ARCHIVES
OF BIOCHEMISTRY
AND
BIOPHYSICS
Vol. 290, No. 1, October, pp. 173-178, 1991
Metabolism of Cyanide by Phanerochaete chrysosporium’ Manish
M. Shah, Thomas
Biotechnology
Center,
Received February
Utah
A. Grover,
State
University,
and Steven Logan,
Utah
D. Aust’
84322-4700
21, 1991, and in revised form June 16, 1991
The oxidation of veratryl alcohol (3,4-dimethoxybenzyl alcohol) by lignin peroxidase H2 (Lip H2) from the white rot fungus Phanerochaete chrysosporium was strongly inhibited by sodium cyanide. The IeO was estimated to be about 2-3 PM. In contrast, sodium cyanide binds to the native enzyme with an apparent sodium cyanide dissociation constant Kd of about 10 jtM. Inhibition of the veratryl alcohol oxidase activity of LiP H2 by cyanide was reversible. Ligninolytic cultures of P. chrysosporium mineralized cyanide at a rate that was proportional to the concentration of cyanide to 2 mM. The N-tert-butyl-a-phenylnitrone-cyanyl radical adduct was observed by ESR spin trapping upon incubation of LiP H2 with HzOz and sodium cyanide. The identity of the spin adduct was confirmed using 13C-labeled cyanide. Six-day-old cultures of the fungus were more tolerant to sodium cyanide toxicity than spores. Toxicity measurements were based on the effect of sodium cyanide on respiration of the fungus as determined by the metabolism of [ “C]glucose to [‘4C]C02. We propose that this tolerance of the mature fungus was due to its ability to mineralize cyanide and that this fungus might be effective in treating environmental pollution sites contaminated with cyanide. 0 1991
Academic
Press,
Inc.
Cyanides have been designated hazardous substances and priority toxic pollutants by the EPA. Discharge of cyanide into the environment is limited by its extreme toxicity to fish rather than man and microorganisms. The electroplating, steel, carbonization, and other important industries produce large quantities of cyanide wastes. To prevent damage to natural ecosystems, the cyanide content of these effluents must be reduced to essentially zero before discharge into the environment. Even for discharge
into a sewer there must be a reduction to no more than 1 to 2 pg of cyanide per milliliter (1). Conventional chemical methods of disposal of cyanide wastes are wasteful of resources, since other chemicals, such as chlorine, are required for detoxification of the cyanide (2). In addition, chemical degradation is relatively expensive and often requires further disposal of the products. The wood rotting fungus Phanerochaete chrysosporium possesses remarkable biodegradative properties. This fungus is one of the relatively few microorganisms known to degrade lignin to carbon dioxide (3). P. chrysosporium is also able to degrade a wide variety of environmentally persistent xenobiotics such as DDT, lindane, benzo[a]pyrene, and polychlorinated biphenyls to carbon dioxide (4). Recent evidence demonstrates that the lignin degrading system is responsible, at least in part, for the ability of P. chrysosporium to degrade xenobiotics (5-7). P. chrysosporium secretes a number of peroxidases; over 10 have been detected in extracellular fluid of lignolytic cultures (8). Kirk et al. have named these proteins Hl to HlO in respect to their order of elution from a Mono Q column (8). The lignin peroxidases catalyze the one-electron oxidation of a number of different substrates. Oneelectron oxidation of cyanide to cyanyl radical has been detected by spin trapping in chemical (9) and electrochemical (10-12) oxidizing systems. In addition, a number of peroxidases also catalyze the one-electron oxidation of cyanide (13) and a variety of other inorganic substrates such as sulfite, nitrite, and azide (14-16). With this in mind, we investigated the metabolism of cyanide by P. chrysosporium and its oxidation by lignin peroxidase H2 (Lip H2). In this communication, we show that lignin peroxidase H2 oxidizes cyanide to the cyanyl radical in the presence of Hz02 and that lignolytic cultures of P. chrysosporium mineralized cyanide to carbon dioxide. MATERIALS
1 Supported in part by NIH Grant ES04922. * To whom correspondence should be addressed.
0003.9861/91 $3.00 Copyright 0 1991 by Academic Press, Al1 rights of reproduction in any form
Inc. reserved.
AND
METHODS
Fungus. P. chrysosporium (BKM F-1767) was obtained from the United States Department of Agriculture, Forest Products Laboratory
173
174
SHAH,
GROVER,
(Madison, WI). The microorganism was maintained on malt agar slant cultures at room temperature and was subcultured every 30 to 60 days. Chemicals. Hydrogen peroxide and [“CIKCN (51.2 mCi/mmol) were obtained from Sigma (St. Louis, MO). Veratryl alcohol and N-tert-butyla-phenylnitrone (PBN)3 and [i3C]KCN were purchased from Aldrich (Milwaukee, WI), sodium cyanide and potassium cyanide were purchased from Mallinckrodt (Paris, KY), and [U-i4C]glucose (1.88 mCi/mmol) was from Pathfinder Laboratories, Inc. (St. Louis, MO). Sodium phosphate buffer and sodium tartrate buffer were prepared in purified water (Barnstead Nanopure II system; specific resistance, 18.0 Mohm-cm-‘). Enzyme production andpurification. Culture conditions for the production of lignin peroxidases were as previously described (17). The concentrated extracellular fluid was dialyzed overnight against 10 mM sodium acetate buffer, pH 6.0, and the proteins were purified on a Mono Q HR 5/5 column (Pharmacia, Uppsala, Sweden) as previously described (17). The major lignin peroxidase, H2 (17), was then analyzed for purity (99%) by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and was used throughout these studies. Enzyme assay. Veratryl alcohol oxidase activity was determined by following the formation veratryl aldehyde measured at 310 nm. Reaction mixtures (1 ml) contained 100 pM sodium tartrate buffer, pH 3.5, 0.1 +M LIP H2, 500 pM Hz02, and 1.5 mM veratryl alcohol. Reactions were carried out in a closed system at 10 f 2°C. Cyanide solution was freshly prepared and was kept at 4°C. Determination of apparent dissociation constant (Kd) for cyanide. Sample and reference cuvettes contained 2.6 +M LIP H2 and 0.1 M sodium tartrate buffer, pH 3.5. Difference spectra were obtained by addition of cyanide in the sample cuvette. The difference in maxima and minima peak absorbance (approximately Ako9 minus A423nm) were measured at each concentration of sodium cyanide. Effect of cyanide on inactivation of lignin peroxidase by Hz02. We incubated 1 pM LiP H2 with 500 pM Hz02 in 0.1 M sodium tartrate buffer, pH 3.5, and various concentrations of sodium cyanide. After 5 min of incubation, aliquots (20 11 from a l-ml incubation) were removed and assayed for veratryl alcohol oxidase activity as described above. Spin trapping of cyanyl radical. Reaction mixtures contained 0.6 mg/ ml, 50 mM sodium cyanide, 100 mM PBN, and 500 PM HzOz in 200 mM in sodium phosphate buffer, pH 6.5. Reaction mixtures were transferred to the ESR sample cavity within 1 min following the initiation of reaction with H,Os. Spectra were recorded at room temperature using a Varian E-109 spectrometer operating at 9.5 GHz with 100 kHz field modulation. A scan range of 100 G, modulation amplitude of 1.0 G, and micowave power of 50 mW were used for all spectra. Hyperfine splitting constants were determined by comparison with the standard 2,2,6,6-tetramethyl4-hydroxypiperidine-1-oxyl (Tempol) using 17.1 G for aN in water. Toxicity of sodium cyanide. The toxicity of cyanide to the fungus was estimated by determining its effect on glucose metabolism. The fungal culture medium contained 56 mM glucose, 1.2 mM ammonium tartrate, trace metals (18), and thiamine (1 mg) in 20 mM 2,2’-dimethyl sodium succinate buffer, pH 4.2. The culture medium was sterilized by filtration through a cellulose acetate membrane filter (pore size, 0.22 FM). Culture bottles were sterilized by autoclaving at 121°C and 15 lb/ in.* for 20 min. Samples of the culture medium (9 ml) were dispensed into 250.ml Wheaton bottles equipped with gas-exchange manifolds equipped with Teflon seals (4). Cultures were inoculated with a spore suspension of P. chrysosporium (1 ml; Asso = 0.5) and grown at room temperature. Control cultures contained culture medium minus P. chrysosporium. To measure the toxicity of cyanide to spores, NaCN and [i4C]glucose (0.1 &i) were added on Day 0, and 3 days later the headspaces were flushed with oxygen (99%) and the liberated [“C]CO, was trapped in 10 ml of a solution containing ethanolamine-methanol-safety
s Abbreviations peroxidase.
used: PBN, N-tert-butyl-a-phenylnitrone;
LIP, lignin
AND
AUST
solve scintillation cocktail (Research Products International Corp., Mt. Prospect, IL) (1:4:5). The amount of [i4C]C0, trapped was determined by scintillation spectrometry (Beckman LS-5801). The toxicity of NaCN to mature, ligninolytic cultures of P. chrysosporium was measured as described for spores, except the fungus was allowed to grow for 6 days and the culture bottles were flushed with pure oxygen on Days 3 and 6. The culture filtrate contained 95 units of LiP activity (a unit is 1 pmol of veratryl alcohol oxidized per minute). Sodium cyanide and [“Clglucose were added to the 6-day-old cultures and 3 days later the amount of [i4C]C02 was determined as detailed above. Mineralization of cyanide by P. chrysosporium. Cultures of P. chrysosporium were started as described above. On Day 6, various concentrations of [i4C]KCN were added to the cultures. Three days later the headspaces were flushed with oxygen (99%) and the liberated COP was trapped in 1 M Ba(OH),. The radioactivity of [i4C]BaC0s precipitate was quantitated by scintillation spectrometry after the removal of [i4C]Ba(CN), in the supernatant by centrifugation.
RESULTS
Inhibition of veratryl alcohol oxidase activity by sodium cyanide. Inhibition of the veratryl alcohol oxidase activity of lignin peroxidase H2 by sodium cyanide was found to be complex. Cyanide was found to be an effective inhibitor of initial activity (150- 2-3 PM); however, after a lag phase veratryl alcohol activity returned, dependent upon the concentration of cyanide (Fig. 1). Inhibition appeared to be competitive with respect to veratryl alcohol but again the data were difficult to analyze (Fig. 2). In part, the reversal of inhibition could be due to dissociation of cyanide from the enzyme but cyanide was also effective in protecting the enzyme from inactivation by H202 (Fig. 3). A total of 500 PM H202 inactivated about 50% of the enzyme in 5 min. However, the addition of 8 mM cyanide completely protected the enzyme from inactivation. The
0 @.4Cyanide
0.0
-
aptd Cyanide
-
130 @4 Cyanide
-
345 @4 Cyanide
0.2
0.4
0.6
0.8
1.0
Time(min) FIG. 1. Effect of cyanide on veratryl alcohol oxidase activity of lignin peroxidase. Reaction mixtures contained 0.1 PM lignin peroxidase H2, 0.1 M sodium tartrate buffer, pH 3.5, 500 PM HxOr, 1.5 mM veratryl alcohol, and various concentration of cyanide.
METABOLISM
OF CYANIDE
BY Phcznerochaete chrysosporium
0.6 ,
175
I
25 pM veratryi alcohol 35 JJM veratryi aicohol 4.5 gM veratyl alcohol 75 pM veratryl alcohol 150 bM verauyi eJcohol 250 !M verzry
0.0
0.8
0.4
Time
1.6
1.2
alcohol
2.0
(min)
FIG. 2. Effect of veratryl alcohol on its oxidation by lignin peroxidase in the presence of NaCN. Reaction mixtures contained of veratryl alcohol as shown. 0.2 pM lignin peroxidase H2, 0.1 M sodium tartrate buffer, pH 3.5, and various concentrations
Kd for the binding of cyanide to the resting enzyme, measured by difference spectrometry, was found to be approximately 10 PM (Fig. 4). Generation of cyanyl radical was demonstrated in reaction mixtures containing LiP H2, cyanide, and HzOz using ESR spectroscopy and the spin trap PBN (Fig. 5).
110
1
150
pM
NaCN,
No spin adducts were observed when cyanide was eliminated from the reaction mixture. The ESR spectrum consisted of a triplet of doublets with hyperfine constants aN = 16.0 G and a; = 2.4 G. When [13C]KCN was substituted for KCN, the resulting spectrum consisted of 12 lines (aN = 16.0 G, a13CB= 10.7 G, and aSH = 2.4 G). Figure of cyanide by P. chrysosporium. Mineralization 6 demonstrates that cyanide underwent rapid and exten1.2
100 90 : 5 F
s: s
80
70 60 50
i
40 0
2
4
6
a .Ol
[CYANIDE]
.l
1
10
100
1000
mM
FIG. 3. Protection of lignin peroxidase by cyanide from inactivation by HZ02. Reaction incubations (1 ml) contained 1 pM lignin peroxidase H2, 0.1 M sodium tartrate buffer (pH 3.51, 500 pM HzOz, and various concentration of cyanide. After 5 min of incubation, 20-~1 aliquots were taken from each incubation and veratryl alcohol oxidase activity was determined as described under Materials and Methods. The activity before the addition of H,OB was considered 100% activity. Each incubation was conducted in triplicate and the average values were plotted with standard deviations.
[CYANIDE]@l
FIG. 4. Determination of the apparent dissociation constant for cyanide. Sample and reference cuvettes contained 2.6 pM lignin peroxidase H2 in 0.1 M sodium tartrate buffer, pH 3.5. Difference spectra were obtained by addition of cyanide in the sample cuvette. The difference in maxima and minima at each addition of cyanide was recorded (A,,A& and the apparent dissociation constant Kd was determined by plotting AA/AA,, vs the concentration of cyanide. The data are from an experiment conducted in duplicate.
176
SHAH, 18
GROVER,
GA”55
A
k FIG. 5. ESR spectrum of PBN-cyanyl radical adduct formed by lignin peroxidase H2 and Hz02. (A) Reaction mixtures contained 10 pM lignin peroxidase H2,40 mM sodium cyanide, 500 pM H202, and 100 mM PBN in 100 mM sodium phosphate buffer, pH 6.0. (B) [‘%]KCN was substituted for NaCN. The receiver gain was 4 X lo4 in (A) and 2.5 X lo4 in (B).
sive mineralization by ligninolytic cultures of P. chrysosporium. At an initial concentration of 20 PM, 45% of the cyanide was mineralized to CO2 in 3 days. The rate of cyanide mineralization increased with increasing cyanide concentration up to 2 FM. The percentage of the cyanide mineralized dropped to about 5% at 10 mM but the absolute amount mineralized remained about the same as at 2 mM. Toxicity of NaCN to P. chrysosporium. Figures 7 and 8 show the effect of cyanide on fungal respiration (as measured by metabolism of [14C]glucose to [14C]C0,) for cultures started with spores and (j-day-old cultures of fungus, respectively. It can be seen that 50% inhibition of glucose metabolism occurred at approximately 100 /*M for cultures started with spores and at approximately 5 mM cyanide for B-day-old cultures of fungus. DISCUSSION
As expected, cyanide was found to be a rather effective inhibitor of veratryl alcohol oxidase activity catalyzed by lignin peroxidase H2. However, it was also evident that cyanide was oxidized by this enzyme. Veratryl alcohol oxidase activity, initially inhibited by cyanide, returned after several minutes of incubation at room temperature. Inhibition appeared to be competitive with respect to ver-
AND
AUST
atryl alcohol, although it was impossible to determine the type of inhibition. Cyanide has also been considered to be a competitive inhibitor of the oxidation of nitroalkane by horseradish peroxidase (19). Cyanide was effective in protecting LiP H2 from inactivation by H202. It has been reported that reducing substrates protect peroxidases from HzO,-based inactivation (20). About 8 mM cyanide was required to protect LiP H2 from inactivation by 500 pM H202. This is much higher than the amount of cyanide required for inhibition of veratryl alcohol oxidase activity (&N - 2-3 PM). The formation of cyanyl radical from cyanide by LiP/H202 was demonstrated using 13C-labeled cyanide. The spin trapping of the cyanyl radical indicates that cyanide might be a substrate for lignin peroxidase, since free radical formation is the hallmark of peroxidasecatalyzed substrate oxidation. The hyperfine coupling constants agree well with those reported by Moreno et al. (13) for the PBN-cyanyl adduct formed during the oxidation of cyanide by horseradish peroxide, lactoperoxidase, chloroperoxidase, NADH peroxidase, or methemoglobin in the presence of H202. Stolze et al. used tertnitrosobutane to spin trap cyanyl radical produced by horseradish peroxidases/HzOz at a pH above the pK, of hydrocyanic acid (pK, 9.3) and suggested the formation of the formamide radical on hydrolysis of the cyanyl radical (24). Cyanide also inhibited the oxidation of dihydrofumaric acid by Japanese radish peroxidases and the inhibition was dependent on pH and concentration of cyanide (16). All of these results suggest that cyanide is a reductant for several peroxidases. Our studies of cyanide metabolism by P. chrysosporium also suggested that cyanide was rapidly mineralized by the fungus. The amount of cyanide mineralized increased with increasing concentration of cyanide to about 2 mM. Ten mM cyanide was mineralized at almost the same rate as was 2 mM. Thus the metabolism system must have been saturated somewhere between 2 and 10 mM cyanide or toxicity limited the rate of mineralization at 10 mM cyanide. The latter may be possible, as 10 mM cyanide inhibited respiration (metabolism of [14C]glucose to [i4C]C0,) by approximately 70%. However, apparently the rapid rate of metabolism of cyanide provides considerable protection for the fungus against toxicity, as 6-day-old ligninolytic cultures were much less sensitive to inhibition of respiration by cyanide - 5 mM) than were cultures started from spores it: - 0.1 mM). It has been reported that fungus Fusarium solani could oxidize cyanide to ammonia and CO:! (21). Several species of fungi that attack cyanogenic plants can convert cyanide to formamide (22). In neither case was the mechanism of cyanide oxidation elucidated. It has been reported that when cyanide is used as an inhibitor of myeloperoxidase it also acts as a substrate for chlorination (23). Our results indicate that cyanide can be oxidized by LiP H2 and cul-
METABOLISM
OF CYANIDE
BY Phanerochoete
t 10
1 50 -
40-
177
chrysosporium
?i 0 5 s = e i E ,I
-
30 -
z C 3
20 -
'ii
10 -
I E a.
[KCNI
mM
FIG. 6. Mineralization of cyanide by 6-day-old cultures of P. chrysosporium. [14C]KCN was added to g-day-old cultures of P. chryaosporium; 3 Radioactivity of days later evolved [i4C]C0, was trapped using 1 M Ba(OH)z. [i4C]BaC0 3 was separated from [i4C]Ba(CN), by centrifugation. [“C]BaCOa was quantitated using liquid scintillation spectrometry. All incubations were carried out in quadruplicate in a closed system at room temperature and in stationary cultures. Open circles show the percentage of the cyanide mineralized in 3 days and squares show total micromoles of cyanide mineralized. The error bars (standard deviations) are within the data points in some cases.
tures of P. chrysosporium and that this may result in considerable protection of the fungus from the toxicity of cyanide. This may be important if this fungus were to be used for bioremediation of pollution sites that may also
contain cyanide. The mineralization of various cyanide salts, both alone and in the presence of other pollutants, such as polycyclic aromatic hydrocarbons, is currently under investigation.
80
60
20 [Cyanide]mM
FIG. 7. Toxicity of cyanide to spores of I’. chrysosporium. Sodium cyanide and [‘4C]glucose were added to cultures of P. chrysosporium at Day 0, and 3 days later evolved [“C]CO, was trapped and its radioactivity quantitated using liquid scintillation spectrometry. The controls were without cyanide. Glucose metabolism in the controls was considered 100%. All incubations were carried out in quadruplicate in a closed system at room temperature and in stationary cultures. The data are average values with standard deviations, some of which are within the data points.
40
60
[Cyanide]mM
FIG. 8. Toxicity of cyanide to B-day-old cultures of I’. chrysosporium. Sodium cyanide and [‘4C]glucose were added to B-day-old ligninolytic cultures of P. chrysosporium, and 3 days later the evolved CO, was trapped and its radioactivity quantitated using liquid scintillation spectrometry. The controls were without cyanide. Glucose metabolism in the control was considered 100%. All incubations were carried out in quadruplicate in a closed system at room temperature and in stationary cultures. The data are average values with standard deviations, some of which are within the data points.
178
SHAH, GROVER, AND AUST
REFERENCES 1. Knowles, C. J. (1976) Bacterial. Reu. 40, 652-680. 2. Smith, S. E. (1972) J. Water Pollut. Control Fed. 44, 1100-1104. 3. Kirk, T. K., Schultz, E., Connors, W. J., Lorenz, C. F., and Zeikus, J. G. (1978) Arch. Microbial. 117, 277-285. 4. Bumpus, J. A., Tien, M., Wright, D., and Aust, S. D. (1985) Science
228,1434-1436. 5. Hammerli, S. D., Leisola, M. S. A., Sanglard, D., and Fiechter, A. (1986) J. Biol. Chem. 261, 6900-6903. 6. Hammel, K. E., Kalyanaraman,
B., and Kirk, T. K. (1986) J. Biol.
Chem. 261,16,948-16,952. 7. Sanglard, D., Leisola, M. S. A., and Fiechter, A. (1986) Enzyme Microb. Technol. 8, 209-212. 8. Kirk, T. K., Croan, S. C., Tien, M., Murtagh, K. E., and Farrel, R. (1985) Enzyme Microb. Tech&. 8, 27-32. 9. Chawla, 0. P., and Fessenden, R. W. (1975) J. Phys. Chem. 79, 2693-2700. 10. Walter, T. Y. H., Bancroft, E. E., McIntire, G. L., Davis, E. R., Gierasch, L. M., Blount, H. N., Stronks, H. J., and Janzen, E. G. (1982) Can. J. Chem. 60, 1621-1636. 11. Janzen, E. G., Stronks, H. J., Nutter Jr., D. E., Davis, E. R., Blount, H. N., Poyer, J. L., and McCay, P. B. (1980) Con. J. Chem. 58, 1596-1598. 12. Janzen, E. G., and Stronks, H. J. (1981) J. Phys. Chem. 85,39523954.
13. Moreno, S. N. J., Stolze, K., Janzen, E. G., and Mason, R. P. (1988) Arch. Biochem. Biophys. 265, 267-271.
14. Yamazaki, I., and Piette, L. H. (1963) 47-64. 15. Roman, R., and Dunford, H. B. (1973) 16. Kalayanraman, B., Janzen, E. G., and &em. 260,4003-4006. 17. Tuisel, H., Sinclair, R., Bumpus, J. A.,
B&him.
Biophys. Acta 77,
Can. J. Chem. 51,588-596. Mason, R. P. (1985) J. Biol.
Ashbaugh, W., Brock, B. J., and Aust, S. D. (1990) Arch. Biochem. Biophys. 279, 158-166.
18. Kirk, T. K., Schultz, E., Connors, W. J., Lorenz, L. F., and Zeikus, J. G. (1978) Arch. Microbial. 117, 277-285. 19. Porter, D. J., and Bright, 9924.
H. J. (1983) J. Biol. Chem. 258, 9913-
20. Arno, M. B., Acosta, M., de1 Rio, J. A., and Garcia-Canora, Biochim. Biophys. Acta 1038, 85-89.
F. (1990)
21. Shimizu, T., and Taguchi, H. (1969) J. Ferment. Technol. 47,639643. 22. Fry, W. E., and Myers, D. F. (1981) in Cyanide in Biology (Vennesland, B., Conn, E. E., Knowles, C. J., Westley, J., and Wissing, F., Eds.), pp. 321-324, Academic Press, New York.
23. Stelkmaszynska, T. (1986) Znt. J. Biochem. 18, 1107-1114. 24. Stolze, K., Moreno-Silva, N. J., and Mason, R. P. (1989) J. Znorg. Chem. 37,45-53.
OF BIOCHEMISTRY
AND
BIOPHYSICS
Vol. 290, No. 1, October, pp. 173-178, 1991
Metabolism of Cyanide by Phanerochaete chrysosporium’ Manish
M. Shah, Thomas
Biotechnology
Center,
Received February
Utah
A. Grover,
State
University,
and Steven Logan,
Utah
D. Aust’
84322-4700
21, 1991, and in revised form June 16, 1991
The oxidation of veratryl alcohol (3,4-dimethoxybenzyl alcohol) by lignin peroxidase H2 (Lip H2) from the white rot fungus Phanerochaete chrysosporium was strongly inhibited by sodium cyanide. The IeO was estimated to be about 2-3 PM. In contrast, sodium cyanide binds to the native enzyme with an apparent sodium cyanide dissociation constant Kd of about 10 jtM. Inhibition of the veratryl alcohol oxidase activity of LiP H2 by cyanide was reversible. Ligninolytic cultures of P. chrysosporium mineralized cyanide at a rate that was proportional to the concentration of cyanide to 2 mM. The N-tert-butyl-a-phenylnitrone-cyanyl radical adduct was observed by ESR spin trapping upon incubation of LiP H2 with HzOz and sodium cyanide. The identity of the spin adduct was confirmed using 13C-labeled cyanide. Six-day-old cultures of the fungus were more tolerant to sodium cyanide toxicity than spores. Toxicity measurements were based on the effect of sodium cyanide on respiration of the fungus as determined by the metabolism of [ “C]glucose to [‘4C]C02. We propose that this tolerance of the mature fungus was due to its ability to mineralize cyanide and that this fungus might be effective in treating environmental pollution sites contaminated with cyanide. 0 1991
Academic
Press,
Inc.
Cyanides have been designated hazardous substances and priority toxic pollutants by the EPA. Discharge of cyanide into the environment is limited by its extreme toxicity to fish rather than man and microorganisms. The electroplating, steel, carbonization, and other important industries produce large quantities of cyanide wastes. To prevent damage to natural ecosystems, the cyanide content of these effluents must be reduced to essentially zero before discharge into the environment. Even for discharge
into a sewer there must be a reduction to no more than 1 to 2 pg of cyanide per milliliter (1). Conventional chemical methods of disposal of cyanide wastes are wasteful of resources, since other chemicals, such as chlorine, are required for detoxification of the cyanide (2). In addition, chemical degradation is relatively expensive and often requires further disposal of the products. The wood rotting fungus Phanerochaete chrysosporium possesses remarkable biodegradative properties. This fungus is one of the relatively few microorganisms known to degrade lignin to carbon dioxide (3). P. chrysosporium is also able to degrade a wide variety of environmentally persistent xenobiotics such as DDT, lindane, benzo[a]pyrene, and polychlorinated biphenyls to carbon dioxide (4). Recent evidence demonstrates that the lignin degrading system is responsible, at least in part, for the ability of P. chrysosporium to degrade xenobiotics (5-7). P. chrysosporium secretes a number of peroxidases; over 10 have been detected in extracellular fluid of lignolytic cultures (8). Kirk et al. have named these proteins Hl to HlO in respect to their order of elution from a Mono Q column (8). The lignin peroxidases catalyze the one-electron oxidation of a number of different substrates. Oneelectron oxidation of cyanide to cyanyl radical has been detected by spin trapping in chemical (9) and electrochemical (10-12) oxidizing systems. In addition, a number of peroxidases also catalyze the one-electron oxidation of cyanide (13) and a variety of other inorganic substrates such as sulfite, nitrite, and azide (14-16). With this in mind, we investigated the metabolism of cyanide by P. chrysosporium and its oxidation by lignin peroxidase H2 (Lip H2). In this communication, we show that lignin peroxidase H2 oxidizes cyanide to the cyanyl radical in the presence of Hz02 and that lignolytic cultures of P. chrysosporium mineralized cyanide to carbon dioxide. MATERIALS
1 Supported in part by NIH Grant ES04922. * To whom correspondence should be addressed.
0003.9861/91 $3.00 Copyright 0 1991 by Academic Press, Al1 rights of reproduction in any form
Inc. reserved.
AND
METHODS
Fungus. P. chrysosporium (BKM F-1767) was obtained from the United States Department of Agriculture, Forest Products Laboratory
173
174
SHAH,
GROVER,
(Madison, WI). The microorganism was maintained on malt agar slant cultures at room temperature and was subcultured every 30 to 60 days. Chemicals. Hydrogen peroxide and [“CIKCN (51.2 mCi/mmol) were obtained from Sigma (St. Louis, MO). Veratryl alcohol and N-tert-butyla-phenylnitrone (PBN)3 and [i3C]KCN were purchased from Aldrich (Milwaukee, WI), sodium cyanide and potassium cyanide were purchased from Mallinckrodt (Paris, KY), and [U-i4C]glucose (1.88 mCi/mmol) was from Pathfinder Laboratories, Inc. (St. Louis, MO). Sodium phosphate buffer and sodium tartrate buffer were prepared in purified water (Barnstead Nanopure II system; specific resistance, 18.0 Mohm-cm-‘). Enzyme production andpurification. Culture conditions for the production of lignin peroxidases were as previously described (17). The concentrated extracellular fluid was dialyzed overnight against 10 mM sodium acetate buffer, pH 6.0, and the proteins were purified on a Mono Q HR 5/5 column (Pharmacia, Uppsala, Sweden) as previously described (17). The major lignin peroxidase, H2 (17), was then analyzed for purity (99%) by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and was used throughout these studies. Enzyme assay. Veratryl alcohol oxidase activity was determined by following the formation veratryl aldehyde measured at 310 nm. Reaction mixtures (1 ml) contained 100 pM sodium tartrate buffer, pH 3.5, 0.1 +M LIP H2, 500 pM Hz02, and 1.5 mM veratryl alcohol. Reactions were carried out in a closed system at 10 f 2°C. Cyanide solution was freshly prepared and was kept at 4°C. Determination of apparent dissociation constant (Kd) for cyanide. Sample and reference cuvettes contained 2.6 +M LIP H2 and 0.1 M sodium tartrate buffer, pH 3.5. Difference spectra were obtained by addition of cyanide in the sample cuvette. The difference in maxima and minima peak absorbance (approximately Ako9 minus A423nm) were measured at each concentration of sodium cyanide. Effect of cyanide on inactivation of lignin peroxidase by Hz02. We incubated 1 pM LiP H2 with 500 pM Hz02 in 0.1 M sodium tartrate buffer, pH 3.5, and various concentrations of sodium cyanide. After 5 min of incubation, aliquots (20 11 from a l-ml incubation) were removed and assayed for veratryl alcohol oxidase activity as described above. Spin trapping of cyanyl radical. Reaction mixtures contained 0.6 mg/ ml, 50 mM sodium cyanide, 100 mM PBN, and 500 PM HzOz in 200 mM in sodium phosphate buffer, pH 6.5. Reaction mixtures were transferred to the ESR sample cavity within 1 min following the initiation of reaction with H,Os. Spectra were recorded at room temperature using a Varian E-109 spectrometer operating at 9.5 GHz with 100 kHz field modulation. A scan range of 100 G, modulation amplitude of 1.0 G, and micowave power of 50 mW were used for all spectra. Hyperfine splitting constants were determined by comparison with the standard 2,2,6,6-tetramethyl4-hydroxypiperidine-1-oxyl (Tempol) using 17.1 G for aN in water. Toxicity of sodium cyanide. The toxicity of cyanide to the fungus was estimated by determining its effect on glucose metabolism. The fungal culture medium contained 56 mM glucose, 1.2 mM ammonium tartrate, trace metals (18), and thiamine (1 mg) in 20 mM 2,2’-dimethyl sodium succinate buffer, pH 4.2. The culture medium was sterilized by filtration through a cellulose acetate membrane filter (pore size, 0.22 FM). Culture bottles were sterilized by autoclaving at 121°C and 15 lb/ in.* for 20 min. Samples of the culture medium (9 ml) were dispensed into 250.ml Wheaton bottles equipped with gas-exchange manifolds equipped with Teflon seals (4). Cultures were inoculated with a spore suspension of P. chrysosporium (1 ml; Asso = 0.5) and grown at room temperature. Control cultures contained culture medium minus P. chrysosporium. To measure the toxicity of cyanide to spores, NaCN and [i4C]glucose (0.1 &i) were added on Day 0, and 3 days later the headspaces were flushed with oxygen (99%) and the liberated [“C]CO, was trapped in 10 ml of a solution containing ethanolamine-methanol-safety
s Abbreviations peroxidase.
used: PBN, N-tert-butyl-a-phenylnitrone;
LIP, lignin
AND
AUST
solve scintillation cocktail (Research Products International Corp., Mt. Prospect, IL) (1:4:5). The amount of [i4C]C0, trapped was determined by scintillation spectrometry (Beckman LS-5801). The toxicity of NaCN to mature, ligninolytic cultures of P. chrysosporium was measured as described for spores, except the fungus was allowed to grow for 6 days and the culture bottles were flushed with pure oxygen on Days 3 and 6. The culture filtrate contained 95 units of LiP activity (a unit is 1 pmol of veratryl alcohol oxidized per minute). Sodium cyanide and [“Clglucose were added to the 6-day-old cultures and 3 days later the amount of [i4C]C02 was determined as detailed above. Mineralization of cyanide by P. chrysosporium. Cultures of P. chrysosporium were started as described above. On Day 6, various concentrations of [i4C]KCN were added to the cultures. Three days later the headspaces were flushed with oxygen (99%) and the liberated COP was trapped in 1 M Ba(OH),. The radioactivity of [i4C]BaC0s precipitate was quantitated by scintillation spectrometry after the removal of [i4C]Ba(CN), in the supernatant by centrifugation.
RESULTS
Inhibition of veratryl alcohol oxidase activity by sodium cyanide. Inhibition of the veratryl alcohol oxidase activity of lignin peroxidase H2 by sodium cyanide was found to be complex. Cyanide was found to be an effective inhibitor of initial activity (150- 2-3 PM); however, after a lag phase veratryl alcohol activity returned, dependent upon the concentration of cyanide (Fig. 1). Inhibition appeared to be competitive with respect to veratryl alcohol but again the data were difficult to analyze (Fig. 2). In part, the reversal of inhibition could be due to dissociation of cyanide from the enzyme but cyanide was also effective in protecting the enzyme from inactivation by H202 (Fig. 3). A total of 500 PM H202 inactivated about 50% of the enzyme in 5 min. However, the addition of 8 mM cyanide completely protected the enzyme from inactivation. The
0 @.4Cyanide
0.0
-
aptd Cyanide
-
130 @4 Cyanide
-
345 @4 Cyanide
0.2
0.4
0.6
0.8
1.0
Time(min) FIG. 1. Effect of cyanide on veratryl alcohol oxidase activity of lignin peroxidase. Reaction mixtures contained 0.1 PM lignin peroxidase H2, 0.1 M sodium tartrate buffer, pH 3.5, 500 PM HxOr, 1.5 mM veratryl alcohol, and various concentration of cyanide.
METABOLISM
OF CYANIDE
BY Phcznerochaete chrysosporium
0.6 ,
175
I
25 pM veratryi alcohol 35 JJM veratryi aicohol 4.5 gM veratyl alcohol 75 pM veratryl alcohol 150 bM verauyi eJcohol 250 !M verzry
0.0
0.8
0.4
Time
1.6
1.2
alcohol
2.0
(min)
FIG. 2. Effect of veratryl alcohol on its oxidation by lignin peroxidase in the presence of NaCN. Reaction mixtures contained of veratryl alcohol as shown. 0.2 pM lignin peroxidase H2, 0.1 M sodium tartrate buffer, pH 3.5, and various concentrations
Kd for the binding of cyanide to the resting enzyme, measured by difference spectrometry, was found to be approximately 10 PM (Fig. 4). Generation of cyanyl radical was demonstrated in reaction mixtures containing LiP H2, cyanide, and HzOz using ESR spectroscopy and the spin trap PBN (Fig. 5).
110
1
150
pM
NaCN,
No spin adducts were observed when cyanide was eliminated from the reaction mixture. The ESR spectrum consisted of a triplet of doublets with hyperfine constants aN = 16.0 G and a; = 2.4 G. When [13C]KCN was substituted for KCN, the resulting spectrum consisted of 12 lines (aN = 16.0 G, a13CB= 10.7 G, and aSH = 2.4 G). Figure of cyanide by P. chrysosporium. Mineralization 6 demonstrates that cyanide underwent rapid and exten1.2
100 90 : 5 F
s: s
80
70 60 50
i
40 0
2
4
6
a .Ol
[CYANIDE]
.l
1
10
100
1000
mM
FIG. 3. Protection of lignin peroxidase by cyanide from inactivation by HZ02. Reaction incubations (1 ml) contained 1 pM lignin peroxidase H2, 0.1 M sodium tartrate buffer (pH 3.51, 500 pM HzOz, and various concentration of cyanide. After 5 min of incubation, 20-~1 aliquots were taken from each incubation and veratryl alcohol oxidase activity was determined as described under Materials and Methods. The activity before the addition of H,OB was considered 100% activity. Each incubation was conducted in triplicate and the average values were plotted with standard deviations.
[CYANIDE]@l
FIG. 4. Determination of the apparent dissociation constant for cyanide. Sample and reference cuvettes contained 2.6 pM lignin peroxidase H2 in 0.1 M sodium tartrate buffer, pH 3.5. Difference spectra were obtained by addition of cyanide in the sample cuvette. The difference in maxima and minima at each addition of cyanide was recorded (A,,A& and the apparent dissociation constant Kd was determined by plotting AA/AA,, vs the concentration of cyanide. The data are from an experiment conducted in duplicate.
176
SHAH, 18
GROVER,
GA”55
A
k FIG. 5. ESR spectrum of PBN-cyanyl radical adduct formed by lignin peroxidase H2 and Hz02. (A) Reaction mixtures contained 10 pM lignin peroxidase H2,40 mM sodium cyanide, 500 pM H202, and 100 mM PBN in 100 mM sodium phosphate buffer, pH 6.0. (B) [‘%]KCN was substituted for NaCN. The receiver gain was 4 X lo4 in (A) and 2.5 X lo4 in (B).
sive mineralization by ligninolytic cultures of P. chrysosporium. At an initial concentration of 20 PM, 45% of the cyanide was mineralized to CO2 in 3 days. The rate of cyanide mineralization increased with increasing cyanide concentration up to 2 FM. The percentage of the cyanide mineralized dropped to about 5% at 10 mM but the absolute amount mineralized remained about the same as at 2 mM. Toxicity of NaCN to P. chrysosporium. Figures 7 and 8 show the effect of cyanide on fungal respiration (as measured by metabolism of [14C]glucose to [14C]C0,) for cultures started with spores and (j-day-old cultures of fungus, respectively. It can be seen that 50% inhibition of glucose metabolism occurred at approximately 100 /*M for cultures started with spores and at approximately 5 mM cyanide for B-day-old cultures of fungus. DISCUSSION
As expected, cyanide was found to be a rather effective inhibitor of veratryl alcohol oxidase activity catalyzed by lignin peroxidase H2. However, it was also evident that cyanide was oxidized by this enzyme. Veratryl alcohol oxidase activity, initially inhibited by cyanide, returned after several minutes of incubation at room temperature. Inhibition appeared to be competitive with respect to ver-
AND
AUST
atryl alcohol, although it was impossible to determine the type of inhibition. Cyanide has also been considered to be a competitive inhibitor of the oxidation of nitroalkane by horseradish peroxidase (19). Cyanide was effective in protecting LiP H2 from inactivation by H202. It has been reported that reducing substrates protect peroxidases from HzO,-based inactivation (20). About 8 mM cyanide was required to protect LiP H2 from inactivation by 500 pM H202. This is much higher than the amount of cyanide required for inhibition of veratryl alcohol oxidase activity (&N - 2-3 PM). The formation of cyanyl radical from cyanide by LiP/H202 was demonstrated using 13C-labeled cyanide. The spin trapping of the cyanyl radical indicates that cyanide might be a substrate for lignin peroxidase, since free radical formation is the hallmark of peroxidasecatalyzed substrate oxidation. The hyperfine coupling constants agree well with those reported by Moreno et al. (13) for the PBN-cyanyl adduct formed during the oxidation of cyanide by horseradish peroxide, lactoperoxidase, chloroperoxidase, NADH peroxidase, or methemoglobin in the presence of H202. Stolze et al. used tertnitrosobutane to spin trap cyanyl radical produced by horseradish peroxidases/HzOz at a pH above the pK, of hydrocyanic acid (pK, 9.3) and suggested the formation of the formamide radical on hydrolysis of the cyanyl radical (24). Cyanide also inhibited the oxidation of dihydrofumaric acid by Japanese radish peroxidases and the inhibition was dependent on pH and concentration of cyanide (16). All of these results suggest that cyanide is a reductant for several peroxidases. Our studies of cyanide metabolism by P. chrysosporium also suggested that cyanide was rapidly mineralized by the fungus. The amount of cyanide mineralized increased with increasing concentration of cyanide to about 2 mM. Ten mM cyanide was mineralized at almost the same rate as was 2 mM. Thus the metabolism system must have been saturated somewhere between 2 and 10 mM cyanide or toxicity limited the rate of mineralization at 10 mM cyanide. The latter may be possible, as 10 mM cyanide inhibited respiration (metabolism of [14C]glucose to [i4C]C0,) by approximately 70%. However, apparently the rapid rate of metabolism of cyanide provides considerable protection for the fungus against toxicity, as 6-day-old ligninolytic cultures were much less sensitive to inhibition of respiration by cyanide - 5 mM) than were cultures started from spores it: - 0.1 mM). It has been reported that fungus Fusarium solani could oxidize cyanide to ammonia and CO:! (21). Several species of fungi that attack cyanogenic plants can convert cyanide to formamide (22). In neither case was the mechanism of cyanide oxidation elucidated. It has been reported that when cyanide is used as an inhibitor of myeloperoxidase it also acts as a substrate for chlorination (23). Our results indicate that cyanide can be oxidized by LiP H2 and cul-
METABOLISM
OF CYANIDE
BY Phanerochoete
t 10
1 50 -
40-
177
chrysosporium
?i 0 5 s = e i E ,I
-
30 -
z C 3
20 -
'ii
10 -
I E a.
[KCNI
mM
FIG. 6. Mineralization of cyanide by 6-day-old cultures of P. chrysosporium. [14C]KCN was added to g-day-old cultures of P. chryaosporium; 3 Radioactivity of days later evolved [i4C]C0, was trapped using 1 M Ba(OH)z. [i4C]BaC0 3 was separated from [i4C]Ba(CN), by centrifugation. [“C]BaCOa was quantitated using liquid scintillation spectrometry. All incubations were carried out in quadruplicate in a closed system at room temperature and in stationary cultures. Open circles show the percentage of the cyanide mineralized in 3 days and squares show total micromoles of cyanide mineralized. The error bars (standard deviations) are within the data points in some cases.
tures of P. chrysosporium and that this may result in considerable protection of the fungus from the toxicity of cyanide. This may be important if this fungus were to be used for bioremediation of pollution sites that may also
contain cyanide. The mineralization of various cyanide salts, both alone and in the presence of other pollutants, such as polycyclic aromatic hydrocarbons, is currently under investigation.
80
60
20 [Cyanide]mM
FIG. 7. Toxicity of cyanide to spores of I’. chrysosporium. Sodium cyanide and [‘4C]glucose were added to cultures of P. chrysosporium at Day 0, and 3 days later evolved [“C]CO, was trapped and its radioactivity quantitated using liquid scintillation spectrometry. The controls were without cyanide. Glucose metabolism in the controls was considered 100%. All incubations were carried out in quadruplicate in a closed system at room temperature and in stationary cultures. The data are average values with standard deviations, some of which are within the data points.
40
60
[Cyanide]mM
FIG. 8. Toxicity of cyanide to B-day-old cultures of I’. chrysosporium. Sodium cyanide and [‘4C]glucose were added to B-day-old ligninolytic cultures of P. chrysosporium, and 3 days later the evolved CO, was trapped and its radioactivity quantitated using liquid scintillation spectrometry. The controls were without cyanide. Glucose metabolism in the control was considered 100%. All incubations were carried out in quadruplicate in a closed system at room temperature and in stationary cultures. The data are average values with standard deviations, some of which are within the data points.
178
SHAH, GROVER, AND AUST
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B., and Kirk, T. K. (1986) J. Biol.
Chem. 261,16,948-16,952. 7. Sanglard, D., Leisola, M. S. A., and Fiechter, A. (1986) Enzyme Microb. Technol. 8, 209-212. 8. Kirk, T. K., Croan, S. C., Tien, M., Murtagh, K. E., and Farrel, R. (1985) Enzyme Microb. Tech&. 8, 27-32. 9. Chawla, 0. P., and Fessenden, R. W. (1975) J. Phys. Chem. 79, 2693-2700. 10. Walter, T. Y. H., Bancroft, E. E., McIntire, G. L., Davis, E. R., Gierasch, L. M., Blount, H. N., Stronks, H. J., and Janzen, E. G. (1982) Can. J. Chem. 60, 1621-1636. 11. Janzen, E. G., Stronks, H. J., Nutter Jr., D. E., Davis, E. R., Blount, H. N., Poyer, J. L., and McCay, P. B. (1980) Con. J. Chem. 58, 1596-1598. 12. Janzen, E. G., and Stronks, H. J. (1981) J. Phys. Chem. 85,39523954.
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Biophys. Acta 77,
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21. Shimizu, T., and Taguchi, H. (1969) J. Ferment. Technol. 47,639643. 22. Fry, W. E., and Myers, D. F. (1981) in Cyanide in Biology (Vennesland, B., Conn, E. E., Knowles, C. J., Westley, J., and Wissing, F., Eds.), pp. 321-324, Academic Press, New York.
23. Stelkmaszynska, T. (1986) Znt. J. Biochem. 18, 1107-1114. 24. Stolze, K., Moreno-Silva, N. J., and Mason, R. P. (1989) J. Znorg. Chem. 37,45-53.
