Characterization of two isozymes of coniferyl alcohol dehydrogenase from Streptomyces sp. NL15-2K

Article abstract of DOI:10.1271/bbb.110301

We purified two isozymes of coniferyl alcohol dehydrogenase (CADH I and II) to homogeneity from cellfree extracts of Streptomyces sp. NL15-2K. The apparent molecular masses of CADH I and II were determined to be 143 kDa and 151 kDa respectively by gel filtration, whereas their subunit molecular masses were determined to be 35,782.2 Da and 37,597.7 Da respectively by matrix-assisted laser-desorption ionization time-of-flight mass spectrometry (MALDI-TOF-MS). Thus, it is probable that both isozymes are tetramers. The optimum pH and temperature for coniferyl alcohol dehydrogenase activity were pH 9.5 and 45 °C for CADH I and pH 8.5 and 40 °C for CADH II. CADH I oxidized various aromatic alcohols and allyl alcohol, and was most efficient on cinnamyl alcohol, whereas CADH II exhibited high substrate specificity for coniferyl alcohol, and showed no activity as to the other alcohols, except for cinnamyl alcohol and 3-(4-hydroxy-3-methoxyphenyl)- 1-propanol. In the presence of NADH, CADH I and II reduced cinnamaldehyde and coniferyl aldehyde respectively to the corresponding alcohols.

Full text of DOI:10.1271/bbb.110301

Biosci. Biotechnol. Biochem., 75 (9), 1770–1777, 2011
Characterization of Two Isozymes of Coniferyl Alcohol Dehydrogenase
from Streptomyces sp. NL15-2K
1
;y
2
1
1
3
Motohiro NISHIMURA, Kunie KOHNO, Yoshio NISHIMURA, Masanori INAGAKI, and Julian DAVIES
1
Department of Pharmaceutical Chemistry, Faculty of Pharmacy, Yasuda Women’s University,
6
-13-1 Yasuhigashi, Asaminami-ku, Hiroshima 731-0153, Japan
Department of Medical Pharmacy, Faculty of Pharmacy, Yasuda Women’s University,
2
6
-13-1 Yasuhigashi, Asaminami-ku, Hiroshima 731-0153, Japan
Department of Microbiology and Immunology, Life Sciences Institute, University of British Columbia,
3
2
350 Health Sciences Mall, Vancouver, British Columbia V6T 1Z3, Canada
Received April 14, 2011; Accepted June 15, 2011; Online Publication, September 7, 2011
[doi:10.1271/bbb.110301]
We purified two isozymes of coniferyl alcohol dehy-
of these compounds are degraded to protocatechuic acid
or catechol, and further broken down via specific ring
cleavage pathways. For example, in Pseudomonas sp.
HR199, eugenol is degraded by side chain shortening to
protocatechuic acid via coniferyl alcohol, coniferyl
drogenase (CADH I and II) to homogeneity from cell-
free extracts of Streptomyces sp. NL15-2K. The apparent
molecular masses of CADH I and II were determined to
be 143 kDa and 151 kDa respectively by gel filtration,
whereas their subunit molecular masses were deter-
mined to be 35,782.2 Da and 37,597.7 Da respectively by
matrix-assisted laser-desorption ionization time-of-flight
mass spectrometry (MALDI-TOF-MS). Thus, it is
probable that both isozymes are tetramers. The opti-
mum pH and temperature for coniferyl alcohol dehy-
1
)
aldehyde, ferulic acid, vanillin, and vanillic acid, and
protocatechuic acid is further degraded by an ortho-
2
)
cleavage pathway. These sequential degradation path-
ways are found in a variety of bacterial species,
5
)
6)
including Acinetobacter,
,8)
Corynebacterium,
and
7
Streptomyces.
ꢀ
drogenase activity were pH 9.5 and 45 C for CADH I
ꢀ
Streptomycetes are gram-positive bacteria abundant
in soil. They are known to play important roles in
biotransformation and biodegradation in nature, but the
metabolic pathways involved, with the exception of
those for the biosynthesis of bioactive secondary
metabolites, have not been well studied. Studies done
on streptomycetes include several reports on the degra-
and pH 8.5 and 40 C for CADH II. CADH I oxidized
various aromatic alcohols and allyl alcohol, and was
most efficient on cinnamyl alcohol, whereas CADH II
exhibited high substrate specificity for coniferyl alcohol,
and showed no activity as to the other alcohols, except
for cinnamyl alcohol and 3-(4-hydroxy-3-methoxyphen-
yl)-1-propanol. In the presence of NADH, CADH I and
II reduced cinnamaldehyde and coniferyl aldehyde
respectively to the corresponding alcohols.
7
–10)
dation of lignin-related aromatic compounds.
Sutherland et al. described the processes by which
S. setonii catabolizes cinnamic acid, p-coumaric acid,
7
)
and ferulic acid. Bioconversion of vanillin to vanillic
acid and the catabolism of vanillic acid via guaiacol and
Key words: coniferyl alcohol dehydrogenase; cinnamyl
alcohol dehydrogenase; substrate specificity;
Streptomyces sp.
9
)
catechol were identified in S. viridosporus and S.
1
0)
setonii respectively. In addition, we have isolated
Streptomyces sp. NL15-2K from forest soil. We ob-
Lignins are cross-linked phenylpropanoid polymers.
They are the most abundant aromatic compounds in
nature. Partial decay of lignins produces numerous
aromatic monomers that have many applications in the
cosmetics, food, pharmaceutical, and chemical indus-
tries. Hence lignins have attracted considerable attention
as natural resources for the production of chemicals
traditionally derived from petroleum.
served that it can demethylate veratric acid to vanillic
8
)
acid.
Liquid chromatography-mass spectrometry
(LC-MS) analysis indicated that NL15-2K has an
extensive catabolic network for lignin-related aromatic
8
)
compounds (Fig. 1).
Although it is known that streptomycetes catabolize a
variety of lignin-related aromatic compounds, only a
few studies have attempted to characterize the catabolic
1
1–13)
In nature, lignins are assumed to be depolymerized
initially by white rot fungi, followed by mineralization
of the subsequent breakdown products by soil bacteria.
Bacterial degradation pathways for lignin-related aro-
matic compounds have been characterized extensively
for gram-negative bacteria, particularly Pseudomonas
pathways in question enzymatically or genetically.
Our project goal is to identify and elucidate the
characteristics of the enzymes and genes responsible
for the degradation of lignin-related aromatic com-
pounds in Streptomyces sp. NL15-2K.
Coniferyl alcohol is the principal precursor of lignin
biosynthesis in plants, and the sequential degradation of
spp.1 and Sphingomonas paucimobilis SYK-6. Most
–3)
4)
y
To whom correspondence should be addressed. Fax: +81-82-878-9540; E-mail: m-nishi@yasuda-u.ac.jp
Abbreviations: ADH, alcohol dehydrogenase; CAD, cinnamyl alcohol dehydrogenase; CADH, coniferyl alcohol dehydrogenase; DTT,
dithiothreitol; HPLC, high-performance liquid chromatography; LC-MS, liquid chromatography-mass spectrometry; MALDI-TOF-MS, matrix-
assisted laser-desorption ionization time-of-flight mass spectrometry; PAGE, polyacrylamide gel electrophoresis; PVDF, polyvinylidene difluoride

Coniferyl Alcohol Dehydrogenase from Streptomyces sp.
1771
CH=CHCH2OH
CH=CHCHO
CH=CHCOOH
CHO
OCH3
OH
OCH3
OH
OCH3
OH
OCH3
OH
Coniferyl alcohol Coniferyl aldehyde
Ferulic acid
Vanillin
CH=CHCOOH
COOH
OH
Cinnamic acid
OCH3
Isovanillic acid
COOH
COOH
OH
COOH
COOH
OCH3
COOH
OH
OCH3
OCH3
OH
OH
p-Hydroxybenzoic acid
Protocatechuic acid
Vanillic acid
Veratric acid
Benzoic acid
CH=CHCOOH
CH=CHCOOH
Ring fission
OH
OH
OCH3
OH
OH
OH
OH
p-Coumaric acid
Caffeic acid
Guaiacol
Catechol
Fig. 1. Degradation Pathways for Lignin-Related Aromatic Compounds by Streptomyces sp. NL15-2K.
Dotted arrows indicate unidentified steps.
lignin-related aromatic compounds in NL15-2K is
initiated by the oxidation of coniferyl alcohol to
coniferyl aldehyde, which is catalyzed by coniferyl
alcohol dehydrogenase (EC 1.1.1.194). Three CADH
assay conditions. The molar absorption coefficients at 400 nm, used in
ꢁ
1
ꢁ1
coniferyl aldehyde determination, were as follows: 13.9 mM ꢂcm at
ꢁ
1
ꢁ1
ꢁ1
ꢁ1
pH 8.0, 20.7 mM ꢂcm at pH 8.5, and 24.4 mM ꢂcm at pH 9.5.
Specific activity was expressed as units per mg of protein, where the
protein concentration was determined using a Bio-Rad Protein Assay
Kit (Bio-Rad, Hercules, CA) with bovine serum albumin as standard.
The rates of oxidation of the other aromatic alcohols and allyl alcohol
were determined by measuring the rate of production of NADH at
1
4,15)
16)
enzymes have been found in bacteria
and plants,
but there is little information on their properties. In
a preliminary study, we detected CADH activity in a
cell-free extract of NL15-2K. Here we describe the
purification and characterization of two isozymes of
CADH isolated from Streptomyces sp. NL15-2K.
3
40 nm (" ¼ 6:3 mMꢁ ꢂcm ), and the rate of cinnamaldehyde for-
1
ꢁ1
mation was calculated as described previously,17) using molar
absorption coefficients at 340 nm of NADH and cinnamaldehyde
ꢁ1
ꢁ1
ꢁ1
ꢁ1
at pH 9.5). To
(
0.48 mM ꢂcm
at pH 8.5 and 0.34 mM ꢂcm
determine the kinetic parameters of the purified isozymes, the initial
velocities at various substrate concentrations were determined by the
assay procedures described above, except that 0.1 M Tris–HCl buffer
Materials and Methods
(
pH 8.0) in the reaction mixture was replaced with 0.1 M glycine-
Materials and reagents. 3-(4-Hydroxy-3-methoxyphenyl)-1-propa-
nol was purchased from Tokyo Chemical Industry (Tokyo). Coniferyl
alcohol, cinnamyl alcohol, 3-phenyl-1-propanol, benzyl alcohol, allyl
alcohol, and the other aromatic alcohols used were from Wako Pure
Chemical Industries (Osaka, Japan). Coniferyl aldehyde and cinna-
maldehyde were purchased from Sigma-Aldrich (St. Louis, MO) and
NaOH buffer (pH 9.5) for CADH I and 0.1 M Tris–HCl buffer (pH 8.5)
for CADH II. The Km and kcat values were calculated from a Hanes-
Woolf plot.18) The rates of reduction of coniferyl aldehyde and
cinnamaldehyde to the corresponding alcohols were determined by
measuring the rates of decrease in the absorbance at 400 nm of
coniferyl aldehyde and at 340 nm of NADH and cinnamaldehyde,
respectively.
þ
Wako respectively. NAD(P) and NAD(P)H were from Oriental Yeast
(Osaka, Japan).
Microorganism culture conditions. Streptomyces sp. NL15-2K,
Purification of CADH isozymes. All steps of purification were
ꢀ
isolated from forest soil,8) was used throughout the study. The spores
were grown on inorganic salts-starch agar (ISP4; Difco Laboratories,
Detroit, MI), and then inoculated into and cultured in 100 mL of
YEME medium (1% glucose, 0.5% polypeptone, 0.3% yeast extract,
carried out at 4 C. Sodium phosphate buffer (25 mM, pH 7.0)
containing 1 mM dithiothreitol (DTT) was used as standard buffer in
the enzyme purification procedures, unless otherwise stated.
(i) Preparation of cell-free extract: Cells (46 g) harvested from 2 L
of culture were washed once with buffer I (standard buffer with 1 mM
EDTA and 1 M KCl), and then twice with buffer II (standard buffer
with 1 mM EDTA and 1 mM phenylmethylsulfonyl fluoride). The
washed cells were resuspended in 120 mL of buffer II and disrupted
using a sonicator (UD-201; TOMY, Tokyo). Cell debris was removed
by centrifugation at 20;000 ꢃ g for 30 min, and the supernatant was
collected as a cell-free extract.
.
ꢀ
0
4
1
.3% malt extract, and 0.04% MgCl2 6H2O, pH 7.0) at 30 C. After
8 h in liquid culture, 10% of the cultured mycelia was transferred to
00 mL of fresh YEME medium and incubated for an appropriate time
ꢀ
at 30 C with reciprocal shaking.
Assay for CADH activity. CADH activity was determined by
measuring the amount of coniferyl aldehyde formed from coniferyl
alcohol as substrate. The reaction mixture (0.3 mL) containing 0.1 M
(ii) DEAE-Sepharose column chromatography: The cell-free extract
was applied to a column (2:1 ꢃ 20 cm) of DEAE-Sepharose FF (GE
Healthcare Japan, Tokyo) equilibrated with standard buffer. The
column was washed thoroughly with the same buffer, and the proteins
were eluted with a linear gradient of 0 to 1 M NaCl. The active fractions
were pooled and dialyzed against standard buffer containing 0.7 M
þ
Tris–HCl (pH 8.0), 1.5 mM coniferyl alcohol, 2.5 mM NAD , and
ꢀ
variable amounts of enzyme, was incubated at 30 C for 10 min. The
formation of coniferyl aldehyde was monitored spectrophotometrically
at 400 nm. One unit was defined as the amount of the enzyme that led
to the formation of 1 mmol of coniferyl aldehyde per min under the

1772
M. NISHIMURA et al.
ammonium sulfate for CADH I and 1 M ammonium sulfate for CADH
II. The subsequent purification steps for the CADH isozymes were
performed separately with a fast protein liquid chromatography system
solution of sinapinic acid (Bruker Daltonics) in 30% acetonitrile with
0.1% trifluoroacetic acid. The mixture (1 mL) was deposited onto a
dried thin layer of the matrix on a MTP 384 target ground steel TF
plate (Bruker Daltonics) and allowed to dry at room temperature. The
measurements were performed in linear positive-ion mode with a
nitrogen laser under the control of FLEXControl software (version 2.4;
Bruker Daltonics). Spectra were obtained by accumulating up to 1,500
laser shots acquired at the minimum laser power necessary to ionize
the samples. The instrument was calibrated using Protein Calibration
Standard II (Bruker Daltonics).
(AKTApurifier 10; GE Healthcare).
(iii) Purification of CADH I: The dialysate was applied to a
RESOURCE PHE column (6 mL; GE Healthcare) equilibrated with
standard buffer containing 0.7 M ammonium sulfate. The column was
washed with 3 column volumes of the same buffer, and CADH I was
eluted with a 16-column-volume linear gradient of 0.7 to 0.1 M
ammonium sulfate at a flow rate of 1.0 mL/min. The active fractions
were pooled and concentrated by precipitation with 75% ammonium
sulfate, followed by centrifugation at 20;000 ꢃ g for 20 min. The
resulting pellet was resuspended and dialyzed against standard buffer
containing 20% glycerol. The dialysate was applied to a Mono Q 4.6/
N-Terminal amino acid sequencing. After SDS–PAGE, the proteins
in the gel were electroblotted onto a pre-wetted polyvinylidene
difluoride (PVDF) membrane (Sequi-Blot; Bio-Rad) using a Trans-
Blot semi-dry electrophoretic transfer cell (Bio-Rad) and stained with
Coomassie Brilliant Blue R-250. Protein bands of 37 kDa for CADH I
and 39 kDa for CADH II were excised from the membrane and
subjected to protein sequencing. The N-terminal amino acid sequences
of the CADH isozymes were determined by the automated Edman
method with a model 476A protein sequencer (Applied Biosystems,
Foster City, CA).
1
3
2
00 PE column (1.7 mL; GE Healthcare) equilibrated and washed with
column volumes of the same buffer. CADH I was eluted with a
0-column-volume linear gradient of 0 to 0.6 M NaCl at a flow rate
ꢀ
of 0.6 mL/min. The active fractions were pooled and stored at 4 C
until use.
(iv) Purification of CADH II: The dialysate was applied to a
RESOURCE PHE column (6 mL) equilibrated with standard buffer
containing 1 M ammonium sulfate, and washed with 3 column volumes
of the same buffer. After washing, CADH II was eluted with a 20-
column-volume linear gradient of 1 to 0.4 M ammonium sulfate at a
flow rate of 1.0 mL/min. The active fractions were pooled and
concentrated by precipitation with 75% ammonium sulfate. The
precipitate was recovered by centrifugation and dissolved in a small
volume of standard buffer, and the resulting solution was passed
through a HiLoad 16/60 Superdex 200 pg column (120 mL; GE
Healthcare) equilibrated with buffer containing 0.2 M NaCl at a flow
rate of 0.5 mL/min. The active fractions were pooled and supple-
mented with glycerol to 20%. The enzyme solution was then applied to
a Mono Q 4.6/100 PE column equilibrated with buffer containing 20%
glycerol and 0.2 M NaCl. The column was washed with 3 column
volumes of the same buffer, and the enzyme was eluted with a
Effects of pH and temperature on enzyme activity and stability. The
effect of pH on CADH activity was examined by the standard assay
method described above, except that the 0.1 M Tris–HCl buffer (pH 8.0)
in the reaction mixture was replaced with 0.1 M sodium acetate buffer
(
pH 4.0–5.5), 0.1 M sodium phosphate buffer (pH 5.5–7.5), 0.1 M Tris–
HCl buffer (pH 7.5–9.0), or 0.1 M glycine-NaOH buffer (pH 9.0–11.0).
The pH stabilities of CADH isozymes were assessed by measuring
residual activities after the enzymes were pre-incubated in the above
ꢀ
buffers at 4 C for 75 h. To investigate the effect of temperature on
CADH activity, reactions were carried out at temperatures ranging from
ꢀ ꢀ
2
5 C to 55 C for 10 min. The thermal stabilities of the CADH
isozymes were assessed by measuring residual activities after the
enzymes were pre-incubated in 25 mM sodium phosphate buffer
2
0
0-column-volume linear gradient of 0.2 to 0.4 M NaCl at a flow rate of
ꢀ
ꢀ
ꢀ
(
pH 7.0) at temperatures from 25 C to 60 C for 10 min.
.6 mL/min. The active fractions were pooled and stored at ꢁ20 C
until use.
Inhibition studies. The effects of metal ions and chemical reagents
on CADH activity were determined by pre-incubating the enzyme at
HPLC analysis. Oxidation of coniferyl alcohol to coniferyl alde-
ꢀ
3
7 C for 15 min in 25 mM sodium phosphate buffer (pH 7.0)
hyde by CADH isozymes was checked by high-performance liquid
chromatography (HPLC). After incubation for assay, the reaction
mixture was diluted 20-fold with water, and a 5-mL portion of the
dilute solution was subjected to HPLC using a Shim-pack XR-ODS
containing 1 mM of the various metal ions and reagents. Residual
activities after pre-incubation were assayed by the standard assay
method.
column (3:0 ꢃ 100 mm; Shimadzu, Kyoto, Japan) at a flow rate of
ꢀ
Results
0
.40 mL/min at 45 C. Elution was done with 30% methanol
containing 0.1% phosphoric acid, and was monitored by the absorption
at 220 nm and 400 nm. Compounds in the reaction mixture were
identified by comparison of the retention times (3.70 min for coniferyl
alcohol and 5.56 min for coniferyl aldehyde) with authentic standards
and comparison of the chromatograms at 220 nm and 400 nm.
Purification and properties of CADH I and II
Two isozymes of coniferyl alcohol dehydrogenase
CADH I and II) were separated by DEAE-Sepharose
(
ion exchange chromatography. CADH I and II were
eluted at NaCl concentrations of 0.25 M and 0.44 M
respectively (Fig. 2). Both isozymes were confirmed to
catalyze the oxidation of coniferyl alcohol to coniferyl
aldehyde by HPLC analysis. The active fractions from
DEAE-Sepharose column chromatography were purified
independently. CADH I was purified by hydrophobic
interaction chromatography on RESOURCE PHE and
ion exchange chromatography on Mono Q. CADH II
was purified by RESOURCE PHE, followed by gel
filtration on Superdex 200 pg, and then by Mono Q. The
results of the various courses of enzyme purification are
summarized in Table 1. CADH I and II were purified up
to 71-fold and 902-fold, with yields of 9% and 8%
respectively. Approximately 1.6 mg of CADH I and
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS–
PAGE). SDS–PAGE was performed on pre-cast 10% polyacrylamide
slab gels (e-PAGEL; Atto, Tokyo) under reducing conditions, and the
proteins were stained with Coomassie Brilliant Blue G-250 (Bio-Safe
Coomassie Stain; Bio-Rad). The molecular masses of the proteins
observed were estimated using a molecular size marker (EzStandard;
Atto).
Molecular mass determination. The molecular masses and quater-
nary structures of the CADH isozymes were determined by gel
filtration, SDS–PAGE, and MALDI-TOF-MS. The molecular masses
of the native isozymes were estimated by gel filtration on a HiLoad
16/60 Superdex 200 pg column equilibrated with standard buffer
containing 0.2 M NaCl at a flow rate of 0.5 mL/min. A gel filtration
calibration kit (GE Healthcare) for high-molecular-weight proteins,
containing catalase (232 kDa), aldolase (158 kDa), conalbumin
0.12 mg of CADH II were obtained per liter of culture.
(75 kDa), and ovalbumin (44 kDa), was used to provide approximate
In a series of purification steps, hydrophobic chroma-
tography on RESOURCE PHE resulted in high purifi-
cation of both isozymes. The final preparations of
CADH I and II gave single protein bands on SDS–
protein size markers. The MALDI-TOF mass spectra of the intact
isozymes were acquired on an Ultraflex TOF/TOF mass spectrometer
(Bruker Daltonics, Billerica, MA). Each CADH solution was dialyzed
against water and mixed with the matrix solution (1:4 v/v), a saturated

Coniferyl Alcohol Dehydrogenase from Streptomyces sp.
1773
2
1
1
0
5
0
5
0
0.8
0.6
0.4
0.2
0.0
kDa
1
2
3
II
1
9
6
7.2
6.4
I
0.5
0
4
5.0
29.0
0
10
20
30
40
50
Number of fraction
2
1
0.1
4.3
Fig. 2. Separation of CADH Isozymes by DEAE-Sepharose Column
Chromatography.
Cell-free extract from Streptomyces sp. NL15-2K was applied to a
DEAE-Sepharose column. Peaks I and II were collected as CADH I
Fig. 3. SDS–PAGE Analysis of Purified CADH Isozymes.
and CADH II respectively, and were further purified separately.
absorbance at 280 nm; , CADH activity; - - - -, NaCl concentration.
,
The enzyme (6 mg) from the final purification step was denatured,
reduced, and then subjected to SDS–PAGE. Lane 1, standard
molecular mass markers (sizes indicated); lane 2, CADH I; lane 3,
CADH II.
Table 1. Purification of CADH Isozymes from Streptomyces sp. NL15-2K
Total
activity
Total
protein
(mg)
Specific
activity
(units/mg)
Yield
%)
Purification
(-fold)
Step
(
(
units)
Cell-free extract
CADH I
56.2
2400
0.023
100
1
DEAE-Sepharose
RESOURCE PHE
Mono Q
20.8
12.9
5.2
378
8.5
3.2
0.055
1.5
1.6
37
23
9
2
66
71
CADH II
DEAE-Sepharose
RESOURCE PHE
Superdex 200
Mono Q
26.0
12.6
8.9
166
2.2
0.16
5.7
46
22
16
8
7
249
792
902
0.49
0.23
18.2
20.8
4.8
PAGE, with molecular masses of 37 kDa and 39 kDa
respectively (Fig. 3). The apparent molecular masses of
the native isozymes as measured by gel filtration were
accession no. ZP 05021059), and Gemmatimonas aur-
antiaca T-27 (RefSeq accession no. YP 002762200),
which have molecular masses of 35,428, 35,311, and
35,631 Da respectively. The N-terminal amino acid
sequence of CADH II was AQEVRGVIAPGKDEP-
VRM. A homology search revealed that the N-terminal
sequence of CADH II showed 94% similarity to those of
four oxidoreductases, from S. ambofaciens ATCC
23877 (Swiss-Prot accession no. A0ADE4), S. coelicolor
A3(2) (RefSeq accession no. NP 625045), S. ghanaensis
ATCC 14672 (RefSeq accession no. ZP 04684033),
and S. sviceus ATCC 29083 (RefSeq accession no.
ZP 05022334), a mycothiol-dependent formaldehyde
dehydrogenase from S. lividans TK24 (RefSeq acces-
sion no. ZP 06532967), and an ADH from S. coelicolor
A3(2) (Swiss-Prot accession no. Q9ZNB1), the molecu-
lar masses of which ranged from 37,616 to 37,868 Da.
The calculated molecular masses of the CADH homo-
logs were very close to those of the subunits of CADH I
(35,782 Da) and CADH II (37,598 Da), suggesting that
the amino acid sequences of the CADHs subunits are
very similar to those of their homologs. The enzymatic
properties of the homologs have not been described
to date.
1
43 kDa for CADH I and 151 kDa for CADH II (data not
shown), indicating that both isozymes are tetramers,
with four identical subunits. For accurate determination
of the molecular masses of the CADH isozymes, the
purified CADH proteins were analyzed by MALDI-
TOF-MS in linear positive-ion mode. The MALDI-TOF
mass spectrum of CADH I showed a major peak at m=z
3
5,782.197, whereas that of CADH II showed a major
peak at m=z 37,597.729 (Fig. 4). These molecular mass
values are roughly equal to those estimated by SDS–
PAGE, indicating that the two major peaks represented a
single subunit of the various CADH isozymes. A minor
peak at m=z 71,565.673 on the mass spectrum of CADH
I was assumed to be a molecular ion of the dimer form,
containing two subunits of 35,782.197 Da.
The 37- and 39-kDa proteins were blotted onto PVDF
membranes and subjected to N-terminal protein se-
quencing. Seventeen and 18 amino acids were obtained
from the N-terminal regions of CADH I and II
respectively. The N-terminal amino acid sequence of
CADH I was MKAAVVRAFGEPLVIEE. A BLAST
search revealed that the N-terminal sequence of CADH I
was identical to those of alcohol dehydrogenases
Effects of pH and temperature
The influence of pH on CADH activity and stability
was examined over a pH range of 4.0–11.0. The
(ADHs) from S. scabiei 87.22 (RefSeq accession
no. YP 003486809), S. sviceus ATCC29083 (RefSeq

1774
M. NISHIMURA et al.
1
7,964.100
2
,000
,000
0
CADH I
3
5,782.197
1
7
1,565.673
2
0,000
40,000
60,000
80,000
100,000 120,000 140,000 160,000
(
m/z)
4
00
00
0
CADH II
2
3
7,597.729
1
8,783.580
20,000
40,000
60,000
80,000
100,000
120,000
140,000
160,000
(
m/z)
Fig. 4. MALDI-TOF-MS Analysis of CADH I and II.
The purified enzymes were subjected to MALDI-TOF-MS analysis. The peaks at 17,964.100 for CADH I and 18,783.580 for CADH II
correspond to the double-charged molecular ions of their subunit proteins.
1
00
A
100
80
60
40
20
0
B
80
60
40
20
0
3
.0
5.0
7.0
9.0
11.0
3.0
5.0
7.0
9.0
11.0
pH
pH
1
00
100
80
60
40
20
0
D
C
80
60
40
20
0
20
30
40
50
20
30
40
50
60
Temperature (°C)
Temperature (°C)
Fig. 5. Effects of pH and Temperature on CADH Activity and Stability.
A, Optimum pH. CADH activities were measured in 0.1 M buffer solutions at various pH values under otherwise standard assay conditions.
The buffer systems used were sodium acetate buffer (pH 4.0–5.5), sodium phosphate buffer (pH 5.5–7.5), Tris–HCl buffer (pH 7.5–9.0), and
glycine-NaOH buffer (pH 9.0–11.0). After incubation for assay, the pH was adjusted to 8.0 with 1 M Tris–HCl buffer (pH 8.0), and the
absorbance at 400 nm was determined. The maximum activities, obtained at pH 9.5 for CADH I and at pH 8.5 for CADH II, were defined as
ꢀ
100%. B, pH stability. The enzyme solution was stored at 4 C for 75 h in 25 mM buffer solutions at various pH values, and residual activities
were measured by the standard assay method. The activity at the beginning was taken to be 100%. C, Optimum temperature. CADH activities
ꢀ
were measured at various temperatures under otherwise standard assay conditions. The maximum activities, obtained at 45 C for CADH I and at
ꢀ
4
0 C for CADH II, were defined as 100%. D, Thermal stability. The enzyme solution was pre-incubated at various temperatures for 10 min in
5 mM sodium phosphate buffer (pH 7.0), and residual activities were measured by the standard assay method. The activity without pre-
2
incubation was taken to be 100%. , CADH I; , CADH II.
optimum pHs for CADH I and II activities were 9.5 and
8
storing the enzymes at the indicated pH at 4 C and
monitoring changes in enzyme activity every 25 h for
75 h. Figure 5B shows the residual activities after 75 h.
CADH I was stable over a broad pH range of 5.0–9.5,
retaining over 80% of its activity, whereas CADH II was
stable in only a narrow pH range of 6.5–7.5 (Fig. 5B).
The optimum temperatures for CADH I and II activities
were 45 C and 40 C respectively (Fig. 5C). Thermal
stability was studied by measuring residual enzyme
activity after pre-incubation at various temperatures for
.5 respectively (Fig. 5A). pH stability was studied by
ꢀ
ꢀ
ꢀ

Coniferyl Alcohol Dehydrogenase from Streptomyces sp.
1775
Table 2. Effects of Metal Ions on CADH Activity Table 3. Effects of Chemical Reagents on CADH Activity
Residual activity (%)
Residual activity (%)
Metal ions (1.0 mM)
Reagents (1.0 mM)
CADH I
CADH II
CADH I
CADH II
None
100
100
93
99
94
74
80
0
100
98
90
67
23
30
9
None
Diisopropyl fluorophosphate
Dithiothreitol
100
100
105
90
100
100
117
94
81
0
MgCl2
CaCl2
MnCl2
FeCl2
FeCl3
ZnCl2
CuCl2
Phenylmethylsulfonyl fluoride
Ethylenediaminetetraacetic acid
Ethylenediaminetetraacetic acid (10 mM)
N-Ethylmaleimide
98
40
90
56
19
57
3
0
Monoiodoacetic acid
N-Bromosuccinimide
6
Table 4. Substrate Specificity of CADH Isozymes from Streptomyces sp. NL15-2K
CADH I
CADH II
Substrate
Km
kcat
kcat/Km
Km
kcat
ꢁ1
kcat/Km
(s 1)
ꢁ
(sꢁ1ꢂmM
ꢁ1
)
(mM)
(s
)
(sꢁ1ꢂmM
536
ꢁ1
)
(
mM)
Coniferyl alcohol
Cinnamyl alcohol
21.1
0.17
20.3
0.96
36.1
0.50
0.052
17.2
25.9
18.1
9.13
11.2
37.8
32.9
19.9
1.53
1.23
106
0.13
1.70
74.6
n.d.
n.d.
n.d.
0.40
n.d.
5.10
—
69.7
116
68.3
0.29
3
3
-(4-Hydroxy-3-methoxyphenyl)-1-propanol
-Phenyl-1-propanol
0.45
11.6
1.05
66.5
387
21.5
Benzyl alcohol
Allyl alcohol
þ
NAD
NADP
53.7
134
þ
0.089
Coniferyl aldehyde
Cinnamaldehyde
—
0.92
354
69.3
14.4
15.6
A dash indicates that values were not determined. n.d., not detectable.
1
0 min. At pH 7.0, CADH I and II were stable at
ꢀ
benzyl alcohol was recognized as a substrate by CADH
I, whereas vanillyl alcohol (4-hydroxy-3-methoxybenzyl
alcohol) was not oxidized (data not shown). This
suggests that CADH I favors aromatic alcohols without
ꢀ
temperatures of up to 50 C and 40 C, but their
activities decreased rapidly after incubation above
5
ꢀ
ꢀ
0 C and 40 C respectively (Fig. 5D).
4-hydroxy-3-methoxy groups. Allyl alcohol was the
Effects of metal ions and various reagents
We examined the effects of metal ions and other
next-best CADH I substrate after cinnamyl alcohol (3-
phenyl allyl alcohol). On the other hand, CADH II
exhibited a narrow substrate specificity, acting only on
coniferyl alcohol, cinnamyl alcohol, and 3-(4-hydroxy-
3-methoxyphenyl)-1-propanol. In contrast to CADH I,
CADH II displayed a k /K ratio for coniferyl alcohol
reagents on CADH activity. CADH I was completely
2
þ
3þ
inhibited by Cu and mildly inhibited by Fe and
þ
2
2þ
Zn , while CADH II was markedly inhibited by Cu ,
2
þ
2þ
3þ
2þ
Zn , Fe , and Fe , and mildly inhibited by Mn
Table 2). The other metal ions did not significantly
cat
m
(
that was 8-fold higher than that for cinnamyl alcohol.
Neither phenethyl alcohol (2-phenyl-1-ethanol) nor
homovanillyl alcohol (4-hydroxy-3-methoxyphenethyl
alcohol) were oxidized by either enzyme (data not
shown). Both CADH I and II catalyzed the reverse
reaction in the presence of NADH, reducing cinnamal-
dehyde and coniferyl aldehyde respectively to the
corresponding alcohols, but the k /K ratios of these
affect the activity of either CADH. Both enzymes were
strongly inhibited by N-bromosuccinimide and partially
inhibited by monoiodoacetic acid (Table 3). In addition,
CADH II was inhibited by N-ethylmaleimide. The
residual activities of CADH I and II were about 40%
and 0% respectively after pre-incubation with 10 mM
EDTA.
cat
m
enzymes were 7–8-fold lower for the aldehydes than for
the alcohols (Table 4), suggesting that CADH I- and II-
mediated reduction of the aldehydes was less efficient
than oxidation of the alcohols. The differences in
catalytic efficiencies were due mostly to the lower
affinities (increased Km values) of the enzymes for the
substrates.
Substrate specificity
Next we investigated the substrate specificities of
CADH I and II for various aromatic alcohols and allyl
alcohol. The kinetic parameters are summarized in
Table 4. CADH isozymes showed clear differences in
substrate preference. CADH I had higher affinity for
cinnamyl alcohol (Km ¼ 0:17 mM) and 3-phenyl-1-prop-
anol (Km ¼ 0:96 mM) than for coniferyl alcohol
The cofactor specificities of CADH I and II were
investigated with 1.5 mM of cinnamyl alcohol and
þ
(
Km ¼ 21:1 mM) or 3-(4-hydroxy-3-methoxyphenyl)-1-
coniferyl alcohol respectively as substrate. Both NAD
þ
propanol (K ¼ 20:3 mM). The k /K (catalytic effi-
and NADP supported CADH I activity, but a compar-
þ
m
cat
m
ciency) ratios for cinnamyl alcohol and 3-phenyl-1-
propanol were approximately 86- and 26-fold higher
than those for coniferyl alcohol and 3-(4-hydroxy-3-
methoxyphenyl)-1-propanol respectively. Similarly,
ison of the k /K ratios showed that NADP
m
cat
ꢁ1
ꢁ1
(0.089 s ꢂmM ) was a much less efficient cofactor
þ
ꢁ1
ꢁ1
for CADH I than NAD (387 s ꢂmM ). CADH II was
þ
active only in the presence of NAD .

1776
M. NISHIMURA et al.
Discussion
vary among these bacterial enzymes, although the
Rhodococcus CADH subunit has not been identified.
The molecular masses of plant CADHs are unknown,
because purified enzymes are not available. CADH II
exhibited a very narrow substrate specificity, and
oxidized only coniferyl alcohol, cinnamyl alcohol,
and 3-(4-hydroxy-3-methoxyphenyl)-1-propanol. Rho-
dococcus CADH demonstrated a similarly limited
substrate specificity and was active only on coniferyl
alcohol, cinnamyl alcohol, vanillyl alcohol, 4-(4-metho-
xyphenyl)-1-butanol, and 3-(3,4-dimethoxyphenyl)-1-
propanol. Coniferyl alcohol was the best substrate for
the enzymes from both Streptomyces and Rhodococcus,
and its Km values were 0.13 mM for CADH II and
In this study, we purified two isozymes (CADH I and
II) of coniferyl alcohol dehydrogenase from cell-free
extracts of Streptomyces sp. NL15-2K. Substrate speci-
ficity studies with aromatic alcohols and allyl alcohol
revealed that CADH I oxidized many of the alcohols
tested, whereas CADH II oxidized only coniferyl
alcohol, cinnamyl alcohol, and 3-(4-hydroxy-3-metho-
xyphenyl)-1-propanol. A comparison of Km values
showed that the highest substrate affinities of CADH I
and II were for cinnamyl alcohol and coniferyl alcohol
respectively. Similarly, the kcat/Km ratios indicated
that cinnamyl alcohol and coniferyl alcohol were the
preferred substrates of CADH I and II respectively.
These results suggest that CADH I should be considered
a cinnamyl alcohol dehydrogenase rather than a con-
iferyl alcohol dehydrogenase.
0.65 mM for Rhodococcus CADH. Both enzymes were
þ
also NAD -dependent, and the K values of CADH II
m
þ
and Rhodococcus CADH for NAD were 0.40 mM and
0.22 mM respectively. In contrast, plant CADH activ-
þ
Cinnamyl alcohol dehydrogenases (CADs; EC
.1.1.195) also catalyze the reverse reaction, the
ities were detected only with NADP in 86 of the 89
1
6)
1
plants species tested.
The N-terminal amino acid
reduction of cinnamaldehyde to cinnamyl alcohol. A
number of CADs in plants catalyze the reduction of
p-hydroxycinnamaldehydes to their corresponding
alcohols; this is the final step in the biosynthesis of
sequence of Pseudomonas CADH was found to be
MQLTNKKIVVV. This sequence shows no significant
homology with the N-terminal sequence of CADH II.
A homology search revealed that the N-terminal amino
acid sequence of the CADH II subunit showed 94%
similarity to those of four different oxidoreductases,
a mycothiol-dependent formaldehyde dehydrogenase,
and an ADH. These are all from Streptomyces spp.
and have not been enzymatically characterized. How-
ever, as described in ‘‘Results’’ above, the molecular
masses of these homologs were almost equal to that of
CADH II subunit. Judging from these results, CADH II
might be closely related to these homologs. To
evaluate the similarities in amino acid sequence
between CADH II and its homologs, it will be
necessary to analyze further the amino acid sequence
of the CADH II subunit.
monolignols before their polymerization in the cell
Outside of the plant kingdom, CAD enzymes
walls.1
9,20)
have been characterized from three microorganisms,
2
1)
22,23)
Helicobacter pylori, Mycobacterium bovis BCG,
2
4)
and Saccharomyces cerevisiae.
These enzymes as
well as plant CADs show strong substrate preferences
for aldehydes to their corresponding alcohols and high
dependence on NADP(H). For example, the kcat/Km
ratios of the microbial CADs for cinnamaldehyde and
cinnamyl alcohol respectively are as follows: 5,480 and
ꢁ
1
ꢁ1
ꢁ1
ꢁ1
ꢁ1
ꢁ1
1
26 s ꢂmM for H. pylori CAD; 42 and 0.2 s ꢂmM
for M. bovis BCG CAD; and 3,035 and 706 s ꢂmM
for S. cerevisiae CAD. In contrast, the kcat/Km ratios of
CADH I for cinnamaldehyde and cinnamyl alcohol were
CADH I and II were unstable in buffers without
DTT and were sensitive to thiol-blocking reagents such
as N-ethylmaleimide and monoiodoacetic acid, as well
ꢁ1
ꢁ1
15.6 and 106 s ꢂmM
showed a marked preference for NAD , the k /K
respectively. CADH I also
þ
cat
m
þ
2þ
2þ
ratio being 4,348-fold higher than for NADP . These
results indicate that CADH I is different from previously
described CAD enzymes. In addition, the N-terminal
amino acid sequence (17 amino acids) of the CADH I
subunit showed identity with those of three ADHs from
Streptomyces and Gemmatimonas, but no significant
homology with those of plant and microbial CADs.
Hence CADH I was assumed to be an ADH with a high
substrate preference for cinnamyl alcohol.
as to divalent heavy metal ions Zn and Cu . This
suggests that their active sites contain a sulfhydryl
group. In streptomycetes, mycothiol is the major thiol
and plays a key role in maintaining a reducing
2
5)
environment in the cells. Therefore, the high sim-
ilarity in N-terminal amino acid sequence between the
CADH II subunit and a mycothiol-dependent form-
aldehyde dehydrogenase suggests the presence of a
sulfhydryl group in the active site of CADH II. CADH
I and II were also markedly inhibited by N-bromosuc-
cinimide, suggesting that a tryptophan residue is also
present in the active sites of these enzymes. In the
presence of 10 mM EDTA, the activity of CADH I was
reduced to 40% of the baseline, and CADH II was
completely inhibited. Therefore, some metal ions are
assumed to be involved in the enzymatic activities or
structural stabilities of CADH I and II. There are three
major classes of microbial alcohol dehydrogenases as
In contrast to CAD enzymes, little is known about
the characteristics of coniferyl alcohol dehydrogenases.
To date, only three CADH enzymes have been
1
4)
reported, from Rhodococcus erythropolis,
Pseudo-
1
5)
16)
monas sp. HR199, and plants, but a comparison of
enzymatic properties among these CADHs and CADH
II is difficult, because there is little information on the
properties of previously reported CADHs. The mo-
lecular masses of the native CADHs from Streptomyces
2
6)
þ
(
CADH II), Rhodococcus, and Pseudomonas were
51 kDa, 200 kDa, and 54.9 kDa, respectively. The
CADH from Pseudomonas is a dimer composed of two
categorized by cofactor specificities: (i) NAD(P) -
dependent ADHs, which are further sub-divided into
zinc-dependent ADHs, short-chain zinc-independent
1
þ
identical subunits with individual molecular masses of
27 kDa, whereas CADH II is tetrameric. This suggests
that the molecular masses and quaternary structures
ADHs, and iron-activated ADHs; (ii) NAD(P) -inde-
pendent enzymes, which use pyrroloquinoline quinone,
heme, or cofactor F420; and (iii) FAD-dependent

Coniferyl Alcohol Dehydrogenase from Streptomyces sp.
1777
ADHs. Since both CADH I and II showed marked
þ
7) Sutherland JB, Crawford DL, and Pometto III AL, Can. J.
Microbiol., 29, 1253–1257 (1983).
8) Nishimura M, Ooi O, and Davies J, J. Biosci. Bioeng., 102, 124–
preference for NAD , they should be classified as
þ
members of the NAD -dependent group of ADHs.
However, further information on the amino acid
1
27 (2006).
Pometto III AL and Crawford DL, Appl. Environ. Microbiol.,
5, 1582–1585 (1983).
9
)
sequences and/or sensitivities to Zn2 or Fe
CADH I and II are required to assign them to one of
þ
2þ
of
4
10) Pometto III AL, Sutherland JB, and Crawford DL, Can. J.
Microbiol., 27, 636–638 (1981).
þ
the NAD(P) -dependent ADH sub-groups.
1
1
1
1) Chow KT, Pope MK, and Davies J, Microbiology, 145, 2393–
403 (1999).
2) Iwagami SG, Yang K, and Davies J, Appl. Environ. Microbiol.,
6, 1499–1508 (2000).
In this study, we obtained two dehydrogenases that
showed different substrate specificities for aromatic
alcohols and allyl alcohol. These enzymes performed
2
6
2
-way conversions between coniferyl alcohol or cin-
3) Nishimura M, Ishiyama D, and Davies J, Biosci. Biotechnol.
Biochem., 70, 2316–2319 (2006).
14) Jaeger E, Methods Enzymol., 161, 301–306 (1988).
namyl alcohol and the corresponding aldehydes. We are
in the process of cloning the genes encoding the CADH I
and II from the chromosomal DNA of Streptomyces sp.
NL15-2K to characterize these enzymes further.
15) Steinb u¨ chel A and Priefert H, Japan Kokai Tokkyo Koho, H10-
1
55496 (June 16, 1998).
6) Mansell RL, Babbel GR, and Zenk MH, Phytochemistry, 15,
849–1853 (1976).
7) Wyrambik D and Grisebach H, Eur. J. Biochem., 59, 9–15
1975).
1
1
1
Acknowledgments
(
J. Davies thanks the National Science and Engineer-
ing Council of Canada for support of his work at
University of British Columbia.
18) Hanes CS, Biochem. J., 26, 1406–1421 (1932).
19) Barakat A, Bagniewska-Zadworna A, Choi A, Plakkat U,
DiLoreto DS, Yellanki P, and Carlson JE, BMC Plant Biol., 9,
2
0) Ma QH, J. Exp. Bot., 61, 2735–2744 (2010).
6 (2009).
2
2
References
1) Mee B, Kelleher D, Frias J, Malone R, Tipton KF, Henehan GT,
and Windle HJ, FEBS J., 272, 1255–1264 (2005).
1
)
Overhage J, Priefert H, and Steinb u¨ chel A, Appl. Environ.
Microbiol., 65, 4837–4847 (1999).
22) de Bruyn J, Johannes A, Weckx M, and Beumer-Jochmans MP,
J. Gen. Microbiol., 124, 359–363 (1981).
23) Wilkin JM, Soetaert K, St e´ landre M, Buyssens P, Castillo G,
Demoulin V, Bottu G, Laneelle MA, Daffe M, and de Bruyn J,
Eur. J. Biochem., 262, 299–307 (1999).
2
)
Overhage J, Kresse AU, Priefert H, Sommer H, Krammer G,
Rabenhorst J, and Steinb u¨ chel A, Appl. Environ. Microbiol., 65,
9
51–960 (1999).
Bertani I, Kojic M, and Venturi V, Microbiol., 147, 1611–1620
2001).
3)
4)
5)
6)
(
24) Larroy C, Par e´ s X, and Biosca JA, Eur. J. Biochem., 269, 5738–
5745 (2002).
Masai E, Katayama Y, and Fukuda M, Biosci. Biotechnol.
Biochem., 71, 1–15 (2007).
25) Newton GL, Arnold K, Price MS, Sherrill C, delCardayre SB,
Aharonowitz Y, Cohen G, Davies J, Fahey RC, and Davis C,
J. Bacteriol., 178, 1990–1995 (1996).
Delneri D, Degrassi G, Rizzo R, and Bruschi CV, Biochim.
Biophys. Acta, 1244, 363–367 (1995).
Merkens H, Beckers G, Wirtz A, and Burkovski A, Curr.
Microbiol., 51, 59–65 (2005).
26) Reid MF and Fewson CA, Crit. Rev. Microbiol., 20, 13–56
(1994).