Kinetic studies of Haemophilus influenzae malate dehydrogenase

Biochimica et Biophysica Acta, 955 (1988) 10-18 Elsevier

10

BBA33141

Kinetic studies of Haemophilus influenzae m a l a t e d e h y d r o g e n a s e H c e j e o n g Y o o n a n d Bruce M. A n d e r s o n Department of Biochemistry and Nutrition, Virginia Polytechnic Institute and State University, Blacksburg, t'A (U.S.A.) (Received 25 January 1988)

Key words: Malate dehydrogenase; Enzyme kinetics; Meningitis; (H. influenzae)

HaemophYus influenzue malate dehydrogenase ((S)-malate: NAD + oxidoreduetase EC 1.1.1.37) was purified 109-fold with a 26% recovery through a four-step procedure involving salt fractionation, hydrophobic and dye affinity chromatography. The purified enzyme was demonstrated to be a dimer of Mr 61000. Initial velocity studies of all four substrates in the forward and reverse reactions indicated a sequential mechanism for the enzyme. Product and dead-end inhibition studies were consistent with an ordered bi-bi mechanism in which NAD is the first substrate bound to the enzyme and NADH, the second product released. Several analogs of NAD structurally altered in either the pyridine or purine moiety were observed to function as coenzymes in the reaction catalyzed by the purified malate dehydmgenase. Alterations in the purine portion of the dinucleotides had a more pronounced effect on the kinetic parameters observed in malate oxidation. The enzyme was inactivated by incubation with diethylpyrocarbonate, whereas no inactivation was observed with sulfhydryl reagents.

Introduction

Organisms of the genus Haemophilus have been subclassified on the basis of unique requirements for one or both of two growth factors, viz. X-factor (heroin) and V-factor (NAD). Manifestation of the V-factor requirement had been related to the inability of V-factor organisms to synthesize NAD from typical precursors of the de novo biosynthetic pathways as well as from nicotinamide and nicotinic acid [1-3]. As a V-factor organism, Haemophilus influenzae exhibits growth dependency on externally provided NAD, and recent studies [4] have presented information concerning the process by which external NAD is utilized by

Correspondence: B.M. Anderson, Department of Biochemistry and Nutrition, Virginia Polytechnic Institute and State University, Biacksburg, VA 24061, U.S.A.

this organism. A periplasmic nucleotide pyrophosphatase (EC 3.6.1.9) was isolated from H. influenzae and was demonstrated to catalyze the hydrolysis of external NAD to AMP and nicotinamide mononucleotide (NMN), the latter being utilized for the intraceUular resynthesis of NAD [4]. This enzyme was purified to electrophoretic homogeneity, and concentrations of AMP and ADP that selectively inhibited the enzyme were observed to inhibit effectively the growth of the organism. Interest in controlling the growth of H. influenzae arises from the documented involvement of this and related species in numerous diseases in :nan and other animals. H. influenzae type b is the primary cause of bacterial meningitis in humans, responsible for 8000 cases annually in the United States [5]. Treatment of H. influenzae infections has relied predominantly on the use of ampicillin; however, a significant increase in the observed involvement of strains resistant to this antibiotic has been reported [6,7].

0167-4838/88/$03.50 © 1988 Elsevier Science Publishers B.V. (Biome~cal Division)

11

In studies of the periplasmic nudeotide pyrophosphatase, the 3-acetylpyridine and thionicotinamide analogs of NAD served as substrates for the nucleotide pyrophosphatase as well as supporting growth of the organism in the absence of NAD. The observed growth on these analogs would require intracellular dehydrogenases to utilize these unnatural coenzymes in important metabolic processes. In recent studies of glucose utilization in Haemophilus organisms [8], a partial tricarboxylie acid cycle was identified in which an obvious importance for a malate dehydrogenase was indicated. The present study reports the purification and characterization of H. influenzae malate dehydrogenase ((S)-malate: N A D ~ oxidoreductase, EC 1.1.1.37), including an evaluation of coenzyme specificity of this metabolically important enzyme. Experimental procedures

Materials Haemophilus influenzae str ".aJnRd was obtained from Dr. W.L. Albritton of the University of Saskatchewan, Saskatoon. All mono- and dinucleotides were purchased from Sigma, except thionicotinamide adenine dinucleotide, 3-pyridylacryloamide adenine dinucleotide, 3-aminopyridine adenine dinucleotide, and 3-pyridinealdehyde adenine dinucleotide, which were prepared by published procedures [9,10]. Fluorescein mercuric acetate, pyridoxal 5'-phosphate, iodoacetamide, iodoacetic acid, N-ethylmaleimide, diethylpyrocarbonate, and molecular weight standards were from Sigma. Phenyl-Sepharose CL-4B and Sephadex G-100 were purchased from Pharmacia. Matrex Green Gel A and Matrex Blue Gel A were obtained from Amicon. Brain Heart Infusion was purchased from Fisher Scientific.

Methods Purification of malate dehydrogenase. H. influenzae cells were grown in Brain Heart Infusion medium according to the procedure described by Kahn and Anderson [4]. In studies of the malate dehydrogenase, cells were harvested after 10 h (late linear phase) by centrifugation, washed, and then stored at - 1 5 *C in a minimal amount of 50 mM potassium phosphate buffer (pH 7.0) until needed. A 50% (w/v) homogenate of frozen wet

cells in 50 mM potassium ,,~,,,o,,h~)° e,,,,~v,,,,-- buffer (pH 7.0) was prepared at 4 ° C using a glass homogenizer. The homogenate was sonicated in an ice/salt bath using the microprobe tip of an Ultrasonics Sonifier Cell Disruptor and centrifuged at 17000 × g for 20 rain. The sonication was repeated twice using the pellets from centrifugations, and the supemates were combined for the protamine sulfate step. Protamine sulfate was added to the sonicate supernate at 4°C to obtain a 0.2% protamine sulfate solution. This solution was stirred for 30 rain in an ice-bath, and centrifuged at 39000 x g for 20 rain. The pellet was discarded and the supernate was adjusted to 40% ammonium sulfate saturation with continuous stirring at 4°C. After centrifugation at 11000 x g for 30 rain, the resulting supernate was adjusted to 60% ammonium sulfate saturation. The pellet obtained after centrifuging this suspension was resuspended in 10 ml or more of 10 mM potassium phosphate buffer (pH 7.0) to adjust the protein concentration of this solution to 8 mg/rnl. At this step in the purification procedure, the malate dehydrogenase would not bind to affinity chromatography resins at protein concentrations higher than 8 mg/ml. The diluted protein solution was applied to a Matrex Green Gel A column (1.0 × 6 cm) equilibrated in 10 mM potassium phosphatic (pH 7.0). After washing with the equilibration buffer, the malate dehydrogenase was eluted by using a linear salt gradient, 0.05-0.8 M KCI in the same buffer. Fractions (2.4 ml) were assayed for enzyme activity, and the protein concentration was monitored by 280 nm absorbance. Fractions containing enzyme activity were pooled and used in the next ourification step. The pooled sample from the Green A column was adjusted to a final concentration of I M KC1 by addition of solid KC1. This solution was applied to a phenyl-Sepharose hydrophobic column (1.0 × 9.5 cm) previously equilibrated with 10 mM potassium phosphate (pH 7.0)/1 M KC1. Unbound proteins were washed from the column with equilibration buffer. The malate dehydrogenase was then eluted with the lower ionic strength 10 mM potassium phosphate buffer (pH 7.0). Fractions (2.4 ml) containing enzyme activity were pooled for further fractionation.

12 TABLE I PURIFICATION OF H. INFLUENZAE MALATE DEHYDROGENASE Fraction

Cell sonicate Ammonium sulfate pellet Matrix Green Gel A Phenyl-Sepharose Matrex Blue Gel A

Total protein (rag)

Total activity

Yield (%)

(-fold)

(units)

Specific activity (units/mg)

411.4 159.1 15.1 2.1 1.0

391 371 246 153 104

0.95 2.3 16.3 73 104

100 95 63 39 26

1 2.4 17 77 109

The sample from the phenyl-Sepharose colunm was applied to a Matrex Blue Gel A column (1.0 x 5 cm) equilibrated with 10 mM potassium phosphate (pH 7.0). After washing with equilibration buffer, the malate dehydrogenase was eluted by a linear salt gradient, 0.05-0.8 M KCI in the same buffer. Fractions (2.4 ml) containing enzyme activity were pooled and concentrated by ultrafiltration. The results of the purification procedure are shown in Table I. The enzyme was purified 109-fold with a 26~; yield and a final specific activity of 104/~mol/min per mg protein. Assays of malate dehydrogenase activity. Routine assays of malate dehydrogenase activity were performed at 23°C in reaction mixtures containing 45 mM glycine-NaOH buffer, (pH 9.6), 3 mM malate, 0.54 mM NAD and enzyme in a final volume of 1 ml. Initial velocities were determined spectrophotometrically by monitoring NADH formarion at 340 nm. For the determination of kinetic parameters, replotting methods were applied using initial velocities obtained by varying one substrate at several fixed concentrations of the second substrate. Oxalacetate reduction was assayed in 1 ml reaction mixtures containing 50 mM Tris-HCl buffer (pH 8.5), measuring NADH decrease at 340 nm. A unit of enzyme activity was defined as 1 /~mol of coenzyme oxidized or reduced per rain. Spectrophotometric measurements were performed on a Beckman Acta MVI recording spectrophotometer. Fluorescence ~as measured on a Perkin-Elmer 650-40 spectrofluorometer. Protein concentrations were determined by the microprorein assay of the Coomassie blue method [11] using bovine serum albumin as a standard. Ultrafiltration was carried out using Amicon PM-10 membranes.

Purification

Molecular weight determination. Molecular weight of the purified enzyme was determined under nondenaturing conditions using a gel-filtration Sephadex G-100 chromatography column, and high-performance liquid chromatography (HPLC) on a Bio-Sil TSK column. Under denaturing conditions, the molecular weight was determined by SDS-polyacrylamide gel electrophoresis according to established procedures [12,13]. Results

Properties of pvrified malate dehydrogenase The purified enzyme analyzed by polyacrylamide gel electrophoresis at pH 8.3 showed one protein band which corresponded to malate dehydrogenase as determined by activity staining techniques. The molecular weight of the purified enzyme was determined by Sephadex G-100 molecular exclusion chromatography at 4° C. The elution volume of malate dehydrogenase in comparison with protein molecular weight standards (Fig~ 1, line A) indicated an apparent Mr of 62 000 for the native enzyme. An apparent molecular weight of 60000 was observed when the native enzyme was analyzed by HPLC on a Bio-Sil TSK gel column. When the purified enzyme was subjected to SDSpolyacrylamide gel electrophoresis in comparison with protein molecular standards (Fig. 1, line B), an apparent M r of 31000 was obtained. The purified enzyme was routinely stored at - 1 5 ° C in a 50% propylene glycol solution containing 100 mM KCI and 10 mM potassium phosphate (pH 7.0). Under these conditions, the enzyme was stable for at least 3 months with no apparent loss of activity. The enzyme was slightly less stable in the absence of propylene glycol. The

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I I m ~°. i 1.8 2,2 2.4 Ve/Vo Fig. 1. Molecular weight determination of malate dehydrogenase. (A) Gel filtration on Sephadex G-100 column. Molecular weight standards were (1) fl-galactosidase, (2) bovine serum albumin, (3) ovalbumin and (4) lysozyme. (B) Sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Molecular weight standards were (1) lysozyme, (2) soybean trypsin inhibitor, (3) carbomc a,hydrase, (4) ovalbumin and (~) bovine serum albumin. H. influenzae I

1.6

2.0

ThearrowsinAandBdenoteposi~:~onsdeterminedfor malatedehydrogenase.

purified enzyme preparation was, however, denatured at moderate temperatures. When the enzyme activity was assayed at timed intervals during incubation in 10 mM potassium phosphate (pH 7.0) at four different temperatures, the rate of thermal denaturation followed first-order kinetics and at 50 o C, the rate constant for thermal denaturation was 0.051 rain -1. The presence of 0.3 mM NAD gave 46% protection against thermal denaturation. Concentration of malate as high as 4 mM did not provide any protection against thermal denaturation.

Kinetic mechanism of malate dehydrogenase The kinetic constants for both substrates in the forward and reverse directions were determined through initial velocity studies by varying one substrate at several fixed concentrations of the second substrate. The initial velocities obtained by varying malate concentration at four fixed concentrations of NAD are shown in Fig. 2. Slope and intercept replots were used to determine values for K m and Vmax- In a separate experiment using saturating concentrations of malate and NAD, Vm~ values were proportional to enzyme

concentration over a 6-fold concentration range. The Krn values determined for all four substrates in the forward and reverse reactions were: NAD, 57 #M; NADH, 2 #M; malate, 0.29 mM; and oxalacetate, 13 #M. The results obtained by varying each substrate at different fixed concentrations of the appropriate second substrate gave converging line relationships similar to that depicted in Fig. 2, indicating the involvement of a sequential mechanism. In order to determine whether the sequential reaction mechanism was of an ordered or random type, product and dead-end inhibition studies were conducted. In the product inhibition studies, inhibition patterns were determined from double-reciprocal plots using two fixed product concentrations and varying one substrate at unsaturating and saturating concentrations of the second substrate. When using NAD as the variable substrate, inhibition by one product, NADH, at both unsaturating and saturating concentrations of malate was competitive (Ki = 1.3/~M) as shown in Fig. 3. When inhibition by NADH was examined with malate as the variable substrate at an unsaturating concentration of NAD, noncompetitive inhibition was observed. No inhibition by NADH was observed when a saturating concentration of NAD was employed. When using NAD as the variable substrate, product inhibition by oxalacetate was noncompetitive at an unsaturating concentration of malate (Ki = 12.8/~M), and uncompetitive at a saturating concentration of malate. When inhibition by oxalacetate was studied with malate as the variable substrate, noncompetitive inhibition was observed at both unsaturating and saturating concentrations of NAD. Structural analogs of NAD and malate, adenosine diphosphoribose and hydroxymalonate, respectively, were studied as dead-end inhibitors. Inhibition by adenosine diphosphoribose was competitive when NAD was used as the variable substrate and noncompetitive when malate was the variable substrate. Inhibition by hydroxymalonate was uncompetitive when NAD was the variable substrate and competitive when malate was the variable substrate. The Ki values determined for adenosine diphosphoribose and hydroxymalonate were 50 #M and 1.6 mM, respec-

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[.ALATE] raM" Fig. 2. Effect of NAD and malate concentrations on initial velocities. The assay mixtures contained concentrations of malate varying from 0.2 to 0.8 mM at four fixed concentrations of NAD in 45 mM glycine-NaOH buffer (pH 9.6). Reactions were initiated by the addition of 110 ng of malate dehydrogenase. The concentrations of NAD used (~M) were: line 1, 8i; line 2, ]35; fine 3, 270; fine 4, 540. Inset is a replot of slopes (line 1) and intercepts (line 2).

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[NA.]-, Fig. 3. Product inhibition of malate dehydrogenase by NADH as a function of NAD concentration at a saturating concentration (3 mM) of malate. Reaction mixtures contained 45 mM glycine-NaOH buffer (pH 9.6), 41-270/LM NAD, and 50 ng of enzyme. NADH concentrations were: line 1, none; line 2, 2/LM; line 3, 2.5/~M.

15 TABLE II STUDIES OF KINETIC MECHANISM C, Competitive inhibition; NC, noncompetitive inhibition; UC, uncompetitive inhibition.

(A) Product inhibition studies Inhibitor

Variable substrate NAD (A)

NADH Oxalacetate

Malate (B)

unsaturated B

satuxated B

unsaturated A

saturated A

C NC

C UC

NC NC

None NC

(B) Dead-end inhibition studies Inhibitor

1/v

Adenosine diphosphoribose Hydroxymalonate

C UC

1/v

vs. 1/[NAD]

tively. The results of both product and dead-end inhibition studies are summarized in Table II. The results of these studies were consistent with an ordered bi-bi mechanism in which N A D is the first substrate binding to the enzyme and NADH is the second product dissociating from the enzyme.

'TABLE llI

Coenzyme specificity

NAD analog

Several pyridine nucleotide analogs altered in either the pyfidhle or purine moiety were studied with respect to their functioning as coenzymes for the H. influenzae malate dehydrogenase. Kinetic parameters for six analogs exhibiting coenzyme activity are listed in Table III. In comparison to NAD, greater deviations in kinetic parameters were observed with those dinucleotides structurally altered in the purine portion of the molecule and increasing the malate concentration had no further effect on these parameters. NADP, NMN, and the nicotinic acid, pyridylacryloamide, and 3-aminopyridine analogs were not reduced in these reactions, even at 10-times higher enzyme concentration. The addition of millimolar concentrations of AMP did not promote the reduction of nicotinamide mononucleotide.

Chemical modification of the enzyme The chemical modification of H. influenzae malate dehydrogenase was investigated in an attempt to identify functional group involvement in

vs. 1/[malate]

NC C

K m

AND Vma~ VALUES FOR NAD ANALOGS

Initial velocities were determined at 23°C in 1.0 ml reaction ~rfxtures containing 45 mM glycine-NaOH buffer (pH 9.6) and 3 mM malate. Coenzyme analog concentrations were varied from 0.1- to 2-times the observed K m values. -, no reaction detected at millimolar concentrations.

N~icotinamide adenine dinucleotide 3-Acetylpyridine ade~ne dinucleotide Thionicotinamide adenine dinucleotide 3-Pyridinealdehyde adenine dinucleotide Nicotinamide hypoxanthine dinucleotide Nicotinamide guanine dinucleotide Nicotinamidc 1, N6-ethenoadenine dinucleotide Nicotinamide mononucleotide Nicotinic acid adenine dinucleotide Nicotinamide adenine dinucleotide phosphate 3-Pyridylacryloamideadenine dinucleotide 3-Aminopyridine adenine dinucleotide

Km (#M)

Vmax (Fmol/min)

57

23

63

45

83

7

220

16

230

15

4000

3.2

1540 -

4.4 -

-

-

-

-

-

-

-

-

16

enzyme activity. Incubation of the purified enzyme with 5 mM iodoacetamide, iodoacetic acid, N-ethylmaleimide or fluorescein mercuric acetate did not decrease catalytic activity over a 60 rain period. Attempts to mo~fy lysine residues with pyridoxal 5'-phosphate were likewise unsuccessful. A time-dependent inactivation of the enzyme was, however, observed by incubation at 15°C of the enzyme at pH 7.0 with diethylpyrocarbonate. The inactivation followed pseudo-first-order kinetics and first-order rate constants determined were 0.032 rain -1, 0.082 min -~ and 0.143 rain -1 at 1.3 raM, 2.5 mM and 4 mM diethylpyrocarbonate, respectively. Although the sulfhydryl reagents studied did not inactivate the enzyme, it was of interest to determine the presence and availability of free sulfhydryl groups in the malate dehydrogenase. For this purpose, the purified enzyme was titrated with fluorescein mercuric acetate in the presence and absence of 8 M urea. The fluorescence intensity as a function of fluorescein mercuric acetate concentration measured according to Hsu and Lardy [14] is shown in Fig. 4. In t~e presence of 8 M urea, the fluorescence titration revealed the presence of two sulfhydryl groups per 61 kDa.

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Fig, 4, Fluorescence titration of malate dehydrogenase with Nuorescein mercuric acetate, Reaction mixtures contained 8 M urea, 50 m M potassium phosphate buffer (pH 7.5) and fluorescein mercuric acetate as indicated, in a total volume of 3 mL Eaz~nnae additions were: line 1, none; line 2, 1.28.10 -11 tool; line 3, 3.2.10-11 tool. Fluorescence intensity was measured at 525 mr= with excitation at 495 am.

These sulfhydryl groups could not be titrated in the native enzyme in the absence of 8 M urea. Discussion

H. influenzae as a V-factor organism exhibits the unique growth requirement for intact NAD. The inability of this and related organisms to synthesize NAD from the usual precursors and the likelihood that the availability of intact NAD in the environment of these organisms could be somewhat limiting suggest that enzymes involved in NAD metabolism in these organisms may exhibit unique properties. The H. influenzae periplasmic nucleotide pyrophosphatase observed to be responsible for hydrolysis of external NAD, providing the NMN required t'or intraceUular resynthesis of NAD, exhibits negative cooperativity and thereby significant catalysis at very low NAD concentrations [4]. As a nucleotide pyrophosphatase, this enzyme showed an unusual response to structural alterations in the purine portion of the dinucleotide, such as loss of cooperativity and substrate function. It was of interest to study other NAD-dependent enzymes in H. influenzae with respect to unusual processes involved in pyridine nuc!eodde interactions. In this respect, the malate dehydrogenase, which was previously indicated to be of importance in the partial tricarboxylic acid cycle observed in studies of glucose metabolism [8], was chosen for the present investigation. The malate dehydrogenase was isolated from H. influenzae cell sonicates and purified 109-fold through a four-step procedure involving salt fractionation, hydrophobic chromatography and affinity chromatography (Table I). The molecular weight of the purified enzyme was determined by both gel filtration and gel electrophoresis methods. Under nondenaturing conditions, the enzyme exhibited an Mr of 61000. Under denaturing conditions of SDS-polyacrylamide gel electrophoresis, an Mr of 31000 was observed, indicating a dimeric structure in the native enzyme. This is consistent with the reported dimeric structure of malate dehydrogenases from a variety of sources [15-20]; however, larger, tetrameric, malate dehydrogenases have been observed [17,21].

17

In !dnetic studies of the purified H. influenzae malate dehydrogenase, both the oxidation of malate and the reduction of oxalacetate were investigated by varying each substrzte and coenzyme at several fixed concentrations of the second reaction component, h~ all four cases studied, converging line relationstfips similar to that shown in Fig. 2 were obtained, indicating a sequential mechaifism for this enzyme. The question of whether the sequential mechanism for the malate dehydrogenase was of an ordered or random type was investigated through product and dead-end inhibition studies. The uncompetitive inlfibition by oxalacetat¢ when NAD was varied at a saturating concentration of malate (Table II) is characteristic of the ordered bi-bi mechanism. The remaining product inhibition profile was likewise consistent with this mechanism. In the studies of dead-end inhibition, the observed uncompetitlve inhibition by hydro×ymalonate at varying NAD concentrations (Table II) was also indicative of the ordered mechanism, since noncompetitive inhibition would have been expected in a random process. Therefore, the results of both types of inhibition study were consistent with an ordered bi-bi mechanism, with NAD being the first substrate bound and NADH the second product dissociating from the enzyme. Protection of the enzyme against thermal denaturation by NAD but not by malate further supports the ordered mechanism. The purified malate dehydrogenase can be classified strictly as an NAD-dependent dehydrogenase, since no activity with NADP was observed at high enzyme concentrations. The purified enzyme was observed to function with several NAD analogs structurally altered in either the purine or pyridine portion of the dinudeotide molecule (Table III). Of special interest was the functioning of the 3-acetylpyridine and thionicotinamide analogs, both of which were observed previously to support growth of H. influenzae in the absence of NAD [4]. Structural alterations in the purine moiety of the dinucleotide, as in the case of the guanine and ethenoadenine derivatives, resulted in a significant decrease in the functioning of these dinucleotides as coenzymes. Such large effects from purine alterations were also observed in reactions catalyzed by the H. influenzae nucleotide pyrophosphatase

[4]. NMN was completely inactive as a coenzyme, alone and in the presence of adenosine derivatives. NMN was included in these studies, since the reduction of NMN has been observed in a few dehydrogenase reactions [22-25]. Since NMN is a product of the H. influenzae periplasmic nucleotide pyrophosphatase reaction [4] and is known to support growth of the organism, the inability of NMN to function with the malate dehydrogenase confirms the need for intracellular resynthesis of NAD tO meet the coenzymic requirement of at least one metabolically important dehydrogenase. The results of the coenzyme specificity studies and those establishing order of addition of substrates and dissociation of products, provide important information for future studies of selective inhibition of the malate dehydrogenase. It is of interest to correlate inhibition of growth of H. influenzae with selective hflfibition of specific enzymic processes. For example, inhibition of growth by AMP and ADP appears related to the inhibition of the periplasmic nucleotid¢ pyrophosphatase [4]. However, the very potent growth inhibition by 3-aminopyfidine adenine dinucleotide observed in these studies [4] remains unclarified. Preliminary studies of this analog witi~ the malate dehydrogenase (data not shown) indicate a very ineffective inhibition and presumably not a l'~c~or in the effective growth inhibition by this dinucleotide. The purified H. influenzae malate dehydr'.~genase was inactivated by diethylpyrocarbonate in reactions following pseudo-first-order kinetics. Under the conditions used in these reactions, some specificity for the modification of histidine residues has been noted in the inactivation of various other dehydrogenases [26-30]. Essential histidine residues have been reported in studies of a number of malat¢ dehydrogenases of various origins [26,29,31-35]. The inactivation of the H. influenzae malate dehydrogenase by diethylpyrocarbonate and the absence of inactivation by the other functional group reagents studied suggests the possible involvement of a histidine residue in reactions catalyzed by the H. influenzae enzyme. Once sufficient purified enzyme is available for analysis, this possibil;ty will be investigated further. The results of the molecular weight, kinetic mechanism, coenzyme specificity and functional

18

group modification studies of H. influenzae malate dehydrogenase were consistent with those observed with many other malate dehydrogenases. The relatively poor utilization of NAD analogs with minor structural alterations in the purine portion of these dinucleotides appears more pronounced in the case of the Haemophilus enzyme. Acknowledgements These studies were supported by Research Grant DMB 8508930 from the National Science Foundation. References 1 Lwoff, A. and Lwoff, M. (1937) Proc. R. Soc. Lond. Biol. Sci. 122, 352-359. 2 Schlenck, F. and Gingrich, W. (1942) J. Biol. Chem. 143, 295-296, 3 Ginsrich, W. and Schlenck, F. (1944) J. Bactertol. 47, 535-550. 4 Kahn, D.W. and Anderson, B.M. (1986) J. Biol. Chem. 261, 6016-6025. 5 Smith, A.L. (1979) N. Engl. J. Med. 301, 155-156. 6 Gunn, B.A., Woodall, J.B., Jones, J.F. and Thornsberry, C. (1974) Lancet fi, 845. 7 Syriopoulou, V., Scheifele, D., Smith, A.L.o Perry, P.M. and Howil V. (1978) J. Pediatr. 97, 421-424. 8 Tuyau, J.E., Sims, W. and Williams, R.A.D. (1984) J. Gen. Microbiol. 130, 1787-1793. 9 Anderson, B.M., Yost, D.A. and Anderson, C.D. (1986) Methods Enzymol. 122, 173-181. 10 Yost, D.A. and Anderson, B.M. (1982) J. Biol. Chem. 257, 767-772. 11 Bradford, M.M. (i976) Anal. Biochem. 72, 248-254. 12 Davis, BJ. (1964) Ann. N.Y. Acad. Sci. 121, 404-427. 13 Weber, K. and Osborn, M (1969) J. Biol. Chem. 244, 4406-4412.

14 Hsu, R.Y. and Lardy, H.A. (1967) J. Biol. Chem. 242, 527-532. 15 Gerding, R.K. and Wolfe, R.G. (1969) J. Biol. Chem. 244, 1164-1171. 16 Shore, J.D. and Chakrabarti, S.K. (1976) Biochemistry 15, 875-879. 17 Murphy, W.H., Kitto, G.B., Everse, J. and Kaplan, N.O. (1967) Biochemistry 6, 603-609. 18 Ohshima, T. and Sakuraba, H. (1986) Biochim. Biophys. Acta 869, 171-177. 19 Rudolph, R., Fuchs, 1. and Jaenicke, R. (1986) Biochemistry 25, 1662-1669. 20 Wali, A.S. and Mattoo, A.K. (1984) Can. J. Biochem. 62, 559-565. 21 Yoshida, A. (1965) J. Biol. Chem. 240, 1113-1117. 22 Fisher, H.F. and McGregor, L.L. (1969) Biochem. Biophys. Res. Commun. 34, 627-632. 23 Sicsic, S., Durand, P., Langrene, S. and Le Goffic, F. (1984) FEBS Lett. 176, 321-324. 24 Shimizu, M., Suzuki, T., Hosokawa, Y., Nagase, O. and Abiko, Y. (1970) Biochim. Biophys. Acta 222, 307-319. 25 Sicsic, S., Durand, P., Langrene, S. and Le Goffic, F. (1986) Eur. J. Biochem. 155, 403-407. 26 Holbrook, J J and Ingrain, V.A. (1973) Biochem. J. 131, 729-738. 27 Meyer, S.E. and Chromartie, T.H. (1980) Biochemistry 19, 1874-1881. 28 Collier, G.E. and Nishimura, J.S. (1979) J. Biol. Chem. 254, 10925-10930. 29 lijima, S., Oh, M.-J., Saiki, T. and Beppu, T. (1986) J. Biochem. 99, 1667-1672. 30 Topham, C.M. and Dalziel, K. (1986) Eur. J. Biochem. 155, 87-96. 31 Birktoft, J.J. and Banaszak, L.J. (1983) J. Biol. Chem. 258, 472-482. 32 Anderton, B.H. (1970) Eur. J. Biochem. 15, 562-567. 33 Gregory, E.M., Rohrbach, M.S. and Harrison, J.H. (1971) Biochim. Biophys. Acta 243, 489-497. 34 Lodola, A., Parker, D.M., Jeck, R. and Holbrook, J.J. (1978) Biochem. J. 173, 597-605. 35 Holbrook, J.J., Lodola, A. and lllsley, N.P. (1974) Biochem. J. 139, 797-800.