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Am J Physiol Renal Physiol 296: F78-F86, 2009. First published October 22, 2008; doi:10.1152/ajprenal.90518.2008
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Angiotensin II-induced contraction is attenuated by nitric oxide in afferent arterioles from the nonclipped kidney in 2K1C

Frank Helle,1,2 Michael Hultström,1,2 Trude Skogstrand,1,2 Fredrik Palm,3 and Bjarne M. Iversen1,2

1Renal Research Group, Institute of Medicine, University of Bergen, and 2Haukeland University Hospital, Bergen, Norway; and 3Department of Medical Cell Biology, Uppsala University, Uppsala, Sweden

Submitted 26 August 2008 ; accepted in final form 16 October 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Two-kidney, one-clip (2K1C) is a model of renovascular hypertension where we previously found an exaggerated intracellular calcium (CaFormula) response to ANG II in isolated afferent arterioles (AAs) from the clipped kidney (Helle F, Vagnes OB, Iversen BM. Am J Physiol Renal Physiol 291: F140–F147, 2006). To test whether nitric oxide (NO) ameliorates the exaggerated ANG II response in 2K1C, we studied ANG II (10–7 mol/l)-induced calcium signaling and contractility with or without the NO synthase (NOS) inhibitor NG-nitro-L-arginine methyl ester (L-NAME). In AAs from the nonclipped kidney, L-NAME increased the ANG II-induced CaFormula response from 0.28 ± 0.05 to 0.55 ± 0.09 (fura 2, 340 nm/380 nm ratio) and increased contraction from 80 ± 6 to 60 ± 6% of baseline (P < 0.05). In vessels from sham and clipped kidneys, L-NAME had no effect. In diaminofluorescein-FM diacetate-loaded AAs from the nonclipped kidney, ANG II increased NO-derived fluorescence to 145 ± 34% of baseline (P < 0.05 vs. sham), but not in vessels from the sham or clipped kidney. Endothelial NOS (eNOS) mRNA and ser-1177 phosphorylation were unchanged in both kidneys from 2K1C, while eNOS protein was reduced in the clipped kidney compared with sham. Cationic amino acid transferase-1 and 2 mRNAs were increased in 2K1C, indicating increased availability of L-arginine for NO synthesis, but counteracted by decreased scavenging of the eNOS inhibitor asymmetric dimethylarginine by dimethylarginine dimethylaminohydrolase 2. In conclusion, the CaFormula and contractile responses to ANG II are blunted by NO release in the nonclipped kidney. This may protect the nonclipped kidney from the hypertension and elevated ANG II levels in 2K1C.

two-kidney; one-clip; renovascular hypertension

TWO-KIDNEY, ONE-CLIP (2K1C) has been studied as an ANG II-dependent model of renovascular hypertension. Earlier studies have demonstrated elevated circulating levels of ANG II, with high ANG II concentration in the cortical tissue of the clipped and nonclipped kidney (20).

Renal blood flow (RBF) and intrarenal vascular resistance are controlled by autocrine and paracrine factors as well as myogenic and tubuloglomerular feedback responses. The afferent arteriole (AA) is the main site for RBF and glomerular filtration rate (GFR) regulation and plays an important role during development of 2K1C hypertension (12). The clipped kidney has reduced RBF and GFR (18), impaired autoregulation, and dilated AAs (14). Although RBF autoregulation is reset to higher perfusion pressures, the nonclipped kidney has well-maintained RBF and GFR (6) despite high levels of circulating and cortical tissue ANG II.

Previously, we found that the ANG II dose-response curve in AAs from clipped kidneys did not saturate at high doses (10–6 M), while those from sham and nonclipped kidneys did (5). The effect of cyclooxygenase (COX) inhibition, together with mRNA and protein (1) expression for the AT1a receptor, was similar in the nonclipped and clipped kidneys. This indicated that the changed dose-response relationship was not regulated at the receptor level, or by prostaglandins produced by COX.

Earlier studies indicate that nitric oxide (NO) release can be induced by ANG II stimulation (23) and that NO release can buffer the ANG II-induced contraction in vivo in situations with high blood pressure (BP) and increased levels of circulating ANG II, such as in the nonclipped kidney from 2K1C (27) or during systemic ANG II infusion (10). In the present study, we hypothesized that increased endothelial NO release in the nonclipped kidney buffers the ANG II-induced contractile response and that this buffering system is active in the AA. To explore this hypothesis, we studied intracellular free calcium (CaFormula) responses, contractility, and NO release in AAs isolated from sham, nonclipped, and clipped kidneys. To study how NO production was regulated, we measured endothelial NO synthase (eNOS) mRNA and protein expressions as well as eNOS phosphorylation. Furthermore, we assessed mRNA expression of the cationic amino acid transporter 1 (CAT1) and 2 (CAT2), both of which facilitate cellular uptake of the NO precursor L-arginine, mRNA and protein for arginase 1 and 2, which break down intracellular L-arginine, and dimethylarginine dimethylaminohydrolase 2 (DDAH2), which scavenges the potent eNOS inhibitor asymmetric dimethylarginine (ADMA).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals. Ninety-three male Wistar rats, aged 8 wk, were obtained from Taconic M&B. Animals were fed ordinary rat pellets containing 0.5% sodium, 0.6% potassium, 0.71% calcium, and 14.7% crude protein. The rats had free access to tap water. The experiments were performed in accordance with and under the approval of the Norwegian State Board for Biological Experiments with Living Animals.

Experimental groups. Fifty-eight rats were assigned to the 2K1C groups and 35 rats to the sham groups. Clipping of the left renal artery and sham surgery were performed as previously described (5). In short, the renal arteries of rats, weighing 120–130 g, were clipped (0.20-mm internal diameter) during pentobarbital sodium anesthesia (50–70 mg/kg). Body weight and systolic BP were measured weekly. BP was measured after 20- to 30-min preheating at 32°C by the tail-cuff method following the instructions of the manufacturer (CODA, Kent Scientific).

Isolation and preparation of AAs for intracellular calcium, lumen diameter, and NO. AAs for CaFormula and diameter measurements were isolated from 15 sham and 33 2K1C animals using an agarose infusion/enzyme treatment technique (5) adapted from Loutzenhiser and Loutzenhiser (17). Only vessels identified as AAs based on their attachment to the glomerulus (Fig. 1A) were studied. Isolation was performed in Ca2+-free buffer as this improved the viability of the vessels. AAs for CaFormula measurements were loaded with 1.25 µmol/l fura 2 acetoxymethyl ester in Roswell Park Memorial Institute (RPMI) medium at room temperature for 45 min, and the AAs for NO measurements were loaded with 5.0 µg/ml diaminofluorescein-FM diacetate (DAF-FM DA) in RPMI at room temperature for 1 h. Before the experiments, Ca2+ concentration in the medium was increased to 2 mmol/l in three steps (20 µmol/l, 200 µmol/l, and 2 mmol/l) with a 5-min incubation between each step. Thereafter, vessels were incubated for 15 min to ensure complete deesterification of the dyes (fura 2-AM or DAF-FM DA), which were added to the perfusion bath as esters to facilitate passage across the cell membrane. CaFormula transients and luminal vessel diameters were recorded during superfusion (1–2 ml/s) in a 400-µl chamber at 36–37°C (5). Only one experiment was conducted for each arteriole. The microvessels were initially superfused with normal RPMI medium, followed by 10–7 mol/l ANG II in the same solution, and then normal medium again; all periods lasted 150 s. Inhibition of NOS was performed by adding NG-nitro-L-arginine methyl ester (L-NAME; 10–4 M) to the vessel bath for 15 min before the start of the recordings. DAF-FM-loaded vessels for NO recordings were attached to the clean glass surface of a no. 1 coverslip mounted in a petri dish (MatTek P35G-1.5–14-C) containing 3 ml medium. Vessels were placed in the warmed stage (32°C) of a PerkinElmer UltraView RS system. At t = 0 s, 300 µl of medium was aspirated from the dish, and the same volume containing 10–6 mol/l ANG II was added, producing a final concentration of 10–7 mol/l ANG II.


Figure 1
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  		Fig. 1. A: afferent arterioles (AAs, arrows) with attached glomeruli (G) and agarose protruding from the vessel lumen (arrowhead). B: AA displaying well-defined wall-lumen outlines (arrowheads). C: AA visualized under brightfield illumination in a PerkinElmer UltraView spinning-disk microscope. D: the same arteriole as in C, but visualized as a z-projection of 10 confocal images displaying diaminofluorescein-FM (DAF-FM) fluorescence (excitation 488 nm, emission >510 nm). E: single confocal image of a DAF-FM-loaded AA, displaying increased nitric oxide fluorescence along the luminal side (arrowheads).

 
Calcium signaling. CaFormula concentration was measured as fura 2 fluorescence with a PTI imaging system (5). Since the AA consists of both endothelium and smooth muscle, it also contains two independent cytosolic compartments with separate CaFormula transients. We therefore decided it would be most correct to provide fura 2 fluorescence as a 340 nm/380 nm ratio and not CaFormula concentration. The peak fura 2 ratio increase was calculated as the difference between initial and baseline ratio, and the sustained fura 2 ratio increase was calculated as the difference between sustained and baseline ratio.

Diameter measurements. The agarose used to infuse the vessels is flexible, allowing it to be compressed during vessel contraction. When compressed, the elastic intravascular agarose exerts a pressure on the vessels wall, returning it to near-baseline diameter after removal of agonists such as norepinephrine (Helle F, unpublished observations). In the present preparation, however, normalization of the vascular diameter was not seen after vasoconstriction induced by ANG II, indicating that the contractile effect of this agonist was more long lasting. Internal lumen diameter was measured from a series of images (2,576 x 1,932 pixels) taken at 5, 15, 20, 30, 45, 60, 90, 120, and 180 s after stimulation with ANG II (10–7 mol/l). The images were acquired with an UltraView IIIu CCD camera. Lumen outline (Fig. 1B, arrowheads) and length were traced using Olympus DP-Soft 5.0, and the mean lumen diameter was calculated as mean lumen diameter = lumen area/lumen length.

Recording of NO release in isolated AAs. NO was measured as DAF-FM fluorescence, using a Perkin-Elmer UltraView RS spinning-disk system. DAF-FM is a fluorescein compound that reacts irreversibly with an oxidized form of NO (N2O3). Since DAF-FM reacts with a derivate of NO, the dye does not alter the concentration of this signaling molecule in the cells. DAF-FM has a detection limit of 5 nM and does not react with other stable oxidized forms of NO (such as NOFormula) or reactive oxygen species (16). Images of DAF-FM-loaded AAs were acquired with both brightfield illumination (Fig. 1C) and as z-series stacks (Fig. 1D) of confocal images (Fig. 1E) displaying >510-nm fluorescence. After a stable baseline, vessels were stimulated with ANG II (10–7 M), and 10 pictures (972 x 957 pixels) at 10 different z-steps were collected to produce a complete projection of the vessel every 3 s. Using 480-nm excitation, 13% laser power, and no binning, >510-nm fluorescence was collected.

Quantitative real-time PCR. mRNA levels for CAT1, CAT2, arginase 1, arginase 2, and DDAH2 were measured in preglomerular vessels isolated with an iron oxide method (28) from 7 sham and 10 2K1C animals. The same three-step dilution of cDNA standard was used as a reference in all reactions. The results are expressed as the quantity of mRNA relative to 18S ribosomal RNA in the same sample. The isolated tissue was immediately transferred to RNA-later (Qiagen) and frozen at –20°C. Total RNA was extracted with an RNeasy mini-kit (Qiagen) and stored at –80°C until use. Reverse transcription was performed with an RT-core kit (Eurogentec) using random nonomers as primers. Quantitative PCR (qPCR) was performed on an ABI prism (Applied Biosystems) using a qPCR Mastermix for SYBR Green I (Eurogentec). Primers for PCR of 18S were constructed using Primer Express software (ABI), while primers for the cationic amino acid transferases (CAT1, CAT2), inducible NOS (iNOS), and eNOS were ordered as ready-made Gene expression assays (ABI). The primers used for 18S ribosomal RNA (18S) were sense 5'-agtccctgccctttgtacaca-3' and antisense 5'-gatccgagggcctcactaaac-3'.

Western blot analysis. Proteins isolated from AAs from five sham and seven 2K1C animals were extracted in lysis buffer and sonicated before centrifugation at 12,000 rpm for 10 min to remove cell debris and remaining iron oxide. The supernatants were stored at –80°C until further use. Protein was thawed and mixed with running buffer (1:4, Bio-Rad Laboratories) and heated to 95°C for 15 min. Samples were run on 12.5% Tris·HCl gels with Tris/glycine/SDS buffer. The proteins were detected, after transfer to nitrocellulose membranes, with mouse anti-eNOS (1 µg/ml, Zymed Laboratories, Invitrogen, Carlsbad, CA) and horseradish peroxidase (HRP)-conjugated secondary antibodies (goat anti-mouse, 1:5,000, Santa Cruz Biotechnology, Santa Cruz, CA), goat anti-DDAH2 (1:5,000, Abcam, Cambridge, UK) and HRP-conjugated secondary antibodies (goat anti-mouse, 1:5,000, Santa Cruz Biotechnology), and goat anti-rat arginase 2 (1:1,000, Santa Cruz Biotechnology) and HRP-conjugated secondary antibodies (donkey anti-goat, 1:5,000, Santa Cruz Biotechnology) by an ECL camera (Kodak Image station 2000, New Haven, CT). β-Actin was detected with mouse anti-rat β-actin antibody (1:10,000, Sigma-Aldrich) and secondary HRP-conjugated goat-anti mouse antibody (1:30,000, Sigma, St. Louis, MO).

Phosphorylated eNOS/eNOS immunoprecipitation method. To investigate eNOS phosphorylation (P-eNOS), eNOS was immunoprecipitated from isolated preglomerular vessels from eight sham and eight 2K1C animals. The staining for phospho-ser-1177-eNOS was normalized by the total staining for eNOS in each sample. eNOS was immunoprecipitated with a Protein G Immunoprecipitation kit (IP-50, Sigma) according to the manufacturer's instructions, using a polyclonal rabbit anti-eNOS antibody (sc-653, Santa Cruz Biotechnology). The immunoprecipitation was incubated overnight at 4°C with both the antibody and the agarose beads. The immunoprecipitate was eluted and boiled for 5 min at 95°C with sample buffer (XT Sample Buffer, Bio-Rad). The samples were directly separated by SDS-PAGE in a 12% acrylamide ready-made gel (25242, Pierce, Rockford, IL) and transferred to nitrocellulose membranes using the iBlot system (Invitrogen). eNOS was detected using a polyclonal rabbit anti-eNOS antibody (AB16301, Chemicon International), and P-eNOS was detected using monoclonal rabbit anti-phospho-ser-1177-eNOS (Cell Signaling Technology, Danvers, MA). A pooled sample was used to create a standard dilution curve for immunoprecipitated eNOS as well as P-eNOS. The results were normalized for each membrane, and the normalized values for P-eNOS were compared with the total detected eNOS in each sample.

Digital image processing. Scale bar insertions were performed with Olympus DP-Soft 5.0. DAF-FM images were acquired using minimum input grey level = 0, maximum input grey level = 255, background subtraction, and green pseudocolor in UltraView RS. Region selection and rotation of images were performed in ThumbsPlus 6. Assembly of final figures was performed in Adobe Photoshop 8.0.

Chemicals. All chemicals were from Sigma unless otherwise stated. The media for CaFormula, contractility, and NO studies contained (in g/l) 17.65 NaCl, 0.40 KCl, 0.203 MgCl2, 0.20 NaH2PO4, 1.34 HEPES, 1.0 glucose, 0.11 Na pyruvate, 0.35 CaHCO3, and 0.22 CaCl2, as well as RPMI vitamins (R7256, Sigma) and amino acids (R7131, Sigma). Media was added to 2 mmol/l CaCl2 after isolation of vessels. Lysis buffer for actin and eNOS Western blotting contained 20 mmol/l Tris·HCl, 330 mmol/l Sucrose, pH 7.3, supplemented with 1 tablet/10 ml extracting buffer of protease inhibitor cocktail, complete EDTA free (Roche Diagnostics). Blocking buffer for eNOS Western blotting contained Tris-buffered saline (137 mmol/l NaCl, 20 mmol/l Tris), 0.1% Tween 20, 0.2% iblock (Tropix), 3.08 mmol/l NaN3, and 10.72 mmol/l MgCl2*6 H2O.

Statistics. The data are presented as means ± SE. Diameter, fura 2 ratio, and DAF-FM responses were compared using one-way ANOVA with the Student-Newman-Keuls post hoc test (SigmaStat 3.1). Peak and plateau fura 2 ratio responses and contractile responses with or without L-NAME were compared using a two-sided paired t-test; n denotes the number of arterioles used. P < 0.05 was considered statistically significant.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
BP, body weight, and kidney weight. Systolic BP increased continuously after clipping of the kidney and became greater than the control group after 3 wk. Systolic BP measured the same week as isolation of renal vessels for CaFormulai, diameter, and NO measurements was 122 ± 3 mmHg in sham-operated rats (n = 15) and 152 ± 4 (n = 33) in the 2K1C animals at week 6 (P < 0.005). BP values in animals used for real-time PCR and Western blotting were similar. The left and right kidney weights were not different in sham-operated rats (1.26 ± 0.15 vs. 1.19 ± 0.02 g, respectively, P > 0.6). The nonclipped kidneys of 2K1C animals were heavier than the clipped kidneys (1.25 ± 0.06 vs. 1.08 ± 0.05 g, respectively, P < 0.001).

Calcium signaling. Mean baseline fura 2 ratios in the six groups studied were similar, ranging between 0.58 ± 0.03 and 0.77 ± 0.08 (P > 0.3). ANG II stimulation produced an initial peak followed by a smaller sustained response in all groups (P < 0.05, Fig. 2). L-NAME treatment had little effect on the CaFormula response after ANG II stimulation in AAs from sham-operated rats (Fig. 2 left). The peak fura 2 ratios were 0.36 ± 0.08 (n = 8) and 0.32 ± 0.11 (n = 7) with and without L-NAME treatment, respectively (P > 0.7). Vessels from the nonclipped kidney of 2K1C animals displayed an enhanced response to ANG II after NOS inhibition compared with sham (Fig. 2, middle). The peak fura 2 ratio increased from 0.28 ± 0.05 (n = 6) to 0.55 ± 0.09 (n = 7, P < 0.05), while the sustained response increased from 0.11 ± 0.03 to 0.34 ± 0.08 after L-NAME treatment (P < 0.05). AAs from the clipped kidney showed a strong response to ANG II in untreated vessels (0. 51 ± 0.06, n = 7), which did not increase further after L-NAME treatment (0.61 ± 0.07, n = 7). The ANG II response of AAs from the clipped kidney was significantly stronger than in AAs from the sham kidney and untreated vessels from the nonclipped kidney (P < 0.001, Fig. 2, right).


Figure 2
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  		Fig. 2. Top row: representative fura 2 traces (340 nm/380 nm) in AAs with or without NG-nitro-L-arginine methyl ester (L-NAME) after stimulation with 10–7 mol/l ANG II. Bottom row: averaged peak and plateau fura 2 responses. AAs from sham and clipped kidneys did not respond to L-NAME treatment, but AAs from the nonclipped kidney displayed an increased fura 2 ratio after L-NAME treatment. *P < 0.05 vs. corresponding sham. {dagger}P < 0.05 vs. untreated nonclipped.

 
AA contractility. The baseline diameters were 18.0 ± 3.9 (n = 17), 17.0 ± 3.6 (n = 28), and 18.3 ± 4.5 µm (n = 30) in AAs from sham, nonclipped, and clipped kidneys, respectively, and were not significantly different (P > 0.4). The diameter of AAs from sham-operated animals decreased to 88 ± 8% of baseline after ANG II stimulation (n = 8), similar to the response in L-NAME-treated vessels in which the diameter fell to 88 ± 6% of baseline (n = 9, Fig. 3, left). The diameter of AAs from the nonclipped kidney decreased to 80 ± 6% of baseline (n = 13) after ANG II. When AAs from this kidney were pretreated with L-NAME, however, vessel diameter fell to 60 ± 6% of baseline, significantly more than in untreated vessels (t = 240 s, P < 0.05 vs. untreated 2K1C, P < 0.005 vs. sham, n = 15, Fig. 3, middle). The diameter of AAs from the clipped kidney decreased to 59 ± 8% of baseline diameter after ANG II (n = 15, P < 0.005 vs. sham and untreated nonclipped AAs) and did not contract further when pretreated with L-NAME (61 ± 6% of baseline, n = 15, P > 0.001 vs. L-NAME-treated sham, Fig. 3, right).


Figure 3
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  		Fig. 3. Averaged diameter changes shown as percentage of baseline lumen diameter in AAs from 2-kidney, 1-clip hypertensive animals with or without L-NAME after stimulation with 10–7 mol/l ANG II. In the nonclipped kidney, L-NAME induced a significantly increased diameter response. *P < 0.05 vs. corresponding sham. **P < 0.005 vs. corresponding sham. {dagger}P < 0.05 vs. untreated nonclipped. {dagger}{dagger}P < 0.005 vs. untreated nonclipped.

 
NO release. The increase in DAF-FM fluorescence was usually strongest at the inside of the vessel wall (Fig. 1E, arrowheads), likely arising from the endothelium, which is consistent with eNOS tissue distribution (19). Sample images of fluorescence intensities at baseline and 800 s after ANG II are given in Fig. 4, A (0 s) and B (800 s). Typical DAF-FM tracings are given in Fig. 4C, with averaged responses in Fig. 4D. Stimulation with ANG II was only done if the baseline was stable. Tracings show that ANG II stimulation induced oscillations in NO fluorescence at the beginning of the recordings. The oscillations correspond in time with the contractions of the vessels between 0 and 200 s (Fig. 3) and are likely artifacts caused by the change in area from which the fluorescence is collected. DAF-FM fluorescence in AAs from sham (n = 21) and clipped (n = 21) kidneys was not increased by ANG II (Fig. 4D, right and left). In vessels from the nonclipped kidney, however, a steep increase in DAF-FM fluorescence was observed after ANG II stimulation, reaching 145 ± 34% after 800 s (P < 0.05 vs. sham, n = 17, Fig. 4D, middle).


Figure 4
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  		Fig. 4. Representative images of DAF-FM-loaded AAs after 10–7 mol/l ANG II administration visualized at 0 (A) and 800 s (B). C: representative nitric oxide (NO) tracings. D: averaged NO responses shown as percentage of baseline fluorescence. *P < 0.05, **P < 0.01 vs. control.

 
eNOS expression and phosphorylation. eNOS mRNA was similar in preglomerular vessels from sham (n = 13), nonclipped (n = 10), and clipped (n = 10) kidneys (Fig. 5, left). Western blot analysis showed reduced expression of eNOS protein in vessels from the clipped kidney (n = 7) compared with sham (n = 8, P < 0.05). eNOS protein from the nonclipped kidney (n = 6) was not different from sham (Fig. 5, middle). Phosphorylation of eNOS at ser-1177 was similar among groups (Fig. 5, right).


Figure 5
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  		Fig. 5. Endothelial nitric oxide synthase (eNOS) expression in preglomerular resistance vessels. Left: quantitative RT-PCR of eNOS mRNA adjusted for 18S. Middle: Western blot of eNOS protein normalized by β-actin and given as the percentage of the expression in sham. eNOS protein in the clipped kidney was significantly reduced compared with sham. Right: phosphorylation of eNOS (P-eNOS) at ser-1177 normalized by total eNOS expression. *P < 0.05 vs. sham.

 
Expression of CAT1, CAT2, arginase 1, arginase 2, DDAH2, and AT2 receptor. The mRNA expression for the L-arginine transporters CAT1 and CAT2 was increased in both kidneys from 2K1C animals (Fig. 6, A and B). mRNA expression for the L-arginine-hydrolyzing enzyme arginase 2 was increased in 2K1C compared with sham (P < 0.05, Fig. 6C), while protein expression was similar in all groups (Fig. 6D). mRNA for arginase 1 was also increased in 2K1C (P < 0.05 vs. sham), which was 0.69 ± 0.1 (n = 13), 1.21 ± 0.2 (n = 13), and 1.36 ± 0.2 (n = 10) in the sham, nonclipped, and clipped kidney, respectively. DDAH2 mRNA expression was similar in all groups (Fig. 6E), while protein expression was significantly reduced in the nonclipped kidney compared with sham (P < 0.05, Fig. 6F). ANG II AT2 receptor mRNA expression relative to 18S was 100 ± 18 (n = 7), 91 ± 21 (n = 9), and 72 ± 10% (n = 10) in vessels from the sham, nonclipped, and clipped kidney, respectively, and were not significantly different.


Figure 6
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  		Fig. 6. Protein and mRNA expression for genes involved in NO release from preglomerular vessels. A: cationic amino acid transporter 1 (CAT1) mRNA. B: cationic amino acid transporter 2 (CAT2) mRNA. C: arginase 2 mRNA. D: arginase 2 protein. E: dimethylarginine dimethylaminohydrolase 2 (DDAH2) mRNA. F: DDAH2 protein. mRNA expression was adjusted for 18S. Protein expression was normalized by β-actin and given as the percentage of the expression in sham. *P < 0.05 vs. sham. **P < 0.005 vs. sham.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The main result of the present study is that enhanced NO release buffers ANG II-induced contraction of isolated AAs from the nonclipped kidney of 2K1C hypertensive rats. Blockade of NO release with L-NAME increased ANG II-induced CaFormula signaling and contractility in AAs from the nonclipped kidney but had no effect on ANG II responses in AAs from the sham or clipped kidneys. As seen from Figs. 2 and 3, diameter and CaFormula responses in vessels from the nonclipped kidney were similar to controls. If AAs from the nonclipped kidney were treated with L-NAME, however, the vessels responded more strongly, similar to the clipped kidney. This finding indicates that the sensitivity to ANG II was increased in both kidneys from 2K1C but buffered by NO release in the nonclipped kidney. Consistent with this, NO release measured using the dye DAF-FM was increased in vessels from the nonclipped kidney only. As seen from Figs. 2 and 3, it seems that the vasoconstrictor signal induced by ANG II is enhanced in both kidneys from 2K1C but blunted in the nonclipped kidney, due to increased NO release.

Isolation of renal microvessels, as shown in Fig. 1, A and B, was achieved using an agarose infusion/enzyme treatment technique (17). CaFormula transients and diameter changes were measured simultaneously. We have found the lumen diameter measurements to be highly reproducible in this preparation. Baseline diameters are different from what we have previously reported in vivo (14), but this was probably a result of different isolation procedures. The agarose-infused AA has, in contrast to perfused models, no flow during the experiment. This was utilized in the present study to measure agonist-induced NO release alone, without interference from physiological stimuli that alter NO release, such as shear stress.

Removal of vessels from their native environment in the kidney may alter their characteristics, such as those affected by the native endocrine environment and sympathetic nervous activity. We believe that differences observed between vessels from 2K1C and sham kidneys result from different physiological conditions in vivo and are basal characteristics of the vessels that have a functional relevance in the animal.

In vivo studies have demonstrated that RBF and GFR are relatively well maintained in the nonclipped kidney, which has a high perfusion pressure (6, 18). In the clipped kidney, however, these hemodynamic parameters are deteriorated (2), despite normal perfusion pressure behind the clip (13). In other studies, however, GFR in the clipped and nonclipped kidney was similar during control conditions but lowered in the clipped kidney after inhibition of the renin-angiotensin system (8, 9). Dilated AAs (14) and reduced glomerular capillary pressure (25) indicate microvascular hypoperfusion in the clipped kidney, and a site of resistance between the clip and capillary bed has been suggested (25). Because of these observations, different levels of shear stress would be expected in the nonclipped and clipped kidneys (6, 18).

Several reports indicate a direct link between NO release and AT1a receptor stimulation. Blockade of the AT1a receptor with losartan reduced the effect of L-NAME on basal RBF in the nonclipped kidney, demonstrating that AT1a antagonism masked the effect of NO inhibition (27). In normotensive rats, Parekh and Dobrowolski (22) found that L-NAME amplified the RBF response to ANG II, indicating that ANG II generates a NOS-dependent vasodilatory stimulus simultaneously with the vasoconstrictor signal. Consistent with this, ANG II receptor antagonism was found to blunt the vasoconstrictor effect of NO blockade (21). Cervenka et al. (3) recently inhibited NOS in 2K1C mice, which resulted in a BP increase, indicating an increased dependency of NO release during renovascular hypertension. In AT1a-deficient mice, however, the BP increase induced by NOS inhibition was similar in 2K1C and sham, demonstrating that the increased NO release in wild-type 2K1Cs was dependent on a functional AT1a receptor (3). Taken together, these reports indicate that NO release is directly linked to AT1a receptor stimulation.

Two main mechanisms induce endothelial NO release in vasculature: continuous NO release stimulated by shear stress and agonist-induced NO release induced by receptor stimulation. Qiu and Baylis (24) found that NOS inhibition induced a greater increase in AA vascular resistance if ANG II and ET receptors were operative. In rats treated with ET and ANG II receptor blockers before NOS inhibition, only 25% of the increase in AA resistance was retained (24). This indicates that agonist-induced NO release is more abundant than steady-state NO release in the AA.

In sham-operated kidneys, the CaFormula and contractile responses to ANG II were unaffected by L-NAME treatment. This appears to be inconsistent with earlier reports demonstrating increased contractility to ANG II after NO inhibition in normal animals (21), which in microdissected AAs from the mouse were shown to be AT1 receptor dependent (23). In the juxtamedullary nephron preparation, arterioles remain attached to renal tissue, while the present preparation was free of surrounding tissue. Furthermore, vessels in the juxtamedullary nephron preparation and isolated perfused arterioles may have a higher basal release of NO due to perfusion-induced shear stress, which is not present in our preparation. Thus different isolation techniques and the absence of shear stress in our model may explain the different results obtained in the present and the above-mentioned studies.

Previously, we showed that both AT1a receptor mRNA (5) and protein (1) were unchanged during the established phase of 2K1C hypertension, and we proposed that altered CaFormula signaling in the nonclipped kidney was a result of vasodilatory mechanism(s) rather than receptor regulation (5). This view was strengthened in the present study, since mRNA for the AT2 receptor, which has been proposed to play a vasodilatory role in rodent pressure homeostasis (11), was similar among the groups.

We found a considerable reduction of eNOS protein in AAs from the clipped kidney, which may reduce NO release and explain the difference between the clipped and nonclipped kidney. The connection between shear stress and eNOS protein has to our knowledge not been explored in the AA, but in endothelial cell culture from conductance vessels eNOS expression is shear stress dependent (31). The clip-induced pressure drop and compensatory vasodilation (14) strongly suggest that shear stress is reduced in the clipped kidney, which could reduce the stimulus for eNOS synthesis and provide an explanation for the lowered level of eNOS in the clipped kidney. However, localization of eNOS within the endothelial cell might be more important than total protein levels (26) and complicate the estimation of NO synthesis capacity.

In conductance vessels, both eNOS protein and ser-1177 phosphorylation have been reported to increase in 2K1C compared with sham (7). In the nonclipped kidney, however, we and others found no increase in eNOS mRNA, protein, or ser-1177 phosphorylation (29). In the left ventricle of the heart, which also contain eNOS of arteriolar origin, no increase in eNOS mRNA was found in 2K1C compared with sham (15). Taken together, these data point to a differential regulation of eNOS. In conductance vessels, 2K1C hypertension induces an upregulation of eNOS protein and ser-1177 phosphorylation, causing increased NO release. In resistance vessels from the heart and kidney, however, 2K1C hypertension seems to induce no increase in eNOS protein or ser-1177 phosphorylation.

We explored other proteins relevant for NO metabolism to find an explanation for the increased NO release in the nonclipped kidney. The role of L-arginine entry in NO synthesis has been debated, but extracellular transport of L-arginine was shown to influence NO release in rat intestinal arterioles (32). AAs from 2K1C had approximately doubled CAT1 and CAT2 mRNA levels, indicating increased intracellular availability of L-arginine, which may increase the capacity of NO release in 2K1C. Arginases catalyze the conversion of L-arginine to urea and ornithine and are potential competitors with eNOS for L-arginine (30). Arginase 2 protein was similar in all groups, despite increased mRNA levels in 2K1C. It is therefore unlikely that arginases altered the availability of L-arginine locally in the AA. The amino acid ADMA is a potent inhibitor of eNOS and neuronal NOS (4) and is inactivated by the aminohydrolase DDAH2. Despite similar DDAH2 mRNA levels in all groups, protein expression in vessels from the nonclipped kidney was significantly reduced compared with sham. This indicates a reduced scavenging capacity of ADMA, which could reduce the efficiency of eNOS in the nonclipped kidney.

It is interesting to note that mRNAs for CAT1, CAT2, and arginase II were similarly upregulated in the clipped and nonclipped kidneys, despite the different hemodynamic properties of these kidneys. It is therefore reasonable to suggest that the mRNA pools for the enzymes mentioned above were regulated by a systemic signal that was independent of local hemodynamics.

In conclusion, the present study clearly demonstrates that ANG II-induced CaFormula signaling and contractility in AAs from the nonclipped kidney are buffered by NO release. A higher bioavailability of L-arginine due to increased CAT expression is a possible component of the increased capacity of NO production in 2K1C, although this may be offset by a reduction in scavenging of the eNOS inhibitor ADMA, due to a lower DDAH2 level. A lower protein level of eNOS may limit NO release in the clipped kidney and is a possible explanation for the difference in L-NAME sensitivity in the two kidneys from 2K1C.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study has been supported by grants from Western Norway Regional Health Authority funds and by the Strategic Research Program at Haukeland University Hospital. These supporters played no part in development or approval of the present manuscript.


   ACKNOWLEDGMENTS
 
PerkinElmer UltraView RS spinning disk imaging was performed at the Molecular Imaging Center (FUGE, Norwegian Research Council), University of Bergen.


   FOOTNOTES
 

Addres for reprint requests and other correspondence: B. M. Iversen, Renal Research Group, Haukeland Univ. Hospital, N-5021 Bergen, Norway (e-mail: Bjarne.Iversen{at}med.uib.no)

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.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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