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
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ABSTRACT |
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 (
m) by ANG II. ANG II
(10
11 to 10
7 M) consistently depolarized
OMDVR and induced
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
m oscillations.
medulla; kidney; microcirculation; patch clamp; niflumic acid; oscillations
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INTRODUCTION |
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 (
m)
and constriction by ANG II. To provide a detailed investigation of the
temporal variation of
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
m and
whole cell currents. The results verify that transport of
Cl
dominates membrane conductance after ANG II treatment.
Furthermore,
m and Cl
conductance undergo
oscillations, especially at lower ANG II concentrations.
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METHODS |
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.
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
( m) are also shown. Bar= ~8 µm.
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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.
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.
m was recorded with pipettes of 8- to 15-M
resistance.
Whole cell current recording in voltage-clamp mode was accomplished
with 5- to 8-M
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 M
. 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
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)
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(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 (
p
c) is given by
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(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 (
p
c) =
3.3 mV, i.e.,
c is 3.4 mV
greater than
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
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
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,
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
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
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
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.
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RESULTS |
Effect of ANG II on pericyte
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.
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 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.
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.
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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
m for all experiments was then averaged for
display. This facilitates illustration of the overall effects of ANG II
on
m and hides the complex time-dependent variation of
m observed
in some records. Large
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
m variation occurred (Fig. 3).

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Fig. 3.
m oscillations induced by ANG II. Examples of sustained
(A) and spiking oscillations (B) of 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.
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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 m are shown
in which ANG II was introduced into the bath in sequential log-molar
increments at 4-min intervals.
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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
m were
rapidly reversible so that multiple concentrations could be tested per
vessel in random order. Similar concentration-dependent effects on
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
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 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 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. 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 m. Left: the
effect of A9C (1 mM) on 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.
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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.
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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 ( ) and 10.1 mV ( )
in PSS and low-Cl buffer, respectively. Junction
potential corrections have been applied to correct the values on the
abscissa.
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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
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 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.
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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.
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DISCUSSION |
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
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
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
m
oscillations in OMDVR pericytes were common (Figs. 3 and 4). Oscillations of
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
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
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
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
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
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.
 |
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