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Intestinal Intraepithelial Lymphocyte -T Cell-Derived Keratinocyte Growth Factor Modulates Epithelial Growth in the Mouse¹

Section of Pediatric Surgery, Department of Surgery, University of Michigan Medical School, and C. S. Mott Children’s Hospital, Ann Arbor, MI 48109

	Abstract

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Keratinocyte growth factor (KGF) promotes intestinal epithelial growth. To understand the relevance of intraepithelial lymphocyte (IEL)-derived KGF expression on epithelial growth, we used a mouse model of villus atrophy by the administration of total parenteral nutrition, and a model of villus hypertrophy by the creation of a short bowel syndrome. KGF expression was confined to -TCR⁺ IELs. IEL-derived KGF expression was highest in the crypts, somewhat less in the lower portion of villi, and markedly lower in the upper portion of villi. Total parenteral nutrition administration was associated with a down-regulation of IEL-derived KGF expression, and short bowel syndrome was associated with an up-regulation of IEL-derived KGF expression. In the absence of -TCR⁺ IEL, using ^-/- mice, intestinal epithelial cell proliferation decreased in control, and in both mucosal atrophy (22% decline) and mucosal hypertrophy (14%) models. These results show that KGF from IELs is an important factor for maintenance of intestinal epithelial cell proliferation and villus growth.

	Introduction

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Intestinal epithelial cell (EC)³ growth is extremely rapid with new growth within 3–4 days. A number of growth factors support the intestinal epithelium including epidermal growth factor (1), insulin-like growth factor-1 (2), glucagon-like growth factor-2 (3), and keratinocyte growth factor (KGF) (4). KGF is a known mitogenic growth factor, and has been shown to stimulate proliferation of a variety of EC lines (5). KGF is strongly expressed throughout the gastrointestinal tract and embryonic organs. Because KGF receptors have been detected in high numbers in the gastrointestinal tract, it is believed that KGF has a critical role in intestinal epithelial growth and maintenance (6, 7). KGF administration can have a number of beneficial effects. Administration of exogenous KGF can prevent mucositis during the administration of chemotherapy and radiation to the intestine (8), and ameliorates mucosal injury in an experimental model of colitis in rats (9). A recent study from Byrne et al. (10) found that administration of recombinant human KGF (rHuKGF) can improve survival, body weight loss, diarrhea, hematochezia, and histopathology in a dextran sodium sulfate-induced inflammatory bowel disease mouse model. More recently, Fernandez-Estivariz et al. (11) found that KGF administration can enhance trefoil factor 2 expression in the proximal small bowel, as well as result in an increase in goblet cell number and trefoil factor-3 protein content throughout the intestine in a rat starvation model. Additionally, KGF has been shown to be able to prevent mucosal atrophy when given in the same starvation rat model (12). KGF administered to rodents on parenteral nutrition has also been shown to prevent villus atrophy (13). This suggests a relative deficiency of KGF in many of these disorders. Interestingly, it has been shown that -TCR⁺ intraepithelial lymphocytes (IELs) express KGF (14). IELs are the population of immunocytes within the epithelial layer. Despite the recognition that IELs express KGF, the relevance of this expression has not been completely investigated.

Alteration in KGF expression has been studied in a number of intestinal pathological processes. KGF mRNA expression has been shown to be increased in patients with inflammatory bowel disease, in those with colorectal cancer, and in a starvation mouse model (15, 16, 17, 18, 19). Recently, Chen et al. (20) found that -TCR⁺ IELs help preserve the integrity of damaged epithelial surfaces caused by dextran sodium sulfate, by providing a localized delivery of KGF. Additionally, KGF appears to be needed for growth in other epithelial tissue. In a recent report, Jameson et al. (21) showed that dermal -TCR⁺ lymphocytes were found to be needed for adequate epithelial wound healing in a mouse model. Importantly, however, it is difficult to determine whether the observed changes in IEL-derived KGF expression are in response to the inflammatory process, or an adaptive response to the loss of epithelium. It is currently unknown how KGF expression is altered in more controlled states of villus atrophy and hypertrophy. Our study used two models of altered EC growth to study mucosal-derived KGF expression. The first is a mouse model of villus atrophy induced by the administration of total parenteral nutrition (TPN), with the elimination of all enteral feeding. The second is a model of villus hypertrophy (removal of 55% of the mid-small intestine) by creating a short bowel syndrome (SBS). These models permitted the study of increased or decreased villus growth without the associated inflammatory or oncologic processes which were used in previous studies. We hypothesized that KGF expression would be down-regulated in the TPN model, and up-regulated in mice with SBS; and this may contribute to the mechanisms involved in the control of these two processes.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Animals

Studies reported here conformed to the guidelines for the care and use of laboratory animals established by the University Committee on Use and Care of Animals at the University of Michigan (Ann Arbor, MI) and protocols were approved by that committee. Male, specific pathogen-free C57BL/6J mice were used. In some experiments, C57BL/6J-Tcrd^tm1Mom (The Jackson Laboratory, Bar Harbor, ME) were used, as these mice have an induced mutation of the TCR -chain and are deficient of -TCR⁺ T cells. Mice were maintained in temperature-, humidity-, and light-controlled conditions. During the experiments, the mice were housed in metabolic cages.

Operative procedures

TPN model. Administration of TPN was performed as previously described (22). Mice were infused with a crystalloid solution (5% dextrose in 0.45 normal saline with 20 mEq KCl/liter) at 4 ml/day. After 24 h, mice were randomized into two groups (n = 6 each group). The control group received the same i.v. crystalloid solution at 7 ml/24 h, in addition to standard laboratory mouse chow and water ad libitum. The TPN groups received an i.v. TPN solution at 7 ml/24 h. The TPN solution has been described in detail previously (22), and contained a balanced mixture of amino acids, lipids, and dextrose in addition to electrolytes, trace elements, and vitamins. Caloric delivery was based on estimates of caloric intake by the control group and from previous investigators (23), so that caloric delivery was essentially the same in both groups.

SBS model. Anesthetized mice were randomly divided into two groups. A SBS group underwent a 55% mid-small-bowel resection. This consisted of removing all bowel between the point 7-cm distal to the ligament of Treitz and 7-cm proximal to the ileocecal value (24). After resection, an end-to-end jejuno-ileal anastomosis was performed with six interrupted sutures of 7-0 Dexon (Tycohealthcare, Mansfield, MA). Control (n = 6) mice underwent a transection of the mid-small bowel with an immediate reanastomosis. After operation, all mice had free access to water for 12 h. Mice then received standard laboratory mouse liquid diet that improved survival and prevented obstructive problems (micro-stabilized rodent diet; Purina Mills, Richmond, IN) and water ad libitum (25). In both groups, no antibiotics were used.

For all animals in control, TPN, and SBS groups, mice were sacrificed at 7 days using CO₂. Survival in both groups exceeded 85%.

IEL isolation and purification

IEL isolation from the small intestine. IELs were isolated as previously described (26, 27). Importantly, this isolation method does not penetrate into the lamina propria avoiding the contamination of the IEL (27). Briefly, the small bowel was placed in tissue culture medium (RPMI 1640 with 10% FCS; Life Technologies, Gaithersburg, MD). Mesenteric fat and Peyer’s patches were removed. The intestine was then opened longitudinally and agitated to remove mucus and fecal material. The intestine was cut into 5-mm pieces, washed three times in an IEL extraction buffer (l mM EDTA, l mM DTT in PBS), and incubated in the same buffer with continuous brisk stirring at 37°C for 20 min. The supernatant was then filtered rapidly through a glass wool column. Cells were then purified in 40% isotonic Percoll (Amersham Pharmacia Biotech, Piscataway, NJ) by differential centrifugation and reconstituted in tissue culture medium. Viability exceeded 95% using trypan blue exclusion staining. After isolation, the cell suspension contained an enriched lymphocyte population, but approximately one-third of the population was ECs. Because of the relatively scant number of IELs isolated from SBS mice, jejunal and ileal specimens were pooled as one IEL population. These same sections of bowel were taken from the control and TPN mice to insure a similar population of cells was studied in each group.

IEL purification. IEL subpopulations were purified by flow cytometry with cell sorting. Isolated IELs and ECs were initially differentiated by gating on forward scatter and side scatter characteristics. Confirmation of the purity of the IEL and EC populations was done using Ab to the pan-lymphocyte marker (CD45) and an anti-mouse EC mAb (clone G8.8; generously donated by A. Farr (Seattle, WA)) (28). After initial gating, the following Abs were used for isolation of specific IEL subpopulations (BD PharMingen, San Diego, CA): TCR- (GL3) and TCR- (H57). Isotype-matched, irrelevant Abs were used as a negative control. An EPICS Elite (Coulter Pharmaceutical, Miami, FL) flow cytometer was used for cell sorting.

Isolation of the IEL along the crypt-villus axis. Isolation of the IEL was also examined along the crypt-villus axis to gain a better understanding of KGF expression at various levels of epithelial maturation. For this, the intestinal mucosa was dissected by using laser capture microdissection (LCM) techniques (29, 30). Three levels were selected: crypt, lower one-third of villus, and upper one-third of villus. Briefly, small pieces of intestine were embedded in paraffin and underwent histologic sectioning (7 µm) and mounting onto glass slides. Slides were stained with H&E before LCM. Microdissection was performed using a PixCell II Image Archiving Working Station instrument (Arcturus Engineering, Mountain View, CA) (29). Care was taken to limit the microdissection to the epithelium, without extending below the basement membrane, thus avoiding expression of KGF from the lamina propria. After microdissection of each specimen, the thermoplastic film-coated cap containing the captured tissue was placed in a microtube. RNA was extracted by using a Total RNA microprep kit (Stratagene, La Jolla, CA) according to the manufacturer’s instructions. Total RNA (poly(A) positive) was then reversed transcribed into cDNA (see below), and then assayed for KGF mRNA expression using real-time PCR.

KGF gene expression

Reverse transcriptase and qualitative PCR. For identification of which IEL subpopulations express KGF, qualitative PCR was performed. A guanidine isothiocyanate/chloroform extraction method was used with TRIzol (Life Technologies) following manufacturer’s guidelines. Total RNA (poly(A) positive) was then reversed transcribed into cDNA by adding 50 µg/ml of total RNA to the following mixture (manufacturer and final concentrations): nucleotides (ATP, CTP, TTP, and GTP, each at 1 mM; Boehringer Mannheim, Indianapolis, IN,), Moloney murine leukemia virus reverse transcriptase (8 U/µl; Life Technologies), oligo(dT) (2.5 µM; New England Biolabs, Beverly, MA,); and RNase inhibitor (2 U/µl; Boehringer Mannheim). Diethyl pyrocarbonate-treated H₂0 was added to yield an appropriate final concentration. Samples were incubated at 39°C for 1 h and the reaction was stopped by incubating at 95°C for 5 min. Reverse transcriptase product (5 µl) was added to forward and reverse specific oligomers (5 mM); PCR buffer (with 10 mM Tris and 50 mM KCl), MgCl₂ (2.5 mM), and Taq polymerase (0.4 U/sample; PerkinElmer, Foster City, CA), with sufficient diethyl pyrocarbonate-treated H₂O to allow for appropriate concentrations. Specific oligomers were designed using an optimization program (OLIGO 4.1; National Biosciences, Plymouth, MN). GenBank accession numbers were M12481 for -actin, and NM_008008 for KGF. For KGF, the forward primer was 5'-CGCAAATGGATACTGACACG-3', and the reverse primer was 5'-GGGCTGGAACAGTTCACACT-3'. For -actin, the forward primer was GAGGGAAATCGTGCGTGACAT and the reverse primer was AGAAGGAAGGCTGGAAAAGAG. The following thermal cycler (PTC-100; MJ Research, Watertown, MA) settings were used: 94°C for 2 min, followed by 30 cycles of 94°C x 15 s; 55°C x 15 s; and 72°C x 30 s, followed by a 5-min extension time at 72°C. Thermal cycler settings were optimized to insure products were in the linear phase of production. The PCR products were run out on a 2% agarose gel containing ethidium bromide, for 1 h at 170 V and then visualized under UV light. Quantification of cDNA product was completed using image quantitation software (ImageQuant; Molecular Dynamics, Sunnyvale, CA), and a semiquantification of the amount of target PCR product was done by normalizing each sample to the production of -actin.

Real-time PCR. For quantification of KGF expression in control and study groups, real-time PCR was performed. Samples of RNA were studied by using a Smart Cycler thermal cycler (Cepheid, Sunnyvale, CA). For this, a mastermix of the following reaction components was prepared to the indicated end concentration: 2.5 µl of ddH₂O, 6.5 µl of MgCl₂ (25 mM), 2.5 µl of 10x PCR buffer, 1 µl of forward primer (12.5 µM), 1 µl of reverse primer (12.5 µM), 1.5 µl of dNTP, 1 µl of 1/600 SYBR green I, 5 µl of 5x additive reagent, 0.5 µl of AmpliTaq Polymerase, and 2.5 µl of reverse transcriptase product. The following experimental run protocol was used: denaturation program (95°C for 2 min), amplification and quantification program repeated 43 times (95°C for 15 s, 66°C for 15 s, and 72°C for 40 s). Detection of the PCR products was made in real-time by measuring the fluorescent signal emitted by the intercalating of SYBR green into dsDNA at the beginning of each annealing step. Specificity of real time PCR products was documented with gel electrophoresis and resulted in a single product with the desired length (148 bp). Additionally, cDNA was extracted using a centrifugal filter device (Millipore, Bedford, MA), and sequencing of this product showed that it matched the KGF GenBank NM_008008 mRNA sequence. The number of cDNA copies was then calculated using the gene sequence size of each gene amplicon. Serial dilutions of the amplified gene at known concentrations were tested to make a standard curve. Standard curve extrapolation of gene copy number was performed for the KGF gene as well as for the -actin gene. Normalization of values was performed by dividing the number of copies of the KGF gene by the number of copies of -actin. Ratio results are multiplied by 10^-5.

Immunoblot analysis for IEL-derived KGF expression

Briefly, purified IELs by flow cytometry sorting, were homogenized on ice in lysing buffer (31). Protein determination was performed by using a Bio-Rad protein assay kit (Bio-Rad, Hercules, CA). Approximately 80 µg of total protein in loading buffer was loaded per lane, and was separated on a SDS-polyacrylamide-gel electrophoresis (13%). Protein markers (Life Technologies) and human KGF as a positive control were run on the same gel. After electrophoresis, proteins were transferred to a polyvinylidene difluoride membrane (Bio-Rad). The membrane was treated with blocking solution (Zymed Laboratories, San Francisco, CA), and probed with rabbit anti-KGF (purified polyclonal Ab, 1/1000 in blocking solution; PeproTech, Rocky Hill, NJ) for 1 h. Bound Abs were exposed to a peroxidase-conjugated secondary Ab (Zymed Laboratories), detected on x-ray film, and quantified using ImageQuant (Molecular Dynamics).

Immunohistochemistry

At the time of death, a proximal small piece of small intestine was mounted in optimal cutting compound (OCT; Tissue-Tek, Sakura Finetechnical, Tokyo, Japan) and was frozen in liquid nitrogen before storage at -70°C. Frozen tissues were cut (6 µm), and slides were placed in cold acetone for 15 min. Immunohistochemical staining was done by using standard protocol provided by the manufacturer (BD Biosciences). Briefly, endogenous peroxidase was quenched with 3% H₂O₂. Slides were incubated with anti--TCR Ab in a 1/25 dilution (GL3; BD Biosciences) for 1 h, and then incubated with diluted biotinylated secondary Ab (1/100, G70-204; BD Biosciences) for 30 min. Prediluted strepavidin-HPR was then added for 30 min. Slides were exposed to diaminobenzidine substrate (BD Biosciences) and then counterstained with hematoxylin.

IEL were counted according to the method described by German et al. (32). Briefly, -TCR⁺ IELs were determined by counting positive cells, which were located in the epithelial layer of the crypt, the lower one-third of the villus, and the upper one-third of villus. Results are a mean number of IEL in a total of six crypt-villus complexes, and results are expressed as the number of -TCR⁺ IELs per 100 ECs.

Epithelial cell proliferation assay

Mice were injected i.p. with 5-bromo-2-deoxyuridine (BrdU) (50 mg/kg; Roche Diagnostic, Indianapolis, IN) 1 h before mice were sacrificed. Paraffin-embedded sections of 5 µm thickness were deparaffinized with xylene. Immunohistochemistry was done by using a BrdU in situ detection kit according to manufacturer’s guidelines (BD PharMingen). Briefly, endogenous peroxidase was quenched with 3% H₂O₂. Slides were then incubated with biotinylated anti-BrdU Ab in a 1/10 dilution, washed, and then incubated with streptavidin-HRP. Slides were then exposed to diaminobenzidine substrate and counterstained with hematoxylin. An index of the crypt cell proliferation rate was calculated by the ratio of the number of crypt cells incorporating BrdU to the total number of crypt cells. The total number of proliferating cells per crypt was defined as a mean of proliferating cells in 10 crypts (counted at x45 magnification).

Histology

At the time of death, a 5-mm section of mid-small intestine was fixed with 10% formaldehyde for histologic sectioning (5 µm thickness). Tissues were dehydrated, imbedded in paraffin, cut, and then stained with H&E. A calibrated eye objective micrometer was used to measure both villus height and crypt depth. Ten replicate measurements were made per mouse and averaged.

Data analysis

Statistical analysis was conducted using either paired t tests or nonparametric ANOVA with Bonferroni t test for posthoc analysis using SPSS software (SPSS, Chicago, IL). A value of p < 0.05 was considered significant. Unless indicated otherwise, results are expressed as the mean ± SD.

	Results

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General description

Total amount of i.v. fluid was the mean of 7.3 ± 1.4 ml/day for the TPN group and 7.0 ± 1.8 ml/day for the control group (p > 0.05). Mean intake of fluid was similar between TPN and control mice. Mean caloric intake of TPN animals was 9.0 ± 1.2 KCal/day. Body weight was recorded both before the initiation of the study and after mice were euthanized. Body weights at the beginning and end of the study were 25.6 ± 1.4 and 23.8 ± 1.3 g for the TPN group, and 25.0 ± 1.7 and 24.8 ± 1.4 g for the control group, respectively, and were not statistically different between the two groups (p > 0.05). For the SBS group, weight slightly (p > 0.05) declined by 4% in the control group (25.3 ± 3.10 vs 22.3 ± 2.2 g) and 6.3% in the SBS group (24.8 ± 4.90 vs 19.20 ± 4.7 g). An examination of the rate of weight decline in SBS mice showed it to plateau after postoperative day 4, indicating that these mice were in an anabolic phase when examined 7 days after surgery.

TPN led to mucosal villus atrophy and a decline in epithelial cell proliferation

There was a decrease in villus height in the TPN group. Histologic examination demonstrated a 15% decline in jejunal villus height (265 ± 38 vs 310 ± 42 µm) in TPN mice compared with controls (Fig. 1). To examine the change of EC proliferation after TPN, BrdU was injected into mice, which allowed for the detection of ECs that had passed through the S phase of the cell cycle (33). BrdU was incorporated into the proliferating cells of tissues. BrdU-positive cells were found to be distributed in the crypts of Lieberkuhn of the small intestine. Administration of TPN led to a significantly decreased number of BrdU-positive cells (13.5 ± 2.4%) compared with controls (24.4 ± 4.2%) (Fig. 2).

View larger version (115K): [in this window] [in a new window]   FIGURE 1. Histology of jejunal biopsies using H&E staining. Specimens are taken from a representative mouse from control, TPN, and SBS groups (x40). The second row is the respective groups from -/- TCR ( KO) mice. The bar represents a 200-µm length.

View larger version (14K): [in this window] [in a new window]   FIGURE 2. BrdU staining to detect epithelial cell proliferation. BrdU-positive cells in mouse jejunum are detected with immunohistochemical staining. Mice were injected with BrdU, 50 mg/kg body weight via i.p. route 1 h before sacrificing (day 7), n = 6 each group. Crypt cell proliferation rate was calculated by the ratio of the number of crypt cells incorporating BrdU to the total number of crypt cells. *, p < 0.05 for the control group compared with KO controls. **, p < 0.05 TPN compared with KO TPN group, , p < 0.05 SBS compared with KO SBS group.

There was an increase in both villus height and crypt depth in the SBS group. Massive bowel resection led to a 62% increase in jejunal villus height (502 ± 77 µm), as compared with controls (310 ± 42 µm). Crypt depth was also greater in the SBS group than controls (93 ± 18 vs 166 ± 17 µm in the control and SBS groups, respectively) (Fig. 1). SBS significantly increased BrdU-positive cells to 36.3 ± 7.6% compared with controls (24.4 ± 4.2%) (Fig. 2). There was no difference in labeled cell positions between groups of mice injected with BrdU; all positive cells remained in the crypts.

IEL-derived KGF expression

-TCR⁺ IEL express KGF. It has been well established that KGF is expressed in the gastrointestinal tract (5, 14, 34). To determine which mucosal cell was responsible for the increase in KGF expression, IELs and ECs were sorted according to phenotypic cell markers (-TCR and -TCR). After sorting into mucosal subpopulations, cells were analyzed for KGF mRNA expression. As shown in Fig. 3, only -TCR⁺ lymphocytes expressed IEL. KGF was not identified in either -TCR IEL or in ECs. IEL-derived KGF expression was undetected in the ^-/- group.

View larger version (76K): [in this window] [in a new window]   FIGURE 3. Composite gel demonstrating IEL subpopulation expression of KGF mRNA. Samples are from representative specimens in various IEL subpopulations. Cells were purified using flow cytometry sorting. A, -TCR+ IEL. B, -TCR+ IEL. C, IEL from -/- mice. D, Epithelial cells. Note the confinement of KGF mRNA expression to -TCR+ IEL. Results were consistently noted from at least three mice in each group

View larger version (15K): [in this window] [in a new window]   FIGURE 4. Changes in IEL-derived KGF mRNA in intestinal mucosal specimens measured by real-time RT-PCR. Results are expressed as the ratio of the number of copies of the KGF gene to the number of copies of the -actin gene multiplied by 10-5, n = 6 each group. Expression of KGF was markedly increased in SBS mice compared with controls, *, p < 0.05. TPN administration decreased KGF expression when compared with the control group, *, p < 0.05.

View larger version (38K): [in this window] [in a new window]   FIGURE 5. Changes in IEL-derived KGF protein expression in representative control, TPN, and SBS mice. IEL KGF protein expression was markedly increased in the SBS group when compared with the control group, *, p < 0.05, n = 6 per group. TPN administration caused a decline in IEL KGF protein expression when compared with controls, *, p < 0.05. KGF expression was confirmed by using rHuKGF, which is slightly larger than that detected in our mice due to partial proteolysis at the amino-terminal end (54 ).

View larger version (28K): [in this window] [in a new window]   FIGURE 6. Changes in KGF mRNA expression at different levels of the crypt-villus axis. A, Schematic diagram of an intestinal villus showing how the intestinal mucosal levels were divided into three portions using LCM: upper one-third of the villus; lower one-third of the villus; and the crypt portion. B, Results are expressed by dividing the number of KGF gene copies by the number of copies of the -actin gene multiplied by 10-5. Expression of KGF mRNA in the crypts was significantly increased in SBS compared with control mice, *, p < 0.05. TPN administration led to a decreased KGF expression in crypts when compared with controls, *, p < 0.05. There was no significant change in KGF mRNA expression in the upper portion of villi after bowel resection or TPN administration.

View this table: [in this window] [in a new window]   Table II. -TCR+IEL distribution at different portions of intestinal mucosaa

-TCR^-/- knockout (KO) mice receiving TPN or with SBS have significant changes in villus height and epithelial cell proliferation

To further confirm that -TCR⁺ lymphocytes were responsible for the increase in mucosal-derived KGF expression and functional maintenance of EC growth, -TCR^-/- mice were placed on standard mouse chow, received TPN, or underwent bowel resection to create a SBS model. Flow analysis confirmed that no -TCR⁺ lymphocytes were present in the small intestine (data not shown). IEL-derived KGF mRNA expression was undetected in -TCR^-/- mice (Fig. 3). To investigate the role of -TCR⁺ IEL has in promoting EC proliferation, the change in EC proliferation in -TCR^-/- control, TPN, and SBS mice was examined. Mice were injected with BrdU to assess EC proliferation. The number of EC that incorporated BrdU were significantly reduced in control -TCR^-/- mice (20.9 ± 1.1% vs 24.4 ± 3.1% in wild-type control mice, p < 0.05; Fig. 2). In the TPN model, there was a 22% decline in BrdU incorporation in -TCR^-/- TPN mice compared with wild-type TPN mice. Additionally, BrdU-positive cells in -TCR^-/- SBS mice were 20.1% lower than wild-type SBS mice (p < 0.05; Fig. 2).

Fig. 1 shows the alteration in villus height in -TCR^-/- mice compared with wild-type mice. Interestingly, the villus height was significantly (p < 0.05) shorter in -TCR^-/- mice compared with wild-type mice (265 ± 18 vs 310 ± 42 µm). There was a 15% decline in villus height in -TCR^-/- control mice compared with wild-type control mice. In the TPN model, villus height in -TCR^-/- mice was 23% lower than wild-type TPN mice (204 ± 12 vs 265 ± 38 µm, p < 0.05). In the SBS model, villus height in -TCR^-/- mice was 24% lower than wild-type SBS mice (380 ± 21 vs 502 ± 77 µm, p < 0.05). Apparently, the decline in -TCR⁺ lymphocyte-derived KGF expression led to a greater decline in villus height in our TPN (mucosal atrophy) model and resulted in a diminished rise in villus height in the SBS (mucosal hypertrophy) model. These findings further confirm that expression of KGF from -TCR⁺ IEL has an important role in the alteration of villus growth and EC proliferation in both mucosal atrophy and villus hypertrophy models.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
In this study, we found that a decrease in IEL-derived KGF expression is associated with the administration of TPN in a mouse model and an increase in IEL-derived KGF expression occurs in a SBS mouse model. We also report that the up-regulation of mucosal KGF mRNA was localized to -TCR⁺ IELs, which lie in close proximity with intestinal ECs. Additionally, in the absence of -TCR⁺ IEL, EC proliferation decreased in normal controls and in both mucosal atrophy and mucosal hypertrophy models. Moreover, IEL-derived KGF mRNA expression varied widely along the course of the crypt-villus axis, despite a fairly uniform distribution of -TCR⁺ IEL along this axis.

It has been suggested that -TCR⁺ T cells can regulate the generation and differentiation of ECs (16, 35). Komano et al. (35) found a significant decrease in BrdU-labeled ECs in the intestine of -TCR^-/- T cell-deficient mice. This result suggests that -TCR⁺ T cells stimulate the proliferation of crypt stem cells. More recently, a study from Chen et al. (20) found that -TCR⁺ IELs help preserve the integrity of damaged epithelial surfaces induced by dextran sodium sulfate. Using -TCR^-/- mice, it was noted that there were significant defects in the repair of the intestinal mucosa after completion of dextran sodium sulfate treatment. Interestingly, in the absence of KGF (KGF^-/- mice), a similarly impaired intestinal healing occurred. These results suggest that -TCR⁺ IELs play an important role in this process, most likely due to a decreased expression of KGF at the site of intestinal damage. These findings are in agreement with our results, which showed a lower degree of epithelial proliferation as well as villus height in -TCR^-/- mice. Many mechanisms have been reported to be involved in the reciprocal regulation between IEL and EC, including the expression of cytokines and growth factors and cell-to-cell interactions. These intercommunications have been claimed to maintain homeostasis within the gut (14, 35, 36, 37, 38, 39). Evidence to support these claims show that -TCR^-/- T lymphocyte-deficient mice show a decrease in epithelial cell development of the small intestine (35). Interestingly, the villus atrophy in -TCR^-/- TPN mice was more severe than that which developed in wild-type TPN mice, and the degree of villus hypertrophy in -TCR^-/- SBS mice was less than in wild-type SBS mice.

Alterations in KGF expression have been examined in a number of different models. KGF expression appears critical to fetal development of the human intestinal epithelium (15, 34, 40). KGF mRNA expression has been shown to increase in inflammatory bowel disease and in a starvation mouse model (16, 19, 41, 42). KGF mRNA expression has been further shown to be particularly elevated in patients with ulcerative colitis compared with patients with Crohn’s disease (15). These results show an association of increased KGF expression in disease states during the process of increased EC turnover. This suggests that KGF, via -TCR⁺ T cells, plays an important role in maintaining epithelial integrity. Such a function has similarly been shown in dendritic epidermal T cells (21). Such epidermal T cells in murine skin were shown to recognize Ag expressed by adjacent damaged or stressed keratinocytes, with subsequent production of KGF. Because many of the examples given above involve alteration of epithelial growth during an inflammatory process or periods of starvation, we felt that the alteration in KGF expression may not completely reflect the true interrelation of EC growth and IEL-derived KGF expression. We chose to study KGF expression in two controlled models of epithelial growth: administration of TPN and a SBS model.

Administration of TPN results in an altered rate of epithelial turnover, villus atrophy, and loss of epithelial barrier function (43, 44). Our study confirmed that TPN decreased EC proliferation and led to villus atrophy. It has been previously shown that administration of TPN can lead to a number of alterations within the immune system of the small bowel mucosa. Among these changes is an increased expression of IFN- and a loss of cells in the gut-associated lymphoid tissue (22, 23, 45). IELs eliminate infected and senescent cells through a combination of cytotoxicity, and via IFN- and TNF- release (46). This action may insure the rapid renewal of the villi and maintenance of the epithelial barrier function. The increases in IFN- or TNF- may alter the regulation of this barrier function. Mice injected with IFN- show an accelerated epithelial renewal, and TNF- can cause a rapid induction of apoptosis of the villus tips (46). Interestingly, these cytokines can induce KGF expression in fibroblasts. It is possible that a cytokine imbalance prevails with administration of TPN, or with SBS, which may result in the observed altered expression of KGF. In recent studies, exogenous KGF reversed the villus atrophy observed in rodent starvation and TPN models (12, 13, 47, 53). In our mouse TPN model, villus atrophy occurred simultaneous with a 2-fold decline in KGF mRNA expression. This suggests that this lack of IEL-derived KGF expression may play an important role in the maintenance of EC proliferation. It also suggests that exogenous administration of KGF in patients receiving TPN may have a beneficial role in reversing atrophic changes.

The significant increase in villus height in our SBS mice is consistent with previously observed findings of adaptive villus hypertrophy with SBS (24, 48). It was interesting to note that KGF expression increased within a week after the development of SBS, suggesting it was an important early regulator of villus hypertrophy. As shown in our results, -TCR^-/- mice do not express mucosal KGF; however, lamina propria-derived KGF (from nonlymphoid cells) is still present (7, 34). Use of -TCR^-/- mice has given us insight into the physiologic relevance of IEL-derived KGF on normal villus growth, and how it affects adaptive changes with TPN or in SBS. It was also interesting to note that although the villus architecture appears normal in -TCR^-/- control mice, EC proliferation was significantly lower in this group compared with wild-type controls. This finding of a decline in villus height corresponds to the findings of Komano et al. (35), who found a decline in both epithelial proliferation and turnover in ^-/- mice. Clearly, the production of KGF from stromal cells of the lamina propria is present, and still must function to some degree to support epithelial growth through a paracrine action (6, 17, 34). It has been shown that stromal cells in the lamina propria have a strong role in epithelial proliferation (34). Because our study focused on the IEL-derived KGF function, an evaluation of the role of KGF from lamina propria stromal cells in the maintenance of ECs was not performed. Additionally, a number of other growth factors have been shown to have important roles in the adaptation observed in the SBS. Such factors include epidermal growth factor, insulin-like growth factor 1, glucagons-like peptide 2, and hepatocyte growth factor (24, 49, 50). Each of these growth factors, while important by themselves, may have even greater relevance when simultaneously expressed, as the effects are additive (51). The fact that the absence of KGF in our KO mice still allowed for hypertrophy in SBS mice (although at a lower amount), indicates that these other factors were still being expressed.

The expression of IEL-derived KGF changed along the crypt-villus axis. There was a predominate expression in the crypts, with reduced expression in the lower one-third of the villi and marked reduction in the distal villi. Interestingly, the distribution of -TCR⁺ IEL numbers along crypt-villus axis did not parallel these observed changes in KGF expression. This suggests that the functionality of the IEL does not follow the physical distribution of these cells. The reason for this functional difference of -TCR⁺ IEL at different locations along the crypt-villus axis is not clear, but may relate to alterations in signaling between the IEL and EC populations. A number of epithelial functions are differentially expressed from the crypt base to villus tip (52). Although these changes in EC maturation have been well studied, an understanding of how growth factors affect these maturational changes is not well understood. Our observations suggest that the maximal effect of KGF may be in the crypt and lower portion of the villus. This differential expression of KGF within the crypt paralleled the increase in crypt depth observed in our SBS mice. This suggests that KGF is more important for the proliferative effects of ECs, as opposed to the maturational effects of these cells which are found to predominate in the upper portion of the villi (52). Nevertheless, KGF may also result in functional changes to the intestine, as well. We have previously reported that supplementation of rHuKGF to mice receiving TPN resulted in an attenuation of the observed increase in intestinal ion transport typically seen with TPN administration. KGF administration also resulted in an increase in epithelial barrier function (53).

TPN is vital for the nutritional support of patients who cannot take adequate enteral intake. Lack of enteral intake or the administration of TPN results in the development of mucosal atrophy with a decrease in epithelial proliferation, and a loss of absorptive function. Loss of large amounts of intestine in SBS can further cripple patients. Clearly, the potential use of exogenous KGF may alleviate many of the adverse effects that such patients suffer from. A further understanding of KGF expression, and the action of KGF on intestinal villus growth, will hopefully help to address these clinical problems and potentially offer future therapeutic options.

	Acknowledgments

 
We appreciate the cooperation from the University of Michigan-Comprehensive University of Michigan-Biomedical Research Core Facilities Core Flow Cytometry Facility, and the Laser Capture Microdissection Core.

	Footnotes

 
¹ This research was supported by National Institutes of Health Grant AI44076-05 and Abbott Laboratories, Hospital Division. The work was also supported in part by the University of Michigan-Comprehensive Cancer Center National Institutes of Health Grant 5 P30 CA46592, and the University of Michigan-Multipurpose Arthritic Center National Institutes of Health Grant AR20557.

² Address correspondence and reprint requests to Dr. Daniel H. Teitelbaum, Section of Pediatric Surgery, University of Michigan Hospitals, Mott F 3970, Box 0245, Ann Arbor, MI 48109. E-mail address: dttlbm{at}umich.edu

³ Abbreviations used in this paper: EC, intestinal epithelial cell; KGF, keratinocyte growth factor; IEL, intraepithelial lymphocyte; TPN, total parenteral nutrition; SBS, short bowel syndrome; LCM, laser capture microdissection; BrdU, 5-bromo-2-deoxyuridine; KO, knockout.

Received for publication March 19, 2003. Accepted for publication January 8, 2004.

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