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Originally published In Press as doi:10.1074/jbc.M408047200 on July 28, 2004

J. Biol. Chem., Vol. 279, Issue 41, 42867-42874, October 8, 2004
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Intectin, a Novel Small Intestine-specific Glycosylphosphatidylinositol-anchored Protein, Accelerates Apoptosis of Intestinal Epithelial Cells*

Hidefumi Kitazawa,abc Tamao Nishihara,abd Tadahiro Nambu,e Hitoshi Nishizawa,af Masanori Iwaki,a Atsunori Fukuhara,a Toshio Kitamura,g Morihiro Matsuda,adh and Iichiro Shimomuraafij

From the aDepartment of Medicine and Pathophysiology, Graduate School of Frontier Bioscience, Graduate School of Medicine, Osaka University, 2-2 Yamadaoka, Suita, Osaka 565-0871, the eTsukuba Research Institute, Banyu Pharmaceutical Co., Ltd., Okubo 3, Tsukuba 300-2611, the fDepartment of Internal Medicine and Molecular Science, Graduate School of Medicine, Osaka University, 2-2 Yamadaoka, Suita, Osaka 565-0871, the gDepartment of Hematopoietic Factors, Institute of Medical Science, University of Tokyo, Minato-ku, Tokyo 108-8639, iPrecursory Research for Embryonic Science and Technology, Japan Science and Technology Agency, 4-1-8 Honcho, Kawaguchi, Saitama 332-0012, and the d21st Century Center of Excellence Program, the Japan Society for the Promotion of Science, Tokyo 102-8471, Japan

Received for publication, July 16, 2004


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Intestinal epithelial cells undergo rapid turnover and exfoliation especially at the villus tips. This process is modulated by various nutrients especially fat. Apoptosis is one of the important regulatory mechanisms of this turnover. Therefore, identification of the factors that control epithelial cell apoptosis should help us understand the mechanism of intestinal mucosal turnover. Here, we report the identification of a novel small intestine-specific member of the Ly-6 family, intectin, by signal sequence trap method. Intectin mRNA expression was exclusively identified in the intestine and localized at the villus tips of intestinal mucosa, which is known to undergo apoptosis. Intectin mRNA expression was modulated by nutrition. Intestinal epithelial cells expressing intectin were more sensitive to palmitate-induced apoptosis, compared with control intestinal epithelial cells, and such effect was accompanied by increased activity of caspase-3. Intectin expression also reduced cell-cell adhesion of intestinal epithelial cells.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Intestinal epithelial cells originate from stem cells at the base of the crypt and migrate along the crypt-villus axis toward the intestinal lumen. When intestinal epithelial cells reach the luminal surface at the villus tips, they finally exfoliate into the lumen, and their cell cycle is terminated with a life span of only 3–5 days (1, 2). This constant and rapid turnover of intestinal mucosa is essential for maximal nutrient absorption, adaptation to changes in diet, and repair of mucosal injury (3).

Apoptosis plays an important role in maintaining the physiological integrity of many tissues. In the intestine, apoptosis is a key regulator for the turnover of intestinal mucosa, and apoptotic intestinal epithelial cells have been detected at the villus tips of the small intestine and the colonic luminal surface (4–6). However, the one or more underlying mechanisms of this process have not been elucidated. In this regard, it is important to identify the factors that control intestinal epithelial cell apoptosis to understand the mechanism of intestinal mucosal turnover.

The present study was designed to identify a small intestine-derived protein or secretory protein modulated nutritionally that is involved in the control of intestinal epithelial cell apoptosis. Using the efficient signal sequence trap (SST)1 method (7), we identified a novel small intestine-specific GPI-anchored protein, intectin, which showed distinct localization at the villus tips of intestinal mucosa and accelerated fatty acid-induced apoptosis of intestinal epithelial cells.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cloning of Intectin cDNA—Poly(A)+ RNAs were extracted from the small intestinal epithelium of C57BL/6J mice under three feeding conditions; ad libitum, 24-h fasting, and 24-h fasting followed by 24-h feeding, and from the small intestinal epithelium of ad libitum-fed db/db mice. Equal amount of poly(A)+ RNA from each group was pooled and used as the template to synthesize complementary DNA (cDNA). To selectively clone the genes with signal sequence at the N-terminal end of cDNAs, the SST-REX system (signal sequence trap by retrovirus-mediated expression screening system) was introduced as described previously by our laboratories (7). Briefly, cDNA was synthesized from the poly(A)+ RNA by random hexamers, using the SuperScript System (Invitrogen), and was then inserted into BstXI sites of the pMX-SST vector, using BstXI adapters (Invitrogen). The ligated DNA was amplified in DH10B cells (Electromax, Invitrogen) to construct an SST-REX library, and a library DNA was prepared using Qiagen plasmid kits (Qiagen). High titer retroviruses representing the SST-REX library were produced using the packaging cell line PlatE and infected to Ba/F3 cells. After 1-day infection period, selection of factor-independent Ba/F3 cells commenced in the absence of interleukin-3, using 96-well multititer plates. The integrated cDNAs were isolated from the interleukin-3-independent Ba/F3 cells by genomic PCR and sequenced.

Animals and Experimental Protocol—C57BL/6J and obese diabetic db/db mice were obtained from Clea Japan (Tokyo) and kept under a 12-h/12-h dark/light cycle (lights on: 8 am to 8 pm) at constant temperature (22 °C) with free access to food and water. The composition of the diet was as follows: carbohydrate, 54.0%; protein, 23.8%; and fat, 5.1%. For analysis of intectin mRNA expression in various tissues of mouse, 6-week-old male C57BL/6J mice were used. For fasting and refeeding time-course experiments, 3–4 male mice (6-week-old) were used in each group. The groups consisted of mice fed ad libitum with standard chow, mice fasted for 24 h, and mice refed 1, 2, and 4 h after 24-h fasting. Mice were sacrificed under deep anesthesia, and their small intestines were harvested immediately. Experiments designed to determine intectin mRNA expression in the small intestine were conducted in 6-week-old C57BL/6J and db/db mice. All experimental protocols described in this report were approved by the Ethics Review Committee for Animal Experimentation of Osaka University.

RNA Isolation, Northern Blot, and Quantitative Reverse Transcription-PCR Analysis—Total RNA was isolated from various tissues, using RNA STAT-60 kit (Tel-Test "B" Inc., Friendswood, TX) according to the instructions provided by the manufacturer.

Ten micrograms of total RNA was subjected to Northern blotting as described previously (8). First strand cDNA was synthesized using ThermoScriptTM RT-PCR System (Invitrogen). Quantitative RT-PCR was performed on a LightCycler using the FastStart DNA Master SYBR Green I (Roche Diagnostics, Tokyo, Japan) according to the protocol provided by the manufacturer. The following forward and reverse primers were used for quantitative PCR amplifications: mouse intectin, 5'-GTTGCCCCTGATTCTGCTGG-3' and 5'-GCACTATTGCAGAGGT-CCGT-3'; mouse cyclophilin, 5'-CAGACGCCACTGTCGCTTT-3' and 5'-TGTCTTTGGAACTTTGTCTGCAA-3'.

Construction of Stable Expression Vector—To express FLAG-tagged intectin, we constructed an expression vector of FLAG-tagged intectin containing the signal sequence of CD59. We prepared a cDNA of intectin (amino acids 21–111: removed the region corresponding to intectin signal sequence) by PCR using a pair of primers containing HindIII and NotI restriction enzyme sites, respectively, 5'-CCCCAAGCTTTTGAAGTGTCATGAATGC-3', corresponding to amino acids 21–26 and 5'-TTTTTTTTGCGGCCGCCTACTGGCTGAGATAGATATAGC-3', corresponding to amino acids 106–111. The amplified product was digested with HindIII and NotI to isolate a 290-bp fragment. The pME-puro-FLAG-CD59 vector (a kind gift from Dr. T. Kinoshita, Osaka University) was digested with HindIII-SfiI and NotI-SfiI, respectively. The above three isolated fragments were ligated to generate pME-CD59 (signal sequence)-FLAG-intectin.

Cell Culture—IEC-6 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 100 units/ml penicillin, and 100 mg/ml streptomycin in a watersaturated atmosphere with 5% CO2 at 37 °C. To measure palmitate-induced cell death, IEC-6 cells were seeded at a density of 2 x 104 cells/cm2. The medium was removed 36 h later, and the cells were incubated in DMEM supplemented with 1% FBS in the presence of 0.5% bovine serum albumin (BSA) alone or with various concentrations of palmitate (25, 50, 100, and 200 µM) for 12 h.

Establishment of Stable Cell Lines Expressing FLAG-tagged Intectin—To create stable cell lines that express FLAG-tagged intectin, electroporation was performed using GenePulser transfection apparatus (Bio-Rad Laboratories, Richmond, CA). IEC-6 cells (80–90% confluence) were trypsinized and suspended in DMEM. Approximately 6 x 106 cells were resuspended in FBS-free DMEM without antibiotics and with 10 µg of plasmid DNA in a final volume of 400 µl. Electroporation was performed at room temperature (500 microfarads, 250 V, 0.4-cm cuvette). After electroporation, the cells were incubated at room temperature for 10 min, diluted in DMEM with 10% FBS, and then plated onto a 10-cm dish. The transfected cells were selected by growth in a medium containing 2 µg/ml puromycin. Cell clones were obtained from individual puromycin-resistant colonies using the limiting dilution method. FLAG-tagged intectin expression was investigated by flow cytometry.

Cell Fractionation and Western Blotting—IEC-mock cells (IEC-mock) and IEC-6 cells expressing FLAG-tagged intectin (IEC-intectin) were harvested after confluence. The cells were lysed with extraction buffer containing a protease inhibitor mixture as described previously (8) and divided into membrane fraction and cytosolic fraction by centrifugation (15 min, 20,000 x g, 4 °C). Equal amounts of protein were subjected to SDS-14% PAGE and transferred onto polyvinylidene difluoride membrane. The membrane was incubated with a mouse anti-FLAG M2-horseradish peroxidase conjugate (Sigma). Immunoreactive protein bands were visualized by using the ECL kit (Amersham Biosciences).

Flow Cytometry—For selection of FLAG-tagged intectin-expressing cells, cells were stained with anti-FLAG FITC-conjugated antibody. Stained cells were analyzed by FACSort (BD Biosciences). The cellular DNA content was determined by flow cytometric measurement of propidium iodide (PI) binding. In preparation for flow cytometry, both adherent and unattached cells were harvested and combined. The cells were washed twice with ice-cold PBS and fixed with 70% cold ethanol. After treatment with 1 µg/ml DNase-free RNase A in PBS containing 10 µg/ml PI, the cells were analyzed using a FACSort. The cell cycle distribution was quantified by using FlowJO (Tree Star Inc., Ashland, OR) software.

In Situ Hybridization—According to the instructions supplied by the manufacturer (digoxigenin (DIG) RNA labeling kit (SP6/T7), Roche Diagnostics), DIG-labeled RNA probes were synthesized using intectin full-length cDNA, which was cloned into pCRII-TOPO (Invitrogen). Male C57BL/6Cr Slc mice (6-week-old, SLC, Shizuoka, Japan) were deeply anesthetized by sodium pentobarbital and perfused transcardially with PBS and 4% paraformaldehyde in phosphate buffer. Tissues were postfixed for 12–24 h with the same fixative and embedded in paraffin. Tissue sections (4 µm) were treated with 0.2 N HCl for 30 min and 10 µg/ml proteinase K for 15 min at 37 °C followed by re-fixation with 4% paraformaldehyde. Hybridization was performed overnight at 50 °C in a hybridization solution (40% formamide, 0.6 M NaCl, 10 mM Tris-HCl, pH 7.4, 1 mM EDTA, 1x Denhardt's solution, 250 µg/ml tRNA, 125 µg/ml salmon sperm DNA, 10% dextran sulfate, and 200 ng/ml DIG-labeled probe). After treatment with 20 µg/ml RNase A (Sigma) for 30 min at 37 °C, the sections were washed with 0.2x SSC at 55 °C. The sections were immunolabeled with anti-DIG alkaline-phosphatase conjugate (Roche Diagnostics), and color was developed using nitro blue tetrazolium/BCIP (5-bromo, 4-chloro, 3-indoylphosphate), which created black and purple signals. Some specimens were counter-stained with hematoxylin.

Cell Viability Assay—The cells (5 x 103/well) in 96-well plates were cultured for 12 h, then exchanged with medium in the absence or presence of various concentrations of apoptotic inducer, and further incubated for 12 h (palmitate) or 24 h (camptothecin and daunorubicin). The cell viability was determined using Cell Count Reagent SF (Nacalai Tesque, Japan). WST-8 (9), which is based on colorimetric quantification of NADH, was added to the culture. After 1 h, absorbance at 450 and 650 nm was measured using a Viento multi-spectrophotometer (Dainippon, Tokyo, Japan).

In Vitro Caspase-3 Activity Assay—Caspase-3 activity was measured using the ApoAlert caspase assay kit (BD Biosciences), using the instructions supplied by the manufacturer. Briefly, cells were washed twice with ice-cold PBS, lysed for 10 min on ice with cell lysis buffer (BD Biosciences), and centrifuged (10 min, 15,000 x g, 4 °C) to remove debris. After determination of protein concentration using the BCA protein quantification kit (Pierce Chemical Co.) with BSA as a standard, 10 µg of the protein was incubated with caspase-3 substrate, ac-DEVD-7-amino-4-trifluoromethyl coumarin at 37 °C for 1 h. The initial rate of release of free ac-DEVD-7-amino-4-trifluoromethyl coumarin was measured using a Spectra MAX GeminiXS microplate reader (Molecular Devices, Sunnyvale, CA) in fluorescence mode using an excitation filter of 400 nm and an emission filter of 505 nm. Enzyme activity was calculated according to the formula provided by the manufacturer.

Cell Aggregation Assay—Cell aggregation assay was performed as described previously (10). Parental Chinese hamster ovary (CHO) cells and intectin-transfected CHO cells (CHO-intectin) were washed with PBS and incubated with 0.125% trypsin and 0.5 mM EDTA at 37 °C for 3 min. Dispersed cells were suspended in Hanks' balanced salt solution (1 x 106 cells/ml) and then placed on BSA pre-coated 24-well plate (0.5 ml). The cells were incubated under continuous shaking at 70 rpm for the indicated periods of time. The reaction was stopped with the addition of an equal volume of 4% paraformaldehyde. Samples were evaluated by counting single cells and aggregates.

Statistical Analysis—Data are expressed as means ± S.E. Statistical analyses were performed with unpaired t-tests. A p value less than 0.05 denoted the presence of a statistically significant difference.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Identification of Intectin as a Small Intestine-derived Nutritionally Modulated Membrane Protein—We previously developed an efficient SST method using retrovirus-mediated gene transfer (7). To identify small intestine-derived nutritionally modulated membrane protein or secretory protein, we conducted this SST method using the pooled poly(A)+ RNA from the small intestinal epithelium of mice at ad libitum, fasted and refed conditions, and obese diabetic db/db mice. We screened and sequenced 1224 clones. Four clones were selected as unknown proteins with signal sequence or transmembrane region. The mRNA of one clone was exclusively expressed in the small intestine among various tissues of mice as revealed by Northern blotting analysis and quantitative RT-PCR (Fig. 1, A and B). This clone and its full-length cDNA were selected and named intectin. The nucleotide sequence and deduced 111-amino acid sequence of intectin are shown in Fig 1C. A hydrophobicity plot revealed a 20-residue N-terminal and a 13-residue C-terminal signal peptide (Fig. 1, C and D) (11). A potential GPI-anchoring site was identified at Asn-88 based on the published GPI consensus sequences (Fig. 1C) (12). A Basic Local Alignment Search Tool (BLAST) search revealed that intectin sequence is identical to NM_025929 [GenBank] (GenBankTM), whose function is totally unknown, and that intectin has a Ly-6 domain (Fig. 2). All cysteine residues highly conserved among Ly-6 family are present in intectin. The amino acid similarity of intectin to other Ly-6 family is not high, with homology of only 30–40% (Fig. 2).



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  		FIG. 1.
Expression and sequence of intectin. A, Northern blot analysis of intectin expression in various tissues of C57BL/6J mice. B, quantitative RT-PCR analysis of intectin mRNA expression in various tissues of C57BL/6J mice. C, nucleotide and deduced amino acid sequence of intectin. Putative N- and C-terminal signal sequences are underlined, and the potential GPI-anchoring site is Asn-88 (N) (bold). D, hydropathy profiles of intectin drawn by using the Kyte and Doolittle program (11).

 



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  		FIG. 2.
Amino acid alignments between intectin and other mouse Ly-6 proteins. The amino acid sequence of intectin is aligned with other known mouse Ly-6 proteins. The highly conserved 10 cysteines in mouse Ly-6 proteins are indicated by bold boxed letters.

 
We established a stable cell line that expressed intectin by transfecting the intectin-FLAG gene into a normal intestinal epithelial cell line, IEC-6. Expression of intectin protein in intectin-transfected IEC-6 (IEC-intectin) and mock-transfected IEC-6 (IEC-mock) are shown in Fig. 3A. IEC-intectin cells expressed intectin protein in the cell membrane fraction, and intectin protein was not detected in the cytosolic fraction (Fig. 3A). These results suggest that intectin can be considered a membrane-associated protein. As described above, intectin protein has a predicted GPI-anchoring site in Asn-88. To clarify whether intectin is a GPI-anchored protein, we treated the cells with phosphatidylinositol-specific phospholipase C (PI-PLC) exhibiting activity to excise GPI-anchored region. Without PI-PLC treatment, IEC-intectin cells exhibited higher FITC signal on cell membrane, compared with IEC-mock cells (Fig. 3B). PI-PLC treatment of IEC-intectin cells decreased the signal to the level of IEC-mock cells (Fig. 3B). The same results were observed in FLAG-tagged intectin-transfected CHO cells as well as FLAG-tagged CD59, a known GPI-anchored protein, transfected CHO cells (data not shown). These results indicate that intectin is a GPI-anchored protein.



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  		FIG. 3.
Expression of FLAG-tagged intectin on IEC-6 cell surface and sensitivity to PI-PLC. A, location of intectin protein. The membrane fraction and cytosolic fraction of IEC-mock cells and IEC-intectin cells were subjected to Western blotting using anti-FLAG antibody. FLAG-tagged intectin protein is indicated by the solid arrowhead. Nonspecific bands are indicated by the open arrowheads. B, IEC-mock cells and IEC-intectin cells were treated with (dotted line) or without (solid line) PI-PLC for 1 h at 37 °C. Then, the cells were stained with FITC-conjugated anti-FLAG antibody and analyzed by flow cytometry.

 
Exclusive Expression of Intectin in the Small Intestine— Analysis of various tissues of 6-week-old C57BL/6J mice by Northern blotting and quantitative RT-PCR showed that intectin mRNA was expressed exclusively in the small intestine (Fig. 1, A and B). Unlike other known members of the Ly-6 family, which are highly expressed in peripheral blood leukocytes and in lymphoid organs, intectin mRNA expression was not detected in the bone marrow, WR19L (mouse T cell line), WEHI3 (mouse macrophage-like cell line), and BaF3 (mouse pro-B cell line) cells by quantitative RT-PCR (data not shown).

In situ hybridization was performed to determine the distribution of intectin mRNA in the intestine. Remarkably, a strong signal was detected only at the villus tips of the duodenum, jejunum, and ileum (Fig. 4, B, E, H, and N), and only a weak signal was detected in the colon (Fig. 4K), and a trace signal was found in the stomach and esophagus (data not shown). No signal was detected in all other tissues examined, including the brain, spleen, heart, lungs, eyes, lymph nodes, thyroid, skeletal muscles, thymus, liver, WAT, BAT, pituitary, adrenals, urinary bladder, gallbladder, and testes, by in situ hybridization (data not shown). Thus, intectin mRNA was exclusively expressed in the villus tips of the epithelial cells of the mouse small intestine. In this regard, Groos et al. (6) reported the exclusive presence of apoptotic cells at the villus tips in human and rat small intestine epithelium. Thus, it is likely that the intectinpositive region is the same as that where apoptosis was identified by Groos et al. (6).



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  		FIG. 4.
Intectin mRNA expression pattern in the mouse intestine. In situ hybridization (ISH) revealed intectin antisense-specific signal (black-purple) at the villus tips of the duodenum (A–C), jejunum (D–F), and ileum (G–I) and in the covering epithelium of the colon (J–L) of C57BL/6 mouse (arrowheads). A, D, G, and J: hematoxylin-eosin stain; B, E, H, and K: ISH with intectin antisense probe; C, F, I, and L: intectin sense probe. Higher magnification of the villus tips of the ileum (M: H&E stain, N: ISH using intectin antisense probe and counter-stained with hematoxylin (blue)) shows intectin antisense signal (black-purple) at the villus tips (arrowheads). Scale bar in L = 100 µm for A–L sections. Scale bar in N = 20 µm for M and N sections.

 
Intectin mRNA Expression Is Modulated by Nutritional Status—We next examined the nutritional regulation of intectin mRNA expression in vivo (Fig. 5A). Intectin mRNA expression in the small intestine was significantly decreased by 24-h fasting and restored to basal level by 1- to 2-h refeeding (Fig. 5A). These results indicate that intectin mRNA expression is modulated by nutritional stimuli. This conclusion was supported by our finding in db/db mice, whose daily food intake is 1.5- to 2-fold higher than that of age-matched C57BL/6J mice (data not shown); intectin mRNA expression in the small intestine of 6-week-old db/db mice was significantly higher than ad libitum-fed C57BL/6J mice (Fig. 5B).



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  		FIG. 5.
Nutritional modulation of intectin mRNA expression in mice. A, modulation of intectin mRNA by fasting and refeeding. In these experiments, 6-week-old male C57BL/6J mice were fed ad libitum, fasted for 24 h, or fasted for 24 h and then re-fed 1, 2, and 4 h later (n = 3–4 mice for each group). Then, mice were sacrificed under anesthesia, and the small intestines were harvested immediately. Total RNA was extracted for real-time RT-PCR using intectin specific primers. Values are normalized to the level of cyclophilin mRNA and expressed as mean ± S.E. (n = 3–4). *, p < 0.05. B, comparison of intectin mRNA expression between C57BL/6J mice and genetically obese db/db mice. Intestines were excised from 6-week-old db/db mice and age-matched ad libitum fed C57BL6J mice. The mRNA was quantified by real-time RT-PCR as described above. Values are normalized to the level of cyclophilin mRNA and expressed as mean ± S.E. (n = 6). *, p < 0.05, compared with ad libitum fed C57BL/6J mice.

 
Involvement of Intectin in the Rapid Turnover of Intestinal Mucosa—Based on its specific expression at the villus tips of the intestine and active nutritional regulation, we hypothesized that intectin could be involved in the rapid turnover of intestinal mucosa. To test our hypothesis, we first compared the viability of IEC-intectin cells and IEC-mock cells. Cell viability assay was performed after 12-h treatment with palmitate, a nutrient-derived inducer of apoptosis, or 24-h treatment with camptothecin and daunorubicin, chemical inducers of apoptosis. Palmitate treatment decreased cell viability of both IEC-intectin cells and IEC-mock cells, but the effect was significantly more pronounced in IEC-intectin cells than IEC-mock cells (Fig. 6A). For example, 6.3 µM palmitate reduced cell viability of IEC-intectin cells to 50%, whereas a similar change in cell viability of IEC-mock cells required 50 µM palmitate. On the other hand, there was no difference in cell viability between IEC-intectin cells and IEC-mock cells after treatment with camptothecin or daunorubicin (data not shown).



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  		FIG. 6.
Effects of palmitate on apoptosis of IEC-6 cells. A, effect of palmitate on viability of IEC-6 cells. IEC-intectin and IEC-mock cells were treated with palmitate. After 12-h treatment, the cells were treated with WST-8 and incubated for 1 h at 37 °C. The optical densities were measured at 450 and 650 nm. Values are expressed as mean ± S.E. (n = 3). *, p < 0.05. B, effect of palmitate on cell cycle. The cells were incubated with 0, 25, 50, or 100 µM palmitate for 12 h, and fixed with 70% cold ethanol. Then the samples were treated with 10 µg/ml RNase, stained with 10 µg/ml propidium iodide, and analyzed by flow cytometry. The sub-G1 range is indicated by a horizontal bar above which the percentage of nucleicontaining the sub-G1 amount of DNA is indicated. C, IEC-intectin and IEC-mock cells were incubated with 0, 25, 50, 100, or 200 µM palmitate for 12 h. Caspase-3 activity was measured as described under "Experimental Procedures." Values are expressed as mean ± S.E. (n = 4). *, p < 0.05, compared with IEC-mock cells. D, cell aggregation assay. Parental CHO cells and CHO-intectin cells were subjected to aggregation assay as described under "Experimental Procedures." Cells remaining as single cells were counted at indicated periods of time. The number of cells present in aggregates was also counted. Values are expressed as mean ± S.E. (n = 4). *, p < 0.05, compared with parental CHO cells.

 
Intectin Induces Apoptosis of Intestinal Mucosal Cells via Caspase-3 Activation—Next, we compared the effects of palmitate on cell cycle in IEC-intectin cells and IEC-mock cells. For this purpose, the cells were incubated with palmitate and then stained with PI and sorted according to DNA content. As shown in Fig. 6B, cell cycle analysis by flow cytometry indicated that palmitate dose-dependently decreased the G1 population, whereas it increased the sub-G1 population and the cell death fraction, including apoptotic cells. These palmitate-induced changes in the cell cycle toward cell death were more pronounced in IEC-intectin cells compared with IEC-mock cells. For example, the majority of IEC-mock cells were in the G1 or G2 phase, while the majority of IEC-intectin cells were in the sub-G1 phase when these cells were incubated in the presence of 50 or 100 µM palmitate (Fig. 6B).

To further examine the possible involvement of intectin in the rapid turnover of intestinal mucosa through cell death and especially through apoptosis, we measured caspase-3 activity in palmitate-treated IEC-intectin cells and IEC-mock cells (Fig. 6C). Because activation of caspase-3 is a common downstream effector of diverse apoptotic pathways (13), we measured cleavage of colorimetric substrate, specific to caspase-3. Low dose palmitate significantly increased caspase-3 activity in IEC-intectin cells compared with in IEC-mock cells. Taken together, the above results suggest that intectin expression causes activation of caspase-3, which in turn induces apoptotic death of IEC-6 cells.

Intectin Reduces Cell-to-Cell Adhesion—Previous studies suggested that some members of the Ly-6 family are involved in cell-cell adhesion (14, 15). Therefore, we conducted a cell aggregation assay using parental CHO cells (CHO) and CHO cells stably expressing intectin (CHO-intectin). Intectin expression significantly attenuated CHO cell aggregation (Fig. 6D), suggesting that intectin expression weakens the cell-cell adhesion probably to promote the exfoliation of intestinal epithelial cells.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The intestinal mucosa is continuously exposed to various toxic factors, including enteropathogenic microorganisms and food antigens. It also has to face potent digestive enzymes present in the lumen secreted from the liver and pancreas. In such harmful environments, the intestinal mucosa exhibits a rapid turnover, which serves to maintain the essential functions of the intestine and the integrity of the intestinal wall. The regulation of this rapid turnover of intestinal mucosa is complex and controlled by several factors. Apoptosis is one important factor that determines this process (16, 17). Therefore, identification of the regulatory factors of intestinal mucosal apoptosis should help us understand the mechanism of intestinal mucosal turnover.

In the present study, we identified intectin using an efficient SST technique. Although the precise physiological significance of intectin has yet to be determined, several important features were defined in the present study. 1) Intectin is a new member of GPI-anchored Ly-6 family; 2) intectin mRNA is exclusively expressed in the villus tips of the small intestine, which are known to undergo apoptosis; 3) intectin mRNA expression is influenced by nutritional changes; 4) intectin expression increases the sensitivity of intestinal epithelial cells to palmitate-induced apoptosis; and 5) intectin expression significantly attenuates cell aggregation. These findings suggest that intectin might modulate nutrition-dependent apoptosis, weaken cell-cell attachment, and promote the dying cells to exfoliate into the lumen at the final stage of intestinal mucosal turnover.

The intectin gene is located on murine chromosome 15, similar to murine Ly-6 genes (18). The intectin protein is composed of 111 amino acid residues and has an Ly-6 domain, which is defined by a distinct disulfide-binding pattern between 10 cysteine residues. Furthermore, intectin protein has N-terminal and C-terminal signal sequences, and the result of PI-PLC treatment of intectin-expressing cells indicated that intectin was a GPI-anchored protein, a hallmark of Ly-6 family. These findings indicate that intectin is a novel member of the Ly-6 family. Previous studies suggested the involvement of Ly-6A/E in T-cell activation (19, 20) and development (21). Ly-6A/E and Ly-6C have been shown to regulate cell adhesion (14, 15), but the in vivo functions of Ly-6 family are unknown with the exception of CD59 and urokinase-type plasminogen activator receptor. CD59 functions as a membrane inhibitor of reactive lysis, and failure to express CD59 is related to the pathogenesis of paroxysmal nocturnal hemoglobinuria (PNH). In PNH, acquired somatic defect of the PIG-A gene results in a defect in GPI-anchored proteins expression, including CD59, on the cell surface, making blood cells more susceptible to host complement-mediated lysis (22, 23). These patients are more susceptible to leukemia (24) due to resistance to apoptosis caused by PIG-A gene mutations, suggesting that some of GPI-anchored proteins are important in modulating apoptosis (25). Considered together with the results of the present study, it is conceivable that intectin, a member of the GPI-anchored Ly-6 family proteins, is involved in the regulation of intestinal epithelial cell apoptosis.

Induction of apoptosis by palmitate has been reported in various cells, including pancreatic {beta}-cells (26), cardiomyocytes (27), and hematopoietic cells (28). In the present study, we demonstrated that palmitate induced apoptosis of intestinal epithelial cells. Furthermore, recent studies reported nutritional modulation of intestinal mucosal apoptosis (29–31). Raab et al. (29) showed that high energy diet and purines in the diet induced intestinal mucosal apoptosis. In addition, Sukhotnik et al. (30) reported that exposure to low fat diet decreased intestinal epithelial cell apoptosis in a rat model of short bowel syndrome. These reports implicate nutrient-derived fat should induce apoptosis of intestinal epithelial cells. Groos et al. (31) also showed that apoptosis of intestinal epithelial cells at the villus tips was markedly reduced in subjects receiving total parenteral nutrition over 2 weeks compared with enterally nourished subjects, suggesting that food components seem to influence apoptosis of intestinal epithelial cells. The precise molecular mechanism that regulates apoptosis of intestinal mucosa in response to nutritional conditions has not yet been established. Since intectin mRNA expression was restored as fast as 1 h after refeeding following 24-h fasting, intectin expression is more likely regulated by food-related physical stimuli rather than by endocrine factors such as insulin. Palmitate is a major fatty acid in fat diet and is probably involved in fat diet-induced apoptosis of intestinal mucosa. Our finding that intectin accelerated palmitate-induced apoptosis of intestinal epithelial cells suggests that diet-induced expression of intectin might mediate this diet-induced apoptosis of intestinal mucosa.

Apoptotic cells are observed at the villus tips of intestinal mucosa. In certain inflammatory conditions, such as celiac disease, nematode infections, and graft-versus-host disease, the numbers of apoptotic nuclei were increased in villus epithelial cells (32–35). In addition, the involvement of dysregulation of the apoptotic process in intestinal epithelial cells in inflammatory bowel diseases and colon cancer has also been suggested (36, 37). These studies indicate that apoptosis plays some role in pathological conditions as well as in physiological turnover of villus epithelial cells. In this regard, whether intectin is involved in the pathology of inflammatory bowel diseases or colon cancer remains to be elucidated.

In conclusion, we described in this study the identification of intectin, a novel intestine-specific GPI-anchored protein. This protein enhanced palmitate-induced apoptosis of intestinal epithelial cells.


   FOOTNOTES
 
* This work was supported by The Suzuken Memorial Foundation, The Nakajima Foundation, the Kanae Foundation for Life and SocioMedical Science, The Tokyo Biochemical Research Foundation, the Takeda Medical Research Foundation, Uehara Memorial Foundation, the Takeda Science Foundation, the Novartis Foundation (Japan) for the Promotion of Science, The Cell Science Research Foundation, The Mochida Memorial Foundation for Medical and Pharmaceutical Research, a Grant-in-Aid from the Japan Medical Association, The Naito Foundation, a grant from the Japan Heart Foundation Research, the Kato Memorial Bioscience Foundation, the Japan Research Foundation for Clinical Pharmacology, a grant from the Ministry of Health, Labor, and Welfare, Japan, and grants from the Ministry of Education, Culture, Sports, Science, and Technology, Japan. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

b Both authors contributed equally to this work. Back

c Present address: Tsukuba Research Institute, Banyu Pharmaceutical Co., Ltd., Okubo 3, Tsukuba 300-2611, Japan. Back

h To whom correspondence may be addressed: Dept. of Medicine and Pathophysiology, Graduate School of Frontier Bioscience, Graduate School of Medicine, Osaka University, 2-2 Yamadaoka, Suita, Osaka 565-0871, Japan. Tel.: 81-6-6879-3272; Fax: 81-6-6879-3279; E-mail: mmatsuda{at}fbs.osaka-u.ac.jp. j To whom correspondence may be addressed: Dept. of Internal Medicine and Molecular Science, Graduate School of Medicine, Osaka University, 2-2 Yamadaoka, Suita, Osaka 565-0871, Japan. Tel.: 81-6-6879-3730; Fax: 81-6-6879-3739; E-mail: ichi{at}imed2.med.osaka-u.ac.jp.

1 The abbreviations used are: SST-REX, signal sequence trap by retrovirus-mediated expression screening system; BSA, bovine serum albumin; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; GPI, glycosylphosphatidylinositol; PI, propidium iodide; PIG-A, phosphatidylinositolglycan-class A; PI-PLC, phosphatidylinositol-specific phospholipase C; PNH, paroxysmal nocturnal hemoglobinuria; FITC, fluorescein isothiocyanate; PBS, phosphate-buffered saline; DIG, digoxigenin; CHO, Chinese hamster ovary cells; RT, reverse transcription; ISH, in situ hybridization. Back


   ACKNOWLEDGMENTS
 
We thank the members of Shimomura's laboratory for the helpful discussions. We thank Drs. Yusuke Maeda and Taro Kinoshita (Osaka University) for kindly providing the pME-puro-FLAG-CD59 vector and for the helpful suggestions. We also thank Drs. Rikinari Hanayama and Shigekazu Nagata (Osaka University) for kindly providing the expression plasmid pEF-BOS and for the invaluable suggestions.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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