Genetic disruption of atrial natriuretic peptide receptor-A alters renin and angiotensin II levels

Shang-Jin Shi, Huong T. Nguyen, Guru Dutt Sharma, L. Gabriel Navar, and Kailash N. Pandey

Department of Physiology, Tulane University School of Medicine, New Orleans, Louisiana 70112


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

We have studied cardiovascular and renal phenotypes in Npr1 (genetic determinant of natriuretic peptide receptor-A; NPRA) gene-disrupted mutant mouse model. The baseline systolic arterial pressure (SAP) in 0-copy mutant (-/-) mice (143 ± 2 mmHg) was significantly higher than in 2-copy wild-type (+/+) animals (104 ± 2 mmHg); however, the SAP in 1-copy heterozygotes (+/-) was at an intermediate value (120 ± 4 mmHg). To determine whether Npr1 gene function affects the renin-angiotensin-aldosterone system (RAAS), we measured the components of RAAS in plasma, kidney, and adrenal gland of 0-copy, 1-copy, and 2-copy male mice. Newborn (2 days after the birth) 0-copy pups showed 2.5-fold higher intrarenal renin contents compared with 2-copy wild-type counterparts (0-copy 72 ± 12 vs. 2-copy 30 ± 7 µg ANG I · mg protein-1 · h-1, respectively). The intrarenal ANG II level in 0-copy pups was also higher than in 2-copy controls (0-copy 33 ± 5 vs. 2-copy 20 ± 2 pg/mg protein, respectively). However, both young (3 wk) and adult (16 wk) 0-copy mutant mice showed a dramatic 50-80% reduction in plasma renin concentrations (PRCs) and in expression of renal renin message compared with 2-copy control animals. In contrast, the adrenal renin content and mRNA expression levels were 1.5- to 2-fold higher in 0-copy adult mice than in 2-copy animals. The results suggest that inhibition of renal and systemic RAAS is a compensatory response that prevents greater increases in elevated arterial pressures in adult NPRA null mutant mice. However, the greater renin and ANG II levels seen in 0-copy newborn pups provide evidence that the direct effect of NPRA activation on renin is an inhibitory response.

guanylyl cyclase A; gene targeting; renin-angiotensin-aldosterone system; renin activity; natriuretic peptide receptor-A gene-deficient mice


    INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

ATRIAL NATRIURETIC PEPTIDE (ANP) is a cardiac hormone that elicits natriuretic, diuretic, vasorelaxant, and anti-proliferative responses, all of which contribute to lowering arterial blood pressure (2, 7, 27, 32). Natriuretic peptides belong to a family that includes at least three endogenous hormones: ANP, brain natriuretic peptide (BNP), and C-type natriuretic peptide (CNP). Three distinct natriuretic peptide receptors have been identified (9, 30, 41). Two of these receptors containing guanylyl cyclase (GC) activity in the intracellular domain are known as natriuretic peptide receptor-A and -B (NPRA and NPRB, respectively), whereas a third receptor that lacks the GC activity and has been considered to clear natriuretic peptides from circulation is referred to as natriuretic peptide clearance receptor (NPRC). Both ANP and BNP activate NPRA, whereas CNP activates only NPRB; however, all three natriuretic peptides indiscriminately bind to NPRC. NPRA is also designated as guanylyl cyclase A (GC-A) receptor, which produces the second messenger cGMP in response to ANP binding (1, 16, 43, 47).

Experimental data suggest that the ANP/NPRA system plays an important role in blood pressure homeostasis by direct vasodilatory and natriuretic actions as well as by its ability to counteract the renin-angiotensin-aldosterone system (RAAS) (2, 32). There is evidence that the chronic hypotensive effect of ANP is partly mediated by suppression of RAAS and that ANP opposes the angiotensin (ANG) II-mediated vascular and renal effects (2, 22). In the absence of the counterregulatory effect of NPRA signaling, sensitization of arterial blood pressure may result, at least in part, from failure to appropriately overcome the effects of ANG II. Previous experimental data have lead to the notion that ANP plays an important role in regulation of renal function by its direct vasodilatory effect, natriuretic response, and antagonistic action of RAAS (27, 32). Earlier studies with ANP-deficient genetic strains of mice demonstrated that a defect in ANP synthesis can cause hypertension in homozygous mutant mice with no circulating or cardiac ANP (21). Therefore, genetic defects that reduce the activity of the natriuretic peptide system can be considered as candidate contributors to essential hypertension. Previous studies showing elevation in arterial pressures in genetic mouse models with reduced expression of ANP/NPRA system have provided strong support for a physiological role of this hormone receptor system in the regulation of arterial pressures and other physiological functions (28, 39, 40, 45).

To elucidate the compensatory role of NPRA in blood pressure homeostasis and cardiovascular regulation, we performed studies evaluating the changes in hemodynamics and RAAS components in Npr1 (genetic determinant of NPRA) gene-disrupted mutant mice. To obtain an assessment of the extent to which NPRA contributes to the regulation of the RAAS, we measured plasma renin concentrations (PRCs), tissue renin contents, and ANG II and aldosterone levels in Npr1 gene-disrupted (0-copy; -/-), heterozygous (1-copy; +/-), and wild-type (2-copy; +/+) mice. In this communication, we describe a model system in which genetic disruption of Npr1 gene leads to elevated arterial pressures associated with decreased PRC as well as decreased intrarenal renin contents, but with increased adrenal renin and plasma aldosterone levels in adult male mice lacking functional NPRA.


    METHODS
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INTRODUCTION
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Generation of mice and genotyping. Mice lacking NPRA were generated by homologous recombination in embryonic stem cells as previously described (39). Animals were bred and maintained at the animal facility of Tulane University Health Sciences Center. Animals were handled under protocols approved by the Institutional Animal Care and Use Committee. Npr1 genotypes are designated as follows: wild-type allele (+/+; 2-copy), heterozygous allele (+/-; 1-copy), and homozygous mutant allele (-/-; 0-copy). They were determined by polymerase chain reaction (PCR) analyses of DNA isolated from tail biopsies. The NPRA 0-copy, 1-copy, and 2-copy mice used in this study were littermate progenies of a mixed 129/C57Bl6 genetic background. Animals were genotyped by multiple PCR by use of primer A (5'-GCT CTC TTG TCG CCG AAT CT-3'), corresponding to a sequence 5' to the mouse Npr1 gene common to both alleles; primer B (5'-TGT CAC CAT GGT CTG ATC GC-3'), corresponding to an exon 1 sequence only present in the intact mouse allele; and primer C (5'-GCT TCC TCG TGC TTT ACG GT-3'), a sequence in the neomycin resistance cassette only present in the null allele. The PCR reaction from tail DNA included 50 mM Tris · HCl, 20 mM ammonium sulfate, 1.5 mM MgCl2, 10% DMSO, 100 µM dNTPs, 2 units of Taq DNA polymerase, and 40 nM primers. The PCR was performed by use of a 60-s denaturation step at 94°C, a 60-s annealing step at 60°C, and a 60-s extension step at 72°C, respectively, for 35 cycles, using DNA Thermal Cycler 480. PCR products were resolved on a 2% agarose gel. The endogenous band is 500 base pairs (bp), and the targeted band is 200 bp. Mice were generated from littermate crosses of (129 × C57BL/6) Npr1 heterozygote parents. Animals were provided normal chow (Purina Laboratory) and water ad libitum and maintained on a 12:12-h light/dark cycle at 25°C. In the present experiments, different age groups, including newborn pups (2 days after birth) and young (3 wk) and adult (16-24 wk) male mice, were utilized.

Assessment of baseline blood pressure. In 3- and 16-wk-old mice, the blood pressures were measured by a noninvasive computerized tail-cuff method as described previously (26). Blood pressures were calculated as the average of 6-10 sessions/day for 7 consecutive days. Statistical analyses were performed for repeated measures with age and genotypes as variables. Blood pressures were also determined by the cannulated carotid artery method as described before (5). On the day of the experiment, the animals were anesthetized with a combination of inactin (thiobutabarbital sodium salt, 100 mg/kg body wt) and ketamine (10 mg/kg body wt) delivered intraperitoneally. Supplemental dosages of anesthesia (ketamine, 5 mg/kg body wt) were administered intramuscularly. The mouse was placed on a servo-controlled surgical table that maintained body temperature at 37°C, and a tracheostomy was performed. The animals were allowed to breath humidified 95% O2-5% CO2 by placing the exterior end of the tracheal cannula inside a small plastic chamber. The left carotid artery was cannulated (PE-10 tubing connected to PE-50 tubing) for measurements of arterial pressure, recorded on a pressure transducer connected to a Grass polygraph (Grass Instrument, Quincy, MA). Blood pressure determinations were made after the 45-min equilibration period.

Blood and tissue collection. Blood was collected by cardiac puncture under CO2 anesthesia in prechilled tubes containing 5 µl of 0.25 M Na2-EDTA. Plasma was separated by centrifugation at 3,000 rpm for 20 min at 4°C and stored at -80°C until use. Animals were euthanized by a high concentration of CO2. Tissues were dissected, frozen in liquid nitrogen, and stored at -80°C until use.

Assays of plasma renin concentrations. PRC was measured using rat renin substrate as described previously (6). Renin substrate was prepared from rat plasma collected 48 h after nephrectomy. Ten microliters of plasma were incubated with 225 µl of 100 mM sodium phosphate buffer (pH 6.0) containing 1 mM diisopropyl fluorophosphate (DFP); 1 mM phenylmelthylsulfonyl fluoride (PMSF); 5 mM EDTA; 2.5 µg/ml each of leupeptin, aprotinin, and soybean trypsin inhibitor; and 10 µl of maleate generation buffer. The reaction was started by the addition of 25 µl of rat renin substrate at 37°C and continued for 40 min, according to established protocols (42). The cocktail of protease inhibitors added in the reaction mixture protected the generated ANG I as well as renin from proteolytic destruction and permitted >90% recoveries of ANG I, as tested with the exogenously added peptide. The reaction was stopped by heating in boiling water for 10 min, and the generated ANG I was determined by radioimmunoassay (RIA) kit.

As a control for interference by non-renin-like activity, parallel experiments were performed with the lysate that had been pretreated with anti-rat renin antibody. Plasma (10 µl) was incubated with 10 µl of antibody (1:1,000 dilution) in 0.1 M Tris-acetate buffer (pH 7.0) containing 0.1% bovine serum albumin (BSA) (wt/vol) for 24 h at 4°C and was allowed to react with renin substrate at 37°C as described above. Renin-like activity was estimated by subtracting ANG I generated by antibody-treated extracts from the value obtained with untreated samples.

Assays of tissue renin contents. Frozen tissues (kidney and adrenal glands) were homogenized with a Polytron (Brinkmann Instruments, Westbury, NY) at a setting of 4, two to three times for 30 s at 4°C in a buffer containing 10 mM pyrophosphate, pH 6.0, 100 mM NaCl, 1 mM PMSF, and 1 mM EDTA. The supernatants obtained by centrifugation at 40,000 g for 40 min were used for the renin assay. The protein concentration of the extracts was determined by using a protein assay kit (BioRad, Hercules, CA). For renin assays, 10 µl of extract was incubated with 255 µl of 100 mM sodium phosphate buffer (pH 6.0) containing 1 mM DFP; 1 mM PMSF; 5 mM EDTA; 2.5 µg/ml each of leupeptin, aprotinin, and soybean trypsin inhibitor; and 10 µl of maleate generation buffer as previously described (44). The reaction was started by adding 25 µl of rat renin substrate at 37°C for appropriate time periods. The generated ANG I was determined by RIA kit as described above. The optimum time course of ANG I generation for specific tissues was determined by preliminary studies.

Renin expression analysis. Total RNA was extracted by use of the RNeasy kit (Qiagen, Valencia, CA). Briefly, tissues were homogenized in lysate buffer (50 mg/1 ml) by Polytron. The homogenate was passed through a 20-gauge needle fitted to a syringe (10 times) to shear the genomic DNA. The supernatant obtained after centrifugation at 8,000 g for 5 min was mixed with 70% ethanol (1:1, vol/vol), and total RNA was isolated according to the manufacturer's protocols. RT-PCR was performed using the GeneAmp RNA PCR kit (PerkinElmer, Norwalk, CT). Total RNA (1 µg) in a 20-µl reaction mixture containing 2.5 µM primer, 5 mM MgCl2, 1× PCR buffer II (50 mM KCl and 10 mM Tris · HCl, pH 8.3), 1 mM dNTPs, 20 units of RNase inhibitor, and 50 units of reverse transcriptase was incubated at 42°C (15 min), 99°C (5 min), and 5°C (5 min), respectively. Sense (5'-GTG TCT GTG GGG TCT TCC ACC CT-3') and antisense (5'-ATG TCG GGG AGG GTG GGC ACC TG-3') primers for PCR amplification of mouse renin-1d cDNA (30) corresponded to renin gene nucleotides +9488 through +9510 and +9902 through +9924, respectively. Sense (5'-TGG AGA AGA GCT ATG AGC TGC CTG-3') and antisense (5'-GTA CCA CCA GAC AAC ACT GTG TTG-3') primers for PCR amplification of rat beta -actin cDNA corresponded to rat beta -actin gene nucleotides +2501 through +2524 and +2767 through +2790, respectively. The renin and beta -actin cDNA fragments corresponded to 185 bp and 202 bp, respectively. PCR for renin (35 cycles) and beta -actin (25 cycles) was performed using a 60-s denaturation step at 94°C, a 60-s annealing step at 55°C, and a 60-s extension step at 72°C with a final 10-min extension at 72°C, respectively. Parallel samples without reverse transcriptase served as a negative control. The reverse-transcribed cDNA was amplified in 100 µl of reaction mixture containing 2 mM MgCl2, 1× PCR buffer II (pH 8.3), 0.4 mM dNTPs, 1 µCi [alpha -32P]dCTP, 2.5 units of Taq DNA polymerase, and 0.5 µM sense and antisense primers with the use of DNA Thermal Cycler 480. Amplified cDNA was electrophoresed using 12% polyacrylamide gel. The radioactive signals in the dried gel were detected by autoradiography with the use of Hyperfilm (Amersham). The density of the band was determined with the use of the Alpha Innotech (San Leandro, CA) densitometer. The PCR product was sequenced to confirm that it corresponded to mouse renin-1d cDNA sequence (4). For each analysis, tissue samples from five animals were collected and analyzed, and each PCR was repeated three to four times.

Assays of ANG II in plasma and kidney extracts. Four hundred microliters of plasma or kidney extracts were applied to Sep-Pak cartridges (Sep-Pak C-18 column, Waters Associates), which were equilibrated with 0.1% trifluoroacetic acid (TFA). The cartridges were washed with 10 ml of 0.1% TFA-1% NaCl and were eluted with a 2-ml mixture of methanol-water-TFA (80:19.9:0.1) as previously described (35). The eluates were brought to dryness in speed-vac centrifuge. The residues were dissolved into 100 mM Tris-acetate buffer (pH 7.5) containing 2.5 mM EDTA, 1 mM PMSF, 0.02% sodium azide, and 0.1% bovine serum albumin. Immunoreactive ANG II was measured with the use of an RIA kit (Peninsula, Belmont, CA).

Aldosterone and ANP assay. Plasma aldosterone was measured using a direct RIA kit (Diagnostic Systems, Webster, TX). Briefly, 100 µl of plasma were added into antibody-coated tubes and mixed with 500 µl of 125I-labeled aldosterone. All tubes were gently shaken at 180 rpm at room temperature for 3 h, supernatant was aspirated, and 125I-aldosterone radioactivity was determined by RIA. ANP in the plasma was quantitated directly by RIA as previously described (46).

Statistical analysis. Statistical analyses were performed by factorial one-way analysis of variance (ANOVA), followed by Fisher's least-significant difference post hoc test. The results are presented as means ± SE. For all tests, significance was set at P < 0.05.


    RESULTS
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ABSTRACT
INTRODUCTION
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Body weight, kidney weight, and the ratio of body to kidney weight among the mutant (0-copy; -/-), heterozygous (1-copy; +/-), and wild-type (2-copy; +/+) mice did not differ significantly between genotypes at 2 days, 3 wk, or 16 wk of age (Table 1). The blood pressure measurements by the noninvasive computerized tail-cuff method as well as by the cannulated carotid artery method are shown in Fig. 1, A-D. Between 3 and 16 wk of age, systolic arterial pressure (as determined by the computerized tail-cuff method) was 35-40 mmHg higher in 0-copy mutant mice than in 2-copy wild-type control animals (Fig. 1, A and B). In heterozygous 1-copy mice, the systolic arterial pressure was of intermediate value between 0-copy and 2-copy mice (0-copy 143 ± 2, 1-copy 120 ± 4, and 2-copy 104 ± 2 mmHg). As determined by the cannulated carotid artery method (Fig. 1, C and D), the mean arterial pressure (MAP) was ~35 mmHg higher in 0-copy mice than in 2-copy animals (0-copy 130 ± 3 vs. 2-copy 95 ± 4 mmHg).

                              
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Table 1.   Body and kidney weights in different age groups of Npr1 mutant, heterozygous, and wild-type mice



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Fig. 1.   Systolic arterial pressures in conscious state at the age of 3 (A) and 16 wk (B) in Npr1 homozygous mutant (0-copy; n = 5), heterozygous (1-copy; n = 9), and wild-type (2-copy; n = 9) mice. Pulsatile carotid arterial pressures (C) and mean blood pressures (D) at 16 wk of age in 0-copy (n = 5) and 2-copy (n = 5) mice.

The PRCs for different genotypes of mice are presented in Fig. 2. At 3 wk of age, the PRC (in ng ANG I · ml-1 · h-1) in 0-copy mice was approximately one-half of that in 2-copy wild-type controls (0-copy 118 ± 17 vs. 2-copy 259 ± 29). However, at 16 wk of age, the PRC in 0-copy mutant mice was reduced to approximately one-fifth of that in 2-copy wild-type animals (0-copy 23 ± 3 vs. 1-copy 57 ± 9 vs. 2-copy 142 ± 19). As noted in Fig. 2 the PRC decreased with increasing age of the animals up to 16 wk of age. However, after 4 mo, PRC did not decrease significantly and remained at almost similar levels up to 24 wk of age in all three genotypes (0-copy 28 ± 2, 1-copy 56 ± 7, and 2-copy 133 ± 12). Because there was no significant difference in PRC between 16 and 24 wk, the remainder of the experiments were carried out up to 24 wk in these studies. To have an accurate assessment of the renin levels in mutant and wild-type mice, we also determined the kidney renin content (in µg ANG I · mg protein-1 · h-1) at 2 days and 3 and 16 wk of age. As shown in Fig. 3, intrarenal renin content in 0-copy newborns (2 days after birth) was almost 2.5-fold higher than in 2-copy wild-type counterparts (0-copy 72 ± 12 vs. 2-copy 30 ± 7). However, at 3 wk of age, the renal renin content was lower by approximately one-half in 0-copy mice compared with 2-copy animals (0-copy 11 ± 2 vs. 2-copy 17 ± 2). As shown in Fig. 3, the renal renin content in 1-copy heterozygous mice was measured only at 16 wk of age and was significantly lower compared with 2-copy animals (0-copy 3.2 ± 0.3 vs. 1-copy 4.2 ± 0.7 vs. 2-copy 7.5 ± 0.6).


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Fig. 2.   Determination of plasma renin concentrations at 3, 16, and 24 wk in Npr1 homozygous mutant (0-copy; n = 5), heterozygous (1-copy; n = 9), and wild-type control (2-copy; n = 9) mice. Renin concentrations were measured as described in METHODS. Bars represent the mean ± SE values of genotypes as indicated.



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Fig. 3.   Determination of kidney renin contents at 2 days and 3 and 16 wk of age in Npr1 gene-disrupted (0-copy; n = 5), heterozygous (1-copy; n = 9), and wild-type (2-copy; n = 9) mice. Renin activity was assayed as described in METHODS. Bars represent the mean ± SE values of genotypes as indicated.

To address whether the changes in PRC and renal renin contents of 0-copy, 1-copy, and 2-copy mice were due to differential renin gene expression, we also measured the renal renin mRNA contents by RT-PCR (Fig. 4, A-C). Parallel samples with no reverse transcriptase served as a negative control (data not shown). In agreement with the renal renin contents, at 2 days of birth, the ratio of renal renin mRNA expression to beta -actin mRNA was approximately twofold higher in 0-copy mutant infants compared with 2-copy wild-type controls (Fig. 4A). However, at 3 and 16 wk of age, the expression of renal renin mRNA was reduced by approximately one-half in 0-copy mice compared with 2-copy wild-type control animals (Fig. 4, B and C).


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Fig. 4.   Kidney renin mRNA expression by RT-PCR in Npr1 gene-disrupted homozygous mutants (0-copy; n = 5) and wild-type controls (2-copy; n = 5) at 2 days (A), 3 wk (B), and 16 wk (C) of age. Total RNA was isolated from newborn pups (2 days after birth) and young (3 wk) and adult (16 wk) mice, and renin expression levels were determined as described in METHODS. The exposure times for kidney renin cDNA were varied for different age groups: 2 days (6 h), 3 wk (24 h), and 16 wk (24 h). The beta -actin cDNA exposure time was 6 h in all 3 groups. A single band, either renin or beta -actin cDNA, appeared after electrophoresis. The band size only matched renin cDNA (185 bp) or beta -actin cDNA (202 bp), which we expected. Bars represent the ratios of kidney renin to beta -actin mRNAs.

To determine whether a reduction in the renal renin mRNA expression and renin activity in plasma and kidney affects the intrarenal and circulating ANG II levels, we measured ANG II concentrations in plasma and kidneys of 0-copy, 1-copy, and 2-copy mice. As shown in Fig. 5, in 0-copy newborn pups, intrarenal ANG II concentration (in pg/mg protein) was ~1.5-fold higher compared with 2-copy wild-type controls (0-copy 33 ± 5 vs. 2-copy 20 ± 2). However, at 3 and 16 wk of age, intrarenal ANG II concentrations in 0-copy mice were significantly lower compared with the levels in 2-copy control animals (Fig. 5). As shown in Fig. 5, intrarenal ANG II concentration in 1-copy heterozygous mice was between that for 0-copy and 2-copy animals (0-copy 6 ± 0.4, 1-copy 8 ± 1.2, and 2-copy 11 ± 1.1). The levels of circulating ANG II concentrations (in pg/ml) are presented in Fig. 6. At 3 wk of age, plasma ANG II levels were approximately one-half in 0-copy mice compared with 2-copy control animals (0-copy 16 ± 2 vs. 2-copy 29 ± 4). Similarly, at 16 wk of age, plasma ANG II concentration was further reduced to one-fourth in 0-copy mutant mice compared with 2-copy normal counterparts (0-copy 6 ± 1 vs. 2-copy 24 ± 8).


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Fig. 5.   Estimation of kidney ANG II contents at 2 days and 3 and 16 wk of age in Npr1 gene-disrupted (0-copy; n = 5), heterozygous (1-copy; n = 9), and wild-type (2-copy; n = 9) mice. Intrarenal ANG II concentrations in heterozygous (1-copy; n = 9) mice are shown only at 16 wk of age. Bars represent the mean ± SE values of each genotype.



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Fig. 6.   Estimation of plasma ANG II levels at 3 and 16 wk of age in Npr1 gene-disrupted (0-copy; n = 5) and wild-type (2-copy; n = 5) mice. Plasma ANG II concentration in heterozygous (1-copy; n = 9) mice is shown only at 16 wk of age. Bars represent the mean ± SE values of each genotype.

To further examine whether the extrarenal renin levels were also altered in NPRA gene-disrupted mice, we measured the tissue renin contents and renin mRNA expression levels in the adrenal glands of Npr1 gene-disrupted mutant and control mice at 16 wk of age (Fig. 7, A and B). Surprisingly, the adrenal renin content (in ng ANG I · mg protein-1 · h-1) was higher by 1.8-fold in 0-copy mice compared with 2-copy control animals (0-copy 106 ± 14 vs. 2-copy 65 ± 5). Similarly, the adrenal renin mRNA expression level in 0-copy mice was also twofold higher than in 2-copy normal counterparts (Fig. 7B). As shown in Fig. 8, the plasma aldosterone concentration (in pg/ml) in 0-copy mice was almost twofold higher than in 2-copy wild-type control mice (0-copy 245 ± 46 vs. 2-copy 116 ± 11). The data presented in Fig. 9 indicate that the plasma ANP concentration (in pmol/ml) was significantly higher in 0-copy mice compared with 2-copy animals (0-copy 1.48 ± 0.18 vs. 2-copy 0.99 ± 0.06).


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Fig. 7.   Determination of adrenal renin content (A) and renin mRNA expression by RT-PCR (B) in Npr1 gene-disrupted mutant (0-copy; n = 5), heterozygous (1-copy; n = 9), and wild-type (2-copy; n = 9) mice at 16 wk of age. Renin assay and RT-PCR were performed as described in METHODS. Bars indicate the mean ± SE values for indicated genotypes. The exposure times for kidney renin and beta -actin cDNAs were 24 and 2 h, respectively.



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Fig. 8.   Estimation of plasma aldosterone concentration in Npr1 gene-disrupted (0-copy; n = 5) and wild-type (2-copy; n = 9) mice. All determinations were carried out at 16 wk of age as described in METHODS. Bars indicate the mean ± SE values for the representative genotypes.



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Fig. 9.   Estimation of plasma atrial natriuretic peptide (ANP) concentration in Npr1 gene-disrupted (0-copy; n = 5) and wild-type (2-copy; n = 9) mice. All determinations were carried out at 16 wk of age as described in METHODS. Bars indicate the mean ± SE values for the representative genotypes.


    DISCUSSION
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ABSTRACT
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METHODS
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DISCUSSION
REFERENCES

In the present study, we have quantitated the RAAS components in Npr1 gene-disrupted homozygous mutants (0-copy), heterozygotes (1-copy), and wild-type control mice (2-copy) to determine the role of NPRA in the regulation of plasma and tissue levels of renin and ANG II at 2 days and 3 and 16 wk of age. We also examined the transcriptional adaptations of renin mRNA that occurred in the absence or presence of a reduction of Npr1 gene compared with wild-type gene function in both newborn pups and adult animals. The present study shows that in 0-copy newborn pups, renal renin content was ~200% higher compared with 2-copy wild-type controls. However, by 3 and 16 wk of age, renal renin content as well as PRC were dramatically reduced by one-half and one-fifth, respectively, in 0-copy mice compared with 2-copy control animals. Similarly, renal renin mRNA content was also higher in 0-copy newborn pups compared with 2-copy normal counterparts. However, between 3 and 16 wk of age, renal renin mRNA in 0-copy mice was significantly reduced compared with 2-copy control animals. It is evident that renin levels were higher in newborn kidneys compared with adult kidneys, indicating that intrarenal renin markedly changes during maturation as previously described (10, 17, 23). To examine whether a reduction in the renal renin mRNA expression and renin contents affected the intrarenal ANG II levels, we measured ANG II concentrations in kidney homogenates from 0-copy, 1-copy, and 2-copy animals at 2 days and 3 and 16 wk of age. Our results show that at 2 days of birth in 0-copy pups, the intrarenal ANG II concentration was 1.8-fold higher than in 2-copy normal counterparts. However, by 3 and 16 wk of age, intrarenal ANG II concentrations were reduced in 0-copy mice to levels that were ~50% of those measured in 2-copy control animals. Similarly, circulating ANG II concentration was also lower by 50 and 75% in 0-copy mice compared with 2-copy control animals at both 3 and 16 wk of age, respectively.

ANP has been shown to suppress renin and decrease blood pressures (3, 31, 38); accordingly, we anticipated that mutant mice lacking NPRA should exhibit increased renin levels in both plasma and kidney compared with wild-type control animals. Consistent with this concept, our findings indicate that at birth, the absence of the NPRA receptor allows greater renin and ANG II levels and increased renin expression compared with 2-copy control mice (P < 0.05). However, at 3-16 wk of age, both circulating and kidney renin and ANG II levels were decreased dramatically in 0-copy mice compared with 2-copy wild-type control animals. This decrease in the renin activity observed at 3 and 16 wk of age in 0-copy homozygous mutant mice could be due to the progressive elevation in arterial pressure leading to inhibition of renin synthesis and release from the kidney juxtaglomerular cells. Because the aldosterone levels were dissociated from the circulating ANG II levels and were elevated, it is also possible that the synthesis and expression of renin in 0-copy mice might be suppressed as a consequence of aldosterone-mediated sodium retention with associated volume expansion. Elevated arterial pressure would also lead to activation of baroreceptors and reflex neural inhibition of renin release in the absence of functional Npr1 gene. Therefore, at 3 wk of age, when higher arterial pressures developed in 0-copy mice, a reversal resulted with lower plasma and renal renin contents as well as lower renal renin mRNA expression. Our present findings suggest that the elevated blood pressure overcomes the abnormal regulation of renin contents that occurs in the absence of functional Npr1 gene. Similarly, ANG II concentrations in both plasma and kidney were also significantly reduced in 0-copy mice compared with 2-copy control animals. These present observations are consistent with the evidence for baroreceptor mechanisms affecting renin synthesis and release in the kidney, which largely contributes to the regulation of PRC levels (15, 19).

An intriguing finding was that 0-copy hypertensive mutant mice showed increased adrenal renin activity, mRNA expression, and aldosterone levels. Renin expression in the extrarenal tissues seems to be evolutionarily conserved, and its tissue-specific activation is considered functionally important (49-51). These present results indicate that in 0-copy adult mice, despite an elevated arterial pressure, the adrenal renin level can remain elevated, suggesting its independent role from circulating renin. It is plausible that adrenal ANG II might be greatly elevated in 0-copy mice compared with wild-type animals. Interestingly enough, it has been previously demonstrated that the adrenal ANG II level was greatly increased in contrast to renal or plasma ANG II levels in ANG II-infused hypertensive rat model with elevated arterial pressures (56). Indeed, it would be interesting to know whether the adrenal renin levels in young 0-copy mice as well as the ANG II levels in both young and adult mutant mice are also altered. However, in these present studies, small organ size precluded us from these determinations. It has been previously suggested that the expression of renin gene in the adrenal glands is regulated by dietary salt (20, 29, 35); however, the factors that modulate renin gene expression in the adrenal glands have not been clearly identified. Markedly elevated renin protein levels in the adrenal glands have also been reported in spontaneously hypertensive (SHR) and stroke-prone SHR rats as well as in renin transgenic rats, a monogenic hypertensive model (25, 34, 36). These hypertension models may also be useful for interpreting our present results, suggesting that in hypertension, both the kidney and circulatory renin concentrations are decreased; however, as a compensatory event, the adrenal renin is increased. Previous studies have suggested that the synthesis and release of renin in kidney are controlled by both physiological and genetic determinants. Some of the well-known factors that affect renin levels include salt concentrations sensed by macula densa, arterial pressure sensed by baroreceptors in the renal vasculature, sympathetic nerve activity affecting the beta -adrenergic receptors, angiotensin AT1 receptors in the juxtaglomerular cells, and transcriptional activators in response to both endocrine and/or autocrine factors (19, 24, 37, 48, 52-56). In light of such previous observations, the present results indicate that the ANP/NPRA system may play a key regulatory role in the synthesis and maintenance of both systemic and tissue levels of RAAS components in physiological and pathological conditions.

Earlier studies have indicated that mice lacking the ANP gene and kept on a high-salt diet (8% NaCl) showed hypertension with an increased arterial pressure of 22 mmHg. These findings suggested that genetically reduced production of ANP can lead to salt-sensitive hypertension (21). On the other hand, NPRA gene-deficient mice generated by Lopez et al. (28) did not show salt-sensitive hypertension. On the contrary, NPRA gene-deficient mice generated by Oliver et al. (39), used in the present studies, exhibited a higher MAP on a high-salt diet compared with animals kept on medium- or low-salt diet. We were intrigued by the finding that a decreased expression of NPRA in 0-copy and 1-copy mice provoked salt-sensitive increases in blood pressures, whereas an increased expression of NPRA in 3-copy and 4-copy mice was able to lower the blood pressure and protect against high dietary salt intake (40). Indeed, more experimentation is needed to clarify the role of the ANP/NPRA system in establishing a relationship between salt sensitivity and blood pressure. The above-noted discrepancy suggests that the manifestation of salt sensitivity on arterial blood pressure in both ANP gene-deficient and NPRA gene-targeted mice is probably determined by the duration of increased salt intake as well as the state of hypertension in the mutant animals.

Recent findings have suggested that aldosterone contributes to elevated arterial pressures in patients with low plasma renin activity exhibiting essential hypertension (11). It has also been reported that patients with low renin and essential hypertension who consumed a normal salt diet exhibited an augmented aldosterone response to endogenous renin angiotensin as well as to infused ANG II (8, 18). On the contrary, however, on a low-salt diet, the low-renin hypertensive patients showed a blunted aldosterone response to infused ANG II (13, 14). There is the implication that the augmented aldosterone response to ANG II on a normal salt diet may be more relevant to the regulation of aldosterone secretion under normal salt intake. Patients with idiopathetic primary hyperaldosteronism also have shown enhanced aldosterone responses to ANG II (53). It has been suggested that idiopathetic hyperaldosteronism and low renin with essential hypertension are not distinct clinical entities, but they seem to be pathologically related and differ only in the level of aldosterone production. Consistent with this hypothesis, the polymorphism of the aldosterone synthase gene has been observed in patients with idiopathic primary aldosteronism with low renin and essential hypertension, where aldosterone seems to contribute the elevated arterial pressures (11, 12, 33). In light of these previous experimental findings, we speculate that in 0-copy mice, the elevated plasma aldosterone may be a contributing factor for elevated arterial pressure, despite suppressed renin and ANG II in both plasma and kidney compared with 2-copy wild-type mice.

In summary, our results presented herein show that renin and ANG II levels are significantly elevated at birth in NPRA-deficient pups; however, in adult male mutant mice, renal and systemic renin levels are reduced although adrenal renin contents remain increased. These results suggest that inhibition of renal and systemic renin and ANG II levels is a compensatory response linked to the increased arterial pressures; however, the adrenal renin is not affected by high blood pressure. The data support the notion that NPRA might differentially regulate adrenal and renal renin and ANG II levels, resulting in differential absorption of salt and water in kidney, which might be critical for the development of hypertension in NPRA null mice. It is expected that further comparative analyses of the biochemical, physiological, and pathophysiological phenotypes of Npr1 gene-deficient mice will have tremendous potentials for the assessment of the fundamental questions related to the biological importance of the interaction of NPRA with RAAS in control of blood pressure and cardiovascular homeostasis.


    ACKNOWLEDGEMENTS

We are indebted to Oliver Smithies and Joshua Knowles for providing the initial animal breeding pairs, stimulating discussion, and critical reading of this manuscript. We are grateful to Tadashi Inagami for a gift of renin antibody. Our thanks are also due to Lynell Dupepe and Paul J. Spurlock, Jr., for excellent help in animal surgery. We also thank Nicole Hunt and Jing-Shi Shen for laborious assistance in sample preparations. The expert administrative assistance of Bridget Harbor is sincerely acknowledged.


    FOOTNOTES

This work was supported by National Heart, Lung, and Blood Institute Grants HL-62147 (to K. N. Pandey) and HL-26371 (to L. G. Navar).

Address for reprint requests and other correspondence: K. N. Pandey, Dept. of Physiology SL39, Tulane Univ. School of Medicine, New Orleans, LA 70112 (E-mail: kpandey{at}tulane.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.

Received 19 December 2000; accepted in final form 21 May 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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