Tài liệu Báo cáo khoa học: Structural and mechanistic aspects of flavoproteins: probes of hydrogen tunnelling pptx

MINIREVIEW
Structural and mechanistic aspects of flavoproteins:
probes of hydrogen tunnelling
Sam Hay, Christopher R. Pudney and Nigel S. Scrutton
Manchester Interdisciplinary Biocentre and Faculty of Life Science, University of Manchester, UK
Introduction
There is now fairly widespread recognition that
enzyme-catalysed C–H bond cleavage reactions can
occur by quantum mechanical tunnelling [1–5]. The
role of protein dynamics in these reactions is still hotly
debated and it has been proposed that promoting
vibrations, nonequilibrated fast (sub-ps) dynamics,
could modify the reaction barrier and profoundly
influence the reaction rate [4,6–12]. In recent years, we
have investigated H-transfer reactions in a number of
enzymes, primarily quinoprotein [4,13,14] and flavo-
protein [8,15–20] systems. Using a combination of
experimental and computational approaches, we have
shown that H-transfer reactions can occur by ‘deep’
tunnelling and the reaction can be enhanced by local-
ized dynamics in the enzyme active site – putative pro-
moting vibrations. Although it is fairly well established
that enzymatic H transfers often involve tunnelling,
the role of promoting vibrations remains contentious
[21]. In this minireview, we describe experimental
methods we have recently employed and developed to
probe the role of environmental coupling ⁄ promoting
vibrations in H-transfer reactions in the Old Yellow
Enzyme (OYE) family of flavoproteins.
Hydrogen tunnelling
Because of wave ⁄ particle duality, electrons and light

atoms have appreciable (de Broglie) wavelengths. The
Keywords
high pressure; H-tunneling; kinetic isotope
effect; kinetic isotope fractionation; multiple
reactive conformations; Old Yellow Enzyme;
promoting vibration; protein dynamics;
quantum mechanics; stopped-flow kinetics
Correspondence
N. S. Scrutton, Manchester Interdisciplinary
Biocentre and Faculty of Life Science,
University of Manchester, 131 Princess
Street, Manchester M1 7ND, UK
Fax: +44 161 306 8918
Tel: +44 161 306 5152
E-mail:
(Received 23 December 2008, revised 28
April 2009, accepted 1 May 2009)
doi:10.1111/j.1742-4658.2009.07121.x
At least half of all enzyme-catalysed reactions are thought to involve a
hydrogen transfer. In the last 10 years, it has become apparent that many
of these reactions will occur, in part, or in full, by quantum mechanical
tunnelling. We are particularly interested in the role of promoting vibra-
tions on H transfer, and the Old Yellow Enzyme family of flavoproteins
has proven to be an excellent model system with which to examine such
reactions. In this minireview, we describe new and established experimental
methods used to study H-tunnelling in these enzymes and we consider
some practical issues important to such studies. The application of these
methods has provided strong evidence linking protein dynamics and H-tun-
nelling in biological systems.
Abbreviations

DHFR, dihydrofolate reductase; EIE, equilibrium isotope effect; ET, electron transfer; GO, glucose oxidase; KIE, kinetic isotope effect; MR,
morphinone reductase; OYE, Old Yellow Enzyme; PETNR, pentaerythritol tetranitrate reductase; RHR, reductive half-reaction.
3930 FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS
wavelength of H (used here to denote H
+
,H
•
and H
)
)
is $1A
˚
and thus similar to a typical bond length,
whereas the wavelength of deuterium is shorter by a
factor of $
ffiffiffi
2
p
. As a consequence, the position of H
(and to a lesser extent, D) is somewhat diffuse, and
H transfer may involve an appreciable degree of quan-
tum mechanical tunneling, in which H or D transfer
occurs by ‘tunnelling’ through part of the reaction
barrier rather than by passing over the barrier as is the
case in a classical transition-state reaction [22]. It is
accepted that long-range electron transfer (ET) reac-
tions occur by tunnelling [23,24] and we now have
nearly 20 years of both experimental and computa-
tional evidence demonstrating that H-tunnelling reac-
tions can also occur during enzyme-catalysed reactions

[1–5]. It is possible to computationally estimate the
extent to which tunnelling plays a role during an
H-transfer reaction. In dihydrofolate reductase,
hydride transfer occurs by tunnelling $50% of the
time [25,26], with the remainder of the H transfer
occurring by an over-the-barrier mechanism. Con-
versely, in soybean lipoxygenase-1 [27], aromatic amine
dehydrogenase [4] and the flavoprotein morphinone
reductase (MR) [28], calculations have shown that at
least 99% of H transfer occurs by tunnelling. These
reactions can be thought of as ‘deep’ tunnelling reac-
tions because the H tunnels a relatively long way
below the top of the reaction barrier.
The rate of a nonadiabatic (deep tunnelling)
H transfer can be described using modified Marcus
theory [23]:
k
tun
¼
2p
h
ffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
1
4pkk
B
T
r
V
jj
2

F:C:ðÞexp À
DG
z
k
B
T
!
ð1Þ
where V is the electronic coupling, F.C. is the Frank–
Condon nuclear wave function overlap (related to the
de Broglie wavelength of the H or D) and DG
à
is the
Marcus activation energy. The activation energy is
described by the driving force, DG
0
, and reorganization
energy in the standard way:
DG
z
¼ DG
0
þ k
ÀÁ
2
=4k ð2Þ
The driving force dependence of H transfer in the
flavoprotein glucose oxidase (GO) was investigated by
Brinkley & Roth [29]. The endogenous FAD was
substituted with other chemically modified flavins with

differing redox potentials, and these authors showed
that the apparent rate of H transfer obeys Eqns (1)
and (2) and the reorganization energy of this reaction
appears to be large ($280 kJÆmol
)1
). We have since
shown a driving force dependence during the reduction
of the quinoprotein aromatic amine dehydrogenase
with p-substituted phenylethylamine substrates [30]
and estimated the reorganization energy for the reac-
tion of this enzyme with tryptamine to be 250–300 kJÆ
mol
)1
[9]. From the little experimental evidence cur-
rently available, it appears that it is appropriate to
describe H-tunnelling reactions using Marcus theory,
and that a general feature of these reactions may be a
large reorganization energy.
The Frank–Condon term in Eqn (1) has been
described by Kuznetsov & Ulstrup:
F:C:
0;0
¼
Z
r
0
0
exp Àl
i
x

i
Dr
2
=2h
ÀÁÂÃ
exp ÀE
X
=k
B
TðÞdX
ð3Þ
where l and x are the mass and frequency of the
transferred H or D and E
x
is the environmental energy
or promoting vibration, which reduces the H-transfer
distance from an equilibrium distance, r
0
,byDr =
(r
0
) r
X
) [9,31,32]. The kinetic isotope effect (KIE =
k
H
⁄ k
D
) arises because of differences in the mass,
frequency and consequently the transfer distance of H

and D. Experimentally, the identification of promoting
vibrations is extremely challenging and, as yet, there is
no method to directly visualize such vibrations because
they occur in the relatively inaccessible THz region.
The first experimental evidence for a role of environ-
mental coupling during H-tunnelling reactions in
enzymes [5,13] was inferred from observations of KIEs
with aberrant temperature dependencies that do not
conform to the predictions of semiclassical transition
state theory or Bell-type quantum correction models
[33]. However, we have recently shown that the KIE
temperature dependence (DDH
à
, see below) is not a
reliable diagnostic of environmental coupling [9] and
other experiments are required to corroborate predic-
tions based solely on DDH
à
values.
Measurement of H-transfer reactions
in OYEs
The OYE family of flavoproteins comprise a large
group of FMN-containing NAD(P)H-dependent oxid-
ases. We have concentrated our studies on two homol-
ogous OYEs: MR isolated from Pseudomonas putida
M10 and pentaerythritol tetranitrate reductase (PET-
NR) from Enterobacter cloacae. For reference, in
Table 1, we summarize the flavoproteins for which a
specific H-tunnelling study has been performed. Gener-
ally, the KIEs of H transfer in flavoproteins are fairly

modest (< 10) but the temperature dependencies of
these KIEs are quite varied (Table 1).
S. Hay et al. Hydrogen tunnelling in biological systems
FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS 3931
The reductive half-reaction (RHR) of MR and PET-
NR occurs in three steps:
E
ox
þ NADðPÞH À!
binding
½E
ox
Á NADðPÞH
CT
À!
reduction=
HÀtransfer
E
red
Á NADðPÞ
þ
À!
product
release
E
red
þ NADðPÞ
þ
ð4Þ
MR reacts only with NADH, whereas PETNR has a

preference for NADPH. It is sometimes possible to
trap the binary CT complex in OYEs by substituting
NAD(P)H with the nonreactive analogue 1,4,5,6-tetra-
hydroNAD(P)H (NAD(P)H
4
) [19]. MR binds NADH
4
with K
d
= 0.35 mm [34] and we have recently solved
the X-ray crystallographic structure of MR in complex
with NADH
4
to a resolution of 1.3 A
˚
[19]. The struc-
ture of the active site in MR is shown in Fig. 1, as is
the proposed mechanism of hydride transfer ⁄ FMN
reduction. In MR and PETNR, the H transfer is ste-
reospecific because the NAD(P)H nicotinamide moiety
can only bind in one orientation within the active site,
which places the pro-R (transferred) primary hydrogen
(H
p
) directly over the FMN N5 acceptor atom
(labelled in Fig. 1).
The reduced OYEs can be reoxidized by molecular
oxygen (k $ 1s
)1
) [35] or with various oxidative sub-

strates – one generic substrate being cylcohexen-1-one.
With many of the oxidative substrates tested, the oxi-
dative half-reactions of MR and PETNR are fully rate
limiting during steady-state turnover. Consequently,
the steady-state KIE on the RHR hydride transfer is
unity and steady-state analysis is clearly not appropri-
ate to study these reactions. However, in MR, we have
measured a KIE of 3.5 ± 0.2 on k
cat
for the hydride
transfer from the reduced FMN to cylcohexen-1-one
Table 1. Kinetic isotope effects observed in selected flavoproteins. CO, choline oxidase; DD, human class 2 dihydroorotate dehydrogenase;
FDTS, flavin dependent thymidylate synthase; GO, glucose oxidase; MAO A ⁄ B, monoamine oxidase A ⁄ B; MR, morphinone reductase; PAO,
L-phenylalanine oxidase; PETNR, pentaerythritol tetranitrate reductase; TMADH, trimethylamine dehydrogenase; TSOX, heterotetrameric sar-
cosine oxidase; nd, not determined or reported. The KIEs are for pre-steady-state flavin reduction by the denoted substrate unless otherwise
stated. Isotope effects are H ⁄ D unless otherwise stated.
Enzyme Substrate 1° KIE DDH
à
(kJÆmol
)1
) A
àH
: A
àD
Ref.
PETNR b-NADPH 7.0 ± 0.04
a
6.5 ± 2.76
a
0.51 ± 0.04 [15,18]

MR b-NADH 6.8 ± 0.1
a
7.4 ± 1.5
a
0.12 ± 0.09 [15,18]
MR cylohexen-1-one 3.5 ± 0.2 )0.5 ± 1.8 4 ± 2 [15]
GO 2-deoxyglucose $10
b
$0
b
$10
g
[73]
CO choline 10.6 ± 0.6
b
1.0 ± 0.3
b
14 ± 3 [74]
MAO A benzylamine 8.0
e
n.d. n.d. [75]
MAO B p-methoxy-benzylamine 8.9 ± 1.6
c
$9.0
c,e
$0.2
g
[76]
FDTS b-NADPH 25 ± 6
d

n.d. n.d. [77]
DD dihydroorotate 3.77 ± 0.08 n.d. n.d. [78]
TMADH trimethylamine 4.6 ± 0.4 0.5 ± 5.2
f
7.8 ± 1.2 [16]
PAO
L-phenylalanine 5.4 ± 0.3 0.2 ± 0.03 5.2 ± 0.2 [72]
TSOX sarcosine 7.3
e
0.6 ± 2.1 5.4 ± 0.4 [17]
a
Revised from previously reported, manuscript in preparation.
b
Data from k
cat
⁄ K
m
measurements.
c
Data not corrected for the calculated
commitment to catalysis.
d
Data from H ⁄ T isotope effect.
e
No error given.
f
Data for the H172Q mutant.
g
Calculated from the KIE and
DDH

à
values.
A
B
Fig. 1. (A) A model of the active site of NADH-bound morphinone
reductase based on pdb 2R14 [19]. Residues which form hydrogen
bonds (dotted lines) to the bound NADH are shown, as are the
NADH nicotinamide pro-R (H
p
, the transferred H) and pro-S (H
s
)
hydrogens and the FMN N5 (acceptor). (B) The proposed reaction
mechanism of hydride transfer ⁄ FMN reduction during the reductive
half reaction in old yellow enzymes.
Hydrogen tunnelling in biological systems S. Hay et al.
3932 FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS
[15]. There is also a solvent KIE of 2.3 ± 0.3 and a
double isotope effect (measured with deuterated FMN
in D
2
O) of 8.2 ± 1.4. The rule of geometric mean [36–
38] states that multiple KIEs should be described by:
KIE
a;b
¼ KIE
a
 KIE
b
ð5Þ

and in the case of the oxidative half-reaction of MR,
the observed double KIE should be: 3.5 · 2.3 = 8.0,
which is in agreement with the observed value of
8.2 ± 1.4 [15]. Although it has been argued that vio-
lation of the rule of geometric mean may be used as
evidence for H-tunnelling [36–38], the oxidative KIEs
in MR are not measurably temperature dependent – a
diagnostic of ground state H-tunnelling [15].
Using a stopped-flow spectrometer, it is possible to
determine most of the rate constants for the steps in the
OYE RHR reaction (Eqn 4) above. However, care must
be made to keep the samples anaerobic by either using
an anaerobic glove box or by adding glucose ⁄ GO. The
binary complex has a characteristic p-p charge transfer
(CT) absorbance and NAD(P)H binding and dissocia-
tion can be measured by following the formation of this
CT absorbance at, for example, 555 nm while perform-
ing a concentration dependence [35,39]:
k
obs
¼ k
off
þ k
on
½NADðPÞHð6Þ
Similarly, the rate of hydride transfer can be deter-
mined because H transfer is concomitant with FMN
reduction. By following the bleaching of FMN absor-
bance at $465 nm while performing a concentration
dependence [15,18,35,39], it is possible to characterize

the RHR:
k
obs
¼ k
ox
þ k
red
½NADðPÞH
k
off
=k
on
ðÞþ½NADðPÞH
ð7Þ
In both MR and PETNR, at room temperature, k
on
,
the apparent rate of coenzyme binding is $10
6
m
)1
Æs
)1
.
The rate of NAD(P)H dissociation, k
off
,is$10
2
s
)1

and the apparent rate of H transfer, k
red
, is 56 and
33 s
)1
in MR and PETNR, respectively [18,35,39].
Product (NAD(P)
+
) inhibition of MR and PETNR is
very weak suggesting that NAD(P)
+
rapidly dissoci-
ates from the active site once FMN reduction occurs.
We have been unable to measure the reverse rate of
hydride transfer, k
ox
, in either enzyme and it appears
to be close to zero [18,40]. We have also determined
the driving force for hydride transfer during the RHR
of MR with NADH to be $60 kJÆmol
)1
[40], which
is also consistent with an effectively irreversible
H transfer.
We have mutated various amino acid residues within
the active site of MR and PETNR [19,41,42]. In the
wild-type enzymes, FMN reduction occurs as a mono-
exponential process (Fig. 2) – greatly simplifying the
stopped-flow analysis. In the H186A and N189A
active-site mutants in MR (Fig. 1A), FMN reduction

kinetics become more complex – i.e. multi-exponential
[41]. As an extreme example, we have measured at
least four kinetically resolved components of FMN
reduction in the N189A mutant of MR, each with a
significant KIE (Fig. 2) [19], and each able to intercon-
vert [42a]. We have attributed this complexity to
the formation of multiple reactive configurations in the
mutant enzyme because of improper binding of the
NADH nicotinamide moiety within the active site. Of
concern is that, had we performed a steady-state analy-
sis of this mutant, we would not have observed this
heterogeneity and the mutant enzyme would have
appeared to be quite similar to the wild-type enzyme.
Caution is therefore needed when using steady-state
approaches to analyse tunnelling, especially with
mutant enzymes.
During the RHR of MR and PETNR, when pro-R
deuterated NAD(P)H (denoted (R)-[4-
2
H]-NAD(P)H)
is used in place of the protiated coenzyme, a significant
KIE is observed on hydride transfer (k
red
) but not on
k
off
⁄ k
on
(Table 1) [15,18]. Because the H transfer is
effectively irreversible and kinetically resolved from

coenzyme binding (no KIE on k
off
⁄ k
on
), the observed
KIE is essentially the intrinsic KIE. Using stopped-
flow methods, we have measured the temperature
dependence of the rate of H transfer in both MR and
PETNR. For convenience, we tend to measure k
obs
(in
0.20
0.15
0.10
0.05
0.00
0.01
0.0
0.1
0.00
0.05
0.10
0.15
0.20
Absorbance
Absorbance
20 40
0.1
wt
N189A

1 10 100
t·s
–1
t·s
–1
Fig. 2. Multiple reactive conformations during an H-transfer reac-
tion. Stopped-flow traces of the observed FMN reduction during
the reaction of the wild-type (wt) and N189A mutant of MR with
NADH. The wt trace is fit to a single exponential and the N189A
trace to a 4-exponential function – see the main text for more
details. (Inset) The same data on a split-axis linear time scale.
Adapted from Pudney et al. [19].
S. Hay et al. Hydrogen tunnelling in biological systems
FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS 3933
the presence of saturating [NAD(P)H]) rather than k
red
(Eqn 7), although they give equivalent results
[8,15,18,19,35]. We define saturating coenzyme concen-
trations as [NAD(P)H] > 10 · K
S
, where K
S
= k
off
⁄
k
on
. Care must be taken because K
S
can be quite tem-

perature dependent [15,19,35]. We typically analyse
these data in terms of Eyring (transition state) theory:
ln
k
obs
T

¼ ln
k
B
h

þ
DS
z
T
À
DH
z
RT
ð8Þ
with KIE
obs
¼ k
H
obs
=k
D
obs
and the temperature dependence

on the KIE:
DDH
z
¼ DH
z
D
À DH
z
H
¼ DE
a
ð9Þ
In wild-type MR and PETNR it is generally possible
to determine observed rate constants with an accuracy
of $1% (measured with different samples on different
days). It is then possible to determine KIEs to an
accuracy of 2-5% and DDH
à
with an error of 1-2 kJÆ
mol
)1
[18].
The use of KIE analyses relies heavily on the ability
to obtain isotopically pure substrates or coenzymes.
One of the reasons OYEs are particularly attractive to
study is the ability to create stereospecifically labelled
isotopologues of NAD(P)H (described in the next sec-
tion). This allows the measurement of 1° KIEs, 2° KIEs
and double KIEs – where both H
p

and H
s
are deuterated
[18]. We have been able to use stopped-flow methods to
measure quite accurately both the magnitude and tem-
perature dependence of a-2° KIEs during the RHR in
MR and PETNR [18,40], and for hydride transfer in the
thermophilic dihydrofolate reductase (DHFR) from
Thermotoga maritima [43]. The equilibrium isotope
effect (EIE) on NAD(P)H oxidation was measured by
Cook & Cleland to be 1.13 [44]. The observation of a-2°
KIEs values larger than the EIE was rationalized by
Huskey & Schowen [36] because of coupling of the
motion between the 2° hydrogen (labelled in Fig. 1A)
and the 1° (transferred) hydrogen during an H-tunnel-
ling reaction. We have measured identical a-2° KIE val-
ues of $1.2 in MR and PETNR, which are significantly
larger than the EIE [18,40]. We have also measured the
double KIE in MR [18] and shown that in this reaction,
the rule of geometric mean (Eqn 5) is most likely vio-
lated [39]. We have shown computationally that the
H transfer in MR occurs by deep tunnelling [28] so Hus-
key & Schowen’s [36] interpretation of inflated 2° KIEs
would seem plausible. However, we have measured a
normal (KIE £ EIE) and temperature-independent a-2°
KIE in TmDHFR, yet this reaction proceeds by 50-80%
tunneling, depending on the temperature [25,43]. A simi-
lar observation has been observed in the Escherichia coli
DHFR [45]. Consequently, it would appear that inflated
a-2° KIEs may be indicative of a tunnelling contribution

to the H transfer, but normal KIEs do not rule out tun-
nelling [43]. A similar argument can be made for 1°
KIEs – although KIEs £ 7 can be explained using transi-
tion state theory, the KIE arising from a full tunnelling
reaction (described by Eqns 1–3) can be any value
[10,46].
Preparation of coenzymes
The methods of coenzyme deuteration are well
described [47–50], but are typically for microscale syn-
theses, $1 mg. This can be an issue for stopped-flow
experiments, substrate-binding titrations and crystallo-
graphic studies when a very large amount of the sub-
strate may be required; typical NAD(P)H saturation
constants for OYEs are 0.1-1 mm and as an example,
a typical measurement, by stopped-flow, of the temper-
ature dependence of the 1° KIE of an OYE may
require $100 mg of isotopically pure substrate. For
reference, we briefly describe our preferred methods of
synthesis for large-scale ($1 g) preparations of all
three deuterated NADH and NADPH isotopologues:
(R)-[4-
2
H]-NAD(P)H, (S)-[4-
2
H]-NAD(P)H and (R,S)-
[4,4-
2
H
2
]-NAD(P)H. These syntheses typically yield >

95% isotopologue purity (based on
1
H NMR spectra,
see Pudney et al. [18] for examples), with the corre-
sponding impurity being the protiated coenzyme.
Kohen [49] recently developed syntheses for extremely
high-purity NADPH isotopologues and the method of
McCracken et al. [49] has been reported to yield >
99% isotopologue purity – a purity necessary when
performing competitive isotope experiments. We also
describe our synthesis of 1,4,5,6-tetrahydroNAD(P)H.
We typically prepare (R)-[4-
2
H]-NADH by the ste-
reospecific reduction of NAD
+
(500 mg) with 1-[
2
H
6
]-
ethanol (1 g) using yeast alcohol dehydrogenase
(200 U) and aldehyde dehydrogenase (100 U) in
20 mm Taps pH 8.5 (20 mL) at room temperature.
This method is a slight modification of the procedure
reported in Viola et al. [48]. (R)-[4-
2
H]-NADPH is pre-
pared through a stereospecific reduction of NADP
+

(300 mg) with 1-[
2
H
6
]-isopropanol (1 g) using
NADP
+
-dependent alcohol dehydrogenase from Ther-
moanaerobacter brokii (100 U) in 20 mm Taps pH 8.5
(50 mL) at 42 °C. These reactions are deemed com-
plete when A
340
stopped increasing and A
260
:A
340
<3,
typically after 1 h. (S)-[4-
2
H]-NADH and (S)-[4-
2
H]-
NADPH are prepared through stereospecific reduction
of NAD
+
(500 mg) and NADP
+
(500 mg), respec-
tively, with 1-[
2

H]-glucose (150 mg) using glucose
Hydrogen tunnelling in biological systems S. Hay et al.
3934 FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS
dehydrogenase (150 U) in Taps pH 8.5 (10 mL) at
room temperature. This method is a slight modifica-
tion of the procedures reported in Ottolina et al. [50]
and the reactions typically take 3 h. (R,S)-[4,4-
2
H
2
]-
NADH is prepared by stereospecific oxidation of
(S)-[4-
2
H]-NADH (300 mg) with 100 mm cyclohexen-
1-one catalysed by 10 lm MR in Taps pH 8.5
(10 mL). The deuterated NAD
+
is purified in the same
manner as for (R)-[4-
2
H]-NADH. (R,S)-[4,4-
2
H
2
]-
NADH is then prepared through a further stereospe-
cific reduction of [4-
2
H]-NAD

+
with 1-[
2
H]-glucose
(100 mg) and glucose dehydrogenase (100 U) in Taps
pH 8.5 (10 mL). (R,S)-[4,4-
2
H
2
]-NADPH can be pre-
pared in the same manner as (R,S)-[4,4-
2
H
2
]-NADH.
However, an NADPH-specific enzyme such as PETNR
must be used in place of MR. NADH
4
and NADPH
4
are prepared by maintaining a slight pressure
($1.2 bar) of hydrogen (> 99%) over a solution of
NAD(P)H (500 mg) and palladium-activated charcoal
(30 mg) in Tris ⁄ Cl pH 9.0 (5 mL) stirred on ice [19].
The reaction is stopped when no absorbance at
340 nm is observed and typical A
266
⁄ A
288
ratios of

$1.06 are obtained.
We purify the coenzymes using anion-exchange (Q-
Sepharose) chromatography, eluting NADH and
NADPH isotopologues (including the tetrahydro forms)
in $200 and $500 mm ammonium bicarbonate, respec-
tively [18]. All of the enzymes used in these syntheses
(excluding MR) are available from Sigma-Aldrich (St.
Louis, MO, USA) and the coenzymes are available
from Melford Laboratories (Chelsworth, UK). We use
extinction coefficients of 6.22 mm
)1
Æcm
)1
at 340 nm for
NAD(P)H isotopologues and 16.8 mm
)1
Æcm
)1
at 289
nm for NAD(P)H
4
[34]. Usually the enzymatic synthesis
of (R)-[4-
2
H]-NAD(P)H does not proceed to comple-
tion. We have observed that freezing or freeze-drying
the reaction volume before purification usually leads to
the formation of a significant impurity of undeuterated
NAD(P)H. Consequently, on this scale, it is important
to purify the reaction volume as quickly as possible after

synthesis has ceased. Also, one must take care to main-
tain the pH at $8.5 over the course of the enzymatic
synthesis, because acid catalysed decomposition of
NAD(P)H may be a significant contributor to substrate
(in)activity [51].
As an aside, we have recently investigated the effect
of incomplete coenzyme ⁄ substrate deuteration on the
observed KIE measured using stopped-flow methods
[39]. We found that, if there is a reversible chemical
step preceding H transfer and the reverse rate of this
step (k
off
in Eqn 4) is comparable with the rate of
H transfer, then kinetic isotope fractionation can
occur, leading to the formation of more protiated than
deuterated product. This fractionation also leads to an
overestimation of k
D
and consequently an underesti-
mation of the KIE. It appears that it is possible to cor-
rect for incomplete deuteration using a simple linear
relationship in an analogous fashion to that used to
correct steady-state data:
k
D
obs
¼ k
H
1 Àf
D

ðÞþk
D
f
D
ð10Þ
where f
D
is the fraction of substrate deuteration, which
can usually be determined quite accurately by
1
H NMR [18,48,52] or possibly MS [18,53]. In Fig. 3,
we use Eqn (10) to model the effect of partial deutera-
tion on the observed rate and KIE of an H transfer
reaction. If k
D
is underestimated then so too will be
DDH
à
and the effect of f
D
on the apparent temperature
dependence of the KIE is also shown in Fig. 3. We
determined Eqn (10) empirically and this relationship
is quite approximate. Nevertheless, we have been able
to correct the RHRs of MR and PETNR and also the
RHR of aromatic amine dehydrogenase with benzyl-
amine using Eqn (10) [39]. However, further studies
are required to confirm the general validity of this
correction method. That fractionation can occur
emphasizes the need for: (a) care in preparing high-

purity coenzymes, and (b) correction for small isotope
impurities in the analysis of tunnelling kinetics using
stopped-flow single turnover measurements.
Hydrostatic pressure
Hydrostatic pressure offers an alternative or comple-
mentary method to temperature with which to study
6

A

B
5
4
3
2
1
4
2
0
10
K
obs
/s
–1
KIE
obs
8
6
4
2

0
0.0 0.2 0.4 0.6 0.8 1.0
Fr
ac
ti
o
n
deu
t
e
r
a
ti
o
n
ΔΔH
‡
/ kJ·mol
–1
Fig. 3. The effect of substrate isotopic purity on the observed rate
of deuterium transfer (filled squares) and the corresponding KIE
(open circles) (A), and on the temperature dependence (B) of a
modelled H-transfer reaction. The data are modelled using Eqn (10)
with k
H
=5s
)1
, a KIE of 5 and various values of DDH
à
.

S. Hay et al. Hydrogen tunnelling in biological systems
FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS 3935
enzymatic reactions. Semiclassical transition-state the-
ory states that pressure effects on isotope effects arise
because of differences in vibrational frequencies [54–56]
and stretching vibrations are insensitive to pressures of a
few kbar [54]. Consequently, the KIEs of purely transi-
tion state reactions are expected to be insensitive to pres-
sure. Conversely, several chemical systems with inflated
KIEs indicative of a tunnelling contribution to the
H transfer have been shown to exhibit a significant pres-
sure-dependence of both rate and KIE [57]. Thus, in
principle, the pressure-dependence of an isotope effect
provides an excellent method for distinguishing between
transition state and tunnelling reactions.
The use of pressure to study enzymatic H-tunnelling
reactions was pioneered by Northrop, who, 10 years
ago, developed a model [58] for the pressure-depen-
dence of H-transfer reactions based on the Bell correc-
tion [33,58]. This model was then used quite
successfully to model the pressure dependence of
steady-state H-transfer reactions in alcohol dehydroge-
nase and aldehyde dehydrogenase [57–59]. We recently
performed a high-pressure stopped-flow study of the
hydride transfer during the RHR of MR with NADH.
The apparent rate of hydride transfer increased by
approximately twofold per kbar increase in pressure
and the 1° KIE also showed a small but significant
increase in magnitude with pressure (Fig. 4C).
Together, these observations could not be explained

using Northrop’s model [58], nor with a simple non-
adiabatic H-tunnelling model (e.g. Eqns 1–3) when
pressure simply causes a compression of the reaction
barrier [60]. However, we found that we could qualita-
tively model the data by invoking a promoting vibra-
tion that changes frequency with pressure [8]. We have
since refined this analysis and recently described a sim-
ple nonadiabatic H-tunnelling model which explicitly
includes pressure as a variable [61]:
k
H
=k
D
$ exp l
D
x
D
À l
H
x
H
½r
0
þ Dr:p
ÀÁ
2
=2h
no
 exp À l
D

x
D
À l
H
x
H
½k
B
T=h j
0
þ Dj:pðÞ
fg
ð11Þ
where r
0
is the average H-transfer distance, Dr is the
change in this distance with pressure, j
0
is the force
constant describing the promoting vibration and Dj is
the change in this force constant with pressure. Equa-
tion (11) can be used as a fitting function with four
adjustable parameters and the KIE can either increase
or decrease with increasing pressure when Dr and ⁄ or
Dj become significant (Fig. 4). Although this model is
oversimplistic, it is possible to use Eqn (11) to describe
a reaction in which both the apparent rate and KIE
increase (or decrease) with pressure [61]. The model
10
A

B
C
8
6
KIE
KIE
4
2
10
8
6
4
2
10
KIE
obs
8
6
4
2
2.0
1.5
1.0
0.5
0.0
3.2
3.3
3.4
3.5
3.6

1.0
0.5
0.0
–0.5
–1.0
2.0
1.5
1.0
0.5
0.0
0.02
0.01
dr/Å kbar
–1
dκ/J·m
–2
·kbar
–1
10
3
/T·K
–1
0.00
–0.01
–0.02
2.0
1.5
Pressure/kbar
Pressure/kbar
Press

ure/kbar
1.0
0.5
0.0
Fig. 4. A variable pressure H-tunnelling model (Eqn 3) [61]. The KIE
pressure dependence is modeled when (A) the frequency of the
promoting motion or (B) the H-transfer distance changes with pres-
sure. Positive values of Dj and Dr reflect increases in frequency
and distance with pressure, respectively. The data are modeled
with j =5JÆm
)2
, r
0
= 0.52 A
˚
(KIE
0
= 5) and only one parameter in
each plot is varied. It is possible for both Dj and Dr to vary with
pressure (as we have modelled in MR) [61], causing curvature in
the KIE versus pressure plots. We have also plotted (C) the com-
bined pressure and temperature dependence of the observed KIE
on hydride transfer during the reductive half reaction of morphinone
reductase. The data are taken from Hay et al. [8]. We have not plot-
ted error bars for clarity but the average error in the KIE for this
data set is ±5% and the minimum and maximum error is 1% and
18%, respectively.
Hydrogen tunnelling in biological systems S. Hay et al.
3936 FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS
should only be used qualitatively, but is useful to esti-

mate whether: (a) the tunnelling distance changes with
pressure, and (b) to confirm that there is environmen-
tal coupling and to determine whether the frequency of
this vibration is likely to change with pressure.
Although it seems intuitive that hydrostatic pressure
will ‘squeeze’ the enzyme and thus compress the
H-transfer distance (achieved by increasing the popula-
tion of enzyme–substrate conformers with shorter
H-transfer distances in an equilibrium distribution of
conformational states), until recently, this assumption
remained untested. Ewald [62] has shown that increasing
pressure causes a progressive shortening of the CT bond
in synthetic p-p complexes with an accompanying shift
to red wavelengths and increase in absorbance. We have
recently shown that increasing pressure also causes a
shortening of the CT bond (increase in CT absorbance)
in NADH
4
-bound MR, which we have interpreted as
pressure-induced barrier compression [34]. Using vari-
able pressure molecular dynamics simulations of
NADH-bound MR, we were able to corroborate this
finding [34]. We found that the heavy atom transfer dis-
tance has an approximately Gaussian distribution that
both narrows and shifts to shorter distances at elevated
pressures. It appears, at least in MR, that pressure does
not physically squeeze the microscopic reaction barrier,
but rather reduces the average barrier width, the macro-
scopic barrier, by restricting the conformational space
available to the NADH and FMN moieties within the

active site. Further studies are required to determine
whether this is a general phenomenon.
Other experimental probes
In addition to temperature and hydrostatic pressure, it
is possible to experimentally probe enzymatic H-tun-
nelling reactions using additional experimental parame-
ters and we briefly discuss the use of varying the
solvent composition to probe the effect of solvent
dielectric and viscosity on H transfer chemistry.
It is predicted from von Smoluchowski’s theory [63]
that the rate of a diffusion-controlled (bimolecular)
reaction will be inversely proportional to the bulk
solution viscosity. The effect of viscosity on a unimo-
lecular reaction is more complicated but can be
described in combination with the Eyring equation
according to Ansari et al. [64]:
k
obs
¼
k
B
T
h
1 þr
g þr

exp
DS
z
R

!
exp
ÀDH
z
RT
!
ð12Þ
where r, in units of viscosity, is the contribution of
the protein friction to the total friction of the system.
The activation entropy and enthalpy can be deter-
mined independently from the temperature dependence
of the reaction [65]. Solution viscosity has been used to
probe the role of dynamics in interprotein [65–68] and
intraprotein [69] ET reactions and protein rearrange-
ment after carbon monoxide dissociation from myoglo-
bin [64]. In general, the rates of ET reactions that are
conformationally gated decrease upon an increase in
solvent viscosity.
The viscosity dependence of several enzymatic
H-transfer reactions has now been investigated. Protein
dynamics can be affected by surface glycosylation and
this approach has been used by Klinman and cowork-
ers to study the viscosity dependence of the rate of
hydride transfer in GO [70,71]. These authors studied
various glycoforms of the enzyme (varying in the
extent of glycosylation) [70] and also replaced the
native polysaccharide with different polymeric forms
of polyethylene glycol [71]. A decrease in the ‘fitness’
of GO was observed when the apparent surface viscos-
ity increased or decreased relative to the wild-type

enzyme. Fitness was defined as a reduction (away from
unity) in the Arrhenius pre-exponential ratio (A
D
: A
T
)
[70,71]. In a more conventional study, we found that
the magnitude and temperature dependence of the pre-
steady-state rate and KIE for proton tunnelling during
the RHR of the quinoprotein methylamine dehydroge-
nase are unchanged following the addition of 30%
glycerol – an increase in solvent viscosity of approxi-
mately two- to threefold [13]. Conversely, a decrease in
KIE and increase in apparent enthalpy for the RHR
of l-phenylalanine oxidase upon the addition of 30%
glycerol has been reported [72]. In a more systematic
study, we recently showed that the rate of coenzyme
capture decreases, whereas the rate and KIE of
hydride transfer during the RHR in MR are invariant
over a 10-fold increase in solution viscosity [20]. We
found it was possible to use a conventional stopped-
flow to make these measurements by varying the
viscosity between $0.9 and 9 cP at 25 °C with the
addition of 0–60% w ⁄ w glycerol. Addition of > 60%
glycerol leads to mixing artefacts that precluded
further measurements. The addition of glycerol to the
solvent will also reduce the solvent dielectric. We inde-
pendently probed the role of solvent dielectric on the
RHR of MR by measuring the temperature depen-
dence in this reaction in the presence of ethanol.

Neither the rate nor enthalpy significantly changed
upon the addition of 20% v ⁄ v ethanol – a change in
dielectric from $80 to $65, but NADH binding was
significantly compromised with an increase in K
S
from
0.2 to 2.7 mm observed. Unfortunately, no clear trends
have emerged as to the effect of viscosity (or dielectric)
S. Hay et al. Hydrogen tunnelling in biological systems
FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS 3937
on enzymatic H-transfer reactions and more systematic
studies are probably required to determine whether
this is a useful probe of H transfer dynamics.
Future perspectives
It is now fairly well established that many enzyme
H-transfer reactions involve a degree of quantum
mechanical H-tunnelling. The role of promoting vibra-
tions, which couple protein dynamics to the H transfer
reaction coordinate, remains contentious. Although
there is now a growing body of compelling experimental
and computational evidence for such vibrations, the
experimental evidence is all by inference. A combined
temperature and pressure study seems to be the best
experimental probe of environmental coupling to
H-transfer chemistry [8]. Computational studies are also
invaluable because these can determine the extent of H-
tunnelling and also visualize promoting vibrations [9].
The challenge for the future remains the direct measure-
ment of such vibrations. If they are found to exist then
a further challenge is to exploit them for practical gain.

Acknowledgement
This work was funded by the UK Biotechnology and
Biological Sciences Research Council. NSS is a
BBSRC Professorial Fellow.
References
1 Garcia-Viloca M, Gao J, Karplus M & Truhlar DG
(2004) How enzymes work: analysis by modern rate
theory and computer simulations. Science 303, 186–
195.
2 Cha Y, Murray CJ & Klinman JP (1989) Hydrogen
tunneling in enzyme-reactions. Science 243, 1325–1330.
3 Benkovic SJ & Hammes-Schiffer S (2003) A perspec-
tive on enzyme catalysis. Science 301, 1196–1202.
4 Masgrau L, Roujeinikova A, Johannissen LO, Hothi
P, Basran J, Ranaghan KE, Mulholland AJ, Sutcliffe
MJ, Scrutton NS & Leys D (2006) Atomic description
of an enzyme reaction dominated by proton tunneling.
Science 312, 237–241.
5 Kohen A, Cannio R, Bartolucci S & Klinman JP (1999)
Enzyme dynamics and hydrogen tunnelling in a thermo-
philic alcohol dehydrogenase. Nature 399, 496–499.
6 Antoniou D & Schwartz SD (1997) Large kinetic iso-
tope effects in enzymatic proton transfer and the role
of substrate oscillations. Proc Natl Acad Sci USA 94,
12360–12365.
7 Caratzoulas S, Mincer JS & Schwartz SD (2002) Iden-
tification of a protein-promoting vibration in the reac-
tion catalyzed by horse liver alcohol dehydrogenase.
J Am Chem Soc 124, 3270–3276.
8 Hay S, Sutcliffe MJ & Scrutton NS (2007) Promoting

motions in enzyme catalysis probed by pressure studies
of kinetic isotope effects. Proc Natl Acad Sci USA 104,
507–512.
9 Johannissen LO, Hay S, Scrutton NS & Sutcliffe MJ
(2007) Proton tunneling in aromatic amine dehydroge-
nase is driven by a short-range sub-picosecond pro-
moting vibration: consistency of simulation and theory
with experiment. J Phys Chem B 111, 2631–2638.
10 Knapp MJ & Klinman JP (2002) Environmentally cou-
pled hydrogen tunneling – linking catalysis to dynam-
ics. Eur J Biochem 269, 3113–3121.
11 Antoniou D & Schwartz SD (2001) Internal enzyme
motions as a source of catalytic activity: rate-promot-
ing vibrations and hydrogen tunneling. J Phys Chem B
105, 5553–5558.
12 Bruno WJ & Bialek W (1992) Vibrationally enhanced
tunneling as a mechanism for enzymatic hydrogen
transfer. Biophys J 63, 689–699.
13 Basran J, Sutcliffe MJ & Scrutton NS (1999) Enzy-
matic H-transfer requires vibration-driven extreme tun-
neling. Biochemistry 38, 3218–3222.
14 Basran J, Patel S, Sutcliffe MJ & Scrutton NS (2001)
Importance of barrier shape in enzyme-catalyzed reac-
tions – vibrationally assisted hydrogen tunneling in
tryptophan tryptophylquinone-dependent amine dehy-
drogenases. J Biol Chem 276, 6234–6242.
15 Basran J, Harris RJ, Sutcliffe MJ & Scrutton NS
(2003) H-tunneling in the multiple H-transfers of the
catalytic cycle of morphinone reductase and in the
reductive half-reaction of the homologous pentaerythr-

itol tetranitrate reductase. J Biol Chem 278, 43973–
43982.
16 Basran J, Sutcliffe MJ & Scrutton NS (2001) Deute-
rium isotope effects during carbon–hydrogen bond
cleavage by trimethylamine dehydrogenase – implica-
tions for mechanism and vibrationally assisted hydro-
gen tunneling in wild-type and mutant enzymes. J Biol
Chem 276, 24581–24587.
17 Harris RJ, Meskys R, Sutcliffe MJ & Scrutton NS
(2000) Kinetic studies of the mechanism of carbon–
hydrogen bond breakage by the heterotetrameric sar-
cosine oxidase of Arthrobacter sp 1-IN. Biochemistry
39, 1189–1198.
18 Pudney CR, Hay S, Sutcliffe MJ & Scrutton NS
(2006) Alpha-secondary isotope effects as probes of
‘tunneling-ready’ configurations in enzymatic H-tunnel-
ing: insight from environmentally coupled tunneling
models. J Am Chem Soc
128, 14053–14058.
19 Pudney CR, Hay S, Pang JY, Costello C, Leys D,
Sutcliffe MJ & Scrutton NS (2007) Mutagenesis of
morphinone reductase induces multiple reactive
Hydrogen tunnelling in biological systems S. Hay et al.
3938 FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS
configurations and identifies potential ambiguity in
kinetic analysis of enzyme tunneling mechanisms.
J Am Chem Soc 129, 13949–13956.
20 Hay S, Pudney CR, Sutcliffe MJ & Scrutton NS
(2008) Are environmentally coupled enzymatic
hydrogen tunneling reactions influenced by changes

in solution viscosity? Angew Chem Int Ed Engl 47,
537–540.
21 Ball P (2004) Enzymes – by chance, or by design?
Nature 431, 396–397.
22 Gladstone S, Laidler KJ & Eyring H (1941) The Theory
of Rate Processes. McGraw-Hill, New York, NY.
23 Marcus RA & Sutin N (1985) Electron transfers in
chemistry and biology. Biochim Biophys Acta 811, 265–
322.
24 Devault D, Parkes JH & Chance B (1967) Electron
tunnelling in cytochromes. Nature 215, 642–644.
25 Pang J, Pu JZ, Gao JL, Truhlar DG & Allemann RK
(2006) Hydride transfer reaction catalyzed by hyper-
thermophilic dihydrofolate reductase is dominated by
quantum mechanical tunneling and is promoted by
both inter- and intramonomeric correlated motions. J
Am Chem Soc 128, 8015–8023.
26 Garcia-Viloca M, Truhlar DG & Gao JL (2003) Reac-
tion-path energetics and kinetics of the hydride transfer
reaction catalyzed by dihydrofolate reductase. Bio-
chemistry 42, 13558–13575.
27 Tejero I, Garcia-Viloca M, Gonzalez-Lafont A, Lluch
JM & York DM (2006) Enzyme dynamics and tunnel-
ing enhanced by compression in the hydrogen abstrac-
tion catalyzed by soybean lipoxygenase-1. J Phys
Chem B 110, 24708–24719.
28 Pang J, Hay S, Scrutton NS & Sutcliffe MJ (2008)
Deep tunneling dominates the biologically important
hydride transfer reaction from NADH to FMN in
morphinone reductase. J Am Chem Soc 130, 7092–

7097.
29 Brinkley DW & Roth JP (2005) Determination of a
large reorganization energy barrier for hydride abstrac-
tion by glucose oxidase. J Am Chem Soc 127, 15720–
15721.
30 Hothi P, Hay S, Roujeinikova A, Sutcliffe MJ, Lee M,
Leys D, Cullis PM & Scrutton NS (2008) Driving force
analysis of proton tunnelling across a reactivity series
for an enzyme–substrate complex. ChemBioChem 9,
2839–2845.
31 Knapp MJ, Rickert K & Klinman JP (2002) Tempera-
ture-dependent isotope effects in soybean lipoxygenase-
1: correlating hydrogen tunneling with protein dynam-
ics. J Am Chem Soc 124, 3865–3874.
32 Kuznetsov AM & Ulstrup J (1999) Proton and hydro-
gen atom tunnelling in hydrolytic and redox enzyme
catalysis. Can J Chem 77, 1085–1096.
33 Bell R (1980) The Tunnel Effect in Chemistry. Chap-
man and Hall, London.
34 Hay S, Pudney C, McGrory TA, Pan J, Sutcliffe MJ &
Scrutton NS (2009) Barrier compression enhances an
enzymatic H-transfer reaction. Angew Chem 48, 1452–
1454.
35 Craig DH, Moody PCE, Bruce NC & Scrutton NS
(1998) Reductive and oxidative half-reactions of
morphinone reductase from Pseudomonas putida M10:
a kinetic and thermodynamic analysis. Biochemistry
37, 7598–7607.
36 Huskey WP & Schowen RL (1983) Reaction-coordinate
tunneling in hydride-transfer reactions. J Am Chem

Soc 105, 5704–5706.
37 Saunders WH (1985) Calculations of isotope effects in
elimination-reactions – new experimental criteria for
tunneling in slow proton transfers. J Am Chem Soc
107, 164–169.
38 Ostovic D, Roberts RMG & Kreevoy MM (1983)
Isotope effects on hydride transfer between NAD
+
analogs. J Am Chem Soc 105, 7629–7631.
39 Hay S, Pudney CR, Hothi P & Scrutton NS (2008)
Correction of pre-steady state KIEs for isotopic impu-
rities and the consequences of kinetic isotope fraction-
ation. J Phys Chem A 112, 13109–13115.
40 Hay S, Pudney CR, Sutcliffe MJ & Scrutton NS
(2008) Solvent as a probe of active site motion
and chemistry during the hydrogen tunnelling reaction
in morphinone reductase. ChemPhysChem 9, 1875–
1881.
41 Messiha HL, Munro AW, Bruce NC, Barsukov I &
Scrutton NS (2005) Reaction of morphinone reductase
with 2-cyclohexen-1-one and 1-nitrocyclohexene. J Biol
Chem 280, 10695–10709.
42 Khan H, Barna T, Bruce NC, Munro AW, Leys D &
Scrutton NS (2005) Proton transfer in the oxidative
half-reaction of pentaerythritol tetranitrate reductase.
FEBS J 272, 4660–4671.
42a Pudney CR, McGrory T, Lafite P, Pang J, Hay S,
Leys D, Sutcliffe MJ & Scrutton NS (2009) Parallel
pathways and free-energy landscapes for enzymatic
hydride transfer probed by hydrostatic pressure.

ChemBioChem 10, 1379–1384.
43 Hay S, Pang JY, Monaghan PJ, Wang X, Evans RM,
Sutcliffe MJ, Allemann RK & Scrutton NS (2008)
Secondary kinetic isotope effects as probes of environ-
mentally-coupled enzymatic hydrogen tunneling reac-
tions. ChemPhysChem 9, 1536–1539.
44 Cook PF, Blanchard JS & Cleland WW (1980)
Primary and secondary deuterium-isotope effects on
equilibrium-constants for enzyme-catalyzed reactions.
Biochemistry 19, 4853–4858.
45 Pu JZ, Ma SH, Garcia-Viloca M, Gao JL, Truhlar
DG & Kohen A (2005) Nonperfect synchronization of
reaction center rehybridization in the transition state
of the hydride transfer catalyzed by dihydrofolate
reductase. J Am Chem Soc 127, 14879–14886.
S. Hay et al. Hydrogen tunnelling in biological systems
FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS 3939
46 Hay S & Scrutton NS (2008) H-transfers in photosys-
tem II: what can we learn from recent lessons in the
enzyme community? Photosynth Res 98 , 169–177.
47 Markham KA & Kohen A (2006) Analytical proce-
dures for the preparation, isolation, analysis and pres-
ervation of reduced nicotinamides. Curr Anal Chem 2,
379–388.
48 Viola RE, Cook PF & Cleland WW (1979) Stereoselec-
tive preparation of deuterated reduced nicotinamide
adenine-nucleotides and substrates by enzymatic-syn-
thesis. Anal Biochem 96, 334–340.
49 McCracken JA, Wang T & Kohen A (2004) Synthesis
of R and S tritiated reduced beta-nicotinamide

adenine dinucleotide 2¢ phosphate. Anal Biochem 324,
131–136.
50 Ottolina G, Riva S, Carrea G, Danieli B & Buckmann
AF (1989) Enzymatic-synthesis of 4R-H-2NAD(P)H
and 4S-H-2NAD(P)H and determination of the stereo-
specificity of 7-alpha-hydroxysteroid and 12-alpha-hy-
droxysteroid dehydrogenase. Biochim Biophys Acta
998, 173–178.
51 Branlant G, Eiler B & Biellmann JF (1982) A word of
caution – 1,4,5,6-tetrahydronicotinamide adenine-dinu-
cleotide (phosphate) should be used with care in acidic
and neutral media. Anal Biochem 125, 264–268.
52 Arnold LJ, You K, Allison WS & Kaplan NO (1976)
Determination of hydride transfer stereospecificity of
nicotinamide adenine-dinucleotide linked oxidoreducta-
ses by proton magnetic-resonance. Biochemistry 15,
4844–4849.
53 Park H, Girdaukas GG & Northrop DB (2006) Effect
of pressure on a heavy-atom isotope effect of yeast alco-
hol dehydrogenase. J Am Chem Soc 128, 1868–1872.
54 Isaacs NS, Javaid K & Rannala E (1978) Reactions at
high-pressure. 5. Effect of pressure on some primary
kinetic isotope effects. J Chem Soc Perkin Trans 2,
709–711.
55 Isaacs NS (1984) The effect of pressure on kinetic
isotope effects. In Isotope Effects in Organic Chemistry
(Buncel E & Lee CC eds), pp. 67–105. Elsevier,
London.
56 Isaacs NS, Javaid K & Rannala E (1977) Pressure
effects on proton tunnelling. Nature 268, 372.

57 Northrop DB (2006) Unusual origins of isotope effects
in enzyme-catalysed reactions. Phil Trans R Soc Lond
B Biol Sci 361, 1341–1349.
58 Northrop DB (1999) Effects of high pressure on iso-
tope effects and hydrogen tunneling. J Am Chem Soc
121, 3521–3524.
59 Northrop DB (2002) Effects of high pressure on enzy-
matic activity. Biochim Biophys Acta 1595, 71–79.
60 Hay S, Sutcliffe MJ & Scrutton NS (2009) Probing
coupled motions in enzymatic hydrogen tunnelling
reactions: beyond temperature-dependence studies of
kinetic isotope effects. In Quantum Tunnelling in
Enzyme Catalyzed Reactions (Allemann RK & Scrut-
ton NS eds), pp. 199–218. Royal Society of Chemistry,
London.
61 Hay S & Scrutton NS (2008) Incorporation of hydro-
static pressure into models of hydrogen tunneling
highlights a role for pressure-modulated promoting
vibrations. Biochemistry
47, 9880–9887.
62 Ewald AH (1968) Effect of pressure on charge trans-
fer complexes in solution. Trans Faraday Soc 64,
733–743.
63 von Smoluchowski M (1917) Mathematical theory of
the kinetics of the coagulation of colloidal solutions.
Z Phys Chem 92, 129–168.
64 Ansari A, Jones CM, Henry ER, Hofrichter J & Eaton
WA (1992) The role of solvent viscosity in the dynam-
ics of protein conformational-changes. Science 256,
1796–1798.

65 Ivkovic-Jensen MM & Kostic NM (1997) Effects of
viscosity and temperature on the kinetics of the elec-
tron-transfer reaction between the triplet state of zinc
cytochrome c and cupriplastocyanin. Biochemistry 36,
8135–8144.
66 Ivkovic-Jensen MM & Kostic NM (1996) Effects of
temperature on the kinetics of the gated electron-
transfer reaction between zinc cytochrome c and
plastocyanin. Analysis of configurational fluctuation
of the diprotein complex. Biochemistry 35, 15095–
15106.
67 Zhou JS & Kostic NM (1993) Gating of photoinduced
electron-transfer from zinc cytochrome-c and tin cyto-
chrome-c to plastocyanin – effects of solution viscosity
on rearrangement of the metalloprotein complex. JAm
Chem Soc 115, 10796–10804.
68 Zhou JS & Kostic NM (1992) Photoinduced electron-
transfer from zinc cytochrome-c to plastocyanin is
gated by surface-diffusion within the metalloprotein
complex. J Am Chem Soc 114, 3562–3563.
69 Feng CJ, Kedia RV, Hazzard JT, Hurley JK, Tollin G
& Enemark JH (2002) Effect of solution viscosity on
intramolecular electron transfer in sulfite oxidase. Bio-
chemistry 41, 5816–5821.
70 Kohen A, Jonsson T & Klinman JP (1997) Effects of
protein glycosylation on catalysis: changes in hydrogen
tunneling and enthalpy of activation in the glucose oxi-
dase reaction. Biochemistry 36, 2603–2611.
71 Seymour SL & Klinman JP (2002) Comparison of
rates and kinetic isotope effects using PEG-modified

variants and glycoforms of glucose oxidase: the
relationship of modification of the protein envelope
to C–H activation and tunneling. Biochemistry 41,
8747–8758.
72 Ohta Y, Mukouyama EB & Suzuki H (2006)
Kinetic isotope effect of the l-phenylalanine oxidase
from Pseudomonas sp P-501. J Biochem 139,
551–555.
Hydrogen tunnelling in biological systems S. Hay et al.
3940 FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS
73 Nagel ZD & Klinman JP (2006) Tunneling and
dynamics in enzymatic hydride transfer. Chem Rev
106, 3095–3118.
74 Fan F & Gadda G (2005) Oxygen- and temperature-
dependent kinetic isotope effects in choline oxidase:
correlating reversible hydride transfer with environ-
mentally enhanced tunneling. J Am Chem Soc 127,
17954–17961.
75 Dunn RV, Marshall KR, Munro AW & Scrutton NS
(2008) The pH dependence of kinetic isotope effects in
monoamine oxidase A indicates stabilization of the
neutral amine in the enzyme–substrate complex. FEBS
J 275, 3850–3858.
76 Jonsson T, Edmondson DE & Klinman JP (1994)
Hydrogen tunneling in the flavoenzyme monoamine-
oxidase-B. Biochemistry 33, 14871–14878.
77 Agrawal N, Lesley SA, Kuhn P & Kohen A (2004)
Mechanistic studies of a flavin-dependent thymidylate
synthase. Biochemistry 43, 10295–10301.
78 Fagan RL, Nelson MN, Pagano PM & Palfey BA

(2006) Mechanism of flavin reduction in class 2 dihydro-
orotate dehydrogenases. Biochemistry 45, 14926–14932.
S. Hay et al. Hydrogen tunnelling in biological systems
FEBS Journal 276 (2009) 3930–3941 ª 2009 The Authors Journal compilation ª 2009 FEBS 3941