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3 7 9 M&U
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CELL-FREE RECOVERY AND ISOTOPIC IDENTIFICATION
OF CYANIDE DEGRADING ENZYMES FROM
Pseudomonas fluorescens NCIMB 11764
DISSERTATION
Presented to the Graduate Council of the
University of North Texas in Partial
Fulfillment of the Requirements
For the Degree of
DOCTOR OF PHILOSOPHY
By
Chien-Sao Wang, B.S., M.S.
Denton, Texas
December, 1995
3 7 9 M&U
H^sro
CELL-FREE RECOVERY AND ISOTOPIC IDENTIFICATION
OF CYANIDE DEGRADING ENZYMES FROM
Pseudomonas fluorescens NCIMB 11764
DISSERTATION
Presented to the Graduate Council of the
University of North Texas in Partial
Fulfillment of the Requirements
For the Degree of
DOCTOR OF PHILOSOPHY
By
Chien-Sao Wang, B.S., M.S.
Denton, Texas
December, 1995
T>M<
Wang, Chien-Sao, Cell-free recovery and isotopic
identification of cvanide degrading enzymes from Pseudomonas
fluorescens NCIMB 11764. Doctor of Philosophy (Molecular
Biology), December, 1995, 94 pp., 15 tables, 10 illustrations,
references, 79 titles.
Cell-free extracts from Pseudomonas fluorescens NCIMB
11764 catalyzed the degradation of cyanide into products that
included C0 2 , formic acid, formamide and ammonia. Cyanide-
degrading activity was localized to cytosolic cell fractions and was
observed at substrate concentrations as high as 100 mM. Two
cyanide degrading activities were identified by: (i) the
determination of reaction products stoichiometries, (ii)
requirements for NADH and oxygen, and (iii) kinetic analysis. The
first activity produced C0 2 and NH3 as reaction products, was
dependent on oxygen and NADH for activity, and displayed an
apparent Km for cyanide of 1.2 mM. The second activity generated
formic acid (and NH3) pfus formamide as reaction products, was
oxygen independent, and had an apparent Km of 12 mM for cyanide.
The first enzymatic activity was identified as cyanide oxygenase
whereas the second activity consists of two enzymes, a cyanide
nitrilase (dihydratase) and putative cyanide hydratase. In addition
to these enzymes, cyanide-grown cells were also induced for
formate dehydrogenase (FDH), providing a means of recycling NADH
utilized by cyanide oxygenase.
Of the various enzymes elaborated by P. fluorescens NCIMB
11764 putative cyanide oxygenase was shown to be physiologically
important for cyanide assimilation. Therefore, further studies were
carried out to identify this enzyme. The stoichiometry of the
reaction indicated that one mole each of oxygen and NADH were
consumed per mole cyanide degraded, which was consistent with
that expected for an oxygenase type enzyme. Isotopic oxygen-18
labelling experiments indicated that one atom of molecular oxygen
was incorporated when 13C-labelled cyanide was converted to 13CC>2
by high-speed supernatants. These findings confirm the involvement
of a cyanide oxygenase in cyanide metabolism and prove that the
enzyme is a monooxygenase. However, one atom of oxygen from
H2 1 8 0 was also incorporated indicating that an additional step
between cyanide and CO2, involving the possible enzymatic
hydrolysis of a transient intermediate, is also involved. Preliminary
work aimed at the purification of cyanide oxidation enzymes
indicated that activity was localized in two fractions following
ultrafiltration, one containing the bulk of protein (fraction F1) and
the other (fraction F2) containing a small dialyzable protein of as
yet unknown nature.
Results from this work show that P. fluorescens NCIMB 11764
is capable of metabolizing cyanide by several different enzymatic
processes. Of these, oxidative conversion to CO2 and NH3 is
physiologically required for cyanide assimilation. Isotopic 1 8 0 -
labelling results now clearly show that a monooxygenase is involved
in initial cyanide degradative attack. Inhibitor studies suggest that
an essential sulfhydryl group may be involved in a mechanism
paralleling that described for nitrilase-type enzymes, but including
oxygen and water as reactants as opposed to water alone.
11
ACKNOWLEDGMENTS
I would like to express my gratitude to Dr. Daniel Kunz for
providing the opportunity to work in his laboratory and sharing his
experience and knowledge with me. I also thank the members of my
doctoral committee, Dr. Gerard O'Donovan for his support and helpful
comments during the study, Dr. Paul Cook, Dr. Manus Donahue and Dr.
Robert Benjamin, for their discussion and review of this manuscript.
A special thanks to Dr. Barney Venables and Dr. Michael
Richmond for their assistance in GC/MS analysis and isotopic
oxygen-18 labelling methods. I owe special thanks to Juan Silva-
Avalos, Olagappan Nagappan and Jui-Lin Chen for their support,
advice and friendship over the past several years.
I am grateful to my wife, Ai-Fang, for her encouragement in
just the right proportions, and finally, I thank my parents,
Hang-Tong and May, for their sustained support and understanding.
111
ABBREVIATIONS
CDA: Cyanide-degrading activity
CNO: Cyanide oxygenase
CND: Cyanide nitrilase (dihydratase)
CNH: Cyanide hydratase
F1: Ultrafiltration (30K molecular weight cut-off) retentate
fraction
F2: Ultrafiltration (30K molecular weight cut-off) filtrate
fraction
FDH: Formate dehydrogenase
HSSs: High-speed supernatants
KCN: Cyanide (CN7HCN in aqueous solution)
NCIMB 11764: National Collection of Marine and Industrial
Bacteria collection number 11764. Torrey, Scotland
IV
TABLES OF CONTENTS
Page
LIST OF TABLES viii
LIST OF ILLUSTRATIONS x
Chapter
1. INTRODUCTION 1
Cyanide in the microbial world Physical properties of cyanide Detoxification of cyanide by microorganisms Microbial assimilation of cyanide as a growth substrate Assimilation of cyanide as a nitrogenous substrate by
Pseudomonas fluorescens NCIMB 11764
2. METHODS 11
Bacterial strains Growth media and cultivation conditions Preparation of cell-extracts Analytical methods Analytical instrumentation Enzyme assays Oxygen uptake measurement Spectrophotometric measurement of NADH oxidation
by HSSs Identification and quantification of 14C-radiolabelled
cyanide metabolites Identification and quantification of 14C-radiolabelled
formate metabolites GC/MS Identification of cyanide conversion metabolite
Methods for measuring oxygen-18 incorporation Resolution of cyanide oxygenase Chemicals
3. RESULTS 27
PART I. RECOVERY OF CYANIDE-DEGRADING ENZYMATIC ACTIVITIES AT THE CELL-FREE LEVEL 27
Centrifugal fractionation of cyanide-degrading activity in cell-free extracts
Kinetics of cyanide conversion by HSS Reaction products and stoichiometries Conversion of formate to C02 by cyanide-induced
cell-free extracts Spectrophotometric identification of cyanide-induced
formate dehydrogenase activity in HSSs Summary of enzymatic activities induced by cyanide Effects of inhibitors on cell-free cyanide degrading
act iv i ty
PART II. REACTION STOICHIOMETRY AND ISOTOPIC IDENTIFICATION OF CYANIDE OXYGENASE 52
Reaction stoichiometry Evidence for the involvement of formate dehydrogenase
in NADH recycling Enzymatic incorporation of isotopic oxygen-18
PART III. PARTIAL RESOLUTION OF CYANIDE OXYGENASE... 72
Recovery of cyanide oxygenase in ammonium sulfate fractions
Requirement of a small dialyzable component for cyanide oxygenase
Characterization of small dialyzable cyanide oxygenase component
v I
4. SUMMARY 80
REFERENCES 86
V I I
LIST OF TABLES
Page Table
1. Reported mechanisms of cyanide degradation by
microorganisms 3
2. Composition of Lennox medium 12
3. Composition of minimal growth medium 13
4. Conversion of cyanide by cell-free fractions of P. fluorescens NCIMB 11764 29
5. Requirements for cell-free conversion of cyanide by
P. fluorescens NCIMB 11764 30
6. Cyanide degrading activity (CDA) kinetic constants 41
7. Products of cyanide metabolism by HSSs of P. fluorescens NCIMB 11764 43
8. Summary of cyanide-induced enzymatic activities in P. fluorescens NCIMB 11764 49
9. Compound test for as effector on cyanide degrading activity 50
10. Reaction stoichiometry for putative cyanide oxygenase measured in HSSs of P. fluorescens 11764 58
11. Effect of formate and NAD+ on putative cyanide oxygenase activity in P. fluorescens NCIMB 11764. . . 60
V I I I
12. Relative ion intensities of isotopic CO2 species detected by mass spectrometry in reaction mixtures supplied P.
fluorescens NCIMB 11764 HSSs and K13CN 68
13. Recovery of cyanide oxygenase in ammonium sulfate fractions 73
14. Recovery of cyanide oxygenase in ultrafiltration fractions 75
15. Effect of F2 treatment on cyanide oxygenase activity . . 79
i v
LIST OF ILLUSTRATIONS
Page Figure
1. Time course of cyanide consumption by high-speed supernatants from P. fluorescens NCIMB 11764 32
2. Effect of increasing protein concentration on cyanide-degrading activity in high-speed supernatants from P. fluorescens NCIMB 11764 34
3. Kinetics of cyanide degradation by high-speed supernatants from P. fluorescens NCIMB 11764 37
4. Lineweaver-Burk plots for enzyme-catalyzed cyanide degradation 40
5. Spectrophotometric detection of cyanide-induced formate dehydrogenase in HSSs from P.fluorescens NCIMB 11764 48
6. Stoichiometry of O2 uptake and cyanide 55
7. Stoichiometry of cyanide-dependent NADH oxidation. . . . 57
8. A metabolic scheme showing the interrelationship of formate dehydrogenase, putative cyanide oxygenase and cyanide nitrilase in cofactor NADH recycling 62
9. Time course of 1 4C0 2 formation by HSSs from P. fluorescens NCIMB 11764 65
10. Effect of F2 concentration on stimulation of cyanide oxygenase of F1 78
x
CHAPTER I
INTRODUCTION
Cyanide in the microbial world
Cyanide arises in the environment by two means. First it is
formed naturally by a number of bacteria, fungi, and plants in a
process known as cyanogenesis (Knowles, 1976), and second, it can
be generated by various industrial practices such as electroplating,
mining and steel processing (Knowles, 1976; Pettet and Ware, 1955;
and Way, 1981). Interest in cyanide research has grown since a
number of industries have sought to use biological methods for
cyanide waste treatment, which may compete effectively with
chemical processes (Knowles, 1976; Knowles and Bunch, 1986). For
this reason and because of its fundamental value, research directed
at understanding the chemical basis of cyanide metabolism at the
enzymatic level was undertaken.
Physical properties of cyanide
Hydrogen cyanide is a colorless weak acid (pKa=9.3), which
boils at 26°C, and is therefore readily volatilized from aqueous
solution. Cyanide (CN'/HCN) is highly reactive and complexes
tightly to metals such as nickel, copper, zinc, iron and gold (Sharp,
1976). Cyanide also reacts reversibly with keto groups to form
cyanohydrin derivatives. As a result of its reactivity, many enzymes
are strongly inhibited by cyanide. It is therefore often used as a
metabolic inhibitor, especially of cytochrome oxidases. Although
inhibitory, some biological systems elaborate an electron transport
system that is cyanide tolerant. This is thought to occur by the
formation of alternative cytochrome oxidase enzymes that do not
respond to the effects of cyanide (Knowles, 1976).
Detoxification of cyanide by microorganisms
Several chemical and physical treatment process have been
pilot plant evaluated either alone or in combination for the
degradation of cyanide compounds from waste waters (Mudder and
Whitlock, 1984). These processes require large quantities of
expensive chemicals resulting in high operation costs. Therefore,
research directed towards the evaluation and development of
biological treatment processes that generally produce low toxicity
effluent simply and cost effectively are of interest.
A number of cyanide-degrading microorganisms have been
reported and various enzymatic routes of metabolism have been
proposed (Knowles, 1976; Knowles, 1988). These are shown in Table
1 and discussed as follows.
a. Cvanide hydratase (EC 4.2.1.66, formamide hydrolyase)
Phytopathogenic fungi have been shown to elaborate cyanide
hydratase, which catalyzes the conversion of cyanide to non-toxic
formamide. For example, both spores and mycelia of Stemphylium
loti and Gloeocercospora sorghi induce when cultivated in the
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presence of cyanide suggesting that its induction has a
detoxification role (Fry and Millar, 1972; Fry and munch, 1975; Nazly
and Knowles, 1981; and Nazly et al., 1983).
b. Rhodanase (EC 2.8.1.1, thiosulphate: cyanide sulphur
transferase).
Rhodanase catalyses the conversion of cyanide to thiocyanate
in the presence of thiosulphate. It is widely distributed in
biological systems having been described in mammalian tissues,
plants and microorganisms (Sorbo, 1953; Silver and Kelly, 1976;
Ryan and Tilton, 1977; and Alexander and Volini, 1987). In addition
to cyanide detoxification, this enzyme has also been proposed to
function in the transfer of sulphur atoms in oxidative metabolism
(Westley, 1981).
c. B -Cvanoa lan ine synthase [EC 4.4.1.9, L-cysteine hydrogen
sulphide-lyase (adding HCN)].
Incorporation of cyanide into cysteine or o-acetylserine by p-
cyanoalanine synthase is thought to represent a major mechanism by
which cyanide is detoxified in plants (Hendrickson and Conn, 1968;
Ressler et al., 1969; Ting and Zschoche, 1970). This enzyme has
been detected in fungi (Strobel, 1967; Castric, 1981) and some
bacteria including Escherichia coli (Dunhill and Fowden, 1965),
Cromobacterium. violaceum (Brysk et al., 1969; Rodgers, 1981; and
Macadam and Knowles, 1984), and Enterobacter species (Sakai et
a!., 1981; and Yanese et al., 1982).
d. Nitroaenase (EC 1.18.6.1, reduced ferrodoxin: dinitrogen
oxidoreductase [ATP-hydrolysing] ).
Although not explicitly demonstrated in vivo, organisms that
elaborate nitrogenase have also been proposed to be capable of
cyanide detoxification. This is because it has long been recognized
that this enzyme, in addition to catalyzing the reduction of
molecular nitrogen, can also reduce various substrate analogues of
nitrogen such as cyanide (Li et at., 1982). In this process, cyanide
is thought to be reduced to methane and ammonia (a six electron
process), methylamine (four electrons) and possibly also
formaldehyde and ammonia via hydrolysis of a two electron
intermediate. The reduction of cyanide by nitrogenase has been
described for several organisms including Rhodospirillus gelatinosa
(Harris et ai, 1987).
e. Cvanide nitrilase (Cyanide dihydratase)
Cyanide dihydratase catalyses the conversion of cyanide to
formate and ammonia. This enzyme has been detected in the Gram-
positive bacterium Bacillus pumilus (Meyers et ai, 1991; Meyers
et at., 1993) and the Gram-negative organism Alcaligenes
xylosoxidans subsp. denitrificans (Ingvorsen et ai, 1991).
f. Cvanide oxygenase
The oxidative conversion of cyanide to C02 and ammonia by P.
fluorescens NCIMB 11764 was first reported by Harris and Knowles
(1983a), who proposed that this transformation was mediated by an
oxygenase-type enzyme. The importance of this enzyme to cyanide
conversion represents the major focus of this dissertation.
Microbial assimilation of cyanide as a growth substrate
Since cyanide contains two of the essential elements of life it
might be hypothesized that microorganisms could utilize it for
growth. But the toxicity of cyanide presents a problem when
isolating bacteria capable of using it as a substrate for growth.
Thus, a cyanide concentration high enough to serve as a source of
carbon and energy is likely to be too toxic to allow growth and there
have been no reports of organisms that can use cyanide as a
nutritional source of carbon. But several microorganisms capable of
using cyanide as a nitrogen source have been described. These
include, Aspergillus niger (Ivanoff and Zwetkoff, 1936; Rangaswami
et al., 1963); a unidentified OSram-positive, filamentous bacterium
(Ware and Painter, 1955); Pseudomonas fluorescens NCIMB 11764
(Harris and Knowles, 1983a); a pseudomonad reported by White et
ai (1988); seven pseudomonad species and three Klebsiella species
reported by Kunz and coworkers (Silva-Avalos et al., 1990) and
Alcaligenes xylosoxidans subsp. denitrificans reported by
Ingvorsen et al., 1991.
Assimilation of cyanide as a nitrogenous substrate by
Pseudomonas fluorescens NCIMB 11764
P. fluorescens NCIMB 11764 was chosen as the focus of this
study because of its ability to degrade cyanide and use the nitrogen
component as a source of nutrition. Therefore, this organism was
routinely cultivated in a medium where cyanide (KCN) was supplied
as the sole nitrogen source (Harris and Knowles, 1983a; Kunz et ai,
1992). Previous work indicated that growth on cyanide occurs by
conversion to ammonia, which is presumed to be assimilated via
established pathways involving glutamine synthetase (EC
6.3.1.2)/glutamine synthase (EC 1.4.1.13) and glutamate
dehydrogenase (EC 1.4.1.4) enzyme (Neidhardt et ai, 1990).
However, the enzymatic basis of cyanide conversion to NH3 was
poorly understood. Therefore, a major goal of this dissertation was
to learn more about how cyanide (CN* or HCN) was metabolized at the
enzymatic level.
Initial studies with P. fluorescens NCIMB 11764 by Harris and
Knowles (1983a) led them to suggest that enzymatic conversion of
cyanide was oxygenase (CNO) mediated. This suggestion was based
on the following observations: (i) washed cell suspensions and cell-
extracts did not catalyze cyanide conversion under anaerobic
conditions, (ii) cyanide turnover by cell-free preparations appeared
to be correlated with simultaneous oxygen uptake and NADH
oxidation, and (iii) C02 appeared to be formed (along with NH3) as a
metabolic product. These investigators proposed that either a
monooxygenase (1) or a dioxygenase (2) as shown in Scheme 1 acted
on cyanide. The former was thought to produce cyanate, thus
requiring an additional enzyme for its breakdown. This enzyme, it
was proposed, might be a cyanase (EC 3.5.5.3). However, subsequent
studies showed that while 11764 elaborated cyanase after growth
8
v+ HCN + 0 2 + NADH + H+ > HOCN + H20 + NAD'
HOCN + H20 > C02 + NH3 (1)
HCN + 0 2 + NADH + H+ > C02 + NH3 + NAD+ (2)
Scheme 1
on cyanate (Kunz and Nagappan, 1989), it was not coinduced along
with cyanide oxygenase when cells were grown on cyanide.
Conversely, growth on cyanate induced cyanase activity but not
cyanide oxygenase. In addition, a mutant strain unable to grow on
cyanate (phenotypic designation Cnt"), was still able to utilize
cyanide (phenotypic designation Cn+) as a nitrogen source (Nagappan,
1992). These results showed that cyanate and the enzyme cyanase
were not involved in cyanide utilization by P. fluoresceins NCIMB
11764 and therefore, it was proposed that the enzyme responsible
for cyanide conversion to C02 (and NH3) was a dioxygenase.
However, additional investigations by Kunz et a/., (1992) showed
that intact cells of NCIMB 11764 produced not only CO2 as a reaction
product, but also formate and formamide. Since formation of the
latter two products was observed under both aerobic and anaerobic
conditions, it was postulated that other enzymes in addition to a
putative oxygenase were capable of attacking cyanide. Scheme 2
shows three different enzymes that were proposed to be involved in
cyanide metabolism in P. fluorescens NCIMB 11764. These include
in addition to a cyanide oxygenase (CNO), a cyanide nitrilase-type
h o o 2 h + n h 3 ^ -
Assimilation
2H20
JL. CND
HCN
Ofe NADH + H+ NAD+
H>0
I™3
hcosh2
hH3 + C02
Assimilation
Scheme 2
enzyme (dihydratase) (CND) (giving formate and ammonia) and a
cyanide hydratase (CNH) generating formamide. Evidence that the
latter transformation was separate from that leading to formate
was based on the fact that formamide was shown not to undergo
further degradation (Kunz et al., 1992).
The hypothesis that this organism might produce a cyanide
nitrilase was consistent with evidence for related enzymes in other
bacteria. For example, Ingvorsen et al., (1991) showed that this
type of enzyme was made in Alcaligenes xylosoxidans subsp.
denitrificans strain DF3. Complete conversion of cyanide to
formate and ammonia under either aerobic or anaerobic conditions
was demonstrated without the apparent involvement of formamide
as an intermediate. A similar enzyme is thought to be responsible
for hydrolysis of cyanide to formate and ammonia in a pseudomonad
isolated from industrial wastewater by White et a/., (1988).
10
Cyanide degradation by this organism was also shown not to require
oxygen. With regards to cyanide hydratase, this enzyme has thus far
been described in phytopathogenic fungi only (e.g., Stemphylium loti
[Fry and Millar, 1972], and Gloeocercospora sorghi [Nazly et a!.,
1983]), making its possible occurrence in NCIMB 11764 novel. In
addition, no organism other than NCIMB 11764 has ever been
described that contains a putative cyanide oxygenase. Therefore,
biochemical studies were initiated to further identify and
characterize these enzymes at the cell-free level. In particular,
this dissertation had the following objectives: (i) recover cyanide
degrading enzyme activities at the cell-free level, (ii) verify the
actual involvement of cyanide oxygenase by isotopic oxygen-18
labelling experiments, and (iii) initiate the purification of the latter
enzyme.
CHAPTER II
METHODS
Bacterial strains
All experiments were conducted with P. fluorescens NCIMB
11764 which was obtained from the National Collection of Marine
and Industrial Bacteria, Torrey, Scotland. This bacterium was
isolated by R. E. Harris and C. J. Knowles in the United Kingdom
(Harris and Knowles, 1983a). For long-term storage this and other
strains were maintained in 10% dimethyl sulphoxide at -70°C. Cells
were routinely subcultured on Lennox agar (see Table 2 for formula)
plates every 4-6 weeks.
Growth media and cultivation conditions
The formula for minimal medium (MM) used for the growth of
cells is shown in Table 3. The procedure used for routine fed-batch
cultivation of cells on cyanide as a nitrogen source (cyanide-grown
cells) was the same as that described by Kunz et a!., 1992. For
this purpose, the inoculum was cultivated in 200 ml of minimal
medium containing glucose (20 mM) as the carbon source and 1 mM
NH4CI as the source of nitrogen. After 48 hours incubation at 30°C,
the entire 200 ml-inoculum was added to 2 liters of glucose
supplied MM (20 mM) with KCN (0.25 mM) serving as the nitrogen
11
12
Table 2. Composition of Lennox medium {Lennox, 1955)
Lennox broth medium
Bacto tryptone 10.0 g
Yeast-extract 5.0 g
NaCI 5.0 g
Distilled water 1,000 ml
pH 6.8, sterilize at 121°C for 20 min.
For Lennox agar, 20 g of Bacto-agar (Difco) was added.
13
Table 3. Composition of minimal growth medium (Kunz et al., 1981)
Buffer phosphate (P10X^
KH2P04 91.0 g
NaOH 16.8 g
Distilled water to complete 1,000 m I
pH 7.0
R-salts m-200Xl
10 % MgS04 • 7H20 400 ml
1 % FeS04 • 7H20 100 ml
concentrated HCI 2 ml
Mix and sterilize at 121°C for 20 min.
Minimal medium compositions
P1X (sterile) 100 ml
Glucose 1 M (sterile) 2 ml
R-salts 200 X (sterile) 0.5 ml
Glucose and R-salts were added aseptically to sterilized P1X. For
minimal agar, 2 % Bacto-agar (Difco) was added.
14
source. After 24 hours incubation a second addition of KCN (0.25
mM) was made followed by additions of 0.125 mM KCN each at 48 and
60 hours (total cultivation time was 72 hours with KCN additions
totaling 0.75 mM). The cells were then harvested by centrifugation
at 12,000 x g at 4°C and washed twice in 50 mM Na2HP04-KH2PC>4
buffer (pH 7.0) (SP buffer) and used immediately, or were stored at -
70°C. Cells induced for cyanide-degrading activity (CDA) were also
acquired through a service contract with the Fermentation Pilot
Plant in the Department of Biochemistry, University of Wisconsin,
Madison, Wl. In this procedure cells induced for cyanide-degrading
activity were obtained by growing cells starved for nitrogen in
stationary phase as described by Kunz et al., (1994). The large-
scale growth of cells was performed in 360 liters of glucose
supplied MM (20 mM) and NH4CI (1 mM) in a 400 liter fermenter.
After 24 hours the cells were induced by several additions of KCN
(0.1 to 0.2 mM) and cells were harvested 24 hours later by
continuous flow centrifugation. The yield was 300 g [wet weight]
cells, which were stored at -70°C until use.
Preparation of cell-extracts
Cell-extracts were prepared by the breakage of cells in a
French Press. For this purpose, frozen cells were suspended in two
volumes of SP buffer and broken at 20,000 psi. A small amount of
deoxyribonuclease (20 ng ml"1) was added to the resultant viscous
liquid and the preparation was incubated for 15 minutes at room
temperature prior to centrifugation at 30,000 x g for 30 minutes.
15
The supernatant retained as "crude-extract", which was then
further separated into a high-speed pellet (membrane) and a high-
speed supernatant (HSS, cytosolic) fraction by ultracentrifugation at
150,000 x g for 90 minutes at 4°C. The pellet was resuspended in
the original volume of phosphate buffer and washed by an additional
high-speed centrifugation before it and other fractions were
assayed for cyanide conversion activity. In general, enzyme activity
in HSS was stabilized by the addition of 1.5 mM EDTA (disodium
ethylene-diamine tetraacetate) to the buffer when extracts were
prepared. NAD+ was removed from HSSs when necessary by
incubation with 0.2 U ml"1 NADase (NAD+ glycohydrolase [EC 3.2.2.5]
[Sigma Chemical Co]) for 1 h at 30°C.
Analytical methods.
Cyanide
Cyanide was determined by the colorimetric method of
Lambert et al., (1975). For this purpose, 10 jxl of sample was added
to a test tube containing 1.14 ml of water and 50 nl of N-
chlorosuccinimide-succinimide reagent, followed by the addition of
50 |il barbituric acid-pyridine reagent. The absorbance was read at
580 nm. A lavender color indicated the presence of cyanide and the
absorbance was converted to cyanide concentration in unknown
samples using standard curve.
Ammonia
Ammonia was determined using the colorimetric method
described by Fawcett and Scott (1960). Reaction mixtures contained
16
in a total volume of 1.125 ml: 0.125 ml of sample, 0.25 ml of
sodium phenate reagent, 0.375 ml of 0.01% of sodium nitroprusside
and 0.375 ml of 0.02 N sodium hypochlorite. After 30 min, the
absorbance was measured at 630 nm at room temperature. A blue
color indicated the presence of ammonia and the absorbance was
converted to ammonia concentration using standard curve.
Formamide
The quantification of both formamide and formic acid in
reaction mixtures was determined after deproteinization of samples
with ZnSC>4. Also, any unconsumed cyanide present in deproteinized
samples that might interfere either with formamide or formic acid
determinations was first removed by flushing samples with a N2-
gas stream for 5 min. Samples were deproteinized by adding 10 |xl
6 M ZnSC>4 to 100 nl of cyanide-free sample. Precipitated protein
was removed by centrifugation in a microcentrifuge. The formamide
present was then determined by the colorimetric method of Fry and
Millar (1972), which involves conversion of formamide to the
corresponding hydroxamate. The sample (0.1 ml) was incubated for
10 min at 60°C with 0.2 ml of a 1:1 mixture of 3.5 N NaOH and 2.3 M
hydroxylamine hydrochloride. One tenth ml of 4 N HCI was added
followed by 0.1 ml of 1.23 M FeC^ reagent prepared in 0.075 N HCI,
and the absorbance read at 540 nm after 5 minutes. The limit of
formamide detection using this procedure was <0.2 mM. When
present at concentrations below this, formamide was determined by
chemical hydrolysis to formic acid, which was then determined
enzymatically (see below). Formamide hydrolysis was achieved by
17
adding 10 \i\ 3 M H 2 S0 4 to 90-100 | l deproteinized sample in a
sealed microcentrifuge tube and allowing the mixture to incubate at
95°C for 2 h. After this the sample was neutralized with 5 M NaOH
and assayed for formic acid.
Formic acid
Formic acid was determined enzymatically using commercial
formate dehydrogenase (Sigma Chemical Co., St. Louis, MO) as
described by Hopner and Knappe, (1974). Reaction mixtures
contained the following in 0.4 ml : 18-20 |imol KH2P04 (pH 7.5 ), 0.4
jxmol NAD+, 0.16 U formate dehydrogenase, and 10-20 jxl of sample
(0.1-0.2 |imol formic acid). Reactions were initiated by the addition
of enzyme and the change in absorbance at 340 nm was measured
over 2-4 min. Since reactions proceeded only to about 62%
completion, an average extinction value (e) for NADH of 3.85 mM*1
cm - 1 was routinely used to calculate the formate concentration.
Protein determination
The protein content in cell-extracts was determined using the
Lowry procedure (Lowry et a!., 1951). This was carried out by
adding 5 ^l of sample to 0.2 ml water. Just before use reagent A
was prepared by mixing one ml of 0.5% CuS04 • 5H20 prepared in 1%
sodium citrate with 50 ml of 2% Na2C03 prepared in 0.1 N NaOH.
One ml of reagent A was then added to samples, which were
incubated for 10 min at room temperature. This was followed by the
addition of 0.1 ml of 1 N Folin-phenol reagent. After 30 min
incubation the absorbance was read at 750 nm and protein
18
concentration was determined from standard curve using bovine
serum albumin as a standard.
Analytical instrumentation
All spectrophotometric analyses were recorded on either an
LKB Ultraspec II or a Perkin-Elmer Lambda-6 uv/vis
spectrophotometer. The pH of all prepared solutions was measured
on a PHM82 model Radiometer pH meter.
Enzyme assays
Ceil-free cyanide-disappearance assay
The routine determination of cyanide degrading activity was
measured in cell-free extracts in sealed vials (2 ml HPLC screw-cap
autosampler vials sealed with silicon rubber/PTFE septa, Rainin
Instrument Co., Woburn, MA). The standard reaction mixture
contained in 0.25 ml: cell-extract (9-10 mg protein ml"1), 2 mM
KCN, 4 mM NADH (where indicated), and SP buffer (pH 7.0). Reactions
were initiated by the injection of KCN and incubation mixtures were
placed at 30°C on a Gyrotory shaker (250 rpm). Ten |il samples were
withdrawn with a syringe at desired intervals for the determination
of cyanide by the method described in Analytical methods. Separate
reaction vials containing cell-extract incubated in the absence of
cyanide were also included as references for colorimetric
determinations. Where indicated, reaction mixtures were made
anaerobic by flushing vials with oxygen-free N2 for 5 minutes prior
to injecting KCN.
19
Cyanide oxygenase assay (oxygen/NADH-dependent
CDA)
The CDA in HSSs measured under aerobic conditions in the
presence of 4 mM NADH with KCN supplied at 2 mM was taken as a
measure of putative cyanide oxygenase (CNO) activity.
Cyanide dihydratase/cyanide hydratase assay
(oxygen/NADH-independent CDA)
The CDA measured at high concentrations of KCN (25 mM) under
anaerobic conditions in the absence of NADH was taken as a measure
of putative cyanide dihydratase/cyanide hydratase activity.
Reaction mixtures were made anaerobic by flushing vials with
oxygen-free N2 for 5 minutes prior to injecting KCN.
Formate dehydrogenase
Formate dehydrogenase was assayed spectrophotometrically in
reaction mixtures containing the following components in 0.4 ml, 50
mM KH2P04 (pH 7.5), 0.1 mM NAD+, and 1 mM sodium formate.
Reactions were initiated by the injection of HSS (10-20 |il, 0.12-
0.24 mg protein), and NADH formation was monitored at 340 nm.
Specific activities were calculated using a molar extinction
coefficient of 6.22 mM'1 cm"1 for NADH.
Oxygen uptake measurement
Enzyme-dependent oxygen uptake was measured with a
Hansatech Clark-type oxygen-electrode (Hansatech Instruments Ltd.,
Norfolk, England). The electrode was connected to a CB1 control box
20
and Cole Parmer Model 8373-20 recorder (Cole-Parmer Instrument
Company, Chicago, IL). The electrode cathode disc was prepared for
operation by placing a cigarette paper spacer (1" square) over the
disc, followed by moistening with saturated KCI. A one-inch square
piece of electrode membrane was then placed over the paper-covered
disc and held in place with spacer and O ring applied so that the
membrane was stretched smoothly over the surface of the electrode
dome. Calibration of the electrode was carried out as follows. The
range switch for the instrument was set to x1, output control to
maximum (fully clockwise), and back-off switch to "cancel". The
control box switch and recorder were then turned on and one ml of
aerated SP buffer placed into the electrode cuvette chamber. The
maximal output reading was adjusted to 90-95% (with 100%
representing buffer saturated with 0 2 ) with the output control
switch, and a few crystals of sodium dithionite (Na2S204) w a s added
to the chamber. As the dithionite combined with dissolved 0 2 the
electrode signal quickly responded to zero. At this time the "back-
off" control was turned on and the output reading adjusted to 10-
15% (of maximum recorder output) using the "coarse" adjustment.
The electrode chamber was then thoroughly washed with H 2 0 and
filled with 1 ml of aerated SP buffer. The recorder came to 90-95%
of full scale deflection indicating that the instrument was ready for
use. Further calibration was achieved by adding 0.1 nmol of catechol
follwed by 10-25 nl of crude catechol-2,3-dioxygenase (C230)
(prepared from P. putida cells carrying the TOL plasmid and induced
for this enzyme by growth on m-toluate [Kunz & Chapman, 1981]),
21
and measuring pen deflection upon addition of enzyme; since 1 mole
of oxygen is consumed per mole substrate degraded by C230, this
provides a convenient means of making sure the instrument is
calibrated appropriately.
Oxygen uptake by HSSs from P. fluorescens NCIMB 11764 was
measured as follows. One ml of SP buffer was added to the
electrode cuvette maintained at 30°C by means of a water-jacketed
circulator and the instrument allowed to stabilize momentarily.
Two tenths ml of HSS (2.4 mg protein) was then added and the rate
of oxygen uptake recorded for 1-3 min. After this, 1 ^imol of NADH
was added and the rate of 0 2 uptake again recorded for 1-2 minutes
or until a steady rate of consumption was observed. This gave a
measure of the rate of NADH oxidation. To determine whether KCN
could further stimulate oxygen consumption, 0.05 - 0.1 jxmol KCN
was added when the rate of NADH oxidation had reached steady state
and both the stimulation in rate and total magnitude of oxygen
consumption further recorded [see Figure 7 for further description
of the order of addition of substrate(s)].
Spectrophotometric measurement of NADH oxidation by
HSSs
The oxidation of NADH by HSSs was further determined
spectrophotometrically. For this purpose, 0.1 jimol NADH (10 fxl of
10 mM) was added to a quartz cuvette that contained 0.4 ml SP
buffer and 0.1 ml of HSS (1.2 mg protein). After determination of
the endogenous rate of NADH oxidation (NADH oxidase), 5 JLLI 5 mM
22
KCN was added. Another 5 fi.1 5 mM KCN was added when cyanide
became depleted. Specific activities were calculated using a molar
extinction coefficient of 6.22 mM"1 cm"1 for NADH.
identification and quantification of 1 4 C - r a d i o l a b e l l e d
cyanide metabolites
Radiolabeled metabolites generated under cell-free conditions
were categorized as either volatile (containing CO2 and HCO3*) or
non-volatile (containing formamide and formic acid) using BaCI2 as
a bicarbonate trapping reagent according to the procedure described
by Fallon et al. (1991) and Kunz et al.{1992). Reactions were
carried out in 25 ml serum-stoppered flasks fitted with a center
well. The main-compartment of the flask contained, 0.4 ml cell
extract (HSS) (9-10 mg protein ml*1), NADH (where indicated) and
SP buffer in a total volume of 0.5 ml. Reactions were initiated by
the injection of labelled (1-2.5 M-Ci; 37-92.5 kBq) and unlabelled KCN
(2 or 25 mM), and when complete, as a determined by simultaneous
colorimetric assays for cyanide, 0.3 ml of 4 N NaOH was injected
into the center well to trap volatile CO2. The flasks were allowed
to incubate for an additional 15 minutes at which time the contents
of the center well and main compartment of the flask were removed
and fractionated with BaCl2 to recover volatile and nonvolatile
products. One quarter ml of the contents from the main
compartment was treated with 0.12 ml of 0.1 N NaOH plus 0.03 ml of
40% BaCI2 for 5 min, and the mixture separated into fractions
designated main-compartment alkaline barium precipitate
23
(containing bicarbonate) and main-compartment alkaline barium
soluble (containing formamide plus formate) by centrifugation. The
pellet (alkaline barium precipitate) was washed by centrifugation
with 0.2 ml of 0.12 N NaOH and the washing combined with the
supernatant (alkaline barium soluble). The pellet was then
resuspended in 0.5 ml of 0.12 N NaOH and both it and the supernatant
sample were added to scintillation fluid. The contents of the center
well (0.3 ml) were treated in exactly the same fashion, generating
fractions designated center-well alkaline barium precipitate
(containing CO2 and HCO3") and center-well alkaline barium soluble
(containing unconsumed cyanide). All radioactive samples were
added to 8 ml of Aquasol scintillation fluid (Du Pont, NEN Research
Products, Boston, MA) and counted on an LS 7000 liquid scintillation
counter (Beckman, Irving, CA).
Identification and quantification of 1 4 C - r a d i o l abel led
formate metabolites
The identification and quantitation of C02 generated from
formic acid was also performed radioisotopically using the same
procedure employed for 14C02 quantitation from cyanide. Reactions
were again performed in sealed flasks and initiated by the injection
of 1-2.5 |xCi (37-92.5 kBq) sodium [14C]formate (0.5 mM). NAD+
(where supplied) was provided at 1 mM.
GC/MS identification of cyanide conversion metabolites
Carbon-containing reaction products were also determined by
24
GC/MS. GC/MS analysis was performed in collaboration with Dr. B.
Venables, Trac Laboratories Inc., Denton, TX., using a Hewlett
Packard 5992 GCMS equipped with a Teknivent Vector-One Data
Acquisition System. 1 3 C0 2 45 m/z species in the headspace of
incubations conducted with K13CN as the substrate was analyzed by
mass spectrometry. This was accomplished by injecting 10 (AI gas
over a polydimethylsiloxane capillary column (Alltech Associates
SE3-30M). The presence of formic acid was also confirmed by
GC/MS. This was accomplished by injecting 10 jal of the reaction
mixture onto an Alltech Associates AT1000 10m megabore column
(injector 110°C, flow 6.5 ml min"1 He, oven programmed at 40°C for
2 min, then ramped to 110°C at 10°C min"1) fitted to a mass
spectrometer. The reaction time was 9 min and mass of 46 (HCOOH)
were detected in incubations conducted with KCN.
Methods for measuring oxygen-18 incorporation
The conditions for measuring oxygen-18 incorporation were
the same as for the assay of the putative cyanide oxygenase except
that K13CN was used as the substrate. Also, to increase the amount
of enzyme available and reduce the extent of labelled CO2 exchange,
some reactions were conducted with concentrated HSS and with
NaHC03 (final 50 mM) added to the reaction mixtures. HSS was
concentrated by lyophilization in a Speed-Vac and the dried powder
resuspended in water or oxygen-18 water. Reactions with labelled
water were conducted in 2 ml-sealed HPLC vials containing HSS or
concentrated HSS (2.4 -11 mg protein), 120-200 nl H 21 8 0 (final 44-
25
77% enrichment), 5 fxl of 100 mM K13CN (final 2 mM) and 10 p.l of
100 mM NADH (final 4 mM) in a final volume 0.25 ml. Experiments
with 1 8 0 2 were carried out in 15 ml capacity glass vessels to which
a break-seal glass ampoule containing 25 ml of 1 8 0 2 (at 1 atm) was
attached. The vessel was flushed with oxygen-free N2 (99.9 atom%)
for 10 minutes before breaking the seal and gases were allowed to
equilibrate throughout the system. Reactions mixtures contained
the same components as those described for H 21 8 0 experiments. In
each case, reactions were initiated by the injection of NADH and the
headspace gas analyzed as described above at the times where
indicated . The relative abundances of m/z species 45 ( 1 3 C 1 6 0 1 6 0) ,
47 (1 3C1 601 80), 49 (1 3C1 801 80), 44 (1 2C1 601 60), 46 ( 1 2 C 1 6 0 1 8 0)
and 48 ( 1 2 C 1 8 0 1 8 0) were quantified from the relative ion intensities
measured at an ionizing voltage of 70 eV. The isotopic enrichments
of 1 8 0 2 (m/z 32) and H 21 8 0 (m/z 20) were determined respectively,
by analyzing the headspace gas and aqueous reaction solution.
Resolution of cyanide oxygenase
Ammonium sulfate precipitation
HSSs were fractionated by ammonium sulfate precipitation
into three fractions: the 0-35% pellet, the 35-70% pellet and the
70% supernatant fraction. The two pellet fractions were
resuspended in SP buffer and all three fractions were desalted by
dialysis against SP buffer (MW cutoff 6000-8000).
U l t r a f i l t r a t i o n
Ultrafiltration was carried out using Centricon-30 (30,000 MW
26
cutoff) centrifugal microconcentrators. HSSs were loaded on
Centricon and centrifuged at 5,000 x g for an hour at 4°C. After
centrifugation, retentate and filtrate fractions were pooled.
Chemicals
Both K12CN (97%) and K13CN (99 atom%) were obtained from
Aldrich Chemical Company (Milwaukee, Wl). K14CN (54 mCi mmol"1,
2.0 GBq mmol"1 ) and H14COONa (48.7 mCi mmol*1, 1.8 GBq mmol"1)
were purchased from Du Pont (NEN Research Products, Boston, MA).
Isotopic 1802 (99 atom%) and H2180 (96 atom%) were supplied by
Isotec Inc. (Miamisburg, OH). Other chemicals and reagents used
were purchased from Fisher (Springfield, NJ), or Sigma (St. Louis,
MO) and were of the highest available commercial grade.
CHAPTER III
RESULTS
PART I. RECOVERY OF CYANIDE-DEGRADING ENZYMATIC ACTIVITIES AT
THE CELL-FREE LEVEL
Studies by Kunz et al., (1992) showed that intact cells grown
on cyanide catalyzed its time-dependent disappearance from
reaction mixtures at concentrations as high as 100 mM. Since the
product distribution varied with the reaction conditions (aerobic vs
anaerobic) and substrate concentration, it was hypothesized that
more than a single enzyme was responsible for cyanide attack. As a
first step toward the identification and characterization of these
enzymes efforts were made simply to show that the cyanide
conversion activities observed for intact cells could be recovered at
the cell-free level. The results that follow represent the outcome
of these efforts.
Centrifugal fractionation of cyanide-degrading activity in
cell-free extracts
Crude extracts were initially fractionated into cytosolic and
membrane fractions by ultracentrifugation as described in METHODS,
and different fractions assayed for CDA. Since initial studies by
27
28
Knowles and colleagues (1983a,b) indicated that cyanide degradation
using cell-free preparations was both oxygen- and NADH-dependent,
these experimental conditions were repeated in an effort to recover
CDA. Table 4 shows that under these conditions the bulk of CDA
measured at relatively low substrate concentration (2 mM) was
located in the HSS fraction. These findings are similar to those
first reported by Harris and Knowles (1983b). However, as shown in
Table 5, activity could still be observed when NADH was omitted
from reaction mixture (21% of that of complete), or under anaerobic
conditions (7% of complete). F:igure 1 shows the rate at which
cyanide was degraded under various reactions. These data again
show the stimulatory effect of oxygen and NADH on cyanide
degrading activity, but also show the lack of complete dependence on
the two for activity. These results supported the idea that P.
fluorescens NCIMB 11764 was capable of producing additional
enzymes besides a putative cyanide oxygenase. The presence of
these additional enzymatic activities which appeared to function
anaerobically was previously not observed in this organism, but was
inferred from the intact cell studies of Kunz et al., (1992). In order
to show that cyanide disappearance under cell-free conditions were
enzyme-catalyzed, rates of disappearance were measured at
different protein concentrations and the results obtained are
illustrated in Figure 2. These data show that regardless of whether
or not activity was measured aerobically (in the presence of NADH)
or anaerobically (in the absence of NADH) CDA was protein-
concentration dependent. It was further shown that in non-enzyme
29
Table 4. Conversion of cyanide by cell-free fractions of P.
fluorescens NCIMB 11764
Cell fraction Cyanide converted3
(nmol min"1 mg [protein]"1)
crude-extract
30,000 X g supernatant 8
high-speed
150,000 X g supernatant 14
high-speed
150,000 X g pellet
a Reaction mixtures contained: 2 mM KCN, 4 mM NADH, 50 mM SP phosphate buffer (pH 7.0), and 1.6 to 4.0 mg protein in a final volume of 0.25 ml. Measured as initial rates of cyanide disappearance.
30
Table 5. Requirements for cell-free conversion of cyanide by P. fluorescens NCIMB 11764
Reaction Cyanide converted
conditions3 nmol min"1 mg"1 (%)
Complete (+ 02 /+ NADH) 14 (100)
+ 02 / -NADH 3 (21)
- 0 2 /+ NADH 1 (7)
- 0 2 / - NADH 1 (7)
a Reactions conducted with HSS and activity measured as the rate of cyanide disappearance as described in METHODS. Anaerobic reaction vials were flushed with N2 for 10 minutes prior to the addition of NADH and KCN.
31
Figure 1. Time course of cyanide consumption by high-speed
supernatants from P. fluorescens NCIMB 11764. Reactions
conducted at 2 mM KCN and 4 mM NADH where indicated under the
following conditions: Q +02/+NADH; Q +02/-NADH; # , - 0 2 /
-NADH.
32
•8
>» u
\ 10 20 30 40 50
Time (min)
33
Figure 2. Effect of increasing protein concentration on cyanide-
degrading activity in high-speed supernatants from P. fluorescens
NCIMB 11764. Activities measured at 10 mM KCN under aerobic
(+02/+NADH) [Q] and anaerobic (-02 / -NADH)[^] conditions.
34
r—>
C I
§ u 0> H3
u
1 2
Protein (mg)
35
controls, or reaction mixtures supplied boiled HSS (100°C for 15
min), <1% of the cyanide supplied was degraded.
Kinetics of cyanide conversion by HSS
The kinetics of cyanide degradation were investigated under
the different conditions in which CDA was observed. This included:
aerobic reaction mixtures with NADH present, aerobic reaction
mixtures with NADH absent, and anaerobic reaction mixtures with
NADH absent. Figure 3 shows a plot of initial rates of cyanide
consumption observed under the above conditions as a function of
substrate concentration. These data show that cell-free CDA could
be observed at KCN concentrations as high as 100 mM, which is one
hundred-fold greater than that previously reported (Harris and
Knowles, 1983b). Again, the highest rate of cyanide consumption
was observed when both oxygen and NADH were supplied, with
maximal activity recorded at 20 mM cyanide: above this the activity
declined probably due to enzyme inhibition. A second activity
observed in the absence of oxygen (and NADH) was also apparent at
cyanide concentrations as high as 100 mM. Under aerobic conditions
with NADH omitted rates of substrate consumption fell between
those recorded aerobically and anaerobically, suggesting that under
these conditions the total CDA represented the contribution of
possibly two enzymes. Under all three conditions the initial rates of
substrate disappearance appeared to followed Michaelis-Menten
kinetics. In order to further differentiate the various activities,
36
Figure 3. Kinetics of cyanide degradation by high-speed
supernatants from P. fluorescens NCIMB 11764. Reaction
conditions: O . +02/+NADH; D , +02/-NADH; -02/-NADH.
37
e 200 \ %
S
/
r/ a 50
K.CN
38
rates of substrate consumption were compared after the initial
rates recorded under anaerobic conditions (with NADH absent) were
subtracted from those recorded aerobically (with NADH present). In
this way enzymes contributing to the total CDA were distinguished
on the basis of favored reaction conditions and the apparent
(denoting the use of non-purified enzyme preparations) kinetic
constants estimated without individually purifying each enzyme.
Figure 4 shows the results when plot of the data was made in a
Lineweaver-Burk form (Cornish-Bowden and Eisenthal, 1974; Dowd
and Riggs, 1965; Eisenthal and Cornish-Bowden, 1974). These data
show that significantly different Km values were obtained under
aerobic and anaerobic conditions (1.2 versus 12 mM, respectively).
From these observations it was concluded that one enzyme in HSSs
had a low Km and was both oxygen- and NADH-dependent (referred to
CDA1), and a second enzyme had a high apparent Km and required
neither oxygen nor NADH (referred to as CDA2). In contrast to the
differences observed in apparent Km, the observed Vmax values for
the two were found to be identical (125 iM min"1; Figure 4). The
kinetic data for the CDAs described in cell-extracts are further
summarized in Table 6.
Reaction products and stoichiometries
The dependence of CDA1 on oxygen and NADH was found to be
analogous to the findings of Harris and Knowles (1983b), who
proposed that cyanide-degradation was oxygenase-mediated.
However, in order to further demonstrate the involvement of such an
39
Figure 4. Lineweaver-Burk plots for enzyme-catalyzed cyanide
degradation under aerobic (O ) and anaerobic ( 0 ) conditions.
40
0.025
0.020 -
S 0.015 -
0.005 -
1.2 -0.8 -0.4 0 0.4 0.8 1.2 1/[KCN] (mM1)
41
Table 6. Cyanide degrading activity (CDA) kinetic constants
Activity Vmax Km
(nmol min"1mg"1) (mM)
CDA1 (+02/+NADH) 12.5 1.2
CDA2 (-02/-NADH) 12.5 12
42
enzyme, and also to identify the nature of additional enzymes (e.g.,
CDA2), the reaction products formed after incubating HSSs with
cyanide at concentrations favoring one or the other activity were
determined. For this purpose, incubations were conducted at 2 and
25 mM KCN (considered optimal, respectively, for the apparent low
[CDA] and high [CDA] Km activities) and the product balances
determined. Table 7 shows that regardless of the reaction
conditions, greater than 90% of the cyanide carbon could be
recovered as C02 , formic acid and formamide. In contrast, the
accumulation of cyanide-nitrogen equivalents as formamide and
ammonia varied substantially (39-93 molar%). Presumably, this is
because ammonia once formed, is further metabolized. This helps to
explain the molar ratios of formamide : ammonia, which under
aerobic (with NADH present) and anaerobic (without NADH)
conditions were only 10:38 and 32:7 (48 and 39% molar recoveries),
respectively. For comparison, at 25 mM KCN, values of 37:56 and
41:49 were recorded.
Significant similarities were obtained when the carbon-
derived product balances obtained under cell-free conditions (Table
7) were compared with those previously reported for intact cells
(Kunz et a!., 1992). In each case, CO2 was identified as the major
reaction product under aerobic conditions as opposed to formic acid
and formamide under anaerobic conditions. Also, CO2 represented
the major reaction product at low substrate concentrations as
opposed to formic acid and formamide at higher concentrations.
These results when taken together with the kinetic data summarized
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44
in Table 6 suggest that the oxygen- and NADH-dependent activity
(CDA1) was similar to putative cyanide oxygenase (CNO) as first
described by Harris and Knowles (1983b). This enzyme, because of
its lower apparent Km, is believed to be responsible for conversion
of cyanide to C02 in 87% yield when supplied at low substrate
concentration (2 mM). In contrast, at higher concentrations CDA2 is
believed to catalyze the conversion of cyanide to formate (and NH3)
and formamide. These results are analogous to the findings reported
earlier (Kunz et a/., 1992) in which the same products were
identified when intact cells were supplied high concentrations of
cyanide. The production of these products is consistent with an
enzyme reaction mechanism involving putative cyanide nitrilase
(producing formic acid [and NH3 j) and cyanide hydratase (producing
formamide) enzymes as previously proposed (Scheme 2) (Kunz et al.,
1992).
Conversion of formate to C02 by cyanide-induced cell-free
extracts. The detection of C02 as a reaction product regardless of
whether or not HSSs were induced with cyanide under aerobic or
anaerobic conditions (Table 7) raised the possibility that C02 could
arise by two different means as shown in Scheme 3. One of these, it
was presumed, involved the direct conversion of cyanide to C02 (by
the action of CDA1 [putative cyanide oxygenase]) while the second
involved C0 2 production from formate (since the latter was also
identified as a reaction product). To determine whether formate
could be further metabolized, initial experiments were performed in
45
which HSSs were incubated with radiolabeled formate and the
production of 1 4 C0 2 (and H14C03") was measured as described in
METHODS. As much as 25 ± 2% (mean and mean deviations from two
CQ>+NH3
NH3+HCOOH COz
Scheme 3
separate experiments) of the formate supplied to HSSs was found to
be converted to C02 during a 2.5 h incubation. This increased to 88 ±
3% when reaction mixtures were also supplied NAD+, indicating that
formate could in fact be converted to C02, and that the responsible
enzyme was NAD+-dependent. The conversion in the absence of
added NAD+ was somewhat of a surprise until it was discovered that
pretreating HSSs with NADase prior to incubation decreased the
yield of C02 to 4.8 + 1.7%. This indicated that formate conversion in
the absence of added cofactor could be attributed to endogenous
NAD+ present in HSSs.
Spectrophotometric identification of cyanide-induced
formate dehydrogenase activity in HSSs
The observation that conversion of formate to C02 was NAD+-
dependent suggested that the responsible enzyme was a formate
46
dehydrogenase. This was verified by spectrophotometric assays as
shown in Figure 5. These data show that HSSs catalyzed the
formate-dependent reduction of NAD+ and that this activity was
protein-concentration dependent. Other experiments indicated that
activity was present under either aerobic or anaerobic conditions.
Summary of enzymatic activities induced by cyanide
Table 8 summarizes the various enzymatic activities detected
in P. fluorescens NCIMB 11764 when either grown on, or induced
with cyanide. One of these (CDA1) was tentatively identified as
cyanide oxygenase as already discussed while CDA2 is believed be
composed of one or two enzymes, namely CND and CNH. Since it has
not been possible at this point to differentiate the latter two
activities, CDA2 is simply referred to as "CND/CNH", representing
the possible contribution of two different enzymes. In addition to
CDA1 and CDA2, cyanide also induced the formation of formate
dehydrogenase.
Effects of inhibitors on ceil-free cyanide degrading
ac t iv i ty
In order to obtain information on the possible mechanism of
cyanide degradation, various compounds, as shown in Table 9, were
tested as potential inhibitors (or activators) of cyanide degrading
activity. The results show that of all the compounds listed, only
phenylmercuric acetate, p-hydroxymercuri benzoate, and HgCI2
gave any significant inhibition (35-53%). Since these compounds are
47
Figure 5. Spectrophotometric detection of cyanide-induced formate
dehydrogenase in HSSs from P. fluorescens NCIMB 11764-showing
the effect of increasing protein concentration on formate-dependent
formation of NADH.
48
r r
en
CN
o> <*•» 9 C
B w
E H
( u i a ot>£) a o u B q j o s q y
49
Table 8. Summary of cyanide-induced enzymatic activities in P.
fluorescens NCIMB 11764
Specific activity (nmol min"1 mg"1 [protein])
Cultivation
conditions3
CNO CND/CNH Formate
dehydrogenase
Cyanide-grown
Cyanide-induced
Ammonia-grown
12.6
8.1
<0.5
11.8
4.5
<0.5
14.0
8.8
<0.5
a Cultivation conditions are described in METHODS.
50
Table 9. Compound test for as effector on cyanide degrading activity
Compound Concentration Activi ty3
(mM) (%)
Control 100
HgCI2 0.1 65
p-Hydroxymercuri benzoate 0.0005 62
Phenylmercuric acetate 0.005 47
Fe3+ 0.2 98
Ni2 + 0.2 87
Acetylene 1.5 99
Imidazole 1.5 96
Pyrimidine 1.5 91
Histidine 1.5 86
Antimycin A 1 100
Carbonyl cyanide m-chloro-
phenyl hydrazone 1 100
2,2'-Dipy ridyl 1 100
2,4-Dinitrophenol 1 100
Sodium azide 1 100
a Activity measured as rate of cyanide disappearance in the standard cyanide degradation assay described in METHODS with reagents present at the concentration indicated.
51
known to interact with sulfhydryl groups, these data indicate that
this functional group may play a role in the enzymatic mechanism.
Although none of the compounds tested showed any significant
activation, EDTA (1.5 mM) was found to stabilize cyanide degrading
activity in HSSs; it therefore, was routinely added to buffer used to
prepare cell-extracts.
52
PART II. REACTION STOICHIOMETRY AND ISOTOPIC IDENTIFICATION
OF CYANIDE OXYGENASE
Results which showed that C02 (and NH3) were formed as
reaction products when cyanide was oxidatively metabolized by
HSSs (Table 6) supported the idea that an oxygenase-type enzyme as
originally proposed by Harris and Knowles, (1983b) was responsible
for conversion. We further suspected that since the Km towards
cyanide by CDA1 (believed to contain putative cyanide oxygenase)
compared with CDA2 was much lower (1.2 vs 12 mM) than the former
enzyme, it was important physiologically for cyanide utilization.
This idea was reinforced by parallel work conducted by fellow
graduate student Mr. Jui-Lin Chen who showed conclusively that
cyanide oxygenase activity in a mutant having lost the ability to
grow could not be detected (Kunz. et al., 1994). Thus, because of
the importance of this enzyme for cyanide utilization, one of the
major goals of this thesis was to prove unequivocally that an
enzyme fitting the description of an oxygenase was involved. To
accomplish this, experiments aimed at verifying the reaction
stoichiometry as originally reported by Harris and Knowles (1983b)
were repeated and the molecular source of oxygen atoms
incorporated into cyanide was investigated with isotopically
labelled 1 8 0 2 and H21 80.
53
Reaction stoichiometry
Figure 6 shows the results obtained when oxygen consumption
was measured with an oxygen electrode in the presence and absence
of cyanide (and NADH). These data show a stimulation in the rate of
oxygen consumption by NADH and a further stimulation upon the
addition of cyanide, which continued until all of the available
cyanide was consumed. The initial NADH-dependent rate of oxygen
consumption was believed to be attributed to NADH oxidase activity
which was calculated to have a specific activity of 2 nmol min"1
mg"1 (protein). The cyanide stimulated activity was concluded to be
due to cyanide oxygenase. The calculated molar ratio for oxygen
consumed and cyanide degraded was 1:0.97 + 0.12 (n=7). Related
experiments aimed at measuring the stoichiometry between NADH
oxidation and cyanide consumed as determined spectrophoto-
metrically were also conducted. As shown in Figure 7, cyanide
stimulated the rate of NADH oxidation above that attributed to NADH
oxidase, with the amount of NADH consumed continuing until all of
the available cyanide was again degraded. In this case, the molar
ratio between NADH oxidized (based on e, 6.22 mM"1cm"1) and
cyanide degraded was determined to be 1:1.02 ± 0.06 (n=10). Table
10 summarizes the results obtained for the reaction stoichiometry
which are in general agreement with findings initially reported by
Harris and Knowles (1983b).
54
Figure 6. Stoichiometry of 0 2 uptake and cyanide. The oxygen
electrode at 30°C contained 0.7 ml phosphate buffer (pH7.0) and 0.3
ml HSS {3.6 mg protein). NADH (10 |xl, 100 mM) was injected and the
rate of oxygen uptake by NADH oxidase was determined. KCN (10 ]i\,
10 mM) was then added and the stimulation in oxygen uptake was
recorded.
55
0.05 fimol KCN
0.1 jimol KCN
1 jxmol NADH
300 )il HSS (3.6 mg protein)
56
Figure 7. Stoichiometry of cyanide-dependent NADH oxidation. A
cuvette containing 0.4 ml phosphate buffer and 0.1 ml of HSS (1.2
mg protein) was incubated with 10 ]nl of 10 mM NADH. After
determination of the endogenous rate of NADH oxidation (NADH
oxidase), 5 jnl 5 mM KCN was added. Another 5 |il 5 mM KCN was
added when cyanide became depleted.
57
3 e
4> £
(uiu OK) wueqjosqv
58
Table 10. Reaction stoichiometry for putative cyanide oxygenase
measured in HSSs of P. fluorescens 11764a
KCN 0 2 NADH Estimated
Supplied Consumed Consumed molar ratios
(nmol) (nmol) (nmol) KCN: 02 :NADH
25 NDb 25 ± 1 1 : ND : 1.00
50 48 ± 6 50 ± 4 1 : 0.96 : 1.00
100 97 ± 12 ND 1 : 0.97 : ND
a Cyanide was measured colorirnetrically. Oxygen and NADH consumption were determined polarographically and spectrophtometrically as described in METHODS.
b ND indicates not determined.
59
Evidence for the involvement of formate dehydrogenase in
NADH recycling
The discovery of formate dehydrogenase in P. fluorescens
NCIMB 11764 suggested that the NAD+-driven oxidation of formate
might provide a means of generating the NADH utilized by cyanide
oxygenase. To explore this possibility the effect of NAD4" and
formate on both cyanide oxygenase activity and corresponding molar
conversion yield of cyanide into C02 were tested; the rationale being
that if these compounds were involved in NADH recycling, an effect
on both enzymatic activity and production of C02 might be observed.
Table 11 shows that when reactions were conducted with extracts
that were treated with NADase, both cyanide oxygenase activity and
the corresponding C02 yield were substantially lower than in
comparable reactions in which untreated HSSs were employed.
However, when reactions were supplied with NAD+, both cyanide
degrading activity and C02 yield increased to 67 and 46% of
maximal, and when formate was supplied, these increased to 98 and
100%, respectively. In contrast, the addition of formate alone had
no significant effect on either enzymatic activity or C02 production.
A role for both formate and NAD+ in the recycling of NADH was thus
inferred. A metabolic scheme showing the inter-relationship of
formate dehydrogenase, putative cyanide oxygenase and cyanide
nitrilase in cofactor recycling is shown in Figure 8.
60
Table 11. Effect of formate and NAD+ on putative cyanide oxygenase
activity in P. fluorescens NCIMB 11764
Reaction conditions Specific % Molar yield
activity 1 4 C0 2 from
[nmol min"1 K14CNb
(mg protein)"1]3
NADH 12.6 ± 1.4 (100) 87.1 ± 5.1 (100)
No NADH 2.6 ± 0.6 (21) 25.1 ± 2.8 (29)
NADase-treated HSS 1.2 ± 0.3 (10) 3.6 ± 0.8 (4)
NAD+ 8.5 + 1.2 (67) 39.6 ± 0.6 (46)
Formate 2.4 + 0.4 (19) 18.2 ± 2.4 (21)
NAD+ + formate 12.4 + 1.6 (98) 87.2 + 1.3 (100)
a Calculated from initial rates of cyanide consumption by HSSs from cyanide-grown cells.
b Determined as the sum of volatile C02 and non-volatile HC03~
collected after > 90% of the substrate was consumed.
61
Figure 8. A metabolic scheme showing the inter-relationship of
formate dehydrogenase, putative cyanide oxygenase and cyanide
nitrilase in cofactor NADH recycling.
62
0. HCN
NADH + NAD
CNO
FDH
CND COo + NH-
63
Enzymatic incorporation of isotopic oxygen-18
While the above experiments were all consistent with a
reaction mechanism involving an oxygenase, ultimate proof for this
required that incorporation of molecular oxygen into the substrate
be demonstrated (Hayaishi, 1982). In anticipation of these
experiments a time course for the production of C02 was determined
as shown in Figure 9. This experiment shows that HSS catalyzed the
conversion of radioactive K14CN into CO2 in about an 80% yield in 30
minutes. Enzymatic activity was both oxygen and NADH dependent
although a small amount of conversion was also observed under
anaerobic conditions which is consistent with activities expected
respectively, for CNO and CND/CNH as described in Part I of this
dissertation. The specific activity for the putative cyanide
oxygenase determined by measuring the rate of CO2 production was
estimated to be 16 nmole min"1 mg"1. This agrees with values
obtained by other methods including measurement of rates of
cyanide disappearance, oxygen uptake, or NADH oxidation. Once it
was established that we were dealing with what appeared to be an
oxygenase-type enzyme oxygen-18 incorporation experiments were
performed, the procedures for which are outlined in Methods.
The recovery of labelled oxygen in CO2 was complicated since
it readily exchanges its oxygen atoms with water. At pH 7.0, for
example, CO2 is rapidly hydrated and dehydrated (hydration/
dehydration cycle) (Pocker and Bjorkquist, 1977) as shown in
Scheme 4 and scrambling of oxygen atoms occurs. This helps to
explain the results observed when theisotopic content of 1 3C02
64
Figure 9. Time course of 1 4 C0 2 formation by HSSs from P.
fluorescens NCIMB 11764. Reactions conducted with 2 mM K1 4CN
and 4 mM NADH supplied under aerobic (O) and anaerobic ( £ )
conditions. 1 4 C0 2 was determined as described in METHODS.
65
100 "O 0) fc. 0) > c o o
<M S 8 *
o s <D O) <0
0) o I-0) Q.
10 20
Time (minute)
66
formed from K13CN was determined in initial experiments conducted
for prolonged periods as shown in Table 12. Only 0.1% of the CO2
formed was 180-Iabelled when 1802 was supplied (Experiment 1).
C01 80 + H20 " HC00180' + H+ C00 + H21 80
Scheme 4
This can be explained by the fact that any potentially labelled
species would likely be diluted out as a result of exchange with the
large excess of unlabelled water. Conversely, the same explanation
can be used to explain why up to 46.8 and 19.2%, respectively of m/e
species 47 ( 1 3C 1 60 1 80) and 49 (1 3C1 801 80) were detected in
incubations conducted with H2180 (Experiment 1, Table 12). In this
case, unlabelled 13C02 became labelled because of chemical
exchange with the large excess of labelled H2180 (44%) supplied. It
is important to emphasize that the ratio of m/z 45:47:49 observed in
the latter experiment (0.34:0.47:0.19) is consistent with the
expected ratio (0.31:0.49:0.19) assuming complete chemical
exchange. The latter calculation was arrived at from the following
probability expression:
£(x2 + 2xy + y2) = 1
where x2 = probability that both atoms are derived from 1 6 0
and y2 = probability that both atoms are derived from 1 8 0
at 44% H 21 8 0 enrichment
67
Table 12. Legend
a Reaction mixtures (0.25 ml) contained approximately 50 mM SP
buffer, HSS (or concentrated HSS at the protein concentration
shown), and 50 mM NaHC03 (where indicated). Reaction mixtures
supplied H 21 8 0 contained 125 jxl of 96 atom% H2
1 80, and 100 jil
and 95 JLLI respectively, HSS and four-times concentrated HSS in
Experiment 1 and 2. In Experiment 3, 200 ^l of H 21 8 0 was used to
resuspend the dry powder obtained after lyophilizing HSS, which
was added to reaction vessels already supplied NaHC03. Reactions
were initiated by injection of 2 mM K13CN and 4 mM NADH. b Values obtained for 1 8 0 2 experiments were corrected for m/z 46
and 48 generated from 1 2 C 1 6 0 2 (m/z 44) by non-enzymatic
labelling as a result of chemical exchange with 97-99 %
enrichment 1 8 0 2 .
c o V-» o co 0
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68
CM a!
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69
It was therefore concluded from these experiments that because of
the length of incubation it would not be possible to determine the
molecule source of oxygen atoms incorporated.
Several solutions were thought of to overcome the problem of
CO2 exchange. One was to decrease the time between reaction
initiation and analysis of reactions. The second was to increase the
enzyme concentration, and the third was to incorporate bicarbonate
into reaction mixtures. The latter approach it was hypothesized,
would alter the chemical equilibria (hydration/dehydration cycle) to
the point where exchange would effectively be reduced. The latter
approach was based on findings reported over forty years ago by
Mills and Urey (1940) who showed that the rate of CO2 exchange at
25°C in aqueous solution was reduced from 0.22 min*1 ( t h a i f - f i f e > 3
min) to 0.0114 min-1 ( t h a i f - i i f e » 60 m ' n ) when excess bicarbonate
versus CO2 was provided in solution. After initial experiments
showed that the addition of bicarbonate (50 mM) to reaction
mixtures had no effect on enzyme activity, the second set of
labelling experiments shown in Table 12 (in which CO2 was rapidily
analyzed and concentrated enzyme preparations were used
[Experiment 2]) were carried out.
Both m/z species 47 and 49 were again detected in reaction
mixtures supplied H21 80 after 1 minute. This time, however, the
ratio of isotopic species 45:47:49 (0.54:0.33:0.13) was different
from that expected on a theoretical basis assuming complete
exchange at equilibrium (0.28:0.50:0.22, based on 47% isotopic
enrichment). This finding implied that some biological labelling
70
from water had occurred. Parallel experiments with 1 802 suggested
that oxygen incorporation had also occurred. In this case, 10% label
was recovered in m/z species 47 ( 1 3 C 1 6 0 1 8 0) , which represented a
hundred-fold increase over that observed in long-term incubations
(Experiment 1). Thus, these experiments convincingly indicated that
labelling from molecular oxygen had occurred. The data with H2180
were less convincing because it was difficult to distinguish
between enzymatic and non-enzymatic (chemical exchange) labelling
because of the relatively low enrichment of H21sO (47%). Therefore,
experiments were repeated one more time (Experiment 3) with even
more concentrated enzyme preparations and the isotopic content of
CO2 determined after 20 seconds. Also, H2180 in experiment 3 was
enriched to 77%. The results with 1 802 were consistent with those
observed for Experiment 2, with approximately 10% of the labelled
CO2 recovered in m/z 47 species and no m/z 49. Interestingly, very
similar results were now also obtained with H2180, with m/z 47
representing almost 50% of labelled species and no 49 being
detected. These results provided strong evidence that in addition to
the incorporation of a single atom of oxygen from 1 802, one atom
was also incorporated from water. A further comparison of the
ratio for m/z 45:47;49 with those observed in Experiment 1 and 2 (in
which case significant non-enzymatic labelling due to chemical
exchange had occurred because of longer reaction times and the
absence of bicarbonate) revealed a significant difference. In this
case the ratio was 50:50:0, further indicating that m/z 47
(13c16o180) had been formed by enzymatic labelling.
71
The above findings suggested that one atom each of oxygen
from molecular oxygen and water were incorporated into cyanide.
We were concerned somewhat with the rather low recoveries of m/z
47 when 1 802 was supplied, since one might hypothesize that the
45:47:49 ratio should be close to 3:97:0 (at 97% enrichment).
However, the explanation for this may be related to the reaction
mechanism, a proposal for which is shown below. For example, if
initial incorporation from molecular oxygen occursto give an
NADH + H+ + 18O2 NAD+ + H2180 H2
16O NH3
V V V 'J HCN ^ ^ • X-180 ^ ^ • C180160
Scheme 5
unstable intermediate that is further hydrolyzed (by the large
excess of unlabelled water provided in 1 802 experiments), then the
amount of label capable of being recovered in CO2 would be expected
to be quite low as was observed.
72
PART III. PARTIAL RESOLUTION OF CYANIDE OXYGENASE
The identification of a cyanide oxygenase and putative cyanide
dihydratase/cyanide hydratase enzymes in cell-extracts was
consistent with findings reported previously (Kunz et al., 1992),
indicating that cyanide is metabolized by more than a single
mechanism in P. fluorescens NCIMB 11764. Because of the central
importance of CNO in cyanide assimilation, preliminary efforts were
made to purify this enzyme.
Recovery of cyanide oxygenase in ammonium sulfate
f r a c t i o n s
Initial studies reported by Harris and Knowles (1983b)
indicated that cyanide oxygenase activity was present in combined
0-35% and 35-70% ammonium sulfate fractions. These experiments
were repeated and similar results were obtained (Table 13); some
activity (30% of that observed for HSS) was also recovered in the
35-70% fraction. However, over 50% of the initial specific activity
detected in HSS could not be recovered in the combined 0-35% and
35-70% ammonium sulfate fractions. A similar loss in activity was
observed when HSS was subjected to simple dialysis (Table 13),
suggesting that an essential component was removed during
desalting of fractions by dialysis. In examining the data initially
reported by Harris and Knowles (1983b) a similar loss in activity
was noted, however, at the time no explanation for this loss was
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74
provided but we hypothesize that it may have been due to the
removal of the same dialyzable component.
Requirement of a small dialyzable component for cyanide
oxygenase
The involvement of a dialyzable factor in cyanide oxygenase
activity was further investigated by ultrafiltration. For this
purpose, 1 ml of HSS (12 mg ml"1 protein) was subjected to
ultrafiltration through a 30,000 molecular weight cut-off
Centricon® membrane after centrifugation at 5,000 x g for 90
minutes. The retentate (fraction F1, 0.08 ml [150 mg ml"1 protein])
and filtrate (fraction F2, 0.92 ml [0.55 mg ml"1 protein]) were then
assayed separately and in combination for cyanide-degrading
activity. As shown in Table 14, activity was detected in the
retentate (fraction F1), however, the specific activity was only 51%
of that observed for HSS. When the retentate and filtrate were
combined (F1 [1.2 mg protein] + F2 [50 |xg]) activity comparable to
that of HSS was again observed. Similarly, the addition of F2 to the
combined 0-35% and 35-70% fractions also restored complete
activity. These results further implied the involvement of a small
molecular weight (<30K) component in cyanide oxygenase activity.
Characterization of small dialyzable cyanide oxygenase
component
Further experiments were performed to identify and
characterize the F2 component. Since F2, in comparison with F1,
CO c o .mm
o CO
c o CO
CO U. 4— D
c
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76
contained very little protein (0.55 mg ml"1) it was concentrated 6
to 8 fold by lyophilization and rates of cyanide disappearance
measured as a function of F2 concentration. Figure 10 shows that
the rate of F1 -catalyzed cyanide disappearance was F2-
concentration dependent implying a catalytic role.
Table 15 shows a series of experiments performed in an effort
to characterize the F2 component. When F2 was heat treated, over
85% of the initial combined F1 + F2 activity was retained, indicating
that the active component was heat stable. In contrast, the
stimulatory effect of F2 was lost when subjected to acid hydrolysis
or protease treatment. Based on these observations it was
concluded that F2 contained a small molecular weight protein. To
determine its approximate size F2 fractions were subjected to
ultrafiltration and dialysis through 10,000 and 500 MW cut-off
membranes. Various fractions were then assayed in combination
with F1, and based on the results shown in Table 15, it was
concluded that the molecular size of the F2-containing protein was
between 500 and 10,000 Dalton.
77
Figure 10. Effect of F2 concentration on stimulation of cyanide
degrading activity of F1. Reaction mixtures contained 2 mM KCN, 4
mM NADH, F1 (1.2 mg protein) and different amounts of F2 (0-200 ^ig
protein) in a final volume of 0.25 ml.
78
4> O c (0 la.
<0 0) a a (0 (0 ^ =5 =
• I
! l >» w
a
<u (Q CC
100
100 150 200
F2 protein ( | ig)
79
Table 15. Effect of F2 treatment on cyanide oxygenase activity
Specific activity3
Fraction nmol KCN-degraded min"1 mg"1
HSS 18.1 (100)
F1 9.3 (51)
F1 + F2 (heated at 100°C/15 min) 15.5 (86)
F1 + F2 (treated with protease)*3 9.3 (51)
F1 + F2 (Acid hydrolysis)0 10.1 (56)
F1 + F2 (Amicon 10K filtrate) 17.2 (95)
F1 + F2 (500 MW cut-off dialyzed) 17.6 (97)
a Reaction mixtures contained 2 mM KCN, 4 mM NADH, HSS (4.8 mg
ml"1) or F1 (4.8 mg ml"1) and variously treated F2 (0.2 mg ml'1) as
indicated. bOne ml of F2 fraction (0.55 mg) was treated with Protease E
(Sigma) at 37°C for 18 h at which time treatment was stopped by
adding EDTA. c The F2 fraction was subjected to acid hydrolysis with 0.5 N HCI at
95°C for 15 minutes.
CHAPTER IV
SUMMARY
The recovery of several cyanide-degrading enzyme activities
in 150,000 x g cell-free extracts of P. fluorescens NCIMB 11764
was found to be consistent with work published previously from this
laboratory indicating that cyanide was metabolized by several
different pathways (Kunz et al., 1992). Results showing that one of
these activities (CDA1), catalyzed the formation of ammonia and
C0 2 and was both oxygen and NADH dependent was consistent with
earlier work suggesting that the enzyme responsible for this
conversion was a cyanide oxygenase (CNO) (Harris and Knowles,
1983b). However, in addition to this enzyme the present work
showed that a second activity (CDA2) composed of two enzymes, a
cyanide dihydratase (CND) (generating formate and ammonia) and a
cyanide hydratase (CNH) producing formamide was also elaborated by
this organism (since it was not possible to distinguish between one
or two different enzymes this activity was referred to as the
combined CND/CNH activity). CND/CNH activity as expected, could be
detected under anaerobic conditions and without added reduced
pyridine nucleotide factors. A comparison of the apparent Km
values toward cyanide revealed a remarkable difference between the
two, with the putative CNO enzyme showing a 10-fold higher
80
81
affinity for cyanide than CND/CNH. This suggested that the putative
cyanide oxygenase plays a significant role in cyanide assimilation as
a growth substrate. Separate studies in our laboratory showing that
a mutant strain having lost the ability to grow on cyanide could no
longer synthesize this enzyme further supported this conclusion
(Kunz et al., 1994). In addition to putative CNO and CND/CNH
enzymes cyanide also induced the synthesis of formate
dehydrogenase. The formation of this enzyme in response to cyanide
has previously not been described and a role for this enzyme in the
recycling of reduced pyridine nucleotide (NADH) has earlier been
discussed (Figure 8).
Because of the central role of CNO in cyanide assimilation,
particular attention was paid to the identification and unequivocal
verification that this enzyme was a true oxygenase. Since earlier
conclusions (Harris and Knowles, 1983b) that this enzyme was an
oxygenase were based primarily on the reaction stoichiometry,
related experiments were performed. Similar results as those
reported earlier were obtained; that is, one mole each of oxygen and
NADH being consumed per mole cyanide degraded as expected for an
oxygenase enzyme (Hayaishi, 1982). However, to obtain absolute
proof of this it was necessary to show that molecular oxygen was
incorporated during catalytic conversion using isotopically labelled 1 8 0 2 - As summarized in Part II of this dissertation, after
painstaking efforts were taken to limit the extent of isotopic
exchange, convincing evidence that molecular oxygen was
incorporated during catalysis was obtained (Table 12). Parallel
82
experiments showed that oxygen was also incorporated from
labelled water, indicating that one atom each from molecular oxygen
and water are incorporated during cyanide oxidation to CO2 and NH3.
This work is significant for the following reasons. First, it verifies
the actual existence of an oxygenase in cyanide conversion, which
previously could only be inferred from biochemical studies. Second
it shows that this enzyme is a monooxygenase since only one atom
of molecular oxygen is incorporated. Third, since oxygen from water
is incorporated, it infers that an additional step between the
substrate cyanide and the reaction products, CO2 and NH3 must exist.
As discussed in Part II, we hypothesize that this additional step may
involve the hydrolysis of a transient oxygenated intermediate.
Finally, since cyanide disappearance catalyzed by extracts induced
primarily for CNO was observed only when oxygen and NADH were
present it may be concluded that monooxygenation rather than the
addition of water (in which case substrate disappearance would be
expected to occur anaerobically) represents the initial step in
cyanide degradative attack.
Inhibitor studies revealed that cyanide oxidation by high-speed
supernatants was inhibited by classical sulfhydryl-group inhibitors
(see Table 9). This observation is important because the role of an
essential sulfhydryl group in the chemical transformation of related
substrates containing the C=N group have also been proposed (for
example, ricinine nitrilases [E.C. 3.5.5.1] and other nitrilases that
hydrolyze nitriles to carboxylic acids by the addition of 2 moles of
water) (Harper, 1977; Hook and Robinson, 1966; Mahadevan and
83
Thimann, 1964). By analogy, we propose that a related mechanism
involving nucleophilic attack on cyanide by an essential enzyme
sulfhydryl group followed by monooxygenation as shown in equation
may be involved. Oxygenation would be expected to give rise to the
H • C=N • NH
• H " C . + 0 2 + NADH + H*
XE
NH
H O C , 0=C
-H
XE \
( i )
XE
H M * 0=C. \
XE
H.0
H H S k
H - O - C - O H
I? XE
H,0
/O H + A' + BH + 0 = C N ( I I )
XE
H
0 = C &
& ^XE H
0 = C = 0 Carbon Dioxide
+ A" + BH + EXH <NI>
0=C
5 H. U
C»
AS
0=C = N Cyanate
+ A" + BH + EXH (IV)
Scheme 6
corresponding imino ether, which on hydrolysis (equation II) would
give a tetrahedral intermediate. Ammonia would be removed as a
84
consequence of the interaction of adjacent acidic and basic groups
to leave the carboxylated enzyme, which in turn could give rise to
C 0 2 and the regenerated enzyme (equation III). Alternatively, if the
enzyme serves as the initial leaving group then cyanate (HOCN)
would be generated (equation IV). However, no evidence for cyanate
as intermediate was obtained during this work or other
investigations conducted in this laboratory. Therefore, the latter
mechanism seems unlikely. Further experiments with purified
enzyme are, of course, required to confirm the proposed enzyme
mechanism. Part III of this dissertation was directed at resolving
the enzyme(s) responsible for cyanide oxidation. We now think this
system consists of a cyanide monooxygenase (CNO) and a possible
additional enzyme necessary for conversion of an as yet unidentified
oxygenated intermediate. Efforts to resolve this system resulted in
the identification of two protein fractions, one including the bulk
protein (>30K) fraction (F1) and the other a small molecular weight
protein fraction (0.5-1 OK) (F2) of as yet unknown nature. Thus, at
this point we think that at least two protein components are
required. Findings are analogous to other oxygenases which usually
are multicomponent in nature (Gibson, 1988; Whited and Gibson,
1991).
The primary goal of this dissertation was to begin biochemical
investigations of cyanide-degrading enzymes at the cell-free level
in P. fluorescens NCIMB 11764. This has been accomplished with
the recovery and preliminary characterization of enzymes by
identification of reaction products and using isotope-labelling
85
methods. The results show that cyanide metabolism in this
organism is more sophisticated than previously recognized. At least
four different enzymes involved in cyanide metabolism were
discovered in P. fluorescens NCIMB 11764. This includes, CNO,
CND, CNH, formate dehydrogenase, and an enzyme catalyzing the
possible hydrolysis of an oxygenated intermediate generated by
cyanide monooxygenase. Of these enzymes CNO has been shown to be
most important for cyanide growth. Additional studies on this and
the other enzymes synthesized by P. fluorescens NCIMB 11764
should reveal further insight into the unique biochemistry and
physiology of cyanide metabolism in this as well as other potential
cyanide-degrading microorganisms.
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