Control of descending vasa recta pericyte membrane potential by angiotensin II

Thomas L. Pallone and James M.-C. Huang

Division of Nephrology, School of Medicine, University of Maryland, Baltimore, Maryland 21201-1595


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Using nystatin perforated-patch whole cell recording, we investigated the role of Cl- conductance in the modulation of outer medullary descending vasa recta (OMDVR) pericyte membrane potential (Psi m) by ANG II. ANG II (10-11 to 10-7 M) consistently depolarized OMDVR and induced Psi m oscillations at lower concentrations. The Cl- channel blockers anthracene-9-decarboxylate (1 mM) and niflumic acid (10 µM) hyperpolarized resting pericytes and repolarized ANG II-treated pericytes. In voltage-clamp experiments, ANG II-treated pericytes exhibited slowly activating currents that were nearly eliminated by treatment with niflumic acid (10 µM) or removal of extracellular Ca2+. Those currents reversed at -31 and -10 mV when extracellular Cl- concentration was 152 and 34 mM, respectively. In pericytes held at -70 mV, oscillating inward currents were sometimes observed; the reversal potential also shifted with extracellular Cl- concentration. We conclude that ANG II activates a Ca2+-dependent Cl- conductance in OMDVR pericytes to induce membrane depolarization and Psi m oscillations.

medulla; kidney; microcirculation; patch clamp; niflumic acid; oscillations


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

DESCENDING VASA RECTA (DVR) are small resistance vessels that supply blood flow to the medulla of the kidney. Anatomically, DVR arise from juxtamedullary efferent arterioles and traverse the outer medulla sequestered into vascular bundles (21, 34). Those in the bundle center perfuse the inner medulla of the kidney, whereas those on the vascular bundle periphery give rise to the capillary plexus that supplies the outer medullary interbundle region with blood flow. Given that DVR are vasoactive and surrounded by smooth muscle-like pericytes, the parallel arrangement within the bundles suggests that neurohormonal control of their vasomotion serves to regulate both total blood flow and the regional distribution of blood flow within the medulla. Variation of medullary perfusion has been linked to optimization of urinary concentration, salt balance, and control of arterial blood pressure (7).

The inaccessibility of the medulla to experimentation in vivo has hampered detailed investigation of the mechanisms that control DVR pericyte contractile responses. Recently, our laboratory (46) reported the use of microfluorescent and electrophysiological methods to demonstrate that Cl- plays an important role in the control of pericyte membrane potential (Psi m) and constriction by ANG II. To provide a detailed investigation of the temporal variation of Psi m and Cl- conductance, we have now adapted the nystatin perforated-patch technique (14) to access pericytes on isolated outer medullary DVR (OMDVR) for electrophysiological studies. We tested the hypothesis that Cl- dominates pericyte membrane conductance after treatment with ANG II and studied the temporal variation of Psi m and whole cell currents. The results verify that transport of Cl- dominates membrane conductance after ANG II treatment. Furthermore, Psi m and Cl- conductance undergo oscillations, especially at lower ANG II concentrations.


    METHODS
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INTRODUCTION
METHODS
RESULTS
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Vessel isolation. Kidneys were removed from Sprague-Dawley rats (70-150 g; Harlan), sliced into sections along the corticomedullary axis, and stored at 4°C in a physiological saline solution (PSS) containing (in mM) 145 NaCl, 5 KCl, 1 MgSO4, 1 CaCl2, 10 HEPES, and 10 glucose, pH 7.4, at room temperature. Psi m and whole cell currents were monitored by patch-clamp recording from pericyte cell bodies on isolated vessels at room temperature. To accomplish this, OMDVR were digested to remove basement membrane and enable establishment of gigaohm seals between patch pipettes and the cell membrane (46). Small wedges of renal medulla were separated from kidney slices by dissection and transferred to CaCl2-free PSS containing collagenase 1A (0.45 mg/ml, Sigma, St. Louis, MO), protease XIV (0.4 mg/ml, Sigma), and albumin (1 mg/ml). Tissue wedges were incubated at 37°C for 22 min and then transferred back to CaCl2 (1 mM) containing PSS at 4°C for subsequent storage and microdissection. At intervals, vessels were isolated from the digested renal tissue and transferred to a perfusion chamber on the stage of an inverted microscope (Nikon Diaphot). On the microscope stage, OMDVR were captured with a microperfusion-style holding pipette (35), guided into position on a coverslip, and oriented perpendicular to the axis of approach of the patch pipettes. The perfusion chamber was custom machined using a design identical to that of our standard feedback temperature-controlled microperfusion chambers, except that the heating system was eliminated and acrylic was used in place of aluminum for electrical insulation. Two inlets were placed in the chamber. Through one, buffer inflow occurred just upstream of the patch site. Through the other inlet, the Ag-AgCl reference electrode or KCl agar bridge was positioned adjacent to the experimental preparation. All recordings were performed at room temperature. After digestion, pericyte cell bodies on the vessels had a beaded appearance (Fig. 1). The largest cells seen on the side of the vessel facing the patch pipette micromanipulator were generally selected for recording.


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Fig. 1.   Photomicrographs of isolated outer medullary descending vasa recta (OMDVR) captured with differential interference contrast optics. OMDVR were isolated by enzymatic digestion and microdissection. Under these conditions, smooth muscle-like pericyte cell bodies appear as beaded structures protruding from the abluminal surface of the vessel. Patch pipettes of the style used to record membrane potential (Psi m) are also shown. Bar= ~8 µm.

Patch-clamp recording. Patch pipettes were made from borosilicate glass capillaries (PG52151-4; external diameter, 1.5 mm; internal diameter, 1 mm; World Precision Instruments, Sarasota, FL), using a two-stage vertical pipette puller (Narshige PP-830), and subsequently heat polished. For whole cell-permeabilized patch-clamp recording, the pipette solution contained 120 mM potassium aspartate, 20 mM KCl, 10 mM NaCl, 10 mM HEPES, pH 7.2, and nystatin (100 µg/ml with 0.1% DMSO). Nystatin in DMSO was kept frozen at -20°C and renewed weekly. Each day, the nystatin stock was thawed, dispensed into the potassium aspartate pipette solution at 37°C by vigorously vortexing for 1 min, and subsequently protected from light. Under these conditions, nystatin did not completely dissolve, so a saturated or supersaturated solution of uncertain final concentration was actually present in the pipettes. To clear the slight remaining nystatin precipitate, we backfilled the pipettes from a syringe via a 0.2-µm filter. The standard bath solution was PSS (see Vessel isolation). ANG II and pharmacological agents were added as described with individual protocols below. Slight pH correction was needed when anthracene-9-decarboxylate (A9C) and higher concentrations of niflumic acid were used. Psi m was measured using a CV201AU headstage and Axopatch 200A amplifier (Axon Instruments, Foster City, CA) in current clamp mode (I = 0) at a sampling rate of 10 Hz. Psi m was recorded with pipettes of 8- to 15-MOmega resistance.

Whole cell current recording in voltage-clamp mode was accomplished with 5- to 8-MOmega pipettes. Due to the small size of the pericytes, lower resistance pipettes proved technically difficult to use and led to premature loss of seals. Pipettes with nystatin-containing electrode solution were inserted into the bath under positive pressure and positioned near the cell, and the offset of the amplifier was adjusted to null the junction and electrode potentials. The final approach to the cell was controlled with a piezoelectric drive (Burleigh PCS-5000). Gigaseals were established by pressing the pipette tip against the cell and applying light suction. The progress of seal formation was followed on a digital oscilloscope (Hameg M305) by observing the current elicited by test pulses of 5 mV amplitude. Seal formation was facilitated by gradually reducing the holding potential from 0 to -70 mV. After seal formation, the appearance of the cell capacitance transient and the access resistance were monitored using a Digidata analog-to-digital converter and Clampex 7.0 (Axon Instruments, Union City, CA) with 10-mV pulses at a holding potential of -70 mV. Final access resistance was generally between 15 and 40 MOmega . Cellular capacitance and access resistance compensations were applied as appropriate. Whole cell currents were sampled at 2 kHz with filtering at 5 or 10 kHz unless otherwise specified. The reference electrode was an Ag-AgCl wire except when glutamate was substituted for Cl- to lower the bath Cl- concentration (see below).

Corrections for junction potential and Donnan equilibrium effects. The goal of many of these studies was to record Psi m. Uncertainties in those measurements arise from the need to compensate for junction potentials between the bath and electrode (26) and the Donnan equilibrium effects created by the nystatin pore interface between the pipette and cell interior (14). Nystatin produces electrical coupling to the cell interior by establishing pores that conduct monovalent ions with little or no permeation by divalent ions or macromolecules. The presence of immobile ions in the cells establishes a Donnan equilibrium that results in both a transmembrane potential difference and osmolar gradient between the pipette and cell interior. The expected Cl- concentration in the cell can be calculated from the quadratic equation (14)
Cl<SUP>2</SUP><SUB><IT>c</IT></SUB><IT>+</IT>A<SUB>c</SUB>Cl<SUB>c</SUB><IT>−</IT>(Na<SUB>p</SUB><IT>+</IT>K<SUB>p</SUB>) Cl<SUB>p</SUB><IT>=</IT>0 (1)
where Clc and Ac are the concentrations of Cl- and immobile anions in the cell, respectively, and Nap and Kp are sodium and potassium concentrations in the pipette, respectively. In turn, Clp will be reduced and the Donnan effects minimized by the addition of immobile ions to the electrode solution in the pipette. For this purpose, we employed potassium aspartate (120 mM). The transmembrane potential difference between the pipette and cell interior (Psi p - Psi c) is given by
&PSgr;<SUB>p</SUB><IT>−&PSgr;</IT><SUB>c</SUB><IT>=</IT><FENCE><FR><NU><IT>RT</IT></NU><DE><IT>F</IT></DE></FR></FENCE> ln<FENCE><FR><NU>Cl<SUB>p</SUB></NU><DE>Cl<SUB>c</SUB></DE></FR></FENCE> (2)
where R is the gas constant, T is absolute temperature, and F is Faraday's constant. The actual magnitude for Ac cannot be measured. However, using the typical estimate of Ac = 100 mM, we predict that Clc = 34 mM and (Psi p - Psi c) = -3.3 mV, i.e., Psi c is 3.4 mV greater than Psi p (14).

The existence of a liquid junction potential between the pipette and initial bath solution (PSS), nulled before seal formation, also requires correction. Our measurement of this offset, obtained using a flowing 3 M KCl bridge as the bath electrode as described by Neher (26), was -9.7 ± 0.3 mV (n = 4). Thus our true pipette potential is 9.7 mV lower than that specified by the amplifier. Accordingly, we have combined the estimate of the Donnan potential with the liquid junction potential and adjusted the Psi m measurements by subtracting 6.4 mV from the measured value. We recognize that an uncertainty of several millivolts remains due to the need to estimate Ac in the calculation of the Donnan potential.

In some experiments, extracellular Cl- concentration was lowered by substituting NaCl with sodium gluconate. To avoid a large error resulting from a change in the reference electrode/bath interface, we substituted a 3 M KCl, 3% agar bridge for an Ag-AgCl wire as the reference electrode. The low-Cl- buffer was composed of (in mM) 130 sodium gluconate, 25 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose, pH 7.4, at room temperature. This buffer has a final Cl- concentration of 34 mM. After accounting for Donnan effects (see above), this yields a Cl- concentration that is approximately symmetrical across the cell membrane. The predicted offset adjustment resulting from the change in diffusion potential between the 3 M KCl reference electrode and PSS vs. low-Cl- buffer is only -1.4 mV and was neglected.

Experimental protocols. The effect of ANG II and other agents on Psi m was tested by exchanging them into the bath via a manifold from gravity-driven reservoirs. With our chamber design and the bath flow rate employed, exchange of the bath occurs in ~30 s. In an initial series, after baseline recording for 2 min, ANG II (10-8 M) was added for 1 or 5 min and then removed to assess the reversibility of its effects. In separate time controls, Psi m was recorded for 20 min without ANG II, after which ANG II was added to the bath to verify responsiveness. The concentration dependence of the effect of ANG II on Psi m was determined by exchanging ANG II into the bath in log molar increments from 10-11 to 10-7 M at 4-min intervals.

The ability of the Cl- channel blocker niflumic acid to reverse ANG II (10-8 M)-dependent depolarization was examined by exchanging it into the bath for 5 min and then removing it. Multiple concentrations of niflumic acid were tested (1, 10, 25, 50, and 100 µM) in random order. Seals could not be maintained long enough to test all concentrations in each vessel. The effect of niflumic acid on Psi m in the absence of ANG II was examined in a separate series of experiments at 10 and 100 µM. The ability of the Cl- channel blocker A9C (1 mM) to affect Psi m in control and ANG II (10-8 M)-treated vessels was examined using a similar protocol.

Whole cell currents were measured in pericytes that had been exposed to ANG II (10-8 M) for 20 min. The effect of niflumic acid (10 µM) on whole cell Cl- currents was measured with a voltage-clamp, pulse-step protocol executed before and after exposure to niflumic acid. In a separate series of experiments, we examined the ability of changing bath Cl- concentration to shift the reversal potential of the currents. Pericytes were conditioned by depolarizing them from a holding potential of -70 to -25, -10, or 0 mV for 1 s followed by a shift to potentials of -60 to 30 in 10-mV increments for 500 ms. Recordings were obtained in PSS and low-Cl- extracellular solution in random order. Finally, in some ANG II (10-8 M)-treated cells that were undergoing rapid oscillations, whole cell current was monitored at a holding potential of -70 mV and then stepped from -30 through 0 mV in 10-mV increments. This was repeated in both PSS and low-Cl- buffer to determine the effect of Cl- concentration on the direction of the oscillating current. To deplete pericytes of Ca2+, we switched the extracellular buffer from PSS to PSS with nominally zero CaCl2 containing 1 mM EGTA.

Reagents. ANG II, BSA (A2153, Cohn fraction V), nystatin, collagenase 1A, protease XIV, niflumic acid, and A9C were from Sigma. ANG II in water was stored in 200-µl aliquots at -20°C and diluted on the day of an experiment. The Cl- channel blockers niflumic acid and A9C were also stored in aliquots in DMSO and diluted on the day of experimentation. Reagents were frozen and thawed once only. Excess reagents were discarded at the end of each day.

Statistics. Except where otherwise specified, data are given as means ± SE. The significance of differences between means was calculated using Student's t-test (paired or unpaired, as appropriate) and repeated measures ANOVA. In some figures, data sampled at 10 Hz was averaged 10 values at a time for display at 1 Hz. Where specified, most error bars were suppressed to optimize display of the data.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Effect of ANG II on pericyte Psi m. Without exception, all pericytes depolarized when exposed to ANG II. Addition of ANG II (10-8 M) to the extracellular buffer for 8 min (Fig. 2) resulted in depolarization that did not reverse after its removal. In contrast, when ANG II was introduced into the bath for only 1 min, its effects were slowly reversible. Psi m in time controls subjected to sham exchange at 2 min was stable over a subsequent 18-min period. When those cells were subsequently exposed to ANG II, rapid depolarization occurred, verifying that prolonged nystatin patch formation does not interfere with responsiveness.


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Fig. 2.   Depolarization of pericytes by ANG II. Recordings of pericyte Psi m measured using I = 0 current clamp are shown for 3 groups of pericytes (n = 6 each). In 1 group, after 2 min, ANG II (10-8 M) was added to the extracellular buffer for 8 min and then removed. In a second group, ANG II was added for only 1 min and then removed. A group of time controls was sham exchanged at 2 min and exposed to ANG II from 20 to 25 min. Psi m was sampled at 10 Hz and averaged to 1 Hz for display; 1 cell/vessel was studied. Most error bars were suppressed for clarity.

Psi m oscillations. The data points of the records in Fig. 2 were reduced by sampling at 10 Hz and averaging 10 data points/s to yield one value per second. The mean ± SE of Psi m for all experiments was then averaged for display. This facilitates illustration of the overall effects of ANG II on Psi m and hides the complex time-dependent variation of Psi m observed in some records. Large Psi m oscillations occurred in a few cells (excluded from the averaging in Fig. 2). The data from those experiments are shown separately in Fig. 3. Both sustained, regular oscillations and a rapid spiking pattern of Psi m variation occurred (Fig. 3).


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Fig. 3.   Psi m oscillations induced by ANG II. Examples of sustained (A) and spiking oscillations (B) of Psi m are shown. In both records, ANG II (10-8 M) was added to the extracellular solution at 1 min and maintained for the duration of the recording.

ANG II concentration dependence. The ability of ANG II to depolarize the pericytes over a concentration range (10-11 to 10-7 M) was examined by exchanging ANG II into the bath in log molar increments at 4-min intervals (Fig. 4). As shown in Fig. 4, at lower concentrations ANG II consistently elicited oscillations when added to the extracellular buffer. The oscillations tended to be suppressed when ANG II concentration was raised to 10-8 and 10-7 M. Again, similar to the results illustrated in Fig. 2, reversal of depolarization did not occur during the final period after ANG II was removed from the bath.


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Fig. 4.   Concentration dependence of ANG II-induced depolarization. A and B: individual recordings of Psi m are shown in which ANG II was introduced into the bath in sequential log-molar increments at 4-min intervals.

Effect of Cl- channel blockade. With the use of a voltage-sensitive dye loaded into pericytes and endothelia, our laboratory (46) previously reported that Cl- channel blockade prevented ANG II-induced depolarization and reversed vasoconstriction. In the present study, we used the nystatin perforated-patch technique to examine this in single pericytes. The Cl- channel blocker niflumic acid modulated ANG II (10-8 M)-induced depolarization in a biphasic, concentration-dependent manner (Fig. 5). Between 1 and 50 µM, repolarization occurred. At 10 µM, niflumic acid repolarized cells to values below the resting potential before ANG II treatment. However, niflumic acid at 100 µM augmented ANG II-induced depolarization. The effects of niflumic acid on Psi m were rapidly reversible so that multiple concentrations could be tested per vessel in random order. Similar concentration-dependent effects on Psi m were also observed in pericytes that had not been pretreated with ANG II (Fig. 6). Like niflumic acid, the Cl- channel blocker A9C (1 mM) reversed ANG II-induced depolarization and hyperpolarized untreated cells (Fig. 7). A9C was less effective than niflumic acid (10 µM) in repolarizing the cells, and its effects on Psi m reversed more slowly after it was removed from the bath (compare with Figs. 5 and 6).


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Fig. 5.   Effect of the Cl- channel blocker niflumic acid (Nif A) on ANG II-depolarized pericytes. In each panel, the mean ± SE () of the resting Psi m is shown for the group of pericytes tested at the niflumic acid concentration specified (above the solid bar). After ANG II-induced depolarization, the effect of niflumic acid on Psi m was tested by adding it to the extracellular buffer for 5 min. Multiple, but not all, concentrations of niflumic acid were tested in each vessel, because seals could not be sustained for a long enough time period. The order of testing of niflumic acid at the various concentrations was randomized. Niflumic acid repolarized the cells between 1 and 50 µM but augmented depolarization at 100 µM. For display, data were reduced from 10 to 1 Hz by averaging, and most error bars were suppressed for clarity. n = 6-8/group.



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Fig. 6.   Effect of the Cl- channel blocker niflumic acid on untreated pericytes. Psi m was recorded for 1 min, after which niflumic acid was added to and then removed from the extracellular buffer at either 10 (n = 7; left) or 100 µM (n = 5; right) at 5-min intervals. Data were reduced from 10 to 1 Hz by averaging, and most error bars were suppressed for clarity.



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Fig. 7.   Effect of the Cl- channel blocker anthracene-9-decarboxylate (A9C) on pericyte Psi m. Left: the effect of A9C (1 mM) on Psi m was measured in untreated pericytes (n = 5). Right: the ability of A9C to repolarize pericytes pretreated with ANG II (10-8 M) was tested (n = 5). Mean ± SE of the resting potential is indicated by  and error bar. After ANG II depolarization, A9C was added to the extracellular buffer and then removed. Data were reduced from 10 to 1 Hz by averaging, and most error bars were suppressed for clarity.

Whole cell current measurements. In voltage-clamp experiments, depolarization of pericytes from a holding potential of -70 mV to potentials greater than -30 mV induced an outward current that activated with time. Currents observed after performing leak subtraction are illustrated in Fig. 8, A and C. After exposure to 10 µM niflumic acid, the currents were nearly eliminated (Fig. 8, B and C), supporting the notion that the charge carrier is Cl-. The currents elicited by the test pulses in Fig. 8A show activation with time after depolarization. Similarly, the tail currents that occurred on repolarization to the holding potential show deactivation with time.


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Fig. 8.   Effect of niflumic acid on whole cell currents in ANG II-treated pericytes. A and B: whole cell currents (Im) were measured with a voltage-clamp protocol in nystatin patched pericytes exposed to ANG II (10-8 M) in the extracellular buffer for 20 min before recording. The command potential was stepped from a holding level of -70 mV to values ranging from -80 to 50 mV (10-mV increments) with 10 s between steps. Examples of data obtained in the absence or presence of niflumic acid (10 µM) are shown in A and B, respectively. C: Im 10 ms before the end of the pulse was averaged for display (n = 6; * P < 0.05, ANG II vs. ANG II + niflumic acid; # P < 0.05, ANG II vs. ANG II + niflumic acid for tail currents). Junction potential corrections have been applied to the values on the abscissa.

To further establish a role for Cl-, we examined the reversal potential by varying the holding potential of cells bathed in both PSS and low-Cl- buffer. To accomplish this, cells were conditioned to activate the Cl- conductance by first stepping them from -70 to -25 (n = 1), -10 (n = 5), or 0 mV (n = 2) for 1,000 ms. In all cases, cells were then changed to a different holding potential for 500 ms. The second step was varied between -60 and 30 mV in 10-mV increments, and a 10-s recovery time was allowed between steps. An example of the current generated by this protocol is shown in Fig. 9A for a cell conditioned at 0 mV. The cells depolarized to -25, -10, or 0 mV behaved similarly, and the results are summarized in Fig. 9C as the means ± SE of the current present 10 ms after the conditioning pulse (Fig. 9A, point a). After applying liquid junction potential corrections, the change from PSS to symmetrical Cl- shifted the reversal potential from -31.5 to -10.1 mV. The direction of that shift is consistent with Cl- as the charge carrier but the values did not conform to the theoretical equilibrium potential for Cl- in PSS and symmetrical Cl- buffer of -39 and 0 mV, respectively. This suggests incomplete equilibration of the cell interior with the electrode solution (see DISCUSSION).


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Fig. 9.   Reversal of whole cell currents after ANG II treatment. A and B: whole cell current (A) was measured in pericytes previously exposed to ANG II (10-8 M) for 20 min. The cells were preconditioned for 1,000 ms by depolarizing them from a holding level of -70 mV to either -25 (n = 1), -10 (n = 5), or 0 mV (n = 2). In each cell, after the conditioning pulse, the command potential was stepped to values ranging from -60 to 30 mV (10-mV increments), according to the protocol depicted in B. To determine the shift of the reversal potential with variation of extracellular Cl- concentration, the protocol was executed in physiological saline solution (PSS) and low-Cl- buffer. C: mean ± SE of the current present 10 ms after the second step (point a in A) is shown. Currents reversed at -31.5 (open circle ) and -10.1 mV () in PSS and low-Cl- buffer, respectively. Junction potential corrections have been applied to correct the values on the abscissa.

Due to rapid time-dependent variation of membrane conductance, cells that were undergoing oscillations produced erratic results in voltage-clamp, pulse-step protocols like those shown in Figs. 8 and 9. As shown in Fig. 10, A and B, ANG II (10-8 M) induced spontaneous inward currents in some cells. To examine this, we recorded currents in oscillating cells as command potential was stepped sequentially between -30 and 0 mV in 10-mV increments with either PSS or low-Cl- as the extracellular buffer. Command potential changes were manually performed after 5 to 10 oscillations had been recorded at -30, -20, -10, and 0 mV. An example, typical of n = 4 observations, is illustrated in Fig. 10C. The mean ± SE (n = 4 cells) of the peaks of the current spikes is summarized as a function of Psi m, corrected for liquid-junction potential, in Fig. 10D. In PSS, the direction of the current spikes reversed at -40.3 mV, whereas in symmetrical Cl-, reversal occurred at -11.3 mV, supporting the conclusion that the oscillating current is attributable to variation of Cl- conductance.


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Fig. 10.   Spontaneous inward currents after ANG II. A and B: 2 examples of spontaneous inward currents in pericytes held at -70 mV are shown. In both recordings, ANG II (10-8 M) was added to the bath at 1 min (*). Currents were sampled at 10 Hz. C: leak subtracted spontaneous currents are shown in a pericyte bathed in PSS (trace a) and then low-Cl- buffer (trace b). In each buffer, the command potential (Vm) was manually varied between 0 and -30 mV after several peaks had been recorded. D: the mean ± SE (n = 4) of current peaks is shown as a function of Psi m, corrected for liquid junctions. The mean peak currents reversed at -40.3 and -11.3 mV in PSS and low-Cl- buffer, respectively. Data were sampled at 10 Hz.

Recently, our laboratory (46) showed that a 16.8-pS Ca2+-activated Cl- channel exists in OMDVR pericyte cell-attached patches. To demonstrate the Ca2+ dependence of whole cell Cl- currents, we examined the ability of Ca2+ depletion to suppress the post-ANG II Cl- currents. Pulse protocols were executed in PSS, after 5 min of exposure to PSS with nominally zero CaCl2 containing 1 mM EGTA, and after restoration of CaCl2 to the bath during a recovery period. Currents were markedly suppressed by removal of extracellular Ca2+ (Fig. 11, A and B). The results of these experiments (n = 6) are summarized in Fig. 11C in which the means ± SE of currents 10 ms before the end of the pulse are displayed for the control, Ca2+-depletion, and recovery periods.


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Fig. 11.   Effect of Ca2+ removal on whole cell currents in ANG II-treated pericytes. A and B: Im were measured with a voltage-clamp protocol in nystatin patched pericytes exposed to ANG II (10-8 M) in the extracellular buffer for 20 min before recording. The command potential was stepped from a holding level of -70 mV to values ranging from -80 to 50 mV (10-mV increments) with 10 s between steps. Examples of data obtained before and after removal of extracellular Ca2+ are shown in A and B, respectively. C: Im 10 ms before the end of the pulse was averaged for display in C (n = 6; * P < 0.05, PSS vs. 0 CaCl2; # P < 0.05, PSS/recovery period vs. 0 CaCl2 ). Junction potential corrections have been applied to the values on the abscissa.


    DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
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DISCUSSION
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The microcirculation of the kidney is anatomically organized to accommodate the differing requirements of the renal cortex and outer and inner medulla. DVR occupy a unique niche in this scheme because they are the final small generation resistance vessels that supply medullary blood flow. Historically, DVR have been viewed as one limb of a passive countercurrent exchanger, but growing evidence indicates far more complex behavior. In addition to expressing specific endothelial transporters for urea (35, 45) and water (27, 30), they are surrounded by smooth muscle-like pericytes that impart contractile function (32). Studies (29, 34, 40) involving application of vasoactive agents have proven that OMDVR express receptors for both constrictors and dilators. In the outer medulla, OMDVR are radially distributed within vascular bundles so that those in the center perfuse the inner medulla and those on the periphery peel off to supply the outer medullary interbundle region with blood flow. On the basis of these features, it seems likely that OMDVR vasomotor tone is governed by a number of paracrine stimuli that serve to regulate total and regional distribution of blood flow in the medulla.

Despite the key role that the pericyte must play in the regulation of water reabsorption by the kidney and the growing evidence (7, 25) that regional perfusion of the medulla regulates salt balance and blood pressure, little is known of the mechanisms that govern pericyte contraction. This is due to the inaccessibility of the outer medulla in vivo, the lack of cell culture models, and the difficulty of isolating these 10- to 15-µm vessels. In a recent study (46), our laboratory demonstrated the feasibility of using fluorescence microscopy and electrophysiological recording to gain insight into the channel architecture of the OMDVR pericyte. A 16.8-pS Cl- channel that activates on exposure to ANG II or excision into high-Ca2+ buffer was identified. Vasomotion studies (46) verified a pivotal role for Cl- in ANG II-induced vasoconstriction. In the present study, we have shown that the nystatin perforated-patch technique can be used productively to gain electrical access to the pericyte cytoplasm for whole cell recording. This allowed us to examine the temporal variation of Psi m (Figs. 2-6), establish its dependence on ANG II concentration (Fig. 4), and verify the role of Cl- in its determination (Figs. 5-7). Voltage-clamp protocols revealed dramatically that Cl- conductance dominates membrane currents after prolonged ANG II treatment (Figs. 8-10) and that the Cl- current is Ca2+ dependent (Fig. 11).

In all of the measurements taken in the present study, pericytes depolarized when ANG II was added to the extracellular buffer (Figs. 2-7). After ANG II exposure, the dominant current identified in whole cell recordings was carried by Cl- so that a shift in Psi m away from the Nernst potential for K+ to that for Cl- is likely to be the mechanism responsible for depolarization. Evidence of the dominant role of Cl- was provided by repolarization on blockade of Cl- channels (Figs. 5-7), the marked reduction of whole cell currents by Cl- channel blockade, and the shift in the reversal potential with variation of extracellular Cl- concentration (Figs. 9 and 10). The reversal potentials observed for steady state (Fig. 9) and oscillating peak currents (Fig. 10) did not precisely conform to the theoretical equilibrium potentials for Cl- in PSS and low-Cl- buffer. For the reversal potential to be equal to the equilibrium potential for an ion, the conductance pathway has to be perfectly selective. Calculation of the expected reversal potential also assumes knowledge of the intracellular Cl- concentration. The latter demands perfect equilibration between the pipette and cell. In the reversal experiments, we shifted the extracellular buffer from 154 mM Cl- (PSS) to 34 mM Cl- (low-Cl- buffer). If the cell had been completely dialyzed, after accounting for Donnan equilibrium, the pipette solution should have forced intracellular Cl- to be ~34 mM and the equilibrium potential for Cl- would be -39 and 0 mV, respectively. After correction for junction potentials, the observed reversals were at -31.3 and -10.1 mV, respectively (Fig. 9), and -40 and -11 mV, respectively (Fig. 10). Thus the direction of the shift was as expected for Cl- flux but the magnitude of the shift was less than expected. These data are not readily explained by contaminating current from another ion. For example, if a K+ current contributed to the observations, the reversals in PSS and low-Cl- buffer should have differed from predicted levels (-39 and 0 mV) by more negative values. If Na+ or Ca2+ currents were present, both reversal potentials should have differed by more positive values. Instead, the reversal potential in PSS was greater than -39 mV and the reversal in low-Cl- buffer was less than 0 mV. This finding is likely to be explained by incomplete equilibration of the cell interior with the pipette, in which case changes in extracellular Cl- concentration could induce parallel changes in intracellular concentration.

Given that OMDVR are branches of efferent arterioles, Cl--mediated depolarization is a somewhat surprising finding because efferent arteriolar constriction is not dependent on depolarization or affected by Cl- channel blockade (5, 6, 22, 23). Depolarization through activation of Cl- conductance is common in the afferent microcirculation of the kidney (5, 10, 12, 16, 43, 44) and extrarenal microvascular beds (1, 18, 28) in which it serves to activate voltage-dependent Ca2+ entry pathways. Rigorous examination of smooth muscle isolated from larger resistance vessels of the renal cortex identified T- and L-type voltage-gated Ca2+ entry pathways and endothelin-stimulated currents carried by Cl- (10). Given the diverse control mechanisms required by the kidney, including tubuloglomerular feedback, myogenic autoregulation, the renin angiotensin axis, and the need to separately regulate medullary vs. cortical blood flow, regionally specific channel architecture of smooth muscle is to be expected.

Particularly at submaximal concentrations, ANG II-induced Psi m oscillations in OMDVR pericytes were common (Figs. 3 and 4). Oscillations of Psi m have been recognized in smooth muscle cells and may be linked to oscillations of intracellular Ca2+ concentration ([Ca2+]i). Janssen and Daniel (15) showed that carbachol elicited depolarization and Psi m oscillations in bronchial smooth muscle. The oscillations were inhibited by nitrendipine or replacement of extracellular Ca2+ with Sr2+ (15). Sieck and colleagues (17, 39) found that [Ca2+]i oscillations in porcine tracheal smooth muscle elicited by ACh involved uptake and release from ryanodine-dependent stores. Similarly, ANG II-induced [Ca2+]i oscillations in pulmonary artery myocytes were found (11) to be dependent on repetitive release of Ca2+ from inositol trisphosphate-dependent Ca2+ stores. Synchronous oscillations of Psi m and [Ca2+]i were demonstrated by Kohda et al. (19) in ileal smooth muscle. Kohda and co-workers (19) concluded that the oscillations were dependent on cation channel openings and repetitive release of Ca2+ from internal stores. Our laboratory (46) previously identified a Ca2+-dependent Cl- channel in OMDVR pericytes. If that channel is important in the determination of Psi m after ANG II treatment, oscillations of Ca2+ would be expected to stimulate repetitive cycles of openings and closings to explain the oscillations of Cl- conductance and Psi m shown in Figs. 3, 4, and 10. The existence of Ca2+ oscillations in OMDVR pericytes remains to be examined, and our laboratory's (33, 35) past attempts to measure pericyte Ca2+ transients with fura-2 have been frustrated because this indicator loads into the pericyte cytoplasm very poorly.

It is generally accepted that resting cell Psi m is held at negative values near the Nernst potential for K+. Depolarization results when conductances to other cations are enhanced, especially in excitable cells, or to Cl- in other cell types. Thus inhibition of K+ channels also favors depolarization and activation of K+ channels could limit depolarization and vasoconstriction (8). ANG II has been found (9, 41) to activate K+ conductance in mesangial cells and the main renal artery. In contrast, K+ channel inhibition mediated by 20-hydroxyeicosatetraenoic acid occurs after ANG II stimulation of smaller renal arterioles (2, 24, 42) and the cerebral circulation (13). In the present study, K+ channel blockers were not used in either the pipette or extracellular buffer. Despite this, the Cl- channel blocker niflumic acid (10 µM) nearly eliminated membrane currents after 20 min of ANG II exposure (Fig. 8). Therefore, it seems likely that ANG II inhibits K+ channels in OMDVR pericytes, but this remains to be proven. The finding that niflumic acid, when applied at a concentration of 100 µM, favors depolarization (Figs. 5 and 6) is consistent with previous findings (18) that this concentration can block Ca2+-dependent K+ channels. Definition of the classes of K+ channels that exist in pericytes and mechanisms that determine their regulation requires further study.

The ANG II-dependent Cl- current was found to be voltage dependent, activated by depolarization, and inactivated by repolarization of the pericytes (Figs. 8 and 9). Time- and voltage-dependent activation of Cl- currents has been frequently described in smooth muscle (1, 18, 28) and a variety of other cell types, including secretory epithelia (3), pancreatic B cells (20), and Xenopus laevis oocytes (4). When K+ channels are pharmacologically blocked to isolate the Cl- current, a depolarizing pulse applied to a smooth muscle cell often yields a transient inward current followed by a slowly activating outward current (1, 3, 4, 18, 20, 28). The inward current is attributed to Ca2+ influx through voltage-gated channels and the outward current to Cl- influx through a pathway activated by elevation of [Ca2+]i [ICl(Ca)]. Our protocols do not substantially differ from those used by other investigators; however, we failed to identify the early transient inward current in any of the traces leading to the results shown in Figs. 8 and 11. It seems likely that Ca2+ influx occurs after pericyte depolarization but that the current generated cannot be resolved because it is small. A less likely explanation is that voltage-gated Ca2+ channels do not exist in OMDVR pericytes. In the latter case, the mechanism of activation of the Cl- current would be unexplained and the general purpose of depolarization would be enigmatic. Verification of the nature of the Ca2+ dependence of ICl in these cells and definition of the Ca2+ influx pathway require further study.

In these experiments, solute concentrations in the bath and patch pipette govern extracellular and intracellular concentrations, respectively. Those ion concentrations may not simulate values present in vivo. The situation in the vascular bundles is complex because pericytes lie at various levels along the corticomedullary gradient where ion concentrations can vary widely. In addition, the pericytes surround DVR within a few micrometers of adjacent endothelia. OMDVR endothelia strongly express the aquaporin-1 water channel and conduct solute-free transcellular water flux from the OMDVR lumen toward the abluminal pericyte (30, 37). Most likely, a diffusional flux directed toward the pericyte establishes standing solute gradients in the vicinity of the OMDVR wall. As a consequence, the absolute magnitude of the local ion concentrations to which pericytes are exposed cannot be easily predicted. Pericytes surround DVR in the inner medulla as well as the outer medulla (38). In the concentrating rat, Na+ at the papillary tip can exceed 500 mM and K+ concentration in the extracellular fluid exceeds that of peripheral plasma manyfold (31, 37). The transcellular gradients for K+ and Cl+ and associated Nernst potentials governing pericytes in the inner medulla are therefore very uncertain. It seems possible that pericyte channel expression and regulation might vary along the corticomedullary axis and that these studies, performed in OMDVR, might not reflect the behavior of pericytes derived from inner medullary DVR.


    ACKNOWLEDGEMENTS

This study was supported by National Institutes of Health Grants DK-42495, HL-62220, and HL-68686.


    FOOTNOTES

Address for reprint requests and other correspondence: T. L. Pallone, Division of Nephrology, N3W143, Univ. of Maryland, Baltimore, MD 21201-1595 (E-mail: tpallone{at}medicine.umaryland.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.

First published January 29, 2002;10.1152/ajprenal.00306.2001

Received 8 September 2001; accepted in final form 31 December 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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