Assay to mechanically tune and optically probe fibrillar fibronectin
conformations from fully relaxed to breakage
William C. Little, Michael L. Smith, Urs Ebneter, Viola Vogel⁎
Department of Materials, ETH Zurich, CH-8093, Zürich, Switzerland
Received 21 September 2007; received in revised form 4 February 2008; accepted 5 February 2008
In response to growing needs for quantitative biochemical and cellular assays that address whether the extracellular matrix (ECM) acts as a
mechanochemical signal converter to co-regulate cellular mechanotransduction processes, a new assay is presented where plasma fibronectin
fibers are manually deposited onto elastic sheets, while force-induced changes in protein conformation are monitored by fluorescence resonance
energy transfer (FRET). Fully relaxed assay fibers can be stretched at least 5–6 fold, which involves Fn domain unfolding, before the fibers break.
In native fibroblast ECM, this full range of stretch-regulated conformations coexists in every field of view confirming that the assay fibers are
physiologically relevant model systems. Since alterations of protein function will directly correlate with their extension in response to force, the
FRET vs. strain curves presented herein enable the mapping of fibronectin strain distributions in 2D and 3D cell cultures with high spatial
resolution. Finally, cryptic sites for fibronectin's N-terminal 70-kD fragment were found to be exposed at relatively low strain, demonstrating the
assay's potential to analyze stretch-regulated protein-rotein interactions.
© 2008 Elsevier B.V./International Society of Matrix Biology. All rights reserved.
Keywords: Fibronectin; Protein unfolding; Mechanotransduction; Mechanochemical signal conversion; Fluorescence resonance energy transfer; FRET
In addition to soluble biochemical cues such as growth
factors and cytokines, cells are also known to sense and respond
to physical factors, including the mechanical properties of their
environment (as reviewed in Discher et al., 2005; Ingber, 2006;
Vogel and Sheetz, 2006) and the local topography of their
substrates (Dalby et al., 2004; Dalby et al., 2005). Differences in
substrate rigidity (Engler et al., 2006; Galbraith et al., 2002;
Giannone and Sheetz, 2006; Kostic and Sheetz, 2006; Yeung
et al., 2005) as well as micro- and nanoscale features (Dalby
et al., 2005) have been correlated with profound effects on cell
adhesion, morphology, proliferation, and gene regulation. The
rigidity of a substrate, for example, determines whether or not
mammary epithelial cells up-regulate integrin expression and
differentiate into a malignant phenotype (Paszek et al., 2005),
and also dictates whether mesenchymal stem cells differentiate
into bone, muscle, or neuronic tissue (Engler et al., 2006). Yet,
the underlying molecular mechanisms by which physical fac-
tors, including force and the mechanical properties of matrices,
are converted into biochemical signals that ultimately regulate
protein expression are only beginning to be explored (Kostic
and Sheetz, 2006). One focus is to identify proteins whose
structure/function relationship can be altered by mechanical
forces, causing for example the opening of ion channels, a
change in the spatial presentation of binding sites, or the
exposure of otherwise cryptic binding sites (for reviews see
Bustamante et al., 2004; Kung, 2005; Vogel, 2006; Vogel and
Sheetz, 2006; Arcangeli and Becchetti, 2006).
On the intracellular side, mechanochemical signal conver-
sion through protein unfolding has been demonstrated. The
phosphorylation sites of p130Cas become exposed by tensile
Available online at www.sciencedirect.com
Matrix Biology 27 (2008) 451–461
Abbreviations: ECM, extracellular matrix; Fn, Fibronectin; FRET, fluores-
cence resonance energy transfer; Fn-d/a, Fn labeled with donor and acceptor
fluorophores; GdnHCL, guanidine hydrochloride; IA/ID, ratio of the acceptor
intensity divided by the donor intensity; u-Fn, unlabeled Fn.
⁎Corresponding author. Department of Materials, Wolfgang Pauli Strasse 10,
HCI F 443, ETH Zurich, CH-8093, Zürich, Switzerland. Tel.: +41 44 632 0887;
fax: +41 44 632 10 73.
E-mail address: email@example.com (V. Vogel).
0945-053X/$ - see front matter © 2008 Elsevier B.V./International Society of Matrix Biology. All rights reserved.
force (Sawada et al., 2006) which leads to the downstream
activation of Rap1 upon stretch (Tamada et al., 2004). Also, two
isoforms of the cytoskeletal protein spectrin within red blood
cells showed enhanced exposure of otherwise cryptic cysteines
as a function of shear stress and time, as did nonmuscle myosin
IIA and vimentin within mesenchymal stem cells (Johnson
et al., 2007).
On the extracellular side, fibronectin (Fn) can be unfolded by
cell generated tension which causes at least a partial loss of
secondary structure of its domains (Baneyx et al., 2002; Smith
et al., 2007). Fn exists in a broad range of conformations, once it
is harvested by cells from solution and incorporated into their
own matrix as revealed by fluorescence resonance energy
transfer (FRET) (Baneyx et al., 2002; Barker et al., 2005; Smith
et al., 2007).
Deciphering the physiological significance of force-induced
unfolding events of Fn and whether any of Fn's versatile
functions are up- or down-regulated in a mechanoresponsive
manner has been hampered due to the lack of well suited assays.
Fn plays a critical role in a wide variety of processes including
embryogenesis and wound healing (Hynes, 1990; Midwood et
al., 2006; Pankov and Yamada, 2002; Sechler and Schwarz-
bauer, 1998), and its fibrillogenesis into matrix fibers is tightly
regulated (for reviews see Mao and Schwarzbauer, 2005;
Wierzbicka-Patynowski and Schwarzbauer, 2003). This large
440kDmultimodulardimericprotein contains multiple surface-
exposed molecular recognition sites and cryptic binding sites
buried in the fully folded module that regulate fibrillogenesis
and mediate cell adhesion, as well as interactions with other
ECM proteins, and proteolytic activity (Magnusson and
Mosher, 1998; Mao and Schwarzbauer, 2005; Pankov and
Yamada, 2002; Romberger, 1997; Vogel, 2006). First indica-
tions also exist that downstream cell signaling is impacted by
the conformation of Fn (Keselowsky et al., 2005; Lan et al.,
2005). When adsorbed from solution to biomaterial surfaces, its
conformation is for example impacted by the underlying
surface chemistry (Altroff et al., 2003; Antia et al., 2006;
Baugh and Vogel, 2004; Garcia et al., 1999; Halter et al., 2005)
which can then alter integrin binding (Keselowsky et al., 2005).
The conformations Fn assumes on synthetic surfaces, however,
are rather distinct from those of native Fn in soluble and
fibrillar forms (reviewed in Antia et al., 2006; Baugh and
Vogel, 2004; Halter et al., 2005). Fn polymerization into fibers
is thought to proceed as cryptic self-assembly sites are ex-
posed by cell contractility (Baneyx and Vogel, 1999; Mao and
Schwarzbauer, 2005; Ohashi et al., 1999; Zhong et al., 1998).
Single-molecule force-spectroscopy experiments provided the
first indications that Fn might serve as a mechanochemical signal
converter. Major insights into force-induced unfolding pathways
are available (Oberhauser et al., 2002; Samori et al., 2005; Wang
et al., 2001), as well as high-resolution structural models relating
Craig et al., 2004b; Craig et al., 2001; Vogel, 2006).
Native, cell-derived Fn matrices, however, are not ideal
systems for investigating force-induced potentially altered
structure–function relationships. A key complication is the
complex and interwoven nature of ECM fibers, where the
conformation of Fn can vary from fiber to fiber and even within
fibers as they are not freely suspended (Baneyx et al., 2001;
Baneyx et al., 2002; Chen et al., 1997; Peters et al., 1998).
Our aim was thus to develop a mechanical strain assay where
the conformation of Fn can be adjusted externally on demand
and force-induced Fn extensions can be optically measured. To
probe how the alterations of the structure of stretched Fn impact
the displayed biochemistry and cellular behavior, such a strain
assayneedstobeamenable tocellculture environments.Totune
the conformation of Fn, Fn fibers were drawn from a
concentrated Fn solution (Ahmed and Brown, 1999; Ejim et
al., 1993; Wojciak-Stothard et al., 1997) and were deposited
onto a stretchable substrate which was mounted onto a custom-
built, one-dimensional strain device.
2. Results and discussion
2.1. Manually deposited Fn fibers bundle into fiber cables
similar to cell-derived fibers and promote cell adhesion and
While it has been reported previously that manually deposited
fibers pulled from concentrated solutions of soluble Fn resemble
in vivo Fn fibers in diameter and composition (Wojciak-Stothard
et al., 1997), the latter was difficult to confirm because the exact
conformation and the nature of the lateral interactions between
adjacent Fn molecules in fibers is unknown (Mao and
Schwarzbauer, 2005). To briefly characterize the morphology of
our deposited fibers, it should be noted that the average diameter
as observed via fluorescence confocal microscopy, is similar to
matrices (Chen et al., 1978). The average fiber diameter can be
moderately adjusted via the pulling procedure. Our fibers were
typically between 2 and 5 μm in diameter when the fibers were
deposited out of 0.2 mg/mL or 2.6 mg/mL Fn solutions,
respectively. Also the length of a fiber can be controlled by
contact with the PDMS substrate. Fibers used in this study were
generally 1 to 2 cm long. Fibronectin fibers in 24-h Fn matrix
producedby NIH-3T3 fibroblasts incellculture forma mesh-like
network where smaller fibrils merge to form bundles and later
branched again (Chen et al., 1978).
Similarly, fluorescence images of manually deposited fibers
(Fig. 1) show that submicron fibers emerge from the surface of
the drop and bundle together to form larger cables of fibrils
(Fig. 1B,C), in agreement with previously published images of
such artificial Fn fibers (Ejim et al., 1993; Wojciak-Stothard
et al., 1997). Indeed, cryo-scanning electron microscopic
images suggest that Fn fibers exist as ‘cables’ comprised of
individual fibrous strands of ~5–15 nm in diameter and larger
(Chen et al., 1978; Dzamba and Peters, 1991; Peters et al., 1998;
Singer, 1979) which are proposed to be held together by hy-
drogen bonds, intermolecular beta-strand swapping (Briknarova
et al., 2003; Litvinovich et al., 1998), disulfide bonds which are
potentially formed by cryptic disulfide isomerase activity
452 W.C. Little et al. / Matrix Biology 27 (2008) 451–461
(Langenbach and Sottile, 1999), and other weak electrostatic
interactions (Chen and Mosher, 1996; Morla et al., 1994).
Although the exact location and properties of these bonds are
unknown, it has been observed that they are strong enough to
render cell-derived Fn fibers irreversibly insoluble in 1% de-
oxycholate (McKeown-Longo and Mosher, 1983), which is a
phenomenon we observed with manually deposited Fn fibers as
well (data not shown).
Furthermore, Human Foreskin Fibroblasts (HFFs) and
Human Umbilical Vein Endothelial Cells (HUVECs) were ob-
served to adhere and polarize along the axis of the manually
deposited fibers (Fig. 1E,F), a phenomenon shown previously
by others (Ahmed and Brown, 1999; Ahmed et al., 2000;
Wojciak-Stothard et al., 1997), confirming that cell adhesion
sites are exposed on the surface of the fibers.
2.2. Establishing a quantitative FRET vs. strain curve to read
out Fn extensions in 2D and 3D matrices
To conduct a conformational FRET analysis of manually
deposited Fn fibers, plasma Fn dimers were chemically
denatured and site-specifically labeled with acceptors on the
four native and free cysteines (buried within type III7and III15
domains of the Fn dimer (Lai et al., 1984; Mosher and Johnson,
1983; Smith et al., 1982)) and randomly labeled with donors on
free lysines (Fn-d/a, see Materials and methods). Circular
dichroism spectroscopy of FRET-labeled Fn confirmed that
refolding occurs in physiological buffer (Smith et al., 2007).
Ratiometric FRET images of manually deposited fibers show
little color change along a single fiber (visualized as false colors
in Fig. 2B), whereas fibers from a 24-h cell-derived matrix
(Fig. 2C) show more locally heterogeneous conformations
(Fig. 2D). The frequency at which a given ratio between the
intensity of the acceptor channel and the donor channel (IA/ID
ratio) is found within a field of view containing a total number of
pixels (640 pixel×640 pixel) is displayed in the FRET histo-
grams (Fig. 2E).The histogram forthemanually depositedfibers
peaks at an IA/IDof 0.62, which corresponds to the mean value
recorded from Fn in solution at ~1 M GdnHCl. Histograms for
each of 11 different pulled fibers are provided in Supplementary
Fig. S1B. Variations of the pulling rate by which the fibers were
drawn out of solution, from 1.0 mm/s to 0.2 mm/s, had no major
impact on the conformational distribution (data not shown). The
width of the histogram originates from the pixel-to-pixel vari-
noise (as further discussed in Smith et al., 2007). The widths of
the histograms taken from Fn-d/a in solution (1 M GdnHCl;
data not shown) and in manually deposited fibers (Fig. 2E) were
To stretch Fn fibers, a one-dimensional strain device was
constructedtoholdathinrectangularsiliconesheet andstretch it
over four times its length (Fig. S2A-E, see Materials and
methods). Fibers are first deposited onto the silicone sheet
(Fig. S2A), rinsed and incubated with a PBS or BSA solution,
and then stretched to a desired length (Fig. S2B). A holding
chamber is then used to sandwich and capture the strained
substrate (Fig. S2D) before it is removed from the strain device.
Finally, the bottom half of the chamber is removed after the ends
of the substrate are fastened to the side (Fig. S2E) in order to
allow a microscope objective to reach the substrate for imaging.
Fibers can also be first deposited on pre-strained substrates such
as in Fig. S2B and relaxed to a position such as in Fig. S2A to
remove any residual tension that may exist as they are originally
deposited from solution. Shown in Fig. 3A is the resulting IA/ID
histograms from relaxed, strained, and control fibers as they
comparetocell-derived 24-hFnmatrices, andFig.3Bshowsthe
corresponding color and relative length-changes that are pos-
sible to achieve with the strain device.
Thus, in order to begin establishing an absolute IA/IDvs.
strain curve, we first deposited Fn fibers on silicone sheets that
were pre-strained to 4×. Subsequent relaxation of the fibers led
to a uniform increase of their IA/IDratios which indicated pro-
gressively more refolding of fibronectin until the sheet has been
relaxed to about 1/3 of its initial length (Fig. 3C). With further
sheet relaxation, the fibers began to bulge which likely explains
the slightly lower IA/IDvalues observed at 1/4 the substrate
length (Fig. 3C).
The 1/3× relaxation point shown in Fig. 3C represented the
amount of strain release leading to maximal FRET recovery.
Fig. 1. Fabricationand characterization of manually depositedfibronectin fibers.
(A) A pipette tip is submersed slowly into a concentrated solution (0.76 mg/mL)
of fibronectin (Fn) and removed to generate polymerized Fn fibers that were
deposited onto stretchable silicone sheets. (B) Differential Interference Contrast
(DIC) and (C) fluorescence images of a fiber as it extends away from the droplet
(upper-left corner) reveal a bundling behavior that is confirmed via (D) a
Scanning Electron Micrograph taken along the length of the fiber. (E) DIC
image of Human Foreskin Fibroblasts (HFFs) adhered to and oriented along the
length of a fiber. (F) 3D-confocal microscopy reconstruction of a Human
Umbilical Vein Endothelial Cell (HUVEC) spread on a manually deposited
fiber: the cell is fluorescently labeled with a CellTrace™ Far Red dye
(Invitrogen) andculturedfor 1 hafter seedingonto an AlexaFlour® 488-labeled
fiber. A dual-channel z-stack and Imaris™ software (Bitplane) were used to
render the image shown. Grid marks represent 10 μm. The image is representative
ofcells onmanuallydeposited fibers andconfirms thatcell adhesionoccurs onthe
top and sides of a fiber but not onto the underlying substrate.
453 W.C. Little et al. / Matrix Biology 27 (2008) 451–461
After having determined that the drawing process by which
the fibers are pulled out of the droplet causes considerable pre-
straining of fibronectin, for which we have to compensate by
relaxing the substrate, we can now establish an absolute
extension versus FRET curve for fibronectin fibrils (Fig. 3E).
This is significant because it allows us to read out the strain and
thus the conformation of fibronectin by optical means, whether
the cells are cultured in two or three dimensions.
2.3. Fully relaxed fibers do not contain the globular Fn
conformation found in solution
When the Fn fibers are fully relaxed, the average IA/IDratio
is 0.74±0.02, which lies between the 0 M (0.98) and 1 M (0.64)
GdnHCL calibration points, indicating that fully relaxed
fibrillar Fn does not return to its globular solution conformation
where its two dimer arms are folded upon themselves. These
results confirm previous data obtained with fully relaxed cell-
derived fibronectin matrices which also do not contain the
tightly folded quaternary structure otherwise seen in solution
(Smith et al., 2007). These findings thus invalidate a recent
model postulating that relaxed fibronectin fibers are composed
of arrays of compactly folded fibronectins in which the
opposing dimeric arms cross each other (Abu-Lail et al.,
2005; Erickson, 2002).
2.4. Fn fibers show signs of breakage only when stretched over
5–6 times their resting length
To establish the extension and the associated optical
signature at which the fibers begin to rupture, Fn fibers were
deposited onto unstrained silicone sheets (Fig. S2A). After
applying uniaxial strain, the observed IA/IDratios decreased as
expected (Fig. 3C), and beginning at a 1.8× length increase, the
fibers began to fracture at a few sites into randomly distributed
large segments, each hundreds of micrometers long. In the 2×
strain field of the view shown in Fig. 4A (as calibrated using data
shown in Fig. 3C and 4B), a magnified image of one of the
breaks (Fig. 4C,D) and a contrast-enhanced fluorescence image
(Fig. 4E), shows that a small amount of Fn is left attached to the
substrate. Since the IA/IDvalues continued to decrease along the
length of the fiber segments when further strained until the limit
Fig. 2. Spatially resolved conformational mapping by Fluorescence Resonance Energy Transfer (FRET) and comparison of FRET histograms of manually deposited
fibronectinandextracellularmatrixfibers of living fibroblasts.(A) Plasma fibronectin, dual-labeledwithdonorsandacceptors (Fn-d/a—seeMaterialsandmethods),is
subjected to increasing concentrations of guanidine hydrochloride (GdnHCL) to calibrate a ratio of the acceptor intensity divided by the donor intensity (IA/ID) to
known conformational states of fibronectin in solution. In concentrations greater than 1.5 M GdnHCL, circular dicroism indicates that fibronectin undergoes a rapid
and significant loss of secondary structure causing the Fn modules to unfold (Alexander et al., 1979; Baugh and Vogel, 2004). (B) Reproducibility of conformational
distribution: 11 manually deposited Fn fibers containing 5% Fn-d/a were laid onto a transparent sheet of silicone and the pixel-resolved IA/IDvalues were mapped to a
color gradient representing module-unfolded (blue) or dimer-arm separated (red) Fn conformations. Images of individually deposited fibers here were mounted next to
each other for presentation purposes (though it is possible manually, see Fig. 5). (C) A fluorescence image of a 24-h cell-derived Fn matrix derived from NIH-3T3
fibroblasts (Baneyx et al., 2001) is shown beside the (D) corresponding false color FRET map. (E) The top histogram represents IA/IDdata collected from every pixel
analyzed from the eleven fibers shown in (B). The middle histogram shows the IA/IDdata collected from the single fiber in (B) as denoted by the⁎. The bottom
histogram shows the IA/IDdata collected from the cell-derived Fn matrix image in (D). The absolute strain scale represents the average IA/IDvalues of Fn fibers, from
fully relaxed (nodules of quatenary structure are present), to extended, to unfolded, to the point where they begin to break (5.4×), as described in the text and in Fig. 3.
Scale bar in (B) is the same scale as the images in (C) and (D).
454 W.C. Little et al. / Matrix Biology 27 (2008) 451–461
ofthe straindevice was reached at4× (Fig. 3C), thismay indicate
that the adhesion strength of the fiber bundle to the silicone
substrate is considerable and does not allow for major slippage
between the strained fiber and the silicone sheet once the first
breakages have occurred. Atomic force microscopy (AFM)
images of fiber breakage areas reveal two kinds of breaks; one
where a small layer of fibronectin remains attached to the
underlying silicone sheet and the broken fiber ends appear
tapered (Fig. 4F), and another where the break results in
relatively blunt fibers and the fibronectin layer left behind exists
closer to the fractured ends (Fig. 4G). Finally, the IA/IDvalues
continued to decrease homogenously along the length of the
fibrous segments upon further strain until the limit of the strain
device was reached at 4× (Fig. 3C).
Bringing the fiber strain and relaxation data together,
mechanically stretched Fn-fibers began to show signs of
breakage at a relative length increase of 1.8× compared to
fibers deposited onto unstrained silicone sheets, and can thus be
strained at least 5–6 times with respect to their resting state
(5.4=1.8 / 0.333). It is currently believed that Fn fibers can be
stretched three- to four-fold beyond their resting length (Ohashi
et al., 1999). This conclusion was derived from various cell
culture measurements of the lengths of chimeric GFP or YFP-
Fn fibers before and after breakage or detachment of fibers
under cell-dependent strain, from fiber rupture from laser
damage, or after inhibition of cell contractility with cyto-
chalasin B. Since physisorption to a substrate might have an
impact on strain at which the fibers break, future work is needed
to determine the extension at which freely suspended fibers
rupture. The FRET ratios seen at the onset of fiber fracture,
however, are also seen in native cell-made matrix (see Fig. 2D).
2.5. High extensions of Fn fibers involve
These data on the total range of extensibility of manually
deposited Fn fibers further support recentconclusions thatnative,
cell-derived Fn matrices are unfolded by cell traction forces
(Baneyx et al., 2001; Baneyx et al., 2002; Smith et al., 2007).
FRET measurements of dual-labeled Fn matrices supported a
model whereby Fn fiber elasticity involves unfolding of typeIII
modules (Baneyx et al., 2001; Baneyx et al., 2002). The
suggested based on purely theoretical calculations (Erickson,
quaternary structure in resting fibers, and that fibril elasticity
originates from a separation of the overlapping dimer arms
(quaternary structure) followed by an extension into a straight
string of modules (loss of quaternary but not of secondary
structure) (Abu-Lail et al., 2005). If one assumes that module
unfolding does not occur prior to fiber breakage, this quaternary
structural model only allows for a maximum 4-fold extension
from compactly folded (~40 nm) to extended (~130 nm;
Fn fibers can be stretched 5- to 6-fold prior to the onset of fiber
breakage. Furthermore, fully relaxed manually deposited Fn
Fig. 3. FRET vs. strain dependency of manually deposited fibronectin fibers probed by externally stretching a supporting silicone sheet. (A) Histograms of pixel-
resolved acceptor channel / donor channel ratios (IA/ID) from representative fibers shown in (B) and cell-derived fibers from Fig. 2D,E containing 5% dual-labeled
fibronectin. The absolute strain scale represents the average IA/IDvalues of Fn fibers that are fully relaxed or extended to the point where they begin to break (5.4×), as
describedin the text andshownin (C). Thefibers are subjectedto 1/4× releaseof strain (orange), no strain (green),or 4× applied strain(blue) and showthat distinct and
uniform conformational changes can be induced via mechanical strain and each mean IA/IDcan be found within the histogram of cell-derived fibers. For presentation
purposes, the fibers in (B) have been cropped to scale in order to demonstrate the length reductions and extensions achievable via the strain device. (C) A plot of the
average IA/ID±standard deviation versus applied to or released strain from manually deposited Fn fibers (see Materials and methods). Average values were obtained
from at least 10 fibers to compute a single data point. Released strain is acquired by pre-straining the silicone substrate before pulling fibers and then relaxing the strain
device. IA/IDvalues are calibrated to chemical denaturing data (Fig. 2A). The absolute scale of strain is shown as well, assuming that 1/3× is the point offull relaxation.
Scale bar in (B)=50 μm.
455 W.C. Little et al. / Matrix Biology 27 (2008) 451–461
fibers do not contain a measurable population of Fn in the fully
Fn module unfolding must contribute to the extraordinary
extensibility of Fn fibers. While our measurements cannot
directly determine the IA/ID ratio below which mechanically
stretched fibers unfold, CD measurements from Fn in solution
indicate that Fn unfolding occurs at concentrations of GdnHCl
well below 2 M. Khan et al. 1990 reported a 26% loss in
secondary structure for the central cell binding fragment of Fn
(modules FnIII1–11) from a baseline of 5 (0 to 1.2 M GdnHCl) to
~3.7×102deg×cm2×dmol−1(2 M GdnHCl). A 26% loss of
secondary structure within the FnIII1–11fragment would roughly
correspond to complete unfolding of three FnIII modules. If a
FnIII module is unfolded mechanically (not chemically), it
lengthens from ~3.2 nm to a contour length of ~28.5 nm
(Oberhauser et al., 2002). If three modules are unfolded
mechanically, an extended Fn molecule which is estimated to be
130 nm in length prior unfolding (Engel et al., 1981), would thus
increase in length to 206 nm (130+3⁎(28.5−3.2)), or 1.6-fold.
Interestingly,Fig.3shows thattheaverage end-to-enddistanceof
atIA/IDratios that correspond to those measured at 1.5 (0.58) and
2 M GdnHCl in solution (0.52), and are thus in agreement with
our results derived from this strain. Since publishing 360 our first
paper on using FRET to probe conformational alterations of Fn
(Baneyx et al., 2001), the FRET signature of each batch of
progressively denatured Fn in solution was determined. This
allows for a comparison between batch to batch variations of
labelled Fn, and for other groups to see whether their labelling
protocols lead to comparable results. Certainly, Fn has different
conformations in solution and in the fibrillar states. However, if
the two dimeric Fn arms are separated by breaking stabilizing
electrostatic interactions, the labelled domains do not have to
move far to completely eliminate inter-arm FRET. Similarly, if
two FRET probes are on adjacent domains, domain unfolding
increase that exceeds the Förster radius. Experiments are
underway to identify the exact point at which fibrillar Fn starts
to lose secondary structure by force-induced unfolding.
state of fibrils is not composed of Fn dimers whose arms are
already extended into a straightstring offoldedmodules. Instead,
indicated by the finding that the fully relaxed fibers have IA/ID
Fig. 4. Fn fiber breakage for fibers that are adherent to an elastic silicone substrate. (A) A spatially resolved FRET map of manually deposited Fn fibers subjected to a
2× increase in substrate length shows areas of fiber breakage and large areas of mechanically induced unfolded Fn Type III modules, as confirmed via (B) a histogram
of every pixel's acceptor intensity / donor intensity (IA/ID, see Materials andmethods) and calibrated to a guanidinium hydrochloride (GdnHCL) denaturingcurve. The
absolute strain scale represents the average IA/IDvalues of Fn fibers that are fully relaxed or extended to the point where they begin to break (5–6×), as described in the
text and in Fig. 3. (C) Zooming in on an example breakage location in a fiber as noted by the red box in (A). (D) A fluorescence image of the break, especially a (E)
contrast-enhancedimaged, revealsthat a small numberof individualstrandsare beingpulledapart andleft behindin the breakagegapthat are notdetectedin our FRET
measurements due to the necessary analysis thresholding (see Materials and methods). (F) AFM image confirming the presence of a small amount of fibrillar Fn left on
the substrate and a tapered end to the broken fiber. Inset zoomed-in image of a fractured fiber end shows the receding strands. (G) Some breaks also show only a small
amount of Fn left behind on the substrate and show more blunted fibers ends. Scale bar in (A)=30 μm, Scale bar in (D)=15 μm and is the same scale as the images in
(C) and (E).
456 W.C. Little et al. / Matrix Biology 27 (2008) 451–461
IDvalue ofextendedFnfoundat1.5M GdnHCl(0.58)(Figs.2A,
3C). This conclusion agrees with other spectroscopic ob-
electron microscopy images showing that nodules decorate the
otherwise smooth surface of the fibers (Chen et al., 1997; Peters
each nodule can contain ~4 FnIII modules. These nodules are
shown in the Cryo-SEM studies to essentially disappear within
under mechanical tension (Chen et al., 1997; Peters et al., 1998).
Therefore, Fn fibrils upon stretch transition from a nodular to a
their content integrates into the smooth fibers as the end-to-end
distance of the fiber is increased. Future research must address
whether the nodules completely disappear before the first Fn
modules into nodules and a second from loss of secondary
structure at higher tensions.
2.6. The range of conformations observed from relaxed to
highly strained manually deposited Fn fibers simultaneously
exists within one field of view within ECM of living fibroblasts
In addition to the microscopic structural similarities and
similar average IA/IDratios of cell-derived and manually de-
posited Fn fibers (0.59±0.09 vs. 0.62±0.06, respectively),
further evidence that the assay fibers can serve as physiolo-
gically relevant and useful model systems lies in the
observation that they can be strained or relaxed to yield IA/
IDratios that span the range of IA/IDratios found in single
fields of view of ECM (Fig. 2E). When strained or relaxed,
manually deposited fibers maintain a relatively narrow
conformational distribution (Fig. 3C); the data points in the
resulting FRET vs. strain curve for both relaxed and strained
fibers had an average IA/IDstandard deviation of only 0.018
(Fig. 3C), compared to 0.089 found within cell-derived fibers
(Fig. 2E). On absolute scales, cellular fibers had IA/IDratios
between 0.42 and 0.81 (from 4 M to 0.5 M GdnHCL
equivalents) and a few particularly long and thin fibrils
(Fig. 2D) showed IA/IDratios between 0.42 and 0.58 (similar
to Fn in 1.5–4 M GdnHCL).
2.7. Fn fibers can be manually deposited in various orientations
and curves to generate a variety of molecular conformations in
one field of view after simultaneous one-dimensional strain
An advantageous property of a 1D stretch assay is that when
a rectangular sheet is stretched 3× their length in one dimension
the width compresses ~0.59× in the orthogonal direction.
Therefore, depositing “grids” of fibers onto 3× pre-strained
sheet followed by a full relaxation of the sheet results in
horizontal fibers relaxed 1/3× and vertical fibers stretched 1.7×
their end-to-end length, corresponding to the fully relaxed
conformation and near-breakage point on the IA/IDvs. strain
calibration curve, respectively (Fig. 3C). Therefore, this
method of pre-straining the sheet before depositing fibers in
various angles and curves and subsequently relaxing it renders
the highest possible range in homogeneous fibrillar Fn
conformations existing in a single field of view without fiber
2.8. Mechanically strained Fn fibers expose cryptic binding
sites for the N-terminal 70 kD Fn fragment at early extensions
A number of studies have demonstrated previously that either
proteolytic cleavage or heat-, chemical-, or substrate-induced
unfolding of soluble Fn reveals a variety of cryptic protein binding
sites (for reviews see Pankov and Yamada, 2002; Vogel, 2006).
However, to our knowledge only one study has investigated how
Fn conformation (Zhong et al., 1998) showing that the binding of
Fn, the N-terminal 70 kD Fn fragment, and the L8 monoclonal
antibody to cell-made Fn fibers was decreased when cells were
treated with Rho inhibitors. Since Rho stimulates cell contractility,
the authors concluded that cell-generated tension is needed to
expose Fn's binding sites for these proteins. Fig. 5C now shows
that the binding of the 70 kD fragment to Fn is indeed highly
dependent on the Fn conformation. While the Rho inhibition
assay presented above to generate fibers of defined conformations
to examine how the binding of the 70-kd fragment relates to the
conformation of Fn fibers as detected via FRET. Fig. 5C shows a
representative field of view of the far-red channel (645 nm–
633-labeled 70-kd fragment (See Materials and methods). In
strained1.7× clearlyboundmoreof the 70-kdfragmentthanthe 1/
3×-relaxed vertical fibers. Interestingly, the curved fiber in the
image contains both vertical and more horizontal areas along its
70 kDa Fn fragment appeared to begin binding to the curved fiber
to IA/IDvalues calibrated to the ~1 M GdnHCL point, which as
to the extended conformation prior to the loss of major secondary
structure. In the broader context, the 70 kD binding studies show
that the assay is well suited for the study of strain-dependent
A new strain assay is presented which is well suited for
biochemical and cellular assays aimed at investigating if Fn
within the extracellular matrix can act as a mechanotransducer.
Essential features of the assay are the ability of tuning the
conformational distribution of Fn by the application of external
strain, and of probing the strain-induced protein conformational
457 W.C. Little et al. / Matrix Biology 27 (2008) 451–461
changes within a large number of parallel fibers or fiber grids.
This assay has the capability to detect strain-dependent protein–
Future studies need to examine whether integrins or other
proteins involved in outside-in cell signaling might bind to
ECM fibers in a stretch-dependent manner which could then
result in a stretch-dependent regulation of signaling pathways.
Since it has been shown so far only that most cell types are
capable of probing the rigidity of their environment (Discher
et al., 2005; Engler et al., 2006; Galbraith et al., 2002; Giannone
and Sheetz, 2006; Ingber, 2006; Kostic and Sheetz, 2006; Vogel
and Sheetz, 2006; Yeung et al., 2005), the assay introduced
here could play a key role in the design of experiments that
might distinguish between the force-induced functional
changes of ECM molecules and of fiber rigidity on cell
4. Materials and methods
4.1. Isolation of Fn
Human plasma Fn was isolated from human plasma (Swiss
Red Cross) via a previously described procedure involving the
use of gelatin-sepharose chromatography (Engvall and Ruo-
slahti, 1977). Briefly, the serum was first centrifuged at
2000 RPM for 10 min to ensure the removal of all blood cells.
The supernatant was then supplied with 10 mM ethylenediami-
netetraacetic acid (EDTA) and 2 mM phenylmethylsulfonyl
resulting supernatant was then passed through a sepharose 4B
column (Sigma) and then through a gelatin-sepharose 4B
column until the eluant contained no detectable proteins
(absorbance at 280 nm)—washing buffer was PBS with
10 mM EDTA and 2 mM PMSF. The column was then washed
with 1 M NaCl followed by 1 M urea. Finally, the Fn was eluted
using 6 M urea and its purity was confirmed using western
blotting and silver staining (not shown).
4.2. Cell culture and fluorescent cell labeling
NIH-3T3 mouse fibroblasts were obtained from ATCC and
cultured in Dulbecco's Modified Eagle's Medium (DMEM)
with 10% Fetal Calf Serum (FCS) (Gibco). Human Umbilical
Vein Endothelial Cells (HUVECs) were obtained from
PromoCell and cultured in endothelial cell growth medium
plus supplement (PromoCell). Both cell types were incubated
in a humidified 37 °C chamber with 5% CO2. Only cells below
passage 15 were used for the experiments described in this
report. For Fig. 1E, HUVECs were fluorescently labeled with
CellTrace™ Far Red DDAO-SE (Invitrogen) via the manufac-
4.3. Fluorophore conjugation of Fn
Fn was dual-labeled with Alexa Fluor® 488 succinimidyl
ester and Alexa Fluor® 546 maleimide for FRET experiments
via a modified procedure previously described (Baneyx et al.,
2001). Plasma Fn contains two free sulfhydryls per subunit
(Smith et al., 1982) which are buried within the tertiary structure
of the Fn III7and III15modules (Mosher and Johnson, 1983),
which requires that they be exposed by denaturant for
fluorophore conjugation. Briefly, Fn in 6 M urea taken from
the final step of the isolation procedure was further denatured by
the addition of guanidine hydrochloride (GdnHCL, 4 M final
concentration) and incubated at room temperature for 5 min. A
25-fold molar excess of Alexa Fluor® 546 maleimide was then
Fig. 5. Stretch-dependent adsorption of the 70-kd N-terminal Fn fragment to a wide range of fibrillar Fn conformations generated in a single field of view. (A) Fn fibers
were manually deposited onto a 3× pre-strained silicone sheet in curves and anti-parallel orientations and then fully relaxed to simultaneously strain horizontally
oriented fibers in (B) 1.7× and relax vertically oriented fibers in (B) 1/3×. A histogram of the resulting IA/IDfor each pixel analyzed (see Materials and methods) is
shown and color-coded so that the corresponding image map (B) can be used to easily identify highly strained fibers (blue) and fully relaxed fibers (red) and
conformations in between. (C) A solution of Alexa Flour ® 633-labeled 70-kd Fn fragment was adsorbed to the substrate for 10 min, rinsed 4× with PBS, then imaged
in a far-red channel (645 nm–745 nm). Stretched fibers (horizontal orientation) clearly bind more of the protein fragment than relaxed fibers (vertical orientation), and
the curved fiber shows that the transition between relatively low and high binding occurs where the region of the fiber transitions between a vertical and horizontal
458 W.C. Little et al. / Matrix Biology 27 (2008) 451–461
added to the denatured Fn solution, mixed by gentle pipetting,
and incubated at room temperature for 2 h with gentle rocking.
Free dye from the solution was removed via size-exclusion
chromatography with phosphate buffered saline (PBS, pH 7.4)
through a PD10 column and the concentration of remaining Fn
was determined via a spectrophotometer (MBA2000, Perki-
nElmer Instruments). At this point the free cysteine residues
located within modules FnIII7and FnIII15(4 total per Fn dimer)
are specifically labeled with dye.
Next, sodium bicarbonate was added to the solution to a final
concentration of 0.1 M at pH 8.7 for amine labeling. A 110-fold
molar excess of Alexa Fluor® 488 succinimidyl ester was then
added to the solution and allowed to incubate for 1 h at room
temperature with gentle rocking to randomly label amine
residues. Free dye was again removed from the solution via
size-exclusion chromatography and the eluant solution was
determined via spectrophotography to have Fn labeled on
average with 6.7 donors (Alexa Fluor® 488) and 3.7 acceptors
(Alexa Fluor® 546). The point at which fragments contaminated
the eluant was determined via Western blotting (not shown) and
only the volume before this point was collected for experiments.
The Fn solution was stored in frozen 10 μl aliquots and only
used when thawed for less than 5 days. This batch of dual-
labeled Fn (Fn-d/a) was used to collect all the FRET data in
Figs. 2–4 presented in this report. The batch used for Fig. 5 was
prepared by the same methods and had on average 6.0 donors
and 3.6 acceptors.
4.4. Chemical denaturing curve, microscopy, and FRET
To calibrate how FRET relates to the loss of quaternary
(separation of the dimer arms) and tertiary structure, the IA/ID
ratios were determined for Fn in denaturing solutions of 0 M,
1 M, 2 M, and 4 M GdnHCL. Bovine Serum Albumin (BSA)-
coated glass coverslips and glass microscope slides were used to
create 2 mm wide chambers to house the GdnHCL-Fn-d/a
solutions and to prevent adsorption to the glass surfaces.
200 μm-thick silicone sheets (SMI, Saginaw, MI) were used for
the chamber walls and spacer for the coverslip and slide. The
Fn-d/a solution (~2 μL) was drawn into the chamber via
capillary forces and each solution was imaged via scanning laser
confocal microscopy (Olympus FV-1000). The samples were
excited at 488 nm and the two emission detection windows were
set at 514–526 nm (donor channel) and 566–578 nm (acceptor
channel) to capture their peak emissions.
Spatially resolved FRET analysis (by pixel) was then
performed using a custom script written in PHP (Php Hypertext
Preprocessor, www.php.net) or Ruby (www.ruby-lang.org)
together with image analysis software (GD graphics library,
www.boutell.com/gd—ImageMagick Software Suite, www.
imagemagick.org). The script first subtracted a dark current
background from each channel (acquired from each experi-
ment), then applied a 1-pixel radius Gaussian blur (with a 1-
pixel standard deviation) to each channel to smooth errant
intensity values. The intensity of the 12-bit acceptor channel
(values from 0–4095) for each pixel was then divided by the
intensity of the corresponding 12-bit donor channel (as long as
each channel's intensity value was above a threshold of at least
100 and not saturated at 4095) to yield IA/IDratios. Decreasing
IA/IDratios indicate increased nodule extension and domain
unfolding. To generate spatially resolved FRET maps, the IA/ID
ratio for each pixel (0.33 μm×0.33 μm) was mapped to a false
color, with red and blue representing compact and unfolded
4.5. Cell-derived Fn fibers
Fn-d/ -labeled cell-derived matrix was obtained by a mod-
ified procedure previously described (Baneyx et al., 2001).
Briefly, a 10 μg/mL solution of unlabeled Fn (u-Fn) was first
adsorbed onto a glass coverslip for 30 min at room temperature
and before a near-confluent layer of NIH-3T3 cells was
cultured. Thirty minutes later the cells were rinsed with media
and then incubated in media containing 45 μg/mL of u-Fn and
5 μg/mL of Fn-d/a in order to avoid intermolecular FRET
(Baneyx et al., 2001). After 24 h, the sample was rinsed with
PBS at 37 °C and then filled with a 2 mM 6-Hydroxy-2,5,7,8-
tetramethylchroman-2-carboxylic acid (Trolox) solution in PBS
for live imaging of the matrix.
4.6. Manually deposited Fn fibers
Manually deposited Fn fibers were generated by a modified
procedure previously described (Ejim et al., 1993). In all
fibrillar experiments, soluble Fn-d/a was diluted into 95%
unlabeled Fn to prevent intermolecular FRET. At this dilution
factor, the acceptor intensity divided by donor intensity (IA/ID)
ratio was independent of the Fn concentration, ensuring that
the signals reported here originate from intramolecular FRET
(Fig. S3). Briefly, a plastic pipette tip was cut into a sharp
point and dipped into a 0.76 mg/mL 5% Fn-d/a / 95% u-Fn
droplet on a 200 μm-thick silicone sheet (SMI, Saginaw, MI)
that was pre-cleaned by immersion in ethanol for 10 min and
gently drawn upward to induce Fn polymerization. The
resulting fiber was then deposited onto the surface ~1–2 cm
away from the droplet slightly pressing the tip into the elastic
silicone sheet. Once the desired number and pattern of fibers
were deposited, the droplet and sample would be allowed to
dry for 5 min before rinsing four times with a 2% (w/v) bovine
serum albumin (BSA) solution in PBS. Samples were left in
this BSA solution at room temperature no longer than 2 h
Once in contact with glass or silicone substrates, the fibers
attach tightly along their full length, which was confirmed by
using a microneedle to disrupt the fiber at various locations in
both dry and aqueous environments. Instead of detaching and/or
recoiling non-uniformly away from a break, the fiber stayed in
place and only the immediate area around the microneedle was
cut, independent of where the break occurred (not shown). This
observation was also confirmed by the fact that the fibers did
not detach from the silicone substrate when stretched and the
spatially resolved IA/IDratios were homogeneous when strain
was applied or released (Fig. 3).
459W.C. Little et al. / Matrix Biology 27 (2008) 451–461
4.7. Fluorescent labeling and adsorption of the 70-kd Fn
For adsorption studies, Alexa Fluor® 633 was used to fluo-
rescently label the 70-kd N-terminal Fn fragment (Sigma) per the
manufacturer's protocol. After depositing Fn fibers on 3× strained
the substrate was fully relaxed resulting in a 1/3× relaxation of
fibers in the direction of the one-dimensional strain and a 1.7×
70-kd fragment in PBS was introduced to the fibers for 10 min at
room temperature then rinsed 4× with PBS before imaging.
4.8. AFM of manually deposited fibronectin fibers
Surface scans of Fn fibers (Fig. 4I, J) were obtained by first
depositing the fibers onto 4× pre-strained silicone substrates in
an orientation perpendicular to the direction of 1D strain and
rinsing 4× with PBS. After full relaxation of the substrate in
PBS the fibers resulted in being strained 2× to induce fracturing.
AFM of dried samples was used in AC-mode (Asylum
Research, MFP-3D). The cantilever was an Olympus silicon
cantilever AC160TS, operating slightly below the resonance
frequency of 292 kHz at a scan rate of 0.3 Hz.
The authors gratefully acknowledge Jörg Albuschies for
taking the AFM images shown in Fig. 4, Jean Schwarzbauer for
providing us with the Fn isolation, Sheila Luna for the Fn
purifications, Anne Wandrey for aid with SEM imaging,
Kristopher Kubow, Delphine Gourdon, John Saeger, Gretchen
Baneyx, and Meher Antia for their helpful discussions, and
members of the ETH Materials department machine shop for
constructing the custom strain device.This workwas financially
supported by the ETH Zurich (VV), the Human Frontier
Science Program Organization (MLS), and from the Nanome-
dicine Development Center (NDC) “Nanotechnology Center for
Mechanics in Regenerative Medicine” (NIH grant PN2
EY016586), that participates in the NIH Nanomedicine
Development Center Network (NNDCN).
Appendix A. Supplementary data
Supplementary data associated with this article can be found,
in the online version, at doi:10.1016/j.matbio.2008.02.003.
Abu-Lail, N.I., Ohashi, T., Clark, R.L., Erickson, H.P., Zauscher, S., 2005.
Understanding the elasticity offibronectinfibrils: unfolding strengths of FN-
III and GFP domains measured by single molecule force spectroscopy.
Matrix Biol. 25, 175–184.
Ahmed, Z., Brown, R.A., 1999. Adhesion, alignment, and migration of cultured
Schwann cells on ultrathin fibronectin fibres. Cell Motil. Cytoskelet. 42,
Ahmed, Z., Underwood, S., Brown, R.A., 2000. Low concentrations of
cables. Cell Motil. Cytoskelet. 46, 6–16.
Alexander, S.S., Colonna, G., Edelhoch, H., 1979. The structure and stability of
human plasma cold-insoluble globulin. J. Biol. Chem. 254, 1501–1505.
Altroff, H., Choulier, L., Mardon, H.J., 2003. Synergistic activity of the ninth
and tenth FIII domains of human fibronectin depends upon structural
stability. J. Biol. Chem. 278, 491–497.
Antia, M., Islas, L.D., Boness, D.A., Baneyx, G., Vogel, V., 2006. Single
molecule fluorescence studies of surface-adsorbed fibronectin. Biomaterials
Arcangeli, A., Becchetti, A., 2006. Complex functional interaction between
integrin receptors and ion channels. Trends Cell Biol. 16, 631–639.
Baneyx, G., Vogel, V., 1999. Self-assembly of fibronectin into fibrillar networks
underneathdipalmitoyl phosphatidylcholinemonolayers:role of lipidmatrix
and tensile forces. Proc. Natl. Acad. Sci. U. S. A. 96, 12518–12523.
Baneyx,G., Baugh,L.,Vogel, V., 2001.Coexistingconformationsoffibronectin
in cell culture imaged using fluorescence resonance energy transfer. Proc.
Natl. Acad. Sci. U. S. A. 98, 14464–14468.
Baneyx, G., Baugh, L., Vogel, V., 2002. Fibronectin extension and unfolding
within cell matrixfibrils controlled by cytoskeletal tension.Proc.Natl.Acad.
Sci. U. S. A. 99, 5139–5143.
Barker, T.H., Baneyx, G., Cardo-Vila, M., Workman, G.A., Weaver, M., Menon,
2005. SPARC regulates extracellular matrix organization through its modu-
lation of integrin-linked kinase activity. J. Biol. Chem. 280, 36483–36493.
Baugh, L., Vogel, V., 2004. Structural changes of fibronectin adsorbed to model
surfaces probed by fluorescence resonance energy transfer. J. Biomed.
Mater. Res. 69A, 525–534.
Briknarova, K., Akerman, M.E., Hoyt, D.W., Ruoslahti, E., Ely, K.R., 2003.
Anastellin, an FN3 fragment with fibronectin polymerization activity,
resembles amyloid fibril precursors. J. Mol. Biol. 332, 205–215.
Bustamante, C., Chemla, Y.R., Forde, N.R., Izhaky, D., 2004. Mechanical
processes in biochemistry. Annu. Rev. Biochem. 73, 705–748.
Chen, H., Mosher, D.F., 1996. Formation of sodium dodecyl sulfate-stable
fibronectin multimers. Failure to detect products of thiol-disulfide exchange
in cyanogen bromide or limited acid digests of stabilized matrix fibronectin.
J. Biol. Chem. 271, 9084–9089.
Chen, L.B., Murray, A., Segal, R.A., Bushnell, A., Walsh, M.L., 1978. Studies
on intercellular LETS glycoprotein matrices. Cell 14, 377–391.
Chen, Y., Zardi, L., Peters, D.M., 1997. High-resolution cryo-scanning electron
microscopy study of the macromolecular structure of fibronectin fibrils.
Scanning 19, 349–355.
Craig, D., Gao, M., Schulten, K., Vogel, V., 2004a. Structural insights into how
the MIDAS ion stabilizes integrin binding to an RGD peptide under force.
Structure 12, 2049–2058.
Craig, D., Gao, M., Schulten, K., Vogel, V., 2004b. Tuning the mechanical
stability of fibronectin type III modules through sequence variations.
Structure 12, 21–30.
Craig, D., Krammer, A., Schulten, K., Vogel, V., 2001. Comparison of the early
stages of forced unfolding for fibronectin type III modules. Proc. Natl. Acad.
Sci. U. S. A. 98, 5590–5595.
Dalby, M.J., Riehle, M.O., Sutherland, D.S., Agheli, H., Curtis, A.S., 2004. Use
of nanotopography to study mechanotransduction in fibroblasts—methods
and perspectives. Eur. J. Cell Biol. 83, 159–169.
Dalby, M.J., M.O. Riehle, D.S. Sutherland, H. Agheli, and A.S. Curtis. 2005.
Morphological and microarray analysis of human fibroblasts cultured on
nanocolumns produced by colloidal lithography. Eur Cell Mater. 9:1–8;
Discher, D.E., Janmey, P., Wang, Y.L., 2005. Tissue cells feel and respond to the
stiffness of their substrate. Science 310, 1139–1143.
Dzamba, B.J., Peters, D.M., 1991. Arrangement of cellular fibronectin in
noncollagenous fibrils in human fibroblast cultures. J. Cell Sci. 100 (Pt 3),
Ejim, O.S., Blunn, G.W., Brown, R.A., 1993. Production of artificial-orientated
mats and strands from plasma fibronectin: a morphological study.
Biomaterials 14, 743–748.
Engel,J., Odermatt,E.,Engel,A.,Madri,J.A., Furthmayr,H., Rohde,H., Timpl,
R., 1981. Shapes, domain organizations and flexibility of laminin and
fibronectin, two multifunctional proteins of the extracellular matrix. J. Mol.
Biol. 150, 97–120.
460 W.C. Little et al. / Matrix Biology 27 (2008) 451–461
Engler, A.J., Sen, S., Sweeney, H.L., Discher, D.E., 2006. Matrix elasticity Download full-text
directs stem cell lineage specification. Cell 126, 677–689.
Engvall, E., Ruoslahti, E., 1977. Binding of soluble form of fibroblast surface
protein, fibronectin, to collagen. Int. J. Cancer 20, 1–5.
Erickson, H.P., 1994. Reversible unfolding of fibronectin type III and immu-
noglobulin domains provides the structural basis for stretch and elasticity of
titin and fibronectin. Proc. Natl. Acad. Sci. U. S. A. 91, 10114–10118.
Erickson, H.P., 2002. Stretching fibronectin. J. Muscle Res. Cell Motil. 23,
Galbraith, C.G., Yamada, K.M., Sheetz, M.P., 2002. The relationship between
force and focal complex development. J. Cell Biol. 159, 695–705.
Garcia, A.J., Vega, M.D., Boettiger, D., 1999. Modulation of cell proliferation
and differentiation through substrate-dependent changes in fibronectin
conformation. Mol. Biol. Cell 10, 785–798.
Giannone, G., Sheetz, M.P., 2006. Substrate rigidity and force define form
through tyrosine phosphatase and kinase pathways. Trends Cell. Biol. 16,
Halter, M., Antia, M., Vogel, V., 2005. Fibronectin conformational changes
induced by adsorption to liposomes. J. Control. Release 101, 209–222.
Hynes, R.O., 1990. Fibronectins. Springer-Verlag Inc., New York.
Ingber,D.E.,2006.Cellularmechanotransduction:puttingall the piecestogether
again. Faseb J. 20, 811–827.
Johnson, C.P., Tang, H.Y., Carag, C., Speicher, D.W., Discher, D.E., 2007.
Forced unfolding of proteins within cells. Science 317, 663–666.
Keselowsky, B.G., Collard, D.M., Garcia, A.J., 2005. Integrin binding
specificity regulates biomaterial surface chemistry effects on cell differ-
entiation. Proc. Natl. Acad. Sci. U. S. A. 102, 5953–5957.
Khan, M.Y., Medow, M.S., Newman, S.A., 1990. Unfolding transitions of
fibronectin and its domains. Stabilization and structural alteration of the N-
terminal domain by heparin. Biochem. J. 270, 33–38.
Kostic, A., Sheetz, M.P., 2006. Fibronectin rigidity response through Fyn and
p130Cas recruitment to the leading edge. Mol. Biol. Cell 17, 2684–2695.
Kung, C., 2005. A possible unifying principle for mechanosensation. Nature
Lai, C.S., Tooney, N.M., Ankel, E.G., 1984. Structure and flexibility of plasma
fibronectin in solution: electron spin resonance spin-label, circular
dichroism, and sedimentation studies. Biochemistry 23, 6393–6397.
Lan, M.A., Gersbach, C.A., Michael, K.E., Keselowsky, B.G., Garcia, A.J.,
2005. Myoblast proliferation and differentiation on fibronectin-coated self
assembledmonolayerspresentingdifferent surfacechemistries. Biomaterials
Langenbach, K.J., Sottile, J., 1999. Identification of protein-disulfide isomerase
activity in fibronectin. J. Biol. Chem. 274, 7032–7038.
Litvinovich, S.V., Brew, S.A., Aota, S., Akiyama, S.K., Haudenschild, C.,
Ingham, K.C., 1998. Formation of amyloid-like fibrils by self-association of
a partially unfolded fibronectin type III module. J. Mol. Biol. 280, 245–258.
Magnusson, M.K., Mosher, D.F., 1998. Fibronectin: structure, assembly,
and cardiovascular implications. Arterioscler. Thromb. Vasc. Biol. 18,
Mao, Y., Schwarzbauer, J.E., 2005. Fibronectin fibrillogenesis, a cell-mediated
matrix assembly process. Matrix Biol. 24, 389–399.
McKeown-Longo, P.J., Mosher, D.F., 1983. Binding of plasma fibronectin to
cell layers of human skin fibroblasts. J. Cell Biol. 97, 466–472.
Midwood, K.S., Mao, Y., Hsia, H.C., Valenick, L.V., Schwarzbauer, J.E., 2006.
Modulation of cell-fibronectin matrix interactions during tissue repair.
J. Investig. Dermatol. Symp. Proc. 11, 73–78.
Morla, A., Zhang, Z., Ruoslahti, E., 1994. Superfibronectin is a functionally
distinct form of fibronectin. Nature 367, 193–196.
Mosher, D.F., Johnson, R.B., 1983. In vitro formation of disulfide-bonded
fibronectin multimers. J. Biol. Chem. 258, 6595–6601.
2002. The mechanical hierarchies of fibronectin observed with single-
molecule AFM. J. Mol. Biol. 319, 433–447.
Ohashi, T., Kiehart, D.P., Erickson, H.P., 1999. Dynamics and elasticity of the
fibronectin matrix in living cell culture visualized by fibronectin-green
fluorescent protein. Proc. Natl. Acad. Sci. U. S. A. 96, 2153–2158.
Pankov, R., Yamada, K.M., 2002. Fibronectin at a glance. J. Cell. Sci. 115,
Paszek, M.J., Zahir, N., Johnson, K.R., Lakins,J.N., Rozenberg, G.I., Gefen, A.,
Reinhart-King, C.A., Margulies, S.S., Dembo, M., Boettiger, D., Hammer,
D.A., Weaver, V.M., 2005. Tensional homeostasis and the malignant
phenotype. Cancer Cell 8, 241–254.
Peters, D.M.P., Chen, Y., Zardi, L., Brummel, S., 1998. Conformation of
fibronectin fibrils varies: discrete globular domains of type III repeats
detected. Microsc. Microanal. 4, 385–396.
Romberger, D.J., 1997. Fibronectin. Int. J. Biochem. Cell Biol. 29, 939–943.
Samori, B., Zuccheri, G., Baschieri, R., 2005. Protein unfolding and refolding
under force: methodologies for nanomechanics. Chemphyschem 6, 29–34.
Sawada, Y., Tamada, M., Dubin-Thaler, B.J., Cherniavskaya, O., Sakai, R.,
Tanaka, S., Sheetz, M.P., 2006. Force sensing by mechanical extension of
the Src family kinase substrate p130Cas. Cell 127, 1015–1026.
Sechler, J.L., Schwarzbauer, J.E., 1998. Control of cell cycle progression by
fibronectin matrix architecture. J. Biol. Chem. 273, 25533–25536.
Singer, I.I., 1979. The fibronexus: a transmembrane association of fibronectin-
containing fibers and bundles of 5 nm microfilaments in hamster and human
fibroblasts. Cell 16, 675–685.
Smith, D.E., Mosher, D.F., Johnson, R.B., Furcht, L.T., 1982. Immunological
identification of two sulfhydryl-containing fragments of human plasma
fibronectin. J. Biol. Chem. 257, 5831–5838.
Smith, M.L., Gourdon, D., Little, W.C., Kubow, K.E., Eguiluz, R.A., Luna-
Morris, S., Vogel, V., 2007. Force-induced unfolding of fibronectin in the
extracellular matrix of living cells. PLoS Biol. 5, e268.
Tamada, M., Sheetz, M.P., Sawada, Y., 2004. Activation of a signaling cascade
by cytoskeleton stretch. Dev. Cell 7, 709–718.
Vogel, V., 2006. Mechanotransduction involving multimodular proteins:
converting force into biochemical signals. Annu. Rev. Biophys. Biomol.
Struct. 35, 459–488.
Vogel, V., Sheetz, M., 2006. Local force and geometry sensing regulate cell
functions. Nat. Rev. Mol. Cell Biol. 7, 265–275.
Wang, K., Forbes, J.G., Jin, A.J., 2001. Single molecule measurements of titin
elasticity. Prog. Biophys. Mol. Biol. 77, 1–44.
Wierzbicka-Patynowski, I., Schwarzbauer, J.E., 2003. The ins and outs of
fibronectin matrix assembly. J. Cell Sci. 116, 3269–3276.
Wojciak-Stothard, B., Denyer, M., Mishra, M., Brown, R.A., 1997. Adhesion,
orientation,andmovement of cells culturedon ultrathin fibronectin fibers. In
Vitro Cell Dev. Biol. Anim. 33, 110–117.
Yeung,T.,Georges,P.C.,Flanagan, L.A.,Marg,B.,Ortiz, M.,Funaki,M.,Zahir,
N., Ming, W., Weaver, V., Janmey, P.A., 2005. Effects of substrate stiffness
on cell morphology, cytoskeletal structure, and adhesion. Cell Motil.
Cytoskelet. 60, 24–34.
Zhong, C., Chrzanowska-Wodnicka, M., Brown, J., Shaub, A., Belkin, A.M.,
Burridge, K., 1998. Rho-mediated contractility exposes a cryptic site in
fibronectin and induces fibronectin matrix assembly. J. Cell Biol. 141,
461 W.C. Little et al. / Matrix Biology 27 (2008) 451–461