Cultured monolayers of the dog jejunum with the structural and functional properties resembling the normal epithelium

Xing-He Weng, Klaus W. Beyenbach, and Andrea Quaroni

Department of Biomedical Sciences, Cornell University, Ithaca, New York

Submitted 12 December 2003 ; accepted in final form 12 November 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The development of a culture of the normal mammalian jejunum motivated this work. Isolated crypt cells of the dog jejunum were induced to form primary cultures on Snapwell filters. Up to seven subcultures were studied under the electron microscope and in Ussing chambers. Epithelial markers were identified by RT-PCR, Western blot, and immunofluorescent staining. Confluent monolayers exhibit a dense apical brush border, basolateral membrane infoldings, desmosomes, and tight junctions expressing zonula occludens-1, occludin-1, and claudin-3 and -4. In OptiMEM medium fortified with epidermal growth factor, hydrocortisone, and insulin, monolayer transepithelial voltage was –6.8 mV (apical side), transepithelial resistance was 1,050 {Omega}·cm2, and short-circuit current (Isc) was 8.1 µA/cm2. Transcellular and paracellular resistances were estimated as 14.8 and 1.1 k{Omega}·cm2, respectively. Serosal ouabain reduced voltage and current toward zero, as did apical amiloride. The presence of mRNA of {alpha}-epithelial Na+ channel (ENaC) was confirmed. Na-D-glucose cotransport was identified with an antibody to Na+-glucose cotransporter (SGLT) 1. The unidirectional mucosa-to-serosa Na+ flux (19 nmol·min–1·cm–2) was two times as large as the reverse flux, and net transepithelial Na+ flux was nearly double the amiloride-sensitive Isc. In plain Ringer solution, the amiloride-sensitive Isc went toward zero. Under these conditions plus mucosal amiloride, serosal dibutyryl-cAMP elicited a Cl-dependent Isc consistent with the stimulation of transepithelial Cl secretion. In conclusion, primary cultures and subcultures of the normal mammalian jejunum form polarized epithelial monolayers with 1) the properties of a leaky epithelium, 2) claudins specific to the jejunal tight junction, 3) transepithelial Na+ absorption mediated in part by SGLT1 and ENaC, and 4) electrogenic Cl secretion activated by cAMP.

canine jejunal cell culture; epithelial sodium channel; sodium-glucose cotransporter 1; sodium absorption; chloride secretion


OUR UNDERSTANDING OF TRANSPORT across the intestine rests largely on the study of intestinal segments perfused in vivo, excised segments in vitro, organ cultures, epithelial cell cultures, and, most recently, knockout animal models. The details of intestinal transport mechanisms and their regulation have been elucidated most clearly in cultured intestinal cells such as Caco-2 (1, 2, 21), T84 (27), HT-29 (24), COLO 205 (19), SW620 (52), and other epithelial cell lines derived from cancerous tissue. Only a few nontransformed intestinal epithelial cells (IEC) are available, such as IEC-6 and IEC-18 derived from the normal small intestine of the rat (32, 33, 42) and the IPEC-J2 cell line established from newborn piglet jejunum (44).

In the present study, we introduce a new primary culture of the normal jejunum with a focus on the method of producing that culture from crypt cells isolated from the dog jejunum. In addition, we provide a preliminary structural and functional characterization of the epithelial monolayer. Primary cultures and up to seven subcultures consistently formed confluent epithelial monolayers on Snapwell filters. The monolayers exhibit the morphological and functional polarization expected of the normal jejunum, including a prominent apical brush border. The tight junction presents claudin-3 and -4 as in the normal jejunum. The expression of epithelial Na+ channels (ENaC) and an amiloride-sensitive short-circuit current (Isc) can be attributed to the presence of culture-stimulating agents (epidermal growth factor, hydrocortisone, insulin), because this current disappears in plain Ringer solution lacking culture-stimulating agents. Fortuitously, the expression of an amiloride-sensitive Isc allows an estimate of the electrical resistance of transcellular and paracellular pathways. The estimates reveal a leaky epithelium with a paracellular pathway 13 times as conductive as the transcellular pathway. Measures of the unidirectional isotopic Na+ fluxes confirmed the leaky nature of the cultured monolayers and pointed to Na+ transport systems in addition to that mediated by ENaC. One such transporter is the Na+-D-glucose cotransporter SGLT1 identified by Western blot. When electrogenic Na+ absorption via ENaC is minimized by the use of plain Ringer solution containing mucosal amiloride, the addition of dibutyryl-cAMP (DBcAMP) to the serosal side activated a Cl-dependent Isc consistent with the stimulation of transepithelial Cl secretion. The culture can be studied for hours in Ussing chambers, thus affording detailed investigations of jejunal transport across a single layer of epithelial jejunal cells under well-defined experimental conditions.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Epithelial cell isolation and culture. In general, the method to establish primary cultures from the small intestine of adult beagle dogs followed the procedures described for fetal human intestine (39). Segments of the small intestine were obtained from adult beagle dogs (1–2 yr old) that were killed for cardiac electrophysiological studies, as described by Fox et al. (14) and approved by the Center for Research Animal Resources at Cornell University. Segments of the jejunum (30–50 cm long) were removed and placed on ice. The lumen was washed four times with 50 ml ice-cold sterile PBS (in mM: 136.9 NaCl, 2.7 KCl, 1.5 KH2PO4, and 6.5 Na2HPO4), filled with a 1.25% trypsin-0.5 mM EDTA solution, clamped at the two ends, and incubated at room temperature for 10 min. The segments were then cut open longitudinally and laid flat on a sterile glass plate on ice, and the apical surface was scraped gently with a razor blade to remove mucus and most of the villi. The scrapings were discarded. Harder scraping, leaving behind only serosal layers, yielded intact crypts embedded in fragments of surrounding tissue that were collected in 50-ml centrifuge tubes containing serum-free OptiMEM (Invitrogen-GIBCO, Carlsbad, CA) supplemented with 100 U/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml amphotericin B (antibiotic-antimycotic solution; Invitrogen-GIBCO-BRL). After centrifugation at 1,000 rpm (220 g), the supernatant was discarded. The pellet was washed three times with serum-free OptiMEM and then incubated with serum-free OptiMEM containing 0.4 mg/ml collagenase type IV (filter sterilized; Sigma, St. Louis, MO) at 37°C for 45–60 min, with gentle and brief shaking every 10–15 min. The dissociated epithelial cells were spun down at 1,000 rpm (220 g) and washed five times with 40 ml of 10% FBS (Hyclone Laboratories, Logan, UT) in DMEM supplemented with antibiotic-antimycotic solution. After each centrifugation at 1,000 rpm, the pellet was resuspended by pipetting up and down vigorously using a 10-ml wide-bore Falcon pipette (Milian, Gahana, OH). The final pellet was suspended in 100–250 ml "fortified OptiMEM" (vide infra). After large fragments were allowed to sediment within 1–2 min, the upper two-thirds of the suspension were plated in 100-mm-diameter dishes coated with extracellular matrix ECL (Upstate Biotechnology, Lake Placid, NY) and incubated at 37°C and 6% CO2.

For the culture of monolayers and for their initial physiological characterization, we used fortified OptiMEM, which we define as serum-free OptiMEM supplemented with 10 mM HEPES, pH 6.5, 2.5 mM glutamine, 50 U/ml penicillin, 50 µg/ml streptomycin, 2 mM GlutaMAX-I (Invitrogen-GIBCO-BRL), 20 ng/ml epidermal growth factor (human recombinant; Upstate Biotechnology), 150 nM hydrocortisone 21-hemisuccinate sodium salt (Sigma), 10 µg/ml insulin (human recombinant; Sigma), and 4% FBS. After 3 h, fortified OptiMEM was aspirated, and the attached cells (consisting almost exclusively of intact or large portions of crypts) were rinsed four times, refed with fortified OptiMEM, and incubated at 37°C. After 24 h, the incubator temperature was changed to 32–34°C for the rest of the culture period.

After the primary cultures had grown in 100-mm-diameter dishes, between 300,000 and 400,000 cells in 0.5 ml fortified OptiMEM were subcultured onto Snapwell permeable filters (insert growth area 1.13 cm2, 0.4 µm pore size; Costar, Cambridge, MA) and incubated at 32°C and 6% CO2 to form dog intestinal epithelial cell (DIEC) monolayers.

We evaluated the growth of DIEC monolayers by visual inspection under the microscope (Nikon Diaphot) and by daily measurements of the resistance across the filter area with a hand-held Millicell-ERS volt- and ohmmeter (Millipore, Billerica, MA). Because this resistance measurement (between two points with heterogeneous current distribution) is meant to monitor culture growth to confluence, we call this resistance "growth area resistance" (see Fig. 4e). For measurements of the transepithelial resistance (Rt) under more homogeneous conditions of transepithelial current distribution, we used current-voltage (I-V) plots measured across confluent monolayers in the Ussing chamber.



View larger version (132K):
[in this window]
[in a new window]
 
Fig. 4. Scanning electron micrograph of DIEC monolayers grown in fortified OptiMEM at 34°C on plastic dishes (a and b) and on Millicell HA filters (c and d). Monolayers grown on plastic are flat with a sparse apical layer of microvilli, whereas monolayers on Millicell HA filters are tall and covered with a dense brush border. e: Electrical resistance as a measure of DIEC growth to confluence in fortified OptiMEM. Resistance was measured daily across cultures grown on Millicell HA filters and Snapwells. DIEC monolayers were confluent 6–7 days after seeding crypt cells.

 
Immunocytochemistry. Cells of primary cultures or passages 1 to 3 were washed three times with PBS, fixed with 3% formaldehyde, and then directly processed for immunofluorescence staining or permeabilized by one of the following two methods: 1) incubation with acetone-methanol 1:1 at –20°C for 10 min; or 2) lysis with 0.2% Triton X-100 in PBS for 2 min at room temperature. Further processing steps are described by Tian and Quaroni (49). The secondary antibodies were FITC- or rhodamine-conjugated goat anti-mouse or donkey anti-rabbit IgG (Boehringer Manheim, Indianapolis, IN) diluted 1:25 in PBS plus 2% BSA. Cells were counterstained with 0.01% Evans blue and 2 µg/ml DAPI for 2 min. After incubation with antibody and washing, the cells were mounted in glycerol-PBS (9:1) and 2.5% 1,4-diazabicyclo-[2.2.2]octane and covered with coverslips. Stained cells were examined with a Zeiss Axiovert 10 microscope equipped with epifluorescence optics and an Optronics three-chip CCD camera. Digital images were processed with Adobe Photoshop software. We used the following primary antibodies: 1) CY-90 (Sigma) against keratin 18, mouse monoclonal, diluted 1:400; 2) anti-desmosomal protein, mouse monoclonal (Sigma), diluted 1:400; 3) anti-human occludin, mouse monoclonal, OC-3F10 (Zymed catalog no. 33–1500; Zymed), diluted 1:1,000; 4) anti-claudin-1, rabbit polyclonal (JAY.8, catalog no. 51–9000; Zymed), diluted 1:1,000; 5) anti-claudin-2, rabbit polyclonal (MH44, catalog no. 51–6100; Zymed), diluted 1:1,000; 6) anti-claudin-3, rabbit polyclonal (Z23.JM, catalog no. 34–1700; Zymed), diluted 1:1,000; 7) anti-claudin-4, mouse monoclonal (3E2C1, catalog no. 32–9400; Zymed), diluted 1:2,000; 8) anti-claudin-5, rabbit polyclonal (rabbit Z43.JK, catalog no. 35–2500; Zymed), diluted 1:1,000; and 9) anti-claudin-5, mouse monoclonal (4C3C2, catalog no. 34–1600; Zymed), diluted 1:2,000.

Cells were processed for transmission and scanning electron microscopy as previously described (41).

Western blot. Total cell lysates solubilized in SDS-PAGE sample buffer were subjected to SDS-PAGE and Western blot as described previously (49). The DNA concentration was determined using the Hoechst-33258 DNA assay and a mini-fluorometer (Hoefer; Pharmacia Biotech, Piscataway, NJ). The amount of cell lysate applied to each well was normalized to DNA (39). Lysates obtained from 0.2 x 106 cells/sample were subjected to SDS-PAGE on a 7.5–10% acrylamide gel. The proteins were transferred to a nitrocellulose membrane (High-bond nitrocellulose; Amersham Life Science, Arlington Heights, IL) using a transblot system (Bio-Rad, Hercules, CA) at 100 V and 5°C for 90 min. The membranes were then blocked in 80 mM Na2HPO4, 20 mM NaH2PO4·2H2O, 100 mM NaCl, and 0.1% Tween 20 containing 3% BSA at 4°C overnight. Incubation with antibodies, washing, protein detection, antibody stripping, and reprobing were performed according to the ECL-Plus Western blotting protocol from Amersham Life Science. Blots were scanned with a Molecular Dynamics Storm 840 scanner (Amersham Pharmacia Biotech, Sunnyvale, CA) in the fluorescence mode. In other experiments, we employed fluorescent secondary antibodies and the Li-COR Odyssey Infrared Imaging System (Lincoln, NE). After transfer of the proteins to a Hybond-P/polyvinylidene difluoride (PVDF) membrane (Amersham Pharmacia Biotech) and blocking for 4 h in Odyssey Blocker, incubation with primary antibodies was done in Odyssey Blocker containing 0.1% Tween 20 overnight at 4°C. After four washes with Tris-buffered saline (TBS: 20 mM Tris base, 137 mM NaCl, pH 6) containing 0.1% Tween 20, the blots were incubated with Alexa Fluor 680 goat anti-mouse IgG (1:2,500 dilution in Odyssey Blocker, 0.1% Tween 20) for 1 h at room temperature and shielded from light. After being washed, the blots were scanned in the Odyssey Infrared scanner. The primary antibodies used are the same as listed above.

In a separate experiment, the presence of SGLT1 in DIEC cell lysates was examined by Western blot with an antibody specific to rabbit SGLT1. In these studies, SDS-PAGE and the transfer of protein from gel to PVDF membrane was done as described previously (55). After the membrane was incubated for 2 h in blocking solution consisting of TBST (20 mM Tris·HCl, 137 mM NaCl, and 0.1% Tween, pH 7.6) fortified with 3% BSA, the membrane was then treated for 2 h with antibody diluted in TBST plus 1% BSA (1:1,000). To verify immunoreactivity of SGLT1, the antibody 8821 was first preincubated with 0.5 µg/ml of the immunizing peptide for 2 h, which yielded a negative Western blot. Both antibody 8821 and immunizing peptide were gifts of Dr. E. M. Wright (University of California Los Angeles). The antibody 8821 has been used successfully in the past in investigations of SGLT1 in the dog jejunum in vivo (22).

Electrophysiological studies in Ussing chambers. Electrophysiological measurements were performed on DIEC monolayers mounted in Ussing chambers (CHM5; World Precision Instruments, Sarasota, FL). Monolayers of passage 1 through 7 were studied. Finding consistent electrophysiological properties among them, we did most experiments on passages 1 to 3.

In most transport experiments, fortified OptiMEM (10 ml) was present on both sides of the epithelium. Using the methods of atomic absorption spectrophotometry, flame photometry, and coulometry, we measured the following partial composition of the OptiMEM culture medium: Na+ concentration, 157.0 ± 3.7 mM (n = 3); K+ concentration, 4.6 ± 0.3 mM (n = 3); Cl concentration, 123.7 ± 4.7 mM (n = 3); and protein, 0.56 ± 0.01 mg/ml. According to the manufacturer of OptiMEM culture medium, the glucose concentration is 13.9 mM (personal communication); other ingredients of OptiMEM culture medium are proprietary information.

In experiments examining the ion dependence of the measured Isc, confluent monolayers were taken from their cultures in fortified OptiMEM and mounted in plain Ringer solution lacking culture-stimulating agents. Plain Ringer solution contained (in mM): 120 NaCl, 5 KCl, 25 NaHCO3, 1.2 CaCl2, 1.2 MgCl2, and 5 glucose, pH 7.3. Low-Cl Ringer was prepared by replacing 116.8 mM NaCl with sodium isothionate. Glucose-free Ringer solution contained no glucose.

Mucosal and serosal solutions were circulated at a rate of 0.2 ml/s by air-lifting with 95% O2-5% CO2. The pH was maintained between 7.35 and 7.38. The temperature was maintained at 32°C with a water bath (Neslab Instruments, Portsmouth, NH). For voltage and current recording, we used the multichannel voltage/current clamp of World Precision Instruments (EVC-4000) and Ag/AgCl voltage/current electrodes embedded in 4% agar and 150 mM NaCl. Voltage and current were measured with respect to ground in the serosal compartment. To measure the Rt, we clamped monolayers at 5-mV voltage steps from –20 to +20 mV for 5 s each and recorded the transepithelial current. The Rt was calculated as the inverse of the slope of the I-V plot.

After mounting of the DIEC monolayers in the Ussing chamber, the transepithelial voltage (Vt) was monitored for ~20–30 min until Vt had become stable. Thereafter, the bath temperature was gradually increased to 38°C for the next 5 min. The Isc was measured at a Vt clamp of 0 mV. After Isc had stabilized after ~10 min, experiments were begun. Amiloride (Merck, Whitehouse Station, NJ) was applied to the mucosal or serosal side. The effect of mucosal amiloride concentration on electrical variables was examined in detail, since serosal amiloride had no effect. Ouabain and DBcAMP were purchased from Sigma (St. Louis, MO) and presented from the serosal side only. Amiloride and ouabain were dissolved in DMSO, and stored as stock solutions at 1,000 times their final concentration in the experiment. The experimental DMSO concentration of 0.1% had no effect on measured electrical variables.

Transepithelial Na+ flux measurements. We used 22Na+ (Amersham Biosciences) to measure unidirectional Na+ fluxes across DIEC monolayers. Radioactivity in known and unknown samples was measured with a Beckman gamma counter (Gamma 300). In the typical isotopic flux experiment, the DIEC monolayer was mounted in the Ussing chamber and bathed with fortified OptiMEM on both sides. After Vt had reached steady state at 38°C, the monolayer was voltage clamped at 0 mV and maintained in the short-circuit condition for the rest of the experiment. To begin, 10 µCi 22Na+ were added to the mucosal or serosal side. At intervals of 10 min, 1 ml sample solution was taken from the other side for the measurement of 22Na+ that had crossed the DIEC monolayer. The counted sample was then returned to the side from where it had been taken. After the 40-min flux period, amiloride (10 µM) was added to the mucosal solution, and the flux measurements were repeated, again under short-circuit conditions.

When 1 ml was removed from 10 ml of apical or basolateral solution, the water column dropped by 0.4 cm of water, i.e., 1.5%. The transepithelial hydrostatic pressure difference thus introduced did not cause measurable transepithelial water flows, since water levels remained constant. Moreover, there were no transepithelial streaming potentials, and no effects on Isc and Rt. The unidirectional isotopic Na+ flux from mucosa to serosa was measured in one set of DIEC monolayers and the reverse flux in another set. Accordingly, the net transepithelial Na+ flux is the difference between values of two experimental groups (Table 1).


View this table:
[in this window]
[in a new window]
 
Table 1. Effects of amiloride on unidirectional Na+ fluxes and Isc

 
RT-PCR of dog jejunal {alpha}ENaC mRNA. Total RNA was extracted from DIEC monolayers at the third passage by CsCl banding as described by Kinston (28). The integrity of RNA was verified by agarose gel electrophoresis and ethidium bromide staining, and its quantity was determined spectrophotometrically.

For RT-PCR, 1 µl RNA (1.55 µg/µl) sample was used. The mRNA was reverse-transcribed to cDNA and further amplified by PCR reactions (SuperScript III One-Step RT-PCR System; Invitrogen). The 25-µl reaction mixture for RT-PCR included 12.5 µl of 2x reaction buffer, 1 µl total RNA sample, 1 µl forward primer (10 µM), 1 µl reverse primer (10 µM), 1 µl RT/Taq enzyme mixture, and 8.5 µl H2O. The first cDNA strands were reverse-transcribed at 52°C for 30 min. After denaturation at 94°C for 2 min, the cDNA strands were amplified by PCR reactions for 40 cycles. Each cycle entailed incubation at 94°C for 15 s, then 52°C for 30 s, and 72°C for 45 s. Finally, the DNA strands were extended at 72°C for 5 min. During the reactions, the following primers were used: {alpha}ENaC forward: 5'-AACCTGCCTTTATGGACGAT-3' and {alpha}ENaC reverse: 5'-AGGTTGGACAGGAGGGTGAC-3'.

To identify the PCR product, 10 µl of the PCR product were mixed with 2 µl 6x DNA loading buffer (0.25% bromphenol blue, 0.25% xylene cyanol FF, and 15% Ficoll in H2O), applied to 1.5% agarose gel dissolved in 40 mM Tris-acetate and 1 mM EDTA and separated under 100 V with a horizontal elctrophoresis apparatus (MINI SUB DNA CELL; Bio-Rad). The PCR product was purified and submitted for sequencing with the {alpha}ENaC forward primer to the Bioresource Center of Cornell University.

Statistics. All data are presented as means ± SE. The paired Student's t-test was used for the significant difference (P < 0.05) between control and experimental groups, whereas the z-test was used for the significant difference between specific data and a group mean value (P < 0.05).


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell proliferation and differentiation in primary and secondary cultures. In the present study, we produced 17 primary cultures of DIEC. Some primary cultures were taken to up to seven passages before they lost proliferative activity. As long as the cells could be subcultured, they retained the ability to form confluent monolayers with similar morphological and electrophysiological properties.

Although the culture steps producing DIEC are straightforward and reliable, a few steps proved particularly important. First, it was crucial to start cultures with intact or nearly intact crypts because isolated epithelial cells or fully differentiated villus cells did not attach to the substrate, nor did they survive for more than 2–3 days. As shown in Fig. 1a, essentially intact crypt cells attach to the substrate after only 4 h of incubation, and cells at the border of the cluster start to spread out and migrate. The use of ECL-coated dishes significantly improved (2- to 3-fold) the number of crypt cells attaching to substrate, but ECL-coated dishes were not essential, since qualitatively similar results were obtained with uncoated dishes. Nonepithelial cells, large tissue fragments, and clumps of villus cells did not adhere strongly and were washed out. Proliferative (mitotic) cells could be detected by staining mitotic apparatuses with anti-tubulin antibodies after only one night of incubation. However, mitotic apparatuses were relatively rare and flat and therefore difficult to detect by direct examination of unstained cultures.



View larger version (113K):
[in this window]
[in a new window]
 
Fig. 1. Primary cultures of dog intestinal (jejunal) epithelial cells (DIEC) grown in fortified OptiMEM. a: Crypt cells attach to the ECL-coated dish 4 h after seeding and wash out of nonattached cells; b: same culture as in a 1 day later. Epithelial cells have now attached and spread out without a significant increase in the number of cells; c: 1-day-old culture stained with an immunofluorescent antibody to keratin, demonstrating the epithelial nature of the culture; d: 1-day-old culture stained with a monoclonal antibody (TS23) specific to the glycosylated form of Notch-1, identifying crypt stem cells; e: 7-day-old culture maintained at 35°C, showing the presence of clusters of small proliferative cells (*) among older and larger nonproliferative cells; f: same culture as in e incubated with bromodeoxyuridine (BrdU) for 24 h to visualize proliferative cells (green-blue). BrdU incorporated into cellular DNA was visualized by immunofluorescence staining. Bar = 25 µm in a–f.

 
Figure 1b shows that the crypt cells had spread out entirely to occupy a much larger surface after 1 day. The staining of these cultures with anti-keratin antibodies confirmed the epithelial nature of all, or nearly all cells (>99.9% in 12 samples) present in these primary cultures (Fig. 1c). Contamination by fibroblasts and other cell types was negligible. Staining with a monoclonal antibody (TS23) specific for a glycosylated form of Notch-1 expressed in human and rat intestinal stem cells (Quaroni and Beaulieu, unpublished observation) indicated the presence of a subpopulation of stem cells in the primary cultures (Fig. 1d).

A second crucial step proved to be temperature control. When cultures were maintained at 37°C, they remained viable for at least 3–4 wk, but cell proliferation ceased entirely after 1 wk. In contrast, when cultures were grown at 32–34°C starting on day 2, regions containing rapidly proliferative cells became evident after 3–4 days (Fig. 1e). Proliferative cells incorporated bromodeoxyuridine, demonstrating rapid cell division and expansion in the cell population (Fig. 1f). Older, nonproliferative cells became enlarged and apoptotic after ~2 wk. By that time, the smaller proliferative cells represented the main cell type that could also be serially passaged. We have estimated a population doubling time to be 18–27 h in nine cultures. Proliferative cells could be readily subcultured at least six times, yielding epithelial cells that were characterized and used in all subsequent studies. We have called these cells DIEC.

When confluent, DIEC maintained a typical epithelial morphology and formed tightly confluent DIEC monolayers with frequent, obvious domes when grown on solid (plastic) support, indicating transepithelial ion and water transport in the direction of absorption. The presence of tight junctions could be readily demonstrated by microscopic studies and by Western blotting with antibodies to junctional proteins. As shown in Fig. 2a, of the five claudins examined, only claudin-3 and -4 were detected in primary, first-, and second-passage cultures. In some experiments, small amounts of claudin-1 were also seen. Another tight junction protein, occludin-1, could also be identified by Western blot (Fig. 2a). Claudin-2 and claudin-5 were not detected in any primary culture or passage. In these experiments, {beta}-tubulin was used as a loading control (Fig. 2a).



View larger version (89K):
[in this window]
[in a new window]
 
Fig. 2. Expression of paracellular proteins in dog intestinal epithelial cells (DIEC) grown in fortified OptiMEM. a: Claudins-3 and -4 and occludin-1 were detected by Western blot in cell lysates derived from primary cultures (P) and 1st and 2nd passages of DIEC; b: occludin-1 (green); the arrow points to a cell undergoing mitosis; c: zonula occludens (ZO)-1 (green); d: claudin-4 (green); e: desmosomes (green). The tight junction proteins occludin-1, ZO-1, and claudin-4 were visualized in DIEC of the 2nd passage. Bar = 25 µm in b–e.

 
Immunostaining of primary or secondary cultures localized the expressed junctional proteins to the paracellular pathway. Occludin-1 appeared in a discrete pericellular band as expected from its association with tight junctions (Fig. 2b). Likewise, zonula occludens (ZO)-1 was distributed in a pattern consistent with its presence in tight junctions (Fig. 2c). The distribution of claudin-3 and claudin-4 was more diffuse than that of occludin and ZO-1. Claudin-4 and claudin-3 (data not shown) appeared to expand well beyond tight junctions (Fig. 2d). The distribution of desmosomes outlines again the paracellular space of confluent epithelial cells (Fig. 2e).

Electron microscopy. When DIEC were cultured in plastic dishes in the presence of fortified OptiMEM, both primary cultures and subcultures formed monolayers, with the basolateral membrane adhering to the substrate and microvilli on the apical membrane facing the culture medium (Fig. 3a). Next to this polarization, transmission electron micrographs revealed extensive infoldings of the basolateral membrane (Fig. 3, b–e), tight junctions near the apical border (Fig. 3d), and desmosomes near the serosal border (Fig. 3e). Some cells of the DIEC monolayers contain granules in the apical cytoplasm, possibly identifying primitive goblet cells (Fig. 3, a–c). Our attempt to identify goblet cells with antibodies to mucin did not succeed, since the rat and human antibodies apparently did not recognize mucin in the dog. Cells without granules are probably epithelial cells with the properties of transepithelial transport.



View larger version (195K):
[in this window]
[in a new window]
 
Fig. 3. Ultrastructure of DIEC monolayers (2nd passage) grown on Millicell HA filters in fortified OptiMEM. a and b: Monolayer of polarized epithelial cells. Granule-containing cells (arrows) may be pregoblet cells; c: granules in the apical cytoplasm at higher magnification; d: tight junctions (arrow) and basolateral membrane infoldings (*) of two neighboring epithelial cells; e: desmosomes (arrow) in the lower lateral interstitial space near the serosal border and basolateral membrane infoldings (*). a–c were selected to show the presence of two cell types, granulated and nongranulated cells. Other areas of the DIEC monolayers were devoid of granulated cells.

 
Scanning electron micrographs of primary and secondary cultures grown on plastic support for 10 days revealed epithelial cells with sparse apical microvilli (Fig. 4, a and b). Adjacent epithelial cells overlap, hiding their tight junctions below their lateral edges. In contrast, cultures grown on Millicell HA filters or Snapwells yielded taller cells with a prominent brush border (Fig. 4, c and d). DIEC monolayers of this type were used in the transport studies described below.

Daily measurements of the electrical resistance across the insert growth area revealed an exponential rise in resistance as DIEC monolayers became confluent (Fig. 4e). After 3–4 days, the electrical resistance increased sharply, reaching peak resistances 6–7 days after seeding the culture. Thereafter, resistance decreased together with increasing variability. In view of the transient nature of the Rt, we confined our study on transepithelial transport in Ussing chambers to monolayers between 4 and 6 days old.

Electrophysiology of DIEC monolayers in fortified OptiMEM culture medium. Primary cultures of DIEC were not studied. Instead, our electrophysiological observations focused largely on confluent DIEC monolayers of the first, second, and third passage after noting no major differences with subsequent passages. Monolayers showing signs of apoptosis or degeneration were not used in electrophysiological studies. Under control conditions in fortified OptiMEM containing culture-stimulating agents (epidermal growth factor, hydrocortisone, and insulin), the DIEC monolayers had an open-circuit voltage (Vt) of –6.8 ± 0.6 mV (n = 36) with a range from –2.0 to –14.4 mV, apical side negative. Apical side positive voltages were never observed. Rt was 1,050 ± 105 {Omega}·cm2 on average with a range from 429 to 2,173 {Omega}·cm2 (n = 22). When DIEC monolayers were voltage clamped at 0 mV, Isc was 8.1 ± 0.4 µA/cm2 (n = 36), with positive current flowing from the apical to the basolateral side. The Isc ranged from 5.3 to 13.3 µA/cm2.

Amiloride-sensitive Isc and expression of ENaC in DIEC monolayers grown and studied in fortified OptiMEM culture medium. The effects of amiloride were studied in the presence of fortified OptiMEM (Fig. 5). The control Vt was –3.6 ± 0.4 mV (n = 10), the Rt was 694 ± 72 {Omega}·cm2 (n = 7), and the Isc was 8.4 ± 0.6 µA/cm2 (n = 10; Fig. 5A). After amiloride (10 µM) was added to the apical side, Vt dropped immediately and significantly to –0.7 ± 0.2 mV (P < 0.001), and Isc significantly decreased to 1.3 ± 0.4 µA/cm2 (P < 0.001, Fig. 5A). The inhibitory effects of amiloride were immediate, requiring 1–2 s to reach full effect. Even though amiloride significantly inhibited the Vt and Isc, it had no significant effect on Rt, which was 694 ± 72 {Omega}·cm2 under control conditions and 687 ± 62 {Omega}·cm2 (n = 7) in the presence of amiloride (Fig. 5A).



View larger version (27K):
[in this window]
[in a new window]
 
Fig. 5. Effect of amiloride on transepithelial electrical variables of DIEC monolayers in fortified OptiMEM. A: amiloride (10 µM) applied to the mucosal solution significantly inhibited transepithelial voltage (Vt) and the short-circuit current (Isc) without an effect on the transepithelial resistance (Rt). B: amiloride applied to the serosal solution had no effect. *P < 0.001. Nos. in parentheses are no. of determinations.

 
A consistent observation was the subsequent decrease of Rt in the presence of amiloride. After amiloride had inhibited Vt and Isc within 1–2 s of application, these two variables remained unchanged in the presence of amiloride. In contrast, ~10–20 min later, Rt began to decrease gradually yet significantly, dropping to 81.3 ± 4.3% (P < 0.001) of control. The nature of this resistance drop was beyond the scope of the preliminary electrophysiological characterization of DIEC monolayers.

Amiloride (10 µM) added to the basolateral solution had no significant effect on Vt, Rt, and Isc (Fig. 5B). In these nine DIEC monolayers, the control Vt was –6.6 ± 0.8 mV, the Rt was 887 ± 82 {Omega}·cm2, and the Isc was 7.2 ± 0.3 µA/cm2. After amiloride was added to the serosal side, Vt decreased to –5.1 ± 0.6 mV, Rt decreased to 731 ± 80 {Omega}·cm2, and Isc decreased to 7.0 ± 0.3 µA/cm2. None of these changes was significant.

We also examined the effect of amiloride on Vt, Isc, and Rt at 32°C, the temperature at which monolayers were cultured. At this temperature, 10 µM amiloride added to the apical side of the monolayer inhibited Vt and Isc, again with no effects on Rt (data not shown).

Figure 6A shows the linear I-V plot measured across DIEC monolayers in the absence and presence of amiloride. The slope of the I-V plots is the transepithelial conductance (1/Rt), and the y-intercept is the Isc. Apical amiloride (10 µM) significantly reduced the Isc from 7.5 ± 0.3 to 0.9 ± 0.2 µA/cm2 (P < 0.001), without affecting transepithelial conductance (1.49 and 1.42 mS/cm2 before and after amiloride, respectively). Significant effects on Vt and Isc but not on Rt suggest that the paracellular conductance is much greater than the transcellular conductance such that changes in transepithelial conductance are undetectable when a transcellular transport pathway is blocked.



View larger version (18K):
[in this window]
[in a new window]
 
Fig. 6. Effect of apical amiloride on transepithelial electrical variables of DIEC monolayers in fortified OptiMEM. A: linear current (I)/voltage (V) plots in the absence and presence of amiloride (10 µM). Amiloride reduces the transepithelial current without affecting the transepithelial conductance. A significant Isc remains at the short-circuit voltage of 0 mV at an amiloride concentration of 10 µM. Data are means ± SE of 7 monolayers. B: concentration-response curve of the effect of amiloride on Isc. The amiloride concentration at half-maximal inhibition (IC50) is 0.76 µM. No significant Isc remains at the amiloride concentration of 100 µM. Vc, voltage clamp; Ic, clamp current. C: expression of {alpha}-epithelial Na+ channel (ENaC) mRNA in DIEC monolayers grown in fortified OptiMEM. The 547-bp cDNA sequence of {alpha}-ENaC localized at the expected size of the amplified sequence.

 
A concentration-response curve of the effects of apical amiloride on Isc shows that inhibitory effects begin at an amiloride concentration of 0.1 µM (Fig. 6B). A four-parameter sigmoid curve was fitted to mean values of Isc, which drops from ~9 µA/cm2 in the absence of amiloride to 0.3 ± 0.2 µA/cm2 in the presence of an amiloride concentration of 100 µM. The amiloride concentration at half-maximal inhibition (IC50) was 0.76 µM. The correlation coefficient of the regression line was 0.9985.

Figure 6C shows the separation of the RT-PCR product of {alpha}ENaC mRNA in agarose gel, revealing a DNA band of 500 bp. After being sequenced, the band turned out to be a DNA strand with 547 bp, 100% identical to the reported canine {alpha}ENaC cDNA (GenBank accession no. AF209748).

Effects of amiloride on transepithelial Na+ fluxes in monolayers grown in fortified OptiMEM culture medium. To examine whether the amiloride-sensitive Isc is carried by Na+, we measured unidirectional, transepithelial isotopic 22Na+ fluxes in the absence and presence of apical amiloride (Table 1).

Monolayers were bathed on both sides with fortified OptiMEM containing culture-stimulating agents. Under control conditions, the unidirectional Na+ flux from the mucosal to the serosal side (Jm->s) was 18.6 nmol·min–1·cm–2 and the reverse flux, from serosa to mucosa (Js->m) was 9.7 nmol·min–1·cm–2. The net transepithelial Na+ flux was 8.9 nmol·min–1·cm–2 from mucosa to serosa, which is equivalent to a current of 14.3 µA/cm2. Because the Isc measured in parallel was only 8.7 ± 1.1 µA/cm2, the transepithelial Na+ flux significantly exceeds the Isc by a factor of 1.6 (z = 5.09, P < 0.001).

In the presence of amiloride (10 µM), the Jm->s Na+ flux remained near control values, 18.9 ± 2.6 nmol·min–1·cm–2 (n = 6). Likewise, Js->m Na+ flux, 11.6 ± 1.7 nmol·min–1·cm–2, remained near control values (n = 6). Accordingly, the net transepithelial Na+ flux was 7.3 nmol·min–1·cm–2, or 11.7 µA/cm2, which was not significantly different from control (Table 1). In contrast, amiloride caused Isc to drop significantly to 1.1 ± 0.2 µA/cm2. Thus amiloride substantially reduced the measured Isc to 13% of control values without affecting the net transepithelial Na+ absorptive flux (Table 1).

Effects of ouabain on the electrophysiology of monolayers grown in fortified OptiMEM culture medium. In monolayers studied in the presence of fortified OptiMEM containing culture-stimulating agents, the addition of ouabain (1 mM) to the basolateral side inhibited Isc and Vt and decreased Rt (Fig. 7). Unlike the immediate effects of amiloride, the effects of ouabain developed slowly. Under control conditions, Vt was –9.7 ± 1.9 mV, Rt was 1,708 ± 151 {Omega}·cm2, and Isc was 7.2 ± 0.8 µA/cm2 in six DIEC monolayers. After addition of ouabain to the serosal solution (1–2 s), Vt significantly decreased to –8.2 ± 1.4 mV (P < 0.001), and Isc significantly decreased to 5.9 ± 0.8 µA/cm2 (P < 0.001). Later (20 min), Vt had decreased further to –0.5 ± 0.3 mV (P < 0.001, compared with control), Isc had fallen to values not significantly different from zero, and Rt had decreased significantly to 1,106 ± 111 {Omega}·cm2 (P < 0.01).



View larger version (22K):
[in this window]
[in a new window]
 
Fig. 7. Effect of serosal ouabain on transepithelial electrical variables of DIEC monolayers in fortified OptiMEM. Upon the addition of ouabain, Vt and Isc decreased initially by –3.2 mV and 1.3 µA/cm2, respectively. Thereafter, Vt and Rt underwent a slow, gradual decrease until 20 min later when Vt and Isc had decreased to 0, and Rt had decreased significantly as well. NM, not measured; Values are means ± SE. *P < 0.01; nos. in parentheses indicate the no. of determinations.

 
Electrophysiological studies of monolayers in plain Ringer solution. When DIEC monolayers cultured in fortified OptiMEM were bathed in Ussing chambers on both sides with plain Ringer solution lacking culture-stimulating agents, the Vt decayed in parallel with a decrease in the Isc (Fig. 8A). Together with four additional experiments, the Vt dropped from –12.5 ± 4.6 to –1.5 ± 0.9 mV (n = 5) in 10–30 min. The Isc dropped from 6.6 ± 1.6 to 2.0 ± 0.6 µA/cm2 (n = 5).



View larger version (23K):
[in this window]
[in a new window]
 
Fig. 8. DIEC monolayers studied in Ringer solution lacking culture-stimulating agents. A: representative decay of the Vt and Isc upon the transfer of the DIEC monolayer from fortified OptiMEM containing epidermal growth factor, hydrocortisone, and insulin to plain Ringer solution. Upon the transfer a time 0, Vt and Isc decreased with time. B: effect of serosal dibutyryl-cAMP (DBcAMP, 1 mM) on transepithelial electrical variables of DIEC monolayers in plain Ringer solution containing 130 mM Cl on both sides and amiloride (10 µM) on the apical side. C: response to DBcAMP in low-Cl Ringer solution containing 13 mM Cl on both sides and amiloride (10 µM) on the apical side of the monolayer.

 
Effects of cAMP on monolayer electrophysiology in plain Ringer solutions. Responding to a query of a reviewer, we examined DIEC monolayers for transepithelial Cl secretion, a functional hallmark of the mammalian jejunum. In these experiments, monolayers were first transferred to plain Ringer solution lacking culture-stimulating agents, and then amiloride (10 µM) was added to the mucosal solution (Fig. 8B). Both maneuvers served to abolish the transepithelial Na+ current mediated in part by ENaC. Under these conditions, Isc was 0.3 ± 0.2 µA/cm2, Vt was –1.5 ± 0.1 mV, and Rt was 3,073 ± 114 {Omega}·cm2 in six monolayers (Fig. 8B). Upon the addition of 1 mM DBcAMP to the basolateral solution, Vt and Isc increased gradually and significantly, reaching peak values 5 min later. Within in the next 5 min, Isc went to steady-state values of 3.8 ± 0.2 µA/cm2, Vt went to –6.8 ± 0.4 mV, and Rt decreased to 1,718 ± 148 {Omega}·cm2. The effects of DBcAMP on Vt, Rt, and Isc are highly significant (P < 0.001).

The same experiment repeated in the presence of low-Cl Ringer solution (13 mM Cl) revealed a marked decrease in the response to DBcAMP (Fig. 8C). In the presence of low-Cl Ringer on both sides of the monolayer and apical amiloride (10 µM), the addition of DBcAMP (1 mM) to the serosal side significantly increased Isc from zero to 2.5 ± 0.1 µA/cm2 (P < 0.001); it increased Vt from –1.2 ± 0.1 to –3.6 ± 0.3 mV (P < 0.001), and it significantly decreased Rt from 3,883 ± 388 to 1,827 ± 255 {Omega}·cm2 (P < 0.001) in six monolayers. Although the effects of DBcAMP were statistically significant, they were less than those observed in the presence of plain Ringer solution with a Cl concentration of 129.8 mM (Fig. 8, B and C). These results show that the DBcAMP stimulation of Isc and Vt was dependent on the presence of Cl.

The same conclusion was reached in studies of DIEC monolayers in glucose-free Ringer solution and in the presence of apical amiloride (10 µM). Under these conditions, the addition of 1 mM DBcAMP to the serosal Ringer solution significantly increased Vt from –1.3 ± 0.1 to –7.1 ± 0.1 mV (P < 0.001), it significantly increased Isc from 0.1 ± 0.1 to 4.6 ± 0.4 µA/cm2 (P < 0. 0.001), and it significantly reduced Rt from 2,718 ± 229 to 1,532 ± 134 {Omega}·cm2 (P < 0.001), again in six monolayers. Because these effects are quantitatively similar to those observed in the presence of glucose, the significant effects of DBcAMP are not the result of stimulation of electrogenic SGLT.

Expression of SGLT1 in DIEC monolayers. Western blot analysis confirmed the presence of SGLT1 in DIEC cultures (Fig. 9A). In a protein extract of DIEC, the antibody 8821 specific to mammalian SGLT1 recognized a protein band at 75 kDa, where SGLT1 is expected to locate (Fig. 9A, left). Antibody specificity was confirmed by tying up the antibody with immunizing peptide, which prevented it from binding to SGLT1 in the DIEC extract (Fig. 9A, right).



View larger version (34K):
[in this window]
[in a new window]
 
Fig. 9. Presence of the SGLT1 in DIEC monolayers. A: antibody 8821 raised against SGLT1 recognized the Na+-D-glucose cotransporter in an extract of DIEC (Western blot on left). Western blot on right shows nonspecific background when antibody 8821 binding to immunizing peptide is not free to bind to SGLT1. B: effect of 10 mM glucose on Vt, Isc, and Rt in DIEC monolayers bathed in plain Ringer solution lacking glucose. The addition of glucose to both sides of the monolayer tended to increase Isc and to decrease Rt, but not significantly. Rt was determined as the inverse slope of current-voltage plots.

 
The electrophysiological evidence for the presence of Na+-D-glucose cotransport was less clear (Fig. 9B). In this series of experiments, five DIEC monolayers studied in glucose-free Ringer solution had a Vt of –3.6 ± 0.3 mV, an Isc of 3.2 ± 0.5 µA/cm2, and an Rt of 1,061 ± 133 {Omega}·cm2. The addition of 10 mM glucose to both serosal and mucosal solutions decreased Vt to –2.0 ± 1.1 mV, increased Isc to 3.9 ± 0.3 µA/cm2, and decreased Rt to 830 ± 156 {Omega}·m2 (Fig. 9B). Although the changes in Isc and Rt were consistent with the stimulation of electrogenic Na+-D-glucose cotransport, they were not statistically significant.


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cultures of the mammalian intestine. Fully differentiated epithelial cells derived from the normal mammalian intestine are difficult to culture in vitro (40), and most have been obtained in nonproliferative states that do not allow significant expansion beyond the primary culture. Most normal cell lines available today have been derived from stem cells or undifferentiated committed crypt cells but have proved of limited use in studies of nutrient transport across the intestine. The first IEC line was produced by one of us (Quaroni), who started the culture with crypt cells of the newborn rat (42). IEC tends to grow in packed colonies with short and sparse apical microvilli and only limited expression of tight junctions and desmosomes. IEC does not grow well on filter supports, nor does it form tight monolayers suitable for transport studies. Except for aminopeptidase and dipeptidylpeptidase, the usual brush-border digestive enzymes are absent, suggesting the largely undifferentiated nature of the IEC culture (40). Furthermore, the cell line lacks most of the active transport systems and the enzymes of the normal epithelium (53).

In the present study, we introduced an intestinal cell culture derived from the normal, healthy intestinal mucosa of the dog jejunum. The DIEC culture was directly produced from jejunal crypts, apparently from committed crypt cells. The culture could be carried through at least six passages without apparent morphological changes or loss of ability to form polarized monolayers on filters. When monolayers are grown on filters, epithelial cells display long and dense microvilli at the apical membrane, extensive basolateral membrane infoldings, and well-defined tight junctions and desmosomes (Figs. 3 and 4). The expression of keratin no. 18 (Fig. 1c) and the presence of several tight junction proteins confirm the epithelial nature of the culture (Fig. 2). The expression of the epithelial Na+ channel ENaC (Figs. 5 and 6) was unexpected and most likely the result of the presence of epidermal growth factor, corticosterone, and insulin in the culture medium (vide infra).

Proteins of the tight junction and the paracellular pathway. To date, 24 members of the claudin protein family have been identified in mice and men (50). The claudins have four transmembrane domains with NH2 and COOH terminals located in the cytoplasm. Associated with other junctional proteins (occludin, ZO proteins, and junctional adhesion proteins), the claudins define the barrier and permselectivity functions of the paracellular pathway (29, 34). For example, the presence of claudin-16 in tight junctions of the thick ascending limb of the Loop of Henle defines the paracellular Ca2+ and Mg2+ permselectivity in this part of the nephron (46). More than two claudins may coexist in a tight junction, forming heteropolymers that appear to increase the structural and functional diversity of the paracellular pathway (17).

So far, claudin-3 and claudin-4 have been found in the jejunum (43). Significantly, the same claudins are expressed in the DIEC culture of the dog jejunum, indicating a cultured tight junction that resembles the junction of the normal jejunum. A receptor function has been attributed to these two claudins (5, 26). Binding to the enterotoxin of Clostridium perfringens, claudin-3 and claudin-4 are thought to participate in triggering the diarrhea of the Clostridium infection. Apparently, claudins can have functions in addition to permissive paracellular transport.

Occludin is another tight junction protein with four transmembrane domains. The discrete distribution of occludin in a narrow band surrounding the cells of DIEC monolayers is consistent with its location in tight junctions (Fig. 2b). Likewise, the distribution of ZO-1 is limited to the region of the tight junction (Fig. 2c).

On first examination, the distribution of claudin-4 suggests its presence in the cytoplasm of cell (Fig. 2d). However, upon closer inspection, it appears that the microtome cut takes a tangent to tight junctions. Leaving one epithelial cell, the cut encounters a sharp edge of claudins, suggesting its entry into the tight junction. Passing obliquely through the tight junction, the cut gives rise to the diffuse distribution of claudin-4. In other regions of Fig. 2b, claudin is as discretely confined to the cell border, like occludin and ZO-1, consistent with the expression of claudins at plasma membranes (35). Nevertheless, claudins have been observed inside cells. When exogenous claudin-2 and -4 were experimentally expressed in Madin-Darby canine kidney II cells, they were detected in some intracellular, vacuole-like structures in addition to their expression at the cell border, whereas native claudins were restricted to the cell border (9). Studies by Kobayashi et al. (30) suggest that foreign claudins can induce the formation of vacuole-like structures in the cytoplasm.

Desmosomes are "welding spots" that hold cells together and resist tears in epithelial tissues. Desmosomes are abundantly expressed in DIEC monolayers where they outline the lateral interstitial space between epithelial cells and beyond the tight junction (Fig. 2e).

DIEC, a leaky epithelium in spite of appreciable Rt values. In 1972, Frömter and Diamond introduced the concepts of "leaky" and "tight" epithelia (16). In general, leaky epithelia have 1) low values of Vt (from 0 to 11 mV) and Rt (from 6 to 200 {Omega}·cm2), 2) a brush-border apical membrane, and 3) the functional properties of isosmotic fluid transport at high rates. In contrast, tight epithelia have high values of Vt (30–100 mV) and Rt (300–2,000 {Omega}·cm2). Tight epithelia lack a brush border at apical surfaces. The main function of tight epithelia is to generate and maintain high transepithelial concentration differences of solute and water for storage.

In the present study, DIEC monolayers had on average a Rt of 1,050 {Omega}·cm2 consistent with a tight epithelium when compared with 50 {Omega}·cm2, the resistance of the small intestine in vivo (15, 36). On the other hand, a low Vt and the presence of a dense brush border indicate a leaky epithelium. Instead of measures of the Vt and Rt, Boulpaep (6) has proposed a more appropriate criterion of epithelial leakiness or tightness, namely the ratio of transcellular and paracellular transport in general and the ratio of transcellular and paracellular resistance in particular. To estimate this resistance ratio in DIEC monolayers, we have assumed that the electromotive force of the active transport pathway passing through epithelial cells is 120 mV. Because Isc is 8.1 µA/cm2, it follows that the tanscellular resistance is 14,815 {Omega}·cm2. In view of a Rt (transcellular and paracellular resistance in parallel) of 1,050 {Omega}·cm2, the paracellular resistance is 1,131 {Omega}·cm2. Thus the ratio of transcellular and paracellular resistance indicates a paracellular conductance 13 times greater than the transcellular conductance consistent with a leaky epithelium. A high paracellular conductance may obscure the effects of amiloride on the Rt (Figs. 5 and 6).

Functional characterization of DIEC. Absorption of salt, water, and dietary nutrients is the hallmark function of the mammalian small intestine. The human intestine absorbs ~600 mmol Na+ and 6.5 liters H2O/day. The absorption of water is coupled to the absorption of solute. In turn, the absorption of solute relies largely on Na+-dependent transport mechanisms such as Na+-glucose cotransport, Na+-amino acid cotransport, Na+/H+ exchange transport, and NaCl absorption via parallel Na+/H+ and Cl/HCO3 transport. The transport of Na+ across the small intestine varies 1) along the length of the intestine (radial heterogeneity), 2) along the length from crypt to the tip of a villus (axial heterogeneity), and 3) between individual epithelial cells in regions of the villus tip and crypt that express transport systems to varying degree (cellular heterogeneity). DIEC monolayers derived from the normal jejunum appear to illustrate some of these heterogeneities. For one reason, the net absorptive isotopic Na+ flux was nearly two times the Isc measured in parallel (Table 1), suggesting the presence of electroneutral Na+ transport mechanisms such as NaCl absorption via parallel Na+/H+ and Cl/HCO3 transport. For another reason, the inequality of current and flux suggests that flux and current do not derive from the same monolayer area, i.e., the monolayer is heterogeneous like the normal jejunum.

Our preliminary functional characterization of DIEC monolayers illustrated the influence of environmental factors on the expression of transport systems. When DIEC monolayers are grown and studied in OptiMEM solution containing the culture-stimulating agents epidermal growth factor, hydrocortisone, and insulin, the monolayers display an amiloride-sensitive Isc mediated by ENaC (Fig. 6). This current disappears in plain Ringer solution (Fig. 8A). The subsequent addition of cAMP to the serosal side activates a Cl-dependent current consistent with transepithelial secretion of Cl (Fig. 8, B and C). Thus DIEC monolayers can be manipulated to express electrogenic Na+ absorption under one set of conditions and electrogenic Cl secretion under another.

When grown and studied in fortified OptiMEM solution, the inhibition of the Isc with amiloride presented from the mucosal side (and not the serosal side, Fig. 5) and the inhibitory effects of serosal ouabain on voltage and Isc (Fig. 7) outline the rudiments of the Ussing model of epithelial Na+ transport (51). The amiloride concentration-response curve revealed an IC50 of 0.76 µM (Fig. 6B), similar to that measured in other epithelia expressing ENaC (4). The immediate effect of ouabain on Vt and Isc probably results from the blockade of the electrogenic Na+-K+-ATPase operating with a stoichiometric exchange of three Na+ for two K+. The ouabain-sensitive pump current was measured as 1.3 ± 0.2 µA/cm2, which is significantly different from zero (P < 0.001). The gradual decline of Vt and Isc in the presence of ouabain probably reflects the dissipation of Na+, K+, and voltage gradients across cell membranes.

Isotopic Na+ flux measurements in DIEC monolayers grown and studied in fortified OptiMEM show that ENaC-mediated transepithelial Na+ transport accounts for only 18% of the net absorptive transepithelial Na+ transport (Table 1). Because these measurements were made in the absence of Vt and concentration differences, DIEC monolayers must possess additional transepithelial active transport mechanisms for Na+. Indeed, an antibody specific to SGLT1 recognizes a protein in DIEC lysates that localizes at the expected position (Fig. 9A). That the same antibody recognizes SGLT1 in the dog jejunum in vivo indicates that our culture expresses Na+/D-glucose cotransport like the normal jejunum (22). Next to Na+/D-glucose cotransport, DIEC monolayers may express other Na+-dependent solute transport systems that were not investigated in the present study.

Significantly, amiloride-sensitive Vt and Isc disappeared in monolayers when fortified OptiMEM, containing growth factor, hydrocortisone, and insulin, was replaced with plain Ringer solution (Fig. 8A). In the presence of plain Ringer solution on both sides of the monolayer, the Vt and Isc decayed with a time course expected from the endocytic deactivation of ENaC (7, 8). The decay of voltage and current suggests that the presence of growth factor, glucocorticoid, and insulin in the culture medium had activated the expression of ENaC (38, 45). Still, the presence of ENaC in a cell culture of the small intestine may not be so unusual as it first appears. In rats, the surgical removal of the colon (ileoanal anastomosis) induces ENaC expression in the ileum (31). Furthermore, the induction of ENaC activity has been attributed to serum- and glucocorticoid-induced kinase that is present in both jejunum and ileum (10).

Next to transepithelial Na+ absorption, the transepithelial secretion of fluid driven by secretory Cl transport is a functional hallmark of the jejunum. In particular, Cl-driven fluid secretion is thought to take place in the crypt region of villi and is mediated by epithelial cells that harbor Cl channels in the apical membrane and the Na+-K+-2Cl cotransporter in the basolateral membrane (13, 54). Up to now, at least the following three different Cl channels have been found in the small intestine: the stretch-activated Cl channel, the Ca2+-activated Cl channel, and cystic fibrosis transmembrane conductance regulator (CFTR) that is activated by phosphorylation via protein kinase A (PKA) and cAMP (3). DIEC monolayers apparently exhibit Cl secretion via CFTR in view of the effects of DBcAMP (Fig. 8, B and C). The nucleotide induced a significant increase in Isc (from mucosa to serosa) when ENaC channels were blocked with amiloride. The cAMP-stimulated Isc was dependent on the presence of Cl in the bath medium consistent with the stimulation of transepithelial Cl secretion. Furthermore, the sustained stimulation of Isc distinguishes CFTR channels from Ca2+-activated and stretch-activated Cl channels, which respond only transiently to activation (3, 13).

Of note is the significant decrease of the Rt upon stimulation with cAMP (Fig. 8, B and C). The activation of CFTR in the apical membrane of DIEC monolayers was not expected to decrease the Rt too much in view of the low paracellular resistance relative to the transcellular resistance that prevented a significant increase in Rt in the presence of amiloride (Figs. 5A and 6A). Thus cAMP may have affected the paracellular pathway in addition to CFTR in the apical membrane of DIEC. Indeed, the nucleotide is thought to increase tight junction permeability (25). The molecular mechanism is not completely understood, but the PKA phosphorylation of Thr207 of claudin-5, one of the tight junction proteins, is known to trigger the rapid decrease in transendothelial resistance in rat lung endothelium (47).

Na+-dependent glucose cotransport via SGLT1 is another hallmark of the jejunum. Glucose enters the cell from the intestinal lumen against its chemical gradient at the expense of the Na+ electrochemical gradient across the apical membrane. Glucose leaves the cells across the basolateral membrane down its chemical gradient through another sugar transporter of the family of GLUT glucose transporters (48).

Western blot analysis revealed the presence of SGLT1 in DIEC monolayers. The electrophysiological evidence for SGLT1 was less clear. The addition of 10 mM glucose to glucose-free medium tended to increase the Isc, but the change was not statistically significant (Fig. 9B). SGLT1 is also present in other cultures of the mammalian intestine, such as Caco-2, HT29 cl.19A, and HT29-D4. However, it is active only in fully differentiated HT29-D4 cell monolayers (12, 18, 20), suggesting that SGLT1 activity correlates with cell differentiation (11). Indeed, SGLT genes are transcribed and translated only in mature enterocytes at villus tips (23). Thus it is likely that the number of tip cells present in DIEC monolayers may be sufficient to yield a positive Western blot but insufficient to yield significant transepithelial electrical changes when stimulated with glucose.

Na+-D-glucose cotransport can be activated by cAMP, as in HRT-18 cells (37). However, in DIEC monolayers, the cAMP stimulation of the Isc was primarily the result of stimulation of a Cl-dependent current (Figs. 8 and 9).

As a good model for the study of intestinal transport, a monolayer must resemble the in vivo condition physically, morphologically, and biochemically. In the present study, we have examined the morphological and electrophysiological characteristics of a new intestinal culture derived from the normal dog jejunum. DIEC monolayers resemble normal epithelium to a remarkable degree. They display the morphological polarization of the jejunum with a prominent brush border, basolateral membrane infoldings, and the functional polarization of transporters in expected basolateral and apical membrane domains of the jejunum. Likewise, DIEC tight junctions possess the claudin proteins of the normal jejunum. Yet we leave many transport questions open to further studies, and the expression of digestive enzymes has yet to be examined. As a culture of the normal small intestine, DIEC monolayers may find wide application serving basic and applied motivations, from studies of the mechanism and regulation of transepithelial solute and water transport to drug absorption, intestinal clearance of xenobiotics, and high throughput evaluations of pharmaceutical agents.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by Grant IBN 0078058 awarded to K. W. Beyenbach from the National Science Foundation and Grant DK-48331 awarded to A. Quaroni from the National Institute of Diabetes and Digestive and Kidney Diseases.


    ACKNOWLEDGMENTS
 
We thank Dr. Robert Gilmour (Cornell University) for providing dog intestines and Dr. Bruce A. Hirayama and Dr. Ernest M. Wright for the gift of antibody 8821 and immunizing peptide.


    FOOTNOTES
 

Address for reprint requests and other correspondence: K. W. Beyenbach, Dept. of Biomedical Sciences, VRT 8004, Cornell Univ., Ithaca, NY 14853 (E-mail: kwb1{at}cornell.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.


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Adibi SA. Regulation of expression of the intestinal oligopeptide transporter (Pept-1) in health and disease. Am J Physiol Gastrointest Liver Physiol 285: G779–G788, 2003.[Abstract/Free Full Text]
  2. Artursson P, Palm K, and Luthman K. Caco-2 monolayers in experimental and theoretical predictions of drug transport. Adv Drug Delivery Res 46: 27–43, 2001.[CrossRef][ISI][Medline]
  3. Basavappa S, Vulapalli SR, Zhang H, Yule D, Coon S, and Sundaram U. Chloride channels in the small intestinal cell line IEC-18. J Cell Physiol 202: 21–31, 2005.[CrossRef][ISI][Medline]
  4. Benos DJ. Amiloride: a molecular probe of sodium transport in tissues and cells. Am J Physiol Cell Physiol 242: C131–C145, 1982.[Abstract]
  5. Berkes J, Viswanathan VK, Savkovic SD, and Hecht G. Intestinal epithelial responses to enteric pathogens: effects on the tight junction barrier, ion transport, and inflammation. Gut 52: 439–451, 2003.[Abstract/Free Full Text]
  6. Boulpaep EL. In: The Paracellular Pathway, edited by Bradley SE and Purcell EF. New York, NY: Macy Foundation, 1982, p. 131–132.
  7. Butterworth MB, Helman SI, and Els WJ. cAMP-sensitive endocytic trafficking in A6 epithelia. Am J Physiol Cell Physiol 280: C752–C762, 2001.[Abstract/Free Full Text]
  8. Carattino MD, Hill WG, and Kleyman TR. Arachidonic acid regulates surface expression of epithelial sodium channels. J Biol Chem 278: 36202–36213, 2003.[Abstract/Free Full Text]
  9. Colegio OR, Van Itallie C, Rahner C, and Anderson JM. Claudin extracellular domains determine paracellular charge selectivity and resistance but not tight junction fibril architecture. Am J Physiol Cell Physiol 284: C1346–C1354, 2003.[Abstract/Free Full Text]
  10. Coric T, Hernandez N, de la Rosa DA, Shao D, Wang T, and Canessa CM. Expression of ENaC and serum- and glucocorticoid-induced kinase 1 in the rat intestinal epithelium. Am J Physiol Gastrointest Liver Physiol 286: G663–G670, 2004.[Abstract/Free Full Text]
  11. Delezay O, Baghdiguian S, and Fantini J. The development of Na+-dependent glucose transport during differentiation of an intestinal epithelial cell clone is regulated by protein kinase C. J Biol Chem 270: 12536–12541, 1995.[Abstract/Free Full Text]
  12. Delezay O, Verrier B, Mabrouk K, van Rietschoten J, Fantini J, Mauchamp J, and Gerard C. Characterization of an electrogenic sodium/glucose cotransporter in a human colon epithelial cell line. J Cell Physiol 163: 120–128, 1995.[CrossRef][ISI][Medline]
  13. Field M. Intestinal ion transport and the pathophysiology of diarrhea. J Clin Invest 111: 931–943, 2003.[Free Full Text]
  14. Fox JJ, Riccio ML, Hua F, Bodenschatz E, and Gilmour RF Jr. Spatiotemporal transition to conduction block in canine ventricle. Circ Res 90: 289–296, 2002.[Abstract/Free Full Text]
  15. Fromm M, Schulzke JD, and Hegel U. Epithelial and subepithelial contributions to transmural electrical resistance of intact rat jejunum, in vitro. Pflugers Arch 405: 400–402, 1985.[CrossRef][ISI][Medline]
  16. Fromter E and Diamond J. Route of passive ion permeation in epithelia. Nat New Biol 235: 9–13, 1972.[ISI][Medline]
  17. Furuse M, Sasaki H, and Tsukita S. Manner of interaction of heterogeneous claudin species within and between tight junction strands. J Cell Biol 147: 891–903, 1999.[Abstract/Free Full Text]
  18. Grasset E, Bernabeu J, and Pinto M. Epithelial properties of human colonic carcinoma cell line Caco-2: effect of secretagogues. Am J Physiol Cell Physiol 248: C410–C418, 1985.[Abstract]
  19. Hariton-Gazal E, Rosenbluh J, Graessmann A, Gilon C, and Loyter A. Direct translocation of histone molecules across cell membranes. J Cell Sci 116: 4577–4586, 2003.[Abstract/Free Full Text]
  20. Hemlin M and Huang X. Na+/glucose cotransport in the colonic adenocarcinoma cell line HT29 cl.19A: effect of cAMP. Acta Physiol Scand 160: 185–194, 1997.[ISI][Medline]
  21. Hidalgo IJ, Raub TJ, and Borchardt RT. Characterization of the human colon carcinoma cell line (Caco-2) as a model system for intestinal epithelial permeability. Gastroenterology 96: 736–749, 1989.[ISI][Medline]
  22. Hines OJ, Whang EE, Bilchik AJ, Zinner MJ, Welton ML, Lane J, McFadden DW, and Ashley SW. Role of Na+-glucose cotransport in jejunal meal-induced absorption. Dig Dis Sci 45: 1–6, 2000.[CrossRef][ISI][Medline]
  23. Hwang ES, Hirayama BA, and Wright EM. Distribution of the SGLT1 Na+/glucose cotransporter and mRNA along the crypt-villus axis of rabbit small intestine. Biochem Biophys Res Commun 181: 1208–1217, 1991.[CrossRef][ISI][Medline]
  24. Itoh A, Tsujikawa T, Fujiyama Y, and Bamba T. Enhancement of aquaporin-3 by vasoactive intestinal polypeptide in a human colonic epithelial cell line. J Gastroenterol Hepatol 18: 203–210, 2003.[CrossRef][ISI][Medline]
  25. Karczewski J and Groot J. Molecular physiology and pathophysiology of tight junctions III. Tight junction regulation by intracellular messengers: differences in response within and between epithelia. Am J Physiol Gastrointest Liver Physiol 279: G660–G665, 2000.[Abstract/Free Full Text]
  26. Katahira J, Inoue N, Horiguchi Y, Matsuda M, and Sugimoto N. Molecular cloning and functional characterization of the receptor for Clostridium perfringens enterotoxin. J Cell Biol 136: 1239–1247, 1997.[Abstract/Free Full Text]
  27. Keely SJ and Barrett KE. p38 mitogen-activated protein kinase inhibits calcium-dependent chloride secretion in T84 colonic epithelial cells. Am J Physiol Cell Physiol 284: C339–C348, 2003.[Abstract/Free Full Text]
  28. Kinston RE. Preparation and analysis of RNA. In: Current Protocols in Molecular Biology. New York, NY: Wiley, 2002, p. 4.2.3–4.2.5.
  29. Kiuchi-Saishin Y, Gotoh S, Furuse M, Takasuga A, Tano Y, and Tsukita S. Differential expression patterns of claudins, tight junction membrane proteins, in mouse nephron segments. J Am Soc Nephrol 13: 875–886, 2002.[Abstract/Free Full Text]
  30. Kobayashi J, Inai T, and Shibata Y. Formation of tight junction strands by expression of claudin-1 mutants in their ZO-1 binding site in MDCK cells. Histochem Cell Biol 117: 29–39, 2002.[CrossRef][ISI][Medline]
  31. Koyama K, Sasaki I, Naito H, Funayama Y, Fukushima K, Unno M, Matsuno S, Hayashi H, and Suzuki Y. Induction of epithelial Na+ channel in rat ileum after proctocolectomy. Am J Physiol Gastrointest Liver Physiol 276: G975–G984, 1999.[Abstract/Free Full Text]
  32. Li T, Ito K, and Horie T. Transport of fluorescein methotrexate by multidrug resistance-associated protein 3 in IEC-6 cells. Am J Physiol Gastrointest Liver Physiol 285: G602–G610, 2003.[Abstract/Free Full Text]
  33. Ma TY, Hollander D, Bhalla D, Nguyen H, and Krugliak P. IEC-18, a nontransformed small intestinal cell line for studying epithelial permeability. J Lab Clin Med 120: 329–341, 1992.[ISI][Medline]
  34. Mankertz J, Hillenbrand B, Tavalali S, Huber O, Fromm M, and Schulzke JD. Functional crosstalk between Wnt signaling and Cdx-related transcriptional activation in the regulation of the claudin-2 promoter activity. Biochem Biophys Res Commun 314: 1001–1007, 2004.[CrossRef][ISI][Medline]
  35. Morita K, Furuse M, Fujimoto K, and Tsukita S. Claudin multigene family encoding four-transmembrane domain protein components of tight junction strands. Proc Natl Acad Sci USA 96: 511–516, 1999.[Abstract/Free Full Text]
  36. Munck BG and Schultz SG. Properties of the passive conductance pathway across in vitro rat jejunum. J Membr Biol 16: 163–174, 1974.[ISI][Medline]
  37. Nath SK, Rautureau M, Heyman M, Reggio H, L'Helgoualc'h A, and Desjeux JF. Emergence of Na+-glucose cotransport in an epithelial secretory cell line sensitive to cholera toxin. Am J Physiol Gastrointest Liver Physiol 256: G335–G341, 1989.[Abstract/Free Full Text]
  38. Pearce D. SGK1 regulation of epithelial sodium transport. Cell Physiol Biochem 13: 13–20, 2003.[CrossRef][ISI][Medline]
  39. Quaroni A and Beaulieu JF. Cell dynamics and differentiation of conditionally immortalized human intestinal epithelial cells. Gastroenterology 113: 1198–1213, 1997.[ISI][Medline]
  40. Quaroni A and Hochman J. Development of intestinal cell culture models for drug transport and metabolism studies. Adv Drug Delivery Res 22: 3–52, 1996.[CrossRef][ISI]
  41. Quaroni A and Isselbacher KJ. Cytotoxic effects and metabolism of benzo[a]pyrene and 7,12-dimethylbenz[a]anthracene in duodenal and ileal epithelial cell cultures. J Natl Cancer Inst 67: 1353–1362, 1981.[ISI][Medline]
  42. Quaroni A, Wands J, Trelstad RL, and Isselbacher KJ. Epithelioid cell cultures from rat small intestine. Characterization by morphologic and immunologic criteria. J Cell Biol 80: 248–265, 1979.[Abstract]
  43. Rahner C, Mitic LL, and Anderson JM. Heterogeneity in expression and subcellular localization of claudins 2, 3, 4, and 5 in the rat liver, pancreas, and gut. Gastroenterology 120: 411–422, 2001.[ISI][Medline]
  44. Rhoads JM, Chen W, Chu P, Berschneider HM, Argenzio RA, and Paradiso AM. L-Glutamine and L-asparagine stimulate Na+-H+ exchange in porcine jejunal enterocytes. Am J Physiol Gastrointest Liver Physiol 266: G828–G838, 1994.[Abstract/Free Full Text]
  45. Schulz-Baldes A, Berger S, Grahammer F, Warth R, Goldschmidt I, Peters J, Schutz G, Greger R, and Bleich M. Induction of the epithelial Na+ channel via glucocorticoids in mineralocorticoid receptor knockout mice. Pflügers Arch 443: 297–305, 2001.[CrossRef][ISI][Medline]
  46. Simon DB, Lu Y, Choate KA, Velazquez H, Al-Sabban E, Praga M, Casari G, Bettinelli A, Colussi G, Rodriguez-Soriano J, McCredie D, Milford D, Sanjad S, and Lifton RP. Paracellin-1, a renal tight junction protein required for paracellular Mg2+ resorption. Science 285: 103–106, 1999.[Abstract/Free Full Text]
  47. Soma T, Chiba H, Kato-Mori Y, Wada T, Yamashita T, Kojima T, and Sawada N. Thr(207) of claudin-5 is involved in size-selective loosening of the endothelial barrier by cyclic AMP. Exp Cell Res 300: 202–212, 2004.[CrossRef][ISI][Medline]
  48. Thorens B. Facilitated glucose transporters in epithelial cells. Annu Rev Physiol 55: 591–608, 1993.[CrossRef][ISI][Medline]
  49. Tian JQ and Quaroni A. Involvement of p21(WAF1/Cip1) and p27(Kip1) in intestinal epithelial cell differentiation. Am J Physiol Cell Physiol 276: C1245–C1258, 1999.[Abstract/Free Full Text]
  50. Tsukita S and Furuse M. Claudin-based barrier in simple and stratified cellular sheets. Curr Opin Cell Biol 14: 531–536, 2002.[CrossRef][ISI][Medline]
  51. Ussing HH and Zerahn K. Active transport of sodium as the source of electric current in the short-circuited isolated frog skin. Acta Physiol Scand 23: 110–127, 1951.[ISI][Medline]
  52. Varga A, Nugel H, Baehr R, Marx U, Hever A, Nacsa J, Ocsovszky I, and Molnar J. Reversal of multidrug resistance by amitriptyline in vitro. Anticancer Res 16: 209–211, 1996.[ISI][Medline]
  53. Versantvoort CHM, Ondrewater RCA, Duizer E, Van de Sandt JJM, Gilde AJ, and Groten JP. Monolayers of IEC-18 cells as an in vitro model for screening the passive transcellular and paracellular transport across the intestinal barrier: comparison of active and passive transport with the human colon carcinoma Caco-2 cell line. Environ Toxicol Pharmacol 11: 335–344, 2002.[CrossRef][ISI]
  54. Welsh MJ, Smith PL, Fromm M, and Frizzell RA. Crypts are the site of intestinal fluid and electrolyte secretion. Science 218: 1219–1221, 1982.[ISI][Medline]
  55. Weng XH, Huss M, Wieczorek H, and Beyenbach KW. The V-type H+-ATPase in Malpighian tubules of Aedes aegypti: localization and activity. J Exp Biol 206: 2211–2219, 2003.[Abstract/Free Full Text]




This Article
Abstract
Full Text (PDF)
All Versions of this Article:
288/4/G705    most recent
00518.2003v1
Alert me when this article is cited
Alert me if a correction is posted
Citation Map
Services
Email this article to a friend
Similar articles in this journal
Similar articles in ISI Web of Science
Similar articles in PubMed
Alert me to new issues of the journal
Download to citation manager
Search for citing articles in:
ISI Web of Science (2)
Google Scholar
Articles by Weng, X.-H.
Articles by Quaroni, A.
Articles citing this Article
PubMed
PubMed Citation
Articles by Weng, X.-H.
Articles by Quaroni, A.


HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2005 by the American Physiological Society.