Báo cáo khoa học: Crystal structure and solution characterization of the activation domain of human methionine synthase doc

Crystal structure and solution characterization of the
activation domain of human methionine synthase
Kirsten R. Wolthers
1,
*, Helen S. Toogood
1,
*, Thomas A. Jowitt
1
, Ker R. Marshall
2
, David Leys
1
and
Nigel S. Scrutton
1
1 Faculty of Life Sciences, University of Manchester, UK
2 Department of Biochemistry, University of Leicester, UK
Human methionine synthase (EC 2.1.1.13; 5-methyl-
tetrahydrofolate homocysteine methyltransferase, hMS)
plays a vital role in folate metabolism and the recyc-
ling of homocysteine. It is the only enzyme that
liberates tetrahydrofolate (H
4
folate) from methyltetra-
hydrofolate (CH
3
-H
4
folate), which is a key metabolite
for protein and nucleic acid biosynthesis. The enzyme
contains a cobalamin cofactor, and in the highly nucle-

ophilic cob(I)alamin state, the cofactor abstracts a
methyl group from CH
3
-H
4
folate to form H
4
folate
and methylcob(III)alamin (Scheme 1, 1a) [1,2]. The
methyl group is subsequently transferred from
methylcob(III)alamin to homocysteine to generate
methionine and cob(I)alamin (Scheme 1, 1b) [3].
Keywords
activation domain; cobalamin-dependent
enzyme; methionine synthase; methionine
synthase reductase; S-adenosyl-methionine
Correspondence
N. S. Scrutton, Manchester Interdisciplinary
Biocentre and Faculty of Life Sciences,
University of Manchester, 131 Princess
Street, Manchester M1 7ND, UK
Fax: +44 161 306 8918
Tel: +44 161 306 5152
E-mail:
Database
The atomic coordinates and structure fac-
tors (202K) have been deposited in the Pro-
tein Data Bank, Research Collaboratory for
Structural Bioinformatics, Rutgers Univer-
sity, New Brunswick, NJ, USA (http://

www.rcsb.org)
*These authors contributed equally to this
work
(Received 3 October 2006, revised 22
November 2006, accepted 28 November
2006)
doi:10.1111/j.1742-4658.2006.05618.x
Human methionine synthase (hMS) is a multidomain cobalamin-dependent
enzyme that catalyses the conversion of homocysteine to methionine by
methyl group transfer. We report here the 1.6 A
˚
crystal structure of the
C-terminal activation domain of hMS. The structure is C-shaped with the
core comprising mixed a and b regions, dominated by a twisted antiparallel
b sheet with a b-meander region. These features, including the positions of
the active-site residues, are similar to the activation domain of Escheri-
chia coli cobalamin-dependent MS (MetH). Structural and solution studies
suggest a small proportion of hMS activation domain exists in a dimeric
form, which contrasts with the monomeric form of the E. coli homologue.
Fluorescence studies show that human activation domain interacts with the
FMN-binding domain of human methionine synthase reductase (hMSR).
This interaction is enhanced in the presence of S-adenosyl-methionine.
Binding of the D963E ⁄ K1071N mutant activation domain to the FMN
domain of MSR is weaker than with wild-type activation domain. This
suggests that one or both of the residues D963 and K1071 are important in
partner binding. Key differences in the sequences and structures of hMS
and MetH activation domains are recognized and include a major reorien-
tation of an extended 3
10
-containing loop in the human protein. This struc-

tural alteration might reflect differences in their respective reactivation
complexes and ⁄ or potential for dimer formation. The reported structure is
a component of the multidomain hMS : MSR complex, and represents an
important step in understanding the impact of clinical mutations and poly-
morphisms in this key electron transfer complex.
Abbreviations
AdoMet, S-adenosyl-methionine; AUC, analytical ultracentrifugation; FLD, flavodoxin; FNR, ferredoxin-NADP
+
oxidoreductase; hMS, human
methionine synthase; MALLS, multiangle laser light scattering; MS, methionine synthase; MSR, methionine synthase reductase.
738 FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS
The latter half of the reaction highlights additional
roles of MS in cell homeostasis (a) the production of
methionine, which is an essential amino acid and a
precursor in the biosynthesis of S-adenosyl-methionine
(AdoMet); and (b) the recycling of homocysteine,
which is cytotoxic to vascular endothelial cells and an
independent risk factor in coronary arterial disease
[4,5]. High total plasma homocysteine coupled with
diminished folate pools has also been associated with
an increased incidence of neural tube defects in new-
borns and Down’s syndrome [6,7].
These medical conditions may arise from either a
vitamin deficiency or inborn errors in the gene enco-
ding hMS or the gene encoding the enzyme involved in
the reactivation of hMS, human methionine synthase
reductase (hMSR) [8]. The activity of hMS ceases after
% 1–2000 catalytic turnovers with the one-electron oxi-
dation of cob(I)alamin (Scheme 1, 2) [3,9]. Human
MSR binds to hMS forming a ‘reactivation complex’

and an NADPH-derived electron is shuttled to
cob(II)alamin via the FAD and FMN cofactors of
MSR (Scheme 1, 3) [9]. Transfer of a methyl group
from AdoMet accompanies reduction by MSR, thus
converting cob(II)alamin to methylcob(III)alamin and
returning MS to the primary catalytic cycle. Reactiva-
tion of the Escherichia coli homologue of hMS, MetH,
also involves FAD and FMN redox centres; however,
the cofactors are components of individual proteins:
ferredoxin-NADP
+
oxidoreductase (FNR) and flavo-
doxin (FLD), respectively [10].
To date, the majority of information on the struc-
tural and functional behaviour of hMS has been
derived from biochemical and biophysical research on
MetH [11]. Although the 3D structure of the full-
length MetH has not been determined, structures of
three individual functional modules have been solved.
The N-terminal module, determined from MetH
of Thermotoga maritima, consists of two b,a barrels,
which each house substrate-binding pockets for homo-
cysteine and CH
3
-H
4
folate [12]. The cobalamin is
sandwiched between two domains in the central mod-
ule [13]. Finally, the C-terminal domain, termed the
activation domain, binds AdoMet and forms part of

the reactivation complex with hMSR [14]. The struc-
ture of the E. coli MetH activation domain is
C-shaped, with a twisted antiparallel b sheet as a cen-
tral feature. AdoMet binds near the centre of the inner
surface of the domain and is held in place by interac-
tions with both side-chain and backbone atoms [14].
MetH, and by extension MS, are envisioned to be
highly dynamic proteins as both substrate-binding
pockets on the N-terminal module (separated by 50 A
˚
)
and the activation domain have to form discrete com-
plexes with the cobalamin-binding domain in order to
catalyse each of the transmethylation reactions of the
primary catalytic cycle and reactivation process [15,16].
hMS has an added level of complexity compared
with MetH, as its redox partner, MSR is itself a multi-
domain protein; MSR is modelled on the structural
family of diflavin reductases, of which cytochrome
P450 is the prototype [17]. Enzymes belonging to this
class of proteins have a NADPH ⁄ FAD-binding
domain, tethered to an FMN-binding protein, which
are related to bacterial FNR and FLD, respectively.
Studies have shown that with the two-component
system of E coli, FLD forms mutually exclusive com-
plexes with MetH and FNR [18]. If the FMN-binding
region of MSR behaves similarly, this domain would
pivot between hMS and the FAD domain of MSR
to facilitate electron transfer. Alternatively, the FMN
domain is relatively immobile and is sandwiched

between the FAD domain and hMS during electron
transfer to cob(II)alamin. In this case, the binding
interfaces between the activation domain of hMS and
MSR may be sufficiently different from that of the
MetH activation domain and FLD.
Evidence for notable differences in the binding inter-
face between MetH and hMS is supported by the poor
ability of the FNR ⁄ FLD-reducing system to reactivate
hMS and the complete inability of MSR to reactivate
MetH [19]. Structural information on the hMS activa-
tion domain will help establish those key components
on these proteins that make them specific for their
respective redox partners. In addition, structural infor-
mation will help identify how particular clinical and
polymorphic (e.g. P1173L) variations appearing in the
MS activation domain compromise the activity of the
enzyme and lead to various clinical states [20,21].
Here, we provide insight into the biophysical proper-
ties of human MS ⁄ MSR system and we report the
crystal structure of a D963E ⁄ K1071N double-mutant
CH
3
-H
4
-folate
H
4
-folate
Cob(I)alamin
Methylcob(III)alamin

Methionine
Homocysteine
Cob(II)alamin
e
AdoMet
AdoHyc
e
1a
2
3
1b
Scheme 1.
K. R. Wolthers et al. Human MS activation domain structure
FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS 739
of the 38 kDa activation domain of hMS to 1.6 A
˚
resolution. Solution studies demonstrate binding of
both the wild-type and mutant activation domains to
the FMN domain of the physiological partner protein,
MSR. We show that the human activation domain
exists as a distribution of monomeric and dimeric
forms, with the monomer comprising the main compo-
nent in solution. Key structural differences between
the hMS activation domain and the E. coli homologue
are discussed.
Results
Crystal structure determination
The mutant activation domain structure of hMS was
determined by molecular replacement using the struc-
ture of the corresponding domain of MetH as a model

[14]. The asymmetric unit contains two protein mole-
cules, related by near-perfect translational symmetry.
The crystallographic and final refinement statistics are
summarized in Table 1. The last 1–2 residues are not
visible in the electron-density map. Similarly, residues
T1001–G1003 and D1072–A1074, which are located in
flexible loops, are not visible. In humans, residue
Gln1041 in both subunits is in a disallowed region in
the Ramachandran plot. It is one of only two residues
in the sharp turn between b2 and b3 in the b meander.
In several organisms, including E. coli, this residue is
glycine (G1008) which can accommodate this geometry.
The structure of the activation domain monomer of
human MS is C-shaped, the central feature comprising
a twisted antiparallel b sheet (Fig. 1A), similar to the
E. coli structure [14]. This is not surprising as the two
MS proteins share 48% sequence identity in the activa-
tion domain, with the human activation domain con-
taining several insertions (Fig. 1B). The core of the
structure comprises mixed a + b regions, dominated by
an antiparallel b sheet, with an overall topology that
does not resemble any other AdoMet-binding protein
structure [14]. An antiparallel b sheet (b1, b2, b5 and
b8) forms the upper part of the structure along with
strands b3 and b4, which form a b meander. On the
opposite side of the sheets, after the meander, is a
region of six a helices and a 3
10
-helix. Helices a2–a4
connect the strands b1 and b2, whereas helices a5–a7

follow strand b5. The long helix a6 is surrounded by the
central b sheet on one side, b6–b7 and the short helix a7
on the other side. The C-terminus is dominated by two
short helices a10–a11 [14]. The two mutations D963E
and K1071N are located after b1 and b4, respectively,
in regions distant from the AdoMet-binding site. Owing
to their location in flexible surface loops, these muta-
tions are not thought to have a major impact on the
overall structure, but rather have only localized effects.
Structural comparison of the hMS and E. coli
MetH activation domains
Figure 2A shows a superimposition of the structures
of the hMS activation domain and corresponding
MetH domain in E. coli. Using the program dalilite
[22], the rmsd value of the superimposition is 2 A
˚
with
a Z-score of 41.9 (314 residues aligned by C a).
Although the two structures look very similar, there
are significant differences between them, most strik-
ingly in the region of helices a3–a4. In the human
enzyme, helix a3 is extended by a further four amino
acids followed by the insertion of an extra five amino
acids between a3 and the 3
10
2-helix. This has resulted
in a dramatic reorientation of these latter two helices,
relative to the E. coli enzyme, beginning at residue
Leu984 (Leu956 in MetH) and ending at the beginning
of the a4 helix. In the E. coli enzyme, this region is

oriented so the 3
10
-helix is within 4 A
˚
of helix a7, and
within 7 A
˚
of Ile1126 in the AdoMet-binding region.
In the human structure, the longer a3 helix ends at
the beginning of the extra five amino acids. The 3
10
2-
helix is oriented below the beginning of helix a4 and is
Table 1. Crystallographic data and refinement values for hMS
activation domain double-mutant D963E ⁄ K1071N. All numbers in
parentheses represent last outer shell (1.66–1.60 A
˚
).
D963E ⁄ K1071N mutant
Data collection
Resolution limits (A
˚
) 20–1.6 (1.66–1.60)
Space group P212121
Unit cell (A
˚
)a¼ 77.8 b ¼ 90.1 c ¼ 123.0
Total reflections 110417
Unique reflections 106145
Redundancy 5.3

Completeness (%) 96.3 (96.5)
I ⁄ rI 20.1 (3.2)
R
merge
(%) 6.4 (40.2)
Solvent content (%) 58
Refinement
No. of residues 666
No. of water molecules 1205
R
fac
⁄ R
free
(%) 20.9 ⁄ 22.3 (27.0 ⁄ 29.9)
rmsd bond lengths (A
˚
) 0.006
rmsd bond angles (°) 1.0
Average B-factor (A
˚
2
) 25.2
Ramachandran plot
Most favoured regions (%) 94.1
Additionally allowed regions (%) 5.6
Generously allowed regions (%) 0.0
Disallowed regions (%) 0.3 (Q1041 both subunits)
Human MS activation domain structure K. R. Wolthers et al.
740 FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS
close to the three amino acid turn between helices a5–

a6 (Fig. 2A). This positions the 3
10
-helix at one of the
tips of the C-shaped molecule, preventing it from being
in a position near the AdoMet-binding region as in
MetH. This creates a new hydrophobic cluster not
found in the E. coli enzyme between residues Phe993
and Phe997 and the a3 and a6 helices. Other hydro-
phobic residues involved include Trp982, Val1009,
Tyr1121, and Ile1124. Part of this region is disordered
in the human enzyme, with clear electron density lack-
ing for residues Thr1001–Gly1003.
The role of this loop)3
10
region in E. coli is uncer-
tain, but owing to its location near the AdoMet-bind-
ing site it may be involved in interacting with its FLD
partner. Recent cross-linking and NMR studies of
MetH show that Lys959 (Lys987 in hMS) is located
close to Glu61 of FLD [18,23] in the activation
domain–FLD complex, supporting the notion that this
is a key interaction in the formation of the reactivation
complex. hMS interacts with a much larger partner,
MSR, which is a cytochrome P450 reductase-like pro-
tein constructed by the fusion of two genes encoding a
FMN-containing FLD and an FAD-containing ferre-
doxin oxidoreductase separated by a large interdomain
linker [9,17]. The change in position of this loop)3
10
region of the human enzyme in addition to the shift in

position of Lys987 from the equivalent MetH residue
Lys959 of 7.9 A
˚
(Fig. 2A) suggests a different mode of
interaction between human MS and MSR compared
with the E. coli complex. Because no structure is avail-
able for MSR, or any structure of a MS–partner com-
plex, the exact nature of these interactions remains to
be determined.
The region containing b3–b5, known as the b mean-
der [14], contains a six amino acid insertion in the
human structure (Fig. 1A). The b-meander begins with
a twist in b2 at the conserved residue Pro1036 (Pro1003
in E. coli), as indicated by an asterisk in Figs 1B and
2A. This meander is oriented % 90° to the central sheet
[14]. In the human enzyme, three of these extra residues
are located between b3 and b4, forming an additional
solvent-exposed 3
10
-helix (3
10
3), absent in the E. coli
enzyme. The other three extra amino acids are located
in the more disordered region between b4–b5, next to
A
B
Fig. 1. (A) Stereoview of the overall struc-
ture of the hMS activation domain. The car-
toon is drawn as a gradient from blue at the
N-terminus to red at the C-terminus. Red

spheres, location of the mutations D963E
and K1071N; magenta sphere, clinical muta-
tion P1173L. (B) Sequence alignment of
human and E. coli MS activation domains,
confirmed by structural superimposition.
Secondary structure elements were
assigned using
DSSP [28]. Pink, secondary
structure in humans; green, major differ-
ences in secondary structure in E. coli
compared with humans; orange, additional
3
10
-helices in humans; asterisk, Pro1036
(beginning of b-meander); blue, residues
interacting with AdoMet in E. coli.
K. R. Wolthers et al. Human MS activation domain structure
FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS 741
the mutation K1071N. The equivalent lysine residue in
E. coli activation domain (Lys1035) is suggested to be
involved in the interaction between MS and FLD [18].
We note that in the two regions of the protein where
the major differences occur between the E. coli and
human structures, both contain a residue which in
MetH is known to interact with FLD. This suggests
that these differences between both structures of the
activation domains might reflect structural differences
in their respective reactivation complexes.
Like MetH, the structure of the hMS activation
domain contains some well-ordered water molecules in

two cavities which interact with residues in the b-mean-
der region. This region, containing helices a10–a11, is
one of the more conserved regions of the activation
domain. It contains many buried ionic and polar resi-
dues, forming an extended network of salt bridges and
hydrogen bonds similar to the E. coli enzyme [14].
Active-site region
A superimposition of the two structures at the MetH
AdoMet-binding site shows a high similarity of both
residue type and position (Fig. 2B). AdoMet binds
near the centre of the inner surface of the domain
of the E. coli structure (Fig. 2A). It is held in place
by hydrophobic interactions and hydrogen bonds,
through both side chain and backbone interactions,
and is partly solvent exposed [14]. This is one of the
more conserved regions of the activation domain and
contains the consensus sequence Arg-X-X-X-Gly-Tyr
critical for the binding of AdoMet [14]. The human
enzyme structure does not contain AdoMet, although
the positions of the active site residues are strikingly
similar.
Only two of the residues (Tyr1190 and Ala1141)
known to directly interact with AdoMet in the E. coli
protein are different in human activation domain (i.e.
Phe1228 and Ser1179, human numbering). However,
these residues in the E. coli protein interact with Ado-
Met via backbone interactions only. The main struc-
tural difference in the active site residues is the
reorienting of the Tyr1177 side chain away from where
AdoMet would bind. This suggests only minimal chan-

ges in the positions of the active site residues are
required for binding of AdoMet.
A
B
Fig. 2. Stereoview of a superimposition of
human and E. coli MS activation domains.
(A) Overall structures: blue and green car-
toons are human and E. coli proteins,
respectively; AdoMet from the E. coli struc-
ture (1MSK) [14]; is shown as red sticks; Ca
atoms of K987 and K959 of human and
E. coli activation domains, respectively, are
shown as red spheres. (B) AdoMet-binding
region: E. coli residues interacting with Ado-
Met, and the equivalent human residues,
are shown as atom-coloured sticks with
green and orange carbons, respectively;
AdoMet is shown as purple lines.
Human MS activation domain structure K. R. Wolthers et al.
742 FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS
Monomer interactions in the crystalline state
The crystal structure reveals that both monomers
within the asymmetric unit are involved in close inter-
actions with symmetry-related molecules, leading to a
putative dimeric form. Figure 3A shows two views of
the structure of this human MS activation domain
‘dimer’. The contact surface between the two mono-
mers, consisting of two nearly identical interaction
sites, is % 609 A
˚

3
with a shape complementarity (S
c
)
value of 0.769, which is in the range of S
c
values for
surfaces within known dimeric proteins and protein–lig-
and interactions [24]. There were no other interactions
of sufficient size between monomers of the activation
on the structure, and their symmetry-related molecules
to be able to accurately calculate S
c
values. Both the
size of the contact surface, the S
c
value and the fact
both monomers in the asymmetric unit form near iden-
tical dimers suggests dimer formation could be physio-
logically relevant rather than simply a consequence of
the crystallization conditions and ⁄ or crystal packing.
The central cavity of the dimer consists of a large
elliptical-shaped central groove, with extensions at the
base of the dimer forming a cross shape. This exten-
sion of the cavity is due to the side chains of several
residues of the extended a3 helix of both subunits
pointing directly into the cavity. The volume of this
cavity is calculated to be % 7000 A
˚
3

using the program
CASTp [25]. The smallest part of the cavity (24 · 5A
˚
)
is at the lower region of the dimer (Fig. 3A) close to
where AdoMet binds in the E. coli structure [14]. Two
AdoMet molecules can be modelled to bind within the
dimer, with the large size of the cavity easily allowing
substrate entry. However, the side chains of residues
Tyr988 and Lys987 of the second molecule of the
‘dimer’ clash with the region where the adenosyl group
of AdoMet is bound in the MetH structure. While the
side chain of Lys987 can move to accommodate Ado-
Met, the side chain of Tyr988 cannot, suggesting the
dimeric form is not compatible with substrate binding.
Superimposition of the E. coli structure onto the
human activation domain ‘dimer’ shows significant cla-
shes due to the reorientation of the a 3–3
10
loop region
(Fig. 3A). Clearly, the E. coli activation domain is unli-
kely to form a human-like dimer. Figure 3B shows the
residues involved in forming the dimer interface of the
human activation domain. Although there is a signifi-
cant contact surface between the monomers, the number
of direct interactions is small. The key interacting resi-
due is Arg991. The NH2 group of Arg991 forms a
hydrogen bond with the carbonyl oxygen of Glu1077
(2.7 A
˚

). The NH1 and NH2 groups are also within van
A
B
Fig. 3. Dimeric nature of human MS activa-
tion domain. (A) Two views of the human
activation domain dimer, with the mono-
mers shown as green and blue cartoons.
AdoMet is superimposed onto the structure
as red sticks in the equivalent position
found in the E. coli enzyme [11]. E. coli
‘dimer’ superimposed on the human dimer
is represented by an olive cartoon. (B) The
dimer interface of human activation domain
shown as green and blue cartoons for
monomers A and B, respectively. Side
chains of residues directly interacting at the
dimer interface are shown as atom-coloured
sticks. Side chains of residues lining a
hydrophobic pocket are shown as magenta
sticks.
K. R. Wolthers et al. Human MS activation domain structure
FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS 743
der Waal’s distance of the hydroxyl group of Tyr1079,
as is the NE group to the backbone oxygen atoms of
Pro1078 and Glu1077. The OD2 atom of Asp1120
forms a salt bridge with the NH2 group of Arg927
(2.7 A
˚
). The interface is furthermore formed by hydro-
phobic interaction between Tyr988 of one subunit with

Pro1173, Tyr1177, Pro1178 and Tyr1227 of the second
subunit. As the equivalent residues of the E. coli activa-
tion domain are located in significantly different posi-
tions, these dimer interface interactions are not present.
Molecular mass determination and dimer
formation
A key question to address is whether the dimer species
as seen is a significant species in solution. To answer
this question, we carried out multiangle laser light
scattering (MALLS) and analytical ultracentrifugation
(AUC) to determine the proportion of dimer formed
in solution under both high and low salt conditions.
Table 2 shows the results of the sedimentation and
light-scattering analyses. Both wild-type and the dou-
ble-mutant activation domains show near identical
sedimentation profiles (Fig. 4A) with corrected sedi-
mentation coefficients (s
0
20;W
) of 2.87 ± 0.16 and
2.84 ± 0.2, respectively, in 0.4 m sodium acetate buf-
fer (i.e. similar to the crystallization conditions). How-
ever, in NaCl ⁄ P
i
and Tris ⁄ HCl buffers in the absence
of sodium acetate, the sedimentation coefficient was
increased to 2.98 ± 0.2. Thus, the molecular mass of
the two proteins are identical, as shown by light scat-
tering, and there is no detectable difference in shape
between the wild-type and mutant proteins in the same

buffer systems (Fig. 4B). However, differences in the
solution properties are seen for both proteins in differ-
ent buffers. The addition of 0.4 m sodium acetate to
the buffers increases the presence of the dimer species
(14% of the total protein analysed). The estimated
frictional ratios for the monomeric species using the
sedimentation coefficient distribution (c(s)), where
f ⁄ f
0
¼ estimated frictional ratio, for 2D size and shape
distributions showed that there is a difference in
the apparent length or flexibility of the molecules in
the two different buffers. In 0.4 m sodium acetate, the
molecules exhibited a more extended or more flexible
conformation with a frictional ratio of 1.45 compared
with a value of 1.31 representing a more compact
structure in NaCl ⁄ P
i
. This corresponds to a difference
in hydrodynamic radius of % 3A
˚
(Table 2).
Global analysis of mutant and wild-type protein in
high salt buffer at different concentrations was per-
formed to ascertain whether there was evidence of
reversible association occurring between monomeric
and dimeric species. The results fitted well to a mono-
mer–dimer model. However, only a very small amount
of dimer is present. This was true even at very high
concentrations of 2 mgÆmL

)1
, giving a very low associ-
ation constant. This indicates that, although there may
be dimer species present within the samples in 0.4 m
sodium acetate, there is no real evidence for a dynamic
associating system.
The molecular mass obtained throughout these
experiments was lower than expected (Table 2) for the
sequence molecular masses when taking into account
the calculated mass for hydrated protein. This might be
attributed to a small amount of protease cleavage during
purification. The molar mass distribution is near identi-
cal for the wild-type and mutant proteins giving an aver-
age monomeric molecular mass of 41 600 ± 1020 Da.
The high polydispersity detected is similar to the results
gained using sedimentation equilibrium indicating that a
proportion of the molecules are slightly smaller than
sequence molecular mass (Fig. 4C). Some of the dimer
species can also be seen using this technique.
Analysis of dimer interactions by chemical
cross-linking
Inspection of the dimer interface reveals that Lys925
of one subunit of the activation domain ‘dimer’ is
Table 2. Solution studies of wild-type and mutant activation domain to determine oligomeric states in high- and low-salt buffers. NaCl ⁄ P
i
;M,
observed hydrated mass of the monomer; S, sedimentation coefficient; R
H
, radius of the structure; f ⁄ f
o

, estimated frictional ratios; ND, not
determined; NA, not applicable.
R
Experimental
M (kDa)
a
M (kDa)
b
S
0
20,W R
H
a
(nm) R
H
b
(nm) f ⁄ f
0
Bead modelling
Monomer
R
H
(nm) S
0
20,W
Dimer
R
H
(nm) S
0

20,W
WT NaCl ⁄ P
i
ND 43.10 ± 1.5 2.98 ± 0.2 2.83 2.7 ± 0.3 1.27 2.97 3.25 3.56 5.0
Mutant NaCl ⁄ P
i
ND 43.0 ± 1.5 3.0 ± 0.2 2.92 2.6 ± 0.2 1.31 ND ND ND ND
WT HS
c
36.20 ± 2.2 40.80 ± 0.8 2.87 ± 0.2 3.23 2.7 ± 0.1 1.45 NA NA NA NA
Mutant HS
c
35.12 ± 2.0 41.60 ± 1.0 2.84 ± 0.1 3.18 3.0 ± 0.2 1.43 NA NA NA NA
a
Determined by sedimentation studies.
b
Determined by light scattering.
c
100 mM Tris ⁄ HCl pH 7.0 + 400 mM Na acetate.
Human MS activation domain structure K. R. Wolthers et al.
744 FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS
likely to be in close proximity to the N-terminal amino
group of the second subunit. To determine if this spe-
cific surface interaction occurs frequently in solution,
an attempt was made to cross-link these two specific
primary amine groups with a homo-bifunctional imi-
doester cross-linker. Mass analysis of in-gel trypsinized
samples showed that although the 80-kDa band
observed in SDS–PAGE was indeed a dimer of activa-
tion domain molecules, the location of the cross-link

indicated a ‘back-to-back’ orientation for the two
monomers and is thus inconsistent with the crystal
structure (results not shown). This suggests that either:
(1) the concentration of the dimer species under cross-
linking conditions is very small; (2) the location of the
N-terminus in solution is not consistent with models of
the dimer (the N-terminus is not visible in the crystal
structure of the activation domain); and ⁄ or (3) there is
some proteolytic removal of the N-terminus during
purification.
Analysis of the interaction between MS
activation domain and the FMN domain of MSR
Titration of the activation domain of MS (both the
D963E ⁄ K1071N mutant and wild-type proteins) with
the FMN domain of human MSR resulted in a
quenching of the fluorescence emission spectra of the
flavin cofactor. The decrease in fluorescence intensity
showed a hyperbolic dependence on the concentration
of the activation domain (Fig. 5). A fit of Eqn (1) to
the data yielded the apparent dissociation constant
for the complex formed between activation domain
and the FMN-binding region of MSR (Table 3). The
wild-type activation domain has an apparent K
d
value 1.3 lm for the FMN domain, which is 10-fold
lower than the apparent K
d
(11.9 lm) for binding of
the D963E ⁄ K1071N to the flavin-binding protein.
The presence of AdoMet in the fluorescence bind-

ing assays resulted in a twofold decrease in the
apparent dissociation constant for both the wild-
type (0.7 lm) and the mutant (4.7 lm) activation
domains.
Discussion
hMS is important for maintaining adequate levels of
methionine and AdoMet, preventing the accumulation
of cytotoxic homocysteine, and is essential in methi-
onine metabolism. Elevated levels of homocysteine in
the blood have been linked to an increased likelihood
of developing cardiovascular disease, birth defects,
Down’s syndrome and affecting the development of
some types of cancer [4–7,26]. Functional deficiency
of MS or MSR results in diseases such as homocys-
tinuria, hyperhomocysteinemia and hypomethionine-
mia [8,20]. A P1173L mutation (magenta sphere in
A
0246810
0246810
0.0
0.2
0.4
0.6
0.8
1.0
1.2
c(s)
c(s)
Sedimentation coefficient (s)
0.0

0.5
1.0
1.5
2.0
2.5
3.0
Sedimentation coefficient (S)
0 50000 100 000 150000 200 000
0.0
2.0x10
-5
4.0x10
-5
6.0x10
-5
8.0x10
-5
1.0x10
-4
1.2x10
-4
c(M)
Molar Mass (Da)
B
10 12 14 16
0
20 000
40 000
60 000
80 000

10 0000
Refractive Index
Molar Mass (Da)
Volume (ml)
0.0
0.2
0.4
0.6
0.8
1.0
C
Fig. 4. (A) Sedimentation coefficient distribution c(s) of wild-type
(solid line) and mutant (dotted line) in 10 m
M Tris, 150 mM NaCl
pH 7.0. Inset shows the same protein run in 0.4
M sodium acetate.
(B) Wild-type molar mass distribution c(M) using the estimated fric-
tional ratio (f ⁄ f
0
) of 1.45. (C) Wild-type (solid line) and mutant (dot-
ted line) elution from a Superdex 200 gel filtration column in
NaCl ⁄ P
i
+0.4 M sodium acetate, with the absolute molar mass
superimposed. The thick dotted line within the major peak shows
the degree of polydispersion of the enzyme in NaCl ⁄ P
i
.
K. R. Wolthers et al. Human MS activation domain structure
FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS 745

Fig. 1B) in the activation domain of human MS is
commonly found among patients exhibiting hype-
rhomocysteinemia [21]. This residue is located
between Arg1172 and Ala1174, both of which interact
directly with AdoMet in the E. coli structure
(Fig. 2B). P1173L is located at the start of a loop in
the active site and which contains four proline resi-
dues of the sequence P-X-P-X-X-P-X-X-P. This
sequence is highly conserved among MS enzymes sug-
gesting an evolutionary pressure to retain this struc-
ture in the active site of the activation domain. Other
known clinical mutations of the human activation
domain include H920D, E1204X (early termination),
and insertion ⁄ deletion mutations [21].
The suggestion of a dimer of human activation
domain in the crystal structure as well as in some solu-
tion studies raises some important issues. As the acti-
vation domain is the C-terminal domain of MS,
separated from the rest of the protein by a 38-residue
linker region, a key question to be addressed is the
possibility of the full-length enzyme forming a dimeric
structure. Also, the possibility of binding Ado-
Met ± the FMN domain of MSR influencing the like-
lihood of dimer formation needs to be considered.
That the major differences between the hMS and
MetH activation domains occur in those regions
involved in dimer association is intriguing. This might
reflect the need to recognize different redox partners,
i.e. FLD versus MSR, but a role in dictating the oligo-
meric state of MS cannot be ruled out. That said, only

a small proportion of dimer is found in solution stud-
ies, and this is most prevalent under high salt condi-
tions. Clearly, further structural analysis of other
components of the MS holoenzyme is required to
ascertain if full-length hMS possesses higher order
quaternary structure.
Fluorescence titration assays have demonstrated an
interaction between the FMN-binding domain of
MSR and the activation domain of MS. The appar-
ent K
d
value determined from these binding assays is
similar to that reported for the interaction between
E. coli FLD and its redox partner, MetH [23]. The
decrease in dissociation constant for the complex with
the wild-type and mutant form of the activation
domain in the presence of AdoMet suggests that
binding of the substrate for the transmethylation
reaction might effect a conformational change in the
activation domain to increase its affinity for the
FMN domain of MSR.
The 10-fold higher dissociation constant measured
for the mutant activation domain ⁄ FMN domain inter-
action indicates that Asp963 and ⁄ or Lys1071 are
important residues in this interaction. Several lysine
mutants of E coli FLD have also showed a marked
decrease (3- to 70-fold) in affinity towards MetH [23],
suggesting that salt bridges are a key recognition
02468101214
0

2
4
6
8
10
[Act domain] μM
A
0 5 10 15 20 25 30
0.0
0.5
1.0
1.5
2.0
2.5
3.0
3.5
Δ Fluorescence (AU)
Δ Fluorescence (AU)
[Act domain] μM
B
Fig. 5. Fluorescence titration of the FMN domain of MSR with the
wild-type (A) and D963E ⁄ K1071N double-mutant (B) of the MS acti-
vation domain. The FMN domain at 0.25 l
M was titrated with both
forms of the purified activation domain under conditions described
in the Experimental procedures. The binding assays were per-
formed in the absence (d) or presence (s)of1m
M S-adenosyl
methionine. The change in the FMN fluorescence intensity was
plotted versus the concentration of the activation domain and the

curves show the best fit of the data to the quadratic Eqn (1).
Table 3 lists the calculated dissociation constants for the wild-type
and D963E ⁄ K1071N alone and in the presence of AdoMet.
Table 3. Fluorescence titration of the FMN domain of MSR with
the wild-type and D963E ⁄ K1071N double-mutant of the MS activa-
tion domain. The FMN domain at 0.25 l
M was titrated with both
forms of the purified activation domain under conditions described
in the Experimental Procedures.
Enzyme
K
d
(lM)
No AdoMet
1mM
AdoMet
Wild-type 1.3 ± 0.1 0.7 ± 0.1
D963E ⁄ K1071N 11.9 ± 1.5 4.7 ± 0.6
Human MS activation domain structure K. R. Wolthers et al.
746 FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS
feature at the binding interface in both the E. coli and
human systems.
While this study describes the structure of a double-
mutant of the activation domain, the location of the
mutations are on flexible, surface-exposed loops. We
conclude from this observation, combined with the
MALLS and AUC data which showed that the wild-
type and mutant activation domains have identical
shapes, that the structure of the mutant domain resem-
bles closely that of the wild-type activation domain.

To further understand the process of MS reactiva-
tion, a structure of the cob(I)alamin ± activation
domains in complex with at least the FMN domain of
MSR is needed (studies that we are currently pursuing).
The structure of the activation domain reported here
has allowed us to gain insight into the likely mode of
AdoMet binding, and provide atomic level insight into
the effect of clinical mutations on the activity of MS.
Experimental procedures
Cloning and mutagenesis
The cDNA encoding the activation domain of methionine
synthase gene was cloned by PCR amplification, using
nondegenerate oligonucleotides based on the published
sequence. Total RNA was isolated and purified from whole
human blood using a High Pure RNA isolation kit (Roche
Diagnostics, Welwyn Garden City, UK). cDNA was gener-
ated by reverse transcription using the Titan RT PCR sys-
tem (Roche Diagnostics). The MS gene was cloned into the
plasmid vector pET15b, and the 1184 bp PCR product cor-
responding to the 3¢-terminus of the MS gene was gener-
ated with Pfu turbo DNA polymerase (Stratagene, La
Jolla, CA) and ligated into a pGEMT vector (Promega,
Madison, WI) to generate the vector pGEMMB. The
sequence encoding the activation domain (residues 925–
1265) was subsequently amplified by PCR from the
pGEMMB vector, incorporating the restriction sites NcoI
and HindIII into the 5¢- and 3¢-regions, respectively. These
restriction sites were used to subclone the PCR product
into pET23d, generating clones containing a C-terminal
His-tag. Sequencing of the resulting vector, pACT, revealed

two missense mutations that resulted in conversion of an
Asp at position 963 to a Glu and a Lys to an Arg at posi-
tion at 1071. (At this stage we cannot say if the identified
differences to the published sequence of human methionine
synthase (GenBank accession number Q99707) represent
polymorphic variation of the MS gene.) The pACT vector
was subsequently mutated to revert the sequence back to
wild-type. Both the wild-type gene and the uncorrected
mutant D963E ⁄ K1071N were subsequently transformed
into competent E. coli BL21(DE3) (Stratagene). MSR
FMN domain was cloned as a glutathione-S-transferase
fusion protein according to the method of Wolthers et al.
[27].
Protein production and purification
Wild-type and mutant activation domain proteins were
expressed in Terrific broth containing 100 lgÆmL
)1
ampi-
cillin. Cells were lysed by sonication in buffer A (20 mm
K
2
HPO
4
⁄ KH
2
PO
4
buffer, pH 7.4, and 0.5 m NaCl) also
containing 1 mm MgCl
2

, EDTA-free Complete protease
inhibitor tablets (Roche) and Benzonase (Merck Bio-
sciences Ltd., Nottingham, UK). The proteins were puri-
fied by running through Ni-NTA resin contained in 5 mL
HisTrap column (Amersham Biosciences, GE Healthcare,
Little Chalfont, UK) in Buffer A. Proteins were eluted in
a gradient from 0 to 0.5 m imidazole. For the final puri-
fication step, the protein was loaded onto HiPrep Q
Sepharose (Amersham Biosciences, GE Healthcare) in
50 mm Tris ⁄ HCl pH 8.0 (Buffer B) and eluted in a gradi-
ent from 0 to 1 m NaCl.
The expression and purification of the FMN domain of
MSR was carried out according to a modification of the
method of Wolthers et al. [27]. An additional purification
step was performed using Resource Q (Amersham Bio-
sciences, GE Healthcare) in Buffer B. The protein was elut-
ed in a gradient of 0–0.5 m NaCl.
Crystallogenesis and data collection
Crystals of the wild-type and mutant MS activation
domains were grown using the sitting drop vapour diffusion
method at 19 °C. The enzyme (10 mgÆmL
)1
) was desalted
into 10 mm Tris pH 7.0 containing 0.1 mm EDTA and
0.5 mm dithiothreitol. The reservoir solution comprised 0.1
m Tris ⁄ HCl pH 7.5 containing 0.1–0.4 m sodium acetate,
10–12% poly(ethylene glycol) 8000 and 10–12% poly(ethy-
lene glycol) 1000. Crystals appeared between 2 and 14 days.
Crystals were soaked in mother liquor supplemented with
5% poly(ethylene glycol) 200 as a cryorotectant, before

being flash-cooled in liquid nitrogen. A full 1.6 A
˚
data set
was collected on a single crystal of the mutant MS activa-
tion domain at the European Synchrotron Radiation Facil-
ity (Grenoble, France) on ID14-EH1 using an ADSC Q4
CCD detector. Owing to problems associated with twin-
ning, a model could not be obtained for data collected from
wild-type crystals.
Structure determination and refinement
Data were processed and scaled using the HKL package
programs denzo and scalepack [28]. The structure was
solved via molecular replacement using the program
amore [29] and the deposited structure of the MS activa-
tion domain from E. coli MetH [14] as the search model
K. R. Wolthers et al. Human MS activation domain structure
FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS 747
(PDB code 1MSK). Positional and B-factor refinement
was performed using refmac5 [30] with alternate rounds
of manual rebuilding of the model in turbo-frodo [31].
Partial rebuilding of the model and building of waters
was carried out using arp/warp [32]. The quality of the
model was checked using the program procheck [33].
Determination of the oligomeric state of human
activation domain in solution
The absolute hydrated molecular mass of hMS activation
domain was determined by MALLS coupled to gel-filtra-
tion chromatography. Approximately 50 lg of both wild-
type and mutant activation domains were loaded onto a
Superdex 200 24 ⁄ 30 gel filtration column (Amersham Phar-

macia Biotech, Piscataway, NJ) at 0.5 mLÆmin
)1
in either
NaCl ⁄ P
i
buffers pH 7.4 or 10 mm Tris, 0.15 m NaCl
pH 7.4 ± 0.4 m sodium acetate. The samples were passed
through a DAWN EOS 18-detector photometer using a
688 nm laser to induce scattering with a QELS dynamic
light scattering attachment and an Optilab rEX refractome-
ter (Wyatt Technologies, Santa Barbara, CA, USA).
AUC using an Optima XLA ultracentrifuge was used to
assess the overall shape of the molecules and to determine
if there was any reversible association occurring between
oligomeric states. Sedimentation velocity was performed
using two sector cells at 104 936 g collecting 100 scans at
10-min intervals using a wavelength of 230 nm. Protein
samples were run in NaCl ⁄ P
i
as well as 10 mm Tris,
0.15 m NaCl pH 7.4 ± 0.4 m sodium acetate buffers at a
concentration of % 2 lm . Data were analysed using the
continuous size distribution analysis program sedfit [34].
Size-distribution analysis of macromolecules was carried
out by sedimentation velocity ultracentrifugation and
Lamm equation modelling [34]. Sedimentation coefficients
were corrected for standard conditions using a

m value of
0.734 calculated from the amino acid sequence of the acti-

vation domain. Sedimentation equilibrium experiments
were carried out using six-sector cells at speeds of 5881,
16 336 and 32 000 g at concentrations of 1, 2.5 and 5 lm.
Global analysis of the results was performed using sed-
phat [35].
The shape complementarity of the dimer interface was
investigated using the CCP4 program S
c
[24]. The shape
correlation between interacting surfaces A and B can be
defined as:
S
c
¼ðfS
A!B
gþfS
B!A
gÞ=2
Where the braces denote the median (50th percentile) of
the distribution of S
A fi B
and S
B fi A
values over two sur-
faces A and B, respectively. A full discussion of the calcu-
lations can be found in of Lawrence and Colman [24].
These calculations were done on both the dimer interface
and between symmetry-related monomers in a nondimeric
conformation.
Cross-linking of MS activation domain

The homo-bifunctional imidoester cross-linker, dimethyl
pimelimidate 2 HCl (Pierce, Rockford, IL) with a spacer
arm of 9.2 A
˚
, was used in cross-linking reactions with puri-
fied activation domain. A 10- and 25-fold molar excess of
dimethyl pimelimidate was added to the activation domain,
in 0.2 m triethanolamine pH 8.0, at 5.0 and 0.5 mgÆmL
)1
protein, respectively. The cross-linking reaction was allowed
to proceed for 45 min at 25 °C before quenching with gla-
cial acetic acid. Protein samples were electrophoresed in a
12% SDS–PAGE gel. An 80 kDa band, equivalent to a
‘dimer’ of activation domain, as well as the monomeric
40 kDa band, was subject to in-gel digestion by trypsin,
and the peptides were identified by MS.
Analysis of binding between MS activation
domain and MSR FMN domain
Fluorescence quenching experiments were performed in
50 mm Tris ⁄ HCl pH 7.5 at 25 °C in a 3 mL volume on
a Cary Eclipse Fluorescence Spectrophotometer (Varian,
Oxford, UK). The FMN domain (0.25 lm) was excited at
450 nm, the absorbance maxima for the FMN cofactor,
and emission spectra were recorded between 500 and
600 nm with the excitation and emission slit widths set at
10 and 5 nm, respectively. After addition of activation
domain (in 2–10 lL aliquots), the sample was mixed and
incubated for 1 min before the stable fluorescence emission
spectra were recorded. The change in the fluorescence inten-
sity at the emission maxima (529 nm) was plotted against

the concentration of the activation domain and the data
were fit to a quadratic binding Eqn (1) to obtain the disso-
ciation constant for the FMN domain—activation domain
complex.
DF ¼ F
o
þ 2DF
max
ðE
0
þ L þ K
d
ÞÀ
ffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
ðE
0
þ L þ K
d
Þ
2
À 4E
0
L
q
ð1Þ
In Eqn (1), DF represents the change in fluorescence inten-
sity, F
0
is the measure of the fluorescence intensity in the
absence of the activation domain, E

0
is the concentration
of the FMN domain, L denotes the concentration of the
activation domain added to the sample, and K
d
represents
the dissociation constant for the FMN domain—activation
domain complex.
Acknowledgements
This study was funded by the UK Biotechnology
and Biological Sciences Research Council. NSS is
a BBSRC Professorial Research Fellow; DL is an
EMBO Young Investigator and Royal Society Univer-
sity Research Fellow.
Human MS activation domain structure K. R. Wolthers et al.
748 FEBS Journal 274 (2007) 738–750 ª 2006 The Authors Journal compilation ª 2006 FEBS
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