The Rockefeller University Press $30.00
J. Cell Biol. Vol. 197 No. 3 439–455
Correspondence to Ryan J. Petrie: firstname.lastname@example.org; or Kenneth M. Yamada:
Abbreviations used in this paper: CDM, cell-derived matrix; ERK, extracellular
signal–regulated kinase; Fc, corrected FRET; FRET, fluorescence resonance energy
transfer; HFF, human foreskin fibroblast; PBD, p21-binding domain; PH, pleckstrin
homology; PI, polarization index; PIP3, phosphatidylinositol (3,4,5)-trisphosphate;
PIP2, phosphatidylinositol (4,5)-bisphosphate; ROCK, Rho-associated protein
kinase; VASP, vasodilator-stimulated phosphoprotein.
How normal cells move efficiently through chemically and
structurally diverse 3D environments in vivo is not well
understood. In contrast, findings of metazoan cells migrating
on uniform 2D surfaces in vitro have led to a comprehensive
model of cell motility wherein polarized signaling orchestrates
cell movement by directing lamellipodial protrusion at the lead-
ing edge, adhesion to the underlying substrate, and retraction at
the trailing edge (Lauffenburger and Horwitz, 1996; Ridley
et al., 2003). The second messenger phosphatidylinositol (3,4,5)-
trisphosphate (PIP3) is enriched at the leading edge (Haugh
et al., 2000), where it can recruit downstream effectors, such as
guanine exchange factors (Côté et al., 2005) that activate the
Rho family of GTPases. Rho family members Rac1, Cdc42, and
RhoA are active at the leading edge and coordinate protrusion
and adhesion (Kraynov et al., 2000; Nalbant et al., 2004; Pertz
et al., 2006; Machacek et al., 2009).
Disrupting the subcellular localization of Rac1, Cdc42, or
RhoA can lead to defects in adhesion and motility (van Hennik
et al., 2003; ten Klooster et al., 2006; Bass et al., 2007), whereas
the light-mediated activation of photosensitive guanine ex-
change factor, Rac1, or Cdc42 constructs at discrete regions of
the plasma membrane triggers protrusion and directional cell
migration (Levskaya et al., 2009; Wu et al., 2009). Discrepan-
cies in the localization of Rho family GTPase activities during
cell migration in vivo versus on 2D surfaces might reveal dif-
ferences in the mechanisms that drive cell motility. Studies of
cancer cell migration in 3D environments show that metastatic
cells can switch between adhesion-dependent mesenchymal
(elongated) and adhesion-independent amoeboid (rounded) cell
motility (Table S1), driven by actin polymerization and actomy-
osin contraction, respectively (Wolf et al., 2003; Lämmermann
and Sixt, 2009). Although these two different modes of cancer
cell migration have specific requirements for Rho family GTPase
signaling, how that signaling is organized is not known.
Furthermore, it is unclear how the mesenchymal–amoeboid
(3,4,5)-trisphosphate (PIP3) and active Rac1 and Cdc42
in primary fibroblasts migrating within different 3D
environments. In 3D collagen, PIP3 and active Rac1
and Cdc42 were targeted to the leading edge, con-
sistent with lamellipodia-based migration. In contrast,
elongated cells migrating inside dermal explants and
the cell-derived matrix (CDM) formed blunt, cylindri-
cal protrusions, termed lobopodia, and Rac1, Cdc42,
and PIP3 signaling was nonpolarized. Reducing RhoA,
e search in this paper for context-specific
modes of three-dimensional (3D) cell migra-
tion using imaging for phosphatidylinositol
Rho-associated protein kinase (ROCK), or myosin II activity
switched the cells to lamellipodia-based 3D migration.
These modes of 3D migration were regulated by matrix
physical properties. Specifically, experimentally modify-
ing the elasticity of the CDM or collagen gels established
that nonlinear elasticity supported lamellipodia-based
migration, whereas linear elasticity switched cells to
lobopodia-based migration. Thus, the relative polar-
ization of intracellular signaling identifies two distinct
modes of 3D cell migration governed intrinsically by
RhoA, ROCK, and myosin II and extrinsically by the
elastic behavior of the 3D extracellular matrix.
Nonpolarized signaling reveals two distinct modes
of 3D cell migration
Ryan J. Petrie,1 Núria Gavara,2 Richard S. Chadwick,2 and Kenneth M. Yamada1
1Laboratory of Cell and Developmental Biology, National Institute of Dental and Craniofacial Research; and 2Auditory Mechanics Section, National Institute on Deafness
and Other Communication Disorders; National Institutes of Health, Bethesda, MD 20892
This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No
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Noncommercial–Share Alike 3.0 Unported license, as described at http://creativecommons
T H E J O U R N A L O F C E L L B I O L O G Y
JCB • VOLUME 197 • NUMBER 3 • 2012 440
lobopodia (Kudo, 1977), an intracellular pressure-driven
protrusion (Fig. 1 C and Video 1; Yanai et al., 1996). This
morphology of human foreskin fibroblasts (HFFs) in der-
mal explants was similar to that of cells in wounded tissue
(Singer et al., 1984). In addition, cells often displayed small
blebs along their sides that we termed lateral blebs (Fig. 1 C,
arrowheads). Imaging a dual-chain Rac1 biosensor based
on intermolecular fluorescence resonance energy transfer
(FRET) revealed that, contrary to findings using 2D surfaces
(Kraynov et al., 2000), active Rac1 was not targeted to the
leading edge during HFF migration in both proximal or distal
regions of the explant (85%, n = 26; Rac1 polarization index
[PI] = 0.04 ± 0.09, in which 1 = forward polarization,
0 = nonpolarization, and 1 = rear polarization; Figs. 1 D,
S1, and S2). Thus, Rac1 activity was not polarized in elon-
gated HFFs migrating within the structurally heterogeneous,
physiological 3D environment of dermal explants. To study
the mechanistic basis of this apparently novel mode of
lamellipodia-independent 3D cell motility (hereafter referred
to as lobopodia-based migration), we recapitulated it using 3D
in vitro models of the ECM.
Lobopodia and lamellipodia in 3D in vitro
models of the ECM
The CDM contains fibronectin, collagen I and III, hyaluronic
acid, heparan sulfate proteoglycan, and thrombospondin in
parallel fibers 50–500 nm thick (Fig. 2 A; Hedman et al.,
1979; Allio and McKeown-Longo, 1988). Polymerized col-
lagen I forms a nonaligned 3D meshwork (Fig. 2 A) of single
or bundled collagen fibers 2–500 nm in diameter (Elsdale and
Bard, 1972; Gelman et al., 1979). We compared the mechani-
cal properties of dermal explants, the CDM, and collagen by
characterizing their stiffness and elastic behavior. Dermal ex-
plants (6,427 Pa, range of 277–19,400 Pa) and CDMs (627 Pa,
range of 224–2,454 Pa) were stiffer than 1.7 mg/ml collagen
(15 Pa, range of 11–21 Pa), did not undergo strain stiffen-
ing (Ehigh/Emed = 1.01), and were thus linearly elastic (Fig. 2,
B and C). In contrast, 1.7 mg/ml collagen displayed the strain-
stiffening behavior (Ehigh/Emed = 1.12) characteristic of nonlin-
ear elastic materials as previously established (Storm et al.,
2005). Cells migrating in the soft 1.7 mg/ml of 3D collagen
deformed the collagen fibers as described in a previous study
of collagen remodeling by motile cells (Grinnell and Lamke,
1984), whereas the arrangement of fibers within the stiffer
CDM was largely unaffected by migrating HFFs (Fig. 2 A).
Consistent with the similar stiffness and elastic behav-
ior of the CDM and dermal explants, the majority of HFFs
migrating inside the CDM also lacked obvious lamellipodia
and instead displayed the blunt cylindrical protrusions and lat-
eral blebs observed during lamellipodia-independent migra-
tion in dermal explants (77%, n = 31; Fig. 2 D and Video 2).
Lobopodia-based migration appeared specific to 3D because
HFFs migrating on top of the CDM (2D CDM) formed broad
lamellipodia with prominent ruffles (100%, n = 33; Fig. 2 D
and Video 2). In contrast to cells migrating in the 3D CDM,
cells migrating in 1.7 mg/ml of 3D collagen formed multiple
branched protrusions tipped with small lamellipodia (100%,
transition relates to normal 3D cell migration (Sanz-Moreno
and Marshall, 2010).
Some aspects of intracellular signaling organization
during cell migration in vivo can differ from the organization
seen on 2D surfaces. Chemotaxing primordial germ cells display
randomly distributed regions of RhoA activity and a uniform
distribution of PIP3 in the plasma membrane (Dumstrei et al.,
2004; Kardash et al., 2010). However, Rac1 activity is enriched
at the leading edge of migrating border cells and primordial
germ cells during development, and PIP3 is abundant at
the leading edge of neutrophils during interstitial migration
toward wounded tissue (Kardash et al., 2010; Wang et al.,
2010; Yoo et al., 2010). The reason for these differences
is not clear, but they may result from structural differences
in the surrounding ECM (Friedl and Wolf, 2010). Two
structural parameters that characterize the ECM are stiff-
ness, defined by the elastic or Young’s modulus (E; Engler
et al., 2006), and strain stiffening, a measurement of how
the stiffness of a material depends on the magnitude of
force applied to it (here measured as Ehigh/Emed; Storm et al.,
2005; Winer et al., 2009). Strain stiffening (Ehigh/Emed > 1)
is a form of nonlinear elasticity; thus, materials that do not
undergo strain stiffening (Ehigh/Emed = 1) are considered lin-
Tissue explants and in vitro models of the 3D ECM,
such as the cell-derived matrix (CDM) and type I collagen,
can closely mimic different complex tissue environments
(Elsdale and Bard, 1972; Cukierman et al., 2001; Even-Ram
and Yamada, 2005; Ahlfors and Billiar, 2007; Wolf et al.,
2009) and permit high-resolution live-cell imaging to visual-
ize intracellular signaling. We used primary human fibro-
blasts in these models to test the hypothesis that structurally
distinct 3D ECM environments support different modes of
normal cell migration. We find that the degree of polariza-
tion of PIP3 and Rho family GTPase signaling at the leading
edge identifies two distinct modes of normal cell motility
governed intrinsically by RhoA, Rho-associated protein kinase
(ROCK), and myosin II and extrinsically by the elastic behavior
of the ECM.
Lamellipodia-independent 3D migration
in dermal explants
Dermal tissue explants derived from mouse ears contain many
of the structural features found in the human dermis (Fig. 1 A;
Montagna et al., 1992; Lämmermann et al., 2008). Thick bun-
dles of collagen fibers are proximal to the basal explant sur-
face, whereas adipocytes and hair follicles embedded within
reticular collagen and elastic fibers tend to be more distal
(Fig. 1 B). Human fibroblasts migrating in both proximal and
distal regions of the explant were predominantly uniaxial,
consistent with cells migrating inside 3D in vitro models of
the ECM (Fig. 1 C; Bard and Hay, 1975; Cukierman et al.,
2001). The prominent lamellipodia and flat lamellae of cells
migrating in 2D were not apparent, and many cells instead
featured large blunt, cylindrical protrusions characteristic of
441Lobopodia- versus lamellipodia-based 3D motility • Petrie et al.
in the 3D CDM (Fig. 2 E). Similar distributions were observed
for vasodilator-stimulated phosphoprotein (VASP; Bear et al.,
2002) and F-actin (by phalloidin staining; Small, 1981).
Together, these data indicate that normal fibroblasts use at
least two distinct modes of 3D cell migration: lobopodia-based
migration in the stiff, linear elastic environment of dermal
explants and the 3D CDM and lamellipodia-based migration
in soft, nonlinear elastic 3D collagen.
n = 12; Fig. 2 D, right panels; and Video 3), consistent with
a previous study (Bard and Hay, 1975). To confirm that these
morphological differences corresponded to distinct cellular
structures, we compared the distribution of the actin-binding
protein cortactin (Kaksonen et al., 2000) in HFFs migrating
in collagen and the CDM. Although cortactin was enriched at
the leading edge during migration on the 2D CDM and in 3D
collagen, no such enrichment was detected during migration
Figure 1. Lobopodia-based 3D migration occurs in the mammalian dermis. (A) A 3D reconstruction of a mouse ear dermal explant labeled with Alexa
Fluor 633 (grayscale). Stratum corneum (SC), basal keratinocytes (BK), papillary dermis (PD), and reticular dermis (RD) are indicated. Sebaceous gland
(SG) and hair follicle (HF) are outlined in gray. (B) Examples of ECM structures proximal (left) and distal (right) to the basal surface of a dermal explant
labeled with Alexa Fluor 633. Images are from the same confocal stack, 9 µm (left) and 30 µm (right) from the basal surface. AC, adipocyte. (C) 3D
reconstructions of lobopodia-bearing HFFs migrating in proximal and distal collagen; GFP-actin is shown in green, and second harmonic imaging of col-
lagen appears in grayscale. Arrowheads indicate lateral blebs. (D) Active Rac1 is not targeted to the leading edge of HFFs migrating in the mammalian
dermis. Rac1 activity was imaged in HFFs migrating in proximal or distal ECMs; active Rac1, representing the Fc image, was pseudocolored according
to the 16-color scale shown to the right of the figure, and the explant was labeled with Alexa Fluor 633 (grayscale). All cells are oriented with the leading
edge toward the right of the figure. Bars, 5 µm.
JCB • VOLUME 197 • NUMBER 3 • 2012 442
Figure 2. CDM and type I collagen support lobopodia- and lamellipodia-based 3D migration, respectively. (A) HFF-generated CDM has an aligned,
fibrillar structure (top left), whereas polymerized 1.7 mg/ml type I collagen forms a random meshwork (bottom left). Both images are maximum projections
of 30-µm confocal stacks. Collagen is remodeled by migrating HFFs (GFP, green) along the axis of migration (bottom right), whereas the organization of
the CDM is unaffected by migrating HFFs (top right). The CDM was labeled with Alexa Fluor 633, and collagen was visualized by reflection microscopy.
(B) Matrix stiffness (Young’s modulus [E]) of the indicated 3D matrices. (C) Strain-stiffening (Ehigh/Emed) behavior of the indicated 3D matrices. Ehigh/Emed > 1
indicates nonlinear elasticity, whereas Ehigh/Emed = 1 indicates linear elasticity (dashed red line). (D) Collagen and 2D CDM support lamellipodia-based
migration, whereas 3D CDM triggers lobopodia-based motility. (top) Maximum projections of HFFs expressing GFP migrating inside the 3D CDM, on
top of the 2D CDM, or inside type I collagen. LM, lamellipodium; LB, lobopodium. (bottom) The orthogonal views of the corresponding panel above,
with the CDM (Alexa Fluor 633) and type I collagen (reflection microscopy) in red. Arrowheads indicate lateral blebs. (E) Cortactin is not enriched at the
leading edge during lobopodia-based migration. HFFs migrating in the indicated ECM were fixed and immunostained for cortactin. Arrows indicate the
local accumulation of cortactin at the leading edge. Bottom graphs correspond with their respective top images and represent the mean cortactin intensity
measured from the leading edge (0 µm) toward the cell center. Each cortactin intensity profile was averaged from 13 cells, with three measurements per
cell. Bars: (A and E) 5 µm; (D, top left and middle) 10 µm; (D, top right) 20 µm. All cells are oriented with the leading edge toward the right of the figure.
Error bars show means ± SEM. *, P < 0.001 versus the dermal explant. a.u., arbitrary unit.
443Lobopodia- versus lamellipodia-based 3D motility • Petrie et al.
surfaces (Kraynov et al., 2000; Nalbant et al., 2004). However,
during lobopodia-based migration in the 3D CDM, active Rac1
and Cdc42 were no longer restricted to the leading edge; instead,
they were localized to patches of unknown function around the
perimeter of the cell (Rac1: PI = 0.22, n = 44; Cdc42: PI = 0,
n = 21; Fig. 3, C–F), consistent with the nonpolarized distribu-
tion of active Rac1 in HFFs in dermal explants (Fig. 1 D). The
distribution of Rac1 and Cdc42 activity in lobopodia could not
be attributed to failure of the biosensors because nonfunctional
versions reported uniform Fc patterns in HFFs on 2D and in
3D CDMs (100%; Fig. S2 D). Together, these data show that
the canonical polarizations of PIP3, Rac1, and Cdc42 signaling
during 2D migration are not necessary for lobopodia-based 3D
migration in dermal explants and the CDM but are observed
during lamellipodia-based migration in 3D collagen.
RhoA, ROCK, and myosin II are required
for lobopodia-based 3D migration
To define further the mechanistic basis of lobopodia-based 3D
migration, the roles of Rac1, Cdc42, and RhoA in motility in
the 3D CDM were tested by specific siRNA-mediated knock-
down (Figs. 4, A and B; and S4). Reducing Rac1 or Cdc42
protein levels moderately increased or decreased the velocity
of HFFs migrating in and on the CDM, respectively, without
affecting the mode of 3D cell motility (Fig. 4, C and D). Cells
that were inside the 3D CDM continued to migrate without la-
mellipodia (94% for Rac1 siRNA and 92% for Cdc42 siRNA),
and many of the cells retained lateral blebs. In contrast to
Rac1 and Cdc42 siRNA, knockdown of RhoA protein and
activity dramatically switched the mode of migration in the 3D
CDM from lobopodia to lamellipodia based. RhoA-depleted
cells migrated in the 3D CDM with distinct lamellipodia at
the leading edge (100%; Figs. 4 C and S4 A and Video 4)
and without any lateral blebs. The role of RhoA in regulating
the mode of migration was confirmed with independent single
siRNAs (Fig. S4, B–D) and a single siRNA from the original
RhoA siRNA pool (Fig. S4, E–G). Additionally, treating cells
migrating in the 3D CDM with a RhoA inhibitor switched the
cells to lamellipodia-based motility (Fig. S4, H and I). Despite
switching the mode of 3D cell migration in the CDM, knock-
ing down RhoA did not affect the velocity of HFF migration
in or on the CDM (Fig. 4 D).
Chemical inhibition of the RhoA effector ROCK also
switched the mode of HFF migration in the 3D CDM, transi-
tioning cells from lobopodia- to lamellipodia-based migration
within 20 min of treatment (50%, n = 10; Video 5). Similar
to RhoA siRNA, ROCK inhibition switched the mode with-
out affecting the velocity of cells migrating in or on the CDM
(Fig. 4 E). In contrast, ROCK inhibition significantly reduced
HFF migration in 3D collagen, as previously observed for epi-
thelial cells (Provenzano et al., 2008). Thus, ROCK inhibition
functionally distinguishes lobopodia- from lamellipodia-based
3D motility. Inhibition of the ROCK target myosin II by bleb-
bistatin also switched the mode to lamellipodia-based migra-
tion in the 3D CDM (Fig. 4, G and H). In contrast to RhoA and
ROCK inhibition, myosin II inhibition significantly reduced the
velocity of elongated cells migrating in the 3D CDM, apparently
Nonpolarized signaling during
We investigated whether the distinct morphological features of
lobopodia- versus lamellipodia-based 3D migration were linked
to different signaling mechanisms. We compared the canoni-
cal polarization of PIP3, Rac1, and Cdc42 signaling during 2D
lamellipodia-based migration (Haugh et al., 2000; Kraynov
et al., 2000; Nalbant et al., 2004) by imaging the relative distribu-
tion of PIP3 and active Rac1 and Cdc42 in live HFFs migrating
in the CDM and collagen. The intracellular localization of PIP3
was imaged by binding of the pleckstrin homology (PH) domain
of Akt fused to GFP (Haugh et al., 2000). PIP3 was highly en-
riched near the leading edge of the elongated cells migrating on
top of the CDM (PI = 0.69, n = 13; Fig. 3, A and B), as reported
for cells migrating on 2D surfaces using lamellipodia (Haugh
et al., 2000). This enrichment was specific compared with local-
ization of phosphatidylinositol (4,5)-bisphosphate (PIP2) using
GFP-PLC-PH and was confirmed by immunolocalization of
PIP3 (Fig. S3; Várnai and Balla, 1998; van Rheenen and Jalink,
2002). Importantly, PIP3 was not polarized toward the leading
edge of HFFs using lobopodia-based motility in the CDM but
was instead distributed around the cell in clusters at the plasma
membrane in a pattern indistinguishable from PIP2 (PI = 0.05,
n = 18; Figs. 3, A and B; and S3). In contrast, PIP3 was concen-
trated at the leading edge of small lamellipodia during migra-
tion in 3D collagen (PI = 0.47, n = 13; Fig. 3, A and B). Thus,
lobopodia-based migration in the 3D CDM does not require the
polarization of PIP3 at the leading edge.
Active Rac1 and Cdc42 were imaged using dual-chain
FRET-based biosensors (Kraynov et al., 2000; Picard et al.,
2009). Both the single- and dual-chain versions of these bio-
sensors have been used to image the distribution of Rho family
GTPase activities in living cells (Pertz and Hahn, 2004). To
validate the FRET-based biosensors, we determined that their
expression in HFFs did not affect the amount or activity of the
corresponding endogenous GTPase (Boulter et al., 2010), and
the magnitude of the corrected FRET (Fc) signal imaged in
cells expressing constitutively active, nonfunctional, or wild-
type versions of the biosensors correlated with the amounts
of activity detected by a GTPase pull-down assay (Fig. S1, A,
B, D, and E; Benard et al., 1999). Plotting Fc versus donor
intensity for the constitutively active and nonfunctional ver-
sions of each biosensor showed the signal from the wild-type
Rac1 and Cdc42 biosensors could be considered positive when
Fc > 500 arbitrary units (Fig. S1, C and F). We confirmed that
their expression did not significantly affect cell velocity or down-
stream signaling (Fig. S2, A and B). Additionally, the subcellular
localization of the CFP-tagged Rac1 and Cdc42 was predomi-
nantly cytosolic and consistent with the corresponding endog-
enous protein (Fig. S2 C); excessive GTPase expression would
have resulted in inappropriate membrane targeting (Michaelson
et al., 2001).
Active Rac1 and Cdc42 were both polarized near the lead-
ing edge of cells on the 2D CDM (Rac1: PI = 0.73, n = 27;
Cdc42: PI = 0.49, n = 14; Fig. 3, C–F) and in 3D collagen
(Rac1: PI = 0.49, n = 34; Cdc42: PI = 0.38, n = 21; Fig. 3, C–F),
as previously reported for lamellipodia-based migration on 2D
JCB • VOLUME 197 • NUMBER 3 • 2012 444
Figure 3. Nonpolarized PIP3, Rac1, and Cdc42 signaling during lobopodia-based 3D migration. (A) Maximally projected confocal stacks of HFFs express-
ing GFP-AktPH to detect PIP3, migrating on the 2D CDM, in the 3D CDM, or inside 1.7 mg/ml of 3D collagen. Images were pseudocolored according to the
16-color scale. Bars, 10 µm. (B) Mean PI of GFP-AktPH in HFFs migrating in the indicated ECM environments. (C–F) Localization of active Rac1 and Cdc42
away from the leading edge during lobopodia-based 3D migration inside the CDM. (C and E) Maximally projected confocal stacks of HFFs expressing Rac1
(C) or Cdc42 (E) biosensors migrating on the 2D CDM (left), in the 3D CDM (middle), or in 3D collagen (right). The Fc images, representing the total activity of
445Lobopodia- versus lamellipodia-based 3D motility • Petrie et al.
each GTPase, were pseudocolored according to the 16-color scale. Arrowheads indicate regions of intracellular signaling. Bars: (3D CDM) 5 µm; (collagen
and 2D CDM) 10 µm. (D and F) Mean PI of active Rac1 (D) and active Cdc42 (F) in HFFs migrating in the indicated ECM environments. All cells are oriented
with the leading edge toward the top of the figure. Error bars show means ± SEM. *, P < 0.05 versus the 2D CDM; **, P < 0.05 versus the 3D CDM.
Figure 4. RhoA, ROCK, and myosin II are required for lobopodia-based 3D migration inside CDM. (A) A representative Western blot demonstrating the
specificity of siRNA-mediated knockdown of Rac1, Cdc42, or RhoA. HFFs were transfected with the indicated siRNAs and lysed 72 h after transfection,
and the lysates were blotted with the indicated antibodies. (B) Quantification of Western blots represented in A. (C) RhoA siRNA treatment switches HFFs
to lamellipodia-based 3D migration in the CDM. The percentage of lobopodia-bearing HFFs migrating inside the CDM after the indicated treatments.
(A and B) *, P < 0.001 versus the siGLO control. 48 h after siRNA treatment, HFFs were transfected with GFP-actin and imaged migrating in the 3D CDM.
(D) Quantification of the velocity of siRNA-treated HFFs in the CDM 72–84 h after transfection. *, P < 0.05 versus the siGLO control. (E) ROCK dependence
distinguishes HFF migration in collagen from the CDM. Quantification of the velocity of HFFs migrating in the CDM or 1.7 mg/ml collagen treated with
FBS or FBS + 10 µM Y-27632. *, P < 0.03 versus the FBS control. (F) Quantification of HFF velocity in the CDM when treated with FBS or FBS + 25 µM
blebbistatin. Blebbistatin treatment resulted in two subsets of HFFs, rapidly moving spread cells on top of the CDM (2D), and slowly moving elongated cells
inside the CDM (3D). *, P < 0.001 versus the FBS control. (G and H) Myosin II is required for 3D lobopodia-based migration. (G) Cortactin localization
in HFFs in the 3D CDM, either untreated or treated with 25 µM blebbistatin. The arrow indicates the local accumulation of cortactin at the leading edge.
Bars, 5 µm. (H) The mean cortactin intensity profile, measured from the leading edge (0 µm) toward the cell center, of cells treated with FBS or FBS +
25 µM blebbistatin. Each cortactin intensity profile was averaged from 13 cells, with three measurements per cell. All cells are oriented with the leading
edge toward the right of the figure. Error bars show means ± SEM. GAPDH, glyceraldehyde 3-phosphate dehydrogenase; a.u. arbitrary unit.
JCB • VOLUME 197 • NUMBER 3 • 2012 446
Figure 5. Lobopodia-based migration is distinct from cancer cell motility. (A) Amoeboid and mesenchymal HT1080 cells in the CDM. Phase-contrast
image showing amoeboid (rounded) and mesenchymal (elongated) HT1080 cells in the CDM. (B) The percentage of amoeboid and mesenchymal HT1080
cells in the CDM (n = 1,001). (C and D) HT1080 cells do not form lobopodia in the 3D CDM. Round amoeboid cells lack matrix adhesions, whereas
Lobopodia- versus lamellipodia-based 3D motility • Petrie et al.
elongated mesenchymal cells have prominent lamellipodia and matrix adhesions. Images show maximally projected confocal stacks of amoeboid (C) and
mesenchymal (D) HT1080 cells, expressing GFP-actin or YFP-paxillin, migrating inside the CDM (Alexa Fluor 633, grayscale). Arrowheads indicate matrix
adhesions. (E) Matrix adhesions are present during both lamellipodia- and lobopodia-based 3D migration of elongated normal fibroblasts. Maximally
projected confocal stacks of HFFs expressing YFP-paxillin or vinculin–tension sensor (TS) migrating inside 3D collagen or the 3D CDM. (F) Matrix adhesions
contribute to both lamellipodia- and lobopodia-based migration of normal fibroblasts. Blocking v3 and 1 integrins significantly decreased the velocity
and directionality of HFFs migrating in 3D collagen and CDM, indicating that integrin-mediated adhesion contributed to the efficient directional migration
of both cell populations. Quantification of the velocity (top) and directionality (bottom) of HFFs migrating on glass, CDM, or 3D collagen, either in media
or in media with 100 µM cyclic RGD (cRGD; an V3-blocking peptide) plus 500 µg/ml 1 integrin–blocking antibody (mAb13). *, P < 0.05 versus the
untreated control. All cells are oriented with the leading edge toward the right of the figure. Bars: (A) 50 µm; (C–E) 5 µm.
impeding translocation of the nucleus through the matrix,
whereas the migration of cells spread on top of the matrix was
unaffected (Fig. 4 F and Video 6) and was consistent with the
effect of blebbistatin on 2D fibroblast migration (Even-Ram
et al., 2007). Therefore, RhoA, ROCK, and myosin II form part
of the mechanistic basis of lobopodia-based migration. RhoA
and ROCK inhibition switched the mode of migration without
affecting the efficiency of migration, whereas myosin II activity
was required for both lobopodia formation and efficient migra-
tion in the 3D CDM.
3D matrix adhesions and lobopodia-
To determine that amoeboid cancer cell motility (Table S1)
was distinct from the lobopodia-based migration of normal
cells, despite their shared requirement for RhoA activity (Sahai
and Marshall, 2003), we imaged the HT1080 human fibrosar-
coma cell line within the CDM (Wolf et al., 2003). HT1080
cells adopted a mixture of amoeboid and mesenchymal mor-
phologies in aligned, fibrillar regions of the CDM (Fig. 5,
A–D) similar to those reported for breast cancer cells (Deakin
and Turner, 2011). Round amoeboid cells in the CDM lacked
3D matrix adhesions (90%, n = 20) but had large prominent
blebs (100%, n = 20; Fig. 5 C). Elongated mesenchymal cells
exhibited actin stress fibers and fan-shaped protrusions con-
sistent with lamellipodia (100%, n = 20; Fig. 5 D). These cells
also formed 3D matrix adhesions (100%, n = 20; Fig. 5 D). In
contrast to primary fibroblasts in the same 3D environment,
no lobopodia-bearing HT1080 cells were observed migrating
in the CDM. Unlike the two modes of cancer cell motility,
HFFs formed 3D matrix adhesions containing paxillin and
vinculin during both lamellipodia (100%, n = 18)- and lobo-
podia (100%, n = 20)-based 3D migration (Fig. 5 E). Blocking
v3 and 1 integrins significantly decreased the velocity and
directionality of HFFs migrating in and on the CDM and col-
lagen, indicating that integrin-mediated adhesion contributed
to the efficient directional migration of these cell populations
(Fig. 5 F). Therefore, lobopodia-based 3D migration was dis-
tinct from amoeboid and mesenchymal cancer cell motility,
based on the formation of 3D matrix adhesions, its integrin
dependence, morphology, and regulation by the ECM.
Matrix elastic behavior dictates the mode
of 3D migration
To define the relationship of matrix stiffness and/or elastic
behavior to the mode of normal 3D cell migration, we ma-
nipulated these physical characteristics without affecting the
organization of the matrices (Fig. 6, A–D). Trypsinization
decreased the stiffness of the CDM (8 Pa, range of 4–10 Pa)
and rendered it nonlinearly elastic (Ehigh/Emed = 1.23) compared
with the untreated CDM (Fig. 2, B and C) and permitted fiber
remodeling by migrating cells. Cross-linking the trypsinized
CDM increased the stiffness (143 Pa, range of 19–243 Pa;
Fig. 6, B and C), restored the linear elasticity of the matrix
(Ehigh/Emed = 1.03; Fig. 6 D), and prevented cells from remod-
eling the fibers. Although cross-linking the 1.7 mg/ml collagen
gels only marginally increased their stiffness (28 Pa, range of
12–201 Pa [Fig. 6 C] versus 15 Pa, range of 11–21 Pa for
uncross-linked 1.7 mg/ml collagen [Fig. 2 B]), it rendered the
material linearly elastic (Ehigh/Emed = 1.03; Fig. 6 D) compared
with uncross-linked 1.7 mg/ml collagen (Fig. 2 C). Elongated
fibroblasts used lamellipodia-based migration inside the tryp-
sinized CDM (92 versus 23% in the untreated CDM; Fig. 6 G
and Video 7), with Rac1 activity polarized toward the leading
edge (PI = 0.45, n = 17; Fig. 6, E and F). Cells migrated inside
the trypsinized and cross-linked CDM without polarization of
active Rac1 at the leading edge (PI = 0.11, n = 16; Fig. 6,
E and F), consistent with lobopodia-based motility (100%;
Fig. 6 G and Video 8). Importantly, cross-linked 1.7 mg/ml
collagen supported lobopodia-based migration of the major-
ity of cells (59 versus 0% in untreated collagen; Fig. 6 G and
Video 9), with a loss of polarized Rac1 activity in leading
protrusions (PI = 0.17, n = 6; Fig. 6, E and F).
To determine whether matrix stiffness, capacity for re-
modeling, or elastic behavior govern lobopodia formation, we
examined HFF migration in 8.6 mg/ml collagen. Cells in 8.6
mg/ml collagen underwent lamellipodia-based 3D migration,
including enrichment of cortactin at the leading edge (Fig. 6 H).
The 8.6 mg/ml collagen was substantially stiffer than 1.7 mg/ml
collagen (19×, P < 0.001) and was not remodeled during cell
migration, as expected (Fig. 6, B and C; and Video 10; Miron-
Mendoza et al., 2010), similar to the CDM. Significantly, like
1.7 mg/ml collagen, 8.6 mg/ml collagen exhibited nonlinear
elastic behavior, undergoing strain stiffening (Ehigh/Emed = 1.2;
Fig. 6 D). Together, these data show that 3D matrix linear elas-
ticity is a key structural property necessary for nonpolarized
Rac1 activity during lobopodia-based migration.
Lobopodia and efficient migration
in the CDM
Given that RhoA or ROCK inhibition in 10% FBS switched
the mode of HFF migration in the CDM without affecting ve-
locity, it was not clear why more than one mode of migration
was necessary. To test the hypothesis that lobopodial cells
migrated more efficiently in the linear elastic ECM under sub-
optimal environmental conditions, we assessed HFF migration
JCB • VOLUME 197 • NUMBER 3 • 2012 448
Figure 6. Matrix elastic behavior governs the mode of normal 3D migration. (A) HFFs can remodel trypsinized CDM (middle), whereas the structure
of the untreated (left) or the trypsinized (tryp) and cross-linked CDM (XL; right) is unaffected by HFF migration. CFP-Rac1 is shown in green, and Alexa
Fluor 633–labeled matrix is in gray. Bars, 40 µm. (B) Cross-linked 1.7 mg/ml collagen (middle) and 8.6 mg/ml collagen (right) are not remodeled
during HFF migration. GFP-actin is shown in green, and collagen is shown in gray (reflection). Bars, 10 µm. (C) Matrix stiffness (Young’s modulus [E])
of the indicated modified matrices. (D) Strain-stiffening (Ehigh/Emed) behavior of the indicated native and modified matrices. The dashed red line indicates
a value of Ehigh/Emed corresponding to 1 (linear elasticity). (C and D) *, P < 0.05 versus trypsinized CDM; **, P < 0.05 versus cross-linked collagen.
(E–G) Trypsinization and chemical cross-linking redistribute Rac1 activity and switch the mode of cell migration. (E) Maximally projected confocal stacks
of HFFs expressing the Rac1 biosensor migrating in the trypsinized CDM (top), trypsinized and cross-linked CDMs (middle), or cross-linked collagen
(bottom). The Fc images, representing the total activity of each GTPase, were pseudocolored according to the 16-color scale. Arrowheads indicate
regions of intracellular signaling. Bars, 5 µm. (F) Mean PI of active Rac1 in HFFs migrating in the indicated ECM environments. *, P < 0.05 versus
3D trypsinized CDM. (G) The percentage of lobopodia-bearing HFFs migrating inside the CDM or collagen treated as indicated. *, P < 0.001 versus
untreated CDM; **, P < 0.001 versus trypsinized CDM; ***, P < 0.007 versus untreated collagen. (H) 8.6 mg/ml collagen supports lamellipodia-
based 3D cell migration. (top) A representative image of cortactin enrichment at the leading edge of cells migrating in 8.6 mg/ml collagen (arrow).
(bottom) The mean cortactin intensity profile measured from 13 cells, with three measurements per cell. Cells are oriented with their leading edge toward
the top right (A and B) or the right (E and H) of the figure. a.u., arbitrary unit.
449Lobopodia- versus lamellipodia-based 3D motility • Petrie et al.
driven by PDGF in the presence of different levels of glucose.
10 ng/ml PDGF with 25 mM glucose was sufficient to replace
10% FBS for rapid migration in and on the CDM and collagen
compared with DME alone, with the cells in the CDM form-
ing lobopodia (Fig. 7, A–C). However, treatment of cells with
PDGF in glucose-deficient medium reduced cell velocity in
and on the CDM without affecting the velocity of lamellipodia-
based migration in collagen (Fig. 7 A). This loss of migration
efficiency in and on the CDM was associated with a switch to
lamellipodia-based 3D motility (Fig. 7, B and C) and a reduc-
tion of RhoA activity to 51% of control, significantly less than
in cells treated with PDGF with glucose (P < 0.05; Fig. 7 D).
Therefore, lobopodia may be required for efficient motility in
the CDM under low-glucose conditions. Thus, the external
signaling environment can regulate the mode and efficiency
of normal cell migration in the CDM as well as the relative
activation of RhoA.
The major differences in polarization of active Rac1, Cdc42,
and PIP3 identified in this study reveal lobopodial and lamelli-
podial migration as two distinct modes of normal 3D cell mo-
tility governed intrinsically by RhoA, ROCK, and myosin II
and extrinsically by the elastic behavior of the ECM (Fig. 8 A).
The lobopodia we find associated with 3D mesenchymal cell
motility appear to be distinct from other cellular protrusions in ad-
dition to lamellipodia, including filopodia (Nobes and Hall, 1995),
Figure 7. Regulation of the mode and efficiency of normal 3D cell migration by extracellular soluble factors. (A) 10 ng/ml PDGF in glucose-deficient media
specifically reduces cell velocity in the CDM versus collagen. Quantification of cell velocity in the CDM or collagen in response to DME with 25 mM glucose
(control), 10% FBS in DME with 25 mM glucose (FBS), 10 ng/ml PDGF in DME with 25 mM glucose (PDGF), or 10 ng/ml PDGF in glucose-deficient DME
(PDGF glucose). *, P < 0.05 versus FBS. (B) Representative images of cortactin localization in HFFs in the 3D CDM treated as indicated and quantified
in C. Arrows indicate the local accumulation of cortactin at the leading edge. Bottom graphs correspond with their respective top images and represent the
mean cortactin intensity measured from the leading edge (0 µm) toward the cell center. Each cortactin intensity profile was averaged from 13 cells, with
three measurements per cell. (C) Quantification of the percentage of cells without enrichment of cortactin at the leading edge in B. *, P < 0.001 versus
FBS. (D) Treatment of HFFs with 10 ng/ml PDGF in glucose-deficient media reduces cellular RhoA activity by 50%. RhoA activities in HFFs treated on
tissue-culture plastic as indicated were measured using G-LISA activation assays. Absorbance values were normalized to the relative amount of actin in each
sample before comparison with the FBS treatment. *, P < 0.01 versus FBS; **, P < 0.05 versus PDGF. Cells are oriented with their leading edge toward
the right of the figure. Bars, 5 µm. a.u., arbitrary unit.
JCB • VOLUME 197 • NUMBER 3 • 2012 450
and -independent migration in normal mesenchymal cells as
well as its regulation.
We demonstrate that the canonical polarization of PIP3,
Rac1, and Cdc42 activity during 2D migration (Ridley et al.,
2003) is not required for normal mesenchymal cells to migrate
equally efficiently in the 3D ECM. During lobopodia-based
migration in the 3D CDM, nonpolarized Rac1 and Cdc42
activity can modulate the velocity of migration independently
of lamellipodia formation. Enrichment of active Rac1 and
Cdc42, along with PIP3, at the tips of fan-shaped protrusions
during 3D collagen migration identified these structures as
podosomes (Tarone et al., 1985), invadopodia (Mueller et al.,
1992), eupodia (Fukui and Inoué, 1997), and membrane
blebs (Charras et al., 2005). The decision between lamellipodia-
and lobopodia-based motility during normal mesenchymal
cell migration can be conceptualized as a series of three
binary questions evaluated by migrating cells (Fig. 8 B):
what is the dimensionality of the matrix, what is the level
of RhoA activity, and is the 3D matrix linearly elastic? This
study also provides the first description of lobopodia-based
movement coupled to 3D matrix adhesions and the first
demonstration of a transition between lamellipodia-dependent
Figure 8. Dimensionality, matrix elastic behavior, and RhoA–ROCK–myosin II govern the mode of normal cell migration. (A) During cell migration on flat or
fibrillar 2D surfaces, lamellipodia form the leading edge of adherent fibroblasts. The presence of lamellipodia was confirmed by a prominent rim of F-actin
and cortactin, along with active Rac1 and Cdc42 with PIP3 at the leading edge. In a 3D ECM, HFFs can use either lobopodia- or lamellipodia-based 3D
migration. (B) The choice to migrate using lobopodia- or lamellipodia-based migration can be represented by a decision tree consisting of three questions:
what is the dimensionality of the matrix, what is the level of RhoA activity, and is the 3D matrix linearly elastic? Lobopodia-based 3D migration predomi-
nates in linear elastic ECM and may use high actomyosin contraction downstream of RhoA–ROCK–myosin II to increase intracellular pressure and push
the leading edge forward in combination with integrin-mediated adhesion. When RhoA–ROCK–myosin II activity is diminished, either in nonlinear ECM or
through treatment of cells with RhoA siRNA or specific inhibitors, cells form lamellipodia with actin polymerization to advance the leading edge. Both modes
of migration involve elongated cells that form 3D matrix adhesions, but the distribution of active Rac1, Cdc42, and PIP3 distinguishes the two modes.
451Lobopodia- versus lamellipodia-based 3D motility • Petrie et al.
cancer cell migration could represent a regulatory defect pro-
moting metastatic disease.
In summary, the differential organization of intracellular
signaling in lobopodia and lamellipodia identifies these cellular
structures as mediating two distinct modes of normal 3D cell
migration. The transition between these modes can be regulated
externally by the elastic behavior of the ECM via intracellular
RhoA, ROCK, and myosin II, and it may represent a universal
property of 3D cell motility.
Materials and methods
Reagents, cell culture, and transfection
The following reagents were used in this study: rhodamine-phalloidin
(Invitrogen), Y-27632 (EMD), blebbistatin (EMD), cell-permeable C3 trans-
ferase (Cytoskeleton), human recombinant PDGF-BB (Sigma-Aldrich), and
glucose-free DME (Invitrogen). HFFs (used at passages 8–20) and HT1080
cells were maintained in phenol red–free DME (HyClone) containing 10%
FBS (HyClone), 4.5 g/liter glucose, 100 U/ml penicillin, 100 µg/ml
streptomycin (Invitrogen), and 2 mM l-glutamine (Invitrogen) at 37°C and
in 10% CO2. All cDNA constructs were transfected into cells with the
Nucleofector system (Lonza) using the human dermal fibroblast kit
(Lonza) according to the manufacturer’s instructions. siRNA was trans-
fected into HFFs using Lipofectamine 2000 (Invitrogen) as previously
described (Pankov et al., 2005).
cDNA constructs and siRNAs
The pECFP-Rac1 and -Cdc42 constructs (Picard et al., 2009) were gener-
ated by subcloning the full-length sequence into the EcoRI–BamH1 sites
of pECFP-C1 (Takara Bio Inc.). pYPet–p21-binding domain (PBD) was
generated by subcloning the sequence encoding amino acids 65–150
of human Pak1 into the EcoRI–BamH1 sites of pYPet-C1. pYPet-C1 was
constructed by subcloning the sequence corresponding to the fluorescent
protein YPet into the AgeI–XhoI sites of pEGFP-C1 (Takara Bio Inc.). The
YPet sequence was amplified from pCEP4YPet-MAMM (plasmid 14032;
Addgene; Nguyen and Daugherty, 2005). pGFP-actin was purchased
from Takara Bio Inc. pGFP-PLC-PH was a gift from T. Balla (National
Institute of Child Health and Human Development, Bethesda, MD), who
generated it by subcloning the cDNA sequence corresponding to the
PH domain of PLC-1 (amino acids 1–170) into pEGFP-N1 (Takara Bio Inc.;
Várnai and Balla, 1998). To create pGFP-AktPH, the Akt PH domain
(amino acids 1–148) was subcloned into pEGFP-C1 (Pankov et al.,
2005). Vinculin–tension sensor (plasmid 26019; Addgene) was gen-
erated by Grashoff et al. (2010) by inserting a FRET-based sensor
module between the vinculin head and vinculin tail domains of vincu-
lin. pYFP-paxillin was generated by subcloning the sequence encoding
human paxillin into the HindIII–XbaI sites of pEYFP-C1 (Takara Bio Inc.)
that contained a modified multiple cloning site.
Rac1, 5-GAACUGCUAUUUCCUCUAA-3, 5-AUGAAAGUGU-
CACGGGUAA-3, 5-GUAGUUCUCAGAUGCGUAA-3, and 5-GUGAU-
UUCAUAGCGAGUUU-3; Cdc42, 5-CGGAAUAUGUACCGACUGU-3,
and 5-CUGCAGGGCAAGAGGAUUA-3; and RhoA, 5-CGACAG-
CCCUGAUAGUUUA-3, 5-GACCAAAGAUGGAGUGAGA-3, 5-GCA-
GAGAUAUGGCAAACAG-3, and 5-GGAAUGAUGAGCACACAAG-3
ON-TARGETplus SMARTpool siRNAs, along with siGLO RNA-induced silenc-
ing complex–free control siRNA, were purchased from Thermo Fisher Scien-
tific. The specificities of the siRNA pools were confirmed with the following
independent siRNAs: Rac1, 5-GGAACUAAACUUGAUCUUATT-3; Cdc42,
5-UGAGAUAACUCACCACUGUTT-3; RhoA, 5-CACAGUGUUUGAGA-
ACUAUTT-3 (Silencer Select; Invitrogen); and the individual RhoA SMART-
pool siRNA, 5-CGACAGCCCUGAUAGUUUA-3.
Live-cell imaging in dermal explants
Dermal explants (Lämmermann et al., 2008) were prepared by separat-
ing the dorsal and ventral halves of ears from euthanized ICR (Institute of
Cancer Research) mice. The cartilage-free halves were washed extensively
in PBS plus 100 U/ml penicillin and 100 µg/ml streptomycin. For some
experiments, the explant was labeled as previously described for the CDM
(Hakkinen et al., 2011). In brief, explants were incubated with 20 µg of
the succinimidyl ester of Alexa Fluor 633 (Invitrogen) in 2 ml of 50-mM
NaHCO3 for 20 min. The labeled explant was washed with PBS plus
lamellipodia, consistent with a previous morphological study
(Small, 1981). These features in 3D collagen mimic the 2D
signaling and lamellipodial dynamics that control directional
cell movement (Haugh et al., 2000; Kraynov et al., 2000;
Nalbant et al., 2004; Pertz et al., 2006; Petrie et al., 2009).
Despite the nonpolarized distribution of Rac1, Cdc42, and PIP3
intracellular signaling, lobopodial cells can still be considered
polarized based on their elongated morphology and direction-
ally persistent migration in the 3D CDM. Although the mech-
anism that sustains this polarity is not clear, the polarization
of the microtubule-organizing center, Golgi apparatus, and/or
membrane trafficking might be maintained in these cells inde-
pendent of the distribution of PIP3, Rac1, and Cdc42 activity
(Kupfer et al., 1982; Bergmann et al., 1983; Gundersen and
Lobopodia formation was governed intrinsically by RhoA,
ROCK, and myosin II, consistent with other contractility-based
modes of cell migration (Sahai and Marshall, 2003; Klopocka
and Redowicz, 2004; Yoshida and Soldati, 2006; Lämmermann
et al., 2008). In fact, the lateral blebs we often observed dur-
ing lobopodia-based motility, which were absent after RhoA
or ROCK inhibition, may be a manifestation of increased cy-
toplasmic pressure (Charras et al., 2005) because of RhoA-
mediated actomyosin contraction (Chrzanowska-Wodnicka
and Burridge, 1996; Lämmermann and Sixt, 2009). Lobopo-
dial migration was controlled extrinsically by soluble signals
and the elastic behavior of the ECM. Cells in the CDM with-
out motogens present formed lamellipodia, but these structures
were not associated with significant cell movement. Thus,
normal fibroblasts require a motogenic signal, e.g., 10% FBS
or 10 ng/ml PDGF with 25 mM glucose, in combination with
the linear elastic 3D ECM to undergo lobopodia-based migra-
tion. Stimulation with PDGF in glucose-deficient medium or
inhibition of RhoA, ROCK, or myosin II switched the mode of
migration independently of the elastic behavior of the matrix.
Interestingly, we found that lobopodia were associated with
efficient migration in the CDM when motility was triggered in
low glucose, a condition that did not affect lamellipodia-based
migration in 3D collagen. This suggests that lobopodia are
required for rapid movement in the linear elastic environment
of the CDM under certain environmental conditions. However,
rapid motility in the CDM did not depend on a single mode of
migration in 25 mM glucose, suggesting that lobopodia might
also be involved in other aspects of normal fibroblast function,
such as matrix production and remodeling.
An example of migration mode regulation is the amoeboid–
mesenchymal transition during cancer cell migration (Wolf
et al., 2003). Importantly, lobopodia-bearing HFFs differ
from amoeboid cancer cells on the basis of morphology, the
use of 3D matrix adhesions, and their regulation by the ECM
(Table S1; Wolf et al., 2003; Sabeh et al., 2009). Determining
whether the transition between lobopodia- and lamellipodia-
based integrin-dependent migration is restricted to normal
cells may help us to understand what signaling pathways are
subverted to promote metastasis through the mesenchymal–
amoeboid transition (Bissell, 1981; Sanz-Moreno and Marshall,
2010). For example, the integrin independence of amoeboid
JCB • VOLUME 197 • NUMBER 3 • 2012 452
Live-cell imaging in 3D in vitro models
CDMs were prepared from HFFs as follows (Cukierman et al., 2001).
MatTek dishes were coated with 0.2% gelatin for 1 h at 37°C, treated with 1%
glutaraldehyde for 30 min at RT, and incubated with DME for 30 min at RT.
Three washes with PBS followed each treatment. 4 × 105 HFFs were plated
per MatTek dish, which were maintained for 10 d, adding fresh media with
50 µg/ml ascorbic acid every other day. The cells were removed from the
CDM with extraction buffer (20 mM NH4OH and 0.5% Triton X-100 in PBS)
for 5 min at RT and washed with PBS. The cell-free CDM was treated with
10 U/ml DNase (Roche) for 30 min at 37°C, washed, and stored at 4°C in PBS
with 100 U/ml penicillin and 100 µg/ml streptomycin. 105 HFFs, transfected
with the indicated constructs, were plated on the CDM and blocked with 1%
heat-denatured BSA (MP Biomedicals) for imaging the next day. For some
experiments, the CDM was labeled with 5 µg Alexa Fluor 633 in 2 ml of
50-mM NaHCO3 as for dermal explants immediately before the addition of
HFFs. 1.7 mg/ml collagen gel was prepared by combining 3.32 mg/ml rat
tail type I collagen (BD) with 10× reconstitution buffer (0.26 M NaHCO3
and 0.2 M Hepes) and 10× DME (Sigma-Aldrich) at an 8:1:1 ratio, the pH
was adjusted to 7.5, and the collagen was diluted to 1.7 mg/ml with media.
MatTek dishes were coated with 50 µl of 1.7-mg/ml collagen, which was
then polymerized at 37°C for 30 min. 200 µl of 1.7-mg/ml collagen contain-
ing 105 HFFs, transfected the previous day with the indicated constructs, was
added to the first layer and polymerized at 37°C for 30 min. 2 ml of medium
was added to the dish, and cells were imaged the next day.
Time-lapse fluorescence imaging of GFP-actin was performed as
described for dermal explants. Cells were determined to be fully within the
3D CDM when covered with labeled matrix and within 3D collagen when
above the glass surface of the MatTek dish. When determining the percent-
age of amoeboid and mesenchymal HT1080 cells in the CDM, a cell was
considered mesenchymal when its length was at least twice its width, and
amoeboid was determined when less (Sanz-Moreno et al., 2008).
Confocal stacks were captured using the microscope (LSM 510 NLO
META Axiovert 200M) with a Plan Apochromat 40×, 1.0 NA oil iris objec-
tive as described in the Live-cell imaging in dermal explants section, except
reflection of the 514-nm argon laser was used to visualize polymerized type I
collagen, and a 457- and 514-nm argon laser was used to excite the FRET
pair. Maximum projections of the confocal stacks were generated using Zen
software (Carl Zeiss).
Live-cell FRET imaging
The binding of active CFP-Rac1 or -Cdc42 to YPet-PBD was detected by
imaging the FRET-dependent sensitized emission of the acceptor fluorophore
(YPet) in the presence of the donor fluorophore (CFP; Kraynov et al., 2000).
Optimal FRET acquisition settings were independently determined for each
microscope and strictly maintained during all subsequent FRET imaging.
The spectral bleed-through ratios were determined for each microscope by
imaging cells expressing donor or acceptor alone using the microscope’s
optimized acquisition settings. All FRET image processing was performed
using the LSM FRET Tool macro (Carl Zeiss). The Fc image was generated
from the raw FRET image using
Ff Df Fd DdAf Fa Aa
Da Fa Fd_
_ _ Dd
in which Ff is the raw FRET image, Df is the CFP image, Af is the YPet image,
Fd_Dd is the proportion of CFP emission in the FRET image, Fa_Aa is the pro-
portion of YPet emission in the FRET image, Da_Aa is the proportion of YPet
emission in the donor image, Da_Fa is the cross talk coefficient determined by
dividing the donor image by the FRET image of cells expressing YPet alone,
and G is the system constant (set to 1; Gordon et al., 1998). Each channel
image was thresholded before processing, and saturated and zero intensity
pixels were excluded from the calculations. Maximum projections of the Fc con-
focal z stacks were generated using Zen software. Although the output of the
Fc image was linearly adjusted using ImageJ 1.40g before application of the
16-color spectrum to fill the display range and show relative differences in FRET
intensity, positive Rac1 and Cdc42 Fc images (12-bit; 0–4,096 fluorescence
intensity range [arbitrary units]) were typically between 700 and 1,300.
HFFs were transfected as indicated using PolyFect according to the manufac-
turer’s instructions. The next day, FRET images were captured of the live cells, and
the Fc image was calculated as described in the Live-cell FRET imaging section
of Materials and methods. The magnitude of the Fc signal in the transfected
cells was measured in unadjusted Fc images using the LSM FRET Tool macro.
100 U/ml penicillin and 100 µg/ml streptomycin, and any unreacted
Alexa Fluor 633 dye was quenched with 200 mM Tris, pH 7.4, for 20 min
and washed with PBS. 5 × 105 HFFs were transfected with the indicated
constructs and plated in a 60-mm dish. 18 h later, the transfected cells
were added to the unlabeled or Alexa Fluor 633–labeled dermal explants
immediately after explant preparation. The next day, the explants with
adherent HFFs were transferred to a new dish and imaged.
Image stacks of dermal explants in media at 37°C and 10% CO2
were captured using a confocal microscope (LSM 510 NLO META with
Axiovert 200M; Carl Zeiss) with a Plan Apochromat 63×, 1.4 NA oil ob-
jective lens (Carl Zeiss). A 488-nm argon and 633-nm HeNe2 laser (Carl
Zeiss) excited GFP-actin and Alexa Fluor 633, respectively. The two-photon
laser was set to 800 nm for visualizing collagen through second harmonic
generation (Friedl et al., 2007). Image stacks were imported into Volocity
software (PerkinElmer) to generate the 3D reconstructions.
Time-lapse fluorescence imaging of GFP-actin–transfected HFFs
in dermal explants in media at 37°C and 10% CO2 was performed by
spinning-disc confocal microscopy using a microscope (Axiovert 200M)
equipped with a confocal scanning unit (CSU-X1; Yokogawa), a Plan
Apochromat 100×, 1.4 NA oil objective (Carl Zeiss), and an EM charge-
coupled device camera (C9100; Hamamatsu Photonics). Hardware con-
trol, image acquisition, and linear brightness and contrast adjustments
were performed using MetaMorph software (Molecular Devices). GFP-
actin was imaged with a 488-nm argon laser, and a 647-nm krypton-
argon laser was used for Alexa Fluor 633.
For FRET imaging, HFFs were transfected with pYPet-PBD and pCFP-
Rac1 and plated on Alexa Fluor 633–labeled dermal explants. The next day,
the explants with HFFs were transferred to a new dish and imaged in media at
37°C and 10% CO2 using a confocal microscope (LSM 710 Axio Examiner.
Z1; Carl Zeiss) equipped with a C Achroplan 40×, 0.8 NA water objective
lens (Carl Zeiss). A 457- and 514-nm argon laser was used to excite the FRET
pair, and a 633-nm HeNe laser excited Alexa Fluor 633. FRET images were
processed as described in the Live-cell FRET imaging section.
GTPase activation assays
pECFP-Rac1 and -Cdc42 were mutated using the site-directed mutagenesis
kit (QuikChange II; Agilent Technologies) to create the following constructs:
pECFP-Rac1-Q61L (constitutively active) and -Rac1-Y40C (nonfunctional)
and pECFP-Cdc42-Q61L (constitutively active) and -Cdc42-Y40C (nonfunc-
tional). HFFs were either untransfected or transfected with the indicated
constructs using transfection reagent (PolyFect; QIAGEN) according to the
manufacturer’s instructions. Pull-down or G-LISA assays were performed to
determine Rac1 or Cdc42 activity using the appropriate activation assay kit
from Cytoskeleton following the manufacturer’s instructions.
Immunofluorescence labeling of fixed cells
The following antibodies were used: mouse anticortactin (Millipore),
mouse anti-VASP (BD), mouse anti-PIP2 (Echelon), and mouse anti-PIP3
(Echelon). Mouse anti-Rac1 (Millipore) and rabbit anti-Cdc42 (Abcam)
were used for immunofluorescence analysis of endogenous GTPases.
To localize endogenous Rho family GTPases, PIP2, or PIP3, HFFs were
fixed and stained following a published protocol for preservation of the
plasma membrane during immunocytochemistry (Hammond et al., 2009)
but with slight modifications. HFFs were fixed with 4% formaldehyde
and 0.2% glutaraldehyde in PBS for 15 min at RT, rinsed 3× with
50 mM NH4Cl in PBS, placed in an aluminum foil–lined tissue-culture dish
on ice, and chilled for 2 min, with all subsequent steps performed on ice
with prechilled solutions. Fixed HFFs were treated with buffer A (1% BSA,
50 mM NH4Cl, 0.5% saponin, and 0.005% Triton X-100 in PBS) for
45 min. Antibodies were applied in buffer B (1% BSA, 50 mM NH4Cl, and
0.1% saponin in PBS) for 1 h. Cells were washed 2× with buffer C (1%
BSA, 50 mM NH4Cl, and 0.5% saponin in PBS). IgG secondary antibod-
ies (Jackson ImmunoResearch Laboratories, Inc.) were applied in buffer
B for 45 min. Cells were washed 4× with buffer C and postfixed with
2% formaldehyde in PBS for 10 min on ice and then 5 min at RT. Cells
were washed 3× in 50 mM NH4Cl in PBS and 2× in distilled H2O before
drying and mounting with mounting media (Gel/Mount; BioMeda). For
VASP, cortactin, or rhodamine-phalloidin labeling, cells were fixed with
4% formaldehyde in PBS, permeabilized with 0.25% Triton X-100 in PBS,
and blocked with 0.2% BSA in PBS. Rhodamine-phalloidin and primary
and secondary antibodies were applied in 0.2% BSA in PBS and washed
with PBS between each treatment. The fixed cells were imaged using the
microscope (510 NLO META Axiovert 200M) with a Plan Apochromat
63×, 1.4 NA oil objective. Brightness and contrast were linearly adjusted
using ImageJ 1.40g (National Institutes of Health).
453Lobopodia- versus lamellipodia-based 3D motility • Petrie et al.
regions of the force displacement curves corresponding to large indentations
or medium indentations. The ratio of Ehigh/Emed was reported as a measure of
the matrix’s strain-hardening behavior. All computations were performed using
custom-built code written in Matlab (MathWorks).
Modification of the 3D in vitro models
Alexa Fluor 633–labeled CDM was treated with 0.05% trypsin + 0.5 mM
EDTA (Invitrogen) for 10 min at 37°C and then washed extensively with
DME with 10% FBS. 1.7 mg/ml collagen and trypsinized CDM were cross-
linked with 25 mM BS(PEG)9 (bis-N-succinimidyl-(nonaethylene glycol)
ester) and BS(PEG)5 (Thermo Fisher Scientific), respectively, in PBS for 30 min
at RT followed by extensive washing with DME with 10% FBS. 105 HFFs
transfected with pCFP-Rac1 and pYPet-PBD or pGFP-actin were plated on
the trypsinized and/or cross-linked matrix and imaged the next day. GFP-
actin dynamics were analyzed by spinning-disc confocal microscopy, and
Rac1 activity was visualized using the confocal microscope (LSM 710 Axio
Examiner.Z1) as described in Live-cell imaging in dermal explants.
Results are reported as the mean ± SEM. One-way analysis of variance with
Tukey posttests was used to compare three or more variables; otherwise, un-
paired, two-tailed Student’s t tests were performed. Nonparametric Kruskal–
Wallis tests were used with a Dunn’s multiple comparisons posttest when
comparing more than two groups of measurements with bounded values
when analyzing the PI. All comparisons were performed using Prism5
(GraphPad Software). Differences were considered statistically significant
at P < 0.05.
Online supplemental material
Fig. S1 and S2 detail the validation of the Rac1 and Cdc42 FRET-based
biosensors. Fig. S3 shows the relative distribution of GFP-PLC-PH in HFFs
in different environments, along with endogenous PIP2 and PIP3. Fig. S4
provides additional evidence that RhoA activity is required for lobopodia
formation in the 3D CDM. Table S1 compares the lobopodia- and
lamellipodia-based 3D migration of normal cells with previously published
work examining cancer cell motility. Video 1 shows lobopodia-based 3D
migration in the mammalian dermis. Video 2 shows lobopodia-based
3D migration in the CDM and lamellipodia-based 2D migration on top of
the CDM. Video 3 that shows HFFs use lamellipodia-based 3D migration in
type I collagen. Video 4 shows that reduction of RhoA protein switches the
mode to lamellipodia-based 3D migration in the CDM. Video 5 shows
that ROCK activity is required for lobopodia-based migration. Video 6
is a phase-contrast video of HFFs migrating in and on the CDM during
blebbistatin treatment. Video 7 shows that HFFs use lamellipodia-based
3D migration in a pliable CDM. Video 8 shows that cross-linking the
trypsinized CDM restores lobopodia-based 3D migration. Video 9 shows
that HFFs use lobopodia-based 3D migration in cross-linked collagen.
Video 10 shows lamellipodia-based 3D HFF migration in 8.6 mg/ml
collagen. Online supplemental material is available at http://www.jcb
We thank Myungso Chung for helping to develop the glucose-deficient
migration assays, Dr. Carole Parent for helpful discussions, and Dr. Andrew
Doyle, Dr. Sarah Knox, and Matthew Kutys for critical comments on
This work was supported by the Intramural Research Programs of the Na-
tional Institute of Dental and Craniofacial Research and the National Institute on
Deafness and Other Communication Disorders at the National Institutes of Health.
Submitted: 23 January 2012
Accepted: 30 March 2012
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To determine the PI of a cell, its center of mass was placed at the origin of an
x–y axis, with the leading edge perpendicular to the positive x axis. The posi-
tions of discrete regions of Fc (considered positive when >500 but typically
between 700 and 1,300) or GFP-AktPH intensity (considered positive when
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siRNA treatment, Western blotting, and motility assays
2 × 105 HFFs were plated in each of three 60-mm dishes and transfected
the next day with a 20-nM solution of the indicated siRNA preparation.
48 h later, 105 siRNA-treated cells from the first 60-mm dish were trans-
fected with pGFP-actin, plated on the CDM labeled with Alexa Fluor 633,
and imaged the next day using a spinning-disc confocal microscope. 104
HFFs from the second 60-mm dish were plated in 10% FBS in DME on the
CDM. The next day, time-lapse sequences were captured at 37°C and
10% CO2 using a 5×, 0.12 NA A-Plan objective on a microscope (Axio-
vert 40C; Carl Zeiss) with a charge-coupled device camera (INFINITY2;
Lumenera). Cells were tracked every 26 min for 12 h using the Manual
Tracking plugin (F. Cordelieres, Institut Curie, Paris, France) with ImageJ
1.40g. Velocity and directionality (the ratio of Euclidean to accumulated
distance traveled, in which 1 = a straight line) were calculated from the
tracking data using the Chemotaxis and Migration Tool plugin (ibidi) with
ImageJ. For the integrin, ROCK, or myosin II inhibition experiments or
PDGF ± glucose motility assays, 104 HFFs were plated on glass, the CDM,
or in 1.7 mg/ml collagen in 35-mm MatTek dishes and treated the follow-
ing day. Imaging, cell tracking (every 26 min for 12 h), and analysis were
performed as described for the siRNA-treated cells.
72 h after transfection with siRNA, HFFs from the third 60-mm dish were
lysed in lysis buffer (2% IGEPAL, 40 mM NaCl, 10 mM MgCl2, and 50 mM
Tris, pH 7.5) plus protease inhibitors (Cytoskeleton). Cleared lysates were com-
bined with an equal volume of 2× sample buffer (Invitrogen), heated to 95°C
for 5 min, resolved by SDS-PAGE on a 4–12% Tris-glycine polyacrylamide
gel (Invitrogen), and transferred to nitrocellulose (0.2-µm pores; Invitrogen).
Antibodies used for Western blotting were mouse anti-Rac1 (Millipore), mouse
anti-Cdc42 (BD), mouse anti-RhoA (Abcam), mouse antiactin (Sigma-Aldrich),
and mouse anti–glyceraldehyde 3-phosphate dehydrogenase (Fitzgerald
Industries). Blots were developed using ECL reagents (GE Healthcare) and
visualized on a luminescent image analyzer (LAS-4000; Fujifilm). Western
blots were quantified by normalizing the GTPase intensity to the corres-
ponding glyceraldehyde 3-phosphate dehydrogenase signal and deter-
mining the change in expression relative to the siGLO-treated control.
Measurement of Young’s modulus by atomic force microscopy
Experiments were performed using an atomic force microscope (Catalyst;
Bruker AXS) mounted on the stage of an inverted microscope (Axiovert
200) placed on a vibration isolation table (IsoStation). Matrices were
kept at 37°C and buffered with PBS throughout the experiments. Force
displacement curves were obtained in contact mode using pyramidal tip
cantilevers of nominal stiffness, k = 0.03 N/m. For each cantilever used,
stiffness was obtained by the thermal fluctuations method (Butt and
Jaschke, 1995). Force displacement curves were acquired by ramping
the cantilever 12 µm at 1 Hz. To prevent damage to the samples, the
maximum force applied was 0.5 nN for soft matrices and 3 nN for
stiff matrices. Young’s modulus (E) was computed by fitting force dis-
placement curves with Sneddon’s formula for an indenting cone using
in which F is the applied force, is Poisson’s ratio and was assumed to be
0.5, is the indentation, and is the half-opening angle of the pyramidal tip
(Sneddon, 1965). To measure strain stiffening, E was computed fitting only
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