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ARTICLE |
CORRESPONDENCE Anjaparavanda P. Naren: anaren{at}utmem.edu
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Infectious diarrhea disease is second only to cardiovascular disease as cause of death (1) and is one of the major causes of morbidity and mortality among children. It accounts for 21% of deaths of children who are younger than 5 yr of age in the developing world, and causes 2.5 million deaths per year (2). Although incidence rates increase with a country's decreasing socioeconomic status, infectious diarrhea also is a common illness in Western industrialized societies, particularly among vulnerable groups of young children, elderly adults, and people who have underlying diseases (3). Two major organisms that cause infectious diarrhea in humans are Escherichia coli and Vibrio cholerae; their secreted toxins (heat stable or heat labile toxin, and cholera toxin [CTX], respectively) influence gastrointestinal epithelial cell function and induce diarrhea by numerous mechanisms (4). CTX is likely the most recognizable enterotoxin that causes secretory diarrhea by stimulating transepithelial Cl secretion, thereby increasing the osmotic impetus for fluid secretion (5). Diarrhea also can result from an increased immunoinflammatory response that is triggered by cytokine or mediator (e.g., adenosine; ADO) secretion from intestinal mucosal inflammatory cells in response to luminal factors (e.g., dietary or bacterial antigens). This also is a common symptom that is associated with the major gastrointestinal disorders, such as inflammatory bowel disease (IBD) afflicting >1 million Americans (6), and irritable bowel syndrome, which affects 15 million Americans (7). Mucosal inflammation plays an essential role in the pathogenesis of irritable bowel syndrome and IBD (8, 9). ADO, an inflammatory mediator, stimulated chloride secretion in several types of secretory epithelia, including mammalian ileum and colon (10, 11).
Cystic fibrosis transmembrane conductance regulator (CFTR) is a cAMP-regulated chloride channel that is localized primarily at the apical or luminal surfaces of epithelial cells that line the airway, gut, and exocrine glands (12); it is well-established that CFTR plays a pivotal role in CTX-mediated diarrhea (13). Lysophosphatidic acid (LPA), a naturally occurring phospholipid present in blood and foods (14, 15), acts on the LPA receptors (LPA1, LPA2, and LPA3), which are G-protein coupled receptors that belong to the endothelial cell differentiation gene family (1618). LPA was reported to play an essential role in a variety of conditions involving gastrointestinal wound repair, apoptosis, IBD, and diarrhea (1923). In the present study, we investigated how LPA-elicited LPA2-mediated signaling might regulate CFTR function in the gut, and evaluated the usefulness of LPA in preventing CTX-induced secretory diarrhea in mice.
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RESULTS |
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LPA2 binds NHERF2 with the highest affinity
The COOH-terminal tails of LPA1 and LPA2, but not LPA3, contain a consensus for the PDZ motif, the short COOH-terminal sequences that specifically interact with the PDZ-binding domains (usually 7090 amino acid sequences) of PDZ proteins, including NHERF2 (28) (Fig. S2 A, boxed area, available at http://www.jem.org/cgi/content/full/jem.20050421/DC1). Thus, LPA1 and LPA2 are likely candidates with which PDZ-domaincontaining proteins can interact, modulate, and mediate their signals. Pull-down assays demonstrated that LPA2 interacted directly with NHERF2 with the highest affinity (Fig. S2, B and C), as reported previously (25, 26).
LPA inhibits CFTR-dependent iodide efflux through LPA2 receptormediated Gi pathway
Although all three LPA receptors (LPA1, LPA2, and LPA3) respond to LPA, LPA2 has the highest affinity to the lipid (29), and leads to the activation of at least three distinct G-protein pathways: Gi, Gq, and G12/13 (17, 18). Activation of these receptors by LPA results in inhibition of the adenylate cyclase (AC) pathway, which, in turn, decreases cAMP levels (1618). The Gi activation pathway of the receptor is summarized in Fig. S2 D.
Given that LPA2 binds NHERF2 (25, 26) (Fig. S2, B and C) and our previous demonstration that NHERF2 can form a macromolecular complex with CFTR (30), we hypothesized that LPA might regulate CFTR Cl channel function which is regulated by cAMP. To test this, the effect of LPA on CFTR activity was monitored using iodide efflux measurements (31). Calu-3 cells, like HT29-CL19A cells, express all three proteins (CFTR, NHERF2, and LPA2) at the apical surfaces, and show a robust CFTR function because they express a higher level of CFTR compared with HT29-CL19A cells. Therefore, they represent a better model cell line for demonstrating the effects of LPA on CFTR function. LPA pretreatment (20 µM LPA 18:1 for 4 min before ADO addition) reduced the response of Calu-3 cells to cAMP-elevating ligand ADO in a dose-dependent manner (Fig. 2, A and B). CFTR activity was not inhibited by a structurally related lysophospholipid, sphingosine-1-phosphate (S1P; data not shown), whose receptors also are expressed in the gut epithelium but lack PDZ domain, nor by a related lipidphosphatidic acid (PA) (Fig. 2 B)which is not a ligand for LPA receptors (16). Similar results also were observed with HT29-CL19A cells (unpublished data). These results clearly demonstrate that LPA can inhibit CFTR function in epithelial cells.
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LPA inhibits CFTR-dependent short-circuit currents in polarized epithelial cells and mouse intestinal epithelia preparation
The effect of LPA on CFTR-dependent short-circuit currents (Isc) also was monitored on polarized epithelial monolayers (30). LPA substantially inhibited CFTR-dependent Cl currents in response to ADO activation in HT29-CL19A cells (Fig. 3 A) and Calu-3 cells (Fig. 3 B). Isc stimulated by ADO (Fig. 3 C) or 8-(4-chlorophenylthio)-cyclic AMP (cpt-cAMP) (a cell-permeant isoform of cAMP, a submaximal CFTR activating dose was used; Fig. S3 A, available at http://www.jem.org/cgi/content/full/jem.20050421/DC1) also was inhibited, in a dose-dependent fashion, in LPA-pretreated epithelia that were isolated from mouse intestine (34) (Fig. S3 B). The inhibitory effect of LPA on CFTR Cl currents in the intestinal epithelia was more prominent when CFTR was activated at lower concentrations of ADO (Fig. 3 D), whereas LPA failed to inhibit the channel function significantly when CFTR was activated maximally by a cocktail agonist mixture in intestinal epithelia (Fig. 3, C and D) and cultured epithelial cell monolayers (Fig. S3 C). The Isc in response to ADO was inhibited by CFTR blockers, diphenylamine-2-carboxylate (DPC) (Fig. 3 B) and glybenclamide (not depicted). PA and S1P did not inhibit the CFTR Cl channel (unpublished data). ADO is produced at almost 1 µM in unstressed tissue, whereas it can be 100 µM in inflamed or ischemic tissues (35). Under conditions of inflammation (e.g., IBD), ADO is secreted into the gut, which activates CFTR and leads to increased chloride and fluid secretion into the gut (10, 11, 36).
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LPA inhibits CFTR-dependent CTX-induced mouse intestinal fluid secretion in vivo
To provide proof of concept that LPA could be used in a therapeutic setting, we tested the effect of LPA on CTX-induced diarrhea in a mouse model (38). Binding of the cholera enterotoxin from V. cholerae to the intestinal enterocyte leads to ADP-ribosylation of the Gs -subunit, which, in turn, activates AC of enterocytes and leads to an elevation of cAMP (39, 40). Thus, the apical membrane CFTR Cl channel is activated, which results in a voluminous Cl and fluid secretion that can be fatal, if untreated (5, 39). In ligated ileal loops, CTX treatment elicited fluid accumulation in a dose-dependent manner (Fig. 5 A). A linear increase in cAMP accumulation within the loop tissue was elicited by the increasing amounts of CTX (injected into the ileal loops) in the range of 010 µg (Fig. S5 A, available at http://www.jem.org/cgi/content/full/jem.20050421/DC1). These results are consistent with the reports that CTX can induce a dose-dependent increase in intracellular cAMP production in cultured cells (4143) as well as a dose-dependent fluid accumulation in nonligated intestine in vivo (44). LPA treatment significantly reduced the fluid accumulation in the toxin-treated intestinal loops in a dose-dependent fashion (Fig. 5 B). PA (10100 µM) was used as a negative control in these experiments, and showed no effect in reversing CTX action (i.e., fluid accumulation) (unpublished data).
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LPA2 forms a macromolecular complex with CFTR mediated by way of NHERF2
We hypothesized that the interaction between CFTR and LPA2 likely was mediated by NHERF2. To test this, we assembled a macromolecular complex of LPA2, NHERF2, and CFTR in vitro, which is represented schematically in Fig. 6 A (24). A macromolecular complex was formed between maltose binding protein (MBP)-CCFTR, glutathione S-transferase (GST)-NHERF2, and LPA2 (Fig. 6 B). MBP-CCFTR did not bind directly to LPA2 nor did the complex form in the presence of GST or MBP alone (Fig. 6 B). The complex formation was PDZ-motif dependent. Mutating the last amino acid (aa) of LPA2 (L351A) reduced the complex formation, whereas deleting the last three aa's of LPA2 (STL) completely eliminated the macromolecular complex (Fig. 6 C).
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Disruption of the macromolecular complex of LPA2NHERF2CFTR using an LPA2-specific peptide prevents LPA2-mediated CFTR inhibition by LPA
We next determined if a physical association of LPA2 and CFTR by way of NHERF2 was essential for the LPA-elicited inhibition of the Cl channel. To disrupt the complex, we delivered an LPA2 isoform-specific peptide containing the PDZ motif (last 11 aa's of LPA2; aa's 341351), using the Chariot system (45), into polarized epithelial cells before Isc measurements. Isc in response to ADO in the presence of LPA was monitored. LPA2-specific peptide significantly prevented the LPA-elicited inhibition of CFTR-dependent Cl currents in polarized HT29-CL19A (Fig. 7 A) and Calu-3 monolayers (Fig. S6 A, available at http://www.jem.org/cgi/content/full/jem.20050421/DC1). At the end of the experiment, the epithelial cells were fixed and immunostained to confirm the delivered peptide (Fig. 7 B and Fig. S6 B). Other peptides (CFTR peptide aa's 107117) were used as a negative control (unpublished data). We also demonstrated that the LPA2 peptide could compete with MBPLPA2 C-tail for binding to NHERF2 in a dose-dependent manner (Fig. 7 C).
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DISCUSSION |
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We tested the effect of LPA on CFTR channel function under various conditions. For cultured cells or polarized cell monolayers, we used LPA of 040 µM, which is within the physiologic concentrations of LPA in blood serum (range, 550 µM) that was reported previously (47, 48). In the open-loop experiments, the maximal dose of LPA that we delivered into the mouse stomach was 1 mg/kg body weight, which is similar to the suggested dose for various dietary supplements (e.g., vitamins, minerals).
ADO is an endogenous purine nucleoside that is generated at sites of tissue stress and injury, including inflammation, ischemia, and tissue remodeling. Following its production, ADO diffuses to the surrounding cells, where it binds to the ADO receptors, and acts as a paracrine factor with diverse effects on a variety of organ systems. It was reported that in the intestinal lumen, ADO is produced in crypt abscesses during active inflammation by the interaction of neutrophils with the intestinal epithelia, whose ectonucleotidases convert neutrophil-derived ATP into ADO, which acts through the A2b ADO receptor subtype to induce vectorial chloride transport (4951). Based on these observations, it was hypothesized that luminal ADO may be involved in secretory diarrhea that is elicited in pathophysiologic states, such as parasitic or allergic diseases, and hypereosinophilic syndromes (52, 53). ADO is produced at almost 1 µM in unstressed tissue, whereas in inflamed or ischemic tissues it can be 100 µM (35). In light of extensive literature support for the physiologic and pathophysiologic role of ADO, our application of 0.2100 µM of this ligand is well within the (patho)physiologic range. In addition to ADO, we also used cpt-cAMP at submaximal CFTR activating conditions and forskolin (increases cAMP globally at 10 µM concentration) as agonists.
It was demonstrated that cAMP accumulation inside cells is inhibited by LPA treatment (54, 55). In this study, we also monitored cAMP levels under various conditions. We observed that upon stimulation with low concentration of ADO (2 µM), cAMP accumulation inside polarized epithelial cells (Calu-3) was almost indistinguishable from that of unstimulated cells (Fig. 4 A). However, CFTR is functional under these conditions and LPA can inhibit channel activity significantly (Fig. 3 B). Our data are consistent with the published results of Stutts's group (37), who demonstrated a compartmentalized signaling from the receptor (A2b) to the channel (CFTR) using electrophysiologic methods. This group reported that CFTR chloride channel function was observed upon stimulation of the cells by a low concentration of ADO (1 µM); the accumulation of cAMP inside the cell was not significantly higher than that from unstimulated control cells (37). In the present study, we found that LPA efficiently inhibits CFTR function (in response to 2 µM ADO; Fig. 3 B) without causing a decrease in the global cAMP accumulation in the cell (Fig. 4 A). Our results support the notion that cAMP likely is generated in a compartmentalized pocket upon stimulation by receptor-mediated agonists (ADO). However, when the ADO level was increased to 20 µM, we observed that the CFTR function increased slightly compared with 2 µM ADO, but there was a significant increase in cAMP accumulation inside the cell (global cAMP accumulation). Although there is a significant decrease in cAMP accumulation with LPA treatment, the Isc is not significantly different (in the presence or absence of LPA); this suggests that a global increase in cAMP may offset CFTR inhibition that is elicited by LPA.
LPA inhibition of CFTR function is most prominent when tested at a low concentration of ADO in the polarized monolayers (0.22 µM; Fig. 3, A and B) and in intestinal epithelia (240 µM; Fig. 3, C and D), or at a submaximal activating dose of cpt-cAMP (20100 µM) in isolated intestinal epithelia (Fig. S3 A). This is because polarization of epithelia leads to the accurate targeting of the channel (CFTR) and the receptor (LPA2) to the apical surfaces, compared with nonpolarized cells. In addition, in Ussing chamber experiments, the CFTR-mediated Cl currents (short-circuit currents) were monitored in real time, and we could titrate various concentrations of the agonists and monitor the CFTR response that shows maximum inhibition by LPA. In contrast, cells that were used for iodide efflux measurement (Fig. 2, AD) were grown in 60-mm dishes and were not polarized. Due to the experimental limitations, we had to choose a concentration of ADO (2 µM). cpt-cAMP used in this study (Fig. S3 A) activated CFTR submaximally (<50% of maximum activating cocktail). Therefore, external addition of a low concentration of cpt-cAMP would be very similar to cAMP accumulation observed with low concentrations of agonists, such as ADO, although ADO would likely generate a more compartmentalized cAMP accumulation.
Diarrhea is the most common gastrointestinal disorder. It is well-established that CFTR plays a central role in this process (5, 13, 36). Any reagent or factor that augments CFTR activity (e.g., cAMP in CTX-induced diarrhea) at the luminal surface of the colonic epithelial cells is likely to lead to diarrhea. Conversely, factors that inhibit the CFTR Cl channel are likely to be beneficial in cases of CFTR-induced diarrhea. Given that LPA2 is expressed in the gut, localized to the apical surface, and activates the Gi pathway and leads to decreased cAMP accumulation, we propose a novel model (Fig. 8). According to our hypothesis, LPA2 and CFTR are associated physically with NHERF2, which clusters LPA2 and CFTR into a macromolecular complex at the apical plasma membranes of epithelial cells. This macromolecular complex is the foundation of functional coupling between LPA signaling and CFTR-mediated Cl transport. The model predicts that if the physical association is disrupted, functional coupling will be compromised. Upon LPA stimulation of the receptor, AC is inhibited through the Gi pathway, which leads to a decrease in cAMP level. This decreased local or compartmentalized accumulation of cAMP results in the reduced activation of Cl channel in the vicinity by CFTR agonists (e.g., ADO). Our findings shed light on the detailed mechanism that underlies the transduction pathway of the LPA2 agonists, and helps to clarify how LPA inhibits CFTR-dependent Cl channel activity; all of this will lead to the alleviation of diarrhea. Because LPA is richly available in certain forms of foods (14, 15), our proof of concept study might pave the way for the use of certain diets to control diarrhea.
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MATERIALS AND METHODS |
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Antibodies, reagents, and constructs.
Cystic fibrosis transmembrane conductance regulator (CFTR) antibodies, R1104 monoclonal mouse antibody and NBD-R polyclonal rabbit antibody, have been described previously (57). Anti-LPA2 antibody (rabbit-2143) against the last 11 amino acids (aa's 341351) of LPA2 and antiNa+/H+ exchanger regulatory factor (NHERF)-2 antibody (rabbit-2346) against the full-length NHERF2 protein were generated by Genemed Synthesis, CA. Anti-Flag mAb, cholera toxin (CTX), and pertussis toxin (PTX) were obtained from Sigma-Aldrich. Maltose binding protein (MBP)-fusion proteins for COOH-terminal tails of LPA1 (aa's 316364), LPA2 (aa's 298351), and LPA3 (aa's 298353) were generated using pMAL vectors (New England Biolabs, Inc.). Flag-tagged full-length LPA2, LPA2-STL, and LPA2-L351A were generated using pcDNA-3 vector (Invitrogen) and QuickChange mutagenesis kit (Stratagene). The peptide delivery system (Chariot) was procured from Active Motif. LPA 18:1, sphingosine-1-phosphate (S1P), and phosphatidic acid (PA) were purchased from Avanti Polar Lipids, Inc. Other lipids, LPA (20:4) and LPA (18:2), have been described (21). Other reagents were described previously (30).
Pull-down assay.
Pull-down assay was performed as described (30). In brief, BHK or COS-7 cells expressing Flag-tagged LPA receptors (or CFTR) were lysed in lysis buffer (PBS 0.2% Triton X-100 plus protease inhibitors). Cell lysates were mixed at 4°C for 15 min followed by centrifugation at 15,000 g for 10 min at 4°C. Glutathione S-transferase (GST), GSTNHERF1, and GSTNHERF2 (03 µM) were added and mixed at 4°C with the clear supernatant or MBP-CLPA2 for the direct binding between MBP-CLPA2 and GSTfusion proteins. After 3060 min incubation, glutathione sepharose beads (20 µl) were added and mixed for an additional 3 h. Thereafter, the mixture was spun at 800 g for 2 min, and the beads were washed three times with the same lysis buffer before the proteins were eluted from beads with Laemmli sample buffer (containing 2.5% ß-mercaptoethanol). Eluates were separated on 415% gel, and were immunoblotted with anti-FLAG and anti-LPA2 antibodies.
Coimmunoprecipitation and immunoblotting.
Cells were harvested and processed as described previously (30). In brief, BHK cells (cross-linked with dithiobis(succinimidyl)propionate as described before [reference 30]) and epithelial cells (HT29-CL19A and Calu-3) were solubilized in RIPA buffer (plus protease inhibitors) on ice for 20 min, and lysates were spun at 15,000 g for 15 min at 4°C to pellet insoluble material. Protein concentration of the cell lysates was determined by the bicinchoninic assay (Pierce Chemical Co.). For coimmunoprecipitation, CFTR monoclonal antibody (R1104 IgG, 1.0 µg) was cross-linked to 20 µl protein A/G agarose as described previously (57). The supernatant was incubated with the cross-linked beads overnight at 4°C under a constant mixing. The beads were washed three times with RIPA buffer before the immunoprecipitated proteins were eluted with sample buffer containing 2.5% ß-mercaptoethanol. The immunoprecipitated proteins were separated on 415% gel, and blotted using polyclonal anti-NHERF2 IgG, anti-LPA2 IgG, and anti-CFTR IgG (NBD-R IgG). For immunoblotting, cell lysates were separated on 415% SDS-PAGE, transferred to PVDF membranes, and immunoblotted for CFTR, NHERF2, and LPA2 using specific antibodies.
Short-circuit current measurements.
Calu-3 and HT29-CL19A polarized cell monolayers were grown to confluency on Costar Transwell permeable supports (filter area is 0.33 cm2). Filters were mounted in an Ussing chamber, and short-circuit currents (Isc) mediated through CFTR Cl channel were performed as described (30). Epithelia were bathed in Ringer's solution (mM) (serosal/basolateral: 140 NaCl, 5 KCl, 0.36 K2HPO4, 0.44 KH2PO4, 1.3 CaCl2, 0.5 MgCl2, 4.2 NaHCO3, 10 Hepes, 10 glucose, pH 7.2, [Cl] = 149), and low Cl Ringer's solution (mM) (luminal/apical: 133.3 Na-gluconate, 5 K-gluconate, 2.5 NaCl, 0.36 K2HPO4, 0.44 KH2PO4, 5.7 CaCl2, 0.5 MgCl2, 4.2 NaHCO3, 10 Hepes, 10 mannitol, pH 7.2, [Cl] = 14.8) at 37°C, and gassed with 95% O2 and 5% CO2. LPA (20:4) was added into the luminal and serosal sides of the cell monolayers for 30 min before ADO was added into the luminal side. In parallel, some filters also were pretreated for 30 min with PA (035 µM) and S1P (035 µM) before ADO addition. CFTR Cl channel inhibitor DPC (500 µM; added into luminal side) was used to inhibit the Cl currents toward the end of the experiment.
To demonstrate the effect of LPA on Cl transport in the mouse mucosa, the mice were killed, and the distal small intestine (24 cm proximal to the cecum) was removed and immediately placed in ice-cold, oxygenated Ringer solution, and opened along the mesenteric border. Indomethacin (10 µM) was present in the rinse and experimental Ringer solutions to prevent prostanoid generation during tissue manipulation. A patch of small intestine proximal to the cecum was stripped of serosal and smooth muscle layers (34) before being mounted in a Ussing chamber (0.112 cm2 exposed surface area). Any nerve plexus activity in the partially remaining myenteric plexus was blocked by tetrodotoxin acetate (1 µM), and local prostaglandin E production was inhibited by indomethacin (10 µM). LPA (20:4; 035 µM) was added into the luminal and serosal sides of the isolated mouse intestine for 30 min. cpt-cAMP (0100 µM) or ADO (0100 µM) were applied to both sides to elicit a CFTR-dependent Cl current response, followed by luminal addition of diphenylamine-2-carboxylate (DPC; 500 µM) toward the end of the experiment. The epithelial integrity was monitored by passing a 3-mV pulse (for cultured cells) or 0.5-mV pulse (for isolated intestinal preparation) across the epithelia every min during the entire experiments.
Intestinal fluid secretion (in vivo) experiments.
The method of Verkman's group was followed with modifications (38). CD1 mice (body weight 2022 g; obtained from Charles River Laboratories) were held off food for 24 h before inducing anesthesia using pentobarbital (60 mg/kg). Mouse body temperature was maintained at 3638°C during surgery using a circulating water heating pad. A small abdominal incision was made to expose the small intestine. Ileal loops (20 mm) proximal to the cecum were exteriorized and isolated (two loops per mouse: 1 loop PBS, 1 loop PBS plus CTX). The closed-loops were injected with 100 µl PBS alone or PBS containing CTX (1100 µg). The control and CTX-injected loops were tested in the presence and absence of LPA (1040 µM). The abdominal incision and skin incision were closed with wound clips, and the mice were allowed to recover. Intestinal loops were collected in a terminal procedure 6 h later. PA (10100 µM) and S1P (10100 µM) also were used in parallel experiments as described above.
In the open-loop mouse model of secretory diarrhea, mice were given single doses of CTX (1, 10, or 100 µg) dissolved in 100 µl 7% NaHCO3 buffer (or buffer alone) in the presence or absence of LPA 20:4 (BSA-complexed, 75, 250, or 750 µg/kg body weight) delivered by an orogastric feeding needle. In some experiments, oral administration of LPA 20:4 (BSA-complexed, 1,000 µg/kg body weight) was performed at different time points (02 h) after CTX feeding. 6 h later, the mice were killed. The entire small intestine (from pylorus to cecum) was exteriorized and isolated with care to avoid tissue rupture and fluid loss. The associated mesentery and connective tissues were removed, and the weight of the intestinal tissue and the enclosed fluid was determined. The intestine was opened longitudinally to remove luminal fluid by blotting, and was weighed again. Intestinal fluid accumulation was determined from the ratio of intestinal weight before/intestinal weight after luminal fluid removal.
Macromolecular complex assembly.
This assay (24) was performed using maltose binding protein (MBP)-CFTR-C tail fusion protein (1 µM) immobilized on amylose beads (20 µl) and incubated with GSTNHERF2. This step, which is called pairwise binding, was done in 200 µl lysis buffer (PBS-0.2% Triton X-100 plus protease inhibitors) and mixed at 22°C for 2 h. The complex was washed once with the same buffer, and allowed to bind Flag-tagged LPA2 from lysates of BHK cells expressing the receptor. The binding was done at 4°C for 3 h with constant mixing. The complex was washed with lysis buffer, eluted with sample buffer, and analyzed by immunoblotting using anti-Flag mAb.
Delivery of LPA2 receptor-specific peptide and immunostaining.
Delivery of LPA2-specific peptide was performed using the Chariot system according to the manufacturer's instructions (Active Motif) (45). In brief, 1.2 µM peptide containing the PDZ motif of LPA2 (last 11 aa's, 341351, ENGHPLMDSTL) was mixed with Chariot solution (total volume: 400 µl) at room temperature for 30 min. The Chariot-peptide complex was added to luminal and serosal sides of polarized Calu-3 and HT29-CL19A cells grown on permeable supports and incubated for 1 h at 37°C in a humidified atmosphere containing 5% CO2 before mounting in an Ussing chamber. At the end of the experiment, the epithelial cells were fixed and immunostained for the efficiency of the peptide delivery using anti-LPA2 antibody, and were subjected to immunofluorescence confocal microscopy. Other peptide (CFTR peptides, aa's 107117) also was delivered as described before as a control.
Cell-attached single-channel recordings.
Single-channel recordings were obtained from HT29-CL19A cells using the cell-attached configuration (58). Patch-clamp pipettes were obtained using quartz class (Sutter Instrument Co.), and a Sutter model P-2000 puller and had resistance of 68 M. The extracellular (pipette and bath) solution contained 140 mM NMDG-Cl, 2 mM MgCl2, 2 mM CaCl2, 10 mM Hepes, and 200 µM DIDS (to block non-CFTR anion channel pharmacologically) titrated to a pH of 7.4 with NMDG. CFTR channels were activated with 2 µM ADO or 10 µM forskolin included in the pipette or bath solution as indicated. Single-channel currents were recorded continuously for 5 min at a test potential of 60 mV (referenced to the cell interior) delivered from the recording electrode, and were filtered at 1 kHz and sampled at 2 kHz. All experiments were conducted at room temperature (2224°C) using an EPC-9 patch clamp amplifier (HEKA Electronik GmbH) and the Pulse + PulseFit V 8.65 acquisition program (HEKA Electronik GmbH). Data analysis was performed using TacX4.1.5 (Bruxton Corp.).
Measurement of cAMP level in cultured cells and intestine tissue.
Calu-3 or HT29-CL19A cells were cultured (100-µl volumes) in standard 96-well microplates (tissue-culture grade), with cell concentrations of between 104106 cells/ml. The plates were incubated overnight at 37°C (5% CO2 and 95% humidity). On the third day, the medium was decanted and 100 µl PBS (± LPA) was added and incubated for 2030 min before the addition of agonists (ADO, or CFTR agonist cocktail) and incubation for another 5 min. The cells were lysed, and intracellular cAMP levels were measured using a cAMP Biotrak competitive enzyme immunoassay system (GE Healthcare) according to the manufacturer's instructions. For the measurement of cAMP levels in the native intestine tissue, ileal loops (2 cm) were exposed to 0.25100 µg CTX. After 6 h of CTX treatment, the excised tissue sections were frozen rapidly in liquid nitrogen and samples were stored at 20°C until the cAMP assay was performed using the liquid phase extraction method according to the manufacturer's instructions (cAMP Biotrak enzyme immunoassay, GE Healthcare).
Statistical analysis.
Results are presented as mean ± SEM for the indicated number of experiments. Statistical analyses were performed using Student's t test and one-way ANOVA. A value of P < 0.05, P < 0.01, or P < 0.001 was considered to be statistically significant.
Online supplemental material.
Fig. S1 shows that LPA2 mRNA is detected in mouse intestinal tissue and various epithelial cells. Fig. S2 shows that LPA2 interacts preferentially with NHERF2. Fig. S3 shows that LPA inhibits CFTR-dependent Cl currents in isolated intestine epithelia, but failed to inhibit Cl currents when CFTR was activated by cocktail. Fig. S4 shows the single-channel recordings of CFTR in cell-attached configuration in HT29-CL19A cells in response to cpt-cAMP stimulation with or with LPA pretreatment. Fig. S5 shows that CTX induced a dose-dependent increase of cAMP production in gut tissue, and intact LPA is recovered 2 h later from intestine when complexed with BSA. Fig. S6 shows that LPA2-specific peptide prevented the LPA inhibition of CFTR-mediated Isc in polarized Calu-3 cells. Online supplemental material is available at http://www.jem.org/cgi/content/full/jem.20050421/DC1.
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Acknowledgments |
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C. Li is a recipient of the Dorothy K. and Daniel L. Gerwin graduate scholarship, and the Leonard Share Young Investigator Award from the University of Tennessee Health Science Center. This work was supported by grants from the National Institutes of Health (to A.P. Naren , G.J. Tigyi, and D.J. Nelson), and a Career Investigator Award from the American Lung Association (to A.P. Naren).
The authors have no conflicting financial interests.
Submitted: 23 February 2005
Accepted: 9 August 2005
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References |
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