Cineole biodegradation: Molecular cloning, expression and characterisation of (1R)-6β-hydroxycineole dehydrogenase from Citrobacter braakii

Article abstract of DOI:10.1016/j.bioorg.2009.12.003

The first steps in the biodegradation of 1,8-cineole involve the introduction of an alcohol and its subsequent oxidation to a ketone. In Citrobacter braakii, cytochrome P450_cin has previously been demonstrated to perform the first oxidation to produce (1R)-6β-hydroxycineole. In this study, we have cloned cinD from C. braakii and expressed the gene product, which displays significant homology to a number of short-chain alcohol dehydrogenases. It was demonstrated that the gene product of cinD exhibits (1R)-6β-hydroxycineole dehydrogenase activity, the second step in the degradation of 1,8-cineole. All four isomers of 6-hydroxycineole were examined but only (1R)-6β-hydroxycineole was converted to (1R)-6-ketocineole. The (1R)-6β-hydroxycineole dehydrogenase exhibited a strict requirement for NAD(H), with no reaction observed in the presence of NADP(H). The enzyme also catalyses the reverse reaction, reducing (1R)-6-ketocineole to (1R)-6β-hydroxycineole. During this study the N-terminal His-tag used to assist protein purification was found to interfere with NAD(H) binding and lower enzyme activity. This could be recovered by the addition of Ni²⁺ ions or proteolytic removal of the His-tag.

Full text of DOI:10.1016/j.bioorg.2009.12.003

Bioorganic Chemistry 38 (2010) 81–86
Contents lists available at ScienceDirect
Bioorganic Chemistry
journal homepage: www.elsevier.com/locate/bioorg
Cineole biodegradation: Molecular cloning, expression and characterisation
of (1R)-6b-hydroxycineole dehydrogenase from Citrobacter braakii
*
Kate E. Slessor, Jeanette E. Stok, Sonia M. Cavaignac, David B. Hawkes, Younes Ghasemi, James J. De Voss
School of Chemistry and Molecular Biosciences, University of Queensland, St. Lucia, Brisbane QLD 4072, Australia
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 11 November 2009
Available online 16 December 2009
The first steps in the biodegradation of 1,8-cineole involve the introduction of an alcohol and its subse-
quent oxidation to a ketone. In Citrobacter braakii, cytochrome P450cin has previously been demonstrated
to perform the first oxidation to produce (1R)-6b-hydroxycineole. In this study, we have cloned cinD from
C. braakii and expressed the gene product, which displays significant homology to a number of short-
chain alcohol dehydrogenases. It was demonstrated that the gene product of cinD exhibits (1R)-6b-
hydroxycineole dehydrogenase activity, the second step in the degradation of 1,8-cineole. All four iso-
mers of 6-hydroxycineole were examined but only (1R)-6b-hydroxycineole was converted to (1R)-6-
ketocineole. The (1R)-6b-hydroxycineole dehydrogenase exhibited a strict requirement for NAD(H), with
no reaction observed in the presence of NADP(H). The enzyme also catalyses the reverse reaction, reduc-
ing (1R)-6-ketocineole to (1R)-6b-hydroxycineole. During this study the N-terminal His-tag used to assist
protein purification was found to interfere with NAD(H) binding and lower enzyme activity. This could be
recovered by the addition of Ni2+ ions or proteolytic removal of the His-tag.
Keywords:
Oxidoreductase
Cineole
Biodegradation
Enantiospecificity
Terpene
Citrobacter braakii
Histidine-tag
Ó 2009 Elsevier Inc. All rights reserved.
1. Introduction
ways and enzymology of cineole biodegradation however, are not
well documented.
Terpenes constitute a large class of natural products and play
important roles in chemical ecology including acting as repellents,
attractants, toxins and sex or alerting pheromones [1]. Monoter-
penes (C10) are major components of plant oils and are structurally
diverse, encompassing acyclic, monocyclic and bicyclic com-
pounds. The generation of oxygenated terpenoid derivatives is usu-
ally initiated by a cytochrome P450 monooxygenase, with this
functionalization forming part of either biosynthetic or biodegra-
dative pathways [2]. Cineole, a monoterpene, is one of the more
widespread and abundant chemicals in the Australian bush. It is
a component of many essential oils and is the dominant compo-
nent of the oil from many eucalypts. It is estimated that more than
500,000 tonnes of cineole are produced and released into the Aus-
tralian environment annually by the eucalypt population [3]. The
role of cineole remains unclear, although it is believed to have a
number of different functions including as a deterrent to herbi-
vores and pathogens, and attracting pollinators. A large proportion
of the cineole released into the environment is believed to be re-
moved by microbial degradation, with lesser quantities consumed
by native wildlife (koalas, possums) and bushfires [4,5]. The path-
A metabolic pathway for the degradation of cineole was pro-
posed [6] following the analysis of cineole metabolites from two
soil microbes: Pseudomonas flava [4] and a Rhodococcus species
[7] (Fig. 1). Interestingly, the two species are believed to metabo-
lize cineole (Fig. 1, 1) via chemically identical but enantiomerically
different pathways. Thus, the initial step is believed to be hydrox-
ylation at either the pro-S or pro-R carbon of cineole to furnish 6b-
hydroxycineole1 (Fig. 1, 2a). This is then followed by oxidation to
give the corresponding ketone, 6-ketocineole (Fig. 1, (1R) 3a). Fur-
1
Nomenclature footnote: The nomenclature of the hydroxy cineoles is complex
and there is no consistency in the literature for naming such compounds. We use the
descriptors
a and b in order to discuss stereochemistry. The a descriptor describes
substituents below the plane passing through C5, C6, C7 and C8 while b describes
substituents above this plane. As cineole is an achiral, meso compound hydroxylation
at either of the carbons flanking the C1 bridgehead leads to enantiomeric products
and the creation of three new stereogenic centres at C1, C4 and C6. We therefore have
defined the carbon atom that, following oxidation, leads to the R-C1 isomer as the
pro-R carbon and the carbon atom that leads to the S-C1 isomer as the pro-S carbon.
11
10
3
2
O
8
5
4
β-substituent
Pro-R
1
Pro-S
α-substituent
7
Abbreviations: PAGE, Polyacrylamide gel electrophoresis; His-tag, Hexahisti-
dine-tag; P450, Cytochrome P450.
6
9
* Corresponding author. Fax: +61 7 3365 4273.
E-mail address: j.devoss@uq.edu.au (J.J. De Voss).
0045-2068/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved.
doi:10.1016/j.bioorg.2009.12.003

82
K.E. Slessor et al. / Bioorganic Chemistry 38 (2010) 81–86
via subcloning small internal fragments and sequencing using
M13 forward and reverse primers. Any gaps that remained in the
sequence were determined by using appropriate synthetic primers.
All sequencing was analysed using the BLAST algorithm (http://
blast.ncbi.nlm.nih.gov/Blast.cgi). The open reading frame cinD
was sequenced completely in both directions.
O
O
O
HO
O
1
2a
3a
2.3. Cloning cinD into proEX HTa
O
HO
O
O
Primers were designed to either end of cinD containing EcoRI and
NdeI sites at the 50 end (50-CAGAATTCCATATGACCGGCAGAC-
TCGAAGG-30) and EcoRI and HindIII sites at the 50 end (50-AAGAATT-
CAAGCTTCCTACCCGTCGACGACAGC-30). cinD was amplified from
pC1 employing the following conditions: 94 °C for 30 s, 59 °C for
45 s, 72 °C for90 s. The0.7 kbDNA fragmentwasdigestedwithEcoRI
and HindIII and cloned into similarly cut pPROEX HTa (Invitrogen,
Mount Waverley, VIC) to give pPROEX-cinD. This construct was ver-
ified by sequencing (AGRF, Brisbane QLD).
O
O
Central metabolism
O
HO
O
HO
HO
O
O
O
OH
O
O
2.4. Expression and purification of CinD ((1R)-6b-hydroxycineole
dehydrogenase)
Fig. 1. Proposed pathway of 1,8-cineole degradation [6]. The first step has shown to
be catalysed by P450cin in Citrobacter braakii. The step that (1R)-6b-hydroxycineole
dehydrogenase (CinD) is proposed to catalyse is boxed.
Terrific broth containing ampicillin (50
with pPROEX-cinD-transformed E. coli DH5
was incubated at 37 °C until an OD600 of 0.6 was attained. Isopropyl
b- -thiogalactopyranoside (IPTG; 1 mM final concentration) was
l
g/mL) was inoculated
a
cells. The culture
ther transformations are somewhat speculative but are believed to
produce a highly functionalized hydroxy keto acid that can enter
central metabolism, thereby enabling the organism to live on cineole
as its sole source of carbon and energy [7].
D
then added and the culture incubated at 27 °C for 18 h. All subse-
quent procedures were performed at 4 °C unless otherwise stated.
The cells were harvested by centrifugation at 4000g for 20 min and
resuspended in Buffer A (50 mM Tris–HCl, pH 7.4, 1 mM EDTA,
0.5 mM DL-dithiothreitol). Lysis was carried out by sonication on
ice (Branson sonifer 450; 6 ꢀ 1 min intervals, 30% output) and
the cellular debris was removed by centrifugation (10,000g, 1 h).
The supernatant was loaded onto a Ni2+-chelating column (1 mL,
HiTrap Chelating HP; GE Healthcare, Rydalmere, NSW) that had
been equilibrated with Buffer B (20 mM phosphate buffer, 0.5 M
NaCl, 10 mM imidazole, pH 7.4). The column was washed with
10 column volumes of Buffer B and the dehydrogenase eluted in
Buffer B containing 100–250 mM imidazole. Fractions were ana-
lysed by SDS–PAGE (NuPage 4–12% Bis–Tris gels; Invitrogen) and
those containing a single band at ꢁ30 KDa were combined and
concentrated. The dehydrogenase was loaded onto a G-25-desalt-
ing column (1 ꢀ 6 cm) and eluted in buffer (50 mM Tris–HCl pH
7.4). The eluted protein was stored at ꢂ70 °C. Protein concentra-
tion was determined using the Bio-Rad Protein Assay (Bio-Rad,
Gladesville, NSW) with bovine serum albumin as the standard.
To date, only two enzymes implicated in cineole biodegradation
have been successfully cloned, expressed and purified: cytochrome
P450cin and its immediate FMN-containing redox partner, cindox-
in. In the presence of an exogenous electron transport partner
P450cin and cindoxin catalyze the hydroxylation of cineole to
(1R)-6b-hydroxycineole (2a) [8,25]. The genes encoding these pro-
teins were isolated from the CIN operon of the soil microbe, Citro-
bacter braakii, which is capable of living on cineole as its sole
source of carbon and energy. The next step in cineole degradation
is believed to be oxidation of the secondary alcohol to the corre-
sponding ketone (Fig. 1, boxed). It has previously been observed
that proteins encoded by both the CAM operon from Pseudomonas
putida and the TERP operon from Pseudomonas sp. are responsible
for the initial degradation of camphor and a-terpineol, respectively
[9–11]. In both these operons an open reading frame encoding an
alcohol dehydrogenase was found which is responsible for the oxi-
dation of the alcohol produced following the initial P450 reaction.
We thus set out to determine whether a gene encoding a dehydro-
genase was also present in, or adjacent to, the CIN operon and, once
isolated, clone the gene and express and characterise the gene
product in order to investigate its role in cineole metabolism.
2.5. Cleavage of His-tag from (1R)-6b-hydroxycineole dehydrogenase
(1R)-6b-hydroxycineole dehydrogenase (34 lM) was added to a
solution containing Buffer A and AcTEV™ protease (200 U; Invitro-
gen) and incubated at 4 °C for 16 h. The solution was then diluted
5-fold in Buffer B before loading onto a Ni2+-chelating column
(1 mL, HiTrap Chelating HP) that had been equilibrated with Buffer
B. The flow-through (containing cleaved native enzyme) was col-
lected and concentrated. Excess NaCl/imidazole was removed by
loading the protein onto a G-25-desalting column (1 ꢀ 6 cm) and
eluting in buffer (50 mM Tris–HCl pH 7.4). The cleaved protein
was stored at ꢂ70 °C.
2. Materials and methods
2.1. Chemicals
The four isomers of 6-hydroxycineole (2) and two isomers of 6-
ketocineole (3) were prepared from enantiomerically enriched
terpineol according to the method of Carman and Fletcher [12]
with the stereochemistry of the starting -terpineol determining
a-
a
which enantiomer is formed. All other chemicals used in this study
were analytical grade reagents.
2.6. Assay for (1R)-6b-hydroxycineole dehydrogenase activity: NAD(H)
detection/consumption
2.2. Sequencing of cinD
Oxidation of (1R)-6b-hydroxycineole (2a) was detected by mea-
Sequence analysis of clone pC1 (pUC19 containing the original
suring the rate of NADH formation. The assay, containing mixture
4 kb PstI fragment isolated from C. braakii [8]) was carried out
of the dehydrogenase (0.5–10 lM) and (1R)-6b-hydroxycineole

K.E. Slessor et al. / Bioorganic Chemistry 38 (2010) 81–86
83
(2a) (60 mM stock solution in ethanol, final concentration:
600 M) in buffer (50 mM Tris–HCl pH 7.4), was initiated by the
addition of NAD+ (6 mM; 340nm = 6200 Mꢂ1cmꢂ1). The reaction
3. Results
l
e
3.1. Operon analysis
was monitored by UV spectroscopy, measuring the appearance of
NADH at 340 nm. All assays were performed at room temperature
(typically 25 °C) and ethanol concentration never exceeded 1%. The
oxidation of other substrates, such as androsterone was measured
in an analogous fashion. To measure the reverse reaction (reduc-
tion) the rate of NADH consumption was monitored by the de-
crease in absorbance at 340 nm.
Following the sequencing of pC1, a fourth open reading frame
(cinD; Genbank ID: GQ849481) was identified. The cinD gene is
divergently transcribed with respect to cinA (P450cin), cinB and cinC
(cindoxin), three ORFs that were identified during previous analy-
sis of the CIN operon (Fig. 2) [8]. This ORF (cinD) displays signifi-
cant homology to
a
number of short-chain alcohol
The NADH-detection assay was used to examine the effect of
metal ions on the dehydrogenase activity. The assay was run as de-
scribed above with the inclusion of the following metal ions at
1 mM: Zn2+ (ZnCl2), Cu2+ (CuCl2), Co2+ (CoCl2), Ca2+ (CaCl2), Mg2+
(MgCl2), Mn2+ (MnCl2) and Ni2+ (NiSO4).
dehydrogenases (Table 1).
3.2. Subcloning and expression of CinD ((1R)-6b-hydroxycineole
dehydrogenase)
Initially, expression of CinD in E. coli as the wild type enzyme
(i.e. with no His-tag present) was attempted. These efforts were
unsuccessful and consequently, an alternative strategy to express
CinD was evaluated. PCR was used to amplify cinD employing
primers that introduced EcoRI and HindIII restriction sites and
facilitated subsequent cloning of the gene into the expression vec-
tor pPROEX HTa. pPROEX HTa includes a coding region for a hexa-
histidine N-terminal affinity tag (His-tag) to assist in protein
2.7. Assay for (1R)-6b-hydroxycineole dehydrogenase activity:
enantioselective GC
2.7.1. Oxidation of 6-hydroxycineole (2) to 6-ketocineole (3)
All four isomers of 6-hydroxycineole (2a–d) were assayed. 6-
hydroxycineole (2) (6 mM) was added to a solution of (1R)-6b-
hydroxycineole dehydrogenase (8 lM) in buffer (50 mM Tris–HCl
pH 7.4) and stirred at room temperature for 10 min before the
addition of NAD+ (1 mM). The reaction was left stirring for 2 h, ex-
tracted with ethyl acetate and the organic extract dried over
MgSO4. The ethyl acetate was removed in vacuo and the product
resuspended in dichloromethane. Several drops of trifluoroacetic
anhydride were added to the solution and the sample was then
concentrated in vacuo. The derivatized product was dissolved in
diethyl ether and analysed by enantioselective GC. Briefly, this
was performed on a gas chromatograph fitted with a b-cyclodex-
trin enantioselective column under the following conditions:
120 °C for 2 min, 1 °C/min to 200 °C. Retention times were com-
pared with synthetic standards [12] that had been identically
derivatized.
purification. pPROEX-cinD was transformed into E. coli DH5
and protein expression was induced with 1 mM isopropyl b-
galactpyranoside (IPTG). The best expression was obtained at be-
tween 27–30 °C following induction in terrific broth (data not
shown).
a
cells
D-thio-
3.3. Purification and characterisation of the (1R)-6b-hydroxycineole
dehydrogenase
Purification of (1R)-6b-hydroxycineole dehydrogenase was
facilitated by the addition of an N-terminal His-tag, enabling puri-
fication of the protein in a single step. The recombinant protein
was purified by immobilized metal affinity chromatography
(IMAC) employing a HiTrap Chelating column (GE Healthcare)
loaded with Ni2+. This resulted in a single band as evaluated by
SDS–PAGE at approximately 30 KDa (theoretical mass including
His-tag = 29.3 KDa). The purification typically yielded 50 mg of
purified protein per litre of original culture.
2.7.2. Reduction of 6-ketocineole (3) to 6-hydroxycineole (2)
(1R)- and (1S)-6-ketocineole (3a and 3b) were assayed in an
analogous fashion to that described for 6-hydroxycineole (2),
employing NADH (rather than NAD+) to initiate the reaction.
Promotor
cinD
cinA
cinB
cinC
direction of
transcription
Fig. 2. Map of the CIN operon region that contains four open reading frames, cinA, cinB, cinC and cinD.
Table 1
Sequence homology of cinD open reading frame.
Protein
Organism
%Ia (%S)b
Genbank ID
Refs.
Putative 20-b-hydroxysteroid dehydrogenase FabG3_1
3a, 20b-Hydroxysteroid dehydrogenase
Putative short-chain dehydrogenase
Putative oxidoreductase
R-specific alcohol dehydrogenase
3-Ketoacyl reductase
Mycobacterium marinum M
M. tuberculosis
Nocardia facinica IFM 10152
Streptomyces coelicolor A3(2)
Lactobacillus brevis
60 (73)
59 (72)
50 (67)
47 (62)
37 (55)
32 (50)
YP_001851274
P69167
YP_121022
NP_628345
CAD66648
AAC69639
[20]
[13]
[21]
[22]
[23]
[24]
M. tuberculosis H37Rv
a
Percent identity.
b
Percent similarity (identity + substitutions with an amino acid with similar properties).

84
K.E. Slessor et al. / Bioorganic Chemistry 38 (2010) 81–86
To establish the pH profile of (1R)-6b-hydroxycineole dehydro-
cineole (Fig. 3, 2a–d) were incubated with (1R)-6b-
hydroxycineole dehydrogenase and NAD+ and the products ana-
lysed by enantioselective GC. Only (1R)-6b-hydroxycineole (2a)
was observed to be converted to (1R)-6-ketocineole (Fig. 4, 3a);
(2b–d) were unreactive. To establish if the dehydrogenase could
also catalyse the reverse reaction (reduction), both isomers of the
6-ketocineole (3) were incubated with NADH and the enzyme.
Again only one isomer, (1R)-6-ketocineole (3a) was converted to
the corresponding alcohol: (1R)-6b-hydroxycineole (2a). Cofactor
specificity was also examined by incubating the dehydrogenase
and (1R)-6b-hydroxycineole (2a) with either NAD+ or NADP+. No
product could be observed with NADP+ as the cofactor. In addition,
no products were observed when attempting the reduction of (1R)-
6-ketocineole (3a) with the dehydrogenase and NADPH.
genase, specific activity was measured every 0.2 pH units using
50 mM Tris–HCl from pH 7.0 to 8.4 employing the NADH-detection
assay. Below pH 7.2 the enzyme was found to significantly drop in
activity (approximately 30% of maximum activity). Between pH 7.2
and 8.2 only a slight increase in activity was observed. A high con-
centration of sodium chloride is present in the elution buffer for
the Ni2+-chelating column used to purify the His-tagged dehydro-
genase. To determine whether the sodium chloride concentration
had any effect on activity the introduction of sodium chloride at
various concentrations (20–200 mM) was examined. No significant
effects were observed with increased sodium chloride (data not
shown).
Other structurally related terpenes, isoborneol (4) and camphor
(5) (Fig. 3) were incubated with the dehydrogenase to gain insight
into the substrate specificity of the enzyme. Neither (4) nor (5)
were metabolised. Because of the high degree of similarity of the
3.4. Substrate and cofactor specificity
It was postulated that the cinD gene product may be responsible
for catalysing the oxidation of 6-hydroxycineole (2) to 6-ketocine-
ole (3) (Fig. 1, boxed). Individually, all four isomers of 6-hydroxy-
(1R)-6b-hydroxycineole dehydrogenase to a 3a, 20b-hydroxyster-
oid dehydrogenase (Genbank ID: P69167, 59% identity, 72% simi-
larity; [13]), both androsterone and progesterone were examined
as possible substrates. Androsterone was found to be oxidised in
the presence of CinD and NAD+ and have a Vmax of 23 2 minꢂ1
(rate measured in the presence of Ni2+ vide infra). Progesterone
was not observed to be reduced by (1R)-6b-hydroxycineole
dehydrogenase.
O
O
O
O
HO
OH
OH
OH
2c
2a
2b
2d
3.5. Effect of metal ions
To examine whether metal ions are required for activity a num-
ber of divalent metal ions were incubated with (1R)-6b-hydroxy-
cineole dehydrogenase (with the His-tag still present), NAD+ and
(1R)-6b-hydroxycineole (2a). Both the nickel (NiSO4) and cobalt
(CoCl2) ions increased the activity of the dehydrogenase by 10 to
15-fold. Activity did not increase any further with both Ni2+ and
HO
O
O
O
O
O
3a
3b
4
5
Co2+ present together. All other metal ions tested (Zn2+, Cu2+
,
Fig. 3. The four isomers of 6-hydroxycineole (2a–d), the two enantiomers of 6-
ketocineole (3a–b), isoborneol (4) and camphor (5).
Ca2+, Mg2+, Mn2+) had no effect on the enzyme activity. For maxi-
mal activity the optimum concentration of nickel was observed
3b
3a
synthetic standard
2a
(a)
2b
(b)
2c
(c)
2d
(d)
14
16
18
20
Retention Time (min)
Fig. 4. Enantioselective GC analysis of products formed following incubation of 6-hydroxycineole (2a–d) with (1R)-6b-hydroxycineole dehydrogenase. Arrow indicates the
formation of (1R)-6-ketocineole (3a) from (2a) by CinD catalysed oxidation in the presence of NAD+. (Traces of the other isomers are present in each of 2a–2d due to the
synthetic route employed [12]).

K.E. Slessor et al. / Bioorganic Chemistry 38 (2010) 81–86
85
to be between 0.5 and 5 mM metal ion/
centrations above 5 mM had a similar dehydrogenase activity to
that when no nickel is present. Assays were routinely run at
l
M dehydrogenase. Con-
cinB and cinC was discovered. The cinA, cinB and cinC genes encode
the suite of enzymes responsible for the hydroxylation of cineole to
(1R)-6b-hydroxycineole (2a) [8,25]. The cinD gene displays signifi-
cant homology to a number of short-chain dehydrogenases/oxido-
reductases (Table 1). The second step in cineole degradation is
believed to be oxidation of the initially formed secondary alcohol
to the corresponding ketone (Fig. 1). Because the sequence align-
ment indicated that the cinD gene product was likely to be a
short-chain dehydrogenase it was postulated that the cinD gene
product may be responsible for catalysing the oxidation of (1R)-
6b-hydroxycineole (2a) to (1R)-6-ketocineole (3a) in C. braakii.
Following the purification of the His-tagged-dehydrogenase,
incubation of (1R)-6b-hydroxycineole (2a) with the enzyme and
NAD+ as a cofactor indicated (1R)-6b-hydroxycineole dehydroge-
nase did catalyse its oxidation producing (1R)-6-ketocineole
(Fig. 3, 3a). This demonstrates that the cinD gene product is in-
volved in the second step of cineole degradation in C. braakii. As ex-
pected, the reverse reaction, the reduction of (1R)-6-ketocineole
(3a) to (1R)-6b-hydroxycineole (2a), is also catalysed by the (1R)-
6b-hydroxycineole dehydrogenase in the presence of NADH. The
dehydrogenase was found to be highly stereoselective. Of the four
isomers that exist for 6-hydroxycineole (2), only (1R)-6b-hydroxy-
cineole (2a) is oxidised to the corresponding ketone (3a) (Figs. 3
and 4). Of the two enantiomeric ketocineoles (3), only (1R)-6-keto-
cineole (3a) is reduced, specifically yielding 2a. NADP+/NADPH
could not be used as a cofactor in these reactions. These results
are similar to that observed in a Rhodococcus species reported by
Williams et al. [7]. NAD+-dependent ‘6-endo-hydroxycineole dehy-
drogenase’ activity was detected in extracts of cells grown in the
presence of cineole, although the enzyme was not purified to
homogeneity. These extracts were found to catalyse the dehydro-
genation of (1R)-6b-hydroxycineole (2a) to (1R)-6-ketocineole
(3a) and the corresponding reverse reaction. They report a small
amount of activity (<10%) when NADPH is substituted as the elec-
tron donor. This activity is not observed with (1R)-6b-hydroxycine-
ole dehydrogenase isolated from C. braakii. Interestingly, GC
analysis of extracts of P. flava, another soil microbe capable of
employing cineole as an energy source, revealed the formation of
2 mM metal ion/lM dehydrogenase.
3.6. Removal of the His-tag from (1R)-6b-hydroxycineole
dehydrogenase
To examine whether the His-tag was interfering with the natu-
ral activity of the dehydrogenase, AcTEV™ protease was employed
to remove the tag. This protease recognises a seven amino acid se-
quence (ENLYFQG) and cleaves the peptide bond between Q and G
[14]. Following the cleavage only eight amino acids remained be-
fore the native start site of the protein. The successful removal of
the His-tag was confirmed by SDS–PAGE, in addition to the dehy-
drogenase’s behaviour on the Ni2+-chelating column.
3.7. (1R)-6b-hydroxycineole dehydrogenase-kinetic analysis
Kinetic constants (Km and Vmax) were measured for the His-
tagged-dehydrogenase and the dehydrogenase with the His-tag re-
moved with both the substrate (1R)-6b-hydroxycineole (2a) and
the cofactor NAD+ (Table 2). Reliable Km values could not be deter-
mined for the His-tagged-dehydrogenase because Michaelis–Men-
ten conditions could not be satisfied (i.e. substrate
concentration = 10 ꢀ Km could not be attained). Only with the
addition of divalent nickel ions could accurate Km and Vmax values
be measured. In contrast, the addition of nickel to the dehydroge-
nase with the His-tag removed did not change the enzyme activity.
The Km for NAD+ with the dehydrogenase is 3-fold higher than that
of the His-tagged-dehydrogenase in the presence of nickel. The
improvement in Km seen for the His-tagged dehydrogenase in the
presence of nickel is mirrored in the observed Vmax for both these
forms of the dehydrogenase with NAD+ (3-fold increase for His-
tagged-dehydrogenase with nickel). The opposite trend was ob-
served when comparing Km values for (1R)-6b-hydroxycineole
(2a). The Km for the cleaved-dehydrogenase with (1R)-6b-hydroxy-
cineole (2a) was 2-fold lower than for the His-tagged-dehydroge-
nase with nickel.
both (1S)-6a- and (1S)-6b-hydroxycineole (2c and 2d) [4]. This
indicates that either the enzymes responsible for the initial steps
in the oxidative biodegradation of cineole in P. flava are not as ster-
eoselective as those in C. braakii or that there are other, parallel
activities in P. flava that account for the production of both 2c
and 2d.
4. Discussion
Subsequent to the isolation of the genes encoding P450cin (cinA)
and its two redox partners (cinB and cinC) from C. braakii [8], a
fourth gene (cinD) divergently transcribed with respect to cinA,
The (1R)-6b-hydroxycineole dehydrogenase shares a high de-
gree of similarity to a group of enzymes called 3
steroid dehydrogenases (Table 1). Some of these enzymes have
been shown to be capable of the oxidation/reduction of the 3
a, 20b-hydroxy-
Table 2
a
Kinetic constants determined for the purified (1R)-6b-hydroxycineole dehydrogenase.
and 20b-hydroxyl/ketone groups of steroids such as progesterone
[15,16]. One such enzyme from Mycobacterium tuberculosis was
found to have 59% identity and 72% similarity [13] with CinD
and has been demonstrated to oxidise androsterone
NAD+
Km
(1R)-6b-hydroxycineolea
(l
M)
Vmax (minꢂ1
)
Km
(
lM)
Vmax (minꢂ1
)
HCDHb
HIS-HCDH
HIS-HCDH (Ni)
537 39
<6000d
175
26 1c
11
2
12 1c
ꢁ1.8d
9.5 0.3d
88 1c
ꢁ56d
(Vmax = 7.6 minꢂ1
)
and reduce progesterone (Vmax = 1.2 minꢂ1).
6
21
2
101 7c
(1R)-6b-hydroxycineole dehydrogenase was also found to oxidise
androsterone (Vmax = 23 2 minꢂ1) but not reduce progesterone.
Considering the highly selective requirements of the dehydroge-
nase for its proposed substrates ((1R)-6b-hydroxycineole 2a/(1R)-
6-ketocineole 3a) this result was unexpected. This confirms that
assigning function simply based on sequence alignment will not al-
ways be reflective of in vivo function.
Initially, (1R)-6b-hydroxycineole dehydrogenase was found to
have very low rates of activity (1–10 minꢂ1). Two different ap-
proaches were found to be successful in improving the activity. It
was thought that the His-tag could be interfering with the NAD+
binding to the enzyme due to the inability to accurately measure
the Km for NAD+ (Table 2). Two distinctive sequences that consti-
a
Kinetic constants for the conversion of (1R)-6b-hydroxycineole to the (1R)-6-
ketocineole were calculated from the production of NADH from NAD+ (see
Section 2).
b
Abbreviations used in this table: HCDH = cleaved (1R)-6b-hydroxycineole
dehydrogenase; HIS-HCDH = His-tagged (1R)-6b-hydroxycineole dehydrogenase;
HIS-HCDH (Ni) = His-tagged (1R)-6b-hydroxycineole dehydrogenase in the pres-
ence of 2 mM NiSO4/lM enzyme.
c
Ideally, the Vmax values should be the same for each form of the enzyme under
the same conditions. The observed differences may be due to variations in tem-
perature and/or the instability of the enzyme. In addition, the high Km for the HCDH
(NAD+) means that the conditions for measuring Vmax for this form of the enzyme
only just satisfy Michaelis–Menten conditions.
d
Km or Vmax could not be measured accurately as Michaelis–Menten conditions
could not be satisfied.

86
K.E. Slessor et al. / Bioorganic Chemistry 38 (2010) 81–86
tute the Rossmann Fold [GXX(X)GX(X)G and YXXXK] were found in
the protein sequence of (1R)-6b-hydroxycineole dehydrogenase.
The Rossmann Fold allows for the binding of NADH/NAD+ during
the reduction/oxidation of the substrate [17,18]. The first
(GAAQGMG (14-20) is very close to the N-terminus to which the
His-tag was attached. His-tags have previously been demonstrated
to affect enzyme activity in some cases [19]. For example, glucosa-
mine-6-phosphate synthase from Candida albicans was expressed
in E. coli with a His-tag at the N-terminus. It was found to lose
activity completely. Once the His-tag was placed at the C-terminus
rather than the N-terminus the enzyme regained activity [19]. We
hypothesised that divalent nickel ions could coordinate to the His-
tag and remove it from the NAD(H) binding site. It was also possi-
ble that the enzyme may require a metal-ion for catalysis. Another
dehydrogenase, 5-exo-hydroxycamphor dehydrogenase from P.
putida, that catalyses a similar reaction, converting 5-exo-hydroxy-
camphor to 2,5-diketocamphane is believed to have two firmly
bound zinc atoms per subunit [10]. In an attempt to improve the
rates of turnover of (1R)-6b-hydroxycineole dehydrogenase, vari-
ous divalent metal ions were added to the reaction. Interestingly,
only nickel and cobalt were observed to increase the activity
(10–15-fold). This, together with the observation that nickel did
not improve the activity of (1R)-6b-hydroxycineole dehydrogenase
without the His-tag (vide infra), suggested that the His-tag had
been coordinated by the nickel or cobalt, exposing the NAD(H)
binding site. In addition, (1R)-6b-hydroxycineole dehydrogenase
does not have any significant homology with 5-exo-hydroxycam-
phor dehydrogenase, or the zinc-containing dehydrogenases in
general.
The second approach to improving the activity was to remove
the His-tag enzymatically using AcTEV™ protease. This approach
also resulted in an enzyme with increased activity (Table 2;
approximately 2-fold). The additional overnight incubation and
purification of the enzyme is thought to be responsible for the only
moderate increase in activity compared to that observed with the
His-tagged (1R)-6b-hydroxycineole dehydrogenase in the presence
of nickel as we had previously observed loss of activity in the en-
zyme upon storage at room temperature. Interestingly, the Km
(NAD+) for the cleaved-dehydrogenase was not as low as that for
the nickel-coordinated His-tagged-dehydrogenase (Table 2). This
perhaps suggests that the eight amino acids left at the N-terminus
following cleavage are still large enough to interfere with the opti-
mal binding of NAD+ to the enzyme.
ered the activity of the enzyme. This activity could be recovered
via the addition of Ni2+ ions, postulated to coordinate to the His-
tag and thus allow NAD(H) to access its binding site. (1R)-6b-
hydroxycineole dehydrogenase is stereo- and enantioselective,
only accepting (1R)-6b-hydroxycineole (2a). Importantly, this is
the same enantiomer produced by P450cin during the oxidation
of 1,8-cineole linking these two activities in cineole biodegradation
[8].
Acknowledgments
The authors would like to thank Yen-Yin Pan for early work on
the cleavage of the His-tag. This work was supported in part by
ARC Grant DP0881116.
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In summary, we have demonstrated that the gene product of
cinD from C. braakii exhibits (1R)-6b-hydroxycineole dehydroge-
nase activity and also catalyses the reverse reaction, reducing
(1R)-6-ketocineole (3a) to (1R)-6b-hydroxycineole (2a). During
this study it was observed that the N-terminal His-tag used to as-
sist purification interfered with the NAD(H) binding site and low-