©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
The Molybdenum Centers of Xanthine Oxidase and Xanthine Dehydrogenase
DETERMINATION OF THE SPECTRAL CHANGE ASSOCIATED WITH REDUCTION FROM THE Mo(VI) TO THE Mo(IV) STATE (*)

(Received for publication, June 2, 1995; and in revised form, June 16, 1995)

Matthew G. Ryan Kapila Ratnam Russ Hille

From the Department of Medical Biochemistry, Ohio State University, Columbus, Ohio 43210

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES

ABSTRACT

The UV-visible absorbance change associated with reduction of the molybdenum centers of xanthine oxidase and xanthine dehydrogenase has been determined using a double-difference technique. At pH 8.5, the Mo(VI) minus Mo(IV) difference spectrum seen with xanthine oxidase exhibits a positive feature at 420 nm, having an extinction change of 3,000 M cm as well as evidence for a negative feature below 340 nm. In xanthine oxidase this change is found to exhibit a marked pH dependence, implicating protonation/deprotonation events associated with changes in the molybdenum oxidation state. Application of the double-difference protocol to the respective circular dichroism spectra of xanthine oxidase and xanthine dehydrogenase reveals appreciable CD changes at 420 and 580 nm associated with the reduction of the molybdenum center. The present results demonstrate a direct spectroscopic handle on the molybdenum centers of both xanthine oxidase and xanthine dehydrogenase.


INTRODUCTION

Milk xanthine oxidase and chicken liver xanthine dehydrogenase belong to the molybdenum hydroxylase class of enzymes, with each possessing a MoOS catalytic unit in its active site. Both enzymes are homodimers of molecular weight 300,000 with two iron-sulfur centers (of the spinach ferredoxin variety) and one molecule of flavin adenine dinucleotide in addition to the molybdenum center in each subunit (1, 2, 3, 4) . The oxidative hydroxylation of a variety of aromatic heterocycles and simple aldehydes takes place at the molybdenum centers of these enzymes, with reducing equivalents thus introduced passed to the flavin site where they are removed by either molecular oxygen (for the oxidase) or NAD (for the dehydrogenase). In the case of xanthine oxidase, it has been shown that the hydroxylation reaction passes through an intermediate in which the nascent uric acid product is coordinated to the (reduced) molybdenum via the newly introduced hydroxyl group as Mo(IV)-OR. Oxidation of this species by 1 eq yields a Mo(V) species that gives rise to the well characterized ``very rapid'' molybdenum EPR signal(3, 5, 6) . Xanthine dehydrogenase is thought to operate via a similar reaction mechanism.

The kinetics of the reductive half-reaction of these enzymes have been studied extensively by both UV-visible absorption and EPR spectroscopy (7, 8, 9, 10) . Studies monitoring absorbance changes associated with enzyme reduction necessarily focus on the iron-sulfur and flavin centers, as these are responsible for the preponderance of the spectral changes seen upon reduction for both enzymes. Any spectral change attributable to the molybdenum site has remained ill defined because of the much larger absorption changes associated with these other chromophores. Strong evidence exists, however, for transient absorption changes attributable to the molybdenum center of xanthine oxidase in the course of the reaction of enzyme with lumazine (2,6-dihydroxypteridine; Refs. 11 and 12), 2-hydroxy-6-methylpurine(7) , and xanthine(13) . With lumazine, two long wavelength-absorbing species have been identified as intermediates in the reductive half-reaction, the first ascribed to EbulletS (i.e. Mo(VI)bulletlumazine) and the second to EbulletP (Mo(IV)bulletviolapterin). The latter intermediate, having an absorbance maximum at 650 nm with an extinction change of approximately 8000 M cm, is readily formed by anaerobic addition of violapterin to dithionite-reduced enzyme(11) , while the existence of the former has been inferred from the kinetic behavior of the enzyme(12) . The same long wavelength-absorbing species has been observed with xanthine dehydrogenase. (^1)The reaction of xanthine oxidase with 2-hydroxy-6-methylpurine has also been found to possess two successive reaction intermediates detectable by UV-visible spectrophotometry, exhibiting absorption differences (relative to oxidized enzyme) at 470 and 540 nm, respectively, with extinction changes of approximately 410 M cm. These spectral intermediates have been attributed to the molybdenum center of the enzyme in the Mo(IV) and Mo(V) valence states, respectively, the latter corresponding to the species exhibiting the ``very rapid'' Mo(V) EPR signal known to be formed in the course of the reaction(14) . A comparable species has also been seen in the reaction of xanthine oxidase with xanthine(13) , with an associated spectral change above 350 nm very similar to that observed upon addition of product uric acid to reduced enzyme(15) . It has recently been shown that analogous intermediates are also observed in the reaction of xanthine dehydrogenase with either xanthine or xanthopterin(16) .

Despite the above evidence for spectral intermediates attributable to the molybdenum center of xanthine oxidase in the course of its reaction with substrate, there is no unambiguous evidence to date for a spectral change associated with reduction of the molybdenum center. Using a double-difference spectroscopic technique, we report the determination of this spectral change in both xanthine oxidase and xanthine dehydrogenase and its associated circular dichroism change.


MATERIALS AND METHODS

Xanthine oxidase was purified from unpasteurized cow's milk (obtained from the dairy herd of Ohio State University) according to the procedure of Massey et al.(17) . Sephacryl S-300 gel filtration chromatography and CM-52 ion-exchange column chromatography steps at the end of the procedure ensured removal of contaminating lactoperoxidase(18) . The purified enzyme exhibited a ratio of absorbance at 276 to 450 nm of 5.4 and was typically 70% functional. The 30% inactive enzyme found in all conventional preparations of xanthine oxidase lacks a catalytically essential sulfur atom at the molybdenum center (19) and for the purposes of the present studies can be considered to be inert. Routine enzyme assays and the determination of the specific activity of xanthine oxidase were performed as described by Massey et al.(17) . Xanthine dehydrogenase was purified by a method involving homogenization of fresh chicken livers in liquid nitrogen, followed by centrifugation, ammonium sulfate, and butanol fractionation and sequential chromatography on hydroxylapatite, Sephacryl S-300, and a folate affinity column. This procedure avoids use of acetone extraction, thereby minimizing damage to the molybdenum center of this enzyme.

UV-visible absorption spectra were obtained with a Hewlett-Packard 8452 diode-array spectrophotometer interfaced to a Hewlett-Packard Chemstation computer. Circular dichroism spectra were obtained using an AVIV-40DS UV-visible near infrared spectrophotopolarimeter. Alloxanthine (1H-pyrazolo[3,4-d]pyrimidine-4,6-diol) was purchased from Sigma. All other reagents and buffers were of the highest quality commercially available and used without further purification.


RESULTS

The Spectral Change Associated with Reduction of the Molybdenum Centers of Xanthine Oxidase and Xanthine Dehydrogenase

It is known that the alloxanthine complex of xanthine oxidase is very resistant to air reoxidation, the reaction taking place with a t½ of approximately 5 h(20) . By comparison, the iron-sulfur and flavin centers of the complexed enzyme are rapidly reoxidized under these conditions, so that the difference between the spectrum of the initial oxidized enzyme and the reoxidized alloxanthine complex can be obtained, reflecting the spectral difference between the oxidized Mo(VI) center and the reduced Mo(IV)bulletalloxanthine complex. Similarly, it is straightforward to obtain the spectral change associated with the binding of alloxanthine to reduced xanthine oxidase under anaerobic conditions, although this has not been previously reported. The corresponding difference spectrum corresponds to Mo(IV) minus Mo(IV)bulletP. The double-difference spectrum obtained from the subtraction of these two difference spectra (Mo(VI) - Mo(IV)bulletP) minus (Mo(IV) - Mo(IV)bulletP) should give Mo(VI) minus Mo(IV) according to Fig. S1.


Figure S1: Scheme 1.



Fig. 1shows the several difference spectra obtained according to Fig. S1for both xanthine oxidase and xanthine dehydrogenase at pH 8.5. PanelA shows the difference spectra of oxidized enzyme and enzyme that has been reduced, treated with alloxanthine, and reoxidized for 10 min for xanthine oxidase (solidline) and xanthine dehydrogenase (dashedline). The data obtained using xanthine oxidase are in good agreement with the literature(20) . PanelB shows the difference spectrum for reduced xanthine oxidase in the absence and presence of alloxanthine (solidline) and the same for xanthine dehydrogenase (dashedline). In the latter spectrum, some reoxidation of the large amount of sodium dithionite required to achieve full formation of the alloxanthine complex results in a large negative artifact below 360 nm, but above this wavelength the difference spectrum is minimally perturbed. (^2)PanelC of Fig. 1gives the double-difference spectra obtained by subtraction of the appropriate spectra from PanelsA and B for xanthine oxidase (solidline) and xanthine dehydrogenase (dashedline). Again, the difference spectrum obtained with xanthine dehydrogenase is accurate only above 360 nm. It is readily evident in the double-difference spectrum obtained with each enzyme that a spectral change having a difference maximum at 420 nm is observed, and on the basis of the argument summarized above (Fig. S1) this spectral change must correspond to that observed on reduction of the molybdenum center of each enzyme. The extinction change at 420 nm is estimated to be 3,000 M cm in the case of xanthine oxidase; corrections made for uncomplexed alloxanthine resulted in a negligible shift in the molybdenum absorption profile above 380 nm and were considered negligible in the overall treatment. It is noteworthy that the extinction change associated with reduction of the molybdenum center of xanthine oxidase is of a comparable magnitude with that observed upon formation of the Mo(IV)bulletalloxanthine complex (20) and the Mo(IV)bulletviolapterin complex(11) , although the wavelength maxima are very different for the latter two spectral changes.


Figure 1: Determination of spectral change associated with reduction of the molybdenum center of xanthine oxidase and xanthine dehydrogenase. PanelA (difference spectrum 1), solid line, difference spectra obtained after the addition of excess alloxanthine to dithionite-reduced xanthine oxidase under anaerobic conditions; dashed line, the same with xanthine dehydrogenase. The experimental conditions for xanthine oxidase were 68 µM functional enzyme and 125 µM alloxanthine (after mixing) in 0.1 M pyrophosphate, 0.3 mM EDTA, pH 8.5, 25 °C. Spectra were recorded in a diode-array spectrophotometer shortly after the addition of product to dithionite-reduced enzyme. Absorbance changes are given relative to reduced enzyme. Panel B (difference spectrum 2), difference spectrum obtained after introduction of air to the dithionite-reduced enzyme-alloxanthine complex. Spectra are for xanthine oxidase (solid line) and xanthine dehydrogenase (dashed line). Absorbance changes are given relative to oxidized enzyme. Panel C, double-difference spectra obtained after subtraction of difference spectrum 2 from difference spectrum 1 for xanthine oxidase (solid line) and xanthine dehydrogenase (dashed line).



Xanthine oxidase as isolated from cow's milk is known to possess a variable amount (usually 20-30%) of nonfunctional enzyme lacking a catalytically essential Mo=S moiety. It has been shown, however, that alloxanthine binds only to the reduced, functional form of the enzyme (20) (also confirmed in the present studies, data not shown). As a result, the double-difference spectrum for xanthine oxidase shown in Fig. 1C reflects the spectral change associated with reduction of the functional form of xanthine oxidase only. Chicken liver xanthine dehydrogenase, on the other hand, does not possess a substantial amount of the desulfo-form as isolated. Differences in the results obtained with the two enzymes appear to be associated with differences in pKs associated with their molybdenum centers (see below).

Spectral Deconvolution of the Overall Spectral Changes for Reduction of Xanthine Oxidase

The contribution of the flavin center of xanthine oxidase to the overall spectral change observed on reduction of the enzyme has been known for some time from a comparison of the spectral changes seen with native enzyme and enzyme that has had the flavin removed by incubation at high salt concentrations(21, 22) . The present results permit the deconvolution of the spectral change for reduction of the deflavo-form of xanthine oxidase into its molybdenum and iron-sulfur components. Fig. 2shows spectral change for holoenzyme broken down into its several components, and it can be seen that the spectral change associated with reduction of the molybdenum center, while small, is not negligible. It is clear that detection of the spectral change associated with the molybdenum center has been made difficult in previous studies by the fact that its absorption maximum is near an inflection point in the difference spectra for the flavin and iron-sulfur centers of both the oxidase and dehydrogenase.


Figure 2: Deconvolution of absorption profiles attributable to prosthetic centers of xanthine oxidase. Solid line, difference spectrum corresponding to the subtraction of native xanthine oxidase (oxidized) from native xanthine oxidase (reduced), corresponding to the cumulative UV-visible spectral change due to reduction of the flavin, iron/sulfur, and molybdenum centers. Dashed line, difference spectrum corresponding to the subtraction of deflavoxanthine oxidase (oxidized) from deflavoxanthine oxidase (reduced), corresponding to the cumulative spectral changes attributable to the iron/sulfur and the molybdenum centers, respectively. Dotted line, difference spectrum corresponding to the spectral change associated with reduction of the molybdenum center of xanthine oxidase.



pH Dependence of the Spectral Change Associated with Reduction of the Molybdenum Center of Xanthine Oxidase

The alloxanthine-based protocol described above allows the spectral change associated with reduction of the molybdenum site to be extracted despite the strong absorption of the FAD and Fe/S chromophores. Because of the well established association of protonation-deprotonation events with changes in the oxidation state of even the simplest oxomolybdenum complexes (23, 24) the pH dependence of the double-difference spectrum for xanthine oxidase has been investigated. As illustrated in Fig. 3, the double-difference spectrum is in fact found to be influenced markedly by pH: a decrease in the overall magnitude of the molybdenum absorption along with a small red shift in the absorption maximum is observed on going from pH 10 to 6.0. This result suggests the presence of ionizable groups(s) in the immediate vicinity of the molybdenum site whose ionization is intimately associated with reduction of the molybdenum. These may be amino acid side chains and/or atoms directly coordinated to the molybdenum. Unfortunately, no convincing isosbestic point is observed for the three difference spectra shown, undoubtedly a reflection of the involved nature of the double-difference technique and the unavoidable uncertainty associated with subtraction of a large background absorption, particularly in the case of the difference spectrum associated with oxidized enzyme. As a result, no attempt was made to obtain difference spectra at other values of pH, and we conclude simply that an ionization or ionizations in the pH range 6-10 are associated with the observed spectral change.


Figure 3: Effect of pH on the spectral change associated with reduction of the molybdenum center of xanthine oxidase. Solid line, double-difference spectrum obtained under the same experimental conditions as described above, that is in 0.1 M pyrophosphate, 0.3 mM EDTA, pH 8.5; dashed line, double-difference spectrum obtained as above except in 0.1 M CAPS (3-(cyclohexylamino)propanesulfonic acid), 0.1 N KCl, 0.3 mM EDTA, pH 10; dotted line, the double-difference spectrum obtained as above except in 0.1 M MES (4-morpholineethanesulfonic acid), 0.1 N KCl, 0.3 mM EDTA, pH 6.0.



The Circular Dichroism Change Associated with Reduction of the Molybdenum Center

The alloxanthine-based protocol has also been used to determine the circular dichroism change associated with the reduction of the molybdenum center. As shown in Fig. 4, an appreciable CD activity associated with the reduction of the molybdenum center is observed for both the xanthine oxidase and xanthine dehydrogenase, with pronounced CD-active features observed around 420 and 580 nm. It is to be noted that the CD changes around 580 nm do not correspond to any prominent spectral features in the UV-visible absorbance double-difference spectrum. This discrepancy may be explained if the molybdenum center exhibited absorbance in both the oxidized and reduced states, which cancel one another in the double-difference profile yet exhibit different CD. UV-visible spectral changes for both xanthine oxidase and xanthine dehydrogenase were recorded concomitantly with the CD spectra used to generate the difference spectra shown in Fig. 4and yielded difference absorbance changes in good agreement with the data presented in Fig. 1(data not shown).


Figure 4: The circular dichroism change associated with the reduction of the molybdenum center of xanthine oxidase and xanthine dehydrogenase. Solid line, the double-difference CD spectra obtained for xanthine oxidase after subtraction of its oxidized difference spectrum from its reduced difference spectrum as described above for the UV-visible absorbance spectra; dashed line, the same with xanthine dehydrogenase. The experimental conditions were the same as above, and the CD spectra were recorded in an AVIV-40DS CD spectrometer.




DISCUSSION

The results presented here clearly demonstrate a significant but heretofore undetected spectral change associated with simple reduction of the molybdenum centers of xanthine oxidase and xanthine dehydrogenase. This constitutes the first determination of the absorbance change for the MoOS centers found in these enzymes (although as discussed in the introduction there is abundant evidence for other types of spectral changes associated with the formation of several types of reaction intermediates at the molybdenum center). The spectral change appears to reflect a shift in absorbance maximum from 420 nm in the oxidized center to below 300 nm in the reduced center of xanthine oxidase at pH 8.5 (Fig. 1C), although the shift is difficult to quantitate due to the deterioration in signal-to-noise in the 260-310-nm region, where background absorption by the protein is large. The pH dependence of the spectral change attributed to the molybdenum center of xanthine oxidase is not surprising given the known relationship of protonation/deprotonation events to a molybdenum oxidation state in even the simplest inorganic complexes. The present work indicates that at least one ionizable group is associated with the molybdenum center of xanthine oxidase. With xanthine dehydrogenase, it appears that the pK values associated with the molybdenum center are somewhat lower than in the case of the oxidase, since the dehydrogenase exhibits a spectral change for reduction of its molybdenum center at pH 8.5 that is comparable with that observed with the oxidase at pH 6 (compare the dashedspectrum of Fig. 1C and the dottedspectrum of Fig. 3, respectively).

Other oxomolybdenum enzymes possessing a Mo(VI)O(2) center in the oxidized state rather than Mo(VI)OS (e.g. sulfite oxidase and nitrate reductase) have very little absorption in the 200-800-nm region attributable to the molybdenum centers, although the molybdenum domain of sulfite oxidase exhibits an absorption band at 480 nm having an extinction coefficient of 1600 M cm. Only dimethyl sulfoxide reductase exhibits a spectral change that is clearly attributable to the molybdenum center(25) . This indicates a major difference in the electronic structure between the molybdenum centers of sulfite oxidase and nitrate reductase, on the one hand, and dimethyl sulfoxide reductase on the other. Clearly there are important differences between the electronic structures of the molybdenum centers of these enzymes that must ultimately be due to differences in the molybdenum coordination sphere. The present work demonstrates for the first time a spectral change associated with reduction of the molybdenum centers of two MoOS-containing enzymes, xanthine oxidase and xanthine dehydrogenase. This spectral change is pH-dependent with a maximum extinction change at 410 nm in the high pH limit and provides a new experimental probe of the molybdenum center that in the future should make it amenable to techniques such as resonance Raman spectroscopy.


FOOTNOTES

*
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

(^1)
K. Ratnam and R. Hille, unpublished data.

(^2)
A somewhat higher concentration of dithionite was used in the experiments performed with the dehydrogenase than was the case with the oxidase given the greater difficulty in maintaining the redox-active centers of the dehydrogenase fully reduced. Because partial reoxidation of the enzyme in the present experiments is so undesirable, use of additional dithionite in the experiment with the dehydrogenase was deemed necessary to ensure accurate determination of the spectral change above 360 nm, even though this compromised the spectral change below this wavelength.


REFERENCES

  1. Hille, R., and Massey, V. (1985) in Molybdenum Enzymes (Spiro, T. G., ed) pp. 443-518, John Wiley and Sons, New York
  2. Bray, R. C. (1988) Q. Rev. Biophys. 21,299-329 [Medline] [Order article via Infotrieve]
  3. Hille, R. (1994) Biochim. Biophys. Acta 1184,143-169 [Medline] [Order article via Infotrieve]
  4. Hille, R., and Nishino, T. (1995) FASEB J. , in press
  5. Gutteridge, S., and Bray, R. C. (1980) Biochem. J. 189,615-623 [Medline] [Order article via Infotrieve]
  6. Hille, R., and Sprecher, H. (1987) J. Biol. Chem. 262,10914-10917 [Abstract/Free Full Text]
  7. Bray, R. C., Palmer, G., and Beinert, H. (1964) J. Biol. Chem. 239,2667-2676 [Free Full Text]
  8. Edmondson, D., Ballou, D., van Heuvelen, A., Palmer, G., and Massey, V. (1973) J. Biol. Chem. 248,6135-6144 [Abstract/Free Full Text]
  9. Olson, J. S., Ballou, D., Palmer, G., and Massey, V. (1974) J. Biol. Chem. 29,4363-4382
  10. Schopfer, L. M., Massey, V., and Nishino, T. (1988) J. Biol. Chem. 263,13528-13538 [Abstract/Free Full Text]
  11. Davis, M. D., Olson, J. S., and Palmer, G. (1982) J. Biol. Chem. 257,14730-14737 [Abstract/Free Full Text]
  12. Davis, M. D., Olson, J. S., and Palmer, G. (1984) J. Biol. Chem. 259,3526-3533 [Abstract/Free Full Text]
  13. Kim, J. H., and Hille, R. (1993) J. Biol. Chem. 268,44-51 [Abstract/Free Full Text]
  14. McWhirter, R. B., and Hille, R. (1991) J. Biol. Chem. 266,23724-23731 [Abstract/Free Full Text]
  15. Hille, R., and Stewart, R. C. (1984) J. Biol. Chem. 259,1570-1576 [Abstract/Free Full Text]
  16. Hunt, J., and Massey, V. (1994) J. Biol. Chem. 269,18904-18914 [Abstract/Free Full Text]
  17. Massey, V., Brumby, P. E., Komai, H., and Palmer, G. (1969) J. Biol. Chem. 244,1682-1691 [Abstract/Free Full Text]
  18. Morrison, M., and Hultquist, D. E. (1963) J. Biol. Chem. 238,2847-2849 [Free Full Text]
  19. Massey, V., and Edmondson, D. (1970) J. Biol. Chem. 245,6595-6598 [Abstract/Free Full Text]
  20. Massey, V., Komai, H., Palmer, G., and Elion, G. B. (1970) J. Biol. Chem. 245,2837-2844 [Abstract/Free Full Text]
  21. Komai, H., Massey, V., and Palmer, G. (1969) J. Biol. Chem. 244,1692-1700 [Abstract/Free Full Text]
  22. Nishino, T., Nishino, T., Schopfer, L. M., and Massey, V. (1989) J. Biol. Chem. 264,6075-6085 [Abstract/Free Full Text]
  23. Steifel, E. I. (1977) Prog. Inorg. Chem. 21,1-221
  24. Cramer, S. P., and Steifel, E. I., (1985) in Chemistry and Biology of the Molybdenum Cofactor, in Molybdenum Enzymes (Spiro, T. G., ed) pp. 411-441, Wiley-Interscience, New York
  25. Bastian, N. R., Kay, C. J., Barber, M. J., and Rajagopalan, K. V. (1991) J. Biol. Chem. 266,45-51 [Abstract/Free Full Text]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.