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J. Biol. Chem., Vol. 275, Issue 25, 18830-18835, June 23, 2000

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Kinetics of Manganese Lipoxygenase with a Catalytic Mononuclear Redox Center^*

Chao Su, Margareta Sahlin^§, and Ernst H. Oliw^¶

From the  Division of Biochemical Pharmacology, Department of Pharmaceutical Biosciences, Uppsala Biomedical Center, Uppsala University, SE-751 24 Uppsala, Sweden and the ^§ Department of Molecular Biology, Stockholm University, SE-106 91 Stockholm, Sweden

Received for publication, February 21, 2000, and in revised form, March 28, 2000

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Manganese lipoxygenase was isolated from the take-all fungus, Gaeumannomyces graminis, and the oxygenation mechanism was investigated. A kinetic isotope effect, k_H/k_D = 21-24, was observed with [U-²H]linoleic acid as a substrate. The relative biosynthesis of (11S)-hydroperoxylinoleate (11S-HPODE) and (13R)-hydroperoxylinoleate (13R-HPODE) was pH-dependent and changed by [U-²H]linoleic acid. Stopped-flow kinetic traces of linoleic and -linolenic acids indicated catalytic lag times of ~45 ms, which were followed by bursts of enzyme activity for ~60 ms and then by steady state (k_cat ~26 and ~47 s¹, respectively). 11S-HPODE was isomerized by manganese lipoxygenase to 13R-HPODE and formed from linoleic acid at the same rates (k_cat 7-9 s¹). Catalysis was accompanied by collisional quenching of the long wavelength fluorescence (640-685 nm) by fatty acid substrates and 13R-HPODE. Electron paramagnetic resonance (EPR) of native manganese lipoxygenase showed weak 6-fold hyperfine splitting superimposed on a broad resonance indicating two populations of Mn^II bound to protein. The addition of linoleic acid decreased both components, and denaturation of the lipoxygenase liberated ~0.8 Mn²⁺ atoms/lipoxygenase molecule. These observations are consistent with a mononuclear Mn^II center in the native state, which is converted during catalysis to an EPR silent Mn^III state. We propose that manganese lipoxygenase has kinetic and redox properties similar to iron lipoxygenases.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Lipoxygenases oxygenate polyunsaturated fatty acids with one or several (1Z,4Z)-pentadiene units to a hydroperoxy-conjugated diene (1-4). Lipoxygenases are widely distributed in animals and plants, and the lipoxygenase products have a wide range of biological functions as diverse signal molecules, oxidants, and modifiers of membrane structures (1 and 2). Mammalian lipoxygenases insert molecular oxygen at positions C-5, C-8, C-12, or C-15 of arachidonic acid, whereas many plant lipoxygenase oxygenate linoleic acid at positions C-9 or C-13. All lipoxygenases belong to the same gene family. A characteristic feature is a metal center with non-heme iron (1-4). One lipoxygenase with manganese as a prosthetic metal has been discovered (5, 6). Whether manganese lipoxygenase belongs to the lipoxygenase gene family is presently unknown. Manganese lipoxygenase has so far only been identified in a fungal pathogen of wheat, Geaumannomyces graminis.

Lipoxygenases with prosthetic iron or manganese have fundamental properties in common but differ in important details. Both types of enzymes abstract with stereospecificity a bisallylic hydrogen from C-3 of the (1Z,4Z)-pentadiene unit and are postulated to form a delocalized alkyl radical over C-1 to C-5 (6-8). Dioxygen reacts with the radical in different ways. All iron lipoxygenases allow oxygen to react with the alkyl radical in an antarafacial way to form a 1-hydroperoxy-(2E,4Z)-pentadiene (1-4, 7), whereas manganese lipoxygenase allows oxygen to react in a suprafacial way at either C-1 or C-3 in a ~3:1 ratio (6). Manganese lipoxygenase also catalyzes the isomerization of the 3-hydroperoxy-(1Z,3Z)-pentadiene to the end product, the 1-hydroperoxy-(2E,4Z)-pentadiene (6). The differences between iron and manganese lipoxygenases are likely due to steric factors at the active site, which control the position of the alkyl radical and the access of oxygen. The prosthetic iron plays a key role during catalysis. Lipoxygenation starts with oxidation of Fe^II to Fe^III during a short time lag, which is followed by a burst of enzymatic activity and then by steady state catalysis (1-3). The active Fe^III-lipoxygenase (Fe^III-OH) abstracts the bisallylic hydrogen, an alkyl radical is formed, and Fe^III is reduced to Fe^II (Fe^II-OH₂). Dioxygen reacts with the radical and forms a peroxyl radical, which regains a hydrogen, the ferrous iron is re-oxidized to ferric (Fe^III-OH), and the fatty acid hydroperoxide is formed. Experimental evidence of this mechanism was first provided by EPR¹ analysis of the iron center during soybean lipoxygenase-1 catalysis and by detection of the peroxyl radical (10-12). It is also known that the hydroperoxy group can undergo nonenzymatic rearrangement and be exchanged with surrounding molecular oxygen (13). As an alternative, a lipoxygenation mechanism with an organoiron intermediate has been proposed (14), but many of the described experimental observations are consistent with the radical mechanism (1-3, 10-12).

No information on the prosthetic manganese of manganese lipoxygenase is available except a crude estimate of its manganese content and indirectly by inhibition of the enzyme by a 5-lipoxygenase inhibitor with chelating and reducing properties and by GSH peroxidase (5, 6). Experiments with oxygen-18-labeled 11S-HPODE and 13R-HPODE and with stereospecifically deuterated linoleic acids have provided a framework for the catalytic mechanism of manganese lipoxygenase described above. However, investigation of the catalytic role of the prosthetic manganese might contribute to elucidation of the reaction mechanism of this enzyme as well as iron lipoxygenases and other dioxygenases. Our main objective was therefore to use a variety of spectroscopic methods to study the kinetics and the metal center of manganese lipoxygenase and to compare the data with other lipoxygenases.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Materials-- Fatty acids were obtained as described (5). A mixture of perdeuterad unsaturated fatty acids was purchased from Larodon Fine Chemicals (Malmö, Sweden). Equipment for HPLC, SDS-polyacrylamide gel electrophoresis, and media and columns for chromatography were as described previously in Ref. 5. GSH, soybean lipoxygenase-1 (lipoxidase type IV), methyl--D-glucopyranoside, and lyophilized glutathione peroxidase (purified as described (15)) were from Sigma. 11S-HPODE and 13R-HPODE were prepared by biosynthesis and characterized by LC-MS (9). 11S-HPODE was enzymatically converted to 13R-HPODE with purified manganese lipoxygenase for quantification by UV at 235 nm.

Purification of Manganese Lipoxygenase and Metal Analysis-- Inductively coupled plasma atomic emission spectroscopy was performed as described (5). Manganese lipoxygenase was purified as described (5) with the following modifications. Fraction I, which eluted with buffer A (10 mM potassium phosphate buffer (pH 6. 8), 2 mM NaN₃, 0.04% Tween 20) from the phenyl-Sepharose column, was applied to a chelating Sepharose FF column (1.6 × 5.5 cm). The latter was charged with Cu²⁺ and equilibrated with 0.2 M NaCl in buffer A. The run-through fraction was dialyzed against water (dialysis membrane, Union Carbide, Chicago, IL). This material was then pooled with fraction II from the phenyl-Sepharose column and loaded on a CM-Sepharose column. The enzyme eluted with 0.2 M NaCl in buffer A. Active fractions were concentrated (Millipore Ultrafree-15/Biomax-30 centrifugal filter, Bedford, MA) and purified on a Superdex 200 HR column in buffer B (0.05 M potassium phosphate buffer (pH 7. 3), 0.15 M NaCl, 0.05 M methyl--D-glucopyranoside, 3 mM NaN₃, 1 mM EDTA, 0.5 mM CHAPS). The active fraction had a specific activity 8. 2 µM mg¹ min¹ with linoleic acid as a substrate (25 °C) and showed a single band on SDS-polyacrylamide gel electrophoresis. The material was concentrated to 50 mg of protein/ml (Biomax-30 filtration). All glassware was washed overnight with 15% HNO₃ and rinsed with Milli-Q water. As an extra precaution, the buffer of the manganese lipoxygenase sample was replaced five times by ultrafiltration with 1 mM EDTA in buffer C (10 mM triethanolamine-HCl (pH 7. 3), 2 mM NaN₃, 0.04% Tween 20). The run-through of the final Superdex 200 HR column and the flow-through from filtrations were used as controls for EPR and atomic emission spectroscopy.

Manganese Lipoxygenase Assay-- Lipoxygenase activity was assayed by UV analysis at 235 nm at 25 °C (5) (and corrected for biosynthesis of 11-hydroperoxy fatty acids without UV absorption at 235 nm as described below). The linear parts of the curves were used to calculate reaction rates using a molar extinction coefficient of 23 mM cm¹ for conjugated diene formation. The biosynthesis of 11S-HPODE and 13R-HPODE was analyzed after mixing manganese lipoxygenase (20 nM) with excess linoleic or [U-²H]linoleic acids (0.25-0.5 mM) in 0.5 ml of buffer (pH 5-11; 0.04% Tween 20 was added to buffers of pH 5, 6, and 7). The steady increase in absorbance (235 nm) was followed until 5-10% of the substrate was metabolized. Vigorous mixing with acidified ethyl acetate stopped the reactions, and the extracted products were analyzed by LC-MS. Enzyme activity was normalized for the protein content determined by total amino acid analysis.

Amino Acid Analyses-- Manganese lipoxygenase was subject to total amino acid analysis after acidic hydrolysis (6 M HCl, 1 mg/ml phenol, 110 °C under vacuum for 24 and 72 h), whereas tryptophane was determined by hydrolysis in alkali (courtesy Dr. D. Eaker of Uppsala Biomedical Center). C-terminal amino acid analysis was performed at the Karolinska Institute, Stockholm (courtesy of Drs. H. Jörnvall and E. Cederlund).

Electronic and Fluorescence Spectroscopy-- Light absorption was recorded as described (5). Fluorescence was recorded with a spectrofluorophotometer (Hitachi F-4000, Tokyo, Japan) with a temperature-controlled cell holder and a magnetic stirrer. Anaerobic incubations were performed with an anaerobic cell repeatedly flushed with argon. Rapid kinetic stopped-flow assays were carried out at 25 °C as described (15), and formation of conjugated dienes was recorded at 250 nm (0.2-cm light path; extinction coefficient ~7500 M¹ cm¹ in buffer C). Changes in fluorescence during stopped-flow were recorded at 640 nm (excitation 570 nm).

EPR Analysis-- EPR spectra of manganese lipoxygenase were recorded on an EPR spectrometer (Elexsys EPR, Bruker) at 77-100 K or with an Oxford-Instrument helium cryostat at 10 K. EPR spectra were recorded at 9.4 GHz and analyzed with the Xepr software (version 2.0, Bruker).

LC-MS and HPLC-- Equipment for LC-MS analysis was as described (9). The column contained octadecasilane silica (5-µm, 150 × 2 mm; Chromasil 5 C₁₈ 100 A, Phenomenex, Torrance, CA) and was eluted at 0.2 ml/min with methanol/water/acetic acid, 80/20/0.01 (v/v). The effluent first passed a UV detector (235 nm) and was then subject to negative ion electrospray ionization in an ion trap mass spectrometer (LCQ, ThermoQuest, San Jose, CA). The source voltage was 2.4 kV, the capillary temperature was 170 °C, and the collision energy in arbitrary units was 30%. PGF₁ was used for tuning. Off-line nanoelectrospray was performed with a 2-3-µl sample (~30 µM) in disposable gold-coated capillary probes (Protana A/S, Odense, Denmark) with a spray voltage of 0.8 kV. The perdeuterated fatty acid mixture was resolved by HPLC on a µBondapak C18 column (Waters, 8. 4 × 250 mm) with methanol/water/acetic acid, 92.5/7.5/0.01, at 2 ml/min with UV monitoring at 205 nm. [U-²H]Linoleic acid was characterized by nanoelectrospray-MS and by gas chromatography-MS (5). The apparent isotope distribution of [U-²H]linoleic acid was d₂₉, 46%; d₂₈, 30%; d₂₇, 17%; and d₂₆, 7% according to gas chromatography-MS analysis.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Analysis of Amino Acid Residues and Prosthetic Metal-- The amino acid compositions of manganese lipoxygenase, soybean lipoxygenase-1, and human 5-lipoxygenase-1 are shown in Table I. The C-terminal residue of manganese lipoxygenase was valine, whereas the C-terminal residue of most other lipoxygenases is isoleucine (1-3). The apparent molar extinction coefficient at 280 nm was 9.27 × 10⁴ M¹. The manganese content was determined by atomic emission spectroscopy to be 0.94 mol of manganese/mol of enzyme protein. The iron content was below the detection limit. Concentrated solutions of manganese lipoxygenase were yellow.

Glutathione Peroxidase-- Soybean lipoxygenase-1 (1 µg/ml with 0.24 mM linoleic acid in buffer C; 25 °C) was completely and constantly inhibited by GSH peroxidase (0.1 unit/ml) and 1 mM GSH. In contrast, the reaction rate of manganese lipoxygenase (1.3 µg/ml) was only reduced by 50% with a 10 times larger concentration of GSH peroxidase, and the enzyme regained full enzyme activity after a few minutes. The manganese lipoxygenase could thus be catalytically active even in the presence of hydroperoxide-reducing agents.

Fluorescence of Manganese Lipoxygenase-- The emission spectrum of manganese lipoxygenase during excitation at 280 nm is shown in Fig. 1A. Soybean lipoxygenase-1 gives a similar emission spectrum. The fluorescence of manganese lipoxygenase centered at 345 nm decreased in the presence of fatty acids, which were not substrates of manganese lipoxygenase. The long wavelength fluorescence centered at 640 nm was quenched by linoleic acid (Fig. 1B). 13R-HPODE had the same effect. The quenching of 13R-HPODE was linear up to ~0.25 µM (Fig. 1C). The quench increased with temperature, and a Stern-Volmer plot (16) of the effect of temperature suggested collisional quenching (Fig. 1C). The quench was reversed by removal of linoleic acid and its metabolites from the reaction mixture. Oleic acid did not quench the long wavelength fluorescence, but linoleic and linolenic acids did as shown by the inset in Fig. 1C. Stopped-flow with 10-s traces was used to calculate the first-order decay rate constants of the fluorescence. The rate constants were 2.3 s¹ and 4.8 s¹ for linoleic and -linolenic acids, respectively, which were proportional to their turnover numbers.

View larger version (23K): [in this window] [in a new window]   Fig. 1.   Fluorescence spectra of manganese lipoxygenase and quenching by fatty acids and 13R-HPODE. A, emission spectrum of manganese lipoxygenase (exc = 280 nm; 0.17 µM enzyme in buffer B at 25 °C). The peak centered at 335 nm is typical protein fluorescence, and the narrow intense band centered at 565 nm corresponds to the second order Rayleigh peak also present in the buffer blank. The broad band centered at 640 nm was explored in B and C. B, excitation spectra (em = 685 nm) of 0.17 µM manganese lipoxygenase, which were recorded before and with 2-min intervals after mixing with 0.4 mM linoleic acid. C, a Stern-Volmer plot. The effect of 0.1-0.6 µM 13R-HPODE on the fluorescence of 0.3 µM manganese lipoxygenase at 8, 25, and 37 °C (top, middle, and bottom traces, respectively) was recorded. The inset shows time courses of the fluorescence changes when 0.17 µM manganese lipoxygenase was mixed with 0.4 mM fatty acids: trace a, oleic acid; trace b, -linolenic acid; trace c, linoleic acid; trace d, -linolenic acid. The horizontal bar marks 1 min. The fluorescence was recorded at 25 °C in buffer B under aerobic conditions. Each point shows the mean and standard deviation of triplicate determinations. Fo is the initial fluorescence and F is the fluorescence with 13R-HPODE. Excitation 575 nm, emission 685 nm.

No quenching was noticed under anaerobic conditions. Preincubation of manganese lipoxygenase with linoleic acid for 30 min under anaerobic condition led to enzyme inactivation, and there was no quenching when aerobic conditions were restored. The long wavelength fluorescence of heat-inactivated enzyme was not quenched by linoleic acid. We conclude that the quench of the long wavelength fluorescence can be related to catalysis of manganese lipoxygenase. Similar findings have also been reported for soybean lipoxygenase-1 (17).

Formation of HPODE-- Under steady state, linoleic acid was metabolized to 29% 11S-HPODE and 71% 13R-HPODE at pH 7. 3. The relative amounts of 11S-HPODE and 13R-HPODE at different temperatures and different pH values were determined. Temperature changes from 15 to 45 °C were without apparent effect but pH changes had a marked influence. Biosynthesis of 11S-HPODE increased from 3% at pH 5, to 29% at pH 7 and reached 40% at pH 11 (Fig. 2). As previously reported, manganese lipoxygenase has a broad pH optimum centered at pH 7 with over 60% enzyme activity at pH 5 and at pH 11 (5). 11S-HPODE is chemically unstable at acidic pH (6), but this could not explain the low formation at pH 5, as 11S-HPODE was found to be chemically stable under these conditions. However, linoleic acid has a pK_a between pH 7 and 8 (1), and the charge of the carboxyl group may affect substrate binding and the biosynthesis of products.

View larger version (27K): [in this window] [in a new window]   Fig. 2.   Effect of pH on biosynthesis of 11S- and 13R-HPODE by manganese lipoxygenase. Linoleic acid was incubated with manganese lipoxygenase at pH 5-11 and 25 °C, and the reaction was stopped at steady state. The following buffers were used: 0.1 M sodium citrate (pH 5), 0.1 M sodium phosphate (pH 6-8), and 0.1 M glycine-NaOH (pH 9-11). The products were quantified by LC-MS by monitoring the intensity of the carboxylate anions (m/z 311). Mean ± S.D. of repeated determinations of duplicate incubations.

Oxidation of [11S-²H]linoleic acid by manganese lipoxygenase occurred with a large kinetic isotope effect, k_H/k_D = 15-22 (6). UV and LC-MS analysis showed that [U-²H]linoleic acid was oxidized by manganese lipoxygenase and by soybean lipoxygenase-1 with a similar kinetic isotope effect. The kinetic isotope effect of manganese lipoxygenase was determined from duplicate determinations at 25 °C to yield k_H/k_D = 21-24 (with correction for 11S-HPODE formation), whereas soybean lipoxygenase-1 yielded k_H/k_D = 20-23. The kinetic isotope effect of manganese lipoxygenase was not temperature-dependent (5-45 °C), which also has been reported for the kinetic isotope effect of soybean lipoxygenase-1 (18). [U-²H]Linoleic acid was metabolized by manganese lipoxygenase to 15 and 85% of [U-²H]11S-HPODE and [U-²H]13R-HPODE, respectively, during steady state (pH 7.3), possibly because of steric effects of the perdeuterated substrate.

Kinetic Analysis-- Stopped-flow with kinetic traces of 0.5 and 20 s was used to study the oxygenation of -linolenic acid to 13R-HPOTrE by manganese lipoxygenase. The 20-s trace showed that the reaction rates increased for a few seconds, reached steady state, and then leveled off due to the exhaust of substrate (Fig. 3A). The 0.5-s trace demonstrated that catalysis started after a lag phase of ~40 ms, which was followed by a burst of enzyme activity for ~55 ms and then by steady state catalysis (Fig. 3A, inset). The k_cat for oxidation of -linolenic acid to 13R-HPOTrE at the burst of enzyme activity was 97 s¹ at 25 °C, which increased to 209 s¹ at 35 °C. Steady state values were 34 and 126 s¹, respectively. Because manganese lipoxygenase forms ~27% 11S-HPOTrE and ~73% 13R-HPOTrE at steady state (9), the apparent k_cat for oxygenation of -linolenic acid to these two metabolites at 25 °C was calculated to be 47 s¹. As previously reported, -linolenic acid had the highest turnover of the unsaturated C18 fatty acids and the lowest K_m (2.2 µM) (5).

View larger version (25K): [in this window] [in a new window]   Fig. 3.   Stopped-flow analysis of manganese lipoxygenase with -linolenic acid and with 11-HPODE. A, a 20-s kinetic trace, which shows the increase in absorption at 250 nm due to formation of 13R-HPOTrE from 35 µM -linolenic acid. The inset shows a 0.5-s kinetic trace (mean of three shots). B, a 20-s kinetic trace, which shows the increase in absorption at 250 nm due to formation 13R-HPODE from 13 µM 11S-HPODE. The inset shows a 0.5-s kinetic trace (mean of three shots). 1 µM manganese lipoxygenase was used in all experiments.

Stopped-flow with linoleic acid yielded similar traces, but linoleic acid was metabolized at only half the rate of -linolenic acid. Quantitative stopped-flow data with linoleic acid and with 11S-HPODE are summarized in Table II. The lag phase of linoleic acid was not reduced by preincubation of manganese lipoxygenase with 13R-HPODE (Table II). 11S-HPODE was also converted to 13R-HPODE after a lag phase, which was followed by a relatively long lasting burst of enzyme activity and then by steady state biosynthesis at a perfectly linear rate for 6-7 s (Fig. 3B). 11S-HPODE was metabolized to 13R-HPODE at less than half the rate of linoleic acid oxidation (Table II).

Using conventional spectroscopy, the K_m and V_max for isomerization of 11S-HPODE were found to be 8.1 µM and 7.5 µmol min¹ mg¹, respectively, at pH 7.3 and 25 °C. The V_max corresponded to k_cat = 9.1 s¹.

EPR Analysis-- EPR spectra of manganese lipoxygenase (0.6 mM, 100 K) revealed 6-fold hyperfine splitting on a broad singlet (Fig. 4, trace a). Protein-bound Mn^II characteristically shows weak EPR signals (19). The signals were not due to unbound Mn^II for two reasons. First, atomic emission spectroscopy of the buffer, which was obtained by filtration of the enzyme, contained no detectable manganese. Second, EPR analysis was performed after mixing this enzyme preparation with 4 mM linoleic acid. The reaction was quenched (105 °C) after incubation for ~1 s and for 10 min at room temperature. The EPR spectra after 1 s now showed that the 6-fold hyperfine splitting had almost disappeared together with a prominent decrease of the broad singlet and that a weak radical signal appeared at g = 2.005 (Fig. 4, trace b). After 10 min the Mn^II characteristics had completely disappeared, and the radical at g = 2.005 had decreased to 1/3 of the initial value (Fig. 4, trace c). EPR analysis with 11S-HPODE as a substrate yielded essentially similar results, i.e. the Mn^II-related signals decreased and the radical signal was apparent. The manganese signal of the native enzyme was therefore likely due to two different populations of Mn^II, one with octahedral coordination and the other with tetrahedral coordination (20), which were oxidized during catalysis to an EPR silent Mn^III state. The radical signal showed peak to trough width of 1.0 millitesla centered at g = 2.005. The saturation behavior indicated that it was not interacting with any magnetic relaxing species. The quantity of the radical before and 1 s after the addition of substrates yielded similar results, about 0.3 µM or 0.05% of the enzyme concentration, but decreased to about 0.1 µM 10 min after the addition of linoleic acid. The radical was not present in the buffer (Fig. 4, trace d). It seems possible that the disappearance of the six manganese signals made this signal appear (cf. traces a and b of Fig. 4). The nature of the radical was not further investigated.

View larger version (22K): [in this window] [in a new window]   Fig. 4.   EPR spectra of manganese lipoxygenase. Trace a shows the EPR spectrum of native manganese lipoxygenase. Trace b shows an EPR spectrum of manganese lipoxygenase incubated with linoleic acid for 1 s. Trace c shows EPR spectrum of manganese lipoxygenase incubated for 10 min with linoleic acid. Manganese lipoxygenase (0.6 mM) and linoleic acid (4 mM; or solvent only) were incubated for 1 s and 10 min and then immediately quenched by rapid freezing (105 °C). Trace d shows EPR spectrum of a buffer B blank from the last purification step of manganese lipoxygenase. Trace e shows EPR spectrum of manganese lipoxygenase (0.1 mM) after denaturation of the protein with 15% HNO3. Traces a-d represents averages of 16 scans and trace e is the average of 4 scans. All spectra were recorded at 9.4 GHz, 100 K, microwave power 20 mW, and modulation amplitude 0.5 millitesla.

The transition from an Mn^II to an Mn^III center during catalysis was obvious from the color of the EPR samples, which were yellow in the resting state and colorless in the presence of linoleic acid. This indicates again that the majority of Mn²⁺ ions are bound in the tetrahedral coordination (20).

Manganese lipoxygenase was also analyzed by EPR after denaturation of protein with 15% HNO₃, which released the bound metal and increased the manganese signals. The EPR spectrum exhibited a nearly isotropic g = 2.0 signal having 6-fold hyperfine splitting (A = 9. 5 millitesla) typical of octahedrally coordinated Mn²⁺ in solution (Fig. 4, trace e) (19). This spectrum was identical with an aqueous standard of MnCl₂. The amount of Mn²⁺ in the denaturated sample was estimated from spin quantitation to be 0.8 ± 0.08 atoms/lipoxygenase molecule. This was slightly lower than the manganese content according to atomic emission spectroscopy.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The oxygenation of linoleic acid by manganese lipoxygenase and iron lipoxygenases differs with respect to the stereochemistry of hydrogen abstraction and oxygen insertion at C-11 and/or C-13 of the delocalized alkyl radical, but the metal centers of both manganese and iron enzymes were nevertheless found to be mechanistically similar. Soybean lipoxygenase-1 and human 5-lipoxygenase contain a mononuclear Fe^II center, which is oxidized to Fe^III in the active enzyme (1-3, 10-11, 21). Atomic emission spectroscopy and EPR now demonstrate that manganese lipoxygenase also contains a mononuclear metal center. EPR of resting manganese lipoxygenase was consistent with two different populations of Mn^II bound to the apoprotein in tetrahedral and octahedral coordination (19, 20). The EPR signals of Mn^II decreased during catalysis suggesting that the metal was oxidized to a nonparamagnetic state, presumably Mn^III. The Mn^II oxidation state is a common form of manganese in biological systems and Mn^II  Mn^III redox cycling is observed in superoxide dismutase and catalase (22).

To obtain readily detectable signals with EPR we used 600-fold higher concentration of manganese lipoxygenase than during stopped-flow, which might influence the comparison of EPR spectra with stopped-flow kinetics. Nevertheless, the oxidation of Fe^II and Mn^II in lipoxygenases might be related to the lag phase, the inhibitory effect of GSH peroxidase (although it had a relatively weak and transient inhibitory effect on manganese lipoxygenase), and to inhibition of manganese lipoxygenase with a lipoxygenase inhibitor with reducing properties, BW4AC (5). A hypothetical reaction mechanism of manganese lipoxygenase is outlined in Fig. 5.

View larger version (13K): [in this window] [in a new window]   Fig. 5.   A hypothetical reaction mechanism of manganese lipoxygenase. Resting enzyme is in the MnII oxidation state and oxidized to the active form, MnIII, which is proposed to undergo redox changes as indicated. The alkyl and peroxyl radicals within brackets have not been identified but are likely formed according to isotope experiments (6). Glutathione peroxidase (GSH-PX) inhibits the reaction. The 3-hydroperoxypentadiene can be isomerized to the cis-trans conjugated hydroperoxypentadiene end product by the enzyme (as indicated by the arrows).

The turnover of -linolenic and linoleic acids by manganese lipoxygenase is ~47 and ~26 s¹, respectively, whereas the turnover of linoleic acid by soybean lipoxygenase-1 is ~260 s¹ (23). A peroxyl radical can be readily detected by EPR during soybean lipoxygenase catalysis, and an alkyl radical has been tentatively identified (12). According to isotope experiments (6), these radicals are also likely formed both during oxidation of linoleic acid and isomerization of 11S-HPODE to 13R-HPODE by manganese lipoxygenase. Substrate-induced peroxyl or alkyl radicals could not be detected by EPR under our experimental conditions. It is conceivable that these transient radicals were formed in too low a concentration due to the relatively low turnover of manganese lipoxygenase. The origin of the weak radical signal we observed at g = 2.005 is unclear.

The initial step of manganese lipoxygenase and iron lipoxygenase catalysis is removal of the pro-S hydrogen at C-11, which is accompanied by a large isotope effect (k_H/k_D > 15 for manganese lipoxygenase (6) and k_H/k_D = 20-80 for soybean lipoxygenase-1 (8, 18, 24)). The theoretically expected value of k_H/k_D is 7-10 (18). Perdeuterated linoleic acid yielded a similar kinetic isotope effect as [11S-²H]linoleic acid, which implies a small secondary isotope effect. The reason for the large primary kinetic isotope effect is unknown but might be related to proton leakage through a potential barrier (18). The de Broglie wavelengths are 0.5 Å for ¹H and 0.3 Å for ²H at 10 kJ mol¹, and the probability of proton tunneling is a function of these wavelengths and the barrier width (25, 26). When hydrogen abstraction has occurred, 11S-HPODE and 13R-HPODE are formed in parallel in a pH-dependent ratio. It is interesting to compare the rate of biosynthesis of 11S-HPODE and 13R-HPODE from linoleic acid and the rate for isomerization of 11S-HPODE to 13R-HPODE (Table II). The data suggest that biosynthesis of 11S-HPODE from linoleic acid and the isomerization of 11S-HPODE to 13R-HPODE occur at a lower rate (k_cat = 7 and 9 s¹, respectively) than the biosynthesis of 13R-HPODE (k_cat = 19 s¹) from linoleic acid.

Catalysis by manganese lipoxygenase and by soybean lipoxygenase-1 is accompanied by quenching of the long wavelength fluorescence (17). This might be because of interaction with aromatic amino acid residues (e.g. tryptophane or tyrosine) near the active site of the enzymes (17). Manganese in metalloenzymes is often ligated to histidine, aspartate, glutamate, and tyrosine residues (22). It is known that superoxide dismutase and catechol-2,3-dioxygenases use identical coordinating residues for their iron- and manganese-dependent enzymes (22, 27, 28). For catechol-2,3-dioxygenases, the metal is ligated to two histidines and a glutamic acid residue. We have no information on the metal ligands of manganese lipoxygenase, but Fe of lipoxygenases is ligated to three conserved histidines, the carboxyl group of the C-terminal isoleucine, a variable residue, and water (1-4). The C-terminal amino acid of manganese lipoxygenase was valine. It is possible that manganese might use the C-terminal valine acid as a metal ligand in analogy with isoleucine of lipoxygenases. A site-directed mutagenesis study, where the C-terminal isoleucine of a murine 12-lipoxygenase was substituted for many different amino acids, showed that only substitution with valine retained significant lipoxygenase activity (29). It will be of phylogenetic and mechanistic interest to determine whether manganese lipoxygenase belongs to the lipoxygenase gene family.

Are there any advantages with a manganese lipoxygenase? We can only speculate, as the biological function of manganese lipoxygenase is unknown. In oxidation state III of manganese and iron lipoxygenases, the two metals are expected to be similar in binding and other physicochemical properties (30). In oxidation state II of resting enzymes, however, the two metals may differ significantly. Mn^II is chemically stable, whereas Fe^II can be easily oxidized in the aerobic world. The different activation properties observed for manganese lipoxygenase and soybean lipoxygenase-1 may reflect the different redox potentials for transition from oxidation state II to oxidation state III. Manganese lipoxygenase is secreted by G. graminis, which is a fungal pathogen of wheat roots. The mycelia penetrate and devastate the roots and manganese lipoxygenase products might cause oxidative damage to the plant cells. To this respect, manganese lipoxygenase is heavily glycosylated, quite stable and has a broad pH optimum (5). In contrast to other lipoxygenases, manganese lipoxygenase oxidizes linoleic and -linolenic acids to two products, 11S-hydroperoxy and 13R-hydroperoxy fatty acids, but whether these products have any unique biological properties or can be further transformed to biological mediators are unknown. Disruption of the gene of manganese lipoxygenase might be needed to reveal the biological function of manganese lipoxygenase.

In summary, we report as the main finding that manganese lipoxygenase contains a mononuclear metal center, which likely undergoes Mn^II  Mn^III redox changes in analogy with Fe^II  Fe^III redox changes of other lipoxygenases. In agreement with this mechanism, manganese and iron lipoxygenases were found to have many kinetic properties in common.

	FOOTNOTES

^* This work was supported by the Swedish Medical Research Council (6523), the Swedish Society for Medical Research, and the Wallenberg Foundation.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

^¶ To whom correspondence should be addressed: Dept. of Pharmaceutical Biosciences, Uppsala Biomedical Cntr., P. O. Box 591, SE-751 24 Uppsala, Sweden. Tel.: 46 18 471 44 55; Fax: 46 18 55 29 36; E-mail: Ernst.Oliw@farmbio.uu.se.

Published, JBC Papers in Press, April 5, 2000, DOI 10.1074/jbc.M001408200

	ABBREVIATIONS

The abbreviations used are: EPR, electron paramagnetic resonance; 13R-HPODE, (13R)-hydroperoxy-(9Z,11E)-octadecadienoic acid; 11S-HPODE, (11S)-hydroperoxy-(9Z,12Z)-octadecadienoic acid; HPLC, high performance liquid chromatography; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid; 13R-HPOTrE, 13R-hydroperoxy-(9Z,11E,15Z)-octadecatrienoic acid; LC-MS, liquid chromatography-mass spectrometry.

	REFERENCES

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