Structural basis for protein–protein interactions
in the 14-3-3 protein family
Xiaowen Yang*, Wen Hwa Lee*, Frank Sobott*, Evangelos Papagrigoriou*, Carol V. Robinson†, J. Gu ¨nter Grossmann‡,
Michael Sundstro ¨m*, Declan A. Doyle*§, and Jonathan M. Elkins*
*Structural Genomics Consortium, University of Oxford, Botnar Research Centre, Oxford OX3 7LD, United Kingdom;†Department of Chemistry, University of
Cambridge, Cambridge CB2 1EW, United Kingdom; and‡Molecular Biophysics Group, Council for the Central Laboratory of the Research Councils Daresbury
Laboratory, Warrington WA4 4AD, United Kingdom
Edited by Tony Hunter, The Salk Institute for Biological Studies, La Jolla, CA, and approved October 2, 2006 (received for review July 10, 2006)
The seven members of the human 14-3-3 protein family regulate a
diverse range of cell signaling pathways by formation of protein–
protein complexes with signaling proteins that contain phosphor-
crystal structures of three 14-3-3 isoforms (zeta, sigma, and tau)
have been reported, with structural data for two isoforms depos-
ited in the Protein Data Bank (zeta and sigma). In this study, we
provide structural detail for five 14-3-3 isoforms bound to ligands,
providing structural coverage for all isoforms of a human protein
family. A comparative structural analysis of the seven 14-3-3
proteins revealed specificity determinants for binding of phos-
phopeptides in a specific orientation, target domain interaction
movements. Specifically, the structures of the beta isoform in its
apo and peptide bound forms showed that its binding site can
exhibit structural flexibility to facilitate binding of its protein and
peptide partners. In addition, the complex of 14-3-3 beta with the
exoenzyme S peptide displayed a secondary structural element in
the 14-3-3 peptide binding groove. These results show that the
is likely to facilitate recognition and binding of their interaction
phosphorylation ? signaling
the human genome revealed seven isoforms: beta (?), epsilon (?),
eta (?), gamma (?), tau (?), sigma (?), and zeta (?), each encoded
by a distinct gene. The 14-3-3 proteins form homo- and het-
erodimers (2, 3) that have been shown to interact with a large
number of proteins, e.g., ? was shown in a yeast two-hybrid study
to have as many as 130 potential binding partners (4). The 14-3-3
proteins are involved in the regulation of metabolism, signal
transduction, cell-cycle control, apoptosis, protein trafficking, tran-
scription, stress responses, and malignant transformation (5, 6)
mainly through binding to phosphopeptides, thus modulating sig-
naling events. Examples of well validated 14-3-3 interaction part-
ners are the proto-oncogene RAF-1 (7–9) and the cell-cycle
regulatory phosphatases Cdc25C?B (10, 11).
All isoforms recognize two high-affinity phosphorylation-
dependent 14-3-3 binding motifs: RSXpSXP (mode 1) and
RXXXpSXP (mode 2), where pS represents a phosphoserine
(12–14). The 14-3-3 proteins can also recognize unmodified
proteins such as the Pseudomonas aeruginosa virulence factor
exoenzyme S (ExoS), p190RhoGEF, and the phage display-
derived R18 peptide inhibitor (15–19). Recently, a small number
of proteins that interact with 14-3-3 proteins via a C-terminal
phosphorylation motif have also been identified (20, 21). Phos-
phorylation-dependent and -independent binding have been
shown to be targeted to the same site of the 14-3-3 proteins (18).
The 14-3-3 proteins have been found to be up- or down-
regulated in human disease, but their direct role in disease
progression has not been clearly established. Examples of such
conserved and ubiquitously expressed (1). Later, sequencing of
disease conditions are as follows: (i) the ? isoform has been
implicated in breast cancer (22); (ii) ? has been implicated in
neurological disorders (23); and (iii) ? in the cerebrospinal fluid
(CSF) can be used as a marker for sporadic Creutzfeldt-Jakob
disease (CJD) (24, 25), although studies on a ?-deficient mutant
mouse showed that it is unlikely to play a causal role. In addition,
?, ?, and ? are also found in the CSF of CJD patients (26); (iv)
?, ?, and ? expression levels are increased in lung cancer as
compared with the equivalent normal tissues (27); (v) ? has been
shown to be a specific marker of lung cancer (28); and (vi) ? is
necessary for proper G2checkpoint function (29).
Further evidence of the importance of 14-3-3 proteins has
been provided in mouse studies. (i) Mice deficient in ? have
it has been shown in humans to be absent in certain human
neuronal migration disorders such as the Miller–Dieker syn-
drome (30). (ii) Dominant-negative mutant forms of ? were
shown to inhibit ERK MAPK activation but increased the
activation of JNK1 and p38 MAPK leading to increased apo-
ptosis (31), and (iii) a dominant-negative ? increased the basal
activation of JNK1 and p38 MAPK and affected the ability of
mice to compensate for pressure overload resulting in increased
mortality, cardiomyopathy, and massive cardiomyocyte apopto-
sis (31). In contrast, the survival rate, anatomy, and cage
behavior of ?-deficient mice were normal (32). A recent example
displaying the physiological importance of 14-3-3 proteins dem-
onstrated that they are required for lifespan extensions in C.
elegans when promoted by extra copies of the sir-2.1 gene (33).
The mechanisms of 14-3-3 action have been suggested to
target protein, (ii) physically occluding sequence-specific or
structural features, (iii) scaffolding, and (iv) changing cellular
localization (34). Previously, a number of three-dimensional
structures of 14-3-3 isoforms have been determined and depos-
ited in the Protein Data Bank (PDB; www.pdb.org): ?, in its apo
form (35), as peptide complexes (14, 17, 18) and in complex with
serotonin N-acetyltransferase (AANAT) (36); and ? in apo- and
peptide-bound states (37, 38). In addition, the structure of ? was
published (39) but not deposited in the PDB. These structural
data showed that the 14-3-3 proteins adopted a similar confor-
mation in the unliganded and peptide?protein-bound structures,
Author contributions: X.Y. and W.H.L. contributed equally to this work; D.A.D. and J.M.E.
F.S., M.S., D.A.D., and J.M.E. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS direct submission.
Abbreviations: ExoS, exoenzyme S; ODA, optimal docking area.
Data deposition: The crystal structures reported in this paper have been deposited in the
Protein Data Bank, www.pdb.org (PDB ID codes 2BR9, 2BTP, 2B05, 2C74, 2C63, 2C23, and
§To whom correspondence should be addressed. E-mail: email@example.com.
© 2006 by The National Academy of Sciences of the USA
November 14, 2006 ?
vol. 103 ?
no. 46 ?
to which the interaction partner adapts, and led to the hypothesis
that the 14-3-3 proteins may act as ‘‘molecular anvils’’ (40).
We set out to determine the structures of all remaining 14-3-3
isoforms (including the ? isoform, PDB ID 2BTP) to be able to
conduct a thorough comparative structural analysis and in
addition performed a study on dimerization preferences. Here
we report the structures of apo-?, ? in complex with an ExoS
peptide, ? as a mode 2 peptide complex, ? as both mode 1 and
2 peptide complexes, ? as a mode 2 complex, and ? as a mode
1 peptide complex, and we analyze them together with the
available ? and ? structures.
Results and Discussion
Structure Overview. The overall structural features of the 14-3-3
protein family members were recently reviewed (41), and thus
only a brief description is provided here. The determined 14-3-3
crystal structures were all homodimers. Each monomer consists
of a bundle of nine ?-helices (?A to ?I) organized into groups
of two, two, two, and three helices (39, 42). The first four are
essential for formation of the dimer, which has a sizeable
aperture at the subunit interface (Fig. 1). Helices ?C, ?E, ?G,
and ?I form a conserved peptide-binding groove, which has a
positively charged patch on one side and a hydrophobic patch on
the other (39, 42). The positively charged patch is formed by a
conserved triad of two arginines and a tyrosine residue (Arg-57,
Arg-130, and Tyr-131; residue numbers will all refer to the ?
isoform unless otherwise specified) that bind the phosphate
group of the interacting phosphopeptide?protein. Tables 1 and
2, which are published as supporting information on the PNAS
web site, contain the crystallization and data collection details
for the seven structures generated in this study.
Dimerization Preferences. Previous studies have shown that, apart
from ?, several of the isoforms are able to form heterodimers (2,
3). The presence of unique residues at the ? dimer interface
provided a clear structural explanation for its homodimerization
preference (38). Using an in vivo-based approach, Chaudhri et al.
(2) found that the ? isoform preferentially formed heterodimers
with no observable homodimer formation. In addition phos-
phorylation influences the dimerization process (43, 44).
In this study, we investigated the dimerization equilibria of the
?, ?, ?, ?, and ? isoforms using an in vitro-based approach
allowing a quantitative description of both the homo- and
Dimerization Equilibria of 14-3-3 Proteins. Dimerization equilibria
of ?, ?, ?, ?, and ? were investigated by nanoelectrospray
ionization mass spectrometry under conditions where noncova-
lent complexes are preserved, i.e., gentle ionization and desol-
vation conditions and the appropriate pressure settings (see ref.
45). All five isoforms were detected as dimers (Fig. 6A, which is
published as supporting information on the PNAS web site). The
? and ? isoforms were almost entirely dimeric, ? was mostly
dimeric, and ? and ? were found to be in equilibrium with the
monomeric form (70:30 ratio, dimer?monomer) under the same
experimental conditions. This ratio reflects the relative stability
of the dimeric interaction in solution, which appears to be higher
for ? and ? as compared with ? and ?.
We also studied the formation of heterodimers between four
overnight before measurements were taken. Shorter incubation
periods, for instance, of 1 h, did not allow the subunit exchange
to proceed to completion. The results of this interaction study,
presented as percentage of observed heterodimers, are as fol-
lows: ???, ?95%; ???, 50%; ???, 50%; ???, ?95%; ???, ?95%;
and ???, 50%. The ? isoform showed a preference for binding of
?, ?, and ? to an extent that virtually no homodimers were
detected (Fig. 6B). Thus, the ? subunit has a higher affinity for
the other subunits than for itself. In all of the other cases that do
not involve the ? isoform, a statistical distribution of XX?
XY?YY with a 1:2:1 ratio was formed with 50% heterodimer in
the mixture (Fig. 6). The statistical formation of heterodimers
between the ?, ?, and ? isoforms suggests that their binding
affinity to each other is similar, whereas the ? isoform shows a
strong preference for heterodimerization.
Although 14-3-3 proteins commonly form dimers, the mono-
mers are sometimes functional, depending on the target protein.
For example, monomeric ? is able to modulate the activity of a
potassium channel (46) but not Raf kinase (47). Although these
results were based on the use of a mutant ? form that is unable
to dimerize, our analysis shows that the native protein exists in
both monomeric and dimeric states and suggests that these
14-3-3 proteins may use the dimerization process to control their
The molecular mechanism for dimerization and the basis for
the preference for homo- and heterodimer formation can in part
be deduced from the panel of crystal structures available.
Recently, Gardino et al. (41) reviewed the structural determi-
nants for dimer formation, and thus only a brief description of
the main features is presented here. The dimerization interface
around the aperture of all human 14-3-3 isoforms involves a
conserved salt bridge between Arg-19 and Glu-92. Besides this
interaction, two residues (?: Asp-21 and Lys-85) are conserved
in all human isoforms except ? (equivalent ? residues being
Glu-22 and Met-88). In all of the homodimer structures, the side
chains at these positions do not interact. Only with ? as one
subunit of a heterodimer is an additional hydrogen-bonding
opportunity created, as illustrated schematically in Fig. 2, which
is likely the contributing factor for its preference to form
Binding of 14-3-3 Proteins to Interaction Partners. Generally, pro-
domain with an unstructured region that contains a phosphor-
ylated Ser?Thr residue within a mode 1 or 2 binding motif. In the
binding event, two processes likely occur: binding of the phos-
phorylated peptide to the conserved groove (?C, ?E, ?G, and
?I), which we define as the primary interaction and interaction
of the 14-3-3, which we define as secondary interaction. The
availability of three-dimensional structures for all human iso-
forms now allows us to rationalize this concept at a detailed
from the N to C terminus. An aperture exists at the central dimeric interface,
which is marked with a circle.
Overview of the dimeric 14-3-3 structure. Helices and loops involved
www.pnas.org?cgi?doi?10.1073?pnas.0605779103Yang et al.
to interact with a large but distinct number of phosphorylated
proteins. In the phosphopeptide-interacting motif, the orienta-
tion of the phosphate group is fixed once bound to the Arg–
Arg–Tyr triad; however, the phosphoserine C?OC? bond is free
to rotate. Thus, for an extended peptide that binds along the
groove, two orientations are possible, differing by a twofold
rotation around the C?OC? bond. When analyzing the ligand
complexes (?, ?, ?, ?, ?, and ?), it is apparent that there is
specificity in the relative orientation of the bound peptide. All
of the phosphorylated peptides bind with the N to C termini
being orientated similarly and interacting with three conserved
features. (i) The Asn-176 side chain has an orientation that is
fixed as a result of a hydrogen bond with the conserved Asp-127
as well as hydrogen bonds with an amide on the peptide
backbone of the ligand. This interaction occurs only in the
observed orientation that also allows the conserved Asn-227 to
make hydrogen bonds with the ligand backbone (Fig. 3A). (ii)
The extended phosphopeptide has most of its side chains point-
ing away from the 14-3-3 molecule, thus explaining the absolute
requirement for peptide direction as if it were reversed (by 180°)
steric clashes of the side chains would occur (Fig. 3B). (iii) A
conserved hydrophobic patch (Leu-175, Leu-219, and Ile-220)
within the 14-3-3 binding groove complements the hydrophobic
character of the peptide on the C-terminal side of the phospho-
serine (Fig. 3C).
In conclusion, general features of 14-3-3–phosphopeptide
interactions are that it relies on fixed positions of the binding
pocket side chains and orientation of the peptide. Interactions
are formed only with the main chain of the bound peptide and
its phosphate group. For example, the hydrogen bond from
Asn-176 to the backbone amide nitrogen in the ?2 main chain
atom position of the peptide would not be available with a
peptide in the reversed orientation. In contrast, interactions with
unphosphorylated peptides such as R18 and ExoS that have a
reversed orientation are dependent on sequence-specific inter-
actions with the peptide side chains.
Secondary Interaction. According to this hypothesis, once the
phosphopeptide has bound, the target protein would bind to the
remaining sections of the 14-3-3 protein. Determination of
the crystal structure of ? in complex with AANAT (36) permit-
ted the identification of the loop between helices ?H and ?I (HI
loop) as being critical for binding. This interaction, as well as
others potentially critical for binding, was recently further ana-
To complement the observations from the crystallographic
data, we conducted a further analysis by applying the optimal
docking area (ODA) methodology, which is based on atomic
desolvation parameters adjusted for protein–protein docking
(48). This method identifies continuous surface patches likely to
be involved in protein–protein interactions and was initially
validated by using a set of 66 unbound protein structures (48)
and, later in the prediction ‘‘competition’’ CAPRI (critical
assessment of predicted interactions), where it accurately pre-
dicted eight of nine protein–protein complexes (49)
The results from the ODA analysis are shown in Fig. 4, where
red patches represent likely protein–protein interaction inter-
faces. No low-desolvation patches were observed for any of the
isoforms in the conserved peptide binding groove or on the
opposite face of each monomer. Two major ODA sites occur
consistently for all 14-3-3 family members: site S1, located at the
observed dimerization interface, and site S2 in the solvent
accessible side of the helices ?H and ?I, located close to the
previously identified specificity region (37).
Based on the analysis of a few 14-3-3 interactions, the se-
quence variability within the HI loop was postulated to be of
critical importance for isoform?target specificity. The S1 patch
constitutes a recognition motif involved in the dimerization
interface. However, analysis of target structures revealed that
the S1 patch could be distinguished into two contributing
‘‘subsites.’’ We suggest that the first (S1a), is directly involved in
indicate specific interactions.
Schematic representation of the heterodimerization process involv-
N- to C-terminal orientation is the same in all other 14-3-3 structures with phosphopeptides. (B) Same view as in A except with the ? peptide binding groove
surface colored yellow. Reverse orientation of the same peptide (blue wire frame) about the phosphoserine C?–C? bond results in major clashes. (C) Same view
as in A, now with the character of the residues that make up the peptide binding groove color coded onto the surface as yellow (hydrophobic), red (negatively
charged), blue (positively charged), and gray (neutral).
The selective nature of the primary interaction site. (A) Close-up view with side chain interactions highlighted for Pep1 in the binding groove of ?. The
Yang et al.
November 14, 2006 ?
vol. 103 ?
no. 46 ?
the dimerization process, whereas the second site (S1b) contrib-
utes to target protein interactions. The S1b region has several
similarities to S2: (i) the residues responsible for the low
desolvation energy are conserved; (ii) the immediate vicinity of
the desolvation patch consists of flexible loops; (iii) the residues
in the loops are not well conserved; and (iv) there is a preference
for charged residues in the loops. Validating this approach, the
S1b and S2 patches are extensively involved in forming protein–
protein contacts within the crystals (Table 3, which is published
as supporting information on the PNAS web site).
We suggest that the target protein is attracted first by
general protein–protein interaction motifs (the desolvation
patches) followed by chain?loop rearrangements that contrib-
ute to the formation of specific contacts. In keeping with this
hypothesis, the CD and HI loops appear to be flexible: in all
crystal structures of the human 14-3-3 proteins, they appear
either as regions with high thermal motion or as unstructured
Binding Mechanism. The crystal structures of the first three 14-3-3
proteins (?, ?, and ?) adopted a similar conformation in both
apo- and ligand-bound forms (here denoted as the ‘‘closed
state’’). These initial data gave rise to the hypothesis that the
14-3-3 proteins may have a fairly rigid conformation and that the
interacting protein had to adapt to enable the interaction.
However, the structure determination of the apo-? isoform in
this study clearly shows that the peptide-binding site on the
14-3-3 protein allows flexible adaptation. In the apo-? structure,
one of the monomers was captured in the closed state confor-
mation, whereas the phosphopeptide binding site in the opposite
subunit had an open conformation following a ?20° rotation of
the ?G to ?I helices, producing a shallow and exposed groove
(Fig. 5A). Because ? does not contain any unusual sequence
motifs, it is possible that such flexibility is a feature of all
isoforms and the variation in size of the peptide binding groove
allows binding to peptides of diverse sequence and differing
secondary structures. Based on the structural data, another
suggested mechanism by which the 14-3-3 proteins can facilitate
binding of diverse target proteins is through conformational
heterogeneity to alter the angle between the two subunits.
Superimposition of one subunit for all of the closed state 14-3-3
dimers shows that there is a significant difference in relative
position of the partner subunit (Fig. 5B). This is particularly
evident for the ? (PDB ID 2C23; blue) and ? (PDB ID 2BTP;
To test whether the ? isoform would undergo a conforma-
tional change upon binding of a nonphosphorylated target, we
determined the structure of ? in complex with a peptide based
on the sequence of the P. aeruginosa ExoS. In this structure, the
14-3-3 subunits adopting a closed conformation. We obtained
this structure by soaking crystals of apo-? with the ExoS peptide.
The conformational change took place in the crystalline form
14-3-3 subunits are identical. The structure of this complex also
allowed us to further rationalize target specificity dependent on
the phosphorylation status of the interaction partner. Both ExoS
(this study) and R18 (17, 18) bind with their N to C termini in
the opposite direction compared with phosphorylated peptides.
In addition, when ExoS is bound to the 14-3-3 partner, it forms
a helical structure (Fig. 7, which is published as supporting
information on the PNAS web site), showing a secondary
structural element in the 14-3-3 peptide binding groove, as
predicted for the general mode of 14-3-3 binding by Liu et al.
(35), before the phosphorylated 14-3-3 binding sequence motifs
The orientation and structure of the ExoS helix are stabi-
lized by the interaction of ? Asn-175 with backbone carbonyl
(Asp-427) and amide (Ala-429) groups at the C terminus of the
ExoS fragment peptide. These interactions define the end of
the helix while the C terminus is in an extended conformation.
The interaction is stabilized by a hydrogen bond between Tyr
in the conserved 14-3-3 Arg–Arg–Tyr triad and the negatively
charged Asp-427 of ExoS, a hydrogen bond between the
conserved Lys-51 and Asp-424 of ExoS, and interactions
between hydrophobic side chains in the ExoS peptide and the
conserved hydrophobic patch on the surface of the 14-3-3
peptide binding groove. A superimposition of the closed state
representations for each of the seven human isoforms. All views look down
onto the dimerization interface (Left), central binding groove, and the ?G–?I
helices (Right). The desolvation energies are color coded from high (blue) to
the ? isoform, color coded from green (100%) to white. The variable CD and
HI loops are labeled V1 and V2, respectively, and the two conserved low-
energy desolvation sites are labeled S1 (S1a and S1b) and S2, respectively.
apo-? isoform looking down the peptide binding grooves, which are labeled
closed state 14-3-3 isoforms using only one monomer as the reference, with ?
shown in blue and ? in green. The other 14-3-3 monomers, which have
intermediate positions, are colored transparent gray.
Dynamic nature of the 14-3-3 dimers. (A) Crystal structure of the
www.pnas.org?cgi?doi?10.1073?pnas.0605779103 Yang et al.
of apo-? and the closed state of ExoS-? showed that the
ExoS-? interface was rotated ?4°, displaying further flexibility
most likely incurred by the specific binding partner, but we
cannot exclude that this is due to crystal packing effects.
However, the helical conformation of the ExoS peptide ap-
pears to push the 14-3-3 ?H and ?I helices backwards, which
results in the 4° rotation difference.
To analyze any change in conformation on peptide binding we
measured the radius of gyration (Rg) of ? and ? with and without
the phosphoserine peptide Pep2 by small angle x-ray scattering.
The observed changes in Rgupon peptide binding were from
30.7 ? 0.3 Å to 30.2 ? 0.3 Å and 31.3 ? 0.2 Å to 31.1 ? 0.2 Å
for ? and ?, respectively. A change from a fully open to a closed
conformation as indicated by the apo-? and peptide-bound ?
structures would correspond to a reduction in Rgof 7%. The
1–2% reduction observed is smaller, although still significant,
and consistent with a dynamic 14-3-3 dimer in solution. The apo
measurements represent a conformational average, with Rgin
the presence of Pep2 only slightly smaller than this average.
Comparing the experimental scattering data with the crystal
structures shows that the peptide-bound closed conformations
also provide acceptable fits to the scattering curves of the
apo-forms (Fig. 8, which is published as supporting information
on the PNAS web site).
Based on the results from this structural comparison of all
14-3-3 members, we suggest that conformational flexibility in
both the peptide binding site and in the dimer interface
facilitates binding of 14-3-3 proteins to diverse peptides and
proteins of varying size and sequence. Specifically, we propose
that the initial interaction involves conformational changes of
the three C-terminal helices (?G to ?I). Through these
changes, the 14-3-3 protein could adapt to the type of bound
peptide whether it is phosphorylated and extended or non-
phosphorylated and helical. The flexibility at the dimerization
interface provides a mechanism for the 14-3-3 proteins to bind
proteins of varying size. This versatility is most likely enhanced
by the presence of desolvation sites (S1b and S2) providing
nonspecific protein–protein interaction motifs, with specificity
features primarily from the variable V1 and V2 regions (see
Fig. 4). Further specificity in 14-3-3 regulation of intracellular
signaling is most likely provided by its tissue (50, 51), temporal
distribution (52, 53), and subcellular localization (41, 54),
allowing this small protein family to be a central and specific
regulator through its ability to adapt its conformation to
interact with a variety of binding partners.
Materials and Methods
Protein Expression and Purification. DNA for ?, ?, ?, ?, and ? was
PCR amplified from clones in the Mammalian Gene Collection
(I.M.A.G.E. Consortium Clone IDs 3051079, 2900956, 3915246,
3543571, and 6164592, respectively) and inserted into vectors
protease tag cleavage site (Table 4, which is published as
supporting information on the PNAS web site). The resulting
plasmids were transformed into E. coli BL21 (DE3) cells for
expression. Cultures were grown in Terrific Broth media at 37°C
until an OD600of 0.6 was reached. The temperature was then
reduced to 25°C for 1 h before induction by addition of 1 mM
isopropyl-?-D-thiogalactopyranoside. Protein expression was al-
The cells were resuspended in 20–50 mM Tris?HCl (pH 8.0),
200–500 mM NaCl, 5% glycerol, 10 mM imidazole, 0.5 mM
TCEP, and Complete EDTA-free protease inhibitor mixture
(Roche Applied Science, Burgess Hill, U.K.). The cells were
lysed by high-pressure homogenization, and the insoluble debris
was removed by centrifugation.
then washed with 50 mM Tris?HCl (pH 8.0), 150–500 mM NaCl,
5% glycerol, 25 mM imidazole (pH 8.0), and 0.5 mM TCEP.
Proteins were eluted with 50 mM Hepes (pH 8.0), 150–500 mM
NaCl, 5% glycerol, 250 mM imidazole (pH 8.0), and 0.5 mM
TCEP. The eluted fractions were purified by gel filtration in 50
mM Hepes (pH 8.0), 100–500 mM NaCl, and 0.5 mM TCEP.
Removal of the hexahistidine tag was accomplished by using
TEV protease at 4°C overnight followed by passing the solution
over Ni2?resin. Purified proteins were concentrated and stored
at ?80°C before crystallization.
Peptides Used in Crystallization. All peptides were synthesized by
Thermo Electron Corporation (Ulm, Germany), dissolved to a
concentration of 40 mM in water, and stored at ?20°C. The
sequences of phosphoserine peptide I (Pep I) and II (Pep II) are
RRQRpSAP and RSIpSLP, respectively, where pS represents a
phosphorylated serine. The sequence of the ExoS fragment is
GLLDALDLASK. The peptides and 14-3-3 proteins were mixed
to a molar ratio of 3:1 peptide?protein for crystallization.
Crystallization and Data Collection. Crystals were grown at 20°C by
using the sitting drop method (Table 1). All crystals were
mounted in loops and cryo-cooled before data collection. X-ray
data were collected at 100 K. Diffraction data were collected to
2.5 and 2.8 Å resolution on a Rigaku?MSC FR-E rotating anode
generator equipped with an R-AXIS HTC image plate for
14-3-3? and ?, respectively. The data sets for ? and ? were
collected to 1.6- and 2.5-Å resolution, respectively, at the Swiss
MOSFLM (55) and the CCP4 suite (56). Structures were solved
by molecular replacement using PHASER (57), and crystallo-
graphic models were rebuilt by using O (58) or COOT (59).
Refinements were performed by using Refmac5 (60) (Table 2).
Mass Spectrometry of Intact Complexes. Mass spectra were re-
corded by using either an oTOF (Waters, Manchester, U.K.,
LCT) or a tandem-mass spectrometer (Waters Q-Tof 2)
modified for high mass operation (61). Both instruments were
fitted with a standard offline nanoelectrospray source. Sample
solutions were sprayed from borosilicate glass capillaries with
a tapered tip that was cut under a microscope to an inner
diameter of ?2–5 ?m. Approximately 2 ?l of protein solution
(10–30 ?M) was loaded per capillary. Protein samples were
buffer exchanged into 20 mM aqueous ammonium acetate (pH
7.0) using gel-filtration spin columns (microbiospin; Bio-Rad,
A low backing pressure of nitrogen gas was used to initiate and
maintain flow through the capillary, and a nitrogen cone gas
flow was used to aid desolvation of the spray. The conditions
within the mass spectrometer were adjusted to preserve nonco-
valent interactions (61). All data were acquired in positive ion
mode and processed by using MassLynx software (Waters).
Experimental conditions for the monomer?dimer study were:
Waters Q-Tof 2; capillary voltage 1.4 kV, cone 100–120 V,
extractor 0–2 V, source temperature 25°C, drying gas rate 150
liters?h and Pirani pressure 5 ? 10?3mbar. Experimental
conditions for the heterodimerization study were: Waters LCT
oTOF mass spectrometer, capillary voltage 1.4 kV, cone 120 V,
extractor 0 V, source temperature 25°C, drying gas rate 150
liters?h, and Pirani pressure 6 ? 10?1mbar.
Small Angle X-Ray Scattering. Data were collected at station 2.1 of
the Synchrotron Radiation Source (Daresbury, U.K.) of ? and ?
isoforms (at concentrations between 2 and 3 mg?ml) and buffer
in the momentum transfer interval 0.02 Å?1? q ? 0.19 Å?1
where q ? 4?sin???, (2? is the scattering angle and ? is the x-ray
wavelength 1.5 Å) according to published procedures (62).
Measurements with peptides were performed at a 9-fold excess
of peptide. The calculation of the Rgas well as scattering pattern
Yang et al.
November 14, 2006 ?
vol. 103 ?
no. 46 ?
simulations based on crystal structure information were carried Download full-text
out with the program CRYSOL (63).
ODA Calculations. The default ODA algorithm as described by
Fernandez-Recio et al. (48) and implemented in the program ICM
version 3.4-3 (www.molsoft.com) was used to calculate the optimal
docking areas for all members of the human 14-3-3 family.
We thank members of the Structural Genomics Consortium (SGC) for
assistance with plasmid preparation and diffraction data collection. The
SGC is a registered charity (no. 1097737) funded by the Wellcome Trust,
GlaxoSmithKline, Genome Canada, the Canadian Institutes of Health
Research, the Ontario Innovation Trust, the Ontario Research and
Development Challenge Fund, the Canadian Foundation for Innovation,
VINNOVA, The Knut and Alice Wallenberg Foundation, The Swedish
Foundation for Strategic Research, and Karolinska Institutet.
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www.pnas.org?cgi?doi?10.1073?pnas.0605779103Yang et al.