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Silencing profilin-1 inhibits endothelial cell proliferation, migration and cord
morphogenesis
Zhijie Ding, Anja Lambrechts, Mayur Parepally and Partha Roy
Journal of Cell Science 119, 4366 (2006) doi:10.1242/jcs.03268
There was an error published in J. Cell Sci. 119, 4127-4137.
In the e-press version of this paper that was published on 12 September 2006, the labels in Fig. 5B were incorrect. Both the
published print and online versions of this article are correct.
The correct figure is shown below.
The authors apologise for this mistake.
Journal of Cell Science
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Research Article
Introduction
Angiogenesis, a process of capillary outgrowth from
preexisting vessels, plays a pivotal role in various physiological
adaptations and pathological conditions including wound
healing, coronary heart disease, diabetic retinopathy and
growth of solid tumors (Carmeliet and Jain, 2000; Yancopoulos
et al., 2000). During angiogenesis, endothelial cells (ECs)
organize to form three-dimensional (3D) capillary networks,
which encompasses a range of cellular processes including
activation, protease production, migration, proliferation and
differentiation of ECs (Kanda et al., 2004; Whelan and Senger,
2003). The exact molecular details how ECs physically
assemble into capillary structures are not completely
understood. Over the years, studies using in vitro models where
ECs plated either on reconstituted basement membrane or
embedded in 3D collagen matrices form polygonal network of
pre-capillary cords (akin to the capillary structures in vivo)
have been utilized to investigate different aspects of EC
morphogenesis (Davis et al., 2002). Those studies indicate that
extracellular matrix (ECM)-integrin interactions and signaling
events involving cytoskeletal elements that control EC
migration and shape change, now collectively termed as MIC
(matrix-integrin-cytoskeleton) signaling axis, drive the
capillary morphogenesis of ECs (Davis et al., 2002; Davis et
al., 2000; Davis and Camarillo, 1995).
Marked changes in cell shape and migration that accompany
capillary morphogenesis of ECs imply active reorganization of
the actin cytoskeleton (Davis et al., 2002). Remodeling of actin
cytoskeleton is controlled by several different classes of actin-
binding proteins (ABPs) including those involved in monomer
sequestering, nucleating, filament-severing, depolymerizing,
and capping activities (Pollard and Borisy, 2003). However,
only a handful of studies have been reported to date that
directly examined the role of various ABPs in morphogenetic
events of ECs. Subtractive cDNA cloning of ECs plated on
matrigel first detected increased expression of thymosin 4 [a
G (globular)-actin sequestering protein] (Grant et al., 1995),
which was later shown to be a potent stimulator of
angiogenesis (Cha et al., 2003; Grant et al., 1999; Malinda et
al., 1997; Philp et al., 2004). Similarly, a previous study
reported up-regulation in the expression of profilin-1 (Pfn1, a
ubiquitously expressed ABP encoded by the Pfn family of
genes) during capillary morphogenesis (Salazar et al., 1999),
the biological significance of which, however, remains
unknown.
Four distinct Pfn genes have been identified so far: Pfn1,
Pfn2 [two splice variants of Pfn2 exist that are mainly found
in nerve cells (Di Nardo et al., 2000; Kwiatkowski and Bruns,
1988; Lambrechts et al., 2000)]; Pfn3 [kidney and testis
specific (Hu et al., 2001)]; and Pfn4 [testis specific (Obermann
et al., 2005)]. Although Pfn1 was originally identified as a G-
actin sequestering protein (Carlsson et al., 1977), depending
upon the conditions, it can either sequester G-actin and hence
inhibit actin polymerization, or promote actin assembly
Expression of several actin-binding proteins including
profilin-1 is up-regulated during capillary morphogenesis
of endothelial cells, the biological significance of which
remains unknown. Specifically, we hypothesized that
profilin-1 is important for endothelial migration and
proliferation. In this study, we suppressed profilin-1
expression in human umbilical vein endothelial cells by
RNA-interference. Gene silencing of profilin-1 led to
significant reduction in the formation of actin filaments and
focal adhesions. Loss of profilin-1 expression was also
associated with reduced dynamics of cell-cell adhesion.
Data from both wound-healing experiments and time-lapse
imaging of individual cells showed inhibition of cell
migration when profilin-1 expression was suppressed.
Cells lacking profilin-1 exhibited defects in membrane
protrusion, both in terms of its magnitude and directional
persistence. Furthermore, loss of profilin-1 expression
inhibited cell growth without compromising cell survival,
at least in the short-term, thus suggesting that profilin-1
also plays an important role in endothelial proliferation
as hypothesized. Finally, silencing profilin-1 expression
suppressed matrigel-induced early cord morphogenesis of
endothelial cells. Taken together, our data suggest that
profilin-1 may play important role in biological events
that involve endothelial proliferation, migration and
morphogenesis.
Key words: Profilin-1, Endothelial cells, Morphogenesis, Migration,
Proliferation, Adhesion
Summary
Silencing profilin-1 inhibits endothelial cell
proliferation, migration and cord morphogenesis
Zhijie Ding1, Anja Lambrechts2, Mayur Parepally3and Partha Roy1,4,*
1Department of Bioengineering, University of Pittsburgh, 749 Benedum Hall, 3700 O’Hara Street, Pittsburgh, PA 15261, USA
2Department of Biochemistry, Ghent University, Albert Baertsoenkaai 3, 9000 Ghent, Belgium
3Department of Biological Sciences, Carnegie Mellon University, 4400 Fifth Avenue, Pittsburgh, PA 15213, USA
4Department of Pathology, University of Pittsburgh, School of Medicine, S-417 BST, 200 Lothrup Street, Pittsburgh, PA 15261, USA
*Author for correspondence (e-mail: proy@engr.pitt.edu)
Accepted 20 July 2006
Journal of Cell Science 119, 4127-4137 Published by The Company of Biologists 2006
doi:10.1242/jcs.03178
Journal of Cell Science
4128
(Schluter et al., 1997). Since the intracellular concentration of
Pfn1 does not appear to be sufficient to account for the high
G-actin content in most cells, its role as a promoter of actin
polymerization is currently favored. Pfn1 promotes actin
assembly via its ability to accelerate nucleotide exchange
(ADP to ATP) on G-actin and shuttle Pfn-actin (ATP-bound)
complex to free barbed ends of actin filaments (Witke,
2004). Besides actin, Pfn1 binds to phosphoinositides
[mainly phosphatidylinositol-4,5-bisphosphate (PIP2) and
phosphatidylinositol-3,4,5-triphosphate (PIP3)] and a plethora
of proline-rich ligands ranging from those that directly
stimulate actin assembly in response to extracellular signals
[example: proteins belonging to Vasodilator Stimulated
Phosphoprotein or VASP (Reinhard et al., 1995), Wiskott-
Aldrich Syndrome Protein or WASP (Suetsugu et al., 1998)
and Diaphanous (Watanabe et al., 1997) families] to ones
involved in endocytosis, gene splicing and transcription
(Witke, 2004).
Gene deletion of both Pfn1 and Pfn2 causing impaired
motility and cytokinesis of Dictyostelium amebae produced the
first direct evidence of Pfn’s involvement in migration and
proliferation of lower eukaryotic cells (Haugwitz et al., 1994).
Defects in cell proliferation and migration were also evident in
chickadee (a Pfn1-homolog)-null mutants of Drososphila
where a late-stage embryonic lethality was observed (Verheyen
and Cooley, 1994). Due to a very early-stage embryonic
lethality resulting from Pfn1-gene knockout in mice (Witke et
al., 2001), similar studies have not been performed to date to
explore whether lack of Pfn1 has any overall impact on the
migration of mammalian cells. Lack of viability of Pfn1-null
embryos nevertheless suggests Pfn1’s role in proliferation
and/or survival for mammalian cells. Pfn1’s function has been
extensively studied in pathogen-based model systems where it
plays an important role in promoting the intracellular
movement of bacterial pathogens (Grenklo et al., 2003;
Laurent et al., 1999; Mimuro et al., 2000; Sanger et al., 1995;
Theriot et al., 1994). Since host-cell induced movement of
pathogens is a molecular mimicry of actin polymerization at
the leading edge of migrating cells, it is thought that one of
Pfn1’s function is to facilitate actin assembly during cell
protrusion. This notion is further supported by several other
studies demonstrating Pfn1’s involvement in the formation of
actin-based protrusive structures (Hajkova et al., 2000;
Suetsugu et al., 1998; Suetsugu et al., 1999) as well as its
preferential localization at the protrusive edge (Buss et al.,
1992; Chou et al., 2002; Neely and Macaluso, 1997; Neuhoff
et al., 2005). Given Pfn1’s importance in cell migration and
proliferation, it is thus intriguing that Pfn1 expression is
conspicuously down-regulated in aggressive mammary and
pancreatic carcinoma cells when compared with their normal
counterparts (Gronborg et al., 2006; Janke et al., 2000).
Suppression of growth (Janke et al., 2000) and migration (Roy
and Jacobson, 2004) of mammary carcinoma cells induced by
overexpression of Pfn1 further suggest that Pfn1’s role in
mammalian cell migration and proliferation may be complex
and cell-type dependent.
Based on significant up-regulation of Pfn1 expression in
human umbilical vein endothelial cells (HUVECs) during
capillary morphogenesis (Salazar et al., 1999), one envisions
that Pfn1 might play an instrumental role during endothelial
morphogenesis. Related to this, we specifically hypothesized
Journal of Cell Science 119 (19)
that Pfn1’s function is important for EC migration and
proliferation. In the present study, we tested this hypothesis by
examining the effects of suppression of Pfn1 expression on EC
proliferation and migration. We further assessed whether loss
of Pfn1 expression affects ECM-induced early morphogenesis
of ECs.
Results
Silencing Pfn1 alters the actin cytoskeleton, cell-matrix
and cell-cell adhesions in HUVECs
To suppress Pfn1 expression, we adopted transient transfection
of HUVECs with either a non-targeting control (C) or Pfn1-
specific (P) siRNA construct. Initially, to determine the
specificity of Pfn1-siRNA, we transiently transfected the
siRNA constructs into MDA-MB-231 breast carcinoma cells,
which were genetically engineered by us to provide stable
expression of different point-mutants of GFP-Pfn1. These
mutants involved a 2 base-pair alteration either within (mutant-
1) or outside (mutant-2) the region targeted by Pfn1-siRNA.
As judged by the decrease in GFP-fluorescence, our Pfn1-
siRNA down-regulated the expression of mutant-2 as expected,
but was ineffective in suppressing the expression of mutant-1.
In negative control experiments, Pfn1-siRNA treatment did not
non-specifically decrease the fluorescence of GFP-expressing
MDA-MB-231 cells (Fig. 1A). Taken together, these data thus
demonstrate the specificity of action of our Pfn1-siRNA. The
bar graph in Fig. 1B displays the silencing efficiency of Pfn1-
siRNA in HUVECs as a function of time (48-96 hours), which
shows a time-dependent progressive loss of Pfn1 expression
with ~97% gene-silencing achieved 96 hours after transfection.
The representative Pfn1 immunoblots at different time-points
after transfection are shown in the inset of Fig. 1B (the GAPDH
blot serves as the loading control). We have also confirmed that
the polyclonal Pfn1 antibody used in this study is specific and
does not cross-react with Pfn2 (data not shown).We could not
detect Pfn2 expression by immunoblotting under either of the
experimental conditions (inset of Fig. 1B) thus suggesting that
there is no compensatory up-regulation of Pfn2 when Pfn1
expression is suppressed in HUVECs.
The effect of silencing Pfn1 on endothelial actin
cytoskeleton was next evaluated by rhodamine-phalloidin
staining of HUVECs 96 hours after transfection, which
showed that cells bearing Pfn1-siRNA have significantly
less actin filaments, particularly those comprising stress-
fibers, compared with the control-siRNA transfected cells
(Fig. 1C). Quantification of phalloidin-fluorescence showed
approximately 29% reduction in the overall F-actin content of
HUVECs due to loss of Pfn1 expression (Fig. 1D). Since Pfn1
has been recently implicated in gene transcription (Lederer et
al., 2005), we next asked whether expression levels of actin and
some of the ABPs that are important for actin assembly in
response to growth factor signaling such as VASP, N-WASP
and mDia1 are altered as a result of silencing Pfn1.
Immunoblots of whole cell lysates showed no appreciable
change in the expression levels of actin and the indicated ABPs
at any time-point after transfection (Fig. 1E). Taken together,
these data suggest that loss of Pfn1 expression alters actin
cytoskeleton via direct modulation of actin polymerization
and/or bundling of actin filaments.
Since both cell-matrix and cell-cell adhesion complexes
physically associate with actin cytoskeleton, we next asked
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Profilin in endothelial morphogenesis
whether loss of Pfn1 expression has any impact on either kind
of adhesive structures in ECs. Previous data showed that Pfn1-
overexpression causes human aortic ECs to form more focal
contacts and display increased adhesion on fibronectin-
substrate (Moldovan et al., 1997) thus suggesting a possible
role of Pfn1 in regulating cell-matrix adhesion. To further this
line of inquiry, we performed vinculin [a marker for focal
adhesion (FA)] immunostaining of HUVECs, which showed a
marked inhibition of FA assembly when Pfn1 expression was
silenced (Fig. 2A). A bar graph in Fig. 2B summarizes the
results from quantitative analyses of vinculin-staining data,
which showed significantly lower FA density (defined as the
number of FA plaques per 100 m2of cell area) in Pfn1-
deficient cells (2.7±1.9) when compared with control cells
(6.1±2.1). The FAs observed for the control cells also appeared
to be larger in size compared with those in Pfn1-deficient cells.
Overall, these data demonstrate that Pfn1 plays an important
role in regulating cell-matrix adhesions in ECs.
Whether Pfn1 is involved in regulating cell-cell adhesion has
not been explored at all. Interestingly, yeast-two hybrid
screening technique has previously identified Pfn1 as a binding
partner for AF-6, a multidomain protein that is localized at cell-
cell junctions (Boettner et al., 2000). However, the functional
significance of such interaction remains unknown. To determine
whether loss of Pfn1 expression affects endothelial cell-cell
junctions, confluent monolayers of serum-starved HUVECs,
carrying either control or Pfn1-siRNA, were subjected to VEGF
(a potent disruptor of endothelial cell-cell junctions) stimulation
and immunostaining of VE-cadherin (a marker for adherence
junction) and ZO-1 (a marker for tight junction) were
performed. As expected, VEGF stimulation caused significant
loss of junctional staining of both VE-cadherin and ZO-1
(arrows) with concomitant creation of paracellular holes
(arrowheads) in the monolayer of control cells (Fig. 2C,D).
Interestingly, VEGF-induced delocalization of both VE-
cadherin and ZO-1 from the cell-cell junctions was significantly
inhibited in Pfn1-deficient cells (Fig. 2C,D). Although the base-
line (i.e. under serum-starved state) staining pattern of VE-
cadherin was similar for both control and Pfn1-deficient cells,
subtle differences in ZO-1 staining were apparent between the
two experimental conditions. For example, control cells had
somewhat more fragmented junctional and higher nuclear
Fig. 1. Silencing Pfn1 expression affects the actin cytoskeleton in HUVECs: (A) Target specificity of Pfn1-siRNA was initially demonstrated by
transfecting either control (C) or Pfn1-specific (P) siRNAs in MDA-MB-231 breast cancer cells that stably express either GFP or different
point-mutants of GFP-Pfn1 (designated as mutant-1 and mutant-2 that carried a 2-base-pair alteration both within and outside of the targeting
region of Pfn1-siRNA, respectively). Fluorescence micrographs of cells show that Pfn1-siRNA suppresses the expression of mutant-2, but not
of mutant-1. Fluorescence of control GFP-expressing cells is not affected by Pfn1-siRNA treatment. Bar, 150 m. (B) A bar graph shows time-
dependent progressive loss of Pfn1 expression with nearly 97% suppression of Pfn1 expression 96 hours after Pfn1-siRNA transfection (data
summarized based on immunoblot analyses of HUVEC extracts from three independent experiments). The inset shows the actual representative
Pfn1-immunoblots of HUVEC extracts prepared at different time-points after siRNA transfection with GAPDH blot serving as the loading
control. Additional immunoblot data shows no detectable Pfn2 expression under either experimental condition (purified Pfn2 serves the positive
control for the immunoblot). (C) Rhodamine-phalloidin staining of HUVECs shows that silencing Pfn1 dramatically inhibits the formation of
actin stress-fibers. Bar, 20 m. (D) A bar graph displaying the relative (normalized with respect to the control cells) fluorescence intensity of
phalloidin shows a 29% decrease in the average level of F-actin in Pfn1-deficient cells. These data were obtained from analyses of 640 control
and 584 Pfn1-deficient cells from two independent experiments, the difference of which was found to be statistically significant (the asterisk
indicates P<0.0001). (E) Immunoblots show comparable expression levels of actin and several ABPs such as VASP, mDia1, and N-WASP at
48, 72 and 96 hours after transfection.
Journal of Cell Science
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staining of ZO-1. Immunoblot data showed no appreciable
change in the expression level of either VE-cadherin or ZO-1
between the two experimental conditions (Fig. 2E). These
results suggest that loss of Pfn1 expression suppresses VEGF-
induced dynamics of cell-cell adhesions in ECs.
Pfn1 regulates HUVEC proliferation
To test our hypothesis that Pfn1 plays a role in EC
proliferation, we next compared cell growth at different time-
points (48-96 hours) after siRNA transfection, which showed
that Pfn1-siRNA treatment led to nearly 42% inhibition in
HUVEC growth when compared with the control transfection
condition (Fig. 3A). Inhibition in cell growth was evident
within 48 hours after transfection. From nuclear staining of
cells with DAPI, we did not detect any ECs with multinuclei
(>2) phenotype thus suggesting no gross defect in cytokinesis
as a result of lack of Pfn1 expression. We also wanted to see
whether a difference in the number of apoptotic cells (display
nuclear fragmentation in DAPI staining) between the two
culture conditions might explain the differential cell-growth.
The nuclear morphology of both control and Pfn1-deficient
cells appeared completely normal (Fig. 3B), and the number
of apoptotic cells in the culture was absolutely negligible
(<0.5%) in either case. These data suggest that loss of Pfn1
expression does not compromise HUVEC survival, at least in
the short term, and hence, reduced cell growth in Pfn1-siRNA
treated culture was most likely due to diminished proliferative
capacity of these cells.
Loss of Pfn1 inhibits HUVEC spreading
Since Pfn1 has been shown to play an important role in forming
actin-based protrusive structures, we next evaluated whether
early cell spreading (an event that involves active cell
protrusion), on ECM-coated substrate is affected by loss of
Journal of Cell Science 119 (19)
Fig. 2. Loss of Pfn1 expression alters cell-matrix and cell-cell adhesions in HUVECs: (A) Vinculin-immunostaining shows a dramatic
reduction in FA formation when Pfn1 expression is silenced (C, control siRNA; P, Pfn1-siRNA). Bar, 20 m. (B) A bar graph shows a
significantly (P<0.001) higher FA density (number of FA/100 m2of cell area) in control cells (6.1±2.1) than in Pfn1-deficient cells (2.7±1.9).
These data are based on analyses of 56 control and 69 Pfn1-deficient cells that were randomly selected from two independent experiments.
(C,D) VE-cadherin and ZO-1 immunostaining show that VEGF stimulation causes loss of junctional staining of these proteins (arrow) and
creation of paracellular holes (arrowhead) in control cells. VEGF-induced delocalization of VE-cadherin and ZO-1 from cell-cell junctions is
significantly inhibited in Pfn1-deficient ECs. Bar, 20 m. (E) Immunoblot data show no appreciable change in the expression of VE-cadherin
and ZO-1 between control and Pfn1-siRNA treated cells (the GAPDH blot serves as the loading controls).
Fig. 3. Silencing Pfn1 expression inhibits cell
growth: (A) A line graph shows the ratio of the
number of control (denoted by ‘C’) to that of
Pfn1-deficient cells (denoted by ‘P’) in culture at
48, 72 and 96 hours after transfection. (These
data were summarized from three independent
experiments and P<0.0002.) (B) DAPI staining
of cells shows normal nuclear morphology under
either experimental condition. Bar, 20 m.
Journal of Cell Science
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Profilin in endothelial morphogenesis
Pfn1 expression. Fig. 4A shows the morphology of HUVECs
on matrigel (100 g/ml)-coated substrate within 1 hour after
plating. From phase contrast images where spreading cells
appear darker (phase-dense), it is evident that Pfn1-deficient
cells are much less efficient in spreading when compared with
the control cells. Fig. 4B summarizes the data in the form of a
bar graph plotting the percentage of spreading cells on
substrates coated with different concentrations of matrigel.
Although a higher concentration of matrigel generally
facilitated cell spreading, a 3-fold decrease in the spreading
efficiency was observed when Pfn1 expression was silenced,
thus further confirming that Pfn1 plays a key role in regulating
EC protrusion. To further determine whether loss of Pfn1
actually inhibits or only delays cell spreading, we compared
HUVEC-morphology on matrigel–coated substrates at later
time-points. Even at 22 hours after cell-seeding, the extent of
spreading of Pn1-deficient cells was clearly much less
compared with that of control cells (Fig. 4C), therefore
meaning that silencing Pfn1 actually inhibits cell spreading.
Pfn1 is important for HUVEC migration
To test our hypothesis that Pfn1’s function is important for EC
migration, we next examined whether directed migration of
HUVECs is affected by loss of Pfn1 expression using a
standard wound-healing assay. Fig. 5A depicts the results of a
typical wound-healing experiment performed 96 hours after
siRNA transfection, where Pfn1-deficient cells clearly showed
significant impairment in wound closure when compared with
the control cells. At the wound margins, control ECs were seen
as isolated cells, whereas Pfn1-deficient cells appeared to
Fig. 4. Loss of Pfn1 expression affects HUVEC spreading: (A) Phase contrast images of HUVECs seeded 1 hour after plating on a substrate
that is pre-coated with 100 g/ml matrigel show higher proportion of spreading cells (appear phase dense) in the control group (denoted by
‘C’). Impaired cell spreading was evident from round morphology of majority of Pfn1-deficient cells (denoted by ‘P’). Bar, 100 m. (B) A bar
graph plotting the percentage of spreading cells at two different time-points (30 minutes and 1 hour) and for two different coating
concentrations of matrigel clearly shows increased spreading efficiency of the control cells (these data were pooled from the analyses of
approximately 800-1000 cells for each experimental condition from two independent experiments). The asterisk indicates P<0.001. (C) Pfn1-
deficient cells were still found to be much less flat and spread-out compared with the control cells at 22 hours after cell-seeding on matrigel-
coated substrates. Bar, 50 m.
Fig. 5. Loss of Pfn1 inhibits HUVEC migration in wound healing assay. (A) Representative images of the wound margins immediately and 12
hours after wounding show significant impairment in wound closure by HUVECs due to loss of Pfn1 expression (C, control siRNA; P, Pfn1-
siRNA). Bar, 100 m. (B) A higher magnification of the wound margin indicates closer association of Pfn1-deficient cells. Bar, 100 m. (C) A
bar graph plotting the relative efficiency of wound closure shows Pfn1-siRNA treatment inhibited wound-closure by 25%, 36% and 47% when
evaluated at 48, 72 and 96 hours after transfection, respectively. (These data are summarized from three independent experiments and the
asterisk indicates P<0.002.)
Journal of Cell Science
4132 Journal of Cell Science 119 (19)
Fig. 6. Effect of silencing Pfn1 expression on single cell migration: (A) A typical time-lapse imaging experiment shows directed migration of
control cells (denoted by ‘C’) involving directed protrusion and significant net cell translocation. By contrast, Pfn1-deficient cells (denoted by
‘P’) produce small, randomly directed protrusion with much less net cell translocation (the direction of protrusion is indicated by the arrow).
Bar, 30 m. (B) Trajectories of individual cells from the frame-by-frame analyses of the centroid of cell nuclei show a significantly (P<0.001)
larger net distance traveled by the control cells (43.6±26.0 m) compared with the Pfn1-deficient cells (11.3±9.6 m) during the 90-minute
observation period (migration data of 27 control and 21 Pfn1-deficient cells from a total of three independent experiments were pooled for the
analysis). (C) Representative plots of change in protrusion direction between successive image frames (⌬) from a typical experiment are
shown side-by-side to display much larger oscillation of ⌬ in Pfn1-deficient cells (these plots were generated based on motility data of five
control and six Pfn1-deficient cells in one experiment). (D) The bar graph compares the standard deviation values of ⌬ for the entire 90-
minute observation period (these data are based on the average values calculated for 14 control and 19 Pfn1-deficient cells – those cells that
either barely protruded or came in contact with a neighboring cell at any point during the course of the experiment were excluded from the
analysis). The asterisk indicates P<0.0001.
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Profilin in endothelial morphogenesis
maintain connections with the neighboring cells (Fig. 5B). This
distinctive feature also seems to be consistent with reduced
dynamics of cell-cell adhesions in Pfn1-deficient cells as
previously demonstrated in Fig. 2C,D. A bar graph in Fig. 5C
summarizes the wound-healing data as a function of time
where Pfn1-siRNA treatment inhibited wound-closure by 25%,
36%, and 47% at 48, 72 and 96 hours after transfection,
respectively. These data thus demonstrate increasing inhibition
of HUVEC migration with progressive loss of Pfn1 expression.
Cell migration in a monolayer set-up, as in a wound-healing
assay, can be affected by the strength of cell-cell adhesion.
Since our data suggested Pfn1-dependent alteration in cell-cell
adhesion, we additionally performed time-lapse imaging
of individual HUVECs to determine whether intrinsic EC
migration is affected by loss of Pfn1 expression. Fig. 6A shows
the results of a typical set of time-lapse experiments where
HUVECs bearing control-siRNA displayed directed migration
involving significant translocation of their cell bodies during
the course of the experiment (upper panel). Consistent with our
findings from wound-healing experiments, a marked inhibition
in random cell migration was observed in the case of Pfn1-
deficient cells (lower panel). The single-cell migration data are
summarized in Fig. 6B, which depicts the trajectories of
individual cells obtained by frame-by-frame analyses of
the centroid positions of cell-nuclei (assumed to be the
representations of cell-bodies in this case). The average net
distances translocated by the control and Pfn1-deficient cells
in the 90-minute observation period were 43.6±26.0 m (n=27
cells) and 11.3±9.6 m (n=21 cells), respectively, the
difference of which was found to be statistically significant
(P<0.001). Overall, the results from wound-healing and time-
lapse imaging experiments clearly demonstrate that Pfn1 plays
an important role in EC migration as hypothesized.
Our time-lapse imaging data also showed a difference in the
protrusive activity between the control and Pfn1-siRNA treated
cells. While control cells mostly maintained persistence in the
direction of protrusion, cells bearing Pfn1-siRNA typically
exhibited smaller and random protrusions that lacked any
directional bias (the direction of protrusion at any instance is
marked by an arrow; see Fig. 6A). To represent the differences
in the protrusive behavior between the two experimental
conditions in a quantitative fashion, we then analyzed and
plotted the change in the direction of protrusion between
successive image frames (⌬i=i-i-1, where idenotes the
direction of protrusion at the i-th image frame) as a function of
time during the entire 90-minute course of experiment for each
cell. Representative plots based on data analyses of control and
Pfn1-deficient cells from a typical experiment are shown side-
by-side in Fig. 6C, where clearly a much larger oscillation of
⌬, thus meaning a decreased directional persistence of
protrusion, was seen when Pfn1 expression was suppressed. To
summarize the persistence data, the standard deviation of ⌬
for the 90-minute time-lapse period (a higher value of such
index would correlate with decreased directional persistence of
protrusion) was calculated for individual cells, pooled and then
averaged based on the total number of cells analyzed from three
independent experiments. A nearly 2.3-fold higher value of this
index determined for Pfn1-deficient cells (Fig. 6D) further
confirms a decreased ability of these cells to sustain directed
protrusion when compared with the control cells.
It has been previously shown that membrane targeting of
VASP to the leading edge decreases the persistence of
protrusion in migrating cells (Bear et al., 2002). To further
study whether VASP localization in migrating HUVECs is
altered when Pfn1 expression is suppressed, we performed
VASP-immunostaining of HUVECs in a wound-healing set-up,
the results of which are shown in Fig. 7. We were specifically
interested in VASP localization for cells situated at the wound-
edge, since those cells are capable of executing directed
migration with least obstructed membrane protrusion. In
control HUVECs, VASP predominantly localized at the FAs
and actin stress-fibers. At the leading edge, a much stronger
VASP staining was observed in Pfn1-deficient cells when
compared with the control cells, thus suggesting that loss of
Pfn1 expression in HUVECs is associated with increased
membrane targeting of VASP.
Silencing Pfn1 inhibits ECM-induced cord formation by
HUVECs
To finally determine whether Pfn1 plays any role in the early
Fig. 7. Silencing Pfn1 alters the localization of VASP in migrating HUVECs. In control cells (denoted by ‘C’), VASP is predominantly
localized at the focal adhesions and actin stress-fibers, and has a punctate distribution at the leading edge. Pfn1-deficient cells (denoted by ‘P’)
have much stronger localization of VASP at the leading edge. The inset in each panel shows a magnified view of the boxed region depicting
differences in VASP localization at the leading edge between the two experimental conditions. Bar, 10 m.
Journal of Cell Science
4134
morphogenetic events of ECs, we examined the effect of
silencing Pfn1 on matrigel-induced cord formation of ECs.
HUVECs bearing the control siRNA started spreading as early
as 1 hour after seeding on matrigel (data not shown) and formed
prominent cord-like structures by 8 hours (Fig. 8A). However,
cord formation was significantly inhibited when Pfn1
expression was silenced as evident from the round morphology
of majority of Pfn1-deficient cells at the indicated time-point.
Quantitative analyses showed that early cord morphogenesis
was inhibited by nearly 92% due to loss of Pfn1 expression (Fig.
8B, data summarized from four independent experiments).
Even at later time-points (18-22 hours after cell-seeding), we
also found the number of cords formed by Pfn1-deficient cells
to be still significantly less than that formed in the control
culture (data not shown). These data suggest that Pfn1 is a key
player in ECM-induced cord morphogenesis of ECs.
Discussion
In the present work, we tested a hypothesis that Pfn1’s function
is important for EC proliferation and migration. Specifically,
we examined the effects of silencing Pfn1 expression on actin
cytoskeleton, cell-cell and cell-matrix adhesion, proliferation,
migration and consequently, on ECM-induced early cord
morphogenesis in vitro.
Consistent with previous findings reported for smooth
muscle (Tang and Tan, 2003) and alveolar epithelial (Bitko et
al., 2003) cells, we observed a significant reduction of F-actin
level in HUVECs when Pfn1 expression was silenced thus
implying that Pfn1 promotes actin polymerization in ECs. By
contrast, a previous study had shown that gene deletion of Pfn1
and Pfn2 resulted in increased actin polymerization in
Dictyostelium amoeba thus meaning Pfn’s function as G-
actin sequestering proteins in this organism (Haugwitz et al.,
1994). Cell-specific difference in Pfn’s net action on actin
cytoskeleton is not surprising since whether Pfn1 would
function as a promoter of actin polymerization or a G-actin
sequester depends on its concentration relative to that of
available G-actin and free barbed ends of actin filaments. These
parameters are controlled by other ABPs (sequestering,
severing and capping), expression of which can vary between
different cell types. Also, in mammalian cells, the intracellular
concentration of Pfn1 does not appear to be sufficient for G-
actin sequestration, which is primarily regulated by proteins
belonging to the thymosin-family. From reduced actin stress-
fibers in Pfn1-deficient cells, it is not immediately clear
whether lack of Pfn1 inhibits only actin polymerization or
affects the bundling of actin stress-fibers as well. It is known
that Diaphanous-family proteins, such as mDia1, utilize Pfn1
to polymerize actin to form stress-fibers (Romero et al., 2004),
which might partly explain reduced actin stress-fibers found in
Pfn1-deficient cells. Bundling of actin stress-fibers, on the
other hand, is facilitated by cell-contractility that requires
actomyosin interactions. Pfn1-depletion may down-regulate
EC-contractility by partially suppressing actin polymerization
[also shown previously for smooth muscle cells (Tang and Tan,
2003)] and therefore affect the bundling of actin stress-fibers.
Consistent with the loss of actin stress-fibers, silencing Pfn1
expression also suppressed FA assembly in HUVECs. This
data seems to be in qualitative agreement with previous
findings by us and others where Pfn1-overexpression caused
increased substrate-adhesion of breast cancer cells (Roy and
Jacobson, 2004) and human aortic ECs (Moldovan et al.,
1997). FA formation is initiated through integrin clustering, a
process that is driven by both ECM-binding and contractility
(Schoenwaelder and Burridge, 1999). Further assembly of FAs
involves signaling through FAK (Focal adhesion kinase) and
Src that recruits other molecular components to this adhesive
structure. Integrin clustering can be diminished in Pfn1-
deficient cells because of possible down-regulation of cell
contractility. Surface recruitment of integrins can also be
affected by Pfn1-dependent changes in cytoskeletal
organization as postulated earlier (Moldovan et al., 1997).
Furthermore, we earlier showed that Pfn1-overexpression up-
regulates tyrosine phosphorylation of FAK and paxillin in
breast cancer cells thus suggesting dependence of FAK-
signaling on Pfn1’s function (Roy and Jacobson, 2004). Thus,
recruitment of molecular components to FA may also be
influenced by altered FAK signaling in Pfn1-deficient ECs.
An interesting finding of the present study is that in addition
to affecting cell-matrix adhesion, loss of Pfn1 expression also
influences cell-cell adhesion where, in particular, VEGF-
induced dynamics of intercellular junctions is suppressed.
Previous studies showed that agonist-induced disruption of
intercellular junctions requires Rho-based EC contraction
Journal of Cell Science 119 (19)
Fig. 8. Effect of silencing Pfn1 on early cord morphogenesis of HUVECs. (A) Control HUVECs (denoted by ‘C’) form prominent cord-like
structures on polymerized matrigel by 8 hours after plating. Cord formation is significantly inhibited in the case of Pfn1-deficient cells (denoted
by ‘P’). Bar, 100 m. (B) A bar graph shows significantly higher number of nodes involving at least three branches in the control cells compared
with the same in Pfn1-deficient cells (the graph summarizes data from a total of four independent experiments; the asterisk indicates P<0.001).
Journal of Cell Science
4135
Profilin in endothelial morphogenesis
(Alexander and Elrod, 2002; Bates et al., 2002). If depletion of
Pfn1 reduces EC contractility as observed previously for
smooth muscle cells (Tang and Tan, 2003), ECs will become
resistant to VEGF-induced disruption of cell-cell adhesion. It
has been shown that activation of receptor tyrosine kinases, such
as VEGF receptor, is stimulated by integrin clustering and cell-
contraction (Gingras et al., 2000; Sundberg and Rubin, 1996).
Thus, reduced contractility of Pfn1-deficient cells may also
modulate the spatial distribution of VEGF-receptors and hence,
suppress VEGF signaling by decreasing the receptor activation.
Interestingly, we observed increased nuclear staining of ZO-1
in control HUVECs. It was suggested that nuclear localization
of ZO-1 inversely correlates with the maturation of tight
junction (Gottardi et al., 1996). This may also explain why the
baseline ZO-1 staining in control HUVECs appears somewhat
more fragmented than the same in Pfn1-deficient cells.
We found significant inhibition of EC migration as a result
of loss of Pfn1 expression thus supporting our hypothesis that
Pfn1 is an important player of EC migration. The only previous
study that directly evaluated how lack of Pfn1 affects the
overall cell migration was performed in dictyostelium and thus,
the present work is the first demonstration of the effect of loss
of Pfn1 expression on the overall migration of any mammalian
cell. In dictyostelium, deletion of both Pfn1 and Pfn2 genes
resulted in impaired cell motility, and knocking out Pfn1 gene
alone failed to produce a phenotype because of functional
compensation by Pfn2 gene product (Haugwitz et al., 1994).
Since no Pfn2 expression was detected in HUVECs, we were
able to see progressive inhibition of EC migration with loss of
Pfn1 expression. Data from our spreading and time-lapse
experiments confirm previous findings regarding Pfn1’s
involvement in cell-protrusion (Hajkova et al., 2000; Suetsugu
et al., 1998; Suetsugu et al., 1999). Interestingly, a listeria-
motility study showed that although Pfn1 increases the
efficiency of listeria-induced actin polymerization and hence,
the velocity of pathogen movement, it is not one of the essential
cellular components needed for initiating motility of pathogens
(Loisel et al., 1999). Similarly, our experiments showed that
Pfn1-deficient cells are still able to protrude and migrate, but
clearly not to the same extent as displayed by the control cells.
Detailed analyses of protrusion revealed that ECs lacking Pfn1
are significantly less efficient in maintaining directional
persistence of protrusion compared with the control cells.
Since stabilization of membrane protrusion requires cell
adhesion, significantly reduced formation of adhesion
complexes might partly explain the lack of sustenance of
protrusion in Pfn1-deficient cells. Previously, membrane
targeting of VASP was inversely correlated with the persistence
of cell protrusion (Bear et al., 2002). Interestingly, we also
observed a much stronger localization of VASP at the leading
edge in Pfn1-deficient cells. Because of reduced actin stress-
fibers and FAs in Pfn1-deficient cells, one possibility is that
VASP is less sequestered to those cellular structures; therefore,
there might be an increased availability of VASP for targeting
to cell membrane. Despite consistency in observation, without
further experiments it is presently not clear whether there is an
actual causal relationship between increased membrane
targeting of VASP and decreased persistence of protrusion in
our case. Our time-lapse data showed that EC translocation is
also inhibited when Pfn1 expression is suppressed. Since
actomyosin-based contraction drives cell translocation during
motility, our data further suggests a possible down-regulation
of contractility in Pfn1-deficient cells.
Besides affecting cell migration, loss of Pfn1 expression also
inhibited EC growth by 42% without compromising cell
survival, at least in the short-term, thus supporting our
hypothesis that Pfn1 plays an important role in EC
proliferation. Previous studies showed that Pfn1 localizes at the
cleavage furrow during cytokinesis and its function is
important for cleavage furrow regression (Dean et al., 2005;
Suetsugu et al., 1999). Since Pfn1-deficient cells did not
display multinuclei (>2) phenotype, our data suggests that
Pfn1’s function is not essential for cytokinesis of mammalian
ECs as it is for some other species including yeast and amoeba
(Haugwitz et al., 1994; Lu and Pollard, 2001). The fact that we
observed only a partial inhibition of EC proliferation after near
complete silencing of Pfn1 gene is intriguing since it has been
shown that gene deletion of Pfn1 arrests developing mouse
embryo at the two-cell stage and produces embryonic lethality
(Witke et al., 2001). Although it is tempting to speculate that
embryonic stem cells might be more sensitive to loss of Pfn1
expression, a more detailed work is necessary to address
whether persistent suppression of Pfn1 expression completely
arrests cell-growth and/or affects the long-term survival of
ECs. Future work is also needed to determine the effect of loss
of Pfn1 expression on different phases of cell cycle.
Finally, we demonstrated that silencing Pfn1 expression
significantly inhibits ECM-induced early cord morphogenesis
of ECs. Network formation by ECs on ECM proceeds through
several stages including (1) cell adhesion on ECM, (2) cell
migration, (3) cell-induced mechanical remodeling of ECM that
further defines matrix guidance tracks to allow directed EC
migration to the neighboring cells, and (4) cell proliferation
(Cascone et al., 2003; Davis et al., 2002; Davis and Camarillo,
1995; Liu and Senger, 2004; Whelan and Senger, 2003). Early
impairment in cord-forming ability of Pfn1-deficient ECs is
most likely a result of defect in cell migration and not due to
delayed cell proliferation. We observed less cord formation by
Pfn1-deficient cells also at later time-points (18-22 hours after
cell seeding – data not shown) in which case, however,
additional contribution of inhibited cell proliferation cannot be
ruled out. Directed EC migration on ECM is not only influenced
by the intrinsic migratory ability of cells, but also governed by
the ECM-remodeling capacity of cells. Reduced formation of
actin stress-fibers and FAs might render Pfn1-deficient cells less
efficient in ECM remodeling because of possible down-
regulation of contractility. One also needs to consider a possible
influence of Pfn1-dependent modulation of cell-cell adhesion
on cord formation. Although cadherin-mediated junctions are
important for endothelial assembly and capillary formation,
early events of capillary morphogenesis of EC involve
disruptions of cell-cell contacts that are presumably critical for
both EC migration and alignment to form pre-capillary cords
(Carmeliet, 2000; Liu and Senger, 2004). Our data showed that
both control and Pfn1-deficient cells are capable of forming
cell-cell adhesions; however, the latter display reduced
dynamics of cell-cell adhesions when challenged with VEGF.
Thus, altered dynamics of cell-cell adhesion due to loss of Pfn1
expression can have an inhibitory effect on EC migration.
Finally, our wound-healing data showed that cell migration is
only partially inhibited by silencing Pfn1 expression. Thus, it is
not surprising that cells lacking Pfn1 are still able to form cords
Journal of Cell Science
4136
but to a much lesser extent compared with the control cells. In
conclusion, since cord morphogenesis is an early endothelial
rearrangement necessary for capillary formation by ECs, results
of the present work justify further studies to explore whether
Pfn1 plays a role in capillary morphogenesis of ECs.
Materials and Methods
Antibodies and reagents
Polyclonal antibodies specific for Pfn1 and Pfn2 were generous gifts of Drs Sally
Zigmond (University of Pennsylvania) and Walter Witke (European Molecular
Biology Laboratory, Italy), respectively. Polyclonal antibody for N-WASP was
kindly provided by Dr Hideki Yamaguchi (Albert Einstein College of Medicine).
Monoclonal antibodies for VASP and ZO-1 (zonula occludens-1) were obtained
from Pharmingen (San Diego, CA). Monoclonal antibodies for GAPDH and actin
are products of Chemicon (Temecula, CA). Monoclonal antibody for vinculin is a
product of Sigma (St Louis, MO). Polyclonal antibody for mDia1 was obtained from
Abcam (Cambridge, MA). Monoclonal antibody for VE-cadherin (vascular
endothelial cadherin) was obtained from Santa Cruz Biotechnology (Santa Cruz,
CA). For immunoblotting, the antibodies were used at the following concentrations:
Pfn1 (1:500), Pfn2 (1:1000), VASP (1:500), GAPDH (1:200), actin (1:1000),
ZO-1 (1:500), VE-cadherin (1:1000), N-WASP (1:1000), and mDia1 (1:2500).
Rhodamine-phalloidin and DAPI were purchased from Molecular Probes (Carlsbad,
CA). Collagen type I and growth-factor-reduced matrigel are products of BD
Biosciences (Bedford, MA).
Cell culture and transfection
HUVECs (source: Cambrex Biosciences, Walkersville, MD) were cultured in the
complete EBM2 growth media (also commercially available from the same source).
To silence Pfn1 expression, a custom-designed siRNA (sense strand: 5⬘-AGA AGG
UGU CCA CGG UGG UUU-3⬘; antisense-strand: 5⬘-ACC ACC GUG GAC ACC
UUC UUU-3⬘) was synthesized by Dharmacon (Lafayette, CO) and was transfected
into HUVECs using a proprietary reagent according to the manufacturer’s protocol.
A control siRNA (sense strand 5⬘-UAG CGA CUA AAC ACA UCA AUU-3⬘;
antisense strand: 5⬘-UUG AUG UGU UUA GUC GCU AUU-3⬘) that bears no
significant homology with any known mouse or human gene and commercially
available from the same source was used for the control experiments. Briefly, ECs
were transfected with 100 nM of siRNA for 24 hours. The transfection media was
then replaced with the regular growth media and cells were cultured for another 24-
72 hours before performing the experiments.
Protein extraction and immunoblotting
For protein extraction, cells were washed twice with cold PBS and lysed on ice for
30 minutes in modified RIPA buffer containing 50 mM Tris-HCl (pH 7.5), 150 mM
NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 2 mM EDTA, 50 mM NaF,
1 mM sodium pervanadate, and protease inhibitors (10 g/ml of leupeptin, aprotinin,
pepstatin and 1 mM phenylmethylsulfonyl fluoride). The lysates were clarified at
~17,000 gfor 15 minutes at 4°C and the protein concentration was determined using
a Coomassie-based protein assay kit (Pierce; Rockford, IL). Proteins separated on
SDS-PAGE were transferred onto a nitrocellulose membrane. After blocking the
membrane with 5% non-fat dry milk in TBST for 1 hour at room temperature,
immunoblotting was performed by overnight incubation with the appropriate primary
antibodies. After extensive washing with TBST, the blot was incubated with the
appropriate secondary antibody (Pharmingen, San Diego, CA) and washed three times
with TBST before performing chemiluminescence for visualization of protein bands.
Immunostaining
For VASP and vinculin immunostaining, cells cultured on collagen-coated cover-
slips were washed 3 times with PBS, fixed with 3.7% formaldehyde for 15 minutes,
permeablized with 0.5% Triton X-100 for 5 minutes and then blocked with 10%
goat-serum for 30 minutes. After incubating with the primary antibody at a 1:100
dilution for 1 hour at room temperature, cells were washed four times (first twice
with PBS containing 0.02% tween and then twice with PBS), each of 3-minute
duration, before incubating with a FITC-conjugated anti-mouse secondary antibody
(1:100 dilution). For VE-cadherin and ZO-1 staining, cells were fixed and
permeabilized using cold methanol at –20°C for 20 minutes and then blocked with
5% BSA (containing 15% glycine) for 45 minutes at room temperature. After
incubating with either VE-cadherin (1:200 dilution) or ZO-1 (1:250 dilution)
antibody for 1 hour at room temperature, cells were washed five times with 5% BSA
(containing 15% glycine) followed by washing five times with PBS before
incubating with the appropriate secondary antibody. Stained cells were then washed
five times using similar procedures. Fluorescence microscopy of immunostained
cells was performed on an IX-71 Olympus inverted microscope where all images
were acquired using the Metamorph imaging software.
F-actin quantification
To visualize F-actin, cells were stained with rhodamine-phalloidin using standard
protocol and imaged using a 60⫻objective. For quantification of fluorescence, we
acquired images of phalloidin-stained cells at 6-12 random fields of observation in
each experiment using a 20⫻objective. After performing background subtraction
of the images, we calculated the average fluorescence intensity per cell for each
field of observation. These values were then normalized with respect to the average
fluorescence value calculated for the control cells for a given experiment.
Normalized fluorescence data of control and Pfn1-siRNA treated cells were pooled
from two experiments, the average values of which were then statistically compared
using a Student’s t-test.
Cell-proliferation assay
HUVECs were plated in triplicate at a density of 20,000 cells per well of a 24-well
plate 24 hours after transfection with the appropriate siRNAs. At different time-
points after transfection (48-96 hours), cells were washed with PBS and stained with
0.5% crystal violet for 15 minutes. After washing cells three times with PBS, dye
was eluted from cells by adding 300 l of 100% ethanol in each well and its
absorbance was measured using a plate-reader. Absorbance data based on triplicate
set of samples for each experimental condition from a total of three independent
experiments were then averaged for statistical comparison using a Student’s t-test.
Cell spreading assay
HUVECs were plated on the wells of a 24-well plate that were pre-coated with
different concentrations of matrigel (25 and 100 g/ml) and the percentages of cells
that showed spreading morphology (appears phase-dense) were determined at
different time-points (30 and 60 minutes) after cell seeding. Percentage of spreading
cells was calculated in each of the three random fields of observation and averaged
based on data from two independent experiments for statistical comparison using a
Student’s t-test.
Wound healing assay
Confluent monolayers of HUVECs cultured in the wells of a 24-well plate was
mechanically scratched using a pipette tip. Cell debris was removed by washing
with PBS before adding complete growth media to the cells. Images of the wound
edges were acquired at three random locations first immediately after wounding and
then at the same locations after 12 hours to assess the wound closure by migrating
HUVECs. Wound closure was quantified by the percentage change in the wound
area per unit time and averaged for three locations per well from a triplicate set of
samples for each experimental condition. This assay was performed 48, 72 and 96
hours after siRNA transfection.
Single-cell migration assay
HUVECs transfected with the appropriate siRNAs were sparsely plated on a 35 mm
tissue-culture dish and after an overnight incubation, time-lapse videomicroscopy
of three random fields were simultaneously performed at an interval of 3 minutes
for a total duration of 90 minutes. The acquired images were analyzed using ImageJ
and Metamorph softwares.
Cord formation assay
Two-hundred microliters of matrigel was polymerized in the wells of a 48-well plate
at 37°C for 30 minutes prior to seeding 25,000 HUVECs on top of matrigel, and
early cord morphogenesis of ECs was assessed after 8 hours by phase-contrast
microscopy. Cord-formation data was quantified by counting the number of nodes
in a given field of observation that had at least three branches (Shen et al., 2005),
which was then averaged for three fields per well from a duplicate set of samples
for each experimental condition. These data were statistically compared using a
Student’s t-test.
We thank Anna DiRienzo for critically reading the manuscript. This
work is supported by grants from the National Institute of Health
(CA108607-01), American Heart Association, Central Research
Development and Competitive Medical Research funds of the
University of Pittsburgh (to P.R.).
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