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Bromoperoxidase from the marine snail, Murex trunculus
Comp. Biochem. PhysioL Vol. 88B, No. 3, pp. 917-922, 1987 Printed in Great Britain
0305-0491/87 $3.00+ 0.00 © 1987 Pergamon Journals Ltd
BROMOPEROXIDASE FROM THE MARINE SNAIL, M U R E X TRUNCULUS RANDA JANNUN* and ELMON L ~ COE? Biochemistry Department, American University of Beirut, Beirut, Lebanon (Received 5 January 1987)
Abstraet--l. Extracts of a 25,000 g sediment of Murex trunculus hypobranchial gland homogenate contain an enzyme which brominates monochlorodimedon in the presence of bromide and HzO2. 2. Dependence of activity on H z02 concentration exhibits a sharp optimum which increases from 20/~M at pH 7.4 to 400/~M at pH 4.0. 3. Dependence on Br- concentration follows a simple saturation curve with Km values of 7 mM at pH 7.4 and 30 mM at pH 5.0. 4. The pH optima depend on H20: concentration, ranging from pH 6.6 at 10 #M to pH 4.8 at 200/~M. 5. The enzyme is totally inactive with CI- and F-, although both these ions inhibit bromination.
INTRODUCTION
Carnivorous marine snails of the genera M u r e x and Purpura are known to release compounds which, when exposed to air and sunlight, form a fast purple dye. Friedlandcr (1909) first identified the major component in the purple dye as 6,6' dibromoindigo and subsequent work has shown that the snail releases a collection of brominated and unbrominated indoxyl sulfate derivatives which, after hydrolysis by an aryl sulfatase, spontaneously dimerize in the presence of oxygen and light to form indigo derivatives (Baker, 1974). Occurrence of the brominated intermediates implied the presence of a brominating enzyme and prompted an investigation of Murex gland extracts for bromoperoxidase activity. Such activity was demonstrated, as has been reported in preliminary communications (Jannun et al., 1981). Exclusive of the well-known iodoperoxidase activities of several classical peroxidases, the first enzyme especially identified as a haloperoxidase was the chloroperoxidase from Caldariomyces fumago. This soluble enzyme, which has been highly purified and crystallized, requires H2Oz, acts on bromide, iodide and chloride (Hager et al, 1966) and can act as a catalase and a "classical peroxidase" (Thomas et al., 1970). More recently, another enzyme which has been classified with chloroperoxidase as a C I - - H 2 0 :oxidoreductase, the so-called "myeloperoxidase", has been highly purified from human neutrophils (Matheson et al., 1981). Mydoperoxidase exhibits catalytic properties similar to those of fungal chloroperoxidase (Bakkenist et al., 1980; Clark and Klebanoff, 1979). Bromoperoxidase activity was first reported in marine algae (Theiler et al., 1978). Subsequently, it has been identified and characterized in many species of marine algae (Baden and Corbett, 1980; Hewson and Hager, 1980; A h e m et al., 1980; De Boer et aL, *Present Address: Institute for Molecular Virology, St Louis University, School of Medicine, St Louis, MO, USA. 1"Present Address: Arabian Gulf University, Medical College Manama, Bahrain.
1986; Itoh et al., 1986) and in several species of bacteria (Van Pe'e and Lingens, 1985a, b; Weisner et al., 1985). Bromoperoxidase activity has also been detected in a marine annelid and in a hemichordate (Ahem et al., 1980), although the enzymes have not been characterized. The bromoperoxidase from the green alga Penicillus capitatus, the most extensively purified and characterized of the group (Manthey and Hager, 1981, 1985), catalyzes an array of reactions comparable to that of the fungal chloroperoxidase; however, it does not decompose organic peroxides and in general, its catalytic properties more closely resembles the "classical" plant peroxidase. Thus, haloperoxidases appear to span a spectrum of peroxidative types, from the plant peroxidases to the human myeloperoxidase. Substantial loss of activity during fractionation has prevented purification of M u r e x bromoperoxidase and quantitative comparison of different proxidative activities. However, the properties of the brominating activity of the crude enzyme have been characterized and these properties resemble those reported for other bromoperoxidases. MATERIALS AND METHODS
Preparation of extract The shells of 1-2 kg of Murex trunculus snails were
crushed in a vise; the hypobranchial gland, a light-brown "vein" in a mantle above the foot, was excised with scissors. The collected glands, ca 5-10 g, were homogenized in 50 ml of 100 mM Tris-acetate buffer, pH 7.4, first in a Waring blender and then in aliquots of 10--15ml in a Con-Torque Eberbach homogenizer (Teflon-glass). The homogenate was centrifuged at 480g for 10min to remove a heavy load of fibrous matter. The aspirated supernatant was recentrifuged at 27,000 g at 4°C for I hr in a Sorvall superspeed refrigerated centrifuge (model RC-2-B). The 27,000 g superuatant contained high aryl sulfatase activity but no bromoperoxidase activity. The sediment was resuspended in 1-2ml of buffer containing 1% Triton X-100 and was cleared by centrifugation at 480g for 10min to yield a slightly turbid, opalescent solution and a precipitate, both of which are purple. The solution contained all the de. tectable bromoperoxidase activity. Omission of Triton X100 from the suspending medium effected a decrease in yield 917
918
RAND^ JASNUNand ELMONLEE Coe
of activity, although a lower detergent concentration (0.3%) was also effective in releasing activity. The Tritonsolubilized sediment was used directly as the "enzyme" in most assays. The combined presence of turbidity and purple indigo dye interfered with standard methods of protein determination. Absorption at 280 nm indicated a protein concentration of ca 20 mg/ml, based on the albumin extinction coefficient; however, this approx, value may be high. Activation of enzyme Early assays showed an initial lag, partly attributable to the high H202 level used; This lag was shortened by a preincubation procedure in which 0.2 mi of solution conraining 0.1 rul of Triton-solubflized sediment and a final concentration of 90 mM Tris-acetate, pH 7.4, 5 mM KBr and 0.5mM H202 was allowed to stand at room temperature for 30 min, after which H202 was added to increase the concentration by 0.5 mM and this solution was allowed to stand for 30 min or longer. Enzyme subjected to this preincubation, designated "activated enzyme", exhibited shorter lag periods. Incubation with KBr alone had no effect; H202 alone caused loss of activity. Activated enzyme was stable at room temperature for at least 24 hr. Assay procedures Bromoperoxidase assays depend on change in absorbance of artificial acceptor molecules with bromination. The decrease in monochlorodimedon (MCD) absorbance at 278 nm on bromination has been used ~'s a standard assay procedure for bromoperoxidase activity (Hewson and Hager, 1980). A second method, more specific for bromination but limited to a neutral pH range, depends on a shift in absorption spectrum as cresol red (CR) is brominated to bromocresol purple. At pH 7.5, the two spectra differ maximally at 600 nm and the increase in absorbance at 600 nm is proportional to the analogous decrease at 278 nm in the MCD assay. Typical assay mixtures contained: Tris-acetate buffer, pH4.0-8.4 (100mM); H202 (0.01-2.0 mM); KBr or KC1 (0.5-20 mM) and either cresol red (0.10 raM) or monochlorodimedon (0.05 raM). Following addition of 5-20 p 1 of enzyme preparation to 1.0 ml of assay mixture, absorbance was recorded at 278 nm (MCD) or 600 nm (CR) with a Bausch and Lomb Spectronic 700 spectrophotometer coupled to a Perkin-Elmer Model 56 recorder. In some instances, H202 was replaced with a H202-generating system in which 50mM glucose was included in the assay mixture and 100-300/~g of glucose oxidase was added 5-10rain prior to addition of enzyme. All assays were conducted at 23-25°C. Iodoperoxidase activity was estimated from the increase in the 350 nm absorption of Ii-, as described by Theiler et aL (1978) and Ahem et aL (1980). Enzyme was omitted from the blank cuvette and absorbance was measured with a Beckman UV-5260 double-beam spectrophotometer. The absorbance-difference curve at 350nm showed a rapid rise and then approached a steady-state level which represented balanced I ; generation and consumption. The maximum rate of increase, 1-2 min after addition of enzyme, was used as an estimate of iodoperoxidase activity. Typical reaction mixtures contained: Tris-acetate buffer (100raM), H202 (0;0i-2.0 mM) and KI (1-20 raM). Changes in bromide ion concentrations were detected with a Ag:AgBr electrode against an identical electrode immersed in a I mM KBr reference solution, using a Sargent-Welch DRLG recorder as a voltmeter.. Disappearance of bromide was clearly discernible despite the small percentage of change in Br- concentration and the logarithmic response. Materials
Monoclilorodimedon, glucose and Triton X-100 were obtained from Sigma Chemical; hydrogen peroxide (30%),
from Baker; Tris (hydroxymethyl) amino methane, from Fisher Scientific; o-cresolsulfophthalein (cresol red), from Eastman Kodak and glucose oxidase (mold, tech. grade), from Fiuka AG. Inorganic salts and acetic acid were analytical reagent grade. RESULTS Initial detection o f activity
First attempts to detect bromoperoxidase activity employed reaction mixtures comparable to those described for chloroperoxidase (Hager et al., 1966) and contained: KBr (10 raM); H202 (0,2 raM); either MCD (0.05 raM) or CR (0.1 raM) and either "Iris or Na acetate buffer (100 mM). Results generalized from several experiments are illustrated in Fig. 1. Curve A represents a typical response in the initial assay in which the enzyme was dormant, became active and then lost activity even though cresol red was yet in excess. The MCD assay system yielded similar results. Measurements with the Ag:AgBr electrode revealed that bromide ion remained constant during the lag period and began to decline coincidentally with the appearance of activity in the other assays. Increasing the enzyme or decreasing the H202 concentration to 0.1 mM decreased the lag but also diminished the total reaction (Curve B). Addition of additional enzyme or KBr after the reaction had ceased in (B) produced no effect, but addition of more H202 (0.1 raM) at time P caused renewed activity (Curve C), which indicated that lack of H202 had produced inactivity. However, the shorter lag at a lower initial H202 concentration also suggested inhibition by high levels of peroxide. Preincubation of the concentrated enzyme in 0.5mMH202 and 5 mM KBr both reduced the lag and increased the velocity. This preincubated enzyme in the presence of lower levels of H202 or of a H202-generating system yielded a nearly immediate, linear reaction (Curve D). In subsequent kinetic studies, only those preparations yielding immediate reaction were used.
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Fig. 1. Illustratioli of bromoperoxidase assay with cresol red. Reaction in 100raM Tris-acetate buffer, pH7.4, 0.1 ram CR, I0 mM KBr and variable level of H202; results generalized from several experiments. Curves: (A) Typical response of freshly prepared enzyme on 0.2raM H202; (13) Effect of either doubling enzyme concentration or decreasing H202 concentration; (C) Effect of adding more H202 to preparation B at time P; (D) Response of activated enzyme at optimal H202 concentration.
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Velocity as a function of enzyme and bromide concentration At lower concentrations of enzyme, the initial velocity was proportional to the amount of enzyme added. In the presence of the optimal level of H20: (20pM at pH 7.4; see below), the rate decreased with time as the peroxide level decreased. A glucoseglucose oxidase peroxide-generating system provided a steady peroxide concentration but required some balancing of the two enzymes. By varying the amount of bromoperoxidase at constant levels of substrate and glucose oxidase, it was possible to establish a
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Fig. 3. Brominating activity as a function of H202 concentration at different pH values. Reaction mixture was 1 ml containing 100 mM Tris-acetate, adjusted to pH values of 4.0-8.4; 10 mM KBr; 0.05 mM monochlordimedon; and H202 concentrations ranging from 10-300 pM. pH values are designated by numbers next to curves; dashed fines indicate single determinations and solid lines, averages of 2 or 3 assays. Optimum peroxide concentrations at pH 5.0 and 7.4 are indicated by arrows. Protein concentration: ca 10 pglml.
range over which the initial velocity was proportional to bromoperoxidase concentration. Subsequently, a bromoperoxidase level was chosen such that measurements would perforce fall within the linear range. A comparison of velocities as a function of KBr concentration (pH 7.4) obtained by three techniques is shown in Fig. 2A. The generating system with the cresol red assay (©) gave the most reproducible values at low activities; higher activities were more easily estimated using the optimal H202 concentration with either the CR assay (X) or the MCD assay (A). A double reciprocal plot of the data in Fig. 2A indicates simple saturation kinetics for bromide ion (Fig. 2B). Application of an unweighted non-linear regression analysis to fit these data to the Michealis-Menten equation, using the program of Oestreicher and Pinto (1983) modified for use wih a TRS-80 Model 100 computer, yields Km= 7.2 _+ 1.1 (SE) and Vm= 34.8 + 3.1. Optimal hydrogen peroxide concentration as a function of p H Inhibition by high concentrations of H202 leads to a definite optimum in the H2Oz concentration. The activity as a logarithmic function of g M HzOz is shown for pH values from 4.0 to 8.4 in Fig. 3. The sharp optimum concentration at each pH increases with decreasing pH. At pH 5.0, the optimum HzO2 concentration is near 125/~M; at pH 7.4, it is only 20tiM. The shifting 1-1202 optimum obscures definition of the pH optimum for the enzyme. In Fig. 4 (upper frame), maximum velocities from Fig. 3 are plotted against pH and an optimum between pH 5 and 6 is observed. However, the pH-activity profile changes with fixed peroxide concentration (Fig. 4, lower frame). At the optimum H202 concentration for pH 5 (125 #M), a sharp pH optimum about pH 5 is observed, whereas at H202 concentration of 20 # M there is a broad pH optimum hetwocn pH 6 and 7. Thus, at lower H202 concentrations, the enzyme is most active in the neutral pH range, but at higher
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Fig. 4. Brominating activity as a function of pH at optimal and at fixed concentrations ofHzO2. Upper frame shows the maximal activities from the curves in Fig. 3. Lower frame gives activity vs pH profile at two fixed H202 concentrations, 20 (optimum for pH 7.4) and 125 #M (optimum for pH 5.0).
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Fig. 5. Inhibition of brominating activity by chloride and fluoride at pH 5.0. Reaction mixture contained 100raM acetate buffer, pHS.0, 200pMH202 and 50pM monochlorodimedon. Double reciprocal plot of curves: (V -- pM/min) ( 0 - - 0 ) with bromide alone, ( x - - u ) in the presence of 100 mM chloride, ( + - - + ) in the presence of 200 mM chloride, (C)---C)) in the presence of 25 mM fluoride, ( & - - - & ) in the presence of 50 mM fluoride.
H 2 0 : concentrations, it is active only at pH 6 or less (Figs 3 and 4). Since all velocities shown in Figs 3 and 4 were at fixed bromide concentration (10 mM), which is in the range of the Km for bromide at pH 7.4 (Fig. 2) and which is less than the Km at pH 5.0 (Fig. 5), these curves would shift towards higher values with increasing bromide levels.
much more complex pattern of inhibition. The K m for ( B r - ) at pH 5.0 is 23-30 mM and the Ki for (C1-) is about 100 mM. At low concentrations (10 raM, not shown) fluoride appears competitive with bromide; however, as fluoride is increased, inhibition shifts first to a non-competitive mode (25 raM) and finally to an uncompetitive or mixed inhibition (50 raM). A similar pattern of fluoride inhibition has been reported for the fungal chloroperoxidase (Hager et ai., 1966).
Inhibition of bromination by chloride and fluoride ions Chloride and fluoride ions were totally inactive as halogenating agents in the M C D assay; however, both inhibited bromination, the more powerful inhibitor being fluoride. Double reciprocal plots (Fig. 5) indicate that chloride acts as a non-competitive or as a mixed non-competitive/uncompetitive inhibitor with respect to bromide, while fluoride exhibits a
Iodoperoxidase activity The occurrence of multiple reactions in the presence of H~ O~, I - , I f , and protein precludes precise measurement of iodoperoxidase. However, the data indicate semiquantitatively that the rate of I~- formation is dependent on I - concentration (Fig. 6B)
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Fig. 6. Comparison of bromoperoxidas¢ and iodoperoxidas¢ activities at pH 7.4. Reaction mixtures: I mi containing 100raM Tris-acetate pH7.4 and variable amounts of H:O 2 or halide ion plus enzyme equivalent to 2 pl (ca 40 pg). Bromoperoxidase was assayed by the decrease in A27s in the presence of 0.05 mM monuehlorodimedon; iodoperoxidase, by the increase in A3~odue to formation of IF in excess of the spontaneous generation of I~-. Bromoperoxidase activities were multiplied by I0. (A) Variable H 202 concentration; KBr or KI were constant at l0 raM. (B) Variable halide (X-) concentrations; H202 was constant at 20pM in Br- or 50pM I-. Assays were at 23-25°C.
Snail bromoperoxidase and that the iodoperoxidase also has a sharp optimum for H20~ concentration (Fig. 6A) which is higher than that for bromoperoxidase. At pH 7.4 and 10mM halide, the iodoperoxidase exhibits 20-fold the activity of the bromoperoxidase when these activities are compared at their respective optimal H202 levels. Activities at higher levels of iodide ion or at lower pH were difficult to estimate because the rate of the spontaneous reactions was high. These rates approached values limited by the assay system rather than by enzyme activity. The similarities in H202 optima suggest that a single enzyme is responsible for both the bromo- and iodoperoxidase activities. Verification of this presumption awaits improved purification. DISCUSSION
Comparison of Murex bromoperoxidase with other bromoperoxidases The terminology used in this paper follows that used in the extensive papers by L. P. Hager and his associates. In this system, haloperoxidases are identified with the most electronegative halide which they will oxidize, even though the relative activity is in the order I - > B r - > CI-. Thus, the fungal (C. fumago) enzyme is a "chloroperoxidase" despite higher activity with bromide and iodide (Hager et al., 1966). Algal (P. capitatus) enzyme is a "bromoperoxidase" because it utilizes bromide but not chloride or fluoride, even though it oxidizes iodide ten times faster than bromide (Manthey and Hager, 1981). Hence, human granulocyte "myeloperoxidase" can be considered a chloroperoxidase and Murex enzyme may be termed a bromoperoxidase. The first bromoperoxidase identified was in a marine red alga, B. hamifera (Theiler et al., 1978). Similar enzymes have since been detected in a wide range of red, green and brown algae, including the extensively characterized enzyme from the green marine alga, P. capitatus (Manthey and Hager, 1981, 1985). Recently, an array of prokaryotes have been found to possess bromoperoxidases (Van Pe'e and Lingens, 1985a, b). The Murex enzyme now provides evidence for a bromoperoxidase from an animal (molluscan) source. Several features common to all known bromoperoxidases may be noted. Firstly, they have pH optima in the mildly acid range (pH 4-6) and exhibit considerable brominating activity around pH 7 (see Baden and Corbett, 1980; Hewson and Hager, 1980; Van Pe'e and Lingens, 1985a, for example). In contrast, the chloroperoxidases (either fungal or human) have lower optima (pH 2.5-5) and are virtually inactive in the neutral pH range (Hager et al., 1966; Bakkenist et al., 1980). Secondly, both bromoperoxidases and cloroperoxidases are inhibited by high levels of H2 O2 although the H2 O5 concentration required varies with the enzyme, pH and halide concentration. The peroxide concentration optima reported for algal bromoperoxidases are near those found for the Murex enzyme at pH values around 5, i.e. 100-200#M, although much higher levels of peroxide ( > 2 mm) may be required for full suppression of activity in some algal systems (Baden and
921
Corbett, 1980; Ahem et al., 1980). Judging from the assay procedures used, the bacterial bromoperoxidases seem insensitive to high levels of H20~ (Van Pe'e and Lingens, 1985a, b). Although Murex enzyme resembles other bromoperoxidases in its general interactions with halides, details vary. Murex enzyme follows a simple saturation curve below 20 mM bromide. Penicillus capitatus behave similarly in this range, but its velocity continues to increase at very high Br- concentration without reaching saturation (Manthey and Hager, 1981). In contrast to these two enzymes, the red alga (R. larix) enzyme exhibits a sharp optimum Brconcentration between 1 and 2 mM and is reduced to less than 10% of maximal activity at 20mM Br(Ahem et al., 1980). By definition, none of the bromoperoxidase utilize chloride, but some, like the Murex enzyme, are inhibited by chloride, whereas others, like the P. capitatus enzyme is not inhibited by even 1 M C1- concentration. All haloperoxidases examined are inhibited by fluoride. The complicated pattern of fluoride inhibition in the Murex enzyme, which progresses from competitive to non-competitive to complex, with respect to bromide, as concentration of fluoride is increased, resembles the pattern of fluoride inhibition with respect to H202 seen in the C. fumago chloroperoxidase (Hager et al., 1966). The latter pattern was interpreted to mean that F - binds to both the Cland the HeO2 sites of the chloroperoxidase. A comparable multiple binding may also occur with the Murex bromoperoxidase. One peculiar property of the snail bromoperoxidase is its activation by preincubation with H202 together with Br-. This characteristic suggests either that some bromination of the enzyme itself may be necessary for full activity, or that some inhibitor is being inactivated. As with other haloperoxidases, preincubation with H202 alone causes a slow, irreversible inactivation. Interrelationships among halide, hydrogen peroxide and hydrogen ion concentrations The dependence of pH optimum on H2 O2 concentration noted with Murex bromoperoxidase (Fig. 4) has not been reported for other bromoperoxidases, althoug h a comparable dependence has been observed with myeloperoxidase (Bakkenist et al., 1980). A similar shift in pH optimum with halide concentration has been reported for both fungal chloroperoxidase (Thomas et al., 1970) and myeloperoxidase. Generally, increasing halide concentration increases the pH optimum, whereas increasing HeO2 lowers the pH optimum. Other peroxidase activities of the Murex enzyme Other haloperoxidases have peroxidase and catalase activities as well as halogenating activity. These have been most extensively studied in fungal chloroperoxidases (Thomas et al., 1970), although algal bromoperoxidases catalyze a comparable array of reactions (Manthey and Hager, 1981). Catalase activity is present in the Murex preparation and it is higher at pH 7.4 than at pH 4 or 5. This may be compared to the higher pH optimum of the catalase activity in the purified fungal chloroperoxidase (Thomas et al., 1970) and bacterial bromo-
RANDAJANNUN and ELMON LEE COE
922
peroxidases (Van Pe'e and Lingens, 1985a). However, further purification of the crude Murex enzyme will be required before the catalase activity can be attributed to the bromoperoxidase. Biological aspects A brominated natural product in a sea snail necessitates selection of bromide from a vast excess of chloride without confusion with iodide. In sea-water the concentration of bromide (0.8raM) is three orders of magnitude greater than that of iodide (0.4/~M) and three orders of magnitudes less than that of chloride (0.54 M). Yet, the enzyme shows no detectable activity with chloride. Even though it may be 20 times more active in oxidizing iodide ion, the lower concentration of iodide in sea-water would reduce the iodination rate to only 1% of the broruination rate. The advantage reaped by the snail for synthesizing exotic indoxyl derivatives remains enigmatic; indeed, at the height of the fashion of Tyrian Purple, this unusual skill had a negative survival value. Acknowledgements--The authors thank Mr Joseph Doumet for his provision of fresh Murex trunculus and for his continued interest in this project, Dr Naziha Nuwayhid for technical advice and assistance and Dr Chiadao Chen for sharing his expertise on haloperoxidase assays and oxygenase mechanisms. They are also grateful for Mr Habib Abu Diwan for crunching snail shells and to Mr W. Steven Ward for his creative suggestions. This research was supported by grants from the A.U.B. Medical Research Committee and the Diana Tamari Sabbagh Fund. REFERENCES
Ahem T. J., Allan G. G. and Medcalf D. G. (1980) New bromoperoxidases of marine origin: partial purification and characterization. Biochim. biophys. Acta 616, 329-339. Baden D. G. and Corbctt M. D. (1980) Bromoperoxidases from Penicillus capitatus, Penicillus lamourouxii, and Rhipocephalus phoenix. Biochem. J. 187, 205-211. Baker J. T. (1974) Tyrian purple: an ancient dye, a modern problem. Endeavour 33, 11-17. Bakkenist A. R., De Boer J. E., Plat H. and Wever R. (1980) The halide complexes of myeloperoxidase and the mechanism of the halogenation reactions. Biochim. biophys. Acta 613, 337-348.
Clark R. A. and Klebanoff S. J. (1979) Role of the myeloperoxidase-H202-halide system in concanavalin A induced tumor cell killing by human neutrophils. J. lmmun. 122, 2605-2610. De Boer E., Van Kooyk Y., Tromp M. G. M., Plat H. and Wever R. (1986) Bromoperoxidase from Ascophyllum nodosum: a novel class of enzymes containing vanadium as a prosthetic group? Biochim. biophys. Acta 869, 48-53. Friedlander P. (1909) Uber den Farbstoff des antiken Purpurs ans Murex brandaris. Ber. Dtsch. Chem. Ges. 42, 765-770. Hager L. P., Morris D. R., Brown F. S. and Eberwcin H. (1966) Chloroperoxidase---II. Utilization of halogen anions. J. biol. Chem. 241, 1769-1777. Hewson W. D. and Hager L. P. (1980) Bromoperoxidases and halogenated lipids in marine algae. Y. Phycol. 16, 340-345. Itoh N., Izumi Y. and Yamada H. (1986) Characterization of nonheme type bromoperoxidase in Corallina pilulifera. J. biol. Chem. 261, 5195-5200. Jannun R., Nuwayhid N. and Coe E. L. (1981) Biological bromination: bromopcroxidase activity in the Murex sea snail. Fed. Proc. 40, 1774. Manthey J. A. and Hager L. P. (1981) Purification and properties of bromoperoxidase from Penicillus capitatus. J. biol. Chem. 256, 11232-11238. Manthey J. A. and Hager L. P. (1985) Characterization of the oxidized states of bromoperoxidase. J. biol. Chem. 260, 9654-9659. Matheson N. R., Wong P. S. and Travis J. (1981) Isolation and properties of human neutrophil myeloperoxidase. Biochemistry 20, 325-330. Oestreicher E. G. and Pinto G. F. (1983) Pocket computer program for fitting the Michaelis-Mentenequation. Cornput. biol. Med. 13, 309-315. Theiler R. F., Cook J. C. and Hager L. P. (1978) Halohydrocarbon synthesis by bromoperoxidase. Science 202, 1094-1096. Thomas J. A., Morris D. R. and Hager L. P. (1970) Chloroperoxidase---VIII. Classical peroxidatic, catalytic, and halogenating forms of the enzyme. J. biol. Chem. 245, 3129-3134. Van Pe'e K. H. and Lingens F. (1985a) Purification of bromoperoxidase from Pseudomonas aureofaciens. J. Bact. 161, 1171-1175. Van Pe'e K. H. and Lingens F. (1985b) Purification and molecular and catalytic properties of bromoperoxidase from Streptomyces phaeochromogenes. J. Gen. Microb. 131, 1911-1916. Wcisner W., Van Pe'e K. H. and Lingens H. (1985) Purification and properties of bromoperoxidase from Pseudomonas pyrrocinia. Biol. Chem. Hoppe-Seyler 366, 1085-1091.
0305-0491/87 $3.00+ 0.00 © 1987 Pergamon Journals Ltd
BROMOPEROXIDASE FROM THE MARINE SNAIL, M U R E X TRUNCULUS RANDA JANNUN* and ELMON L ~ COE? Biochemistry Department, American University of Beirut, Beirut, Lebanon (Received 5 January 1987)
Abstraet--l. Extracts of a 25,000 g sediment of Murex trunculus hypobranchial gland homogenate contain an enzyme which brominates monochlorodimedon in the presence of bromide and HzO2. 2. Dependence of activity on H z02 concentration exhibits a sharp optimum which increases from 20/~M at pH 7.4 to 400/~M at pH 4.0. 3. Dependence on Br- concentration follows a simple saturation curve with Km values of 7 mM at pH 7.4 and 30 mM at pH 5.0. 4. The pH optima depend on H20: concentration, ranging from pH 6.6 at 10 #M to pH 4.8 at 200/~M. 5. The enzyme is totally inactive with CI- and F-, although both these ions inhibit bromination.
INTRODUCTION
Carnivorous marine snails of the genera M u r e x and Purpura are known to release compounds which, when exposed to air and sunlight, form a fast purple dye. Friedlandcr (1909) first identified the major component in the purple dye as 6,6' dibromoindigo and subsequent work has shown that the snail releases a collection of brominated and unbrominated indoxyl sulfate derivatives which, after hydrolysis by an aryl sulfatase, spontaneously dimerize in the presence of oxygen and light to form indigo derivatives (Baker, 1974). Occurrence of the brominated intermediates implied the presence of a brominating enzyme and prompted an investigation of Murex gland extracts for bromoperoxidase activity. Such activity was demonstrated, as has been reported in preliminary communications (Jannun et al., 1981). Exclusive of the well-known iodoperoxidase activities of several classical peroxidases, the first enzyme especially identified as a haloperoxidase was the chloroperoxidase from Caldariomyces fumago. This soluble enzyme, which has been highly purified and crystallized, requires H2Oz, acts on bromide, iodide and chloride (Hager et al, 1966) and can act as a catalase and a "classical peroxidase" (Thomas et al., 1970). More recently, another enzyme which has been classified with chloroperoxidase as a C I - - H 2 0 :oxidoreductase, the so-called "myeloperoxidase", has been highly purified from human neutrophils (Matheson et al., 1981). Mydoperoxidase exhibits catalytic properties similar to those of fungal chloroperoxidase (Bakkenist et al., 1980; Clark and Klebanoff, 1979). Bromoperoxidase activity was first reported in marine algae (Theiler et al., 1978). Subsequently, it has been identified and characterized in many species of marine algae (Baden and Corbett, 1980; Hewson and Hager, 1980; A h e m et al., 1980; De Boer et aL, *Present Address: Institute for Molecular Virology, St Louis University, School of Medicine, St Louis, MO, USA. 1"Present Address: Arabian Gulf University, Medical College Manama, Bahrain.
1986; Itoh et al., 1986) and in several species of bacteria (Van Pe'e and Lingens, 1985a, b; Weisner et al., 1985). Bromoperoxidase activity has also been detected in a marine annelid and in a hemichordate (Ahem et al., 1980), although the enzymes have not been characterized. The bromoperoxidase from the green alga Penicillus capitatus, the most extensively purified and characterized of the group (Manthey and Hager, 1981, 1985), catalyzes an array of reactions comparable to that of the fungal chloroperoxidase; however, it does not decompose organic peroxides and in general, its catalytic properties more closely resembles the "classical" plant peroxidase. Thus, haloperoxidases appear to span a spectrum of peroxidative types, from the plant peroxidases to the human myeloperoxidase. Substantial loss of activity during fractionation has prevented purification of M u r e x bromoperoxidase and quantitative comparison of different proxidative activities. However, the properties of the brominating activity of the crude enzyme have been characterized and these properties resemble those reported for other bromoperoxidases. MATERIALS AND METHODS
Preparation of extract The shells of 1-2 kg of Murex trunculus snails were
crushed in a vise; the hypobranchial gland, a light-brown "vein" in a mantle above the foot, was excised with scissors. The collected glands, ca 5-10 g, were homogenized in 50 ml of 100 mM Tris-acetate buffer, pH 7.4, first in a Waring blender and then in aliquots of 10--15ml in a Con-Torque Eberbach homogenizer (Teflon-glass). The homogenate was centrifuged at 480g for 10min to remove a heavy load of fibrous matter. The aspirated supernatant was recentrifuged at 27,000 g at 4°C for I hr in a Sorvall superspeed refrigerated centrifuge (model RC-2-B). The 27,000 g superuatant contained high aryl sulfatase activity but no bromoperoxidase activity. The sediment was resuspended in 1-2ml of buffer containing 1% Triton X-100 and was cleared by centrifugation at 480g for 10min to yield a slightly turbid, opalescent solution and a precipitate, both of which are purple. The solution contained all the de. tectable bromoperoxidase activity. Omission of Triton X100 from the suspending medium effected a decrease in yield 917
918
RAND^ JASNUNand ELMONLEE Coe
of activity, although a lower detergent concentration (0.3%) was also effective in releasing activity. The Tritonsolubilized sediment was used directly as the "enzyme" in most assays. The combined presence of turbidity and purple indigo dye interfered with standard methods of protein determination. Absorption at 280 nm indicated a protein concentration of ca 20 mg/ml, based on the albumin extinction coefficient; however, this approx, value may be high. Activation of enzyme Early assays showed an initial lag, partly attributable to the high H202 level used; This lag was shortened by a preincubation procedure in which 0.2 mi of solution conraining 0.1 rul of Triton-solubflized sediment and a final concentration of 90 mM Tris-acetate, pH 7.4, 5 mM KBr and 0.5mM H202 was allowed to stand at room temperature for 30 min, after which H202 was added to increase the concentration by 0.5 mM and this solution was allowed to stand for 30 min or longer. Enzyme subjected to this preincubation, designated "activated enzyme", exhibited shorter lag periods. Incubation with KBr alone had no effect; H202 alone caused loss of activity. Activated enzyme was stable at room temperature for at least 24 hr. Assay procedures Bromoperoxidase assays depend on change in absorbance of artificial acceptor molecules with bromination. The decrease in monochlorodimedon (MCD) absorbance at 278 nm on bromination has been used ~'s a standard assay procedure for bromoperoxidase activity (Hewson and Hager, 1980). A second method, more specific for bromination but limited to a neutral pH range, depends on a shift in absorption spectrum as cresol red (CR) is brominated to bromocresol purple. At pH 7.5, the two spectra differ maximally at 600 nm and the increase in absorbance at 600 nm is proportional to the analogous decrease at 278 nm in the MCD assay. Typical assay mixtures contained: Tris-acetate buffer, pH4.0-8.4 (100mM); H202 (0.01-2.0 mM); KBr or KC1 (0.5-20 mM) and either cresol red (0.10 raM) or monochlorodimedon (0.05 raM). Following addition of 5-20 p 1 of enzyme preparation to 1.0 ml of assay mixture, absorbance was recorded at 278 nm (MCD) or 600 nm (CR) with a Bausch and Lomb Spectronic 700 spectrophotometer coupled to a Perkin-Elmer Model 56 recorder. In some instances, H202 was replaced with a H202-generating system in which 50mM glucose was included in the assay mixture and 100-300/~g of glucose oxidase was added 5-10rain prior to addition of enzyme. All assays were conducted at 23-25°C. Iodoperoxidase activity was estimated from the increase in the 350 nm absorption of Ii-, as described by Theiler et aL (1978) and Ahem et aL (1980). Enzyme was omitted from the blank cuvette and absorbance was measured with a Beckman UV-5260 double-beam spectrophotometer. The absorbance-difference curve at 350nm showed a rapid rise and then approached a steady-state level which represented balanced I ; generation and consumption. The maximum rate of increase, 1-2 min after addition of enzyme, was used as an estimate of iodoperoxidase activity. Typical reaction mixtures contained: Tris-acetate buffer (100raM), H202 (0;0i-2.0 mM) and KI (1-20 raM). Changes in bromide ion concentrations were detected with a Ag:AgBr electrode against an identical electrode immersed in a I mM KBr reference solution, using a Sargent-Welch DRLG recorder as a voltmeter.. Disappearance of bromide was clearly discernible despite the small percentage of change in Br- concentration and the logarithmic response. Materials
Monoclilorodimedon, glucose and Triton X-100 were obtained from Sigma Chemical; hydrogen peroxide (30%),
from Baker; Tris (hydroxymethyl) amino methane, from Fisher Scientific; o-cresolsulfophthalein (cresol red), from Eastman Kodak and glucose oxidase (mold, tech. grade), from Fiuka AG. Inorganic salts and acetic acid were analytical reagent grade. RESULTS Initial detection o f activity
First attempts to detect bromoperoxidase activity employed reaction mixtures comparable to those described for chloroperoxidase (Hager et al., 1966) and contained: KBr (10 raM); H202 (0,2 raM); either MCD (0.05 raM) or CR (0.1 raM) and either "Iris or Na acetate buffer (100 mM). Results generalized from several experiments are illustrated in Fig. 1. Curve A represents a typical response in the initial assay in which the enzyme was dormant, became active and then lost activity even though cresol red was yet in excess. The MCD assay system yielded similar results. Measurements with the Ag:AgBr electrode revealed that bromide ion remained constant during the lag period and began to decline coincidentally with the appearance of activity in the other assays. Increasing the enzyme or decreasing the H202 concentration to 0.1 mM decreased the lag but also diminished the total reaction (Curve B). Addition of additional enzyme or KBr after the reaction had ceased in (B) produced no effect, but addition of more H202 (0.1 raM) at time P caused renewed activity (Curve C), which indicated that lack of H202 had produced inactivity. However, the shorter lag at a lower initial H202 concentration also suggested inhibition by high levels of peroxide. Preincubation of the concentrated enzyme in 0.5mMH202 and 5 mM KBr both reduced the lag and increased the velocity. This preincubated enzyme in the presence of lower levels of H202 or of a H202-generating system yielded a nearly immediate, linear reaction (Curve D). In subsequent kinetic studies, only those preparations yielding immediate reaction were used.
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Fig. 1. Illustratioli of bromoperoxidase assay with cresol red. Reaction in 100raM Tris-acetate buffer, pH7.4, 0.1 ram CR, I0 mM KBr and variable level of H202; results generalized from several experiments. Curves: (A) Typical response of freshly prepared enzyme on 0.2raM H202; (13) Effect of either doubling enzyme concentration or decreasing H202 concentration; (C) Effect of adding more H202 to preparation B at time P; (D) Response of activated enzyme at optimal H202 concentration.
919
Snail bromoperoxidase x
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I/ K B r ] mM-' Fig. 2. Initial velocity of bromination at pH 7.4 as a function of bromide concentration. (A) Comparison of velocities obtained by three different procedures: ( 0 ) assay with glucose oxidase H202-generating system and cresol red, as described in text; (X), assay with CR in the presence of 20 pM H202, (A), assay with 0.05 mM monochlorodimedon in the presence of 20 pM H202. All reactions were carried out at 23-25°C. (B) Double reciprocal plot of curve shown in frame (A). Protein concentration: ca 100 l~g/mi. [KBr
]
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Velocity as a function of enzyme and bromide concentration At lower concentrations of enzyme, the initial velocity was proportional to the amount of enzyme added. In the presence of the optimal level of H20: (20pM at pH 7.4; see below), the rate decreased with time as the peroxide level decreased. A glucoseglucose oxidase peroxide-generating system provided a steady peroxide concentration but required some balancing of the two enzymes. By varying the amount of bromoperoxidase at constant levels of substrate and glucose oxidase, it was possible to establish a
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Fig. 3. Brominating activity as a function of H202 concentration at different pH values. Reaction mixture was 1 ml containing 100 mM Tris-acetate, adjusted to pH values of 4.0-8.4; 10 mM KBr; 0.05 mM monochlordimedon; and H202 concentrations ranging from 10-300 pM. pH values are designated by numbers next to curves; dashed fines indicate single determinations and solid lines, averages of 2 or 3 assays. Optimum peroxide concentrations at pH 5.0 and 7.4 are indicated by arrows. Protein concentration: ca 10 pglml.
range over which the initial velocity was proportional to bromoperoxidase concentration. Subsequently, a bromoperoxidase level was chosen such that measurements would perforce fall within the linear range. A comparison of velocities as a function of KBr concentration (pH 7.4) obtained by three techniques is shown in Fig. 2A. The generating system with the cresol red assay (©) gave the most reproducible values at low activities; higher activities were more easily estimated using the optimal H202 concentration with either the CR assay (X) or the MCD assay (A). A double reciprocal plot of the data in Fig. 2A indicates simple saturation kinetics for bromide ion (Fig. 2B). Application of an unweighted non-linear regression analysis to fit these data to the Michealis-Menten equation, using the program of Oestreicher and Pinto (1983) modified for use wih a TRS-80 Model 100 computer, yields Km= 7.2 _+ 1.1 (SE) and Vm= 34.8 + 3.1. Optimal hydrogen peroxide concentration as a function of p H Inhibition by high concentrations of H202 leads to a definite optimum in the H2Oz concentration. The activity as a logarithmic function of g M HzOz is shown for pH values from 4.0 to 8.4 in Fig. 3. The sharp optimum concentration at each pH increases with decreasing pH. At pH 5.0, the optimum HzO2 concentration is near 125/~M; at pH 7.4, it is only 20tiM. The shifting 1-1202 optimum obscures definition of the pH optimum for the enzyme. In Fig. 4 (upper frame), maximum velocities from Fig. 3 are plotted against pH and an optimum between pH 5 and 6 is observed. However, the pH-activity profile changes with fixed peroxide concentration (Fig. 4, lower frame). At the optimum H202 concentration for pH 5 (125 #M), a sharp pH optimum about pH 5 is observed, whereas at H202 concentration of 20 # M there is a broad pH optimum hetwocn pH 6 and 7. Thus, at lower H202 concentrations, the enzyme is most active in the neutral pH range, but at higher
920
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Fig. 4. Brominating activity as a function of pH at optimal and at fixed concentrations ofHzO2. Upper frame shows the maximal activities from the curves in Fig. 3. Lower frame gives activity vs pH profile at two fixed H202 concentrations, 20 (optimum for pH 7.4) and 125 #M (optimum for pH 5.0).
,,,
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Fig. 5. Inhibition of brominating activity by chloride and fluoride at pH 5.0. Reaction mixture contained 100raM acetate buffer, pHS.0, 200pMH202 and 50pM monochlorodimedon. Double reciprocal plot of curves: (V -- pM/min) ( 0 - - 0 ) with bromide alone, ( x - - u ) in the presence of 100 mM chloride, ( + - - + ) in the presence of 200 mM chloride, (C)---C)) in the presence of 25 mM fluoride, ( & - - - & ) in the presence of 50 mM fluoride.
H 2 0 : concentrations, it is active only at pH 6 or less (Figs 3 and 4). Since all velocities shown in Figs 3 and 4 were at fixed bromide concentration (10 mM), which is in the range of the Km for bromide at pH 7.4 (Fig. 2) and which is less than the Km at pH 5.0 (Fig. 5), these curves would shift towards higher values with increasing bromide levels.
much more complex pattern of inhibition. The K m for ( B r - ) at pH 5.0 is 23-30 mM and the Ki for (C1-) is about 100 mM. At low concentrations (10 raM, not shown) fluoride appears competitive with bromide; however, as fluoride is increased, inhibition shifts first to a non-competitive mode (25 raM) and finally to an uncompetitive or mixed inhibition (50 raM). A similar pattern of fluoride inhibition has been reported for the fungal chloroperoxidase (Hager et ai., 1966).
Inhibition of bromination by chloride and fluoride ions Chloride and fluoride ions were totally inactive as halogenating agents in the M C D assay; however, both inhibited bromination, the more powerful inhibitor being fluoride. Double reciprocal plots (Fig. 5) indicate that chloride acts as a non-competitive or as a mixed non-competitive/uncompetitive inhibitor with respect to bromide, while fluoride exhibits a
Iodoperoxidase activity The occurrence of multiple reactions in the presence of H~ O~, I - , I f , and protein precludes precise measurement of iodoperoxidase. However, the data indicate semiquantitatively that the rate of I~- formation is dependent on I - concentration (Fig. 6B)
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Fig. 6. Comparison of bromoperoxidas¢ and iodoperoxidas¢ activities at pH 7.4. Reaction mixtures: I mi containing 100raM Tris-acetate pH7.4 and variable amounts of H:O 2 or halide ion plus enzyme equivalent to 2 pl (ca 40 pg). Bromoperoxidase was assayed by the decrease in A27s in the presence of 0.05 mM monuehlorodimedon; iodoperoxidase, by the increase in A3~odue to formation of IF in excess of the spontaneous generation of I~-. Bromoperoxidase activities were multiplied by I0. (A) Variable H 202 concentration; KBr or KI were constant at l0 raM. (B) Variable halide (X-) concentrations; H202 was constant at 20pM in Br- or 50pM I-. Assays were at 23-25°C.
Snail bromoperoxidase and that the iodoperoxidase also has a sharp optimum for H20~ concentration (Fig. 6A) which is higher than that for bromoperoxidase. At pH 7.4 and 10mM halide, the iodoperoxidase exhibits 20-fold the activity of the bromoperoxidase when these activities are compared at their respective optimal H202 levels. Activities at higher levels of iodide ion or at lower pH were difficult to estimate because the rate of the spontaneous reactions was high. These rates approached values limited by the assay system rather than by enzyme activity. The similarities in H202 optima suggest that a single enzyme is responsible for both the bromo- and iodoperoxidase activities. Verification of this presumption awaits improved purification. DISCUSSION
Comparison of Murex bromoperoxidase with other bromoperoxidases The terminology used in this paper follows that used in the extensive papers by L. P. Hager and his associates. In this system, haloperoxidases are identified with the most electronegative halide which they will oxidize, even though the relative activity is in the order I - > B r - > CI-. Thus, the fungal (C. fumago) enzyme is a "chloroperoxidase" despite higher activity with bromide and iodide (Hager et al., 1966). Algal (P. capitatus) enzyme is a "bromoperoxidase" because it utilizes bromide but not chloride or fluoride, even though it oxidizes iodide ten times faster than bromide (Manthey and Hager, 1981). Hence, human granulocyte "myeloperoxidase" can be considered a chloroperoxidase and Murex enzyme may be termed a bromoperoxidase. The first bromoperoxidase identified was in a marine red alga, B. hamifera (Theiler et al., 1978). Similar enzymes have since been detected in a wide range of red, green and brown algae, including the extensively characterized enzyme from the green marine alga, P. capitatus (Manthey and Hager, 1981, 1985). Recently, an array of prokaryotes have been found to possess bromoperoxidases (Van Pe'e and Lingens, 1985a, b). The Murex enzyme now provides evidence for a bromoperoxidase from an animal (molluscan) source. Several features common to all known bromoperoxidases may be noted. Firstly, they have pH optima in the mildly acid range (pH 4-6) and exhibit considerable brominating activity around pH 7 (see Baden and Corbett, 1980; Hewson and Hager, 1980; Van Pe'e and Lingens, 1985a, for example). In contrast, the chloroperoxidases (either fungal or human) have lower optima (pH 2.5-5) and are virtually inactive in the neutral pH range (Hager et al., 1966; Bakkenist et al., 1980). Secondly, both bromoperoxidases and cloroperoxidases are inhibited by high levels of H2 O2 although the H2 O5 concentration required varies with the enzyme, pH and halide concentration. The peroxide concentration optima reported for algal bromoperoxidases are near those found for the Murex enzyme at pH values around 5, i.e. 100-200#M, although much higher levels of peroxide ( > 2 mm) may be required for full suppression of activity in some algal systems (Baden and
921
Corbett, 1980; Ahem et al., 1980). Judging from the assay procedures used, the bacterial bromoperoxidases seem insensitive to high levels of H20~ (Van Pe'e and Lingens, 1985a, b). Although Murex enzyme resembles other bromoperoxidases in its general interactions with halides, details vary. Murex enzyme follows a simple saturation curve below 20 mM bromide. Penicillus capitatus behave similarly in this range, but its velocity continues to increase at very high Br- concentration without reaching saturation (Manthey and Hager, 1981). In contrast to these two enzymes, the red alga (R. larix) enzyme exhibits a sharp optimum Brconcentration between 1 and 2 mM and is reduced to less than 10% of maximal activity at 20mM Br(Ahem et al., 1980). By definition, none of the bromoperoxidase utilize chloride, but some, like the Murex enzyme, are inhibited by chloride, whereas others, like the P. capitatus enzyme is not inhibited by even 1 M C1- concentration. All haloperoxidases examined are inhibited by fluoride. The complicated pattern of fluoride inhibition in the Murex enzyme, which progresses from competitive to non-competitive to complex, with respect to bromide, as concentration of fluoride is increased, resembles the pattern of fluoride inhibition with respect to H202 seen in the C. fumago chloroperoxidase (Hager et al., 1966). The latter pattern was interpreted to mean that F - binds to both the Cland the HeO2 sites of the chloroperoxidase. A comparable multiple binding may also occur with the Murex bromoperoxidase. One peculiar property of the snail bromoperoxidase is its activation by preincubation with H202 together with Br-. This characteristic suggests either that some bromination of the enzyme itself may be necessary for full activity, or that some inhibitor is being inactivated. As with other haloperoxidases, preincubation with H202 alone causes a slow, irreversible inactivation. Interrelationships among halide, hydrogen peroxide and hydrogen ion concentrations The dependence of pH optimum on H2 O2 concentration noted with Murex bromoperoxidase (Fig. 4) has not been reported for other bromoperoxidases, althoug h a comparable dependence has been observed with myeloperoxidase (Bakkenist et al., 1980). A similar shift in pH optimum with halide concentration has been reported for both fungal chloroperoxidase (Thomas et al., 1970) and myeloperoxidase. Generally, increasing halide concentration increases the pH optimum, whereas increasing HeO2 lowers the pH optimum. Other peroxidase activities of the Murex enzyme Other haloperoxidases have peroxidase and catalase activities as well as halogenating activity. These have been most extensively studied in fungal chloroperoxidases (Thomas et al., 1970), although algal bromoperoxidases catalyze a comparable array of reactions (Manthey and Hager, 1981). Catalase activity is present in the Murex preparation and it is higher at pH 7.4 than at pH 4 or 5. This may be compared to the higher pH optimum of the catalase activity in the purified fungal chloroperoxidase (Thomas et al., 1970) and bacterial bromo-
RANDAJANNUN and ELMON LEE COE
922
peroxidases (Van Pe'e and Lingens, 1985a). However, further purification of the crude Murex enzyme will be required before the catalase activity can be attributed to the bromoperoxidase. Biological aspects A brominated natural product in a sea snail necessitates selection of bromide from a vast excess of chloride without confusion with iodide. In sea-water the concentration of bromide (0.8raM) is three orders of magnitude greater than that of iodide (0.4/~M) and three orders of magnitudes less than that of chloride (0.54 M). Yet, the enzyme shows no detectable activity with chloride. Even though it may be 20 times more active in oxidizing iodide ion, the lower concentration of iodide in sea-water would reduce the iodination rate to only 1% of the broruination rate. The advantage reaped by the snail for synthesizing exotic indoxyl derivatives remains enigmatic; indeed, at the height of the fashion of Tyrian Purple, this unusual skill had a negative survival value. Acknowledgements--The authors thank Mr Joseph Doumet for his provision of fresh Murex trunculus and for his continued interest in this project, Dr Naziha Nuwayhid for technical advice and assistance and Dr Chiadao Chen for sharing his expertise on haloperoxidase assays and oxygenase mechanisms. They are also grateful for Mr Habib Abu Diwan for crunching snail shells and to Mr W. Steven Ward for his creative suggestions. This research was supported by grants from the A.U.B. Medical Research Committee and the Diana Tamari Sabbagh Fund. REFERENCES
Ahem T. J., Allan G. G. and Medcalf D. G. (1980) New bromoperoxidases of marine origin: partial purification and characterization. Biochim. biophys. Acta 616, 329-339. Baden D. G. and Corbctt M. D. (1980) Bromoperoxidases from Penicillus capitatus, Penicillus lamourouxii, and Rhipocephalus phoenix. Biochem. J. 187, 205-211. Baker J. T. (1974) Tyrian purple: an ancient dye, a modern problem. Endeavour 33, 11-17. Bakkenist A. R., De Boer J. E., Plat H. and Wever R. (1980) The halide complexes of myeloperoxidase and the mechanism of the halogenation reactions. Biochim. biophys. Acta 613, 337-348.
Clark R. A. and Klebanoff S. J. (1979) Role of the myeloperoxidase-H202-halide system in concanavalin A induced tumor cell killing by human neutrophils. J. lmmun. 122, 2605-2610. De Boer E., Van Kooyk Y., Tromp M. G. M., Plat H. and Wever R. (1986) Bromoperoxidase from Ascophyllum nodosum: a novel class of enzymes containing vanadium as a prosthetic group? Biochim. biophys. Acta 869, 48-53. Friedlander P. (1909) Uber den Farbstoff des antiken Purpurs ans Murex brandaris. Ber. Dtsch. Chem. Ges. 42, 765-770. Hager L. P., Morris D. R., Brown F. S. and Eberwcin H. (1966) Chloroperoxidase---II. Utilization of halogen anions. J. biol. Chem. 241, 1769-1777. Hewson W. D. and Hager L. P. (1980) Bromoperoxidases and halogenated lipids in marine algae. Y. Phycol. 16, 340-345. Itoh N., Izumi Y. and Yamada H. (1986) Characterization of nonheme type bromoperoxidase in Corallina pilulifera. J. biol. Chem. 261, 5195-5200. Jannun R., Nuwayhid N. and Coe E. L. (1981) Biological bromination: bromopcroxidase activity in the Murex sea snail. Fed. Proc. 40, 1774. Manthey J. A. and Hager L. P. (1981) Purification and properties of bromoperoxidase from Penicillus capitatus. J. biol. Chem. 256, 11232-11238. Manthey J. A. and Hager L. P. (1985) Characterization of the oxidized states of bromoperoxidase. J. biol. Chem. 260, 9654-9659. Matheson N. R., Wong P. S. and Travis J. (1981) Isolation and properties of human neutrophil myeloperoxidase. Biochemistry 20, 325-330. Oestreicher E. G. and Pinto G. F. (1983) Pocket computer program for fitting the Michaelis-Mentenequation. Cornput. biol. Med. 13, 309-315. Theiler R. F., Cook J. C. and Hager L. P. (1978) Halohydrocarbon synthesis by bromoperoxidase. Science 202, 1094-1096. Thomas J. A., Morris D. R. and Hager L. P. (1970) Chloroperoxidase---VIII. Classical peroxidatic, catalytic, and halogenating forms of the enzyme. J. biol. Chem. 245, 3129-3134. Van Pe'e K. H. and Lingens F. (1985a) Purification of bromoperoxidase from Pseudomonas aureofaciens. J. Bact. 161, 1171-1175. Van Pe'e K. H. and Lingens F. (1985b) Purification and molecular and catalytic properties of bromoperoxidase from Streptomyces phaeochromogenes. J. Gen. Microb. 131, 1911-1916. Wcisner W., Van Pe'e K. H. and Lingens H. (1985) Purification and properties of bromoperoxidase from Pseudomonas pyrrocinia. Biol. Chem. Hoppe-Seyler 366, 1085-1091.