1 Department of Pharmacology and 2 Center for Clinical Pharmacology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15213
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ABSTRACT |
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The exaggerated
sensitivity of spontaneously hypertensive rat (SHR) renal
microvasculature to angiotensin II (ANG II) may be due to an imbalance
between the effectiveness of
Gs-utilizing vasodilator
pathways and vasoconstrictor pathways activated by ANG II (mediated by
G
i-1,
G
i-2,
G
i-3, and
G
q). Because the alteration
appears to be distal to the hormone receptors and proximal to the
effector adenylyl cyclase, we hypothesized that SHR have altered
amounts of signal-transducing G proteins. This was examined by
quantifying the steady-state mRNA levels of specific G
subunits in
renal microvessels of 12- to 14-wk-old SHR and control Wistar-Kyoto (WKY) rats, using a quantitative-competitive polymerase chain reaction
technique coupled to reverse transcription. No significant differences
were detected in the absolute levels of
G
s (0.96 ± 0.35 vs. 0.74 ± 0.25 amol/50 ng RNA) or in the relative levels of
G
i-1 (0.44 ± 0.05 vs. 0.48 ± 0.13), G
i-2 (40.9 ± 7.8 vs. 45.2 ± 8.9), or
G
i-3 (0.79 ± 0.05 vs. 0.82 ± 0.15) normalized to the level of
G
s for WKY vs. SHR,
respectively. The ratio of G
q
to G
s tended to be higher in
SHR, but this difference did not achieve statistical significance (0.41 ± 0.08 vs. 1.04 ± 0.32, P = 0.08). In conclusion, the steady-state levels of
G
s,
G
i-1, G
i-2,
G
i-3, and
G
q are similar in SHR and WKY
renal microvasculature, suggesting that other components of the ANG II
signal transduction mechanism are responsible for the enhanced renal
vascular responsiveness in SHR.
quantitative polymerase chain reaction; angiotensin II; enzyme-linked oligonucleotide-sorbent assay; signal transduction
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INTRODUCTION |
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SPONTANEOUSLY HYPERTENSIVE rats (SHR), compared with
the control Wistar-Kyoto (WKY) strain, possess an intrinsically greater sensitivity of the renal vasculature to angiotensin II (ANG II) (10,
18-21, 30). This abnormality in renal vascular regulation appears
to be due at least in part to an imbalance between the effectiveness of
Gs-utilizing
signal-transduction pathways mediating vasodilatation and ANG
II-mediated pathways causing vasoconstriction. This imbalance has been
demonstrated not only for regulation of vascular resistance (3-5,
30) but also with regard to synthesis of adenosine
3',5'-cyclic monophosphate (cAMP) (31). The preglomerular arterioles are key resistance vessels of the kidney and mediate renovascular responses to various stimuli, including ANG II.
Importantly, Kost et al. (19) showed that medial hypertrophy in
response to ANG II is more pronounced in SHR preglomerular vessels
(e.g., interlobular and arcuate arteries), suggesting that
hyperresponsiveness to ANG II of preglomerular microvessels mediates
the enhanced renovascular responses of SHR.
The number and subtype composition of ANG II receptor subtypes is
similar in the preglomerular vessels of the two strains (2). Moreover,
pretreatment with pertussis toxin, which inactivates Gi, normalizes enhanced
renovascular responses to ANG II in SHR (14). Additionally, experiments
using a cell-permeable analog of cAMP, a direct activator of adenylyl
cyclase (5), and direct activators of
G
s (4) suggest
that renovascular responses are equivalent in SHR and WKY when the cAMP
signal-transduction pathway is activated downstream of the ANG II
receptor-G protein coupling step. These observations strongly suggest a
role for G proteins in altered signal transduction in the renal
microvasculature of SHR.
Accordingly, the goal of the present study was to test the hypothesis
that SHR have altered expression of the G proteins involved in ANG II
signal transduction and/or of
Gs, which is the common signal
transducer for several vasodilators that have defective interaction
with ANG II in SHR. In this regard, steady-state mRNA levels were
measured, because quantification of low-abundance proteins using
Western blotting presents difficulties related to specificity and
sensitivity. Moreover, in the literature pertaining to SHR
and/or ANG II effects in vascular smooth muscle cells, mRNA
levels for G proteins correlate well with protein levels (11, 17, 29).
The ANG II type 1 receptor (AT1
receptor) is a seven-transmembrane domain receptor that is linked to G
proteins. The various G proteins known to be associated with the ANG II
receptor are the three subtypes of
Gi
(G
i-1,
G
i-2, and
G
i-3) and
G
q. ANG II acts through a
G
i-type protein to inhibit
production of cAMP by adenylyl cyclase (15). This can oppose the
effects of vasodilator agents that act via
G
s such as dopamine (5),
prostaglandin I2 (3), and
2-adrenoceptor agonists (31).
Linkage of the AT1 receptor to all
three subtypes of G
i has been
demonstrated in the case of recombinant
G
i-1,
G
i-2, and
G
i-3 (9). Also, Pobiner et al.
(26) have shown linkage of the AT1
receptor with G
i-3 in
hepatocytes. Another G
i
isoform, G
i-2, is the most abundant G
subunit in most systems, and the role of
G
i-1, while unknown, may be
inhibitory to adenylyl cyclase activity and thereby mediate the
vasoconstrictor effects of ANG II. Thus we measured the mRNA for all
three isoforms of G
i.
ANG II also acts through Gq to
activate phospholipase C, thereby generating inositol
1,4,5-trisphosphate and diacylglycerol (12). One
consequence of activating this cascade is the induction of smooth
muscle contraction. With this in mind, the levels of G
q mRNA were measured.
Furthermore, since SHR may have defective interaction between
G
s-coupled vasodilatory and ANG
II-coupled vasoconstrictor pathways, the levels of mRNA for
G
s were quantified as well.
To accomplish the goal of quantifying mRNA for five different G
subunits in the same microvessel sample, we developed a quantitative polymerase chain reaction (PCR) assay for individual G protein
-subunit mRNAs (25). The assay utilizes an internal competitor that
is identical to the target sequence of interest except for a 20-base
cassette in its midportion. Thus the primer recognition areas are
identical for both target and competitor, whereas the altered
midportion allows differentiation of PCR products of the target from
that of the competitor. Quantification of PCR products is accomplished
by means of a colorimetric plate assay, the enzyme-linked oligonucleotide-sorbent assay (ELOSA), which is similar in format to an
enzyme-linked immunosorbent assay (ELISA), with the exception that a
biotin-linked oligo is used (in place of the ELISA primary antibody)
for the specific detection of complementary PCR products. We previously
demonstrated that this assay can resolve twofold differences in initial
analyte amount from 2,000 × 10
21 mol to 7 × 10
21 mol with extreme
specificity conferred at two levels: first, by specific PCR primers,
and second, by specific ELOSA capture probes. Moreover, the assay is
highly precise (11.9% intra-assay and 14.7% inter-assay coefficients
of variation), because of the use of an internal standard very similar
to the analyte.
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METHODS |
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Male SHR and WKY rats 12-14 wk of age from Taconic Farms (Germantown, NY) were housed at the University of Pittsburgh Animal Facility with controlled temperature, relative humidity, and light cycle (22°C, 55%, and 7 AM to 7 PM). Animal care was in accordance with institutional guidelines. The animals were maintained on Wayne Rodent Blox 8604 (Madison, WI). Studies had prior approval of the Institutional Animal Care and Use Committee.
Renal preglomerular microvessels were isolated using the method
described by Dubey et al. (8), with modification. Briefly, rats were
anesthetized with pentobarbital sodium, and the abdomen was opened to
expose the aorta and superior mesenteric artery. The aorta was
cannulated with polyethylene tubing (Becton Dickinson, Parsippany, NJ),
the superior mesenteric artery and proximal aorta were ligated, the
inferior vena cava was cut, and the kidneys were perfused with 20 ml of
phosphate-buffered saline (PBS) to flush out blood. Twenty milliliters
of a 1% suspension of iron oxide particles (Aldrich Chemical,
Milwaukee, WI) in PBS was infused, and the kidneys were obtained. The
medulla was removed, and the cortex was pressed through a wire mesh,
thus separating microvessels from cortical tissue. Larger vessels, if
seen, were removed using a pair of fine-pointed scissors. The
preparation was washed repeatedly in ice-cold PBS while using a magnet
to retain the iron-laden vessels. Next, the samples were digested with
collagenase at 37°C for 15-30 min to free vessels from
surrounding structures and connective tissue. After this, they were
passed through a 20-gauge needle to shear off glomeruli, then washed
repeatedly until the wash fluid was clear. Samples were then frozen at
70°C until RNA extraction was performed.
Preparation of cDNA. RNA was isolated
using a modified guanidinium phenol-chloroform Tri-Reagent extraction
method (6) with the addition of polysaccharide gel carrier (Microgel;
Molecular Research Center, Cincinnati, OH) followed by
deoxyribonuclease (DNase) digestion using ribonuclease-free
DNase (Stratagene, La Jolla, CA). Contamination with genomic DNA was
tested using 40 cycles of PCR with
Gi-2 primers, and DNase
digestion was repeated if necessary.
To reduce intersample variations in the efficiency of reverse transcription (RT), RT was carried out in at least three replicates for each sample, and the products were pooled before use in quantitative competitive-PCR. RT of total RNA was performed as described (25) using random hexamers and reverse transcriptase enzyme Superscript (GIBCO-BRL, Grand Island, NY). The procedure included an RT control to which no RNA was added; this reaction product was later subjected to PCR to verify the purity of RT reagents.
Quantitative competitive PCR assay. We developed an assay that utilizes, as an internal standard, a synthetic oligonucleotide that competes with the wild-type analyte during PCR amplification (25). For clarity, the segment of the wild-type template that is amplified by PCR is denoted the target template (TT), and the competing internal standard is termed the competitor template (CT). The CT was designed to be identical to the TT at both ends, to maximize the possibility that it would be amplified with the same efficiency and specificity as the TT and be affected in the same way by variations in reaction conditions. Differentiation of CT from TT in the PCR products is accomplished by altering a 20-base-length region in the midportion of the CT. This region carries a scrambled version of the sequence in the corresponding portion of the TT.
PCR products are quantified using a colorimetric 96-well microplate assay similar to the ELISA. PCR products are bound to streptavidin-coated plates by hybridization with biotin-labeled oligos, which serve the same function as the primary antibody in ELISAs. These oligos are complementary to the midportion of the CTs and TTs, allowing specificity of detection. PCR products bound to the plates are then incubated with an antibody that recognizes an antigen, fluorescein, that is tagged to the primers and thereby incorporated in the PCR products. The antibody is linked to an enzyme (horseradish peroxidase) that produces a colored compound from a substrate that is added in the final step. This assay is termed an ELOSA.
Competitive PCR was performed by amplifying the sample with several known dilutions of CT in parallel reactions. The amplified CT and TT were measured at the end of PCR by ELOSA. The results were quantified as ratio of CT signal to TT signal, or CT/TT. This ratio was plotted as a function of the amount of CT added, and a linear regression equation for the relation was derived using a least-squares fit. In theory, if the assay detection system is linear with respect to the amount of CT added, then, since the amount of TT is constant within a sample, the relation between CT/TT vs. amount of added CT should be linear with a y-intercept at zero, i.e., CT/TT = slope × CT or CT = CT/TT × 1/slope. Moreover, if the amplification and detection procedures are equally efficient for CT vs. TT, then, when the amount of added CT is equal to the amount of TT in the sample, the CT/TT ratio should be equal to unity (i.e., CT/TT = 1). Thus, when CT/TT = 1, then CT = TT = 1/slope. This relation would hold even when the y-intercept is nonzero, because of trace contamination of assay components by CT, since such contamination would elevate the relationship between CT/TT and added CT without changing the slope. Thus the level of TT was measured as the inverse of the slope of the relationship between CT/TT vs. added CT. We empirically confirmed the validity of this approach by extensive testing (25).
G protein primers and probes. Primers
and probes for PCR amplification (Table 1)
were selected from the published cDNA sequence for rat
Gs (13),
G
i-1,
G
i-2,
G
i-3 (16), and
G
q (28) using MacVector 4.1 software (Kodak International Biotechnologies). CT and 5' primers
were obtained from Midland Certified Reagent (Midland, TX). The
5' end labels of biotin and fluorescein were added to the capture
probes and 3' primers, respectively (Ransom Hill Biosciences,
Ramona, CA; and Promega, Madison, WI), during oligo synthesis.
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PCR. PCR conditions were optimized for each primer pair so that optimal yield of a single band product of the appropriate size was obtained. These conditions were determined for a conventional thermal block PCR cycler (Perkin-Elmer, Norwalk, CT) and a rapid air thermal cycler (model 1605; Idaho Technology, Idaho Falls, ID). Both types of cyclers yielded similar sensitivity, specificity, accuracy, and reproducibility of results; however, most of the reported data were obtained from the air thermal cycler. Optimized PCR conditions are shown in Table 1.
For the conventional cycler, a final concentration of 50 mM KCl, 10 mM tris(hydroxymethyl)aminomethane (Tris) hydrochloride (pH 8.3), 2.5 mM MgCl2, and 0.2 mM each of dATP, dGTP, dCTP, and dTTP were used. The unlabeled 5' primer and the fluorescein-labeled 3' primer (400 ng each), 10 µl of dimethyl sulfoxide, the requisite amount of cDNA, and varying amounts of CT were added and the mixture heated to 94°C for 5 min. The temperature was then reduced to 86°C, and 2.5 U of DNA polymerase were added (AmpliTaq, Perkin-Elmer, Foster City, CA). Optimal cycling conditions were determined empirically using a PCR optimizer kit (Invitrogen, San Diego, CA). PCR cycles consisted of 94°C for 1 min, annealing temperature (see Table 1) for 1 min, and 72°C for 3 min.
Reactions in the rapid cycler were performed in glass capillary tubes in a total volume of 10-12 µl. Each reaction received 40-48 ng of the 5' and 3' primers, 1-1.2 µl of 10× PCR reaction buffer [Tris 500 mM (pH 8.3), bovine serum albumin (BSA) 2.5 mg/ml, and 10, 20, or 30 mM MgCl2], 1-1.2 µl enzyme diluent (10 mM Tris and 2.5 mg/ml BSA), a final concentration of 100 mM of each dNTP, and 0.4 U of DNA polymerase. Reaction conditions were optimized using the startup kit provided by the manufacturer. Samples were denatured for 15 s to 2 min, followed by rapid cycling through 94°C, the annealing temperature (see Table 1), and 72°C, with a final extension at 72°C for 2 min.
Each set of reactions included the following three controls: 1) a negative control that had no wild-type template or CT, 2) a TT-positive control that had TT but no CT, and 3) another positive control that had CT but no TT.
In each measurement, the microvessel cDNA was amplified with five
different concentrations of CT. Briefly, reactions were performed using
12.5 to 100 ng of RNA (depending on the abundance of the specific
target) equivalent RT product. Figure 1,
which depicts assay results for all five G isoforms in the same
sample, illustrates this procedure.
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SHR and WKY samples were batched together at all stages of processing,
starting with the day of surgery until final assay by ELOSA. Absolute
quantities as measured by ELOSA were then expressed as a ratio to the
level of Gs. Not only does this
use G
s as a housekeeping
message to normalize for variability in RNA quality and RT efficiency,
but it measures possible imbalances between G
i or
G
q (mediating ANG II effects)
and G
s (mediator of several vasodilators that appear to be subnormally effective in buffering ANG
II-mediated effects in SHR).
Data analysis and statistical tests.
Quantifications of specific isoforms of G TT in each sample were
performed at least in duplicate. These values were averaged, and the
ratio of the specific TT to that of
G
s in the same sample was
calculated. The average of this ratio for each G
isoform was
determined for each strain. Statistical comparisons between SHR and WKY
samples were made using Student's
t-test for unpaired samples.
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RESULTS |
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Quantification of G protein mRNA for
Gs,
G
i-1,
G
i-2,
G
i-3, and
G
q was performed in renal
microvessels of 12- to 14-wk-old SHR and WKY rats. Levels of
G
s and
G
i-2 message were measured in
22 samples each (9 WKY and 13 SHR).
G
q was measurable in 19 samples
(7 WKY and 12 SHR), and G
i-1
and G
i-3 were measurable in 8 each (3 WKY and 5 SHR). All five subunits were quantifiable in eight
samples. These data are shown in Fig. 2.
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Gi-2 was by far the most
abundant of the G
subunits measured, and
G
q was the least. The abundance
rank order for the various isoforms was
G
i-2 >>
G
i-3
G
s
G
i-1 >>
G
q. When data were expressed as
10
18 mol (1 amol) per 50 ng
of RNA equivalent cDNA, the amount of G
i-2 (25.3 ± 6.3 amol) was
about 100-fold greater than that of G
i-3 (0.95 ± 0.08 amol),
G
s (0.83 ± 0.19 amol), or
G
i-1 (0.51 ± 0.04 amol),
and the amount of G
q was the
lowest of all the isoforms (3.45 ± 0.5 ×10
21 mol, or 3.45 ± 0.5 zmol).
No differences were detected between SHR and WKY with regard to the
levels of Gs assayed after RT
from equal amounts of RNA (0.96 ± 0.35 for WKY vs. 0.74 ± 0.25 amol for SHR; P = 0.617) (Fig. 2,
inset).
Figure 2 also shows that the ratios of the levels of
Gi-1,
G
i-2,
G
i-3, and
G
q, normalized to the level of
G
s, showed no significant
differences between SHR and WKY (0.44 ± 0.05 vs. 0.48 ± 0.13 for G
i-1, 40.9 ± 7.8 vs.
45.2 ± 8.9 for G
i-2, and 0.79 ± 0.05 vs. 0.82 ± 0.15 for
G
i-3, in SHR vs. WKY,
respectively). Study of the G
q-to-G
s
ratio suggested, however, that G
q may be more abundant
in SHR (WKY 0.41 ± 0.08 vs. SHR 1.04 ± 0.32, P = 0.08).
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DISCUSSION |
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The hypothesis that the altered renovascular sensitivity of SHR to ANG
II and/or Gs-mediated
vasodilators is associated with alterations at the G protein level was
studied in terms of quantifying steady-state mRNA levels for specific
isoforms of G
in freshly isolated renal microvessels.
Commercially available antibodies for G protein -subunits present
problems with sensitivity, specificity, and quantification issues.
These difficulties are exacerbated in the case of low-abundance proteins like G
i-1,
G
i-3, and
G
q. On the other hand, the use of PCR followed by a specific DNA capture probe allowed us to quantify
individual G
subunit messages with a high degree of sensitivity and
specificity (25). To our knowledge, this is the first report of G
protein mRNA quantifications in renal preglomerular microvessels of SHR
and WKY rats. The assay revealed no significant strain differences in
the levels of G
s or of
individual G
i subunits normalized to G
s levels. The
level of G
q normalized to
G
s, however, tended to be
higher in SHR.
The isolation procedure used for microvessels yielded vessels predominantly in the size range of 25-200 µm. The size and arborization pattern indicate that these samples are composed of afferent arterioles as well as interlobular and arcuate arteries. This range of vessel distribution was preferred in view of the study by Kost et al. (19) showing that arcuate and interlobular arteries are major sites of increased sensitivity to ANG II in SHR.
Another important consideration in the rationale for this study was
whether increased RNA levels necessarily imply an increased level of
protein. A review of the literature on SHR and G proteins suggests that
there is usually a good correlation. Anand-Srivastava (1) showed
enhanced levels of Gi and
unaltered G
s in SHR heart and
aorta and in a later report demonstrated a corresponding increase in
G
i mRNA, with no change in
G
s, in the same tissues (29).
Gurich et al. (11) showed by Northern analysis and ADP ribosylation
studies that equivalent amounts of
G
s mRNA and protein were
present in both strains. Kai et al. (17) showed in cultured aortic
vascular smooth muscle cells from Sprague-Dawley rats that prolonged
incubation with ANG II leads to a reduction of protein levels of
Gq/G11
-subunits as well as a
decrease in G
q mRNA levels.
Thus mRNA levels should provide an indication of the level of the
corresponding G proteins.
Inasmuch as the mechanism of vascular hyperreactivity to ANG II in SHR
involves either an exaggerated decrease in agonist-induced cAMP in
response to ANG II (31) or a subnormal response to
Gs-mediated vasodilators
(3-5) that act by increasing cAMP, we expected to see any or some
of the following: 1) a lower level
of G
s mRNA in SHR,
2) a higher level of any
G
i in SHR, or
3) a higher level of
G
q in SHR. In the present
study, we normalized the levels of
G
i and
G
q to the level of
G
s. Because we were exploring the imbalance between ANG II-mediated and
G
s-mediated pathways, we
reasoned that this ratio would be a sensitive indicator of such an
imbalance.
Our finding that there are similar levels of
Gs in 12- to 14-wk-old SHR and
WKY is at variance with the results of Ruan and Arendshorst (27), who
found a slight increase in the quantity of
G
s protein in 6- to 8-wk-old
SHR preglomerular vessels. Possible reasons for the discrepancy are
differences in the ages of rats in the two studies and the fact that
our samples included arcuate arteries in addition to the interlobular
and afferent arterioles studied by Ruan and Arendshorst (27). It is
important to note that studies of renal G protein expression at
different ages in SHR and WKY (24) showed no alteration of any G
protein at 3 wk, whereas
G
s-long and
G
q were reduced in 28-wk-old
SHR, with no alteration of
G
s-short or
G
i. Our findings are consistent with those of Gurich et al. (11), who demonstrated equivalent amounts
of G
s protein in renal cortex
by cholera toxin labeling and of mRNA by Northern
blotting. Measurements of
G
s protein in extrarenal
tissues have in most cases shown no differences between SHR and WKY (1,
7, 22, 23).
We found that the levels of mRNA for
Gi-1,
G
i-2, and
G
i-3, analyzed in terms of
their ratio to the level of G
s
in the same sample, were similar in both strains. Similarly, Ruan and Arendshorst (27) found no differences in
G
i-1,
G
i-2, or
G
i-3 protein in renal afferent
and interlobular arterioles. These match the findings of Michel et al.
(24) in renal membranes at various ages and McLellan et al. (22) in
renal cortical plasma membranes. In membranes from freshly isolated
mesenteric vessels (7), the levels of
G
i-2 and
G
i-3 were unaltered. Myocardial
membranes from SHR have been shown to have increased
Gi by pertussis toxin and
immunolabeling (1) and Northern analysis (29). However, other studies
have failed to demonstrate such increases in myocardial (22, 23) or
renal plasma membranes (22).
With regard to Gq, our finding
was an increased level of mRNA expression in SHR (WKY 0.41 ± 0.08 vs. SHR 1.04 ± 0.32), which, however, was not statistically
significant (P = 0.08). Ruan and Arendshorst (27) have reported no strain-related differences in
G
q protein in preglomerular
vessels from 6- to 8-wk-old rats. Renal membranes of 28-wk-old but not
3-wk-old SHR have been reported to have reduced levels of
G
q (24). Quantification of
G
q/11 in cells derived from SHR
and WKY mesenteric arteries or
G
q in myocardium (23) showed no
strain-specific differences.
In conclusion, the findings of the present study demonstrate that
Gs,
G
i, and
G
q mRNA levels in the renal
microvasculature do not differ in SHR vs. WKY rats. Furthermore, these
data suggest that the enhanced renovascular sensitivity of SHR to
pressor agents such as ANG II may not be directly related to
alterations in steady-state levels of these GTP-binding regulatory
proteins. Studies characterizing strain-specific differences in
functional responses in freshly isolated SHR and WKY renal microvessels
are currently in progress.
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ACKNOWLEDGEMENTS |
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This work was supported by National Heart, Lung, and Blood Institute Grants HL-35909 and HL-55314.
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FOOTNOTES |
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Preliminary data and a description of the assay have been published in abstract form (J. Am. Soc. Nephrol. 6: 2240, 1995).
Address for reprint requests: E. K. Jackson, Center for Clinical Pharmacology, 623 Scaife Hall, 200 Lothrop St., Pittsburgh, PA 15213-2582.
Received 11 April 1997; accepted in final form 24 July 1997.
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