Spatial distribution and functional significance of activated vinculin in living cells.
ABSTRACT Conformational change is believed to be important to vinculin's function at sites of cell adhesion. However, nothing is known about vinculin's conformation in living cells. Using a Forster resonance energy transfer probe that reports on changes in vinculin's conformation, we find that vinculin is in the actin-binding conformation in a peripheral band of adhesive puncta in spreading cells. However, in fully spread cells with established polarity, vinculin's conformation is variable at focal adhesions. Time-lapse imaging reveals a gradient of conformational change that precedes loss of vinculin from focal adhesions in retracting regions. At stable or protruding regions, recruitment of vinculin is not necessarily coupled to the actin-binding conformation. However, a different measure of vinculin conformation, the recruitment of vinexin beta by activated vinculin, shows that autoinhibition of endogenous vinculin is relaxed at focal adhesions. Beyond providing direct evidence that vinculin is activated at focal adhesions, this study shows that the specific functional conformation correlates with regional cellular dynamics.
Methods in Enzymology 02/1992; 211:353-88. · 2.04 Impact Factor
Article: Intramolecular interactions in vinculin control alpha-actinin binding to the vinculin head.[show abstract] [hide abstract]
ABSTRACT: Using blot overlay techniques we have investigated the interaction of vinculin with alpha-actinin. We show that an alpha-actinin binding site is located in the 90 kDa vinculin head and confirm a vinculin binding site in the C-terminal rod of alpha-actinin, as recently reported by McGregor et al. [(1994) Biochem. J. 310, 225-233]. The isolated vinculin head binds much more strongly to alpha-actinin than intact vinculin. Using a proteolytic 81 kDa head fragment, we show that vinculin residues 1-107 are required for alpha-actinin binding. Antibodies directed against vinculin residues 808-850 inhibit the vinculin-alpha-actinin binding, suggesting that this sequence is directly involved in, or topographically related to, the alpha-actinin binding site.FEBS Letters 01/1995; 355(3):259-62. · 3.54 Impact Factor
T H E J O U R N A L O F C E L L B I O L O G Y
The Journal of Cell Biology, Vol. 169, No. 3, May 9, 2005 459–470
The Rockefeller University Press$8.00
Spatial distribution and functional significance of
activated vinculin in living cells
Daniel M. Cohen,
Dilshad M. Choudhury,
and Susan W. Craig
Department of Biological Chemistry, Johns Hopkins University School of Medicine, Baltimore, MD 21205
Division of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Sakyo-ku, Kyoto 606-8502, Japan
onformational change is believed to be important
to vinculin’s function at sites of cell adhesion.
However, nothing is known about vinculin’s con-
formation in living cells. Using a Forster resonance energy
transfer probe that reports on changes in vinculin’s
conformation, we find that vinculin is in the actin-binding
conformation in a peripheral band of adhesive puncta in
spreading cells. However, in fully spread cells with estab-
lished polarity, vinculin’s conformation is variable at focal
adhesions. Time-lapse imaging reveals a gradient of
conformational change that precedes loss of vinculin from
focal adhesions in retracting regions. At stable or pro-
truding regions, recruitment of vinculin is not necessarily
coupled to the actin-binding conformation. However, a
different measure of vinculin conformation, the recruit-
ment of vinexin
by activated vinculin, shows that auto-
inhibition of endogenous vinculin is relaxed at focal
adhesions. Beyond providing direct evidence that vinculin
is activated at focal adhesions, this study shows that the
specific functional conformation correlates with regional
Vinculin is a 116-kD, cytoskeleton-associated protein that is
essential for brain and heart development in mice (Xu et al.,
1998a) and for muscle contraction in nematodes (Barstead and
Waterston, 1991). Vinculin is expressed in most cell types and
tissues (Otto, 1990), but its localization in muscle is particularly
informative. In skeletal and cardiac muscle, vinculin describes
a subsarcolemmal lattice of transmembrane connections or
costameres (Craig and Pardo, 1983; Pardo et al., 1983a,b;
Shear and Bloch, 1985) that attach the myofibrils to the sarco-
lemma (Pierobon-Bormioli, 1981) and transduce force laterally
through the extracellular matrix to neighboring myocytes
(Street, 1983; Ervasti, 2003). Vinculin is also enriched at myo-
tendinous junctions (Shear and Bloch, 1985) and intercalated
discs (Koteliansky and Gneushev, 1983; Pardo et al., 1983a),
intercellular adhesion structures involved in longitudinal trans-
mission of forces between adjacent muscle cells. Vinculin
likely plays a role in muscle structure and stability to mechanical
forces. Recent data shows that hearts of vin
posed to stress-induced cardiomyopathy (Zemljic-Harpf et al.,
2004). In several cases, mutations and deletion of metavinculin,
mice are predis-
the muscle-specific spliceform of vinculin (Byrne et al., 1992),
are correlated with idiopathic dilated cardiomyopathy (Maeda
et al., 1997; Olson et al., 2002), offering additional support for
the importance of these proteins in muscle function.
Evidence from cells in culture suggests that vinculin
functions in transducing force across cell membranes (Danowski
et al., 1992; Alenghat et al., 2000), in regulating cell adhesion
and motility (Rodriguez Fernandez et al., 1993; Xu et al.,
1998b; DeMali et al., 2002), in controlling cell survival
(Subauste et al., 2004), and in executing rac-mediated signaling
events (DeMali et al., 2002; Goldmann and Ingber, 2002). In
cultured cells, vinculin is enriched at cell–cell and cell–matrix
junctions (Geiger, 1979) but is in equilibrium with a large cyto-
plasmic pool (Lee and Otto, 1997). When cells adhere and
spread on ECM, a portion of the cytoplasmic vinculin is recruited
to specialized sites on the plasma membrane called focal adhe-
sions and focal contacts. At these sites, dynamic connections
are made between the actin cytoskeleton and ECM through
transmembrane integrin and syndecan receptors for extracellular
matrix molecules. These connections relay force across the
membrane (Balaban et al., 2001; Beningo et al., 2001; Galbraith
et al., 2002; Tan et al., 2003) and are essential for regulation of
cell motility (Palecek et al., 1997; Priddle et al., 1998). Focal
adhesions appear to be compositionally, structurally, and func-
tionally analogous to the costameres of skeletal and cardiac
muscle and the dense plaques of smooth muscle cells and are
Address correspondence to Susan W. Craig: email@example.com
Abbreviations used in this paper: FRET, Forster resonance energy transfer; SE,
sensitized YFP emission; Vh, vinculin head domain; vin
mouse embryo cells; Vt, vinculin tail domain.
The online version of this article includes supplemental material.
MEC, vinculin null
JCB • VOLUME 169 • NUMBER 3 • 2005 460
therefore a good model system to examine the mechanism and
significance of vinculin action.
Because vinculin lacks intrinsic enzymatic activity, it
must exert its functions through interaction with other proteins.
Indeed, multiple proteins, including F-actin, talin,
-catenin, vinexin, VASP, ponsin, CAP, arp2/3 (DeMali et al.,
2002), Raver-1 (Huttelmaier et al., 2001), PKC (Tigges et al.,
2003), and paxillin, interact with specific domains of vinculin
in vitro and colocalize with vinculin at ECM contacts in vivo
(for reviews see Critchley, 2000; Zamir and Geiger, 2001).
However, in contrast to isolated domains of vinculin and to
vinculin immobilized on nitrocellulose, full-length vinculin in
solution binds poorly or undetectably to many of these ligands
(Johnson and Craig, 1994, 1995; Kroemker et al., 1994; Huttel-
maier et al., 1998; Bakolitsa et al., 2004). The inert state of vin-
culin is caused by high affinity intramolecular binding between
the vinculin head domain (Vh; residues 1–851 or 1–857) and
tail domain (Vt; residues 884–1066) of vinculin (Johnson and
Craig, 1994). In bimolecular assays, the
50 nM (Johnson and Craig, 1994); in an intramolecu-
lar context the interaction is estimated, but not directly mea-
sured, to be
10 (Bakolitsa et al., 2004). Thus, it is
implicit that disruption of the Vh–Vt interaction is required for
vinculin activation and subsequent assembly of vinculin-con-
taining protein complexes at adhesion junctions (Johnson and
Nevertheless, the model for vinculin activation and func-
tion is supported solely by in vitro biochemistry using purified
proteins and their domains. Nothing is known about vinculin
conformation in cells, its relevance to focal adhesion composi-
tion, or its relationship to cellular dynamics. Here, we present
data providing new insights on all three of these issues.
of the Vh–Vt com-
Construction of vinculin Forster
resonance energy transfer (FRET)
To monitor activation of vinculin, we developed FRET probes
using CFP and YFP as the donor-acceptor pair (Miyawaki and
Tsien, 2000). Because the crystal structure of full-length vincu-
lin was not known, we began by positioning ECFP and EYFP
on the NH
- and COOH-terminal residues of vinculin (CVY;
Fig. 1 A). In cell lysates prepared from HEK 293 cells trans-
fected with CVY, the corrected FRET emission ratio (see Ma-
terials and methods) of CVY was 0.05, only slightly greater
than the baseline for CFP alone which was set to zero for the
calculation. The calculated FRET efficiency (see Materials and
methods) for CVY was only 3%, indicating that the NH
COOH termini are not in proximity. In addition, there was little
change in FRET of the CVY probe upon activation of vinculin
by IpaA and binding to actin (unpublished data).
Because CVY was not a suitable FRET probe, we ex-
plored internal placements for one of the fluorescent proteins.
The intramolecular distance between the NH
mini of Vt measured from the crystal structure of Vt (Bakolitsa
et al., 1999) is
14 Å. Thus, we anticipated strong FRET from
and COOH ter-
a construct in which CFP and YFP are positioned at the two
ends of Vt. Indeed, an EYFP/Vt/ECFP fusion protein exhibited
robust FRET (emission ratio of 1.3; efficiency of 43%) and
maintained the ability to bind to Vh (unpublished data). There-
fore, to construct a full-length vinculin FRET probe we in-
serted EYFP into vinculin at the beginning of the tail domain
and placed an ECFP at the COOH terminus of vinculin to make
the FRET probe, referred to as tail probe (Fig. 1, A and C).
Characterization of tail probe
Tail probe and various control constructs were expressed in
HEK293 cells, and lysates were analyzed by spectrofluorime-
try. Tail probe exhibited a strong FRET signal (Fig. 1 B), with
a corrected emission ratio of 1.48 and an efficiency of 46% (see
Fig. 4, A and B). To determine whether or not intermolecular
FRET contributes to the FRET measurement, we compared a
structure of vinculin FRET probes. (B) The emission spectra of vinculin1-883-
EYFP-vinculin884-1066-ECFP (Tail Probe), ECFP-vinculin-EYFP (CVY), EYFP-
ECFP-vinculin1-400 (control probe), and vinculin-ECFP (VC). Numbers refer
to amino acid residues in chicken vinculin (Coutu and Craig, 1988). Spectra
were normalized to the emission of VC at 475 nm. (C) The structures of
vinculin and GFP showing the size of each molecule. The arrow marks the
site of YFP insertion between residues 883 and 884.
Structure and spectral properties of the FRET probes. (A) Schematic
DETECTING VINCULIN ACTIVATION IN LIVING CELLS • CHEN ET AL. 461
lysate containing tail probe at 5, 10, and 20 nM with mixtures
of lysates containing vinculin/CFP and vinculin/YFP at 5, 10,
or 20 nM each in the mixture. At all concentrations tested, the
corrected FRET ratio of the mixed probes was close to that of
CFP only, whereas the corrected FRET ratio of tail probe was
insensitive to dilution. Thus, the FRET values obtained for tail
probe report specifically on intramolecular FRET.
SDS-PAGE of HEK 293 cell lysates and Western blot-
ting revealed that tail probe shows three closely spaced bands
near the expected molecular weight. All species were recog-
nized by both antivinculin and anti-GFP, which cross-reacted
with CFP and YFP (Fig. S1, available at http://www.jcb.org/
cgi/content/full/jcb.200410100/DC1). No evidence of pro-
teolytic cleavage of CFP or YFP was detected in the form of
GFP-sized bands. Because there is SDS-resistant structure in
the CFP and/or YFP moieties (Fig. S2, available at http://
www.jcb.org/cgi/content/full/jcb.200410100/DC1), the hetero-
geneity in migration of the FRET probe likely represents a
combination of the fully denatured form (slowest migrating)
and faster migrating, partially folded intermediates.
To determine the conformational state of tail probe and to
assess its ability to report on activation of vinculin, we mea-
sured FRET response as a function of ligand binding to vincu-
lin. Neither tail probe nor endogenous vinculin cosedimented
with F-actin, indicating that the probe and endogenous vinculin
were in a conformationally closed and inactive state (Fig. 2 B).
As expected, addition of F-actin caused no change in the FRET
signal of the probe (Figs. 2 A). To stimulate vinculin to bind
F-actin, we treated lysates with IpaA. IpaA is a
virulence protein that binds to the D1 domain (residues 1–258;
see Bakolitsa et al.  for nomenclature reflecting new
structure-based subdivisions of vinculin domains) of vinculin
head and exposes the actin binding activity of vinculin tail
(Bourdet-Sicard et al., 1999). Addition of IpaA alone did not
cause a change in FRET, but subsequent addition of F-actin
caused a 45% decrease in the corrected FRET ratio (1.48 to
0.81) of tail probe and 14% decrease in FRET efficiency (46%
to 32%), reflecting a change in the conformation of vinculin
(Fig. 2 A; and see Fig. 4, A and B). Tail probe cosedimented
with actin filaments only in the presence of IpaA, demonstrat-
ing that the loss of FRET reports on binding of IpaA-activated
tail probe to F-actin (Fig. 2 B).
Tail probe and endogenous vinculin differ in their sensi-
tivity to IpaA (Fig. 2 B). This difference is abolished by inclu-
sion of 1% Triton X-100 in the lysate (unpublished data). The
requirement of Triton X-100 for IpaA activation of endogenous
vinculin in cell lysates is unexpected because IpaA can activate
at least 40% of purified smooth muscle vinculin and recombi-
nant vinculin in vitro without the presence of Triton X-100
(Bourdet-Sicard et al., 1999; see Fig. 9). The differential re-
sponse between the tail probe and endogenous vinculin reflects
a more tightly closed conformation in endogenous vinculin,
which may be mediated by a Triton X-100–sensitive compo-
nent in the lysate. Despite this difference, the inability of the
tail probe to cosediment with actin filaments shows that like
endogenous vinculin, it adopts an autoinhibited conformation.
Furthermore, tail probe and vinculin localize similarly in cells
to IpaA-activated vinculin tail probe induced FRET loss, indicating a
conformational change of vinculin in the tail domain. (A) Normalized
fluorescence emission spectra of cell lysate from HEK 293 cells transfected
with tail probe in the absence or presence of 1 ?M IpaA or 5 ?M actin or
both. Spectra were normalized to the emission of tail probe at 475 nm.
(B) Samples from A were spun in an Airfuge (Beckman Coulter) at 25 psi
(130,000 g) for 35 min. Equivalent amounts of total sample before spin
(T), supernatant (S), and pellet (P) fractions were subjected to SDS-PAGE
and immunoblotted with hVIN1 and C4 mAbs (Sigma-Aldrich) to vinculin
and actin, respectively.
Response of tail probe to ligands. The binding of actin filaments
Normalized fluorescence emission spectra of cell lysates from HEK 293
cells transfected with the control FRET probe in the absence or in the pres-
ence of 1 ?M IpaA or 5 ?M actin or both. Spectra were normalized to
emission at 475 nm of control probe alone. The control probe preserves
the IpaA binding site and a focal adhesion targeting signal of vinculin but
lacks the actin binding site. It does not display FRET change in response to
IpaA binding. (B) Actin cosedimentation assay, performed under the same
conditions as in Fig. 2 B, showed that the control probe did not bind to
actin filaments under conditions in which tail probe did bind.
Response of the control FRET probe YC-V1-400 to ligands. (A)
JCB • VOLUME 169 • NUMBER 3 • 2005462
and are equally able to rescue spreading defects in vinculin null
cells (see Fig. 5).
For a control probe, we constructed an EYFP-ECFP chi-
mera fused in frame to vinculin residues 1–400 (Fig. 1 A). This
probe contains the binding site for IpaA (Bourdet-Sicard et al.,
1999) and a focal adhesion targeting motif (Bendori et al.,
1989) but lacks F-actin binding capacity (Menkel et al., 1994).
Control probe had a corrected FRET ratio of 1.4 and a FRET
efficiency of 44% (see Fig. 4, A and B). These values are simi-
lar, fortuitously, to unstimulated tail probe in cell lysates. This
property was useful because it allowed us to use the FRET sig-
nal of control probe observed in cells to define the baseline
FRET for the closed conformation of tail probe. There was no
significant change in FRET for the control probe in cell lysates
either before or after treatment with IpaA, actin, or both ligands
together (Fig. 3 A; and Fig. 4, A and B); nor did the control
probe cosediment with actin (Fig. 3 B). Therefore, we conclude
that tail probe reports on conformational changes in vinculin
that reflect its activation and binding to actin filaments,
whereas control probe is insensitive to F-actin and to IpaA, an
activator of the Vh.
When transfected into vinculin null mouse embryo cells
MEC; Xu et al., 1998a), both the tail probe and un-
tagged vinculin showed a diffuse cytoplasmic pool and were
similarly enriched at focal adhesions (Fig. 5 A). Tail probe and
untagged vinculin were equally able to rescue the spreading de-
fect (Xu et al., 1998a,b) and change in cell shape (DeMali et
al., 2002) as shown in Fig. 5 (C and E and B and D, respec-
tively). Thus, the vinculin tail FRET probe is suitable for anal-
ysis of vinculin conformation in living cells.
Detection of tail probe FRET in living cells
To determine if vinculin conformation correlates with subcellu-
lar localization, we transfected vin
MEC with tail probe and
calculated a FRET image from the digital data as described in
Materials and methods. The global emission ratio (averaged over
all pixels above threshold) observed in these cells was 1.43
0.18 SD (
8, compiled from two experiments), which was
distinguishable from the baseline of 0.6 obtained from cells
transfected with vinculin/CFP. To confirm that the emission ra-
tio reflects FRET, the average FRET efficiency of tail probe in
cells was determined by fluorescence recovery after acceptor
photobleaching and found to be
15% (Fig. S3, available at
probes. The mean emission ratios corrected for spectral cross talk (A), and
the extracted FRET efficiencies (B) were obtained as described in Materials
and methods. n ? 3. Error bars are the SEM.
Corrected emission ratios and FRET efficiencies of vinculin FRET
on fibronectin. (A) Vinculin null cells expressing a control CFP-YFP chimera,
tail probe, or untagged vinculin were allowed to spread onto 20 ?g/ml of
FN for 2 h at 37?C. Cells expressing untagged vinculin were stained with
Vin11-5 antibody and rhodamine-conjugated secondary antibody. Cells
expressing CFP-YFP and tail probe were examined by GFP fluorescence.
(B–D) The extent of spreading was quantified by measuring the ratio of
major/minor axis of cells (B and D) and cell areas (C and E). (B and C)
Mean of the axial ratio and cell area, respectively. Error bars are the
SEM. (D and E) Box plots (Chase and Brown, 1997) of B and C. Each box
encloses 50% of the data with median value displayed as a horizontal
line. Top and bottom of box represent the limits of ?25% of the popula-
tion. Lines extending from top and bottom of boxes mark the minimum and
maximum values of the data set that fall within an acceptable range.
Open circles denote outliers, points whose value is either ?UQ ? 1.5 ?
IQD or ?LQ ? 1.5 ? IQD (UQ, upper quartile; IQD, inter-quartile dis-
tance; LQ, lower quartile). Asterisks mark histograms that are statistically
different from the corresponding control. The tail probe (n ? 56) or un-
tagged vinculin (n ? 49) reexpressing cells are significantly (P ? 0.01)
more spread than vin?/? cells (n ? 44).
Vinculin tail probe rescues spreading and lamellipodial extension
DETECTING VINCULIN ACTIVATION IN LIVING CELLS • CHEN ET AL. 463
plating. (A and G) Localization of tail probe and control probe in MECs imaged through CFP channel. (D and J) Pseudocolored ratio (FRET/CFP) image of the
cells shown in A and G. (B, E, H, and K) Enlargement of boxed region in A, D, G, and J, respectively. (C and I) The average FRET ratio measured from segmented
regions of cytoplasm or focal adhesions; all segmentable focal adhesions were included. (F and L) Histograms of FRET ratios measured from the segmented focal
adhesions and cytoplasm. Notably, the tail probe gave a much lower average FRET ratio (corresponding to actin-binding conformation of vinculin) in focal
adhesions (B and E, boxed region) than in cytoplasm even though not all focal adhesions are distinguishable from cytoplasm. The control probe did not
distinguish between the two locations (H and K, boxed region). Similar results were obtained by analysis of three other cells from a separate experiment.
Spatial distribution of activated vinculin in living cells. Vin?/? MEC transfected with tail probe (A–F) or control probe (G–L) were imaged 1 h after
JCB • VOLUME 169 • NUMBER 3 • 2005464
Spatial resolution of vinculin
conformations in living cells
When focal adhesions were examined in vin
fected with tail probe, we found that the average FRET ratio is
significantly lower in focal adhesions than in cytoplasm (Fig.
6, A–F), indicating enrichment of the actin-binding conforma-
tion in focal adhesions. In contrast, in vin
with the control probe (YC/V 1–400), the FRET ratio is similar
in focal adhesions and in cytoplasm (Fig. 6, G–L
ratio of tail probe in cytoplasm is comparable to the global
FRET ratio of control probe (Fig. 6, D, F, J, and L), indicating
that vinculin is in the nonactin binding conformation in the cy-
toplasm. Although the actin-binding conformation of vinculin
is enriched in focal adhesions, there were regions of the cell in
which the conformation of vinculin in the focal adhesions was
not readily distinguished from that in cytoplasm (Fig. 6, com-
pare A with D). Similar results were obtained from analysis of
three other cells in separate experiments.
Correlation of vinculin conformation with
To explore the heterogeneity of vinculin conformation in focal
adhesions, we asked whether or not the average conformation
during cell spreading is associated with conformational activation of vinculin.
Vin?/? MECs transfected with tail probe were replated into a dish (Bioptechs)
coated with 20 ?g/ml fibronectin heated at 37?C. (A and B) Images of a
representative cell at the initial attachment stage ?5 min after plating. (C–F)
Images of two representative cells at ?45–60 min after plating. (A, C, and E)
Localization of tail probe in each Vin?/? MEC imaged through CFP channel.
(B, D, and F) Pseudocolored ratio (FRET/CFP) image of the cells shown in A,
C, and E. Notably, the adherent rounded cell (A and B) gave a high FRET
ratio, similar to that of cells containing control probe (Fig. 6), indicating that
vinculin was largely closed in conformation at the earliest stages of spreading.
During spreading, tail probe showed lower FRET ratios correlating with actin-
binding conformation in adhesion structures (C–F).
Recruitment of vinculin to the peripheral belt of adhesion puncta
after plating, a fully spread smooth muscle cell was imaged at time 0, 10, 40,
and 45 min. Images were corrected for photobleaching before calculation of
the FRET ratio image. (A) The positions of the cell at later time points (green)
relative to 0 time point (red) were displayed as color joins of CFP images.
(B) Enlargement of the retraction zone from region 1 in A. Notably, as mature
focal adhesions disassemble, vinculin loses the actin-bound conformation in a
gradient from the tip to the base of the focal adhesions. (C) Enlargement of the
focal adhesion assembly zone from region 2 in A. As focal adhesions mature,
recruited vinculin does not always adopt the actin-bound conformation.
The conformation of vinculin during focal adhesion dynamics. 24 h
DETECTING VINCULIN ACTIVATION IN LIVING CELLS • CHEN ET AL.465
correlated with cellular activity. In vin
tached to fibronectin but have not initiated spreading, vinculin
is uniformly in the nonactin binding conformation (Fig. 7, A
and B). At early phases of isotropic spreading, vinculin is re-
cruited to puncta in the peripheral adhesion ring and to short
central adhesions where it is largely in the actin-binding con-
formation (Fig. 7, C–F). Some cells (Fig. 7, C and D) showed
the nonactin binding conformation of vinculin in a segment of
the adhesion belt puncta (see bottom edge of cell in Fig. 7, C
and D). Unfortunately, phototoxicity associated with acquiring
the FRET image precluded correlating these asymmetric re-
gions with subsequent events in cell spreading.
Fully spread smooth muscle cells were more photoresis-
tant, enabling limited time-lapse analysis in cells that had
spread for 24 h and were undergoing localized, asymmetric cell
shape changes. We found that in retracting/contracting regions
of the cell there is loss of the actin-binding conformation be-
fore loss of vinculin from focal adhesions (Fig. 8, A and B, re-
gion 1, compare 0- and 10-min time points). Interestingly, a
gradient of vinculin conformation can be observed in which the
MECs that have at-
actin binding conformation is found at the proximal edge of the
gliding or disassembling focal adhesion even out to 45 min.
In contrast, a different region of the same cell shown in
Fig. 8 showed recruitment of vinculin to growing focal adhe-
sions (Fig. 8, A and C, region 2). The recruited vinculin was
not in the actin-binding conformation at the times observed.
This result indicates an additional basis for the heterogeneity of
the vinculin FRET ratio in fully spread MECs and also indi-
cates that vinculin recruitment and conformational activation
are separate processes.
Activation of vinculin is required for
assembly of vinculin–vinexin
in focal adhesions
The images of the FRET probe in living cells indicate that vin-
culin can exist in a conformationally open state in focal adhe-
sions. To confirm that the FRET probe reports faithfully on the
conformation of native, endogenous vinculin at focal adhesions,
we took advantage of our finding that vinexin
focal adhesions in vinculin null cells (Fig. S4, available at http://
fails to target to
Vinculin null cells were permeabilized with 0.05% digitonin and incu-
bated with 10 ?g/mL GST-vinexin ? (residues 1–329, encoding full-length
vinexin) and 25 ?g/mL vinculin (A and B) or Vh (C and D) in 25 mM
MES, pH 6.0, 3 mM MgCl2, and 1 mM EGTA. Vinculin localization was
visualized by staining with 5 ?g/mL of 3.24 monoclonal antivinculin,
followed by Rhodamine red-X–conjugated donkey anti–mouse IgG (A and
C). Vinexin was visualized by staining with 5 ?g/mL of polyclonal anti-
GST followed by Oregon green–conjugated donkey anti–rabbit IgG (B
and D). In the presence of full-length vinculin, vinexin ? becomes strongly
enriched in focal adhesions. However, vinexin ? fails to target to focal
adhesions in the presence of Vh, which lacks the polyproline region required
for a direct interaction with vinexin.
Vinculin mediates vinexin ? recruitment to focal adhesions.
regulated. (A) Vinculin, at a 1-?M concentration, was incubated in 20 mM
Pipes, pH 6.9, 100 mM KCl, and 0.1% Triton X-100 with GST-vinexin
(residues 42–115, encoding the first two SH3 domains of vinexin) immo-
bilized on glutathione-agarose beads in the presence of varying amounts
of IpaA. After an overnight incubation at 4?C, supernatant (S) and pellet
(P) were fractionated by centrifugation for 2 min at 10,000 g. The resin
was washed twice with binding buffer before elution in Laemmli sample
buffer. Equal loading of pellets and supernatants represent 10% of total
reaction. Samples were analyzed by SDS-PAGE and Coomassie staining.
(B) Densitometry-based quantification of vinculin–vinexin interaction based
on digitized Coomassie blue–stained gel analyzed in NIH Image. (C)
Coomassie-stained gel of negative controls for binding experiment shown
in A. IpaA was incubated with GST-vinexin in the absence of vinculin,
demonstrating that no direct interaction occurs. Furthermore, the vinculin–
IpaA complex does not co-sediment with GST alone, demonstrating the
specificity of the ternary complex with vinexin.
Vinculin binding to SH3 domains of vinexin ? is conformationally
JCB • VOLUME 169 • NUMBER 3 • 2005466
uses its first and second SH3 domains to bind to the proline-rich
region of vinculin (Kioka et al., 1999). However, we found that,
in vitro, full-length vinculin binds poorly to the SH3(1–2) of
. Upon addition of IpaA, the amount of vinculin bound
to vinexin increased linearly until binding of IpaA to vinculin
reached saturation at ?40% of the vinculin (Fig. 9, A and B).
Thus, the activated conformation of vinculin is required to bind
vinexin ?. We then used this property to confirm the presence of
the activated conformation of vinculin in focal adhesions and to
establish its functional relevance.
Using a permeabilized cell model prepared from vin?/?
MEC, we found that recruitment of exogenous vinexin ? to fo-
cal adhesions was dependent on the presence of vinculin in the
focal adhesions (Fig. 10, A and B). Consistent with a direct in-
teraction between the two proteins, recruitment of vinexin ? to
focal adhesions depends on the presence of the vinexin binding
site because Vh1-851, which lacks the site (Kioka et al., 1999),
was unable to recruit vinexin (Fig. 10, C and D). Because vin-
culin must be activated to bind vinexin ? (Fig. 9, A and B), we
conclude that some of the endogenous vinculin at the focal ad-
hesions must be in the open or activated conformation and that
one functional consequence of this activated conformation is
assembly of vinculin–vinexin ? complexes at focal adhesions.
Development of FRET probes for vinculin
conformation illustrates the value of
modular protein structure
FRET efficiency (E) declines rapidly as the inverse of the sixth
power of the distance between the chromophores (r) according
to the relationship E ? 1 / 1 ? (r6/R06), in which R0 is the cal-
culated Forster distance for a particular donor and acceptor
chromophore pair (Clegg, 1992). Therefore, to construct a
FRET probe with a high FRET efficiency, the chromophores of
CFP and YFP need to be positioned within 2R0 of each other,
where R0 is the distance between donor and acceptor at which
the FRET efficiency is 50%. The calculated R0 for the CFP/
YFP pair is 49 Å (Patterson and Piston, 2000), assuming ran-
dom orientation between donor emission and acceptor absor-
Initially we placed the donor and acceptor GFP variants
at NH2 and COOH termini of vinculin to monitor the confor-
mation of the whole molecule. This construct (CVY) gave a
barely detectable FRET signal due to long distance or unfa-
vorable angle (or both) between donor and acceptor chro-
mophores. The GFP proteins are ? barrels with dimensions of
?40 ? 30 Å and the chromophore sits in the middle of the bar-
rel that has a radius of ?15 Å (Ormo et al., 1996; Yang et al.,
1996). In a FRET pair dimer, the two chromophores are sepa-
rated by ?30 Å, reducing maximum FRET efficiency to 95%.
Based on recent crystal structures of intact vinculin (Bakolitsa
et al., 2004; Borgon et al., 2004) and vinculin domains (Izard et
al., 2004), there is ?40 Å from the amino- to carboxy-terminal
end of vinculin. The very low FRET of the CVY construct is
consistent with the possibility that as much as 70 Å could sepa-
rate the chromophore centers. Thus, the FRET of CVY indi-
cates that the crystal structure is a good representation of the
inactive conformation of vinculin in solution phase.
Because vinculin is a modular protein (Coutu and Craig,
1988; Price et al., 1989), it offered the possibility that self-fold-
ing, single-domain proteins such as the GFP variants could be in-
serted between modules of the protein, as was done for fibronec-
tin (Ohashi et al., 1999). The success of this approach is
facilitated by the fact that the NH2- and COOH-terminal ends of
GFP are close to each other and contain short unstructured re-
gions that can serve as linker sequences (Fig. 1 C). Our data show
that it is possible for GFP modules to be inserted in the loop be-
tween two functional domains of a protein, with appropriate
spacers, without interfering significantly with the functions of the
molecule. Such probes should be especially useful for monitoring
signal-induced functional changes in molecules that lack enzy-
matic activity, such as the structural cytoskeletal proteins and ex-
tracellular matrix molecules. With appropriately characterized
probes it may be possible to study directly how cellular mechani-
cal activity affects the structure and function of the ECM and cy-
toskeletal proteins in living cells, and indeed some pioneering
work in this vein has been done (Ohashi et al., 1999).
The conformation of vinculin in
cytoplasm versus focal adhesions:
validation of the recruitment and
Vinculin is proposed to modulate the junction between ECM
and the actin cytoskeleton by linking an integrin and talin com-
plex to the actin network (Horwitz et al., 1986). Because bio-
chemical data show that vinculin can bind neither talin nor ac-
tin unless the intramolecular interaction between head and tail
is released, the prediction is that vinculin at focal adhesions
must adopt an active conformation. Our live cell data shows a
striking concentration of the actin-binding conformation of
vinculin in focal adhesions as compared with cytoplasm, sup-
porting the central prediction of the aforementioned model for
vinculin recruitment and activation. For cytoplasmic vinculin,
the FRET data allows us to conclude that the conformation of
cytoplasmic vinculin is inactive, at least with respect to its ac-
The conformation of vinculin in focal
adhesions: new insights
Detailed inspection of the cellular FRET data show that the
aforementioned model of recruitment and activation of vincu-
lin at focal adhesions is oversimplified. Specifically, in some
cells, there was polarity in distribution of the actin-binding
conformation of vinculin at focal adhesions. To explore this
heterogeneity we asked whether the average conformation of
vinculin in an adhesive structure correlated with membrane dy-
namics. Although our ability to do sequential FRET captures
on a cell is limited by phototoxicity and photolability issues,
we were able to do limited time-lapse sequences on smooth
muscle cells transfected with tail probe. These images revealed
that the heterogeneity of vinculin conformation in focal adhe-
sions correlated with regional cell dynamics in fully spread
DETECTING VINCULIN ACTIVATION IN LIVING CELLS • CHEN ET AL.467
cells undergoing localized changes in cell shape. Peripheral fo-
cal adhesions were enriched for the actin-binding conformation
of vinculin, but showed a loss of that conformation upon mem-
brane retraction and focal adhesion disassembly. Interestingly,
there was a gradient of vinculin conformation that proceeded
from the distal tip toward the proximal edge of the focal adhe-
sions, with the proximal edge being the last to show the nonac-
tin binding conformation. Thus, one source of the heterogene-
ity of vinculin conformation in focal adhesions is related to
regional retraction of the plasma membrane.
When observing retraction events in a cell it is important
to consider whether one is simply observing global retraction
induced by phototoxicity. Thus, we analyzed only cells that
had regions that were stable or protruding at the same time that
another region was retracting. This analysis resulted in finding
another source of heterogeneity in vinculin conformation. In
stable regions of the cell membrane in which vinculin was be-
ing recruited to focal adhesions, the vinculin remained in the
nonactin binding conformation. Thus recruitment is not neces-
sarily synonymous with actin binding and a second signal or
event must be required to link vinculin to actin.
In previous work, it has been observed that recruitment of
vinculin is correlated with adhesive strengthening (Galbraith et al.,
2002) and with localized application of tension to cell membrane
(Balaban et al., 2001), implying vinculin-mediated strengthening
of connections to the cytoskeleton. Because recruitment of vincu-
lin to focal adhesions and binding of vinculin to actin are not al-
ways coupled events, one can envision that modulating the actin-
binding conformation of vinculin may be a cellular response to
changes in the amount of tension experienced by a focal adhesion.
Although we were not able to adequately determine
FRET in the tiny, very dim focal complexes at the leading edge
of lamellipodia, we were able to analyze vinculin conformation
in spreading cells. Before initiation of spreading, vinculin is
uniformly in the nonactin-binding conformation. But at early
stages of spreading, when a band of vinculin-containing puncta
circumscribes the edge of the spreading cell, vinculin in the
puncta is largely in the actin-binding conformation. To the ex-
tent that this spreading edge mimics an advancing lamellipo-
dium, the result suggests that vinculin in focal complexes at the
leading edge would be in the actin-binding conformation, as
predicted from the work of DeMali et al. (2002).
The significance of vinculin conformation
at focal adhesions
We have presented biochemical and cellular evidence that con-
formational change of vinculin at focal adhesions is function-
ally correlated with ligand binding. Not only does localization
of vinexin ? to focal adhesions require vinculin but this recruit-
ment results from selective binding of vinexin ? to the confor-
mationally open state of vinculin. These data provide direct ev-
idence that the conformation of vinculin regulates focal
adhesion plaque composition by direct protein–protein interac-
tions. Moreover, in establishing that endogenous vinculin also
exists in a distinct ligand-binding conformation in focal adhe-
sions, these data confirm that the vinculin FRET probe reports
faithfully on sites of vinculin activation in living cells.
Although the physiological function of the vinculin–vin-
exin ? complex is unknown, it is interesting that ectopic ex-
pression of vinexin ? stimulates cell spreading in C2C12 cells
(Kioka et al., 1999), as does reexpression of vinculin in vin?/
? cells (Xu et al., 1998b). Given the requirement for activated
vinculin to localize vinexin ? to adhesion sites, it is intriguing
to speculate that integrin-stimulated recruitment and formation
of the vinculin–vinexin ? complex at focal adhesions may be
part of the machinery that links growth factor–stimulated pro-
cesses to cell adhesion.
In summary, the vinculin FRET probe and the vinexin re-
cruitment experiment have enabled us to demonstrate that vin-
culin becomes activated when it gets recruited to plasma mem-
brane and that activation is required for particular protein–
protein interactions at the focal adhesion. These results estab-
lish the relevance of the in vitro biochemical insights to actual
cellular events. In addition, the FRET analysis reveals that, in
vivo, conformational regulation of vinculin is more complex
than the original model (Johnson and Craig, 1995). Specifi-
cally, vinculin’s conformation varies amongst focal adhesions
in a way that correlates with regional membrane dynamics.
This result adds another layer to the heterogeneity of focal ad-
hesions; not only do they vary in the amounts and spatial distri-
bution of components (Zamir et al., 1999) but also in the func-
tional conformation of the vinculin that they contain.
Materials and methods
Reagents and proteins
Actin was extracted from chicken skeletal muscle acetone powder, pro-
cessed through one cycle of polymerization and depolymerization, and
gel filtered through a Sephadex G-150 column according to Pardee and
Spudich (1982). Recombinant 6?-His-tagged chicken vinculin was puri-
fied. GST-vinexin ? was expressed in bacteria and purified on glutathione
agarose (Smith and Johnson, 1988). pCXN2 encoding murine vinculin
was provided by E. Adamson (Burnham Institute, La Jolla, CA). Details on
cloning, expression, and purification of IpaA can be found in the online
Construction of FRET probes
To generate vinculin tail probe, first a 9-bp fragment encoding a NotI site
was introduced by mutagenesis into pEGFP-C1/vinculin cDNA (chicken)
immediately after the codon for aa 883 (Coutu and Craig, 1988) using
the Quick Change kit (Stratagene). The cDNA of EYFP (CLONTECH Labo-
ratories, Inc.) minus the stop codon and flanked by NotI was inserted into
NotI-digested vinculin. pECFP-N3 vector was constructed as an intermedi-
ate vector for generating tail probe. ECFP was PCR amplified with 5?-GGT-
ACCatggtgagcaagggc-3? and 5?-GCGGCCGCTttacttgtacagctc-3? to gen-
erate 5? KpnI and 3? NotI. The product was subcloned into TOPO pCRII
and sequenced. The KpnI and NotI fragment of pCRII/ECFP was used to
replace the EGFP fragment of pEGFP-N3 to generate pECFP-N3. Finally,
the EcoRI–SalI fragment containing vinculin1-883-YFP-vinculin884-1066 was
subcloned into pECFP-N3 to generate tail probe (p ECFP-N3/V1-883 GGR-
To make the control FRET probe pEYFP-C1/CFPV1-400, a HindIII site
was engineered before the ATG site of pET15b/CFP-V1-851 and a KpnI site
after codon 400 by PCR amplification with 5?-CAAGCTTCGatggtgag-
caagggc-3? and 5?-GGTACCTCAtgcaactttccttgc-3?. The PCR product was
introduced into TOPO pCRII and sequenced. The HindIII–KpnI fragment of
CFP-vinculin1-400 was subcloned into pEYFP-C1 to generate the control
Cell culture and transfection
Cells were cultured on 0.1% gelatin-coated tissue culture plates in DME
with high glucose and glutamine (MediaTech) supplemented with 10%
FCS in a 5% CO2 incubator at 37?C. For cell imaging and FRET analysis,
vin?/? MECs, isolated from embryo #54?/?, were cultured with home-
JCB • VOLUME 169 • NUMBER 3 • 2005468
made phenol red–free DME (same as aforementioned DME except phenol
red–free, 4750 mg/l NaCl, 370 mg/l NaHCO3, 5958 mg/l Hepes, and
one-fourth the concentration of vitamins). These media modifications re-
duced background and autofluorescence. HEK 293 cells were seeded on
0.1% gelatin-coated 100-mm dishes at 3 million per plate; transfection
was performed the next day with 3 ?g of plasmid DNA using Lipo-
fectAMINE/Plus reagent (Invitrogen). HEK 293 cells were lysed 2 d after
transfection. Vin?/? MECs were seeded on 20 ?g/ml of fibronectin-coated
35-mm tissue culture dish at 120,000 cells; transfection was performed
the next day with 1 to ?1.5 ?g of plasmid DNA using LipofectAMINE/
Cell spreading assay
Vin?/? MEFs were transfected with tail probe, vinculin, or CFP-YFP chi-
mera. 24 h after transfection, ?400,000 cells were seeded on coverslips
coated with polylysine and 20 ?g/ml of human fibronectin and incubated
in 10% FCS/90% DME (MediaTech) at 37?C for 2 h. Cells transfected
with tail probe and CFP-YFP chimera were fixed in 4% PFA in PBS for 20
min, washed twice with PBS, and mounted on a slide with Prolong Gold
antifade reagent (Molecular Probes). For cells transfected with untagged
vinculin, coverslips were fixed and immunostained with Vin11-5 (Sigma-
Aldrich) and rhodamine-conjugated donkey anti–mouse IgG (Jackson Im-
munoResearch Laboratories). Axial ratios and cell areas of transfected
cells were measured using the segmentation and quantitation tools in IP-
Lab (Scanalytics). Multinucleated cells were excluded from the analysis.
FRET assay of cell lysates
HEK 293 cells were detached with 1 mM EDTA in calcium- and magne-
sium-free PBS at 37?C for 20 min. The pelleted cells were resuspended in
ice-cold hypotonic buffer (20 mM Tris, pH 7.5, 2 mM MgCl2, 0.2 mM
EGTA, 0.5 mM ATP, 0.5 mM DTT, and 2? protease inhibitor cocktails I
and II [Siliciano and Craig, 1986]) at a density of 2 to ?4 ? 106 cells/
ml, incubated on ice for 20 min, and homogenized manually for 5 min in
a DUALL 21 conical ground glass homogenizer (Kontes Glass Co.). The ly-
sate was cleared by centrifugation at 4?C, 16,000 g for 10 min. The hy-
potonic lysate was supplemented with KCl to a final concentration of 100
mM for fluorimetric and actin sedimentation assays. The emission spec-
trum of fluorescent proteins in the lysate was acquired with a Fluoromax-3
spectrofluorimeter (Jobin Yvon). CFP emission was traced from 460 to 600
nm with excitation at 440 nm, and YFP emission was traced from 510 to
600 nm with excitation at 490 nm. The increment was 1 nm and integra-
tion was 0.2 s. The excitation and emission slit widths were 3 and 5 mm,
respectively. Lysate from an equal number of untransfected HEK293 cells
was used to obtain a background emission spectrum. After subtraction of
background, the spectra comprising a single experiment were normalized
to the CFP emission of a reference spectrum, as specified in the figure leg-
Determination of the corrected FRET emission ratio and FRET efficiencies
To obtain a number for the corrected FRET emission ratio (a value related
to FRET efficiency by Eq. 5) and an estimate of the FRET efficiency itself (as
reported in Fig. 4), the sensitized YFP emission (SE) due to FRET was ex-
tracted from the raw FRET signal. The raw FRET signal is the EYFP emission
(peak at 525) stimulated by excitation of ECFP at 440 nm. It consists of
the SE, the emission from direct excitation of EYFP by 440 nm, and the
overlap of the ECFP emission spectrum with the EYFP emission spectrum
(Erickson et al., 2001). The latter two components of the raw FRET signal
are referred to as “spectral cross talk.”
To determine the amount of spectral cross talk contributed by di-
rect excitation of EYFP by excitation at 440 nm, the value RY was deter-
mined from a sample containing just EYFP by the ratios of the emissions
at 525 nm after excitation at 440 and 490 nm. RY was 0.11 in this
study. The EYFP emission at 525 nm of the FRET probe after excitation at
490 nm was then multiplied by RY to obtain the YFP cross talk compo-
nent. The YFP cross talk at 525 nm was subtracted from the raw FRET
emission at 525 nm to correct for direct excitation of EYFP in the FRET
probe by 440-nm excitation.
To determine the contribution of CFP emission to the signal at 525
nm after excitation of the FRET probe at 440 nm, the value RC was deter-
mined from a sample containing just ECFP by ratioing the emission at 525
nm to that at 475 nm after excitation at 440 nm. RC was 0.43 in this
study. The corrected FRET emission ratio (ER in Eqs. 4 and 5) is the emis-
sion at 525 nm/emission at 475 nm, after correcting for the EYFP and
ECFP cross talk. Corrected FRET emission ratio is SE/FDA. Corrected FRET
emission ratio correlates with FRET efficiency; the higher the corrected
FRET emission ratio, the stronger the FRET. The expression for FRET effi-
ciency (E%), E% ? (FD ? FDA)/FD (Miyawaki and Tsien, 2000), is trans-
formed to Eqs. 1–5 to express FRET efficiency in terms of SE, FDA, and the
quantum efficiencies of EYFP and ECFP, which are the experimentally de-
FD and FDA are the donor ECFP emission in the absence or presence of ac-
ceptor EYFP, respectively. Because FD ? FDA/Qc ? SE/QY, the net FRET
(nF) ? (FD ? FDA) can be approximated as SE ? QC/QY. FD, the fluores-
cence of the donor, is approximated as nF ? FDA. QY and QC are the
quantum efficiencies of EYFP and ECFP. QY is 0.7 and QC is 0.4 (Gries-
beck et al., 2001). The quantity SE/FDA is measured as the corrected emis-
sion ratio (ER) described above.
Fluorescence microscopy and image processing
2 d after transfection, vin?/? MECs were detached with trypsin and re-
seeded on a 20 ?g/ml of fibronectin-coated delta T dish (Bioptechs) equil-
ibrated to 37?C. Images were captured at 37?C with a fluorescence micro-
scope (model Axiovert 135TV; Carl Zeiss MicroImaging, Inc.) equipped
with a stage and objective heater (Bioptechs), a Cool SNAP HQ camera
(Photometrix), and Chroma filters. We used a 100? Plan Neofluor objec-
tive (Carl Zeiss MicroImaging, Inc.) with an NA of 1.3 and collected the
images with 2 ? 2 binning. The excitation filter used for CFP was D436/
10 nm. The emission filter used for CFP was D470/30 nm. The emission
filter used for YFP was HQ535/30 nm. The beam splitter used was JP4
PC. FRET images were captured with the CFP excitation filter and YFP
emission filter. Manipulations of numerical files were done using IPLab
software (Scanalytics). Background images of CFP and FRET channels
were captured from areas lacking cells on the experimental dish, using the
same exposure times as for acquisition of the cell images.
The matched background images were subtracted from the fluores-
cent images to remove background and uneven illumination. After regis-
tration of the images, an empirically determined arithmetic averaging filter
(blurring) was applied to the CFP and FRET images. This was done to min-
imize the presence of artificially high pixel ratios at the edges of focal ad-
hesions. This artifact arises in part because the Airy disc of the FRET image
is 11% larger than that of the CFP image. The pixel size on the CoolSnap
HQ camera is 6.45 ?m. In a 2 ? 2 binned image, the diffraction limited
spots at 100? for the objective are 3.9 pixels for YFP image and 3.4 pix-
els for the CFP image; these values are close to the width of the focal ad-
hesions. Thus, a 3 ? 3 or 5 ? 5 pixel averaging filter was used to mini-
mize the artifact generated by ratioing two different sized images (the
FRET and the CFP image) of the same focal adhesions. Empirical selection
of the filter was made by comparing the histograms of line segments
drawn perpendicular to the long axis of the same focal adhesion region in
both the CFP and FRET images. The averaging filters were adjusted such
that the shapes of these histograms were as closely congruent as possible.
After the aforementioned manipulations, an image of the FRET ratio
at each pixel (FR; Miyawaki and Tsien, 2000) was obtained by arithmetic
manipulations of the CFP and FRET images according to the equation FR ?
IFRET(probe)/ICFP(probe). IFRET(probe) and ICFP(probe) denote the fluorescent
intensity of FRET and CFP images at each pixel.
To determine the position of the cell edge, a threshold was selected
empirically in the registered CFP image such that it included most of the
cell boundary but excluded stray light at the cell edges. The segmentation
and analysis tools in IPLab were used to estimate this threshold from the in-
flection point in the slope of a plot of pixel intensity versus pixel number
along a line segment made perpendicular to the cell membrane. The sub-
---------------------- - =
DETECTING VINCULIN ACTIVATION IN LIVING CELLS • CHEN ET AL.469
threshold region will appear white in a pseudocolor ratio image, the same
as background. The segmented CFP image was converted to a binary
mask such that all pixels above the threshold were assigned a value of
one and all those below were assigned a value of zero. The FRET ratio im-
age was multiplied by the binary mask to remove pixels that had an artifi-
cially high ratio. The final ratio image was pseudocolored by assigning
color values to the ratios. A linear scale from blue (low ratio and activated
vinculin) to green/red (high ratio and closed vinculin) was constructed
and a ? of 0.75 was applied to the display. The background and sub-
threshold regions are color-coded white in the final FRET images.
To differentiate cytoplasm from focal adhesions for separate quanti-
tation, two segmented images were generated, one for cytoplasm and
one for adhesions. Image segmentation was performed in the registered
CFP image. Each segmented image was converted to a binary image with
the segmented region assigned a value of 1 and nonsegmented region as-
signed a value of zero. The FRET ratio image was then multiplied by each
binary mask image to generate a FRET ratio image for focal adhesion or
Permeabilized cell assay
Vin?/? MECs were cultured on glass coverslides coated with PLL and 20
?g/ml FN. After 16-h growth in 10% FCS/90% DME, the cells were
washed briefly in assay buffer (5 mM MES, pH 6.1, 2 mM MgCl2, 0.5
mM CaCl2, 137 mM NaCl, 5.4 mM KCl, 4.2 mM NaHCO3, and 0.1%
glucose) and extracted for 1 min with ice-cold assay buffer plus 0.05% ul-
trapure digitonin (Calbiochem). After a 30-s rinse in ice-cold assay buffer,
coverslips were incubated with the indicated proteins for 15 min at 4?C.
Vinculin and Vh (residues 1–851) were used at a concentration of 25 ?g/
ml and GST-vinexin (residues 1–329, encoding full length vinexin) at 10
?g/ml in 25 mM MES, pH 6.0, 3 mM MgCl2, and 1 mM EGTA. After two
wash steps, cells were fixed in 4% PFA in PBS and stained with a mono-
clonal antivinculin and affinity-purified anti-GST antibodies. Oregon green
donkey anti–mouse and RRX donkey anti–rabbit antibodies were used for
Online supplemental material
Fig. S1 shows immunoblots of cell lysates demonstrating integrity of FRET
probes. Fig. S2 shows that an SDS-resistant structure in GFP causes aber-
rant migration in SDS-PAGE. Fig. S3 illustrates FRET in living cells as de-
tected by the acceptor photobleach method. Fig. S4 demonstrates that vin-
culin is required for the recruitment of vinexin to focal adhesions in
vinculin null cells. The protocol for cloning, expression, and purification of
IpaA can be found in the online supplemental material. Online supple-
mental material is available at http://www.jcb.org/cgi/content/full/
We thank Dr. Douglas Murphy, Director of the Johns Hopkins University Mi-
croscope Facility, for help with setting up hardware and software for FRET im-
aging, Dr. Eileen Adamson for the vin?/? cells, Dr. Jim Bear (University of
North Carolina, Chapel Hill, NC) for advice on live cell imaging, Dr. David
Shortle for use of the fluorimeter, and Drs. Chris Janetopoulos, Peter Devreotes,
Mike Erickson, David Yue, Thomas Hofmann, and Hiro Okuno for helpful dis-
cussions on FRET.
This work was supported by an American Heart Association Postdoc-
toral Fellowship to H. Chen, Howard Hughes Medical Institute Predoctoral
Fellowship to D. Cohen, and National Institutes of Health grant GM41605 to
Submitted: 19 October 2004
Accepted: 18 March 2005
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