Expression of adenosine receptors in the preglomerular
microcirculation
Edwin K.
Jackson1,2,3,
Chongxue
Zhu1,2, and
Stevan P.
Tofovic1,2
1 Center for Clinical Pharmacology and Departments
of 2 Medicine and 3 Pharmacology,
University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania
15261
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ABSTRACT |
The purpose of
this study was to systematically investigate the abundance of each of
the adenosine receptor subtypes in the preglomerular microcirculation
vs. other vascular segments and vs. the renal cortex and medulla. Rat
preglomerular microvessels (PGMVs) were isolated by iron oxide loading
followed by magnetic separation. For comparison, mesenteric
microvessels, segments of the aorta (thoracic, middle abdominal, and
lower abdominal), renal cortex, and renal medulla were obtained by
dissection. Adenosine receptor protein and mRNA expression were
examined by Western blotting, Northern blotting, and RT-PCR. Our
results indicate that compared with other vascular segments and renal
tissues, A1 and A2B receptor protein and mRNA
are abundantly expressed in the preglomerular microcirculation, whereas
A2A and A3 receptor protein and mRNA are barely
detectable or undetectable in PGMVs. We conclude that, relative to
other vascular and renal tissues, A1 and A2B
receptors are well expressed in PGMVs, whereas A2A and
A3 receptors are notably deficient. Thus A1 and
A2B receptors, but not A2A or A3
receptors, may importantly regulate the preglomerular microcirculation.
kidney; renal microcirculation; mesenteric microcirculation; aorta
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INTRODUCTION |
PHARMACOLOGICAL
STUDIES (for a review, see Ref. 14), and more
recently studies in mice with null mutations in one of the adenosine
receptors (35), suggest that adenosine receptors in the
preglomerular microcirculation importantly regulate renal function.
Specifically, evidence exists supporting biologically significant roles
for adenosine receptors in the preglomerular microcirculation with
regard to the regulation of renal blood flow and renal vascular
resistance (14), glomerular filtration rate
(25), renin release (19, 26),
tubuloglomerular feedback (35, 36), and the renal vascular
response to angiotensin II (42). Moreover, it is
conceivable that adenosine receptors may be involved in modulating
vascular smooth muscle cell growth and extracellular matrix production
in the preglomerular microcirculation because adenosine receptors
appear to participate in these processes in vascular smooth muscle
cells from conduit arteries (5, 7).
Despite the potential physiological importance of adenosine receptors
in the preglomerular microcirculation, there are no published studies
characterizing the abundance of the four known adenosine receptor
subtypes in the preglomerular microcirculation. As summarized in Table
1, adenosine receptors in the kidney have been investigated in at least 14 different studies employing a variety
of methods, including ligand binding in isolated membranes, autoradiography, Northern blotting, in situ hybridization, RT-PCR, immunocytochemistry and Western blotting. However, none of these studies defined the relative expression of adenosine receptors in the
preglomerular microcirculation.
Given the probable physiological importance of adenosine receptors in
the preglomerular microcirculation and the recently accelerated efforts
to develop specific and potent adenosine-receptor agonists and
antagonists for the treatment of renal diseases (23, 24)
and as diuretics (10), we felt that it was critical to investigate carefully and thoroughly the expression of adenosine receptor subtypes in the preglomerular microcirculation. To accomplish this objective, we isolated renal preglomerular microvessels (PGMVs) using iron oxide loading with magnetic separation and measured the
expression of A1, A2A, A2B, and
A3 receptor protein using Western blotting with
quantitative densitometry. We also measured adenosine receptor mRNA
expression with Northern blotting and/or RT-PCR. As a reference, we
calibrated the quantities of the various adenosine receptor subtypes in
the preglomerular microcirculation to the expression of the receptors
in the renal cortex, renal medulla, mesenteric microvessels, and
various segments of the aorta (thoracic, middle abdominal, and lower abdominal).
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METHODS |
Isolation of PGMVs.
Adult male Wistar rats were obtained from Taconic Farms (Germantown,
NY), housed at the University of Pittsburgh Animal Facility, and fed
Prolab RMH 3000 (PMI Feeds, St. Louis, MO) containing 0.26% sodium and
0.82% potassium. All studies received prior approval by the University
of Pittsburgh Animal Care and Use Committee. Rats were anesthetized
with Inactin (100 mg/kg ip), and the aorta below the renal artery was
cannulated with polyethylene-190 tubing. The proximal aorta, mesenteric
artery, and small side branches of the aorta were ligated, and the
kidneys were flushed (10 ml) with room temperature L-15 medium
(Leibovitz medium; Sigma, St. Louis, MO). A 1% suspension of iron
oxide (Aldrich Chemical, Milwaukee, WA) in L-15 medium (10 ml) was
flushed into the kidneys. The kidneys were harvested, placed in
ice-cold L-15 medium, and dissected by removing the renal medulla and
interlobar arteries. The cortex was sliced into small pieces, suspended
in ice-cold L-15 medium, and dispersed by pushing the cortical material
through a series of increasingly small needle hubs (3 times through 16, 18, and 21 gauge and 6 times through 23 gauge). The dispersed cortical material was suspended in ice-cold L-15 medium, and a magnet was applied to the tube to retrieve the iron oxide-laden PGMVs while the
unwanted material was decanted. The glomeruli were removed from the
microvessels by filtering the suspension of PGMVs through a 149-µm
nylon mesh. The PGMVs were retrieved from the nylon mesh and
placed in ice-cold PBS. A sample of the PGMVs was examined by
phase-contrast microscopy to confirm that the preparation consisted of
interlobular, accurate, and afferent arterioles without contaminating glomeruli, efferent arterioles, or tubules.
Isolation of other tissues.
Microvessels (second- and third-order branches) from the mesenteric
vascular bed and renal cortical tissue, renal medullary tissue, and
total aorta and segments from the thoracic, midabdominal, and lower
abdominal aorta were removed, placed in ice-cold PBS, and dissected
from surrounding adipose and connective tissue.
Protein extraction.
Immediately after the tissues were obtained, the tissues were frozen in
liquid nitrogen and ground into a powder on liquid nitrogen with a
mortar and pestle. The ground tissues were placed in a tube with 0.5 ml
SDS buffer (50 mM Tris, pH 7.0, 2% SDS, 10% glycerol) containing
protease inhibitors (2 µg/ml antipain, 1 µg/ml aprotinin, 2 µg/ml
leupeptin, 1 mg/ml phenylmethylsulfonyl fluoride) and homogenized in a
tight glass homogenizer with a Teflon pestle. The homogenate was
centrifuged at 12,000 rpm at 4°C for 10 min, and the supernatant was
recovered. Protein in the supernatant was determined by the copper
bicinchoninic acid method, and samples were stored at
20°C.
Western blotting.
Laemmli buffer was added to samples, and they were placed in boiling
water for 5 min and then chilled immediately on ice. Samples (30 µg
protein/well) were loaded onto a 7.5-10% acrylamide gel and
subjected to SDS-PAGE using the Bio-Rad minigel system. Proteins were
then electroblotted onto a polyvinylidene difluoride membrane
(Millipore, Bedford, MA). The membrane was blocked with 5% milk for
1 h and incubated for 3 h at room temperature or at 4°C
overnight with the first antibody [anti-A1 receptor
antibody (catalogue no. A-268, diluted 1:1,000 in PBS containing 0.5%
Tween 20, Sigma); anti-A2A receptor, anti-A2B
receptor and anti-A3 receptor antibodies (catalogue nos.
AB1559P, AB1589P, and AB1590P, respectively, diluted 1:500 in PBS
containing 0.5% Tween 20, Chemicon, Temecula, CA)]. Membranes were
washed three times in PBS containing 0.5% Tween 20 solution and then
incubated at room temperature for 1 h with horseradish
peroxidase-conjugated donkey anti-rabbit IgG secondary antibody
(Amersham, Arlington Heights, IL) at 1:5,000 dilution. Membranes were
exposed to films, and the signals were detected by a Supersignal
Substrate kit (Pierce, Rockford, IL).
Extraction of RNA, RT-PCR, preparation of cDNA probes, and
Northern blotting.
Total RNA was isolated from rat brains using TRIzol reagent solution
(GIBCO Life Technologies, Carlsbad, CA), and this material was used to
prepare cDNA probes for rat A1, A2A,
A2B, and A3 receptor mRNA and for rat
-actin mRNA. By using the primer sequences listed in
Table 2, RNA (0.5 µg) was reverse
transcribed and amplified using a Titanium One-Step RT-PCR Kit
(Clontech, Palo Alto, CA). Each PCR cycle (a total of 30 cycles)
consisted of denaturing at 94°C for 30 s, annealing at 65°C
for 30 s, and extension at 72°C for 60 s. RT-PCR products
were separated on a 1.2% agarose gel. The RT-PCR products were
extracted from the agarose gel and immediately ligated into pCR II
vector (Invitrogen, Carlsbad, CA), which was used to transform
competent bacteria. Gel electrophoresis of specific restriction enzyme
digests (Table 2) of the vector containing the inserted cDNA was
consistent with the expected fragmentation pattern. Furthemore, the
inserted cDNA in the subcloned plasmids was sequenced to confirm that
our procedure did indeed yield the appropriate cDNA probes. For probe
preparation, plasmids extracted from transformed bacteria were digested
with EcoRI restriction enzyme, and the expected cDNA
fragments were harvested from the agarose gel and labeled with
32P using the method of random priming (Roche Diagnostics,
Indianapolis, IN).
Total RNA (10 µg) from kidney tissues and vascular tissues was
isolated with TRIzol reagent, denatured, and loaded onto a 1.2%
agarose-formaldehyde gel. After electrophoresis, RNA was transferred to
a nylon membrane and fixed by exposure to ultraviolet light. Membranes
were prehybridized for 30 min at 60°C with ExpressHyb hybridization
solution (Clontech). Denatured cDNA probe was added to prewarmed
hybridization solution to give a final concentration of 1,000,000 counts · min
1 · ml
1.
Membranes were incubated with the probe solution with continuous rotation at 63°C for 1 h. After incubation with probe solution, membranes were washed three times with solution I (2×
standard sodium citrate, 0.05% SDS) at room temperature, and then
washed in solution II (0.1% standard sodium citrate, 0.1%
SDS) at 50°C. Blots were developed using the PhosphorImage system
(Molecular Dynamics, Sunnyvale, CA). Blots then were stripped and
rehybridized with
-actin cDNA probe and redeveloped.
Data analysis.
For Western blot analysis, band densities were quantitatively measured
using Scion-image software. Background signals were obtained in each
lane and subtracted from the band densities to correct for the
background signal. For Northern blot analysis, band radioactivity was
determined using the PhosphorImage system (Molecular Dynamics), and
values were normalized to (i.e., divided by) the signal for
-actin.
Data were analyzed with a one-factor analysis of variance followed by a
Fisher's least significant difference test if the overall P
value from the analysis of variance was significant. The criterion of
significance was P < 0.05, and all data are expressed
as means ± SE.
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RESULTS |
Figures 1 and
2 show Western blot analysis and
quantitative densitometry results using the anti-A1
receptor antibody. Western blot analysis revealed a single, dense band
at ~38 kDa that corresponded closely to the nominal mass of the
A1 receptor (36.5 kDa) (16) and to the 38-kDa
signal obtained by Rivkees et al. (29) and Cheng et al.
(3) using anti-A1 receptor antibodies. All
vascular tissues examined (thoracic aorta, middle abdominal aorta,
lower abdominal aorta, mesenteric microvessels, and PGMVs) gave a
strong 38-kDa signal; however, quantitative densitometry demonstrated that PGMVs expressed significantly (P < 0.05), albeit
only modestly, more A1 receptors per milligram protein
compared with all other vascular tissues (Fig. 1). Although the three
aortic segments expressed similar levels of A1 receptors,
the mesenteric microvessels expressed significantly, but only slightly,
more A1 receptors than did the middle abdominal and lower
abdominal aorta (Fig. 1). As shown in Fig. 2, PGMVs and renal cortical
tissue expressed similar numbers of A1 receptors, whereas
renal medullary tissue expressed approximately twice the number of
A1 receptors per milligram protein compared with either the
renal cortex or PGMVs (P < 0.05). The approximate
doubling of A1 receptor protein expression in the
renal medulla compared with the renal cortex or PGMVs was associated
with an approximate doubling of the expression of A1 receptor mRNA in the renal medulla compared with the cortex and PGMVs
(Fig. 3). However, unlike receptor
protein expression, receptor mRNA expression was similar in the aorta
(thoracic + middle abdominal + lower abdominal) compared with
PGMVs.

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Fig. 1.
Top: Western blot analysis of proteins (30 µg/lane) using an anti-A1 receptor antibody. Samples were
from the thoracic aorta (lanes 1-3), middle abdominal
aorta (lanes 4-6), lower abdominal aorta (lanes
7-9), mesenteric microvessels (lanes 10-12),
and preglomerular microvessels (PGMVs; lanes 13-15)
from 3 different rats. Bottom: bands shown in top
panel were subjected to quantitative densitometry and analyzed by
1-factor ANOVA followed by Fisher's least significant difference test.
Values are means ± SE (n = 3). MW, molecular
weight.
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Fig. 2.
Top: Western blot analysis of proteins (30 µg/lane)
using an anti-A1 receptor antibody. Samples were from the
renal medulla (lanes 1-4), renal cortex (lanes
5-8), and PGMVs (lanes 9-12) from 4 different
rats. Bottom: bands shown in top panel were
subjected to quantitative densitometry and analyzed by 1-factor ANOVA
followed by Fisher's least significant difference test. Values are
means ± SE (n = 4).
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Fig. 3.
Top: Northern blot analysis using
A1 receptor cDNA probe. Samples were from the total aorta
(lanes 1 and 2), renal medulla (lanes
3 and 4), renal cortex (lanes 5 and
6), and PGMVs (lanes 7 and 8) from 2 different rats. Bottom: radioactivity in bands shown in
top panel was quantified, normalized to the -actin
signal, and analyzed by 1-factor ANOVA followed by Fisher's least
significant difference test. Values are means ± SE
(n = 2).
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Multiple bands were detected when vascular samples were stained with
the anti-A2A receptor antibody (Fig.
4). However, a dense, dominant band was
detected at ~45 kDa in all of the vascular tissues except the PGMVs
in which the 45-kDa band was barely detectable. Most likely, this
45-kDa band represents staining of the A2A receptor because
the nominal mass of the A2A receptor is at 44.7 kDa
(16) and because a similar band was reported by Rosin et
al. (43-48 kDa) (30), Marala and Mustafa (45 kDa)
(17) and Piersen et al. (44 kDa) (27) using
anti-A2A receptor antibodies. Importantly, compared with
either all other vascular tissues (Fig. 4) or with the renal cortex
(Fig. 5), PGMVs expressed few
A2A receptors. It is noteworthy, however, that the
expression of A2A receptors per milligram protein was
significantly greater in the mesenteric microvessels compared with the
three segments of the aorta (Fig. 4) and that the renal cortex, but not
the renal medulla, gave a strong A2A receptor signal.
A2A receptor mRNA could not be detected in PGMVs using
Northern blot analysis (data not shown). Moreover, even when mRNA from
PGMVs was reverse transcribed and subjected to 30 cycles of
amplification by PCR, ethidium bromide staining failed to detect a
signal for A2A receptor cDNA (Fig.
6), whereas a clear signal was obtained
when renal cortical tissue similarly was subjected to RT-PCR.

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Fig. 4.
Top: Western blot analysis of proteins (30 µg/lane) using an anti-A2A receptor antibody. Samples
were from the thoracic aorta (lanes 1-3), middle
abdominal aorta (lanes 4-6), lower abdominal aorta
(lanes 7-9), mesenteric microvessels (lanes
10-12), and PGMVs (lanes 13-15) from 3 different rats. Bottom: bands shown in top panel
were subjected to quantitative densitometry and analyzed by 1-factor
ANOVA followed by Fisher's least significant difference test. Values
are means ± SE (n = 3).
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Fig. 5.
Top: Western blot analysis of proteins (30 µg/lane)
using an anti-A2A receptor antibody. Samples were from the
renal medulla (lanes 1-4), renal cortex (lanes
5-8), and PGMVs (lanes 9-12) from 4 different
rats. Bottom: bands shown in top panel were
subjected to quantitative densitometry and analyzed by 1-factor ANOVA
followed by Fisher's least significant difference test. Values are
means ± SE (n = 4).
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Fig. 6.
RNA samples from renal cortex (lanes 1 and 3)
and PGMVs (lanes 2 and 4) were subjected to
RT-PCR using primers to either A2A receptor mRNA
(lanes 1 and 2) or A3 receptor mRNA
(lanes 3 and 4). cDNA was run on a 1.2% agarose
gel and visualized with ethidium bromide staining.
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Staining vascular (Fig. 7) or renal (Fig.
8) samples with the
anti-A2B receptor antibody gave rise to only one clear-cut
band at ~52 kDa, which is more massive than the nominal molecular
mass of the A2B receptor (36.3 kDa)
(16). However, the Western blot analysis appeared
convincing, and the 52-kDa protein detected in this study corresponded
closely to the 50- to 55-kDa protein detected by Puffinbarger et al.
(28) using an anti-A2B receptor antibody.
Also, preabsorption of the anti-A2B receptor antibody with
a blocking peptide to the anti-A2B receptor antibody
abolished the 52-kDa signal (data not shown). Most likely, this
high-molecular-mass species represents a glycosylated form of the
A2B receptor. Importantly, the expression of the
A2B receptor was similar among all the vascular tissues
examined (Fig. 7) and was similar among renal cortical tissue, renal
medullary tissue, and PGMVs (Fig. 8). Northern blot analysis revealed a
slightly, albeit significantly (P < 0.05), lower
A2B receptor mRNA expression in PGMVs compared with aorta (total), renal medulla, or renal cortex (Fig.
9).

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Fig. 7.
Top: Western blot analysis of proteins (30 µg/lane)
using an anti-A2B receptor antibody. Samples were from the
thoracic aorta (lanes 1-3), middle abdominal aorta
(lanes 4-6), lower abdominal aorta (lanes
7-9), mesenteric microvessels (lanes 10-12),
and PGMVs (lanes 13-15) from 3 different rats.
Bottom: bands shown in top panel were subjected
to quantitative densitometry. Values are means ± SE
(n = 3).
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Fig. 8.
Top: Western blot analysis of proteins (30 µg/lane)
using an anti-A2B receptor antibody. Samples were from the
renal medulla (lanes 1-4), renal cortex (lanes
5-8), and PGMVs (lanes 9-12) from 4 different
rats. Bottom: bands shown in top panel were
subjected to quantitative densitometry. Values are means ± SE
(n = 4).
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Fig. 9.
Top: Northern blot analysis using A2B
receptor cDNA probe. Samples were from the total aorta (lanes
1 and 2), renal medulla (lanes 3 and
4), renal cortex (lanes 5 and 6), and
PGMVs (lanes 7 and 8) from 2 different rats.
Bottom: Radioactivity in bands shown in top panel
was quantified, normalized to the -actin signal, and analyzed by
1-factor ANOVA followed by Fisher's least significant difference test.
Values are means ± SE (n = 2).
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Staining vascular (Fig. 10) or renal
(Fig. 11) samples with the
anti-A3 receptor antibody gave rise to only one clear-cut
band at ~52 kDa, which is more massive than the nominal molecular
mass of the A3 receptor (36.2 kDa) (16).
However, Western blot analysis appeared convincing, and the 52-kDa
protein detected in this study corresponded closely to the 52-kDa
protein detected by Zou et al. (46) using an
anti-A3 receptor antibody. Most likely, this high-molecular-mass species represents a glycosylated form of the
A3 receptor. Importantly, compared with either all other
vascular tissues (Fig. 10) or the renal medulla or renal cortex (Fig.
11), PGMVs expressed few A3 receptors. A3
receptor mRNA could not be detected in PGMVs using Northern blotting
(data not shown). Moreover, even when mRNA from PGMVs was reverse
transcribed and subjected to 30 cycles of amplification by PCR,
ethidium bromide staining failed to detect a signal for A3
receptor cDNA (Fig. 6), whereas a clear signal was obtained when renal
cortical tissue similarly was subjected to RT-PCR.

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Fig. 10.
Top: Western blot analysis of proteins (30 µg/lane) using an anti-A3 receptor antibody. Samples were
from the thoracic aorta (lanes 1-3), middle abdominal
aorta (lanes 4-6), lower abdominal aorta (lanes
7-9), mesenteric microvessels (lanes 10-12),
and PGMVs (lanes 13-15) from 3 different rats.
Bottom: bands shown in top panel were subjected
to quantitative densitometry and analyzed by 1-factor ANOVA followed by
Fisher's least significant difference test. Values are means ± SE (n = 3).
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Fig. 11.
Top: Western blot analysis of proteins (30 µg/lane)
using an anti-A3 receptor antibody. Samples were from the
renal medulla (lanes 1-4), renal cortex (lanes
5-8), and PGMVs (lanes 9-12) from 4 different
rats. Bottom: bands shown in top panel were
subjected to quantitative densitometry. Values are means ± SE
(n = 4).
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DISCUSSION |
An important finding of this study is the abundant level of
expression of A1 receptors in PGMVs. In this regard,
relative to other vascular segments, PGMVs express slightly more
A1 receptor protein per milligram total protein, and the
expression of A1 receptor protein by PGMVs is similar to
that of the renal cortex. It is notable, however, that the most
abundant expression of A1 receptor protein was observed in
the renal medulla, which expressed approximately twice the level of
A1 receptor protein compared with the renal cortex or
PGMVs. The similar levels of expression of A1 receptor
protein in vascular elements and renal cortex and the higher levels of
expression of A1 receptor protein in the renal medulla is
consistent with the Northern blot studies. In this regard, the medulla
expressed twice as much A1 receptor mRNA compared with the
aorta, cortex, or PGMVs, all of which expressed similar levels of
A1 receptor mRNA.
These results imply an important physiological role for A1
receptors in the preglomerular microcirculation. In agreement with this
interpretation are the reports that A1-receptor agonists vasoconstrict the PGMVs (31), potentiate angiotensin
II-induced changes in renal vascular resistance (42),
reduce the glomerular filtration rate (20), and inhibit
renin release (21). Also consistent with this inference
are the findings that A1-receptor antagonists inhibit
tubuloglomerular feedback (31), stimulate renin release
(26), and reduce vasoconstriction induced by certain nephrotoxins (14). Finally, recent studies indicate a
total lack of tubuloglomerular feedback responses in A1
receptor knockout mice (35).
The role of A1 receptors in other vascular segments, such
as the aorta and mesentery, is presently unclear; however, the present results suggest that A1 receptors may play yet-to-be
determined physiological roles in other segments of the vascular tree.
Our results also imply an important physiological role for
A1 receptors in the renal medulla. Indeed, it is
conceivable that the lack of effect of A1-receptor
antagonists on potassium excretion (14) may be because of
blockade of A1 receptors in the collecting tubules of the
renal medulla, a site where potassium secretion is under strong regulation.
In contrast to the A1 receptor, the A2A
receptor is markedly underexpressed in the preglomerular
microcirculation. In this regard, A2A receptors are either
undetectable or only barely detectable by Western blotting, whereas
Western blotting for A2A receptors gives rise to strong
signals in other vascular segments, as well as in the renal cortex.
This contrast is particularly striking when the levels of expression of
A2A receptors in the mesenteric microvessels vs. PGMVs
(Fig. 4) and in the renal cortex vs. PGMVs (Fig. 5) are compared. The
results of the studies using Western blotting are confirmed by the
inability to detect A2A receptor mRNA with either Northern
blotting or RT-PCR (Fig. 6). These results imply an unimportant role
for A2A receptors in the preglomerular microcirculation.
Consistent with this interpretation are the reports that
A2A-receptor agonists have little or no effect on the
preglomerular microcirculation (1, 41), while strongly dilating efferent arterioles (22) and the vasa recta
(1, 45). However, there is evidence that A2A
receptors mediate dilation of preglomerular vessels in the small
population of juxtamedullary nephrons (22).
A2A receptors are particularly densely expressed in
mesenteric microvessels, suggesting a quantitatively significant role for A2A receptors in the regulation of intestinal blood
flow. Indeed, pharmacological studies indicate that
A2A-receptor agonists markedly vasodilate the intestinal
circulation (13). The high level of expression of
A2A receptors in the renal cortex suggests an important
role for A2A receptors in renal physiology, an implication that is supported by recent findings that A2A receptors
inhibit intrarenal inflammation and renal ischemia-reperfusion
injury (24).
A2B receptors are abundantly expressed in the preglomerular
microcirculation. In fact, A2B receptors are widely and
evenly expressed throughout the vascular system and kidneys, including segments of the aorta, mesenteric microvessels, PGMVs, renal cortex, and renal medulla. A similar conclusion is reached when A2B
receptor mRNA abundance is measured by Northern blotting (Fig. 9). In
light of recent investigations into the role of A2B
receptors in vascular biology, our finding that A2B
receptors are present throughout the circulation is of considerable
importance. In cultured vascular smooth muscle cells (5,
7), cardiac fibroblasts (4, 8), and mesangial cells
(6), A2B receptors strongly suppress cell proliferation, extracellular matrix production, and MAP kinase activation. Thus it appears that A2B receptors negatively
regulate vascular remodeling. If this hypothesis is correct,
A2B receptors may importantly suppress inappropriate
vascular remodeling and fibrosis in the vascular tree, including PGMVs.
Therefore, A2B receptors in the preglomerular
microcirculation may be targets for drug development aimed at
ameliorating the progression of chronic renal failure by inhibiting the
abnormal growth of vascular elements and, perhaps, mesangial cells in
the kidney.
Like the A2A receptor, but unlike A1 receptors
or A2B receptors, the A3 receptor is
underexpressed in the preglomerular microcirculation. Figure 10
demonstrates strong signals for the A3 receptor in the thoracic aorta, middle abdominal aorta, lower abdominal aorta, and
mesenteric microvessels, but barely detectable signals for the
A3 receptor in PGMVs. Similarly, Western blotting of renal cortex, medulla, and PGMVs indicates low expression of A3
receptor protein in PGMVs compared with the renal medulla and
cortex. The low signal for the A3 receptor in PGMVs vs.
aortic tissue cannot be attributed to the fact that PGMVs are
resistance vessels and the aorta is a conduit vessel because the
A3 receptor signal is strong in the mesenteric
microvessels. The results of the studies using Western blotting are
confirmed by the inability to detect A3 receptor mRNA in
PGMVs with either Northern blotting or RT-PCR (Fig. 6). The low
expression of A3 receptors in the preglomerular microcirculation is consistent with the lack of effects of
A3-receptor agonists and antagonists in the kidney
(18, 45). However, the abundant expression of
A3 receptors in the aorta and mesenteric microvessels
relative to PGMVs suggests an as yet unrecognized role for
A3 receptors in the extrarenal circulation.
In summary, our results indicate that both A1 receptors and
A2B receptors are highly expressed in the preglomerular
microcirculation and may be important targets for the development of
new drugs to treat renal disease and regulate renal function.
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ACKNOWLEDGEMENTS |
This work was supported by National Heart, Lung, and Blood
Institute Grant HL-55314.
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FOOTNOTES |
Address for reprint requests and other correspondence:
E. K. Jackson, Ctr. for Clinical Pharmacology, 623 Scaife
Hall, 3550 Terrace St., Pittsburgh, PA 15261 (E-mail:
edj+{at}pitt.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.
10.1152/ajprenal.00232.2001
Received 26 July 2001; accepted in final form 11 February 2002.
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