Kinetics of urothelial ATP release

【摘要】  Recent reports have proposed that the urothelium can sense mechanical stretch and communicate this information to sensory afferent neurons by the release of ATP into the vicinity of P2X-containing neurons. This report investigates the bidirectional release of ATP by in vitro rabbit urothelium. ATP was measured using the luciferin-luciferase assay. Immediately after washing of both sides of the epithelium, there was a linear increase in ATP content in the mucosal compartment with a rate of 23 ± 6.5 fmol·min -1 ·cm -2 ( n = 18). Serosal ATP content increased as a saturating exponential function, suggesting a constant rate of release and degradation of ATP by ectonucleotidases/exonucleotidases. The presence of a serosal ectonucleotidase/exonucleotidases was demonstrated by the time-dependent decrease in exogenously added ATP. The maximum rate of hydrolysis was 11 pmol·min -1 ·cm -2 with a K m of 0.49 µM. The time course of serosal ATP release was modeled as a constant rate of release ( d : mol·min -1 ·cm -2 ) and rate constant of hydrolysis ( k h : min - ). In control conditions d was 18 fmol·min -1 ·cm -2 and k h of 0.056 ± 0.01 min - ( n = 18). Steady-state serosal chamber content is 370 ± 90 fmol/cm 2, and concentration is 50 ± 1.2 x 10 -12 M. Stretching the tissue resulted in a transient fivefold increase in the rate of mucosal ATP release and a transient sixfold increase in serosal ATP release. Half-osmotic strength solutions increased mucosal release by 10-fold and serosal release by 5-fold. Tissue damage resulted in a step-increase in mucosal chamber ATP content by 6.6 ± 1 pmol/cm 2 and serosal chamber ATP by 0.1 ± 0.06 pmol/cm 2 ( n = 5).

【关键词】  cell swelling cell stretch urothelium cell damage

HISTOLOGICAL STUDIES OF MAMMALIAN urinary bladder have demonstrated that afferent sensory nerve fibers (which contain calcitonin gene-related peptide and substance P) are immunoreactive to P2X 3 (purinergic receptors that are ligand-gated ion channels) and lie in close proximity to the basement membrane of the urothelial cells ( 31 ) as well as being found between urothelial cells ( 2, 4, 10 ). Recent studies suggest that there may be bidirectional communication between the urothelium, underlying neurons, and myofibroblast-like cells ( 24 ). Because ATP is released from the urothelium during bladder distension ( 9 ), this raises the possibility that afferent sensory neurons and myofibroblast-like cells sense stretch via the ATP release from the urothelium ( 30 ). It has been postulated that these myofibroblasts can then modulate the activity of urothelial cells and afferent neurons.

There have been some recent studies on urothelial ATP release. Vlasovska and colleagues ( 30 ) used an isolated intravesical perfused mouse urinary tract (including the lower vertebrae). Using this preparation, these authors reported that there was a basal release of 25 fmol·min -1 ·cm -2, it is important to note that this release is into the lumen of the bladder and not the serosal compartment. In isolated perfused ureters the basal rate of release was 160 pmol·min -1 ·cm -2 into the lumen of the ureters ( 16 ), and this release was increased 70-fold by increasing intraluminal pressure to 300-700 cmH 2 O. In rabbit urinary bladder, Ferguson and colleagues ( 9 ) reported a steady-state basal concentration of ATP in the in vitro bladder of between 0.71 and 6.3 nM and a serosal concentration of 0.22 to 2.2 nM. Stretching the bladder increased the steady-state ATP concentration in the serosal compartment by 224% but not the mucosal compartment ( 9 ). Of interest is that this group also reported that short circuiting the bladder eliminated the stretch-induced ATP release, whereas mucosal amiloride (10 µM) stimulated serosal ATP release. Birder and colleagues ( 5 ) demonstrated that stretching the mouse bladder resulted in an increase in ATP release into the serosal solution (however, they did not report a basal line value), but they did not report whether there was ATP release into the mucosal compartment. Most recently, Wang and colleagues ( 33 ) measured ATP release by rabbit urothelium and demonstrated that stretch increased both mucosal and serosal release and that this ATP release was involved in the movement of cytoplasmic vesicles into the apical membrane during stretch.

This paper will quantify the time course of the basal release of ATP by the urothelium into both the mucosal and serosal compartments, investigate the possible existence of ectonucleotidase/exonucleotidase, and measure the effect of urothelial stretch, cell volume changes, and tissue damage on bidirectional ATP release.

METHODS

Epithelium and electrical measurements. Intact rabbit bladder epithelia (smooth muscle removed) were mounted on a ring with an area of 2 cm 2 and placed vertically in modified Ussing chambers with a mesh support on the basolateral side of the epithelium ( 20 ). Apical and basolateral sides were bathed in a Ringer solution as described below. The apical chamber provided slight hydrostatic pressure, allowing the epithelium to be supported by the mesh. The chambers were fitted with two pairs of Ag-AgCl electrodes; the current-passing electrodes were in the back of each chamber, and the voltage-measuring electrodes were 3 mm from the tissue on either side. The electrodes were connected to an epithelial voltage clamp (EC-800, Warner Instruments, West Haven, CT). This system was control by a computer with a Labmaster DMA A/D board (Scientific Solutions, Solon, OH) and allowed a continuous monitoring of the transepithelial voltage ( V t ), transepithelial conductance ( G t ), and short-circuit current ( I sc ). These parameters along with the time of acquisition were stored digitally and displayed on a monitor and a printer. The analog signal was displayed on an oscilloscope and strip-chart recorder.

Solutions. The Ringer solution contained (in mM) 111.2 NaCl, 25 NaHCO 3, 10 glucose, 5.8 KCl, 2 CaCl 2, 1.2 KH 2 PO 4, and 1.2 MgSO 4. The half-osmotic strength Ringer was diluted 1:1 with deionized water.

The ATP standard (0.9 mg or 1.8 x 10 -6 mol, Sigma, St. Louis, MO) was diluted to 1 x 10 -7 mol/ml. ATP assay mix and assay mix dilution buffer (Sigma) were dissolved as per instructions. Depending on the sensitivity needed for the standard curve, either 4- or 25-fold dilutions of the assay mix were made with the assay mix dilution buffer.

ATP measurements. ATP was measured using an Analytical Luminescence Laboratory luminometer (Monolight 2010, San Diego, CA). Standard ATP curves were measured daily, using five serial dilutions between 10 -9 and 10 -15 mol/assay, over the range of expected ATP concentration values.

ATP (in moles) was calculated using linear regression analysis of the standard curve. A typical standard curve is shown in Fig. 1. The range was 10 -11 to 10 -15 mol, with both axes plotted on a logarithmic scale. The curve is linear over this range.

Fig. 1. Relative light intensity is a linear function of ATP concentration. This shows a typical standard curve over the range of 1 x 10 -11 to 1 x 10 -15 mol ATP. Curves were made for all experimental conditions, or when a new ATP reagent was used.

Standard protocols. In the stretch experiments, the mesh supporting the tissue from the basolateral side was removed, and the basolateral solution decreased by one-third of its volume and the apical solution increased to one and one-third of its volume, causing the tissue to stretch into the basolateral chamber. Solution ATP content calculations were made with these volume corrections. Half-osmotic strength experiments were done by diluting the Ringer solution by half with deionized water. In the epithelial damage (scratch) experiments, the tissue was lightly scraped with a 1-mm-diameter metal rod. Electrical parameters were continually recorded.

Transepithelial capacitance. The apical membrane surface area was estimated by calculating the transepithelial capacitance, where 1 µF = 1 cm 2 of actual membrane area, using the method of Lewis and Moura ( 22 ). In brief, a current step was applied across the epithelium. The time-dependent transepithelial voltage response was digitized at 1-ms intervals. The voltage response was fitted by the sum of two exponentials, yielding two resistor values and two capacitor values. As demonstrated by Lewis and Moura ( 22 ) for the urinary bladder epithelium, the product of the two capacitors divided by their sum yields the effective capacitance (C t ) for the epithelium. Because the apical capacitance is one-fifth the basolateral membrane capacitance ( 7 ), the C t underestimates the apical capacitance by 20%. A change in C t represents the minimal change in the apical membrane surface area.

Statistics. All data are expressed as the means ± SE. Paired and unpaired Students t -tests were used to determine significance (INSTAT, GraphPAD Software, San Diego, CA). P < 0.05 is considered statistically significant. Curve fitting of theoretical functions to the data was performed using a Pentium computer running the nonlinear curve-fitting program NFit (Island Products, Galveston, TX).

RESULTS

Urothelium secretes ATP. Bladder tissues were mounted between modified Ussing chambers, and the tissue was allowed to reach a steady-state G t and V t. The tissues were then washed on both sides with Ringer solution, and mucosal and serosal samples were taken immediately following the wash and then at 5-, 10-, 30-, and 60-min time points. Figure 2, A and B, shows the time course of ATP release into both the mucosal and serosal solutions ( Fig. 2 B is an expanded view of the serosal ATP release shown in Fig. 2 A ). Apical release of ATP into the mucosal compartment increases in a near-linear manner, suggesting a constant rate of ATP release at the apical membrane. This rate of apical release is determined by curve fitting a linear equation to the data; the slope is equal to the rate of release ( d ). In contrast, the quantity of ATP in the serosal solution plateaus. The time course of the serosal fluid ATP content can be modeled as a constant rate of release ( d : mol·min -1 ·cm -2 ) into the bath, and a rate of hydrolysis by ectonucleotidase/exonucleotidase ( k h : min -1 ) ( 12, 14 ). Thus serosal ATP content as a function of time ( t ) is described by the following equation

Both d and k h were determined by curve fitting the above equation to the data.

Fig. 2. A : example of apical and basolateral release of ATP as a function of time. At time 0 and at the arrow, both mucosal and serosal chambers were washed with Ringer. Note that the ATP release into the mucosa is linear over time, suggesting a constant rate of release and no hydrolysis of ATP. The rate of release for the first 60 min (1.35 x 10 -14 mol·min -1 ·cm -2 ) is the same as for the last 145 min (1.45 x 10 -14 mol·min -1 ·cm -2 ). B : detailed view of serosal ATP release in A. Serosal ATP release reaches a plateau after 40 min, suggesting ATP hydrolysis.

The rate of ATP release into the mucosal and the serosal solutions is not significantly different; the mucosal rate is 23 ± 6.5 x 10 -15 mol·min -1 ·cm -2 ( n = 18), and the serosal rate is 18 ± 4.8 x 10 -15 mol·min -1 ·cm -2 ( n = 18, P = 0.54). These data suggest that the urothelium releases ATP on both the mucosal and serosal sides. However, the increase in serosal content did not continue; the content reached a plateau of 368 ± 90 x 10 -15 mol/cm 2 ( n = 18) with k h = 0.0555 ± 0.01 min -1 ( n = 14; 4 of the 18 had no measurable release), suggesting there is ectonucleotidase/exonucleotidase activity on the serosal side. This rate constant suggests that following a wash the serosal ATP content will plateau in 60 min.

There is no change in C t, G t, or I sc over the time period in which ATP release was measured (data not shown).

Presence of an ectonucleotidase/exonucleotidase. The data from Fig. 2 B suggest the presence of an ectonucleotidase/exonucleotidase in the serosal side of the tissue. Because all muscle layers have been removed, this activity mostly likely resides in the urothelium; however, we cannot rule out the possibility that some cells such as blood vessels, nerve fibers, or mast cells are still present after the dissection and contribute to the nucleotidase activity. To further assess ectonucleotidase/exonucleotidase activity, exogenous ATP was added to the mucosal and serosal solutions and the decrease in bath ATP concentration was measured over a 3-h period ( Fig. 3 A ). As shown in Fig. 3 A, there is a decrease in serosal ATP concentration. The small decrease in mucosal ATP is not significantly different from the decrease in ATP in the absence of tissue (data not shown). Figure 3 B shows the relationship between the added bath ATP concentration and the rate of hydrolysis. Mucosal and Ringer curves were best described by a straight line, suggesting that spontaneous loss of ATP content was not associated with nucleotidase activity. Fitting the serosal data by the Michaelis-Menten equation gave a K m of 0.49 µM and a maximum rate of hydrolysis of 11 x 10 -12 mol·min -1 ·cm -2 (see figure legend for details).

Fig. 3. A : time course of decrease in mucosal and serosal bathing solution ATP content for 5 different tissues. ATP was added to both solutions to a final concentration of 1.0 x 10 -6 M. Note that the rate of decrease in the ATP content of the serosal compartment is greater than for the mucosal compartment. The rate of decrease in mucosal ATP was not different from the rate of decrease in ATP in the absence of tissue (not shown). This suggests that the serosal surface but not mucosal surface of the urothelium has ectonucleotidase/exonucleotidase activity. B : rate of ATP hydrolysis as a function of bath ATP concentration. The concentration dependence of the rate of decrease in ATP in the mucosal chamber ( ) and in the absence of tissue (Ringer, ) was not different from each other and represents a spontaneous hydrolysis of ATP over time. Mucosa and Ringer data were best described by a straight line with a slope of 7.5 x 10 -6 l·m -1 ·cm -2. Serosal data were fit by the sum of a straight line (spontaneous hydrolysis) and the Michaelis-Menten equation. During curve fitting of the data, the slope of the straight line was held constant at 7.5 x 10 -6 l·min -1 ·cm -2. Best fit values for serosa ( ) gave a K m of 0.49 µM/l and a V max of 11 pmol·min -1 ·cm -2.

We also monitored the effect of exogenous ATP on C t, G t, and I sc. Over a 180-min period, at ATP concentrations of 10 -8 and 10 -7 M there was not a significant increase in C t (from 1.23 ± 0.28 to 1.07 ± 0.09 µF/cm 2 and from 1.07 ± 0.04 to 1.037 ± 0.043 µF/cm 2, respectively). However, at an ATP concentration of 10 -6 M, there was a significant increase (from 1.071 ± 0.081 to 1.177 ± 0.072 µF/cm 2, n = 4, P = 0.042). At all three ATP concentrations, there was a significant increase in G t from 47.2 ± 4.5 to 55.2 ± 4.3 µS/cm 2 ( n = 9) and an increase in I sc from 2.25 ± 0.26 to 2.7 ± 0.26 µA/cm 2 ( n = 9) after 180 min. The relationship between G t and I sc was linear, suggesting that ATP was increasing the apical membrane conductance. Using the method of Yonath and Civan ( 34 ), the inverse slope of the plot of conductance vs. current is equal to the cell electromotive force of 59 ± 8.8 mV ( n = 9) and suggests that the increase in conductance is cation selective.

Stretch increases ATP release. To test the effect of stretch on urothelial ATP release, we first measured the release of ATP into both the mucosal and serosal solution for 30 min. After this control period, the volume of the serosal chamber was reduced to two-thirds of its normal volume, whereas the mucosal chamber was increased to one and one-third of its normal volume, causing the epithelium to stretch into the serosal chamber. This generated a constant 2 cm of water pressure gradient from mucosa to serosa. Mucosal and serosal ATP release was then measured at 5, 15, 20, 35, and 65 min during stretch. The rate of secretion as a function of time was calculated as the slope between successive data points. The rate of mucosal ATP release increased from a control rate of 100 ± 82 x 10 -15 mol·min -1 ·cm -2 ( n = 5) to a peak of 510 ± 188 x 10 -15 mol·min -1 ·cm -2 ( n = 5); these rates are significantly different ( P = 0.048). The time to the peak rate of secretion is 16 ± 2.4 min. The normalized rate of mucosal ATP release is shown in Fig. 4 A. Immediately following stretch, there is an initial decrease in the rate of release followed by an increase, which peaks at 20 min and then decays to the prestretch value. The rate of serosal ATP release changed from a control rate of 2.34 ± 0.9 x 10 -15 mol·min -1 ·cm -2 ( n = 5) to a peak rate of 18 ± 8.5 x 10 -15 mol·min -1 ·cm -2 ( n = 5, P = 0.048). The time to the peak rate of release was 24 ± 10.7 min for serosal release. The serosal time course for the normalized rate of secretion is similar to the mucosal release ( Fig. 4 B ). The times to peak rate of release for mucosal and serosal ATP are not significantly different from each other ( P = 0.6).

Fig. 4. A and B : mucosa ( A ) and serosa ( B ). Stretching the tissue by hydrostatic pressure resulted in a significant but transient increase in the rate of ATP release into both the mucosal and serosal compartments. The rate of release for a given tissue was normalized to the maximum rate for that tissue to reduce the variability among the tissues. Note that the increase in the rate of release into the mucosal compartment is greater than into the serosal compartment. The peak rate of release occurred 20 min after bowing the tissue into the serosal compartment.

Figure 5, A - C, shows the time-dependent effects of stretch on C t, G t, and I sc. As in previous studies, bowing the tissue into the serosal compartment results in a time-dependent increase in the transepithelial capacitance of 21 ± 4.6% ( n = 5) over 60 min (from 1.18 ± 0.103 to 1.42 ± 0.13 µF/cm 2, n = 5; this increase was significant; P = 0.01; Fig. 5A ) ( 19 ). G t increased from 66 ± 14 to 220 ± 50 µS/cm 2 in the first 60 s ( n = 5), reaching a maximum at 15 min and then decreased to a new plateau of 187 ± 45 µS/cm 2 ( Fig. 5 B ), which was significantly different from the prestretch value. Last, I sc increased from 1.86 ± 0.4 to 5.86 ± 1.84 µA/cm 2 in the first 60 s and then decreased over time to a new plateau value of 2.43 ± 0.32 µA/cm 2, which was significantly different from the prestretch value ( Fig. 5 C ). Similar increases in G t and I sc on initial exposure to a hydrostatic gradient has been previously reported for the urothelium ( 32 ). The initial increase in I sc suggests that all or part of the initial increase in G t is at the apical membrane and not due to cell damage. Of interest is that the initial change in C t, G t, and I sc occurs before there was a measurable change in ATP release.

Fig. 5. Effect of stretching the urothelium into the serosal chamber on normalized transepithelial capacitance (C t; A ), normalized transepithelial conductance ( G t; B ), and normalized short-circuit current ( I sc; C ). All time courses were normalized to the respective value immediately before stretching of the tissue.

Hyposmotic solutions stimulate ATP release. To determine the effects of hyposmotic solutions on the rate of ATP release, the mucosal and serosal chambers were washed with a half-osmotic strength Ringer solution. ATP release increased into both the mucosal and serosal compartments (see Fig. 6, A and B, and Table 1 ). Mucosal ATP release increased from 12.4 ± 8.5 x 10 -15 ( n = 5) to 133 ± 79 x 10 -15 mol·min -1 ·cm -2, a 10-fold increase ( Fig. 6 A ). When the chamber solutions were replaced with regular-strength Ringer, the release rate returned to 23.0 ± 13 x 10 -15 mol·min -1 ·cm -2. Release into the serosal solution was fit to an exponential. The initial rate of increase in ATP in the serosa was 10-fold higher when the tissue was exposed to the hyposmotic solution than control ( Fig. 6 B and Table 1 ). The difference in the k h value between the regular and hyposmotic solutions was not significant. Plateau ( d / k h ) values between the two conditions were significantly different. Two of the five control tissues did not have a measurable serosal ATP release under control conditions; thus we could not estimate a value for k h for these two tissues.

Fig. 6. A : effect of solution osmolality on ATP release. Baseline ATP release is measured from time 0 to 60 min. After the 60-min measurement, both mucosal and serosal solutions were replaced with half-osmolality solution. Note that there is a rapid increase in ATP release in both mucosal and serosal solutions. Returning to an isosmotic solution returns ATP release to baseline. B : detailed view of ATP release into the serosal compartment. Note the rapid increase in ATP when the solution osmolality is lowered. ATP release returns to normal on return to normal solution osmolalit y.

Table 1. Effect of hyposmotic solutions on serosal ATP release

C t increased 60% over the 60-min exposure to half-osmotic strength Ringer (from 1.05 ± 0.106 to 1.54 ± 0.15 µF/cm 2, n = 6, P = 0.004) and returned to control after return of tissue to isosmotic Ringer ( Fig. 7 A ). This time course was similar to that previously reported ( 22 ). G t initially decreased on exposure to half-osmotic strength Ringer and returned to control values over the following hour. On returning of tissue to isosmotic Ringer, there was a small decrease in G t followed by a return to the control value ( Fig. 7 B ). The time course of the I sc ( Fig. 7 C ) closely follows that of G t, suggesting that the changes in G t are mostly at the apical cell membrane and not due to cell damage. This suggests that ATP release is not due to tissue damage.

Fig. 7. Effect of half-osmotic strength Ringer on C t ( A ), G t ( B ), and I sc ( C ). C t, G t, and I sc were normalized to the respective value immediately before half-osmotic strength Ringer was added to both mucosal and serosal chambers.

ATP release by cell damage. Tissues were damaged by lightly scraping the mucosal surface with a blunt 1-mm-diameter metal rod. Tissue damage results in a significant release of ATP into the mucosal chamber but not the serosal chamber ( Fig. 8, A and B ). Mucosal solution ATP content increased from 1.7 ± 0.5 x 10 -13 ( n = 5) to 68 ± 12 x 10 -13 mol/cm 2 ( n = 5, P = 0.0002), giving a mean increase of 66 ± 11 x 10 -13 mol/cm 2. Serosal ATP content increased from 0.043 ± 0.014 x 10 -13 ( n = 5) to 0.87 ± 0.60 x 10 -13 mol/cm 2 after 5 min ( n = 5, P = 0.1). Although there was not a significant increase in serosal ATP release caused by damage, in every instance there was an increase in serosal ATP release, with a mean value of 0.96 ± 0.57 x 10 -13 mol/cm 2.

Fig. 8. Increase in ATP release and G t after scraping of the apical surface with a metal rod in mucosal ( A ) and serosal compartments ( B ). Control ( T = 0) was immediately after a wash, so ATP is barely measurable. The ATP concentration in both chambers then increased over the following 30 min. Damage ( T = 2), immediately after scraping, shows a 5-fold increase in the mucosal and a 10-fold increase in the serosal ATP concentration ( n = 5).

Figure 8, A and B, shows the change in G t along with the time course of the release of ATP into both the mucosal ( Fig. 8 A ) and serosal ( Fig. 8 B ) compartments. As is evident, even minor damage to the tissue results in a large increase in bath ATP levels.

DISCUSSION

Kinetics of ATP release. In the rabbit urothelium, apical membrane ATP release increases in a near-linear manner, whereas basolateral ATP release reaches a plateau after 1 h. The difference in time courses represents a loss of ATP activity most likely due to ectonucleotidase/exonucleotidase as reported for other tissues ( 12, 14 ). The time-dependent change in serosal fluid ATP content was modeled as a constant rate of release (i.e., d : mol·min -1 ·cm -2 ) into the bath and a rate of hydrolysis by ectonucleotidase/exonucleotidase (i.e., k h : min -1 ). The rate of control mucosal ATP release was 23 fmol·min -1 ·cm -2 and serosal ATP release was 18 fmol·min -1 ·cm -2 of tissue, the rate of hydrolysis was 0.06 min -1, and the predicated plateau value for serosal ATP is then 368 fmol/cm 2. How do these numbers compare with other reports of urothelial ATP release? One of the problems in addressing this question is that there is no uniformity in the literature in the units used for ATP release. As examples, urothelial ATP release has been reported either as a percent increase from control ( 28 ) for tissue-cultured human bladder cells or as picomolar per gram tissue wet weight of human or porcine bladder strips, which includes urothelium and muscle ( 17 ).

Some of the values listed below have been calculated based on estimates of tissue surface area, sampling interval, and solution volume. Mucosal release of ATP from mouse bladder ( 30 ) was estimated (by us) at 25 fmol·min -1 ·cm -2. ATP release from primary cultures of cat bladder epithelial cells was estimated at 40 fmol·min -1 ·cm -2 ( 3 ). From a recent study on rabbit urinary bladder ( 33 ), we estimated a basal rate of mucosal and serosal release of 1.25 fmol·min -1 ·cm -2. These values are in reasonable agreement with those reported above.

In contrast, we estimate that guinea pig ureters release ATP into the lumen at a rate of 160 pmol·min -1 ·cm -2 ( 16 ), 7,000-fold greater than bladder urothelium. Ferguson and colleagues ( 9 ) were the first to report ATP release by rabbit urinary bladder epithelium. They found that after washing of both compartments, mucosal ATP concentration reached a steady-state value between 0.7 and 6.3 nM and serosal ATP between 0.2 and 2.2 nM. The serosal ATP content was then calculated as 4-44 pmol/cm 2 of tissue. Our data suggest that the mucosal content should not reach a plateau and that the serosal content should be 0.4 pmol/cm 2 of tissue area, a value significantly lower than that reported by Ferguson et al. A possible explanation for this difference is that our tissues had all muscle removed, whereas Ferguson and colleagues mounted the urothelium with muscle attached. In our experience, leaving the muscle attached results in rhythmic contraction of the urothelium, which might result in stretch-induced ATP release or cell damage. Similarly, the high rate of release by the ureters reported by Knight and colleagues ( 16 ) might be a consequence of mechanical stress due to the intrinsic peristaltic contractions of the ureters.

The basal rate of release by other nonneural cells is similar to that reported in this paper. Transformed bovine nonpigmented epithelial cells (NPE) and pigmented epithelial cells (PE) release ATP at a basal rate of 70 and 30 fmol·min -1 ·cm -2 ( 23 ). A6 cells (renal cell line from Xenopus laevis ) have a serosal rate of release of 40 fmol·min -1 ·cm -2 ( 14 ). Calu 3, T84 cells, or 9HTEo cells grown on plastic had no measurable ATP release ( 12 ); however, mechanical stress caused a large increase in release. Human umbilical vein endothelial cells had a spontaneous release rate of 800 fmol/min for 10 6 cells ( 6 ), and aortic endothelial cells released ATP at a rate of 2,900 fmol/min for 10 6 cells ( 26 ).

Ectonucleotidase/exonucleotidase. Serosal release of ATP was modeled as a constant rate of release and a rate of hydrolysis by either ectonucleotidase or exonucleotidase. To test for the possibility of ectonucleotidase/exonucleotidase, the rate of disappearance of exogenously added ATP was measured. In the absence of tissue, there was a time-dependent decrease in bath ATP. Mucosal ATP decreased in a time-dependent manner and was the same as in the absence of tissue, suggesting the absence of mucosal ectonucleotidase/exonucleotidase activity. Serosal ATP decreased in a time-dependent manner at a rate greater than predicted by spontaneous hydrolysis, suggesting the presence of an ectonucleotidase/exonucleotidase. In the present study, the rate of hydrolysis as a function of concentration was fit by the Michaelis-Menten equation and yielded a dissociation constant of 0.49 µM and a maximum rate of hydrolysis of 11 pmol·min -1 ·cm -2. The presence of serosal ectonucleotidase/exonucleotidase activity was recently demonstrated by Wang and colleagues ( 33 ). They reported that over 300 min, there was a 40% decrease in the concentration of added ATP (50 µM) that was not due to spontaneous degradation. This translates into a rate of hydrolysis of 433 pmol·min -1 ·cm -2, which is 40 times greater than our maximum reported value. Possible explanations include the degree of urothelial stretch (a more highly stretch urothelium will have fewer cells) and/or completeness of the dissection. In this regard, guinea pig urinary bladder (muscle and epithelium) had a maximum rate of hydrolysis of 0.5 pmol·s -1 ·mg wet wt -1 and a dissociation constant of 0.8 mM ( 35 ). The difference in dissociation constant for the rabbit urothelium and guinea pig urinary bladder suggests that the nonurothelial components of bladder (predominantly the detrusor) has an ectonucleotidase with a different dissociation constant, as was recently reported for guinea pig detrusor and human detrusor, which have a dissociation constant of 0.9 and 1.5 mM, respectively ( 13 ). Thus the urothelium has high-affinity ectonucleotidase activity, which will then control the extracellular ATP concentration in the physiological range for purinergic receptors, whereas the detrusor has a low-affinity, but high throughput ectonucleotidase, which will help control toxic levels of extracellular ATP.

Both human and porcine bladder strips (which contain both muscle and urothelium) were demonstrated to have ectonucleotidase/exonucleotidase activity ( 17 ) as was nonpigmented epithelium and pigmented epithelium from the ciliary body ( 23 ) and A6 cells ( 14 ); however, neither the dissociation constant nor maximum rate of hydrolysis was determined for these tissues. The epithelial tissue culture cell lines Calu 3 and 9HTEo demonstrate ectonucleotidase/exonucleotidase activity with a dissociation constant of 2.4 and 0.5 µM, respectively, and a maximum rate of hydrolysis of 11 and 1 pmol·min -1 ·10 6 cells -1, respectively ( 12 ). These values are similar to that determined for rabbit urothelium in this study.

Stretch, swelling, and ATP release. Mechanical stretch is a known stimulus for ATP release by many different cell types. As an example, Calu 3, T84, and 9HTEo cells all increase ATP release from zero to significant levels depending on the degree and frequency of the stretch-relaxation cycle ( 12 ). The urothelium is a tissue that undergoes a number of stretch-relaxation cycles in a 24-h period. Ferguson and colleagues ( 9 ) first demonstrated that stretching the urothelium in Ussing chambers increases serosal ATP release by 224% with no effect on mucosal release, in bladder strips a 50% increase in length increased ATP release by 500% ( 17 ), an eightfold increase in mucosal release in mouse bladder ( 30 ), and an increase in ATP release by the ureters as the intraluminal pressure was increased. Urothelial cells cultured on flexible supports also increased the rate of ATP release ( 28 ). The present study also demonstrated that there was an initial five- and eightfold increase in the rate of mucosal and serosal ATP release, respectively, when the tissue was stretched. The serosal release is most likely an underestimate due to the presence of ectonucleotidase/exonucleotidase. After the initial increase in the rate of release, the rate then decreased over time to baseline values, suggesting either that the tension on the cells decreased or that the signaling cascade desensitized. Of interest is that the apical membrane surface area of the epithelium increases during stretch; this increase in area should decrease the membrane tension. Wang and colleagues ( 33 ) also reported that stretching the rabbit urothelium resulted in an increase in both mucosal and serosal ATP release to 65 fmol·min -1 ·cm -2 for serosa and 4,333 fmol·min -1 ·cm -2 for mucosa compared with 18 fmol·min -1 ·cm -2 for serosa and 510 fmol·min -1 ·cm -2 for muocsa. The difference is most likely due to the magnitude of the applied pressure, which is 2 cm of water in this paper and 8 cm of water in the other report ( 33 ).

Cell swelling (by decreasing the osmolality of the bathing solution) is a commonly used method to induce cell ATP release. The percent increase in ATP release for different preparations is outlined in Table 2. Of interest is that all of these cells release ATP in response to a hyposmotic challenge and that the percent increase is similar for all of the cells, suggesting that a similar mechanism resulting in ATP release might be shared among these cells.

Table 2. Tissue dependence of osmolality on ATP release

Damage from the mucosal side of the urothelium resulted in a dramatic increase in mucosal ATP release and a smaller serosal ATP release. G t increased from 50 to 1,500 µS/cm 2, and mucosal ATP content increased from 0.17 to 6.8 pmol/cm 2. If the cell ATP concentration is 5 mM ( 12 ), then scratching the surface destroyed urothelial cells with an equivalent volume of 1.3 x 10 -6 cm 3. If cell height is 40 x 10 -4 cm, then scratching destroyed cells occupying an area of 330 x 10 -6 cm 2 or about nine cells (assuming that each cell is a square with a side = 60 x 10 -4 cm). Based on the damaged surface area, cell height, and free solution resistivity of 60 ·cm, one can estimate that the tissue conductance would increase by 1,375 µS/cm 2, a value very close to 1,500 µS/cm 2 measured. A word of caution is that if the cell height is decreased by half, the conductance estimate will be increase to 5,500 µS/cm 2, whereas a doubling of cell height will decrease the conductance to 344 µS/cm 2.

Can either mucosal or serosal ATP release be due to cell lysis? If a surface cell is 60-µm square and 40 µm in height, then the ATP content will be 720 fmol, and one might expect to see bath ATP increase with a step of approximately this value. This was not observed. Also, based on the above calculation, one would also expect to measure a change in conductance of 160 µS for every cell that lyses. Neither cell stretch not swelling resulted in a significant change in G t, suggesting that ATP release is not a consequence of cell lysis. Wang and colleagues ( 33 ) investigated the mechanism of mucosal and serosal ATP release in rabbit urothelium using a pharmacological approach. Due to the multiple sites of action of each pharmacological agent, the precise mechanism, e.g., vesicular release, ABC transport protein, nucleoside transporters, or gap junction hemichannels, could not be ascertained; however, quinicrine staining demonstrated punctate staining, suggesting vesicles as a possible mechanism.

Urothelial damage could occur by cystoscopy, bacterial infection, presence of xenobiotics, bladder outlet obstruction, or generation of reactive oxygen species. In addition to the immediate release of ATP due to damage, endogenous ATP in the bladder lumen will also diffuse into the serosa and activate sensory neurons. In this regard, a recent report demonstrated that in the presence of luminal ATP, the loss of urothelial barrier function resulted in detrusor overactivity ( 25 ).

Physiological significance. Urothelial cells contain numerous receptors, among them purinergic receptors ( 8, 18, 27 ) and vanilloid receptors that are involved in exocytosis and endocytosis of apical vesicles during bladder filling and collapse (see Ref. 1 for a review). Ferguson and colleagues ( 9 ) suggested a possible communication between the urothelium and sensory afferent neurons (which are immunoreactive to P2X 3 and contain calcitonin gene-related peptide and substance P) that have been localized close to the basement membrane of the urothelial cells ( 31 ) as well as between the urothelial cells ( 4, 10 ). Afferent fibers from P2X 3 (-/-) mice were reported to have a decreased nerve firing rate during bladder distension even though the ATP release was similar to that of wild-type mice ( 30 ). Thus urothelial cells may be involved in sensing distension and conveying that information to afferent neurons. More recently, it has been suggested that there may be bidirectional communication between the underlying neurons and myofibroblast-like cells and the urothelium ( 24 ). Given that during bladder distension ATP is released from the urothelium ( 9 ), this raises the possibility that afferent sensory neurons and myofibroblast-like cells sense stretch via ATP release from the urothelium ( 30 ). As a consequence, these myofibroblasts might then modulate the activity of urothelial cells and afferent neurons ( 24 ).

In addition to the purinergic receptors, the vanilloid receptor (TRPV1) is involved in the sensory circuitry. TRPV1 knockout mice have different voiding patterns (larger bladder capacity, absence of large voiding contractions, and diminished nerve firing rate to low pH and capsaicin) from wild-type mice ( 5 ). The bladders from the TRPV1 knockout mice did not release ATP. Thus the vanilloid receptor is involved in communicating stretch to the afferent sensory neurons perhaps by altering urothelial ATP release ( 5 ).

Of interest is that the vanilloid knockout mice did not increase apical membrane surface area in response to stretch nor did they release ATP ( 5 ), suggesting an effect of extracellular ATP on pressure-induced vesicle insertion. In addition, P2X 2 and P2X 3 knockout mice or addition of apyrase (an exonucleotidase) to the serosal compartment also inhibited stretch-induced membrane area increase ( 1 ). This raises the possibility that extracellular ATP is required and sufficient for membrane area increase. Our results on the effect of exogenous ATP on capacitance changes are in disagreement with a previous report ( 33 ). Those authors reported that exogenous ATP at concentrations as low as 10 -9 M were sufficient to increase C t. In contrast, this paper demonstrated that neither 10 -8 or 10 -7 M increased C t, whereas 10 -6 M caused a modest 10% increase over 180 min. In addition, during stretch, capacitance increased before there was a measurable change in bath ATP concentration, again suggesting that ATP might not be required for an increase in membrane area.

Two recent studies have suggested that urothelial ATP release might be involved in interstitial cystitis (IC). IC is a chronic inflammation of unknown etiology and has the following symptoms: diminished urinary capacity, hematuria, frequency, diffuse abdominal pain, and painful urination ( 15 ). Cultured urothelial cells from cats with feline interstitial cystitis (proposed to be the feline equivalent of interstitial cystitis) had an increased release of ATP compared with cells from control cats ( 3 ). Similarly, cultured urothelial cells from bladder biopsies of patients with interstitial cystitis also had an increased rate of ATP release ( 28 ). Whether this is due to an increase in cell release or the presence of different densities of ectonucleotidase/exonucleotidase on the cell surface was not investigated.

It is not known what other physiological factors (in addition to stretch and hyposmotic solutions) can modulate ATP release. For example, alterations in the apical membrane permeability of the bladder might stimulate ATP release via cell volume changes and a similar cellular response might occur as a function of urinary constituents. Thus the urothelium may have a chemical sensory role in addition to the mechanosensory role (sensing epithelial stretch) as recently proposed ( 30 ).

A crucial question is, What is the concentration of ATP near the afferent sensory neurons? A number of factors will influence the concentration of ATP in the lateral intercellular space (LIS) of the urothelium, and include: the rate of ATP release, the volume of the space in to which ATP is released, the density of the ectonucleotidase/exonucleotidase, the thickness of the underlying tissue, and steric hindrance offered by the connective tissue. These latter factors will alter the rate of diffusion of ATP away from the site of release. A recent study developed a computer model to estimate the time course of ATP concentration changes in the LIS of A6 monolayers grown on permeable supports during an exposure to a hyposmotic challenge ( 11 ). The time course of the LIS ATP concentration was based on measured changes in ATP concentration of the basolateral bathing solution and assumed that the ATP measured was only of basolateral membrane origin, this membrane both released and degraded ATP, and ATP diffusion into the basolateral compartment depends on the concentration gradients across the permeable support and the diffusion rate in the filter. This model demonstrates that the LIS ATP concentration is much higher than in the basolateral compartment (by a factor of at least 5) and that the time-dependent changes in ATP concentration in the bath are much slower than in the LIS. Thus our measurements underestimate the magnitude and overestimate the time course of ATP release into the urothelial LIS.

GRANTS

This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-51382.

ACKNOWLEDGMENTS

We thank Dr. O. P. Hamill and Dr. R. Maroto for advice on the ATP assay.

These studies have been published in abstract form ( FASEB J 19: A761, 2005).

【参考文献】
  Apodaca G. The urothelium: not just a passive barrier. Traffic 5: 117-128, 2004.

Birder L. Involvement of the urinary bladder urothelium in signaling the lower urinary tract. Proc West Pharmacol Soc 44: 85-86, 2001.

Birder L, Barrick S, Roppolo JR, Kanai A, De Groat WC, Kiss S, and Buffington CA. Feline interstitial cystitis results in mechanical hypersensitivity and altered ATP release from bladder urothelium. Am J Physiol Renal Physiol 285: F423-F429, 2003.

Birder LA, Kanai AJ, De Groat WC, Kiss S, Nealen ML, Burke NE, Dineley KE, Watkins S, Reynolds IJ, and Caterina MJ. Vanilloid receptor expression suggests a sensory role for urinary bladder epithelial cells. Proc Natl Acad Sci USA 98: 13396-13401, 2001.

Birder LA, Nakamura Y, Kiss S, Nealen ML, Barrick S, Kanai AJ, Wang E, Ruiz G, De Groat WC, Apodaca G, Watkins S, and Caterina MJ. Altered urinary bladder function in mice lacking the vanilloid receptor TRPV1. Nat Neurosci 5: 856-860, 2002.

Bodin P and Burnstock G. ATP-stimulated release of ATP by human endothelial cells. J Cardiovasc Pharmacol 27: 872-875, 1996.

Clausen C, Lewis SA, and Diamond JM. Impedance analysis of a tight epithelium using a distributed resistance model. Biophys J 26: 291-318, 1979.

Elneil S, Skepper JN, Kidd J, and Williamson GG. Distribution of P2X1 and P2X3 receptors in the rat and human urinary bladder. Pharmacology 63: 120-128, 2001.

Ferguson DR, Kennedy I, and Burton TJ. ATP is released from rabbit urinary bladder epithelial cells by hydrostatic pressure changes - a possible sensory mechanism? J Physiol 505: 503-511, 1997.

Gabella G and Davis C. Distribution of afferent axons in the bladder of rats. J Neurocytol 27: 141-155, 1998.

Gheorghiu M and Van Driessche W. Modeling of basolateral ATP release by hypotonic treatment in A6 cells. Eur Biophys J 33: 412-420, 2004.

Grygorczyk R and Hanrahan JW. CFTR-independent ATP release from epithelial cells triggered by mechanical stimuli. Am J Physiol Cell Physiol 272: C1058-C1066, 1997.

Harvey RA, Skennerton DE, Newgreen D, and Fry CH. The contractile potency of adenosine triphosphate and ecto-adenosine triphosphatase activity in guinea pig detrusor and detrusor from patients with a stable, unstable or obstructed bladder. J Urol 168: 1235-1239, 2002.

Jans D, Srinivas SP, Waelkens E, Segal A, Larivière E, Simaels J, and Van Driessche W. Hypotonic treatment evokes biphasic ATP release across the basolateral membrane of cultured renal epithelia (A6). J Physiol 545: 543-555, 2002.

Jones CA and Nyberg L Jr. Epidemiology of interstitial cystitis. Urology 49: 2-9, 1997.

Knight GE, Bodin P, De Groat WC, and Burnstock G. ATP is released from guinea pig ureter epithelium on distension. Am J Physiol Renal Physiol 282: F281-F288, 2002.

Kumar V, Chapple CC, and Chess-Williams R. Characteristics of adenosine triphosphate release from porcine and human normal bladder. J Urol 172: 744-747, 2004.

Lee HY, Bardini M, and Burnstock G. Distribution of P2X receptors in the urinary bladder and the ureter of the rat. J Urol 163: 2002-2007, 2000.

Lewis SA and de Moura JL. Incorporation of cytoplasmic vesicles into apical membrane of mammalian urinary bladder epithelium. Nature 297: 685-688, 1982.

Lewis SA, Eaton DC, Clausen C, and Diamond J. Nystatin as a probe for investigating the electrical properties of a tight epithelium. J Gen Physiol 70: 427-440, 1977.

Lewis SA and Lewis JR. Bidirectional ATP secretion by urothelium (Abstract). FASEB J 19: A761, 2005.

Lewis SA and Moura JLC. Apical membrane area of rabbit urinary bladder increases by fusion of intracellular vesicles: an electrophysiological study. J Membr Biol 82: 123-136, 1984.

Mitchell CH, Carre DA, McGlinn AM, Stone RA, and Civan MM. A release mechanism for stored ATP in ocular ciliary epithelial cells. Proc Natl Acad Sci USA 95: 7174-7178, 1998.

Morrison J, Birder L, Craggs M, De Groat W, Downe J, Drake M, Fowler C, and Thor K. Cell biology. In: Incontinence, edited by Abrams P, Cardozo L, Khoury S, and Wein A. Paris, France: Health Publ., 2005, p. 363-422.

Nishiguchi J, Hayashi Y, Chancellor MB, de Miguel F, deGroat WC, Kumon H, and Yoshimura N. Detrusor activity induced by intravesical application of adenosine 5'-triphosphate under different delivery conditions in rats. Urology 66: 1332-1337, 2005.

Oike M, Kimura C, Koyama T, Yoshikawa M, and Ito Y. Hypotonic stress-induced dual Ca 2+ responses in bovine aortic endothelial cells. Am J Physiol Heart Circ Physiol 279: H630-H638, 2000.

Sun Y and Chai T. Up-regulation of P2X3 receptor during stretch of bladder urothelial cells from patients with interstitial cystitis. J Urol 171: 448-452, 2004.

Sun Y, Keay S, De Deyne PG, and Chai TC. Augmented stretch activated adenosine triphosphate release from bladder uroepithelial cells in patients with interstitial cystitis. J Urol 166: 1951-1956, 2001.

Taylor AL, Kudlow BA, Marrs KL, Gruenert DC, Guggino WB, and Schwiebert EM. Bioluminescence detection of ATP release mechanisms in epithelia. Am J Physiol Cell Physiol 275: C1391-C1406, 1998.

Vlaskovska M, Kasakov L, Rong WF, Bodin R, Bardini M, Cockayne DA, Ford AP, and Burnstock G. P2X 3 knock-out mice reveal a major sensory role for urothelially released ATP. J Neurosci 21: 5670-5677, 2001.

Wakabayashi Y, Tomoyoshi T, Fujimiya M, Ryohachi A, and Maeda T. Substance P-containing axon terminals in the mucosa of the human urinary bladder: pre-embedding immunohistochemistry using cryostat sections for electron microscopy. Histochemistry 100: 401-407, 1993.

Wang ECY, Lee JM, Johnson JP, Kleyman TR, Bridges R, and Apodaca G. Hydrostatic pressure-regulated ion transport in bladder uroepithelium. Am J Physiol Renal Physiol 285: F651-F663, 2003.

Wang ECY, Lee JM, Ruiz WG, Balestreire EM, von Bodungen M, Barrick S, Cockayne DA, Birder L, and Apodaca G. ATP and purinergic receptor-dependent membrane traffic in bladder umbrella cells. J Clin Invest 115: 2412-2422, 2005.

Yonath J and Civan MM. Determination of the driving force of the Na + pump in toad bladder by means of vasopressin. J Membr Biol 5: 366-385, 1971.

Ziganshin AU, Hoyle CHV, Ziganshina LE, and Burnstock G. Effects of cyclopiazonic acid on contractility and ecto-ATPase activity in guinea-pig urinary bladder and vas deferens. Br J Pharmacol 113: 669-674, 1994.

作者单位：Department of Neuroscience and Cell Biology, University of Texas Medical Branch, Galveston, Texas