Swick et al. (1965) and

Orlicky and Ruscak (1976) for the rat liver and heart raAlATs. The small variation with pH in apparent substrate K^s with the amino acid substrates with the gill mAlAT is similar to that found with the gill mAAT activity (Part I). The decrease in apparent for pyruvate with decreasing pH (Table 2) may be of particular physio­ logical importance with the lowered intracellular pH that occurs during short term anaerobiosis (Ellington, 1983).

The mAlAT is freely reversible. The ratio of rates with the gill

AlAT in forward (pyruvate forming) vs the reverse (alanine forming) direction (Table 2) is similar to the ratio of 4.4 (in Tris) reported for the rat liver mAlAT (Swick et al., 1965). Awapara and Campbell

(1964) using homogenates of clam (Rangia) and oyster (Crassostrea)

mantle and snail (Qtala) hepatopancreas at fixed low substrate con­

centrations report lower forward-reverse rate ratios of 1.8, 1.1, and

0.88, respectively.

Aminooxyacetic acid (AOA), L-cycloserine and 6-chloro-L-

alanine but not vinylglycine or serine-o-sulfate inhibit the gill

mAlAT. AOA and L-cycloserine inhibition (I^q concentrations) are •

similar to those found for other AlATs (see Greenwalt and Bishop,

1980; Hopper and Segal, 1962). AOA and L-cycloserine but not

S-chloro-L-alanine are effective inhibitors of the gill AAT 53

activities (Part I).

Previous work with ribbed mussel gill tissue (Part I) has shown that the kinetic properties of the cAAT and mAAT indicate that aspar­ tate synthesis should be favored in the cytosol and that aspartate catabolism should be favored in the mitochondria. Aspartate metab­ olism seems to be linked to alanine or alanine-glycine metabolite

(strombine, alanopine) accumulation in most bivalve tissues when the animals or tissues are subjected to hypoxic or hyperosmotic stress

(Collicutt and Hochachka, 1977; deZwaan, 1977; Baginski and Pierce,

1978; Greenwalt and Bishop, 1980; Bishop et al., 1981; deZwaan et al.,

1982, 1983). In mussels, alanine metabolism (turnover) must be cen­ tered within the mitochondria because the major route of turnover has

been shown to be through an AlAT linked pathway (Greenwalt and Bishop,

1980; Bishop et al., 1981; Greenwalt, 1981; deZwaan et al., 1982,

1983a,b; this study) and most of the mussel tissue AlAT is mitochon­

drial. During hyperosmotic or hypoxic stress, gill tissue alanine

accumulation would require an influx of both amino groups (carried as

aspartate or other amino acids) and carbon skeletons for pyruvate

production (glycolytic products, gluconeogenic precursors) for subse­

quent transamination to alanine. The alanine would then exit the

mitochondria for accumulation in the cytosol. This model suggests a

"tight" regulation of pyruvate metabolism within the mitochondrion and

possibly controlled mitochondrial amino acid transporters. In this

regard it is of considerable interest that pyruvate is a very poor

substrate for oxidative metabolism by bivalve gill tissue mitochondria 54

showing high degrees of respiratory control (Burcham et al., 1984). A similar model linking aspartate and alanine metabolism within the mitochondrion has been proposed for fish liver by Van Waarde et al.

(1982). The pyruvate dehydrogenase complex (PDC) plays a key regula­ tory role in pyruvate metabolism in mammalian tissues. One would predict then that PDC in ribbed mussel mitochondria could play a major role in control of metabolism during hyperosmotic or anaerobic stress. 55

PART III. SUBCELLULAR DISTRIBUTION OF AMINOTRANSFERASES, AND PYRUVATE BRANCH POINT ENZYMES IN GILL TISSUE FROM FOUR BIVALVES

Aspartate aminotransferase (AAT), alanine aminotransferase

(AlAT), malic enzyme (ME), malate dehydrogenase (MDH), pyruvate kinase

(PK), and phosphoenolpyruvate carboxykinase (PEPCK) activities in cytosolic and mitochondrial fractions of gill tissue from Modiolus demissus (ribbed mussel), Mvtilus edulis (sea mussel), Crassostrea virqinica (oyster) and Mercenaria mercenaria (quahog) were determined using enzyme assay and starch gel electrophoresis combined with subcellular fractionation. AAT showed distinct mitochondrial and cytosolic isozymes in gills of all these animals. Although AlAT showed distinct mitochondrial and cytosolic isozymes in the gills of oysters, sea mussels and quahogs, only the mitochondrial AlAT was

evident in ribbed mussel gill tissue. PK and PEPCK were cytosolic in all these preparations. ME was found only in the mitochondrial

fraction of ribbed mussel and quahog gill tissue whereas sea mussel

gills showed distinct cytosolic and mitochondrial ME isozymes. With

oyster gills, the "cytosolic ME" was electrophoretically identical to

the mitochondrial ME indicating that in vivo, the ME is probably

mitochondrial. MDH showed distinct cytosolic and mitochondrial I isozymes in all bivalve gills tested. 56

INTRODUCTION

The subcellular distribution and properties of isozymes associated with the metabolism of the amino acids and the organic acids of the TCA cycle seem important to the metabolic processes regulating the high cellular free amino acid concentrations in tissues from ribbed mussels and other euryhaline bivalves. For instance, the levels of alanine, glycine, proline, and glutamate in ribbed mussel gill tissue appear to be controlled by specific enzymes within the mitochondria. The alanine aminotransferase (AlAT) and the enzymes regulating the catabolism of glutamate (Reiss et al., 1977; Hoffmann et al., 1978), proline and pyruvate (Burcham et al., 1984; Bishop et al., 1981) and glycine (Elllis et al., 1985) are mitochondrial. On the other hand, the lactate and opine dehydrogenases are cytosolic (Bishop et al., 1980; Nicchitta and Ellington, 1984). Part I showed that the kinetic behavior of the cytosolic aspartate aminotransferase (AAT) activity differs considerably from that of the mitochondrial AAT activity.

An essential aspect of the studies from this laboratory is the

combining of subcellular fractionation studies to establish the iden­ tity, heterogeneity within the compartment of origin, the relative

activity and the physiological roles of these isozymic forms. This

part compares the subcellular distribution of six enzyme activities,

malic enzyme (ME), malate dehydrogenase (MDH), pyruvate kinase (PK),

phosphoenolpyruvate carboxykinase (PEPCK), AAT and AlAT in ribbed 57

mussel gill tissues to that in the gill tissue of three other eury- haline bivalves often used in the study of cellular osmoregulation. 58

MATERIALS AND METHODS

Ribbed mussels (Modiolus demissus), purchased from Northeast

Environmental Laboratories (Monument Beach, MA), were kept in artificial sea water (Jungle Laboratories, Inc., Sanford, FL) and maintained as described by Greenwalt and Bishop (1980). Quahogs

(Mercenaria mercenaria) and oysters (Crassostrea virqinica) were obtained from Dr. Michael Castagna (Virginia Marine Institute of

Science, Wachapreague, VA) and kept in the same tank with the ribbed mussels. Sea mussels (Mytilus edulis) were obtained from the New

England Coast through a local seafood supplier and maintained in artificial sea water (850 mOsmols) at approximately 10°C. Except when noted, all reagents were purchased from Sigma Chemical co., St.

Louis, MO, EInzyme grade ammonium sulfate was obtained from

Schwartz-Mann (Orangeburg, NY).

Tissue fractionation

Gills were removed from the bivalves and collected in chilled tared beakers. Approximately 20g of tissue was typically collected.

The tissue was homogenized in a mitochondrial isolation buffer con­ sisting of 0.4M sucrose, 20mM HEPES and ImM EGTA (pH7.5) using an

Ultra-Turrax homogenizer (Tekmar Co., Cincinnati, OH) with three

10-sec bursts on a setting of "40". The homogenates were filtered through Miracloth (Calbiochem) to remove large particulate debris.

The filtrate was centrifuged at 1500g for 8-10 min; the supernatant

collected and recentrifuged at 9000g for 15 min. This procedure has

been used previously (Burcham et al., 1983, 1984; Ellis et al., 1985, 59

Parts I & II). The supernatant from this second centrifugation constituted the cytosolic fraction. The pellet of the second centrifugation was resuspended in mitochondrial isolation buffer (10 ml) and extracted with recrystallized digitonin. Digitonin (sigma grade) was recrystallized according to Kun et al. (1979). The mito­ chondrial suspension was extracted by addition of 1 mg digitonin/mg mitochondrial protein and stirred with a magnetic stirring bar for at least 30 rain on ice. The resulting suspension was centrifuged at

10,000g for 20 min and the supernatant collected as the mitochondrial extract (Part II). Protein was determined using the biuret method

(See Burcham et al., 1984).

Enzyme assays

All enzyme activities were assayed spectrophotometrically by measuring the oxidation/reduction of NADH/NAD as a function of absorption at 340nm. The PK and MAT assays were coupled with LDH, and the PEPCK and AAT assays were coupled with MDH. The six reaction mixtures used for the assays consisted of the following: AAT-100 mM

Tris-HCl (pH 8.0), 20mM aspartate, lOmM a-ketoglutarate, 70yM

NADH, 5 units MDH, and enzyme in 2 ml; AlAT—50mM Tris-HCl (pH 8.0),

20 mM alanine, lOmM a-ketoglutarate, TOuM NADH, 5 units LDH,

and enzyme in 2 ml; PEPCK—50mM Tris-HCl (pH 8.0), 70uM NADH, ImM

MnCl^, l.SmM IDP, ImM MgCl^, 2mM PEP, 20mM NaHCO^, 5 units MDH

and enzyme in 2 ml; ME-ImM Malate, O.lmM NADP, ImM MgClg, 50mM

Tris-HCl (pH 8.0) and enzyme in 2 ml; MDH—lOmM OAA, 70uM NADH,

SOmM Tris-HCl (pHB.O) in 2 ml. All enzyme assays were performed at 60

Starch gel electrophoresis

Starch gels were prepared as described in Part II. Samples of the cytosolic and mitochondrial fractions were loaded into the gel slots and electrophoresed in a 4°C refrigerator at 35mA until the

bromphenol blue marker reached the sponge bridge. The gels were

removed from the refrigerator, sliced in half and immediately stained.

AAT, AlAT, PK, ME, and MDH were stained on individual gel halves using the methods of Harris and Hopkinson (1976). 51

RESULTS

The data for the six enzyme activities in the four bivalve gill tissues are presented in Table 1. The use of the digitonin treatment of the mitochondria appears to have been particularly useful in ob­ taining preparations that showed full activities and no staining at the origin after electrophoresis. The percentages of the total activities in the mitochondrial and cytoslic fractions are presented in parentheses in Table 1. Isozyme patterns are described (Table 1) to indicate the presence or absence of different isozymes between compartments. Fractions exhibiting no electrophoretically common activities were labelled as distinct. Where both fractions exhibited

identical isozyme patterns, the patterns were labelled identical.

Where both fractions exhibited activities which contained both

distinct and identical bands, the patterns were labelled mixed.

Although identical electrophoretic banding patterns may not indicate

identity between the two activities, different banding patterns do

indicate different or modified proteins and hence different isozymes.

The gill tissue AAT activity in pooled samples of tissue from

representatives of the four bivalve species studied exhibited dif­

ferent electromorphs in both cellular compartments of the gills of all

four species. The mitochondrial AAT was the major form in the oyster

gill tissue, whereas the mitochondrial and cytosolic AAT distribution

was more equal in the gill tissue from the other species. Although

the AlAT activity showed distinct mitochondrial and cytosolic forms in

oyster, quahog, and sea mussel gill tissue, only the mitochondrial 62

form was found in the ribbed mussel gill tissue (see Part II). The ME activity showed distinct mitochondrial and cytosolic forms in sea mussel gills; gill tissue from the other bivalves showed the single mitochondrial form.

The MDH activity showed distinct cytosolic and mitochondrial forms in gills from all bivalves tested. The majority of the total

MDH activity was cytosolic in oyster, quahog, and sea mussel gill tissue. The cytosolic and mitochondrial MDHs showed multiple, non- identical electromorphs in gill tissue from all the bivalves tested.

Both the PK and PEPCK activities were cytosolic in the gills from all four bivalves. Table 1. Mitochondrial and cytosolic distribution of enzyme activities in gill tissue from four euryhaline bivalves

Specific activity(percentage of total activity in tissue fraction preparations)'

AAT AlAT ME MDH PK PEPCK

Crassostrea virginica

Mitochondria 1.7(92) 0.10(56) 0.010(49) 0.06(10) 0.008(2) N.D. Cytosol 0.035(8) 0.02(44) 0.002(51) 0.13(90) 0.093(98) 0.11(100) Isozyme pattern Distinct Distinct Identical Mixed Identical

Mercenaria mercenaria

Mitochondria 0.16(43) 0.053(80) .018(100) 0.50(16) 0.09(6) N.D. Cytosol 0.19(57) 0.001(20) N.D. 0.13(84) 0.12(94) 0.004(100) Isozyme pattern Distinct Distinct Distinct Identical

Mvtilus edulis

Mitochondria 0.086(25) 0.034(44) 0.073(76) 0.034(15) 0.084(18) N.D. Cytosol 0.088(75) 0.015(56) 0.080(24) 0.068(85) 0.10(82) 0.065(100) Isozyme pattern Distinct Distinct Distinct Mixed Identical

Modiolus demissus

Mitochondria 0.18(56) 0.10(77) 0.02(100) 0.14(30) 0.031(8) N.D. Cytosol 0.030(44) 0.006(23) N.D. 0.068(70) 0.075(92) 0.007(100) Isozyme pattern Distinct Identical Mixed Identical

^Specific activity in ymol/min/mg protein. Standard error of all determinations was less than 10%. ^Activity below detectable limits. 64

DISCUSSION

AAT activity has been detected in the cytosol and mitochondria of all bivalve species investigated by a number of investigators (see

Bishop et al., 1983; Burcham et al., 1983; Part I). The general distribution of AAT activity in sea mussel, ribbed mussel and quahog gill tissue (Table 1) would seem to fit the pattern seen in most other bivalves. However, the high proportion of mitochondrial AAT compared to cytosolic AAT in oyster gill tissue (Table 1) may be of some importance in terms of population genetics. For instance, Buroker

(1983, 1984) reports two AAT isozyme groups in American oyster populations from the Chesapeake Bay area where these animals were collected. The slower moving isozyme is apparently the mitochondrial form (Table 1) and is essentially monomorphic. The other faster moving cytosolic form can show considerable polymorphism. The relatively low tissue level of the cytosolic AAT isozyme compared to the mitochondrial AAT isozyme may account for some of the difficulties in determining polymorphism for these isozymes in some bivalve tissues

(see Fujio et al., 1983; Kartavtsev and Zaslavskaya, 1983; Johnson and

Utter, 1973; Johnson et al., 1972).

The finding of distinct cytosolic and mitochondrial ME isozymes

in Mi edulis tissues supports other studies on ME in sea mussel tissues (deZwaan and van Marrewijk, 1973; deZwaan, 1977; deZwaan et al., 1981; Fujio et al., 1983; Blanc, 1983). A single monomorphic ME

activity has been reported for the giant clam Tridacna by Ayala et al. 65

(1973). However, Fujio et al. (1983) report both polymorphic and

monomorphic forms of ME in the oyster qiaas and some other bi­

valves. Although the "cytosolic ME" activity from qiqas adductor

muscle has been partially characterized by Hochachka and Mustafa

(1973), there are no studies on the mitochondrial MEs from the tissue

of these other bivalves. The appearance of the mitochondrial ME form

in the cytosolic fraction of the oyster gill preparations (Table 1)

probably indicates a loose binding within the mitochondria and release

to the cytosol during the homogenization. The ME in the other bivalve

gill tissues is apparently more tightly bound within the mitochondria.

The AlAT activity is usually found in both the cytosol and mito­

chondria of bivalve tissues (Chambers et al, 1975; Burcham et al,

1983; deZwann, 1977; Addink and Veenhof, 1975; Bishop et al., 1983).

With oyster, sea mussel and quahog gill tissues these two AlAT activ­

ities are associated with separate isozyme groups. Ribbed mussel gill

tissue lacks the cytosolic AlAT activity (Part II).

The cytosolic localization of PK and PEPCK is common to most bi­

valves (deZwaan, 1977). Although no clear indication of polymorphism

for PK was observed in this study, cytosolic isozymes of PK have been

shown in different sea mussel tissues and some variations in electro-

phoretic migration may occur with phosphorylation or dephosphorylation

(Siebenaller, 1980; Holwerda et al., 1983).

A complex MDH isozyme distribution pattern is found in most

bivalve tissues (Ayala et al, 1973; Tracey et al., 1975; Koehn and

Mitton, 1972; Fujio et al., 1983, Buroker, 1983, 1984). Both the 66

cytosolic and mitochondrial MDH activities in the gill tissues of these bivalves showed polymorphism (data not shown).

From the physiological viewpoint, the mitochondrial localization

of ME in most species and the unusual AAT and AlAT distribution in

oyster gill and ribbed mussel gill, respectively, may be helpful in

rationalizing some of the species and tissue differences in amino acid

accumulation during osmotic and anaerobic stress (see Part I & II). 67

PART IV. PYRUVATE DEHYDROGENASE FROM RIBBED MUSSEL GILL TISSUE

The pyruvate dehydrogenase complex has been demonstrated in high speed (150,000 g) pellet preparations from sonicated ribbed mussel gill mitochondria. The complex is inhibited by low (<100 mM) chloride concentrations, succinate and ATP. ATP inhibition was enhanced by NaT and reversed by high Mg^^ concentrations in the absence of NaT.

Pyruvate and thiamine pyrophosphate inhibited the inactivation by ATP.

The non-hydrolyzable ATP analog AMP-PNP caused inhibition of the overall catalytic activity that was identical to ATP. Factors in­ volved in the ATP inhibition and Mg^^ reversal are lost with freezing or cold storage. The activity of the complex may be regulated by a phosphorylation/dephosphorylation mechanism. The molluscan complex appears to be similar to the mammalian complex. 68

INTRODUCTION

Most estuarine bivalve mollusc tissues accumulate high levels of

intracellular amino acids in response to hyperosmotic stress (see

Pierce, 1982). The accumulation of alanine in mussel tissues is transaminase dependent (Bishop et al., 1981; Greenwalt and Bishop,

1980; deZwaan et al., 1983a,b). In the ribbed mussel gills under

study here, this increase in the alanine levels requires action of the

alanine aminotransferase activity which is localized in the mitochon­

dria (Part II). Therefore, in order to control the accumulation of

the high levels of alanine (0.1-0.2 M) during salt stress, the turn­

over of pyruvate within the mitochondria of these mussel tissues must

be acutely regulated. Results from Parts I & II suggest that there

may be an acute regulation (inhibition) of pyruvate dehydrogenase

complex (PDC) to shunt mitochondrial pyruvate toward alanine during

anaerobic or hyperosmotic stress. This regulation (inhibition) of PDC

is apparently coordinated with the regulation (inhibition) of the

mitochondrial glycine cleavage enzyme to cause coordinate glycine

accumulation (Ellis et al., 1985). From other studies, it appears

that acute regulation or modification of the PDC activity may also

occur in bivalves (Ho and Zubkoff, 1982; Kluytraans et al., 1978) and

helminth parasites (Komuniecki et al., 1981, 1983; Rew and Saz, 1974)

to account for the anaerobic production of acetate from carbohydrate

derived pyruvate.

The pyruvate dehydrogenase complex (PDC) is a multienzyme complex

within the mitochondrion that catalyzes the reaction of pyruvate with 69

CoA and NAD to form acetyl CoA, CO^ and NADH (see Reed, 1974).

Wieland (1983) has recently reviewed the structure and regulation of the mammalian PDC. It is regulated primarily by an ATP dependent phosphorylation/dephosphorylation (protein kinase-phoshatase) mecha­ nism that is mediated by Ca"*"^, Mg"*"*", and/or other intracellular

effectors. Komuniecki et al. (1979, 1983) have investigated the PDC from the muscle tissue mitochondria of the "anaerobic" parasitic roundworm. Ascaris » The properties of this purified Ascaris enzyme appeared to resemble those of the mammalian complex in kinetic terms and showed the protein kinase-phosphatase regulatory characteristics.

Although a small amount of PDC has been reported in mitochondria

from sea mussel tissues (Addink and Veenhof, 1975), there are no stu­

dies on the regulatory or other properties of this important enzyme in

molluscs. In fact, pyruvate added to coupled mollusc gill mitochon­

dria failed to stimulate oxygen consumption (Burcham et al., 1984).

Questions have arisen as to the amount of enzyme in raolluscan tissues

and whether or not regulation of the PDC was similar to the complexes

from other animals. In this final part, a convenient method for

preparation and assay of PDC from bivalve tissue mitochondria is

described, as well as some properties demonstrated by the complex in

this preparation and preliminary evidence for regulation of the

activity. 70

MATERIALS AND METHODS

Ribbed mussels (Modiolus demissus) were obtained from Northeast

Marine Environmental Institute (Monument Beach, MA). The animals were maintained as described by Greenwalt and Bishop (1980). Except where noted, all reagents were purchased from Sigma (St. Louis, MO). Enzyme- grade ammonium sulfate and sucrose were obtained from Schwarz-Mann

(Orangeburg, NY). 1-^^C-Pyruvate was obtained from New England

Nuclear Corp.

The enzyme complexes were partially purified from a mitochondrial lysate by a series of differential centrifugations (Komuniecki et al.,

1979). Mitochondria were prepared as described in Part I and soni­ cated using a Branson sonifier on a setting of "5" with the intermed­ iate size probe. Small batches (1-2 ml) of mitochondrial suspension were sonicated three times for 10 seconds on ice with 10 seconds between each sonic treatment. The sonicated suspension was centri- fugsd for 30 nsin at 20,000 g to remove unbroken mitochondria and debris. The supernatant was centrifuged at 150,000 3 for 90 min. The pellet from this centrifugation was resuspended in mitochondrial isolation buffer, then recentrifuged at 20,000 g for 30 min. The supernatant was then centrifuged a second time at 150,000 g for 90 min. The pellet was again resuspended in mitochondrial isolation buffer and recentrifuged for 30 min at 20,000 3. The supernatant from this final centrifugation constituted the enzyme preparation used for the preliminary studies described in this paper. 71

The enzyme activity was assayed both radiometrically and spectro-

photometrically during preparation (Komuniecki et al., 1979). The

radiometric assay consisted of the incubation of l-^^C-pyruvate (0.5

uCi) in a reaction mixture containing 2.5 mM pyruvate, 0.1 mM NAD,

0.1 mM CoA, 0.1 mM thiamine pyrophosphate (TPP), 1.0 mM dithiothreitol

(DTT) and 100 mM Tris/HCl in 1 ml. The reaction was stopped after an

hr. with 2N HCl and CO^ collected in a filter paper trap with 100

wl of 1 M hyamine hydroxide in methanol. This assay was used in

the early phase of the purification where the spectrophotometric assay

was impossible. The spectrophotometric assay employed the same

reaction mixture (in 1 ml) and the production of NADH (A^^^) was

measured in a recording spectrophotometer. a-Ketoglutarate

dehydrogenase was assayed using the same reaction mixture by

substituting a-keto-glutarate for pyruvate.

ATPases were assayed by incubation with 5 mM phosphoenolpyruvate

(PEP), 5 units pyruvate kinase, 5 units lactate dehydrogenase, 70

yM NADH, and 1 mM ATP in 100 mM Tris/HCl pH 8.3. NADH oxidation

was measured in a spectrophotometer (A^^g). NADH oxidase activity

was measured by incubation of enzyme with 70 uM NADH in 100 mM

Tris/HCl pH 8.3. 32 The molluscan PDC was incubated with y- P-ATP in an attempt

to more directly demonstrate covalent modification of a component of

the complex. Sample (4 mg) was incubated with 100 mM Tris/HCl pH 8.3,

ImM DTT, 10 uM TPP, and 200 uM ATP (specific activity- 300

uCi/ymole ATP) in 1 ml. Specific PDC protein kinase inhibitors 72

pyruvate (5 raM) and TPP (200 yM) were also included in experi­ mental incubations. Twenty ug of purified mammalian PDH (E^) was incubated with 4 mg of the molluscan prep in the same incubation mixture without the protein kinase inhibitors. Incubations were stopped by addition of .5 ml 2x sample buffer (Laemmli, 1970) and heated immediately in a boiling water bath for 1 min. Samples (approx.

6.5 mg protein and 7.5 uCi of radioactivity) were electro-

phoresed on 10% acrylamide gels according to Laemmli (1970). Low

molecular weight standards (10,000-100,000) were obtained from

Bio-Rad. The gels were stained with Coomassie brilliant blue in 25%

isopropanol/10% acetic acid overnight, destained in 10% isopropanol/

10% acetic acid, and fixed in 10% acetic acid/0.1% glycerol. The gels

were dried on a Hoefer Model SE 540 slab gel dryer and exposed at

-70°C for 10 days in the dark using Cronex Lightning Plus

intensifying screens and Kodak X-Omat AR (XAR-5) X-ray film.

Protein was estimated according to Lowry as modified by Miller

(1959) or by the biuret method for mitochondrial suspensions (King,

1967). 73

RESULTS AND DISCUSSION

In crude homogenates, mitochondrial suspensions, and mitochon­ drial lysates, pyruvate dehydrogenase activity was demonstrated using the production of from 1-^^C-pyruvate as the assay procedure. After ultracentrifugation, the complexes pelleting at

150,000 2 were resuspended in buffer yielding a reasonably trans­ parent sample for use with the spectrophotometric assay. This preparation showed CoA and NAD dependency using the radiometric procedure. Pyruvate and CoA dependency was demonstrated using the spectrophotometric method. The rate of the reaction increased with increasing enzyme concentration (Figure 1). There was considerably more KGDC than PDC in gill mitochondria.

Table 1. Pyruvate dehydrogenase complex (PDC) and a-keto- glutarate dehydrogenase complex (KDGC) activities in mitochondria from ribbed mussel gill tissue^

Activity Reaction (nmol/min/mg protein) Mixture PDC KGDC

Complete 12. 62. Without CoA <0.01 <0.01 Without ketoacid <0.01 <0.01 Plus Ars (1 mM) 0.01 0.60 Plus ATP (1 mM) 3. 40. Plus NaF (10 mM) 12. 62. Plus NaF + ATP (1 mM + 10 mM) 0.67 38.

"^The assay procedure and reaction mixture are described in Materials and Methods. Where appropriate, enzyme activity was deter­ mined after a two minute preincubation with inhibitor. In the NaF + ATP assays, enzyme was incubated first with NaF for two minutes, then assayed. Dithiothreitol was present during all incubations except in the arsenite (Ars) experiment. 74

The activities of the PDC and KGDC with and without substrates or effectors are presented in Table 1. EDTA (1 mM) inhibited the PDC activity 100% but did not affect the KGDC activity. Addition of 10 mM

MgClj - 1 mM CaCl^ in buffer (Tris) restored full activity to preparations containing ImM EDTA. Arsenite (Ars) effectively inhib­ ited both enzymes at a 1 mM. ATP addition caused almost complete inhibiton of the PDC but only 35% inhibition of the KGDC. NaF had no effect on either enzyme in the absence of ATP. Addition of both ATP and NaF caused complete inhibition of PDC and a 40% reduction of the

KGDC activity. NADH oxidase activity was <0.5 nmol/min/mg protein.

Preliminary kinetic investigations on this crude preparation indi­ cated that the substrate K s for the molluscan PDC were similar to m those of the mammalian and Ascaris complexes. Apparent Michaelis constants for substrates were generated using Cleland's (1979) kinetic analysis. The apparent K^s for pyruvate, CoA, and NAD were 300 yM, 5.8 yM, and 96 yM, respectively. Standard error for all of the determinations was less than 20%. The apparent K^ for pyruvate seemed higher than those reported for many mammalian complexes whereas the values for CoA and NAD were similar (Biass and

Lewis, 1973; Roche and Cate, 1977; Wisland, 1983). The high K^ for

pyruvate of the bivalve complex was comparable to the apparent K^

(185 yM) reported for the Ascaris PDC (Komuniecki et al., 1979).

Alanine, aspartate, glycine, taurine, acetate, and proline at 100

mM concentrations had no effect on the catalytic activity of the mol­

luscan PDC. Succinate (28 mM) inhibited the catalytic activity 50%. 75

12

.E 8 I o E c .

0

0 100 200

All

Figure 1. Effect of amount of enzyme on the reaction rate of the pyruvate dehydrogenase complex (PDC) and the a-ketoglutarate dehydrogenase (KGDC) from the mitochondria of ribbed mussel gill tissue. Assay procedure is described in Materials and Methods 76

Figure 2. Effect of ATP on the pyruvate dehydrogenase complex activity. 2A: ATP and MgCl^ were 1 mM and 10 mM, respectively. 2B: NaF and ATP were 10 mM and 1 mM, respectively. ATP, MgCl_, or NaF were added to the reaction mixture at the indicated intervals while the reaction rate was being monitored in the recording' spectrophotometer 77

100

50

0.001 0.01 0.1 10 NUCLEOTIDE{mM)

Figure 3. The effect of TPP concentration on the inhibition of PDC activity by ATP (o,®) or AMP-PNP (A,A). Activity was assayed as described in Materials and Methods in the presence of varying amounts of nucleotide at two different concentrations of TPP; 10 yM (o,A) and 100 wM (®,A). TPP clearly blocks the inhibitory effects of ATP on PDC activity These properties were similar to those of the mammalian complex (Biass and Lewis, 1973; Wieland, 1983). Chloride had an inhibitory effect on the catalytic activity, in that low (<100 mM) concentrations of KCl,

NaCl, or choline chloride significantly inhibited the PDC with values of 63 mM, 63 mM, and 40 mM, respectively. The catalytic activ­ ity was lost rapidly if the preparations were stored in chloride con­ taining buffers. NaF (50 mM) or Na- or K-acetate (100 mM) showed no inhibitory effects. This molluscan complex also showed no changes in activity between pH 6.8 and 9.0 when assayed under standard substrate conditions.

The NAD/NADH ratio in cells or mitochondria has been thought to affect the metabolism of those cells and has been shown to have sig­ nificant effects on the pyruvate dehydrogenase activities from mammals

(Biass and Lewis, 1973; Roche and Gate, 1977; Wieland, 1983) and

Ascaris (Komuniecki et al., 1979, 1983). Although this effect may not

be physiologically significant in mammals (Siess et al., 1978), the

role of the NAD/NADH ratio may be very important in the anaerobic

metabolism of adult Ascaris (Komuniecki et al., 1979). The activity

of the molluscan complex was markedly effected by a change in the

NAD/NADH ratio (Fig. 4). The greatest changes in the activity of the

molluscan complex occurred at NAD/NADH ratios between 2 and 6 when the

concentrations of NAD were 100, 200, 600 yM. The Ascaris complex

shows sensitivity at a much lower NAD/NADH ratio (<1) which would

allow the helminth complex to function in a more reduced state (see

Komuniecki et al., 1979). 79

ATP inhibition of the PDC was investigated more closely. ATP completely inhibited the PDC activity in a time dependent fashion

(Figure 2A); this inhibition was reversed by subsequent addition of lOmM MgClj (Figure 2A). When NaF was included in the reaction mixture with ATP, the inhibition was more rapid and complete (Figure

2B). This experiment supported the possibility of a kinase-phospha­ tase regulatory system for PDC.

The preparation was assayed for ATPase activity to test the possibility that a Mg^"*" stimulated ATPase was hydrolyzing ATP and thereby releasing the inhibitory effect of the ATP with the addition of Mg^"*". The ATPase activity was relatively insignificant in the absence of additional Mg (0.50 nmol/rain/mg protein), but increased to a high level (11.05 nmol/min/mg protein) in the presence of lOmM

Mg"""*". The ATPase activity was insensitive to oligomycin indicating that if the activity was the ATP synthase, the regulatory subunit was lost during preparation. Given that the level of ATP (1 mM) in the inhibition experiment (Fig. 2), this Mg"'"'" stimulated ATPase would have reduced the ATP level less than 5% and would not have affected

ATP inhibition or reversal of inhibition with addition.

Other nucleotide phosphates were tested for inhibition of the PDC activity to further investigate the possibility of a kinase-phospha­ tase system. Although GTP and ADP showed some effects at high (1 mM)

concentrations, they showed no effect in the range that ATP was

effective. The non-hydrolyzable ATP analogs, B,y-Methylene-

adenosine 5'-triphosphate (AMP-PCP), and 5-adenylylimidodiphosphate 80

(AMP-PNP) were used in place of ATP. AMP-PCP inhibited only at very high (> ImM) concentrations whereas inhibition by AMP-PNP was nearly identical to that exhibited by ATP at the same concentrations.

High levels of both pyruvate and TPP have been shown to interfere with the protein kinase activity in PDC from mammalian mitochondria

(Wieland, 1983). Pyruvate and TPP concentrations,effect the inactiva- tion of the gill PDC activity'by ATP or AMP-PNP. A ten-fold decrease in TPP concentration causes an approximate five-fold decrease in the amount of ATP required for inhibition of the PDC activity (Figure 3).

Pyruvate also blocked the inhibitory effect of ATP but was not as effective as TPP. These results supported the hypothesis that the protein kinase was associated with the raolluscan PDC and was inhibited by pyruvate and TPP. The inhibitory effect of AMP-PNP was curious and suggested that there was a direct effect of ATP and AMP-PNP on the PDC that might not be protein kinase-phosphatase related.

In order to test for the protein kinase activity in a more direct 32 fashion, preparations were incubated with y- P-ATP under conditions

in which the PDC activity was inhibited in the spectrophotometric assay and in the presence of the specific PDC protein kinase

inhibitors, pyruvate and TPP at concentrations which blocked ATP

inhibition in the spectrophotometric assay (see Materials and

Methods). Several protein bands were apparent in the gels stained

with Coomassie blue. Purified mammalian electrophoresed in a

lane alone showed two bands of 41 and 37 Kd. These bands were also

apparent in the incubations containing the molluscan preparation with NAD o 642 uM A 214 uM o 107 uM

10 ç 'E 4 8 12 16 20 NAD/NADH

>-

œ

0 0 50 100 150 200

NADH (JJM)

Figure 4. The effects of NADH on PDC activity in the presence of different amounts of NAD. Activity was assayed as described in Materials and Methods at three indicated concentrations of NAD with varying amounts of NADH. Inset shows the data recalculated to show activity as related to the NAD/NADH ratio 82

mammalian added. Autoradiography of these gels revealed that 32 several proteins were labelled by y- P-ATP. A band at about 41

Kd which corresponded to the a subunit of the mammalian was labelled. In incubations containing mammalian the 41 Kd band was the major band labelled and contained significantly more radioactivity than samoples without the E^ added. This was the

a subunit of the component of mammalian PDC, and is the

subunit which is phosphorylated in the mammalian (Barrera et al.,

1972; Reed, 1974) and Ascaris (Komuniecki et al., 1983) complexes. No

label was detected in the B subunit (37 Kd). These experiments

indicate that the protein kinase was indeed present and will

covalently modify the a subunit of the PDH component of the

complex. The molluscan PDH component, however, may not have been

present in sufficient quantities in this preparation as evidenced by

the faint 41 Kd radioactive band.

It was noted in continuing experiments that while the catalytic

activity was fairly stable, the inhibitory effect of ATP was modified

fay freezing or cold storage of the preparation. For instance, al­

though fresh preparations were inhibited 100% by ATP with NaF present

(Table 1), after several days of freezing or refrigeration the PDC was

less than 50% inhibited under the same conditions with either 1 mM ATP

or 1 mM AMP-PNP. Additionally, Mg^"^ showed no effect on the relief

from the partial ATP or AMP-PNP inhibition of these frozen or refrig­

erated PDC preparations. This inhibitory effect was also insensitive

to pyruvate or TPP concentration, indicating the ATP or AMP-PNP may be 83

exerting a direct inhibitory effect on the catalytic activity of the complex. These PDC preparations showing altered ATP sensitivity could be stored frozen for several weeks with little loss in overall cataly­ tic activity (standard assay).

The preliminary studies reported here indicate that PDC is pres­ ent in bivalve tissue mitochondria and is probably similar to the mammalian complex. As with the mammalian PDC, the ribbed mussel PDC is dependent on CoA, NAD, and pyruvate, and may be controlled by an

ATP dependent regulatory response characteristic of the protein phosphorylation/dephosphorylation scheme originally described for the mammalian PDCs (Linn et al., 1969; Wieland and Jagow-Westermann,

1969). There appears to be a secondary regulatory or inhibitory effect of ATP or ATP analogs on the catalytic activity of the gill

PDC. The characteristics exhibited by the PDC from ribbed mussel gill tissue lend support to the hypothesis that the complex could play an important role in the regulation of alanine, pyruvate and acetate metabolism during osmotic stress and hypoxia. 84

GENERAL DISCUSSION

Two different forms of aspartate aminotransferase, a cytosolic

(cAAT) enzyme and a mitochondrial (mAAT) enzyme, have been found in

Modiolus demissus gill tissue. The two enzymes differ in electro-

phoretic mobility, heat stability, and kinetic properties. The

kinetic differences may be physiologically significant especially

during metabolic adjustment to hyperosmotic or anaerobic stress. The

alanine aminotransferase, however, appears to have only one form in

this tissue, and that form is apparently mitochondrial. Therefore,

metabolically derived alanine must be produced or degraded within the

mitochondria. This would require an acute regulation (inhibition) of

mitochondrial pyruvate catabolism. The pyruvate dehydrogenase complex

(PDC) is the major regulatory enzyme of pyruvate catabolism in mammals

(Denton and Halestrap, 1979) and, although virtually no work has been

published about molluscan PDCs, the complex was considered the most

likely regulatory enzyme of pyruvate metabolism in the mitochondria

from ribbed mussels. PDC from isolated ribbed mussel mitochondria was

prepared by ultracentrifugation, and partially characterized with

respect to kinetic and regulatory properties.

The kinetic characterization of the two forms of aspartate

aminotransferase (mAAT and cAAT) suggested that aspartate synthesis

would be favored in the cytosol, and that aspartate breakdown would be

favored in the mitochondria. As had been shown, the cAAT would be

inhibited at a low anoxic pH (6.5) and hence cytosolic aspartate

production would tend to shut down as the tissue pH falls, whereas 85

aspartate synthesis or breakdown in the mitochondria would remain relatively unaffected by pH change. If this were true, and alanine and succinate production were linked to aspartate catabolism as has been suggested (Collicutt and Hochachka, 1977; deZwaan, 1977; deZwaan et al., 1982), then alanine production from aspartate would be

predicted to be a mitochondrial process. This hypothesis led to

studies on the alanine aminotransferase from ribbed mussel tissues.

Most mammalian tissues possess two different forms (cytosolic and

mitochondrial) of alanine aminotransferase (AlAT) (Hopper and Segal,

1962; Swick et al., 1965). It was demonstrated that ribbed mussel

gill tissue possessed only one form of AlAT, and that form was

mitochondrial. This finding agreed with the prediction from the

aspartate aminotransferase work and proved that alanine turnover must

be mitochondrial. The demonstration of mitochondrial alanine produc­

tion suggested that pyruvate catabolism would have to be inhibited to

shunt pyruvate to alanine. Since glycolytic flux does not change in

this tissue during hyperosmotic or anaerobic stress (Bishop et al.,

1981, Greenwalt, 1981), the pyruvate dehydrogenase complex (PDC) would

have to be inhibited to accumulate alanine from pyruvate.

The PDC studies have demonstrated several important facts.

First, PDC is present and extractable in ribbed mussel gill tissue.

Second, the mussel PDC shows many of the same properties that the

mammalian complex shows. From these findings, one might predict that

the PDC would be controlled in vivo by the redox (NAD/NADH) balance

within the range of the cellular NAD concentration of the cells, by 86

levels within the mitochondria, or possibly by a phosphorylat ion/dephosphorylat ion event.

This work has several important ramifications. First, it demon­ strates that, in at least ribbed mussel gill tissue, the mitochondria must play the major role in metabolic adjustments to anoxia or hyperosmotic stress. Ellis et al. (1985) have demonstrated that glycine levels are also controlled in the mitochondria by regulation of the glycine cleavage system. Since glycine and alanine levels rise and fall under the same circumstances, it appears that the glycine cleavage system and the pyruvate dehydrogenase complex may be coordi- nately regulated. Second, this work demonstrates that mussel tissues contain the pyruvate dehydrogenase complex, that the complex appears to have most if not all of the regulatory properties that the mam­ malian complex exhibits, and that the molluscan PDC probably plays a key role in the metabolic shift of these animals to an anaerobic metabolism.

In conclusion, this work adds to the growing body of data gener­ ated by this lab which demonstrates that the tissues of the molluscs . use the same pathways as mammalian tissues, but they regulate metab­ olite flux through these pathways differently to produce an accumu­ lation of endproducts (alanine, glycine) in the face of a salt stress that might prove detrimental or lethal in mammalian tissues. 87

ACKNOWLEDGEMENTS

I would like to express my sincere appreciation to Dr. Stephen

Bishop for his help and guidance during ray graduate education and to the members of my committee, especially Dr. Richard Hoffmann. Special thanks go to Dr. Lester Reed, who generously supplied the purified component of the mammalian PDC and to Dr. Joel Abramowitz and Mr. 32 Alfonso Campbell, who provided the Y- P-ATP. I would also like to thank Dr. Sheldon Shen, Dr. Lehman Ellis, Kevin Kraus, Tom McMullin, and Tom Davis for the helpful discussions and support, and Dr. Richard

Komuniecki and Julia Thissen for their help, advice and support on the

PDC work. I would especially like to thank my wife, Lynn, not only for her help in the preparation of this thesis, but for her loving support during our stay at Iowa State. 88

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