4 Flavin and Pteridine Monooxygenases

Flavin and Pteridine Monooxygenases VINCENT MASSEY I . Introduction

PETER HEMMERICH

. . . . . . . . . . . . . . . . . 191

I1. Internal Flavoprotein Monooxygenaaes

. . . . . . . . .

A . Lactate Monooxygenase (Lactate Oxidase or Lactate Oxidative Decarboxylase) . . . . . . . . . . . . . . B . Lysine Monooxygenase . . . . . . . . . . . . C Arginine Monooxygenme (Arginine Decarboxylase) . . . . 111. External Flavoprotein Monooxygenases . . . . . . . . A. Salicylate Hydroxylase . . . . . . . . . . . . . B . pHydroxybenzoate Hydroxylase . . . . . . . . . C . Melilotate Hydroxylase . . . . . . . . . . . . D Phenol Hydroxylase . . . . . . . . . . . . . E Orcinol Hydroxylase . . . . . . . . . . . . . F. m-Hydroxybenzoate-6-hydroxylase . . . . . . . . . G. m-Hydroxybenzoate-4-hydroxylase . . . . . . . . . H Imidazolylacetate Monooxygenase . . . . . . . . . I Bacterial Luciferase . . . . . . . . . . . . . J . Microsomal Amine Oxidase . . . . . . . . . . . K . Kynurenine-3-hydroxylase . . . . . . . . . . . I V Pterin-Linked Monooxygenases . . . . . . . . . . . A Phenylalanine Hydroxylase . . . . . . . . . . . B . Tyrosine Hydroxylme . . . . . . . . . . . . . C . Tryptophan Hydroxylase (Tryptophan-5-monooxygenaae) . . V Model Studies and Possible Mechanisms . . . . . . . .

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. . . .

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193 194 199 203 204 206 211 217 221 223 224 225 225 226 229 230 231 232 238 240 241

.

1 Introduction

The reactivity with oxygen of reduced flavins and flavoenzymes is one of the most interesting problems in the field of flavin chemistry and will be reviewed in more detail a t the end of this chapter . Although the reac191

192

VINCENT MASSEY AND PETER HEMMERICH

tion is complex, involving the formation of highly reactive peroxydihydroflavins, flavin (Fl) radicals, and superoxide anion, 0,- (1-3), it is rapid (tl,, < 1 sec) and results is the stoichiometric production of H,O,: Fbed

4-Oz+ FI,,

+ HzOz

In the case of many flavoproteins, which are converted to the reduced form by the specific substrates whose oxidation they catalyze, the high rate of reaction with 0, may be retained or even enhanced, in which case they are classified as oxidases. On the other hand, many flavoenzymes react very slowly with 0, and much more rapidly with one-electron acceptors ; these enzymes are classified as dehydrogenases. Although the route of flavin reoxidation appears to be different in the two classes, involving the intermediate production of 0,- and flavin semiquinone in the case of the dehydrogenases and direct two electron oxidation in the case of the oxidases (4,6), the product of reaction with 0, in both classes, as with the free flavins, is H,O,. In contrast to these H,O,-producing enzymes, another group of flavoproteins has become recognized in recent years, the flavoprotein rnonooxygenases. This group of enzymes, also known as mixed function oxygenases or mixed function oxidases, in reaction with 0, cause the incorporation of one atom of the O2molecule into the substrate to yield an oxygenated product and the conversion of the other oxygen atom to H,O. The flavoprotein monooxygenases can be further conveniently subclassified on the basis of the electron donor involved in the catalytic reaction. In one group of enzymes the substrate itself serves as the electron donor; this group is therefore referred to as internal monooxygenases. I n the second group, in addition to the substrate to be hydroxylated, an external reductant such as NADH or NADPH is required for catalysis ; this group is therefore known as external monooxygenases. Since the reactions catalyzed by this group are hydroxylations, these enzymes are also often referred to as flavoprotein hydroxylases. I n addition to the numerous flavoprotein monooxygenases, there is a small group of enzymes which require tetrahydropteridines as cofactors. In view of the similar chemistry of pteridines and flavins, it is widely considered that the flavoprotein and pteridine-linked monooxygenases 1. Q. H. Gibson and J. W. Hastings, BJ 68, 368 (1962). 2. D. Ballou, G. Palmer, and V. Massey, BBRC 36, 898 (1969). 3. V. Massey, G. Palmer, and D. Ballou, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 5. Univ. Park Press, Baltimore, Maryland, 1973. 4. V. Massey, S. Strickland, S. G. Mayhew, L. G. Howell, P. C. Engel, R. G . Matthews, M. Schuman, and P. A. Sullivan, BBRC 36, 891 (1969). 5. V. Massey, F. Muller, R. Feldberg, M. Schuman, P. A. Sullivan, L. G. Howell, S. G. Mayhew, R. G. Matthews, and G. P. Foust, JBC 244, 3999 (1969).

4.

193

FLAVIN AND PTERIDINE MONOOXYGENASES

may fanction by basically similar mechanisms. The similarity in structure of dihydroflavins and tetrahydropteridines is shown below:

Di hydrof lavin

Tetrahydrobiopterin

Although model studies support the concept that hydroxylation reactions involving dihydroflavins and tetrahydropteridines proceed via similar mechanisms (see Section V) there are two important pieces of experimental evidence which cast some doubt on this hypothesis. First, unlike the rapid reaction of dihydroflavins with 0, (tlIz< 1 sec), tetrahydropteridines in general are only sluggishly oxidized by O2 ( tlIz 5 min) . Unless the combination with protein dramatically enhances the 0, reactivity (for which no experimental evidence exists) it would appear unlikely that this autoxidation could be fast enough to be of catalytic significance. Second, in contrast to the flavoprotein monooxygenases, which have all been shown to have no metal involvement, the present evidence indicates strongly that the pteridine-linked monooxygenases are iron-containing enzymes. Thus, the possibility exists that the function of the tetrahydropteridines is to reduce the iron, and that it is the latter moiety which is responsible for oxygen activation, or that a tetrahydropteridine-Fe (111) complex is responsible. Several reviews concerned with flavoprotein monooxygenases (6-10) and pteridine-linked monooxygenases (11-13) have appeared previously. H

II. Internal Flavoprotein Monooxygenases

Internal flavoprotein monooxygenases catalyze oxidative decarboxylations in which an oxygen atom is incorporated into the substrate and 6. 0. Hayaishi, in. “Oxygenases” (0. Hayaishi, ed.), p. 1. Academic Press, New York, 1962. 7. 0. Hayaishi, Bacteriol. Rev. 30, 720 (1966). 8. 0. Hayaishi and M. Nozaki, Science 164, 389 (1969). 9. 0. Hayaishi, Annu. Rev. Biochem. 38, 21 (1969). 10. M. S. Flashner and V. Massey, in “Molecular Mechanisms of Oxygen Activation” (0.Hayaishi, ed.), p. 245. Academic Press, New York, 1974. 11. S. Kaufman, in “Oxygenases” (0. Hayaishi, ed.), p. 129. Academic Press, New York, 1962. 12. S. Kaufman, Advan. Enzymol. 35, 245 (1971). 13. S. Kaufman and D. B. Fisher, in “Molecular Mechanisms of Oxygen Activation” (0.Hayaishi, ed.), p. 285. Academic Press, New York, 1974.

194

VINCENT MASSEY AND PETER HEMMERICH

CO, and H,O are the other products. The substrate itself serves as a reductant of the flavin. Hence, these reactions in effect involve a double oxidation of the substrate, which is first oxidized at the expense of flavin reduction and then oxidized again by the insertion of oxygen. These reactions may be formalized as follows: H I R-C-COOH I

+

EFbx

[::

R-C-COOH.EFlredHz

R-C-COOH*EFlredH, II

X

XH

1

+

02-

EFlox

+

R-CZO

XH

+

C02

+

%O

It has been suggested that the mechanism by which the oxygen atom is incorporated into the product may be quite different with the internal flavoprotein monooxygenases than with the external monooxygenases (10). In the latter group there is growing evidence for the direct participation in the hydroxylation reaction of covalent adducts of oxygen and dihydroflavin. I n the case of the internal monooxygenases the active form of oxygen involved may be H,O,, the normal product of autoxidation of dihydroflavins. The nonenzymic decarboxylation of keto acids by H,O, is a well-established reaction, whose mechanism has recently been investigated (14). Similar oxidative decarboxylation of imino acids would be expected. As will be discussed in more detail in Sections II,A and B, the possibility exists that the decarboxylation reactions catalyzed by this group of enzymes result from the locally high concentration of H,Oz and keto acid (or imino acid) a t the active site. In this case the flavoprotein internal monooxygenases would be in fact typical flavoprotein oxidases ; the monooxygenase activity would then be an adventitious one because of the nature of the primary oxidase products and their mutual reactivity. A. LACTATE MONOOXYGENASE (LACTATE OXIDASEOR LACTATE OXIDATIVEDECARBOXYLASE) Lactate monooxygenase catalyzes the incorporation of molecular oxygen into lactate. Historically this was the first flavoprotein monooxygenase studied. In 1955, Sutton (15) reported the purification of the enzyme from Mycobactenurn phlei and established the prosthetic group to be FMN. The enzyme was obtained in crystalline form in 1957 (16) and 14. G. A. Hamilton, Progr. Bioorg. Chem. 1, 83 (1971). 15. W. B. Sutton, JBC 216, 749 (1955). 16. W. B. Sutton, JBC 226, 395 (1957).

4. FLAVIN AND PTERIDINE MONOOXYGENASES

195

reported to have a minimum molecular weight per flavin of 125,700. This value was based simply on the ultraviolet absorption; later workers have determined the minimum molecular weight to be 55,000-56,000 on the basis of dry weight and biuret determinations (17 ) . The oxygenase nature of the enzyme was established by Hayaishi and Sutton by ISO studies (18). They found that ISO was incorporated into acetate when ‘*02 gas and H2’“0was used, but not with ISO, gas and H21s0. I n the same year, Sutton demonstrated that pyruvate was the product when the enzyme was reduced anaerobically by lactate. Also, by the use of carbonyl trapping agents he was unable to detect any free pyruvate formed in aerobic catalysis, and concluded that any pyruvate formed must remain enzymebound (16). Beinert and Sands (19),with enzyme from M . phlei and a rapid scanning spectrophotometer, demonstrated the presence of a transient longwavelength intermediate which appeared under both aerobic and anaerobic conditions. I n 0.07 M phosphate buffer, pH 7.5, the intermediate was formed in a biphasic manner and disappeared slowly. Stopped-flow measurements performed in 0.05 M phosphate buffer showed a similar pattern ( 2 0 , d l ). These results, implying considerable mechanistic complexity, have since been demonstrated to occur from the use of phosphate as a buffer (see below). Sullivan (22) has purified and crystallized a lactate oxidase from Myco bacterium smegmatis with properties very similar to the enzyme from M . phlei. The enzyme has a minimum molecular weight of 49,500 per FMN. From sucrose gradient and sedimentation velocity data molecular weights of 300,000 and 341,000 were obtained, indicating that the enzyme is composed of multiple subunits. Using the enzyme from M . smegmatis, Lockridge et a,?. (23) have shown that lactate oxidase binds a large variety of anions, which inhibit enzymic activity. This inhibition is competitive with L-lactate, and most of the anions cause marked perturbations in the visible spectrum of the M at pH 7.0, 25O. I n enzyme. Phosphate binds with a Ka of 1.3 X 17. S. Takemori, K. Nakazowa, Y . Nakai, K. Suzuki, and M. Katagiri, JBC 243, 313 (1968). 18. 0. Hayaishi and W. B. Sutton, JACS 79,4809 (1957). 19. H. Beinert and R. H. Sands, in “Free Radicals in Biological Systems” (M. S. Blois, Jr., et al., eds.), p. 35. Academic Press, New York, 1961. 20. S. Takemori, Y . Nakai, M. Katagiri, and T. Nakamura, FEBS (Fed. Eur. Biochem. Soc.) Lett. 3,214 (1969). 21. M. Katagiri and S. Takemori, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 447. Univ. Park Press, Baltimore, Maryland, 1971. 22. P. A. Sullivan, BJ 110,363 (1968). 23. 0. Lockridge, V. Massey, and P. A. Sullivan, JBC 247, 8097 (1972).

196

VINCENT MASSEY AND PETER HEMMERICH

phosphate buffer the biphasic formation of a long wavelength absorbing intermediate similar to that reported for the M . phlei enzyme was also observed when enzyme was reduced anaerobically with L-lactate. However, in the absence of phosphate (and other inhibiting anions) a rapid monophasic formation of intermediate was observed, followed by a slow decay to fully reduced enzyme. The biphasic formation of intermediate was shown to result from the lactate being able to reduce only enzyme uncomplexed with phosphate (the rapid phase) ; the slow phase was determined by the rate of dissociation of phosphate from the inactive enzymephosphate complex. Lockridge et al. have also shown that the intermediate seen transiently is a complex of reduced enzyme and pymvate and that this is the species reacting with 0, in catalysis. Similar intermediates but of different stability and 0, reactivity were detected with a number of a-hydroxy acid substrates. In each case the spectrum corresponding to that of the intermediate produced transiently on anaerobic reduction could be obtained on addition of the corresponding keto acid to the free reduced enzyme. The rate of reaction of the reduced enzyme with O2 was markedly different depending on whether the reduced enzyme was uncomplexed or complexed with keto acid. I n the case of the reduced enzyme-pyruvate complex the rate of reaction with 0, was more than 200 times greater than with uncomplexed enzyme. On the basis of steadystate and stopped-flow kinetic analysis, the following reaction scheme for the enzyme is proposed (23) (cf. Table I) :

:.,

E * FMN * R-CHOH-

EmFMN

+

R-CHOH-COOH

COOH

/E

R- COOH co* H,O

+ +

*

k-31

E-FMNH,

+

E-FMN

+

R-CO-COOH

E * FMN R-CO-COOH 1

H,O,

I n this reaction scheme the catalytic pathway of oxidative decarboxylation follows the steps with rate constants k,, k,, k,, and k,. The step represented by k , is the slow release of a stoichiometric amount of pyruvate from the catalytic intermediate, in accord with the results first

TABLE I KINETICCONSTANTS OF LACTATE OXIDASE' Const ant Kd

=

k-i/ki

kz ka

ka ks

V,,,, catalytic K , (RCHOHCOOH) K , (02)

ks

(Free enzyme) a

tLactate

8-Phenyl klactate

Ira-Hydroxyisovalerate

5 x M 14,000 min-l 2 . 5 min-1 1 . 1 X 108 M-1 min-1 (observed) 11,300 min-l 6,250 min-' 2.23 X M 7 . 1 x 10-544

0.34 M 6 , 7 0 0 min-I -2 min-1 5 . 7 X 106 M-1 min-1 (calculated) 1,060 min-l 910 min-1 5 x 10-2 M 1 . 6 x 10-4 M

9 x 10-2 M 3,700 min-' - 0 . 2 min-1 3 . 3 x 106 M-1 min-1 (calculated) 1,370 min-l 1,000'min-1 2.5 X M 3 x 10-4 M

5 . 4 X 106 M-1 min-1

Values were obtained in 0.01 M imidazole-HCl, pH 7.0, 25". From Lockridge et al. (23).

La-HydroxyB-methylvalerate 3 x M 590 min-' 2 . 7 X lo6 M-' min-l (observed) 350 min-l 220 min-1 1.13 X M 8 X M

198

VINCENT MASSEY AND PETER HEMMERICH

obtained by Sutton (16) and confirmed in the above work. It was also shown by Lockridge et al. that when the keto acid is allowed to dissociate from the reduced enzyme intermediate ( t n w 0.3 min) and 0, is introduced, a stoichiometric amount of H,Oz is produced. On the other hand, when free reduced enzyme was complexed with 2-14C-labeled pyruvate and 0, admitted, little HzO, was detected and a stoichiometric amount of 14CH,COOH was formed. The finding that the form of the enzyme which reacts with 0, is a complex of E-FMNH, and keto acid strongly suggests that the mechanism of decarboxylation may follow simply from the enzyme providing locally high concentrations of reactants. The nonenzymic decarboxylation of keto acids by H,O, has been known for many years; however, high concentrations of the reactants in free solution are required for rapid reaction. With lactate oxidase it seems reasonable to postulate that the E.FMNH,-pyruvate complex in its rapid reaction with 0, yields an ESFMN-pyruvate complex, and that the H20zproduced a t the active site reacts with pyruvate before either product has a chance to dissociate from the enzyme. In this case the primary products of the catalytic reaction, pyruvate and H,O,, are typical oxidase products, similar to those from the amino acid oxidases (imino acid and H,O,). The overall monooxygenase reaction is exhibited only because of the slow release of the products, which at the locally high concentration a t the active center can react rapidly in the secondary decarboxylation reaction. Thus, an “uncoupling” of monooxygenase function could result either from an enhanced rate of product dissociation from the reduced enzyme-product complex (k, in the above scheme) or by enhancement of the rate of release of either H,O, or product from the ternary complex of oxidized enzyme, product and H,O,. As will be discussed in the following section, numerous examples of such “uncoupling” have been documented. It is intriguing to speculate that the amino acid oxidases (which have been shown to operate by mechanisms basically similar to that of lactate oxidase) would also demonstrate such adventitious monooxygenase activity if the imino acid and H,O, products were only released slowly from the enzymes. It should be emphasized that this concept in no way detracts from the probability of a peroxydihydroflavin intermediate preceding the production of H,O, (cf. Section V ) . The postulate is made simply on the grounds that the same oxidative decarboxylation reaction occurs in the absence of enzyme. The possibility exists that a peroxydihydroflavin form of the enzyme is the active oxygenating agent reacting with keto acid; however, in that case step k, must result in the production of an E.FAD.acetate.COe.HzO complex, and the partially rate-limiting step k, must result in the slow release of at least one of the products.

4. FLAVIN

AND PTERIDINE MONOOXYGENASES

199

In further keeping with the basic similarity of lactate oxidase and Damino acid oxidase, i t has also been shown that proton abstraction from the a-carbon atom of the substrate by an enzyme base is an early step in the reaction mechanism of lactate oxidase. Thus, p-chlorolactate is a substrate for lactate oxidase under anerobic conditions resulting in the catalytic elimination of chloride ion and the formation of pyruvate ( 2 4 ) . Under aerobic conditions, as well as the elimination reaction, p-chlorolactate also behaves as a normal substrate, yielding chloroacetate, CO,, and H,O as products. Like the analogous reactions of D-amino acid oxidase with p-chloroalanine (65) the partition between the two catalytic pathways of elimination and oxidation depends strongly on the oxygen concentration, indicating competition of the two pathways for a common intermediate. This intermediate may very well be the complex of reduced enzyme and /3-chloropyruvate. Further evidence for the proton abstraction mechanism comes from recent studies with an acetylenic substrate. Walsh et al. (26) have shown that L-2-hydroxy-3-butynoic acid is a substrate for lactate oxidase as well as an inactivator. The rate of inactivation varies inversely with oxygen concentration, suggesting a partitioning of an enzyme-substrate intermediate between two reaction pathways, one leading to normal catalytic turnover, the other to inactivation. The inactivation has been shown to result from the formation of the covalent addition of a substrate moiety to the reduced flavin. Experiments with the acetylenic substrate labeled with tritium a t the a-carbon atom showed that the inactivated enzyme contained no tritium, indicating that the substrate must lose its a-hydrogen atom before inactivation occurs. When the label was a t carbon-4, inactivated enzyme contained 1 mole of tritium, associated with the flavin. Since inactivation may also be achieved by incubating reduced enzyme with the corresponding acetylenic keto acid (27), it appears likely that the partitioning between catalysis and inactivation occurs a t the level of the E.FMNH,-keto acid complex.

B. LYSINEMONOOXYGENASE Lysine monooxygenase has been isolated in crystalline form from Pseudomonas fluorescens (28).It ,catalyzes the oxidative decarboxylation of L-lysine to 8-aminovaleramide: 24. C. T. Walsh, 0. Lockridge, V. Massep, and R. H. Abeles, JBC 248, 7049 (1973). 25. C. T. Walsh, A . Schonbrunn, and R. H. Abeles, JBC 246, 6855 (1971). 26. C. T. Walsh, A. Schonbrunn, 0. Lockridge, V. Massey, and R. H. Abeles, JBC 247, 6004 (1972). 27. S. Ghisla, V. Massey, C. T. Walsh, and R. H. Abeles, unpublished observations. 28. H. Takeda and 0. Hayaishi, JBC 241, 2733 (1966).

200

VINCENT MASSEY AND PETER HEMMERICH

L- Lysine

6-Aminovaleramide

Studies with l*OZshowed that molecular oxygen is indeed incorporated into the product ( 2 9 ) . L-Arginine is also a substrate, being decarboxylated to y-guanidinobutyramide a t a rate approximately 10% that of L-lysine (SO). The crystalline enzyme exhibits a typical flavoprotein absorption spectrum with maxima at 274, 385, and 460 nm, and was found to contain FAD as prosthetic group ( 2 8 ) . Careful studies with metal chelating agents and metal analyses (31) indicate that metals play no role in catalysis and that the enzyme is a simple flavoprotein with FAD as the sole prosthetic group. The molecular weight has been calculated as 191,000 from sedimentation velocity experiments, and the FAD content was estimated to be two equivalents per molecule of protein (SO). However, a recent study indicates that this analysis may have been in error through contamination with nonflavoprotein impurities ; Flashner and Massey (S2) estimated the molecular weight to be 240,000-246,000with four FAD residues per molecule of protein. Lysine monooxygenase has several unusual properties as compared with other flavoprotein monooxygenases ; for example, the anaerobic reduction of the enzyme-bound FAD by equimolar concentrations of lysine is an extremely slow process requiring 7-9 hr for full reduction (SO). An explanation of this puzzling phenomenon comes from the finding (33) that lysine plays the role of an effector as well as that of a substrate. Evidence was found for there being a regulatory site as well as a catalytic site; i t was necessary for the regulatory site to be occupied by lysine (or nonsubstrate effectors such as c-aminocaproate) for the enzyme to be in a catalytically active form. These findings will be discussed further later in this section. Under anaerobic conditions, the reduction of the enzyme by lysine results in the stoichiometric formation of A1-piperideine-2-carboxylate, presumably by the primary dehydrogenation of the substrate followed by 29. N. Itada, A. Ichihara, T. Makita, 0. Hayaishi, M. Suda, and N. Sasaki, J . Biochem. ( T o k y o ) 50, 118 (1961). 30. H. Takeda, S. Yamamoto, Y. Kojima, and 0. Hayaishi, JBC 244, 2935 (1969). 31. S. Yamamoto, H. Takeda, Y. Maki, and 0. Hayaishi, JBC 244,2951 (1969). 32. M. I. S. Flashner and V. Massey, JBC 249, 2579 (1974). 33. M. I. S. Flashner and V. Massey, JBC 249, 2587 (1974).

4.

201

FLAVIN AND PTERIDINE MONOOXYGENASES

hydrolysis of the resulting imino acid and cyclization of the intermediate a-keto r-aminocaproate (34-36‘).

L-Lysine

a-Imino, €-amino

caproic acid

a -Keto, €-amino

caproic acid

A’- Piperideine2-carboxylic acid

The last two steps of this reaction are presumably nonenzymic. The anaerobic formation of a-imino r-aminocaproate is analogous t o the anaerobic formation of pyruvate with lactate oxidase (16,dS) and suggests that a basically similar reaction pathway may exist for both enzymes. This suggestion is strengthened by the recent report of Nakazawa et al. (37) that aerobically lysine monooxygenase catalyzes a typical oxidase reaction with ornithine or 2,8-diaminooctanoate. I n these reactions the products are the corresponding keto acids and H,O,, without formation of CO,, analogous to the anaerobic reaction with lysine described above. Both the oxygenase-type reaction with lysine or arginine as substrate and the oxidase-type reaction with ornithine are kinetically complex, displaying sigmoidal substrate saturation curves (33,37). This phenomenon appears to result from the presence of two different types of substrate binding sites in the enzyme, a regulatory or effector site which must be occupied before the enzyme can react efficiently with substrate a t the catalytic site ( 3 3 ) . Flashner and Massey have shown that the regulator site may be occupied by lysine, arginine (both oxygenase substrates), L-ornithine (an oxidase substrate), or r-aminocaproate and L-lysine hydroxamate (both nonsubstrate effectors) to abolish the sigmoidal kinetic behavior with L-lysine concentration. It was also shown that lysine could act as an effector to permit the more rapid oxidation of c-N,N-dimethyl-L-ly sine. Recent work of Yamauchi et al. (38) showed that preincubation of 34. T. Nakazawa, S. Yamamoto, Y. Maki, H. Takeda, Y. Kajita, M. Nosaki, and 0. Hayaishi, in “Flavins and Flavoproteins” (K. Yagi, ed.), p. 214. Univ. of Tokyo Press, Tokyo, 1968. 35. S. Yamamoto, T. Nakarawa, and 0. Hayaishi, JBC 247, 3434 (1972). 36. S. Yamamoto, Y. Maki, T. Nakazawa, Y. Kajita, H. Takeda, M. Nosaki, and 0. Hayaishi, Advan. Chem. Ser. 77, 177 (1968). 37. T. Nakazawa, K. Hori, and 0. Hayaishi, JBC 247, 3439 (1972). 38. T. Yamauchi, S. Yamamoto, and 0. Hayaishi, JBC 248, 3750 (1973).

202

VINCENT MASSEY AND PETER HEMMERICH

the enzyme with sulfhydryl reagents transforms the activity toward L-Iysine or L-arginine from an oxygenase to an oxidase. Further work from the same laboratory (39) pertaining to the nature of the active site has shown that L-alanine, itself without activity, is made an efficient substrate if the other moiety of the L-lysine molecule, n-propylamine, is added. The activity so induced is an oxidase one; i.e., the products are pyruvate and H,O,. Other a-monoamino acids were also oxidized in the presence of alkylamines of various chain length; the highest oxidase activity was observed when the total chain length of both amino acid and amine was nearly identical to that of lysine. These striking results have been interpreted by Hayaishi and his colleagues as indicating two types of enzyme-substrate complexes: one of the correct type which leads to oxygenation, the other of an approximate fit in which the dehydrogenation of the substrate by FAD does not couple with oxygen activation, and the reduced flavin is simply oxidized by 0, to produce H,02. While the synergistic effect of a-monoamino acids and alkylamines certainly indicates a requirement for both the a-amino and a terminal basic residue, the results are capable of other interpretations. Perhaps the major effect is on the regulatory site and that a-amino acids could be oxidized a t the catalytic site without simultaneous occupation of this site by an alkylamine. An alternative hypothesis to explain the “uncoupling” phenomenon, i.e., the ability of the one enzyme to catalyze either oxygenase or oxidase reactions, has already been considered in detail in the previous section. According to this iiadventitious” monooxygenase hypothesis, the reaction with all substrates may be regarded as proceeding in two distinct halfreactions. The first of these involves the dehydrogenation of the substrate to produce an enzyme: FADH,.imino acid complex. The second step involves the reaction of this complex with 0, to produce an enzymeFAD-imino acid-HzOz complex. If both the imino acid and H,O, are only slowly dissociated from the complex, oxidative decarboxylation occurs (oxygenase activity). On the other hand, oxidase activity would be exhibited if either product dissociated rapidly from the quaternary complex, or alternatively if imino acid dissociated rapidly from the enzymeFADH,-imino acid complex. Yamamoto et al. (40) have used rapid reaction techniques to investigate the transient intermediates involved in both the aerobic and anaerobic reactions with lysine as substrate. They observed, during the anaerobic 39. S. Yamamoto, T. Yamauchi, and 0. Hayaishi, Proc. N u t . Acad. Sci. U . S. 69, 3723 (1972). 40. S. Yamamoto, F. Hirata, T. Yamauchi, M. Nozaki, K. Hiromi, and 0. Hayaishi, JBC 246, 5540 (1971).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

203

reduction of the enzyme, a transient long wavelength absorbing species which appeared in a biphasic manner and disappeared very slowly to yield reduced enzyme. The spectral characteristics of the intermediate are reminiscent of those first reported for D- and L-amino acid oxidases, i.e., complexes of E-F A D H , and imino acid (41,42). The significance of the biphasic appearance of the intermediate is unclear. However, this behavior is also similar to that found with D-amino acid oxidase in the presence of inhibitors (41) and of lactate oxidase when inhibitory ions are present ( 2 3 ) . Indeed, lysine monooxygenase has been found to be very susceptible to inhibitory ions ( 3 2 ) .Hence, the possibility is very real that the observed complexities in the kinetics of formation of the intermediate are related to buffer inhibition effects.

c. ARGININE MONOOXYGENASE (ARGININE DECARBOXYLASE) Arginine monooxygenase and lysine monooxygenase are closely related in properties and function. Both enzymes catalyze the oxidative decarboxylation of basic amino acids. However, substrate specificity appears to be stricter for arginine monooxygenase, which does not attack L-lysine ( 4 3 ) .On the other hand, lysine monooxygenase will also use L-arginine as an oxygenase substrate at 10% the rate of its reaction with L-lysine. Both enzymes exhibit sigmoidal saturation curves with their substrates and both have pH optima in the region of p H 9. The enzyme was partially purified from the washed mycelium of Xtreptomyces griseus (43) and shown to convert L-arginine to 7 -guanidinobutyramide.

F C=NH CHNH, I COOH L

- Arginine

y

2

C=NH

C -NH, II

0

y-Guanidinobutyramide

Two other compounds similar in structure to arginine were also found to be substrates (canavanine and homoarginine), also being converted to their corresponding amides but a t lower rates (4). 41. V. Massey and Q. H. Gibson, Fed. Proc., Fed. Amer. SOC.E x p . Biol. 23, 18 (1964). 42. V. Massey and B. Curti, JBC 242, 1259 (1967). 43. N. V. Thoai and A. Olornucki, BBA 59, 533 (1962). 44. N. V. Thoai and A. Olomucki, BBA 59,545 (1962).

204

VINCENT

MABSEY

AND PETER HEMMERICH

The oxygenase nature of the enzyme was shown by

1802

experiments

(46). Further studies on a more purified preparation revealed the enzyme to be a flavoprotein with FAD as prosthetic group (46). ThomB-Beau et al. (47) have shown that treatment with diethylpyrocarbonate inactivates the enzyme with the modification of one histidyl residue per mole of enzyme-bound flavin. Arginine protects the enzyme completely from this inactivation. These data, together with a study of the pH dependence of the I(, for L-arginine, indicate that one histidyl residue per flavin is required for catalytic activity, and that this histidyl residue is part of the active center of the enzyme. In light of the finding that proton abstraction from substrate appears to be an early step in catalysis by several flavoproteins (24-26) , the possibility exists that the essential histidyl residue of arginine monooxygenase may function by abstracting a proton from the a-carbon atom of the substrate.

111. External Flavoprotein Monooxygenases

With the external flavoprotein monooxygenases, an external reductant such as NADH or NADPH is required for catalytic activity. A large number of enzymes has been found to belong to this group, and a t least for those cases which have received considerable experimental attention, remarkable similarities in properties have emerged. Most of the enzymes are of bacterial origin and may be induced by growing the microorganism in cultures containing the substrate as sole carbon source. Most of the enzymes catalyze hydroxylation reactions (and hence are often called hydroxylases) which utilize an aromatic compound as substrate. I n most cases the hydroxylated product is either more soluble or more readily metabolized than the substrate ; hence, these enzymes aid considerably in the detoxification process of aromatic compounds and constitute an important part of pollution control and detoxification carried out by soil microorganisms. On the molecular level, the chief and remarkable common property of this group of enzymes is the dual role played by the substrate, which a t the same time is an effector as well as a substrate. The effector role is exhibited chiefly in the enormous stimulation of the rate of reduction of the enzyme flavin by NADH or NADPH when the substrate is bound. 45. D. B. Pho, A. Olomucki, and N. V. Thoai, BBA 118,299 (1966). 46. A. Olomucki, D. B. Pho, R. Lebar, L. Delcambe, and N. V. Thoai, BBA 151, 353 (1968). 47. F. ThomCBeau, Le-Thi-Lan, A. Olomucki, and N. V. Thoai, Eur. J . Biochem. IS, 270 (1971).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

205

This simulation in rate is generally of the order of 103 to lo4 over that of the reduction of free enzyme by the pyridine nucleotide. In a few cases an increased rate of reaction of the reduced enzyme with 0, has also been reported when the substrate is complexed to the enzyme. The effector role has been corroborated by the finding of many examples of substrate analogs which increase the rate of reduction of the enzymebound flavin by pyridine nucleotide but do not serve as hydroxylatable substrates and remain unchanged during the reaction. I n these cases the effector role is displayed by an increased rate of the reduced pyridine nucleotide-oxygen reductase activity. The reduction product of 0, in these cases is H,O, rather than H,O, the normal product with hydroxylatable substrates. Thus an increased rate of 0, consumption and reduced pyridine nucleotide oxidation, coupled with the production of H,O,, is a diagnostic test for nonsubstrate effectors. Some effectors have been found which combine both substrate and nonsubstrate properties, i.e., they may be hydroxylated but not quantitatively, also leading to increased pyridine nucleotide oxidation with H,O, production. I n nearly all cases it has been shown that the combination of the oxidized enzyme and the effector (be it a substrate or a nonsubstrate) results in marked changes in the physical properties of the enzymes (absorption, fluorescence, and circular dichroism spectra). Such changes have permitted the ready determination of the dissociation constants for the cornplexes, which are in all cases one to one complexes. While not so much attention has been paid to this aspect, it is also clear that complexing of the reduced enzyme with substrate also results in similarly detectable spectral changes. It is often found that complexing the oxidized enzyme with the substrate leads to a markedly decreased efficiency of the photochemical reduction of the enzyme flavin by EDTA ; hence, in experiments where the enzyme is photochemically reduced it is the usual practice to add the substrate anaerobically after reduction. The effector role of the substrate may be expected t o play a n important role in the metabolic control of the cell. If these flavoenzymes could be rapidly reduced by NADH or NADPH in the absence of substrate, pyridine nucleotide oxidation would be continuously fast and uncontrolled. The fact that rapid reaction between reduced pyridine nucleotide and enzyme flavin is permitted only when the substrate to be hydroxylated is present thus provides an elegant control mechanism of pyridine nucleotide oxidation. While the physiological reductant in all cases is either NADH or NADPH it is important from a mechanistic viewpoint to state that hydroxylation reactions can be accomplished with these enzymes using nonphysiological reductants. For the oxygenation reaction the important

206

VINCENT MASSEY AND PETER HEMMERICH

thing is to have a complex of reduced enzyme flavin and the substrate. This can be accomplished also by reducing the enzyme with chemical reductants such as dithionite, or photochemically with EDTA. Another feature of the external flavoprotein hydroxylases as a class appears to be emerging, although there are not yet sufficient examples to make it a firm generalization. When these enzymes, in their reduced form and complexed with their specific substrates, are mixed with 0, there appears very rapidly an intermediate with distinctive spectral characteristics which can be ascribed to that of a peroxydihydroflavin, i.e., a covalent adduct of molecular oxygen and reduced flavin. It is this species which is undoubtedly responsible for the insertion of one atom of oxygen into the substrate to form the hydroxylated product. The role of such adducts will be considered in more detail in the appropriate sections following and in the final section dealing with model studies. A. SALICYLATE HYDROXYLASE Salicylate hydroxylase was the second flavoprotein monooxygenase to be discovered and the first example of the group of external monooxygenases (48). It was first purified by Katagiri and co-workers from Pseudomonas putida (49,50) and shown to have a molecular weight of 57,000 with one molecule of FAD per molecule of protein. It was also shown to contain no significant amounts of metals. By the use of 1 8 0 2 it was shown (51) that the reaction catalyzed by the enzyme is + K

O

H

NADH

+

H++

180,-

a’:+ NAD’

f

CO,

+

H,”O

The enzyme is thus unique among this group in that it is the only one which simultaneously with introducing a new hydroxyl function also results in the elimination of CO,. Recently, White-Stevens and Kamin (52-54) have reported on a salicylate hydroxylase isolated from a soil micoorganism which they obtained by enrichment culture. This enzyme differs significantly in many of its 48. M. Katagiri, S. Yamamoto, and 0. Hayaishi, JBC 237, PC2413 (1962). 49. S. Yamamoto, M. Katagiri, H. Maeno, and 0. Hayaishi, JBC 240, 3408 (1965). 50. S. Takemori, H., Yasuda, K . Mihara, K. Suzuki, and M. Katagiri, BBA 191, 58 (1969). 51. M. Katagiri, H. Maeno, S. Yamamoto, 0. Hayaishi, T. Kitao, and S. Oae, JBC 240, 3414 (1965). 52. R. H.White-Stevens and H. Kamin, BBRC 38, 882 (1970). 53. R. H. White-Stevens and H. Kamin, JBC 247, 2358 (1972). 54. R. H. White-Stevens, H. Kamin, and Q. H. Gibson, JBC 247, 2371 (1972).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

207

properties from that isolated from Pseudomonas putida. Chief among these differences are its molecular weight of 91,000 and its content of two molecules of FAD and two subunits of molecular weight 46,000. This enzyme provided the first example of a nonsubstrate effector; benzoate was found to greatly stimulate the NADH-0, reductase activity (producing H,O,), the benzoate not being hydroxylated but serving as a substratelike effector (52,53).It was originally thought that the P. putida enzyme did not exhibit this property ( 4 9 ) ,but more recent studies have shown that a t high concentrations benzoate does indeed act as a nonsubstrate effector ( 5 5 ) . The Pseudomonas putida enzyme is quite specific in its requirement for NADH as external reductant; NADPH is ineffective (49).In addition to salicylate, the following hydroxybenzoates were reported as substrates: 2,3-dihydroxybenzoate, 2,4-dihydroxybenzoate, 2,5-dihydroxybenzoate, 2,6-dihydroxybenzoate, p-amino salicylate, 1-hydroxy-2-naphthoate ( 4 9 ) , and 3-methyl salicylate ( 5 0 ) . However, of this series only 3-methyl salicylate was shown by product analysis to be hydroxylated ; the remaining compounds were tested merely by stimulation of NADH-0, reductase activity and so could be either true substrates or nonsubstrate effectors. On the other hand, White-Stevens and Kamin have made this distinction with their enzyme ( 5 3 ) .I n addition to benzoate, they found that o-nitrobenzoate, rn-hydroxybenzoate, p-hydroxybenzoate, and salicylamide were nonsubstrate effectors. A number of substituted salicylates were found to act both as hydroxylatable substrates and nonsubstrate effectors. This list includes p-amino salicylate, 3-methyl salicylate, 2,3-, 2,4-, 2,5-, and 2,6-dihydroxybenzoates ( 5 3 ) . WhiteStevens and Kamin have also reported that their enzyme will utilize NADPH as electron donor with the same V,,, value as NADH; however, the K , value for NADPH is an order of magnitude higher than that for NADH (53). On mixing with salicylate, a marked perturbation of the absorption spectrum of salicylate hydroxylase is obtained ( 5 0 ) . Titration experiments using this property established that a 1:1 complex of enzyme and M at salicylate was formed, with a dissociation constant of 3.5 X pH 7.0 and room temperature. Spectral changes were also observed in the complexing of the enzyme with substituted salicylates. Similarly, complex formation of the holoenzyme with salicylate was shown by fluorescence studies, where decreases in the fluorescence of tryptophan residues of the protein and of salicylate were detected on binding salicylate, 55. M. Katagiri, S. Taltemori, M. Nakamura, and T. Nakamura, in “Oxidases and

Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 163. Univ. Park Press, Baltimore, Maryland, 1973.

208

VINCENT MASSEY AND PETER HEMMERICH

and where increases in the flavin fluorescence of the enzyme were also followed ( 5 6 ) .These results provided a Kd of 3.2 X M , in excellent agreement with that calculated from absorbance studies. It was also shown that dramatic fluorescence changes result on mixing apoprotein with salicylate and various substituted salicylates. The dissociation conM ) was even stant for the apoprotein-salicylate complex (1.8 X lower than that for the holoenzyme-salicylate complex. The fluorescence technique was also used to determine the dissociation constant for FAD ; a value of 4.5 X M was found. It was also found that NADH and NADPH formed complexes with the apoenzyme with Kd values of 1.1 X M and 1.5 X M , respectively. Product analysis from reactions involving stoichiometric amounts of enzyme demonstrated in a very clear fashion that the hydroxylation reaction is a result of reaction of the reduced enzyme-salicylate complex with 0,. The role of NADH was shown simply to be the reduction of the enzyme-bound FAD ; in these stoichiometric studies it could be replaced by dithionite or by photochemical reduction of the flavin with EDTA. The production of catechol was independent of the order of addition of reducing agent and salicylate, but was dependent upon 0, being added last (50,57). A reaction pathway for the catalytic reaction has been proposed on the basis of these results and rapid reaction studies. Takemori et a2. (58) have recently reported in detail on their rapid reaction studies. This publication has clarified a number of incompletely documented preliminary communications. The catalytic pathway is described by the following reactions: E-FAD

+

kon = 1.8

salicylate

-

+ NADH + H+

+ 4

lo7 M-'sec-'

koff = 62 sec-'

E FAD. salicylate

E * FADH,. salicylate

X

b e d = 230 sec-'

XI

&OX

P

x

,

= 21 sec-'

E FAD salicylate

salicylate - E - FADH,. + NAD'

-

E FAD t catechol CO,

+

H,O

56. K. Suauki, S. Takemori, and M. Katagiri, BBA 191,77 (1969). 57. S. Takemori, H. Yasuda, K. Mihara, K. Suauki, and M. Katagiri, BBA 191, 69 (1969).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

209

The rate-limiting step in catalysis has been found to be in the reoxidation of the reduced enzyme-salicylate complex with 0,, which was found to reach a limiting value of 21 sec-I as the 0, concentration was increased. This is identical with the value of V,,, found in the catalytic reaction. The finding of a limiting rate in flavin reoxidation indicates the formation of an intermediate X,, which may well be a complex (or compound) between the reduced flavin and 0,. Unfortunately, no spectral characterization of this intermediate was attempted; however, it is clear from the results that i t has comparatively low absorbance a t 450 nm and so may be the same type of intermediate found with p-hydroxybenzoate hydroxylase, melilotate hydroxylase, and phenol hydroxylase (see later sections on the individual enzymes). Similarly, an intermediate, XI, was detected in the reduction of the oxidized enzyme-salicylate complex by NADH; this intermediate was converted to the reduced enzyme-salicylate complex a t a rate of 230 sec-I. By pulsed-flow studies the rate of combination of oxidized enzyme with salicylate was estimated to be 1.8 X lo7 M-l sec-l. Using the value of X d of this complex nf 3.5 X M , obtained from static titration experiments, the off velocity constant was therefore calculated as 62 sec-l. Fluorescence quenching experiments were used to determine the K a value of binding of salicylate to reduced enzyme; a value of 1.7 x 10-5 M was found. Thus, salicylate binds more weakly to reduced enzyme than to the oxidized form by a factor of approximately 5-fold. Although a complete study was not made, it was evident from rapid reaction studies that salicylate binds much more slowly, by a factor of approximately 2000, to the reduced enzyme than to the oxidized form. This result rules out the possibility that reduction of enzyme by NADH might precede salicylate binding; salicylate binds to reduced enzyme at a rate only one-seventieth of that required to sustain the catalytic velocity of 21 sec-'. It was further shown in these studies that NADH reacts with enzyme in the absence of salicylate a t a comparatively slow rate and in an apparently second-order fashion, k:ed = 2.5 x lo3 M-l sec-'. At the standard concentration of NADH employed in the catalytic assays of 3X M , this reduction rate is some 2000 times slower than that of the enzyme-salicylate complex. It was also found that when the oxidized enzyme was complexed with benzoate, a nonsubstrate effector, rate stimulation of the reduction of the enzyme flavin by NADH was similar to that with salicylate. When uncomplexed reduced enzyme was mixed with 0, the reaction appeared to be second order with a rate constant of 2.9 X lo4 M-' sec-I. Complexing with benzoate did little to change this pattern; a rate constant of 4.5 x lo4 M-l sec-1 was found. These results should be contrasted

210

VINCENT MASSEY AND PETER HEMMERICH

to the limiting first-order rate of 21 sec-l following a rapid equlibrium described for the reduced enzyme salicylate complex. White-Stevens et al. (54) have also reported rapid reaction studies with their enzyme. As with the P. putida enzyme, marked spectral perturbations were observed on mixing the oxidized enzyme with substrate or nonsubstrate effectors, permitting the evaluation of dissociation constants for these complexes. I n general, the same basic reaction scheme as that of Takemori et al. (58) was proposed. However, there were many quantitative differences between the two enzymes. The forward and reverse constants for the formation of the oxidized enzyme-salicylate complex were determined independently as 9 X lo7 M-I sec-I and 130 sec-l, yielding M . This compares favorably with the Kd of a Kd value of 1.5 X 2-5 X M estimated from static titration experiments. The reduction by NADH of the E-FAD-salicylate complex was found t o be a multistep M ) followed process: the formation of a ternary complex (& = by two first-order reactions with rate constants of 150-600 sec-l and 42 sec-l. No long wavelength absorbance changes were found to be associated with reduction, and hence the two first-order processes were ascribed to formation of fully reduced flavin (the fast step) followed by an isomerization reaction (the slow step). The reaction of molecular oxygen with the reduced enzyme was also studied. In marked distinction to the results obtained with the P. putida enzyme, no limiting first-order rate was detected when salicylate was present. With free reduced enzyme, or in the presence of salicylate, benzoate, p-hydroxysalicylate, or p-aminosalicylate, the rate of reaction was the same and describable by a second-order rate constant of 1.7 X lo4 M-l 6ec-I. At atmospheric oxygen equilibration, this corresponds to a pseudo-first-order rate constant of 23 sec-l. While this rate constant is sufficient to account for the catalytic V,, with salicylate as substrate, it appears inadequate to account for the 3.6-fold greater rate found catalytically with p-hydroxysalicylate. The finding of a constant reoxidation rate led White-Stevens et al. (54) to propose the formation of a “nascent H20,” bound to enzyme, which could either react further with bound substrate to yield H,O and hydroxylated product or to dissociate yielding H,O, and unmodified effector. Flashner and Massey (10) have proposed a structure for the “nascent H,O,”; thus it could be a peroxydihydroflavin intermediate as has been observed with other flavoprotein hydroxylases, and that this was not detected experimentally possibly because the reduced enzyme was not snturated with substrate. This objection now seems to be removed by the finding with the P. putida enzyme that salicylate, while bound less tightly 58. S. Takemori, M. Nakamura, K . Suauki, M. Katagiri, and T. Nakamura, BBA 284, 382 (1972).

4. FLAVIN

21 1

AND PTFBIDINE MONOOXYGENASES

to reduced enzyme than to oxidized, has a K d value which is only five times greater for reduced than oxidized enzyme. If the same sort of relationship applies with the White-Stevens and Kamin enzyme, the concentrations of salicylate used should have been sufficient to ensure that most of the enzyme was complexed with substrate. Clearly more work is required to elucidate this problem. The finding of a limiting rate of reoxidation with the P. putida enzyme offers the clearest possibility of determining the nature of the oxygenating species since rapid reaction studies could reveal whether the intermediate determining the rate-limiting reaction has the spectrum of a peroxydihydroflavin.

B. p-HYDROXYBENZOATE HYDROXYLASE p-Hydroxybenzoate hydroxylase enzyme has been obtained in crystalline form from four different species of Pseudomonas: P. desmolytica (59), P. putida A 3.12 (60),P. putida M-6 (61), and P. fluorescens (62). In all cases the enzyme was obtained from p-hydroxybenzoate-adapted cells. The enzymes from these sources show great similarities to each other in terms of the strict requirement for NADPH as physiological external reductant, the Michaelis constant for p-hydroxybenzoate, excess substrate inhibition, and the content of 1 mole of FAD per mole of protein. The molecular weight has been estimated a t 65,000 for the P. fluorexens enzyme (62), 68,000 for the P. desmolytica enzyme (59), and 83,000-93,000 for the P. putida enzyme (60,sl). An important difference, however, has been found in the stability of the enzymes. The crystalline preparations from both P. putida species are extremely unstable requiring addition of p-hydroxybenzoate, EDTA, and a small molecular weight thiol to protect against inactivation (60,61). On the other hand, the enzyme from P. fluorescens is stable for months, even in the absence of stabilizing agents, greatly facilitating experimental work ( 6 2 ) . Working with the P. desmolytica enzyme, Yano et al. (63) demonstrated the following stoichiometry for the catalytic reaction: COOH

COOH I

I

+ NADPH + H' + OzOH

59. 60. 61. 62. 63.

QOH

f

NADP'

f

H,O

I

OH

K. Yano, N. Higashi, and K. Arima, BBRC 34, 1 (1969). K. Hosokawa and R. Y. Stanier, JBC 241, 2453 (1966). B. Hesp, M. Calvin, and K. Hosokawa, JBC 244, 5644 (1969). L. G. Howell, T. Spector, and V. Massey, JBC 247,4340 (1972). K. Yano, N. Higashi, S. Nakamura, and K. Arima, BBRC 34, 277 (1969).

212

VINCENT MASSEY AND PETER HEMMERICH

By spectral titration experiments stoichiometric binding of p-hydroxybenzoate to the holoenzyme was shown (63). Convincing evidence was also given that enzyme-bound FADH, was the direct electron donor to 0, in the oxygenation reaction ; amounts of 3,4-dihydroxybenzoate were found stoichiometric with the amount of E .FADH, taken, independently of whether the latter had been produced by NADPH or dithionite. I n anaerobic stopped-flow experiments performed with enzyme containing less than a stoichiometric amount of p-hydroxybenzoate, reduction by NADPH occurred in a distinctIy biphasic fashion, related to the mole ratio of p-hydroxybenzoate to enzyme. The value for k r e d for the fast reaction was 96 sec-', some lo4 times larger than that for the slow reaction. These results demonstrate the activator or effector role of p-hydroxybenzoate and that the species reacting in catalysis with NADPH is the oxidized enzyme-p-hydroxybenzoate complex. With enzyme isolated from P. fluorescens, Howell et al. (62) have also demonstrated a perturbation of the flavin absorption spectrum by p-hydroxybenzoate. A K d of 2.9 X M was calculated from such spectral perturbations. Addition of p-hydroxybenzoate also resulted in a 75% decrease in the flavin fluorescence of the enzyme. The Kd and 1 :1 stoichiometry of binding from fluorescence titrations were similar to those found spectrally. As with the enzyme from other species, the rate of anaerobic reduction of the enzyme flavin by NADPH is greatly enhanced in the presence of p-hydroxybenzoate. With or without p-hydroxybenzoate the reduction rate with NADPH shows saturation kinetics indicating the formation of oxidized enzyme-NADPH complexes prior to reduction; the limiting first-order rate constant was 0.41 min-l in the absence and 1.52 X lo4 min-l in the presence of p-hydroxybenzoate, representing a rate stimulation of approximately 40,000 ( 6 2 ) . Another effect of prior complexing of the enzyme with p-hydroxybenzoate is to decrease the dissociation constant of the enzyme-NADPH complex by approximately 13-fold. While it is generally considered that the effector role of the substrate in facilitating the reduction of the enzyme by NADPH must result from some conformational change in the enzyme on binding the substrate, such changes must be rather small and restricted since only minor changes have been detected in the circular dichroism spectra of the enzyme on binding either substrate or competitive inhibitors (61,64). A large number of analogs of p-hydroxybenzoate has been screened as possible substrates or inhibitors. Teng et al. (64) drew a positive correlation between the strength of inhibition and the Hammett substituent constant, U , of 64. N. Teng, G . Kotowycz, M. Calvin, and K. Hosokawa, JBC 246, 5448 (1971).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

213

the substituent on the inhibitory benzoate compound. Recently, however, Spector and Massey (65) examined a larger group of inhibitors of the P . fluorescens enzyme and found that many inhibitors deviate greatly from this type of correlation. Halogen ions, in particular C1- and I-, have been found to be inhibitors of the enzyme (66). The inhibition was found to be competitive with NADPH, and a mixture of competitive and noncompetitive with p-hydroxybenzoate. Of special interest in this study was the finding that C1binds more strongly to the enzyme-p-hydroxybenzoate complex (Ed = 8 x M ) than to the free enzyme (I& = 0.11 M ) . Several compounds as well as p-hydroxybenzoate have been found to enhance the rate of flavin reduction by NADPH and hence activate the NADPH oxidase activity. Howell and Massey showed that 6-hydroxynicotinate is an acfivator without itself being a substrate (66‘,67).This was the second example of “uncoupling” of flavin reduction from substrate hydroxylation found with the external monooxygenases ; a similar uncoupling was first described for salicylate hydroxylase (52) (see Section II1,A) and has since been shown for practically every enzyme in this class. Activation of NADPH oxidation has also been found with 2,4dihydrobenzoate, 3,4-dihydroxybenzoate, and benzoate ( 6 5 ) . Product analysis showed that 2,4-dihydroxybenzoate is a substrate effector, being hydroxylated to 2,3,4-hydroxybenzoate. On the other hand, 3,4-dihydroxybenzoate, the product of hydroxylation of p-hydroxybenzoate, is not hydroxylated further, and is therefore a nonsubstrate effector. It was originally thought that benzoate was hydroxylated to a small extent to m-hydroxybenzoate, but recent work has not substantiated this (68).Interestingly, p-amino benzoate, a strong competitive inhibitor of p-hydroxybenzoate, has also been found to be a substrate, being converted to 3-hydroxy 4-amino benzoate (69). All of the effectors mentioned above perturb the absorption and fluorescence spectra of the enzyme, permitting titration experiments to determine the various dissociation constants. Rapid reaction studies (62,70)with the P. fluorescens enzyme in the anaerobic reduction of the enzyme by NADPH in the presence of p-hydroxybenzoate or 2,4-dihydroxybenzoate have allowed the characterization of several intermediates. These intermediates appear to be of the 65. T. Spector and V. Massey, JBC 247, 4679 (1972). 66. P. J. Steennis, M. M. Cordes, J. G. H. Hilkens, and F. Muller, FEBS (Fed. Eur. Biochem. Soc.) Lett. 36,177 (1973). 67. L. G. Howell and V. Massey, BBRC 40,887 (1970). 68. S. Strickland, L. Schopfer, B. Entsch, and V. Massey, unpublished observations. 69. B. Entsch, D. P. Ballou, and V. Massey, unpublished. 70. T. Spector and V. Massey, JBC 247,5632 (1972).

214

VINCENT MASSEY AND PETER HEMMERICH

charge transfer type, exhibiting long wavelength bands and probably representing charge transfer complexes between the oxidized flavin and NADPH and between the reduced flavin and NADP'. Rapid reaction studies with the enzyme from P. desmolytica did not detect such intermediates (71), possibly indicating that with this enzyme these intermediates are kinetically less stable than with the P. fluorescens enzyme. Rapid reaction studies with the P. fluorescem enzyme have permitted the spectral characterization of oxygenated intermediates when the reduced enzyme-substrate complex is reacted with molecular oxygen. Distinctly different intermediates vere detected with p-hydroxybenzoate (72) and 2,4-dih;.droxybenzoate (70) as substrates. No oxygenated intermediates were detected with the free reduced enzyme or enzyme complexed with a nonsubstrate effector such as 6-hydroxynicotinate or 3,4-dihydroxybenzoate. These results were initially interpreted as possibly indicating a different site of addition of 0, to the dihydroflavin depending on the particular substrate complexed to the enzyme. However, more recent studies have shown that when 2,4-dihydroxybenzoate is the substrate, three different intermediates can be detected (73). The first intermediate formed, whose formation rate is directly proportional to 0, concentration, has a spectrum very similar to that formed with p-hydroxy370 nm, E 8000 M-l cm-I). I n the presbenzoate as substrate (A,,, ence of 2,4-dihydroxybenzoate this intermediate is converted rapidly to a second intermediate with a maximum at 410 nm and a very high extinction coefficient, c = 13,600 M-l cm-l. The rate of formation of this intermediate is much less dependent on 0, concentration, and saturates a t high 0, concentrations, with a limiting rate constant (at pH 8.5, temperature 1.5O) of 5300 min-'. I n turn, this intermediate is converted to a third a t a rate independent of 0, concentration, lc, = 21 min-l. Finally, this intermediate disappears with the reformation of the spectrum of oxidized enzyme, with the 0,-independent rate lc, = 8.3 min-'. Figure 1 shows the spectra of the above intermediates. Studies in which the reaction of the reduced enzyme 2,4-dihydroxybenzoate complex with 0, was quenched rapidly with HC1 after finite time periods and then analyses performed for the product 2,3,4-trihydroxybenzoate revealed that oxygen atom insertion into the substrate occurs at the stage of conversion of intermediate I to intermediate 11. While the mechanistic details are still far from clear, i t is tempting r~

-

71. S. Nakamura, Y. Qgura, K. Yano, N. Higashi, and K. Arima, Biochemistry 9, 3235 (1970). 72. T. Spector and V. Massey, JBC 247, 7123 (1972). 73. B. Entsch, V. Massey, and D. P. Ballou, BBRC 57, 1018 (1974).

4.

2 15

FLAVIN AND PTERIDINE MONOOXYGENASES

I

I

05

I

c.-

.

I1

- 10000

- 8000

03 a,

E

- 6000

0 C

n L

0.2 -

0

In

- 4000

n

a

140000

- 12000

0.4 -

m

I

o.,

-Reduced enzyrn$,,

- 2000

*dihyroxybenzoate

Wavelength (nm)

FIQ.1. Intermediates in the reaction of oxygen with reduced p-hydroxybenzoate hydroxylase complexed with 2,4-dihydroxybenzoate. Data of Entsch et al. (79).

to speculate on the chemical nature of the intermediates. Intermediate I, with an absorption maximum a t 385 nm and an extinction coefficient of 8500 M-l cm-l, together with the second-order dependence on O2 concentration of its formation, would most logically represent a covalent adduct of reduced flavin and 0,. The important question of the site or sites of addition in the isoalloxazine ring system cannot be answered a t this time. However, comparison with the spectral properties of model compounds suggests that the intermediate may have the structure of a substituted C (4a)-N ( 5 )-dihydroflavin (see Section V) ; for example,

H

C(4a) - Peroxydihydroflavin

The rapid quench experiments show that oxygen transfer to the substrate is accomplished coincident with the conversion of intermediate I to intermediate 11. Hence, it is tempting to speculate that the unusual

216

VINCENT MASSEY AND PETER HEMMERICH

spectral characteristics of intermediate I1 result from a complex of the enzyme with the flavin now in a hydroxydihydroflavin form and with the product not yet in its final form; for example, this could be some dienonole-type intermediate such as

0

which should aromatize readily to yield 2,3,4-trihydroxybenzoate. Thus, intermediate 111 could represent a complex of 2,3,4-trihydroxybenzoate with the hydroxydihydroflavin form of the enzyme. The latter, by dehydration of the hydroxydihydroflavin and dissociation of the product, would then return the enzyme to the oxidized form ready for the next catalytic cycle. Extension of such studies to other substrates (69) has revealed that the sequence of intermediates described above is not unique to 2,4-dihydroxybenzoate; for example, p-aminobenzoate, previously considered to be merely a competitive inhibitor, has been found to be a substrate for the enzyme, and elicits a similar sequence of intermediates. Studies with the reduced enzyme p-hydroxybenzoate complex a t low pH values and low temperature have also permitted the positive identification of two intermediates in the reaction of this complex with 0,, with spectral characteristics similar to those of I and I11 in the reaction with 2,4-dihydroxybenzoate as substrate (69). It would thus appear likely that the following reaction scheme must be a minimal one for the oxygen reaction: EFHzArH

+

ki

0 2 -+

I

kz --*

I1

2 TI1 2 EF + ArOH + HzO

In the case of 2,4-dihydroxybenzoate and p-aminobenzoate as substrates, the various rate constants are such that three intermediates can be seen. However, if k , were slower than Ic, and kq, only intermediate I would be expected to be detected. This would appear to be the case for p-hydroxybenzoate as a substrate a t pH 8.5. If k , were slower than lc, but not slower than Ic,, then intermediates I and 111 would be expected to be detected. This would appear to be the case for p-hydroxybenzoate as substrate a t pH 6.5.

4.

217

FLAVIN AND PTERIDINE MONOOXYGENASES

C. MELILOTATE HYDROXYLASE In 1965,in the course of investigating the metabolism of coumarin by an Arthrobacter species, Levy and Frost discovered an enzyme which uses NADH and 0, to hydroxylate melilotate to 2,3-dihydroxyphenylpropionate (74): FH,CH,COOH

FH,CH&OOH

+ NAD+ + q o

+ NADH + H+ + o*-

The enzyme was subsequently purified and found to be a flavoprotein with a molecular weight of 65,000 containing one molecule of FAD per molecule protein (75). More recently, Strickland and Massey (76) have reported the isolation of melilotate hydroxylase from a Pseudomonas species. Like the Arthrobacter enzyme this was also found to contain FAD as prosthetic group (1 molar equivalent per 65,000 g protein) but differs from the Arthrobacter enzyme in having a considerably higher molecular weight (238,000-250,000) and in having four protein subunits. This enzyme has been the subject of a comprehensive kinetic study ( 7 7 ) .I n a conventional steady-state analysis, in which the concentrations of each of the three substrates, melilotate, NADH, and 0, were varied systematically, the following unique pathway was deduced: NADH E . FAD + mel.-.

kl

/FAD

E,

k,

E-FADH, k4

me 1

A .~

/FADH,

k,

E

'me1

+ NAD+

The validity of this reaction scheme was demonstrated by rapid reaction studies in which changes in the enzyme itself were monitored by stoppedflow spectrophotometry and fluorimetry, which permitted determination 74. C. C. Levy and P. Frost, JBC 241, (1967). 75. C. C. Levy, JBC 242, 747 (1968). 76. S.Strickland and V. Massey, JBC 248, 2944 (1973). 77. S. Strickland and V. Massey, JBC 248,2953 (1973).

218

VINCENT MASSEY AND PETER HEMMERICH

of most of the individual rate constants. The formation of an oxidized enzyme-melilotate complex was readily detected by perturbation of the flavin absorption spectrum of the enzyme ; titration experiments showed that under standard conditions (pH 7.3, l o ) the Kd for this complex was 3.8 x M . That the reaction pathway follows the ordered sequence shown above was demonstrated by the fact that enzyme in the absence of melilotate is reduced very slowly by NADH (limiting rate 1.4 min-l extrapolated to infinite NADH concentration). This is slower by more than two orders of magnitude than the catalytic turnover number of the enzyme (735 min-I). I n contrast the E. FAD-melilotate complex reacts extremely rapidly with NADH, in an apparently second-order reaction, k , = 1.4 x lo8 M-I min-'. Determination of the absorption spectrum showed that the product of this reaction was a ternary complex of reduced enzyme, melilotate, and NAD', which proceeded to dissociate the NAD+ a t a rate k , = 1300 min-I. The ternary complex was shown to exhibit a charge transfer absorption spectrum (with a long wavelength band centered round 750 nm) as a result of interaction between EaFADH, (donor) and NAD' (acceptor). The same ternary complex could be formed in static equilibrium experiments in which the E.FADH,-melilotate complex was titrated anaerobically with NAD'. These titrations permitted the determination of the dissociation constant (Kd = 1.45 X M ) and hence the calculation of k,. Stopped-flow turnover data (in which all three substrates are mixed with the enzyme) also indicate that NAD+ dissociates from the ternary complex prior to reaction with 0, (77). As well as the dramatic enhancement of the reduction rate, complexing the enzyme with melilotate also results in a significant increase in the rate of reaction of the reduced flavin with 0,. I n the absence of melilotate the rate is 9.7 X lo5 M-l min-l; in the presence of saturating melilotate this is increased some 16fold to 1.6 x lo7 M-l min-'. Of particular interest is the detection of an oxygenated flavin intermediate in the reaction of the reduced enzymemelilotate complex with 02. This intermediate has a spectrum similar to that of intermediate I found with p-hydroxybenzoate hydroxylase, as described in the preceding section. The rate of appearance of the intermediate ( k , = 1.6 X lo7 M-l min-l is directly dependent on 0, concentration; its subsequent breakdown to yield oxidized enzyme and products is independent of 0, concentration and proceeds a t a rate k , = 1400 min-l. These independently measured rate constants were then used to predict values for the Michaelis constants of the substrates and the maximum catalytic velocity of the hydroxylation reaction, using kinetic equivalents appropriate for the reaction mechanism shown. Table I1 lists the individual rate constants determined from stopped-flow and equilibrium studies, the predicted Vn,,, and K , values, and those actually observed in the

4. FLAVIN

219

AND PTERIDINE MONOOXYGENASES

TABLE I1 KINETICANALYSISOF MELILOTATE HYDROXYLASE (pH 7.3, 1”) Step ki k2

ka kr

Rate constant

Method of determination

5 . 7 X lo8 M-I min-1 2 . 2 X l o 4 min-1 1 . 4 X 108 M-1 min-1

From Vmsxand K m (mel) From k, and Kd (mel) Stopped flow Stopped flow Stopped flow From k5 and Kd (NADf) Stopped flow Stopped flow Stopped flow

0 1.3

k7

x 103 min-1 9 . 0 X lo5 M-I min-1 1.6 X l o 7 M-l min-1

ks ko k I o,ki 1,k I 2

1 . 4 X 103 min-1 Not determined

ks ka

Kinetic constant

0

Kinetic equivalent

Observed steady-state value 735 min-1

K , (NADH)

+

(k4 k5) kaks . 40

4.7

x 10V M

Predicted from rate constants 680 min-l

4.8 X

M

steady-state analysis. The predicted and observed values are in remarkably good agreement and provide very strong evidence for the validity of the proposed mechanism. The intramolecular migration of ring substituents during hydroxylation of aromatic substrates (the NIH shift) is a phenomenon which has been investigated extensively by Jerina, Daly, and co-workers (78). Their incisive experiments have led to the mechanistic conclusion that hydroxylations catalyzed by the liver microsomal P 450 system proceed via an arene oxide intermediate (79). It is the subsequent protonation of this arene oxide followed by rearrangement which leads to the final aromatic 78. D. M. Jerina and J. W. Daly, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 143. Univ. Park Press, Baltimore, Maryland, 1973. 79. D. M. Jerina, J. W. Daly, B. Witkop, P. Zaltzman-Nirenberg, and S. Udenfriend, Biochemistry 9, 147 (1969).

220

VINCENT MASSEY AND PETER HEMMERICH

hydroxylated product. One of the most important questions concerning these results is whether the arene oxide pathway is an obligatory route for aromatic hydroxylations occurring via reductive activation of molecular oxygen. Confirming the presence or absence of the NIH shift in flavoprotein-catalyzed hydroxylations would thus lead to important extrapolations concerning similarities in the mechanism of hydroxylations effected by different biological systems. In a recent study, Strickland et at. (80) have employed combined gas chromatography-mass spectrometry to analyze the reaction products when 3,5-dideuteromelilotate was employed as a substrate with melilotate hydroxylase. No retention of deuterium indicative of the NIH shift could be detected (limits of detection 0.5%). Unfortunately, these results cannot rule out definitely the occurrence of the arene oxide pathway since it has been shown that substituents on phenols generally show little or no migration during aromatic hydroxylations with systems known to give the NIH shift with other substrates (81).This is presumed to result from a catenoid intermediate being stabilized effectively through loss of a proton from the phenolic-OH to yield a stable dienone without migration of a substituent (Scheme 1). R

R

R

~

,

& 1

d

R

dienonc stabilization

*He

0

-Y@orD"

-De

NI H SHIFT

1

D

NO NIH SHIFT

SCHEME 1 80. S. Strickland, L. Schopfer, and V. Maasey, Biochemistry 14,2230 (1975). 81. J. W. Daly, G. Guroff, 5. Udenfriend, and B. Witkop, ABB 122, 218 (1967).

4. FLAVIN AND PTERIDINE MONOOXYGENASES

221

While such stabilization effects would make it difficult to interpret a negative result, it should be pointed out that small amounts of migration, 1-5%, have been observed during hydroxylation of phenols (81). Thus, it is possible from the complete absence of migration observed with melilotate hydroxylase that the arene oxide pathway is not operational in this flavoprotein-catalyzed reaction. In the above study it was also found that use of 3,5-dideuteromelilotate instead of the normal protium form led to no changes in the rate of formation ( k , ) of the oxygenated intermediate or in its subsequent breakdown (k9).As discussed in the previous section, recent work with a similar flavoprotein p-hydroxybenzoate hydroxylase (73) has demonstrated the existence of a t least three intermediate steps when reduced enzyme-substrate complex is reacted with molecular oxygen. This suggests the possibility that similar intermediates exist with melilotate hydroxylase which are however invisible for kinetic reasons. It could be one of these latter intermediates which is responsible for the fission of the carbon-hydrogen bond, thus accounting for the lack of an observed isotope effect.

D. PHENOL HYDROXYLASE Recently, Neujahr and Gaal (82) have reported the purification of a phenol hydroxylase from a yeast, Trichosporon cutaneum. This yeast strain was induced to produce the enzyme in response to growth on phenol or resorcinol as the major carbon source. The enzyme was obtained as a protein homogeneous by disc gel electrophoresis. It has a molecular weight of 148,000and was reported to contain one molecule of FAD per molecule of protein. However, this value was probably low, as a result of some loss of FAD from the enzyme during purification; more recent analyses of 17 different preparations of the enzyme gave an average FAD content of 1.74 molecules per 148,000molecular weight. Sodium dodecyl sulfate (SDS) gel electrophoresis also indicates the presence of two subunits of molecular weight approximately 76,000 (Dr. H. Y. Neujahr, personal communication). The enzyme has an unusually broad substrate specificity, although the requirement for NADPH as external electron donor is strict. Phenol is hydroxylated to yield catechol. Catechol is also hydroxylated to yield

82. H. Y. Neujahr and A. Gaal, Eur. J . Bbchem. 35, 386 (1973).

222

VINCENT MASSEY AND PETER HEMMERICH

pyrogallol (1,2,3-trihydroxybenzene). Other dihydroxybenzenes are also substrates ; resorcinol (1,3-dihydroxybenzene) and quinol (1,4-dihydroxybenzene) are both hydroxylated to yield hydroxyquinol ( 1,2,4-trihydroxybenzene) . Various substituted phenols also act as substrates, with rates between 10 and 100% that of phenol. Thus, 2-F-, 3-F-, and 4-Fphenols; 2-C1-, 3-C1-, and 4-Cl-phenols ; 2-methyl, 3-methyl, and 4methyl phenols; and 2-amino, 3-amino, and 4-amino phenols are substrates. While complete stoichiometric studies have not been carried out in all cases, it is evident that substantial hydroxylation of these substrates occurred. Thus, the possibility exists that the enhanced rate of NADPH oxidation with these substances may in part result from their also acting as nonsubstrate effectors (cf. Section II1,A on salicylate hydroxylase) . The enzyme is susceptible to pronounced inhibition by chloride ions,

0.25

1

0.20

E

0.15

m

n

5

2

0.10

0.05

I

350

LOO Anrn L50

500

FIG.2. Phenol hydroxylase, 2.13 x 10.' M with respect, to enzyme-bound flavin, in 0.1 M phosphate pH 7.6, containing 10 mM dithiothreitol, 15 mM EDTA, and 5 mM phenol, was reduced photochemically in an atmosphere of N, and mixed in the stopped-flow apparatus. with an equal volume of 02-saturated buffer mixture. The absorbance changes (path length 2 crn) were followed with time a t the wavelengths shown 6-10 nm intervals from 330 to 550 nrn). The biphasic changes in absorbance were analyzed to determine the spectrum of the intermediate. Temperature of observation, 2" (V. Massey and H. Y. Neujahr, unpublished results).

4.

223

FLAVIN AND PTERIDINE MONOOXYGENASES

this inhibition effect being much more marked on the acid limb of the pH-activity profile than on the alkaline limb. The catalytic reaction is also inhibited by high concentrations of phenol. Metal chelators are without effect on activity; however, a requirement for intact thiol residue (s) is indicated by the susceptibility of the enzyme t o inhibition by heavy metal ions and p-mercuribenzoate ( 82) . Like the other flavoprotein external monooxygenases so far investigated, phenol hydroxylase also shows distinctive changes in the visible absorption spectrum on complexing with its phenolic substrates (82). Preliminary studies also indicate that with such complexes the rate of reduction of the enzyme flavin by NADPH is considerably faster than with uncomplexed enzyme (H. Y. Neujahr and V. Massey, unpublished results). Preliminary rapid reaction studies have shown that phenol hydroxylase, like p-hydroxybenzoate hydroxylase and melilotate hydroxylase (see previous sections), forms a transient peroxydihydroflavin species when the reduced enzyme-phenol complex is mixed with oxygen. The spectrum of this species, together with those of the complexes of phenol with the oxidized and reduced enzyme, is shown in Fig. 2.

E. ORCINOLHYDROXYLASE Orcinol hydroxylase has been isolated in crystalline form from Pseudomonas putida and shown to be a flavoprotein consisting of a single polypeptide chain of molecular weight 60,000-70,000 and containing one molecule of FAD per molecule of protein (83).The reaction catalyzed by the enzyme is

+ HO

NADH

+

-

H+ + 0,

+ NAD+ + H,O

HO

Orcinol

2,3,5-Trihydroxytoluene

The weak NADH oxidase activity of the enzyme is stimulated by resorcino1 and m-cresol. Product analysis showed that these compounds were functioning chiefly as nonsubstrate effectors. When resorcinol is used some hydroxylated product (hydroxyquinol) is formed but not in sufficient quantities to account for the NADH and O2 consumption. Furthermore, addition of catalase to the assay mixture results in the return of 40% 83. Y . Ohta and D. W. Ribbons, FEBS (Fed. Eur. Bioehem. Soc.) Lett. 11, 189 (1970).

224

VINCENT MASSEY AND PETER HEMMERICH

of the 0, consumed. With m-cresol, no hydroxylation occurs, and catalase returns 50% of the consumed 0,. Addition of catalase has no effect on the NADH oxidation in the presence of either substrate analog. These results show that orcinol is the only true substrate with its hydroxylation being tightly coupled to NADH oxidation, while resorcinol is hydroxylated to only a limited extent and m-cresol not a t all. Orcinol hydroxylase can utilize both NADH and NADPH as well as reduced 3-acetylpyridine nucleotide as electron donors, although NADH is the best donor (84). Ribbons et al. have studied the stereospecificity of hydride transfer from NADH to the enzyme-bound FAD using 4Rand 4S-[SH]-NADH (84). The pro-R protium (A side) of NADH is stereospecifically transferred when orcinol is the substrate. With m-cresol as effector, transfer of hydride rather than tritide appears to be preferred from either side of the reduced pyridine nucleotide, while with resorcinol the specificity for the pro-R tritium is less than with orcinol yet more than with m-cresol.

F.

m-HYDROXYBENZOATE-6-HYDROXYLASE

m-Hydroxybenzoate-6-hydroxylaseis an inducible enzyme which has been purified from Pseudomom aeruginosa. It catalyzes the initial hydroxylation reaction in the gentisate pathway for the metabolism of mhydroxybenzoate (85): COOH

rn-Hydroxybenzoic acid

COOH

Gentisic acid

It is a flavoprotein using either NADH or NADPH as electron donor. As with the other external flavoprotein monooxygenases previously discussed, reduced pyridine nucleotide oxidation is greatly facilitated by the presence of the substrate effector, m-hydroxybenzoate (D. W. Ribbons, personal communication). The unusual feature of the reaction catalyzed by this enzyme is the site of hydroxylation in the aromatic ring. With all other flavoprotein hydroxylases, the new hydroxyl function is introduced in the position 84. D. W. Ribbons, Y. Ohta, and I. J. Higgins, in "The Molecular Basis of Electron Transport" (J. Schultz and B . F. Cameron, eds.), p. 251. Academic Press, New York, 1972. 86. E. E. Groseclose and D. W. Ribbons, Bacteriol. Proc. p. 273 (1972).

4. FLAVIN

225

AND PTERIDINE MONOOXYGENASES

ortho to the existing hydroxyl group. This feature has been used by Hamilton (86) t o propose a mechanism for flavoprotein-catalyzed hydroxylation reactions involving a 4a-peroxydihydroflavin which is converted to a ring-opened form, the latter then being attacked by the nucleophilic center ortho to the phenolic group of the substrate. While this mechanism lacked experimental basis, it would seem unable to accommodate the para insertion of a hydroxyl function catalyzed by m-hydroxybenzoate-6-hydroxylase.

G.

m-HYDROXYBENZOATE-4-HYDROXYLASE

Prema Kumar et al. (87) have reported the partial purification from Aspergillus niger of a m-hydroxybenzoate hydroxylase, utilizing NADPH

as electron donor and producing protocatechuate: COOH

COOH

OH

The prosthetic group was found to be FAD. The same workers (88) have shown that the above reaction can be completely inhibited by reasonably low levels of superoxide disrnutase. This result is surprising in view of the lack of inhibition by superoxide dismutase of other flavoprotein hydroxylases (4,SS). The positive effect of superoxide dismutase found with this enzyme indicates the possibility of involvement of the superoxide anion, 02-, in the hydroxylation reaction. This possibility will be considered further in the last section. m-Hydroxybenzoate-4-hydroxylase has also been obtained recently fr0m.P. testosteroni. In addition, this enzyme uses NADPH as the preferred electron donor, although NADH will also function less efficiently (D. W. Ribbons, personal communication).

H. IMIDAZOLYLACETATE MONOOXYGENASE Imidazolylacetate monooxygenase has been isolated from a pseudomonad species and forms part of the histidine catabolic pathway in which 86. G. A. Hamilton, i n “Molecular Mechanisms of Oxygen Activation” (0.Hayaishi, ed.), p. 405. Academic Press, New York, 1974. 87. R. Prema Kumar, P. V. Subba Rao, N. S. Sreeleela, and C. S. Vaidyanathan, Can. J. Bioehem. 47, 825 (1969). 88. R. Prema Kumar, S. D. Ravindranath, C. S. Vaidyanathan and N. Appaji Rao, BBRC 49, 1422 (1972).

226

VINCENT MASSEY AND PETER HEMMERICH

imidazalonylacetate is converted to aspartic acid (89). The reaction catalyzed by the enzyme is HC =C I 1 HN,C+N

,CH,COOH

%

+NADH+H++O,-

H

Imidazolylacetate

C

&

H

,

~

~

Z

1 I HKC’,N H

~ f

~

~

NAD’

+ H,O

Imidazolonylacetate

The monooxygenase nature of the enzyme was shown by Rothberg and Hayaishi with l8O experiments (90). Crystallization of imidazolylacetate monoxygenase (89,91) revealed the flavoprotein nature of the enzyme with FAD as the only prosthetic group. The molecular weight was estimated to be between 87,000 and 90,000, with one molecule of FAD per molecule of protein. NADH is the preferred electron donor, although NADPH can also function in a less effective manner, Okamoto et al. have studied the role of thiol groups in the enzyme (92). They found that two thiol groups could be titrated in the native enzyme by silver nitrate or by p-mercuribenzoate. I n the presence of imidazolylacetate only one thiol group could be titrated and the presence of the substrate protected against inactivation by the mercurial. While the monooxygenase activity was lost completely on reaction with mercurial (in the absence of substrate) the weak NADH oxidase activity was not; in fact, a two- to threefold stimulation of this activity (which results in H,O, formation) was observed. In addition to imidazolylacetate, imidazolylpropionate and imidazolyllactate were found to stimulate the rate of NADH oxidation; however, in the absence of product analysis it is not clear whether these compounds function as true substrates or nonsubstrate effectors (91). In common with other external flavoprotein monooxygenases, imidazolylacetate monooxygenase is not inhibited by metal chelators nor does it contain any significant quantities of trace metals. It is unusual among this group of enzymes however inasmuch as the addition of substrate is without effect on the visible absorption spectrum and on the CD and ORD spectra (911.

I. BACTERIAL LUCIFERASE The phenomenon of bioluminescence emission exhibited by many bacteria has attracted much experimental attention. No attempt will be made 89. Y. Maki, S. Yamamoto, M. Nosaki, and 0. Hayaishi, BBRC 25,609 (1966). 90.S. Rothberg and 0. Hayaishi, JBC 229, 897 (1957). 91. Y . Maki, S. Yamamoto, M. Nosaki, and 0. Hayaishi, JBC 244,2942 (1969). 92. H. Okamoto, M. Noraki, and 0. Hayaishi, BBRC 32, 30 (1968).

4.

227

FLAVIN AND PTERIDINE MONOOXYGENASES

here to review the rather extensive literature on the subject or to consider the possible mechanisms of the light emitting process. The reader is referred to a recent review article for such information (93). It has long been known that the light emission depends on the presence with the bacterial luciferase of FMNH,, 0,, and a long-chain aldehyde (94,95). Although the aldehyde is essential for high quantum yields, its fate during the reaction remained obscure until recently. It was proposed by McElroy and Green in 1955 (96) that the aldehyde is converted to the corresponding long-chain acid since this reaction would provide sufficient energy for the emission of a quantum of light of 490 nm. Experimental support for this proposal came recently from the detection of acid production by mass spectrometry (97). Further definitive proof of acid production was obtained independently by McCapra and Hysert (98) and by Dunn et d.(99). These studies have thus established the luciferase reaction as a hydroxylation of the long-chain aldehyde: FMNHz

+ RCHO +

0

2

4

FMN

+ RCDOH + HzO

Luciferase may therefore be classified as a somewhat unusual flavoprotein monooxygenase, its unique feature being that FMNH, appears to function as a substrate rather than a prosthetic group. However, this is clearly a matter of semantics. The dissociation constant for binding of FMNH, to the enzyme from P. fischeri has been estimated as 9.7 X lo-? M ; a value of 8.0 x lo-? has been found for the so-called MAV-luciferase (100). Baldwin has recently reported that F M N also binds to the latter enzyme, although much less strongly; a Kd value of 2.4 X M was calculated (101). Although most experiments with the isolated enzyme have employed added FMNH, as the electron donor, the luminescent bacteria utilize a NADH-FMN reductase to produce the FMNH, used in the luciferase reaction (93). Thus, the combined effects of the two enzymes is formally the same as for all of the external flavoprotein monooxygenases so far discussed. 93. J. W. Hastings, Annu. Rev. Biochem. 37,597 (1968). 94. M. J. Cormier and B. L. Strehler, JACS 75, 4864 (1953). 95. B. L. Strehler, E. N. Harvey, J. J. Chang, and M. J. Cormier, Proc. Nut. Acad. Sci. U . S . 40, 10 (1954). 96. W. D. McElroy and A. A. Green, ABB 56, 240 (1955). 97. 0. Shimomura, F. H. Johnson, and Y. Kohama, Proc. Nut. Acad. Sci. U . S . 69, 2086 (1972). 98. F. MeCapra and D. W. Hysert, BBRC 52, 298 (1973). 99. D. K. Dunn, G. A. Michaliszyn, I. G. Bogacki, and E. A. Meighen, Biochemistry 12, 4911 (1973). 100. E. A. Meighen and J. W. Hastings, JBC 246, 7666 (1971). 101. T. 0. Baldwin, BBRC 57, lo00 (1974).

228

VINCENT MASSEY AND PETER HEMMERICH

From their early work employing rapid reaction spectrophotometry, Hastings and Gibson (102) concluded that a long-lived intermediate is formed on mixing luciferase, FMNH,, and 02,which reacts further, with concomitant light emission, in the presence of a long-chain aldehyde. Competing with the formation of this intermediate is the rapid reaction of free FMNH, and 0, to yield F M N and H,O,. I n a recent paper the existence and spectral characterization of the proposed enzyme intermediate was shown in a very elegant fashion (103).A mixture of luciferase, FMNH,, and 0, in 50% ethylene glycol-phosphate buffer was allowed to react at 4 O for 10 sec and then the temperature was lowered rapidly to -2OO. The mixture was then chromatographed on a Sephadex L H 20 column a t -20° to separate enzyme-bound intermediates cleanly from free FMN. The results are very nicely consistent with the following scheme:

FMN

.+1

H,O,

E

lkd

+ FMN + H,O,

+

Ikb

E FMN+ H,O +acid hu

+

The intermediate I1 isolated by low temperature chromatography was shown to have an absorption spectrum with a maximum a t 370 nm and comparatively little absorption at 450 nm. Its spectral characteristics are in fact very similar to those of the oxygenated flavin intermediates found with p-hydroxybenzoate hydroxylase and phenol hydroxylase described in the previous sections. The intermediate is stable for long periods a t -20° ; on warming in the absence of aldehyde, enzyme, FMN, and H20, are formed (pathway k d ). I n the presence of aldehyde however the characteristic light emission of the luciferase reaction (490 nm) is observed, consistent with the formation of a further intermediate I I a and breakdown to products via the pathway labeled kb. The physical isolation of an oxygenated flavin intermediate in this case, as opposed to the kinetic “isolation” with the other hydroxylases, offers possibilities of incisive mechanistic studies. Extension of this isolation technique t o other enzymes, particularly with substrates being turned over only slowly, is 102. J. W. Hastings and Q. H. Gibson, JBC 238,2537 (1963). 103. J. W. Hastings, C. Balny, C. Le Peuch, and P. Douznu, Proc. N a t . Acad. Sci. U.8. 70, 3468 (1973).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

229

clearly an attractive experimental approach to investigating further the reaction mechanisms of these enzymes.

J. MICROSOMAL AMINEOXIDASE All of the flavoprotein monooxygenases so far discussed are of microbial origin. I n recent years, Ziegler and his colleagues have reported on a monooxygenase present in liver microsomes of many vertebrate species (104-107).This enzyme appears to be a simple flavoprotein containing no significant quantities of metal ions or heme residues. The enzyme has been isolated from pig liver microsomes and is reported to have a molecular weight of approximately 500,000 (106). The minimum molecular weight per FAD prosthetic group is 71,000. Thus, the enzyme would appear to he a polymeric, aggregating species. The enzyme catalyzes the NADPH- and 0,-dependent N-oxidation of a variety of secondary and tertiary amines. The secondary amines are oxidized to the corresponding hydroxylamines and the tertiary amines to amine oxides, e.g.,

N-Methylaniline

N, N-Dimethylaniline

N - Methylpheny Lhydroxylamine

N, N-Dimethylaniline - N-oxide

The primary amines, 1-naphthylamine and 2-naphthylamine, are also oxidized but a t lower rates (107). With one substrate, dimethyloctylamine, sigmoidal kinetics are found, suggesting the existence of an effector 104. J. M. Machinist, E. W. Dehner, and D. M. Ziegler, ABB 125, 858 (1968). 105. D. M. Ziegler, D. Jollow, and D. E. Cook, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 507. Univ. Park Press, Baltimore, Maryland, 1971. 106. D. M. Ziegler and C. H. Mitchell, ABB 150, 116 (1972). 107. D. M. Ziegler, L. L. Poulson, and E. M. McKee, Xenobiotica 1, 523 (1971).

230

VINCENT MASSEY AND PETER HEMMERICH

site in addition to the catalytic site. This idea is supported by the finding of a variety of compounds, which are themselves not substrates, but which increase the rate of oxidation of true substrates (106).

K.

KYNURENINE-3-HYDROXYLASE

Kynurenine-hydroxylase has been purified partially from rat liver mitochondria (108).Okamoto et al. (109) showed that the enzyme is localized in the outer membrane of the mitochondrion. Several workers, have, as a result, used kynurenine-hydroxylase activity as a mitochondria1 marker (see, for example, 110,111) . Experiments with lsO2 have established the enzyme as a true monooxygenase (112,115): 0 a I ; CII H r

CH-COOH I NH,

+

NADPH

+

HC

+

NADP’

+

H,O

+

0,

~-Kynurenine

qC!Z,C&0 II

CH-CCOOH I

NH*

OH 3 - Hydroxy - ~-kynurenine

The enzyme can use NADH as an alternative electron donor (114). Acid ammonium sulfate treatment of the partially purified enzyme resulted in a decrease in activity to 60% the original level. Addition of FAD restored almost completely the initial activity, while FMN was ineffective (109,11$1l5). There is thus circumstantial evidence that the 108. H. Okamoto, “Methods in Enzymology,” Vol. 174, p. 460,1970. 109. H. Okamoto, S. Yamamoto, M. Nozaki, and 0. Hayaishi, BBRC 26, 309 (1967). 110. D. S. Beattie, BBRC 31, 901 (1968). 111. C. A. Schnactman and J. W. Greenawalt, J. Cell Biol. 38, 158 (1968). 112. Y. Saiton, 0. Hayaishi, and S. Rothberg, JBC 229, 921 (1957). 113. H. Okamoto, in “Flavins and Flavoproteins” (K. Yagi, ed.), p. 223. Univ. of Tokyo Press, Tokyo, 1968. 114. 0. Hayaishi and H. Okamoto, Amer. J . Clin. Nutr. 24, 805 (1971). 115. .H. Okamoto and 0. Hayaishi, BBRC 29, 394 (1967).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

23 1

enzyme is a FAD-containing flavoprotein. Whether or not FAD is the only prosthetic group will obviously have to await more extensive purification of the enzyme. IV. Pterin-Linked Monooxygenases

A .small group of metabolically important enzymes has been shown to require a reduced unconjugated pterin cofactor as electron donor in the aromatic ring hydroxylation reactions which they catalyze. Because of the rather similar chemistry of flavins and pteridines i t is a widely considered possibility that these enzymes may function by mechanisms basically similar to those of the flavoprotein monooxygenases. While this possibility is real, there are several important distinctions which may be made between the properties of the pteridine and flavin monooxygenases and between the properties of tetrahydropteridines and dihydroflavins. 1. While the flavoprotein monooxygenases have been shown to be free of trace metal involvement, the pteridine monooxygenases all seem to contain and require protein-bound iron as an additional cofactor. 2. I n the case of the flavoprotein monooxygenases the flavin is tightly bound to the protein, and the protein is very selective in binding only FAD or FMN. With the pteridine enzymes on the other hand, the pteridine is bound only weakly to the protein, and hence can be considered more as a mobile substrate than as a prosthetic group. I n addition, a large number of unsubstituted tetrahydropteridines will function in the catalytic reaction. In this connection the pteridine monooxygenases are more akin to bacterial luciferase (see Section II1,I) than to the rest of the flavoprotein monooxygenases. 3. With the exception of bacterial luciferase, the reactions catalyzed by the flavoprotein monooxygenases involve a single enzyme. In the case of the pteridine-linked hydroxylations a t least two enzymes are involved in the catalytic reaction, the hydroxylase proper, where the tetrahydropteridine is oxidized to a dihydropteridine in the couse of the hydroxylation reaction, and a tetrahydropteridine regenerating enzyme, NADHdihydropteridine reductase. 4. The reactivity of dihydroflavins with molecular oxygen is rather high (tM < 1 sec) while that of tetrahydropteridines is rather low ( t ~ in the order of minutes). Unless there is a dramatic enhancement of this rate on binding the tetrahydropteridine to the protein, such rates would appear unlikely to be great enough to sustain catalysis.

232

VINCENT MASSEY AND PETER HEMMERICH

A. PHENYLALANINE HYDROXYLASE Phenylalanine hydroxylase was the first discovered and most widely studied enzyme of this group. Studies from several laboratories in the early 1950’s showed the conversion of phenylalanine to tyrosine in rat liver extracts supplemented with pyridine nucleotides (116,117). I n 1957 Kaufman (118) established the following stoichiometry in a partially purified system: GPhenylalanine

+ NADPH + H+ + O2+ Gtyrosine + NADPf + H20

This stoichiometry suggested that the reaction is of the monooxygenase type, i.e., that the phenolic group of tyrosine is derived from 0, rather than H,O. This conclusion was shown to be correct by 1802 studies; when the hydroxylation reaction was carried out in the presence of lRO2the phenolic group was found to be labeled, but no labeling was found when the reaction was carried out with l6OZand H2180 (119). The specificity of this reaction for NADPH rather than NADH remained a matter of uncertainty for many years. Kaufman and his colleagues found that NADPH was the more efficient electron donor in his system (f18), whereas Mitoma reported NADH to be more effective (117). This discrepancy now appears to be satisfactorily explained by the recognition of two enzymes being involved in the overall reaction in addition to the actual phenylalanine hydroxylase. Kaufman established that an unconjugated pterin, tetrahydrobiopterin, is an essential cofactor in the hydroxylation reaction (120).In the course of the reaction this is converted to an oxidized form, identified as a quinonoid isomer of dihydrobiopterin (lal).

Tetrahydrobiopterin

o-Quinonoid dihydrobiopterin

116. S.Udenfriend and J. R. Cooper, JBC 194, 503 (1952). 117. C.Mitoma, A B B 60, 476 (1956). 118. 8. Kaufman, JBC 226, Fill (1957). 119. S.Kaufman, W.F. Bridgers, F. Eisenberg, and S. Friedman, BBRC 9, 497 (1962). 120. S.Kaufman, Proc. Nat. Acad. Sci. U . S. 50, 1085 (1963). 121. S.Kaufman, JBC 239, 332 (1964).

4.

233

FLAVIN AND PTERIDINE MONOOXYGENASES

Hence, the primary hydroxylase reaction may be written : H I CH,- C-COOH

H I CH,- C-COOH +

pheny lalanine tetrahydrobiopterin + 0, hydroxylase

OH L-Pheny lalanine

L-Tyrosine

+

quinonoid dihydrobiopterin

+

H,O

In order for this enzyme to function catalytically, the quinonoid dihydrobiopterin has to be reduced again. This is accomplished by dihydropteridine reductase, an enzyme long recognized t o participate in the overall reaction but only recently purified (122). This enzyme has been found to function much more efficiently with NADH (lower K , values and higher V,,, values) than with NADPH: Quinonoid dihydrobiopterin

+ NADH + H+ dihydropteridine reductase >

tetrahydrobiopterin

+N A P

The quinonoid dihydrobiopterin can rearrange nonenzymically to 7,8-dihydrobiopterin, which is the form of the cofactor isolated from rat liver (120).

7'

cH$-c-c'?XN&N,, I

H

I

H

I

NyNH2 0

When this form of the cofactor is present a third enzyme, identified as dihydrofolate reductase, is required for reduction to the physiologically active tetrahydrobiopterin (13). This enzyme is NADPH-specific: 7,ti-Dihydrobiopterin

-

+ NADPH + H+ dihydrofolate tetrahydrobiopterin + NADP+ reductase

Hence, the pyridine nucleotide specificity depends on the form of the cofactor present. However, it should be pointed out that the role of dihydrofolate reductase is as a scavenger of any dihydropteridine which has 122. J. E. Craine, E. S. Hall, and S. Kaufman, JBC 247, 6082 (1972).

234

VINCENT MASSEY AND PETER HEMMERICH

escaped from the quinonoid form and that in the presence of sufficient dihydropteridine reductase it plays only a minor role (13).It would therefore be more proper to restate the overall stoichiometry carried out hy phenylalanine hydroxylase and dihydropteridine reductase to be Phenylalanine

+ N A D H + Hf + 0 %+ tyrosine + N A D + + H20

The substrate specificity of phenylalanine hydroxylase is complicated by the fact that the rate of substrate oxidation is dependent on the nature of the tetrahydropteridine employed as cofactor (123), on the presence or absence of lysolecithin, whose effect is also dependent on the tetrahydropteridine cofactor employed ( l a d ) , and on the presence or absence of another protein recently isolated from liver known as the phenylalanine hydroxylase stimulating factor (PHS) (125,126).The effect of this protein also depends on the nature of the pteridine cofactor used. An additional complication arises from the fact that with several substrates increased rates of pyridine nucleotide oxidation occur (in the two enzyme system) without complete coupling to hydroxylation of the substrate. I n this case the product of 0, reduction is H,O,, a situation analogous to that found with several flavoprotein monooxygenases (see previous sections). I n fact, the phenomenon of “uncoupling” hydroxylation from oxidation with the phenylalanine hydroxylating system predated by several years (127) the discovery of this phenomenon with the flavoprotein monooxygenases. Again this effect depends on the nature of the substrate, the tetrahydropteridine, the presence or absence of lysolecithin, or the presence or absence of the PHS protein (128). The following compounds have been shown to be hydroxylated, a t least partially : tryptophan (129) p-2-thienylalanine (130), 4-chlorophenylalanine, 2-fluorophenalalanine1 3-fluorophenylalanine1 and 4-fluorophenylalanine (131). With the 4-fluorophenylalanine the products have been identified as tyrosine and F- (131). In addition, a number of p-substituted phenylalanines have been found to be hydroxylated with migration and retention of the p substituent (13%’).This demonstration of the “NIH 123.C.B. Storm and S. Kaufman, BBRC 32, 788 (1968). 124.D.B. Fisher and S. Kaufman, JBC 248,4345 (1973). 125. C.Y.Huang, E. E. Max, and S. Kaufman, JBC 248,4235 (1973). 126. C.Y.Huang and S. Kaufman, JBC 248, 4242 (1973). 127. S. Kaufman, BBA 51, 619 (1961). 128. D.€9. Fisher and S. Kaufman, JBC 248, 4300 (1973). 129. R.A. Freedland, I. M. Wadzinski, and H.A. Waisman, BBRC 5, 94 (1961). 130. S. Kaufman, “Methods in Enzymology,” Vol. 5, p. 802, 1962. 131. S. Kaufman, BBA 51, 619 (1961). 132. G. Guroff, C. A . Reifsnyder, and J. W. Daly, BBRC 24, 720 (1966).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

235

shift” phenomenon (133) strongly implies the formation of arene oxide intermediates in the hydroxylation reaction. This will be considered further in the final section. B y varying the structure of the tetrahydropteridine cofactor, the extent of coupling the pteridine oxidation and hydroxylation of phenylalanine was determined. I n addition to the natural cofactor, 6-methyl and 6,7-dimethyl tetrahydropterin show tight coupling. I n contrast, 7-methyl, 7-phenyl, or unsubstituted tetrahydropterin show loose coupling (123). These results might imply that an alkyl group a t the 6 position is necessary for tight coupling to occur. However, substantial uncoupling of hydroxylation is observed with 4-fluorophenylalanine or tryptophan when 6-methyl tetrahydropterin is used as cofactor (123), making such generalizations somewhat untenable. When 6,7-dimethyl tetrahydropterin is employed as cofactor the initial rate of hydroxylation vs. phenylalanine concentration is hyperbolic. I n contrast, when the natural tetrahydrobiopterin is used as cofactor, the substrate saturation curve is sigmoidal ( l a d ) , Similar sigmoidal kinetics were reported with tryptophan as substrate, and the abolition of this complex behavior by 1-propanol ( 1 3 4 ) . This observation led Fisher and Kaufman (12-4) to explore the effects of various fatty acids and derivatives on the rates and kinetics of various reactions catalyzed by the hydroxylase. It was found that 1-propanol, 1-butanol, and a variety of fatty acids containing 16 carbon atoms or more gave substantial increases in catalytic activity when tetrahydrobiopterin was the cofactor, but were without effect with 6,7-dimethylpterin as cofactor. Long-chain acyl-CoA derivatives were also effective, as well as mixtures of bile salts and fatty acids. The greatest stimulations (of the order of 20-fold) were observed with phospholipids such as lysolecithin and lysophosphatidylserine. While the stimulating effect of lysolecithin on phenylalanine hydroxylation was exhibited only when tetrahydrobiopterin was used as cofactor, increased hydroxylation rates of other substrates were found even with 6,7-dimethyl tetrahydropterin as cofactor. Thus tryptophan hydroxylation is increased dramatically, especially a t low concentration of tryptophan ( 1 2 4 ) . The extent of uncoupling of hydroxylation and oxidation with this substrate remains unchanged, however. I n the presence of lysolecithin, with either tetrahydropteridine cofactor, the rate of hydroxylation of m-tyrosine was increased some 40-fold. Dopa, the product of 133. G. Guroff, D. Jerina, J. Rensen, S. Udenfriend, and B. Witkop, Science 157, 1524 (1967). 134. P. A . Sullivan, N. Kester, and S. J. Norton, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 30, 1067 (1971).

236

VINCENT MASSEY AND PETER HEMMERICH

this hydroxylation reaction, was formed stoichiometrically with respect to oxidation of NADPH, and the reaction is therefore tightly coupled (124). I n contrast, when (p)-tyrosine was tested as a substrate in the presence of lysolecithin, the rate of NADPH oxidation was greatly stimulated, but the tyrosine remained unchanged. Hence, in the presence of lysolecithin, tyrosine behaves as a nonsubstrate effector. This effect was shown to be the direct result of an increased rate of oxidation (producing H202)of the tetrahydropteridine cofactor (128). The stimulating effects of lysolecithin can also be mimicked by preincubation of phenalalanine hydroxylase with chymotrypsin (134).Lysolecithin exposes a thiol group of the enzyme which is unreactive to DTNB in the untreated enzyme (124). Fisher and Kaufman have interpreted these results as indicating that the hydroxylase contains a polypeptide portion which can act as an internal regulator of enzymic activity. It was proposed that the polypeptide can be either displaced reversibly from its inhibitory site by the detergent action of a lipid or can be irreversibly removed by chymotrypsin. If such a regulatory role does indeed operate it must be rather subtle, in view of the big differences in effects found depending on the nature of the tetrahydropteridine cofactor and the substrate used. Further insight into the details of the hydroxylation reaction is promised by the finding of yet another regulator of the enzymic activity. Studies by Kaufman and his colleagues have shown that even in the presence of lysolecithin, the specific activity of phenylalanine hydroxylation decreases sharply if the concentration of enzyme is increased (124). This effect (which was exhibited only with tetrahydrobiopterin as cofactor) was shown to be abolished by a phenylalanine hydroxylase stimulator (PHS) present in liver extracts. This factor has now been purified and shown to be a protein of molecular weight 51,500 (125). Evidence has been presented that PHS is an enzyme which catalyzes the breakdown of an intermediate in the hydroxylation reaction (126). The evidence presented, mainly of a kinetic nature, shows that the effect of PHS is unlikely to result from removal of an inhibitor in the hydroxylase preparations or from effects on the state of aggregation of the hydroxylase. Persuasive evidence is given for the reversible release from the hydroxylase of an intermediate which is subsequently converted nonenzymically to the products of the hydroxylation reaction. In this model PHS serves as an auxiliary enzyme catalyzing the breakdown of the intermediate : ki

ki

E+S=ES=E+S’ k-i

S’

k-z

k 2 p’

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

237

Under the experimental conditions where the phenomenon is observed (approximately equimolar concentrations of enzyme and tetrahydrobiopterin) the reverse step k-, can become significant, thus accounting for the observed decrease in specific activity as the hydroxylase concentration is increased. The fact that this effect is not found with 6,7-dimethyl tetrahydropterin would be explained simply by a lower value of k-, and/or an increased value of k , for this cofactor. Such arguments clearly rule out the possibility that S’ is a simple arene oxide intermediate of phenylalanine/tyrosine and suggest that S’ represents some complex or compound of the pterin, substrate, and O2 which is an intermediate in the hydroxylation reaction. The surprising conclusions from these studies are that such an intermediate is released from the enzyme before undergoing conversion to products, and the slow rate of its breakdown ( k , estimated as approximately 2.4 min-l). This latter value is almost 20 times slower than the calculated rate of release of S’ from the enzyme. Under suitable conditions it might therefore be possible to test this hypothesis directly ; for example, by initiating the reaction and then cooling rapidly, followed by low temperature chromatography (see results with luciferase, Section II1,I) it would be possible to isolate the intermediate and study its properties. Phenylalanine hydroxylase has been purified from rat liver to a state close t o homogeneity (135). The enzyme appears t o have a molecular weight around 100,000, and to be composed of two subunits of molecular weight 55,000. The enzyme has recently shown to contain iron a t the level of 1-2 atoms per molecule of protein (136‘). The enzyme is inhibited by metal chelators and substantial removal of the iron was achieved by incubation with o-phenanthroline and precipitation with ammonium sulfate. The activity was restored only by the addition of FeCl,. The enzyme was found to exhibit an EPR signal a t g = 4.23 attributable to high-spin ferric iron. This signal was substantially abolished when all three substrates were present under turnover conditions, suggesting that the iron may be reduced in the catalytic cycle, or, less probably, that a change in its ligand field results in a change from high- to low-spin state of the ferric iron (136). Unfortunately, no studies were carried out with tetrahydropteridine or phenylalanine added separately, which would be expected to better define the role of the iron. A steady-state kinetic study of the enzyme by Kaufman and Fisher (IS) indicated that a random order quaternary complex mechanism was operational (i.e., that a complex of all three substrates with the enzyme is 135. S. Kaufman and D. B. Fisher, JBC 245,4745 (1970). 136. D. B. Fisher, R. Kirkwood, and S. Kaufman, JBC 247, 5161 (1972).

238

VINCENT MASSEY AND PETER HEMMERICH

formed in the catalytic cycle). It should be noted, however, that these results are a t variance with other studies on the enzyme, which indicated a ping-pong type of mechanism (137).Kaufinan and Fisher (13) have reasoned that their kinetic analysis is inconsistent with reduction of the enzyme. However, their results would be consistent with a mechanism in which the quaternary complex was composed of reduced enzyme, an oxidized form of tetrahydropteridine (perhaps the semiquinone) , phenylalanine, and 02.Clearly, rapid reaction studies, to follow the rate of disappearance of the EPR signal in the presence of various combinations of the substrates, would be very desirable. Equally desirable would be experiments in which enzyme, tetrahydropteridine, and phenylalanine would be mixed with oxygenated buffer, and spectral and EPR changes monitored by rapid reaction techniques. Such experiments have proved very useful in investigating the mechanisms of flavoprotein monooxygenases and should be applicable to the present system.

B. TYROSINE HYDROXYLASE I n 1964 the enzymic conversion of L-tyrosine to 3,4-dihydroxyphenyI-

alanine (dopa) was demonstrated in particles isolated from adrenal medulla, brain, and other sympathetically innervated tissues (138). With partially purified preparations from adrenal medulla, the requirement in the catalytic activity of a tetrahydropteridine cofactor was demonstrated (158,139).Direct proof of the monooxygenase nature of the enzyme came from lSO studies by Daly et al. (140), who showed that the oxygen atom inserted at the 3 position of the benzene ring is derived from molecular oxygen : CH,-

C -COOH

CH,- C-COOH ~

OH

tetrahydropteridine

+

+

O2-

OH

dihydropteridine

+ H,O

OH

The fact that tyrosine hydroxylase can be coupled to NADH oxidation by dihydropteridine reductase (139) strongly suggests that the dihydropteridine product is the quinonoid form (see Section IV,A) . The intracellular location of the enzyme has been a matter of some 137. V. G. Zannoni, I. Rivkin, and B. N. LaDu, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 26, 840 (1967). 138. T.Nagatsu, M.Levitt, and S. Udenfriend, JBC 239,2910 (1964). 139. A. R.Brennemann and S. Kaufman, BBRC 17, 177 (1964). 140. J. W. Dsly, M. Levitt, G. Guroff, and S. Udenfriend, ABB 126, 593 (1968).

4. FLAVIN

AND PTERIDINE MONOOXYGENASES

239

controversy. Nagatsu e t al. (138) , in their pioneering studies, concluded that in guinea pig brain and beef adrenal medulla, most if not all of the enzyme was particle bound, and that soluble enzyme found was the result of release by the homogenization method employed. More recent evidence indicates that in rat brain the enzyme exists in two forms, a soluble and a membrane-bound form (141). Soluble tyrosine hydroxylase has never been purified to any great extent: Almost all studies on the enzyme have been carried out with partially purified enzyme from bovine adrenal medulla which was solubilized by limited proteolysis with either trypsin (142) or chymotrypsin (143). The enzyme is inhibited by iron chelators such as ap-dipyridyl (138) and o-phenanthroline (1&) , but not by the nonchelating analog, m-phenanthroline (145). In addition, the enzymic activity is stimulated by Fe2+ (138). Shiman et al. (143) have pointed out that this stimulation is no valid evidence for participation of iron in the catalytic reaction since catalase has a similar effect. The activation by Fez+was therefore considered to result from removal of H,O, formed by autoxidation of the tetrahydropteridine. However, more recent studies, while admitting the validity of this argument, have produced convincing evidence that iron atoms are required in the catalytic function of the enzyme. Petrack e t al. (142,146') have demonstrated activation of the enzyme by ferrous iron; no other metal ion tested had a similar effect, nor did catalase or peroxidase. I n analogy with phenylalanine hydroxylase, a number of tetrahydropteridines will serve as electron donors with tetrahydrobiopterin being the most efficient in terms of lower K , value and higher V,,,, value. 6-Methyl and 6-7-dimethyl tetrahydropterin are both good donors (139). As would be expected, N(5)-substituted pteridines are inactive as cofactors; the tetrahydropteridine also has to be substituted with either a 2-amino or a 4-hydroxy group in order to function in the hydroxylation reaction (144). Like phenylalanine hydroxylase, the substrate specificity of tyrosine hydroxylase is fairly broad with relative rates depending on the pteridine cofactor used ; for example, phenylalanine is also hydroxylated to 141. 142. 143. 144. 145.

R. T. Kucaenski and A. J. Mandell, JBC 247, 3114 (1972). B. Petrack, F. Sheppy, and V. Fetaer, JBC 243, 743 (1968). R. Shiman, M. Akino, and S. Kaufman, JBC 246, 1330 (1971). L. Ellenbogen, R. J. Taylor, and G. B. Brundage, BBRC 19, 708 (1965). R. J. Taylor, C. S. Stubbs, and L. Ellenbogen, Biochem. Pharmacol. 18, 587

(1966). 146. B. Petrack, F. Sheppy, V, Fetaer, T. Manning, H. Chertock, and D. Ma, JBC 247, 4872 (1972).

240

VINCENT MASSEY AND PETER HEMMERICH

tyrosine a t a rate approximately the same as that of tyrosine hydroxylation when tetrahydrobiopterin is employed as the electron donor (143). However, with 6,7-dimethyltetrahydropterinthe rate is only about onetwentieth that of tyrosine hydroxylation (143). Tong et al. (147,148) have also reported that the conversion of m-tyrosine to dopa occurs at about 50% the rate of conversion of L-tyrosine to dopa. In addition, they found that phenylalanine hydroxylation resulted in approximately 15% formation of m-tyrosine and 85% of p-tyrosine. Studies with isotopically labeled substrate have demonstrated the N I H shift to be operational with this enzyme ( I . @ ) . When [4-aH]phenylalanine was employed as substrate, the tyrosine formed was found to have the tritium retained, indicating migration to the 3 and 5 positions. I n the subsequent conversion of [3,5JH] tyrosine to dopa, however, 50% of the tritium was lost (cf. Scheme 1, Section 111,~). Partial steady-state kinetic studies have been carried out with the enzyme. Using a partially purified soluble enzyme from adrenal medulla, Ikeda et al. (149) have found the kinetic behavior to be of the Ping Pong type, and concluded that a reduced form of the enzyme was involved in catalysis (presumably iron in the ferrous state). Omitting intermediate complexes, their results indicate the following sequence : E,.

+ tetrahydropteridine -+ Ered + dihydropteridine Ered

+ tyrosine + Oz+ E,, + dopa + H20

Working with a solubilieed enzyme, Joh et al. (150) concluded that the mechanism did not involve a reduced enzyme intermediate but rather a quaternary complex. Subsequent work by Shiman and Kaufman (quoted in 13) on both the particulate enzyme from bovine adrenal medulla and a purified solubilized enzyme also showed kinetic behavior indicative of the participation of a quaternary complex. While these experimental discrepancies are hard to rationalize, it should be pointed out that the finding of participation of a quaternary complex does not rule out reduction of the enzyme in the course of catalysis; such a complex could consist of reduced enzyme, oxidized pteridine, tyrosine, and 0,.

c. TRYPTOPHAN HYDROXYLASE (TRYPTOPHAN-5-MONOOXYGENASE) Much less information concerning the properties of tryptophan hydroxylase is available than with phenylalanine and tyrosine hydroxylases. Much of the literature deals with controversies about the subcellular loca147. J. H. Tong, A. D’Iorio, and N. L. Benoiton, BBRC 43, 819 (1971). 148. J. H. Tong, A. D’Iorio, and N. L. Benoiton, BBRC 44, 229 (1971). 149. M. Ikeda, K. A. Fahien, and S. Udenfriend, JBC 241,4452 (1966). 150. T. H. Joh, R. Kapit, and M. Goldstein, BBA 171, 378 (1969).

4.

241

FLAVIN AND PTERIDINE MONOOXYGENASES

tion of the enzyme and about whether a tetrahydropterin is indeed a cofactor. These aspects have been discussed extensively in a recent review by Kaufman and Fisher (13). The enzyme was first detected in brain by Grahame-Smith (151),who also partially purified the enzyme and demonstrated the requirement for a tetrahydropteridine cofactor (168). The relative paucity of information on the enzyme is the result of difficulties encountered in its assay and purification; the most highly purified preparation so far reported has been enriched only 10-fold over the starting material (153).Friedman et al. demonstrated convincingly that in the presence of reduced pyridine nucleotide and dihydropteridine reductase, tetrahydropteridines function catalytically in the reaction. They established the following stoichiometry (153): Tetrahydropteridine

+ tryptophan + 02

4

dihydropteridine 5-hydroxytryptophm

+

+ HtO

Like the other pterin-linked hydroxylases, several tetrahydropteridines were found to function as electron donors. However, tetrahydrobiopterin, the naturally occurring cofactor for phenylalanine hydroxylase, seems to be the most efficient donor, having itself a lower K , value than other tetrahydropteridines and also exhibiting lower K,, values for O2 and tryptophan than found in the presence of other pteridine cofactors (153,154). Although a definitive answer will have to await the availability of a more highly purified enzyme, there is evidence that iron atoms may be associated with the enzymic activity. Friedman et al. (153)have demonstrated that stimulation of the activity by Fez+is the result of removal of deleterious H,O,, but other workers have demonstrated substantial inhibition of the enzyme by the iron chelators, Tiron, a,a-dipyridyl, and o-phenanathroline (154,155). V. Model Studies and Possible Mechanisms

In a series of papers, Mager and Berends (151-161) have proposed a common pathway for hydroxylation reactions catalyzed by flavopro151. D.G.Grahame-Smith, BBRC 16, 586 (1964). 152. D.G.Grahame-Smith, BJ 105, 351 (1967). 153. P. A. Friedman, A. H. Kappelman, and S. Kaufman, JBC 247, 4165 (1972). 154. E.Jequier, D.S. Robinson, W. Lovenberg, and A. Sjoerdsrna, Biochem. Pharmacol. 18, 1071 (1969). 155. A. Ichiyama, S.Nakamura, Y.Nishisuka, and 0. Hayaishi, JBC 245, 1699 (1970). 156. H.I. X. Mager and W. Berends, Rec. Trav. Chim. Pays-Bas 84, 1329 (1965). 157. H. I. X. Mager, R. Addink, and W. Berends, Rec. Trav. Chim. Pays-Bas 86, 833 (1967).

242

VINCENT MASSEY AND PETER HEMMERICH

teins and the pteridine-linked hydroxylases. Their proposal involves the formation of an intermediate hydroperoxide on reaction of a tetrahydropteridine or dihydroflavin with 02: H

I n the above general formulation the N(8) of the pteridines is equivalent to the N(10) of the flavins. The evidence of Mager and Berends for this formulation came from measurements of the stoichiometry of 0, consumption in oxidation of N (8)-substituted tetrahydropteridines and N (10)-substituted dihydroisoallaxazines. Less than stoichiometric 0, consumption was observed, which was formulated to result from the following reactions :

+ +

AH2 02 4 AHOOH AH2 AHOOH -+ 2 A 2 HzO (nonpolar medium) AHOOH + [AH+ OOH-] + A H a 2 (polar medium)

+

+

+

(1) (2)

(3) where AH, represents either tetrahydropteridine or dihydroflavin. In this formulation a complete coupling of reactions (1) and (2) would lead to a stoichiometry of one molecule of 0, consumed for each two molecules of AH,, and a coupling of reactions (1) and (3) to a stoichiometry of one molecule of 0, for each molecule of AH, oxidized. While such a formulation is consistent with their results, the observed stoichiometries could also be explained : AH2 AH1

+ Oz+ A + HzOz + HzOz+

A

+ 2 HzO

(4)

(5) Indeed, there are several reports in the literature that at physiological pH values H,O, is a comparable or even more efficient oxidant of tetrahydropteridines than is 0, (13,f62). However, with dihydroflavins, H,O, is a much poorer oxidant than molecular oxygen (163); thus, a t least 158. H. I. X. Mager and W. Berends, Rec. Trav. Chim. Pays-Bas 91,611 (1972). 159. H. I. X. Mager and W. Berends, Ree. Trav. Chim. Pays-Bas 91, 630 (1972). 160. H. I. X. Mager and W. Berends, Rec. Trav. Chim.Pays-Bas 91, 1137 (1972). 161. H. I. X. Mager and W. Berends, Tetrahedron 30, 917 (1974). 162. J. A. Blair and A. J. Pearson, JCS, Perkin Trans. I p. 80 (1974). 163. M. Dixon, BBA 226,2.59 (1971).

4.

243

FLAVIN AND PTERIDINE MONOOXYGENASES

with flavins, reactions (1)-(3) provide a reasonable interpretation of t,he results. Mager and Berends (157) proposed that the hydroperoxide might act. as a hydroxylating agent in the presence of a suitable acceptor:

'AHOH' (AHOH)

-H*O

Evidence in favor of a peroxydihydro intermediate bearing the OOH residue a t the bridge carbon between two nitrogen atoms was claimed by Mager and Berends ( 1 5 7 ) , using an N(1)-alkylated flavin model ( l-RFlredH,Scheme 2) as starting material for autoxidation. First, such an alkylated intermediate can be handled safely in aprotic solution be-

244

VINCENT MASSEY AND PETER HEMMERICH

cause of enhanced solubility. Second, spontaneous splitting of HzOz is overcome in this case, since this would require a proton a t N (1). In the presence of even slight amounts of water, the reaction sequence in this autoxidation will be HZO RFlredH

+

0,-

RF1- ?

-

O

O

HzO2

H

u RF1-10a-OH-

spirohydrantoin

while in the absence of water the decay of peroxydihydroflavin may be assumed to be RFl-?-OOH

2

[RFl-lOa-OH]-

spirohydrantoin

Henbe, the spirohydantoin (SPH) is the final product of l-alkyl-dihydroflavin autoxidation in any case (164). It should be pointed out that the SPH rearrangement is an entirely irreversible reaction which destroys the fla‘vin system. N (1)-C (10a) cleavage in a potential biocatalytic intermediate RF1-lOa-XH, XH being any protic nuelophile, must, therefore be strictly excluded biologically. In the above formulation a question mark has been put as to the position of OOH fixation in the 1-RFl,,-nucleus since (cf. Schemes 2 and 3) obviously any of the vinylogous positions 6, 8, 9a, and 10a might do. If there exists a rapid equilibrium between those possible isomers, i.e., a low activation barrier for shifts of nucleophiles like OOH- and OHon the flavin surface, the product SPH arising from irreversible decay of RF1-lOa-OH does not give any answer as to the structure of the actual oxygenating intermediate RFl-?-OOH. Such a low activation barrier for group migrations on the flavin surface has indeed been demonstrated by numerous papers of the Hemmerich group (for review, cf. 166,166). Since SPH, as mentioned above, is formed in any case as final product, the incorporation of lSO from lSO2 in the SPH-carbonyl could only then have evidential value for a RF1-10a-00H isomer if it were occurring efficiently in the presence of water. Under aqueous conditions, however, Blair and Pearson (162) demonstrated clearly that the carbonyl of the SPH is derived from water, not from 02. Under anhydrous conditions, as used by Mager and Berends (167),lSO from lSOz must trivially be found in SPH since no other relevant source of oxygen atoms is available. 164. K. H. Dudley and P. Hemmerich, J . Org. Chem. 32, 3049 (1967). 165. P. Hemmerich and W. Haas, in “Structure and Properties of Reduced Flavins”

(K. Yagi, ed.). Univ. of Tokyo Press, Tokyo (in preas).

166. P. Hemmerich and M. Schuman-Jorns, in “Enzymes: Structure and Function” (C. Veeger, J. Drenth, and R. A. Oosterbaan, eds.), p. 95. North-Holland Publ., Amsterdam, 1973; FEBS Symp. 29,95 (1973).

4.

245

FLAVIN AND PTERIDINE MONOOXYGENASES

,

OXIDASES

I

DEHYDROGENASES

nonessential {I; red

essential w k , t i u e

PEROXIDE heterolytic cleavage

H*

+G

SUPEROXIDE

I

N(I) blocked

N(51 blocked

SCHEME 3

Hence, the question as to the isomer structure of RF1-?-00H remains open : Miiller (personal communication) has meanwhile excluded positions 6 and 8 to be involved be means of thorough NMR studies of his “alcohol adducts” Rl-?-OR’(168), for which he initially claimed a 10a structure in line with Mager and Berends, but again without substantial evidence. Unfortunately, no facile differentiation between positions 9a and 10a can be made by NMR. A proposal as to the solution of this problem will be made below. Mager and Berends have found hydroxylation of phenylalanine using either tetrahydropteridines or dihydroflavins and molecular oxygen or hydrogen peroxide (I61) . They postulated the hydroxylating species to be hydroxy radicals. However, no convincing evidence is given for the 167. H. I. X. Mager and W. Berends, Tetrahedron Lett. 41, 4051 (1973). 168. F. Miiller, in “Flavins and Flavoproteins” (H. Kamin ed.), p. 363. Univ. Park

Press, Baltimore, Maryland, 1971.

246

VINCENT MASSEY AND PETER HEMMERICH

involvement of hydroxy radicals. But if it is assumed that the OH hypothesis is correct, two flavin molecules would be required for OH generation from molecular oxygen:

+

RFhH 02 + RFl-?-00H RFI-7-00H RFlredH + RFlOH

+

+ RFI + OH

This stoichiometry, involving stoichiometric flavin radical formation and “interflavin contact,” would make unlikely any biological importance of this “model” reaction. Furthermore, OH is such a reactive species that biocatalytic specificity of OH-involving processes could only be maintained if the radical was not allowed to diffuse away from the center of formation. But in that case it would be difficult to judge whether spin decoupling was indeed occurring in the catalytic pathway. Hence, oxygenation must occur in a quaternary complex made up from flavin, oxygen, substrate, and apoprotein, and the differentiation of OH- transfer on the one hand and oxygen at,om (“oxene” or OH+) transfer on the other hand turns into semantics. The true problem is as to the chemical structure of this active complex. The first experimental evidence indicating formation of oxygenated flavins came from rapid kinetic studies on the chemical reaction of reduced flavins with 0, (1-3), which was found to be an unexpectedly complex process involving the formation of peroxydihydroflavin and its decay into flavin radical and superoxide anion. The latter was found to be an even beter oxidant of reduced flavin than 0,, resulting in an autocatalytic reaction. The following series of steps was found to be minimal in describing the overall reaction (1,3,169) :

In a study of the reaction of various flavoproteins with 0, it has been found (4,s) that all dehydrogenases tested yield 0,- and the neutral flavin radical. On the other hand, no evidence for intermediate flavin 169. P. Hemmerich, A. P. Bhaduri, G. Blankenhorn, M. Brustlein, W. Haaa, and

W.-R. Knappe, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 3. Univ. Park Press, Baltimore, Maryland, 1973.

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

247

radicals or of 0,- could be found with flavoprotein oxidases, suggesting that with this group of enzymes the peroxydihydroflavin must undergo a heterolytic cleavage to yield directly the oxidized flavin and H,O,. This group of flavoproteins can also be characterized by their ability to stabilize the red-colored flavin semiquinone anion a t artificial half-reduction (170). The possibility of participation of 0,- in flavoprotein-catalyzed hydroxylation reactions has been considered. Massey and co-workers could find no evidence for 02-production with p-hydroxybenzoate hydroxylase or melilotate hydroxylase ; furthermore, superoxide dismutase had no effect on the hydroxylation reaction ( 4 ) .Similarly, White-Stevens and Kamin found no evidence with salicylate hydroxylase for the participation of 0,- or of hydroxy radicals ( 5 3 ) .On the other hand, Prema Kumar et al. (88) found that m-hydroxybenzoate-4-hydroxylase was inhibited completely a t fairly low levels of superoxide dismutase. While their interpretation of this result was that 0,- in some way functions as a hydroxylating agent we want to emphasize the possibility that superoxide dismutase may serve to catalyze the breakdown of a peroxydihydroflavin intermediate, e.g., by displacing the equilibrium (b) above. I n this case one would expect little or no change in the rate of NADPH oxidation, which should then be “uncoupled” from the hydroxylation reaction. Unfortunately, no information was given concerning this point. Strickland and Massey (171) found that hydroxylation of aromatic compounds such as p-hydroxybenzoate could be achieved a t physiological pH in the presence of model dihydroflavins and 0,. This reaction was inhibited substantially (75-100%) by superoxide dismutase. It seems unlikely that 0,- was the active hydroxylating agent however, since no hydroxylation was obtained by infusion of electrolytically generated 0,into a solution of p-hydroxybenzoate. These results again suggest that the active hydroxylating species must be the peroxydihydroflavin and that the inhibiting effect of superoxide dismutase in some way results from the destruction of this species, e.g., by removing 0,- from the equilibrium reaction (b) shown above for the autoxidation of dihydroflavins. Whatever the mechanism, there is little doubt that peroxydihydroflavins are active participants in flavoprotein-catalyzed hydroxylation reactions. As detailed in previous sections, transient species with very similar spectral properties have now been detected with three flavoprotein hydroxylases and shown to participate in the hydroxylation reactions. 170. V. Massey and G. Palmer, Biochemistry 5, 3181 (1966). 171. S. Strickland and V. Massey, in “Oxidases and Related Redox Systems” (T. E. King, H. S.Mason, and M. Morrison, eds.), Vol 1, p. 189. Univ. Park Press, Baltimore, Maryland, 1973.

248

VINCENT MASSEY AND PETER HEMMERICH

I n addition, a similar oxygenated flavin derivative has actually been isolated by low temperature chromatography in the case of bacterial luciferase (103). The important question yet to be answered definitely is the position of substitution in the flavin ring system and the mechanism by which oxygen is withdrawn from this intermediate and inserted into the aromatic substrate. Hamilton (14,86) has proposed a “vinylogous ozone” mechanism involving a nucleophilic attack of the aromatic substrate to a ring-opened form of a 4a-peroxydihydroflavin. No experimental evidence exists to support this hypothesis. While it is consistent with the introduction of a hydroxyl residue ortho to the original hydroxy group of the substrate (a phenomenon found with most flavoprotein hydroxylases) it could not account for the para substitution occurring with m-hydroxybenzoate hydroxylase (see Section II1,F). The theoretically possible HF1-00H isomers (cf. Scheme 3) must ab initio be separated into two subgroups, according to whether the position of proton fixation is N(5) or N ( l ) , i.e., one subgroup containing only the isomer 5-HF1-4a-OOH and the second containing four isomers l-HF1-6,8,9a,lOa-OOH. The first group isomer yields upon homolytic cleavage the blue (172,173), chemically stable, and biologically essential radical 5-HFlH, which is clearly associated with the “dehydrogenase” subclass of flavoproteins (5,170). This homolytic cleavage is easy because the spin density a t C(4a) is high (174) in the radical 5-HF1. The isomers of the second group would yield upon homolytic cleavage the tautomeric red (175,176), chemically unstable, and biologically nonessential radical l-HF1 or-the latter being strongly acidic-its anion F1-. This HF1-00H subgroup must, therefore, be associated with the oxidase and oxygenase subclasses of flavoproteins in keeping with their stabilization of the red flavin radical (5,170) arising from artificial l-e- oxidation or reduction, but not from reaction of the reduced enzymes with 0,. Furthermore, the question of which possible HF1-00H isomer is actually involved in flavin-dependent oxygenation requires at first the characterization of alkylated “model” flavin derivatives substituted in the respective positions 4a, 6, 8, 9a, 10a (cf. Scheme 3) by nucleophiles less 172. F. Muller, P. Hemmerich, A. Ehrenberg, G . Palmer, and V. Massey, Eur. J . Biochem. 14, 185 (1970). 173. F. Miiller, M. Briistlein, P. Hemmerich, V. Massey, and W. H. Walker, Eur. J. Biochem. 25, 573 (1972). 174. W. H. Walker, A. Ehrenberg, and J. M. Lhoste, BBA 215, 166 (1970). 175. A. Ehrenberg, F. Muller, and P. Hemmerich, Eur. J. Biochem. 2, 286 (1967). 176. F. Miiller, P. Hemmerich, and A. Ehrenberg, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 107. Univ. Park Press, Baltimore, Maryland, 1971.

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

249

reactive than OOH-. Such model compounds RF1-X will exhibit chromophores practically identical with the respective HF1-00H chromophores and allow their structural assignment by comparison of absorption and fluorescence spectra. Walker et al. (177) were first to find OH- as a stable model nucleo350-370 nm. Rephile, characterizing 5-RF1-4a-OH ( R = benzyl) A, placement of OH- by mercaptide RS- indeed gives rise to facile homolytic C (4a)-SR cleave, yielding the blue radical 5-RFl, as required for a flavoprotein dehydrogenase model. Muller (168) used CH,O- as a model nucleophile characterizing 1-RFl?-OCH, (arising from 1-RFlox+ OCH,-, Scheme 2) hmax 410430 nm, as mentioned above. After F . Muller’s (personal communication) exclusion of positions 6 and 8 by NMR (cf. above) the question mark stands for either 9a or 10a. Experimental evidence for discrimination between these two possibilities is not available presently, but the following chemical reasoning should apply. Hemmerich and Miiller (178) pointed out that in l-RFlos+ the azomethine-type center 9a will be thermodynamically favored for nucleophile attack over the amidine-type center 1Oa. Furthermore, it is clear from the spin density map (179) of flavin radicals that C(10a) has a negligible spin density as compared to C (4a) , since 14N(1) does not contribute by spin polarization to the E P R hyperfine pattern with a splitting of more than 1 Hz. The same should be true for reasons of symmetry with C( 9a ) , though direct EPR-evidence is hampered by the lack of a magnetically active nucleus at C (9). Closer inspection of the 1-HFl-9aOOH structure reveals an acidic center at N ( l ) H with a pK estimated to be <7. Hence, the 9a isomer should be present under physiological conditions as the anion HOO-ga-Fl-, thus giving rise to facile polar elimination of OOH-, which, a t least theoretically, would suggest the 9a-dihydroperoxide t o be the isomer associated with flavoprotein oddases. Flavoprotein oxygenases, however,-requiring a “peracid-type” active intermediate-should involve a HF1-00H intermediate where the carbon center bearing the OOH group is sp2 rather than sp3, as pointed out by McCapra and Hysert (98). Such an intermediate can most easily arise from HF1-lOa-OOH, e.g., by reversible opening of one of the C-N bonds adjacent to C (10a). Since, as mentioned previously, the C (10a)-N (1) bond can only be opened irreversibly, because of spirohydantoin forma-

+

177. W. H. Walker, P. Hemmerich, and V. Massey, Eur. J. Bwchem. 13, 258 (1970). 178. P. Hemmerich and F. Muller, Ann. N. Y. Acad. Sci. 212, 13 (1973). 179. F. Muller, P. Hemmerich, and A. Ehrenberg, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 107. Univ. Park Press, Baltimore, Maryalnd, 1971.

250

VINCENT MASSEY AND PETER HEMMERICH

tion (cf. Scheme 2 ) , the ring opening must occur a t the C (10a)-N (10) bond. Such ring opening is known to be strictly reversible from the mechanism of chemical flavin synthesis (180). Thus, the oxygenation reaction is envisaged to occur as shown in Scheme 4. “Oxene” insertion into the aromatic substrate (ArH) yields

AroH-l 0

”PER ACID”

- Ar H-complex

ArH

SCHEME 4

finally the hydroxylated product (ArOH), presumably through a series of intermediates (see Sections II1,B and C ) . The flavin is left in an “alloxane-aminoanil” state of overall composition HFlOH ; such compounds are known to recyclize very rapidly, with elimination of water, to yield reoxidieed flavin catalyst (180). Hemmerich and Miiller (178)have collected arguments for a bZue color of the l-HF1-10a-OOH chromophore, and indeed a blue (Amax 600 nm) derivative arising upon aprotic autoxidation of l-RFlredHhas been demonstrated as early as 1960 by Hemmerich et al. (181), and has been 180. P. Hemmerich, C. Veeger, and H. 4, 671 (1965).

C. 5. Wood, Angew. Chem., Int. Ed. Engl.

4.

FLAVLN AND PTERIDINE MONOOXYGENASES

251

readily mistaken for the radical 1-RF1 (181,182), later on shown to be red (cf. above). Unpublished measurements of magnetic susceptibility with Ehrenberg proved this very labile blue intermediate to be diamagnetic, and very recent unpublished data of Hemmerich revealed a composition RF1-O-O-FIR, where the peroxo-linkage between the two flavin halves is thought to occur through C (10a). This chemical reasoning has been summarized in Scheme 3 and it would indeed allow the differentiation of the dehydrogenase, oxidase and oxygenase subclasses of flavoproteins according not only to their radical forms but also to their respective flavin oxygen “complexes” HF100H. It must be borne in mind, however, that isomerization of the three possibly “relevant” dihydroperoxides, l-HF1-9a/lOa-OOH and 5-HF1-4a-O0H, may be an intrinsically activationless process governed solely by apoprotein conformation. Hence, if HFlOOH stabilization is reached in an enzyme process experimentally, for example, at low temperature (103) or by complexing the active site with a “nonsubstrate effector’’ (cf. above), the intermediate detected might not be the true active HF1-00H but a ‘‘storage” isomer. While model studies indicate that hydroxylation reactions mediated by dihydroflavins and tetrahydropteridines may be the same (157), there are rather big differences between the properties of flavoprotein hydroxylases and pteridine-linked hydroxylases. The NIH shift has been demonstrated with the latter enzymes (see Section IV) , while there is no evidence yet produced for this happening with the flavoproteins. The pteridine-linked enzymes appear to contain iron, and a t least with phenylalanine hydroxylase there is evidence that the iron functions in the catalysis (136). The rate of reaction of 0, with tetrahydropteridines, while sufficiently fast t o complicate experimental work, is embarrassingly slow from an enzyme point of view, with half-lives a t physiological pH values in the order of minutes (161,157,162).This latter objection may be removed if further experimental work substantiates the conclusions of Kaufman and colleagues that an intermediate is liberated relatively rapidly from the enzyme and then decays slowly to yield products (196). However, this intermediate would have to possess unusual chemical properties consisting of a stable complex made up from oxygenated tetrahydropteridine and phenylalanine. If this were indeed the case it is difficult to envisage what the role of the iron might be. An alternative possibility is that it is the iron in the ferrous state which 181. P.Hemmerich, B.Prijs, and H. Erlenmeyer, Helv. Chim. Acta 43, 372 (1960). 182. K.H.Dudley, A. Ehrenberg, P. Hemmerich, and F. Miiller, Helv. Chim. Acta 47, 1354 (1964).

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VINCENT MASSEY AND PETER HEMMERICH

serves to activate molecular oxygen for the hydroxylation reaction. I n this case the tetrahydropteridine may function simply to reduce the iron, in the same way as the dihydronicotinamide coenzymes serve t o reduce the flavin in flavoprotein hydroxylases. This possibility would eliminate the objection raised above concerning the slow rate of autoxidation of tetrahydropteridines but would not offer an explantion of the slowly decaying intermediate released from the enzyme. A third possibility suggested by Viscontini (183) is that the tetrahydropteridine complexes with the enzyme-bound Fe3+to form a Fez+-pteridine radical complex, which then reacts with 0, to form an oxygenated complex in which the 0, is liganded to the iron atom. The latter is then proposed to yield hydroxy radicals, the active hydroxylating species. While the concept of an iron-tetrahydropteridine complex being the active hydroxylating species is an interesting speculation, there does not appear to be any substantial evidence in favor oi hydroxy radical involvement since this would fail to provide any explanation for the NIH shift, which requires the formation of an arene oxide intermediate. In conclusion, it is clear that the detailed mechanisms of hydroxylation catalyzed by either the flavoproteins or the pteridine-linked enzymes are far from being understood despite the large amount of experimental effort so far expended. However, the most attractive explanation would seem to be that in both classes, whatever the nature of the primary oxygenated species may be, an oxene residue is transferred to the substrate with subsequent rearrangement to the hydroxylated product. ACKNOWLEDGMENT This review was written while Vincent Massey was on sabbatical leave a t the University of Konstanz. He wishes to acknowledge with gratitude a Senior U. S. Scientist Award of the Alexander von Humboldt Foundation. 183. M. Viscontini, in “Chemistry and Biology of Pteridines” p. 217. Int. Acad. Printing Co., Tokyo, 1970.

(K. Iwai et al., eds.),