THE JOURNAL OF BIOLOGICAL CHEMISTRY 0 1993 by The American Society for Biochemintry and Molecular Biology, Inc

Vol. 268, No. 34, Issue of December 5, pp. 26004-26010. 1993 Printed in U.S.A.

A New Paradigm for Biochemical Energy Coupling SALMONELLA TYPHIMURIUM NICOTINATE PHOSPHORIBOSYLTRANSFERASE*

(Received for publication, February 23, 1993)

Alexander VinitskyS and Charles Grubmeyersll From the Department of Biology, New York University, New York, New York 10003 and the §Department of Biochemistry, Temple University School of Medicine, Philadelphia, Pennsylvania 19140

ThepncB gene of Salmonella typhimurium was used to develop an overexpression system for nicotinate phosphoribosyltransferase (NAPRTase, EC 2.4.2.11), which forms nicotinate mononucleotide (NAMN) and PPI from nicotinate and a-~-S-phosphoribosyl- l-py- rophosphate (PRPP). NAPRTase hydrolyzes ATP in 1:l molar stoichiometry to NAMN synthesis. Hydrol- ysis of ATP alters the ratio of products/substrates for the reaction nicotinate + PRPP G= NAMN + PPI from its equilibrium value of 0.67 to a steady-state value of 1100. The energy for the maintenance of this ratio must come from ATP hydrolysis. However, in contrast to other ATP-utilizing enzymes, when all ATP is hy- drolyzed the unfavorable product/substrate ratio col- lapses. ATP/ADP exchange results suggest that the overall reaction involves a phosphoenzyme (E-P) aris- ing from E . ATP. K,,, values for nicotinate and PRPP each decreased by 200-fold when ATP was present to phosphorylate the enzyme. PPi stimulated the ATPase activity of the enzyme to V,, values, suggesting that PPI formation during catalysis provides a trigger for cleavage of the putative E-P in the overall reaction and regenerates the low affinity form of the enzyme. A model is presented in which alternation of high and low affinity forms of NAPRTase provides a ”steady- state” coupling between ATP hydrolysis and NAMN formation.

Free energy coupling in biological systems has been the subject of many reviews and speculation. In many cases where ATP hydrolysis drives a chemical reaction, the linkage is mediated by phosphorylated chemical intermediates; for ex- ample the reactions catalyzed by glutamine (Midelfort and Rose, 1976) and CTP synthetases (von der Saal et al., 1985). Such chemically coupled systems are relatively well-explored and understood since coupling is intrinsic to the chemical process. In contrast, in the case of ATP-dependent transport- ers, it is believed that the linkage between the chemical

* This research was supported by Grant DMB-9103029 from the National Science Foundation. Acquisition of a Pharmacia fast protein liquid chromatography system was made possible by a grant from the New York University Research Challenge Fund. The costs of publi- cation of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertise- ment” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

$ This work was in partial fulfillment of a doctoral dissertation (Biology Department, New York University). Current address: Dept. of Pharmacology, Mount Sinai School of Medicine, 1 Gustav Levy Place, New York, NY 10029.

11 To whom correspondence should be addressed Dept. of Biochem- istry, Temple University School of Medicine, 3400 N. Broad St., Philadelphia, PA 19140. Tel.: 215-221-4495; Fax: 215-221-7536.

processes of ion transport and ATP hydrolysis is less direct and may involve conformational changes (Tanford, 1983). These systems have proven difficult to study and also to understand (Jencks, 1980). In many cases the enzymes are membrane-bound, frequently very complex structurally, and the transported ion is chemically unchanged by the reaction, making it difficult to determine stoichiometries of transport and ATP hydrolysis. The questions remaining to be answered with such systems are both important and timely: how is ATP hydrolysis prevented in the absence of transport? How is transport prevented in the absence of ATP hydrolysis? Can one identify specific steps which constitute the use of ATP hydrolysis energy? These questions have increased in their timeliness with the discovery of a multitude of other systems that apparently proceed via conformational coupling, such as ATP-dependent proteases (Armon et al., 1990), chaperonin groEL (Martin et al., 1991), recA (West, 1992), periplasmic bacterial transport systems (Ames, 1990), the interaction of helicases (Matson and Kaiser-Rogers, 1990), and gyrases (Cullis et al., 1992) with DNA, and the role of ATP hydrolysis in expelling RNA from rho factor (Stitt, 1988).

In this work we have investigated a soluble monomeric enzyme, nicotinate phosphoribosyltransferase (NAPRTase; EC 2.4.2.11),’ whose coupling of NAMN synthesis and ATP hydrolysis provides a novel and important model for energy coupling. NAPRTase catalyzes the formation of NAMN and PPi from nicotinate and a-D-5-phosphoribosyl-1-pyrophos- phate (PRPP). The reaction catalyzed by NAPRTase serves a 2-fold role in uiuo: to recycle the pyridine moiety of pyridine nucleotides derived from NAD-degrading reactions such as that of DNA ligase or poly-ADP-ribose synthetase (Ferro and Olivera, 1987) and to enable cells to utilize external sources of NA for the synthesis of pyridine nucleotides.

The enzyme has been purified previously in low yield from several sources. Recently, our laboratory has cloned the Sal- monella typhimurium pncB gene, coding for NAPRTase, and provided the sequence of the enzyme (Vinitsky et al., 1991). The sequence of Escherichia coli pncB was also recently presented (Wubbolts et al., 1990).

Although the reaction catalyzed by NAPRTase is in every other respect a typical phosphoribosyltransferase reaction, the activities of NAPRTases from yeast and bacterial sources were claimed to be absolutely dependent on the presence of ATP (Imsande, 1964; Ogasawara and Gholson, 1966). With the use of isotopically labeled ATP, Honjo et al. (1966) dem- onstrated hydrolysis of ATP stoichiometric with synthesis of NAMN, with Pi product arising from the y position of ATP.

The abbreviations used are: NAPRTase, nicotinate phosphori- bosyltransferase; ATPrS, adenosine 5’-y-thio-triphosphonucleotide; NA, nicotinic acid; NAMN, nicotinate mononucleotide; PAGE, poly- acrylamide gel electrophoresis; PRPP, a-5-phosphoribosyl-1-pyro- phosphate; DTT, dithiothreitol.

26004

NAPRTase Energy Coupling 26005

NAPRTases from beef liver and human erythrocytes are also stimulated by ATP but are measurably active in its absence (Imsande and Handler, 1961; Smith and Gholson, 1969; Nie- del and Dietrich, 1973). For both of these mammalian en- zymes, ATP decreases the K,,, values for both substrates approximately 10-fold and does not affect the V,, of the reaction detectably. When present, ATP is hydrolyzed stoi- chiometrically with formation of NAMN (Niedel and Die- trich, 1973). This ATP hydrolysis may be unique to NA- PRTase among the PRTases, although one report suggests that the less well characterized nicotinamide PRTase may exhibit a similar usage of ATP (Kasarov and Moat, 1973).

The coupling between the ATP hydrolysis and NAMN synthesis reactions catalyzed by NAPRTase presents a unique and previously unexploited bioenergetics problem. Since eight other phosphoribosyltransferases catalyze their reactions without the use of ATP hydrolysis (in ATP-PRTase, ATP is the phosphoribosylated substrate), it is unlikely that ATP participates chemically in the reaction. Thus, NAPRTase may present an example of an ATPase reaction conformationally coupled to a biosynthetic reaction and provides an exciting model for similar reactions in which ATP hydrolysis is linked with a cycle of affinity changes toward a second substrate.

EXPERIMENTAL PROCEDURES

Materials-["C]Nicotinic acid, obtained from Sigma, was used directly for enzyme assays. For measurement of product/substrate ratios, the compound was first purified by paper chromatography in 8l:l isopropanokl N NH40Hwater. Three radioactive spots were localized by autoradiography, one at the origin, a second migrating with an RF of 0.35, and a third co-migrating with authentic nicotinic acid at an RF of 0.5. The NA spot, containing about 98% of the radioactivity, was eluted with HZO.

Bacterial growth media were from Difco. Biochemicals were from Sigma, and lactate dehydrogenase and pyruvate kinase were from Boehringer Mannheim. NaCN and other chemicals were from Fisher.

Overexpression of pwB-Previously, pC18 carrying a 4.7-kilobase pair insert of S. typhirnuriurn chromosomal DNA was shown to confer the pncB+ phenotype on the nudA pncB host JF1483 and was used in DNA sequencing studies (Vinitsky et aL, 1990). To induce higher expression of pncB, a subclone was made by restricting pC18 with SalI followed by religation to give pSC186 (Fig. 1). pSC186 was also found to confer a pncB+ phenotype on JF1483. pSC186 was used to transform the S. typhimurium strain RM926 (SGSC452, donated by Russell Maurer, Case Western Reserve; RM926 has a wild-type chromosomal pncB gene). RM926 carrying pSC186 is designated VG003.

Purification of NAPRTase-A purification was developed starting

p o R 1 0.00

pSCl86 pUCl9 6.10 Kb

FIG. 1. Map of pSCl86.

10

with 45 g of S. typhimurium strain VG003. Bacteria were grown in shaker cultures overnight in 12 liters of tryptone broth (Davis et al., 1980) containing 50 pg/ml ampicillin at 37 "C. Cells were harvested by centrifugation and resuspended in 2 ml/g of cells of 200 mM sodium phosphate (Napi) buffer, pH 8.0, containing 1 mM phenyl- methylsulfonyl fluoride.

Resuspended cells were disrupted by two 2-min pulses from a Heat Systems W185 sonicator at full power. Debris was removed by cen- trifugation at 15,000 X g for 20 min. To the supernatant was added 50 pl/ml of 10% Polymin-P (polyethyleneimine, Sigma product P- 3143, brought to pH 8.0 with HCI). The resultant suspension was centrifuged to remove the Polymin-P-nucleic acid complex.

The supernatant was brought to 30% saturation with ammonium sulfate, the precipitate removed by centrifugation (10 min, at 10,000 X g, at 4 "C), and the supernatant brought to 50% saturation with (NH&SO, and centrifuged as above. The resultant pellet containing 900 units of NAPRTase activity was redissolved in 10 ml of 200 mM Napi buffer, containing 10% glycerol. Coupled spectrophotometric ATPase assays at this point showed a high basal level ATPase activity in the absence of NA and additional NA-dependent activity. Salt concentration was reduced by gel-filtration chromatography on a Sephadex G-50 column (2.5 X 20 cm) preequilibrated with Buffer A, consisting of 10 mM Tris phosphate, pH 8.0, 1 mM DTT, 0.2 mM MgS04, 0.1 mM EDTA, and 10% glycerol.

The protein sample was applied to a DEAE-Toyopearl650 column (2.5 X 28 cm), preequilibrated with Buffer A, and the column was washed with 100 ml of Buffer A. Using a Pharmacia LKB Biotech- nology Inc. fast protein liquid chromatography apparatus, the enzyme was eluted with a linear gradient of 0-20% Buffer B (Buffer A containing 0.5 M NazS04) over 400 ml. The enzyme eluted at approx- imately 35 mM NazS04. Fractions containing enzymatic activity were pooled and precipitated with 60% saturated ammonium sulfate. 400 units were recovered at this stage, and negligible basal ATPase activity was observed.

The enzyme suspension was centrifuged, redissolved in Buffer A, and applied to a Cibacron blue F3GA-TSK affinity column (1.6 X 36 cm). The column was washed with 75 ml of Buffer A. The elution gradient was 0-100% Buffer B in 300 ml. Active enzyme eluted at 0.1 M Na2S04. After reprecipitation with 60% saturation (NH4),S04. 350 units of NAPRTase activity were present.

After centrifugation, the precipitated enzyme was redissolved in 2 ml of Buffer C (Buffer A plus 100 mM NazS04), applied to a Sephacryl HR200 gel-filtration column (2.5 X 75 cm), and eluted with Buffer C. The fractions were pooled, precipitated with 60% saturated ammo- nium sulfate, and resuspended in 200 mM Napi, pH 8.0, containing 10% glycerol. 300 units of NAPRTase activity were recovered, rep- resenting 33% of the starting activity. The protein was stored as a precipitate in 60% saturated ammonium sulfate, containing 200 mM NaPi, pH 8.0, 5 mM DTT, and 10% glycerol. The activity of the enzyme was stable for a t least 2 months under these conditions.

Enzyme Assays-Three assays for NAPRTase were employed. The ["CINA label transfer procedure of Preiss and Handler (1958) was performed in a 0.1-ml total volume at 30 "C. The reaction mixture contained 200 mM monopotassium glutamate, 20 mM Tris base, pH 8.3, with 3 mM ATP, 1 mM PRPP, 5 mM D m , 7 D M MgSO,, 0.1 mM ["CINA (specific activity, lo6 cpm/nmol). The reaction was started by addition of enzyme and allowed to proceed for 5 min, after which 10-p1 samples were applied to Whatman No. 3MM filter paper pre- spotted with 30% perchloric acid to quench the reaction. NA and NAMN were separated by developing with 3:7 1 M ammonium acetate, pH 5.0:95% ethanol. The chromatogram was cut into 10 equal size strips, and the radioactivity on each strip was determined by liquid scintillation. To study the uncoupled NAMN synthesis reaction, ATP was omitted, and MgSO4 was decreased to 4 mM.

A colorimetric assay based on the complexation of NAMN with cyanide (Packman and Jakoby, 1967) followed the label transfer protocol, except that 0.3-ml reaction mixtures were quenched by addition of 0.8 ml of 7 M NaCN. The concentration of the NAMN. CN complex was then determined at 315 nm. The extinction coeffi- cient of the NAMN.CN complex was determined to be 5.1 mM".

A coupled spectrophotometric assay was based on the previously unexploited ATPase activity of NAPRTase. The assay was carried out in 1-ml quartz cuvettes at 30 "C. The reaction mixture contained 200 mM monopotassium glutamate, 20 mM Tris, pH 8.3 with 5 mM DTT, 3 mM phosphoenolpyruvate, 3 mM ATP, 7 mM MgS04, 0.3 mM NADH, 1 mM PRPP, 1 mM NA, 50 pg pyruvate kinase, 25 pg lactate dehydrogenase, and 0.005-0.02 units of NAPRTase. The activity was followed in the spectrophotometer as the loss of absorbance at 340

26006 NAPRTase Energy Coupling

FIG. 2. Analvsis of thd Durification Drocedure bv SDS- PAGE. A , crude homogenate; B, homogenate after ammonium sulfate fractionation; C, pooled fractions containing NAPRTase activity after anion exchange chromatography; D, pooled fractions containing NA- PRTase activity after dye-affinity chromatography.

nm. This assay could not be used with crude extracts since they contained nonspecific ATPase activity, obscuring the coupled ATP- ase activity of NAPRTase.

T o measure the ATP/ADP exchange, reactions were carried out in 200 mM monopotassium glutamate, 20 mM Tris base, pH 8.3 with 4 mM MgSO,, 5 mM DTT, at 30 "C. ADP (containing lo5 cpm of ['HI ADP) and ATP were present in the indicated concentrations as equimolar mixtures with MgSO,. The reactions were started by addition of 6.5 pg of NAPRTase. Samples of the reaction (1 pl) were taken before the addition of enzyme, and after 1 min, applied to polyethyleneimine-cellulose plates that had been prespotted with 1 pl of solution containing 50 mM ADP and 50 mM ATP as carrier. The chromatograms were developed with 1 M LiCl in 0.5 M formic acid. The nucleotides were visualized on the dried plate with short wave uv light and gave RF values of 0.25 (ATP) and 0.5 (ADP). The spots were scraped into liquid scintillation vials and 0.3 ml of 1 N HCl added. After incubation to elute nucleotides, the radioactivity was determined by liquid scintillation after addition of 3.0 ml of Liquiscint (National Diagnostics). Rates of the exchange reaction were calculated using the equations of Purich and Allison (1983) and are expressed in micromoles of ATP formed per min. mg protein.

Protein was measured by absorbance at. 280 nm. An extinction coefficient for a 1 mg/ml solution of NAPRTase E2wnm = 1.27, determined by correlating the A2mnm with biuret protein assay, was used in this work. A unit of NAMN synthesis activity is defined as that amount catalyzing the formation of 1 pmol of NAMN/min. When ATP hydrolysis activity is being measured, a unit of ATPase activity is that amount catalyzing the production of 1 pmol of ADP/ min.

RESULTS

Overexpression and Purification-Cloning of the pncB gene on a multicopy pUC19 plasmid allowed for 1000-fold overex- pression compared to wild-type S. typhimurium LT-2.* The purification procedure yielded about 80 mg of homogeneous protein from 45 g cells (Fig. 2). Anion exchange chromatog- raphy was a particularly useful step. Edman degradation on the purified enzyme, as well as on tryptic peptides of NA- PRTase, was used previously to obtain sequence data (Vinit- sky et al., 1991) and also suggested a high degree of homoge- neity. A subunit M, of 45,000 was calculated based on the mobility of the protein on SDS-PAGE. This value is in good agreement with the DNA sequence-deduced M , of 45,512 (Vinitsky et al., 1991).

Reactions and Stoichiometry-The rates of the ATPase reactions catalyzed by NAPRTase are presented in Table I. Previous work suggested a Pi requirement for NAPRTase (Kosaka et al., 1971). Following studies by Leirmo et al. (1987)

A. Vinitsky, unpublished data.

TABLE I Coupled and uncoupled ATP hydrolysis

Spectrophotometric assays were carried out in 200 mM monopo- tassium glutamate, 20 mM Tris base, pH 8.3, with 7 mM MgSO4, 5 mM DTT, and 3 mM ATP, unless indicated otherwise. Reactions were started with 3 pg of NAPRTase.

ATP hydrolysis

Additives Rate unitslmg

None 0.02 1 mM NA, 1 mM PRPP 3.00 1 mM PPi 3.00 0.1 mM PRPP ND" 0.1 mM NA 0.02 0.1 mM NAMN 0.03 1 mM PPi, 1 mM PRPP 0.02 1 mM P N P 2.83 1 mM PCP 0.02

a ND, not detectable, rate below 0.005 units/mg.

on the anion content of bacteria, we successfully replaced Pi with monopotassium glutamate. In complete assay mixture the ATPase activity of NAPRTase was 3.0 units/mg. In the absence of substrates for the NAMN synthesis reaction, the ATP hydrolysis reaction proceeded a t a slow but measurable rate of 0.02 units/mg. This apparent uncoupled ATPase ac- tivity of NAPRTase might be the result of a contaminating ATPase or some denatured form of NAPRTase. However, the uncoupled ATP hydrolysis activity was reduced to less than 0.005 units/mg by the presence of 0.1 mM PRPP. The addition of NA or NAMN to uncoupled ATPase assays had no appar- ent effect on the rate of the reaction. In contrast, the addition of NaPPi to uncoupled ATP hydrolysis assays activated an ATPase activity equivalent to that observed under V,,, con- ditions for the coupled reaction. NaP(NH)P, but not NaP(CH2)P, could stimulate a similar uncoupled ATPase activity. The PPi-stimulated uncoupled ATPase activity was severely inhibited by the presence of 1 mM PRPP.

Initial observations suggested that S. typhimurium NA- PRTase catalyzed NAMN synthesis a t rates about the same as ATP hydrolysis when assayed under standard conditions and about 10% of that rate when assayed in the absence of ATP. The existence of this uncoupled NAMN synthesis ac- tivity was in contrast to the yeast enzyme, for which uncou- pled NAMN synthesis was not observed (Hanna et al., 1983). To learn about the relationship between NAMN synthesis and ATP hydrolysis the coupled and uncoupled reactions were further characterized.

Although separate observations had suggested that ATP hydrolysis and NAMN synthesis proceeded at about the same rate, it was of interest to determine the stoichiometry of ATPase and NAMN synthesis reactions under coupled con- ditions. Fig. 3 shows that the two reactions proceed at a 1:l ratio throughout the early reaction course.

Coupling of NAMN Synthesis and ATP Hydrolysis-From the stoichiometry results of Fig. 3 it was not clear whether the reactions of NAMN synthesis and ATP hydrolysis are linked, or fortuitously proceeded at the same rate. If the two reactions are in fact coupled, then the hydrolysis of ATP would be expected to drive the NAMN synthesis reaction. To explore this possibility we determined the final ratio of the substrates and products of the NAMN synthesis reaction in the absence and presence of ATP.

The K,, for the uncoupled NAMN synthesis reaction was readily evaluated using the graphical approach of Purich and Allison (1980) and employing the cyanide complexation assay to determine the amount of NAMN. The results (Fig. 4)

NAPRTase Energy Coupling 26007

2 1 I

I / 1.5

= 0.5 I E c 0 ' I

I I I 0 2 4 6 8 1 0

Time (mln) FIG. 3. Stoichiometry of ATP hydrolysis and NAMN syn-

thesis. The products of the coupled reaction were quantitated at various points over the course of the reaction. The reaction mixture contained 2 mM PRPP, 1 mM NA, 3 mM ATP, 6 mM magnesium acetate, 5 mM DTT, 50 mM sodium phosphate, pH 8.0, in a total volume of 5 ml. The enzyme (60 pg) was added to the reaction mixture prewarmed to 30 "C. At times shown an 810-pl sample was taken and the enzymatic reaction quenched on ice with the addition of 10 pmol of EDTA. Samples of the quenched reaction were used in the coupled enzyme assay to determine the amount of ADP produced and for the determination of NAMN in the reaction by the cyanide complexation assay. Results of a representative experiment are shown.

-0.2 , I

0.1 0.6 1.1 1.6 2.1 2.6

[substrates]/[products]

FIG. 4. Determination of equilibrium constant for uncou- pled NAMN synthesis. Change in NAMN concentration is ex- pressed as the change in A 3 1 6 ~ . The ratio of NA to NAMN was varied from 0.1 to 2.6 as indicated, and PRPP and PPI were each 0.5 mM.

indicated a value of 0.67 for the ratio [NAMN][PPi]/[NA] [PRPP] under our conditions of pH and Mg2+ concentration. This value is close to that of 0.11 determined for the orotate PRTase reaction (Bhatia et al., 1990).

The same measurement was also made under conditions where ATP was present and being hydrolyzed. The NAMN synthesis reaction was followed using the radiolabel transfer procedure and separating ["GINA and ["CINAMN using thin layer chromatography. For these experiments, commercial [14C]NA was not of adequate purity, and [14C]NA repurified from the Sigma product was used as the substrate for the ATP-coupled NAMN synthesis (Table 11). When a plateau in NAMN conversion had been reached (10-15 min), 0.4% of the starting NA apparently remained. This value allowed the

TABLE I1 Conversion of purified ["GINA to P'CINAMN in the presence of

ATP Reaction was carried out in 200 mM monopotassium glutamate, 20

mM Tris base, pH 8.3 with 5 mM DTT, 0.5 mM PRPP, 2.0 mM sodium pyrophosphate, 0.05 mM ["CINA (lo6 cpm/nmol), 3 mM ATP, 8.5 mM MgSO, in 0.1 ml at 30 "C, with 0.7 pg of NAPRTase.

Time NAMN NA rnin n m l

0 0 4.102 1 0.661 3.843 2 1.235 3.141 5 3.170 1.421

10 4.211 0.017 15 4.232 0.017 20 4.241 0.022

calculation of a product-to-substrate ratio for coupled NAMN synthesis of 1100.

There still remained the question of whether the small amount of radioactive material which chromatographed as NA at the end of reaction (Table 11) was NA and not an impurity or breakdown product produced in the reaction mixture. If this were the case, then the product/substrate ratio of 1100 would underestimate the true value. The radio- active material was eluted from the chromatogram with water and lyophilized. The lyophilized material was dissolved in 200 mM monopotassium glutamate, 20 mM Tris base, pH 8.3 with 1 mM PRPP, 5 mM DTT, 8.5 mM MgSO4, and 3 mM ATP. Unlabeled NA was added to the redissolved material to a final concentration of 0.05 mM. Upon addition of enzyme, the reaction proceeded to a nearly complete (95%) conversion of [14C]substrate to NAMN. This result demonstrated that the radioactive material behaving as [14C]NA was in fact NA and not a contaminant.

The value of 1100 obtained for the ratio [NAMN][PPi]/ [NA] [PRPP] is not a K,, since the calculation excluded ATP and its breakdown products ADP and Pi. Instead, the ratio is a measure of the extent of the NAMN synthesis reaction in the presence of ATP and during ATP hydrolysis. Comparison of the K, for NAMN synthesis in the absence of ATP with the extent of the reaction under ATP hydrolysis conditions demonstrates a shift of 1640-fold, representing a difference of about 4.4 kcal/mol. Since the shift can only be caused by ATP hydrolysis, it is clear that about 4.4 kcal/mol of energy from ATP hydrolysis has been captured in the form of a thermo- dynamically unfavorable product/substrate ratio.

Reversal of NAMN Accumulation following ATP Exhaus- tion-With ion-pumping ATPases the coupling between pumping and ATP hydrolysis is very tight, and neither ion flow in the absence of ATP, nor ATP hydrolysis in the absence of ion pumping, is detected (Trevorrow and Maynes, 1984; Stahl and Jencks, 1987). In the case of NAPRTase this is not so, as shown by the ATPase levels measured in Table I. This leakiness of coupling is seen dramatically in Fig. 5, in which ATP hydrolysis was allowed to proceed to completion. The unfavorable product/substrate ratio for NAMN synthesis quickly reversed under these conditions. The leakiness of coupling was also manifested in a greatly altered reaction stoichiometry as product levels increased. For example in Fig. 5, in the first 5 min of the reaction shown, 260 nmol ATP was hydrolyzed, whereas no net conversion of NA to NAMN has occurred. Although these results follow from the obser- vation of the PPi-activated ATPase activity of NAPRTase, they indicate that energy coupling in NAPRTase is not an equilibrium phenomenon, but rather a steady state in which

NAPRTase Energy Coupling 26008 300

p 2 0 0

s E - Q)

E 100

0

1 .5

1

9 z Q) - C

0.5

0

0 5 1 0 1 5 2 0 Time (min)

FIG. 5. Reversal of NAMN synthesis following hydrolysis of ATP. The reaction was carried out in 200 mM monopotassium glutamate, 20 mM Tris base, pH 8.3, with 5 mM DTT, 0.05 mM [“C] NA, 0.5 mM PRPP, 2 mM sodium pyrophosphate, 3 mM ATP, and 8.5 mM MgSO, in a total volume of 0.2 ml at 30 “C. The reaction was started by the addition of 120 pg of NAPRTase. The zero time point was taken approximately 1 min after enzyme addition, at which time greater than 95% of the NA had been converted to NAMN, and 50% of the ATP was converted to ADP. Vertical axes show NA and ATP content calculated for the 200-pl reaction mixture. At each time point shown, ATP concentration was assayed by the coupled enzyme assay. Interconversion of [“CINAMN and [I4C]NA was monitored chro- matographically (see “Experimental Procedures”). Results shown are from a representative experiment.

ATP hydrolysis maintains an unfavorable product/substrate ratio.

Coupling Supported by ATPyS-The compound ATPyS was tested for its ability to support a coupled NAMN synthesis reaction. In the presence of 3 mM ATPyS, the rate of NAMN synthesis was 0.15 units/mg, about one-half the rate observed in its absence. Stoichiometry studies showed that ATPyS hydrolysis occurred with each cycle of NAMN synthesis dur- ing the early reaction time course (not shown). When ATPyS was present in reaction mixtures, the final ratio of products to reactants was 1000, very similar to that in reactions sup- ported by ATP. Thus, ATPyS hydrolysis is coupled to NAMN synthesis, and a slow step in the reaction causes the slowing of the entire sequence.

ATPIADP Exchanges Catalyzed by NAPRTase-The yeast NAPRTase reaction is thought to proceed via a phosphate- containing enzyme form, which has been isolated previously (Kosaka et al., 1977) but whose covalency was not determined. We have performed studies of ATP/ADP exchanges in the absence of substrates for NAMN synthesis to learn whether a catalytically relevant phosphoenzyme exists in S. typhimu- rium NAPRTase. Table I11 shows the results of the kinetic study of the exchange reaction. The maximal velocity was 3.6 pmol of ATP per min. mg, and substrate concentrations re- quired for half-maximal exchange activity were 0.22 mM for ATP and 0.54 mM for ADP. The enzyme is highly selective in that excess ATP and excess ADP did not detectably inhibit the exchange (i.e. ATP does not bind well to E-P and ADP binds only weakly to E). Since the exchanges are performed under conditions where no net catalysis occurs, the near identity of the rate of exchange with that for overall catalysis suggests that a step in the formation of the phosphoenzyme

TABLE I11 Kinetics of PHIADPIATP exchange

mate, 20 mM Tris base, pH 8.3, with 4 mM MgS04, 5 mM DTT, at The exchanges were carried out in 200 mM monopotassium gluta-

30 “C. The reactions were started by addition of 6.5 pg of NAPRTase. ADP (containing lo6 cpm of [3H]ADP) and ATP were added as equimolar mixtures with MgSO4. For the determination of the Kc for ATP, ADP was held constant at 2 mM, and ATP varied between 0.13 and 0.90 mM. For the determination of the K, for ADP, ATP was held constant at 0.9 mM, and ADP varied between 0.33 and 4.0 mM.

Inhibitor K, V ,

P‘U units/mg ATP 215 * 40 ADP

3.6 -t 0.24 536 * 90 3.6 k 0.24

may be rate-limiting for the overall reaction. Such a step could be E-P formation from the binary E. ATP complex, ADP release from the E-P.ADP complex, or a protein con- formational change. PRPP was an excellent inhibitor of the ATP/ADP exchange. The inhibition was competitive versus ADP (Kr = 1 pM, data not shown).

ATP-induced Changes in K , Values-To determine the possible effects of enzyme phosphorylation on substrate affin- ities, we performed kinetic studies on both the ATP-coupled and uncoupled NAMN synthesis reactions. The coupled re- action was studied with the radiolabel transfer assay and revealed K, values of 1.5 ? 0.3 p~ for NA (at 0.5 mM PRPP) and 22 f 3.3 pM for PRPP (at 0.1 mM NA). In the case of the uncoupled reaction, poor substrate affinities made it conven- ient to employ the cyanide complexation assay. A K , value of 0.29 f 0.07 mM for NA was observed in the presence of 4 mM PRPP, and a K , of 4.5 f 2.2 mM for PRPP was calculated from measurements carried out in the presence of either 1 or 0.1 mM NA. Vmax values were 2.9 ? 0.1 units/mg for the coupled reaction and 0.29 f 0.07 units/mg for the uncoupled reaction.

The kinetics of PPi-stimulated ATPase activity were also studied. Linear double-reciprocal plots were observed (not shown). The K , for PPi was 80 p~ and the V,, of PPi- stimulated ATPase was 3 units/mg.

DISCUSSION

S. typhimurium NAPRTase is a valuable and unique tool for the investigation of energy coupling in biochemical sys- tems. Here we discuss our findings in two parts, first reviewing the enzymology of NAPRTase and second discussing the bioenergetic consequences.

The pncB gene had previously been cloned from a X library of the S. typhimurium chromosome and the nucleic acid sequence determined (Vinitsky et al., 1991). The overpro- duction of the protein was accomplished in this work by coupling the pncB gene to the lac promoter. Purification of NAPRTase to homogeneity was readily accomplished using high-resolution ion exchange on TSK-DEAE resin, followed by chromatography on Cibacron blue-agarose. In some cases additional purification was achieved with gel filtration on Sepharose HR200. The enzyme was judged homogeneous by

Several workers have previously purified NAPRTase to homogeneity from a variety of sources, with specific activities similar to those reported here. The function of the enzyme in NAD recycling suggests that it is ubiquitous. Key observations on NAPRTase have included its ATP requirement, which was documented with the yeast (Kosaka et al., 19711, mammalian (Imsande and Handler, 1961) and Bacillus subtilis (Imsande, 1964) enzymes, the existence of an E-P whose covalency is not yet determined (Kosaka et al., 1977), and its kinetic

SDS-PAGE.

NAPRTase Energy Coupling 26009

mechanism, which was delineated by studies on the yeast enzyme (Kosaka et al., 1977; Hanna et al., 1983). Like the yeast enzyme, the NAPRTase from S. typhimurium was highly dependent on ATP for reaction. However, like the mammalian enzyme, and unlike that from yeast, the S. ty- phimurium NAPRTase was not fully ATP-dependent, show- ing 10% activity in the absence of ATP.

In the kinetic mechanisms proposed for the yeast NA- PRTase reaction (Hanna et al., 1983; Kosaka et aZ., 1977), the enzyme binds ATP, becomes phosphorylated, and may then release ADP. The E-P then binds PRPP, followed by NA. After catalysis of the phosphoribosyltransferase step, produc- ing NAMN and PPi, the NAMN synthesis products are re- leased in random order followed by Pi release. The exact location of the E-P cleavage step was not identified.

The robust ATP/ADP exchange catalyzed by the S. typhi- murium enzyme in the absence of the other substrates sug- gests that as in the yeast enzyme, the unliganded enzyme can bind ATP, become phosphorylated, and release ADP at rates as fast as overall catalysis. The E-P is relatively stable as judged by the slow ATPase reaction in the absence of NA or PRPP (uncoupled ATPase reaction). PRPP is able to bind to E-P, as demonstrated by its ability to inhibit the ATPase activity and the ADP/ATP exchanges.

The presence of ATP dramatically changes the interaction of NAPRTase with its substrates. The K , values for both PRPP and NA were each lowered 200-fold, and Vmsx catalysis was increased 10-fold. The role of ATP could be exerted by the bound nucleotide or might be a property of E-P. The competitive inhibition of ADP/ATP exchange reactions by PRPP uersus ADP strongly suggests that E-P, and not E-P. ADP, is the species with high affinity for PRPP. Why does the affinity of E-P for PRPP become higher? The most likely possibility is an enzyme conformational change occurring on phosphorylation. An alternative is that the covalently bound phosphoryl group forms a part of the PRPP binding site and directly interacts with PRPP.

After high affinity binding of PRPP, and binding of NA, phosphoribosyl transfer results in formation of enzyme-bound NAMN and PPi. The ability of added PPi to stimulate a V,, ATPase activity (Table I, text) suggests that E-P cleavage is probably triggered by formation of PPi in the catalytic reac- tion, although it remains to be shown that nascent PPi has the same effect as added medium PPi.

The rate equivalence of ATP-stimulated NAMN synthesis, ADP/ATP exchange, and PPi-triggered ATP hydrolysis sug- gests that a normally rate-limiting step in the mechanism occurs during formation of E-P. This observation suggests that subsequent steps (NAMN formation, E-P hydrolysis, and three product release steps) are relatively fast.

Catalysis in the presence of ATPyS is 20-fold slower than that supported by ATP. In all other respects, catalysis sup- ported by ATPyS is like that supported by ATP: the initial stoichiometry of NAMN production and ATPyS hydrolysis is one, and the NAMN synthesis reaction is driven far to the right. The stoichiometric coupling by this slow substrate suggests it will be a key tool in unravelling the coupling energetics of NAPRTase.

Energy Coupling in NAPRTase-It is clear from the results reported here that the ATPase and NAMN synthesis activi- ties are energetically coupled in the NAPRTase reaction. Two distinct mechanisms might account for such coupling. The first of these, chemical coupling, occurs when ATP is chemi- cally involved in the coupled reaction, by phosphorylation of an intermediate. Well described examples include succinyl CoA synthetase, in which phosphorylation activates succinate

toward thioester formation, and glutamine synthetase in which a glutamyl acyl phosphate allows for attack of ammo- nia. There is no obvious way in which such phosphorylation would be useful in the NAMN synthesis reaction. In addition, the phosphoribosyltransferase reaction of NAPRTase pro- ceeds in the absence of ATP. Finally, the other nine PRTases do not interact with ATP, with the exception of ATP-PRTase, for which ATP is the phosphoribosylated substrate, and the poorly characterized nicotinamide PRTase, for which ATP involvement may be similar to NAPRTase (Kasarov and Moat, 1973).

The alternative mechanism, conformational coupling (Tan- ford, 1983), occurs when ATP use results in a new form of the enzyme, with altered substrate binding properties. Catal- ysis, interlinked with energy use, regenerates the original enzyme form. This type of coupling is believed to be followed by E-P type ion-pumping ATPases, such as Ca-ATPase, for which a phosphorylated enzyme exists, systems such as actin- myosin in which an activated E. ADP. Pi complex occurs and FoF1-type ATPases in which ATP hydrolysis at one active site causes H+ extrusion at a structurally distinct site. In all these cases it has proven difficult to learn about coupling because of the fact that the enzymes are integral membrane proteins, are structurally complex, or have an ion as both product and reactant.

The extent of coupling is best measured by the effect of ATP hydrolysis on the ratio of products over reactants for the NAMN synthesis reaction. It is clear from Table I1 that the presence of ATP has a profound effect on the distribution. It is equally clear that although ATP hydrolysis generates a new steady state in which an unfavorable products/substrates ratio is maintained, this state persists only while ATP is present. The situation is analogous to that of a leaky boat, which is maintained afloat only as long as a pump is operating and sinks when the pump stops.

Can we construct a model for conformational coupling in NAPRTase that accounts for the major observations reported here? Factors which need to be included are the observation that energy coupling is not an equilibrium phenomenon and fails when ATP is exhausted and the observation that K,,, values for PRPP and NA decrease dramatically in the pres- ence of ATP.

The reversal of reaction clearly comes from the existence of an ATP-independent pathway for catalysis, which was readily demonstrated in our work. This fact is unusual among other energy-coupling ATPases, which usually demonstrate high efficiency, and can perform reverse reactions in which ATP is synthesized. In the current case, a coupled reverse reaction from Pi, ADP, NAMN, and PPi would be unlikely, since the key coupling intermediate in that pathway, E-P, is hydrolyzed rapidly in the presence of PPi. Because of the one- way nature of the energy utilization, coupling is clearly not an equilibrium effect, but rather a steady-state process.

It is also clear that a central feature of coupling in NA- PRTase is the dramatic decrease in substrate K,,, values associated with E-P formation. It is tempting to think that a selective lowering of substrate dissociation constants would account for coupling. Thus, we assume the existence of two enzyme forms, E and E-P, with different affinities for NA and PRPP.

To move from the phosphorylation-induced affinity changes toward an explanation of the energetics, we invoke a PPi-triggered phosphohydrolysis of the E-P products complex which would lead to a noncovalent phosphate-containing products complex from which products dissociate readily. Thus, in our model we could draw a pathway in which E-P

NAPRTase Energy Coupling 26010

E

A En- In low alRnlly tam ia phaphorYkt .dbYATp.ADeb r.l.uod.

anlnlty form. parllbnlng doponds B Phorphocylalod bnsynu b hlgh

on r d a h concmtnths of PRPP .nd P P I . C In the produdhn buqubnm, PRPP blnda wlth hbh afRnky, tollowed by Mk D Phaphorlboryl group tranbhr occurs. E N.sant PPI UWI phsphowzymb hydrolysis. F ma IOW m n 4 form of the

a In m altomalive MI. pathway. products In unknown ordbr.

PPI blnds to tha hbh dflnky tam of

.nZymb b rbgbIIbr.1.d Urd rdeaBbs

th@ m s h Phcrphanzymb hydrdpia, M d rqpnbratlng tha low dinky nonphosphorylalod form.

SCHEME I

formation results in high-affinity substrate binding, followed by phosphoribosyl transfer to form PPi and NAMN at the active site. PPi-triggered E-P phosphohydrolysis then ensues, resulting in an E . Pi. PPi . NAMN complex with low affinity for products, which dissociate to regenerate free enzyme. This model is attractive in that the energy of ATP hydrolysis is captured as affinity changes that seem to be able to drive product formation. This reasoning has a major flaw, however, arising from the nonselectivity of the proposed affinity changes. It is not likely that the increased binding affinities of ,E-P are selective for substrate only. Thus, newly formed E-P should be able to bind tightly either reactants or products, preventing any thermodynamic preference for the forward reaction. A route around this problem arises in the fact that PPi binding to E-P or E-P.NAMN is followed by E-P hy- drolysis, regenerating the low affinity E, and releasing the products complex. Since the E-P substrates complex is pro- ductive, but the products complex returns the enzyme to free E, selectivity for forward catalysis is maintained. However, as PPi builds up in the reaction mixture, the energetic cost of the forward reaction would increase, expressed as the number of ATP required for each cycle of net NAMN formation. The enzyme thus functions as a Maxwell's demon, using ATP energy to select substrate complexes and reject products complexes. The model developed here is diagrammed in Scheme 1.

Clearly, much remains to be learned about the molecular mechanism of energy coupling in NAPRTase, but the exist- ence of readily discriminated enzyme forms, chemical differ- ences between substrates and products, and meaningful iso- tope exchange reactions suggest that the system will provide important data on energy coupling. It also remains to be determined whether the mechanism outlined here is relevant to other ATP-coupled systems. A common theme in these systems is that ATP use is linked with an affinity cycle of

binding and release of the other substrate. For example, a model for rho action, based on thermodynamic data and calculations, proposes that an E. ATP complex binds RNA with high affinity and that ATP hydrolysis is followed by a 104-fold decrease in RNA affinity and dissociation of the products complex (Stitt, 1988). Similar proposals have been made for chaperonins (Martin et al., 1991) and ATP-depend- ent proteases (Armon et aZ., 1990).

The physiological significance of energy coupling in NA- PRTase is not apparent at this time. It is thought that PPi levels in cells are kept low, so that coupling to ATP hydrolysis is not likely needed to drive the reaction. The normally high intracellular level of ATP also means that the coupled route will normally be followed, so that the existence of an uncou- pled route, of poor efficiency, is of limited usefulness. During times of energy stress, it may be that competition for PRPP could be limiting and that use of ATP to pull PRPP into synthesis of a key energy-acquiring molecule like NAD is of value.

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