Departments of 1 Surgery, 4 Medicine, 5 Biochemistry and Biophysics, and 3 Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina 27599-7210; and 2 Bayer Pharmaceuticals, Wupertal 20000, Germany
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ABSTRACT |
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The aim of this study was to determine the differential effects of
latent and activated transforming growth factor (TGF)-1 in growth control of normal and proliferating hepatocytes in vivo. Rats
were injected with adenoviruses expressing control transgenes (Ctrl),
latent TGF-
1 [TGF-
(L)], or activated
TGF-
1 [TGF-
(A)]. Additional animals underwent
two-thirds partial hepatectomy (PH) 24 h after injection.
Increased hepatocyte apoptosis was observed in TGF-
(A)-injected but
not TGF-
(L)-injected animals 24 h postinjection (10.5%)
compared with Ctrl animals (0.37%). The percent of apoptotic cells
increased to 32.1% in TGF-
(A)-injected animals 48 h after injection. Furthermore, TGF-
(A)-injected rats did not survive 24 h after PH. Four hours after PH, 0.25 and 14.1% apoptotic
hepatocytes were seen in Ctrl- and TGF-
(A)-injected rats,
respectively. TGF-
(A)-induced apoptosis in primary rat hepatocytes
was blocked with a pancaspase inhibitor. Thus autocrine expression of
TGF-
(A) but not TGF-
(L) induces hepatocyte apoptosis in the
normal rat liver. Rats overexpressing TGF-
(A) do not survive
two-thirds PH due to hepatic apoptosis. Thus activation of
TGF-
1 may be a critical step in the growth control of
normal and proliferating rat hepatocytes.
hepatic regeneration; caspase activity; growth factors; adenovirus; partial hepatectomy
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INTRODUCTION |
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IN THE HEALTHY
LIVER hepatocytes divide rarely, but after chemical or physical
injury hepatocytes progress from the G0 phase to the
G1 phase of the cell cycle. Growth factors regulate this process through both stimulatory and inhibitory signaling. Epidermal growth factor (31), transforming growth factor (TGF)-
(32), and hepatocyte growth factor (34, 53)
stimulate hepatocyte DNA synthesis, whereas TGF-
1 and
interleukin-1
inhibit hepatocyte replication (16).
TGF-
controls hepatocyte growth after partial hepatectomy (PH)
through an autocrine feedback mechanism (5). TGF-
has
been investigated as a growth inhibitor of both hepatocytes in culture
and in the regenerating liver, but the mechanism of growth control and
the importance of relative abundance of activated [TGF-
(A)] vs.
latent [TGF-
(L)] TGF-
in normal and proliferating hepatocytes
has not been examined thoroughly.
TGF-(A) inhibits hepatocyte DNA synthesis both in culture (35,
48) and in vivo (40). In primary hepatocyte
cultures, TGF-
1 inhibits DNA synthesis from normal and
regenerating livers by blocking the transition from the G1
to the S phase of the cell cycle (35, 48). After a
two-thirds PH, TGF-
1 mRNA expression increases
(27), and TGF-
is the ostensible inhibitory peptide for
hepatocyte replication and liver regeneration. Other in vivo studies,
however, have demonstrated that mature TGF-
administered intravenously reduced [3H]thymidine incorporation in
hepatocytes after a PH, but inhibition of hepatocyte DNA synthesis was
transient because complete regeneration of the liver still occurred by
8 days (40). In a transgenic mouse model overexpressing
bioactive TGF-
1, administration of the
latency-associated peptide (LAP) region of TGF-
1
prevented inhibition of hepatocyte DNA synthesis (6).
Both cell cycle arrest and apoptosis have been implicated as mechanisms
of TGF--induced hepatocyte growth arrest. TGF-
has been shown to
inhibit progression of the cell cycle by decreasing the expression of
G1 cyclins and cyclin-dependent kinases (18). In rat liver epithelial cell lines, TGF-
impairs DNA synthesis by
preventing hyperphosphorylation of the retinoblastoma gene product
(pRb; see Ref. 50). Moreover, in vivo administration of
TGF-
in the regenerating rat liver decreased pRb expression, suggesting that this growth-inhibiting cytokine played a prominent role
in growth control of proliferating hepatocytes (15).
Inhibition of hepatocyte proliferation, however, has also been
attributed to apoptosis (37, 38). Only the bioactive form
of TGF-
1, and not the latent form, induces apoptosis
(37). However, this induction of apoptosis by TGF-
(A)
in vivo was shown only in liver undergoing cyproterone-induced
hyperplasia (36) or in diseased liver (42).
Therefore, the in vivo mechanism of hepatocyte growth control in the
normal and regenerating liver remains to be determined.
TGF-1 is translated as a latent dimeric precursor, and
upon cleavage and dissociation of the amino terminal portion, known as
LAP, a mature, biologically active protein is formed (19). TGF-
1 is secreted by hepatocytes in the latent form and
is activated extracellularly by a poorly understood mechanism(s).
Potential in vivo mechanisms of TGF-
activation include plasmin
(29), thrombospondin (TSP)-1 (44, 45),
binding to the mannose 6-phosphate/insulin-like growth factor type II
receptor (14), and the integrin
v
6 (33), but these
activation pathways may be tissue specific. Expression of
TGF-
1 is regulated presumably through an autocrine
feedback mechanism (5). TGF-
signals through two active
receptor subtypes, TGF-
R1 and TGF-
R2. TGF-
binds to TGF-
R2,
which phosphorylates and activates TGF-
R1. TGF-
R1 acts as a
serine-threonine kinase that activates the intracellular Smad family of
signal transducers (23, 51).
Our study investigated the differential effect of autocrine expression
of TGF-(L) and TGF-
(A), delivered by adenoviral vectors, on
hepatocyte growth control in the normal and proliferating hepatocytes in vivo. In this study, TGF-
(A) induced hepatocyte apoptosis in the
normal and regenerating liver, and overexpression of bioactive TGF-
1 was lethal to rats subjected to PH. Additional
experiments showed that TGF-
(A)-induced apoptosis in primary rat
hepatocytes can be inhibited with a pancaspase inhibitor, suggesting
that hepatocyte apoptosis and not cell cycle arrest is the major
mechanism of TGF-
1-induced hepatocyte growth control.
TGF-
(L), however, did not alter hepatocyte DNA synthesis or induce
apoptosis. These findings suggest that activation of
TGF-
1 is a critical step in rat hepatocyte growth
regulation after PH.
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MATERIALS AND METHODS |
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Animals. Adult male Sprague-Dawley rats (200 g) were fed a standard chow diet, had access to water ad libitum, and were kept under 12:12-h light/dark cycles. All animal procedures and protocols were in accordance with the National Institutes of Health guidelines and were approved by The University of North Carolina's Institutional Animal Care and Use Committee.
Purification of adenovirus.
The recombinant replication-deficient adenoviruses [adenoviruses
expressing -galactoside or luciferase (Ctrl), TGF-
(L), or
TGF-
(A)] were grown in 293 cells and were purified by CsCl gradients (52). TGF-
(L) adenovirus contains a 2.1-kb
human TGF-
1 transgene and is converted to the bioactive
TGF-
1 as demonstrated by the mink lung cell growth
inhibition assay (49). TGF-
(A) contained the
full-length porcine TGF-
cDNA with cysteine-to-serine mutations at
positions 223 and 225, resulting in expression of biologically active
TGF-
1 (46). Viral titers were determined by
optical density and plaque assay (30). The recombinant
virus was stored in 25% (vol/vol) glycerol at
80°C.
Adenoviral infection and PH.
Adenoviruses were dialyzed against three changes of 1× PBS containing
1 mM MgCl2 at 4°C. The adenoviral vectors were
injected via the tail vein [1 × 1010
plaque-forming units (PFU)] or intraportally (6.0 × 109 PFU) into 200-g adult male Sprague-Dawley rats.
Twenty-four hours after adenoviral injection, a two-thirds PH was
performed (25) under ketamine/acepromazine anesthesia. The
remnant livers were harvested 4 or 24 h after PH. Livers were
snap-frozen in liquid nitrogen or fixed in 10% neutral buffered
formalin for histological analysis. Additional rats received TGF-(A)
but did not undergo PH, and livers were harvested 6, 12, 18, 24, and
48 h after injection.
Infection of primary rat hepatocytes.
Primary rat hepatocytes were isolated from adult male Sprague-Dawley
rats by collagenase perfusion (22). Hepatocytes were plated in 60-mm tissue culture dishes (1.5 × 106
cells/plate) or 100-mm dishes (4 × 106 cells/plate)
and were cultured in Waymouth's media containing 10% FCS, 5 µg/ml
insulin, and 107 M dexamethasone in a 5%
CO2-95% air atmosphere for 4 h. Cells were serum
starved in hormonally defined medium (HDM; RPMI 1640 containing 5 µg/ml insulin, 10 µg/ml transferrin, 3 × 10
8 M
selenium, and 10 nM free fatty acids) for 2 h. Cells were infected with either Ctrl or TGF-
(A) adenoviruses at a multiplicity of infection of 30 for 2 h. Media was changed to fresh HDM with or without 5 µM Z-VAD.FMK (Calbiochem, La Jolla, CA). Cells were harvested at 18, 24, and 36 h for total RNA to determine
TGF-
(A) expression, fluorescence-activated cell sorter (FACS) and
DNA isolation for apoptotic analysis, and whole cell extracts for caspase activity.
RNA isolation, Northern blot analysis, and -galactosidase and
luciferase assays.
Total RNA from the snap-frozen liver was isolated by CsCl
ultracentrifugation (41). RNA samples (10 µg) were run
on a formaldehyde gel (41) and transferred to a magna
charge nitrocellulose membrane (MSI, Westborough, MA) by capillary
transfer overnight. RNA was fixed to the membranes by ultraviolet
cross-linking using a Stratalinker (Stratagene, La Jolla, CA) and was
prehybridized [5× saline-sodium citrate (SSC), 5× Denhardt's, 50 mM
sodium phosphate, pH 6.5, 0.1% SDS, 250 µg/ml fish sperm DNA, and
50% formamide] for at least 2 h at 42°C. Human
TGF-
1 cDNA was labeled using the Rediprime Kit
(Amersham, Arlington Heights, IL) as recommended by the manufacturer, 106
counts · min
1 · ml
1 was
added to the prehybridization buffer, and incubation was continued for
15 h at 42°C. Blots were washed with 2× SSC and 0.1% SDS two
times at room temperature for 5 min and then with 0.1× SSC and 0.1%
SDS at 65°C for 20 min. Membranes were exposed to Biomax-MR film
(Eastman Kodak, Rochester, NY) overnight at
80°C with an
intensifying screen. Additionally, total RNA was isolated from
hepatocytes by the phenol-guanidinium method (39) and was
analyzed for TGF-
(A) expression as described above.
RNase protection assay.
Total RNA was harvested from liver. Radiolabeled riboprobes for the
RNase protection assay were derived from the 375-nt Pst I-Ava I fragment of the rat 1(I) collagen
cDNA. The riboprobe for the rat glyceraldehyde-3-phosphate
dehydrogenase (GAPDH) gene was generated from the plasmid
pTRI-GAPDH-Rat (Ambion, Austin, TX), which was linearized using
Hind III. The radiolabeled probes were mixed with 50 µg of
total liver RNA, and the samples were dried. Dried pellets were
suspended in 30 µl hybridization buffer (100 mM PIPES, pH 6.7, 400 mM
NaCl, 2 mM EDTA, and 80% formamide), heated at 85°C for 10 min, and
incubated overnight at 45°C. The hybridization reaction was then
incubated with 350 µl RNase buffer (300 mM NaCl, 10 mM
Tris · HCl, pH 7.6, 40 mg/ml RNase A, and 2 mg/ml RNase
T1) at 30°C for 1 h. Afterwards, 10 µl of 20% SDS and 5 µl of 10 mg/ml proteinase K were added, and the reaction mixture was incubated at 37°C for 15 min. The reaction mixture was
phenol extracted and precipitated with the addition of 1 µg of yeast
tRNA and 1.0 ml of 100% ethanol. The samples were suspended in
formamide dye (95% formamide, 20 mM EDTA, 0.05% bromphenol blue, and
0.05% xylene cyanol FF) and heated at 95°C for 5 min and were loaded
onto a standard 5% sequencing gel. After electrophoresis, bands were
visualized by autoradiography and quantitated by PhosphorImager analysis (Molecular Dynamics, Sunnyvale, CA).
Histology and immunohistochemistry. Paraffin-embedded tissues were sectioned at 5 µm thickness and stained with hematoxylin and eosin (H&E) for identification of necrosis and inflammation. For immunohistochemistry, paraffin sections were deparaffinized by three 5-min incubations in xylene followed by 5-min incubations in 100, 95, and 70% ethanol and rehydrated in water and PBS. The terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) assay was performed using the in situ cell death detection kit (Boehringer Mannheim, Indianapolis, IN). Briefly, DNA ends were tagged with fluorescein-labeled dUTP using terminal deoxynucleotidyl transferase by incubating the samples at 37°C in a humidified chamber. Liver sections were then incubated with anti-fluorescein-alkaline phosohatase (AP) conjugate for 30 min in a humidified chamber. Slides were incubated with 5-bromo-4-chloro-3-indolyl phosphate/nitro blue tetrazolium (Sigma, St. Louis, MO) for 20 min at room temperature and were counterstained with hematoxylin. AP-positive cells were counted in five high-power fields (×400) per section, and the number of stained hepatocytes was expressed as a percentage of the total number of hepatocytes.
Caspase assay. Caspase 3-like and caspase 8-like activities were determined by measuring the in vitro fluorogenic peptide substrate carbobenzoxy-Asp-Glu-Val-Asp-7-amino-4-trifluoromethyl coumarin and carbobenzoxy-Ile-Glu-Thr-Asp-7-amino-4-trifluoromethyl coumarin, respectively, as described by the manufacturer (Bio-Rad Laboratories, Hercules, CA). Caspases enzymatically cleave the substrate and release free amino-4-trifluoromethylcoumarin that produces a blue-green fluorescence, which is measured fluorometrically. Liver samples were homogenized in 100 µl of lysis buffer (10 mM HEPES, pH 7.4, 2 mM EDTA, 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate, 5 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml pepstatin A, 10 µg/ml aprotinin, and 20 µg/ml leupeptin), and hepatocytes were lysed in 50 µl of lysis buffer. Liver and cell lysates were prepared by four to five freeze and thaw cycles. Samples were centrifuged at 16,000 g in a microfuge for 10 min at 4°C. Supernatants (10 µl for hepatocyte extracts and 30 µl for liver extracts) were assayed for caspase activity by measuring fluorescence using a Perkin-Elmer luminescence spectrometer LS50B (Perkin-Elmer, Norwalk, CT) and normalized to protein using the Bradford assay (Bio-Rad).
Flow cytometric analysis.
Primary rat hepatocytes (4.0 × 106 cells/plate) were
plated in 100-mm tissue culture dishes. Cells were allowed to adhere to the plates in growth medium as described above. After 4 h of
incubation, the cells were serum starved in HDM for 2 h and then
were infected for 2 h with Ctrl or TGF-(A) adenoviruses. Cells
were trypsinized and collected by centrifugation at 1,000 rpm for 5 min. Cell pellets were suspended in 300 µl of PBS, and 5 ml ice-cold
75% ethanol was added dropwise under gentle agitation to fix the
cells. The cells were washed two times with PBS and were treated with
50 µg/ml RNase A in 1.12% sodium citrate at 37°C for 30 min. Cells were stained with 50 µg/ml propidium iodide in 1.12% sodium citrate overnight at 4°C, and samples were filtered through nitex membrane 3-60/45 (Sefar, Kansas City, MO). Samples were analyzed using a
Becton-Dickinson Flow Cytometer (Becton-Dickinson, Fullerton, CA).
Additionally, hepatocytes plated on 60-mm dishes were fixed with
methanol-acetic acid (3:1) for 10 min at 4°C, washed two times with
H2O, and dried. Cells were stained with propidium iodide (200 ng/µl) and were analyzed by ultraviolet fluorescence.
DNA ladder analysis.
DNA was isolated from hepatocytes of Ctrl or TGF-(A)-infected cells
36 h after infection in the absence or presence of Z-VAD.FMK. Cells were scraped and suspended in 1 ml of HDM and lysed with 200 µl
of 5× lysis buffer (2 M NaCl, 50 mM Tris · HCl, and 10 mM
EDTA, pH 8.0), 50 µl 20% SDS, and 20 µg/ml proteinase K. Samples were incubated at 55°C for 45 min followed by the addition of 400 µl of 5 M NaCl. Samples were agitated well and centrifuged for 30 min
at 3,500 rpm. The clear supernatant was removed, and 5 ml of 96%
ethanol were added to precipitate the DNA. DNA was removed with a
long-hooked Pasteur pipette and was washed in 1 ml of 70% ethanol.
Samples were centrifuged at 16,000 g for 10 min at room
temperature. The DNA was dried and suspended in 40 µl of Tris-EDTA
containing 20 µg/ml RNase A and was incubated for 1 h at 37°C.
Samples (50 µg) were run in a 1.5% agarose gel in 1× 0.04 M
Tris-acetate-0.001 M EDTA running buffer.
Liver function analysis.
Blood was obtained from animals at the time of death by direct inferior
vena cava puncture and was serum stored at 80°C. The University of
North Carolina Division of Laboratory Animal Medicine measured total
bilirubin and alanine aminotransferase (ALT) concentrations from the
serum, and growth media from infected hepatocytes with or without
Z-VAD.FMK were also analyzed for lactate dehydrogenase (LDH) concentrations.
TGF- bioassay.
A stably transfected fibroblast cell line harboring a TGF-
response
element controlling the expression of a luciferase reporter gene was
seeded at a density of 2,000 cells/well in a 384-well microtiter plate.
The medium consisted of DMEM plus 10% FCS (GIBCO-BRL), 2% (vol/vol)
HEPES (GIBCO-BRL), and 1% (vol/vol) gentamicin (GIBCO-BRL). The total
incubation volume was 1,000 µl/well. Cells were incubated for 1 day
at 37°C and 5% CO2.
Data analysis. Data are presented as means ± SE.
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RESULTS |
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In vivo expression of TGF-1.
After injection of the adenoviruses, expression of the transgenes
was confirmed by
-galactosidase or luciferase activity or Northern
blot analysis for TGF-
1 mRNA. Expression of
-galactosidase or luciferase was demonstrated in all animals by
increased enzyme activity in Ctrl-injected rats compared with
TGF-
1-injected rats (data not shown). Twenty-four hours
after injection, without PH, low levels of TGF-
1 mRNA
were observed at 6, 12, and 18 h in TGF-
(A) animals, but a
dramatic increase was seen at 24 and 48 h in the rats injected
with TGF-
(L) or TGF-
(A) (Fig.
1A). Translation of activated
TGF-
protein was confirmed by TGF-
bioassay, which demonstrated
an 8- to 10-fold increase in activated TGF-
in TGF-
(A)-injected rats compared with Ctrl- and TGF-
(L)-injected rats (Fig.
1B). No animals (0 of 7) injected with TGF-
(A) survived
24 h after PH; however, all animals survived 4 h after PH.
Even a dose reduction to 1 × 109 PFU TGF-
(A) was
lethal in animals 24 h after PH. The survival rate of Ctrl- and
TGF-
(L)-injected rats was 100% after PH.
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Functional assessment of TGF-1 in vivo.
TGF-
1 increases
1(I) collagen mRNA in the
liver (7). We measured
1(I) collagen mRNA
by ribonuclease protection assay as a surrogate marker for functionally
active TGF-
1 protein derived from adenoviral
expression. TGF-
(A)-injected animals showed a 2.7 ± 0.36-fold increase in
1(I) collagen mRNA compared with Ctrl-injected animals 48 h after injection (Fig.
2, A and B). TGF-
(L)-treated animals also showed a 3.5 ± 1.0-fold increase in
1(I) collagen mRNA. This result suggests that
functionally TGF-
(A) protein is being made in vivo and that enough
TGF-
(L) is converted to the active form to increase hepatic stellate
cell production of
1(I) collagen mRNA.
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Hepatocyte DNA synthesis.
DNA synthesis was assessed by proliferating cell nuclear antigen
(PCNA) staining in uninfected (saline), Ctrl, and TGF-(L)-injected livers. DNA synthesis could not be assessed in TGF-
(A)-injected rats
because no animals survived 24 h after PH. Saline-, Ctrl, and
TGF-
(L)-injected rats demonstrated 1.7 ± 0.3, 1.5 ± 0.2, and 4.3 ± 2.1% hepatocytes in the S phase before PH,
respectively. Twenty-four hours after PH, however, saline-, Ctrl, and
TGF-
(L)-injected animals demonstrated 31.9 ± 1.0, 26.2 ± 2.0, and 26.8 ± 3.1% replicating hepatocytes, respectively.
TGF-(A) induces apoptosis in hepatocytes.
TUNEL assay of Ctrl and TGF-
(L)-injected rat livers showed little
evidence of apoptosis, but TGF-
(A)-injected rat livers demonstrated
apoptotic hepatocytes (Fig. 3,
A and B). Rats that did not undergo PH showed
0.4 ± 0.2 and 10.5 ± 1.8% apoptotic hepatocytes in Ctrl
and TGF-
(A)-injected animals 24 h after injection, respectively. The percentage of apoptotic hepatocytes increased to
32.1 ± 9.1% in TGF-
(A)-injected animals 48 h after
injection (Fig. 3, A and B). Four hours after PH,
the percentage of apoptotic cells was 0.25 ± 0.2% in
Ctrl-injected animals compared with 14.1 ± 1.3% in
TGF-
(A)-injected animals. Therefore, these results indicate that
expression of TGF-
(A) induces significant apoptosis in the normal
adult liver.
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TGF-(A) increases caspase 3-like and caspase 8-like activity.
Caspase 3-like and caspase 8-like activities were measured to
assess apoptosis signaling in Ctrl- and TGF-
(A)-injected animals. Caspase 3-like and caspase 8-like activities were increased in TGF-
(A)-injected rats 24 h after vector administration compared with Ctrl (Fig. 4). PH and TGF-
(A)
further increased caspase 3-like and caspase 8-like activities with
nearly a fivefold increase in caspase 3-like activity and a fourfold
increase in caspase 8-like activity (Fig. 4).
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Analysis of hepatic function and histological analysis.
Serum obtained at the time of death was used to assess hepatic function
by measuring total bilirubin and ALT concentrations. Twenty-four hours
after vector administration, bilirubin concentrations increased in
TGF-(A)-injected rats compared with Ctrl animals, and further
increases in bilirubin in TGF-
(A)-injected rats were seen 4 h
after a PH (Fig. 5A). Liver
enzyme analysis also demonstrated increases in ALT concentrations in
TGF-
(A)-injected rats compared with Ctrl animals, and PH again
further increased ALT (Fig. 5B). Collectively, these
findings suggest that adenoviral vector administration may result in
slight cytopathic effects but that TGF-
(A) and PH produce marked
alterations in hepatic function.
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Z-VAD.FMK blocks TGF-(A)-induced apoptosis in primary
hepatocytes.
Primary rat hepatocytes were isolated and infected with either
TGF-
(A) or Ctrl adenoviruses in the presence and absence of the
pancaspase inhibitor Z-VAD.FMK. Hepatocytes stained with propidium iodide showed increased membrane blebbing and nuclear condensation, characteristic of apoptosis, in TGF-
(A)-infected cells compared with
Ctrl cells (Fig. 6A,
left). The addition of Z-VAD.FMK blocked apoptosis in
TGF-
(A)-infected cells (Fig. 6A, bottom
right). Similar results were also observed by FACS analysis.
TGF-
(A)-infected cells showed an increase in the percentage of
apoptotic hepatocytes compared with Ctrl cells (18.3 ± 0.35 vs.
10.4 ± 0.45%), and the addition of Z-VAD.FMK inhibited
TGF-
-induced apoptosis (18.3 ± 0.35 vs. 4.9 ± 0.3%;
Fig. 6B). FACS data also showed similar percentages of
hepatocytes in each phase of the cell cycle between Ctrl and
TGF-
(A)-infected cells, suggesting that TGF-
(A) is not inducing
cell cycle arrest but rather apoptosis (data not shown).
TGF-
(A)-infected cells resulted in DNA ladder formation (Fig.
6C, lane 3), whereas Ctrl cells demonstrated no
ladder (Fig. 6C, lane 1). The inhibitor decreased
DNA ladder formation in the TGF-
(A)-infected cells (Fig.
6C, lane 4) and decreased both caspase 3-like and
caspase 8-like activities in TGF-
(A)-infected cells (Fig.
6D). LDH concentrations in media from TGF-
(A)-infected cells were also increased compared with Ctrl cells (1,065 ± 3 vs.
500 ± 38 U/l, respectively), and diminished LDH concentrations were observed with the caspase inhibitor (1,065 ± 3 vs. 458 ± 13 U/l; Fig. 6E). Similar results were observed
with ALT concentrations (data not shown). ALT concentrations increased
in TGF-
(A)-infected cells compared with Ctrl cells (45 ± 0 vs.
30.5 ± 0.5, respectively), and the caspase inhibitor blocked
TGF-
(A) induction of ALT (45 ± 0 vs. 32 ± 1). Expression
of TGF-
(A) in hepatocytes was confirmed by Northern blots (data not
shown).
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DISCUSSION |
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In this study, we investigated whether autocrine expression of
TGF-(L) or TGF-
(A) would control hepatocyte growth in normal and
regenerating liver. We hypothesized that TGF-
(A) would inhibit hepatocyte replication after PH by inducing apoptosis and that these
effects would not be observed in animals treated with TGF-
(L). Surprisingly, we demonstrated that TGF-
(A) but not TGF-
(L)
induced significant apoptosis in the normal liver, independent of PH. This is the first such study to show that autocrine expression of
bioactive TGF-
can modulate hepatocyte survival in normal liver. In
addition, TGF-
(A) was lethal to rats that underwent PH after
TGF-
(A) administration. Lethality was likely due to hepatic failure
from apoptosis-induced insufficient liver volume. Increased
caspase 3-like and caspase 8-like activity confirmed the apoptotic
mechanism of death. These findings, however, were not present in rats
treated with an adenoviral vector expressing TGF-
(L). These findings
suggest that TGF-
(A) exhibits potent growth control of normal and
proliferating hepatocytes by inducing apoptosis and that the conversion
of TGF-
(L) to TGF-
(A) is a rate-limiting step in
TGF-
1 hepatocyte growth control.
The mechanism(s) through which TGF- regulates hepatocyte growth
control may be either cell cycle arrest or apoptosis. Previous studies
have demonstrated that TGF-
induces cell cycle arrest (1, 18,
47) in the G1 phase by preventing the
hyperphosphorylation of pRb, which arrests cells at the
G1/S phase checkpoint (15, 28). Further work
has demonstrated that TGF-
alters expression of
G1-associated cyclins (18), cyclin-dependent
kinases (18), and cyclin-dependent kinase inhibitors
(47). Alternatively, TGF-
controls epithelial cell
growth, specifically hepatocytes, by inducing apoptosis (2, 4, 8,
12, 13, 20, 21, 36-38). Our study is the first to show that
expression of bioactive TGF-
can induce apoptosis in the normal
liver. Our data suggest that TGF-
(A) infection resulted in apoptosis
and not cell cycle arrest, since the caspase inhibitor Z-VAD.FMK
blocked the apoptotic response seen in primary hepatocytes (Fig. 6,
A-E). Additionally, FACS analysis showed little change in
the phases of the cell cycle when cells were infected with TGF-
(A).
The signaling pathways through which TGF-
induces apoptosis are not
known, but Gressner et al. (20) showed that calpain
inhibitors decreased TGF-
-induced apoptosis. Others have suggested
that the transcriptional factor DPC4 and the stress-activated protein
kinase pathway may mediate TGF-
-induced apoptotic cell death
(2). We have shown that caspase 3-like and caspase 8-like
activity is increased markedly after administration of TGF-
(A).
TGF-
-induced apoptosis is associated with the activation of caspase
3 and a cytokine response modifier A (CrmA)-inhibitable caspase
(possibly caspases 1 or 8) in Hep 3B cells (10). This
study (10) also showed that inhibition by CrmA is specific
for the apoptotic effect of TGF-
because CrmA did not alter the
expression of TGF-
-regulated promoters of the plasminogen activator
inhibitor or cyclin A genes. The authors hypothesize that the function
of CrmA is independent of the anti-proliferative and extracellular
matrix-inducing effects of TGF-
. Additionally, treatment of rat
hepatocytes with TGF-
resulted in apoptosis and increased CPP32-like
protease activity that preceded the onset of apoptosis
(26). Additionally, treatment of FAO rat hepatoma cells
with TGF-
induced caspase 2, but not caspase 1, activity
(11). These authors surmised that caspase activity might
be stimuli specific. The mechanisms of caspase activation, caspases
involved, and role of mitochondria in TGF-
-induced apoptosis require
further investigation.
TGF-1 is an inhibitor of epithelial cell growth, yet no
models of exogenously or endogenously produced TGF-
1
growth arrest have demonstrated the lethality present in this study. We
showed that administration of TGF-
1 alone resulted in
apoptosis but was not lethal. The combination of TGF-
(A) and PH,
however, was uniformly lethal by 24 h. Even dose reduction to
TGF-
(A) reliably caused mortality in this model. Death was likely
related to hepatic failure from insufficient liver volume because 10%
of the hepatocytes in the hepatic remnant were apoptotic. Russell et
al. (40) showed that exogenous delivery of
TGF-
1 at the time of or after PH decreased hepatocyte
DNA synthesis but did not inhibit liver regeneration and had no
apparent cytopathic effect. Furthermore, in transgenic mice expressing
bioactive TGF-
1, PH was not lethal, but hepatocyte DNA
synthesis was reduced compared with wild-type mice (6). The findings in the current study demonstrate that autocrine expression of bioactive TGF-
1 and PH significantly alters
hepatocyte growth control, and lethality is due to marked hepatic
dysfunction as seen by increases in total bilirubin and ALT.
Interestingly, release of ALT is usually associated with necrosis and
not apoptosis. This study, however, clearly demonstrates increases in
ALT concentrations with TGF-
(A) infection (Fig. 5B), and
these increases can be blocked by a caspase inhibitor in primary rat
hepatocytes (data not shown), suggesting that release of ALT is
associated with apoptotic cell death in hepatocytes.
Growth control of the liver has been examined extensively, with most
investigations focusing on inducers of hepatocyte proliferation and
hepatic reconstitution. TGF-1 is the most well-studied
inhibitor of hepatocyte growth, yet reports of TGF-
1 as
a potent inhibitor of hepatocyte growth and regeneration remain
unconvincing. Although TGF-
inhibits hepatocyte DNA synthesis, the
liver regenerates completely within days (40).
TGF-
1 transcripts are not present in hepatocytes in the
normal quiescent liver (5, 9) but are present after PH
when TGF-
1 expression increases from 1 to 7 days
(5). TGF-
1 is secreted in the
latent form (17, 29), and bioactive TGF-
1
is required for growth inhibition (19). The current study
shows that autocrine expression of TGF-
(L) delivered to hepatocytes
by an adenoviral vector did not inhibit hepatocyte proliferation or
liver regeneration after PH. These findings suggest that the activation
of TGF-
from a latent molecule to a bioactive epithelial growth
inhibitor represents a critical step of cellular growth control. In
vitro mechanisms of TGF-
activation include heat, extremes of pH,
plasmin (29), and reactive oxygen species
(3). TSP-1 has been shown to bind to LAP and convert
TGF-
(L) to an active molecule (44, 45). Recently, the
integrin
v
6 has been shown to activate
TGF-
in pulmonary inflammation and fibrosis (33), but
this mechanism is unlikely in the liver since little
v
6 is expressed in the liver.
Although the TGF-(L) failed to produce growth control in normal or
proliferating hepatocytes,
1(I) collagen production was increased in response to TGF-
(L). TGF-
is a potent stimulus for
hepatic fibrogenesis (7), and this study demonstrates that the concentration of TGF-
required for
1(I) collagen
production is substantially lower than that required to induce
apoptosis and below the level of detection of our TGF-
bioassay. Our
previous study confirmed that administration of TGF-
(L) was
effective in increasing hepatic stellate cell
1(I)
collagen production 15-fold over control virus-injected mice and
attests to the sensitivity of TGF-
-induced collagen production
(24).
We showed that bioactive TGF- was lethal to rats that underwent
hepatectomy, and histological examination revealed hepatic failure from
massive apoptosis. We were, however, unable to address the effects of
autocrine expression of small amounts of bioactive TGF-
. Although we
decreased the dose by 60%, TGF-
remained lethal. Presumably,
infection with a smaller dose of TGF-
(A) would result in survival,
but whether this dose would permit substantial hepatocyte infection and
allow detection of the transgene is unknown.
We demonstrated that autocrine expression of bioactive TGF- induces
marked apoptosis in the normal liver and is associated with death after
PH. This study suggests that bioactive TGF-
may have important
hepatic growth regulatory functions in both the normal liver and after
PH. Apoptosis, via caspase activation, appears to be the mechanism of
hepatocyte growth control, and activation of TGF-
(L) may be an
important regulatory step in hepatocyte growth control.
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ACKNOWLEDGEMENTS |
---|
We thank Angela Glover and Ellen Hughes for assistance with the preparation of the manuscript.
![]() |
FOOTNOTES |
---|
This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-34987 and by funds from the Center for Gastrointestinal Biology and Disease, University of North Carolina-Chapel Hill.
Address for reprint requests and other correspondence: K. E. Behrns, Dept. of Surgery, CB no. 7210, Chapel Hill, NC 27599-7210 (E-mail: Kevin_Behrns{at}med.unc.edu).
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. Section 1734 solely to indicate this fact.
Received 15 November 1999; accepted in final form 18 August 2000.
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