Vasopressin Increases Intracellular NO Concentration via
Ca2ⴙ Signaling in Inner Medullary Collecting Duct
Takefumi Mori, Jeffrey G. Dickhout, Allen W. Cowley, Jr
Downloaded from http://hyper.ahajournals.org/ by guest on July 31, 2017
Abstract—The present study was designed to determine whether arginine vasopressin (AVP) stimulates NO production in
the epithelial collecting duct cells of the inner medulla (IMCDs) and if this is mediated through Ca2⫹ signaling. Thin
tissue layers containing IMCDs were dissected from Sprague-Dawley rats. Intracellular Ca2⫹ concentration ([Ca2⫹]i) and
NO production were measured in IMCDs by a fluorescence imaging system with the use of fura 2-AM and the
cell-permeable form of the NO-sensitive dye 4,5-diaminofluorescein (DAF-2), respectively. AVP (100 nmol/L)
produced a rapid peak increase in [Ca2⫹]i of 320⫾70 nmol/L within a few seconds and a sustained increase of 120⫾62
nmol/L. The peak increase in [Ca2⫹]i was followed by a significant increase of NO production (34⫾7 U). This was
similar to that produced by 20 ␮mol/L of the NO donor DETA-NONOate (42⫾11 U). The NO scavenger carboxy-PTIO
(100 ␮mol/L) or depletion of [Ca2⫹]i by preincubation with 5 ␮mol/L of the Ca2⫹-ATPase inhibitor thapsigargin in
Ca2⫹-free buffer abolished the NO response to AVP. We conclude that AVP mobilizes Ca2⫹ to produce NO in IMCDs.
(Hypertension. 2002;39[part 2]:465-469.)
Key Words: vasopressins 䡲 nitric oxide 䡲 calcium 䡲 kidney 䡲 signal transduction
S
whether AVP can stimulate NO production in IMCDs and to
clarify whether this is a Ca2⫹-dependent pathway. Kojima et
al8 recently established the use of the NO-sensitive dye
DAF-2, and the cell-permeable form of this dye (DAF-2DA)
has allowed NO detection in individual cells. Therefore, an
imaging technique applied to these renal medullary tissue
slices that enables the application of the NO-sensitive dye
DAF-2DA and the Ca2⫹-sensitive dye fura 2-AM has been
developed. The present study reports the temporal relationship of changes in intracellular Ca2⫹ and NO production that
are due to AVP stimulation in IMCDs.
everal observations in our laboratory have suggested
indirectly that arginine vasopressin (AVP) may stimulate
the production of NO in the epithelial collecting duct cells of
the inner medulla (IMCDs). Using an oxyhemoglobintrapping microdialysis technique,1 we have found that renal
medullary interstitial administration of AVP increases NO in
the renal medulla. NO synthase (NOS), measured by
L-arginine/L-citrulline conversion, was also observed to be
highest in IMCDs, as determined from microdissected tubular
and vascular segments.2 In addition, we have reported that
AVP stimulates medullary NO production only through
vasopressin V2–like receptors1 and that the mRNA for V2
receptors was present only in tubules such as medullary thick
ascending limbs and collecting ducts3 but not in the renal
vasculature or in the vasa recta. On the basis of these
observations, we have proposed that IMCDs may play the
largest part in the renal medullary production of NO in
response to AVP.
The well-recognized classical pathway transducing the
effects of AVP on IMCDs is through V2 receptor activation of
adenylyl cyclase, resulting in increased cAMP4 and an
increase in the hydraulic conductivity of IMCDs via the
aquaporin shuttle system.5 AVP has also been shown to
increase intracellular Ca2⫹ in IMCDs possibly via V2 receptors.6 However, the effects of this release of Ca2⫹ have been
largely unexplored, and the effect of AVP stimulation on NO
production in IMCDs is unknown.
Constitutive NOS is known to be modulated by intracellular Ca2⫹,7 and the aim of the present study was to determine
Methods
Experimental Animals
The present study used male Sprague-Dawley rats weighing 250 to
350 g (Harlan Inc, Madison, Wis) maintained ad libitum on water
and a standard pellet diet (Purina Mills) in the Animal Resource
Center of the Medical College of Wisconsin.
Preparation of Inner Medullary Collecting Duct
Rats were anesthetized with pentobarbital (50 mg/kg IP), and the
aorta distal to the origin of the left renal artery was cannulated with
PE-50 tubing and ligated above the left renal artery. The left kidney
was perfused with 10 mL Hanks’ balanced salt solution (HBSS, Life
Technologies) with 20 mmol/L HEPES buffer (Sigma Chemical Co)
and 1 mg/mL BSA (Sigma) at 3 mL/min. The renal pedical was
ligated, and the kidneys were excised and cut sagittally for the
removal of the inner medulla. The inner medulla was then transferred
into a Petri dish filled with ice-cold HBSS with HEPES buffer and
1 mg/mL BSA. The Petri dish was mounted on a cooling microscope
Received September 23, 2001; first decision October 30, 2001; revision accepted November 12, 2001.
From the Department of Physiology, Medical College of Wisconsin, Milwaukee.
Correspondence to Takefumi Mori, MD, PhD, Department of Physiology, Medical College of Wisconsin, 8701 Watertown Plank Rd, Milwaukee, WI
53226. E-mail [email protected]
© 2002 American Heart Association, Inc.
Hypertension is available at http://www.hypertensionaha.org
465
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Hypertension
February 2002 Part II
stage and maintained at 4°C during dissection. Microdissection was
performed under a Leica M3Z stereomicroscope. Single layers of
collecting ducts were dissected from the inner medulla by stripping
along the collecting ducts with the use of 30-gauge needles.
Approximately 200⫻500-␮m–sized tissue strips containing 2 or 3
collecting ducts were cut out with the tip of the 30-gauge fine needle.
Care was taken not to collapse the lumen. These tissues strips
contained segments of the inner medullary collecting duct, vasa
recta, and thin limbs of Henle. The inner medullary collecting duct
was readily distinguishable from the other segments by its significantly larger diameter and rough lumen appearance. Dissected
tissues were placed on round glass coverslips coated with the tissue
adhesive Cell-tak (BD Biosciences) for fluorescent imaging. Dissection was performed within 30 minutes after the removal of the
kidney. The experimental buffer was HBSS with 20 mmol/L HEPES
and 1 mmol/L L-arginine (Sigma), adjusted to pH 7.4.
NO Measurement in IMCDs
Fluorescence measurements were made by using a Nikon Diaphot
inverted microscope with a ⫻25 (numerical aperture 0.65) objective.
The signal was detected by using a CCD video camera (Hamamatsu
Co) coupled to a Gen II image intensifier (Hamamatsu Co), and
excitation was provided by a Sutter DG-4 175-W xenon arc lamp
(Inovision Co), which allowed high-speed excitation wavelength
switching. For the experiments, coverslips were placed in an imaging
chamber (maintained at 37°C) mounted on the stage of the inverted
microscope, which allowed the superfusion of the experimental
buffer and buffer containing the agonist. Regions, ⬇50 ␮m in length,
were selected along the inner medulla collecting duct to quantify
changes in fluorescence intensity by using MetaFluor imaging
software (Universal Imaging Co).
NO production in the IMCDs was measured by using the NOsensitive dye DAF-2DA, as established by Kojima et al.8 DAF-2DA
is a cell-permeable molecule and is hydrolyzed to an impermeable
nonfluorescent active form, DAF-2, in cells by the action of
esterases. On reaction with NO, DAF-2 is then converted to a
cell-impermeable fluorescent form, DAF-2T, whose presence can be
measured as a specific increase in fluorescence signal.8 IMCDs were
incubated on coverslips in 10 ␮mol/L of the NO-specific dye
DAF-2DA (Calbiochem-Novabiochem Co) for 60 minutes at room
temperature and then washed and incubated for another 15 minutes
in the experimental buffer. DAF-2T was excited at 480 nm and
collected through a 535/40-nm bandpass emission filter at 3.5second intervals. Preliminary data showed that the excitation of the
dye in this condition did not bleach the DAF-2T fluorescence during
the course of the experiment. To compare the intracellular NO
production between IMCDs during the experiment, the change in
DAF-2T fluorescence was measured continuously for 500 seconds
after exchanging the bath solution with the agonist vehicle, 100
nmol/L AVP, 20 ␮mol/L of the NO donor DETA-NONOate (Cayman Chemical Co), and 5 ␮mol/L of the Ca2⫹ ionophore 4-bromoA23187. The agonists remained in the bathing solution during the
500-second time course of fluorescence measurement. To confirm
DAF-2 NO detection specificity, all experiments were also performed with 100 ␮mol/L of the NO scavenger carboxy-PTIO
(Dojindo Molecular Technologies Inc) added to the preincubation
buffer for 15 minutes after dye loading but before agonist stimulation
and also added to the agonist-containing experimental buffers. NO
measurements were expressed as relative units of fluorescence
intensity under constant conditions of microscopic signal acquisition
between experiments.
Intracellular Ca2ⴙ Measurement in IMCDs
NO Detection With Intracellular Ca2ⴙ Depletion
Fluorescence Detection
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The IMCDs isolated on coverslips were incubated in 5 ␮mol/L fura
2-AM (Molecular Probes Inc) for 30 minutes at room temperature
and then washed to remove excess dye. Pluronic F127 (Molecular
Probes Inc) was used to dissolve the fura 2-AM dye to prevent dye
compartmentalization on loading. The coverslips were again incubated in the experimental buffer for 15 minutes before the experiments. Fura 2 signal fluorescence was stimulated by dualwavelength excitation at 340 nm and 380 nm. A 510/40-nm bandpass
emission filter was used to collect fura 2 signals at 0.25-second
intervals. Ratios between the fluorescence intensity stimulated by
340/380-nm excitation were calculated, and the excitation intensity
was adjusted on the DG-4 xenon arc lamp to prevent fura 2
fluorescence bleaching and to balance 340/380-nm excitation
intensities.
Fluorescent signals were recorded in the experimental buffer and
for 500 seconds after the bath was exchanged to the agonist vehicle
or the vehicle containing 100 nmol/L AVP. During the 500-second
period after the solution exchange, fluorescent signals were recorded
in the presence of the agonists. Intracellular Ca2⫹ concentration
([Ca2⫹]i) was calibrated from maximum and minimum fura 2 signals
at the end of each experiment. Specifically, the tissue bath solution
was exchanged to 5 ␮mol/L of the Ca2⫹ ionophore 4-bromo-A23187
(Molecular Probes Inc) with 2.5 mmol/L Ca2⫹ in the experimental
buffer to establish maximum fura 2 signals and also to Ca2⫹-free
experimental buffer with 100 ␮mol/L EGTA (Sigma) and 5 ␮mol/L
of 4-bromo-A23187 to establish minimum fura 2 signals. [Ca2⫹]i was
calculated as follows9: [Ca2⫹]i⫽Kd · (R⫺Rmin)/(Rmax⫺R) · F, where
Kd is the dissociation constant of fura 2, R is the actual ratio of
intensities at excitation wavelengths 340 and 380 nm, Rmax and Rmin
are the maximal and minimal fura 2 ratios in the presence and
absence of Ca2⫹, and F is the ratio of fura 2 intensities at 380 nm in
the presence and absence of Ca2⫹. Kd of fura 2 at 37°C was 224
nmol/L, as reported.9 To compare agonist responses from different
rats, [Ca2⫹]is during the peak and at 500 seconds after the agonist
vehicle and 100 nmol/L AVP (Sigma) were used.
IMCDs were loaded with 5 ␮mol/L fura 2-AM or 10 ␮mol/L
DAF-2DA as described above and preincubated for 15 minutes in
Ca2⫹-free perfusion solution containing 1 mmol/L L-arginine and 10
␮mol/L of the Ca2⫹-ATPase inhibitor thapsigargin (Sigma). Fura 2
signal was measured as described above in response to vehicle
Ca2⫹-free perfusion solution and 100 nmol/L AVP. At the end of
each experiment, maximum and minimum fura 2 signals were
collected. The DAF-2T signal was measured in response to vehicle
Ca2⫹-free perfusion solution containing 1 mmol/L L-arginine and 100
nmol/L AVP with 1 mmol/L L -arginine or 20 ␮ mol/L
DETA-NONOate.
Statistical Analysis
Data are expressed as mean⫾SE. The time responses of [Ca2⫹]i and
NO production were evaluated with a 1-way ANOVA. For [Ca2⫹]i,
the Dunn multiple range test was carried out as a post hoc test to
baseline, peak, and 500 seconds after AVP stimulation. The Bonferroni multiple range test was carried out as a post hoc test for increase
in NO production. To examine the differences in response to vehicle
and agonists, t tests were used. The level of significance was set at
a value of P⬍0.05.
Results
Intracellular Ca2ⴙ Response to AVP
Intracellular Ca2⫹ responses of the IMCDs were measured
after 100 nmol/L AVP stimulation. AVP produced a statistically significant peak increase in [Ca2⫹]i (344.5⫾53 nmol/L)
25 seconds after stimulation, which was followed by a
sustained increase, as represented by the average value at 500
seconds (126.2⫾45 nmol/L) compared with that of the
control vehicle (Figure 1).
Mori et al
Figure 1. [Ca2⫹]i response of IMCDs to 100 nmol/L AVP. IMCDs
were loaded with 5 ␮mol/L of the Ca2⫹ indicator fura 2-AM for
30 minutes. [Ca2⫹]i is shown by calibration with maximum and
minimum fura 2 fluorescence in IMCDs. Response in [Ca2⫹]i is
compared at peak and 500 seconds after bath solution
exchange with vehicle or AVP. Values are mean increase of
[Ca2⫹]i in IMCDs (n⫽7). *P⬍0.05 vs control vehicle.
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NO Response of IMCDs to AVP and Effects of NO
Scavenger Carboxy-PTIO
The intracellular NO production ([NO]i) brought about by
AVP (100 nmol/L) in IMCDs is summarized in Figure 2.
[NO]i was determined by changes of DAF-2T intensity 10
seconds before AVP and 500 seconds after AVP stimulation.
AVP administration (100 nmol/L) in the presence of
1 mmol/L L-arginine significantly increased the NO production to an average of 34⫾7 U (Figure 2A). This increase was
similar to that produced by 20 ␮mol/L of the slowly releasing
NO donor DETA-NONOate. To confirm the presence of
Ca2⫹-dependent NOS activity in IMCDs, we examined the
Vasopressin Stimulation of NO Release in IMCDs
467
Figure 3. Time course of Ca2⫹ and NO response of IMCDs to
100 nmol/L AVP. IMCDs were loaded with 5 ␮mol/L of Ca2⫹
indicator fura 2-AM for 30 minutes or 10 ␮mol/L of NO indicator
DAF-2DA for 60 minutes. Ca2⫹ response is shown as mean of
absolute [Ca2⫹]i for 10-second intervals from baseline to 200
seconds and for 30-second interval from 200 to 500 seconds
after AVP stimulation (dotted line, n⫽7). NO response is shown
as mean increase of [NO]i units for 10-second intervals at the
baseline and for 30-second intervals from 0 to 500 seconds
after AVP stimulation (bar, n⫽6). *P⬍0.05 for increase in [Ca2⫹]i
vs baseline; †P⬍0.05 for increase in [NO]i vs baseline.
[NO]i response to increased [Ca2⫹]i by using 5 ␮mol/L of the
Ca2⫹ ionophore 4-bromo-A23187 and found a significant
increase of [NO]i compared with [NO]i in the control vehicle
(P⬍0.05). To confirm DAF-2DA [NO]i specificity, [NO]i
responses of IMCDs to the agonists in the presence of the NO
scavenger carboxy-PTIO were measured. This compound
completely abolished the changes in [NO]i in response to 100
nmol/L AVP, 20 ␮mol/L DETA-NONOate, and 5 ␮mol/L of
the Ca2⫹ ionophore 4-bromo-A23187 (Figure 2B).
Time Course of Intracellular Ca2ⴙ and NO
Response of IMCDs to AVP
To determine the temporal relationships between the averaged sequential increases in [Ca2⫹]i and the elevations of
[NO]i, the[Ca2⫹]i and [NO]i signals were combined on a
single time line (Figure 3). [Ca2⫹]i showed a rapid peak
increase (466.4⫾70 nmol/L) within a few seconds and a
sustained increase (248.1⫾51 nmol/L) lasting 500 seconds.
The rise of [NO]i followed the peak increase in [Ca2⫹]i and
achieved a maximum plateau response in 300 to 500 seconds.
Abolition of NO Production by Depletion
of [Ca2ⴙ]i
Figure 2. [NO]i response of IMCDs to 100 nmol/L AVP. IMCDs
were loaded with 10 ␮mol/L of NO indicator DAF-2DA for 60
minutes. Responses in [NO]i were compared 500 seconds after
the bath solution exchange with vehicle, 100 nmol/L AVP (n⫽6),
20 ␮mol/L of the NO donor DETA-NONOate (n⫽6), and 5
␮mol/L of the Ca2⫹ ionophore 4-bromo-A23187 (n⫽6). A,
Responses in the experimental buffer with 1 mmol/L L-arginine.
B, Responses in the experimental buffer with 100 ␮mol/L NO
scavenger carboxy-PTIO. Values are mean increase of [NO]i
units. *P⬍0.05 vs control vehicle.
To examine whether increased [Ca2⫹]i was responsible for
increased NO production in IMCDs, we depleted [Ca2⫹]i with
the Ca2⫹-ATPase inhibitor thapsigargin (5 ␮mol/L) in Ca2⫹free buffer. This fully abolished the increase in [Ca2⫹]i to 100
nmol/L of AVP and abolished increases in [NO]i by 100
nmol/L AVP stimulation while preserving the NO donor
response (Figure 4).
Discussion
The present study addressed 2 issues: (1) whether AVP could
increase intracellular NO in IMCDs and (2) whether production of NO in IMCDs by AVP stimulation was mediated by
the intracellular Ca2⫹ signal transduction pathway.
468
Hypertension
February 2002 Part II
Figure 4. NO response of IMCDs to AVP in Ca2⫹-depletion
state. IMCDs were loaded with 10 ␮mol/L DAF-2DA for 60 minutes and preincubated with 5 ␮mol/L of the Ca2⫹-ATPase inhibitor thapsigargin in Ca2⫹-free buffer for 15 minutes. Response in
[NO]i was compared 500 seconds after the bath solution
exchange with vehicle, 100 nmol/L AVP, or 20 ␮mol/L DETANONOate. Values are mean increase of [NO]i units (n⫽6).
*P⬍0.05 vs control vehicle.
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Direct Evidence of Intracellular NO Increase by
AVP Stimulation in IMCDs
Previous studies in which urinary cGMP concentrations were
measured have suggested that AVP can increase the renal
production of NO.10 It has also been shown that stimulation of
vasopressin V2 receptors can result in the dilation of renal
blood vessels,10,11 especially in the medullary vasculature.12
These responses were shown to be mediated by NO production.1,10 –12 Recently, studies from our laboratory found that if
the medullary production of NO was blunted by reducing
NOS activity, even small elevations of circulating levels of
AVP produced sustained reductions of medullary blood flow
(MBF) and hypertension.13 AVP-mediated stimulation of
medullary NO production thereby protects against AVPinduced hypertension. However, the specific sites and the
mechanisms of NO production by AVP stimulation in the
medulla have remained unclear. Moreover, the mRNA for
vasopressin V2 receptors appears absent in renal arteries,
arterioles, and the vasa recta.3 Because NOS enzyme activity
is highest in the inner medullary collecting duct relative to
other tubular or vascular segments2 and because IMCDs
contain abundant vasopressin V2 receptors,14,15 which are
known to stimulate NO production,1 it was logical to search
for these responses in the IMCDs. The results of the present
study provide direct evidence that AVP produces an increase
of [NO]i in IMCDs.
Ca2ⴙ-Dependent Pathway of NO Production After
AVP Stimulation
Star et al4 have reported that Ca2⫹ and cAMP both serve as
second messengers for vasopressin in the rat inner medullary
collecting duct. Because of the identification of vasopressin
receptors in IMCDs,14,15 a number of studies have shown that
AVP increases intracellular Ca2⫹ in IMCDs by V2-like receptors.6,15 However, the physiological responses of these
changes of intracellular Ca2⫹ caused by AVP stimulation are
not fully known. Because IMCDs are rich in Ca2⫹/calmodulin-dependent constitutive NOS isoforms and exhibit the
highest levels of NOS activity within the nephrons,2 we
hypothesized that AVP would stimulate NO through a Ca2⫹
signal transduction pathway. The results of the present study
show that an increase of [Ca2⫹]i brought about by AVP was
followed by NO production. NO production was brought
about by increases in [Ca2⫹]i alone with the Ca2⫹ ionophore
4-bromo-A23187. Abolition of the [Ca2⫹]i increases in response to AVP stimulation by IMCD Ca2⫹ depletion with
thapsigargin in Ca2⫹-free buffer also abolished the NO
response. These results, combined with the knowledge derived from our previous studies of the presence of Ca2⫹/calmodulin-dependent constitutive NOS isoforms in IMCDs,2
suggest a Ca2⫹-dependent pathway for NO production with
AVP stimulation in IMCDs.
NO Detection by NO-Sensitive Dye DAF-2 DA
NO production in IMCDs was measured by using an NOsensitive dye, DAF-2DA, as established by Kojima et al.8 In
the present study, the dye was loaded into IMCDs, and it
enabled the detection of changes in [NO]i with AVP stimulation. The highest level of basal DAF-2T fluorescence
intensity was observed in IMCDs compared with the surrounding tissue, consistent with our previous report, which
found the highest NOS activity in IMCDs.2 The DAF-2DA
technique enabled the characterization of the NO responses
and the temporal relationships related to Ca2⫹ responses but
did not enable absolute quantification of NO, because the
dye-loading condition may differ between the cell types and
experimental conditions. For example, DAF-2T fluorescent
signal responses to equal molar concentrations of the NO
donor DETA-NONOate were ⬇8-fold higher for experiments
performed in the Ca2⫹-free buffer compared with the
1.5 mmol/L Ca2⫹– containing buffer. We assume that these
different experimental treatments of the cells modified the
dye-loading condition, leading to the differences in baseline
NO sensitivity of DAF-2. Therefore, we hesitated to compare
the absolute change in DAF-2 intensity between experimental
groups, and instead, we compared the DAF-2 response from
AVP stimulation or other agonists with that of the NO donor
response in the same experimental conditions. DAF-2T has a
long half-life and is not reversible,8 so the increase in
DAF-2T signal shows the production of NO, and it is these
responses that were evaluated in the present study.
Time-Dependent Relationship of Intracellular Ca2ⴙ
Changes and NO Production
[Ca2⫹]i and [NO]i were determined from IMCDs by loading 2
different fluorescent indicators, which enabled us to characterize the temporal relationship between [Ca2⫹]i and [NO]i.
Simultaneous measurements of [Ca2⫹]i and [NO]i were also
made in several single IMCDs and revealed the same pattern
of response (data not shown). The results clearly demonstrated that the rapid rise of Ca2⫹ precedes the increase of
[NO]i, indicating that increased Ca2⫹ is activating Ca2⫹dependent NOS.
Second Pathway for AVP Signal Transduction
in IMCDs
It is well known that AVP activates adenylyl cyclase via V2
receptors16 and increases cAMP4 in IMCDs. This results in
increased water permeability of the cell membrane and
activates the aquaporin shuttle system.5 Recently, Chou et al17
have shown that AVP-enhanced water permeability in IM-
Mori et al
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CDs due to increased aquaporin-2 trafficking is stimulated by
a Ca2⫹-calmodulin pathway via Ca2⫹ release from ryanodinesensitive stores, indicating that [Ca2⫹]i plays a critical role in
water reabsorption in IMCDs. However, long-term administration of AVP results in only a transient increase in water
retention and urine osmolality18 because of a phenomenon
called vasopressin escape. Many investigators have shown
that this escape is driven by body fluid expansion18 –20 and
increased renal perfusion pressure.21 The escape is also
associated with an increase in renal prostaglandins,19 a
decrease in vasopressin receptors,22 and a decrease in aquaporin water channels.23
The present study reveals another pathway whereby AVP
can affect IMCDs by increasing intracellular Ca2⫹ and activating NOS and NO production. NO-mediated increases in
soluble guanylyl cyclase activity and cGMP24 have been
shown to decrease water permeability via stimulation of
protein kinase G in the collecting duct,25 suggesting that this
response may attenuate the Ca2⫹-dependent aquaporin trafficking described above.17 An increased NO level in IMCDs
may also activate the guanylyl cyclase in the surrounding
vasa recta vessels and buffer the direct vasoconstrictive effect
of AVP. Taken together, these events help to explain the
phenomenon of vasopressin escape and explain why only V1
receptor stimulation results in sustained reduction of medullary flow and hypertension.
In summary, we have shown that AVP mobilizes Ca2⫹,
which stimulates the production of NO in IMCDs. These data
support our hypothesis that NO production in the renal
medulla may be the counterregulatory mechanism that prevents hypertension during chronic elevations of plasma AVP.
Vasopressin Stimulation of NO Release in IMCDs
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
Acknowledgments
19.
This work was supported by National Heart, Lung, and Blood
Institute grants HL-29847 and HL-49219. The authors thank
Meredith M. Skelton for careful review of the manuscript.
20.
References
21.
1. Park F, Zou AP, Cowley AW Jr. Arginine vasopressin-mediated stimulation of nitric oxide within the rat renal medulla. Hypertension. 1998;
32:896 –901.
2. Wu F, Park F, Cowley AW Jr, Mattson DL. Quantification of nitric oxide
synthase activity in microdissected segments of the rat kidney. Am J
Physiol. 1999;276:F874 –F881.
3. Park F, Mattson DL, Skelton MM, Cowley AW Jr. Localization of the
vasopressin V1a and V2 receptors within the renal cortical and medullary
circulation. Am J Physiol. 1997;273:R243–R251.
4. Star RA, Nonoguchi H, Balaban R, Knepper MA. Calcium and cyclic
adenosine monophosphate as second messengers for vasopressin in the rat
inner medullary collecting duct. J Clin Invest. 1988;81:1879 –1888.
5. Sasaki S, Fushimi K, Saito H, Saito F, Uchida S, Ishibashi K, Kuwahara
M, Ikeuchi T, Inui K, Nakajima K, et al. Cloning, characterization, and
22.
23.
24.
25.
469
chromosomal mapping of human aquaporin of collecting duct. J Clin
Invest. 1994;93:1250 –1256.
Maeda Y, Han JS, Gibson CC, Knepper MA. Vasopressin and oxytocin
receptors coupled to Ca2⫹ mobilization in rat inner medullary collecting
duct. Am J Physiol. 1993;265:F15–F25.
Schmidt HH, Pollock JS, Nakane M, Forstermann U, Murad F. Ca2⫹/calmodulin-regulated nitric oxide synthases. Cell Calcium. 1992;13:
427– 434.
Kojima H, Nakatsubo N, Kikuchi K, Kawahara S, Kirino Y, Nagoshi H,
Hirata Y, Nagano T. Detection and imaging of nitric oxide with novel
fluorescent indicators: diaminofluoresceins. Anal Chem. 1998;70:
2446 –2453.
Grynkiewicz G, Poenie M, Tsien RY. A new generation of Ca2⫹ indicators with greatly improved fluorescence properties. J Biol Chem. 1985;
260:3440 –3450.
Rudichenko VM, Beierwaltes WH. Arginine vasopressin-induced renal
vasodilation mediated by nitric oxide. J Vasc Res. 1995;32:100 –105.
Naitoh M, Suzuki H, Murakami M, Matsumoto A, Ichihara A, Nakamoto
H, Yamamura Y, Saruta T. Arginine vasopressin produces renal vasodilation via V2 receptors in conscious dogs. Am J Physiol. 1993;265:
R934 –R942.
Nakanishi K, Mattson DL, Gross V, Roman RJ, Cowley AW Jr. Control
of renal medullary blood flow by vasopressin V1 and V2 receptors. Am J
Physiol. 1995;269:R193–R200.
Szentivanyi M Jr, Park F, Maeda CY, Cowley AW Jr. Nitric oxide in the
renal medulla protects from vasopressin-induced hypertension. Hypertension. 2000;35:740 –745.
Kirk KL. Binding and internalization of a fluorescent vasopressin
analogue by collecting duct cells. Am J Physiol. 1988;255:C622–C632.
Champigneulle A, Siga E, Vassent G, Imbert-Teboul M. V2-like vasopressin receptor mobilizes intracellular Ca2⫹ in rat medullary collecting
tubules. Am J Physiol. 1993;265:F35–F45.
Breyer MD, Ando Y. Hormonal signaling and regulation of salt and water
transport in the collecting duct. Annu Rev Physiol. 1994;56:711–739.
Chou CL, Yip KP, Michea L, Kador K, Ferraris JD, Wade JB, Knepper
MA. Regulation of aquaporin-2 trafficking by vasopressin in the renal
collecting duct. J Biol Chem. 2000;275:36839 –36846.
Chan WY. A study on the mechanism of vasopressin escape: effects of
chronic vasopressin and overhydration on renal tissue osmolality and
electrolytes in dogs. J Pharmacol Exp Ther. 1973;184:244 –251.
Gross PA, Kim JK, Anderson RJ. Mechanisms of escape from desmopressin in the rat. Circ Res. 1983;53:794 – 804.
Cowley AW, Jr, Merrill DC, Quillen EW Jr, Skelton MM. Long-term
blood pressure and metabolic effects of vasopressin with servo-controlled
fluid volume. Am J Physiol. 1984;247:R537–R545.
Hall JE, Montani JP, Woods LL, Mizelle HL. Renal escape from vasopressin: role of pressure diuresis. Am J Physiol. 1986;250:F907–F916.
Tian Y, Sandberg K, Murase T, Baker EA, Speth RC, Verbalis JG.
Vasopressin V2 receptor binding is down-regulated during renal escape
from vasopressin-induced antidiuresis. Endocrinology. 2000;141:
307–314.
Ecelbarger CA, Nielsen S, Olson BR, Murase T, Baker EA, Knepper MA,
Verbalis JG. Role of renal aquaporins in escape from vasopressin-induced
antidiuresis in rat. J Clin Invest. 1997;99:1852–1863.
Murad F, Waldman S, Molina C, Bennett B, Leitman D. Regulation and
role of guanylate cyclase-cyclic GMP in vascular relaxation. Prog Clin
Biol Res. 1987;249:65–76.
Garcia NH, Stoos BA, Carretero OA, Garvin JL. Mechanism of the nitric
oxide-induced blockade of collecting duct water permeability. Hypertension. 1996;27:679 – 683.
Vasopressin Increases Intracellular NO Concentration via Ca2+ Signaling in Inner
Medullary Collecting Duct
Takefumi Mori, Jeffrey G. Dickhout and Allen W. Cowley, Jr
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Hypertension. 2002;39:465-469
doi: 10.1161/hy02t2.102908
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