The Rockefeller University Press $30.00
J. Cell Biol. Vol. 186 No. 4 589–600
Correspondence to M. Amin Arnaout: email@example.com
Parts of this work were presented at the Gordon Research Conference on Fibronectin,
Integrins, and Related Molecules in Ventura, CA on 1 February 2009.
Abbreviations used in this paper: TD, -tail domain; FLIM, fluorescent lifetime
imaging microscopy; FN, fibronectin; FRET, fluorescence resonance energy
transfer; LIMBS, ligand-associated metal-binding site; MIDAS, metal ion–dependent
adhesion site; NMR, nuclear magnetic resonance; PSI, plexin-semaphorin-integrin;
TM, transmembrane; WT, wild type.
Integrins are / heterodimeric type I membrane receptors that
mediate divalent cation–dependent interactions with components
of the extracellular environment (cells and soluble and matrix
proteins) leading to changes in cell shape, movement, growth,
differentiation, and survival (Hynes, 2002). Determination of the
crystal structure of the ectodomain of V3 (∆TM-V3) in
the absence and presence of a prototypical RGD ligand revealed
the modular nature of integrins and clarified the basis of its
divalent cation–mediated interaction with extracellular ligands
(Xiong et al., 2001, 2002). The 12 extracellular integrin domains
are assembled into a head segment mounted on top of two leg
segments. The integrin head comprises a seven-bladed -propeller
domain from V and a vWFA (A or I-like) domain from 3 and
contains the RGD-binding site. The V leg is composed of an
upper Ig-like thigh domain and a lower calf module consisting of
two large -sandwich domains, Calf-1 and -2. The 3 leg con-
sists of an upper leg segment comprising a plexin-semaphorin-
integrin (PSI), an Ig-like hybrid, and integrin IE1 (EGF-like 1)
domain, followed by a lower leg segment made up of three IE do-
mains (IE2–4) and a novel -tail domain (TD). ∆TM-V3 is
bent in half at two knee-like joints, the - and -genu, the former
between the thigh and Calf-2 domains and the latter predicted
between IE1 and -2 (Xiong et al., 2001), such that the head and
upper leg segments of the heterodimer contact the lower leg seg-
ments of the same molecule.
Binding of the F3 subdomain of the talin head to the cyto-
plasmic tail of the subunit breaks an / salt bridge (that nor-
mally stabilizes the inactive state; Wegener et al., 2007), triggering
a conformational wave that travels through the / transmem-
brane (TM) and lower leg domains to switch the conformationally
sensitive A domain to a high affinity state, which is a process
brane stretches of the V and 3 subunits. 1TM-V3 is
more compact and less active in solution when compared
with ∆TM-V3, which lacks the short C-terminal stretches.
The structure reveals a bent conformation and defines the
– interface between IE2 (EGF-like 2) and the thigh
domains. Modifying this interface by site-directed muta-
genesis leads to robust integrin activation. Fluorescent life-
e determined the crystal structure of 1TM-V3,
which represents the complete unconstrained
ectodomain plus short C-terminal transmem-
time imaging microscopy of inactive full-length V3 on
live cells yields a donor–membrane acceptor distance,
which is consistent with the bent conformation and does
not change in the activated integrin. These data are the
first direct demonstration of conformational coupling of the
integrin leg and head domains, identify the IE2–thigh inter-
face as a critical steric barrier in integrin activation, and
suggest that inside-out activation in intact cells may in-
volve conformational changes other than the postulated
switch to a genu-linear state.
Crystal structure of the complete integrin V3
ectodomain plus an / transmembrane fragment
Jian-Ping Xiong,1,2 Bhuvaneshwari Mahalingham,1,2 Jose Luis Alonso,1,2 Laura Ann Borrelli,3 Xianliang Rui,1,2
Saurabh Anand,1,2 Bradley T. Hyman,3 Thomas Rysiok,4 Dirk Müller-Pompalla,5 Simon L. Goodman,6
and M. Amin Arnaout1,2
1Program in Leukocyte Biology and Inflammation and 2Program in Structural Biology, Nephrology Division, Department of Medicine and 3Institute for Neurodegenerative
Disease, Massachusetts General Hospital, Harvard Medical School, Charlestown, MA 02129
4Biologicals: Protein and Cell Science, 5Biologicals: Protein Purification, and 6Therapeutic Area Oncology: Biochemistry and Cellular Pharmacology, Merck-Serono
Research, 64293 Darmstadt, Germany
© 2009 Xiong et al. This article is distributed under the terms of an Attribution–
Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publica-
tion date (see http://www.jcb.org/misc/terms.shtml). After six months it is available under a
Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license,
as described at http://creativecommons.org/licenses/by-nc-sa/3.0/).
T H E J O U R N A L O F C E L L B I O L O G Y
JCB • VOLUME 186 • NUMBER 4 • 2009 590
complete TM domains (V3-cTM). Functional experiments
showed that modifying the defined IE2–thigh interface leads to
constitutive activation. Fluorescent lifetime imaging microscopy
(FLIM) using ligated Fab fragment of the anti–V propeller mAb
17E6 (Mitjans et al. 1995) as fluorescence donor and the plasma
membrane dye FM4-64 FX as acceptor yielded donor–membrane
acceptor separation distances consistent with the V3-cTM
structure model, which did not vary between the inactive and
active states of the full-length integrin in live cells. These data
demonstrate a conformational link between the lower leg and
head domains, define the crystal structure of a near native com-
plete ectodomain, identify a critical steric barrier to activation at
the IE2–thigh interface, and suggest that the genu-linear state is
not an obligate feature of inside-out activation.
Expression and biophysical characterization
Published structures of ∆TM-V3 (Xiong et al., 2001) and
∆TM-IIb3 (Zhu et al., 2008) lacked the last seven extracellu-
lar residues from the subunit and P691-D692 from the 3
subunit (Fig. 1 A). We expressed in insect cells an V3 ecto-
domain, 1TM-V3, encoding all of the missing extracellular
residues in addition to a four-residue TM extension of each sub-
unit. 1TM-V3 was secreted into the culture supernatant of
virus-infected High-Five insect cells (Fig. 1 B) and was purified
by affinity chromatography (Fig. 1 C).
The isocratic elution profiles of purified 1TM-V3 and
∆TM-V3 were compared on molecular sieve chromatography
columns in neutral buffer containing the physiological divalent
cations 1 mM Ca2+ + 1 mM Mg2+ (Ca2+/Mg2+) or 1 mM of the
activating cation Mn2. The apparent hydrodynamic radius (Stokes
called inside-out activation (Hynes, 2002). Direct evidence for
the conformational coupling of the lower leg and head domains
is indirect. And how inside-out activation converts the ecto-
domain to the high affinity state is still ill defined. Modeling the
lower integrin legs of the ectodomain perpendicular to the plasma
membrane placed the ligand-binding head near the lipid bilayer,
seemingly blocking access to macromolecular ligands (for re-
view see Arnaout et al., 2005). A switchblade model (Takagi
et al., 2002) proposed that disruption of the / cytoplasmic–TM
domain interfaces converts the bent ectodomain to a genu-linear
conformation, providing ligand access and allowing A to switch
to high affinity. An alternate model, the deadbolt, proposed that
lifting the constraints exerted on A by the lower leg TD can
induce activation without linearity (Xiong et al., 2003). Com-
plete structural information on the remaining / residues in the
lower leg segments and the conformationally sensitive IE1/IE2
region are missing or incomplete in current structures of the
integrin ectodomains (Xiong et al., 2001; Zhu et al., 2008), and
knowledge of the distance separating the ligand-binding integrin
head from the plasma membrane in the inactive and active states
is crucial to elucidate the conformational transitions and energetics
of affinity switching in integrins.
In this communication, we engineered a new construct,
1TM-V3, encoding the complete sequence of the V3
ectodomain plus the first four TM residues of each subunit.
1TM-V3 was water soluble at neutral pH and was monomeric,
which allowed characterization of its biophysical and functional
properties and a determination of its crystal structure. 1TM-V3
was hydrodynamically more compact and less active than ∆TM-
V3 in solution. Its crystal structure revealed a bent conforma-
tion and defined the complete structure of the ectodomain, including
that of the IE1/IE2 region and TM extensions, permitting us to
build a structure model comprising the bent ectodomain plus the
Figure 1. Construction, generation, and purification
of 1TM-V3. (A) Primary sequence of the exofacial
residues and TM domain (underlined) of human V
and 3. The crystal structure of ∆TM-V3 (Xiong
et al., 2001) ends at Q956 in V and G690 in 3
(closed arrows); ∆TM-IIb3 terminates at the cor-
responding IIb residue A958 and at G690 in 3
(Zhu et al., 2008). Open arrows point to the C-
terminal ends of 1TM-V3 (I967 and V696). G989
in the GFF motif is labeled. (B) Western blot after
fractionation on 4–12% gradient SDS gels of LM609
mAb immunoprecipitates from insect cell-free super-
natant under nonreducing (NR) and reducing (R)
conditions. ∆TM and 1TM indicate ∆TM-V3 and
1TM-V3, respectively. Lanes 1 and 4 show mo-
lecular mass (mm) markers. The 1TM-V and 1TM-
3 subunits migrated with those from ∆TM-V3,
as expected for the small (<1%) increase in mass
caused by the additional C-terminal sequences.
(C) Coomassie-stained purified 1TM-V3 after frac-
tionation on 4–12% gradient SDS gels under non-
reducing conditions. Molecular mass markers in lane
1 are indicated.
591CRYSTAL STRUCTURE 1TM-V3 • Xiong et al.
V3 in its active or RGD-bound states but not in its inactive
state (Honda et al., 1995; Faccio et al., 2002).
Crystal structure of 1TM-V3
A new native dataset derived from one 1TM-V3 crystal let us
determine the structure of the 1TM-V3, including the TM ex-
tensions, at 2.9-Å resolution (Fig. 4). The structure was super-
posable on the ∆TM-V3 structure (Xiong et al., 2001, 2004)
except for those regions newly expressed, the C-terminal exofacial,
and the TM residues (Fig. 4 A) and now resolved the IE1 and -2
domains (Fig. 4, B–D). The nearly parallel exofacial extensions
I955-P959 (in V) and P688-G690 (in 3) remain close through
A958 and G690 (C–C = 6.2 Å) and then diverge at an extended
Pro-rich loop structure in V (P959APMPVP963) before the first
putative TM residue V964. Structure search of this Pro-rich
sequence yielded such loops in two contexts (Protein Data Bank
numbers 2w55 and 2q0s), suggesting that this extended structure is
not a crystal artifact but also exists in the native integrin. The first
TM residues V964 (in V) and I963 (in 3) occupy similar posi-
tions in the structure, but the TM extensions do not assume the
-helical turns found in the nuclear magnetic resonance (NMR)
structure of the / TM domains (Lau et al., 2009) and do not
interact as the result of unfavorable crystal contacts.
The three tandem integrin domains IE2–4 are related by
an approximately twofold screw axis symmetry and form an ex-
tended module (Fig. 4 C). IE1 is roughly antiparallel to IE2, as
a result of the bend at the -genu (Fig. 4 C). Each IE domain has
three disulphides (a–c) having the same connectivity aN-aC, bN-bC,
cN-cC as that found in other EGF domains (Fig. 4 E; Wouters
et al., 2005). IE1 lacks disulphide a (aN-aC), thus potentially avoid-
ing a clash with the adjacent PSI domain. A characteristic forth
N-terminal intradomain disulphide d (dN-dC) occupies an analo-
gous site to the Ca coordination site found in the Ca-binding
subset of EGF domains (Wouters et al., 2005). IE1–4 each contain
radius [Rs]) of 1TM-V3 was calculated by reference to the
elution position of standard proteins (Fig. 2). Rs of 1TM-V3
changed from 55 ± 0.41 Å (mean ± SD; n = 4) in Ca2+/Mg2+ to 58 ±
0.3 Å in Mn2+. The Rs values for ∆TM-V3 in Ca2+/Mg2+
and Mn2+ were 57 ± 1.1 Å and 60 ± 1.1 Å, respectively. Thus,
the mean Rs value for 1TM-V3 in Mn2+ differed little from
that of ∆TM-V3 in Ca2+/Mg2+. The small amounts of oligo-
mers in the ∆TM-V3 preparations in the presence of Mn2+
were largely absent in 1TM-V3, and the peak width at half
height, which is a measure of molecular heterogeneity, was nar-
rower for 1TM than for ∆TM. These data suggest that the short
C-terminal / extensions introduced in 1TM result in a more
compact and homogenous molecule in solution.
Binding of soluble 1TM-V3 to
physiological ligands and to the
activation-sensitive mAb AP5
We performed dose–response curves to quantify the binding
of increasing concentrations of 1TM-V3 to immobilized
FN7–10 in Mn2+ and in Ca2+/Mg2+ (Fig. 3, A and B). Half-maximal
binding was achieved at 0.1 µg/ml and 6.0 µg/ml, respectively
(n = 2). For soluble TM-V3, half-maximal binding was
achieved at 0.15 µg/ml and 1.6 µg/ml, respectively. No binding
took place to uncoated wells or fibronectin (FN)-coated wells in
the presence of cilengitide (unpublished data). Soluble 1TM-V3
formed a stable complex with either FN7–10 (Fig. 3 C) or na-
tive full-length FN (Fig. 3 D and Table S1) in solution in Mn2+-
containing buffer (Fig. 3 D and Table S1). Also consistent with
the solid phase binding assays, soluble 1TM-V3 did not form
a stable complex with either FN7–10 or full-length FN in Ca2+/
Mg2+ buffer (Fig. 3 D, inset; Table S1; and not depicted). How-
ever, soluble 1TM-V3 formed a stable complex in 2 mM
of CaCl2 (or Ca2+/Mg2+) buffer with the Fab fragment of AP5
(Fig. 3 E), a ligand-inducible binding site mAb which binds
Figure 2. Hydrodynamic analyses of 1TM-
V3 and ∆TM-V3 by molecular sieve
chromatography. A representative experiment
(one of four) is shown. Purified ∆TM-V3 or
1TM-V3 in TBS, pH 7.4, containing 1 mM
MgCl2 + 1 mM CaCl2 or 0.2 mM MnCl2 was
analyzed. Each integrin was applied onto a
precalibrated Superdex S-200 GL column, and
Stokes radii were derived as described previously
(Adair et al., 2005). Values next to the major
peaks indicate peak elution volumes (in milli-
liters). A dashed line was added to emphasize
the peak shifts. mAU, milli–absorbance unit.
JCB • VOLUME 186 • NUMBER 4 • 2009 592
three strands (A–C), with the first two antiparallel strands
(A and B) forming a major sheet found in classical EGF domains.
Strand C is hydrogen bonded to strand D, which is contributed in
part by the consecutive domain, with strands C and D forming a
minor sheet–like conformation. A short strand D caps the IE4
domain, which is stabilized by the C601-C604 disulphide linker
that precedes the start of the TD. In IE1–4, strand D has a charac-
teristic bulge at a conserved E/N residue (E472, E522, N559 and
E599, respectively) to allow accommodation of disulphide c; its
cross-strand partner in strand C is an invariant Gly (G468, G518,
G555, G595, respectively), which is found typically in class II EGF
domains (Wouters et al., 2005) and some laminin-type epidermal
growth factor–like domains (Stetefeld et al., 1996).
IE2 has two characteristically long loops, c (between cN and
cC) and d (between dN and aN; Fig. 4 E), with the latter housing the
-genu (Fig. 4 B). The -genu is clearly visible in the electron
density map despite lack of crystal contacts from symmetry-
related molecules, which is stabilized by hydrogen bonds to the
major sheet of IE2. The two long loops and strand A of IE2 face
the bottom of the thigh domain, making mainly electrostatic con-
tacts with its CC and EF loops (Fig. 4 D). The minor strand of
IE2 faces the N-terminal segment of PSI and forms ionic and van
der Waals contacts involving mainly T7-R8 of the PSI domain.
Extending the 1TM structure into the / TM fragment,
which overlaps with structured residues in the NMR structure
of IIb3 TM domains (Lau et al., 2009), allowed us to build
a structure model of the inactive ectodomain plus the com-
plete TM domains (Fig. 4, F–H). In this model, the ligand-
binding site is accessible to macromolecular ligands, and the
extracellular membrane proximal segment is structured. By
comparison, in the recent structure model of the IIb3
Figure 3. Binding of 1TM-V3 to physiological ligands and to the
Fab fragment of the activation-sensitive mAb AP5. (A and B) Receptor-
binding assay to immobilized ligand. Dose–response curves showing
binding increasing concentrations of ∆TM- or 1TM-V3 to wells coated with
FN7–10 in the presence of 1 mM Mn2+ (A) or 1 mM Ca2+ + 1 mM Mg2+
(B). The data shown are from a representative experiment, one of two con-
ducted. Each point was taken at the end of the assay, and the amount of
integrin present was measured by quantitative ELISA (see Materials and
methods for details). No binding took place to uncoated wells run in parallel
(not depicted). (C) Molecular sieve chromatogram showing the stable bind-
ing of 1TM-V3 to FN7–10 in solution containing 0.2 mM MnCl2. Peak elu-
tion volumes for the 1TM-V3–FN complex and FN7–10 are 10.82 ml and
14.75 ml, respectively. (inset) Coomassie-stained SDS-PAGE under non-
reducing conditions. Lane 1, molecular mass markers (in kilodaltons); lane 2,
1TM-V3; lane 3, FN7–10; lane 4, blank; lane 5, FN7–10 from the faster
peak; lane 6, blank; lane 7, 1TM-V3–FN7–10 complex in the slower
peak. An 1:1 integrin/FN molar ratio was calculated from the scanned
gel (see Materials and methods), which is in agreement with previous results
(Adair et al., 2005). (D) Molecular sieve chromatography of 1TM-V3
with intact plasma FN in the presence of 0.2 mM Mn2+ or 1 mM Ca2+ + 1 mM
Mg2+. In Mn2+-containing buffers, 1TM (Kav of 0.229; 11.58 ml) forms a
complex with intact FN, which elutes at a Kav of 0.66 (9.0 ml). FN alone
elutes as a discrete peak at a Kav of 0.127 (10.0 ml). In Ca2+/Mg2+ buf-
fers, 1TM runs as a more compact molecule, and coelution with FN reveals
no indication of complex formation (inset). (E) Molecular sieve chromato-
gram showing complex formation of 1TM-V3 with the Fab fragment of
mAb AP5 in TBS containing 2 mM CaCl2. The dashed line shows the peak
elution volume of purified 1TM-V3 alone run on the same column and in
the same buffer. (inset) Coomassie-stained 12% SDS-PAGE under nonreduc-
ing conditions. Lane 1, molecular mass markers (in kilodaltons); lane 2,
blank; lanes 3 and 5, purified 1TM-V3 and AP5 Fab, respectively, before
mixing; lane 4, 1TM-V3–AP5 Fab complex from the slower peak. A 1:1
integrin/AP5 Fab molar ratio was calculated from the scanned gel (see
Materials and methods). mAU, milli–absorbance unit.
593CRYSTAL STRUCTURE 1TM-V3 • Xiong et al.
Functional effects on activity of full-
length V3 after mutation of features
structurally defined in the 1TM structure
We assessed the effects on cellular V3 caused by modifying
the newly defined IE2–thigh interface through a -genu dele-
tion (-genu), altering the TD–hybrid–IE3 interfaces by
breaking two salt bridges (R404A + R633A), and changing the
A–TD–hybrid interfaces by deleting the TD CD loop plus
ectodomain plus TM domains (Zhu et al., 2009), this region is
not structurally defined and assumed to be very flexible in the
inactive integrin model. However, inserting a flexible linker
into the extracellular membrane proximal Pro-rich sequence
of the 2 integrin subunit CD11b leads to a constitutively
active integrin (Kamata et al., 2005), suggesting that some ri-
gidity in this region is essential to maintain the integrin in its
Figure 4. New features of 1TM-V3 crystal structure and hypothetical model of V3 plus the complete TM domains (V3-cTM). (A) Density map (in
blue; contoured at 1.0 ) of the exofacial and TM extensions. The superposed main chains of 1TM-V3 and ∆TM-V3 (only the lower parts of Calf-2 and
TD shown from each; Xiong et al., 2004) are in light and dark gray, respectively, except for the new exofacial and TM extensions of 1TM-V3, which are
shown in red, and the last residues in the ∆TM-V3 structure (Q956 and G690), which are shown in green. (B) Density map (in blue; contoured at 1.0 )
and main chain tracing of the -genu (in red) in IE2. The orange sphere in this panel and in C represents the metal ion at the -genu. (C) Ribbon diagram
of the IE1–IE2 region in a similar orientation to that shown in B. Main chain tracings of IE1, IE2, and -genu are in green, blue, and red, respectively.
(D) Ribbon diagram showing electrostatic interactions at the IE2–thigh interface. Residues (shown in ball and stick representation) forming a salt bridge, main
chain, or side chain H-bonds are labeled. Thigh and IE2 are labeled in gray and blue, respectively, with the -genu in IE2 shown in red. Oxygen, nitrogen,
and carbon atoms are in red, blue and green, respectively. Hydrogen bonds and salt bridges (distance cutoff, 3.5 Å) are represented with red dotted
lines. (E) Structure alignment and Cys pairing of IE domains. The sequence housing the -genu in IE2 is in blue. The secondary structure elements (strands
are underlined, and a helix is represented by a cylinder) are shown. The atomic coordinates are deposited in the Protein Data Bank (3IJE). (F–H) Structure
model of V3 ectodomain plus the complete TM domains (V3-cTM). The model is built by releasing the C termini in the 1TM-V3 structure from their
respective crystal contacts, such that extracellular P691 and P963 initiate the respective 3 and V TM helices, which is consistent with the known propen-
sity of prolines to strongly stabilize -helical conformations (Senes et al., 2004). (F and G) The resulting movements included a 2.9-Å inward movement
of P691 of 3 (F) and a rotation of V’s Pro-rich loop at Q956, such that P963 initiates the V TM helix (modeled after IIb’s TM NMR structure; G; Lau
et al., 2009). The structure model was energy minimized with Modeller (Fiser and Sali, 2003), and the V3 TM side chains were optimized and repacked
using Rosetta (Rohl et al., 2004). The ribbon diagrams in F and G were generated using PyMOL (DeLano Scientific LLC). (H) A ribbon diagram, which was
generated using Chimera, of the V3-cTM model showing the orientation of the ectodomain relative to the TM domains. The model predicts that the TM
domains are at an 30° angle relative to the long axis of Calf-2, with ADMIDAS (adjacent to MIDAS) metal ion (green sphere) at an 45-Å distance from
the plane parallel to the hypothetical membrane drawn at the C of 3s Pro691. The -genu and propeller metal ions are in orange.
JCB • VOLUME 186 • NUMBER 4 • 2009 594
Mn2+-activated WT V3 did not affect binding of fluorescently
labeled soluble monomeric FN10, either as WT or as a high affin-
ity form (Fig. 6 A). The RGD ligand cilengitide also triggered an
increase in hydrodynamic radius of the 17E6–1TM-V3 com-
plex (Fig. 6 B), indicating that 17E6 Fab did not freeze either
membrane-bound or soluble V3 in an inactive state. The
plasma membrane was labeled with FM4-64 FX (FM) as accep-
tor. This dye preferentially inserts into the outer leaflet of the
membrane of live cells in 1–5 min at 4°C, fluoresces brightly, and
can be rapidly fixed with ice-cold paraformaldehyde, eliminating
diffusion on the time scale of FLIM measurements. The standard
equation to calculate FRET efficiency applies to populations of
fluorophores in such circumstances and is frequently used in bio-
logical systems to report mean donor–acceptor distances.
Alexa Fluor 488 fluorescence lifetime was first measured
in inactive WT V3–expressing K562 cells stained with Alexa
Fluor 488–Fab in 1 mM of CaCl2-containing buffer. A lifetime
of 2,306 ± 52 ps (mean ± SD) was determined in the absence of
acceptor (Fig. 6, C and D). When the Alexa Fluor 488–Fab-
labeled K562 cells were further labeled with the FM membrane
dye, Alexa Fluor 488 lifetime decreased to 2,056 ± 118 ps (P <
0.0001), corresponding to a FRET efficiency of 10.8% and a
mean donor–acceptor separation distance (r) of 88 Å. The re-
spective values for the Mn2+-activated V3 were 2,322 ± 43 ps
in the absence of acceptor and 2,087 ± 118 ps (P < 0.0001) after
addition of the FM dye, corresponding to a FRET efficiency of
10.1% and a mean donor–acceptor separation distance (r) of 89 Å.
This distance is consistent with the hypothetical structure model
depicted in Fig. 4 H, which predicts an 90-Å distance from
the centroid of the integrin-bound 17E6 Fab to the plane of the
membrane parallel drawn at the C of 3’s P691 residue. Mean
donor–acceptor separation distances of 78 ± 9 Å (mean ± SD)
and 83 ± 3 Å were obtained with unliganded V3 and with
high affinity FN10-bound V3, respectively, each in 1 mM of
Mn2+-containing buffer. FLIM measurements conducted on
transiently transfected HEK 293T cells expressing WT, G989FF/
GAA, or -genu V3 in 1 mM of Mn2+-containing buffer
an R633A substitution (CD + R633A). The activating G989FF/
GAA mutation (Zhu et al., 2007) served as a positive control.
None of the mutations impaired expression or heterodimer for-
mation compared with wild type (WT), as judged by reactivity
with the 3-specific AP3 mAb and the heterodimer-specific
LM609 mAb (unpublished data). Mn2+ increased binding of the
WT receptor to soluble Alexa Fluor 488–labeled FN9–10 from
7% in Ca2+/Mg2+ buffer to 48% of the AP3-positive cell popula-
tion in Mn2+. The -genu mutation induced constitutive acti-
vation that was significantly more robust than the G989FF/GAA
mutant (Fig. 5), with a more modest activation induced by the
R404A + R633A mutation. The activating effect of the CD +
R633A mutation was small but significant (P < 0.005; Fig. 5).
FLIM analysis of V3 in live cells
We used FLIM to assess the orientation of the V3 ectodomain
relative to the plasma membrane in K562 cells stably expressing
WT V3. For lifetime calculations, FLIM collects only photons
emitted from the donor fluorophores, thus avoiding the problem
of mis-excitation of the acceptor and the analogous problem
of spectral break through bleeding of the donor signal into the
acceptor spectral window, which are common concerns in spec-
tral fluorescence resonance energy transfer (FRET) applications
(Chigaev et al., 2001; Kim et al., 2003; Coutinho et al., 2007).
Importantly, FLIM allows picosecond measurements and is inde-
pendent of concentration of the fluorophores.
As fluorescence energy transfer between a donor–acceptor
pair described by the Förster equation depends both on the dis-
tance and the relative orientation of donor and acceptor (Jares-
Erijman and Jovin, 2003; Giepmans et al., 2006), it is critical that
a structurally defined probe be used as donor. Therefore, we used
the Fab fragment of mAb 17E6 whose low resolution epitope
mapping (Mould et al., 2000) corresponds to its crystal structure
bound to the -propeller of ∆TM-V3, which we have deter-
mined (17E6 binds at the top of the propeller contacting the DA
loop between blades 2 and 3 and the CD loop in blade 3; unpub-
lished data). Binding of unlabeled 17E6 Fab to the cell surface
Figure 5. Effect of mutations based on the TM-V3
structure on integrin activation. The histogram (mean ±
SD; n = 3) shows binding of Alexa Fluor 488–FN9–10 to
WT and mutant surface-expressed V3. The percentage
of FN9–10-bound cells was expressed as a percentage of
AP3-bound cells. Binding of WT and each of the mutants
in Ca2+/Mg2+ was then expressed as a percentage of that
obtained for the Mn2+-activated WT integrin, with the lat-
ter set at 100.
595CRYSTAL STRUCTURE 1TM-V3 • Xiong et al.
TM domains, (c) functional analyses using the 1TM-V3 crys-
tal structure identified a critical role for the newly defined
IE2–thigh interface in integrin activation, and (d) FLIM of WT
V3 in live cells revealed that the apparent distance between
the integrin head and the plasma membrane changes little in
inactive, Mn2+-activated, constitutively active, or FN10-bound
We found that 1TM-V3 is recognized in solution by the
activation-sensitive mAb AP5 in 2 mM Ca2+ but did not form a sta-
ble complex in solution with FN7–10 or full-length FN, unless the
integrin is activated by Mn2+. Quantitative binding assays showed
that Kd(app) values of 1TM-V3 and ∆TM-V3 binding to im-
mobilized FN7–10 in Mn2+ are comparable (0.15 and 0.1 µg/ml).
However, binding of 1TM-V3 and ∆TM-V3 in Ca2+/Mg2+
buffer yielded Kd(app) values that were 60-fold and 10-fold
lower, respectively, than in Mn2+ (Fig. 3, A and B), reflecting the
proportion of the active species in each preparation. Thus, 1TM,
which differs only in the added / lower leg, is less active and
yielded mean donor–acceptor separation distances of 96 Å, 94 Å,
and 90 Å, respectively. Thus, the donor–acceptor separation dis-
tances are comparable in the inactive, Mn2+-activated, constitu-
tively active, or FN10-bound states of the full-length V3
expressed on the surface of live cells. It is appropriate to men-
tion here that the calculated length of the genu-linear V3
molecule is >200 Å (Xiong et al., 2001).
Our major findings in this study are that (a) the C-terminal ex-
tensions of the V3 ectodomain by the remaining exofacial
residues and four / TM residues gave a water soluble and con-
formationally stable integrin that is predominantly inactive in
Ca2+/Mg2+ buffer in solution, (b) crystal structure of the near na-
tive 1TM-V3 in Ca2+ defined the first complete structure of an
integrin ectodomain plus an / TM fragment and allowed us
to build a structure model of the ectodomain plus the complete
Figure 6. FLIM analysis of V3 in live cells. (A) Histogram (mean ± SEM; n = 2) showing binding of subsaturating Alexa Fluor 488–labeled WT (open
bars) and high affinity (h; shaded bars) FN10 to full-length WT V3 stably expressed on K562 in the absence () or presence (+) of saturating amounts
of unlabeled 17E6 Fab in 1 mM Mn2+ (see Materials and methods). (B) Isocratic molecular sieve elution profiles in Mn2+-containing TBS buffer. 1TM-V3
(black) and its complexes with cilengitide (red), 17E6 Fab (blue), and 17E6 Fab complex followed by the addition of cilengitide to 10 µM (green) were
resolved. The elution profile of 17E6 Fab alone is also shown (violet). Cilengitide runs in the column volume. Cilengitide triggers an increase of the apparent
Stokes radius of the 1TM-V3–17E6 Fab complex. mAU, milli–absorbance unit. (C) Histogram (mean ± SD) showing lifetimes (in picoseconds) of Alexa
Fluor 488 fluorescence determined by FLIM in inactive (Ca2+) and active (Mn2+) full-length V3. *, P < 0.0001 versus donor only. (D) Representative Alexa
Fluor 488 fluorescence intensity of the unliganded WT integrin. The pseudocolored FLIM images represent donor fluorescence lifetimes on a pixel by pixel
basis, where shorter lifetimes are located toward the red area of the spectrum and longer lifetimes toward the blue area. Bar, 8 µm.
JCB • VOLUME 186 • NUMBER 4 • 2009 596
on the metal ion coordination in A. In the unliganded ∆TM-
IIb3 structure, the LIMBS residue D217 points toward LIMBS
(and not away from it as in 1TM-V3) to avoid a clash with a
hydrophobic residue (Phe191, which is a Trp in all other sub-
units; Fig. S1). This orientation provides five coordination sites
for a LIMBS Ca2+, with the sixth coordination site completed by
OE1 of E220, which is thus pulled out of the MIDAS pocket,
allowing coordination of an Mg2+ at the MIDAS.
As 1TM-V3 is the fourth integrin crystal structure re-
solved in a bent conformation under conditions that activate li-
gand binding in biochemical and cell biological assays, we once
again addressed the contentious issue of the conformation of the
integrin V3 at the cell surface. We positioned a FLIM donor
on the integrin head and an acceptor in the outer face of the
plasma membrane. FLIM measurements revealed no significant
change in mean separation distance of the integrin head relative
to the plasma membrane outer face in Mn2+-activated WT V3
compared with the inactive integrin (Fig. 6, C and D). A previ-
ous FRET study measured the distance between an FITC-
labeled ligand-mimetic peptide as donor and a plasma membrane
dye as acceptor (Chigaev et al., 2003). There, the change in
mean distance of closest approach was 50 Å between resting
and Mn2+-activated 41. However, as stated by the authors, the
membrane dye used, R18, has a tendency to flip-flop between
the outer and inner leaflets of the plasma membrane, introducing
uncertainty in the measurements. The FLIM methodology we
have used minimizes this problem and supports a more recent
cryoelectron tomography study of liposome-embedded and
Mn2+-activated IIb3 (Ye et al., 2008). Interestingly, high af-
finity soluble monomeric FN10 bound to the WT V3 in Mn2+
or the constitutive activation of V3 by G989FF/GAA or
∆-genu mutants also failed to trigger an increase in the mean
donor–membrane acceptor separation distance compared with
the inactive WT integrin. The simplest explanation for these
data is that a switch of the integrin from the inactive to the active
state or its binding to soluble FN10 in live cells can occur with
little or no genu extension, which is consistent with a recent
modeling study (Rocco et al., 2008).
The functional experiments presented in this study identify
a previously unappreciated but critical role for the IE2–thigh
interface in stabilizing the inactive state (Fig. 7). This is reflected
by the robust constitutive activation introduced in the surface-
expressed receptor upon deletion of the -genu, which contrib-
utes to this interface. Our results also show that stability of the
IE2–thigh interface appears to be conformationally linked to the
lower leg extensions introduced in 1TM-V3. Other mutations
in the lower leg domain that interrupt two salt bridges linking the
TD and IE3 to the top and bottom of the hybrid domain, respec-
tively (Fig. 7), revealed that these make a modest contribution to
stability of the inactive state: in V3, the steric barrier is mainly
mediated by the IE3–hybrid contact, whereas the TD–hybrid
contact appears to predominate in IIb3 (Matsumoto et al.,
2008). Our functional experiments also show that TD contacts
with both A and hybrid domains contribute to stability of the
inactive conformation, although this contribution is minor in
V3 and absent in IIb3 when the TD–A contact alone
is removed by deleting the CD loop in TD (Zhu et al., 2007).
hydrodynamically more compact than ∆T, reflecting a conforma-
tional coupling between the lower leg and head domains that regu-
lates integrin activation. Our present findings are also consistent
with the integrin existing in equilibrium between two major quater-
nary states, inactive (T state) and active (R state; Perutz, 1989),
with the T state predominating in soluble 1TM-V3 in Ca2+/Mg2+
buffer. mAb AP5 binds preferentially in Ca2+ or Ca2+/Mg2+ buffer
to the minor active species (estimated at 2% and 10% in 1TM-
and ∆TM-V3 preparations, respectively), driving the quaternary
equilibrium toward the active quaternary state (s) and stabilizing it
there (Fig. 3 E). The same shift can be accomplished by FN7–10 or
by native FN but requires Mn2+ not Ca2+, presumably because of
the higher affinity of Mn2+ to the metal ion–dependent adhesion
site (MIDAS) that mediates V3 interaction with ligands. Using
the measured Kd(app) values as approximations of binding affinities,
the mean difference in energy between the two major quaternary
states of 1TM- and ∆TM-V3 is estimated at 2.4 kcal/mol and
1.4 kcal/mol, respectively. It is expected that this value will be even
higher for the full-length integrin.
The crystal structure of ∆TM-IIb3 at 2.55-Å resolution
reported (Zhu et al., 2008) while this work was under review ends
at the equivalent residues to ∆TM-V3 and thus lacks those lower
leg residues and TM fragments defined in the current 1TM-V3
structure and shown to regulate affinity. Superposition of the
∆TM-IIb3 structure onto that of 1TM-V3 using Match-
maker in the Chimera software suite (Pettersen et al., 2004)
revealed several interesting differences between these two struc-
tures. First, a significant inward rotation of Calf-1 and -2 at the
-genu in ∆TM-IIb3 resulting from crystal contacts with sym-
metry-related molecules was associated with translational move-
ments in the IE2–4 domains. Second, the -genu is missing in the
∆TM-IIb3 structure, where it is presumed disordered (Zhu
et al., 2008). Yet, significant changes appear in the main chain
flanking the -genu in ∆TM-IIb3 compared with 1TM-V3,
suggesting proteolytic cleavage within this site in ∆TM-IIb3
as another likely explanation. Third, the TD of ∆TM-IIb3
veers toward IE4 and away from the hybrid/A domains, break-
ing the R633-coordinated salt bridge, which stabilizes the
inactive state of full-length IIb3 (Matsumoto et al., 2008).
The engineered disulphide linking TD to Calf-2 in ∆TM-IIb3
may impose these quaternary changes in the TD environment.
Fourth, the A domain of unliganded ∆TM-IIb3 contained the
MIDAS and the stimulatory ligand-associated metal-binding site
(LIMBS) metal ions (Mg2+ and Ca2+, respectively), but no metal
ions were present in the 1TM-V3 2.9-Å structure formed in
the presence of Ca2+ (Fig. S1) or in our published 3.1-Å unliganded
∆TM-V3 structure (Xiong et al., 2001), even when Mn2+ was
diffused into the crystals. Lower resolution (Zhu et al., 2008)
is an unlikely explanation for this difference, as both metal ions
are overt in the 3.2-Å structure of the ∆TM-V3–cRGD com-
plex (Xiong et al., 2002). Furthermore, in native A isolated from
the 3 subunit, LIMBS did not bind Ca2+ (Pesho et al., 2006), so
the absence of a LIMBS Ca2+ in unliganded 3 is independently
supported. Whether clasping the legs covalently by an artificial
disulphide permits metal occupancy at LIMBS through allosteric
mechanisms remains an open question. A likely interpretation is
that the difference reflects an influence of the associated subunit
CRYSTAL STRUCTURE 1TM-V3 • Xiong et al.
The shape of the quaternary R (active) state of the integrin
ectodomain remains to be defined structurally. But it includes the
high affinity state of A and a proposed genu-linear conformation,
which, it is argued, is required for switching A to the active state
(Takagi et al., 2002). In the homologous A domain, the high affin-
ity state is characterized by an inward movement of the N-terminal
1 helix and a two-turn downward movement of the C-terminal
7 helix, which is permitted by a flexible long linker distally (Lee
et al., 1995). The former movement was observed in the crystal
structure of the bent cRGD-bound ∆TM-V3 (Xiong et al., 2002)
but in the absence of the axial movement of 7 helix, which is
constrained distally in the bent crystal structure by the hybrid
domain. The activating effect of releasing constraints on the
hybrid in the R633 + R404 mutant is consistent with previous work
performed on integrin ectodomains with modified, truncated, or
entirely amputated legs (Mould et al., 2003; Xiao et al., 2004), which
found a correlation between opening the A–hybrid hinge and the
high affinity state of A. An 70° hinge opening in the legless
ectodomain structure (Xiao et al., 2004), which is associated with a
one-turn downward movement of the C-terminal 7 helix, led to the
conclusion that this feature characterizes the high affinity state of
A. However, a recent study (Chigaev et al., 2009) found that the
hybrid domain movement and the high affinity state are regulated
separately and independently of each other. And an EM study of
the intact ∆TM-V3 ectodomain complexed with FN7–extra do-
main B–10 detected only a small 11 ± 4° opening of the A–hybrid
hinge angle (Adair et al., 2005), which is sufficient to break the
IE2–thigh interface, suggesting that a rather modest hinge opening
may be all that is needed to switch A to high affinity. Consistently,
a molecular dynamics study of the legless IIb3 ectodomain de-
tected a spontaneous A–hybrid hinge opening of 20° when a
single N303-K417 bond linking A to hybrid is broken (Puklin-
Faucher et al., 2006), but over the course of this increase, a lateral
rather than the axial shift of the A domain C-terminal 7 helix was
observed. The same study found that subsequent pulling on bound
FN10 caused the A–hybrid domain hinge to further increase to 70°
in the legless ectodomain, suggesting that the wider hinge opening
requires force applied on a ligand-bound integrin. Whether such an
extreme movement requires preconversion of the ectodomain to a
genu-linear structure or can occur in a genu-bent conformation re-
quires further study. Our data suggest that overcoming the critical
IE2–thigh steric barrier through the activating -genu deletion does
not require a genu-linear conversion. It may also be relevant that
binding of soluble monomeric high affinity FN10 did not induce
genu linearity in the full-length V3, suggesting that the high af-
finity state and genu linearity are not conformationally linked events,
an interpretation which is consistent with experimental observations
in WT and modified integrins (Takagi et al., 2003; Coutinho et al.,
2007; Gupta et al., 2007). It has been proposed that genu linearity
requires force applied by the cytoskeleton to the cytoplasmic tails of
the unliganded integrin (Zhu et al., 2008). However, our FLIM data
on constitutively active V3 suggest that this remains as bent as
the inactive molecule. It is possible that the transition to the genu-
linear state requires that force be exerted on both ends of an integrin,
as when it is bound simultaneously to an immobilized ECM ligand
and to the cytoskeleton in mechanically stressed tissues (for review
see Arnaout et al., 2005).
Previous studies have shown that disulphides introduced at the
TD–A or TD–Calf-2 interfaces blocked integrin activation
(Takagi et al., 2002; Kamata et al., 2005), suggesting that the
small barriers at these nodes must also be overcome in the con-
formational switch of the integrin to the active state. Collectively,
these data suggest that binding of the F3 subdomain of talin to
the 3 cytoplasmic tail (mimicked by the G989FF/GAA mutant;
Wegener et al., 2007) unravels the / TM packing, triggering
movement of the membrane-proximal lower leg domains, reduc-
ing the energy cost of destabilizing the critical IE2–thigh inter-
face (Fig. 7), which then drives the conformational equilibrium
toward the quaternary R state almost completely (Fig. 5).
Figure 7. Ribbon diagram showing functionally relevant ionic contacts
in the 1TM-V3 structure. IE2 makes several electrostatic contacts with
two conserved loops at the bottom of the thigh domain. These include
E500 from IE2 making a salt bridge with the conserved CC loop residue
K503 and an H-bond with the invariant EF loop residue D550 (both
from the thigh domain) and the -genu residues E476 and D477 making
H-bonds with the EF residue E547. Two salt bridges (R633–D393 and
R404–D550) link the top and bottom of the hybrid domain to TD and IE3,
respectively. The small TD–A interface (S674–V332, in gray) and an
H-bond between the TD (D606) and Calf-2 (S749; Kamata et al., 2005)
are also shown.
JCB • VOLUME 186 • NUMBER 4 • 2009 598
Coomassie staining. Using ImageJ software (National Institutes of Health),
the measured intensities of the stained 1TM-V3 and FN7–10 gel bands
(Fig. 3 C) were 16,514 arbitrary units and 3,468 arbitrary units, respec-
tively, which is a 4.76:1 ratio, yielding a 1:1 stoichiometry, when the
molecular masses of 1TM-V3 (180.451 kD) and FN7–10 (39.846 kD)
are considered (a ratio of 4.53:1).
Binding of 1TM-V3 to the activation-sensitive mAb AP5
Binding of soluble 1TM-V3 to the Fab fragment of the ligand-inducible
binding site mAb AP5 (provided by P.J. Newman, Medical College of
Wisconsin, Milwaukee, WI; Honda et al., 1995) was performed by mixing
the integrin with the Fab in a 1:1.4 stoichiometric ratio for 30 min on ice
in TBS containing 2 mM CaCl2. The complex was detected by molecular
sieve chromatography in TBS plus 2 mM CaCl2, and the eluted peaks were
analyzed on 12% SDS-PAGE followed by Coomassie staining. Band inten-
sities for the unreduced stained gels corresponding to 1TM-V3 and AP5
Fab (Fig. 3 E) were 7,479.5 arbitrary units (for combined V and 3
bands) and 2,074.7 arbitrary units, respectively (integrin/Fab ratio of
3.6:1), yielding a 1:1 stoichiometry, with a molecular mass ratio of 3.6:1
(the calculated molecular mass of Fab is 50 kD).
mAb and soluble ligand binding to cell surface–expressed V3
WT V3–expressing K562 or transiently transfected HEK cells (6 × 105
cells per sample) were stained in suspension with 10 µg/ml LM609 and
10 µg/ml AP3 mAbs or 20 µg/ml 17E6 Fab in 100 µl TBS containing
0.5% BSA plus 1 mM CaCl2 + 1 mM MgCl2 or 1 mM MnCl2 for 30 min at
RT. After washing, cells were incubated with the fluorophore-conjugated
anti–mouse IgG or the anti-Fab antibody at 0°C for 30 min, washed, fixed,
and then analyzed by flow cytometry using the CellQuest software (BD).
FN binding was assessed in parallel by incubating V3-expressing K562
or batches of transient-transfected HEK 293T with mAb AP3 at 0°C for 45
min and then with 10 µg/ml of Alexa Fluor 488–labeled FN9–10 or FN10
and with Cy3-labeled goat anti–mouse IgG1 in TBS buffer containing 1 mM
CaCl2 + 1 mM MgCl2 or 1 mM MnCl2 at 20°C for 30 min. After washing,
cells were fixed and analyzed by flow cytometry, and the percentage of
AP3-positive cells bound to FN was determined. To assess the effect of un-
labeled 17E6 Fab prebound to cellular V3 on subsequent binding of
Alexa Fluor 488–FN10, K562 cells were first incubated in the absence or
presence of a saturating amount of Fab (at 30 µg/ml) as in the previous
section, washed, incubated at 20°C for 30 min with limiting concentrations
of WT or high affinity Alexa Fluor 488–FN10 (at 10 µg/ml and 2 µg/ml,
respectively), and then analyzed by FACS, and binding was expressed as
the percentage of the LM609-positive cells stained in parallel.
Hydrodynamic shift assay for locked integrin
To investigate whether 17E6 Fab binding locked 1TM-V3 in an inactive
conformation, 17E6 Fab was incubated in TBS containing 0.2 mM Mn2+
(MBB buffer) with 1TM-V3 (1.5:1 Fab/integrin molar ratio) at 20°C for
1.5 h. Aliquots were taken and chromatographed at this time point, and
the preformed complex was challenged by adding cilengitide to an end
concentration of 10 µM and by incubating at 20°C for 1 h and then re-
solved by molecular sieve chromatography in MBB buffer. The unligated
and 17E6 Fab–ligated ± cilengitide-treated forms of the 1TM-V3 com-
plex were resolved unequivocally as discrete symmetrical peaks on the mo-
lecular sieve column. The excess 17E6 Fab, eluting at Kav 0.52, served as
an internal standard. Each column run was repeated at least three times,
and the mean and SDs for the Kav and elution volumes were calculated
and compared using a Student’s t test.
Crystallography, structure determination, and refinement
1TM-V3 was crystallized at 4°C by vapor diffusion using the hanging-
drop method. The precipitating solution contained 12% PEG 4K, 0.6 M
NaCl, 5 mM CaCl2, and 0.1 M Na acetate at pH 5.5. Hexagonal crystals
grew over 4–5 d to dimensions of 0.3 × 0.4 × 0.15 mm3. A native x-ray
diffraction dataset from a single cryocooled crystal was collected using
the ID19 beamline fitted with a charge-coupled device detector at the
Advanced Photon Source Facility. All data were indexed, integrated, and
scaled with the HKL2000 program (Otwinowski and Minor, 1997). Data
collection statistics are presented in Table I. The structure was solved by
molecular replacement using the structure of the ∆TM-V3 as the search
model. The model was refined with the crystallography and NMR system
(CNS; Brünger et al., 1998) and manually adjusted with O (Jones et al.,
1991). After initial simulated annealing, the model was further refined with
macromolecular refinement by the maximum-likelihood method (REFMAC;
Winn et al., 2001). The complete structure of IE1 and -2, the linker between
Materials and methods
Construction, expression, and purification of 1TM-V3
We generated and cloned a new construct encoding human V residues
F1-I967 and 3 residues G1-V696 (with each ending with the fourth putative
TM residue, I967 and V696 of V and 3, respectively; Fig. 1 A) into the
pacUW31vector, and the resulting 1TM-V3 pacUW31 vector was used to
produce high titer recombinant 1TM-V3 virus, which was used to infect
High-Five cells (Invitrogen; Mehta et al., 1998). To assess expression of the sol-
uble recombinant integrin, sterile filtered supernatant was immunoprecipitated
with the V3 heterodimer-specific antibody LM609 (Cheresh and Spiro,
1987) conjugated to Affigel-10 beads, and after extraction in SDS-PAGE sam-
ple buffer, 200-µl supernatant equivalents were run on 4–12% gradient SDS
gels under reducing and nonreducing conditions. After Western blotting, the
membrane was blocked with skim milk–Tween and probed with mixed bio-
tinylated 20H9 and 2A5 antibodies, directed against the 3 and V subunits,
respectively, before detection with anti–biotin-HRP and ECL visualization.
62 ng of purified ∆TM-V3 was run in parallel as a standard. The hetero-
dimer could also be confirmed directly by capture ELISA using 17E6 (anti-V)
as capture reagent and AP3 (anti-3) as detection reagent. The 1TM-V3 re-
ceptor was purified for biochemistry and crystallography as previously
described for the ∆TM-V3 recombinant (Mehta et al., 1998) except that the
mAb 14D9 (Mitjans et al., 1995) was used in affinity chromatography.
Molecular sieve chromatography of purified 1TM-V3
All molecular sieve chromatography analyses were performed as previ-
ously described (Adair et al., 2005) using precalibrated Superdex 200
(10/300 GL) columns (GE Healthcare) on an Akta fast protein liquid chro-
matography system (GE Healthcare) running Unicorn 5.01 software (GE
Healthcare) at a flow rate of 0.4 ml min1 at 20°C. The elution profiles
were monitored in-line by UV adsorption at 280 nm. TBS buffer (145 mM
NaCl and 25 mM Tris-HCl, pH 7.4) containing either Ca and Mg (1 mM
MgCl2 and 1 mM CaCl2) or Mn (0.2 mM MnCl2) was used throughout.
5–10 µg of purified ∆TM-V3 or 1TM- V3 was fractionated, and the
Stokes radii were derived by substituting their peak elution volumes (Ve) in
the fitted standard curve equation (one-phase exponential fit; Prism; Graph-
Pad Software, Inc.). Identity of resolved peaks was formally confirmed by
SDS-PAGE and by ELISA using mAb LM609.
Plasmids and stable and transient transfections
Plasmids containing cDNAs encoding WT human full-length V and 3 were
each subcloned in pcDNA3. -genu deletion (-genu; residues
E472DYRPSQ482), R404A + R633A, and R633A + CD (residues D672-K676)
mutants in 3 and G989FF/GAA mutant in V were made using PCR-based
mutagenesis with the QuikChange kit (Agilent Technologies), and authenticity
was confirmed by DNA sequencing. Plasmids encoding WT human FN7–10
(P1142 to T1509; provided by H.P. Erickson, Duke University Medical Center,
Durham, NC; Leahy et al., 1996), FN9–10 (G1326 to T1509), FN10 (S1417
to T1509), or the high affinity and V3-specific form of FN10 (Richards
et al., 2003) were expressed in bacteria and purified as described previously
(Aukhil et al., 1993). pcDNA3 plasmids encoding full-length WT V and 3
were electroporated into K562 cells (American Type Culture Collection; Gupta
et al., 2007). Stable K562 clones expressing WT V3 were maintained in
Iscove’s modified Dulbecco’s medium plus 10% heat-inactivated fetal bovine
serum, 50 IU/ml penicillin and streptomycin, and 0.5–1.0 mg/ml G418.
V or 3 mutant pcDNA3 plasmids were cotransfected with plasmids encod-
ing the respective WT subunit into HEK 293T cells using Lipofectamine 2000
reagent (Invitrogen) according to the manufacturer’s protocol.
Binding of 1TM-V3 to immobilized and soluble ligands
Binding of increasing concentrations (0.001–10 µg/ml) of soluble 1TM-
V3 or ∆TM-V3 to immobilized purified FN type III domains 7–10
(FN7–10; at 3 µg/ml) was performed in the absence or presence of cyclic-
RGD (cyclo-[Arg-Gly-Asp-d-Phe-(N-Me)Val]-cilengitide) or a control cyclic
RAD peptide in 1 mM CaCl2 + 1 mM MgCl2 or 1 mM MnCl2 (in TBS; 0.1%
BSA for 3 h at 37°C; Kraft et al., 1999). Binding to uncoated wells was
performed in parallel. The bound integrin was detected using biotinylated
LM142 mAb (Millipore) at 2 µg/ml and 1:20,000 HRP-labeled anti-biotin
antibody (Sigma-Aldrich). Binding of 1TM-V3 to soluble FN7–10 was
also performed in solution (Adair et al., 2005) by mixing 1TM-V3 with
FN7–10 (in a 1:1.5 stoichiometric ratio) or with native full-length FN (in a
5:1 stoichiometric ratio) for 1 h at 37°C in 145 mM NaCl and 25 mM Tris-
HCl, pH 7.4, buffer (TBS) containing 0.2 mM MnCl2 or 1 mM CaCl2 and
1 mM MgCl2. The presence of the 1TM-V3–FN7–10 complex was de-
tected after molecular sieve chromatography, 8% SDS-PAGE analysis, and
599 CRYSTAL STRUCTURE 1TM-V3 • Xiong et al.
NaCl (TBS) containing 1 mM CaCl2, 1 mM CaCl2 and 1 mM MgCl2, or 1 mM
MnCl2 for 20 min at 37°C. After two washes, some cells were labeled with
6–12 µM FM4-64 FX (FM) in TBS containing 1 mM CaCl2, 1 mM CaCl2 and
1 mM MgCl2, or 1 mM MnCl2 for 5 min on ice, washed once, immediately
fixed with ice-cold 4% paraformaldehyde, washed, and mounted with GVA
mount (Invitrogen) under a coverslip. The GVA-mounted slides were kept in
the dark and used the next day for FLIM acquisition (Bacskai et al., 2003).
In some experiments, adherent K562 expressing WT V3 were pre-
incubated with saturating amounts (10 µg/ml) of unlabeled high affinity
FN10 followed by the addition of Alexa Fluor 488–Fab and then processed
as in the previous section.
FLIM measurements were made on a two-photon microscope (Radi-
ance 2000; Bio-Rad Laboratories) with a femtosecond-pulsed Ti:Sapphire
Laser (Mai Tai; Spectra-Physics) at 800-nm excitation. Photons were detected
by microchannel plate photomultiplier tube (MCP R3809; Hamamatsu Pho-
tonics) with time-correlated single photon counting (SPC830; Becker & Hickl)
to measure fluorescent decay profiles. Decay curves were best fit into mono-
exponential curves using SPCImage software (version 188.8.131.5211; Becker &
Hickl). Lifetimes for multiple cells (n = 9–14) for each experimental condition
were compared and evaluated for statistically significant differences from
control cells (labeled only with donor fluorophore) by analysis of variance
with Fisher’s post-hoc correction. Percent lifetime decrease, or FRET efficiency
(E), was calculated as the difference between the excited state of the donor
in absence of acceptor (D) and in presence of acceptor fluorophore (DA) ac-
cording to the equation E = 1 DA/D. The distance between donor (Fab)
and acceptor fluorophores, r, was calculated using the following equation
(Duncan et al., 2004): r = R0(D/DA 1)1/6, where R0 is the Förster radius,
the distance at which energy transfer is 50%. We assumed random orienta-
tion for the fluorophore (orientation factor, 2 = 2/3) in the Fab molecule.
We approximated R0 as 62 Å based on spectral similarities to the published
Alexa Fluor 488–Alexa Fluor 568 pair (Invitrogen).
Online supplemental material
Fig. S1 shows a ribbon diagram of the MIDAS face in the 1TM-V3
and TM-IIb3 structures. Table S1 shows the elution profiles after mo-
lecular sieve chromatography of 1TM-V3 or full-length FN alone and
in complex. Online supplemental material is available at http://www.jcb
We are grateful to Dr. Brian Adair for helpful discussions, Dr. Kay Gottschalk
for assistance with building the structure model, Dr. Paul Mould for suggesting
FLIM on the FN10-bound integrin, Jutta Welge for valuable technical assis-
tance, and the reviewers for their helpful comments.
This work was support by grants DK07447 and HL070219 from the
National Institutes of Health.
Submitted: 15 May 2009
Accepted: 16 July 2009
Adair, B.D., J.P. Xiong, C. Maddock, S.L. Goodman, M.A. Arnaout, and M.
Yeager. 2005. Three-dimensional EM structure of the ectodomain of inte-
grin V3 in a complex with fibronectin. J. Cell Biol. 168:1109–1118.
Andrew, S.A. 2002. Enzymatic digestion of monoclonal antibodies. In The Protein
Protocols Handbook. Second edition. J. M. Walker, editor. Humana Press,
Totowa, NJ. 1047–1052.
Arnaout, M.A., B. Mahalingam, and J.-P. Xiong. 2005. Integrin structure, allostery,
and bidirectional signaling. Annu. Rev. Cell Dev. Biol. 21:381–410.
Aukhil, I., P. Joshi, Y. Yan, and H.P. Erickson. 1993. Cell- and heparin-binding
domains of the hexabrachion arm identified by tenascin expression pro-
teins. J. Biol. Chem. 268:2542–2553.
Bacskai, B.J., J. Skoch, G.A. Hickey, R. Allen, and B.T. Hyman. 2003.
Fluorescence resonance energy transfer determinations using multiphoton
fluorescence lifetime imaging microscopy to characterize amyloid-beta
plaques. J. Biomed. Opt. 8:368–375.
Brünger, A.T., P.D. Adams, and L.M. Rice. 1998. Recent developments for the
efficient crystallographic refinement of macromolecular structures. Curr.
Opin. Struct. Biol. 8:606–611.
Cheresh, D.A., and R.C. Spiro. 1987. Biosynthetic and functional properties of an
Arg-Gly-Asp-directed receptor involved in human melanoma cell attach-
ment to vitronectin, fibrinogen, and von Willebrand factor. J. Biol. Chem.
Chigaev, A., A.M. Blenc, J.V. Braaten, N. Kumaraswamy, C.L. Kepley, R.P.
Andrews, J.M. Oliver, B.S. Edwards, E.R. Prossnitz, R.S. Larson, and
PSI and hybrid domains (residues P51ES53), which were not included in the
∆TM-V3 structure (Protein Data Bank number 1U8C; Xiong et al., 2004),
the remaining exofacial residues of the ectodomain (V’s A957-P963 and
3s P691D692), and seven of the eight TM residues were traced and in-
cluded in the final model. 14 N-linked carbohydrate structures modeled at
GlcNAc residues with additional branched sugars, and six Ca ions were
also placed. The final model converged to an Rwork value of 0.24 and Rfree
value of 0.28 (calculated with 5% of reflections omitted from refinement).
97% of the residues from the model fall into favorable or allowed regions
of the Ramachandran diagram using the PROCHECK software in CCP4
(Collaborative Computational Project Number 4, 1994).
Fluorescent labeling of 17E6 Fab and FN
The Fab fragment of the V-specific mAb 17E6 (Mitjans et al., 1995) was
prepared by papain digestion followed by anion exchange and size-exclusion
chromatography (Andrew, 2002), and its purity was confirmed by SDS-
PAGE followed by Coomassie staining. Fab 17E6 and high affinity
V3-specific FN10 were labeled with Alexa Fluor 488 N-hydroxysuccin-
imidyl ester dye using the Alexa Fluor 488 protein labeling kit (Invitrogen)
according to the manufacturer’s instructions. The final antibody and FN con-
centrations and the dye to protein molar ratios (F/P) were determined spec-
trophotometrically, giving F/P molar ratios of 3–7 (for Fab) and 1 (for
FN9–10 and FN10; FN9 has no lysines). Binding of the fluorophore-labeled
FN to V3-expressing cells was evaluated by flow cytometry, and dose–
response curves were established to determine the optimal (saturating or
subsaturating) concentration of ligand used in subsequent experiments.
Time-correlated single photon counting–FLIM acquisition and analysis
Wells of nonfluorescent Labtek II four-chamber microscope slides (Thermo
Fisher Scientific) were coated with poly-l-Lys (Sigma-Aldrich) overnight at
4°C. 25,000–50,000 V3-expressing K562 or HEK 293T cells were trans-
ferred in serum-free Iscove’s modified Dulbecco’s medium to each well and
incubated for 30 min at 37°C in a total volume of 200 µl. Nonspecific sites
were blocked by incubation with 10% serum-rich medium for 10 min at RT
and then washed twice to remove nonadherent cells. Adherent live cells
were stained with 20 µg/ml 17E6 labeled with the fluorescence donor
Alexa Fluor 488 (Alexa Fluor 488–Fab) in 25 mM Tris, pH 7.4, and 145 mM
Table I. Data collection, refinement, and model statistics
Data collection statistics
Unit cell dimensions (Å)
Resolution range (Å)
Number of unique reflections
Resolution range (Å)
Rfactor (work set) (%)b
Rfactor (free set) (%)
Mean B factors (Å2)
Atoms in the model
Number of GlcNAc
Number of Ca2+
(RMSD from ideality)
a = b = 130.256, c = 305.982
Bond lengths (Å)
Bond angles (°)
RMSD, root mean square deviation. Values in parentheses are for the highest
resolution shell (0.1 Å).
aRsym = |I <I>|/I, where I is the observed intensity and <I> is the mean
intensity from multiple observations of symmetry-related reflections.
bRfactor = hkl|Fo(hkl) Fc(hkl)|/hkl Fo(hkl).
JCB • VOLUME 186 • NUMBER 4 • 2009 600 Download full-text
Mould, A.P., S.J. Barton, J.A. Askari, P.A. McEwan, P.A. Buckley, S.E. Craig,
and M.J. Humphries. 2003. Conformational changes in the integrin beta
A domain provide a mechanism for signal transduction via hybrid domain
movement. J. Biol. Chem. 278:17028–17035.
Otwinowski, Z., and W. Minor. 1997. Processing of X-ray diffraction data col-
lected in oscillation mode. In Macromolecular Crystallography. Methods
in Enzymology, vol. 276. J.N. Abelson, M.I. Simon, C.W. Carter Jr., and
R.M. Sweet, editors. Academic Press, New York. 307–326.
Perutz, M.F. 1989. Mechanisms of cooperativity and allosteric regulation in
proteins. Q. Rev. Biophys. 22:139–237.
Pesho, M.M., K. Bledzka, L. Michalec, C.S. Cierniewski, and E.F. Plow. 2006.
The specificity and function of the metal-binding sites in the integrin
beta3 A-domain. J. Biol. Chem. 281:23034–23041.
Pettersen, E.F., T.D. Goddard, C.C. Huang, G.S. Couch, D.M. Greenblatt, E.C.
Meng, and T.E. Ferrin. 2004. UCSF Chimera—a visualization system for
exploratory research and analysis. J. Comput. Chem. 25:1605–1612.
Puklin-Faucher, E., M. Gao, K. Schulten, and V. Vogel. 2006. How the headpiece
hinge angle is opened: new insights into the dynamics of integrin activa-
tion. J. Cell Biol. 175:349–360.
Richards, J., M. Miller, J. Abend, A. Koide, S. Koide, and S. Dewhurst. 2003.
Engineered fibronectin type III domain with a RGDWXE sequence binds
with enhanced affinity and specificity to human alphavbeta3 integrin.
J. Mol. Biol. 326:1475–1488.
Rocco, M., C. Rosano, J.W. Weisel, D.A. Horita, and R.R. Hantgan. 2008. Integrin
conformational regulation: uncoupling extension/tail separation from changes
in the head region by a multiresolution approach. Structure. 16:954–964.
Rohl, C.A., C.E. Strauss, K.M. Misura, and D. Baker. 2004. Protein structure
prediction using Rosetta. Methods Enzymol. 383:66–93.
Senes, A., D.E. Engel, and W.F. DeGrado. 2004. Folding of helical membrane
proteins: the role of polar, GxxxG-like and proline motifs. Curr. Opin.
Struct. Biol. 14:465–479.
Stetefeld, J., U. Mayer, R. Timpl, and R. Huber. 1996. Crystal structure of three
consecutive laminin-type epidermal growth factor-like (LE) modules of
laminin gamma1 chain harboring the nidogen binding site. J. Mol. Biol.
Takagi, J., B.M. Petre, T. Walz, and T.A. Springer. 2002. Global conformational
rearrangements in integrin extracellular domains in outside-in and inside-
out signaling. Cell. 110:599–11.
Takagi, J., K. Strokovich, T.A. Springer, and T. Walz. 2003. Structure of integrin
alpha5beta1 in complex with fibronectin. EMBO J. 22:4607–4615.
Wegener, K.L., A.W. Partridge, J. Han, A.R. Pickford, R.C. Liddington, M.H.
Ginsberg, and I.D. Campbell. 2007. Structural basis of integrin activation
by talin. Cell. 128:171–182.
Winn, M.D., M.N. Isupov, and G.N. Murshudov. 2001. Use of TLS parameters
to model anisotropic displacements in macromolecular refinement. Acta
Crystallogr. D Biol. Crystallogr. 57:122–133.
Wouters, M.A., I. Rigoutsos, C.K. Chu, L.L. Feng, D.B. Sparrow, and S.L.
Dunwoodie. 2005. Evolution of distinct EGF domains with specific
functions. Protein Sci. 14:1091–1103.
Xiao, T., J. Takagi, B.S. Coller, J.H. Wang, and T.A. Springer. 2004. Structural
basis for allostery in integrins and binding to fibrinogen-mimetic thera-
peutics. Nature. 432:59–67.
Xiong, J.P., T. Stehle, B. Diefenbach, R. Zhang, R. Dunker, D.L. Scott, A.
Joachimiak, S.L. Goodman, and M.A. Arnaout. 2001. Crystal structure of
the extracellular segment of integrin alpha Vbeta3. Science. 294:339–345.
Xiong, J.P., T. Stehle, R. Zhang, A. Joachimiak, M. Frech, S.L. Goodman, and M.A.
Arnaout. 2002. Crystal structure of the extracellular segment of integrin alpha
Vbeta3 in complex with an Arg-Gly-Asp ligand. Science. 296:151–155.
Xiong, J.P., T. Stehle, S.L. Goodman, and M.A. Arnaout. 2003. New insights into
the structural basis of integrin activation. Blood. 102:1155–1159.
Xiong, J.P., T. Stehle, S.L. Goodman, and M.A. Arnaout. 2004. A novel adapta-
tion of the integrin PSI domain revealed from its crystal structure. J. Biol.
Ye, F., J. Liu, H. Winkler, and K.A. Taylor. 2008. Integrin alpha IIb beta 3 in a
membrane environment remains the same height after Mn2+ activation
when observed by cryoelectron tomography. J. Mol. Biol. 378:976–986.
Zhu, J., B. Boylan, B.H. Luo, P.J. Newman, and T.A. Springer. 2007. Tests of
the extension and deadbolt models of integrin activation. J. Biol. Chem.
Zhu, J., B.H. Luo, T. Xiao, C. Zhang, N. Nishida, and T.A. Springer. 2008. Structure
of a complete integrin ectodomain in a physiologic resting state and activa-
tion and deactivation by applied forces. Mol. Cell. 32:849–861.
Zhu, J., B.H. Luo, P. Barth, J. Schonbrun, D. Baker, and T.A. Springer. 2009. The
structure of a receptor with two associating transmembrane domains on
the cell surface: integrin alphaIIbbeta3. Mol. Cell. 34:234–249.
L.A. Sklar. 2001. Real time analysis of the affinity regulation of alpha
4-integrin. The physiologically activated receptor is intermediate in af-
finity between resting and Mn(2+) or antibody activation. J. Biol. Chem.
Chigaev, A., T. Buranda, D.C. Dwyer, E.R. Prossnitz, and L.A. Sklar. 2003.
FRET detection of cellular alpha4-integrin conformational activation.
Biophys. J. 85:3951–3962.
Chigaev, A., A. Waller, O. Amit, L. Halip, C.G. Bologa, and L.A. Sklar. 2009.
Real-time analysis of conformation-sensitive antibody binding provides
new insights into integrin conformational regulation. J. Biol. Chem.
Collaborative Computational Project Number 4. 1994. The CCP4 suite: programs for
protein crystallography. Acta Crystallogr. D Biol. Crystallogr. 50:760–763.
Coutinho, A., C. García, J. González-Rodríguez, and M.P. Lillo. 2007.
Conformational changes in human integrin alphaIIbbeta3 after platelet
activation, monitored by FRET. Biophys. Chem. 130:76–87.
Duncan, R.R., A. Bergmann, M.A. Cousin, D.K. Apps, and M.J. Shipston. 2004.
Multi-dimensional time-correlated single photon counting (TCSPC) fluor-
escence lifetime imaging microscopy (FLIM) to detect FRET in cells.
J. Microsc. 215:1–12.
Faccio, R., M. Grano, S. Colucci, A. Villa, G. Giannelli, V. Quaranta, and A.
Zallone. 2002. Localization and possible role of two different alpha v
beta 3 integrin conformations in resting and resorbing osteoclasts. J. Cell
Fiser, A., and A. Sali. 2003. Modeller: generation and refinement of homology-
based protein structure models. Methods Enzymol. 374:461–491.
Giepmans, B.N., S.R. Adams, M.H. Ellisman, and R.Y. Tsien. 2006. The fluor-
escent toolbox for assessing protein location and function. Science.
Gupta, V., A. Gylling, J.L. Alonso, T. Sugimori, P. Ianakiev, J.P. Xiong, and M.A.
Arnaout. 2007. The beta-tail domain (betaTD) regulates physiologic li-
gand binding to integrin CD11b/CD18. Blood. 109:3513–3520.
Honda, S., Y. Tomiyama, A.J. Pelletier, D. Annis, Y. Honda, R. Orchekowski, Z.
Ruggeri, and T.J. Kunicki. 1995. Topography of ligand-induced binding
sites, including a novel cation-sensitive epitope (AP5) at the amino terminus,
of the human integrin beta 3 subunit. J. Biol. Chem. 270:11947–11954.
Hynes, R.O. 2002. Integrins: bidirectional, allosteric signaling machines. Cell.
Jares-Erijman, E.A., and T.M. Jovin. 2003. FRET imaging. Nat. Biotechnol.
Jones, T.A., J.Y. Zou, S.W. Cowan, and M. Kjeldgaard. 1991. Improved methods
for building protein models in electron density maps and the location of
errors in these models. Acta Crystallogr. A. 47:110–119.
Kamata, T., M. Handa, Y. Sato, Y. Ikeda, and S. Aiso. 2005. Membrane-proximal
alpha/beta stalk interactions differentially regulate integrin activation.
J. Biol. Chem. 280:24775–24783.
Kim, M., C.V. Carman, and T.A. Springer. 2003. Bidirectional transmembrane signal-
ing by cytoplasmic domain separation in integrins. Science. 301:1720–1725.
Kraft, S., B. Diefenbach, R. Mehta, A. Jonczyk, G.A. Luckenbach, and S.L.
Goodman. 1999. Definition of an unexpected ligand recognition motif for
alphav beta6 integrin. J. Biol. Chem. 274:1979–1985.
Lau, T.L., C. Kim, M.H. Ginsberg, and T.S. Ulmer. 2009. The structure of the
integrin alphaIIbbeta3 transmembrane complex explains integrin trans-
membrane signalling. EMBO J. 28:1351–1361.
Leahy, D.J., I. Aukhil, and H.P. Erickson. 1996. 2.0 A crystal structure of a four-
domain segment of human fibronectin encompassing the RGD loop and
synergy region. Cell. 84:155–164.
Lee, J.O., L.A. Bankston, M.A. Arnaout, and R.C. Liddington. 1995. Two con-
formations of the integrin A-domain (I-domain): a pathway for activa-
tion? Structure. 3:1333–1340.
Matsumoto, A., T. Kamata, J. Takagi, K. Iwasaki, and K. Yura. 2008. Key in-
teractions in integrin ectodomain responsible for global conformational
change detected by elastic network normal-mode analysis. Biophys. J.
Mehta, R.J., B. Diefenbach, A. Brown, E. Cullen, A. Jonczyk, D. Güssow,
G.A. Luckenbach, and S.L. Goodman. 1998. Transmembrane-truncated
alphavbeta3 integrin retains high affinity for ligand binding: evidence for
an ‘inside-out’ suppressor? Biochem. J. 330:861–869.
Mitjans, F., D. Sander, J. Adán, A. Sutter, J.M. Martinez, C.S. Jäggle, J.M.
Moyano, H.G. Kreysch, J. Piulats, and S.L. Goodman. 1995. An anti-
alpha v-integrin antibody that blocks integrin function inhibits the devel-
opment of a human melanoma in nude mice. J. Cell Sci. 108:2825–2838.
Mould, A.P., J.A. Askari, and M.J. Humphries. 2000. Molecular basis of ligand
recognition by integrin alpha 5beta 1. I. Specificity of ligand binding is
determined by amino acid sequences in the second and third NH2-terminal
repeats of the alpha subunit. J. Biol. Chem. 275:20324–20336.