Vol. 127, No. 3 Printed in U.S.A.

JOURNAL OF BACTERIOLOGY, Sept. 1976, p. 1197-1207 Copyright © 1976 American Society for Microbiology

Isolation and Characterization of Dihydropteridine Reductase from Pseudomonas Species CAROLYN D. WILLIAMS, GENEVA DICKENS, CAROL H. LETENDRE,* GORDON GUROFF, CORALIE HAINES, AND TETSUO SHIOTA Laboratory ofBiomedical Sciences, National Institute of Child Health and Human Development, Bethesda, Maryland 20014,* and Department of Microbiology, The University of Alabama in Birmingham, Birmingham, Alabama 35294

Received for publication 12 April 1976

Dihydropteridine reductase isolated from the bacterium Pseudomonas species (ATCC 11299a) has been purified approximately 450-fold by ammonium sulfate precipitation and diethylaminoethyl-cellulose chromatographic procedures. The preparation is at least 80% pure as judged by polyacrylamide gels. Its molecular weight was determined to be about 44,000. Both dihydropteridine reductase and phenylalanine hydroxylase activities were found to be higher in cells adapted to a medium containing L-phenylalanine or L-tyrosine as the sole carbon source than in those grown in L-asparagine. The substrate of the reductase is quinonoid dihydropteridine, and the product is tentatively identified as a tetrahydropteridine through its ability to serve as a cofactor for phenylalanine hydroxylase. The enzyme shows no marked specificity for the pteridine cofactor that occurs naturally in this organism, L-threo-neopterin. The pH optimum for the reductase is 7.2, and nicotinamide adenine dinucleotide, reduced form, is the preferred cosubstrate. Inhibition of the reduced and untreated enzyme by several sulfhydryl reagents was observed. A metal requirement for the reductase could not be demonstrated. Dihydropteridine reductase was found to be inhibited by aminopterin in a competitive manner with respect to the quinonoid dihydro form of 2-amino-4-hydroxy-6,7-dimethyl-5,6,7,8-tetrahydropteridine.

Reduced pteridines have been found to act as cofactors in mixed-function oxygenase reactions such as phenylalanine hydroxylation. They function catalytically and do so because of the existence of mechanisms for their regeneration. The role of dihydropteridine reductase in the enzymatic conversion of L-phenylalanine to L-tyrosine in mammals was first demonstrated by Kaufman (11-14). Two enzymes were found to be necessary for this conversion, one isolated from rat liver (pheniylalanine hydroxylase) and the other from sheep liver (dihydropteridine reductase). During the hydroxylation of phenylalanine by phenylalanine hydroxylase, tetrahydropteridine is converted to quinonoid dihydropteridine. Dihydropteridine reductase catalyzes the reduction of quinonoid dihydropteridine back to its fully reduced state by a reduced pyridine nucleotide (Fig. 1). Phenylalanine hydroxylase isolated from the bacterium Pseudomonas species (ATCC 11299a) was found to utilize 2-amino-4-hydroxy6, 7 - dimethyl - 5, 6, 7, 8 - tetrahydropteridine (DMPH4) as cofactor, suggesting that the naturally occurring cofactor of the enzyme was a pteridine (8). (The eighth edition [1968] of the

American Type Culture Collection Catalogue gives the preferred designation ofthis organism as Comamonas species. According to the latest issue of the catalogue [llth ed., 1974], Pseudomonas species is the name of choice. Therefore, previous publications from this laboratory reporting studies with Comamonas species deal with the same organism as will be discussed in this paper.) This was later confirmed when the bacterial cofactor was isolated and identified as a reduced form of L-threo-neopterin (9). Because of the requirement of the bacterial hydroxylase for a reduced pteridine cofactor, it seemed likely that dihydropteridine reductase was present, and played an important role, in this organism. Although dihydropteridine reductase from a variety of mammalian sources has been purified and studied in detail (4, 5, 21, 22, 24), the enzyme has not yet been described in bacteria. It seemed of interest to investigate this enzyme in a cell adapted to a hydroxylatable substrate. Further, such a system might offer an opportunity to study physical, biochemical, and adaptive relationships between the reductase and its related hydroxylase. As a necessary first step,

1197

1198

WILLIAMS ET AL.

J. BACTERIOL.

OH

nucdof.'id%

Tyrosine + H20

uinonoid -dihydropt rin

ll

Phenylsilnine hydroxylm

|

HN

N N

OH

N\

wvdroperidine Reductss

~

0xidind pyridin

Phonylolanine + 02 .-H

nuclow kW

N H

5,6,7,8-Tet,ahy&dopt+.

FIG. 1. Reactions catalyzed by dihycdropteridine

reductase and phenylalanine

hydroxylas e.

the purification and characterizatic)n nfdihyof dlhydropteridine reductase from Pseudo omonas sp(ATCC 11299a) was undertaken. D; ata on the characteristics of the enzyme as we,11 as some initial studies on its induction by grc)wth of the organism on L-phenylalanine are r eported in

this

paper.

MATERIALS AND METHOI DS Cultures of Pseudomonas species (ATFCC 11299a) from the American Type Culture coll ection were grown in the medium described by Guroff and Rhoads (9). DMPH4 was obtained from the Aldrich Chemical Co., Inc. L-threo-Neopterin w as prepared according to the method of Rembold a Lnd Metzger (26), and biopterin was a product of Reg is Chemical Co. Samples of aminopterin were purc hased from American Cyanamide Co. and from Nutiritional Biochemicals Corp. Nicotinamide adenine d inucleotide, reduced form, (NADH) and dithiothr eitol .(DTT) were obtained from Sigma Chemical Co. . Peroxidase (grade II) was obtained from Boehringer ^-Mannheim Corp. L-Asparagine, L-phenylalanine, re Mduced NAD phosphate (NADPH), NAD phosphate ( NADP+), f8mercaptoethanol, dichlorophenolindophEenol, and N-

ethylmaleimide (NEM) were from (Talbiochem. Tris(hydroxymethyl)aminomethane (Ti ris; enzyme and buffer grade) and chymotrypsinog ren A (beef pancreas) were obtained from Schwarz /Mann. Hydrogen peroxide was from Allied Chemi ical. Malate dehydrogenase was obtained from Calb iochem and from Worthington Biochemicals Corp. Iodosuccinimide was purchased from K and K Liaboratories, Inc., and ammonium sulfate (enzyme grade) was from Mann Research Laboratories. Bo vine serum albumin was from Metrix and yeast Eextract was from Difco Laboratories. Whatman diEethylaminoethyl-cellulose (DE52, microgranular an Lion exchanger) was used. Ultrogels AcA-34, -44, and -54 are products of LKB, Inc. Thin-layer chrormatography was done on cellulose sheets (Eastman C] hromagram 6064) obtained from the Eastman Ko idak Co. All salts used were at least reagent grade. E xcept in the case of the sucrose gradients, centrifugiations were done at 0°C in a Sorvall model RC2-B ccantrifuge. Growth of cells and preparation of c zell-free extracts. Nutrient agar slants were inociulated with cultures obtained from the American T3ype Culture no.

Collection, incubated for 24 h at room temperature, and then stored at 3°C. Large quantities of cells for the isolation and purification of dihydropteridine reductase were prepared by using the following procedure. Two 20-liter carboys, each containing 8 liters of growth medium with L-phenylalanine (0.2%) as the sole carbon source, were inoculated from nutrient agar slants. Ferrous ammonium sulfate was added (8 mg/carboy) at 0 time, and cultures were incubated at room temperature with aeration for 22 h. The cell suspension was used to inoculate a fermenter, which contained 300 liters of the L-phenylalanine growth medium. Ferrous ammonium sulfate was added (300 mg at 0 time and 210 mg at 6 h), and the cultures were grown for 22 h at room temperature with aeration. Cells were sedimented by centrifugation, washed with 0.05 M sodium acetate buffer, pH 6.0, containing 0.1 M NaCl, and then taken up in this buffer (2.5 ml of buffer per g [wet weight] of cells). Cell-free extracts were made by passing the cell suspension twice through a 40-ml French pressure cell (American Instrument Co.) and then diluting the resulting material with 2 volumes of buffer and sedimenting the cellular debris by centrifugation at 27,000 x g for 10 min. Two hundred and fifteen grams (wet weight) of cells gave 1 liter of cell-free extract. The extracts were stored at - 15°C. For induction studies, cell suspensions were grown and treated as follows. Two-liter Erlenmeyer flasks, each containing 500 ml of growth medium with 0.2% L-phenylalanine, 0.2% L-asparagine, or

0.1% L-tyrosine as carbon source, were inoculated from nutrient agar slants. Flasks were shaken at 30°C in a gyratory shaker (New Brunswick Scientific Co.) for 22 h. Cells were collected by centrifugation, washed with 0.01 M Tris-hydrochloride buffer, pH 6.8, and then taken up in this buffer (4.0 ml of buffer per g [wet weight] of cells). The cell suspension was passed once through a 10-ml French pressure cell, and cell debris was removed by centrifugation at 27,000 x g for 10 min. Cell-free extracts were stored at -15°C. Enzyme purification. Dihydropteridine reductase was purified from French press extracts prepared from 30 g of Pseudomonas sp. cells grown as described above. The summary of a typical preparation is given in Table 1. All procedures were done at 2 to 4°C. A 0 to 35% ammonium sulfate fraction was made by adding 199 g of solid ammonium sulfate per liter of extract over a 30-min period with mechanical stirring. After this, the solution was stirred for another 30 min. The resulting precipitate was sedimented by centrifugation (27,000 x g, 10 min), and the pellet was taken up in 20 ml of 0.01 M Trishydrochloride, pH 7.4, in 0.05 M KCI (starting buffer) and dialyzed overnight against 2 liters of the same buffer. Enzyme was then applied to a DE52 column (1.5 by 10 cm), after being adjusted to the conductivity of starting buffer with water if necessary. After application, the column was washed with starting buffer until the absorbancy at 280 nm (A280) of the effluent was less than 0.1, after which the gradient elution was begun. A gradient between 375 ml each of 0.01 M Tris-hydrochloride, pH 7.4, in

1199

BACTERIAL DIHYDROPTERIDINE REDUCTASE

VOL. 127, 1976

TABLE 1. Purification of dihydropteridine reductase Step

Vol (ml)

Total(U) activity

Cell-free extract Ammonium sulfate fractionation DE52 AcA-34 AcA-44

117 31

127.0 126.8

7' 5 1.5

115.8 52.8 26.5

0.05 M KCl and 0.01 M Tris-hydrochloride in 0.2 M KCl was used to elute the enzyme, which appears at about 0.1 M KCl. Fractions of 7.5 ml were collected in tubes that contain sufficient NADH and DTT so that the final concentrations were 0.02 and 2 mM, respectively. The column was run at 60 ml/h. Column fractions were analyzed for conductivity, A280, and dihydropteridine reductase activity. After pooling of appropriate fractions, enzyme was concentrated to 7 ml or less using an Amicon ultrafiltration device containing a UM-2 membrane at 60-lb/in2 pressure. Enzyme was then applied to a column of AcA-34 (2.5 by 72 cm) equilibrated in 0.01 M Trishydrochloride, pH 7.4, containing 0.2 M KCl, 0.02 mM NADH, and 2 mM DTT. This column would be expected to fractionate globular proteins in the size range of 20,000 to 400,000 daltons. The AcA-44 column used subsequently (see below) fractionates globular proteins in the size range of 12,000 to 130,000 daltons. The column was run with the same buffer at a speed of 18 ml/h using a peristaltic pump. Fractions of 6.25 ml were collected and assayed for dihydropteridine reductase activity, which usually appears between 280 and 315 ml. Appropriate fractions were pooled and concentrated as before to 5 ml or less and applied to an AcA-44 column (1.6 by 72 cm). The column was equilibrated and run in 0.01 M Tris-hydrochloride, pH 7.4, 0.2 M KCl, 0.02 mM NADH, and 2 mM DTT. Fractions of 2 ml each were collected at a rate of 18 ml/h, maintained by a peristaltic pump. Appropriate fractions, usually between 60 and 80 ml, were pooled and concentrated as above. Enzyme was stored in small samples in liquid nitrogen. Recovery of activity was 20%, with a purification of 440-fold. Acrylamide gel electrophoresis showed one major band (R, = 0.58) and two minor bands (Rf = 0.44 and 0.53). The major band was estimated to contain at least 80% of the protein as judged from the intensity of the Coomassie blue staining. When gel slices were eluted and assayed for dihydropteridine reductase, activity corresponded to the major band. Enzyme determinations. Dihydropteridine reductase activity was determined by a modification of the method of Nielson et al. (24). In this assay, substrate for the enzyme, the quinonoid form of 2-

amino-4-hydroxy-6,7-dimethyldihydropteridine, QDMPH2, is continuously generated by the peroxidase-catalyzed oxidation of DMPH4 by hydrogen peroxide. Q-DMPH2 concentration and peroxidase activity were determined according to the procedures of Nielson et al. (24). The dihydropteridine

reductase assay was done at 25°C by incubating 100

Total protein (mg) Ttlpoen(g 1,743.0 151.9

19.3 3.0 0.8

act (U/mg) Sppat(/g 0.07 0.83

6.02 17.60 33.2

Yield il (%

100 100 91 42 21

,umol of Tris-hydrochloride buffer, pH 7.2, 0.2 ,umol of NADH, 0.2 ,umol of DMPH4, 40 ,ug of peroxidase, and 10 ,umol of hydrogen peroxide with a sample containing reductase, in a total volume of 1.0 ml. The rate of NADH oxidation, indicated by the decrease in absorbance at 340 nm as compared with a water blank, was followed in a Beckman DU spectrophotometer, model 2400, equipped with a Gilford model 6040 recorder. Blank rates (the non-enzymatic reduction of Q-DMPH2 by NADH) were measured in the same way except that reductase was omitted from the assay. Blank values generally did not exceed 40% of the enzymatic rate and were constant in those experiments in which enzyme or pteridine concentration was varied. Unless otherwise specified, the non-enzymatic rate has been subtracted from each experiment presented. One unit of enzyme activity is defined as that amount of enzyme that will catalyze the oxidation of 1.0 ,umol of NADH per min under the conditions of the assay. A molar absorption coefficient of 6,200 M/cm for NADH at 340 nm was used (10). Kinetic data were calculated using the weighted least-squares linear regression method of Wilkinson (27). The reaction velocity was linearly dependent on enzyme concentration over at least a 20-fold range, from 0.006 ,umol of NADH oxidized per min to 0.14 ,umol of NADH oxidized per min. In practical terms, measurements of rates higher than this are not reliable, since the amount of NADH present, 0.2 ,umol, would be sufficient for a rate-recording period of 1 min or less. Phenylalanine hydroxylase activity was determined by either the isotopic method of Guroff and Abramowitz (7) or by thin-layer chromatography of the products on cellulose in chloroform-methanolammonia-water (29:16:4:1, vol/vol/vol/vol) (1). In the latter method, tyrosine and phenylalanine were located by scanning in a Packard model 7201 radiochromatogram with a Packard model 385 recording ratemeter and were then cut out and counted. In both methods, radioactivity was measured in a Nuclear-Chicago Isocap liquid scintillation counter with Aquasol (New England Nuclear Corp.) as the counting solution. Preparation of tetrahydropteridines. Pteridines were dissolved in 0.01 N HCI (2.0 mg/ml) and reduced to their tetrahydro forms in a hydrogen atmosphere using PtO2 as the catalyst (3). Tetrahydropteridine concentrations were determined spectrophotometrically from their absorption at 300 nm before and after use. Extinction coefficients of 5,400 M-1 cm-' in 0.1 M Tris-hydrochloride, pH 6.8, for biopterin and the four isomers of neopterin and of

1200

WILLIAMS ET AL.

10,000 M-1 cm-' in 0.1 M Tris-hydrochloride, pH 7.4, for DMPH4 (3) were used. Preparation of Q-DMPH2. Q-DMPH2 was prepared from DMPH4 by a slight modification of the procedure described by Musacchio et al. (22). A 2;tmol amount of DMPH4 and 2.4 ,mol of 2,6-dichlorophenolindophenol, each dissolved in 1.0 ml of water, were rapidly mixed and extracted immediately with four 10-ml portions of ether to remove the dye. The aqueous phase, which contained the Q-DMPH2, was freed of the last traces of ether by bubbling argon or nitrogen through the solution. For certain experiments it was desirable to estimate the concentration of Q-DMPH2 remaining in the dye-generated mixture. This was possible because of the existence of a linear relationship between the Q-DMPH2 concentration and the ratio of A305 to A2M.. This relationship was proved by constructing a series of standard solutions containing varying proportions of Q-DMPH2 and 7,8-DMPH2 and measuring the corresponding A30/Am. DMPH2 was prepared by the peroxidase-catalyzed oxidation of DMPH4 by I02 described above, and 7,8-DMPH2 was made by allowing a dilute solution of DMPH4 in 0.1 M Tris-hydrochloride buffer, pH 7.4, to stand at room temperature for about an hour until a spectrum characteristic of 7,8-DMPH2 was observed. Protein determinations. Proteins were determined according to the method of Lowry et al. (20). Bovine serum albumin was used as a standard. Molecular weight determination. The molecular weight of the bacterial reductase was estimated by using enzyme purified through the DE52 column and again following more extensive purification through the two subsequent Ultrogel columns. The molecular weight of the enzyme purified through the DE52 column step was determined on a Sephadex G-150 column (1 by 70 cm) equilibrated and run in 0.01 M Tris-hydrochloride, pH 7.4, containing 0.02 mM NADH with or without 0.05 M KCl or 0.2 M KCl. The molecular weight of the most purified preparation was determined on an AcA-54 column (1.3 by 25 cm) in 0.01 M Tris-hydrochloride, pH 7.4, buffer containing 0.2 M KCl, 0.02 mM NADH, and 2 mM DI¶T. Protein standards, myoglobin, chymotrypsinogen A, ovalbumin, bovine serum albumin, and gamma globulin, 1 mg each, were run on each column in the appropriate buffers for calibration. The A215 or A28. was determined for each standard. Dihydropteridine reductase was located by enzymatic activity as described above. A plot of molecular weight of the known compounds versus the logarithm of the volume of the half-height of the leading edge of each compound was made. The elution volume of the reductase was then compared to the straight line generated by the standards. Alternatively, sucrose gradients, between 5 and 20% sucrose, were prepared in the Sephadex G-150 buffers mentioned above. Centrifugations were carried out at 2°C in an SW41 rotor at either 40,000 rpm for 41 h, or 32,000 rpm for 36 h, and in an SW65 rotor for 14 h at 65,000 rpm. Beckman ultracentrifuges, models L3-50 and LS65, were used, respectively, for the two centrifugations. Malate dehydrogenase was used as a marker. Coupling of dihydropteridine reductase and

J. BACTECRIOL.

phenylalanine hydroxylase. Partially purified phenylalanine hydroxylase was prepared and assayed as previously reported (18). Four micrograms of phenylalanine hydroxylase, with a specific activity of 3.3 ,umol of tyrosine produced per 10 min per A280, was activated in the usual fashion. Then a reaction mixture with or without reductase and/or DMPH4 as indicated was added. After an additional 10 min of incubation at 300C, tyrosine production was determined. Effect of sulfhydryl reagents. Reduced enzyme for use in studies of the role of sulfhydryl groups in dihydropteridine reductase activity was prepared by the following method. Enzyme, 2.6 U (0.08 mg of protein), was incubated at room temperature for 30 min in 5 mM DTT in a total volume of 0.15 ml. Enzyme was freed of DTT by passage through an Ultrogel AcA-54 column (1.3 by 25 cm) equilibrated in 0.01 M Tris-hydrochloride, pH 7.4, 0.2 M KCl, and 0.02 mM NADH. Fractions containing enzyme were preincubated at room temperature with the sulfhydryl agents at the concentrations indicated. 5,5'Dithiobis-(2-nitrobenzoic acid) (DTNB) and NEM were prepared and incubated with the enzyme in 0.1 M Tris-hydrochloride, pH 7.4, and iodoacetate and HgCl2 were prepared and incubated with the enzyme in 0.1 M Tris-hydrochloride, pH 8.0. Samples were removed at the times shown for assay of enzymatic activity.

RESULTS Induction of enzyme activity. Cell-free extracts of cells grown on medium containing either L-phenylalanine or L-asparagine were assayed for dihydropteridine reductase and phenylalanine hydroxylase activities. Cells grown in the presence of L-phenylalanine showed an increase in reductase as well as in hydroxylase activity as compared with those grown with L-asparagine (Table 2). Hydroxylase activity was stimulated to a greater extent, 55-fold compared with 8-fold, than was reductase activity. A similar experiment was done comparing cells grown on L-tyrosine, in which phenylalanine hydroxylase is also induced (8). The specific activity of dihydropteridine reductase was increased 4.5-fold and that of phenylalanine hydroxylase was increased 17-fold as compared to an asparagine-grown control (Table 2). It was possible that dihydropteridine reductase was more responsive to changes in cellular pteridine levels than to changes in levels of L-

phenylalanine. To determine if this were the case, bacterial cultures were grown for 21 h in medium containing 0.2% L-asparagine or Lphenylalanine and then for an additional 19 h in either unsupplemented L-asparagine or Lphenylalanine medium or in identical media supplemented with 1.0 mM DMPH4. Cell-free extracts were prepared in 0.01 M Tris-hydro-

VOL. 127, 1976

BACTERIAL DIHYDROPTERIDINE REDUCTASE

TABLE 2. Dihydropteridine reductase and phenylalanine hydroxylase activities in cell-free extracts from Pseudomonas sp. cells grown in media containing L-asparagine, L-phenylalanine, or Ltyrosine

7 6

U/mg of protein Medium

Dihydrop- Phenylalateridine reductase activity

nine hydroxylase activity

Experiment 1 L-Asparagine (0.2%) 0.065 1.04 L-Phenylalanine (0.2%) 0.520 55.0 Experiment 2a L-Asparagine (0.2%) 0.21 1.3 L-Tyrosine (0.1%) 0.68 16.7 a Medium also contained 0.25 mg of ferrous am-c monium sulfate.

1201

5

E 0)

E

4

iI

3

0

o

2

w

O

6.6

-

6.8

7.0

7.2

7.4

7.6

8.0

pH

FIG. 2. Dihydropteridine reductase activity as a chloride, pH 7.4, and then assayed for reductase function Activity was determined as described activity. The results showed that growth of in the textofpH. except that Tris-hydrochloride buffers precells with DMPH4 had no significant effect on pared as indicated in the text at the indicated pH the magnitude of the stimulation of reductase values were used. Activity was determined after coractivity by growth on L-phenylalanine. recting for a blank containing all components of the pH optimum. The optimum pH of the Pseu- assay except enzyme. A 5.8-pg portion of enzyme domonas sp. dihydropteridine reductase assay protein was used for each determination.

was determined using the reductase assay described in Materials and Methods. Results are shown in Fig. 2. The buffers were 100 mM in Tris, which had been adjusted to the indicated pH values with HCI. They are uncorrected for possible ionic-strength differences. The enzymatic activity shown has been corrected for the non-enzymatic rate of NADH oxidation. Maximal enzymatic activity can be seen to occur between pH 7.0 and 7.3. We chose to perform enzymatic assays at pH 7.2. The rate of nonenzymatic NADH oxidation increases dramatically below pH 6.6. This rate is dependent on the presence of Q-DMPH2 since, in its absence, the non-enzymatic rate of NADH oxidation is very low at the pH values tested. The nonenzymatic rate of NADH oxidation is further accelerated by replacing Tris with acetate buffer at pH values below 6.2. The standard assay at pH 7.2, as described in Materials and Methods, usually gave a blank rate, i.e., all assay components except enzyme, equivalent to 0.01 ,umol of NADH oxidized per min. Experiments in which NADH or Q-DMPH2 was varied or in which pterin analogues such as aminopterin were included required, in general, a blank determination for each concentration. Enzyme substrate and requirements. Table 3 shows that the reductase activity requires NADH, Q-DMPH2, and hydrogen peroxide and peroxidase, the latter two being necessary for the continuous generation of Q-DMPH2. To show that Q-DMPH2 actually was the sub-

TABLE 3. Requirements for dihydropteridine reductase activitya NADH oxidized

Omission

(,umol/min/ml of enzyme)

None ............................ 8.48 2.65 Dihydropteridase reductase ........ 0.28 Q-DMPH2 ...................... . NADH ........................... 0 0.55 H202 and peroxidase .............. a For each determination, 0.04 U of dihydropteridine reductase activity containing 1.8 ,g of protein was used. Reaction velocities are presented with correction for blank rates.

strate produced by the peroxidase-catalyzed oxidation of DMPH4 by H202, Q-DMPH2 was prepared by another method (see Materials and Methods), and its concentration and ability to act as substrate for the reductase were determined at intervals after its formation. The absorption spectrum of the Q-DMPH2 solution 1 min after preparation was similar to that reported for the quinonoid form of dihydropteridine (Fig. 3A) (2, 13). On standing at room temperature, the Q-DMPH2 isomerized to 7,8DMPH2, as indicated by spectral changes, so that by 90 min the dihydropteridine preparation contained predominantly 7,8-DMPH2. The Q-DMPH2 concentration remaining at any time was determined by using the ratio A305 to A280 as described in Materials and Methods. The

1202

J. BACTERIOL.

WILLIAMS ET AL. 0.6

A

-

-

T

isomer has an additional meaning. Since the 7,8-isomer is a substrate for dihydrofolate reductase, this experiment shows that the enzyme studied here is not a dihydrofolate reduc-

7

' '

I

0.5 -

/

z

.--

m cr

tase.

Minutes

0.4LLJ

5

10

15 -20 -30

0.3 f

0

co Cl)

m

0.2w

0.1-

260

280

300

320

340

360

380 400

WAVELENGTH (nm) T

0.05

/

0.04 O

0.03 -

/

0.02 /

0.01

-

0

0

0.04

0.08

0.12

0.16

0.20

0.24

0.28

0.32

Nature of the product. To show that the product of the action of this enzyme is, in fact, tetrahydropteridine, experiments were designed in which the reductase was coupled with the tetrahydropteridine-requiring phenylalanine hydroxylase from this organism. Table 4 shows two such experiments (A and B), using a partially purified phenylalanine hydroxylase preparation (18). Experiment A was carried out in 4 mM NADH, optimal for the hydroxylase assay, and that in B was carried out in 0.2 mM NADH, the usual conditions for assay of the reductase. As can be seen from both A and B, addition of reductase increased hydroxylation of phenylalanine at limiting cofactor concentrations, 4.3 ,M. In experiment B stimulation of hydroxylase activity by reductase did not occur in the presence of saturating DMPH4 (22 ,uM), ruling out a direct effect of the reductase protein on the hydroxylase protein. The possibility that the reductase preparation was contributing DMPH4 to the reaction mixture was ruled out by results .such as those seen in experiment A, in which the stimulation of hydroxylase activity was not observed when DMPH4 was omit-

Q-DMPH2 (mM)

FIG. 3. Non-enzymatic production of Q-DMPH2 and its use as a substrate for dihydropteridine reductase. (A) Effect of time on the absorption spectrum of Q-DMPH2 preparation. Q-DMPH2 was prepared as described in the text. Spectra were determined in 0.01 M Tris-hydrochloride, pH 7.4. (B) Reaction velocity versus remaining concentration of Q-DMPH2. The assay mixture contained 100 ,umol of Tris-hydrochloride, pH 7.2,0.2 ,umol ofNADH, 0.2 ml of Q-DMPH2 preparation, and dihydropteridine reductase (3.8 ig of protein).

ability of this preparation to serve as substrate for the reductase was determined by the usual spectrophotometric assay except that the QDMPH2-generating system, i.e., DMPH4, hydrogen peroxide, and peroxidase, was replaced by samples from the Q-DMPH2 preparation. The velocity of the reaction varied with QDMPH2 concentration, indicating that QDMPH2 was a substrate for the dihydropteridine reductase (Fig. 3B). Extrapolation of the data to 0.2 mM Q-DMPH2 gives a velocity almost identical to that estimated using the standard (H202 and peroxidase) method for determining dihydropteridine reductase activity. The observation that the rate of reduction decreases as Q-DMPH2 is converted to the 7,8-

TABLE 4. Effect of dihydropteridine reductase on the rate of phenylalanine hydroxylationa Addition Reductase (eM) 4 (U)

Expt

DMPH4

A

0 0 0 0 0 4.3 4.3 4.3 4.3 4.3 22.0

0 0.022 0.044 0.088 0.131 0 0.022 0.044 0.088 0.131 0

4.3 4.3 4.3 4.3 22.0 22.0

0 0.076 0.152 0.304 0 0.304

B

Tyrosine produced (nmol) 0

0 0 0 0 1.84 2.30 3.54 4.20 3.01 10.58 0.14 0.81 0.97 1.04 4.88 4.98

a NADH concentration was 4 mM in experiment A and 0.2 mM in experiment B. Dihydropteridine reductase was purified through the DE52 step (see Table 1) and had a specific activity of 6.3 U/mg of protein.

VOL. 127, 1976

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BACTERIAL DIHYDROPTERIDINE REDUCTASE

ted entirely. We have not, as yet, been able to stimulate phenylalanine hydroxylase maximally with the available reductase preparations in the presence of limiting DMPH4. Several possibilities for this were investigated and shown not to be the cause. Reductase was incubated in hydroxylase assay conditions and shown to be fully active afterwards. Phenylalanine hydroxylase was added to the reductase assay mixture, as well as being preincubated with reductase assay components before addition of the reductase, and was shown to have no effect on reductase activity as measured by NADH oxidation. Independent experiments (Letendre, unpublished data) suggest strongly that, in the presence of the reductase, DMPH4 becomes unavailable to the hydroxylase. These problems have been observed in the mammalian system (5). However, variation of the ratio of the two enzymes did not increase coupling beyond that reported here. Overall, the experiments show that the reductase stimulates hydroxylation by increasing the rate of recycling of the reduced pteridine cofactor, suggesting that the product of the action of reductase must be tetrahydropteridine. Kinetics. The rate of reduction of Q-DMPH2 by dihydropteridine reductase depended upon the concentration of both NADH and QDMPH2. Figure 4 shows both the direct and double-reciprocal plot of the enzymatic activity as a function of NADH in the presence of QDMPH2. The enzymatic activity increased with NADH until a concentration of 0.2 mM was reached, after which a sharp decrease in activity was observed. The Km calculated from the double-reciprocal plot is 0.09 mM. The decrease of activity above 0.2 mM is reflected in the double-reciprocal plot by the upward curve at higher NADH concentrations. Dihydropteridine reductase was essentially inactive with NADPH. Only 3% of expected activity could be observed at NADPH concentrations up to 0.2 mM. Figure 5 shows the dependence of dihydropteridine reductase activity on Q-DMPH2 concentration, as varied from 0.02 to 0.85 mM, in both direct and double-reciprocal plot. The apparent Km for Q-DMPH2 is 0.34 mM. Substrate specificity. The ability of other quinonoid dihydropteridines to serve as substrate for the reductase was determined. The tetrahydro forms of i-threo-neopterin, the naturally occurring pteridine in Pseudomonas sp. (9), and biopterin were prepared. They were then compared, along with 2-amino-4-hydroxy6,7-dimethylpteridine (DMP), as to their ability to serve as substrate for the reductase. Table 5 shows the results of these experiments. Of the

0

E 0 QM

E

N

0

z

z

NADH ImM) T--T

B 0.08

0

0.07 0.06 0.05

-I>

0

0.04 a

0.03

0.02 0.01

-10

-5

0

5

10

15

20

25

30

35

40

NADH (mM)

FIG. 4. Dihydropteridine reductase activity as a function of NADH. (A) Enzyme activity was determined as described in the text except that the NADH concentration was varied from 0.025 mM to 0.3 M. Q-DMPH2 concentration was maintained at 0.2 mM. Each point was corrected by a blank value obtained in the absence of enzyme at the given NADH concentration. A 1.6- pg amount ofenzyme protein was used for each determination. (B) Double-reciprocal plot of data presented in (A).

three compounds tested, biopterin had the lowest Ki, 0.16 mM, whereas L-threo-neopterin had the highest, 1.45 mM. i-threo-Neopterin was distinguished by its high V.ax, some 7 times higher than biopterin and 2.5 times higher than DMP. However, at the physiological cofactor concentration, 0.1 mM (9), this would not seem to confer any advantage for utilization of the pteridine. Sulfhydryl and metal effects. The possible role of sulfhydryl groups and metal ions on the

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WILLIAMS ET AL.

J. BACTERIOL.

TABLE 5. Tetrahydropteridine specificity of dihydropteridine reductase activity Vmax ( mo1 of TetrahydropteriK (mM) NADH oxidized/ dine min/mi of enzyme) Biopterin 0.16 8.92 DMP 0.34 21.56 1.45 51.76 L-threo-Neopterin

A

15 14 13 12

11-

10 9g876543-

TABLE 6. Effect of sulfhydryl reagents on reduced and untreated dihydropteridine reductasea

I-~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~0.1

02

0.3

0.5

0.4

0.6

0.7

Preincu-

0.8

Enzyme

Q-DMPH2 (mM)

B

0.5 0.4-

-1>

0.30.201

0

5

10

15

Agent

Concn bation 22'C Inhibition (% (mM) time, (min)

Untreated Reduced Untreated Reduced Untreated

DTNB DTNB HgCl2

Reduced

lodoacetate

0.57 0.57 0.4 0.4 HgCl2 NEM 2.5 10.0 Reduced NEM 2.0 5.0 Untreated Iodoacetate 7.0

20

7.0

15 15 15 15 15 15 15 15 30 75 5 15

98 93 50 40 0 57 36 100 9 8

0 17

FIG. 5. Dihydropteridine reductase activity as a function ofQ-DMPH2 concentration. (A) Activity was determined as described in the text except that the QDMPH, concentration was varied as indicated. A 3.8-pg amount of protein was used for each determination. (B) Double-reciprocal plot of data presented in (A).

a Enzyme, untreated or reduced (see Materials and Methods), was preincubated with sulfhydryl agents at the concentrations indicated. The preincubation was at 22°C in 0.11 M Tris-hydrochloride, pH 8.0, in a volume of 0.92 ml. Portions were removed at the desired time and, after addition of peroxidase, hydrogen peroxide, NADH, and DMPH4 to a total volume of 1.0 ml, enzyme activity was determined as usual. Each activity determination contained 2.7 Ag of untreated or 1.7 Ag of reduced protein.

bacterial dihydropteridine reductase was investigated. Enzyme was reduced as described in Materials and Methods and incubated at room temperature with the sulfhydryl agent at the concentrations and for the times indicated in Table 6. DTNB was the most inhibitory of the sulfhydryl compounds tested, causing essentially complete inhibition with both untreated and reduced enzyme. Additional studies showed that DTNB completely inhibited the enzyme at the earliest time point studied, i.e., 1 min. HgCl2 was found to inhibit both untreated and reduced forms of the enzyme, as does DTNB; inhibition, however, was less than complete. NEM inhibited both forms of the enzyme, although inhibition was greater with the reduced than with the untreated form. NEM was the only agent of those tested that inhibited the reduced enzyme more extensively than the untreated form. Iodoacetate was found to be with-

out substantial effect on either the reduced or untreated enzyme. The effect of metals was determined by incubating portions of the enzyme with the compound of interest at 40°C for 5 min. Samples were then removed for determination of enzymatic activity. Under these conditions, enzyme alone lost 80% of its activity. MgCl2, MnCl2, and CdCl2 at 0.1 mM caused complete loss of activity even before incubation at 40°C was begun. CoCl2 was without the inhibitory effect of the other metals, but its presence at 0.1 mM did not prevent the 80% loss of activity on incubation. Ethylenediaminetetraacetic acid at 10 mM substantially protected against loss of activity seen at 40°C. Only 10% of the activity was lost when ethylenediaminetetraacetic acid was included in the 40°C incubation procedure. Enzyme stability. Enzyme activity in ammonium sulfate precipitates is stable for at least a

Q-DMPH2

(mM)

BACTERIAL DIHYDROPTERIDINE REDUCTASE

VOL. 127, 1976

year at -20°C. Cells frozen upon collection and stored at -20°C retain reductase activity for about 3 months. Enzyme purified as above and stored in liquid nitrogen in Ultrogel column buffer is stable for at least 2 months. Enzyme partially purified by passage through diethylaminoethyl-cellulose without addition of NADH and D1T lost 50% of its activity in 6 weeks at -20°C. This preparation was used as a means of determining the ability of various agents to stabilize the enzyme against heat inactivation at 400C. Enzyme and the agent to be tested were preincubated at 40°C for 5 min, a condition usually leading to a 50% loss of activity by the enzyme alone. Both NADH and NADPH protected the enzyme from inactivation under these conditions. Partial protection was seen with 1 ,uM and complete protection with 0.01 mM NADH or NADPH. Neither NAD+ nor NADP+ were effective. Other agents tested that did not alter the course of heat inactivation of the diethylaminoethyl-cellulosepurified enzyme were DMPH4 (0.2 mM), ir phenylalanine (40 mM), L-tyrosine (1.0 mM), L-tryptophan (13 mM), and bovine serum albumin (20 ,g/ml). Molecular weight. The molecular weight of the Pseudomonas bacterial dihydropteridine reductase as determined on Sephadex columns was between 43,500 and 49,000 in three separate determinations. Identical values were obtained in 0.01 M Tris-hydrochloride, pH 7.4, in 0.2 M KCI in the absence or presence of 0.02 mM NADH. It was observed that addition of NADH to the buffer in the absence of salt produced an elution pattern containing two peaks of reductase activity, one at a position corresponding to a molecular weight of 44,000 and

EC 0

18 16 14 12

1205

the other at 60,000. Recovery of activity under these conditions was the same as that found in the absence of NADH, about 80%. Addition of 0.2 M KCO to the NADH-containing buffer abolished the component eluting with an apparent molecular weight of 60,000. We have observed this.same phenomenon with the partially puri-. fled sheep liver reductase. (Sheep liver dihydropteridine reductase, purified through the second ammonium sulfate step [5] was a gift from Ingeborg Hanbauer.) Effect of aminopterin. Aminopterin has been found to be an inhibitor of Pseudomonas dihydropteridine reductase. As shown in Fig. 6 in a double-reciprocal plot of the data, the inhibition is competitive with respect of Q-DMPH2. An apparent Ki of 0.21 mM was calculated (6, 23). It was essential in making these measurements that each experimental point be corrected for the value obtained with the appropriate level of Q-DMPH2 in the absence of enzyme.

DISCUSSION Dihydropteridine reductase from Pseudomonas has been purified and some of its properties have been described. Of particular interest is its induction after growth of cells in media that also induce the enzyme phenylalanine hydroxylase (8). This is not completely unexpected, since both enzymes are necessary for the conversion of phenylalanine to tyrosine in this organism. Since dihydropteridine reductase might be expected to provide reduced pteridine cofactors for reactions other than phenylalanine hydroxylation, it may exist at relatively high concentrations in cells grown under noninducing conditions. This would provide an explanation for the smaller induction of dihydropter-

0-0

0

-A,n*iopternn.

10

03-

0.3mM

2

00

0.05

0.10

0.15

0.20

0.25

-10

0

10

20

30

40

1 Q-DMPH2 ImM) Q-DMPH2 (mM) FIG. 6. Inhibition of dihydropteridine reductase by aminopterin. Dihydropteridine reductase activity was determined as described in the text in the presence and absence of aminopterin. The range of Q-DMPH2 concentration was from 0 to 0.125 mM in the absence of aminopterin and from 0 to 0.25 mM in the presence of 0.3 mM aminopterin. Each experimental value has been corrected for the NADH oxidation observed in the absence of enzyme and in the presence of the appropriate level of Q-DMPH2. A 2.7-pg amount of enzyme protein was used for each determination.

1206

WILLIAMS ET AL.

idine reductase activity, compared with phenylalanine hydroxylase activity, produced by growth on L-phenylalanine or L-tyrosine. A third enzyme, dihydrofolate reductase, has been shown to increase in activity in Pseudomonas sp. cells grown with L-phenylalanine compared with cells grown on L-asparagine (Shiota, unpublished data). The increase is usually about twofold, which is lower than that seen for the dihydropteridine reductase. This enzyme is also involved, though indirectly, in phenylalanine hydroxylation by virtue of its ability to convert 7,8-dihydropteridine, the expected product of pteridine biosynthesis (3), to its fully reduced or tetrahydro form. Whether these enzymes, phenylalanine hydroxylase and dihydropteridine reductase, and possibly even dihydrofolate reductase, are coordinately regulated in this organism remains to be studied. The enzyme showed no striking specificity in terms of a lowered K,,, for the naturally occurring bacterial cofactor. The V,,,,, was higher with L-threo-neopterin than with either DMP or biopterin; however, this would seem to confer no advantage since L-threo-neopterin has been estimated to occur in the bacterium at 0.1 mM. Phenylalanine hydroxylase from Pseudomonas sp. also shows a lack of pteridine specificity (C. H. Letendre, T. Kato, G. Dickens, and G. Guroff, Fed. Proc. 34:624, 1975.), as does the tryptophan hydroxylase from Chromobacter violaceum (17). The mammalian enzymes phenylalanine hydroxylase and dihydropteridine reductase, on the other hand, have been reported to show a clear preference for the natural cofactor, tetrahydrobiopterin (5, 25). The purified bacterial reductase is specific for NADH. Activity with NADPH is less than 5% of that with NADH. Mammalian dihydropteridine reductases are more active with NADH but are able to utilize NADPH to a greater extent than seen in the bacterium (4, 5, 24). Beef adrenal medulla dihydropterin reductase has been shown in one report to be more active with NADPH (21), but this observation has not been confirmed by other investigators (5). Partially purified dihydropteridine reductase from Pseudomonas sp. is protected from heat inactivation by NADH or NADPH, but not by NAD+, NADP+, or DMPH4. These findings are similar to those of Lind (19), who showed that NADH partially protected rat liver dihydropteridine reductase from inactivation by 4 M urea but that NAD+, NADP+, and 2-amino-4hydroxy-6-methyltetrahydropteridine were ineffective. The fact that NADH and NADPH are equally effective in protecting the bacterial reductase from heat inactivation suggests that

J. BACTERIOL.

there might be a similar degree of binding between NADH or NADPH and the enzyme. In spite of this, NADPH is a very poor substrate for the bacterial enzyme. Studies of inhibition by sulfhydryl reagents strongly suggest involvement of one or more sulfhydryl groups in the activity of dihydropteridine reductase. NEM was the only one of the sulfhydryl reagents tested whose inhibitory pattern suggested the involvement of a disulfide bond as well. Since the enzyme is purified and stored in the presence of DTT, it is expected that easily accessible sulfhydryl groups will be in the reduced form. The inhibitory effect of Mg2+, Mn2+, and Cd2+ and the protective effect of ethylenediaminetetraacetic acid against heat inactivation further implicate sulfhydryl groups essential to the activity of the enzyme. The evidence implicating sulfhydryl groups in the activity of the mammalian dihydropteridine reductase is contradictory. The data of Cheema et al. (4) suggest a sulfhydryl involvement, whereas Craine et al. (5) have concluded that such an involvement is unlikely. The bacterial reductase is inhibited by aminopterin in a competitive manner with respect to Q-DMPH,. The Ki of 0.2 mM is about 10-fold higher than that reported for inhibition of the sheep liver reductase by aminopterin (11, 22), indicating that the bacterial enzyme is less sensitive to aminopterin than is the mammalian enzyme. At this point we feel that the type of inhibition, i.e., competitive or noncompetitive, may depend on the pterin used for the study. Using biopterin, Craine et al. (5) have reported competitive inhibition of sheep liver dihydropteridine reductase by another folate antagonist, methotrexate. Kinetic parameters for aminopterin using the bacterial enzyme in the presence of its natural cofactor L-threo-neopterin will be carried out as soon as sufficient quantities of the pterin are available. The increase in pterin-catalyzed non-enzymatic NADH oxidation at low pH values was probably not observed by Nielsen et al. (24) since they used a double-beam spectrophotometer. It is likely, based on several previous reports (14, 28), that a peroxide derivative of tetrahydropterin is formed. The reduction of this product by NADH, a reaction usually catalyzed by dihydropteridine reductase, would account for the rapid NADH oxidation we observe. The molecular weights of all the mammalian dihydropteridine reductases described to date, and of the bacterial reductase studied here, are about the same, between 42,000 and 52,000. The observation that NADH in low salt causes an alteration in the elution pattern of Sephadex

VOL. 127, 1976

BACTERIAL DIHYDROPTERIDINE REDUCTASE

columns for both bacterial and liver enzymes is an interesting one. The physical basis for this alteration is currently being explored. Whereas some problems remain in the demonstration of stoichiometric coupling of dihydropteridine reductase and phenylalanine hydroxylase from Pseudomonas sp., the experiments presented here show that the reductase can regenerate reduced pteridine cofactors for phenylalanine hydroxylation. It thus appears that the bacterial dihydropteridine reductase functions like its mammalian counterpart. ACKNOWLEDGMENTS We are indebted to David Rogerson and the National Institute of Arthritis, Metabolism, and Digestive Diseases for the use of their pilot plant facilities. The help of Walter Lovenberg and Eleanor Bruckwick in the performance of pteridine reductions is appreciated. This study was supported by grant GB-35538 from the National Science Foundation, grant BC-107 from the American Cancer Society, and Public Health Service grant DE02670 from the National Institute of Dental Research (to T.S.). LITERATURE CITED 1. Abita, J. P., C. Dorche, and S. Kaufman. 1974. Further studies on the nature of phenylalanine hydroxylation in brain. Pediatr. Res. 8:714-717. 2. Archer, M. C., and K. G. Scrimgeour. 1970. Rearrangement of quinonoid dihydropteridines to 7,8-dihydropteridines. Can. J. Biochem. 48:278-287. 3. Blakley, R. L. 1969. The biochemistry of folic acid and related pteridines, p. 66-7.1. In A. Neubergen and E. L. Tatum (ed.), Frontiers of biology, vol. 13. NorthHolland, Amsterdam. 4. Cheema, S., S. J. Soldin, A. Knapp, T. Hofmann, and K. G. Scrimgeour. 1973. Properties of purified quinonoid dihydropteridine reductase. Can. J. Biochem. 51:1229-1239. 5. Craine, J. E., E. S. Hall, and S. Kaufman. 1972. The isolation and characterization of dihydropteridine reductase from sheep liver. J. Biol. Chem. 247:60826091. 6. Dixon, M. 1953. The determination of enzyme inhibitor constants. Biochem. J. 55:170-171. 7. Guroff, G., and A. Abramowitz. 1967. A simple radioisotope assay for phenylalanine hydroxylase. Anal. Biochem. 19:548-555. 8. Guroff, G., and T. Ito. 1965. Phenylalanine hydroxylation by Pseudomonas Species (ATCC 11299a). J. Biol. Chem. 240:1175-1184. 9. Guroff, G., and C. A. Rhoads. 1969. Phenylalanine hydroxylation by Pseudomonas species (ATCC 11299a). Nature of the cofactor. J. Biol. Chem. 244:142-146. 10. Horecker, B. L., and A. Kornberg. 1948. The extinction coefficients of the reduced band of pyridine nucleo-

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tides. J. Biol. Chem. 175:385-390. 11. Kaufman, S. 1957. The enzymatic conversion of phenylalanine to tyrosine. J. Biol. Chem. 226:511-524. 12. Kaufman, S. 1959. Studies on the mechanism of the enzymatic conversion of phenylalanine to tyrosine. J. Biol. Chem. 234:2677-2682. 13. Kaufman, S. 1961. The nature of the primary oxidation product formed from tetrahydropteridines during phenylalanine hydroxylation. J. Biol. Chem. 236:804-810. 14. Kaufman, S. 1971. The phenylalanine hydroxylating system from mammalian liver. Adv. Enzymol. 35:245-319. 15. Kaufman, S. 1967. Pteridine cofactors. Annu. Rev. Biochem. 36:171-184. 16. Kaufman, S., and D. B. Fisher. 1974. Pterin-requiring aromatic amino acid hydroxylases, p. 288. In 0. Hayaishi (ed.), The molecular mechanisms of oxygen activation. Academic Press Inc., New York. 17. Letendre, C., G. Dickens, and G. Guroff. 1974. The tryptophan hydroxylase of Chromobacterium violaccem. J. Biol. Chem. 249:7186-7191. 18. Letendre, C. H., G. Dickens, and G. Guroff. 1975. Phenylalanine hydroxylase from Pseudomonas species (ATCC 11299a). Purification, molecular weight, and influence of tyrosine metabolites on activation and hydroxylation. J. Biol. Chem. 250:6672-6678. 19. Lind, K. E. 1973. Dihydropteridine reductase. Measurement of dissociation complexes of enzyme and ligands. Eur. J. Biochem. 33:67-70. 20. Lowry, 0. H., N. J. Rosebrough, A. L. Farr, and R. J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275. 21. Musacchio, J. M. 1969. Beef adrenal medulla dihydropteridine reductase. Biochim. Biophys. Acta 191:485487. 22. Musacchio, J. M., G. L. ]'Angelo, and C. A. McQueen. 1971. Dihydropteridine reductase: implication on the regulation of catecholamine biosynthesis. Proc. Natl. Acad. Sci. U.S.A. 68:2087-2091. 23. Neilands, J. B., and P. K. Stumpf. 1955. Outlines of enzyme chemistry, p. 88. John Wiley and Sons, Inc., New York. 24. Nielsen, K. H., V. Simonsen, and K. E. Lind. 1969. Dihydropteridine reductase. A method for the measurement of activity, and investigations ofthe specificity for NADH and NADPH. Eur. J. Biochem. 9:497502. 25. Osanai, M., and H. Rembold. 1971. Cofactor specificity of L-erythro-tetrahydrobiopterin for rat liver phenylalanine-4-hydroxylase. Hoppe-Seyler's Z. Physiol. Chem. 352:1359-1362. 26. Rembold, H., and H. Metzger. 1963. Synthese und chromatographische Trennung von [8a-'4C]Biopterin und [8a-'4C]7-Biopterin. Chem. Ber. 96:1395-1405. 27. Wilkinson, G. N. 1961. Statistical estimations in enzyme kinetics. Biochem. J. 80:324-332. 28. Woolf, L. I., A. Jakubovic, and E. Chan-Henry. 1971. The non-enzymic hydroxylation of phenylalanine to tyrosine by 2-amino-4-hydroxy-5,6,7,8-tetrahydropteridine. Biochem. J. 125:569-574.