Kinetic characterization of Rhodococcus ruber DSM 44541 alcohol dehydrogenase A

Article abstract of DOI:10.1016/j.molcatb.2013.10.023

An increasing interest in biocatalysis and the use of stereoselective alcohol dehydrogenases in synthetic asymmetric catalysis motivates detailed studies of potentially useful enzymes such as alcohol dehydrogenase A (ADH-A) from Rhodococcus ruber. This enzyme is capable of catalyzing enantio-, and regioselective production of phenyl-substituted α-hydroxy ketones (acyloins) which are precursors for the synthesis of a range of biologically active compounds. In this study, we have determined the enzyme activity for a selection of phenyl-substituted vicinal diols and other aryl- or alkyl-substituted alcohols and ketones. In addition, the kinetic mechanism for the oxidation of (R)- and (S)-1-phenylethanol and the reduction of acetophenone has been identified as an Iso Theorell-Chance (hit and run) mechanism with conformational changes of the enzyme-coenzyme binary complexes as rate-determining for the oxidation of (S)-1-phenylethanol and the reduction of acetophenone. The underlying cause of the 270-fold enantiopreference for the (S)-enantiomer of 1-phenylethanol has been attributed to non-productive binding of the R-enantiomer. We have also shown that it is possible to tune the direction of the redox chemistry by adjusting pH with the oxidative reaction being favored at pH values above 7.

Full text of DOI:10.1016/j.molcatb.2013.10.023

Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Kinetic characterization of Rhodococcus ruber DSM 44541 alcohol
dehydrogenase A
Emil Hamnevik, Cecilia Blikstad, Sara Norrehed, Mikael Widersten∗
Department of Chemistry – BMC, Uppsala University, Box 576, SE-751 23 Uppsala, Sweden
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 8 October 2013
Received in revised form 30 October 2013
Accepted 30 October 2013
Available online 7 November 2013
An increasing interest in biocatalysis and the use of stereoselective alcohol dehydrogenases in synthetic
asymmetric catalysis motivates detailed studies of potentially useful enzymes such as alcohol dehydroge-
nase A (ADH-A) from Rhodococcus ruber. This enzyme is capable of catalyzing enantio-, and regioselective
production of phenyl-substituted ␣-hydroxy ketones (acyloins) which are precursors for the synthesis
of a range of biologically active compounds. In this study, we have determined the enzyme activity for
a selection of phenyl-substituted vicinal diols and other aryl- or alkyl-substituted alcohols and ketones.
In addition, the kinetic mechanism for the oxidation of (R)- and (S)-1-phenylethanol and the reduction
of acetophenone has been identified as an Iso Theorell–Chance (hit and run) mechanism with confor-
mational changes of the enzyme-coenzyme binary complexes as rate-determining for the oxidation of
(S)-1-phenylethanol and the reduction of acetophenone. The underlying cause of the 270-fold enantio-
preference for the (S)-enantiomer of 1-phenylethanol has been attributed to non-productive binding of
the R-enantiomer. We have also shown that it is possible to tune the direction of the redox chemistry by
adjusting pH with the oxidative reaction being favored at pH values above 7.
Keywords:
Alcohol dehydrogenase
Kinetic mechanism
Pre-steady state kinetics
Product inhibition
© 2013 Elsevier B.V. All rights reserved.
1. Introduction
Stereoselectivity is a key feature also in asymmetric catalysis
applied to synthetic chemistry. There is therefore an increas-
In cells, NAD(P)H-dependent reactions, catalyzed by dehydro-
genases (EC 1.1.1.1), are instrumental in all aspects of metabolism.
The coenzymes act as carriers of reductive power affording
energy transformations and biosynthesis. The spectroscopic sig-
nals, changes in absorption or fluorescence, reflecting the redox
state and local environment of the coenzyme provide sensitive
and selective probes of discrete reaction steps during catalysis.
This has enabled gathering of an immense volume of experimental
data describing various aspects of dehydrogenase functions. Hence,
these reactions have been applied as model systems in studies of
fundamental enzymology since the beginning of the research field
[1], also increasing our understanding of enzyme-afforded catalysis
in general. Typical NAD(P)H-dependent reactions involve reduction
of carbonyl compounds (ketones or aldehydes) into the corre-
sponding secondary or primary alcohols, respectively. The carbonyl
carbon of a substituted ketone is prochiral and its reduction will
therefore result in a chiral product. This added structural complex-
ity requires stereoselective substrate recognition of the cognate
enzyme catalysts (Scheme 1A).
ing interest in applying stereoselective dehydrogenase enzymes
as biocatalysts in reversible oxidation/reduction of alcohols and
ketones [2–10]. We are exploring possibilities to exchange tra-
ditional organic and organometallic catalysis with biocatalysis to
afford synthesis of chiral and prochiral compounds. The starting
materials are aryl-substituted epoxides which are hydrolyzed to
yield chiral vicinal diols in reactions catalyzed by epoxide hydro-
lase (Scheme 1B) [11–13]. The diol products may subsequently be
derivatized further. Oxidation of a secondary alcohol produces the
corresponding ␣-hydroxy ketone (acyloin). Acyloins are important
building blocks for the synthesis of a range of biologically active
compounds such as anti-depressants, inhibitors of amyloid protein
production and anti-tumor antibiotics [14].
Alcohol dehydrogenase A (ADH-A) from Rhodococcus ruber DSM
44541 catalyzes the reversible reduction/oxidation of ketones and
sec-alcohols, respectively [15,16]. The enzyme is a classical zinc-
dependent group I alcohol dehydrogenase [17,18] structurally
similar to the, in detail studied, horse liver isoenzyme [19,20].
ADH-A displays favorable physicochemical properties, such as tol-
erance to high concentrations of isopropanol and acetone [21]
which allows for facile cofactor regeneration. In addition, ADH-
A catalyzes the oxidation of bulky aryl-substituted alcohols and
was therefore considered a good candidate catalyst for the oxi-
dation of vicinal diols as outlined in Scheme 1B. Our aims with
∗
Corresponding author. Tel.: +46 18 471 4992; fax: +46 18 55 8431.
E-mail address: mikael.widersten@kemi.uu.se (M. Widersten).
1381-1177/$ – see front matter © 2013 Elsevier B.V. All rights reserved.
http://dx.doi.org/10.1016/j.molcatb.2013.10.023

E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
69
Scheme 1. (A) Scheme of a generic dehydrogenase catalyzed oxidation of a sec-
alcohol into the corresponding ketone. (B) Mini-pathway of enzyme-catalyzed
synthesis of acyloins from substituted epoxides. An asterisk indicates a chiral center.
EH, epoxide hydrolase, ADH, alcohol dehydrogenase.
the present work were as follows: (1) to establish the substrate-
, enantio-, and regioselectivities of ADH-A regarding oxidation of
aryl-substituted vicinal diols and other alcohol and ketone sub-
strates (Fig. 1), (2) to characterize the kinetic mechanism of the
enzyme. The gathered information will form the baseline guiding
future directed-evolution efforts aiming to generate more efficient
biocatalysts for synthesis of new acyloins and other substituted
ketones.
2. Experimental
2.1. Chemicals and reagents
Standard reagents for protein expression and all enzyme sub-
strates (except 10) were purchased from Sigma–Aldrich, Merck
and VWR. The (S)- and (R)-3-phenyl-1,2-propane diols (10) were
obtained after enzyme-catalyzed hydrolysis of the correspond-
ing (S)- and (R)-benzyloxiranes (Tokyo Chemical Industry Co)
according to the protocol in ref. [11]. Restriction enzymes DNA
polymerases and other DNA modifying enzymes were purchased
from Fermentas.
2.2. ADH-A gene cloning
The gene sequence encoding alcohol dehydrogenase A (ADH-
A) from R. ruber DSM 44541 was optimized for expression in
Escherichia coli, synthesized and purchased from Invitrogen using
their GeneArt® service. The final gene construct was flanked by
the restriction sites for XhoI and SpeI at the 5ꢀ- and 3ꢀ-ends of
the gene, respectively. The gene was subcloned into a modified
pGT7YNR064c-5H vector [22] (inserting codons for a penta-histidyl
tag at the 3ꢀ-end of the open reading frame) using XhoI and SpeI.
An unwanted BamHI site present in the vector was removed by
Klenow fill-in prior to subcloning of the ADH-A synthetic gene. The
formed construct was denoted as pGT7ADHA-5H. The ADH-A gene
was sequenced in full to ensure sequence integrity.
Fig. 1. Alcohols and ketones tested as substrates. 1, 1-propanol, 2, 2-propanol, 3,
acetone, 4, benzyl alcohol, 5, 1-phenylethanol, 6, acetophenone, 7, 2-phenylethanol,
8, 1-phenyl-1,2-ethanediol, 9, 2-hydroxyacetophenone, and 10, 3-phenyl-1,2-
propanediol.
2.4. Protein expression and purification
The expression constructs were transformed into chemically
competent E. coli BL21-AI cells (Invitrogen) pre-transformed with
the expression plasmid pREP4-groEL/ES [23] allowing for con-
comitant over-expression of chaperonins GroEL/ES. BL21-AI cells
containing pGT7ADHA-5H, or the corresponding construct encod-
ing the H39N mutant, (and pREP4-groEL/ES) were incubated in 2 ml
2TY media (1.6% (w/v) tryptone, 1.0% (w/v) yeast extract, 0.5% (w/v)
NaCl) containing 100 ␮g/ml ampicillin and 30 ␮g/ml kanamycin at
30 ◦C for 6 h. The culture was subsequently diluted 1:50 in 50 ml
2TY containing 100 ␮g/ml ampicillin and 30 ␮g/ml kanamycin and
incubated for additional 16 h at 30 ◦C. Finally, the culture was
diluted 1:100 in 6 × 500 ml 2TY containing 50 ␮g/ml ampicillin and
15 ␮g/ml kanamycin and incubated at 30 ◦C until an OD600 of 0.4
was reached (approximately 2.5 h). Expressions of the ADH-A and
GroEL/ES genes were induced by addition of 0.2% (w/v) l-arabinose
and 1 mM isopropyl ␤-d-1-thiogalactopyranoside, respectively.
The culture was incubated for additional 3 h at 30 ◦C. Cells were
2.3. Site-directed mutagenesis of His39 to Asn
His39 was mutated into an Asn by site-directed mutagenesis
using the QuikChange method (Agilent Technologies). The muta-
tion was inserted into pGT7ADHA-5H using PCR in the presence
of the following mutagenic primers: (Fwd His39Asn) CTG TGT AAT
AGC GAT ATT TTT GTT ATG GAT ATG CCT GCA GCA CAG and (Rev
His39Asn) C GCT ATT ACA CAG ACC TGC TGC GGT AAC TTT CAG
CAG AAT TTC. The product was purified and digested with DpnI
after which it was transformed into electrocompetent E. coli XL1-
blue cells by electroporation. The H39N expression construct was
purified and the ADH-A H39N gene was sequenced in full to ensure
that no unwanted mutations had been introduced.

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E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
harvested by centrifugation at 3800 × g for 13 min at 4 ◦C. Bacteria
were resuspended in 20 ml Binding Buffer (20 mM sodium phos-
phate, 20 mM imidazole, 500 mM NaCl, 0.02% sodium azide, pH
7.5) fortified with protease inhibitors (EDTA-free Complete, Roche)
and 20 ␮g/ml DNaseI. Cells were lysed using a Cell disruptor Z
plus series (Constant Systems LTD) and debris was removed by
centrifugation at 27,200 × g for 1 h at 4 ◦C. The supernatant was
transferred to a conical 50 ml tube and 2.5 ml Ni2+-immobilized
metal ion affinity chromatography (IMAC) gel solution (Chelating
SepharoseTM fast flow, GE Healthcare) equilibrated with Bind-
ing Buffer (75% v/v gel) was added to the lysate and the slurry
was incubated at 4 ◦C for 1 h. The gel/lysate slurry was subse-
quently centrifuged at 100 × g at 4 ◦C and the supernatant was
discarded. Washing Buffer (Binding Buffer containing 100 mM imi-
dazole) was added to a total volume of 20 ml and gel slurry was
incubated at 4 ◦C for 10 min before centrifuged as above. The wash-
ing step was repeated two additional times. Elution of ADH-A was
achieved by adding 5 ml of Elution buffer (Binding Buffer con-
taining 300 mM imidazole) to the IMAC gel. The gel slurry was
transferred to a conical 15 ml tube and incubated for 20 min at
4 ◦C. The gel–buffer mix was centrifuged at 140 × g at 4 ◦C and
the supernatant was transferred to a new 15 ml conical tube.
This elution step was performed twice. The eluted fractions were
pooled and concentrated down to 2.5 ml using a spin-column
(Vivaspin 20, 10,000 MW cut-off, Sartorius Stedim Biotech). The
concentrated sample was desalted using a PD-10 column (GE
Healthcare) equilibrated with Desalting Buffer (0.1 M sodium phos-
phate, 10 ␮M ZnSO4, 0.02% sodium azide, pH 7.4) according to
the manufacturer’s protocol. The purified enzyme sample was
stored at 4 ◦C. Enzyme purity was controlled by SDS–PAGE and
staining with Coomassie Brilliant Blue R-250. The molar extinc-
tion coefficient at 280 nm was determined to be 31,500 cm−1 M−1
by measuring the absorbance in a UV-1700 Pharmaspec UV–vis
spectrophotometer (Shimadzu) with a concomitant determination
of the ADH-A concentration by quantitative amino acid analy-
sis.
2.6. Zinc-dependent functional integrity
Purified ADH-A was stored at 4 ◦C in either Desalting Buffer or
Desalting Buffer devoid of ZnSO4 during 40 and 60 days, respec-
tively. Aliquots were removed at different time intervals and
remaining enzyme activity was determined. Initial reaction veloci-
ties were measured in 0.1 M sodium phosphate, pH 7.0 at 30 ◦C and
1% (v/v) acetonitrile with final concentrations of 5 mM acetophe-
none and 0.4 mM NADH.
2.7. pH-dependence of enzyme activity
The pH dependencies of kcat and kcat/Km for wt ADH-A and
ADH-A H39N-catalyzed oxidation of S-5 and reduction of 6 were
determined in the pH range of 5–9. Reactions were performed in
96-well polystyrene plates (Nunc) in 0.1 M sodium phosphate (pH
5–8.5) or 0.1 M glycine (pH 8.5–9.0) at 30 ◦C and 1% (v/v) acetoni-
trile with final concentrations of 0.2–7.5 mM S-5 and 1.6 mM NAD+
in the oxidation reactions and, 0.18–7.0 mM 6 and 0.4 mM NADH
in the reduction reactions. Reduction or oxidation of the coenzyme
was monitored at 340 nm in a Spectra MAX 190 UV–vis spectropho-
tometer (Molecular Devices) using an extinction coefficient for
NADH of 3440 M−1 cm−1. Kinetic parameters were determined as
described above. Apparent acid constants were extracted by fitting
expressions for titration of one- or two-proton systems (Equations
(1) and (2), respectively), where LH is the titrated parameter kcat or
kcat/Km. The extra parameters for a two-proton system were only
included if justified by an F-test. Curve fitting was performed with
program RFFIT in SIMFIT.
LHA[H+] + LA−Ka1
LH
=
=
(1)
Ka1 + [H+]
LH2A[H+]2 + LHA−[H+]Ka1 + Ka1Ka2LA2−
LH
(2)
K
a1Ka2 + [H+]Ka1 + [H+]2
2.8. Product inhibition
2.5. Steady state kinetics
The steady state kinetics was analyzed in the presence of
increasing concentrations of either product (6 and NADH) in the
oxidative direction i.e. S-5 oxidation and NAD+ reduction. Mea-
surements were performed as described above in the paragraph
describing the steady state kinetics measurements. Final con-
centrations of alcohol were 0.12–7.5 mM, and 1.6 mM NAD+ and
0–0.2 mM NADH or 0–3 mM 6. Final concentrations for coen-
zyme reduction were 0.015–3.6 mM NAD+ with 7.5 mM S-5 and
0–0.12 mM NADH or 0–5 mM 6. The Michaelis–Menten equation
was fitted to the velocity data using MMFIT and the fits were graph-
ically presented as double-reciprocal plots.
Steady state kinetic parameters for ADH-A, or the H39N mutant,
were obtained by measuring initial velocities in the presence of
varying concentrations of 1-propanol (1, Fig. 1), 2-propanol (2),
acetone (3), benzyl alcohol (4), (S)-1-phenylethanol (S-5), (R)-
1-phenylethanol (R-5), acetophenone (6), 2-phenylethanol (7),
(S)-1-phenyl-1,2-ethanediol (S-8), (R)-1-phenyl-1,2-ethanediol (R-
9), 2-hydroxy acetophenone (9), (S)-3-phenyl-1,2-propanediol
(S-10), (R)-3-phenyl-1,2-propanediol (R-10), and in the presence
of saturating concentrations of NAD+ or NADH. All reactions were
performed in 0.1 M sodium phosphate, pH 8.0 at 30 ◦C and 1% (v/v)
acetonitrile. Final concentrations were 6–100 mM 1–3, 1.8–30 mM
4, 0.11–7.5 mM S-5, 0.3–10 mM R-5, 0.1–7 mM 6, 0.38–12.5 mM 7,
1.7–28 mM S-8, 0.1–28 mM R-8, 0.18–12.5 mM 9, 0.1–50 mM S-10
and R-10, in the presence of 1.6 mM NAD+ (18 × Km) in the oxidation
of alcohols and 0.4 mM NADH (10 × Km) when reducing ketones.
The kinetic parameters in the presence of varied concentrations of
coenzymes were determined in 0.015–1.6 mM NAD+ and 7.5 mM
S-5 (12 × Km) and 0.005–0.4 mM NADH and 5 mM 6 (4.2 × Km).
Reduction of NAD+ or oxidation of NADH was monitored at 340 nm
in a UV-1700 Pharmaspec UV–vis spectrophotometer (Shimadzu).
The kinetic parameters, kcat and Km were extracted after fitting the
Michaelis–Menten equation by non-linear regression to the ini-
tial velocity data with program MMFIT (in the SIMFIT package,
www.simfit.uk.org) and kcat/Km was determined from the same
2.9. Regioselectivity
R-5 or S-5 (60 mM) were mixed with 1.6 mM NAD+, 60 mM
sodium pyruvate, 2 U/ml l-lactic dehydrogenase (Sigma) and 3 ␮M
of ADH-A in 0.1 M ammonium bicarbonate to a total volume of
10 ml. The reactions were incubated at 30 ◦C for 10 h in the dark.
CDCl3 (2.4 ml) was added to each reaction vial, forming a two-phase
system. The phases were mixed by vortexing for 5 s and were subse-
quently allowed to separate. The CDCl3 phases were transferred to
new vials. Aliquots from the CDCl3 phases were analyzed by reverse
phase HPLC on an Ascentis C-18 column (Supelco, 25 cm × 4.6 mm,
5 ␮m) using an isocratic elution of 37% (v/v) methanol in water and
0.1% (v/v) of formic acid.
raw data after fitting the equation v0 = ((kcat/Km)[S])/(1 + [S]/Km
)
NMR investigations of the reaction products were carried out
on a Varian Unity Inova (1H at 499.94 MHz, 13C at 125.7 MHz)
with program RFFIT.

E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
71
spectrometer. Chemical shifts were reported in ppm referenced to
3. Results and discussion
tetramethylsilane via the residual solvent signal (CDCl3, 1H at 7.26
and 13C at 77 ppm). Signal assignments were derived from gHSQC
[24], gHMBC [25], and TOCSY [26] spectra.
3.1. Substrate-, enantio- and regioselectivity
It has been shown previously that R. ruber ADH-A accepts a
broad range of secondary alcohols and ketones as substrates and
a wide variety of reaction conditions and co-solvent systems have
been tested to assess the potential of this enzyme as a biocata-
lyst of redox reactions [15,16,27–33]. Our aim with the current
work was to complement this already present bulk of important
applied data by conducting a detailed kinetic characterization. We
have determined the steady state kinetic parameters for a range
of aliphatic and aryl-substituted alcohols, diols and ketones in
order to provide a quantitative assessment of the enzyme’s sub-
strate preferences (Table 1). Our main motivation for the study
was to assess the suitability and potential of this enzyme to act
as a catalyst for the oxidation of vicinal diols into the correspond-
ing acyloins (cf. Scheme 1B) The data shows that ADH-A strongly
prefers secondary alcohols over primary as demonstrated by the
ratio of 5000 in the kcat/Km values for S-5 and 7. ADH-A also cat-
alyzes the oxidation reaction at pH 8.0, with a 4-fold higher catalytic
efficiency as compared to the reduction direction, as seen from a
comparison of the catalytic efficiencies with S-5 and 6. Further-
more, the enzyme prefers bulkier phenyl-substituted substrates
over smaller aliphatic compounds as shown by comparing the
enzyme efficiencies with 2 and S-5 where kcat/Km drops 170-fold
going from S-5 to 2, the same behavior is observed if compar-
ing 1 and 7, or 3 and 6. The phenyl substituent is assumed to
improve both affinity and selectivity in binding to the enzyme
active site during formation of a productive ternary complex.
One parameter that supports this notion is Km, which is affected
by binding affinity. Km2 is a 100-fold larger as compared to KmS-5
while the respective turnover numbers are comparable suggesting
poorer binding of isopropanol at the active site by approximately
3 kcal/mol.
A major reason for the choice of R. ruber ADH-A as model
enzyme was its putative catalytic function with aryl-substituted
vicinal diols. We therefore tested diols 8 and 10 which can
be considered hydrolysis products of styrene oxide and (2,3-
epoxypropyl)benzene. ADH-A does indeed display activity with
these vicinal diols but with lower efficiencies as compared to mono-
hydroxy substituted derivatives; ADH-A displays 2900-fold higher
catalytic efficiency with S-5 as compared to R-8, which is the stere-
ochemically related diol. The probable reason for this decrease in
activity is simply steric hindrance and introduction of a polar group
by the primary alcohol. The same pattern is observed if comparing
the enzyme activities with ketones 6 and 9, although the effect is
less dramatic. The activities with diols R/S-10 are 3–10-folds higher
as compared to the corresponding 8 enantiomers. A reason may be
that the introduction of the extra methylene group in 10 provides
increased flexibility in the enzyme–substrate interactions, as com-
pared to the more rigid structure of diol 8, facilitating formation of
productive ternary complexes.
2.10. Pre-steady state kinetics
Transient kinetics experiments were performed on an Applied
Photophysics (Leatherhead, UK) SX.20MV sequential stopped-
flow spectrophotometer/fluorometer. All measurements were
performed at 30 ◦C in 0.1 M sodium phosphate, pH 8.0. Averages
of 8–10 progression curves were used in all cases and the appar-
ent rates, kobs, were determined by fitting Equation (3), a function
of a single exponential with floating endpoint, to the progression
curves. F is the recorded fluorescence signal, A, the signal amplitude,
t, time and C, the floating end point.
F = Aexp(−kobst) + C
(3)
2.10.1. Coenzyme binding
Binding of coenzyme to ADH-A was detected by monitoring the
change in intrinsic protein tryptophan fluorescence which occur
upon binding of NADH/NAD+ to the enzyme active site. An exci-
tation wavelength of 290 nm was used and fluorescence light was
collected through a 320 nm cut-off filter. Concentration of NADH
ranged from 10 to 75 ␮M and for NAD+ from 1.7 to 67 ␮M. Enzyme
concentrations for NADH measurements were between 0.8 and
2.2 ␮M and for NAD+ measurements between 0.14 and 1.1 ␮M.
Nucleotide concentrations were always kept in at least 10-fold
excess as compared to the enzyme concentrations. The apparent
rates, kobs, were determined as described above. The kinetic param-
eters were determined by fitting the dependence of kobs on the
coenzyme concentration to a linear (Equation (4)) or a hyperbolic
function (Equation (5)) using LINFIT or QNFIT, respectively. Model
discrimination was decided from F-tests of the fitting results with
an accepted significance level of p ≤ 0.05.
kobs = kon[NAD(H)] + koff
(4)
kf [NAD(H)]
KSNAD(H) + [NAD(H)]
kobs
=
+ kr
(5)
2.10.2. The redox chemistry
The alcohol oxidation was monitored by following the change
in fluorescence upon increase (alcohol oxidation) of the NADH con-
centration. Change in fluorescence was detected using an excitation
wavelength of 340 nm and collecting the fluorescent light after
passage through a 395 nm cut-off filter. The oxidation reaction
was measured with S-5 and R-5 with NAD+ as electron acceptor.
For measurements with S-5 the substrate concentrations varied
between 0.6 and 10 mM and the enzyme concentration was 0.6 ␮M.
For R-5, the substrate concentrations were varied between 1.5 and
10 mM and the enzyme concentration was 2.3 ␮M. In both cases
[NAD+] was kept constant at 0.4 mM. Before measurements, the
fluorescence of the NAD+-enzyme complex was monitored and
the voltage was thereafter adjusted to 8.0 V. Averaged progression
curves were fitted to the function of single exponential with float-
ing endpoint, as described above, to determine the apparent rates,
kobs. The kinetic parameters, k4 and K3 were extracted by fitting
the obtained rates to Equation (6) by non-linear regression using
MMFIT.
When comparing the substrate selectivities of ADH-A from R.
ruber with that of horse liver alcohol dehydrogenase [34–37] it is
clear that both enzymes are able to catalyze oxidation of primary
and secondary alcohols, aliphatic and aryl-substituted alcohols.
The two isoenzymes, however, do display differences in their sub-
strate preferences. The horse liver enzyme prefers small primary
alcohols, while ADH-A shows highest activity with bulkier sec-
ondary alcohols. ADH-A is also more efficient in catalyzing diol
oxidations as compared to the horse liver enzyme [37]. These dif-
ferences can be explained by the respective active-site structures
(Fig. 2A). The horse enzyme active site is more narrow, restricted
and channel-like while the active site of the Rhodococcus isoen-
zyme forms an open cleft allowing for binding of bulkier substrates
k4[alcohol]
K3 + [alcohol]
kobs
=
(6)

72
E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
Table 1
Substrate selectivity. Steady state kinetic parameters for ADH-A catalyzed redox reactions with a panel of alcohols and ketones. See Fig. 1 for chemical formulas. Enantiose-
lectivity, E, was calculated from the ratios of kcat/Km for the respective enantiomers. Values refer to the preferred substrate enantiomer with the configuration shown within
brackets. Data for the H39N mutant of ADH-A have been included for comparison.
Substrate
kcat (s−1
)
Km (mM)
kcat/Km (s−1 mM−1
)
E (fold)
1-propanol (1)
2-propanol (2)
Acetone (3)
Benzyl alcohol (4)
–
56
–
–
74
–
0.0055 0.0002
0.76 0.03
0.54 0.02
2
6
2
0.022 0.001
13
0.0018 0.0001
S-1-phenylethanol (S-5)
R-1-phenylethanol (R-5)
Acetophenone (6)
2-phenylethanol (7)
S-1-phenyl-1,2-ethanediol (S-8)
R-1-phenyl-1,2-ethanediol (R-8)
2-hydroxy acetophenone (9)
S-3-phenyl-1,2-propanediol (S-10)
R-3-phenyl-1,2-propanediol (R-10)
NAD+
80 20 (H39N: 61 0.8)
0.45 0.005
36 0.8 (H39N: 15 0.6)
0.025 0.00007
0.0094 0.0005
0.73 0.01
2.0 0.08
0.15 0.001
0.56 0.005
110 10(H39N: 38 0.2)
37 0.5
0.63 0.05 (H39N: 1.7 0.06)
0.94 0.04
130 30 (H39 N: 36 0.9)
0.48 0.01
270 (S)
1.2 0.09
0.97 0.1
3.0 0.7
17 0.6
(H39N: 1.2 0.1)
30
2
(H39N: 12 0.9)
0.026 0.002
0.0031 0.0006
0.044 0.0008
0.55 0.03
0.053 0.001
0.12 0.003
14 (R)
2.3 (R)
3.7 0.4
3.0 0.09
4.4 0.2
0.086 0.006(H39N: 0.083 0.002)
0.039 0.002
1200 200(H39N: 450 9)
950 40
NADH
(Fig. 2B). The kinetic parameters for reactions with different enan-
tiomers of alcohols used in this study show a preference of ADH-A
for the S-enantiomer comparing S- and R-5, as have been reported
previously [16], with an E-value of 270. In the oxidation of diols
R/S-8 and R/S-10 ADH-A shows a preference for the R-enantiomer
with E-values of 14 and 2.3, respectively. The reason for the appar-
ent shift in enantiopreferences, from S-preference to R for the diols,
is due to the change in priority order of substituents resulting from
the extra hydroxyl group; the actual configuration of the reacting
functional groups is identical. The preference for oxidation of the
secondary alcohol of the R-8 vicinal diol was confirmed both by
the migration of the product in an HPLC reverse phase system in
which 2-hydroxy acetophenone was included as control substance
(Fig. SI1 Supplementary Information) and by NMR. NMR studies
of the product resulted in signals also supporting 2-hydroxy ace-
tophenone as the only reaction product. The NMR analysis of the
oxidation reaction of S-8 was unable to detect any ketone, only
starting material was present, probably due to the low turnover
number of the enzyme with this substrate (Table 1). Chemical shifts
and assignments are presented in the Supplementary Information.
the bound nucleotide (Fig. 2E) by the amide nitrogen, but to remove
the expected positive charge of the protonated imidazolium at
lower pH. The effects on the pH-dependence, however, were not
dramatic (Fig. 3, Table 2) although the mutation did cause changes
in the apparent pKa values. The pKa values determined for kcat/Km
were shifted by approximately 1.7 and 1 pH units towards more
basic values for the oxidation and reduction reactions, respectively.
The effects on kcat for either reaction were modest. Even though the
effect on the pKa values is relatively small, the change in the shapes
of the pH-profiles indicates that His39 is indeed contributing to the
titration of the wild-type enzyme and may be involved in a proton
relay similar to His51 in the horse liver enzyme.
3.3. Kinetics of ADH-A H39N
The ADH-A H39N mutant exhibited a slight decrease in kcat and
an increase of 2.7-fold in Km with S-5 (Table 1, values in brackets),
resulting in a 3.6-fold decrease in kcat/Km. For NAD+, the Km was
unaffected, but the kcat and kcat/Km values were decreased by 2.9-
and 2.7-fold, respectively. These values, however, should be viewed
as apparent and represent lower limit values, since the increase in
KmS-5 prevents the enzyme to become fully saturated with S-5 during
these measurements.
3.2. pH-dependence of catalysis
According to the crystal structure (Fig. 2E), His39 line the
edge of the nucleotide binding site and interacts directly with the
nucleotide. So the fact that the H39N mutation caused no effect
on the Km for NAD+ is noteworthy and indicates that the replacing
amide side-chain can substitute well for the N-␦ of His39. The main
effect by the mutation was the 3-fold increase in Km for S-5 by the
mutation which may indicate a cross-talk between the two binding
sites, may be affected by some small change in the orientation of
the bound nucleotide caused by the mutation.
The catalytic activity, expressed as kcat or kcat/Km, increases with
increasing pH in the oxidation reaction, but decreases with increas-
ing pH in the reduction reaction (Fig. 3). At pH 8, which is the assay
pH during the survey of substrate selectivity and the pre-steady
state kinetics, the oxidation reaction is favored. The shift in reac-
tion direction preference, from reduction to oxidation, occurs at
pH 7.2 for kcat and 6.2 for kcat/Km. The catalyzed oxidation of S-5
displays a higher degree of pH dependency than does the reduc-
tion of ketone 6. In order to model the titration profiles of kcat and
kcat/Km a two-proton model had to be invoked in the cases of kc6at
and (kcat/Km)
S-5 (Table 2). The values of pKa1 ranges from 6.2 to 7.3,
3.4. Zinc-dependence
thus, indicating a role for a histidine residue in the rate-limiting
catalytic steps of the wild-type enzyme. By looking at the structure
of ADH-A (Fig. 2E) and comparing it to the proposed mechanism for
proton relay in the horse liver enzyme [38], His39 was considered
to be a candidate. Although His39 do not have exactly the same
position as His51 in the horse liver enzyme, it can be aligned in a
putative proton relay in the Rhodococcus enzyme.
To determine if His39 is (partly) responsible for the pH-profiles
a H39N mutant was constructed and characterized. The rationale to
mutate His39 into an Asn, was to retain the hydrogen bonding inter-
action between the N-␦ of His39 and the pyrophosphate moiety of
ADH-A depends on two zinc ions, one catalytic and one struc-
tural. At least one of these ions, although we have not identified
which one, is not tightly bound to the enzyme but appears to be
equilibrated with solution ions. Incubating ADH-A over a period of
time in a zinc-free buffer resulted in a decrease of activity over time
(Fig. 4). In the experiment without added zinc the enzyme concen-
tration was 40 ␮M and under these conditions the relative activity
leveled off at 50%. When incubated over the same time range in
buffer fortified with 10 ␮M zinc sulfate, the enzyme was practically
stable retaining 90–95% of initial activity.

E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
73
Fig. 2. Structure comparisons between R. ruber ADH-A and horse liver ADH. The active-site entry is more restricted in the horse liver enzyme (A) as compared to the R. ruber
ADH-A (B) explaining the wider substrate scope of the latter. The ligands shown in stick are in (A) N-formyl piperidine and in (B) (4S)-2-methyl-2,4-pentanediol. The NAD+
cofactor is also shown in stick representation. (C) The active-site loops in the “closedc¨onformation of the horse liver enzyme (gray) and the R. ruber enzyme (black). The
functionality of Val294, which is in van der Waals contact with the nicotinamide ring in this conformation, is conserved in Ile271 in the bacterial enzyme. (D) The interaction
between the non-conserved His272 and the main-chain carbonyl oxygen of Val247 may contribute to stabilization of the closed conformation of the ADH-A enzyme. (E)
His39 in R. ruber ADH-A is replaced by Arg47 in horse liver ADH. Both these residues interact with the coenzyme through electrostatic and hydrogen bonds. Image was
created with Pymol 1.5 (www.pymol.org) from the atomic coordinates in 1lde [46] for the horse liver enzyme and 3jv7 [20] for the ADH-A structure.
3.5. Kinetic mechanism
at pH 7.0 (Table 3, Fig. 6A), in which NADH (Q) acts as a competi-
tive inhibitor of NAD+ (A) and 6 (P) acts as a competitive inhibitor
of S-5 (B) (Fig. SI2). In an ordered Bi Bi mechanism with an accu-
mulation of the ternary EAB complex, which has also been invoked
for the same horse liver isoenzyme [40], one would expect P to
act as a mixed-type inhibitor of B [41]. The distinction is not
The data suggests a Theorell–Chance (hit-and-run) mechanism
(Fig. 5B), as was proposed for the horse liver enzyme [39] with
a low degree of ternary complex accumulation during the steady
state. This is based on the product inhibition patterns, determined
Table 2
Apparent pKa values of kcat and kcat/Km in catalyzed oxidation of S-1-phenylethanol or reduction of acetophenone. Apparent pKa values were extracted after curve-fitting of
functions for one or two titratable groups. See the Section 2 for details.
Oxidation of S-5
Wild-type
pKa1
Reduction of 6
Wild-type
pKa1
H39N
H39N
Parameter
pKa2
pKa1
pKa2
pKa1
kcat
kcat/Km
7.3 0.06
6.2 0.06
7.7 0.04
7.9 0.04
6.2 0.7
7.3 0.3
7.4 0.5
7.3 0.1
8.3 0.3
8.4 0.2

74
E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
Fig. 3. pH-dependence of kcat and kcat/Km in the oxidation of S-1-phenylethanol or reduction of acetophenone. The wild-type ADH-A is represented by filled symbols and
the H39N ADH-A mutant by unfilled symbols. See Table 2 for determined pKa values. (A) pH-dependence of kcat for S-5. The turnover numbers increase with increasing pH
in the oxidation for both enzyme forms, with a slight shift in the pH-dependence towards higher pH values by the H39N mutation. (B) The kcat of the reduction of 6 displays
a maximum around pH 7 for the wild-type catalyzed reactions resulting in two apparent acid constants. The more acidic of these pKa values appears to be lost as a result
of the H39N mutation. (C) kcat/Km for the oxidation reaction increases with increasing pH similar to kcat, but again, the wild-type ADH-A displays two pKa values while the
H39N mutant only appears to depend on one titrated group in the tested pH range. The remaining pKa in the H39N-catalyzed reaction seems to be shifted towards higher
pH. (D) For the reduction reaction kcat/Km decreases with higher pH, with the pKa value of the H39N mutant shifted towards higher pH as compared to the wild type.
straight-forward, however, in that the order of substrate binding (A
before B or possibly, vice versa) has not been unequivocally deter-
mined from direct binding studies. The relative affinities, however,
suggest that A does bind before B in ADH-A (compare K1 and K3 in
Table 4).
Increasing the pH to 8.0 changes the inhibition pattern so that
NADH now acts as a mixed-type inhibitor of NAD+ (Table 3, Fig. 6B).
In order to explain this observation an isomerization step of the free
enzyme (E ↔ E*) has to be invoked and the resulting model may
be described by an Iso Theorell–Chance mechanism [41] (Fig. 5C).
Clues to the actual structural meaning of this invoked isomerization
could be found from comparisons with the horse liver isoenzyme.
It has been firmly established that the horse liver ADH partitions
Table 3
Product inhibition at pH 7 and 8. A, B, Q and P are represented by NAD+, S-5, NADH
and 6 respectively. Schematic representations of the proposed model mechanisms
are presented in Fig. 5B and C. The double-reciprocal plots are presented in the
Supplementary Information.
pH 7
pH 8
Fig. 4. Effect of zinc addition on retaining enzyme activity. The activity of wild-
type ADH-A decreases over time when stored in 4 ◦C in zinc-free buffer to a level
of approximately 50% over a time period of 60 days (filled symbols). Storing ADH-A
in buffer containing 10 ␮M zinc sulfate (unfilled symbols) stabilizes the enzyme,
retaining approximately 95% after 30 days of storage.
Varied substrate
A
Varied substrate
A
Inhibitor
B
B
P
Q
Mixed type
Competitive
Competitive
Mixed type
Mixed type
Mixed type
Competitive
Mixed type

E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
75
closed form the binding site for the coenzyme is inaccessible imply-
ing an ordered binding mechanism in which the cofactor binds to
the enzyme in its open state (E) thereby favoring the transition to
the closed state (E*) as schematically drawn in the model in Fig. 5A.
The mechanism model in Fig. 5A is based on the model described
for the horse liver enzyme [44].
How to link these movements in the horse liver enzyme to the
observed shift in the inhibition pattern of the bacterial ADH-A is not
obvious but the crystal structures of ADH-A provide some space
for speculation. There are two available crystal structures deter-
mined at different pH values, 2xaa at pH 8.5 and 3jv7 at pH 5.5
[21]. Both of these structures describe the coenzyme-bound forms
so it follows that there is no structural evidence for an “open”
form of the ADH-A apoenzyme. A comparison of the crystal struc-
tures of ADH-A determined at pH 5.5 (3jv7) with that of the horse
liver enzyme in the closed conformation (1lde) [46] demonstrates
that although the sequence similarity of these two enzymes is
poor in the corresponding regions of the active sites, the positions
of the loops closing over the nucleotide binding sites are virtu-
ally superimposable between the two structures (Fig. 2C). In the
closed conformation of the horse liver enzyme, the side-chain of
Val294 comes in van der Waals contact with the pyridine ring of
the bound coenzyme [45,47]. MD simulations have pinpointed this
residue as part of a segment undergoing anticorrelated motions
to that of the cofactor and thereby contributing to ground state
activation of the substrates [48,49]. In ADH-A, the residue corre-
sponding to Val294 is Ile271 (Fig. 2C). In the R. ruber enzyme this
loop contains a non-conserved His residue, adjacent to Ile271. In
both ADH-A structures, the N-␦ of His272 forms a hydrogen bond
with the backbone carbonyl oxygen of Val247 (Fig. 2D). Although,
the crystallographic B-factors of the imidazole ring of His272 are
relatively high, suggesting that this group is structurally flexible,
it is conceivable that this interaction may contribute to stabilize
the closed form also of the apoenzyme and that this interaction
could be differentially affected by the differences in pH. This would
require a conformational change back to the open state in order to
form a new productive enzyme-cofactor complex. More studies are
clearly needed to clarify the present observations of pH-dependent
product inhibition.
Fig. 5. Kinetic mechanism models based on product inhibition and pre-steady state
kinetics. (A) The full model including enzyme isomerization steps triggered by
nucleotide binding, as previously described for the horse liver enzyme [44]. (B) The
Theorell–Chance (hit-and-run) mechanism which best describes the product inhi-
bition results for the oxidation of S-5 at pH 7.0 where NADH and 6 act as competitive
inhibitors of NAD+ and S-5, respectively. The apparent rate constants k2 , k
are:
ꢀ
ꢀ
−2
ꢀ
ꢁ
ꢀ
ꢁ
k2ꢀ = k3kꢀ /
k
+ kꢀ
−4
and k−ꢀ 2 = k−3k−ꢀ 4
/
k
−3
+ kꢀ
;
k4ꢀ = k k / k + k5 and
(
−4
)
−3
4
5
4
4
4
k−ꢀ 4 = k−4k / k + k5
.
(
)
−5
(C) The Iso Theorell–Chance mechanism which agrees with the product inhibition
data at pH 8.0 where NADH acts as a mixed-type inhibitor of NAD+. The difference
between (B) and (C) is an added enzyme isomerization step that is substrate inde-
pendent. (D) Model (C) with an added step for non-productive binding of the alcohol
substrate. The determined values of rate and thermodynamic constants are given in
Table 4.
The microscopic rates were analyzed by monitoring the tran-
sient changes in intrinsic protein tryptophan fluorescence upon
binding of coenzyme or the NADH fluorescence during the redox
chemistry. NAD+ and NADH binding to ADH-A followed single
between two distinct conformations where a more closed form is
stabilized upon binding of the coenzyme [42–44]. In the closed
form the active site and the catalytic zinc ion are shielded from
solvent rendering the micro-environment more hydrophobic. The
ternary enzyme-substrate complex also stabilizes binding modes
of the second substrate that facilitate hydride transfer [45]. In the
exponential curves (Fig. 7C and D) where the observed rates (kobs
)
were best modeled by a hyperbolic dependence of coenzyme con-
centration as described by Equation (5) (Fig. 7A and B). We also
attempted to fit a linear one-state model (Equation (4)) to the data
Fig. 6. Product inhibition of NADH versus NAD+ at pH 7 and 8. Steady state kinetics of the pseudo-first order reaction of S-5 (saturating concentration) and NAD+ (varied
concentrations) in the presence of increasing concentrations of NADH as inhibitor. The Michaelis–Menten equation was fitted to the experimental data. The fits are presented
as double-reciprocal plots. (A) pH 7.0 results in competitive inhibition. (B) pH 8.0 displays mixed inhibition. Table 3 summarizes the product inhibition pattern of ADH-A at
pH 7 and pH 8 for the reaction in the oxidative direction.

76
E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
Fig. 7. Pre-steady state kinetics. Observed rates, kobs, versus (A), [NAD+] and (B), [NADH] Solid lines represent the fits to a two-state model (Equation (5)) and the dashed
line to a one-state model (Equation (4)). Error bars represent standard deviations, n ≥ 8. (C) and (D) show in grey averages of experimental traces (n ≥ 8) of fluorescence
decay in the presence of (C) 25 ␮M NAD+ and 1.1 ␮M ADH-A, and (D), 25 ␮M NADH and 2.2 ␮M ADH-A. Solid black lines represent a single exponential fit to the data. (E)
Substrate dependence of kobs for the oxidation of S-5 (filled squares) and R-5 (unfilled squares). Solid lines represent the fits to Equation (6). (F) and (G) show in grey averages
of experimental traces (n ≥ 8) of fluorescence increase in the presence of (F) 4.4 mM S-5, 0.4 mM NAD+ and 0.6 ␮M ADH-A, and (G), 5.7 mM R-5, 0.4 mM NAD+ and 2.3 ␮M
ADH-A. Solid lines represent a single exponential fit to the burst phase of the fluorescence traces. See Table 4 for determined values of kinetic parameters. Residual plots of
the respective fits are shown in boxes.
Table 4
Microscopic rates and equilibria during catalyzed oxidation of 1-phenylethanol and reduction of acetophenone. Microscopic rates and thermodynamic constants were
determined at pH 8.0 during pre-steady state or at steady state, as described in the Section 2. Numbering refer to the model mechanism in Fig. 5A. K1 and K7 are the
dissociation constants of E•NAD+ (k−1/k1) and E•NADH (k7/k−7), respectively. K3 is the apparent dissociation constant of S- or R-5 from the E*•NAD+•5 ternary complex and
is the ratio k−3/k3.
Substrate
K1 (␮M)
k2 (s−1
)
k
(s−1
3
)
K3 (mM)
k4 (s−1
)
k6 (s−1
)
k
(s−1
−6
)
K7 (␮M)
kcat (s−1
)
Km (mM)
−2
S-5
R-5
6
240 100
820 300
22
2.5 0.5
630 40
220 30
–
46 200
710 100
22 20
80 20
0.45 0.005
36 0.8
0.63 0.05
0.94 0.04
1.2 0.09
12
–
2

E. Hamnevik et al. / Journal of Molecular Catalysis B: Enzymatic 99 (2014) 68–78
77
which would describe the process of binding of the free enzyme
(E) in the open state and coenzyme to form the binary complex
in the closed states (E*•NAD(H)). This simpler model was, how-
ever, rejected after statistical analysis. The two-state model invokes
rapid pre-equilibria of the binary E•NAD(H) complexes followed by
subsequent isomerization steps from the open to closed conforma-
tions. The extracted parameters describe the dissociation constants
of the binary complexes in the open state(s) (K1 and K7) and the
rates of isomerization from the open to the closed state(s) (k2 and
low (apparent) values of kcat and Km would be increased if R-5 was
bound only in a productive mode.
4. Conclusions
The main reason for our group to study the enzymology of the
R. ruber ADH-A was its potential as a biocatalyst. We have estab-
lished the substrate scope and kinetic mechanism of a potentially
important enzyme for biotechnological applications and the gath-
ered results will provide important guidance in future efforts to
tailor its function to suit a role as an efficient chemical catalyst.
The data adds to previous knowledge on the catalytic function of
class I alcohol dehydrogenases and demonstrates the functional
relatedness between evolutionary distant relatives such as the R.
ruber and horse liver isoenzymes. The fact that the catalytic efficien-
cies expressed as kcat in both these cases are limited by structural
isomerizations, emphasizes the notion that high turnover num-
bers are not strongly selected for in natural evolution but rather
the catalytic efficiencies during non-saturating concentrations of
substrates. Both these enzymes can be considered as efficient if
comparing their kcat/Km values for their preferred substrates which
are in the range of 105–106 s−1 M−1. The findings that ADH-A cat-
alyzes the regioselective oxidation of aryl-substituted vicinal diols
place it as an important addition to the biocatalytic tool-box. The
enantioselectivity between favoring S-5 over R-5 appears to be
due to non-productive binding of the least preferred enantiomer.
It follows that manipulating non-productive substrate binding by
mutagenesis and directed evolution may be a new general route
to enzyme variants displaying improved or altered stereoselectiv-
ities. The proven activity with diols 8 and 10 forms a baseline for
improvement by upcoming directed evolution.
k
−6) and back to the open state (k and k6) (Fig. 5A). It should be
−2
noted that the horse liver isoenzyme exhibits an identical kinetic
behavior in its binding of the nicotinamide coenzymes [19,44]. The
values of the microscopic rates and thermodynamic constants are
presented in Table 4.
The NADH fluorescence increase when E•NAD+ was rapidly
mixed with either enantiomer of 5, also proceeded as single expo-
nential events followed by steady state slopes (Fig. 7F and G). The
burst phase of the curves was analyzed to determine the rates of
alcohol oxidation (k4). The substrate dependencies of kobs were
in both cases hyperbolic allowing for determination of both the
oxidation rates and the apparent dissociation constants (K3) of
the alcohols from the respective ternary complexes (Fig. 7E and
Table 4). We were unable to detect any burst kinetics in the reduc-
tion of 6 preventing determination of k and K5 (cf. Fig. 5A). The
−4
reason for the lack of signal may be due to that the rate of hydride
transfer from NADH to acetophenone is too fast (>1000 s−1) to be
detected outside the dead time of the equipment.
From the combined results from the steady state and transient
kinetics it is clear that the rate-determining steps for the reduc-
tion of 6 and the oxidation of S-5 are the isomerizations from the
respective closed binary complex(es) to the corresponding open
complex(es). The isomerization rate from the closed E*•NADH com-
plex to the open complex (k6) is approximately 50 s−1 which is in
Acknowledgments
S-5
the same range as k
(80 s−1) and k has a value of 22 s−1, also
−2
cat
similar to kc6at (36 s−1). It should be noted at this point that the
corresponding isomerization step of the closed state of the binary
complex of the horse liver enzyme and NAD+ has also been shown
to be rate-limiting in the catalyzed reduction of acetaldehyde [44].
We have, however, not been able to pinpoint the rate-
determining step in the catalyzed oxidation of the R-5 enantiomer
from the transient kinetics studies. The slowest identified step is
the described isomerization from the closed to the open binary
E•NADH complex. This rate, however, is still 100-fold faster than
kcRa-t5. One possible explanation is that this enantiomer can bind
in different modes at the active site, with a distribution of pro-
ductive and non-productive modes (Fig. 5D). The fact that the Km
value is significantly lower than the apparent dissociation constant
(K3) supports the idea of non-productive substrate binding. This
would also affect the turnover number to the same degree but leave
kcat/Km unaffected [50]. The determined kcat value should there-
fore, by this argument, be underestimated and k6 could possibly be
rate-limiting also for the R-5 oxidation. The fact that the same non-
chiral product is formed in the reactions with either enantiomer
of 5 also speaks in favor of that non-productive substrate binding
is a reason to the relatively low KmR-5. A (rate-limiting) step down-
stream of acetophenone formation would affect the reactions with
R- or S-5 equally. Isopropanol (2) is oxidized with approximately
the same efficiency as R-5 by ADH-A, as judged by the respective
kcat/Km values (Table 1). The isopropanol oxidation, however, dis-
plays values of kc2at and Km2 that are 100-fold larger than those for
the R-5 reaction. Isopropanol can be viewed as a phenyl-truncated
derivative of 5. Lacking the phenyl substituent is expected to result
in poorer binding affinity at the ADH-A active site and thereby faster
equilibration of binding modes. The larger turnover number and a
relatively higher Km of 2 indicate poorer stabilization of productive
(and non-productive) ternary complexes. Hence, the determined
Plasmid pREP4-groESL was kindly provided by Dr. Martin
Stieger (F. Hoffman-La Roche Ltd. Basel, Switzerland). The work was
supported by Swedish Research Council (VR 621-2011-6055).
Appendix A. Supplementary data
Supplementary data associated with this article can be found,
in the online version, at http://dx.doi.org/10.1016/j.molcatb.
2013.10.023.
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