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1Department of Medicine, Ottawa Hospital, and the Kidney Research Centre, Ottawa Health Research Institute, University of Ottawa, Ottawa, Ontario; and 2Manitoba Centre for Proteomics, University of Manitoba, Winnipeg, Manitoba, Canada
Submitted 21 April 2004 ; accepted in final form 27 September 2004
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ABSTRACT |
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renin-angiotensin system; angiotensin II; tubule; mass spectrometry
In addition to the systemic RAS, the intrarenal RAS is important in the regulation of renal function. All of the components required for ANG II formation are detected within the kidney, and ANG II is generated in relatively high concentrations in the proximal tubule (19, 20). In contrast, although ANG 1–7 has been detected in renal tubules by immunostaining (1, 5), generation of ANG 1–7 by renal tubules remains to be directly demonstrated. However, ACE and ACE2 have been reported to be involved in the generation of ANG 1–7 in other tissues (4, 23). In perfused, intact human heart, generation of ANG 1–7 from ANG I was blocked by ACE inhibitor (ACEI), suggesting that the ACE pathway plays a central role in the generation of ANG 1–7 (31). In human cardiac membrane preparations, ANG 1–7 production occurs in the presence of ANG II, an effect blocked by inhibition of ACE2 (30). Of interest, both ACE and ACE2 have been reported to be present in renal tubules (1, 10, 14, 18), but their roles in the generation of ANG 1–7 remain unclear.
The present studies determined the intrarenal distribution of ACE2 mRNA and protein, examined if ANG 1–7 was produced in freshly dissected rat proximal straight tubules (PSTs), and defined the role of ACE and ACE2 in the production of ANG 1–7. The results reveal that ACE2 mRNA is expressed in all nephron segments, except the medullary thick ascending limb (mTAL), with relatively high expression in PST, inner medullary collecting duct (IMCD), and vasa rectae. Furthermore, using mass spectrometry (MS), we show that ACE2 is involved in the conversion of ANG 1 to ANG 1–7 in isolated PSTs.
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MATERIALS AND METHODS |
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Semiquantitative RT-PCR analysis of ACE and ACE2 in microdissected nephron segments. RT-PCR was performed to determine mRNA expression of ACE and ACE2 in microdissected nephron segments. Total RNA from microdissected nephron segments and vasa rectae was isolated by using a commercial kit (RNeasy, Qiagen, Chatsworth, CA). RNA was treated with DNase I and then reverse transcribed (Gene Amp RNA PCR core kit, Applied Biosystems Roche Molecular System, Branchburg, NJ). To control for possible genomic DNA contamination, all experiments included a reaction in which reverse transcriptase was omitted from the transcription buffer. Negative control RT-PCRs with dissection solution but no nephron segments were also performed in parallel. PCR amplification was carried out in 50 µl of a solution containing 1.5 U ampliTaq DNA polymerase, 2 mM MgCl2, 1x PCR buffer II, and 1 µM of the sense and antisense oligonucleotide DNA primers for the cDNA of interest. The primers were designed with the mRNA sequences of rat ACE and ACE2 (GenBank) by using a primer design program (primer3, www.basic.nwu.edu/biotools/Primer3.html). For ACE, the sense primer was 5'-GCCACATCCAGTATTTCATGCAGT-3', the antisense primer was 5'-AACTGGAACTGGATGATGAAGCTGA-3', and the PCR product corresponded to bases 3,013–3,454 of the rat ACE cDNA (442-bp PCR product) (26). For ACE2, the sense primer was 5'-ggagaatgcccaaaagatga-3', the antisense primer was 5'-CGTCCAATCCTGGTTCAAGT-3', and the PCR product corresponded to bases 387–667 of the rat ACE2 cDNA (281-bp PCR product). PCR was routinely performed for 35 cycles, with a hot start at 95°C for 5 min, followed by cycles at 95°C for 30 s, 60°C for 30 s, and 72°C for 45 s, followed by extension at 72°C for 10 min. RT-PCR products were separated by gel electrophoresis on 2% agarose gels and visualized by ethidium bromide staining. The confirmation of PCR product sequence was performed by the DNA Sequencing Facility, University of Ottawa.
For the semiquantitative analysis of mRNA expression of ACE and ACE2, the fluorescence intensity of the ethidium bromide-stained RT-PCR products was analyzed by using a Kodak imaging system (Kodak EDAS 290 gel doc system, using Kodak 1D version 3.5 image analysis software). The linear range of PCR cycles was first determined, and the relative quantification of ACE and ACE2 mRNA was expressed as the ratio of fluorescence intensity of ACE or ACE2 corrected for the intensity of 
-actin PCR products (26, 28).
Immunohistochemical analysis for ACE2 in rat kidney. Kidneys were removed, cut longitudinally, and immediately placed in Zamboni's fixative (2% paraformaldehyde, 15% picric acid in PBS) for 2 h. The solution was replaced with fresh fixative and incubated at 4°C overnight. The following day, the tissue was washed with 10% sucrose in PBS. The sucrose phosphate buffer was replaced daily on each of the following 7 days. Kidneys were then embedded in paraffin, and 4-µm sections were cut. Before staining, sections were deparaffinized, and endogenous peroxidase activity was blocked by incubating the slides in 0.3% H2O2 in 100% MeOH for 30 min. To block nonspecific binding, sections were incubated at room temperature for 30 min in PBS containing 1% milk and 3% donkey serum. Sections were then incubated for 48 h at 4°C in a humidified chamber with an antibody against human ACE2 (goat, polyclonal IgG; Santa Cruz Biotechnology, Santa Cruz, CA), diluted 1:10 in PBS containing 1.5% donkey serum and 0.5% milk. After incubation with the ACE2 antibody, the slides were incubated for 60 min at room temperature in a humidified chamber with a biotinylated anti-goat IgG, diluted 1:50 in PBS as a secondary antibody. The slides were subsequently placed in 3% H2O2 for 10 min before incubation with streptavidin-horseradish peroxidase, diluted 1:50 in PBS for 30 min at room temperature in a humidified chamber. Finally, the slides were incubated with 50 µl of diaminobenzadine (BioGenex, San Ramon, CA) as substrate. The slides were counterstained with hematoxylin (Sigma), dehydrated, fixed with Permount histological mounting medium (Fisher Scientific, Ottawa, ON), and viewed with a Zeiss Axiophot microscope. To exclude nonspecific binding, experiments included a control in which the primary antibody was preincubated with a fivefold excess of immunizing peptide for 16 h at 4°C.
Western blotting.
Kidney tissue was dissected into cortex, outer medulla, and inner medulla. The tissues were immediately frozen in liquid nitrogen and stored at –80°C until use. Microdissected PSTs (500 mm) were also stored at –80°C for Western blot analysis. Frozen tissue samples were homogenized for up to 30 s with a cell disrupter in lysis buffer, containing 0.5% Nonidet P-40, 50 mM NaCl, 10 mM Tris·HCl (pH 7.4), 2 mM EDTA, 2 mM EGTA, 0.5% sodium deoxycholate, 0.1% SDS, 100 µM Na3VO4, 100 mM NaF, 30 mM sodium pyrophosphate, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, and 10 µg/ml leupeptin. Western blot analysis was performed as previously described (28). Briefly, the lysate was boiled for 10 min, followed by centrifugation at 12,000 g for 2 min to remove insoluble debris. Protein concentrations in the supernatant were determined by the Bradford method (Bio-Rad, Montreal, Quebec) by using BSA as standard. Proteins (25–100 µg) were separated on 7.5% SDS-polyacrylamide gels and transferred onto nitrocellulose membranes (Bio-Rad). The membranes were blocked with 10% skimmed milk in Tris-buffered saline (pH 7.6) containing 0.1% Tween 20 and 0.01% sodium azide, for 1 h at room temperature. The membranes were then incubated for 18 h at 4°C with 1:100 to 1:500 dilution of antibody against human ACE2 (goat polyclonal IgG, Santa Cruz Biotechnology), followed by incubation with 1:2,000–1:5,000 dilution of anti-goat secondary antibody conjugated to horseradish peroxidase (Amersham, Oakville, ON). This antibody has previously been utilized in immunoblot analysis of mouse cardiac tissues (27). Proteins were detected by enhanced chemiluminescence (Amersham) on Hyperfilm (Amersham), according to the manufacturer's instructions. Prestained standards were used as molecular weight markers (Bio-Rad). To control for protein loading, all membranes were stripped and probed with a monoclonal anti--actin antibody (mouse ascites fluid; Sigma) that recognizes the -actin protein at 
43 kDa.
Incubation of PSTs with ANG peptides. Freshly dissected PSTs (50 mm) were incubated with either ANG 1–10, ANG 1–9, or ANG 1–8 (Bachem, Torrance, CA) in 50 µl PBS at 37°C for up to 2 h. At the end of the incubation period, 5.5 µl of 10% trifluoroacetic acid (Sigma-Aldrich Canada) were added, and the samples were centrifuged at 10,000 g for 5 min at 4°C. The supernatant was stored at –80°C until the detection of ANGs by MS. To determine the role of ACE or ACE2 in the generation of ANG 1–7 by PSTs, the ACEIs captopril (7) (10–4 M; Sigma-Aldrich) or enalaprilat (17) (10–5 M; Merck Sharp & Dohme, Montreal, Quebec), or the ACE2 inhibitor DX600 (11) (10–6 M; Phoenix Pharmaceuticals, Belmont, CA) was added to the incubation solution 20 min before the addition of ANG peptides as substrates.
In separate experiments, freshly dissected rat PSTs were microperfused, essentially as described for IMCD segments, with modifications (29). Briefly, male Sprague-Dawley rats (100–150 g), fed regular chow and with unlimited access to water, were killed by decapitation, and the kidneys were rapidly removed. Coronal sections were cut and placed in a chilled perfusion solution (solution A) consisting of (in mM) 105 NaCl, 24 NaHCO3, 2 Na2HPO4, 5 KCl, 1.0 MgSO4, 1.5 CaCl2, 4 lactic acid, 5 glucose, 1 alanine, and 10 HEPES (pH 7.3). PST segments were dissected and isolated from renal cortex. Once dissected, the tubule was transferred to a thermostatically controlled perfusion chamber containing bathing solution (solution A), on the stage of an inverted microscope, and mounted on glass pipettes that suspended the tubule in the bath. The temperature of the bath was maintained at 37°C. The inner pipette contained solution A, and perfusion was initiated via a 1-ml syringe connected to PE-10 tubing that was mounted behind the taper at the tip of the perfusion pipette. The perfusate accumulated in the tip of the opposite pipette was collected (3 µl during a 2-h period). In the present study, the perfusion solution also contained ANG II (10–6 M). At the end of the protocol, the perfusion solution and collected perfusate were stored at –80°C until the assay of ANGs using MS.
Assay of ANG peptides by MS. Samples for the assay of ANG peptides were analyzed by matrix-assisted laser desorption/ionization (MALDI)-quadropole time-of-flight (QqTOF) MS, as has previously been performed for ANGs (4). Aliquots (1 µl) of each sample were mixed with an equal volume of 2,5-dihydroxybenzoic acid (160 mg/ml, in a 1:1 mixture of acetonitrile and water). One microliter of the mixture was applied to the surface of a MALDI plate. The plate was then air-dried and inserted into the sample introduction port of the self-designed QqTOF MS (16). For tandem MS sequencing, product ion spectra were acquired with the same equipment, and the spectra were collected.
Data presentation. Semiquantitative RT-PCR analysis of ACE and ACE2 mRNA expression is presented as means ± SE. Other data are presented as representative results from at least four to five individual experiments.
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RESULTS |
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90 kDa in renal cortex, outer medulla, and inner medulla, as well as in microdissected PSTs and rat cardiac tissue (Fig. 2). To further define the protein expression of ACE2 in the kidney, immunohistochemistry was performed. Similar to its mRNA distribution, ACE2 immunostaining was found in all nephron segments except mTALs (Fig. 3). ACE2 immunostaining was visualized in the cytoplasm of renal tubular cells. In addition, ACE2 staining was observed in endothelial cells of blood vessels and within Glom. Preabsorption of the primary antibody with the immunizing peptide antigen blocked any positive immunostaining in these studies.
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Absence of detectable ANG 1–7 generation by PSTs with ANG II as substrate. In cell-free systems, ANG II is converted to ANG 1–7 by ACE2 (4, 23, 30). Incubation of PSTs with ANG II as substrate (10–6 M) for up to 2 h did not lead to any detectable production of ANG 1–7. In PSTs, there was no significant decrease in the spectral peak for ANG II after 2-h incubation, indicating no significant degradation (Fig. 8A). To rule out the possibility that ACE2 might be restricted to the apical membranes of tubules, and thereby inaccessible to ANG II administered in the incubation solution, luminal microperfusion of freshly dissected PSTs was utilized. Intraluminal perfusion of ANG II (10–6 M) in isolated PSTs for up to 1 h did not lead to generation of detectable ANG 1–7 in the collected perfusate (Fig. 8B).
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DISCUSSION |
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There are several reports showing the presence of ACE2 in kidney (1, 3, 4, 9, 13, 22, 23, 27). Using immunohistochemistry, Donoghue et al. (4) and Brosnihan et al. (1) showed that ACE2 was present in rat renal tubule cells, with no significant expression in Glom. Tikellis et al. (22), however, used in situ hybridization and immunohistochemistry to demonstrate ACE2 in both rat renal tubules and Glom. Our data, consisting of RT-PCR analysis in microdissected nephron segments, immunoblots, and immunostaining of kidney slices, demonstrated that ACE2 was expressed in all nephron segments except mTAL. The immunostaining demonstrated a predominantly cytoplasmic pattern in renal tubular cells, including the proximal tubule. In diabetic mice (db/db) and their control littermates, Ye et al. (27) observed ACE2 expression in the brush border of cortical tubules, a site at which ACE expression has previously been localized (14). In contrast, Hamming et al. (9) demonstrated ACE2 immunostaining in both brush border and cytoplasm of human proximal tubular cells. The precise intracellular localization of ACE2 in proximal tubule will require further study. In this regard, ANG II concentrations are compartmentalized within the kidney, with high concentrations in both interstitial fluid and proximal tubular lumen (20). Renal endosomes also contain ACE and ANG I and ANG II (12), and it is possible that, in a similar fashion, intracellular localization of ACE2 could lead to generation of ANG 1–7, with trafficking through endosomal compartments. Thus proximal tubular ACE2 could contribute to compartmentalization of ANG 1–7 in the kidney cortex.
It has been shown that ACE and ACE2 are involved in the conversion of ANG I to ANG 1–7 (4, 23, 30, 31). To determine the enzyme pathway by which ANG 1–7 was produced in isolated PSTs, we utilized inhibitors of enzymes potentially responsible for the formation of ANG 1–7 from ANG I. The technique of MALDI-QqTOF MS demonstrated highly reproducible, distinct m/z spectra for exogenously added ANG peptides, and for peptide products generated by incubation with isolated PST segments. DX600, a specific ACE2 inhibitor (11), completely blocked production of ANG 1–7 or ANG 1–9 from ANG I. ACEIs, on the other hand, blocked conversion of ANG I or ANG 1–9 to ANG 1–7. Because DX600 did not block the conversion of ANG 1–9 to ANG 1–7, the results suggest that, in isolated proximal tubular segments, sequential conversion of ANG I to ANG 1–9 occurs via ACE2 activity, followed by conversion to ANG 1–7 by ACE. In this regard, use of recombinant ACE and ACE2 proteins has revealed their involvement in this sequential pathway for conversion of ANG I to ANG 1–7 (4, 23, 25). In addition to this pathway, studies reveal that ANG I may be converted to ANG II by ACE, followed by ACE2-mediated conversion to ANG 1–7 (4, 23, 25). In proximal tubular segments, ANG 1–7 generation was not detected with ANG II as a substrate. Furthermore, experiments provided a source of ANG II at relatively high concentrations to both the apical and basolateral membranes of proximal tubule cells, indicating that lack of detectable formation of ANG 1–7 was not likely due to inaccessibility of ANG II to ACE2, which might be expressed on these membrane surfaces. However, we cannot rule out the possibility that lower levels of ANG 1–7 were generated by incubation of proximal tubular segments with ANG II, because the MS assay was only able to detect ANG 1–7 at concentrations exceeding 1 x 10–9 M. In our experiments, we used 50 mm of tubules (0.5–1.0 mm/tubule), diluted 250-fold in 50 µl of PBS. Although concentrations of ANG II in renal interstitial fluid and proximal tubular lumen have been reported to be in the nanomolar range (20), even if similar concentrations of ANG 1–7 were produced endogenously or in the presence of ANG II as substrate, the dilution would have resulted in final concentrations below the detectable limit of our assay. Furthermore, our study only focused on proximal tubular segments, whereas we did observe expression of ACE2 in all nephron segments, except the mTAL. It is conceivable that luminal ANG II could be delivered and converted to ANG 1–7 in segments downstream of the proximal tubule, a possibility that awaits further study.
Nonetheless, failure to detect ANG 1–7 generation with incubation of segments with ANG II (in contrast to the reliable detection of ANG 1–7 when ANG I was used as substrate) suggests that proximal tubule ACE2 may have unique regulatory properties. Indeed, the role of ACE2 in the conversion of ANG II to ANG 1–7 remains controversial. In cell-free systems, isolated recombinant ACE2 protein has been shown to convert ANG II to ANG 1–7 with a catalytic efficiency 400-fold higher than its ability to convert ANG I to ANG 1–9 (25). Other studies reveal that the relative enzyme activity of ACE2 is only approximately fivefold higher with ANG II as substrate compared with the activity with ANG I as substrate (24). However, whereas ANG I is converted to ANG 1–7 in perfused rat hindlimb and in human heart (25, 31), there is currently a lack of evidence for ACE2-dependent ANG II conversion to ANG 1–7 in living cells. It is important to note that the kinetic character of ACE2 changes dramatically under different conditions. For example, in the presence of molar concentrations of chloride, recombinant ACE2 is activated with ANG I as substrate, but inhibited with ANG II as substrate (8). We, therefore, speculate that, in isolated proximal tubular segments, ACE2 may function differently in terms of enzyme activity and substrate specificity, compared with the isolated ACE2 protein. Thus the inability of ACE2 to convert ANG II to ANG 1–7 in vivo may be due to altered enzyme activity related to its conformation, intracellular localization, or cofactor requirements in living cells.
In addition to the ACE and ACE2 enzyme pathways, neutral endopeptidase (NEP) has been reported to mediate the formation of ANG 1–7 (2, 5, 6, 30). Although we did not examine the role of NEP in the generation of ANG 1–7 in the present study, our data suggest that NEP may not be an important enzyme in the generation of ANG 1–7 by freshly isolated PSTs, because inhibition of ACE or ACE2 completely blocked ANG 1–7 production with ANG I as the substrate. Indeed, it appears that enzyme pathways responsible for ANG 1–7 formation are tissue specific and vary according to the experimental model. In isolated human cardiac membranes, thiorphan, an inhibitor of NEP, blocked the conversion of ANG I to ANG 1–7, whereas enalaprilat had no effect (30). However, in perfused human heart, enalaprilat significantly inhibited the conversion of ANG I to ANG 1–7, suggesting ACE dependence (31). These varying results might be due to cell-specific enzyme pathways for ANG 1–7 generation, because the enzyme for conversion of ANG I to ANG 1–7 in isolated cardiac membrane proteins is mainly derived from cardiac myocytes, whereas, in perfused human heart, there may be contributions of enzyme activity from endothelial cells as well as myocytes. It is also possible that the properties of ACE or NEP may differ with the isolated membrane preparation compared with intact cells. Our data indicate that ACE2 and ACE are the major enzymes responsible for conversion of ANG I to ANG 1–7 in proximal tubule. However, we cannot completely exclude a role for NEP in the conversion of ANG I to ANG 1–7 in the intact proximal tubule in vivo.
In summary, the present studies show that ACE2 mRNA and protein are present in all rat nephron segments except mTAL. In freshly isolated rat PSTs, ACE and ACE2 play important roles in the generation of ANG 1–7 from ANG I. We suggest that formation of ANG 1–7 in PSTs may serve to antagonize the effects of locally produced ANG II, thereby participating in the regulation of renal tubular function.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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