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Am J Physiol Gastrointest Liver Physiol 293: G438-G445, 2007. First published May 17, 2007; doi:10.1152/ajpgi.00037.2007
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NEUROREGULATION AND MOTILITY

Membrane potential gradient is carbon monoxide-dependent in mouse and human small intestine

Lei Sha,1 Gianrico Farrugia,1 W. Scott Harmsen,2 and Joseph H. Szurszewski1

1Enteric NeuroScience Program, Mayo Clinic College of Medicine; and 2Division of Biostatistics, Mayo Clinic, Rochester, Minnesota

Submitted 19 January 2007 ; accepted in final form 15 May 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The aims of this study were to quantify the change in resting membrane potential (RMP) across the thickness of the circular muscle layer in the mouse and human small intestine and to determine whether the gradient in RMP is dependent on the endogenous production of carbon monoxide (CO). Conventional sharp glass microelectrodes were used to record the RMPs of circular smooth muscle cells at different depths in the human small intestine and in wild-type, HO2-KO, and W/WV mutant mouse small intestine. In the wild-type mouse and human intestine, the RMP of circular smooth muscle cells near the myenteric plexus was –65.3 ± 2 mV and –58.4 ± 2 mV, respectively, and –60.1 ± 2 mV and –49.1 ± 1 mV, respectively, in circular smooth muscle cells at the submucosal border. Oxyhemoglobin (20 µM), a trapping agent for CO, and chromium mesoporphyrin IX, an inhibitor of heme oxygenase, abolished the transwall gradient. The RMP gradients in mouse and human small intestine were not altered by NG-nitro-L-arginine (200 µM). No transwall RMP gradient was found in HO2-KO mice and W/WV mutant mice. TTX (1 µM) and 1H-[1,2,4-]oxadiazolo[4,3-a]quinoxalin-1-one (10 µM) had no effect on the RMP gradient. These data suggest that the gradient in RMP across the thickness of the circular muscle layer of mouse and human small intestine is CO dependent.

resting membrane potential; smooth muscle; heme oxygenase 2

IN ALL REGIONS OF THE CANINE gastrointestinal tract, with the exception of the gastric fundus, there is a 5- to 10-mV difference in the resting membrane potential (RMP) between circular smooth muscle cells located near the myenteric plexus and circular smooth muscle cells located at the submucosal border (2–4, 11, 12, 20–23, 26–29, 31). In the canine gastric antrum and small intestine, the RMP of circular smooth muscle cells near the myenteric plexus is more hyperpolarized by 5–10 mV compared with the RMP of muscle cells near the submucosal border (2–4, 11, 12, 31). Although a similar difference in RMP exists between these two regions in the canine colon, it is the muscle cells near the submucosal border that are more hyperpolarized compared with muscle cells near the myenteric plexus (20–23, 26–29). Recently, we developed a method for recording intracellularly from smooth muscle cells located at different depths of the circular muscle layer of the mouse gastric antrum (9), a decidedly thinner muscle layer compared with the canine antrum. We found that the RMP progressively declined (i.e., became more depolarized) the further away smooth muscle cells were from the myenteric plexus. There is no information available on whether a similar gradient exists in the mouse and human small intestine. Defining the gradient in these two species was one of the purposes of this study.

A number of observations support the hypothesis that the basis for the greater hyperpolarization of the RMP of circular muscle cells located near the myenteric plexus is due to the activity of heme oxygenase. In the dog gastric antrum, heme oxygenase activity and carbon monoxide (CO) production are greater in regions where the RMP is more hyperpolarized than in regions where the RMP is more depolarized. The difference in RMP and the RMP gradient are abolished in heme oxygenase 2 (HO2) knockout (HO2-KO) mice (9). The source of CO appears to be the network of interstitial cells of Cajal (ICC) located in the myenteric region (9, 17). ICC are immunopositive for HO2, and the RMP gradient is abolished in W/WV mutant mice that lack ICC in the myenteric region of the gastric antrum (9). There is no information available on whether RMPs in the circular smooth muscle layer of the mouse and human small intestine are modulated by endogenously produced CO. Thus, the second objective of this study was to determine whether CO plays a role in setting the RMP of muscle cells located at the different depths of the circular muscle layer. Parts of this study have been communicated in abstract form (24).


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Adult (6 wk or older) wild-type mice (WT, SJL/J), HO2-KO mice, and W/WV mutant mice were used. WT mice, HO2-KO mice, and W/WV mice were obtained from The Jackson Laboratory. Animals were killed with CO2 gas with the prior approval of the Mayo Institutional Animal Care and Use Committee. After the abdomen was opened, a segment of small intestine wall, ~6 cm from the pylorus, was removed and placed in preoxygenated normal Krebs solution (NKS) at room temperature. The segments were opened along the antimesenteric border and transferred to a petri dish filled with fresh oxygenated NKS. The mucosa was removed under direct vision by using a binocular microscope, and muscle strips (4 x 8 mm) were cut with the long axis parallel to the longitudinal muscle layer. Muscle strips were placed in a recording chamber and pinned with the serosal side down to a Sylgard-coated floor so that the innermost layer of the circular muscle faced upward. The recording chamber had a volume of 1 ml and was perfused continuously with oxygenized NKS at 37°C at a rate of 2 ml/min. The composition of the solution was (in mM) 137.4 Na+, 5.9 K+, 2.5 Ca2+, 1.2 Mg2+, 124 Cl–, 15.5 HCO3–, 1.2 H2PO4–, and 11.5 glucose. It was continuously bubbled with 97% O2-3% CO2 and maintained at pH 7.4.

Whole-thickness biopsies of human small intestine were obtained from adult patients undergoing gastric bypass surgery. The use of human biopsy specimens was approved by Institutional Review Board. After the mucosa was removed, strips of muscle (1 x 8 mm) were cut with the long axis of the muscle strip parallel to the circular muscle layer. Muscle strips were pinned on their sides to the floor of a recording chamber so that the full thickness of the circular and longitudinal muscle layers faced upward. Thus, when viewed through a microscope, the long axis of the circular muscle layer was seen whereas the longitudinal muscle layer was seen in cross section. The recording chamber was continuously perfused with oxygenated NKS at 37°C and at a rate of 2 ml/min.

Sharp glass microelectrodes filled with 3 M KCl (with input resistances ranging from 40 to 70 M{Omega}) were used to record intracellularly the membrane potential of smooth muscle cells. Approximately 30 min before recording, 1 µM nifedipine was added to reduce contractile activity of the muscle strip, thereby facilitating long-term recordings from single cells. Recorded signals were amplified through an amplifier (Intra 767, WPI), digitized (Digidata 1322A, Axon Instruments), analyzed, and stored in a computer. When spontaneous electrical slow waves occurred, the membrane potential recorded between slow waves was considered the RMP. For recording from preparations of the mouse intestine, a microelectrode manipulator (LSS-8000, EXFO Burleigh) was used to advance the microelectrode into various depths of the thickness of the circular muscle layer. The meter of the Burleigh manipulator was set at 0 when the tip of the microelectrode touched the surface of the innermost layer of the circular muscle. The maximum depth at which a recording could be made without entering the longitudinal muscle layer at the bottom of the strip was considered the thickness of the circular muscle layer. The location of a recorded cell relative to the mucosal surface of the innermost layer of the circular muscle was obtained from the readout on the meter of the manipulator. In some experiments, Lucifer yellow-containing microelectrodes were used to mark cells from which the recordings were made. Preparations with Lucifer yellow-filled cells were later viewed using confocal microscopy (Olympus FluoView FV300). The distance of the marked cell to the mucosal surface was measured and compared with the readout of the meter of the manipulator. There was excellent correlation between these two measurements. Because of the greater thickness of the circular muscle layer in human preparations compared with the mouse intestine and because preparations of human intestine exhibited spontaneous contractions in spite of the presence of nifedipine, stretch was applied along the longitudinal and circular muscle axes when pinning the strips to the floor of the recording chamber so as to obtain stable and long-term intracellular recording (>20 min). As a result, the apparent thickness of the circular muscle layer was greater than the in vivo thickness. In eight human preparations, the distance from myenteric border to mucosal border before and after stretch was 0.67 ± 0.02 mm (mean ± SE) and 0.93 ± 0.05 mm (mean ± SE), respectively. Thus, unlike the experiments using mouse tissue, we are unable to indicate in basic units of linear measurement where a recording was made in the human circular muscle layer. Therefore, in reporting results from human tissue, we treated the stretched whole-thickness circular muscle layer as 100% (0 was at mucosal border and 100% was at myenteric border) and report the location of recorded cells as percent distance from the mucosal border. When recording from the human small intestine, the microelectrode was moved across the circular muscle layer from the mucosal border to the myenteric plexus region.

Values for RMP at different depths of the circular muscle layer obtained from both preparations are reported in RESULTS in two ways. In the first analysis, values for RMP were placed in three groups based on the cell's location in the circular muscle layer: an outer group, a middle group, and an inner group. Each group represented one-third of the measured thickness of the circular muscle layer. In each experiment, multiple impalements were made within a group and the RMP values were averaged for that group. The mean values (± SE) within groups across all preparations used are reported in tabular form. In the second analysis, the RMPs in the mouse intestine were plotted against the depth (µm) from which the recordings were made. For the human intestine, the RMPs were plotted against the percent thickness of the circular muscle layer. This difference in the x-axis reflects the difference between the way mouse and human preparations were pinned out for recording. A random-effect model was used to model RMPs of recorded cells vs. the location of the cell in the circular muscle layer (7). This method of analysis accounts for the correlation of RMPs and their related locations in mouse and human preparations within control (NKS) and drug-treatment groups. RMP gradients and gradient changes due to drug treatment were compared by using this analysis. Slopes of regression are given as mean ± SE. Using this second analysis method, we plotted the regression slopes (RMPs across the circular muscle layer) without individual data points. A P value ≤0.05 was considered a significant difference.

Slow waves, recorded in mouse and human preparations, are reported as mean slow-wave frequency ± SE. The Student's t-test was used to compare mean slow-wave frequency in control conditions and during drug treatment. A P value ≤0.05 was considered a significant difference.

Chromium mesoporphyrin IX, (CrMP), was from Frontier Scientific. All other chemicals used in this study were from Sigma.


   RESULTS
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 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
RMPs in WT Mouse Small Intestine and Human Small Intestine

Intracellular recordings were made in 91 cells from seven WT mice. A gradient in RMP was found in each preparation studied. An example from one preparation is shown in Fig. 1A. In this example, the RMP of circular smooth muscle cells was approximately –60 mV at the submucosal border increasing to approximately –75 mV at the myenteric border. The mean (± SE, n = 7 preparations) values for RMP in circular smooth muscle cells in outer, middle, and inner groups are given in Table 1. The RMP gradient for each of the seven preparations is shown as dashed lines in Fig. 2A. The mean slope ± SE for all seven preparations was –0.9 ± 0.2 mV/10 µm (Fig. 2A, solid line). The mean slope was significantly different (P < 0.01) from zero slope, indicating the presence of a RMP gradient. Slow waves were recorded from all cells impaled. The mean slow-wave frequency was 0.77 ± 0.02 cycles/s. In each preparation studied, the slow-wave frequency was the same at different depths of the circular muscle layer.


Figure 1
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  		Fig. 1. Resting membrane potential (RMP) gradient across the thickness of the circular muscle layer in wild-type (WT) mice (A) and human small intestine (B). Individual data points indicate the RMP recorded at the indicated site in the circular muscle layer. The slope of the regression line for the mouse is –1.8 mV/10 µm and –1.5 mV per 10% increase in the thickness of the muscle layer for the human small intestine.

 

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  		Table 1. Summary of RMPs of smooth muscle cells in mouse and human small intestine preparations

 

Figure 2
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  		Fig. 2. RMP gradient across the thickness of the circular muscle layer of WT mice (A) and human small intestine (B). Dashed lines represent regression analysis of data obtained from individual experiments; solid lines represent mean regression for all cells in all experiments. The slope of the regression line for all experiments (n = 7) in WT mouse was –0.9 ± 0.2 mV/10 µm and is significantly different from zero slope (P < 0.01). The slope for the regression line for all experiments in human preparations (n = 10) was –1.5 ± 0.3 mV per 10% increase in the thickness of the circular muscle layer and is significantly different from zero slope (P < 0.01). The mucosal border is at 0 µm in A and at 0% in B. The myenteric border of the circular muscle is at 120 µm in A and at 100% in B.

 
Intracellular recordings were made from 86 cells in 10 preparations of the human small intestine. A gradient in RMP was present in each of the 10 preparations. An example from one preparation is shown in Fig. 1B. The RMP was approximately –50 mV in cells near the submucosal border increasing to approximately –65 mV at the myenteric border. The mean (± SE, n = 10 preparations) values for RMP in circular smooth muscle cells in outer, middle, and inner groups are given in Table 1. The RMP gradient for each of the 10 preparations is shown as dashed lines in Fig. 2B. The mean (± SE) slope for all 10 preparations was –1.5 ± 0.3 mV for each 10% increase in the thickness of the circular muscle layer. The mean slope was significantly different (P < 0.01) from zero slope, indicating the presence of a RMP gradient. Slow waves were observed in 63 of 86 cells. The mean frequency was 7.87 ± 0.19 cycles/min.

Effect of Oxyhemoglobin

Mouse and human intestine. Oxyhemoglobin, a molecular trapping agent for CO (19), was used to determine whether the RMP gradient was CO dependent. Intracellular recordings were made from 67 cells from four WT mice before and 30 min after treatment with oxyhemoglobin (20 µM). The mean (± SE) values for RMP for cells in the outer, middle, and inner groups of the circular muscle layer before and after treatment with oxyhemoglobin are given in Table 1. Note that treatment with oxyhemoglobin had a greater effect on the most hyperpolarized regions (outer and middle groups) of the muscle layer (Table 1). The mean slope of the RMP gradient in the four preparations used is shown in Fig. 3A. The mean (± SE) slope in the presence of oxyhemoglobin was 0.1 ± 0.2 mV/10 µm (Fig. 3A, solid lines). The P value, which was 0.65 when the slope was compared with zero slope, suggests that there was no RMP gradient across the thickness of the muscle layer when the preparations were pretreated with oxyhemoglobin. The mean slope of the RMP gradient obtained in NKS containing oxyhemoglobin was significantly different (P < 0.01) compared with the mean slope obtained in NKS. Oxyhemoglobin also significantly (P < 0.05) decreased slow-wave frequency from 0.78 ± 0.02 to 0.69 ± 0.06 cycles/s.


Figure 3
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  		Fig. 3. Effect of oxyhemoglobin on the RMP gradient in WT mice (A) and human small intestine (B). In both the WT mouse and human small intestine, there was a significant difference (P < 0.01) between the slope of the regression line for RMP under control conditions (normal Krebs solution) and in preparations pretreated with oxyhemoglobin. For both the WT mouse and human small intestine, the slope of the regression line (0.1 ± 0.2 mV/10 µm from 67 cells in 4 mouse preparations; 0.01 ± 0.2 mV per 10% thickness from 33 cells in 4 human preparations) when oxyhemoglobin was present in the bathing solution was not significantly different from zero slope (for WT mouse P = 0.65; for human P = 0.96). In this figure, Fig. 4, and Fig. 5, the mean regression lines for WT mouse and human small intestine under control conditions are the same as the mean regression lines for all experiments for WT mouse and human small intestine shown in Fig. 2.

 
Intracellular recordings were made from 33 cells from four preparations of the human small intestine in NKS and 30 min after treatment with oxyhemoglobin (20 µM). Results similar to those observed for the mouse intestine were obtained (Table 1 and Fig. 3B).

Effect of L-NNA

Nitric oxide (NO), an inhibitory neurotransmitter in the gastrointestinal tract (25), hyperpolarizes the RMP of smooth muscle cells (16, 30). Oxyhemoglobin traps not only CO but also NO. To determine whether the abolishment of the RMP gradient by oxyhemoglobin was the result of trapping CO and not the result of also trapping NO, we tested the effect of NG-nitro-L-arginine (L-NNA), a NO synthase inhibitor, on both WT mouse and human muscle strips.

Mouse and human intestine. Intracellular recordings were made from 61 cells in four WT mouse small intestine preparations pretreated for 30 min with 200 µM L-NNA. The mean (± SE) values for RMP for cells in the outer, middle, and inner groups are given in Table 1. The mean (± SE) slope of RMP gradient across the thickness of the circular muscle layer was –1.1 ± 0.4 mV/10 µm. The P value was less than 0.05 when the slope was compared with zero slope, which suggested that L-NNA did not abolish the RMP gradient. The mean slope was not different from the slope obtained from cells not exposed to L-NNA (P = 0.66), suggesting that NO does not contribute to the RMP gradient (Fig. 4A). L-NNA had no effect on slow-wave frequency (0.77 ± 0.02 cycles/s in NKS; 0.76 ± 0.02 cycles/s in L-NNA, P > 0.05).


Figure 4
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  		Fig. 4. Lack of effect of NG-nitro-L-arginine (L-NNA) on the RMP gradient across the thickness of the circular muscle in WT mice (A) and human small intestine (B). For both the WT mouse and human small intestine, the slope of the regression line (–1.1 ± 0.4 mV/10 µm from 61 cells in 4 WT mice and –1.1 ± 0.3 mV per 10% thickness from 33 cells in 4 human preparations) when L-NNA was present in the bathing solution was significantly different from zero slope (P < 0.05). The regression lines for the RMP gradient in L-NNA were not significantly different from the regression lines under control conditions (for WT mouse P = 0.66; for human small intestine P = 0.20).

 
Thirty-two cells were recorded from four human small intestine preparations pretreated for 30 min with L-NNA (200 µM). Results similar to those observed for the WT mouse intestine were obtained (Table 1, Fig. 4B).

Effect of CrMP

To further examine the role of CO in generating the RMP gradient, we tested the effect of CrMP, a selective inhibitor of heme oxygenase activity (1). Intracellular recordings were made from 105 circular smooth muscle cells in 8 WT mouse preparations superfused with 5 µM CrMP. The mean (± SE) values for RMP for cells in the outer, middle, and inner groups of the circular muscle layer in the presence of CrMP are given in Table 1. CrMP, like oxyhemoglobin, abolished the RMP gradient and had a greater effect on the most hyperpolarized regions (outer and middle groups) of the muscle layer (Table 1). The mean (± SE) slope of the regression line with CrMP in the eight preparations was –0.04 ± 0.3 mV/10 µm. The P value is less than 0.05 compared with the slope observed in NKS (Fig. 5). When comparing the slope of the RMP gradient in CrMP-containing Krebs solution to zero slope, the P value was 0.9. These results show that the RMP gradient across the thickness of the muscle layer was abolished by CrMP. CrMP had no significant effect (P > 0.05) on the slow-wave frequency.


Figure 5
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  		Fig. 5. Effect of chromium mesoporphyrin IX (CrMP) on the RMP gradient in WT mice. The slope (–0.9 ± 0.2 mV/10 µm) of the regression line for the RMP under control conditions (normal Krebs solution) was significantly different (P < 0.05) from the slope (–0.04 ± 0.3 mV/10 µm) of the regression line when CrMP was present in the bathing solution.

 
RMP in HO2-KO and W/WV Mutant Mice

Ninety-six circular smooth muscle cells were recorded from the small intestine of 6 HO2-KO mice. There was no RMP gradient across the thickness of the circular muscle (Table 1), and the mean (± SE) slope of RMPs across the circular muscle layer was –0.1 ± 0.2 mV/10 µm. Compared with zero slope, the P value was 0.54 (Fig. 6A). There was a significant difference (P < 0.01) between the mean slope observed in the HO2-KO mice and the mean slope observed in NKS in WT mice. There was no significant (P > 0.05) difference in slow-wave frequency between HO2-KO mice and WT mice.


Figure 6
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  		Fig. 6. RMP gradient across the thickness of the circular muscle layer of the small intestine of heme oxygenase 2 knockout (HO2-KO) mouse (A) and W/WV mutant mouse (B). Dashed lines represent regression analysis of data obtained from individual experiments; solid lines represent mean regression lines for all cells in all experiments. The slope of the regression line for all experiments in HO2-KO mice (n = 6) and in 7 W/WV mutant mice were not significantly different from zero slope (for HO2-KO, P = 0.54, for W/WV, P = 0.85). For additional details, see text.

 
Ninety-four circular smooth muscle cells were recorded from the small intestine of 7 W/WV mutant mice. The RMPs of the muscle cells in the outer, middle, and inner group were depolarized compared with RMPs observed in WT mice. The mean (± SE) slope of the RMP gradient was –0.04 ± 0.2 mV/10 µm. The P value was 0.85 when the slope was compared with zero slope. These data suggested that there was no RMP gradient across the circular muscle layer of the W/WV mouse (Fig. 6B). Compared with the mean slope observed in NKS from WT mouse, the P value was less than 0.05. In all seven mutant mice, no slow waves were recorded.

Effect of TTX and ODQ

Recordings were made from 53 cells from four preparations from WT mice after pretreatment with TTX (1 µM) for 30 min. There was no effect of TTX on the RMP gradient (Table 1). The mean (± SE) slope of RMP across the circular muscle layer was –1.2 ± 0.4 mV/10 µm. The P value was less than 0.01 when the slope was compared with zero slope. The mean slopes of RMP gradient in untreated and TTX-treated preparations were not significantly different from each other (P = 0.38) (Fig. 7A). The slow-wave frequency in preparations pretreated with TTX was significantly (P < 0.05) slower compared with the slow-wave frequency observed in untreated preparations (0.66 ± 0.06 vs. 0.78 ± 0.01 cycles/s).


Figure 7
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  		Fig. 7. Lack of effect of TTX (A) and 1H-[1,2,4-]oxadiazolo[4,3-a]-quinoxalin-1-one (ODQ; B) on the RMP gradient in WT mice. Dashed lines represent regression analysis of data obtained from individual experiments; solid lines represent regression lines for all experiments. The mean (± SE) slope of the regression line for all experiments in the preparations treated with TTX (n = 4, 1 µM) and with ODQ (n = 4, 10 µM, B) was –1.2 ± 0.4 and –1.1 ± 0.4 mV/10 µm, respectively. Both were significantly different from zero slope (P < 0.01). For additional details, see text.

 
CO can directly activate ion transporters as well as activate guanylyl cyclase resulting in cGMP production (14, 33). To test whether inhibition of the soluble guanylate cyclase pathway abolished the RMP gradient, 1H-[1,2,4-]oxadiazolo[4,3-a]quinoxalin-1-one (ODQ), a soluble guanylate cyclase inhibitor, was used. In four preparations, muscle strips were pretreated for 30 min with ODQ and recordings were made from 54 cells. ODQ had no effect on the RMP gradient across the circular muscle layer (Table 1, Fig. 7B). The mean (± SE) slope of RMP gradient was –1.1 ± 0.4 mV/10 µm. The P value was less than 0.01 when the slope was compared with zero slope. The slopes for the mean RMP gradient in untreated and ODQ treated preparations were not significantly different from each other (P = 0.59). The slow-wave frequency in untreated and treated preparations was 0.77 ± 0.02 and 0.71 ± 0.08 cycles/s, respectively (P > 0.05).


   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The results of this study show that a gradient in RMP exists across the circular smooth muscle layer of the murine small intestine as well as across the circular muscle layer of the human small intestine. These results extend the original observation made in the murine gastric antrum (9) and canine stomach and intestines (2, 3, 26, 27).

The existence of a gradient in RMP across the circular muscle layer may be central to the ability of the circular muscle layer to regulate the strength of phasic contractions. It is well known that the strength of phasic contractile activity in the gut wall triggered by ongoing electrical slow-wave activity can vary from weak contractions that barely indent the wall to strong and propulsive contractions that can occlude the lumen, with gradations in the strength of contraction between these two extremes. The existence of a voltage gradient is very likely the basis for these variations in the strength of contraction. In the circular muscle layer, the strength of phasic contractions is related to the amplitude of the slow wave and activation of L-type calcium channels (31). Thus, for a weak excitatory stimulus such as during the release of acetylcholine, smooth muscle cells that are more depolarized in the inner layer of the circular muscle and closer to the threshold for activation of L-type calcium channels will contract when the slow wave crosses the threshold for activation of the L-type calcium channels. In contrast, muscle cells in the outer layer of the circular muscle with a more hyperpolarized RMP will not. The entire thickness of the circular muscle layer will contract only when the excitatory stimulus is strong enough and the amplitude of the slow wave crosses the threshold for activation of the L-type calcium channels in the most hyperpolarized circular muscle cells. In essence, the RMP gradient functions as a biological rheostat regulating how much of the thickness of the circular muscle layer contracts during each electrical slow wave.

A number of observations support the hypothesis that CO is the hyperpolarizing agent and that it is released from ICC. Firstly, oxyhemoglobin, which traps CO, abolished the RMP gradient in both the mouse and human small intestine. Since oxyhemoglobin can also trap NO, it might be argued that the hyperpolarizing effect of NO might also be contributing to the RMP gradient. However, the RMP gradients in the mouse and human small intestine were not affected by the NO synthase inhibitor L-NNA, arguing against involvement of NO. Secondly, CrMP, a selective inhibitor of heme oxygenase activity (1), abolished the RMP gradient in the WT mouse. Thirdly, the gradient was abolished in HO2-KO mice.

Although the HO2 enzyme is present in both enteric neurons and ICC, the source of CO is likely to be ICC. The RMP gradient in the mouse small intestine was abolished in W/WV mutant mice that lack ICC in the myenteric region. Further support for the hypothesis that CO was in fact of ICC origin comes from the finding that TTX, a blocker of neural transmission, had no effect on the RMP gradient in the mouse small intestine. Additionally, the RMP of intestinal muscle cells in glial cell line-derived neurotrophic factor knockout mice that lack an enteric nervous system is not depolarized compared with WT controls (34).

Although not studied in any great detail, the mechanism of action by which CO caused hyperpolarization does not appear to depend on stimulation of soluble guanylate cyclase. Like NO, CO can increase soluble guanylyl cyclase activity to stimulate the formation of cGMP (13, 14, 36). CO can also directly increase the channel open probability of KCa in a reversible manner and it increases the activity of other families of K+ channels (15, 32, 33, 35). The results in the present study show that inhibition of soluble guanylyl cyclase activity by ODQ had no effect on the RMP gradient, thereby suggesting that the maintenance of RMP gradient was cGMP independent. There are at least two potential mechanisms by which CO might cause hyperpolarization. It has been suggested that the difference in RMP between muscle cells located at the submucosal border and myenteric region in the dog colon may be related to the Na+-K+-ATPase electrogenic pump (5, 6) or an inward rectifying K+ current (10). The Na+-K+-ATPase pump is the site of action of CO for long-term modulation for cellular activity (18). In a previous study, we showed that CO increases a voltage-dependent outward current by 285% in freshly dispersed circular smooth muscle cells from the dog jejunum and that CO hyperpolarizes the RMP by 15–20 mV (8). The mechanism(s) by which CO hyperpolarizes the membrane potential in the mouse and human small intestine remains to be determined.

In conclusion, the data in this study show that a gradient in RMP exists across the circular smooth muscle layer of the mouse and human small intestine and that the generation and maintenance of the gradient in both mouse and human small intestine is CO dependent. CO, generated by HO2 in ICC, is the likely source.


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 ABSTRACT
 METHODS
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 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by the National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-17238 and by DK-52766.


   ACKNOWLEDGMENTS
 
We thank Jan Applequist for secretarial assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. H. Szurszewski, Dept. of Physiology and Biomedical Engineering, Mayo Clinic College of Medicine, 200 First St. SW, Rochester, MN 55905 (e-mail: gijoe{at}mayo.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
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