1Institut National de la Santé et de la Recherche Médicale U539 and 2Department of Gastroenterology, Hôtel-Dieu Hospital, 44035 Nantes, France; 3Technische Universität München, Department of Human Biology, 85350 Freising; 4Department of Toxicology, Hannover Medical School, D-30623 Hannover, Germany
Submitted 8 November 2002 ; accepted in final form 30 May 2003
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ABSTRACT |
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c-Fos; voltage-sensitive dyes; cytokines; human submucosal plexus
C. difficile can cause antibiotic-associated diarrhea and colitis in humans (27) due to the secretion of two enterotoxins, toxins A (308 kDa) and B (270 kDa) (15, 16). Toxin A induces the synthesis of proinflammatory cytokines such as IL-1, TNF-
, or IL-8 in macrophages, lymphocytes, or intestinal epithelial cells (5, 11). In animal models, toxin A is responsible for neurally mediated gut inflammation and diarrhea. It has been determined that inflammation induced by toxin A is mediated via tachykinergic pathways (6, 20, 21). Diarrhea induced by toxin A was significantly reduced by the blockade of neuronal nicotinergic pathways (6, 35). Finally, Xia et al. (42) showed that local application of toxin A depolarized guinea pig submucosal neurons directly and blocked electrically induced slow inhibitory postsynaptic potentials.
Toxin B of C. difficile is more potent than toxin A (11, 28), but few data are available concerning its effects on the human gut. Toxin B can increase intestinal epithelial barrier permeability (28) and induce the synthesis of proinflammatory cytokines in monocytes (11). Nevertheless, it is presently unknown whether toxin B can cause the synthesis of proinflammatory cytokines or activate enteric neuronal pathways in the human colon. Until recently, it was not possible to conduct electrophysiological studies of human submucosal neurons, mainly because of technical difficulties. However, recording of membrane potential (Vm) in the human ENS can now be performed by a multisite optical recording technique (MSORT) using voltage-sensitive dye so that electrophysiological characterization of human submucosal enteric pathways is in progress (31). A complementary approach using immunohistochemical methods allows detection of the induction of immediate-early genes, such as c-Fos or c-Jun, in activated neurons. This approach has been used successfully in the ENS to identify enteric neurons activated by various stimuli (19, 29, 32). The purpose of the present study was to characterize the response of human submucosal neurons to toxin B of C. difficile and identify the mediators involved in these effects by immunohistochemical methods and optical recording of Vm.
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MATERIALS AND METHODS |
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Tissues were placed in 4°C oxygenated sterile Krebs solution containing (in mM): 117 NaCl, 4.7 KCl, 1.2 MgCl2 6H2O, 1.2 NaH2PO4, 25 NaHCO3, 2.5 CaCl2, 2 H2O, and 11 glucose and then rapidly transported to the laboratory for experiments. The specimen was pinned flat with the mucosa down in a dissection dish containing ice-cold sterile oxygenated Krebs solution that was changed every 10-15 min. Preparations taken from intertaenial regions were 4-7 cm in circumferential direction and 4-5 cm in longitudinal direction. The longitudinal muscle was then carefully removed under a dissection microscope, washed four times with sterile Krebs solution, and pinned back in a sterile Sylgard-coated petri dish before the addition of sterile culture medium (35 ml). The culture medium (DMEM/F-12; Sigma, St. Louis, MO) was supplemented with 10% heat-inactivated fetal calf serum, 100 IU/ml penicillin, 100 µg/ml streptomycin, 1.1 µg/ml amphotericin B, 20 µg/ml gentamicin, glutamine (all from Sigma) and 2.1 g/l NaHCO3.
After 30 min of incubation to allow tissue equilibration, toxin B of C. difficile was added to the medium. Toxin B from the VPI 10463 strain was purified as previously described (17). The tissue was then maintained for 3 h at 37°C in a humidified incubator containing 5% CO2 and air and continuously shaken on a rocking tray. At the end of organotypic culture, the supernatant was removed and stored at -80°C. Finally, the tissue was fixed for 4-6 h in 0.1 M PBS containing 4% paraformaldehyde at room temperature.
Immunohistochemistry and identification of neuronal cell populations. After the fixation procedure, the tissue was washed in PBS and pinned in a dissection dish. The mucosa and remaining circular muscle were then removed under a dissection microscope. The tissue was pinned in the dish with the mucosa up, and the Meissner's plexus (directly under the mucosa) was carefully dissected with fine scissors. The plexus was then permeabilized for 1-2 h in PBS/NaN3 containing 0.5% Triton X-100 and 4% horse serum before being incubated with a combination of the following primary antibodies diluted in PBS/NaN3, 4% horse serum, and 0.5% Triton X-100 for 18-20 h at room temperature: rat anti-substance P (anti-SP; 1:1,000; Fitzgerald, Concord, CA), mouse anti-VIP (1:1,000; AbCys, Paris, France), and rabbit anti-c-Fos (1:10,000 Ab-5; Oncogene Research Products, San Diego, CA). After incubation with primary antisera, the tissue was washed three times with PBS and incubated for 2-3 h with the following secondary antibodies coupled to fluorophores: donkey anti-rabbit IgG conjugated to carboxymethylindocyanine (1:500), donkey anti-rat IgG conjugated to 7-amino-4-indodicarbocyanin (1:500), and donkey anti-mouse IgG conjugated to 7-amino-4-methyl-coumarin-3-acetate (1: 50) (from Jackson Laboratories, purchased from Euromedex, Mundolsheim, France).
In the next step, tissue was labeled with rabbit anti-NSE (1:3,000; Polysciences, Eppelheim, Germany) for 18-20 h. After incubation with the primary antisera, the tissue was washed with PBS and incubated for 2-3 h with donkey anti-rabbit IgG conjugated to FITC (1:200).
Specimens were viewed under a fluorescence microscope (Olympus IX 50; Olympus, Hamburg, Germany) fitted with adequate filter cubes. Pictures were acquired with a black and white video camera (model 4910, Cohu; SL Microtest, Jena, Germany) connected to a Macintosh computer via a frame-grabber card (Scion Image; SL Microtest). The immunohistochemical study was performed by analyzing at least 15 submucosal ganglia in each preparation. Anti-NSE antibody was used as a general neuronal marker to determine the proportion of the population of VIP and SP immunoreactive c-Fos neurons.
Real-time RT-PCR analysis of IL-1 mRNA levels. For RT-PCR experiments, tissues were placed in Tri Reagent (Euromedex) and maintained at -80°C. Total RNA extraction from tissue cells was performed with Tri Reagent (Euromedex) according to the manufacturer's instructions. For reverse transcription, RNA (1 µg) was combined with 0.5 µg of random hexamers (Amersham, Piscataway, NJ), transcription buffer (50 mM Tris · HCl, pH 8.3, 75 mM KCl, 3 mM MgCl2, 10 mM DTT), dNTPs (10 mM each), RNasin (20 units; Promega, Madison, WI), and RNaseH- Maloney murine leukemia virus reverse transcriptase (200 units; Promega) in a total volume of 20 µl. Incubation was performed at 42°C for 60 min. The amplification conditions of the IL-1
and
-actin templates were optimized for the RotorGene 2000 instrument (Ozyme, Saint Quentin Yvelines Cedex, France). PCR amplifications were performed by using titanium Taq DNA polymerase (Clontech, Ozyme). The reaction mixture contained 2 µl of the supplied 10x titanium Taq PCR buffer, 1 µl of a 1:1,000 dilution of SYBRgreen I (Roche Diagnostic Systems, Basel, Switzerland), 1 µl of each primer (10 pM each), 0.4 µl of titanium Taq DNA polymerase, 0.5 µl of dNTPs (10 mM each), and PCR-grade water to a volume of 18 µl. Microtubes (0.2 ml) were loaded with 18 µl of this master mix and 2 µl of the template (cDNA diluted 1:50 for
-actin and undiluted for IL-1
), and the run was then initiated. Cycling conditions were as follows: denaturation for 5 min at 95°C, amplification for 35 cycles with denaturation for 5 s at 95°C, annealing for 15 s at 63°C for
-actin and 68°C for IL-1
, and extension for 20 s at 72°C. At the end of each cycle, the fluorescence emitted by the SYBRgreen I dye was measured. After completion of the cycling process, samples were subjected to a temperature ramp (from 60 to 99°C) with continuous fluorescence monitoring for melting curve analysis. Primers were chosen on separate exons to amplify cDNA, but not genomic DNA. The following primers were used: IL-1
forward 5'-TGC CCG TCT TCC TGG GAG GG-3', reverse 5'-GGC TGG GGA TTG GCC CTG AA-3';
-actin forward 5'-CCT TCC TGG GCA TGG AGT CCTG-3', reverse 5'-GGA GCA ATG ATC TTG ATC TTC-3'.
For each PCR product, a single narrow peak was obtained by melting curve analysis at the specific melting temperature, indicating specific amplifications. An external standard curve was generated with three serial fivefold dilutions for -actin and IL-1
. The reference curve was constructed by plotting the relative amounts of these dilutions vs. the corresponding threshold cycle values. The correlation coefficient of these curves was >0.99. The amount of IL-1
or
-actin transcripts was calculated from these standard curves by using RotorGene software (Ozyme). Samples were tested in triplicate, and average values were used for quantification. For each sample, the ratio between the relative amounts of IL-1
and
-actin was calculated to compensate for variations in the quantity or quality of starting mRNA as well as for differences in reverse transcriptase efficiency.
MSORT. MSORT was previously described in detail (23, 31). Briefly, human tissue is prepared by careful removal of mucosal and circular muscle to obtain a preparation of Meissner's plexus. The preparation is then pinned in a recording chamber with a 42-mm-diameter glass bottom (130- to 170-µm thickness; Sauer, Reutlingen, Germany), continuously perfused with 37°C Krebs solution gassed with carbogen (5% CO2-95% O2), and equilibrated at pH 7.4. The chamber is mounted onto an epifluorescence Olympus IX 50 microscope equipped with a 150-W xenon arc lamp (Osram, Munich, Germany) connected to a stabilized power supply (OptiQuip, Highland Mills, NY). Controlled illumination of the preparation is achieved by a software-controlled shutter (Uniblitz D122; Vincent Associates). Illumination periods of 1.3 to 4 s were used in most experiments. The preparations were stained with the fluorescent voltage-sensitive dye 1-(3-sulfonatopropyl)-4-{[2-(di-n-octylamino)-6-naphthyl]vinyl}pyridinium betaine (Di-8-ANEPPS). The ganglion identified under the microscope was stained by local pressure application through a microejection pipette loaded with 200 µM Di-8-ANEPPS dissolved in DMSO and pluronic F-127 (4.1 mg/ml) containing Krebs solution (final concentration of DMSO in the pipette: 0.2 M). The glass pipette was manipulated within 100 µm of the ganglia, and constant ejection pressure (2 psi) was used to apply the dye. Dye microejection lasted between 9 and 15 s (ejection volume, 10-100 nl) followed by a 60-s incubation period during which the perfusion system was stopped. The perfusion system was then restarted, and the ganglion was allowed to stain for an additional 5-10 min before experiments were started. During the staining period, it was possible to monitor and assess the progress of the labeling by brief illumination of the ganglion. Di-8-ANEPPS-stained neurons were visualized with an x40 objective (model UAPO/340; numerical aperature = 1.4; Olympus) by using a filter cube consisting of a 545 ± 15-nm excitation interference filter, a 565-nm dichroic mirror, and a 590-nm barrier filter (Olympus). Fluorescent images were acquired and processed by the neuro-charge-coupled device (neuro-CCD) system (RedShirtImaging, Fairfield, CT). The neuro-CCD system consisted of a specialized CCD-camera with 70 x 70 pixel resolution and a Windowsbased computer system to control the experiment and acquire, store, and process data. Frames were acquired at a frequency of 1.6 kHz. To reduce noise and minimize the effects of bleaching, data for individual pixels were filtered with a 3-300 Hz band pass. Spatial resolution of 4.7 µm was achieved by using an x40 objective. The software used made it possible to bin pixels corresponding to the surface of individual neurons, thereby reducing noise even more. This setup measures fluorescence intensity in arbitrary units and calculates the relative change in fluorescence as related linearly to changes in Vm.
At the end of the experiments, the total number of neurons in a given ganglion was counted in the fluorescence image of the ganglion. Neuron viability was tested by random electrical stimulation of the interganglionic fiber tracts that linked the ganglion directly. An average of 55 ± 22% of the neurons within a ganglion responded to electrical stimulation. Because not all interganglionic fiber tracts were stimulated during the experiment, this average represents only the lower limit of the viable neurons of the ganglia. Therefore, the number of neurons in this section is based on the total number of neurons identified with the fluorescence image of the ganglion.
Pharmacological study. IL-1 and the natural IL-1 receptor antagonist (IL-1ra) (both from Peprotech, Tebu, Le Perray en Yvelines, France) were diluted in distilled water. The IL-1
level in tissue supernatant was determined by ELISA (Tebu). Results are expressed in picograms per milliliter per gram of tissue.
Statistics. Data are expressed as the means ± SD when normally distributed or as the median (25th-75th percentiles) when nonnormally distributed. A paired or unpaired Student's t-test, a Mann-Whitney U-test, and one-way ANOVA followed by a Bonferroni t-test or two-way repeatedmeasures ANOVA were performed to compare different populations. Differences were considered as significant for P < 0.05.
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RESULTS |
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Analysis of the neurochemical code of the c-Fos-positive neurons revealed that a significantly larger proportion of neurons expressing c-Fos was immunoreactive for VIP compared with the control (79.8 ± 22.5% of c-Fos-positive neurons vs. 55.0 ± 19.3%; P < 0.02; n = 6) (Fig. 2). The proportion of c-Fos-positive neurons expressing SP was similar in the presence or absence of toxin B (19.4 ± 14.0 vs. 12.3 ± 12.9%; P = 0.62; n = 6).
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Toxin B of C. difficile induced c-Fos expression not only in enteric neurons but also in other cells of the preparation containing Meissner's plexus, i.e., in smooth muscle or blood vessels (Fig. 2).
Toxin B-induced expression of c-Fos is mediated, in part, by IL-1. After 3-h incubation of submucosa/mucosa with toxin B of C. difficile, a sixfold increase was observed in the IL-1
level in supernatant comapared with control conditions without toxin B [16.0 (3-37) vs. 81 pg/ml (37-273); P < 0.0001; n = 10]. Moreover, a rapid, transient increase in IL-1
mRNA occurred 30 min after the addition of toxin B of C. difficile, compared with the control (absence of toxin B) (Fig. 3).
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When the tissue was incubated with IL-1 (10 ng/ml), c-Fos expression was significantly increased in submucosal neurons compared with the control (34.2 ± 10.1 vs. 5.1 ± 1.3% of NSE-positive neurons; P = 0.003; n = 5). In contrast to toxin B of C. difficile, c-Fos induced by IL-1
was localized mainly in enteric neurons. As with toxin B of C. difficile, more c-Fos-positive neurons were VIP-immunoreactive in the presence of IL-1
than in control tissue (80.9 ± 8.8 vs. 42.0 ± 12.4%; P = 0.01; n = 5) (Fig. 4). The proportion of c-Fos-positive neurons expressing SP was similar in the presence or absence of IL-1
(7.1 ± 7.7 vs. 7.5 ± 11.2% respectively; n = 5).
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Finally, c-Fos expression induced by toxin B of C. difficile was reduced dose dependently after tissue preincubation with IL-1ra (Fig. 5).
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Multisite optical recording of the membrane potential response of human submucosal neurons to local application of toxin B of C. difficile. Optical methods were used to determine whether toxin B of C. difficile could modify the membrane potential or synaptic properties of enteric neurons. Microejection of toxin B (100-ms to 20-s pulse duration; 100 ng-500 ng/ml) had no excitatory effects on human submucosal neurons (0/28 neurons in 6 ganglia from 3 preparations) (Fig. 6, A and B). In addition, microejection of IL-1 in most neurons tested (54/59 neurons in 12 ganglia from 5 preparations) had no excitatory effect. However, a slowly developing excitatory effect was observed in 5 of 59 neurons, consisting of an ongoing discharge of action potentials (Fig. 6, C and D). Owing to the unstable nature of this effect, it was not possible to determine whether it was of pre- or postsynaptic origin.
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DISCUSSION |
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Our experiments showed that toxin B of C. difficile induced an inflammatory response in ex vivo human colonic fragments, characterized by an increase in IL-1 mRNA level and IL-1
secretion. Although the cells releasing IL-1
were not identified in the present study, toxin B of C. difficile is known to stimulate IL-1
release in human monocytes and human THP-1 monocytic cells (11, 40). The rapid increase in IL-1
mRNA induced by toxin B can be compared with the rapid (30-min) increase in SP mRNA observed in dorsal root ganglia in response to intraluminal injection of toxin A of C. difficile in the rat (7). The mechanisms involved in the regulation of IL-1
mRNA by toxin B are currently unknown but could involve MAPK pathways, as shown in THP-1 after exposure to toxin A (39). It is also noteworthy that preincubation of thymoma cells with toxin B can inhibit JNK and p38 MAPK activation (9), which could account, in part, for the reduced increase over time of IL-1
mRNA induced by toxin B, compared with the control (Fig. 3).
Toxin B increased c-Fos expression dose dependently in human submucosal neurons. At a concentration of 10 ng/ml, this increase was reduced significantly and dose dependently by IL-1ra and was partly reproduced by IL-1. However, IL-1
(10 ng/ml) did not appear to be as potent as toxin B in inducing c-Fos expression in enteric neurons. This lower potency might also have been due to the presence of other cytokines released after incubation of the tissue with toxin B of C. difficile, which could in turn potentiate the effects of IL-1
. In fact, c-Fos was also expressed in smooth muscle and blood vessels in the presence of toxin B but not IL-1
.
IL-1-mediated induction of c-Fos in enteric neurons could be due, in part, to a direct effect of this cytokine on the Vm of enteric neurons (
8% were depolarized). Indeed, IL-1
depolarized and induced action potential discharge directly in enteric neurons in the guinea pig (18, 43), although in a larger proportion than in our study in the human ENS. MSORT may be unable to detect depolarizations with a low amplitude and slow time course, which could account, in part, for this lower proportion. However, other mechanisms may be responsible for IL-1
-mediated induction of c-Fos expression in human enteric neurons. For example, Sharkey and Kroese (34) have shown that IL-1
-induced c-Fos expression in guinea pig submucosal neurons is blocked by indomethacin, which suggests that prostaglandins are involved. In addition to being associated with increased excitability, enhanced c-Fos expression could also play a role in regulating the synthesis of neurotransmitters such as VIP or acetylcholine (25, 37). In our study in human tissue, toxin B- and IL-1
-induced c-Fos expression occurred preferentially in VIP-positive neurons. In the guinea pig, inflammatory mediators such as IL-1
(4) or prostaglandins (8, 12) also depolarized enteric neurons or induced c-Fos expression preferentially in VIP-positive submucosal neurons. Interestingly, toxin A depolarized a majority of submucosal neurons in the guinea pig. Most of these neurons were probably VIP positive, because a large proportion of submucosal neurons are VIP positive, and only a minority are SP-positive (24, 41).
In MSORT experiments, we were unable to detect any effect of toxin B in human submucosal neurons. This is in contrast to toxin A, which depolarized the vast majority of submucosal neurons directly in the guinea pig and blocked electrically induced slow inhibitory postsynaptic potentials (42). These differences may be related to specific mechanisms of action of toxins A and B, the toxin concentrations tested, and/or interspecies differences. For example, Sörensson et al. (35) showed that luminal injection of a concentration of toxin A (1 µg/ml) induced an intrinsic, neurally mediated inflammatory response, whereas at a higher concentration (15 µg/ml) no neuronal component was detected, except for fluid secretion. Interspecies differences have also been reported, because toxin B induced IL-1 production in human but not rabbit macrophages (30). Although toxin B is usually considered not to harm intestinal mucosa in vivo in animals, it is actually able to damage human colonic epithelium (11) and is even more potent than toxin A (11). Furthermore, the role of toxin B in the pathogenesis of C. difficile-associated diarrhea has been reinforced by the detection of toxin A-negative toxin B-positive strains in patients with C. difficile-associated diarrhea (2).
What functional effects could result from the activation of VIP-positive neuronal pathways in response to toxin B of C. difficile? Whereas toxin A-induced inflammation is mediated by SP released from neurons of extrinsic origin (20), the activation by toxin of C. diffi-cile of VIP neuronal pathways in the ENS could modulate the inflammatory response. Indeed, a recent study (1) showed that VIP has anti-inflammatory effects in experimental models of intestinal inflammation. In addition to the prosecretory effect in the human gut of VIP (13), which is contained in enteric neurones projecting to the mucosa (26), activation of VIP-positive pathways could lead to an increase in intestinal epithelial barrier resistance and therefore be involved in its protection/repair. It has been shown that human submucosal neurons are involved in the maintenance of the intestinal epithelial barrier via VIPergic pathways (38), and preliminary results from our group (22) suggest that VIP decreases intestinal permeability by modulating the expression of tightjunction-associated proteins. Therefore, neuronal activation of VIP-positive pathways in the ENS by pathogens such as toxin B of C. difficile or cholera toxin (14) could represent a protective adaptive response to aggression of the intestinal mucosa by a pathogenic agent.
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DISCLOSURES |
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This work was presented, in part, at the Annual Meeting of the American Gastroenterlolgical Association, San Francisco, CA in May 2002 and published in abstract form (3).
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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REFERENCES |
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