Department of Pediatrics, University of California, San Francisco, San Francisco, California 94143
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ABSTRACT |
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Intestinal muscle undergoes stretch
intermittently during peristalsis and persistently proximal to
obstruction. The influence of this pervasive biomechanical force on
developing smooth muscle cell function remains unknown. We adapted a
novel in vitro system to study whether stretch modulates transforming
growth factor-1 (TGF-
1) and type I collagen
protein and component
1 chain [
1(I) collagen] expression in fetal human intestinal
smooth muscle cells. Primary confluent cells at 20-wk gestation,
cultured on flexible silicone membranes, were subjected to two brief
stretches or to 18 h tonic stretch. Nonstretched cultures served as
controls. TGF-
1 protein was measured by ELISA and type I collagen
protein was assayed by Western blot. TGF-
1 and
1(I) collagen mRNA abundance was
determined by Northern blot analysis, quantitated by phosphorimaging, and normalized to 18S rRNA. Transcription was examined by nuclear run-on assay. Tonic stretch increased TGF-
1 protein 40%, type I
collagen protein 100%, TGF-
1 mRNA content 2.16-fold, and
1(I) collagen mRNA 3.80-fold and
enhanced transcription of TGF-
1 and
1(I) collagen by 3.1- and 4.25-fold,
respectively. Brief stretch stimulated a 50% increase in TGF-
1 mRNA
content but no change in
1(I)
collagen. Neutralizing anti-TGF-
1 ablated stretch-mediated effects
on
1(I) collagen. Therefore, stretch
upregulates transcription for TGF-
1, which stimulates
1(I) collagen gene expression in smooth muscle from developing gut.
intestine; transforming growth factor-; collagen
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INTRODUCTION |
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FORMATION AND MAINTENANCE of the intestinal wall require a specific orientation and proportion of smooth muscle and muscle-derived collagens. Derangements are associated with intestinal stenosis, obstruction, and/or perforation, which significantly increases the morbidity and mortality associated with congenital malformations, injury, or inflammation.
During intestinal organogenesis and postnatal life, types I, III, and V
collagens predominate, with type I collagen 1
[
1(I) collagen] chains being the most
abundant (12, 13, 31). Each collagen type and chain possesses unique
biochemical and biomechanical properties that influence not only smooth
muscle cells but inflammatory cells and epithelial cells as well. The
matrix exerts effects on cell attachment (26, 28), migration (2, 8, 21, 24, 25, 26, 28, 43), proliferation (7), differentiation (2, 10), and
storage of growth factors (5, 10). Each collagen type possesses
different tensile strength. Therefore, the three collagens together
serve as regulatory signals as well as provide a scaffold for
intestinal tissue remodeling and muscle contraction. We have previously
shown that collagen synthesis in human fetal intestinal smooth muscle
cells approximates mature patterns by 20-wk gestation, coinciding with
establishment of mature histological architecture (31).
Transforming growth factor (TGF)-1 is a key mediator regulating
collagen deposition during gut morphogenesis, growth, and wound
healing. TGF-
1 is a homologous secreted disulfide-bonded dimer that
mediates cellular growth, matrix production, and cell differentiation
(3, 37). TGF-
1 immunolocalizes to all epithelial and mesenchymal
components of the intestine, excluding the crypts, and it is secreted
by human intestinal smooth muscle cells throughout fetal development
and postnatal life (20, 32). Its targeted gene disruption
alters gut wall integrity (42). In cultures of differentiated human
intestinal smooth muscle cells, fetal or adult, TGF-
1 stimulates
collagen synthesis (11, 32). In many tissues, TGF-
1 induces collagen
accumulation by upregulating collagen gene expression (33, 38, 44),
enhancing production of protease inhibitors and suppression of
proteolytic enzymes (4, 29).
Although much is known regarding regulatory cytokines and matrices
influencing muscle cell behavior during intestinal tissue remodeling,
no prior study accounts for the presence of biomechanical forces on
muscle in gut. There is mounting evidence that biomechanical forces act
in conjunction with diffusible cytokines and local matrix to modulate
cell behavior. Intestinal muscle undergoes stretch intermittently
during peristalsis and persistently proximal to obstruction. The
current study is the first to demonstrate that stretch participates in
regulating TGF-1 and collagen gene expression in fetal human
intestinal smooth muscle cells.
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METHODS |
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Cell isolation and culture. Intestine was obtained from 20-wk abortuses as described previously (31). The protocol for this research was approved by the Committee for Human Research at the University of California at San Francisco according to the policies of the National Institutes of Health.
Muscle cells for culture were harvested from human fetal intestine by dissection and collagenase digestion as described previously (31). Cultures were maintained in DMEM containing 25 mmol/l tricine (pH 7.4), 10% FBS, 100 U/ml penicillin, and 100 µg/ml streptomycin (GIBCO BRL, Gaithersburg, MD) in 37°C incubators with circulating room air. After establishment of primary cultures, cells were passaged once in 0.1% trypsin within 1-3 days after isolation from the tissue. These cultured cells have been shown to possess the same phenotype and synthetic, proliferative, and contractile functions of freshly isolated cells (31).Mechanical distension of cultured cells. Cells (8 × 105) were cultured onto silicone membranes precoated with fibronectin and grown to confluence. Twenty-four hours were allowed for cell attachment before applying stretch stimuli. For the purposes of these studies, the term "stretch" refers to mechanical distension of cultures. Briefly, membranes were bracketed within an acrylic frame overlying a fluid-filled chamber. Cultures were distended to stretch cells 20% by applying hydrostatic pressure beneath the membranes as previously described (14). Cultures undergoing brief stretch were distended for 30 s, relaxed 30 s, distended again for 30 s, and relaxed for 4 h before harvest for mRNA. Cultures undergoing tonic stretch were distended for 18 h. Maintenance of distension was assessed by evaluating the devices for leaks as well as the amount of fluid removal required to return the membranes to their relaxed state.
Cell viability and cytotoxicity. Cell viability and cytotoxicity were determined by measuring intracellular esterase activity with calcein AM and assaying plasma membrane integrity with ethidium homodimer (LIVE/DEAD viability/cytotoxicity kit; Molecular Probes, Eugene, OR). Nonstretched cultures served as controls. Values are expressed as percent of live cells and represent the means ± SD (n = 6).
ELISA for TGF-1 protein.
Both soluble and latent TGF-
1 protein were measured by enzyme-linked
immunosorbent assay (R & D Systems, Minneapolis, MN). Aliquots (1 ml)
of serum-free conditioned medium were mixed with 0.2 ml of 1 N HCL,
incubated at 25°C for 10 min, and neutralized by 1.2 N NaOH and 0.5 mol/l HEPES. TGF-
1 protein was measured by ELISA using an antibody
raised in chickens that reacts fully with human TGF-
1. The limit of
detection is 5 pg/ml with a linear range response over the range of
30-2,000 pg/ml.
Western blot determination of type I collagen protein. Cells were harvested in PBS with protease inhibitors (CalBiochem, San Diego, CA). Type I collagen protein was measured by the bicinchoninic acid method (Pierce, Rockford, IL). Samples (10 µg) and prestained molecular weight standards (Bio-Rad, Hercules, CA) were electrophoresed through a 4% acrylamide stacking gel and subsequent 10% SDS-polyacrylamide gel in the presence of 10% dithiothreitol (DTT). Proteins were electrophoretically transferred to nitrocellulose paper. Gel protein transfer was confirmed by absent Coomassie blue staining. Western blots were blocked 2 h in 1% nonfat dried milk, 0.4% gelatin, and 0.1% BSA in 150 mM NaCl/10 mM Tris (pH 7.2) and incubated 1 h in 20 mM Tris-buffered saline (TBS), pH 7.4, containing 5 µg/ml of a monoclonal antibody against human type I collagen (Oncogene Research Products, Cambridge, MA). After 20 washes with TBS, the blots were incubated for 30 min in a 1:10,000 dilution of horseradish peroxidase-labeled affinity-purified sheep anti-mouse IgG (Cappel, ICN Pharmaceuticals, Aurora, OH). Blots were washed again before incubation in luminol (Amersham) for 1 min. After autoradiography, protein was quantitated by densitometry.
Preparation of RNA, Northern blotting, and hybridization.
After 18 h of tonic distension or 4 h after two brief stretches, tissue
culture medium was aspirated and membranes were washed twice with
sterile PBS at 4°C. Total cellular RNA was extracted directly from
cultured cells on membranes using RNA-STAT (Tel-Test, Friendswood, TX)
and was quantitated spectrophotometrically. RNA integrity was assessed
by electrophoresis and ethidium bromide staining for rRNA. Total RNA
(10 µg/sample) was separated electrophoretically on 1% agarose gels.
Total RNA was transferred to nylon membranes under positive pressure
(Stratagene Posiblotter; Stratagene, La Jolla, CA) and cross-linked
with ultraviolet light (UV Stratalinker 2400; Stratagene). Filters were
probed with full-length cDNAs for human TGF-1 (generous gift of Dr.
Stephan E. Gitelman, University of California at San Francisco),
1(I) collagen (generous gift of Dr.
David A. Brenner and Dr. Richard A. Rippe, University of North Carolina
at Chapel Hill), and 18S rRNA and labeled with [
-32P]dCTP
(DuPont NEN, Boston, MA) by random primer second-strand synthesis
(random primer labeling kit; GIBCO BRL). Filters were prehybridized for
10 min in QuikHyb hybridization solution (Stratagene) at 68°C.
Filters were hybridized in 10 ml of QuikHyb solution containing 1.25 × 106 dpm/ml
for 18 h at 68°C. Hybridized filters were washed three times for 15 min at room temperature in a solution of
2× standard saline citrate (SSC) and 0.1% SDS (Sigma, St. Louis,
MO) and then once for 30 min at 60°C in a solution of 0.1×
SSC and 0.1% SDS. Filters were then subjected to autoradiography
(Hyperfilm, Amersham) before radiolabeled bands were quantified by
volume integration of pixels measured by phosphorimage analysis
(Imagequant software, Molecular Dynamics, Sunnyvale, CA). Equal loading
was ensured by using 18S rRNA as control.
Nuclear run-on transcription assay.
The assay was performed as previously described (14). Briefly, nuclei
were prepared from 12 stretched and 12 control membranes by
homogenization and centrifugation. A number of nuclei
(12 × 106) were
resuspended in reaction buffer containing 10 mM
Tris · HCl, 5 mM
MgCl2, 300 mM KCl (Sigma), 10 mM
ATP, 10 mM CTP, 10 mM GTP (Boerhinger Mannheim, Indianapolis, IN), 10 mM DTT (Promega, Madison, WI), 10 units RNasin (GIBCO BRL), and 200 µCi
[-32P]UTP
(DuPont NEN) and incubated at 37°C for 20 min. The nuclei were then
successively digested with RNase-free DNase I and proteinase K. Unincorporated nucleotides were removed using a Centricon 100 concentrator (Amicon, Beverly, MA).
Neutralizing antibody.
To determine if stretch-induced effects on
1(I) collagen were mediated through
TGF-
1, experiments were repeated in the presence and absence of a
neutralizing antibody, anti-TGF-
1 (Becton Dickinson, Bedford, MA).
After 24 h to permit cell attachment, cultures were washed three times
with serum-free medium and then exposed to antibody or vehicle for 18 h. A concentration of 200 µg/ml was used and has previously been
shown to inhibit the amount of TGF-
1 secreted by these cells at
20-wk gestation (32). Stretched and nonstretched cultures with vehicle
alone (0.1 M
NaH2PO4,
pH 5.5, and 1% BSA) served as controls.
Statistical analysis. Results are expressed as the percent of change from nonstretched controls and represent the means ± SD. Stretched cultures were compared with nonstretched controls by Student's t-test. P < 0.05 was considered significant.
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RESULTS |
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Stretch is not cytotoxic. A tonic stretch of 18 h produced no changes in cell phenotype or adherence by visual inspection and no changes in intracellular esterase or membrane integrity. Tonically stretched cultures demonstrated 98.5 ± 1.9% viability (P < 0.30) compared with nonstretched controls (99.25 ± 1.0% viability; P < 0.26).
Tonic stretch increases TGF-1 and type I collagen
protein.
Tonic stretch increased TGF-
1 protein 140 ± 9.3% (n = 5, P < 0.05; Fig.
1) and type I collagen protein 200 ± 25% (n = 6, P < 0.05; Fig.
2).
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Tonic stretch increases TGF-1 and
1(I) collagen mRNA abundance.
Tonic stretch increased mRNA content of TGF-
1 by 216 ± 7.6% (P < 0.005; Fig.
3) and
1(I) collagen by 380 ± 32%
(P < 0.05; Fig.
4) compared with nonstretched controls.
There was no change in the mRNA content of 18S rRNA (Figs. 3 and 4).
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Stretch enhances transcription of TGF-1 and
1(I) collagen.
Nuclear run-on assays were performed to determine whether tonic stretch
changes mRNA content at transcriptional or posttranscriptional levels.
Tonic stretch increased the transcription of TGF-
1 by 310 ± 34%
(P < 0.005) and of
1(I) collagen by 425 ± 85%
(P < 0.05). Transcription of 18S
rRNA did not change (Fig. 5).
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Brief stretch increases TGF-1 but not
1(I) collagen mRNA content.
To determine whether stretch induced TGF-
1 expression before that of
1(I) collagen, studies were done
showing that brief stretch increased the mRNA content of TGF-
1 by 50 ± 10% without changing mRNA content of
1(I) collagen or 18S rRNA
(Fig. 6).
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Tonic stretch increases 1(I) collagen mRNA content
through TGF-
1.
To determine whether the increase in mRNA content of
1(I) collagen observed after 18-h
tonic stretch was mediated through an earlier increase in TGF-
1, the
relationship between TGF-
1 and collagen was evaluated. Exogenous
TGF-
1 increased
1(I) collagen mRNA in a dose-dependent manner (Fig.
7). Anti-TGF-
1 ablated the
stretch-induced increase in
1(I)
collagen mRNA abundance (Fig. 8).
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DISCUSSION |
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These studies are the first to demonstrate that stretch modulates
smooth muscle gene expression fundamental to intestinal development and
remodeling. Brief stretch increased TGF-1 but not
1(I) collagen expression. Tonic
stretch enhanced both TGF-
1 and
1(I) collagen protein and mRNA
abundance, in part through increased transcription. Anti-TGF-
1
abolished stretch-mediated increases in
1(I) collagen mRNA content,
suggesting that stretch initially increases TGF-
1, which, in turn,
stimulates collagen expression. These findings were obtained through
the novel adaptation of a hydrostatic pressure device to produce
distension in primary cultures of human fetal intestinal smooth muscle cells.
Prior knowledge that biomechanical force influences smooth muscle cell synthetic function is largely based on cells and stretch patterns derived from and relevant to vasculature. Stretch-mediated collagen expression was shown to vary with the degree of stretch, number of stretches per unit time, and anatomic origin of the vessel or cultured smooth muscle cells (6, 18, 22). However, those experiments were designed to examine stretch forces mimicking pulsatile pressures (e.g., 52 cycles/min), normal transmural pressure in different vessels (e.g., 120/80 mmHg in conduit vessels, 40/30 mmHg in precapillary vessels), sustained hypertension, or shear stress (i.e., the friction of flowing blood). The applicability to intestinal smooth muscle is limited due to the organ specificity of smooth muscle cell collagen synthetic response to a given stimulus (30) and the type of stretch. In contrast to blood vessels, the small intestine undergoes different types and patterns of stretch that vary with the fed or fasting state and anatomic location. The only study to address stretch-mediated cell function in gut used a transformed epithelial cell line subjected to mature patterns of distension (1). The current study examines primary cultures of fetal human intestinal smooth muscle cells. This is the only existing in vitro model to study species-specific and developmentally specific aspects of the biology of these cells. Furthermore, the precise patterns of developing intestinal motility and stretch have yet to be elucidated. Therefore, it is not currently feasible to simulate stretch patterns of fetal human intestine in vitro. Our approach characterizes the developing muscle cell response to specific stretch stimuli as an initial means to determine how changing stretch force may modulate collagen synthesis in these fetal cells.
We demonstrate that fetal human intestinal smooth muscle cells respond
to stretch by increasing TGF-1 and
1(I) collagen expression. Other
studies have been corroborative but utilized different models. Cultured
rat mesangial cells demonstrated stretch-stimulated TGF-
1 synthesis,
release, and activation (35). The increased TGF-
1 may underlie the
stretch-enhanced type I collagen accumulation in these cells (36) but
was not specifically addressed. In those studies, TGF-
1 was measured
after 48- and 72-h cyclic stretch. Our studies uniquely examine brief
and sustained stretch stimuli and smooth muscle cell responses that are
specifically fetal, human, and intestinal. Use of an anti-TGF-
1
antibody in this system ablated stretch-mediated increases in
1(I) collagen expression. Therefore,
stretch-induced increases in collagen are mediated through TGF-
1 in
fetal human intestinal smooth muscle cells.
Potential mechanisms whereby stretch regulates soluble cytokines and
local matrix in intestinal smooth muscle are suggested by studies
documenting alterations of signaling pathways and gene expression in
response to single stretch, changes in cell shape, or shear stress.
Maniotis et al. (23) have demonstrated that the extracellular and
intracellular environments are physically linked through the
extracellular matrix, cell surface integrins, cytoskeletal filaments,
chromosomes, and nucleoli. These structures appear to transduce
information from the cell membrane to the nucleus as well as provide
physical support. In cardiac myocytes, a single stretch has been shown
to activate specific signal transduction pathways (40) and to induce
certain immediate-early genes (19, 40, 41). Kheradmand et al. (16)
demonstrated that changes in cell shape can induce collagenase 1 gene
expression. Signaling pathways modulate expression of mechanosensitive
genes through cis-active elements in
the promotor region. The promotor region of TGF-1 contains a
sequence now known to be a shear stress response element (17, 34). In
the current study, tonic stretch increases TGF-
1 transcription in
human fetal intestinal muscle cells. A similar increase in
transcription occurs in vascular endothelial cells subjected to shear
stress (27). Whether stretch-induced changes in muscle transcription
are mediated through the same or similar mechanosensitive elements
within the promotor region remains to be elucidated.
That intestinal smooth muscle cells respond to mechanical stimuli with
altered synthesis of TGF-1 and
1(I) collagen poses significant
ramifications for normal or abnormal gut development. The formation,
integrity, and compliance of the gut wall largely reflect the
proportion and distribution of muscle mass and collagen. TGF-
1
inhibits the proliferation of intestinal smooth muscle cells and
stimulates or inhibits collagen synthesis is a manner that is specific
to the collagen type and stage of fetal development (32). The resulting
collagenous matrix, in turn, influences the migration, proliferation,
and attachment of muscle cells, epithelial cells, and immune cells,
behaviors fundamental to organogenesis, tissue growth, and repair. In
other developing tubes, such as lung and aorta, inadequate distension
results in hypoplasia (15, 39). In gut, sustained distension may occur
proximal to obstruction arising from atresias, volvulous, malrotation,
or aganglionosis. Intestinal obstruction has been associated with
muscle hypertrophy and collagen accumulation, resulting in wall
thickening (9). Fetal human intestinal smooth muscle cells respond to
stretch by altering expression of mRNAs known to affect matrix
deposition, cell proliferation, and cell differentiation. Therefore, it
is possible that the development of normal peristalsis influences subsequent muscle development and that aberrations in patterns of
stretch, such as obstruction, may interfere with subsequent formation
of the intestinal wall.
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ACKNOWLEDGEMENTS |
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We thank Wen Zhou and Karen Matsukuma for technical assistance and Dr. Donna Ferriero for encouragement and editorial review.
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FOOTNOTES |
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This work was funded by the Research Evaluation and Allocation Committee Springer Award and Robert Wood Johnson grant RWJ 030805. Both authors contributed equally to this work.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: H. A. Perr, Dept. of Pediatrics, Box 0136, 500 Parnassus Ave., Univ. of California, San Francisco, San Francisco, California 94143-0136 (E-mail: hperr{at}itsa.ucsf.edu).
Received 15 September 1998; accepted in final form 26 July 1999.
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