Shriners Hospitals for Children and Department of Surgery, University of Texas Medical Branch, Galveston, Texas 77550
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ABSTRACT |
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Gut epithelial
cell death by apoptosis is increased in the gut epithelium
after severe burn associated with mucosal atrophy. We hypothesized that
tumor necrosis factor (TNF)--TNF receptor (TNFR) interaction
activates apoptosis in small bowel mucosal cells after severe
burn. C57BL6 mice received a 30% total body surface area scald burn
and were treated with neutralizing anti-TNF-
. The proximal small
bowel was assessed for mucosal atrophy. Proliferation and
apoptosis of mucosal cells were assessed by proliferative cell
nuclear antigen-immunostaining and terminal deoxyuridine nick-end
labeling assay, respectively. Mucosal height and mucosal cell number
decreased after burn. Anti-TNF-
-treated mice showed significantly
less mucosal atrophy. Proliferation of intestinal cells was not changed
with burn or anti-TNF-
treatment. An over threefold increase in
apoptotic cell number was seen after burn, which was diminished by
anti-TNF-
treatment. Changes in gut mucosal homeostasis after severe
burn are affected, in part, by the activation of apoptosis by
TNF-
-TNFR interaction.
small bowel mucosa; mucosal atrophy; tumor necrosis factor-; apoptosis; proliferation
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INTRODUCTION |
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GUT MUCOSAL
HOMEOSTASIS, and thus the morphological and functional integrity
of the gut, is maintained by a balance between epithelial cell
proliferation and cell death. Proliferation of gut epithelium occurs by
mitosis in the intestinal crypts, whereas cell death occurs throughout
the crypt and villus (19). Apoptosis is programmed
death and removal of senescent or otherwise dysfunctional cells without
inflammation. It can be considered an antagonistic and regulatory
process to cellular proliferation by mitosis. Both apoptosis
and mitosis are continually ongoing in live gut epithelium to maintain
mucosal cellular balance. This delicate balance of mucosal cell mass
can be influenced by exogenous and endogenous factors, such as
nutritional depletion, chronic disease, or severe trauma (i.e., severe
burn). We (19) previously showed increased gut epithelial
cell death by apoptosis in the gut epithelium after severe
burn, which was associated with mucosal atrophy. Potential mechanisms
for these effects include the induction of epithelial cell death
indirectly by relative hypoperfusion or directly through interaction of
inflammatory mediators and their receptors located on gut epithelial
cells. We (14) recently showed that burn-induced hypoperfusion of the gut is insufficient to induce apoptosis of gut epithelial cells. In other organs, programmed cell death can be
induced by several membrane-bound ligand-receptor interactions, such as through Fas ligand (FasL)-Fas interaction or tumor necrosis factor--TNF receptor (TNF-
-TNFR) interaction (3). We
therefore hypothesized that TNF-
-TNFR interaction activates
apoptosis in small bowel mucosal cells after severe burn, and
thus is a crucial element in changes in gut mucosa after severe burn.
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MATERIALS AND METHODS |
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Adult male C57BL6 mice (Harlan Sprague-Dawley, Houston, TX) weighing 23 ± 2 g were housed individually in a temperature-controlled cubicle with a 12:12-h light-dark cycle. Mice were fed and received water ad libitum. The study was approved by the Animal Care and Use Committee of The University of Texas Medical Branch, Galveston, TX.
After 1 wk, the mice were randomly assigned to sham burn control, 30%
total body surface area (TBSA) scald burn, and 30% TBSA scald burn
with treatment by neutralizing hamster-anti-mouse-TNF- antibody.
Mice were anesthetized with methoxyflurane as inhalational agent
(1-4%) and buprenorphine hydrochloride (0.1 mg/kg) given subcutaneously. The dorsum of the trunk was shaved, and a 30% TBSA
burn was administered by placing the animals in a mold exposing an area
of 4.2 × 2.9 cm of the back. The mold was placed in a column of
95 to 99°C steam for 6 s, which delivered a full thickness cutaneous burn demonstrated by histologic sections. Sham control animals were anesthetized, shaved, and placed in the mold without exposure to steam. Burned animals were resuscitated with 1 ml of 0.9%
NaCl solution sq and 1 ml NaCl solution ip. Anti-TNF-
-treated animals received 200 µg neutralizing hamster anti-mouse-TNF-
antibody (BD Pharmingen, San Diego, CA) in 1 ml saline ip in addition to 1 ml resuscitation fluid sq. Mice were then returned to their cages.
This time point was chosen in reference to our previously published
findings (19) in which the maximum apoptotic reaction in small bowel was seen at 12 h. After sham burn or burn, all groups were given water ad libitum and fasted to avoid the confounding variable of different food intake in burned and unburned animals. Animals were killed at 12 h after injury by decapitation. The entire small bowel was excised, measured for length and divided in
half. The proximal segment was flushed with ice cold saline, opened
longitudinally, blotted dry on paper and immediately weighed. Thereafter, a 2-cm segment of the proximal end of the small bowel was
taken and immediately fixed in 10% buffered formalin. This piece was
used for histology, immunohistochemistry, and terminal deoxyuridine
nick-end labeling (TUNEL) assay. Of the remaining 16 to 20 cm lengths,
a 3-cm section was used for the determination of dry weight after
desiccation at 50°C for 48 h. The rest of the bowel sample was
snap frozen in liquid nitrogen and stored at
70°C. Additionally,
measures of body weight and liver weights were obtained.
Formalin-fixed tissues were processed and embedded in paraffin. Three 3-µm sections were obtained of each tissue block at 40-µm intervals, deparaffinized, rehydrated in graded alcohol (100, 95, and 70%), and washed with deionized water. Hematoxylin and eosin staining was performed, and mucosal height, crypt depth, and villus height was determined by randomly selecting 10 complete villi from each section and measuring the distance from the base of the crypt to the villus tip, the base of the crypt to the crypt-villus junction, or from the crypt-villus junction to the villus tip, respectively. Values from the measured villi were averaged to reach individual mucosal height, crypt depth, or villus height measurements. For identification of apoptotic cells the TUNEL method (ApoTag; Oncor, San Francisco, CA) was used. Prepared sections were treated with proteinase K (20 µl/ml in PBS) to digest proteins; endogenous peroxidase activity was quenched with 2% H2O2 in PBS. Seventy-five microliters of equilibration buffer was placed on each section, and diluted TdT enzyme solution was applied and incubated at 37°C for 1 h. After incubation, the slides were placed in stop/wash buffer. Then 55 µl of antidigoxigenin peroxidase was added, and the slides were incubated for 30 min at room temperature. Sections were again washed, and diaminobenzidine-hydrogen peroxide was used for color development. Sections were then counterstained with 2% hematoxylin and mounted for examination. In each section, 10 full-length villi were randomly selected to count TUNEL-positive cells. Apoptotic cells were identified as cells with brown-stained nuclei, or as apoptotic bodies, which are fragments of apoptotic cells engulfed by neighboring epithelial cells. Intraepithelial lymphocytes were excluded by morphology. All epithelial cells within the villi were counted, and apoptosis was expressed as percentage of apoptotic cells of the total cells for each section. Values for all of the three sections were averaged to reach a percentage of apoptosis for the proximal gut of each animal.
Proliferation was quantified in a similar way with immunostaining for proliferative cell nuclear antigen (PCNA). Deparaffinized and rehydrated sections were incubated with a horseradish peroxidase conjugated PCNA-antibody (SC-56, Santa Cruz Biotechnology, Santa Cruz, CA) at a 1:50 dilution overnight at 4°C, followed by washing in PBS, and diaminobenzidine-hydrogen peroxide for color development. After counterstaining and mounting, PCNA-positive cells were counted on three sections for each animal as described earlier. All examinations were carried out by blinded observers (M. Spies and V. L. Chappell).
Serum collected at death was assayed for TNF- by ELISA following the
manufacturer's instructions (Biosource International, Camarillo, CA).
For RT-PCR, total cellular RNA was isolated from small bowel samples by
acid guanidinium thiocyanate-phenol-chloroform extraction using TRIzol
reagent (GIBCO-BRL, Rockville, MD), Samples were homogenized in TRIzol
reagent on ice, and total RNA was extracted following the
manufacturers' instructions. Extracted RNA was quantitated by
ultraviolet (UV) spectrophotometry and stored at 80°C for future
analysis. The cDNA reaction as well as the PCR were performed with an
optimized buffer and enzyme system (Titan One Tube RT-PCR System;
Roche, Indianapolis, IN) according to the manufacturer's instruction.
This system is designed to use avian myeloblastosis virus (AMV) RT for
first-strand synthesis and the Expand high-fidelity blend of
thermostabile DNA polymerases, which consists of Taq DNA
polymerase and a proof reading polymerase, for the PCR part. The
reaction was carried out in 50-µl volume containing 50-100 ng of
the total RNA, 10 pM of forward and reverse primers specific for
TNF-
(GenBank accession no. M11731, forward: 5'-AGC AAA CCA CCA AGT
GGA GG-3' and reverse: 5'-CAA GGT ACA ACC CAT CGG CT-3'), 1× PCR
buffer with Mg2+, 0.2 mM 2-deoxynucleotide 5'-triphosphate,
5 mM dithiothreitol solution, 5-10 units of RNAse inhibitor, and
0.05 U/µl reaction of the enzyme mix (high-fidelity enzyme mix, RT,
and AMV in storage buffer). An initial RT step was performed at 50°C
for 30 min and 94°C for 2 min for one cycle, followed by 35 cycles
(denaturation 94°C for 30 s, annealing at 59°C for 45 s,
and extension at 68°C for 30 s), and finally, one cycle at
68°C for 7 min. In addition, a pair of primers was designed to
amplify a portion of the mouse
-actin transcript that spans an
exon/exon boundary (GenBank accession no. W82269, forward: 5'-CCT TCA
ACA CCC AGC CAT GT-3' and reverse: 5'-TGT GGA CCA CCA GAG GCA TAC-3').
-Actin was used as a "housekeeping gene" to provide an internal
marker for mRNA integrity within the experiment. PCR products were
separated on (1% wt/vol) agarose gels, visualized by ethidium bromide
staining under UV light. Image capture and density analysis of bands
were done with the SynGene gel documentation system (SynGene-Synoptics,
Cambridge, UK).
For evaluation of caspase-8 presence and activity, tissue samples were homogenized in lysis buffer. For Western blot, 25-30 µg of total protein from the tissue extract were separated on a 10% SDS-polyacrylamide gels under reducing conditions and transferred to nitrocellulose membranes (Hybond-C; Amersham Pharmacia Biotech, Piscataway, NJ) in a semidry blotting chamber. After blockage of nonspecific binding sites with 5% nonfat milk in PBS containing 0.1% Tween-20, membranes were incubated in 1:1,000 dilution of anticaspase-8 rabbit polyclonal antibody (Santa Cruz Biotechnology) for 2 h at room temperature. After extensive washing, the nitrocellulose membrane was incubated with anti-rabbit IgG conjugated with horseradish peroxidase (final concentration 1:2,000) for 90 min at room temperature. Bound antibodies were detected with enhanced chemiluminescence Western blotting detection reagents (Amersham Pharmacia Biotech) according to manufacturer's instructions. Image capturing and density analysis of bands were again performed using the SynGene gel documentation system (SynGene-Synoptics).
Caspase-8 activity was determined by a colorimetric assay (R&D Systems, Minneapolis, MN) according to the manufacturer's instructions. In brief, tissue extract of 100 to 200 µg of total protein were incubated with 5 µl of a caspase-8 specific peptide conjugated with IETD-p-nitroaniline in reaction buffer at 37°C for 2 h; then light absorption was read at a wavelength of 405 nm together with controls. Results were compared between groups.
Statistical analysis of data was performed by one-way ANOVA with Tukey's test using a statistical software package (SigmaStat 2.03, SPSS, San Rafael, CA). Significance was accepted at P < 0.05.
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RESULTS |
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Over the 12-h study period, no significant changes in body weights
were seen. Total liver weights were not different between groups. Wet
weights of the proximal small bowel decreased in burned and
anti-TNF- antibody-treated animals compared with controls (Table
1). Dry weights of the proximal small
bowel showed a similar decrease.
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Histologic measurements showed significantly decreased mucosal height
after severe burn (447 ± 30 vs. 602 ± 20 µm;
P < 0.05). Animals treated with anti-TNF- showed
significantly less mucosal atrophy (533 ± 12 vs. 447 ± 30 µm, P < 0.05) than untreated animals (Table
2 and Fig.
1). Treatment with neutralizing
antibody alone in unburned animals did not show morphological changes
compared with unburned controls. When villus heights and crypt depth
were analyzed separately, only villus height was decreased by burn and
again was partially restored by anti-TNF-
, whereas crypt depth was
not affected (Fig. 2). A similar pattern
was seen in the total mucosal cell number. In burned animals, total
mucosal cell number was significantly decreased. This effect is
partially, but not completely, restored by anti-TNF-
treatment (Fig.
3). Again, the main changes were seen in
the villus and not in the crypt.
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Proliferating intestinal cells, as identified by PCNA staining, were
found in the crypt. However, the number of PCNA positive stained cells
per crypt was not different among groups (Fig.
4). TUNEL-positive cells and
apoptotic bodies were much more likely to be found in sections of
burned animals. An over threefold increase in apoptotic cell number
was seen after burn. This response was diminished by anti-TNF-
treatment. However, this did not result in a complete return to
unburned control values (Fig. 5).
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Systemic TNF- levels showed no significant differences among groups
when tested 12 h after burn (data not shown). Gene
expression levels for TNF-
in small bowel, determined by RT-PCR,
showed no differences among groups (Fig.
6). Caspase-8 expression and activity by
Western blot and activity assay also showed no differences among groups
(Fig. 7).
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DISCUSSION |
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Our results show that the response of gut epithelium to severe
burn is a decrease in small bowel weight, which occurs within 12 h
of injury. These changes take place without concomitant changes in
total body weight. This weight loss in the proximal small bowel is an
indication that injury induces gut mucosal atrophy and that this
atrophy occurs very early after injury (within 12 h). Histologic changes associated with this finding are loss of mucosal height and
cell number. The changes after injury occur predominantly in the villus
region. In this study severe burn did not induce any changes in
intestinal cell proliferation. However, intestinal cell
apoptosis occurred at a threefold higher rate in burned
animals. Additionally, we found that atrophic changes in mucosal height and cell mass could be partially restored by treatment with a neutralizing anti-TNF- antibody, implicating TNF as an effector of
gut mucosal changes seen after severe burn. The effect is clearly more
pronounced in the villus region and does not seem to occur in the
crypt. Increased apoptosis in burned animals was diminished by
anti-TNF-
treatment without changes in proliferation.
As previously shown by our group, the loss of gut epithelial cells
after severe burn is due to an increase in apoptotic cell death
(19). This was coupled with an increase in proliferation, indicating increased cell turnover after injury. Maximum response in
gut epithelium in this model was seen at 12 h after burn, after which the response quickly diminishes (19). Although
similar effects may also be seen at earlier or later time points, the most prominent changes were seen at 12 h, making this a valuable time point to investigate the role of TNF- in gut epithelial apoptosis. Potential mechanisms for these events are changes in gut perfusion with burn or the systemic effect of inflammatory mediators released immediately following injury. Studies have shown
that in rat models, ischemia alone is able to induce
apoptosis in jejunal and ileal mucosa, which is further
exacerbated by reperfusion effects (6, 10, 17). However,
this appears not to be the cause for the changes observed in severe
burn. When addressing this issue we showed in a recent study that
significant gut hypoperfusion occurs with severe burn injury within the
first 4 h (14). When inducing a comparable
hypoperfusion in the gut of unburned animals, the apoptotic index
did not differ from normal controls, indicating that hypoperfusion
induced by burns is not sufficient by itself to cause the observed
increase in apoptosis. This confirms that, in fact, the
atrophic changes are induced by increased apoptosis of gut
epithelial cells and shifts the point of interest to other potential
initiation mechanisms of apoptosis.
Several soluble and membrane-bound factors are known to induce
apoptosis. Activation of members of the TNF-receptor
superfamily as well as associated proteins (TNFR, TRAIL, TRADD, FADD,
etc.) has been implicated as a potential mechanism in many cell systems as an inductor of apoptosis (1). Garside et al.
(4) showed that a single dose of TNF- alone is able to
induce typical small bowel pathology, which is manifested as crypt
hyperplasia and villus atrophy with 15 min of application. In a study
by Piguet et al. (13) neutralization of TNF-
in a
murine acute graft-versus-host-disease (GVHD) model reduced target
organ damage. Recently, Stuber et al. (16) showed in a
murine GVHD-model that FasL-Fas interaction is not involved in the
induction of apoptosis of the small intestinal mucosa. The
neutralization of TNF-
reduced the amount of apoptosis and
the extent of mucosal atrophy. Another study by Guy-Grand et al.
(5) showed increased villous cell apoptosis in
TNF-
treated normal animals, which appears to be dependent on the
presence of intraepithelial lymphocytes. This differs from increased
apoptosis in lymphatic tissues such as spleen and thymus found
in reaction to burn injury, which was linked to increased FasL mRNA
expression and increased caspase-3 activity (2, 3).
Apparently the apoptotic response is triggered differently in
distinct tissues.
Triggering of the apoptotic response by TNF- in gut epithelium
may be initiated by either increased systemic levels acting locally in
the epithelium or by increased local production of TNF-
in the gut
mucosa itself. Both mechanisms are valid options and could explain the
observed phenomenon of increased gut epithelial apoptosis.
Systemic levels of TNF- have been found to be elevated after burns
by several authors (7, 8, 21, 22) with burn injury alone
as well as with an additional second hit. Systemic effects of TNF-
neutralization were investigated by O'Riordain et al.
(12) who described a time and dose dependency of survival in a burn and septic challenge model. Efficacy of treatment depended on
intrinsic TNF-
production, showing greatest effects of
neutralization with maximum TNF-
levels. Applying these results to
our study led us to an early application of anti-TNF-
with knowledge
of sharp and transient increase in systemic TNF-
levels. However, we
were not able to detect differing TNF-
levels with burn injury at
the 12-h time point. This leads to the assertion that only early
neutralization might be effective and, secondly, that TNF-
may only
act as a transient trigger of apoptosis in the small bowel by
activating a cascade of events, which in turn, consolidates the tissue response.
On the other hand Ogle et al. (11) and Wu et al.
(20) showed increased local production of TNF- in
enterocytes and hepatocytes of guinea pigs after thermal injury. This
increase in local expression of TNF-
in small bowel may also induce
apoptotic changes. These changes in local TNF-
levels could be
due to increased transcription or increased translation. However, we
were not able to detect specific differences in local TNF-
mRNA
expression. Here again, TNF-
may only act as a trigger mechanism for
induction of apoptosis at an early time point after burn and
not be expressed at elevated levels at the 12-h time point. The
reported findings do exclude either of these mechanisms but also are
not able to support either of these hypotheses. However, the presence
of increased TNF-
levels may be irrelevant, as effects may also be
realized with increased activity below the sensitivity of our methods.
To elucidate the effective source of the TNF-
trigger requires
further investigation.
The activation phase of cell death includes a variety of transduction
pathways with signals implicating FasL-receptor interactions (FasL-Fas,
TNF--TNFR) and the additional proteins (FADD, RIP, TRADD) (15,
18). These proteins, acting as intermediaries with "death
domains," in turn activate procaspase-8 and consequent caspase-3 and
-6. Although we were not able to show differences in caspase-8
activation with anti-TNF-
treatment, this does not preclude a
TNF-
-mediated mechanism for increased gut epithelial apoptosis. Because the absolute number of cells undergoing
apoptosis at a certain time point is 3-8 cell per 1,000 mucosal cells, differences on the protein expression levels would be
predicted to be small, if at all detectable. However, the effects at a
physiological level, here as mucosal atrophy, are clearly demonstrated
and suggest that TNF-
is an effector, at some level, of diminished
gut mucosal integrity.
It is speculated that the administration of anti-TNF- in the early
phase of injury binds to systemic serum TNF-
or locally abundant
TNF-
. The early neutralization of circulating soluble and
transmembrane forms of TNF-
thus removes the initiating factor for
the apoptotic cascade in gut mucosa. A prevention of TNF-
/TNFR interaction would reduce effective signaling with caspase activation. An alternative mechanism might be the inhibition of signals activating the mitogen-activated protein kinase pathway (9).
From this study, we conclude that changes of gut mucosal homeostasis
seen after severe burn are associated with activation of
apoptosis by TNF--TNFR interaction. The effect of TNF-
neutralization on gut mucosal homeostasis partially reversed mucosal
atrophy after burns in this study. It remains open for discussion
whether this is, in fact, beneficial on the gut or systemic level.
Anti-TNF-
strategies were indeed successful to improve survival and
outcomes in injured animals (12). However, similar use in
patients was unsuccessful. Only in defined studies in other animal
models and perhaps in patients will it be possible to determine whether
TNF-
neutralization may be of benefit. Whether these findings can be used to clinical advantage will require further study to elucidate the
source of TNF-
involved in the response and, secondly, to determine
whether inhibiting mucosal atrophy after burn is of clinical benefit.
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ACKNOWLEDGEMENTS |
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This study was supported by Shriners Hospitals for Children Grant 8580.
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FOOTNOTES |
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This study was presented, in part, at the Surgical Forum of the Annual Meeting of the American College of Surgeons, Chicago, IL, October 22-28, 2000.
Address for reprint requests and other correspondence: Steven E. Wolf, Shriners Hospitals for Children, 815 Market St., Galveston, Texas 77550 (E-mail: swolf{at}utmb.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.
March 28, 2002;10.1152/ajpgi.00149.2001
Received 10 April 2001; accepted in final form 13 March 2002.
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