Tissue Factor Signals Airway Epithelial Basal Cell Survival
via Coagulation and Protease-Activated Receptor
Isoforms 1 and 2
Shama Ahmad1,2, Aftab Ahmad1,2, Raymond C. Rancourt1,2, Keith B. Neeves3, Joan E. Loader1,2,
Tara Hendry-Hofer1,2, Jorge Di Paola4, Susan D. Reynolds1*, and Carl W. White1,2*
1Department of Pediatrics, National Jewish Health, Denver;2Department of Pediatrics, and4Human Medical Genetics Program, University of
Colorado, Denver, Colorado; and3Department of Chemical and Biological Engineering, Colorado School of Mines, Golden, Colorado
Tissue factor (TF) initiates the extrinsic coagulation cascade and is a
high-affinity receptor for coagulation factor VII. TF also participates
in protease-activated receptor (PAR)1 and PAR2 activation. Human
epithelial basal cells were previously purified on the basis of TF
expression. The purpose of this study was to determine if tracheo-
bronchialepithelial basalcell–associated TF drivescoagulation and/
or activates PARs to promote basal cell functions. We used human
tracheobronchial tissues to isolate human airway epithelial cells us-
ing specific cell surface markers by flow cytometry and studied TF
knockdown was done using short hairpin RNA, and TF mRNA was
passage human tracheobronchial epithelial cells were basal cells,
and 100% of these basal cells expressed TF. Basal cell–associated
TF was active, but TF activity was dependent on added extrinsic co-
agulation cascade factors. TF inhibition caused basal cell apoptosis
and necrosis. This was due to two parallel but interdependent TF-
regulated processes: failure to generate a basal cell–associated fi-
brin network and suboptimal PAR1 and PAR2 activity. The data
cell attachment, which maintains cell survival and mitotic poten-
tial. The implications of these findings are discussed in the context
of the potential for repair of airway epithelium in lung disease.
Keywords: tissue factor; epithelium; clotting; fibrin; tracheobronchial
Tissue factor (TF) is a transmembrane cell receptor and cofactor
for factor VII (FVII) and serves biological functions beyond
its established role in blood coagulation (1). TF may act as
a transmembrane signaling molecule by activating and deliver-
ing coagulation factors for signaling, and the TF-coagulation
FVIIa complex activates G protein–coupled receptors, includ-
ing the protease-activated receptors (PARs) (2). Under patho-
logic conditions, TF can be expressed by circulating blood
elements (monocytes, neutrophils, and platelets) and by an as-
sociated elevated number of cell-derived circulating micropar-
ticles. The role of TF in epithelial cell function is much less well
understood than its role in the circulation and in hemostasis.
In acute lung injury, increased coagulation and decreased fi-
brinolysis result in diffuse alveolar fibrin deposition, potentially
increasing lung inflammation (3). Recent evidence indicates that
the damaged alveolar epithelium expresses TF (4) and that
FVII-activating protein expression is increased (5). Alveolar
procoagulant microparticles are also elevated (6), and intrinsic
elevation of tissue factor pathway inhibitor (TFPI) expression is
insufficient to inhibit this procoagulant effect (7). Coagulation
pathologies also occur in conducting airways. For example, the
sulfur mustard analog 2-chloroethyl-ethylsulfide causes produc-
tion of fibrin-rich, airway-occlusive casts after acute inhalation
(8), and the TF pathway is activated in this process (9). Fibrin
cast formation in proximal airways also occurs in burns and in
smoke inhalation, with postextubation airway injuries, and in
children with congenital heart disease (post-Fontan procedure)
(10, 11). Thus, excessive fibrin formation by TF pathway acti-
vation can occur throughout airways, potentially causing ob-
structive and fibrotic pathologies.
Despite its role in tissue damage, TF and its activation of co-
agulation may have protective functions. Coagulation and fibrin
deposition in wounds or other sites of injury (e.g., skin) is critical
to maintain epithelial barrier integrity and to protect against
invading microbes. Recently, low TF–expressing mice demon-
strated delayed wound healing, difficulty in responding to infec-
tion, and the potential for pathologic bleeding (12). Inactivation
of the TF gene or the gene encoding its endogenous inhibitor
(i.e., TFPI) causes embryonic lethality (13). Thus, TF may pro-
vide a homeostatic envelope that protects organisms against
bleeding after injury (14, 15).
The normal bronchial epithelium maintains a barrier against
inhaled harmful agents by efficiently and rapidly repairing
(Received in original form May 24, 2012 and in final form September 17, 2012)
*These authors contributed equally to this work.
This work was supported by National Institutes of Health grants RC1HL099461
(S.D.R. and C.W.W.), R01-ES014448 (C.W.W.), and KL2RR025779 (S.A.) and by
the CounterACT Program, National Institutes of Health, Office of the Director,
and the National Institute of Environmental Health Sciences grant U54 ES015678
(C.W.W.). Foundation support was received from Max and Yetta Karasik Family
Foundation (C.W.W.) and by American Heart Association grant 0830418N (A.A).
Imaging experiments were performed in the University of Colorado Anschutz
Medical Campus Advanced Light Microscopy Core, which is supported in part
by National Institutes of Health/NCRR Colorado CTSI grant UL1 RR025780.
Author Contributions: Concept and design: A.A., S.A., S.D.R., and C.W.W.; anal-
ysis and interpretation: A.A., S.A., R.C.R., K.B.N., J.E.L., T.H.H., J.D.P., S.D.R., and
C.W.W.; manuscript preparation: S.A., S.D.R., and C.W.W.
Correspondence and requests for reprints should be addressed to Shama Ahmad,
Ph.D., Department of Pediatrics, Pediatric Airway Research Center, Mailstop
8615, P15-4016, University of Colorado Denver, 12700 E. 19th Ave, Aurora,
CO 80045. E-mail: email@example.com
This article has an online supplement, which is accessible from this issue’s table of
contents at www.atsjournals.org
Am J Respir Cell Mol Biol
Copyright ª 2013 by the American Thoracic Society
Originally Published in Press as DOI: 10.1165/rcmb.2012-0189OC on October 11, 2012
Internet address: www.atsjournals.org
Vol 48, Iss. 1, pp 94–104, Jan 2013
Although tissue factor (TF) has been previously localized to
the basal cell region of airways, the role of TF activity in
tracheobronchial epithelial basal cell function is unknown.
TF activity on the surface of tracheobronchial epithelial
basal cells causes fibrin formation, leading to provisional
matrix formation, basal cell attachment, survival, and pro-
liferation. We demonstrate that TF activity is essential for
airway epithelial repair after injury.
affected areas. Epithelial wounding exposes the basement mem-
brane and extracellular matrix and causes TF activation (16). TF
is expressed by airway epithelial basal cells (16–18) andhas been
used as a marker for basal cell isolation and purification (16, 18).
Fibrin formation may be required for repair in airway epithelial
cell lines (19). We have demonstrated that human airway epithe-
lial basal cells use cell surface TF to form fibrin networks on their
surface and that this fibrin network formation was critical for their
survival. We also established a role for TF in PAR activation and
showed that this activity enhanced basal cell survival and prolifer-
ation. Together, these findings suggest that TF activity may be
a critical component of the epithelial repair process.
MATERIALS AND METHODS
Human Airway Epithelial Cell Isolation
Human tracheobronchial tissue was procured from National Disease Re-
search Interchange under National Jewish Institutional Review Board–
approved protocols. Cell harvest and culture were performed using
established procedures (20). Briefly, epithelial cells were removed from
lower trachea and bronchi by protease XIV digestion and plated in
bronchial epithelial growth medium on collagen-coated dishes.
Fluorescence-Activated Cell Sorting Analysis
One million passage 1 cells were incubated with anti–TF-FITC (Amer-
ican Diagnostica, Stamford, CT), anti-CD31-APC (eBioscience, San
Diego CA) (to exclude endothelial cells), anti–CD45-APC (eBio-
science) (to exclude hematopoietic cells), and anti–CD90-APC (eBio-
science) (to exclude fibroblasts) antibodies diluted in PBS containing
BSA (1%) for 30 minutes at 48C. Nonimmune IgG1k-APC and IgG1-
FITC were used as isotype controls. Cells were washed twice and sus-
pended in PBS-BSA. The DNA-intercalating dye 7-aminoactinomycin
D (7-AAD) was added to stain dead cells. Cell sorting was performed
with a Moflo-XDP high-speed cell sorter (Beckman Coulter, Brea, CA)
equipped with a CyCLONE automated cloner. Compensation for spec-
tral overlap was done using single-color positive and negative compen-
sation beads (BD Biosciences, San Jose, CA). Gates were set according
to unstained and isotype controls. Doublets were excluded by side
scatter versus forward scatter (FS) and FS-INT area versus FS dot
plots. Cell count versus 7-AAD plot defined dead cells. TF-positive
basal cells were then sorted at a rate of 5,000 events per second. Sorted
cells were reanalyzed for purity and viability.
Knockdown of TF was performed using lentiviral-mediated short hair-
pin RNA (shRNA) (Sigma, St. Louis, MO) transduction as described
previously (21). The details of RT-PCR to quantify mRNA are pro-
vided in the online supplement.
Cells were cultured on collagen-coated coverslip chamber slides. Thirty
minutes before imaging, FITC-labeled fibrinogen was added (22). Con-
focal imaging was performed as described previously (8, 23). A two-
photon LSM 510 confocal microscope (Zeiss, Thornwood, NY) was
used to obtain images and Z-stacks.
Scanning Electron Microscopy
Human airway epithelial basal cells were cultured at approximately
10,000 cells/cm2on 14-mm coverslips. Twenty-four hours later, treat-
ments were performed, and cells were fixed in 2.5% glutaraldehyde in
0.1 M sodium cacodylate buffer (pH 7.4) for 4 hours, rinsed four times,
and stored in 0.1 M sodium cacodylate buffer (pH 7.4). Samples were
dehydrated in graded ethanol solutions (50%, 70%, 80%, 90%, 100%,
and 100%) for 5 minutes each and chemically dried once in 50% and
then twice in 100% hexamethyldisilazane for 5 minutes each time.
Samples were air dried overnight at room temperature in a desiccator.
A thin film (z 10 nm) of gold was sputtered on samples before imaging.
Images were taken with an accelerating voltage of 5 KV and at a 6-mm
working distance on a JEOL 7000 FE (JEOL, Peabody, MA) scanning
electron microscope. Each experimental condition was imaged in 10
positions at magnifications of 2,0003 and 25,0003. Brightness and con-
trast were adjusted on scanning electron microscopy images to opti-
mize viewing. No other image manipulation was performed.
ANOVA with Tukey comparisons was calculated using JMP Software
(SAS, Cary, NC). P , 0.05 was considered statistically significant.
Human Tracheobronchial Airway Epithelial Cells Express
TF In Vitro
To define TF function, we first determined if isolated human tra-
cheobronchial airway epithelial cells (HAECs) expressed this
marker. HAECs were isolated from tracheas and bronchi of
patients dying from nonlung disease (20). Passage 1 HAECs
were harvested at 80% confluence. Immunostaining indicated
that the majority expressed TF (see Figure E1A in the online
supplement). Next, TF expression was quantified in passage 1
HAECs using flow cytometry. An iterative gating strategy (Fig-
ure 1A) was used. Dead cells were excluded by 7-AAD staining,
and nonepithelial cells (hematopoietic cells, endothelial cells,
and fibroblasts) were excluded by CD45, CD31, and CD90
staining (Figure 1A). TF expression on live epithelial cells
was present in 97 6 5% (n ¼ 14 donors) (Figure 1B). TF fluo-
rescence intensity was uniformly distributed about the mean,
indicating that TF positivity defined a single cell population.
To determine the phenotype of live/epithelial/TF1 cells, cyto-
spin preparations were stained for cell type–specific markers. Be-
cause motile cilia can be sheared during FLOW cytometry,
ciliated cells were detected by staining for g tubulin. Basal bodies
expressing this antigen are located at the terminal plate of ciliated
cells and in the centrosomes of all cells. Mucus cells were detected
by staining for mucins Muc5AC and Muc5B. Basal cells were
detected by staining for keratin 5. All TF-positive cells expressed
keratin 5 (Figure 1C). Thus, passage 1 HAECs were basal cells
and are henceforth called “‘basal cells.”
Basal Cells Express Active TF
TF initiates clotting by interacting with Factor VII in the pres-
ence of calcium, resulting in activation of Factor X. Factor Xa
verts fibrinogen to fibrin. Thus, TF activity can be evaluated us-
ingan assay detecting cleavage of a Factor X substrate (9).Initial
experiments demonstrated that basal cell cultures expressed
active TF. This activity was proportional to basal cell number
(Figure E1B). TF activity was detected only when Factor VII
and Factor X were added (Figures E1C and E1D).
To determine if Factor Xa generation was TF dependent, vary-
ing numbers of basal cells were plated at low cell density and in-
cubated for 24 hours (Figure 1D). Cultures were then treated with
an isotype-matched control antibody or polyclonal anti–TF-
antibody for 30 minutes, and TF activity was assayed (Figure
1E). TF was detected in isotype IgG-treated cultures, and this
activity was proportional to the number of cells plated (Figure
1E). In contrast, cultures treated with TF antibody exhibited sig-
nificantly less TF activity at all cell inputs. To further evaluate TF
activity, basal cells were treated with human recombinant TFPI.
Vehicle-treated cultures exhibited cell-number–dependent TF ac-
tivity (Figure 1F). In contrast, TFPI-treated cultures exhibited no
detectable TF activity. The basal cells expressed active TF. We
suggest that fibrin formed from TF activity on basal cell surface
Ahmad, Ahmad, Rancourt, et al.: Tissue Factor Regulates Basal Cell Survival95
serves as a cell attachment matrix. We also determined whether
the collagen matrix in these cultures was capable of affecting
TF-dependent FX cleavage. Collagen at concentrations used in
coating the plastic tissue culture dishes (coated or in solution)
did not affect TF activity (Figure E2).
TF Is Necessary for Basal Cell Survival
vector. Treatment with TF-shRNA decreased TF mRNA (Figure
2B) and protein (Figure 2C) expression by approximately 50%.
To determine ifTF knockdown promoted celldeath, caspase 3/7
release was measured in supernatant from shRNA-treated cultures.
This did not differ between control and nontarget shRNA-treated
cells (Figure 2D). However, caspase 3/7 levels increased approxi-
mately 2-fold in TF-shRNA–treated cells. To determine if cells
underwent apoptosis or necrosis, the PI/annexin V flow cytometry
method was used (24). Treatment with nontarget shRNA increased
apoptotic cells 1.5-fold but did not increase the necrotic cell
frequency (Figure 2E). In contrast, treatment with TF-shRNA in-
creased apoptosis and necrosis 2-fold.
Proliferation of shRNA-treated cells was measured by [3H]
thymidine incorporation 48 hours after shRNA treatment. The
rate of proliferation did not differ between untreated and non-
target shRNA-treated cells but decreased by approximately
30% in cells treated with TF-shRNA (Figure 2F). These data
indicated a direct TF effect on basal cell survival and a second-
ary effect on basal cell proliferation.
Active TF Is Necessary for Basal Cell Survival
To determine if TF activity was required for basal cell survival,
cells adhered for 24 hours were treated with diluent or TFPI for
24 hours, and cell number was analyzed by the MTT assay (25)
(Figure 2G). TFPI treatment decreased cell number by 75% at
all cell inputs (Figure 2H), indicating that TF activity was nec-
essary for basal cell survival.
To investigate the mechanism of TFPI-induced basal cell at-
trition, basal cells adhered for 24 hours were treated with vehicle
Figure 1. Human tracheobronchial basal cells (HAECs) express tissue factor (TF). Passage 1 HAECs were cultured for 5 days on collagen-coated
plates as described in MATERIALS AND METHODS. Cells were harvested and stained, and fluorescence-activated cell sorting was performed. (A) Live cells
were defined as those that excluded 7-aminoactinomycin D (7-AAD), and epithelial cells were identified as those that did not stain with allophy-
cocyanin (APC)-tagged CD31 (endothelial cells), CD45 (hematopoietic cells), or CD90 (fibroblasts cells). (B) TF-positive cells were identified by
comparison of live epithelial cells that were stained with an isotype control (gray) and FITC-conjugated TF antibody (black). The data represent three
independent experiments performed with cells isolated from three different donors. (C) To determine if purified HAECs are basal cells, live,
epithelial, TF1, passage 1 HAECs were recovered by flow cytometry and deposited onto microscope slides using a cytocentrifuge. Because cilia
tend to be sheered during flow cytometry, g-tubulin was used to detect basal bodies located at the terminal plate of ciliated cells. Mucus cells were
detected by staining for MUC5AC and MUC5B. Basal cells were detected by staining for keratin 5 (K5, purified basal cells, differentiated epithelial
cell cultures, and fixed tracheal tissues were used as positive controls). Positive controls for MUC5B and g-tubulin were also used. Images repre-
sentative of TF-positive cells from three donors are shown. (A) Secondary antibodies only. (B) g-Tubulin (green). (C) Muc5AC (red) and Muc5B
(green). (D) Keratin 5 (red). (D) Basal cells were allowed to adhere to collagen-coated plates for 24 hours and then treated for 30 minutes with an
isotype control antibody or antitissue factor antibody or with diluent or tissue factor pathway inhibitor (TFPI). TF activity was determined by the
S-2765 FXa substrate assay in the presence of Factor VII, Factor X, and CaCl2. (D) Effect of anti-TF antibody on TF activity. Squares, isotype control;
triangles, anti-TF antibody-treated cells. (E) Effect of TFPI treatment on TF activity. Squares, diluent control; triangles, TFPI treatment. Data are
presented as mean 6 SEM (n ¼ 6). *P , 0.05 for control versus treated cultures.
96 AMERICAN JOURNAL OF RESPIRATORY CELL AND MOLECULAR BIOLOGYVOL 482013
or TFPI and assayed for apoptosis and necrosis (Figures 2I and
2J). Treatment with TFPI increased the necrotic cell index
at each time point and the apoptotic cell index at the 4- and
18-hour time points as compared with diluent, which had no
effect. Similar results were obtained when cell death was
assayed by caspase release or annexin binding (Figure E3).
These data confirmed that TF activity was necessary for basal
Figure 2. TF expression is nec-
essary for basal cell survival. (A)
Basal cells were cultured on
transduced with nontarget or
TF-specific short hairpin RNA
(shRNA). TF knockdown was
assayed 48 hours after transduc-
tion. Cell death was assayed
48 hours after transduction,
and proliferation was assayed
72 hours after transduction.
(B) TF mRNA abundance was
analyzed using quantitative re-
al-time RT-PCR. Fold TF indi-
cates the ratio of TF mRNA to
18 s rRNA normalized to 1 for
nontarget shRNA. (C) Western
blot analysis of TF protein abun-
dance. Lane 1, nontransduced
transduced cells and b-actin
served as the loading standard.
(D) Caspase 3/7 activity was
measured in culture medium
using the Caspase-Glo 3/7 as-
say. Caspase 3/7 release indi-
with the background signal
subtracted. (E) The mode of
cell death was evaluated in at-
tached cells and floating cells
using the annexin V (apopto-
sis) and Sytox Green (necrosis)
assay. The frequency of cells
that were apoptotic
bar) or necrotic (black bar) is
presented as a percent of all
cells. (F) Proliferation was
evaluated by analysis of (3H)-
thymidine incorporation. Fold
indicates the counts per min-
ute for the test sample relative
Each data set represents three
independent experiments per-
formed with cells isolated from
three different donors. The
data are presented as mean 6
SEM (n ¼ 6). *P , 0.05 relative
to the nontransduced control.
#P , 0.05 relative to the non-
target shRNA treated control.
(G) To determine if active tis-
sue factor is required for basal
cell survival, basal cells were plated on collagen-coated plates and allowed to adhere for 24 hours. Cells were then treated with diluent or TFPI for
18 hours, and cell number, necrosis, and apoptosis were evaluated. (H) Cell number was evaluated using the MTT assay. (I) The frequency of
necrotic cells was determined by Sytox green assay. (J) The frequency of apoptotic cells was determined by annexin V apoptosis assay. The inset in
I is a flow cytometry image for analysis of alive (peak A unstained), apoptotic (peak B, Annexin FITC), and necrotic (peak C, Sytox green) cells. Each
data set represents three independent experiments performed with cells isolated from three different donors. Data are presented as mean 6 SEM
(n ¼ 6). *P , 0.05 relative to diluent controls. White bar, diluent; black bar, TFPI.
Ahmad, Ahmad, Rancourt, et al.: Tissue Factor Regulates Basal Cell Survival 97
Because caspase 3 is a key enzyme of final pathway of apo-
ptosis, to distinguish between extrinsic and intrinsic apoptotic
pathway, the role of caspase 8 and 9 in TFPI-induced basal cell
apoptosis was investigated. At 6 and 18 hours after TFPI treat-
ment, caspase 8 was significantly elevated (Figure E4A). How-
ever, caspase 9 was not significantly different (not shown),
indicating a major role of the extrinsic pathway.
Phosphorylation of the cytoplasmic domain may play an im-
portant role in TF-mediated cellular adhesion (26). We there-
fore investigated if TFPI treatment altered TF phosphorylation.
TFPI treatment did not affect TF phosphorylation (Figures E4B
and E4C). PARs phosphorylate the cytoplasmic domain of TF
(27). PAR agonists were used as positive controls in these stud-
ies, and they did increase phosphorylation of TF.
Basal Cells Express a Subset of Clotting
TF initiates the extrinsic clotting cascade that requires multiple fac-
tors for propagation (28).These clotting factors may be supplied by
TF-expressing cells or by the surrounding environment. Thus, we
determined which clotting factors and related molecules were
expressed by TF-positive basal cells. Gene expression was assayed
by quantitative RT-PCR using cDNA substrate prepared from
basal cells and human liver, a positive control (Figure E5A).
mRNA abundance in basal cells was expressed as percentage of
that in liver. Gene expression was also evaluated in scratch-
wounded basal cells to determine potential coagulation factor
induction by injury. Basal cells expressed high levels of TF, TFPI
(Figures E5B and E5C), and fibrinogen g chain mRNA (Figure
E5H). In contrast, basal cells expressed very low levels of
thrombin, antithrombin, and Factor V and X mRNA (Figure
E5D–E5G). Basal cells did not express detectable Factor VII or
IX mRNA (not shown). Thus, basal cells expressed an incom-
plete repertoire of clotting factors.
Basal Cells Form a TF-Dependent Fibrin Network
on their Surface
We sought to determine how TF activity promotes basal cell sur-
vival. It is known that activation of coagulation causes formation
of fibrin networks and that these networks can seal epithelial
defects (14, 15). We hypothesized that fibrin networks serve
as substrates for attachment of reparative cells. To test this
concept, basal cells were cultured at 80% confluence on glass
coverslips (submerged) or on air–liquid interfaces for 10 days.
FITC-labeled fibrinogen was used to determine if fibrinogen
was converted to fibrin (Figure 3A). Fibrin networks were
detected 30 minutes after adding labeled fibrinogen to sub-
merged or air–liquid interface cultures (Figure 3B, panels B
and G). Fibrin network formations were enhanced by pro-
thrombin, Factor VII, and Factor X (Figure 3B, panels C and
H). The addition of TFPI decreased cell surface fibrin networks
(Figure 3B, panels D and I), and the specific fibrin cross-linking
inhibitor peptide Gly-Pro-Arg-Pro (GPRP) (29) eliminated fibrin
networks (Figure 3B, panels E and J). Thus, basal cell–associated
Figure 3. Clotting cascade gene
expression and fibrin formation
in basal cells. (A) To determine if
basal cell–associated TF medi-
ates fibrin network formation,
basal cells were cultured on
collagen-coated glass coverslip
chambers (submerged mono-
layers) for 24 hours or on
collagen-coated snapwells at
the air–liquid interface for 7 days.
Cells were treated with TFPI or
peptide Gly-Pro-Arg-Pro (GPRP)
for 30 minutes before addition
of FITC-fibrinogen (Fbn) and
CaCl2 or FITC-fibrinogen, pro-
VII (FVII), Factor X (FX), and
CaCl2. Cells were incubated
for 30 minutes, and fibrin net-
work (green) formation was
evaluated using a fluorescence
microscope. Arrows indicate fi-
brin networks, white arrowheads indicate large fibrin networks, and open arrowheads indicate areas without fibrin networks. (B) Fibrin network
formation under various conditions. (A–E) Submerged cultures. (F–J) Air–liquid interface cultures. Treatment conditions: A and F, controls; B and G,
Fbn; C and H, Fbn1Prothro1FVII1FX; D and I, Fbn1TFPI; E and J, Fbn1GPRP. (C) Confocal microscopy of fibrin networks formed by submerged
basal cells treated with FITC-fibrinogen and CaCl2in HBSA for 30 minutes (please see online supplement). A merge of the differential interference
contrast (transmitted light) and green fluorescence image is shown. Inset shows a three-dimensional reconstruction of fibrin network. A video
demonstrating the z stacks is shown in online supplement (Figure 3 video1.mov). (D) Scanning electron microscopic analysis of fibrin networks on
the surface of basal cells. Basal cells were cultured on collagen-coated glass coverslips for 24 hours and treated with diluent, GPRG, or TFPI for
30 minutes. A second set of cultures was grown on collagen-coated plastic and transduced with nontarget or TF shRNA for 24 hours. Cultures were
then treated with CaCl2in HBSA and various additives: no factors (control); fibrinogen (Fbn); or Fbn, prothrombin (prothro), Factor VII (FVII), Factor
X (FX, 1 nM). Cells were then fixed and processed for scanning electron microscopy. Low magnification (mag) images of basal cells treated with
agent(s) indicated at the top of each column. Low Mag: 32,000; scale bar ¼ 10 mm. High Mag: 325,000; scale bar ¼ 1 mm. White arrows, tight
network fibrin fibers on cell; white arrowheads, smooth cell surface; open arrowheads, fibrin networks of fibrin formed from exogenously added
fibrinogen; open arrows, larger fibrin aggregates. (E) Scanning electron microscopy analysis of fibrin upon TF inhibition using TFPI and TF shRNA in
basal cells. Arrows as indicated in D. Scale bar ¼ 10 mm.
98 AMERICAN JOURNAL OF RESPIRATORY CELL AND MOLECULAR BIOLOGYVOL 48 2013
TF-initiated conversion of fibrinogen to fibrin was enhanced by
added coagulation factors and was dependent on active TF. The
location of the fibrin network was examined by confocal micros-
copy. Incubation of basal cells with FITC-conjugated fibrinogen
in serum-free medium caused formation of dense fibrin networks
(Figure 3C, and see video 1.mov in the online supplement). Fibrin
network formation was inhibited by the thrombin inhibitors
(hirudin and PPACK) and indicated that this process required
thrombin (not shown). Finally, fibrin networks required calcium
and were fully inhibited by 1 mM EDTA (not shown). Thus,
TF-mediated formation of fibrin networks occurred at the basal
Ultrastructural Characteristics of the Fibrin Network
To evaluate cell surface fibrin networks in greater detail, basal cell
basal cells grown in serum-free medium, ruffled fibrin filaments
were detected (Figure 3D, panels 1 and 2). These extended to
neighboring cells and covered surfaces of untreated control cells.
Cells treated with fibrinogen had larger, thicker fibrin networks
that frequently formed floating spheres (Figure 3D, panels 3 and
4). The addition of Factor VII, Factor X, and prothrombin resulted
in more extensive networks (Figure 3D, panels 5 and 6). Under
these conditions, the cell surface appeared to be more ruffled and
was coated with thicker fibrin filaments. Treatment with cross-
linking inhibitor GPRP decreased fibrin filament density (Figure
3D, panels 7 and 8), and the diminished networks resembled short-
ened stalks (more images in Figure E6). Thus, basal cells formed
fibrin networks at their surfaces, the extent and complexity of
which were increased by the addition of coagulation cascade fac-
tors and components.
To determine if TF activity was necessary for fibrin network
formation, basal cells were treated with TFPI and evaluated by
scanning electron microscopy. TFPI decreased fibrin network
formation and resulted in a smoother cell surface (Figure 3D,
panels 1 and 2). Knockdown of TF by shRNA decreased fibrin
network relative to cells transfected with nontarget shRNA
(Figure 3E, panels 3 and 4). Thus, basal cell–derived TF expres-
sion and activity are required for cell surface fibrin network
Fibrin Network Formation Is Necessary for Basal
To further determine if fibrin network was necessary for basal
cell survival, cells were allowed to adhere for 24 hours and trea-
ted with diluent or tissue-type plasminogen activator (tPA) (Fig-
ure 4A). tPA is a serine protease that is found in a variety of
mammalian tissues, including airway epithelial cells (30). It con-
verts plasminogen to plasmin, which is a key activity involved in
fibrinolysis. Caspase 3/7 release and cell number were measured
18 hour after tPA treatment. tPA treatment increased apoptosis
(Figure 4B) and decreased cell number (Figure 4C) at each cell
input. The extent of cell death induced by tPA was similar to
that caused by TFPI, as shown by experiments that used an
equivalent basal cell to tPA molecule ratio to those used in
studies involving TFPI-induced cell death (1:7 for TFPI and
1:9 for tPA at 280 nM) (Figures E7A and E7B). We also used
Gly-Pro-Arg-Pro (GPRP), a peptide inhibitor of fibrin cross-
linking, to further assess the role of fibrin network in basal cell
survival. Treatment with GPRP enhanced basal cell apoptosis
and decreased proliferation (not shown). The loss of cell viabil-
ity by disruption of fibrin network using tPA and/or GPRP
confirmed that TF-mediated formation of cell surface fibrin net-
works promoted basal cell survival.
TF Activates Protease-Activated Receptors Isoforms
1 and 2, Allowing Basal Cell Survival
TF also activates protease-activated receptor isoforms 1 and 2
(PAR1 and PAR2) (31). PAR1 and PAR2 expression in airway
epithelial cells has been described (32). We confirmed expres-
sion of PAR1 and PAR2 mRNAs in basal cells by qRT-PCR
analysis (Figure 5A). These were not significantly modified by
scratch injury (Figures 5B and 5E).
PARs mediate their effects by mobilizing cytosolic calcium.
We used PAR1 and PAR2 peptide antagonists and the cyto-
solic calcium-sensing dye Fluo-4AM to determine if basal cells
expressed active PAR1 and PAR2. Treatment with the PAR1
with the PAR2 antagonist FSLLRY-NH2 demonstrated that
basal cells expressed active PAR1 (Figures 5C and 5D) and
PAR2 (Figures 5F and 5G). To determine if baseline basal
cell PAR1 or PAR2 could be activated further, we treated
basal cells with PAR1 agonist TFLLRN or PAR2 agonist
SLIGRL-NH2. Both stimulated calcium mobilization (Fig-
ures 5C, 5D, 5F, and 5G). Agonist-induced PAR1 activity was
specifically inhibited by PAR1 antagonist and partially inhibited
by PAR2 antagonist. Similarly, agonist-induced PAR2 activity
was inhibited by the PAR2 antagonist and partially inhibited by
Figure 4. The fibrin network is necessary for basal cell survival. (A) Basal
cells were allowed to adhere for 24 hours and then treated for 18 hours
with diluent or tissue plasminogen activator (tPA). Apoptosis and cell
number were assayed. (B) Caspase 3/7 activity was measured in culture
medium and using Caspase-Glo 3/7 assay. Caspase 3/7 release indicates
luminescence (in relative luminescence units) with the background signal
subtracted. White bars, control (diluent); black bars, tPA. (C) Cell number
was determined using the MTT assay. White bars, control (diluent); black
bars, tPA. Each data set represents three independent experiments per-
formed with cells isolated from three different donors. Data are presented
as mean 6 SEM (n ¼ 8). *P , 0.05 for diluent versus tPA.
Ahmad, Ahmad, Rancourt, et al.: Tissue Factor Regulates Basal Cell Survival 99
the PAR1 antagonist. Thus, basal cells expressed active PARs,
and their baseline activity was submaximal.
assayed 30 minutes later (Figure 6A). TFPI completely inhibited
agonist-mediated PAR1- or PAR2-dependent cytosolic calcium mo-
bilization (Figure 6B). To determine if agonist or antagonist-treated
cells were viable and if PAR inhibition was specific, ATP-induced
cytosolic calcium signaling was evaluated. This response was unaf-
fected upon TF inhibition (not shown). These data indicated that
active TF was necessary for PAR1 and PAR2 activation.
TF Functions Upstream of PAR1 and PAR2 to Promote Basal
The previous data sets indicated that TF activated coagulation and
tioned, we first determined if pretreatment with PAR agonists or
antagonists could overcome the TFPI-mediated decrease in basal
treated with PAR agonists or antagonists for 8 hours. Cells were
then treated with diluent or TFPI apoptosis, and cell number was
assayed 18 hours after treatment (Figure 6C). TFPI treatment
increased apoptosis as indicated by caspase 3/7 release (Figure
6D). Prior agonist-mediated activation of PAR1 or PAR2 re-
versed TFPI-mediated apoptosis. Similar results were obtained
when basal cells were incubated with a combination of PAR1
and PAR2 agonists (not shown). These data indicated that TF
functioned upstream of PAR1 and PAR2.
We next evaluated the effect of diluent/TFPI after antago-
nism of PAR1 or PAR2. Inhibition of PAR1 or PAR2 by their
antagonists did not alter TFPI-induced caspase release (Figure
6D). However, antagonist treatment followed by TFPI treat-
ment increased caspase release 2.5-fold (Figure 6D). The basal
levels of caspases may be nonspecific, pertinent to the culture
system and methods, and not affected by PARs. The additive
effect of PAR antagonism and TF inhibition suggested that TF
functioned in two processes—PAR activation and fibrin net-
work formation—and that the function of both pathways was
necessary for basal cell survival.
We then determined if pretreatment with PAR agonists or
antagonists overcame the TFPI-induced decrease in basal cell
proliferation. As indicated, basal cells were treated with agonist
or antagonist followed by diluent or TFPI. Cell survival was de-
termined using MTT assay 18 hours later (Figure 6E). As shown
Figure 5. Expression and activity
of protease-activated receptors
(PAR1 and PAR2) in basal cells.
(A) Passage 1 basal cells were
allowed to adhere to collagen-
coated plates for 24 hours. One
set of cultures was mechanically
stimulus to observe potential in-
duction), and all cultures were in-
total RNA recovery. An additional
set of cultures was loaded with
dye Fluo-4AM and then treated
with PAR agonist or antagonist
for 2 hours. Calcium flux was
then measured. (B and E) Expres-
sion of PAR1 (B) and PAR2 (E)
mRNA was determined by quan-
titative RT-PCR. Individual mRNAs
were normalized to 18S rRNA,
and fold change relative to hu-
man liver mRNA (a positive con-
trol) was calculated. Data are
presented as the mean 6 SEM
(n ¼ 3). (C and F) The time
course forcytosoliccalcium mobi-
lization under various treatment
conditions. (C) PAR1 agonist
(Ag, 10 or 100 mM TFLLRN)
100 mM, 3-mercaptopropionyl-
PAR2 agonist (30 or 60 mM of
SLIGRL-NH2) and PAR 2 antag-
onist (100 mM, FSLLRY-Amide).
(D and G) The mean peak values
for the curves presented in C and
F, respectively. Data are presented
as the mean6 SEM (n ¼ 3). *P ,
0.05 for buffer versus treated cells.
100AMERICAN JOURNAL OF RESPIRATORY CELL AND MOLECULAR BIOLOGY VOL 482013
previously, TFPI-treatment decreased basal cell number relative
to vehicle treatment (Figure 6E). Prior activation of PAR1 or
PAR2 receptors by their respective agonists prevented the
TFPI-mediated decrease in cell number. Preinhibition of PAR1
or PAR2 by treatment with their respective antagonists resulted
in decreased basal cell number. This effect was exaggerated by
TFPI treatment. We also evaluated if PAR activation protected
against TF knockdown–induced basal cell apoptosis. PAR acti-
vation significantly decreased caspase release in TF-shRNA–
transduced basal cells (Figure E8). These data also supported
the conclusion that TF played a role in multiple basal cell sur-
TF-Dependent Fibrin Network Formation and PAR Activation
Cooperate To Promote Basal Cell Survival
To determine if TF-dependent activation of coagulation and of
PAR receptors occurred in parallel or sequence, basal cells were
adhered for 24 hours and treated with PAR1 or PAR2 antagonist.
Two hours later, fibrin networks were evaluated in the presence of
fibrinogen and calcium chloride (Figures E9B, E9E, and E9H). Fi-
brin network formation was not altered by PAR1 or PAR2 antag-
onism. Similarly, the extensive fibrin network that formed in the
presence of added prothrombin, Factor VII, and Factor X was
not altered by PAR inhibition (Figures E9C, E9F, and E9I).These
data suggested that the basal cell–associated TF functioned as part
of two distinct cascades: (1) PAR1 and PAR2 activation and (2)
initiation of coagulation (Figure 7).
The purpose of this study was to determine the impact of TF-
dependent coagulation and PAR activation on basal cell survival.
Cell surface TF-dependent coagulation was previously demonstrated
in the human lung, transformed tracheal epithelial cells, and
bronchial epithelial cells or cell lines (19, 33). Antibodies
against TF, fibrinogen, and FXIIIa inhibited repair (19). De-
spite these important findings, their significance has not been
extended beyond those in transformed or malignant cells.
TF is a marker for nasal and embryonic basal cells (16–18). We
showed that all first-passage human tracheobronchial epithelial cells
were keratin 5–positive basal cells that also expressed TF. Thus, TF
is also a marker for tracheobronchial basal cells. Basal cells play
dual roles in airways by attaching epithelium to the airways and by
serving as stem or progenitor cells (16, 34–36). Tracheobronchial
basal cells can restore a fully differentiated epithelium in vitro (16,
36). TF-expressing basal cells from nasal airways had several-fold
greater colony forming ability than TF-negative columnar epithelial
cells (16). Thus, TF expression in human tracheal basal cells may
have critical functions beyond coagulation.
We showed that basal cell–associated TF was active, whereas
TF activity was regulated by the availability of coagulation cas-
cade factors VII and X. Our data contrast with those from
bronchial epithelial cell lines wherein mechanical injury caused
fibrin matrix formation and rapid generation of FVII, VX,
FXIIIa, D-dimers, fibrinogen, and soluble fibrin. These data
suggest that an important difference between primary basal cell
isolates and cell lines is the selection of cells that express a more
complete spectrum of preformed coagulation proteins (19). The
limited detectability of some of the clotting factors, such as
FVII in primary basal cell isolates, could be due to short mRNA
half-lives (37). However, our functional assays and kinetic anal-
yses indicate that basal cell–associated TF activity is dependent
on added FVII and FX. The present study suggests that human
tracheobronchial basal cells contribute to fibrin formation, but
this activity is dependent upon additional plasma-derived fac-
tors. This codependence may be critical to regulate fibrin for-
mation in airways such that this does not occur except when
vascular integrity is compromised.
Figure 6. Basal cell–associated TF is necessary for PAR1
and PAR2 activity and functions upstream of PAR1 and
PAR2 to promote basal cell survival. (A) Basal cells were
grown to confluence on collagen-coated, clear–
bottom, black-walled, 96-well plates. Cells were loaded
with cytosolic calcium–sensing dye Fluo-4AM Cells for
90 minutes and then incubated with diluent or TFPI for
30 minutes. Cells were then stimulated with buffer
or agonist. The PAR1 agonist (PAR1 Ag) was TFLLRN,
and the PAR 2 agonist (PAR2 Ag) was FSLLRY-amide.
(B) PAR activity. The mean peak Fluor-4AM (Fluo) value
and fold change relative to control were determined.
Each data set represents three independent experi-
ments performed with cells isolated from three differ-
ent donors. Data are presented as mean 6 SEM (n ¼
3). *P , 0.05 for buffer versus agonist treatment.#P ,
0.05 for buffer versus TFPI and agonist. (C) Basal cells
were allowed to adhere for 24 hours and treated with
PAR agonists (Ag) (PAR1: TFLLRN; PAR: 2 FSLLRY-amide)
or antagonists (Ant) (PAR 1: 3-mercaptopropionyl-
F-Cha-Cha-RKNDK-amide; PAR 2: antagonist FSLLRY-
amide) for 8 hours. Cells were then treated with diluent
or TFPI, and caspase release or cell number were mea-
sured 18 hours later. (D) Caspase 3/7 activity was
measured in culture medium and using the Caspase-
Glo 3/7 assay. Caspase 3/7 release indicates lumines-
cence with background signal subtracted. White bar,
diluent; black bar, TFPI. (E) Cell number was evalu-
ated using MTT. White bar, diluent; black bar, TFPI.
Each data set represents three independent experi-
ments performed with cells isolated from three different donors. Data are presented as mean 6 SEM (n ¼ 3). *P , 0.05 for comparisons to
untreated control.#P . 0.05 for comparisons to TFPI-treated control.
Ahmad, Ahmad, Rancourt, et al.: Tissue Factor Regulates Basal Cell Survival 101
sis and necrosis. Thus, TF affects cell functions that may not be
related to TF’s clotting function. The TF–FVIIa complex is known
to activate PARs, promoting cell survival and proliferation (38).
The antiapoptotic role of TF was demonstrated previously in
TF-positive BHK cells where FVIIa binding to TF protected
against apoptosis induced by growth factor deprivation (39).
The TF/FVIIa complex in another study also activated antiapop-
totic proteins like Bcl2, and TF inhibition enhanced apoptotic cell
death in certain tumor cells (40). Furthermore, TF overexpression
in cardiomyocytes protected against TNF-induced apoptosis (41).
By contrast, TF inhibition via TFPI also caused FAS ligand–
mediated apoptotic cell death in macrophages (42). In vascu-
lar smooth muscle cells, TFPI induction caused inhibition of
proliferation and enhanced apoptosis (43). Although TF-mediated
fibrin deposition and consequent pathogenesis of airway disease
has been reported, we now establish a role of TF in promoting
airway epithelial cell and basal cell survival.
We showed that TF was necessary to form basal cell–associated
fibrin networks. Indeed, fibrin network formation by blood vessel–
associated cell types has been previously noted. For example, cell
surface fibrin formation from exogenously added fibrinogen was
found in fibroblasts, where thrombin, FVII, and FX additives were
required (22). TF expression and fibrin on smooth muscle cell sur-
faces promotes proliferation via a thrombin- and PAR-dependent
mechanism (44). Although human bronchial epithelial cell line
16HBE can up-regulate fibrinogen upon wounding, direct dem-
onstration of fibrin network formation by primary basal cells is
a new finding. This study is also unique in using scanning elec-
tron microscopy to demonstrate TF-dependent fibrin networks
on basal cell surfaces. Inhibition by GPRP confirmed that the
ruffled structure on epithelial cell surfaces was fibrin. In sum,
human airway epithelial basal cells support formation of fibrin
networks on their surfaces, and this process favors proliferation.
Given the role of PARs in coagulation-dependent signaling,
we suspected that PARs might be active in basal cells. PARs
contribute to tissue responses to injury, including repair and cell
survival (45). PARs are widely expressed within the respiratory
tract, and PAR1 and PAR2 are the dominant isoforms in tracheal
epithelium (46). Previous studies suggested that PARs are impor-
tant in downstream TF-dependent effects (47). We report here
that TF was necessary for optimal PAR1 and PAR2 activity.
This study indicates that PARs confer prosurvival and pro-
proliferative effects of TF. PAR1 and PAR2 activation, via li-
gand binding, may also activate TF (27). Thus, PAR signaling
may “self-amplify” or feed forward. Binding of TF to FVIIa
activates PARs and regulates prosurvival proteins, thereby
influencing proliferation and survival (48). Binding of active
Factor VII causes cytosolic calcium mobilization and signal
transduction via mitogen-activated protein kinase phosphoryla-
tion (49). Factor VII–dependent PAR1 signaling promotes bar-
rier function in endothelium (50). PAR2 can also be protective
against virus-induced lung injury (51). By contrast, PAR2-
dependent increases in TF-dependent fibrin formation and
myofibroblast differentiation may promote pulmonary fibro-
sis (52). Here we report that PARs confer the prosurvival and
proproliferative effects of TF in airway epithelial basal cells.
Downstream signaling and procoagulant effects seem to be re-
quired. Because TF, fibrin networks, and PAR activation were
important for basal cell proliferation, we suggest that TF medi-
ates basal cell attachment and survival, leading to proliferation
(Figure 7). We propose, therefore, that TF activity contributes
to repair of mucosal surfaces by promoting extravascular provi-
sional fibrin matrix formation, allowing survival and prolifera-
tion of airway epithelial basal cells.
These data may indicate a new direction for tissue engineer-
ing in the large airways. Recent studies indicated that fibrin gels
and delivery of gene therapy, slowly release and deliver growth
factors, and store and/or deliver cell-based therapies (53). Fibrin
can direct cell migration and homing of circulation-derived cells
and may direct proliferation, differentiation, and gene expres-
sion (54, 55). The current study enhances our understanding of
the potential utility of fibrin in the regeneration or replacement
of large airway epithelium and advances a TF-centered ap-
proach to scaffolds, provisional matrices, and cell-based treat-
ment of large airways.
Author disclosures are available with the text of this article at www.atsjournals.org.
Acknowledgments: The authors thank Dr. Alisa Wolberg for many very useful dis-
cussions and for providing FITC-labeled fibrinogen for the studies and Abhilasha
nical assistance. FLOW cytometry analysis was conducted at the University of Col-
orado Cancer Center Flow Core.
1. Rao LV, Mackman N. Factor VIIa and tissue factor: from cell biology to
animal models. Thromb Res 2010;125:S1–S3.
2. Fan L, Yotov WV, Zhu T, Esmailzadeh L, Joyal JS, Sennlaub F,
Heveker N, Chemtob S, Rivard GE. Tissue factor enhances protease-
activated receptor-2-mediated factor VIIa cell proliferative proper-
ties. J Thromb Haemost 2005;3:1056–1063.
3. Idell S. Coagulation, fibrinolysis, and fibrin deposition in acute lung in-
jury. Crit Care Med 2003;31:S213–S220.
4. Bastarache JA, Wang L, Geiser T, Wang Z, Albertine KH, Matthay MA,
Ware LB. The alveolar epithelium can initiate the extrinsic coagulation
cascade through expression of tissue factor. Thorax 2007;62:608–616.
5. Wygrecka M, Markart P, Fink L, Guenther A, Preissner KT. Raised
protein levels and altered cellular expression of factor VII activating
protease (FSAP) in the lungs of patients with acute respiratory dis-
tress syndrome (ARDS). Thorax 2007;62:880–888.
6. Bastarache JA, Fremont RD, Kropski JA, Bossert FR, Ware LB. Pro-
coagulant alveolar microparticles in the lungs of patients with acute
Figure 7. Roles played by TF in basal cell survival. Schematic represen-
tation of TF-mediated fibrin network formation and PAR activation pro-
viding basal cell survival attachment and proliferation to potentially
restore injured airway epithelium.
102 AMERICAN JOURNAL OF RESPIRATORY CELL AND MOLECULAR BIOLOGYVOL 482013
respiratory distress syndrome. Am J Physiol Lung Cell Mol Physiol
7. Bastarache JA, Wang L, Wang Z, Albertine KH, Matthay MA, Ware
LB. Intra-alveolar tissue factor pathway inhibitor is not sufficient to
block tissue factor procoagulant activity. Am J Physiol Lung Cell Mol
8. Veress LA, O’Neill HC, Hendry-Hofer TB, Loader JE, Rancourt RC,
White CW. Airway obstruction due to bronchial vascular injury after
sulfur mustard analog inhalation. Am J Respir Crit Care Med 2010;
9. Rancourt RC, Veress LA, Guo X, Jones TN, Hendry-Hofer TB, White
CW. Airway tissue factor-dependent coagulation activity in response
to sulfur mustard analog 2-chloroethyl ethyl sulfide. Am J Physiol
Lung Cell Mol Physiol 2011;302:L82–L92.
10. Enkhbaatar P, Murakami K, Cox R, Westphal M, Morita N, Brantley K,
Burke A, Hawkins H, Schmalstieg F, Traber L, et al. Aerosolized
tissue plasminogen inhibitor improves pulmonary function in sheep
with burn and smoke inhalation. Shock 2004;22:70–75.
11. Heath L, Ling S, Racz J, Mane G, Schmidt L, Myers JL, Tsai WC,
Caruthers RL, Hirsch JC, Stringer KA. Prospective, longitudinal
study of plastic bronchitis cast pathology and responsiveness to tissue
plasminogen activator. Pediatr Cardiol 2011;32:1182–1189.
12. Monroe DM, Mackman N, Hoffman M. Wound healing in hemophilia
b mice and low tissue factor mice. Thromb Res 2010;125:S74–S77.
13. Pedersen B, Holscher T, Sato Y, Pawlinski R, Mackman N. A balance
between tissue factor and tissue factor pathway inhibitor is required
for embryonic development and hemostasis in adult mice. Blood 2005;
14. Drake TA, Morrissey JH, Edgington TS. Selective cellular expression of
tissue factor in human tissues: implications for disorders of hemostasis
and thrombosis. Am J Pathol 1989;134:1087–1097.
15. Flossel C, Luther T, Muller M, Albrecht S, Kasper M. Immunohisto-
chemical detection of tissue factor (TF) on paraffin sections of rou-
tinely fixed human tissue. Histochemistry 1994;101:449–453.
16. Hajj R, Baranek T, Le Naour R, Lesimple P, Puchelle E, Coraux C.
Basal cells of the human adult airway surface epithelium retain
transit-amplifying cell properties. Stem Cells 2007;25:139–148.
17. Imokawa S, Sato A, Hayakawa H, Kotani M, Urano T, Takada A. Tissue
factor expression and fibrin deposition in the lungs of patients with
idiopathic pulmonary fibrosis and systemic sclerosis. Am J Respir Crit
Care Med 1997;156:631–636.
18. Hackett TL, Shaheen F, Johnson A, Wadsworth S, Pechkovsky DV,
Jacoby DB, Kicic A, Stick SM, Knight DA. Characterization of side
population cells from human airway epithelium. Stem Cells 2008;26:
19. Perrio MJ, Ewen D, Trevethick MA, Salmon GP, Shute JK. Fibrin
formation by wounded bronchial epithelial cell layers in vitro is es-
sential for normal epithelial repair and independent of plasma pro-
teins. Clin Exp Allergy 2007;37:1688–1700.
20. Ahmad S, Ahmad A, Dremina ES, Sharov VS, Guo X, Jones TN,
Loader JE, Tatreau JR, Perraud AL, Schoneich C, et al. Bcl-2 sup-
presses sarcoplasmic/endoplasmic reticulum Ca21-atpase expression
in cystic fibrosis airways: role in oxidant-mediated cell death. Am J
Respir Crit Care Med 2009;179:816–826.
21. Ahmad S, Nichols DP, Strand M, Rancourt RC, Randell SH, White CW,
Ahmad A. Serca2 regulates non-CF and CF airway epithelial cell
response to ozone. PLoS ONE 2011;6:e27451.
22. Campbell RA, Overmyer KA, Bagnell CR, Wolberg AS. Cellular pro-
coagulant activity dictates clot structure and stability as a function of
distance from the cell surface. Arterioscler Thromb Vasc Biol 2008;28:
23. Ahmad S, Raemy DO, Loader JE, Kailey JM, Neeves KB, White CW,
Ahmad A, Gehr P, Rothen-Rutishauser BM. Interaction and locali-
zation of synthetic nanoparticles in healthy and cystic fibrosis airway
epithelial cells: effect of ozone exposure. J Aerosol Med Pulm Drug
24. Ahmad S, Ahmad A, McConville G, Schneider BK, Allen CB, Manzer R,
Mason RJ, White CW. Lung epithelial cells release ATP during ozone
exposure: signaling for cell survival. Free Radic Biol Med 2005;39:213–226.
25. Ahmad S, Ahmad A, Schneider KB, White CW. Cholesterol interferes
with the MTT assay in human epithelial-like (A549) and endothelial
(HLMVE and HCAE) cells. Int J Toxicol 2006;25:17–23.
26. Dorfleutner A, Hintermann E, Tarui T, Takada Y, Ruf W. Cross-talk of
integrin alpha3beta1 and tissue factor in cell migration. Mol Biol Cell
27. Ahamed J, Ruf W. Protease-activated receptor 2-dependent phosphor-
ylation of the tissue factor cytoplasmic domain. J Biol Chem 2004;279:
28. Key NS, Mackman N. Tissue factor and its measurement in whole blood,
plasma, and microparticles. Semin Thromb Hemost 2010;36:865–875.
29. Kaczmarek E, Lee MH, McDonagh J. Initial interaction between fibrin
and tissue plasminogen activator (t-PA): the gly-pro-arg-pro binding
site on fibrin(ogen) is important for t-PA activity. J Biol Chem 1993;
30. Idell S, Kumar A, Zwieb C, Holiday D, Koenig KB, Johnson AR.
Effects of TGF-beta and TNF-alpha on procoagulant and fibrinolytic
pathways of human tracheal epithelial cells. Am J Physiol 1994;267:
31. Mackman N, Taubman M. Tissue factor: past, present, and future.
Arterioscler Thromb Vasc Biol 2009;29:1986–1988.
32. Ostrowska E, Sokolova E, Reiser G. PAR-2 activation and LPS syner-
gistically enhance inflammatory signaling in airway epithelial cells by
raising PAR expression level and interleukin-8 release. Am J Physiol
Lung Cell Mol Physiol 2007;293:L1208–L1218.
33. Shetty S, Bhandary YP, Shetty SK, Velusamy T, Shetty P, Bdeir K,
Gyetko MR, Cines DB, Idell S, Neuenschwander PF, et al. Induction
of tissue factor by urokinase in lung epithelial cells and in the lungs.
Am J Respir Crit Care Med 2010;181:1355–1366.
34. Boers JE, Ambergen AW, Thunnissen FB. Number and proliferation of
basal and parabasal cells in normal human airway epithelium. Am J
Respir Crit Care Med 1998;157:2000–2006.
35. Evans MJ, Van Winkle LS, Fanucchi MV, Plopper CG. Cellular and
molecular characteristics of basal cells in airway epithelium. Exp
Lung Res 2001;27:401–415.
36. Engelhardt JF, Schlossberg H, Yankaskas JR, Dudus L. Progenitor cells
of the adult human airway involved in submucosal gland develop-
ment. Development 1995;121:2031–2046.
37. Prydz H, Gaudernack G. Studies on the biosynthesis of factor VII
(proconvertin): the mode of action of warfarin. Biochim Biophys Acta
38. van den Hengel LG, Versteeg HH. Tissue factor signaling: a multi-
faceted function in biological processes. Front Biosci 2011;3:1500–
1510. (Schol Ed).
39. Sorensen BB, Rao LV, Tornehave D, Gammeltoft S, Petersen LC. Anti-
apoptotic effect of coagulation factor VIIa. Blood 2003;102:1708–1715.
40. Fang J, Tang H, Xia L, Zhou M, Chen Y, Wei W, Hu Y, Song S, Hong
M. Down-regulation of tissue factor by siRNA increased doxorubicin-
induced apoptosis in human neuroblastoma. J Huazhong Univ Sci
Technolog Med Sci 2008;28:42–45.
41. Boltzen U, Eisenreich A, Antoniak S, Weithaeuser A, Fechner H, Poller
W, Schultheiss HP, Mackman N, Rauch U. Alternatively spliced tis-
sue factor and full-length tissue factor protect cardiomyocytes against
TNF-alpha-induced apoptosis. J Mol Cell Cardiol 2012;52:1056–1065.
42. Pan JJ, Shi HM, Luo XP, Ma D, Li Y, Zhu J, Liang W, Mu JG, Li J.
Recombinant TFPI-2 enhances macrophage apoptosis through up-
regulation of FAS/FASL. Eur J Pharmacol 2011;654:135–141.
43. Ekstrand J, Razuvaev A, Folkersen L, Roy J, Hedin U. Tissue factor
pathway inhibitor-2 is induced by fluid shear stress in vascular smooth
muscle cells and affects cell proliferation and survival. J Vasc Surg
44. Marutsuka K, Hatakeyama K, Sato Y, Yamashita A, Sumiyoshi A, Asada
Y. Protease-activated receptor 2 (PAR2) mediates vascular smooth
muscle cell migration induced by tissue factor/factor VIIa complex.
Thromb Res 2002;107:271–276.
45. Ossovskaya VS, Bunnett NW. Protease-activated receptors: contribution
to physiology and disease. Physiol Rev 2004;84:579–621.
46. Lan RS, Stewart GA, Henry PJ. Role of protease-activated receptors in
airway function: a target for therapeutic intervention? Pharmacol
47. Camerer E, Huang W, Coughlin SR. Tissue factor- and factor X-dependent
activation of protease-activated receptor 2 by factor VIIa. Proc Natl
Acad Sci USA 2000;97:5255–5260.
48. Versteeg HH, Ruf W. Emerging insights in tissue factor-dependent
signaling events. Semin Thromb Hemost 2006;32:24–32.
Ahmad, Ahmad, Rancourt, et al.: Tissue Factor Regulates Basal Cell Survival 103
49. Poulsen LK, Jacobsen N, Sorensen BB, Bergenhem NC, Kelly JD, Foster Download full-text
DC, Thastrup O, Ezban M, Petersen LC. Signal transduction via the
mitogen-activated protein kinase pathway induced by binding of coag-
ulation factor VIIa to tissue factor. J Biol Chem 1998;273:6228–6232.
50. Sen P, Gopalakrishnan R, Kothari H, Keshava S, Clark CA, Esmon CT,
Pendurthi UR, Rao LV. Factor VIIa bound to endothelial cell protein
C receptor activates protease activated receptor-1 and mediates cell
signaling and barrier protection. Blood 2011;117:3199–3208.
51. Khoufache K, LeBouder F, Morello E, Laurent F, Riffault S, Andrade-
Gordon P, Boullier S, Rousset P, Vergnolle N, Riteau B. Protective role
for protease-activated receptor-2 against influenza virus pathogenesis via
an IFN-gamma-dependent pathway. J Immunol 2009;182:7795–7802.
52. Borensztajn K, Bresser P, van der Loos C, Bot I, van den Blink B, den
Bakker MA, Daalhuisen J, Groot AP, Peppelenbosch MP, von der
Thusen JH, et al. Protease-activated receptor-2 induces myofibroblast
differentiation and tissue factor up-regulation during bleomycin-induced
lung injury: otential role in pulmonary fibrosis. Am J Pathol 2010;177:
53. Ahmed TA, Dare EV, Hincke M. Fibrin: a versatile scaffold for tissue
engineering applications. Tissue Eng Part B Rev 2008;14:199–215.
54. de Boer HC, Verseyden C, Ulfman LH, Zwaginga JJ, Bot I, Biessen EA,
Rabelink TJ, van Zonneveld AJ. Fibrin and activated platelets co-
operatively guide stem cells to a vascular injury and promote differ-
entiation towards an endothelial cell phenotype. Arterioscler Thromb
Vasc Biol 2006;26:1653–1659.
55. Langer HF, May AE, Vestweber D, De Boer HC, Hatzopoulos AK,
Gawaz M. Platelet-induced differentiation of endothelial progenitor
cells. Semin Thromb Hemost 2007;33:136–143.
104AMERICAN JOURNAL OF RESPIRATORY CELL AND MOLECULAR BIOLOGYVOL 48 2013