1Departments of Medicine and 2Surgery, Rhode Island Hospital and Brown Medical School, Providence, Rhode Island 02903; 3Department of Medicine, Division of Biomolecular Medicine, Boston University School of Medicine, Boston, Massachusetts 02118; and 4Division of Gastroenterology, Case Western Reserve University School of Medicine, Cleveland, Ohio 44106
Submitted 24 September 2003 ; accepted in final form 10 December 2003
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ABSTRACT |
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neurokinin A; calcium; smooth muscle; human; colon
We (5) have previously shown that neurokinin A (NKA) is an important excitatory neurotransmitter in human sigmoid circular muscle, because contraction induced by electrical field (i.e., neural) stimulation is abolished by NK-2 receptor antagonists and not by NK-1 antagonists or atropine. We therefore used NKA as an agonist to examine contraction of sigmoid circular muscle.
The pathogenesis of UC is not well understood. It is thought that UC may depend on inappropriate and ongoing activation of the mucosal immune system initiated by normal luminal flora or by their products (34). Genetic factors determine differential susceptibility to the development of the disease and proinflammatory cytokines, such as TNF- and IL-1 and -6, and potentiate the inflammatory processes that eventually cause many of the clinical manifestations (33).
Inflammation in UC has been better characterized in the mucosa than in the muscularis propria. It has been reported that the production of proinflammatory cytokines, including IL-1, TNF-
, IL-6, and IL-8, is increased in the colonic mucosa of patients with UC (6, 10, 18, 19, 25, 29, 51). Elevated levels of other inflammatory mediators, including lipid mediators such as platelet activating factor, prostaglandin E2, leukotriene B4, thromboxane B2, and neuropeptides and reactive oxygen species (ROS) such as hydrogen peroxide (H2O2), have also been reported in mucosal tissue samples from patients with UC (37, 38). The diarrhea commonly observed in patients with UC is almost invariably related to the degree of activity of the disease. Motor dysfunction is another frequent abnormality associated with UC, and it has been described in patients as well as animal models of colonic inflammation (7, 22, 27, 45, 46). The effect of inflammatory mediators on human colonic motor function in UC, however, has not been examined.
Increased H2O2 production has been reported in colonic muscularis propria of dextran sodium sulfate-treated rats (11). In addition, H2O2 has been shown to consistently depress the Ca2+-ATPase responsible for uptake of Ca2+ into the endoplasmic reticulum (1214, 26, 35). In pig coronary artery smooth muscle, H2O2 damaged the sarcoplasmic reticulum Ca2+ pump, causing a decrease in the available Ca2+ stores (14). These data are consistent with our own findings in cat lower esophageal sphincter (LES) muscle in which we have shown that H2O2 depletes intracellular Ca2+ stores, reduces LES tone, and plays an important role in motor dysfunction of acute esophagitis (4). We therefore examined whether NKA-induced Ca2+ release from the intracellular Ca2+ stores are affected in UC and whether H2O2 has a role in the observed motor dysfunction and changes in Ca2+ signaling in this condition.
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MATERIALS AND METHODS |
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Preparation of circular muscle strips. After the mucosa was removed by sharp dissection under a microscope, consecutive circular muscle strips (10 mm long, 2 mm wide) of sigmoid colon were cut with razor blades held in a metal block 2 mm apart. The strips were mounted in separate 1-ml muscle chambers as previously described in detail (2). They were initially stretched to 2.5 g of force to bring them near conditions of optimum force development and equilibrated for an additional 30 min after continuous perfusion with oxygenated Krebs solution for 30 min. During the perfusion period, spontaneous phasic contractions developed gradually and stabilized after a 30-min period of equilibration.
Circular muscle strips from normal or UC colons were randomly divided into control and catalase groups, and a cumulative dose-response to NKA was obtained after strips were incubated with vehicles (Krebs solution) or Krebs solution containing catalase (78 U/ml for 50 min). Contractile forces measured in grams above the basal levels in response to NKA were compared between the two groups.
Isolation of smooth muscle cells. Sigmoid circular smooth muscle strips (1 mm wide) were isolated by enzymatic digestion in HEPES-buffered collagenase solution, as described previously (5, 41). Briefly, the collagenase solution (pH 7.2) contained 0.5 mg/ml collagenase Sigma type F, 1 mg/ml papain, 1 mg/ml BSA, and in mM: 1 CaCl2, 0.25 EDTA, 10 glucose, 10 HEPES (sodium salt), 4 KCl, 125 NaCl, 1 MgCl2, and 10 taurine. The tissue was kept in enzyme solution at 4°C for
16 h, warmed up at room temperature for 30 min, and incubated in a water bath at 31°C for
30 min. At the end of the digestion period, the tissue was poured out over a 200-µm Nitex mesh (Tetko, Elmsford, NY), rinsed in collagenase-free HEPES-buffered solution to remove any trace of collagenase, and incubated in this solution at 31°C and gassed with 100% O2. Collagenase-free HEPES-buffered solution (pH 7.4) contained (in mM) 112.5 NaCl, 3.1 KCl, 2.0 KH2PO4, 10.8 glucose, 24.0 HEPES (sodium salt), 1.9 CaCl2, and 0.6 MgCl2, with 0.3 mg/ml basal medium Eagle (BME) amino acid supplement and 0.08 mg/ml soybean trypsin inhibitor. Gentle agitation was used to release single cells.
Agonist-induced contraction of isolated muscle cells. Cell contraction was induced by exposure to NKA (10-13 to 10-9 M) for 30 s. For thapsigargin treatment, cells were exposed to HEPES-buffered solution without (control) or with thapsigargin (3 µM) for 15 s, 30 s, or 1, 5, 10, or 20 min. When the H2O2 scavenger catalase was used, cells from patients with UC were incubated in HEPES-buffered solution without (control) or with catalase 78 U/ml for 50 min before stimulation with NKA or thapsigargin. When the role of H2O2 in the intracellular Ca2+ stores was tested, normal muscle cells were incubated in HEPES-buffered solution without (control) or with H2O2 (70 µM) for 30 min.
After exposure to NKA or thapsigargin, the cells were fixed in acrolein at a 1% final concentration and kept refrigerated. For cell length measurement, a drop of the cell-containing medium was placed on a glass slide and 30 consecutive cells from each slide were observed through a phase-contrast microscope (Carl Zeiss) and a closed-circuit television camera (model WV-CD51; Panasonic, Secaucus, NJ) connected to a Macintosh computer (Apple, Cupertino, CA). An image software program (National Institutes of Health, Bethesda, MD) was used to acquire images and measure cell length. The average length of 30 cells, measured in the absence of agonists, was taken as the control length and compared with length measured after the addition of test agents. Shortening was defined as percent decrease in average length after agonists compared with the control length.
Cytosolic Ca2+ measurements. Freshly isolated cells were loaded with 1.25 µM fura-2 AM for 40 min and placed in a 5-ml chamber mounted on the stage of an inverted microscope (Carl Zeiss). The cells were allowed to settle onto a coverslip at the bottom of the chamber. The bathing solution was collagenase-free HEPES-buffered solution (normal Ca2+ medium) or the one without CaCl2 but with 200 µM BAPTA (Ca2+-free medium). When Ca2+-free medium was used, after settling to the bottom of the chamber, the cells were rinsed twice with Ca2+-free medium before the experiments.
NKA (1 µM), KCl (1 M), or H2O2 (100 mM) was applied directly to the cells by using a pressure ejection micropipette system. Solutions in the pressure ejection micropipettes were identical to the bathing solutions except for the addition of NKA or H2O2. When KCl was used, it was dissolved in distilled water.
Concentration of agents in the micropipette were considerably higher than those used in cell suspensions. The pipette tip was very small, and it was expected that the solution ejected from the tip may be diluted several times by the buffer surrounding the cells. Thus the concentration of the agonists reaching the cells was much lower than that present in the micropipette. For instance, to cause maximal cell shortening 10-9 M NKA was used for cell suspensions and 1 µM NKA was used in a puffing pipette. Thus the concentrations of NKA in these two preparations were 1,000 times different. Similarly, for H2O2 a 100-mM micropipette concentration was needed to elicit a measurable Ca2+ signal, because 5 and 10 mM H2O2 did not cause visible cytosolic Ca2+ changes.
Ca2+ measurements were obtained by using a modified dual excitation wavelength imaging system (IonOptix, Milton, MA). The Ca2+ concentrations were measured from the ratios of fluorescence elicited by 340-nm excitation to 380-nm excitation using standard techniques (15). Ratiometric images were masked in the region outside the borders of the cell, because low photon counts give unreliable ratios near the edges. We developed a method for generating an adaptive mask that follows the borders of the cell as Ca2+ changes and as the cell contracts. A pseudoisobestic image (i.e., an image insensitive to Ca2+ changes) was formed in computer memory from a weighted sum of the images generated by 340- and 380-nm excitation. This image was then thresholded, i.e., values below a selected level were considered to be outside the cell and assigned a value of zero. For each ratiometric image, the outline of the cell was determined, and the generated mask was applied to the ratiometric image. This method allows the simultaneous imaging of the changes in Ca2+ and in cell length. Our algorithm has been incorporated into the IonOptix software. After the experiment, the cell images were copied into a Microsoft Powerpoint file, which was converted into a .jpg file. The cell length was then measured by using NIH Image software.
H2O2 measurement. Sigmoid circular smooth muscle squares (100 mg) were homogenized in PBS. Homogenization consists of a 20-s burst with a Tissue Tearer (Biospec, Racine, WI) followed by 50 strokes with a Dounce tissue grinder (Wheaton, Melville, NJ). An aliquot of homogenate was taken for protein measurement. The homogenate was centrifuged at 15,000 rpm for 15 min at 4°C in a model J221 centrifuge with a fixed-angle model JA-20 rotor (Beckman, Palo Alto, CA), and the supernatant was collected.
H2O2 content was measured by Bioxytech H2O2-560 Quantitative Hydrogen Peroxide Assay Kit (Oxis International, Portland, OR). This assay is based on the oxidation of ferrous ions (Fe2+) to ferric ions (Fe3+) by H2O2 under acidic conditions. The ferric ion binds with the indicator dye xylenol orange 3,3'-bis[N,N-di(carboxymethyl)-aminomethyl]-o-cresoisulfone-phthalein sodium salt to form a stable, colored complex that can be measured at 560 nm.
Fluorescence microscopic measurement of intracellular ROS. Intracellular ROS were measured according to the methods described previously (28, 49). Briefly, freshly isolated muscle cells were incubated in HEPES-buffered solution with or without catalase 78 U/ml for 50 min and then loaded with 2.5 µM 5-(and 6-)chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) for 30 min at room temperature in the dark. During loading, the acetate groups on CM-H2DCFDA are cleaved by intracellular esterase, trapping the probe inside the muscle cells. Several dihydrofluorescein derivatives have been used for measuring intracellular ROS generation (36, 43, 47). CM-H2DCFDA was chosen, because it showed better retention in cells than other derivatives. After being loaded, the cells were fixed in PBS buffer containing 4% paraformaldehyde and kept at 4°C in the dark.
Production of ROS was measured by changes in fluorescence, because subsequent oxidation of CM-H2DCFDA produced a fluorescent product in sigmoid muscle cells. The fluorescence was detected on a fluorescent microscope (Eclipse model E800; Nikon, Mellville, NY) at an excitation wavelength of 488 nm and emission at 520 nm, and the cell images were collected into a Macintosh computer with identical parameters for all samples. The intensity of fluorescence of each cell was measured by using the NIH image software. At least 10 cells from each patient were measured.
Drugs and chemicals. Soybean trypsin inhibitor was from Worthington Biochemicals (Freehold, NJ); fura-2 AM, CM-H2DCFDA, and BAPTA were from Molecular Probes (Eugene, OR). NKA, H2O2, collagenase type F, papain, catalase, BME amino acid supplement, HEPES sodium, paraformaldehyde, and other reagents were purchased from Sigma (St. Louis, MO).
Statistical analysis. Data are expressed as means ± SE. Statistical differences between two groups were determined by Student's t-test. Differences among multiple groups were tested by using ANOVA and checked for significance using Fisher's protected least significant difference test.
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RESULTS |
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After a 10-min incubation of UC cells in Ca2+-free medium, the KCl-induced Ca2+ signal was reduced to 25.4 ± 3.6 nM (n = 3 subjects, 7 cells), significantly lower than in normal Ca2+ medium (P < 0.01, unpaired t-test) (Fig. 2). Because KCl-induced Ca2+ changes are mediated by Ca2+ influx (16, 21, 30, 39), our data establish a Ca2+-free medium incubation protocol to selectively block influx of extracellular Ca2+ without affecting signals mediated by release of Ca2+ from intracellular stores, as shown in Fig. 3A.
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The unstimulated length of normal muscle cells was 90.7 ± 5.6 µm (4 patients, 120 cells). Figure 3 shows that in normal sigmoid circular muscle cells in normal Ca2+ medium, NKA (1 µM) increased cytosolic Ca2+ levels by 336.1 ± 24.6 nM (n = 3 patients, 14 cells) and caused 26.5 ± 2.4% cell shortening. In Ca2+-free medium, NKA (1 µM) caused a 326 ± 23.9 nM Ca2+ increase (n = 4 patients, 17 cells) and caused 26.1 ± 2.4% cell shortening.
Therefore, in normal sigmoid circular muscle cells, shortening and Ca2+ signals were not different in normal Ca2+ and in Ca2+-free medium, confirming previously reported findings (5, 17) that an NKA-induced calcium signal is mainly due to calcium release from intracellular calcium stores.
In UC muscle cells, the resting cell length was 98.7 ± 7.6 µm (3 patients, 90 cells). On average, the UC muscle cells were slightly longer than normal, but the difference was not statistically significant. NKA (1 µM) caused a 372.2 ± 42.7 nM Ca2+ increase (n = 5 patients, 30 cells) and 18.1 ± 1.3% cell shortening in normal Ca2+ medium. Thus in UC cells, NKA-induced shortening was significantly lower than in normal cells, but the Ca2+ increase was not different from normal cells.
In UC cells in Ca2+-free medium, however, NKA (1 µM) caused a 187.4 ± 30 nM Ca2+ increase (n = 5 patients, 35 cells) and a 7.6 ± 1.7% cell shortening. Thus in UC cells in Ca2+-free medium, the Ca2+ signal was significantly lower than in normal Ca2+ medium and in normal cells (Fig. 3). Similarly, in UC cells in Ca2+-free medium shortening was significantly lower than in normal Ca2+ medium and in normal cells in Ca2+-free medium (Fig. 3).
The data suggest that in normal cells, the NKA-induced Ca2+ signal depends only on release of Ca2+ from intracellular stores, because it is not affected when influx is abolished by incubation in Ca2+-free medium.
In contrast, in UC cells the NKA-induced Ca2+ signal depends in part on influx of extracellular Ca2+ and in part on release of Ca2+ from intracellular stores, because the Ca2+ signal is diminished when influx is abolished by incubation in Ca2+-free medium. In normal Ca2+, however, the Ca2+ signal is the same in UC as in normal cells.
In UC cells, influx of extracellular Ca2+ must compensate for the reduction in release of Ca2+ from intracellular stores. The finding that in normal Ca2+ medium, contraction of UC cells is less than in normal cells despite equal amplitude of Ca2+ signal suggests that in addition to alteration of Ca2+ release, other mechanisms of contractile signal transduction might also be affected in UC.
H2O2 and UC motor dysfunction. Elevated levels of H2O2 have been demonstrated in colonic mucosa of patients with UC (9, 40). To test whether H2O2 may also be present in the circular smooth muscle and contribute to motor dysfunction, UC muscle strips and cells were exposed to the H2O2 scavenger catalase.
Figure 4A shows that NKA caused concentration-dependent contractions of normal and UC circular sigmoid muscle strips. Consistent with the cell data shown in Fig. 3, UC strips contracted less than normal strips at all NKA concentrations tested (P < 0.0001) (Fig. 4A), and incubation in catalase (78 U/ml for 50 min) significantly increased contraction of UC strips (P < 0.05) but did not increase contraction of normal strips.
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Similarly, in enzymatically isolated circular muscle cells, NKA induced a concentration-dependent contraction (Fig. 4B) that was significantly lower in UC than in normal cells. Incubation in catalase (78 U/ml for 50 min) significantly increased contraction of UC cells (P < 0.0001, ANOVA). The resting cell length after catalase treatment was 104.9 ± 8.1 µm (3 patients, 90 cells), and this length was not significantly different from untreated UC cells. In normal cells, catalase had no effect on cell shortening. These data strongly suggest that H2O2 is present in the circular smooth muscle, because preparations of both strips and isolated cells are free of mucosa.
Because NKA-induced release of Ca2+ from intracellular stores may be impaired in UC (Fig. 3), we then examined thapsigargin-induced contraction of circular muscle cells (Fig. 5). Thapsigargin inhibits uptake of Ca2+ into stores, causing a net release of Ca2+ and contraction. Continuous release of Ca2+ in the absence of Ca2+ uptake results eventually in depletion of stores.
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Similarly to NKA-induced contraction, thapsigargin induced a time-dependent contraction (Fig. 5) that was significantly lower in UC than in normal cells, supporting the proposition that UC reduces releasable Ca2+ stores. Incubation in catalase (78 U/ml for 50 min) did not affect contraction of normal cells but significantly increased contraction of UC cells (P < 0.001, ANOVA), confirming presence of H2O2 in circular smooth muscle cells from UC patients.
To directly examine Ca2+ release from intracellular stores, enzymatically isolated circular muscle cells were loaded with the calcium indicator fura-2 AM, and NKA-induced calcium signal and shortening were examined in Ca2+-free medium (i.e., in the absence of Ca2+ influx) in the same cells. Ca2+ signal and shortening of the sigmoid circular muscle cell in Ca2+-free medium are shown in Fig. 6. The figure shows that in Ca2+-free medium, a normal cell exhibits a strong Ca2+ signal and shortens 34.6%. In contrast, a UC cell in Ca2+-free medium exhibits little or no Ca2+ signal and shortening, but both Ca2+ signal and shortening are augmented after the cell is incubated in catalase. Although in UC cells incubated in catalase the Ca2+ signal is greater than in normal cells, the cell shortening is slightly lower than in normal cells. Average values for shortening and amplitude of Ca2+ signal are shown in Fig. 7. However, catalase treatment had no effect on NKA-induced Ca2+ signal and cell contraction in normal cells.
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The finding that the H2O2 scavenger catalase increases amplitude of NKA-induced Ca2+ signal and shortening in UC circular muscle suggests that H2O2 is produced by and present in circular muscle and that it contributes to motor dysfunction in UC by reducing NKA-induced Ca2+ release from intracellular stores.
H2O2 and intracellular calcium stores in normal sigmoid circular smooth muscle cells. To test the hypothesis that H2O2 may be present in the circular muscle layer and responsible for impaired release of Ca2+ from intracellular stores, we measured H2O2 content in normal and UC sigmoid circular muscle.
H2O2 levels were significantly elevated in UC, compared with normal muscle (Fig. 8), supporting the possibility that H2O2 may be involved in the observed changes in muscle contraction. To test whether isolated sigmoid muscle cells contain H2O2, we measured intracellular H2O2 by using fluorescence of CM-H2DCFDA (Fig. 9). The cells were loaded with 2.5 µM CM-H2DCFDA for 30 min at room temperature in the dark. During loading, the acetate groups on CM-H2DCFDA are removed by intracellular esterase, trapping the probe inside the muscle cells. When oxidized in situ by ROS, DCF generates a signal that can be visualized by using a fluorescent microscope (1). A stronger oxidant signal was detected by DCF fluorescence in UC muscle cells than in normal cells. This oxidant signal was significantly reduced by preincubation of the muscle cells with catalase (78 U/ml, 50 min), suggesting that H2O2 is the main oxidant generated (Fig. 9). The data suggest that isolated UC muscle cells contain excess H2O2, which may affect cell contraction. Catalase did not completely remove the oxidant signal, suggesting that other ROS may be present in UC muscle cells.
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To test whether H2O2 may induce release of Ca2+ from intracellular stores, a high concentration of H2O2 was directly applied to fura-2 AM-loaded cells (Fig. 10) through a pressure ejection micropipette to permit visualization of Ca2+ release.
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Application of a high-concentration H2O2 to enzymatically isolated smooth muscle cells caused a gradual Ca2+ increase, which reached a stable level in 1 min. The rate of Ca2+ increase was much lower than that of NKA-induced Ca2+ increase where peak Ca2+ concentration was reached in a few seconds, as shown in Fig. 6B. Figure 10 shows that in normal calcium, H2O2 caused a 194 ± 11.6 nM Ca2+ increase (from 96.6 ± 4.1 to 290.5 ± 9.5 nM, n = 10). In Ca2+-free medium, H2O2 caused a 190.2 ± 23.7 nM Ca2+ increase (from 72.3 ± 6.2 to 262.6 ± 25.1 nM, n = 6) (Fig. 10). There was no difference in H2O2-induced Ca2+ increase in normal Ca2+ medium and in Ca2+-free medium, indicating that H2O2 causes direct release of Ca2+ from intracellular stores. The recorded Ca2+ changes were not associated with any change in cell length within 3 min and were consistent with the finding that a similar Ca2+ increase was insufficient to cause contraction (Fig. 6).
To test whether prolonged exposure of normal cells to a relatively low concentration of H2O2 may result in depletion of intracellular Ca2+ stores, we examined NKA-induced Ca2+ signal in fura-2 AM-loaded normal cells incubated with 70 µM H2O2 for 30 min. The cells were placed in Ca2+-free medium with 200 µM BAPTA to ensure that the Ca2+ signal was entirely produced by release of Ca2+ from intracellular stores.
H2O2 treatment did not change resting calcium levels in unstimulated cells but significantly decreased the NKA-induced Ca2+ signal (78.6 ± 73.3 nM) when compared with untreated cells (326 ± 24 nM) (Fig. 11), demonstrating that H2O2 may be directly responsible for decreasing releasable Ca2+ from intracellular stores and for the reduced Ca2+ signal in response to NKA.
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DISCUSSION |
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To distinguish the contribution of Ca2+ release in the overall Ca2+ signal, we developed a protocol to ensure abolition of Ca2+ influx without affecting Ca2+ release. Isolated smooth muscle cells were incubated in Ca2+-free medium containing 200 µM BAPTA for no longer than 10 min before starting the experiment. This protocol abolished KCl-induced contraction, which is mediated by influx of extracellular Ca2+ without affecting NKA-induced contraction that is mediated by release of Ca2+ from intracellular stores. Establishing an appropriate protocol is important, because longer incubations or higher BAPTA concentrations may cause depletion of intracellular stores.
Using this protocol, we confirmed that, although in normal cells the NKA-induced Ca2+ signal is mediated entirely by the release of Ca2+ from intracellular stores (Fig. 3), in UC cells the Ca2+ signal is only in part mediated by Ca2+ release and releasable Ca2+ stores are reduced. In UC cells, the Ca2+ signal was reduced when the cells were incubated in Ca2+-free medium, supporting damage to Ca2+-release mechanisms. In normal Ca2+, however, the Ca2+ signal was normal, indicating that Ca2+ influx compensated for the reduced Ca2+ release. Despite the normal Ca2+ signal in normal Ca2+ medium, however, contraction in response to NKA was reduced. Similarly, KCl-induced Ca2+ signal in UC cells was not significantly different from that in normal cells, whereas KCl-induced contraction was reduced in UC cells, which is consistent with the literature (42). The data suggest that mechanisms of Ca2+ release and other contractile signal transduction pathways might be impaired in UC. KCl-induced cell shortening in normal cells is lower than NKA despite a similar Ca2+ signal. This is probably due to the fact that KCl and NKA activate different contractile signal transduction pathways. For instance, NKA activates G protein-coupled receptors that may magnify the signal but KCl does not.
Because reduction of releasable Ca2+ and cell shortening is a feature of UC, we tested H2O2 as a possible factor contributing to depletion of releasable Ca2+ stores and sigmoid motor dysfunction. ROS have been shown to consistently depress the Ca2+-ATPase responsible for uptake of Ca2+ into the endoplasmic reticulum (1214, 26, 35). In addition to inhibiting Ca2+ uptake into the endoplasmic reticulum, ROS cause release of Ca2+ stores through both ryanodine- and inositol 1,4,5-trisphosphate-sensitive Ca2+ channels (24). It was recently reported (32) that H2O2 may inhibit actomyosin ATPase, and thus H2O2 might inhibit NKA-induced contraction despite a normal Ca2+ signal in normal Ca2+ medium in UC.
We found that UC sigmoid circular muscle contains three times as much H2O2 as normal muscle, which is consistent with findings in dextran sodium sulfate-treated rats that H2O2 production increases in the muscularis of the inflamed colon (11). We also found that UC sigmoid muscle cells had much higher levels of intracellular H2O2 than normal cells and that this intracellular H2O2 can be removed by extracellular catalase (Fig. 9). This observation is consistent with the literature showing that extracellular catalase can neutralize intracellular H2O2 in adipocytes and HepG2 cells (28). The mechanism of removal of intracellular H2O2 by catalase is not clear. Because H2O2 can diffuse across biological membranes (44), it is possible that removal of extracellular H2O2 by catalase may facilitate diffusion of intracellular H2O2 into the extracellular medium, resulting, eventually, in removal of both intracellular and extracellular H2O2.
Catalase treatment significantly increased NKA-induced contraction in UC cells and muscle strips (Fig. 4) and restored NKA-induced Ca2+ signal in Ca2+-free medium, indicating that H2O2 is present both in circular muscle strips and in enzymatically isolated smooth muscle cells and contributing to UC-associated motor dysfunction.
H2O2-associated depletion of releasable Ca2+ was confirmed by testing thapsigargin-induced contraction of isolated smooth muscle cells. Thapsigargin inhibits uptake of Ca2+ into stores (8), shifting the uptake-release balance toward a net release of Ca2+ and contraction. The finding that thapsigargin-induced contraction is reduced in UC confirms again an UC-associated depletion of releasable Ca2+ stores. Neutralization of H2O2 by catalase restored thapsigargin-induced contraction, indicating that the presence of H2O2 in enzymatically isolated smooth muscle cells may be directly responsible for inhibiting the refilling of Ca2+ stores and that Ca2+ stores quickly return to normal when H2O2 is neutralized.
Catalase-induced restoration of the Ca2+ stores is directly demonstrated in Fig. 6 in which cytosolic Ca2+ can be visualized before and after application of catalase to an enzymatically isolated circular muscle cell.
These data suggest that H2O2 contributes to motor dysfunction and to reduced intracellular Ca2+ signal in UC. Catalase or related compounds may be useful tools in the treatment of UC, as reported by others (3, 50) who have shown that pretreatment with catalase decreased the extent of colonic inflammation in a rat model. It is likely that removal of intracellular H2O2 may allow replenishment of intracellular Ca2+ stores. In fact, after muscle cells from UC patients were exposed to catalase, NKA-induced Ca2+ changes were higher than in normal cells, even in Ca2+-free medium (Fig. 7). The mechanism responsible for this Ca2+ rebound remains to be explored.
Much of the H2O2 produced by eukaryotic cells is derived from reduction of the superoxide anion , normally produced in the respiratory process (44).
in aqueous solution is short-lived and rapidly reduced to the much more stable molecule H2O2. Thus in most biological systems, generation of
usually results in the formation of H2O2. However, the source of excess H2O2 in the circular muscle layer in UC is not clear.
The findings that intracellular H2O2 is present at much higher levels in UC smooth muscle cells than in normal cells and that catalase restores NKA-induced calcium signal and cell shortening in UC cells suggest that UC muscle cells may also be a source of H2O2. H2O2 production has been previously reported in rat aortic smooth muscle cells (23).
To demonstrate that H2O2 may directly cause release of Ca2+ from intracellular stores, H2O2 was directly applied by a pressure ejection micropipette onto fura-2 AM-loaded cells. Applying H2O2 directly onto the cells caused a gradual increase of cytosolic Ca2+ even when cells were maintained in Ca2+-free medium, demonstrating direct H2O2-induced release of Ca2+ from intracellular Ca2+ stores, consistent with previous data (4) in acute esophagitis. Although H2O2 caused Ca2+ increase, it did not cause cell contraction. This may be due to a slower rate of Ca2+ increase and to possible inhibition of actomyosin ATPase (32) induced by H2O2.
To produce a measurable Ca2+ release in a short time, we used a relatively high concentration of H2O2. Over a prolonged period, however, lower H2O2 concentrations are sufficient to cause a reduction in releasable Ca2+ in normal muscle cells, as shown in Fig. 11, where 30-min incubation with 70 µM H2O2 almost abolished NKA-induced Ca2+ release in Ca2+-free medium, suggesting that over a long period, exposure to H2O2 may cause a reduction in releasable Ca2+ stores as previously demonstrated (1214, 26, 35). Thus the presence of increased levels of H2O2 may explain the reduced intracellular calcium release observed in UC.
The finding that catalase only in part restored muscle contraction may perhaps be explained by the existence of other inflammatory mediators such as nitric oxide and arachidonic acid metabolites (prostaglandins, leukotrienes, isoprostanes, etc.) that are not neutralized by catalase.
In conclusion, our data clearly demonstrate that H2O2 is produced not only in the mucosa as generally thought, but also in the muscle layer of UC colon. H2O2 produced in the muscle layer may account for at least some of the motor disturbances observed in UC, and neutralization of H2O2 may result in improvement of UC-associated dysmotility.
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DISCLOSURE |
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ACKNOWLEDGMENTS |
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This work was supported by Lifespan Research Funds (to W. Cao) and National Institute of Diabetes and Digestive and Kidney Diseases Grant R21-DK-62775-01 (to W. Cao).
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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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