Báo cáo khoa học: Mechanistic aspects and redox properties of hyperthermophilic L-proline dehydrogenase from Pyrococcus furiosus related to dimethylglycine dehydrogenase⁄oxidase potx

Mechanistic aspects and redox properties of
hyperthermophilic
L-proline dehydrogenase from
Pyrococcus furiosus related to dimethylglycine
dehydrogenase⁄ oxidase
Phillip J. Monaghan, David Leys and Nigel S. Scrutton
Manchester Interdisciplinary Biocentre and Faculty of Life Sciences, University of Manchester, UK
The membrane-associated flavoprotein PutA in enteric
bacteria is a proline catabolic enzyme that catalyzes
the oxidation of proline to glutamate in a two-step
reaction to form glutamate (Fig. 1). The protein is also
a transcriptional repressor of the proline utilization
(put) genes [1–3]. Cytoplasmic PutA represses tran-
scription from its own gene and also from the
Na
+
⁄ proline transporter PutP [4–6]. Proline catabolism
enables enteric bacteria to use l-proline as a source of
carbon, nitrogen and electrons, and the reaction is
initiated in the FAD-binding domain by two-electron
oxidation of l-proline to form D
1
-pyrroline-5-carboxy-
late (P5C) [5,6]. Following oxidation of l-proline, the
two-electron reduced FAD cofactor passes electrons to
an acceptor in the electron transfer chain. The interme-
diate P5C is hydrolyzed to glutamate 5-semialdehyde,
which is then oxidized to glutamate by the P5C
dehydrogenase domain, with NAD
+
acting as electron

Keywords
amine oxidation; flavoprotein;
hyperthermophile; mechanism; proline
dehydrogenase
Correspondence
N. S. Scrutton, Manchester Interdisciplinary
Biocentre and Faculty of Life Sciences,
University of Manchester, 131 Princess
Street, Manchester M1 7DN, UK
Fax: +44 161 275 5586
Tel: +44 161 275 5632
E-mail:
(Received 4 January 2007, revised 9
February 2007, accepted 19 February 2007)
doi:10.1111/j.1742-4658.2007.05750.x
Two ORFs encoding a protein related to bacterial dimethylglycine oxidase
were cloned from Pyrococcus furiosus DSM 3638. The protein was
expressed in Escherichia coli, purified, and shown to be a flavoprotein
amine dehydrogenase. The enzyme oxidizes the secondary amines l-proline,
l-pipecolic acid and sarcosine, with optimal catalytic activity towards
l-proline. The holoenzyme contains one FAD, FMN and ATP per ab
complex, is not reduced by sulfite, and reoxidizes slowly following reduc-
tion, which is typical of flavoprotein dehydrogenases. Isolation of the
enzyme in a form containing only FAD cofactor allowed detailed pH
dependence studies of the reaction with l-proline, for which a bell-shaped
dependence (pK
a
values 7.0 ± 0.2 and 7.6 ± 0.2) for k
cat
⁄ K

m
as a function
of pH was observed. The pH dependence of k
cat
is sigmoidal, described by
a single macroscopic pK
a
of 7.7 ± 0.1, tentatively attributed to ionization
of l-proline in the Michaelis complex. The preliminary crystal structure of
the enzyme revealed active site residues conserved in related amine dehy-
drogenases and potentially implicated in catalysis. Studies with H225A,
H225Q and Y251F mutants ruled out participation of these residues in a
carbanion-type mechanism. The midpoint potential of enzyme-bound FAD
has a linear temperature dependence () 3.1 ± 0.05 mVÆC°
)1
), and extra-
polation to physiologic growth temperature for P. furiosus (100 °C) yields
a value of ) 407 ± 5 mV for the two-electron reduction of enzyme-bound
FAD. These studies provide the first detailed account of the kinetic ⁄ redox
properties of this hyperthermophilic l -proline dehydrogenase. Implications
for its mechanism of action are discussed.
Abbreviations
DMGO, dimethylglycine oxidase; P5C, D
1
-pyrroline-5-carboxylate; PRODH, L-proline dehydrogenase; TAPSO, 3-{[tris(hydroxymethyl)-
methyl]amino}-2-hydroxypropane sulfonic acid; TMADH, trimethylamine dehydrogenase.
2070 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
acceptor (Fig. 1). The structure of a truncated form of
PutA comprising the FAD-containing proline dehy-
drogenase domain has been elucidated, and reveals a

domain-swapped dimer, with each subunit containing
three domains [7]. These domains comprise a helical
dimerization arm, a three-helix bundle, and a b ⁄ a
barrel l-proline dehydrogenase (PRODH) domain.
A model of the enzyme–substrate complex has been
constructed from which a mechanism for the oxidation
of l-proline has been proposed [7]. In eukaryotes, dis-
tinct enzymes encoded by separate genes catalyze the
oxidation of l-proline to glutamate. A mitochondrial
l-proline oxidase is the human homolog of the PutA
protein of enteric bacteria, and it has roles in p53-
mediated apoptosis, the production of reactive oxygen
species, and also schizophrenia [8,9].
A new type of dye-linked PRODH was recently
identified in crude extracts of the hyperthermophile
Thermococcus profundus [10]. The enzyme is a hetero-
tetramer, contains 2 moles of FAD per mole of
enzyme, and is bifunctional, having proline dehydro-
genase and dye-linked NADH dehydrogenase activities
[10,11]. The enzyme is a complex iron–sulfur flavopro-
tein, with the a subunit containing a 2Fe)2S center.
Additionally, the c subunit contains a broad absorp-
tion peak around 420 nm typical of an 8Fe)8S ferre-
doxin. The a and b subunits show sequence similarity
with other putative amine-oxidizing proteins, including
the putative sarcosine oxidase a and b subunits of
P. furiosus. Recent studies have also identified a dye-
linked d-proline dehydrogenase from Pyrobaculum
islandicum [12]. In searching for other new classes of
PRODH, we have identified two ORFs [annotated

gi_18977617 (a subunit) and gi_18977618 (b subunit),
in the protein extraction, description and analysis tool
(pedant) database] in the genome of P. furiosus
DSM 3638 that show sequence homology to the a and
b subunits of the flavoprotein amine oxidoreductase,
tetrameric sarcosine oxidase [13]. The translated amino
acid sequence of gi_18977618 (b subunit) also aligns
with another member of the amine oxidoreductase
family, dimethylglycine oxidase (DMGO) from
Arthrobacter globiformis [13], for which the crystal
structure has been determined to 1.6 A
˚
resolution [14].
This alignment indicates the conservation of three resi-
dues (His225, Tyr251 and Gly262 in the gi_18977618
translation), known to reside in the active site of
DMGO, that have been implicated in the catalytic
mechanism of dimethylglycine oxidation [14]. The crys-
tal structure of a related enzyme from Pyrococcus hori-
koshii OT-3 has been elucidated [15], and its basic
solution properties have been analyzed [16], but, to
date, detailed mechanistic studies of the activity of this
or related enzymes have not been reported. With this
aim in mind, we present an analysis of recombinant
protein expressed from the two ORFs found in
P. furiosus DSM 3638 that share sequence similarity
with the gene encoding DMGO. We show that these
genes encode a new member of the hyperthermo-
philic class of PRODHs that couples the oxidation of
l-proline to the reduction of a noncovalently bound

FAD and the production of P5C. Oxidation of
enzyme-bound FADH
2
is accomplished using the
artificial electron acceptor ferricenium hexafluorophos-
phate, but not molecular oxygen, NAD
+
, or pyrococ-
cal 4Fe)4S ferredoxin. This article represents the first
detailed report of the kinetic and spectroscopic proper-
ties of this novel type of PRODH.
Results
Analysis of a and b subunit sequences
Sequence analysis of both subunits suggests the pres-
ence of an ADP-binding motif in the N-terminal
region of the a subunit, with the 11 participating
Fig. 1. Reactions catalyzed by the bifunc-
tional PutA protein of enteric bacteria.
Hydrolysis of P5C is nonenzymatic. In euk-
aryotes, distinct enzymes encoded by separ-
ate genes catalyze these reactions.
P. J. Monaghan et al.
L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2071
residues satisfying the physicochemical parameters of
the consensus [17]. The a subunit also contains a con-
served GG doublet five nucleotides downstream of the
dinucleotide-binding domain. An ATG motif is also
evident in the a subunit. This motif has been found in
both FAD-binding and NADPH-binding proteins,

where it forms the fourth b-strand of the Rossman
fold and the connecting loop. In flavoproteins, the
ATG motif has a defined function, in that it is always
present at the junction with the substrate-binding
domain, and not within a domain, as in NADPH-
binding proteins [18]. The b subunit also contains an
ADP-binding motif, and shares 27% sequence identity
with the b subunit of tetrameric sarcosine oxidase from
Corynebacterium sp. P-1, and 26% sequence identity
with the N-terminal half of human dimethylglycine
dehydrogenase. Of particular note is the finding that
the b subunit of PRODH shows sequence conservation
with active site residues His225, Tyr259 and Gly270 of
DMGO from A. globiformis and mouse lung dimethyl-
glycine dehydrogenase (Fig. 2), residues that are pre-
sent in a number of sarcosine dehydrogenase-like
proteins. Given these sequence similarities, we conjec-
tured that the protein encoded by the two identified
A
B
Fig. 2. Multiple sequence alignments of amino acid sequences of the a and b subunits of PRODH. (A) Multiple sequence alignment for the
a subunit, showing 18% sequence identity with the N-terminal region of the a subunit of tetrameric sarcosine oxidase from both Corynebac-
terium sp. P-1 [20] and Arthrobacter sp. 1-IN [13]. The 11 residues that comprise the ADP-binding motif are highlighted in bold, and shaded
where residues are conserved. All 11 residues in PRODHa obey the physicochemical requirements established by Wierenga et al. [17]. The
conserved GG doublet and ATG motif are also shaded. (B) Multiple sequence alignment deduced for the b subunit of PRODH, showing 24%
sequence identity with DMGO from A. globiformis [13] and 26% sequence identity with the cDNA translation product of dimethylglycine
dehydrogenase from M. musculus lung tissue [21]. The N-terminal ADP-binding motif is highlighted in bold, and shaded where residues are
conserved. Again, all 11 residues satisfy the consensus sequence, with the exception of the glutamate residue at position 1, although this
hydrophilic residue has the correct physicochemical requirements for this position. DMGO active site residues His225, Tyr259 and Gly270,
identified from the crystal structure, align with conserved residues in both dimethylglycine dehydrogenase and PRODHb, and are highlighted

with bold type and shading. Additional conserved residues are marked with an asterisk.
L-proline dehydrogenase from P. furiosus P. J. Monaghan et al.
2072 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
ORFs in Pyrococcus furiosus encode a new type of
amine dehydrogenase ⁄ oxidase, a hypothesis that we
addressed through detailed characterization of the pro-
tein as a new type of (PRODH), described below.
Purification of recombinant enzyme and general
properties
Recombinant wild-type enzyme was expressed at high
levels in Escherichia coli strain Rosetta(DE3)pLysS
transformed with either plasmid pPRODH1 or plasmid
pPRODH2. The enzyme was purified to homogeneity
in three steps and in the oxidized form (Fig. 3A). The
holoenzyme form of PRODH was selected for crystall-
ogenesis and X-ray diffraction studies, and was puri-
fied as previously described [19]. PRODH for
mechanistic studies was further exchanged into
100 mm potassium phosphate buffer (pH 7.5). This
treatment releases the ATP and FMN cofactors from
the protein, but leaves the FAD-bound form. This is a
convenient form of the enzyme for simplified analysis
of FAD reduction (see below). The protein yield was
typically $ 10 mg of purified enzyme per liter of
recombinant culture. The purified enzyme was found
to be yellow in color, and had a typical flavoprotein
absorbance spectrum characterized by two flavin peaks
with absorption maxima at 367 and 450 nm (Fig. 3B).
N-terminal sequence analysis of the a and b subunits
purified from pPRODH1 indicated that a subpopula-

tion (approximately $ 13%) of the a subunit was trun-
cated. The sequence MKVQRQ was obtained for the
truncated a subunit N-terminus, indicating that trunca-
tion in the a subunit is located 83 amino acids from
the initiating methionine in the full-length a subunit.
Enzyme expressed from plasmid pPRODH2 lacked a
truncated a subunit, consistent with removal of the
internal ribosome-binding site by mutagenesis. The a
and b subunits were coexpressed in a molar ratio of
$ 1 : 1 from pPRODH2, as judged by SDS⁄ PAGE
peak area image scanning. Analysis of purified enzyme
by MALDI-TOF MS gave a molecular mass of
42 437.5 Da for the b subunit, comparable to the pre-
dicted molecular mass of 42 481.2 Da from the gene
sequence. Electrospray mass data for the b subunit
gave a molecular mass of 42 474.0 Da. Mass data for
the larger a subunit could not be obtained using the
MALDI-TOF or electrospray methods. Purification of
H225A, H225Q and Y251F mutant enzymes was as
described for wild-type PRODH. Mutagenesis of these
active site residues had no effect on recombinant pro-
tein yield, and all mutants were purified in FAD-
bound form. Despite the close proximity of both
His225 and Tyr251 to the isoalloxazine ring moiety of
FAD, no major perturbations in the absorption prop-
erties of the enzyme-bound flavin were evident as a
consequence of mutagenesis.
Holoenzyme cofactor content
Our preliminary crystallographic analysis of the
enzyme indicates a heterooctomer (ab)

4
structure for
PRODH, as initially suggested from the computed
self-rotation of diffraction data [19]. It is evident from
A
B
Fig. 3. (A) SDS ⁄ PAGE analysis of the purification of recombinant
wild-type PRODH from E. coli strain Rosetta(DE3)pLysS trans-
formed with pPRODH2. Lane 1: molecular mass marker (97, 66,
45, 30 and 20.1 kDa from top to bottom of the gel). Lane 2: cell
lysate. Lane 3: sample after heat denaturation at 80 °C and clarifica-
tion by centrifugation. Lane 4: pooled fractions following anion
exchange chromatography (Q-Sepharose). Lane 5: pooled fractions
following size-exclusion chromatography (Superdex 75) showing
the pure a and b subunits of PRODH. (B) UV-visible absorption
spectrum and reductive titration of recombinant wild-type PRODH
with sodium dithionite. The absorption spectrum recorded between
300 and 600 nm is typical of a flavoprotein spectrum. Flavin peaks
are at 367 and 450 nm, and the arrow indicates the direction of
absorption change. A single isosbestic point was observed at
340 nm. Inset: a plot of absorbance at 450 nm versus electron
equivalents, which demonstrates that reduction of FAD-bound
PRODH is complete following addition of two electrons. Condi-
tions: 100 m
M potassium phosphate buffer, pH 7.5; 25 °C; enzyme
concentration 16 l
M.
P. J. Monaghan et al.
L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2073

the preliminary X-ray crystal structure of PRODH
that one molecule of ATP cofactor is bound in the
ADP-binding motif of the a subunit (Fig. 4A). This
cofactor has no obvious function from a mechanistic
perspective, but may play a stabilizing role under the
harsh physiologic conditions that P. furiosus is subject
to. The ADP-binding motif in the b subunit binds one
molecule of noncovalent FAD (Fig. 4B). FMN is
located at the interface of the a and b subunits
(Fig. 4A).
Reductive titration of PRODH highlighted a single
isosbestic point at 340 nm, with no evidence for a
semiquinone species obtained during titration with
sodium dithionite (Fig. 3B). FAD-bound PRODH for
mechanistic studies was confirmed by MALDI-TOF
MS after heat treatment to remove cofactor. A
mass ⁄ charge peak at 787 corresponded to the posi-
tively charged quasimolecular ion ([M +H]
+
)of
FAD. Partial hydrolysis of FAD during heat treatment
was revealed by mass ⁄ charge peaks at 348 and 458,
assigned to AMP and FMN ([M +H]
+
) hydrolysis
products, respectively. ATP cofactor was not detected
(Fig. 5), although the preliminary crystal structure of
PRODH indicates its presence in the enzyme.
Reduction with amine substrates
Alignment of the a and b subunit sequences with well-

characterized enzymes suggests that the purified
enzyme is an amine-specific oxidoreductase (Fig. 2). In
particular, the a subunit shows 18% identity with the
N-terminal region of the a subunit of tetrameric sarco-
sine oxidase from Corynebacterium sp. P-1 [20] and
Arthrobacter sp. 1-IN [13]. The b subunit shows 24%
identity with DMGO from A. globiformis [13] and
26% sequence identity with dimethylglycine dehydro-
genase from Mus musculus [21]. Given these sequence
similarities, amine compounds were analyzed as
potential substrates by mixing with enzyme (19.4 lm)
at 80 °C under anaerobic conditions to preclude
potential oxidase chemistry. Enzyme–substrate reactiv-
ity was established by following bleaching of the flavin
spectrum on addition of the amine compound
(20 mm). The enzyme was found to oxidize only
secondary amine compounds, namely sarcosine,
l-proline, and l-pipecolic acid (Fig. 6); l-proline was
most effective as reducing substrate (t
1 ⁄ 2
¼$105 s),
followed by l-pipecolic acid (t
1 ⁄ 2
¼$110.5 s; a
structural analog of l-proline), and sarcosine
(t
1 ⁄ 2
¼$654 s). Glycine betaine, glycine, dimethylgly-
cine and d-proline did not lead to significant flavin
reduction. The common structural link between these

identified substrates for PRODH is that they are
all secondary a-amino acids (Fig. 6). The ability of
P. furiosus PRODH to oxidize multiple amine com-
pounds is in stark contrast to the catalytic properties
reported for dye-linked proline dehydrogenase 1 of
Pyrococcus horikoshii OT-3, which has been shown to
act exclusively on l-proline, with l-pipecolic acid and
sarcosine being inert as substrates [16]. A spectral fea-
ture is apparent at $ 550 nm upon kinetic reduction
of PRODH with each of the three identified amine
substrates. This signal may represent a minor transient
population of a charge-transfer species during the cat-
alytic reaction. Addition of sodium sulfite (50 mm)to
purified enzyme did not perturb the flavin absorption
spectrum, indicating that a flavin–N5–sulfite adduct
does not form. This suggests that the enzyme is not a
flavoprotein oxidase, as reactivity with sulfite is a
characteristic of this class of flavoenzyme [22].
Steady-state turnover analysis with
L-proline
and
L-pipecolic acid
In developing a suitable and continuous turnover
assay for wild-type enzyme at an elevated temperature
H225β
Y251β
FAD
FAD
FMN
ATP

BA
Fig. 4. Preliminary P. furiosus PRODH crys-
tal structure. (A) Omit maps (in blue) super-
imposed on the bound FMN (green sticks),
FAD (yellow sticks) and ATP (magenta
sticks) cofactors of the heterotetrameric
PRODH (represented by gray ribbons). The
electron density map is contoured at 3r.
(B) Position of the active site residues
His225b and Tyr251b with respect to the
FAD isoalloxazine group.
L-proline dehydrogenase from P. furiosus P. J. Monaghan et al.
2074 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
(80 °C), a number of issues were taken into account.
A continuous assay was chosen, because coupled assays
are usually inappropriate, owing to a lack of suitable
accessory enzymes that are stable at elevated tempera-
ture. Assays were performed in 100 mm potassium
phosphate buffer (pH 7.5), which has a low tempera-
ture coefficient [d(pK
a
) ⁄ dT ¼ ) 0.0028] [23,24], and
ferricenium hexafluorophosphate (200 lm) was used as
electron acceptor. Steady-state assays were performed
at 80 °C with both l-proline and l-pipecolic acid as
substrate. A comparison of steady-state turnover under
aerobic and anaerobic conditions indicated that oxygen
did not affect turnover reaction rates, consistent
with the enzyme not being of the oxidase class.
Consequently, steady-state kinetic parameters were

determined under aerobic conditions. Analysis of hyper-
bolic plots of initial velocity as a function of substrate
concentration yielded apparent K
m
values for the wild-
type enzyme of 30.8 ± 1.1 mm and 212.3 ± 17.0 mm
for l-proline and l-pipecolic acid, respectively. The cor-
responding apparent k
cat
values were 18.1 ± 0.2 s
)1
and
0.4 ± 0.02 s
)1
for l-proline and l-pipecolic acid,
respectively, and the calculated specificity constants
(k
cat
⁄ K
m
) were 0.59 ± 0.03 s
)1
Æmm
)1
(l-proline) and
0.002 ± 0.0002 s
)1
mm
)1
(l-pipecolic acid). We infer

that l-proline is the preferred substrate, and that the
enzyme therefore represents a new member of the class
of PRODHs that is distinct from E. coli PRODH.
Unlike what was observed for the E. coli enzyme, we
were unable to show any NAD
+
reduction activity for
P. furiosus PRODH in either multiple-turnover or sin-
gle-turnover assays, reinforcing functional differences
between the P. furiosus and E. coli enzymes. Exogenous
FMN did not act as electron acceptor in steady-state
A
B
C
D
Fig. 5. MALDI-TOF MS of flavin cofactor released from PRODH (FAD-bound form). (A) Authentic FAD showing positively charged quasi-
molecular ion of FAD ([M +H]
+
) corresponding to the m ⁄ z peak of 787 and the FAD–Na
+
adduct with an m ⁄ z peak at 809. (B) Authentic
FMN showing both [M +H]
+
ion and FMN–Na
+
adduct m ⁄ z peaks at 458 and 480, respectively. (C) Authentic AMP showing both the
[M +H]
+
ion and AMP–Na
+

adduct m ⁄ z peaks at 348 and 370, respectively. (D) Released cofactor of heat-denatured PRODH showing
the [M +H]
+
ion of FAD cofactor with an m ⁄ z peak of 787 identical to that of the authentic FAD standard. The m ⁄ z peak at 825 repre-
sents the K
+
adduct of released FAD cofactor. The m ⁄ z peaks of 348 and 458 represent the [M +H]
+
ion of both AMP and FMN,
respectively, which result from partial hydrolysis of FAD cofactor during protein heat treatment. The m ⁄ z peak of 496 represents the K
+
adduct of the FAD cofactor heat hydrolysis product, FMN. K
+
ions are present from the purification buffers. Conditions: samples of
authentic FAD, FMN and AMP were prepared as 1 mgÆmL
)1
stock solutions in double deionized H
2
O, and filtered using a 0.22 lm Acro-
disc; PRODH was exchanged into double deionized H
2
O.
P. J. Monaghan et al.
L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2075
anaerobic assays. Additionally, we were unable to
show electron transfer from l-proline-reduced PRODH
to P. furiosus ferredoxin under anaerobic turnover
conditions, either in the presence or in the absence of
exogenous FMN.

Steady-state assays were performed over the tem-
perature range 40–90 °C. Plots of initial velocity as
a function of l-proline concentration were hyperbolic
at all temperatures, and kinetic parameters were calcu-
lated by fitting to the Michaelis–Menten equation.
Both k
cat
and K
m
increase with temperature (Table 1).
The temperature dependence of PRODH-catalyzed
l-proline oxidation was investigated at a saturating
l-proline concentration (200 mm) (Fig. 7A). Thermo-
dynamic parameters were obtained by fitting to the
Eyring equation (Eqn 1).
Inðk=TÞ¼In k
B
=h þ DS
z
=R À DH
z
=RT ð1Þ
where k
B
and h are the Boltzmann and Planck constants,
respectively. Initial velocity was strongly dependent
on temperature (Fig. 7B), and analysis of the data
using the Eyring plot gave thermodynamic parameters
DH
à

¼ 83.4 ± 2.9 kJÆmol
)1
, DS
à
¼ 27.2 ± 1.0 JÆmol
)1
Æ
K
)1
,andDG
à
¼ 73.3 kJÆmol
)1
(at 373 K). Incubation of
PRODH at elevated temperatures prior to activity assay
showed that the enzyme is extremely stable, with no loss
of activity being evident up to 100 °C. Above this tem-
perature, thermal denaturation of PRODH is apparent,
with complete loss of activity after 10 min of incubation
in glycerol buffer at temperatures ‡ 115 °C (data not
shown). Thus, PRODH from P. furiosus is the most ther-
mostable PRODH described to date.
A
B
C
D
E
Fig. 6. Absorption changes as a function of
reaction time accompanying reduction of
wild-type PRODH with sarcosine,

L-pipecolic
acid, and
L-proline. (A) Absorption changes
following reduction of PRODH (19.4 l
M)
with sarcosine (20 m
M). (B) As for (A), but
with
L-pipecolic acid (20 mM). (C) As for (A),
but with
L-proline (20 mM). (D) Plot of
absorbance change at 450 nm as a function
of time for each of the spectral changes
shown in (A)–(C). Symbols: d, sarcosine; m,
L-pipecolic; ., L-proline. Conditions: 100 mM
potassium phosphate buffer, pH 7.5; 80 °C.
(E) Structural formulae for the three secon-
dary amine molecules, sarcosine,
L-pipecolic
acid, and
L-proline, which show substrate
reactivity with PRODH. Boxed areas illus-
trate the common moiety suggested to be
important for binding in the PRODH active
site.
Table 1. Steady-state kinetic parameters for the reaction of
PRODH with
L-proline determined at different temperatures. Condi-
tions: 100 m
M potassium phosphate buffer, pH 7.5, at each assay

temperature.
Temperature (°C) K
m
(mM) k
cat
(s
)1
) k
cat
⁄ K
m
(s
)1
ÆmM
)1
)
40 5.6 ± 0.4 0.4 ± 0.005 0.08 ± 0.006
45 5.6 ± 0.2 0.8 ± 0.005 0.1 ± 0.005
50 5.3 ± 0.4 1.3 ± 0.02 0.2 ± 0.02
55 8.0 ± 0.4 2.5 ± 0.02 0.3 ± 0.02
60 9.9 ± 0.4 4.0 ± 0.04 0.4 ± 0.02
65 11.6 ± 1.0 5.9 ± 0.1 0.5 ± 0.05
70 19.9 ± 2.3 10.4 ± 0.4 0.5 ± 0.08
75 25.9 ± 1.3 14.8 ± 0.2 0.6 ± 0.04
80 30.8 ± 1.1 18.01 ± 0.2 0.6 ± 0.03
L-proline dehydrogenase from P. furiosus P. J. Monaghan et al.
2076 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
For mechanistic analyses, steady-state turnover
assays were performed aerobically with l-proline at
60 °C over the pH range 5.5–10.0, to identify

kinetically influential ionizations. Ionic strength
across the pH range was kept constant using a
three-component buffer system (see Experimental
procedures). Kinetic parameters were calculated by
fitting to the Michaelis–Menten equation (Table 2).
For wild-type PRODH, the pH dependence of
k
cat
⁄ K
m
was found to be bell-shaped, and fitting of
the data using Eqn (4) (see Experimental proce-
dures) yielded macroscopic pK
a
values of 7.0 ± 0.2
(acid limb) and 7.6 ± 0.2 (alkali limb) (Fig. 8A).
Assuming no change in rate-limiting step across the
pH range, these macroscopic pK
a
values most likely
represent ionization of residues in the free enzyme.
By analogy with other amine oxidases ⁄ dehydrogenases
that share similarity at the sequence level with
PRODH (Fig. 2), we speculated that the pK
a
of
7.0 ± 0.2 may be attributed to ionization of the
conserved His225, a potential active site base residue.
However, this proposal was later refuted in light of
kinetic analyses performed with both H225A and

H225Q mutant forms (see below). The pH depend-
ence of k
cat
exhibited a simple sigmoid behavior that,
when analyzed by fitting to Eqn (3) (see Experimen-
tal procedures) (Fig. 8B), produced a macroscopic
pK
a
value of 7.7 ± 0.1. The pK
a
value for the
protonation of free proline is 10.6, but this might be
lowered on binding to enzyme in the Michaelis
complex by ) 2.9 pH units. A precedent for stabiliza-
tion of the free base form of amine substrates at
physiologic pH values is available from studies with
trimethylamine dehydrogenase (TMADH) [25], and is
consistent with mechanistic proposals that require
the unprotonated amine substrate species to react
with the enzyme-bound flavin [26].
B
A
Fig. 7. Temperature dependence and Eyring analysis of initial velo-
city data for wild-type PRODH reacting with
L-proline. (A) Three-
dimensional plot showing initial velocity (y-axis) versus time (x-axis)
versus temperature (z-axis). Reactions were performed in the pres-
ence of saturating
L-proline (200 mM) over the temperature range
40–90 °C. The dimension of time demonstrates any potential loss

of activity due to enzyme thermal denaturation at elevated temper-
atures (not observed in the case of PRODH-catalyzed oxidation of
L-proline). The three-dimensional plot was generated using SIGMA-
PLOT
v9.0 for Windows. Curve-fitting used the Loess transformation
to smooth data based on local regression, which applies a tricube
weight function to elicit trends from noisy data [45]. The trend elici-
ted from the smoothing process was then used to extrapolate data
back to time-point zero to compensate for the time lag between
addition of enzyme to initiate the reaction and the start of data col-
lection. Conditions: 100 m
M potassium phosphate buffer, pH 7.5
(pH corrected at each assay temperature). (B) Eyring plot of initial
velocity data for PRODH with
L-proline as substrate. Thermody-
namic parameters derived from fitting of data to the Eyring equa-
tion are DH
à
¼83.4 ± 2.9 kJÆmol
)1
, DS
à
¼ 27.2 ± 1.0 JÆmol
)1
ÆK
)1
and DG
z
373
¼73.3 kJÆmol

)1
at 100 °C.
Table 2. Steady-state kinetic parameters determined for the reac-
tion of PRODH with
L-proline at different pH values and at constant
ionic strength. Assays were performed at 60 °C.
pH k
cat
(s
)1
) K
m
(mM) k
cat
⁄ K
m
(s
)1
ÆmM
)1
)
5.5 1.10 ± 0.06 9.96 ± 2.20 0.11 ± 0.03
6.0 1.67 ± 0.03 5.76 ± 0.51 0.29 ± 0.03
6.5 2.40 ± 0.02 1.08 ± 0.08 2.22 ± 0.18
7.0 4.08 ± 0.04 1.27 ± 0.11 3.21 ± 0.31
7.5 6.83 ± 0.14 1.81 ± 0.32 3.76 ± 0.74
8.0 11.68 ± 0.25 5.82 ± 0.65 2.01 ± 0.27
8.5 16.36 ± 0.46 19.54 ± 1.73 0.84 ± 0.10
9.0 16.36 ± 0.25 42.35 ± 1.56 0.39 ± 0.02
9.5 16.54 ± 0.41 87.21 ± 4.19 0.19 ± 0.01

10.0 18.00 ± 1.14 137.97 ± 15.11 0.13 ± 0.02
P. J. Monaghan et al.
L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2077
Properties of mutant enzymes altered in the
active site
From analysis of the preliminary crystal structure of
P. furiosus PRODH (Fig. 4A; a more complete struc-
tural analysis is to be published elsewhere), residues
His225 and Tyr251 are situated on the re face of the
isoalloxazine ring of FAD, forming part of the sub-
strate-binding site (Fig. 4B). To assess the potential
role of these two residues as active site bases, pH
dependence studies were performed with H225A,
H225Q and Y251F mutant enzymes, as described for
wild-type PRODH. Initial steady-state experiments
using the three-component buffer system at pH 7.5
showed major perturbations in the apparent kinetic
parameters calculated for each mutant in comparison
to wild-type PRODH (Table 3). Unlike those for the
wild-type enzyme, initial velocities recorded for the
Y251F mutant enzyme were subject to inhibition at
high l-proline concentrations in the acid-to-neutral
solution pH region (supplementary Fig. S1). The
apparent kinetic parameters of the Y251F mutant
enzyme for l-proline were derived by fitting data to a
steady-state rate expression that incorporates substrate
inhibition (Eqn 2).
v ¼
V

max
1 þ
K
m
½S
þ
½S
K
i
ð2Þ
where v is the initial velocity, V
max
is the maximum
value of the initial velocity, [S] is the substrate concen-
tration, K
m
is the substrate concentration at half the
maximal velocity, and K
i
is the equilibrium constant
for inhibitor binding. Marked inhibition has also been
reported in studies of mutant forms of the flavoprotein
morphinone reductase under conditions of high sub-
strate concentration [27,28]. pH dependence studies
revealed that the H225A mutant enzyme was unstable
and precipitated from solution below pH 7.0. This
consequently compromised the accuracy of data analy-
sis in the acid solution pH region. The H225Q mutant
was somewhat more stable, displaying activity down to
solution pH 6.0. The pH dependence of k

cat
was sig-
moidal for both the H225A and H225Q mutant forms,
and when fitted to Eqn (3) (see Experimental proce-
dures) produced a macroscopic pK
a
value of 7.1 ± 0.1
for each mutant. The Y251F mutant enzyme was sta-
ble over the entire pH range of study, with k
cat
values
again showing a simple sigmoidal dependence on solu-
tion pH; fitting data to Eqn (3) (see Experimental pro-
cedures) gave a macroscopic pK
a
value of 7.3 ± 0.1,
which compares favorably with the values determined
for the wild-type and His225 mutant. In light of the
high degree of similarity between pK
a
values deter-
mined from fitting of the k
cat
data plots for wild-type
and mutant enzymes, the macroscopic pK
a
of 7.7 for
A
B
Fig. 8. Dependence of steady-state kinetic parameters on solution

pH for the PRODH-catalyzed oxidation of
L-proline. (A) pH depen-
dence of k
cat
⁄ K
m
following ionizations in the free enzyme and sub-
strate. Fitting of data to Eqn (4) gave two pK
a
values of 7.0 ± 0.2
and 7.6 ± 0.2. (B) pH dependence of k
cat
following the pK
a
of the
enzyme–substrate complex. Fitting of data to Eqn (3) showed a
simple sigmoid relationship, giving a pK
a
of 7.7 ± 0.1. This value is
tentatively assigned to deprotonation of the substrate
L-proline.
Conditions: three-component buffer system comprising 0.052,
0.052 and 0.1
M Mes, TAPSO, and diethanolamine, respectively;
60 °C.
Table 3. Steady-state kinetic parameters determined for the H225A,
H225Q and Y251F mutant PRODH forms. Conditions: buffer
composed of Mes, TAPSO and diethanolamine at final concentra-
tions of 0.052, 0.052 and 0.1
M, respectively, pH 7.5, at an assay

temperature of 60 °C.
Mutant k
cat
(s
)1
) K
m
(mM) k
cat
⁄ K
m
(s
)1
ÆmM
)1
)
H225A 1.40 ± 0.01 19.67 ± 0.75 0.07 ± 0.003
H225Q 8.97 ± 0.12 14.46 ± 0.80 0.62 ± 0.042
Y251F 37.17 ± 1.16 1.95 ± 0.23 19.11 ± 2.887
L-proline dehydrogenase from P. furiosus P. J. Monaghan et al.
2078 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
wild-type PRODH has been tentatively attributed to
ionization of l-proline in the Michaelis complex. Like
wild-type PRODH, all mutant enzyme forms displayed
a bell-shaped dependence of k
cat
⁄ K
m
as a function of
solution pH, and analysis of the data by fitting to Eqn

(4) (see Experimental procedures) gave macroscopic
pK
a
values of 6.8 ± 0.1 and 9.9 ± 0.1 (H225A),
6.8 ± 0.1 and 9.4 ± 0.2 (H225Q), and 6.0 ± 0.1 and
7.4 ± 0.1 (Y251F) (supplementary Fig. S2). The initial
idea that the pK
a
of 7.0 ± 0.1 determined for the
wild-type enzyme might represent ionization of the
conserved His225 active site residue in the free enzyme
has been rejected, as mutant pH dependence data
reveal that this ionization is not lost, but is apparent
from the acid limb of the bell-shaped fits in the
k
cat
⁄ K
m
plots for both H225A and H225Q mutant
forms. This analysis has revealed that His225 and
Tyr251 are not active site base residues, and are not
essential for catalysis.
The results obtained suggest that PRODH stabilizes
the deprotonated form of l-proline substrate in the
Michaelis complex, analogously to the substrate activa-
tion mechanisms observed in TMADH [25] and mono-
meric sarcosine oxidase [29]. Given this finding and the
absence of an active site residue that acts as base dur-
ing oxidation of l -proline, the data suggest that
PRODH-catalyzed amine oxidation may occur by

addition of a deprotonated l-proline at the C4 position
of the FAD cofactor and abstraction of a substrate
proton by the N5 atom of the flavin, a contemporary
mechanism of flavoprotein-catalyzed amine oxidation
proposed for TMADH [30].
Identification of product
The product of the enzyme-catalyzed oxidation of
l-proline was determined by monitoring the develop-
ment of the o-aminobenzaldehyde–P5C complex at
443 nm [31] and by MS. The rate constant for the
reaction of P5C with o-aminobenzaldehyde was a
direct function of o-aminobenzaldehyde concentration.
The value of the second-order rate constant was
42.4 s
)1
Æm
)1
, and analysis of the stoichiometry of con-
version indicated that the ratio of l-proline oxidized to
o-aminobenzaldehyde–P5C formed was 0.59, the spon-
taneous hydrolysis of P5C to glutamate–5-semialde-
hyde at the elevated assay temperature used (60 °C)
and rapid polymerization of free o-aminobenzaldehyde
accounting for the remaining 0.41 fraction not detected
as o-aminobenzaldehyde–P5C chromophore.
MALDI-TOF MS was employed for direct analysis
of product. Following enzyme turnover, a peak corres-
ponding to a mass ⁄ charge ratio of 114.1 was observed,
corresponding to the positively charged ion [M +H]
+

of P5C (supplementary Fig. S3). Additionally, we also
analyzed the product of o-aminobenzaldehyde reaction
with P5C using electrospray MS. In this case, a single
peak with a mass ⁄ charge ratio of 217 was observed,
corresponding to the positive ion of the P5C–o-amino-
benzaldehyde condensation product (Fig. 9).
Reduction potential of the enzyme-bound FAD
at physiologic temperature
The midpoint potential (E
m
) of FAD–PRODH was
determined by potentiometric redox titration with
sodium dithionite at ambient temperature and pH 7.0.
During the course of reductive titration, the oxidized
flavin was reduced directly to the dihydroflavin form,
without a visible population of a flavin semiquinone
species, indicating that the potential of the oxidized ⁄
semiquinone flavin couple is much lower than that of
the semiquinone ⁄ hydroquinone couple (Fig. 10A).
Data were fitted to the two-electron Nernst function
(Eqn 5) (see Experimental procedures) by least-squares
regression analysis, and gave a midpoint two-electron
potential value of ) 192 ± 3 mV and a corresponding
unrestricted RT/nF value of 28.9 ± 0.4 mV (where R
is gas constant, T is temperature, n is number of elec-
trons and F is Faraday constant), consistent with the
expected value (29.5 mV) for two-electron reduction of
the enzyme-bound FAD (Fig. 10B). The temperature
dependence of the two-electron midpoint potential was
measured within the range 7.5–31 °C (the limits

imposed by the performance of the electrode), and a
‘normal’ linear temperature dependence was found
(Fig. 10C). The temperature dependence of the mid-
point potential was calculated to be ) 3.1 ±
0.05 mVÆC°
)1
from the plot gradient. Extrapolation to
physiologic temperature (i.e. 100 °C for P. furiosus)
indicated an operational midpoint potential for
PRODH of ) 407 ± 5 mV. Thermodynamic para-
meters for the reduction of PRODH by sodium dithi-
onite were calculated from the temperature dependence
of the midpoint potential, and shown to be DH°¢ ¼
) 127.6 kJÆmol
)1
, DS°¢ ¼ ) 290.4 JÆmol
)1
ÆK
)1
, and
DG°¢
298
¼ ) 41.1 kJÆmol
)1
. Potentiometric redox titra-
tions of the H225A, H225Q and Y251F mutants at
25 °C all showed reduction of oxidized flavin directly
to the dihydroflavin form without a visible population
of a flavin semiquinone species. Mutant datasets were
fitted to the two-electron Nernst function (Eqn 5) (see

Experimental procedures) by least-squares regression
analysis, and gave midpoint potential values of
) 169 ± 3 mV (H225A), ) 155 ± 3 mV (H225Q),
and ) 157 ± 3 mV (Y251F) (supplementary Fig. S4),
P. J. Monaghan et al. L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2079
which compares with ) 174 ± 3 mV for the wild-type
enzyme at the same temperature (25 °C).
Discussion
The mechanism of substrate oxidation by flavoprotein
amine dehydrogenase ⁄ oxidases remains contentious.
In recent years, however, substrate oxidation by
members of this enzyme class has been shown to
occur by quantum mechanical tunneling [30,32]. We
and others have demonstrated that analysis of the
temperature dependence of kinetic isotope effects sug-
gests that H-transfer by quantum tunneling occurs
during substrate C–H bond cleavage, and that the
temperature effects are consistent with contemporary
environmentally coupled tunneling models [33]. A
major restriction in the use of variable temperature to
study tunneling reactions is the limited temperature
range available in experimental studies. For this rea-
son, in this study we have targeted a hyperthermo-
philic amine dehydrogenase, with a view to extending
in future work the temperature window available for
detailed physicochemical analysis of the enzyme
chemistry. Thus, we have cloned and expressed
two ORFs from P. furiosus that show some sequence
similarity with bacterial amine-specific flavoprotein

Fig. 9. Electrospray MS analysis of the o-aminobenzaldehyde adduct of the catalytic reaction product P5C generated during the oxidation of
L-proline by PRODH. The ionization mode used produced the positively charged quasimolecular ion ([M +H]
+
). The electrospray spectrum
showing an m ⁄ z peak of 217 represents the stable adduct formed from the reaction of P5C with o-aminobenzaldehyde. Inset: the proposed
mechanism for the reaction of P5C (1a) and o-aminobenzaldehyde (2). Following addition of activated P5C (1b) C4 carbon to the aldehyde
function of o-aminobenzaldehyde, redolent of a Knoevenagel condensation whereby the imine moiety of activated P5C is comparable to the
carbonyl in a conventional ketone–aldehyde reaction, the adduct precursor (3), following condensation and elimination of water, forms a sta-
ble adduct (4). The reaction is analogous to that of P5C with pyridoxal phosphate [46].
L-proline dehydrogenase from P. furiosus P. J. Monaghan et al.
2080 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
oxidases ⁄ dehydrogenases, including tetrameric sarco-
sine oxidase and DMGO. Conservation of residues
His225 and Tyr251 between PRODH and DMGO
provoked a mutagenesis strategy for PRODH to tar-
get these residues, which are suspected to prime sub-
strate during DMGO catalysis through deprotonation
of the substrate amine function prior to FAD reduc-
tion [34].
PRODH is a heterooctomer (ab)
4
, as confirmed by
determination of a preliminary crystal structure. Purifi-
cation of PRODH for mechanistic and redox studies
enabled the isolation of a form that contains a single
FAD cofactor noncovalently bound to the b subunit of
the protein, and that shows reactivity with l-proline,
generating P5C as product. l-pipecolic acid and sarco-
sine are slow substrates, and are not physiologic sub-
strates of PRODH, as judged from the observed

kinetic parameters. The enzyme is not an oxidase, and
does not use nicotinamide cofactors as electron accep-
tors. It also does not show any sequence homology
with the PRODH (PutA protein) of E. coli [7]. The
enzyme therefore represents a new member of a
recently identified class of PRODHs of hyperthermo-
philic origin.
We have shown that the PRODH active site is
situated in the b subunit and that these residues, as
suggested from structural and kinetic studies of
A. globiformis DMGO [14], play a role in the mechan-
ism of amine substrate oxidation but do not partici-
pate as catalytic base residues. The presence of iron
coordination in the Cys-clustered domain of the a sub-
unit could not be confirmed from the structure
(Fig. 4A). The anaerobic environment that P. furiosus
inhabits led to the possibility that any Fe–S cluster in
the PRODH a subunit may be aerobically labile, so
wild-type enzyme was expressed and purified under
strict anaerobic conditions (see Experimental pro-
cedures), and anaerobic assays were repeated with
P. furiosus ferredoxin and NADP
+
to measure elec-
tron transfer to putative physiologic acceptors via the
Fe–S cluster. No evidence of an Fe–S cluster was
apparent from the UV-visible absorption spectrum of
anaerobic PRODH, and no activity towards ferredoxin
and ⁄ or NADP
+

was observed.
Our kinetic studies indicate a strong pH dependence
on the kinetic parameters k
cat
and k
cat
⁄ K
m
. A single
Wavelength (nm)
300 400 500 600
Absorbance
0
0.2
0.4
0.6
0.8
Temperature (°C)
0 5 10 15 20 25 30 35
Midpoint Potential (mV)
-200
-180
-160
-140
-120
-100
-400 -300 -200 -100 1000
0.2
0.3
0.4

0.5
0.6
0.7
0.8
0.9
Abs (450 nm)
(Potential versus NHE) (mV)
A
B
C
Fig. 10. Redox potentiometric titration of wild-type monoflavinyl-
ated PRODH with sodium dithionite, and temperature dependence
of the midpoint potential. (A) Spectral changes accompanying
reductive titration of PRODH with sodium dithionite at 25 °C. (B)
Plot of absorbance at 450 nm versus the observed potential (cor-
rected against the normal hydrogen electrode). The data are fitted
to the Nernst equation (Eqn 5) for a two-electron reduction pro-
cess, giving a midpoint reduction potential, E
m
,of) 192 ± 3 mV,
and an RTF value of 28.9 ± 0.4 mV. (C) Plot of E
m
versus tempera-
ture, illustrating the linear dependence in the range 7.5–31 °C; gra-
dient, ) 3.1 ± 0.05 mVÆC°
)1
). The plot extrapolates to an operational
midpoint potential at physiologic temperature (100 °C for P. furio-
sus)of) 407 ± 5 mV. Conditions: 100 m
M potassium phosphate

buffer, pH 7.0. Mediator dyes used: methyl viologen (0.3 l
M), ben-
zyl viologen (1 l
M), 2-hydroxy-1,4-napthaquinone (7 lM), and phena-
zine methosulfate (2 l
M).
P. J. Monaghan et al.
L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2081
macroscopic pK
a
of 7.7 can be seen in the plot of k
cat
versus pH, and maximum activity is realized on the
alkaline side of this ionization. This is consistent with
kinetic studies of other amine-oxidizing flavoproteins,
such as TMADH [25], where the ionization has been
attributed to the deprotonation of the cationic form of
the substrate. This implies that the unprotonated form
of the substrate is reactive, consistent with proposed
mechanisms of amine substrate oxidation by flavopro-
tein enzymes [26]. On binding of substrate, the pK
a
for
the substrate ionization is sufficiently perturbed to fa-
vor formation of the free base form at physiologic pH.
In the case of TMADH, stopped-flow studies indicate
a shift in pK
a
of about ) 3.5 pH units (from pH 9.8

to $ pH 6.5) for substrate ionization on formation of
the enzyme–substrate complex [25]. Our data for
P. furiosus PRODH suggest that a similar shift in pK
a
of ) 2.9 pH units (from 10.6 to 7.7) occurs on binding
of l-proline to the enzyme, although unequivocal
demonstration of this must await detailed stopped-flow
studies with protiated and deuterated substrates. A sig-
nificant change in the pK
a
value for the ionization of
the alkaline limb of the k
cat
⁄ K
m
plot is seen following
mutagenesis (Fig. 8 and supplementary Fig. S2). At
this stage, we are unable to tentatively assign this ion-
ization to a functional group, but its location is most
likely in the active site. Mutagenesis could affect the
pK
a
of this neighboring group substantially. At this
stage, we cannot unequivocally rule out changes in the
rate-limiting step, or contributions from other ionizable
groups, as the origin of the observed dependence of k
cat
on solution pH. Mutagenesis studies of TMADH have
indicated a role for residues His172 and Tyr60 in per-
turbing the pK

a
of the substrate, and structural studies
of DMGO suggest that residues His225 and Tyr259,
which are conserved in PRODH, might also facilitate
deprotonation of substrate close to physiologic pH
values. In this regard, it is important to note that the
value of the Michaelis constant for l-proline is based
on the total concentration of substrate (i.e. cationic,
zwitterionic and deprotonated anionic forms). At
pH 7.5, the K
m
for substrate expressed in terms of the
unprotonated form only is 24.5 lm.
Concluding remarks
We have identified a new member of a recently identi-
fied class of PRODHs of hyperthermophilic origin. In
terms of cofactor content and redox chemistry, this
flavoprotein is simpler than the heterotetrameric dye-
linked PRODH from the hyperthermophilic archaeon,
T. profundus, an analog of which is also encoded in
the genome sequence of P. furiosus. Our work demon-
strates structural diversity among l-proline dehydro-
genase ⁄ oxidase enzymes. Our studies are now focused
on: (a) delineating the role of the two types of
PRODH in P. furiosus; and (b) addressing the mecha-
nisms of substrate oxidation and electron transfer.
Experimental procedures
Cloning and expression of genes encoding a
novel flavoenzyme amine dehydrogenase
Two ORFs [gi_18977617 and gi_18977618; protein extrac-

tion description and analysis tool (pedant) database] were
identified in the genome of P. furiousus DSM 3638 that
encode a putative flavoenzyme amine dehydrogenase ⁄ oxid-
ase. These genes were amplified by PCR using the oligonu-
cleotides 5¢-GTG AGA AAC TTG AGG CCA CTA GAC
TTA ACG G-3¢ and 5¢-TCA ACC CAT TTG AAG AGC
AAC AGT TCT TAA TTC TCC C-3¢. The PCR product
was purified by agarose gel electrophoresis, and used as
template DNA in a second PCR reaction to incorporate
flanking restriction sites (5¢ XbaI and 3¢ BamHI) for direc-
tional cloning, and a 5¢-ribosome-binding site to allow
expression of the cloned DNA. Primers used during this
PCR reaction were: 5¢-GGG GGG TCT AGA AAG GAG
ATA AAG AGA TGA GAA ACT TGA G-3¢,and5¢-GGG
GGG GGA TCC TCA ACC CAT TTG A AG AGC A-3 ¢.
The PCR product was digested with XbaI and BamHI,
ligated with vector pET11d, previously made end-compat-
ible with the same restriction endonucleases, and trans-
formed into Novablue cells (Novagen, Merck Chemicals
Ltd., Nottingham, UK). Restriction analysis confirmed pos-
itive recombinants, and DNA sequencing confirmed the
correct sequence of the recombinant clone, designated
pPRODH1.
A silent mutation was incorporated into pPRODH1 to
eradicate an internal ribosome-binding site responsible for
translation initiation within the gene, forming a truncated
population of the a subunit in E. coli. The following prim-
ers were used in the mutagenesis protocol marketed by
Stratagene (Amsterdam, the Netherlands) (QuikChange):
5¢-GGT GTC GAT GCT AGG AAA ACA AA

A GTT
AAA GAT GGA ATG AAA GTA C-3¢, and 5¢-GTA
CTT TCA TTC CAT CTT TAA C
TT TTG TTT TCC
TAG CAT CGA CAC C-3¢. The underlined letters indicate
the mismatch with the template DNA. The new expression
construct was designated pPRODH2.
The H225Ab, H225Qb and Y251Fb mutant enzymes
(mutations on b subunit) were also isolated using the
QuikChange mutagenesis protocol, using the pPRODH2
expression construct as template for primers: 5¢-CCA ATT
GAG CCC TAC AAG
GCT CAA GCA GTG ATA ACC-3¢
and 5¢-GGT TAT CAC TGC TTG A
GC CTT GTA GGG
CTC AAT TGG-3¢ (H225A); 5¢-CCA ATT GAG CCC
L-proline dehydrogenase from P. furiosus P. J. Monaghan et al.
2082 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
TAC AAG CAG CAA GCA G TG ATA ACC-3¢ an d 5¢-GGT
TAT CAC TGC TTG
CTG CTT GTA GGG CTC AAT
TGG-3¢ (H225Q); and 5¢-CAA GTA TGG TCA CGC
TT
T TTT AAC ACA AAC TGC GC-3¢ and 5¢-GCG CAG
TTT GTG TTA AA
A AAG CGT GAC CAT ACT TG-3¢
(Y251F). The underlined letters indicate the mismatch with
the template DNA. All mutant constructs were identified by
DNA sequencing, which also confirmed sequence integrity.
Wild-type PRODH was expressed and purified as described

previously [19]. Mutant forms were purified using the purifi-
cation protocol described for the wild-type enzyme. The pro-
tein concentration was determined by the method of
Bradford [35].
Anaerobic expression and partial purification
of wild-type PRODH
Anaerobic 2xYT media containing 20 mm sodium fumarate
was supplemented with ampicillin (50 lg ÆmL
)1
) and chlo-
ramphenicol (34 lgÆmL
)1
), and inoculated with E. coli
Rosetta(DE3)pLysS cells transformed with plasmid
pPRODH2. Cells were grown at 37 °C until the attenuance
at 600 nm reached $ 0.8. Cells were induced with isopropyl
thio-b-d-galactoside at a final concentration of 1 mm, and
grown for a further 48 h at 37 °C. Cells were harvested
under anaerobic conditions, resuspended in 50 mm anaero-
bic Mops buffer (pH 7.9), and sealed in an airtight centri-
fuge tube. Cells were lysed anaerobically by three cycles of
freeze–thaw treatment, and cell debris was removed by
centrifugation (12 100 g for 50 min using a Beckman
Avanti J-25 centrifuge, rotor type JA 25.50) following
DNA hydrolysis. The supernatant was transferred to an
airtight tube inside a glovebox, and the solution was subjec-
ted to a heat denaturation step at 80 °C for 1 h, with dena-
tured protein being removed by centrifugation (27 200 g for
30 min using a Beckman Avanti J-25 centrifuge, rotor type
JA 25.50). The supernatant was loaded onto an anaerobic

DE52 anion exchange column equilibrated with anaerobic
50 mm Mops buffer (pH 7.9), and anaerobic wild-type
PRODH was eluted with a 0–0.5 m NaCl gradient.
N-terminal protein sequence analysis and MS
N-terminal sequence analysis was performed using a ABI 476
Protein Sequencer (Applied Biosystems, Foster City, CA,
USA). Protein samples were electrophoresed by SDS-PAGE,
and electroblotted onto a poly(vinylidene difluoride) mem-
brane. Protein was stained with Coomassie Brilliant Blue
R250 to identify protein bands prior to excision from the
membrane for automated N-terminal sequence analysis.
MALDI-TOF MS of the purified enzyme was performed
with a Kratos Kompact MALDI-TOF III mass spectrometer
(Kratos, Manchester, UK), using sinapinic acid as matrix.
Electrospray MS was performed using a Waters Micromass
LCT TOF mass spectrometer (Waters Ltd., Elstree, UK).
Preparation of anaerobic samples
Buffers were made anaerobic by bubbling humidified oxy-
gen-free argon gas at 5 lbÆin
)2
through solutions for $ 2h
with stirring. Solutions were then transferred to an anaer-
obic glovebox, which was left open overnight to remove
residual oxygen. Samples of PRODH were made anaerobic
by passing them through a 10-DG column (Bio-Rad
Laboratories Ltd., Hemel Hempstead, UK) equilibrated
with anaerobic buffer in an anaerobic glovebox. Solutions
of substrates, dithionite, potassium ferricyanide and medi-
ator dyes were made by dissolving the appropriate solid in
anaerobic buffer. All anaerobic experiments were performed

in a Belle Technology (Portesham, UK) glovebox under a
positive pressure atmosphere of nitrogen, with residual oxy-
gen levels being maintained at < 0.05 p.p.m. with a BASF
(Cheadle, UK) R3-11 oxygen-scavenging catalyst.
Identification of flavin cofactor in PRODH
Following release from the enzyme, the chemical identity of
the flavin cofactor in PRODH was determined by MALDI-
TOF MS. Cofactor was released from the enzyme by heat
denaturation, and precipitated protein was removed by cen-
trifugation (17 000 g for 10 min at 4 °C in the dark using a
Microcentrifuge Fisherbrand accuSpin Micro 17R). Cofac-
tor was mass analyzed in double deionized H
2
O alongside
authentic FAD, FMN, ATP, ADP and AMP treated in the
same way. Analysis was performed using a Bruker Biflex
mass spectrometer Bruker Daltonics Ltd., Coventry, UK,
calibrated with a combination of peptides of known mass,
and the matrix was dihydroxybenzoic acid.
Optical titrations with reducing substrates
and steady-state kinetic analysis
PRODH (800 lL; 19.4 lm in 100 mm potassium phosphate
buffer, pH 7.5) was incubated in a quartz cuvette at 80 °C
for 10 min to allow temperature equilibration. Anaerobic
stocks (2 m) of dimethylglycine, sarcosine, glycine, glycine
betaine, l-proline, d-proline, l-pipecolic acid and sodium
sulfite were mixed with enzyme to a final concentration of
20 mm. Enzyme reduction was monitored by time-depend-
ent spectral acquisition in the region 300–600 nm.
Steady-state kinetic measurements with identified sub-

strates were performed spectrophotometrically in a 1 cm
light path in 100 mm potassium phosphate buffer (pH 7.5),
at 80 °C in a final reaction cell volume of 800 lL. Ferrice-
nium hexafluorophosphate was used as electron acceptor.
Buffer and substrate were equilibrated at the assay tem-
perature for 10 min prior to addition of ferricenium
(200 lm) followed by addition of 75 nm enzyme to initiate
the reaction. Initial velocities were measured by following
reduction of ferricenium (De
300
¼ 4300 m
)1
Æcm
)1
[36]), and
expressed as the concentration of ferricenium reduced
P. J. Monaghan et al. L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2083
per unit time (lmÆmin
)1
). Initial velocities as a function of
substrate concentration were analyzed by fitting to the
Michaelis–Menten equation. Similar reaction conditions
were employed when studying the activity of PRODH with
P. furiosus 4Fe)4S ferredoxin (the ferredoxin was supplied
by W Hagen, Delft University). Reduction of the 4Fe)4S
ferredoxin was monitored at 390 nm in an anaerobic glove-
box, as described previously for aldehyde ferredoxin oxido-
reductase activity [37], but using l-proline as substrate.
For the acquisition of temperature-dependent steady-

state data, enzyme activity was measured in a continuous
assay using the ferricenium ion as an artificial electron
acceptor. Substrate concentrations were maintained at 10
times K
m
. Reaction temperatures were recorded by direct
measurement inside the cuvette. The temperature was
recorded during temperature equilibration prior to the
reaction at both the top and the bottom of the cuvette to
detect any temperature gradient in the assay mixture.
Temperatures were also recorded immediately after assay
completion, and reactions were repeated if temperature
fluctuations exceeded 0.1 C°. Assays were initiated by
addition of microliter volumes of enzyme to ensure that
there was no significant effect on overall reaction tempera-
ture. Initial velocity data were plotted as a complete data-
set of rate versus time versus temperature, analogously to
the studies performed on thermophilic enzymes by Peter-
son et al. [38], who described an equilibrium model to
determine the thermal parameter T
eq
that represents a
submillisecond timescale-reversible temperature-dependent
equilibrium between active enzyme and inactive (or less
active) forms. The effect of a decrease in enzyme activity
above the optimal temperature occurring due to a shift in
T
eq
is up to two orders of magnitude greater than the
contribution of thermal denaturation. It is important to

note, however, that application of this proposed equilib-
rium model may be restricted to monomeric enzymes, as
the model does not account for the complicating effects of
thermally induced subunit dissociation of oligomeric
enzymes. The function of time in the three-dimensional
dataset is necessary to detect thermal inactivation ⁄ denatur-
ation of enzyme at temperatures up to and exceeding the
source (evolved) temperature.
For studies on pH dependence, steady-state assays were
performed over the pH range 5.5–10.0 in increments of
0.5 pH units, at an assay temperature of 60 °C. To keep
the ionic strength of the assay solutions constant over
the experimental pH range, a three-component buffer
system was employed, comprising Mes (pK
a
6.02),
3-{[tris(hydroxymethyl)-methyl]amino}-2-hydroxypropane
sulfonic acid (TAPSO; pK
a
7.49), and diethanolamine
(pK
a
8.88), at final concentrations of 0.052, 0.052 and
0.1 m, respectively [39]. pH profiles for the kinetic param-
eters k
cat
and K
m
were constructed, and the data were fit-
ted to Eqn (3) and Eqn (4), respectively, to obtain the

relevant pK
a
values.
k
cat
¼
EH Â 10
ðÀpHÞ
þ E Â 10
ðÀpK
a
Þ
10
ðÀpHÞ
þ 10
ðÀpK
a
Þ
ð3Þ
k
cat
K
m
¼
T
max
1 þ 10
ðpK
a1
ÀpHÞ

þ 10
ðpHÀpK
a2
Þ
ð4Þ
where EH and E are the catalytic activities of the proto-
nated and unprotonated forms of the ionization group,
respectively, and T
max
is the theoretical maximal value of
k
cat
⁄ K
m
.
Product identification and quantification
The product of the enzyme-catalyzed oxidation of l-proline
was identified using a modified method described for PutA
protein of enteric bacteria [31] and by MS. Recombinant
enzyme (75 nm) was used to oxidize l-proline (200 mm)ina
final reaction volume of 800 lL, using limiting ferricenium
ion (200 lm) as artificial electron acceptor in 100 mm potas-
sium phosphate buffer (pH 7.5). Assays were performed
at 60 °C, and allowed to progress until complete reduction
of ferricenium. The assay mixture was then incubated at
4 °C, and 100 lL of this mixture was added to
o-aminobenzaldehyde (0.5–4 mm) in the same assay buffer.
Formation of the o-aminobenzaldehyde–P5C chromophore
was followed at 443 nm at 25 °C. o-Aminobenzaldehyde was
made as a 50 mm stock solution in 20% ethanol, and stored

at ) 20 ° C. The extinction coefficient used for quantifying
the complex was e
443
¼ 2.71 mm
)1
Æcm
)1
. Baseline controls
were performed with each assay component in the absence
of enzyme. The o-aminobenzaldehyde–P5C chromophore
was analyzed using electrospray MS. Ten-microliter samples
were injected via a Rheodyne valve into a mobile phase of
methanol flowing at 0.2 mLÆmin
)1
into the electrospray
source. The source temperature was maintained at 80 °C,
and the needle voltage was $ 3.0 kV. The sample cone was
operated at 20 V, and nitrogen was used as the desolvation
and sheath gas at 600 and 100 LÆh
)1
, respectively. The spec-
trometer was calibrated with a solution of sodium iodide.
Direct analysis of enzyme reaction product was also per-
formed using MALDI-TOF MS. Following enzyme turn-
over, a 1 lL sample of reaction mixture was plated for
analysis on a Bruker Biflex mass spectrometer in positive
ion mode, using dihydroxybenzoic acid as matrix. A
1mgÆmL
)1
standard solution of authentic l-proline was

also prepared in double deionized H
2
O and filtered through
a 0.22 lm Millex-GP Acrodisc filter (Millipore Ltd.,
Watford, UK) for MALDI-TOF MS analysis. The spectro-
meter was externally calibrated using a combination of pep-
tides of known mass.
Redox potentiometry
Anaerobic redox titrations were performed between 5 and
31 °C in 100 mm potassium phosphate buffer (pH 7.0).
L-proline dehydrogenase from P. furiosus P. J. Monaghan et al.
2084 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
Enzyme (5 mL; $ 16.3 mgÆmL
)1
) was electrochemically
titrated following the method of Dutton [40], using sodium
dithionite as reductant and potassium ferricyanide as oxi-
dant. The mediator dyes methyl viologen (0.3 lm), benzyl
viologen (1 lm), 2-hydroxy-1,4-napthaquinone (7 lm) and
phenazine methosulfate (2 lm) were added immediately
prior to titration to facilitate equilibration across the range
+ 100 mV to ) 480 mV [41,42]. Equilibration times after
microliter addition of reductant were typically between 10
and 15 min, after which the potential was noted and the
corresponding absorbance spectra were measured between
280 and 800 nm. Complete titration curves typically consis-
ted of $ 40 different potential measurements. Plots of
absorbance against redox potential were fitted to Eqn (5)
[42].
A

obs
¼
ða þ b10
ðE
12
ÀEÞ=29:5
Þ
1 þ 10
ðE
12
ÀEÞ=29:5
ð5Þ
In Eqn (5), A
obs
is the absorbance value at the peak for oxi-
dized flavin at the electrode potential E, and a and b are the
absorbance values of the fully oxidized and reduced enzyme
at this wavelength, respectively. E
12
is the midpoint potential
for the concerted two-electron reduction of the flavin. Data
manipulation and analysis were performed using origin
software package version 6.0 (Microcal, OriginLab Corpora-
tion, Northampton, MA, USA). All redox potentials are
given relative to the standard hydrogen electrode.
Structure determination
Crystals were obtained as described previously [19].
Molecular replacement using the PRODH from Pyrococcus
horikoshii OT-3 as a model (75% and 90% identical to
P. furiosus PRODH a and b subunits, respectively) was per-

formed using AMoRe [43]. Initial electron density maps
were calculated using refmac5.0 [44], following several
rounds of positional refinement with strict noncrystallo-
graphic symmetry restraints imposed, leading to a R ⁄ R
free
ratio of 24.7 ⁄ 28.9 for all data (30–3.25 A
˚
).
Materials
Redox mediators, amine substrates, FAD, FMN, ATP,
ADP, AMP, sodium fumarate and buffer components were
obtained from Sigma-Aldrich (Gillingham, UK). Sodium
dithionite was obtained from FSA Laboratory Supplies
(Loughborough, UK), and complete protease inhibitor
cocktail tablets were purchased from Roche Diagnostics
(Burgess Hill, UK). Isopropyl thio-b-d-galactoside was
obtained from Melford Laboratories (Ipswich, UK). Oxy-
gen-free nitrogen and Pureshield brand argon were pur-
chased from the British Oxygen Company (BOC gases,
Guildford, UK). Broth components were purchased from
Oxoid (Basingstoke, UK). The plasmid pET11d and E. coli
strains Novablue and Rosetta (DE3)pLysS were obtained
from Novagen. Restriction enzymes were obtained from
New England Biolabs (Hitchin, UK). P. furiosus genomic
DNA was obtained from the American Type Culture Col-
lection (ATCC 43587). Ferricenium hexafluorophosphate
was synthesized as previously described [36].
Acknowledgements
We thank Mr Peter Ashton of the University of Bir-
mingham, UK for assistance with MS. We also thank

Mr Nahid Hasan and Professor W. Hagen of Delft
University, The Netherlands for supplying a sample of
P. furiosus 4Fe)4S ferredoxin. P. J. Monaghan thanks
the BBSRC for the award of a studentship. D. Leys
is a Royal Society University Research Fellow and
an EMBO Young Investigator. This work was funded
by the UK Biotechnology and Biological Sciences
Research Council and The Royal Society.
References
1 Brown ED & Wood JM (1992) Redesigned purification
yields a fully functional PutA protein dimer from
Escherichia coli. J Biol Chem 267, 13086–13092.
2 Menzel R & Roth J (1981) Purification of the putA
gene product. A bifunctional membrane-bound protein
from Salmonella typhimurium responsible for the two-
step oxidation of proline to glutamate. J Biol Chem 256,
9755–9761.
3 Scarpulla RC & Soffer RL (1978) Membrane-bound
proline dehydrogenase from Escherichia coli. Solubiliza-
tion, purification, and characterization. J Biol Chem
253, 5997–6001.
4 Maloy SR & Roth JR (1983) Regulation of proline uti-
lization in Salmonella typhimurium : characterization of
put::Mu d(Ap, lac) operon fusions. J Bacteriol 154,
561–568.
5 Menzel R & Roth J (1981) Regulation of genes for pro-
line utilization in Salmonella typhimurium: autogenous
repression by the putA gene product. J Mol Biol 148,
21–44.
6 Wood JM (1981) Genetics of l-proline utilization in

Escherichia coli. J Bacteriol 146, 895–901.
7 Lee YH, Nadaraia S, Gu D, Becker DF & Tanner JJ
(2003) Structure of the proline dehydrogenase domain
of the multifunctional PutA flavoprotein. Nat Struct
Biol 10, 109–114.
8 Polyak K, Xia Y, Zweier JL, Kinzler KW & Vogelstein
B (1997) A model for p53-induced apoptosis. Nature
389, 300–305.
9 Donald SP, Sun XY, Hu CA, Yu J, Mei JM, Valle D &
Phang JM (2001) Proline oxidase, encoded by p53-
induced gene-6, catalyzes the generation of proline-
P. J. Monaghan et al. L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2085
dependent reactive oxygen species. Cancer Res 61,
1810–1815.
10 Sakuraba H, Takamatsu Y, Satomura T, Kawakami R
& Ohshima T (2001) Purification, characterization, and
application of a novel dye-linked 1-proline dehydrogen-
ase from a hyperthermophilic archaeon, Thermococcus
profundus. Appl Environ Microbiol 67, 1470–1475.
11 Kawakami R, Sakuraba H & Ohshima T (2004) Gene
and primary structures of dye-linked l-proline dehydro-
genase from the hyperthermophilic archaeon Thermo-
coccus profundus show the presence of a novel
heterotetrameric amino acid dehydrogenase complex.
Extremophiles 8, 99–108.
12 Satomura T, Kawakami R, Sakuraba H & Ohshima T
(2002) Dye-linked D-proline dehydrogenase from
hyperthermophilic archaeon Pyrobaculum islandicum is a
novel FAD-dependent amino acid dehydrogenase. J Biol

Chem 277, 12861–12867.
13 Meskys R, Harris RJ, Casaite V, Basran J & Scrutton
NS (2001) Organization of the genes involved in
dimethylglycine and sarcosine degradation in Arthrobac-
ter spp.: implications for glycine betaine catabolism. Eur
J Biochem 268, 3390–3398.
14 Leys D, Basran J & Scrutton NS (2003) Channelling
and formation of ‘active’ formaldehyde in dimethyl-
glycine oxidase. EMBO J 22, 4038–4048.
15 Tsuge H, Kawakami R, Sakuraba H, Ago H, Miyano
M, Aki K, Katunuma N & Ohshima T (2005) Crystal
structure of a novel FAD-, FMN-, and ATP-containing
l-proline dehydrogenase complex from Pyrococcus hori-
koshii. J Biol Chem 280, 31045–31049.
16 Kawakami R, Sakuraba H, Tsuge H, Goda S,
Katunuma N & Ohshima T (2005) A second novel
dye-linked l-proline dehydrogenase complex is present
in the hyperthermophilic archaeon Pyrococcus horikoshii
OT-3. FEBS J 272, 4044–4054.
17 Wierenga R, Terpstra P & Hol W (1986) Prediction of
the occurrence of the ADP-binding aaa-fold in proteins,
using an amino acid sequence fingerprint. J Mol Biol
187, 101–107.
18 Vallon O (2000) New sequence motifs in flavoproteins:
evidence for common ancestry and tools to predict
structure. Proteins: Structure, Function Genet 38 ,
95–114.
19 Monaghan PJ, Leys D & Scrutton NS (2005) Crystalli-
zation and preliminary X-ray diffraction analysis of a
flavoenzyme amine dehydrogenase ⁄ oxidase from

P. furiosus DSM 3638. Acta Crystallogr F61, 756–758.
20 Chlumsky L, Zhang L & Jorns M (1995) Sequence ana-
lysis of sarcosine oxidase and nearby genes reveals
homologies with key enzymes of folate one-carbon
metabolism. J Biol Chem 270, 18252–18259.
21 Okazaki Y, Furuno M, Kasukawa T, Adachi J, Bono
H, Kondo S, Nikaido I, Osato N, Saito R, Suzuki H
et al. (2002) Analysis of the mouse transcriptome based
on functional annotation of 60,770 full-length cDNAs.
Nature 420, 563–573.
22 Massey V, Muller F, Feldberg R, Schuman M, Sullivan
PA, Howell LG, Mayhew SG, Matthews RG & Foust
GP (1969) The reactivity of flavoproteins with sulfite.
Possible relevance to the problem of oxygen reactivity.
J Biol Chem 244, 3999–4006.
23 Stevens L (1992) Buffers and the determination of pro-
tein concentration. In Enzyme Assays, a Practical
Approach (Eisenthal R & Danson MJ, eds), pp. 316–
335. IRL Press, Oxford.
24 Beynon R & Easterby J (1996) Buffer Solutions: the
Basics. BIOS Scientific Publishers, Oxford.
25 Basran J, Sutcliffe MJ & Scrutton NS (2001) Optimi-
zing the Michaelis complex of trimethylamine dehydro-
genase: identification of interactions that perturb the
ionization of substrate and facilitate catalysis with
trimethylamine base. J Biol Chem 276, 42887–42892.
26 Scrutton NS (2004) Chemical aspects of amine oxida-
tion by flavoprotein enzymes. Nat Prod Report 21,
722–730.
27 Messiha HL, Munro AW, Bruce NC, Barsukov I &

Scrutton NS (2005) Reaction of morphinone reductase
with 2-cyclohexen-1-one and 1-nitrocyclohexene: proton
donation, ligand binding, and the role of residues histi-
dine 186 and asparagine 189. J Biol Chem 280, 10695–
10709.
28 Messiha HL, Bruce NC, Sattelle BM, Sutcliffe MJ,
Munro AW & Scrutton NS (2005) Role of active site
residues and solvent in proton transfer and the modula-
tion of flavin reduction potential in bacterial morphi-
none reductase. J Biol Chem 280, 27103–27110.
29 Zhao G & Jorns MS (2005) Ionization of zwitterionic
amine substrates bound to monomeric sarcosine oxi-
dase. Biochemistry 44, 16866–16874.
30 Basran J, Sutcliffe MJ & Scrutton NS (2001) Deuterium
isotope effects during carbon–hydrogen bond cleavage
by trimethylamine dehydrogenase. Implications for
mechanism and vibrationally assisted hydrogen tunnel-
ing in wild-type and mutant enzymes. J Biol Chem 276,
24581–24587.
31 Wood JM (1987) Membrane association of proline
dehydrogenase in Escherichia coli is redox dependent.
Proc Natl Acad Sci USA 84, 373–377.
32 Harris RJ, Meskys R, Sutcliffe MJ & Scrutton NS
(2000) Kinetic studies of the mechanism of C–H
bond breakage by the heterotetrameric sarcosine oxid-
ase of Arthrobacter sp-1-IN. Biochemistry 39, 1189–
1198.
33 Sutcliffe MJ, Masgrau L, Roujeinikova A, Johannissen
LO, Hothi P, Basran J, Ranaghan KE, Mulholland AJ,
Leys D & Scrutton NS (2006) Hydrogen tunnelling in

enzyme-catalyzed H transfer reactions: flavoprotein and
quinoprotein systems. Phil Trans R Soc Ser B 361,
1375–1386.
L-proline dehydrogenase from P. furiosus P. J. Monaghan et al.
2086 FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS
34 Basran J, Fullerton S, Leys D & Scrutton NS (2006)
Mechanism of FAD reduction and role of active site
residues His-225 and Tyr-259 in Arthrobacter globi-
formis dimethylglycine oxidase: analysis of mutant
structure and catalytic function. Biochemistry 45 ,
11151–11161.
35 Bradford MM (1976) A rapid and sensitive method for
the quantitation of microgram quantities of protein util-
izing the principle of protein-dye binding. Anal Biochem
72, 248–254.
36 Lehman T, Hale D, Bhala A & Thorpe C (1990) An
acyl-coenzyme A dehydrogenase assay utilizing the ferri-
cenium ion. Anal Biochem 186, 280–284.
37 Mukund S & Adams MW (1991) The novel tungsten–
iron–sulfur protein of the hyperthermophilic archae-
bacterium, P. furiosus, is an aldehyde ferredoxin
oxidoreductase. Evidence for its participation in a
unique glycolytic pathway,. J Biol Chem 266, 14208–
14216.
38 Peterson ME, Eisenthal R, Danson MJ, Spence A &
Daniel RM (2004) A new intrinsic thermal parameter
for enzymes reveals true temperature optima. J Biol
Chem 279, 20717–20722.
39 Ellis K & Morrison J (1982) Buffers of constant ionic
strength for studying pH-dependent processes. Methods

Enzymol 87, 405–426.
40 Dutton PL (1978) Redox potentiometry: determination
of midpoint potentials of oxidation–reduction compo-
nents of biological electron-transfer systems. Methods
Enzymol 54, 411–435.
41 Daff SN, Chapman SK, Turner KL, Holt RA,
Govindaraj S, Poulos TL & Munro AW (1997) Redox
control of the catalytic cycle of flavocytochrome P-450
BM3. Biochemistry 36, 13816–13823.
42 Munro AW, Noble MA, Robledo L, Daff SN &
Chapman SK (2001) Determination of the redox
properties of human NADPH-cytochrome P450
reductase. Biochemistry 40, 1956–1963.
43 Navaza J (1994) AMoRe: an automated package for
molecular replacement. Acta Crystallogr A50, 157–163.
44 Murshudov GN, Vagin AA & Dodson EJ (1997)
Refinement of macromolecular structures by the maxi-
mum-likelihood method. Acta Crystallogr D53, 240–255.
45 Cleveland WS (1993) Visualising Data. Hobart Press,
Summit, NJ.
46 Farrant RD, Walker V, Mills GA, Mellor JM & Lang-
ley GJ (2001) Pyridoxal phosphate de-activation by
pyrroline-5-carboxylic acid. Increased risk of vitamin B6
deficiency and seizures in hyperprolinemia type II.
J Biol Chem 276, 15107–15116.
Supplementary material
The following supplementary material is available
online:
Fig. S1. Plot of initial velocity versus l -proline concen-
tration for the Y251F PRODH enzyme.

Fig. S2. Dependence of steady-state kinetic parameters
on solution pH for the oxidation of l-proline substrate
catalyzed by the H225A, H225Q and Y251F PRODH
enzyme forms.
Fig. S3. MALDI-TOF MS identifying the mass of P5C
produced from the oxidation of substrate l-proline cata-
lyzed by PRODH of P. furiosus DSM 3638.
Fig. S4. Redox potentiometric titration of H225A,
H225Q and Y251F mutant PRODH forms with sodium
dithionite.
This material is available as part of the online article
from
Please note: Blackwell Publishing is not responsible
for the content or functionality of any supplementary
materials supplied by the authors. Any queries (other
than missing material) should be directed to the corres-
ponding author for the article.
P. J. Monaghan et al. L-proline dehydrogenase from P. furiosus
FEBS Journal 274 (2007) 2070–2087 ª 2007 The Authors Journal compilation ª 2007 FEBS 2087