Dr Mary H. Perdue, Intestinal Disease Research Program, McMaster University, HSC-3N5C, 1200 Main St W, Hamilton ON, Canada L8N 3Z5. Tel: +1 905 525 9140 x22591; e-mail: email@example.com
Abstract The intestinal epithelium acts as a barrier restricting uptake of luminal macromolecules such as dietary antigens and microbes. Here, we examined the role of cholinergic signalling in the regulation of permeability to macromolecules. Mouse jejunum was mounted in Ussing chambers and permeability was determined by measuring the flux of the antigen-sized protein, horseradish peroxidase (HRP), across the tissue. Baseline HRP permeability was significantly reduced by neural blockade with tetrodotoxin or cholinergic muscarinic antagonism with atropine, suggesting that ongoing release of endogenous acetylcholine from enteric nerves regulates barrier function. Exogenous addition of the muscarinic agonist bethanechol caused significant increases in both HRP flux and the area of HRP-containing endosomes in enterocytes. Bethanechol-enhanced HRP flux was abrogated by the M3 receptor antagonist, 4-diphenylacetoxy-N-methylpiperidine methiodide (4-DAMP), the phospholipase A2 inhibitor quinacrine, and the cyclooxygenase inhibitor indomethacin. Complementary in vitro studies showed direct effects of bethanechol on T84 epithelial cells, where increased HRP uptake was associated with increased F-actin, and increased cytosolic phospholipase A2 (cPLA2) phosphorylation. Taken together, these results provide evidence for cholinergic regulation of transepithelial transport of macromolecules, mainly mediated by activation of M3 receptors with subsequent involvement of phospholipase A2 and cyclooxygenase products.
The intestinal mucosa is lined by a single layer of epithelial cells which forms the barrier between the contents of the intestinal lumen and the internal milieu of the body proper. This polarized epithelial layer must achieve efficient absorption of nutrients and ions while preventing uptake of noxious luminal antigens, microbes and toxins. Penetration of this barrier by antigens does occur to a limited degree under normal physiological conditions,1 and is significantly increased in many intestinal diseases (reviewed in Ref. 2). A permeability defect at the level of the epithelium is thought to play either a primary aetiologic role in the evolution of intestinal disorders such as inflammatory bowel disease (IBD), or to be a secondary consequence of disease which perpetuates intestinal inflammation.2
While barrier dysfunction is associated with intestinal disease, antigen penetration across the epithelium is also important for the initiation of appropriate immune responses, such as tolerance to dietary antigens and commensal flora. Regulation of epithelial permeability is thus a key factor in the balance between immunosurveillance and inflammation of the gut.
Permeation of the intestinal epithelial barrier occurs via the paracellular or transcellular route. For paracellular traffic, molecules must penetrate the size-selective and dynamic tight junctions, located at the apical pole of the lateral intercellular space between enterocytes, while larger molecules, such as macromolecular protein antigens, cross mainly via the transcellular route, enclosed in endosomes which must avoid lysosomal degradation to gain access to the lamina propria (reviewed in Ref. 3).
While the exact mechanisms by which the intestinal epithelial barrier is regulated have not been fully elucidated, numerous stimuli including luminal contents, microorganisms and cytokines are known to affect the permeability of tight junctions (for review see Ref. 4), while the regulation of transcellular permeability to macromolecules is not as well understood.
The role of the enteric nervous system in regulating epithelial permeability has been studied to a limited extent. Evidence suggests that cholinergic signalling increases permeability5 and may mediate stress-induced barrier dysfunction.6 Cholinergic signalling may also play a role in inflammatory processes as muscarinic stimulation can incite production of pro-inflammatory prostanoids and leukotrienes7 and act directly on immune cells.8,9 Acetylcholine (ACh) in the intestine is derived from two sources: neuronal ACh supplied by direct innervation of the mucosa by the enteric nervous system,10 and non-neuronal ACh produced locally within the mucosa by epithelial cells and lymphocytes.9,11
In this study, we tested the hypothesis that cholinergic signalling modulates permeability to macromolecules in mouse small intestine. Specifically, we examined the role of muscarinic stimulation in permeability to intact protein. We conducted parallel studies in cultured colonic epithelial T84 cells, which allowed for examination of cytoskeletal changes and phosphorylation of cytosolic phospholipase A2 (cPLA2), and confirmation of direct effects on the epithelium. We present evidence that muscarinic signalling elevates intestinal permeability to macromolecules via the M3 receptor on epithelial cells resulting in activation of cPLA2 and cyclooxygenase.
Materials and methods
Male Balb/c mice (mean age 9 weeks) were obtained from Harlan Sprague–Dawley Inc. (Indianapolis, IN, USA). Mice were not restricted by diet or activity during housing. After 1 week of acclimatization, mice were killed and the small intestine was removed for Ussing chamber experiments. All animal experiments described in this paper were conducted with approval from the McMaster University Animal Care Committee.
Tetrodotoxin (TTX, 10−6 mol L−1), atropine (ATR, 10−6 mol L−1), bethanechol (BET, 10−4 mol L−1), 4-diphenylacetoxy-N-methylpiperidine methiodide (4-DAMP, 10−7 mol L−1), quinacrine (10−5 mol L−1), indomethacin (10−5 mol L−1) and horseradish peroxidase type II (HRP) were purchased from Sigma (St Louis, MO, USA) (all dissolved in PBS). Working concentrations for all reagents were based on previously published findings (TTX,12 BET,13–16 4-DAMP,16,17 quinacrine,18 and indomethacin19). BODIPY FL Phallicidin was purchased from Molecular Probes (Invitrogen, Burlington, ON, Canada).
Ussing chamber studies
A 12-cm segment of proximal small intestine taken from naïve mice was opened along the mesenteric border and cut into four flat sheets. Full thickness segments (devoid of Peyer's patches) were mounted in modified Ussing chambers and bathed with oxygenated Krebs buffer containing (in mmol L−1): 115 NaCl, 1.25 CaCl2, 1.2 MgCl2, 2.0 KH2PO4 and 25 NaHCO3 (pH 7.35 ± 0.02, at 37 °C). In the Ussing chambers, an area of 0.6 cm2 of tissue was exposed to 8 mL of buffer circulating on each side. In addition, the serosal buffer contained 10 mmol L−1 glucose as an energy source, osmotically balanced by 10 mmol L−1 mannitol in the mucosal buffer. Tissues were short-circuited at 0 V using a World Precision Instruments automated voltage clamp (Narco Scientific, Mississauga, Ont, Canada). Conductance (G) was calculated according to Ohm's law, using potential difference and short-circuit current (Isc) values. Tissues with abnormally high initial G values (>50 mS cm−1) or whose G exceeded this value during the course of the experiment were considered damaged and were excluded from subsequent analysis. After a 15-min equilibration period, pharmacological agonists or antagonists were added to the serosal buffer.
Measurement of permeability
Flux of HRP The transcellular passage of HRP (44 kDa), a model protein antigen, was determined by flux and electron microscopy (see below) as previously described.5 Briefly, type II HRP was added to the mucosal buffer in the Ussing chamber at a concentration of 4.5 × 10−5 mol L−1, and following a 30-min equilibration period, 500 μL samples were obtained (at 30-min intervals for 2 h) from the serosal buffer and replaced with Kreb's buffer to maintain a constant volume in the chambers. The amount of intact HRP crossing the intestine was determined by measuring HRP activity using a modified Worthington method using a kinetic assay in a 96-well plate reader.20 Mucosal-to-serosal fluxes of HRP were calculated according to standard formulae and expressed as pmol cm−2 h−1. Calculations were made based on two stable time periods (60–90 and 90–120 min) after the flux of HRP had achieved steady state. In cases where a number of treatments were compared, values were expressed as percent of the control in each experiment.
Flux of 51Cr-EDTA 51Cr-ethylenediaminetetraacetic acid (EDTA; Perkin-Elmer, Vaudreuil-Dorion, QC, Canada) is a small inert probe (molecular weight 360 Da), generally considered to be a marker of paracellular permeability. Transepithelial flux of this probe was determined using methods previously described.21 Briefly, 6 μCi mL−1 was added to the mucosal buffer bathing jejunal segments mounted in Ussing chambers. Non-radioactive Cr-EDTA was added to the serosal buffer at an equal concentration. Following a 30-min equilibration period, 500 μL samples from the ‘cold’ buffer and 50 μL samples from the ‘hot’ buffer were collected and replaced with the appropriate volume of buffer every 30 min for 2 h. Radioactivity was measured in a gamma counter and flux expressed in pmol cm−2 h−1. In cases where a number of treatments were compared, values were expressed as percent of the control in each experiment.
Quantification of HRP-containing endosomes in enterocytes Following exposure of the intestine in Ussing chambers to BET, TTX or saline for 15 min, followed by luminal HRP exposure for 30 min (at which point steady state was reached), the segment was removed and immersed in 2% glutaraldehyde in 0.1 mol L−1 sodium cacodylate buffer for 2 h at room temperature, transferred to sodium cacodylate buffer, and stored at 4 °C overnight. The tissue was washed three times in 0.05 mol L−1 Tris buffer and then incubated for 30 min in 5 mg of 3,3′-diaminobenzadine tetrahydrochlorine (Sigma) in 10 mL of 0.05 mol L−1 Tris buffer and 0.01% hydrogen peroxide. The tissue was subsequently processed for electron microscopy and photomicrographs of well-oriented epithelial cells were prepared. The total area occupied by HRP-product-containing endosomes within a defined size window in enterocytes was measured in coded photomicrographs by one investigator (HLC, blinded to treatment variables) using a computer-supported image analysis system as previously described.22 Windows were selected at random using computerized random number generation to avoid observer bias. A total of 20 windows (magnification 5000×) for each treatment group were analysed (four per tissue). Results were expressed as area of HRP-containing endosomes per optical window (300 μm2).
Epithelial cell culture
T84 human epithelial cells were cultured at 37 °C with 5% CO2 in a 1 : 1 mixture of Dubecco's modified eagle medium (DMEM) and Ham's F-12 medium supplemented with 2% penicillin–streptomycin, 0.6%l-glutamine, 1.92% NaHCO3 (all from Invitrogen) and 10% foetal calf serum (CanSera, Toronto, ON, Canada). For permeability assays, one million cells were seeded onto 1 cm2 semi-permeable filter supports (pore size 400 nm) in a Transwell system (Corning Costar, Cornell, NY, USA) and cultured until confluent. Cell monolayers were used when transepithelial resistance (TER) was ≥1000 Ω cm−2 as measured by a voltmeter and chopstick electrode (Millipore, Bedford, MA, USA). After 24 h serum starvation, agonists and antagonists were added to the basal compartment, and at the same time, HRP (4.5 × 10−5 mol L−1) was added to the apical compartment of the culture well. Aliquots were collected from the basal compartment 24 h later, and flux expressed as pmol cm−2 h−1 or as percent compared with the control value.23
Epithelial F-actin quantification
For F-actin quantification, BET-treated T84 monolayers grown on semi-permeable filter supports were washed with phosphate-buffered saline (PBS), fixed in 10% neutral-buffered formalin, permeabilized with 0.2% (vol/vol) Triton X-100 in PBS for 5 min, according to the method of Goldblum et al.24 The monolayers were treated with 1.5 × 10−7 mol L−1 BODIPY Phallicidin (Molecular Probes, Eugene, OR, USA) in PBS for 20 min followed by methanol extraction using 100% methanol for 1 h at 37 °C. Fluorescence was measured from the resulting supernatants in a SPECTRAmax Gemini XS microplate spectrofluorometer (Molecular Devices, Sunnyvale, CA, USA) at excitation and emission wavelengths of 490 and 520 nm, respectively. Results were expressed as percentage of control (arbitrary fluorescence units).
Western blot analysis
T84 cells were seeded on 2-cm2 semi-permeable filter supports (2 million cells per well) and cultured until confluent. Cells were serum-starved overnight and treated with BET (10−4 mol L−1) for various timepoints. Whole-cell lysates were prepared by rocking cells in ice-cold radio-immunoprecipitation assay (RIPA) buffer containing protease inhibitors (complete protease inhibitor cocktail; Roche, Indianapolis, IN, USA) and phosphatase inhibitors (100 mmol L−1 NaF, 100 mmol L−1 NaVO3). Lysates were clarified by centrifugation, and the supernatant was stored at −70 °C. Protein concentration was determined using a Bradford microplate assay (Bio-Rad, Hercules, CA, USA). Samples (40 μg protein) in reducing loading buffer were boiled and electrophoresed through 8% (29 : 1 acrylamide/bisacrylamide) sodium dedecyl sulphate (SDS) gels. Separated proteins were electroblotted onto polyvinylidine difluoride (PVDF) membrane (PerkinElmer, Wellesley, MA, USA) and blocked in 5% non-fat Carnation powdered milk/Tris-buffered saline/Tween 20 (TBST) for 1 h. Primary antibodies used were anti-phospho-cPLA2 (Ser505), 1 : 1000; anti-cPLA2, 1 : 1000, (Cell Signaling, Danvers, MA, USA). Blots were washed and incubated with secondary antibody-HRP conjugates for 1 h (goat anti-rabbit at 1 : 3000; Santa Cruz Biotechnology, Santa Cruz, CA, USA), washed extensively and immunoreactive proteins were visualized by using enhanced luminol (PerkinElmer) and by exposing the membrane to Kodak BioMax film (PerkinElmer). The optical density of the immunoreactive band for phospho-cPLA2 was quantified using Image J software (U.S. National Institutes of Health, Bethesda, Maryland, USA). Density was corrected for background and was normalized to the density of the corresponding band for unphosphorylated cPLA2. The data were expressed as a ratio of the optical densities of phosphorylated cPLA2 to unphosphorylated cPLA2 (whose expression was unaffected by treatment with BET).
Results are expressed as mean ± standard error of the mean (SEM). One-way analysis of variance was used with Newman–Keuls as a subsequent multiple-comparison test. P-values <0.05 were considered significant.
Role of muscarinic signalling in intestinal permeability
Compared with untreated tissues, TTX induced a significant decrease (P < 0.01) in the flux of HRP (Fig. 1A), indicating a role for spontaneous release of neurotransmitters in regulating permeability to macromolecules. Treatment with ATR also significantly reduced (P < 0.01) the flux of HRP, suggesting the involvement of ACh and muscarinic receptors mediating this effect. To confirm the involvement of muscarinic signalling in mediating intestinal permeability we applied the parasympathomimetic choline ester BET which selectively stimulates muscarinic receptors. BET caused a significant increase (P < 0.01) in HRP flux which was not affected by pretreatment of tissues with TTX but was inhibited by pretreatment with ATR. This finding implies that muscarinic signalling enhances permeability to macromolecules via receptors not located on nerves, but by receptors on other cell types, likely epithelial muscarinic receptors. In contrast, we observed no changes in baseline flux of the paracellular probe 51Cr-EDTA in response to TTX or ATR compared with untreated tissues (Fig. 1B). Similar to macromolecular permeability, BET increased paracellular 51Cr-EDTA flux (P < 0.01), which was not affected by pretreatment with TTX but was attenuated by pretreatment with ATR.
Transcellular transport of HRP
We used electron microscopy to visualize the transport pathway of HRP across the epithelium. In all tissues, HRP was observed within endosomes in enterocytes (representative electron photomicrograph of tissue shown in Fig. 2A–C). Compared with untreated tissues, TTX significantly reduced (P < 0.01) the area of HRP-containing endosomes in enterocytes (Fig. 2D), suggesting a downregulation of endocytosis. In contrast, BET treatment increased (P < 0.01) the area of HRP-containing endosomes (Fig. 2D), indicating upregulated endocytosis of fluid-phase macromolecules into enterocytes. HRP was not visualized within the paracellular spaces or the tight junctions in any tissue observed.
Signalling pathway of BET-induced epithelial permeability
To examine if BET increased epithelial permeability via M3 receptors on epithelial cells, we treated jejunum with 4-DAMP, a muscarinic antagonist with selectivity for the M3 receptor. Pretreatment with 4-DAMP significantly attenuated (P < 0.01) the BET-induced permeability to HRP (Fig. 3A). We used the phospholipase A2 inhibitor quinacrine to test the hypothesis that BET acted to increase permeability via phospholipase A2 activation. Quinacrine pretreatment significantly inhibited (P < 0.01) the BET-induced hyperpermeability. To examine signalling downstream of phospholipase A2, we used indomethacin to inhibit cyclooxygenase activity. Indomethacin pretreatment significantly reduced (P < 0.01) the BET-induced permeability to HRP. In contrast, pretreatment with 4-DAMP, quinacrine or indomethacin had no effect on the BET-induced increase in 51Cr-EDTA flux (Fig. 3B). 4-DAMP, quinacrine and indomethacin had no significant effect on basal permeability to HRP (data not shown).
Permeability in cultured epithelial monolayers
We used tight junction-forming T84 epithelial monolayers to determine if the effects of BET were mediated at the level of the epithelial cell. BET induced a significant increase (P < 0.01) in HRP flux (Fig. 4A) which was abrogated by pretreatment with 4-DAMP, quinacrine or indomethacin. These findings indicate that BET acts directly on epithelial M3 receptors to enhance permeability to macromolecules, followed by activation of phospholipase A2 and cyclooxygenase within these cells. In T84 monolayers, BET did not have any measurable effect on transepithelial resistance or permeability to the paracellular probe 51Cr-EDTA (data not shown).
To determine if the changes in epithelial transcellular permeability were accompanied by changes in the F-actin cytoskeleton, we treated T84 cell monolayers with BET and observed increased F-actin polymerization, with significantly increased F-actin observed 5 min following BET treatment (Fig. 5). Levels of F-actin continued to increase in a time-dependent manner, with maximal changes observed at 60 min and levels approximating pretreatment at 120 min.
Phosphorylation of cPLA2
To examine the regulation of cPLA2 by phosphorylation following BET treatment, we treated T84 monolayers with BET for 5–120 min followed by cell lysis, protein extraction and Western blot analysis for phosphorylation at Serine505. BET induced a time-dependent increase in cPLA2 phosphorylation at Serine505 (Fig. 6).
The results from this study demonstrate for the first time that cholinergic signalling increases permeability to macromolecules in the mouse jejunum via a pathway mediated by muscarinic receptors on epithelial cells. We have demonstrated that muscarinic stimulation increases intestinal permeation of intact protein antigen transiting via endosomes. Our data suggest that the mechanism for this increased permeability is mediated mainly by epithelial M3 receptors and the subsequent activation of phopholipase A2 and cyclooxygenase. At the level of the epithelial cell, muscarinic stimulation induces F-actin polymerization. Our findings indicate that the mechanism for cholinergic-mediated macromolecular transport via the transcellular pathway appears to be separate from effects on paracellular permeability.
Barrier dysfunction is implicated in the pathogenesis of many intestinal diseases. It has been suggested that impaired barrier function plays a role in either the initiation or exacerbation of inflammation in intestinal disease. For example, intestinal barrier dysfunction is implicated in the pathogenesis of Crohn's disease and is associated with food allergy, celiac disease and necrotizing enterocolitis.2 We speculate that perturbations in cholinergic signalling within the gut could affect intestinal permeability and thus contribute to intestinal dysfunction. ACh can act on both nicotinic and muscarinic receptors in the gut, with nicotinic receptors located on neurons, enterochromaffin cells, macrophages and smooth muscle cells, and muscarinic receptors on enterocytes, enterochromaffin and enteroendocrine L cells, goblet cells, and smooth muscle cells.25–28 In our study, we sought to examine direct effects of cholinergic signalling on epithelial function in mouse intestine known to express both the muscarinic M3 and M1 receptor isoforms on enterocytes29 and human colonic T84 cells expressing only M3 receptors.30
A relationship between cholinergic signalling and intestinal barrier function is suggested by several lines of evidence. First, altered cholinergic metabolism has been reported in the context of intestinal inflammation associated with increased permeability.13,31 Increased levels of choline acetyltransferase were observed in a mouse model of colitis13 and parasite-infected rat jejunum.31 Secondly, cholinergic signalling was found to mediate intestinal barrier dysfunction in a rat model of stress6 providing a link between ACh and intestinal permeability. In the gut, both neuronal and tissue-derived non-neuronal ACh11 may contribute to the regulation of intestinal permeability providing additional complexity to the role of cholinergic signalling in the regulation of intestinal physiology. It is important to consider that altered barrier function may not necessarily be indicative of pathophysiology, but that increased endocytosis of luminal contents could be a physiologically adaptive response to increase luminal antigen sampling.
In the present study, we identified that tonic release of ACh contributes to baseline jejunal permeability to macromolecules, which appears to be mediated via muscarinic receptors. To further investigate the role of muscarinic signalling in the regulation of permeability to macromolecules, we stimulated mouse jejunum with the muscarinic agonist BET which significantly increased the flux of HRP. Electron microscopy demonstrated that BET dramatically enlarged the area of HRP-containing endosomes resulting from increased numbers and sizes of HRP-containing endosomes. Taken together, these findings suggest that muscarinic signalling in the jejunum increases the rate of fluid-phase endocytosis of protein, some of which escapes lysosomal degradation in the enterocyte, thereby increasing the penetration of antigens across the epithelial barrier. HRP was not observed in the paracellular spaces between adjacent enterocytes, suggesting that although BET induces increased transcellular and paracellular permeability, the effects on the tight junctions are not adequate to permit penetration of large macromolecules. We observed that muscarinic inhibition (with ATR) but not neuronal blockade (with TTX) abrogated the increased transcellular and paracellular permeability, indicating that cholinergic effects on permeability are regulated via non-neuronal receptors, probably directly via epithelial muscarinic receptors.
To investigate the signalling pathway for muscarinic-stimulated macromolecular uptake, we pretreated tissues with the M3 receptor antagonist 4-DAMP which attenuated the BET-induced increase in intact HRP transport. We hypothesized that BET acted directly on muscarinic receptors, probably M3R, located on the basolateral surface of epithelial cells and confirmed direct action on epithelial cells in vitro using cultured T84 epithelial cells, which form tight junctions and are a well-established model for epithelial barrier function. We observed that cultured epithelial monolayers showed increased HRP transport when treated basolaterally with BET and that this effect was abrogated by pretreatment with 4-DAMP, suggesting direct action on epithelial M3 receptors. The gastrointestinal phenotype of M3R null mice has not yet been well characterized, although a role for M3R in colonic anion secretion,29 longitudinal muscle contractility,32 and gastric pepsinogen secretion33 are well established in this model. Unfortunately, intestinal permeability was not addressed in these studies and based on the findings of the present study, merits further exploration. While our data suggest that the effects on macromolecular transport are mediated by epithelial M3 receptors, other muscarinic receptor isoforms, namely M1 also probably play a role. Recently it has been shown that M1 receptors on colonic epithelial cells play a role in mediating ion secretion in the mouse colon, a phenomenon previously thought to be mediated solely by epithelial M3 receptors.29 The contribution of more than one muscarinic receptor isoform to the regulation of epithelial permeability may explain why 4-DAMP (selective for M3) did not reduce baseline permeability as did muscarinic inhibition with ATR.
M3 receptor stimulation is known to activate cytoplasmic phospholipase A2 (cPLA2)34 via increased intracellular calcium and activation of protein kinase C (PKC) but independent from inositol triphosphate (IP3)-mediated calcium release.35 Of note, PKC has been shown to play a role in the regulation of paracellular permeability via modulation of tight-junction proteins,36 but with no known direct effects on macromolecular transport. We hypothesized that muscarinic stimulated cPLA2 activity could be a mediator of this effect. PLA2 comprises a family of lipolytic enzymes that catalyse the hydrolysis of membrane phospholipids generating arachidonic acid which undergoes conversion through several metabolic pathways leading to the biosynthesis of lipoxins, prostaglandins and leukotrienes. PLA2 has been implicated in human intestinal disease associated with increased permeability,37 with improvement of colitis observed following PLA2 inhibition in model of IBD.38,39 PLA2 has also been shown to upregulate endocytosis events at the apical membrane of polarized cells, with increased apical endocytosis of both membrane-bound and fluid-phase macromolecules.40,41 This effect appeared to be mediated by prostaglandins40, suggesting that activation of PLA2 and subsequent generation of arachidonic acid metabolites, specifically the prostaglandins, may be key intracellular mediators for cholinergic stimulation-induced permeability. We used quinacrine to inhibit PLA2 activity and observed complete attenuation of the BET-induced HRP-permeability in mouse jejunum and in T84 cells. We also demonstrated that in T84 cells, BET leads to a time-dependent increase in cPLA2 phosphorylation at Ser505– probably via M3R activation followed by sequential activation of PLC-β1, PKC and ERK1/2.
To test whether cyclooxygenase products played a role in mediating BET-induced hyperpermeability, we pretreated tissues with the cyclooxygenase inhibitor indomethacin and observed complete attenuation of the BET-enhanced permeability. This finding is supported by a study by Curtis et al. identifying a role for prostaglandins in mediating mucosal to serosal macromolecular transport of intact protein in rat stomach.42 As the cell cytoskeleton is important for the trafficking of material through the cell, we examined the polymerization of the major cytoskeletal protein F-actin in cultured T84 epithelial cells. We observed a time-dependent increase in total F-actin concentration. Interestingly, phosphorylation of cPLA2 and increased actin polymerization in response to BET showed similar time-dependent increases with a fading-out phenomenon at 120 min. This finding suggests that these events may be self-limiting, and that some endogenous downregulation of this response may be occurring. As apical endocytosis is increased in response to BET stimulation, it is perhaps not surprising that there is also increased polymerization of cytoskeletal transport machinery within the cell. Actin is known to play a role in the internalization of endocytic vesicles, and both actin and the microtubule network are required for efficient transcytosis and delivery of endosome contents to late endosomes and lysosomes (reviewed in Ref. 43). In our working hypothesis, BET acts to alter permeability via arachidonic acid metabolites, which is supported by the evidence that lipoxygenase and cyclooxygenase products act as second messengers in other systems to mediate changes in the cell cytoskeleton40,44 and alter apical endocytosis.40
In mouse jejunum the flux of paracellular probe 51Cr-EDTA was increased upon BET stimulation indicating an effect on tight-junction permeability. In rat ileum, Bijlsma et al. reported carbachol enhanced HRP permeability and proposed that cholinergic stimulation induces large pores in the tight junctions, although it was suggested that this effect did not occur in all tight junctions.5 The discrepancies between that study and ours may possibly be explained by species differences or the pharmacological conditions, because in that study the rat tissues were also treated with reagents (bumetanide and BaCl2) to address the role of chloride secretion in permeability changes. Other reports provide evidence for both increased5,45,46 and reduced47 paracellular permeability in response to cholinergic stimulation. Differing results may also be related to the underlying cholinergic tone of the tissue, the fasted or fed state of the animal,47 or the intestinal region investigated. However, it is clear from our studies that the signalling pathway involved in mediating permeability of the paracellular pathway is distinct from that involved in the endocytic/transcytotic transcellular pathway because the BET-induced increase in 51Cr-EDTA flux was not affected by treatment with 4-DAMP, quinacrine or indomethacin. It is possible that effects of muscarinic stimulation on paracellular permeability are mediated by muscarinic receptor isoforms not inhibited by 4-DAMP (i.e. not M3). The existence of other muscarinic receptor isoforms on the epithelium that regulate paracellular permeability would explain why in contrast to ATR, 4-DAMP does not inhibit BET-induced increases in paracellular in mouse jejunum and would also explain why T84 cells known to express only the M3 isoform, do not demonstrate altered paracellular permeability in response to BET.
The enteric nervous system is involved in the regulation of many intestinal epithelial functions, and here we provide evidence for the involvement of cholinergic signalling in the regulation of intestinal transcellular epithelial permeability to macromolecules. We propose that cholinergic stimulation via the M3 receptor on epithelial cells leads to generation of phospholipase A2 and cyclooxygenase metabolites that act to increase apical endocytosis with concomitant changes in the apical cell cytoskeleton. Our data add to the body of knowledge on the role of cholinergic mechanisms in health and disease by demonstrating that muscarinic signalling regulates macromolecular penetration of the epithelial barrier.
This study was supported by grants from the Crohn's and Colitis Foundation of Canada and Canadian Institutes of Health Research (MHP). HLC is the recipient of a Canadian Institutes of Health Research/Canadian Digestive Health Foundation Doctoral Research Scholarship. The authors would like to thank Dr. Ping-Chang Yang for preparation of samples for electron microscopy, and Colin Reardon for technical assistance.