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Am J Physiol Heart Circ Physiol 294: H1058-H1066, 2008. First published December 21, 2007; doi:10.1152/ajpheart.01103.2007
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Role of asymmetric dimethylarginine for angiotensin II-induced target organ damage in mice

Johannes Jacobi,1 Renke Maas,2 Nada Cordasic,1 Kilian Koch,1 Roland E. Schmieder,1 Rainer H. Böger,2 and Karl F. Hilgers1

1Department of Nephrology and Hypertension, Friedrich Alexander University Erlangen-Nuremberg, Erlangen; and 2Clinical Pharmacology Unit, Institute for Experimental and Clinical Pharmacology and Toxicology, University Hospital Hamburg-Eppendorf, Hamburg, Germany

Submitted 22 September 2007 ; accepted in final form 19 December 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The aim of the present study was to investigate the role of the endogenous nitric oxide synthase inhibitor asymmetric dimethylarginine (ADMA) and its degrading enzyme dimethylarginine dimethylaminohydrolase (DDAH) in angiotensin II (ANG II)-induced hypertension and target organ damage in mice. Mice transgenic for the human DDAH1 gene (TG) and wild-type (WT) mice (each, n = 28) were treated with 1.0 µg·kg–1·min–1 ANG II, 3.0 µg·kg–1·min–1 ANG II, or phosphate-buffered saline over 4 wk via osmotic minipumps. Blood pressure, as measured by tail cuff, was elevated to the same degree in TG and WT mice. Plasma levels of ADMA were lower in TG than WT mice and were not affected after 4 wk by either dose of ANG II in both TG and WT animals. Oxidative stress within the wall of the aorta, measured by fluorescence microscopy using the dye dihydroethidium, was significantly reduced in TG mice. ANG II-induced glomerulosclerosis was similar between WT and TG mice, whereas renal interstitial fibrosis was significantly reduced in TG compared with WT animals. Renal mRNA expression of protein arginine methyltransferase (PRMT)1 and DDAH2 increased during the infusion of ANG II, whereas PRMT3 and endogenous mouse DDAH1 expression remained unaltered. Chronic infusion of ANG II in mice has no effect on the plasma levels of ADMA after 4 wk. However, an overexpression of DDAH1 alleviates ANG II-induced renal interstitial fibrosis and vascular oxidative stress, suggesting a blood pressure-independent effect of ADMA on ANG II-induced target organ damage.

dimethylarginine dimethylaminohydrolase; hypertension; transgenic

ASYMMETRIC DIMETHYLARGININE (ADMA), an endogenous inhibitor of nitric oxide synthase (NOS), is increasingly recognized as a potential risk factor and prognostic biomarker in cardiovascular disease (38). Thus far, elevated ADMA levels have been associated with all established cardiovascular risk factors (38). Furthermore, recent data from genetic mouse models indicate a causal role for ADMA in the pathophysiology of vascular disease (7, 19). ADMA derives from the posttranslational methylation of L-arginine residues within proteins catalyzed by enzymes called protein arginine methyltransferases (PRMTs). Upon proteolysis, methylarginines are released into the circulation. Although ~15% of ADMA are excreted via the urine, the major elimination occurs through enzymatic degradation (~85%) (1). The enzyme dimethylarginine dimethylaminohydrolase (DDAH), of which two isoforms with distinct tissue distribution have been described, catalyzes the hydrolysis of ADMA to L-citrulline and dimethylamine (38).

Thus far, clinical studies have mostly addressed the prognostic value of ADMA in cardiovascular or kidney disease as well as the impact of pharmacological interventions aimed at lowering ADMA levels under such conditions. Pharmacological agents that have been tested for their ability to modify ADMA levels in humans include drugs that interfere with the renin-angiotensin system (RAS) by either blocking the formation [angiotensin-converting enzyme (ACE) inhibitors] or pharmacological action [angiotensin receptor blockers (ARBs)] of angiotensin II (ANG II). Although some investigators found significant reductions of ADMA levels in patients treated with RAS-blocking agents (2, 4, 9, 15, 16, 27, 42), other studies have failed to confirm these findings (10, 40). Sound interpretation of the aforementioned clinical trials is complicated due to differences in study design, study cohorts, treatment regimen, and the use of different bioanalytical methods to determine ADMA levels. Recent in vitro studies indicate that short-term incubation (24 h) of human umbilical vein endothelial cells with ANG II (1 µM) more than doubles ADMA levels in the conditioned medium, whereas coincubation with either losartan or L-arginine almost completely abolished these effects (5).

Upon completion of our studies, Hasegawa et al. (13) reported that ANG II infusions in wild-type (WT) mice (1.0 µg·kg–1·min–1 over 2 wk) elicited a 100% increase in plasma ADMA levels, whereas in DDAH2 transgenic mice this increase was only ~50%.

Here we investigated the effects of ANG II infusion on ADMA levels in vivo in WT animals and DDAH1 transgenic (TG) mice that overexpress the human isoform of this enzyme (7). We hypothesized that ANG II-induced hypertension and target organ damage are associated with an elevation of plasma ADMA levels and that, if this holds true, mice overexpressing DDAH should be, in part, protected from ANG II-induced target organ damage.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals

DDAH1 TG mice (C57Bl/6J background) were kindly provided by John Cooke (Dept. of Cardiovascular Medicine, Stanford University). After acclimatization, animals were transferred to our transgenic core facility to rederive the colony using embryo transfer. Offsprings were genotyped by PCR of DNA obtained from tail-snip biopsies using transgene-specific oligonucleotide primers as described earlier (7).

All experiments were conducted in male, 4-mo-old, heterozygous DDAH1 TG mice, and age and weight matched WT littermates housed in a temperature-controlled animal facility with a 12-h:12-h light-dark cycle and free access to tap water and rodent chow. The study protocol was approved by the Animal Research Ethics Committee of the local government (Bezirksregierung Mittelfranken, AZ 54-2531.31-1/06).

ANG II Infusion

ANG II (Bachem) at a dose of 1.0 µg·kg–1·min–1 (each, n = 13) was delivered over 4 wk via unprimed osmotic minipumps (model 2004; Alzet) that were subcutaneously implanted into the back of WT or TG mice. The dose of 1.0 µg·kg–1·min–1 ANG II is most commonly used in rodents and causes relevant elevations of blood pressure as measured by tail-cuff plethysmography or telemetric devices (13, 41).

Based on our initial results, we decided to also study a higher dose of ANG II, namely 3.0 µg·kg–1·min–1 (WT, n = 9; and TG, n = 10). For control experiments, WT or TG mice were treated with phosphate-buffered saline (PBS) solution delivered via osmotic minipumps (each, n = 7).

Blood Pressure Measurements

Noninvasive blood pressure was measured by tail-cuff plethysmography (TSE 209000; TSE Systems) as previously described (11). Briefly, all animals were trained on alternate days over a period of 2 wk to get accustomed to the device. Final measurements were performed before pump implantation and on days 2, 3, 7, 14, 21, and 28 after surgery. A total of 20 consecutive readings of systolic blood pressure (SBP) and heart rate were recorded and averaged.

Metabolic Cages

Before pump implantation and 3 days before being euthanized, animals were housed in metabolic cages over 24 h for urine collection. Body weight, food intake, water consumption, and urine output were monitored. Albuminuria was measured and expressed as urinary albumine excretion per gram creatinine.

Hormone Measurements

Plasma levels of aldosterone were measured by radioimmunoassay as previously described in detail (14). Samples were run in duplicate, and mean values were computed.

Measurement of ADMA

Plasma ADMA levels were measured with an enzyme-linked immunosorbent assay (DLD Diagnostika) (32). Optical density was measured at 450 nm using a microtiter plate reader (Tecan Sunrise). All samples were measured in duplicate, and mean values were computed. The intraassay coefficient of variation for ADMA was 8.7%. In mice treated with 3.0 µg·kg–1·min–1 ANG II, ADMA levels were measured before and after treatment both by ELISA as well as by liquid chromatography-tandem mass spectrometry (LC-MS) as described elsewhere (33). The interassay accuracy and precision for the determination of ADMA, symmetric dimethlyarginine (SDMA), and L-arginine in plasma using spiked and native samples were in the range of –0.6–1.8% and –4.4–4.8%, respectively, for all analytes.

Baseline blood samples were obtained via retroorbital phlebotomy under inhalation anesthesia with 2% isoflurane, and at death animals were exsanguinated via arterial lines placed in the carotid artery under inhalation anesthesia as described earlier (11). To avoid the possible degradation of ADMA through blood cell components, all blood samples were immediately centrifuged and plasma was stored at –20°C until further processed.

Histomorphology and Immunohistochemistry

The degree of glomerulosclerosis was determined in methylcarnoy-fixed periodic acid Schiff (PAS)-stained paraffin sections (2 µm) using a previously described semiquantitative scoring system ranging from 0 to 4 (12). Scoring was performed in a blinded fashion, and a total of 100 glomeruli were analyzed. Based on only mild signs of glomerulosclerosis in ANG II-treated animals (score 0 to 2), the results were expressed as percentages of injured glomeruli. The expansion of interstitial collagen I (1:1,000; Biogenesis) was measured using a Leitz Aristoplan microscope (Leica) equipped with an integrated ocular grid (10 x 10 fields). A positive staining within 100 fields was counted in 20 nonoverlapping cortical views excluding glomeruli and expressed as a percentage. Glomerular collagen IV staining (1:500; Southern Biotechnology) was measured using image analysis software (MetaVue version 4.6r9; Molecular Devices) in 30 randomly chosen glomeruli per cross section. The stained area was expressed relative to the total area of the glomerular tuft. Macrophage infiltration (F4/80, clone CI:A3-1, 1:100; Serotec) within kidneys (cortical fields) was analyzed using light microscopy and immunofluorescence, and heat-induced antigen retrieval in citrate buffer was used before incubation. A biotinylated (light microscopy) or Alexa 555-labeled (immunofluorescence) goat anti-rat IgG was used as secondary antibody. Macrophages were counted in 20 randomly chosen high-power fields.

Frozen cross sections of the descending aorta were incubated with monocyte/macrophage antibody (MOMA)-2 antibody (BMA Biomedicals), followed by Alexa 555-labeled anti-rat IgG secondary antibody (Molecular Probes). Sections of the descending aorta were also probed with a polyclonal inducible NOS (iNOS) antibody (LabVision), followed by incubation with a biotinylated secondary antibody and subsequent development using a streptavidin horseradish peroxidase and diaminobenzidine detection kit (Linaris). Controls included the omission of primary antibodies. The area-based wall-to-lumen ratio was determined in PAS-stained cross sections of the descending aorta using image analysis software (MetaVue).

Oxidative Fluorescence Microscopy

In situ production of reactive oxygen species (ROS) was assessed by fluorescence microscopy using the fluorescent dye dihydroethidium (DHE; Molecular Probes). In the presence of superoxide, DHE is rapidly oxidized to ethidium bromide, which gets trapped by intercalation with DNA. Ethidium bromide is excited at 488 nm with an emission spectrum of 610 nm. Unfixed frozen tissue sections of the ascending and descending aorta (10 µm) were incubated with DHE (8 µM) in a humidified, light-protected chamber at 37°C for 1 h. In some studies, slides were preincubated with 200 U/ml polyethylene glycol-SOD (Sigma) to verify specificity of the fluorescent dye. Images were taken by use of a Spot RT digital camera (Diagnostic Instruments) that was connected to a fluorescence microscope (Nikon Eclipse 80i; Nikon). Identical image acquisition setup was used for all slides. Fluorescence signal intensities were analyzed using image analysis software (MetaVue). To account for autofluorescence, images were thresholded and fluorescent pixels were expressed relative to the pixel count of the entire vessel circumference.

Real-Time RT-PCR Detection of mRNA

In mice infused with PBS or 1.0 µg·kg–1·min–1 ANG II, the RNA of renal cortical tissue was extracted with TriFast reagent (Peqlab) (6). First-strand cDNA was synthesized with TaqMan RT reagents (Applied Biosystems) using random hexamers. Polymerase chain reaction was performed with an ABI Prism 7000 sequence detector and SYBR green reagents (Applied Biosystems) according to the manufacturer's instructions. All samples were run in duplicate. The amount of the specific mRNA of interest was normalized to 18S rRNA and expressed as relative copies. Dissociation curves were performed to confirm the specificity of the polymerase chain reaction. Primers were designed using Primer Express software (version 2.0), and selected sequences were subjected to the National Center for Biotechnical Information basic local alignment search tool (BLAST) search. Primer sequences for mouse DDAH1, DDAH2, PRMT1, PRMT3, and collagen I were as follows: DDAH1, forward, CATGTCTTGCTGCACCGAAC, and reverse, GACCTTTGCGCTTTCTGG; DDAH2, forward, GGTTGATGGAGTGCGTAAAGC, and reverse, TCCACAATTCGGAGTCCCAA; PRMT1, forward, AATGGGATGAGCCTCCAGC, and reverse, TGCTTGGCCACAGGAAACTT; PRMT3, forward, TTACCCTGAGAACCACAAAGACG, and reverse, AGTACCCAGCAACTGCCGTG; and collagen I, forward, TCACCTACAGCACCCTTGTGG, and reverse, CCCAAGTTCCGGTGTGACTC. The DDAH1 primers were designed to recognize the endogenous mouse DDAH1 sequence but not the human DDAH1 transgene.

Statistical Analysis

Statistical analysis was performed by use of a SPSS software package (version 14.0). Figures and results are given as means ± SE; n equals the number of animals. For boxplot figures, the lower and upper bounds of the boxes indicate the 25th and 75th percentile values, respectively, and the horizontal lines indicate the 50% percentile (i.e., the median). Pearson correlation coefficients were calculated when indicated.

For statistical analysis, a general linear univariate model was applied using genotype (WT vs. TG) and treatment (PBS vs. 1.0 µg·kg–1·min–1 ANG II vs. 3.0 µg·kg–1·min–1 ANG II) as fixed factors. The Bonferroni post hoc test was used to account for multiple comparisons. Statistical significance was accepted at a treatment- or genotype-related P value of P < 0.05.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Animals

Three animals treated with ANG II (n = 1 WT in the 1.0 µg·kg–1·min–1 ANG II group, and n = 2 TG in the 3.0 µg·kg–1·min–1 ANG II group) died following pump implantation. The WT animal died 2 wk after pump implantation, and necropsy revealed no major abnormalities, i.e., no signs of infection or infarction. The two TG mice died 4 and 6 days after the initiation of drug infusion, and necropsy revealed intrathoracic hemorrhage. In PBS-treated mice, a significant increase in body weight (corrected for osmotic minipump) was observed [WT, 2.4 ± 0.4 g, P = 0.001; and TG, 1.2 ± 0.4 g, P = 0.024; WT vs. TG, P = not significant (NS)], whereas the infusion of ANG II was associated with weight loss (1.0 µg·kg–1·min–1 ANG II WT, –1.9 ± 0.6 g, P = 0.013; TG, –0.6 ± 0.6 g, P = NS; WT vs. TG, P = NS; 3.0 µg·kg–1·min–1 ANG II WT, –2.4 ± 0.4 g, P = 0.0003; TG, –2.3 ± 0.6 g, P = 0.005; WT vs. TG, P = NS).

Systemic Effects of ANG II Infusion

Blood pressure and heart rate. Baseline tail-cuff SBP did not differ between WT and TG mice (119.6 ± 2.6 vs. 113.2 ± 1.9 mmHg, each n = 28, P = NS). Infusion of ANG II at a dose of 1.0 or 3.0 µg·kg–1·min–1 caused a rapid, sustained increase of blood pressure to a similar extent in both WT and TG mice (Fig. 1). Interestingly, neither SBP before death (r = 0.016, P = NS) nor the increase in SBP from baseline (r = –0.17, P = NS) correlated with plasma ADMA levels.


Figure 1
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  		Fig. 1. Chronic infusion of ANG II and blood pressure in wild-type (WT) and human dimethylarginine dimethylaminohydrolase (DDAH)1 gene transgenic (TG) mice. SBP, systolic blood pressure.

 
Treatment with ANG II was associated with a decrease in heart rate, and this effect was significant after 2 wk of treatment (WT mice, PBS vs. 1.0 µg·kg–1·min–1 ANG II vs. 3.0 µg·kg–1·min–1 ANG II, 646 ± 13 vs. 537 ± 15 vs. 530 ± 19 beats/min, P < 0.001; and TG mice, PBS vs. 1.0 µg·kg–1·min–1 ANG II vs. 3.0 µg·kg–1·min–1 ANG II, 629 ± 15 vs. 574 ± 14 vs. 541 ± 31 beats/min, P < 0.001). All heart rate data obtained during the entire period of ANG II infusion were evaluated together in a repeated-measures analysis, and significant treatment (P = 0.003) and genotype (P = 0.013) effects were observed.

Metabolic cages. Baseline urine output (48 ± 3 µl·g–1·day–1, n = 56) and albuminuria (195 ± 13 mg/g creatinine, n = 56) did not differ between treatment groups and genotypes. The infusion of ANG II caused marked diuresis in both WT and TG mice (Table 1). Similarly, a significant increase in albuminuria was noted in mice treated with ANG II (Table 1). In PBS-treated animals, no significant change in urine output and urinary albumine excretion was observed.


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  		Table 1. Target organ damage, hormone measurements, and plasma ADMA levels under chronic infusion of ANG II in WT and TG mice

 
Heart weight and vascular hypertrophy. The infusion of ANG II was also associated with myocardial and vascular hypertrophy as indicated by an increase in total wet heart weight and an increase in the wall-to-lumen ratio of the descending aorta (Table 1).

By use of immunofluorescence, scarce macrophage infiltration as indicated by MOMA-2 staining was noted, and the staining was restricted to the adventitia and the perivascular tissue, whereas the intima and media were largely devoid of macrophages (data not shown). Immunostaining for iNOS was barely detectable in PBS-treated mice, whereas a marked and dose-dependent expression was observed in ANG II-treated mice. The staining pattern and intensity did not differ between WT and TG mice (data not shown).

Plasma Levels of Aldosterone

The infusion of ANG II at 1.0 or 3.0 µg·kg–1·min–1 caused a marked and dose-dependent increase in plasma aldosterone levels in WT and TG mice (P < 0.001; Table 1).

ANG II Infusion and Plasma ADMA Levels

Despite profound hemodynamic effects and signs of target organ damage, the chronic infusion of ANG II had no impact on plasma levels of ADMA (Table 1). As expected, TG mice exhibited markedly lower ADMA levels (~37% lower than in WT mice) with virtually no overlap compared with WT animals (Table 1). Plasma aldosterone levels did not correlate with plasma ADMA levels (r = –0.22, P = NS). The lack of effect of ANG II infusion on plasma ADMA levels was corroborated in mice infused with 3.0 µg·kg–1·min–1 ANG II, in which ADMA levels were measured by ELISA and LC-MS both at baseline and after treatment (Fig. 2). There was an excellent correlation between ADMA levels obtained by ELISA versus LC-MS (r = 0.95, P = 0.0006; Fig. 3). In agreement with previous observations (22, 32), ELISA-derived ADMA levels were higher compared with those from LC-MS; the mean difference was 0.12 µM or 17% (0.685 vs. 0.569 µM, P = 0.008). However, a comparison of both methods in a Bland-Altman plot in which the mean of both methods (x-axis) was plotted against the difference of both methods (y-axis) demonstrated that values were within the limits of agreement (values are means ± 2 SD; data not shown). In addition, the concordance correlation coefficient (CCC), which evaluates the accuracy and precision between two measures based on the expected value of the squared distance function (20), suggested reasonably strong concordance (CCC = 0.680, and 95% lower confidence boundary = 0.514).


Figure 2
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  		Fig. 2. ELISA- and liquid chromatography-tandem mass spectrometry (LC-MS)-derived asymmetric dimethylarginine (ADMA) levels in mice under high dose of ANG II (3.0 µg·kg–1·min–1).

 

Figure 3
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  		Fig. 3. Correlation of ELISA- and LC-MS-derived ADMA levels in mice following high-dose ANG II infusions (3.0 µg·kg–1·min–1). n, Number of animals.

 
By means of the ELISA assay, a small increase of plasma ADMA levels was observed in WT mice. However, this finding could not be confirmed by LC-MS (Fig. 2). Baseline SDMA and L-arginine levels were similar in WT and TG mice (SDMA, 0.17 ± 0.02 vs. 0.17 ± 0.01 µM; and L-arginine, 65.6 ± 4.2 vs. 76.0 ± 7.4 µM, P = NS) and remained unaltered by treatment (SDMA, 0.17 ± 0.02 vs. 0.17 ± 0.01 µM; L-arginine, 66.6 ± 8.7 vs. 60.7 ± 5.8 µM). The baseline and treatment-related L-arginine/ADMA ratio significantly differed between genotypes (baseline WT vs. TG, 98.7 ± 9.3 vs. 177.2 ± 26.5, P = 0.01; and treatment WT vs. TG, 95.2 ± 13.1 vs. 177.5 ± 22.6, P = 0.006) but did not change following ANG II infusion.

Renal Histomorphology and Immunohistochemistry

The infusion of ANG II was associated with signs of glomerulosclerosis and interstitial fibrosis. Light microscopic evidence of glomerular damage was mild and similar between genotypes (Fig. 4A). Glomerular collagen IV staining was significantly enhanced in mice infused with 3.0 µg·kg–1·min–1 ANG II, whereas the lower dose of ANG II had no effect on collagen IV content (WT mice, PBS vs. 1.0 µg·kg–1·min–1 ANG II vs. 3.0 µg·kg–1·min–1 ANG II, 10.9 ± 0.8% vs. 11.2 ± 0.9% vs. 18.3 ± 0.8%; and TG mice, PBS vs. 1.0 µg·kg–1·min–1 ANG II vs. 3.0 µg·kg–1·min–1 ANG II, 9.6 ± 0.5% vs. 9.6 ± 0.7% vs. 18.6 ± 0.6%; treatment P value, P < 0.001; and genotype P value, P = NS). ANG II caused marked renal interstitial fibrosis, which was significantly reduced in TG mice (Fig. 4B). Immunohistochemical findings of reduced renal interstitial fibrosis in TG mice were confirmed by RT-PCR of collagen I mRNA (Table 2). ANG II caused marked macrophage infiltration within the kidneys (Fig. 4C). F4/80 immunohistochemistry revealed a perivascular and periglomerular distribution pattern in ANG II-treated mice, and glomeruli were largely devoid of macrophages.


Figure 4
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  		Fig. 4. Chronic infusion of ANG II and renal histomorphology. A: chronic infusion of ANG II and glomerulosclerosis in WT and TG mice. B: chronic infusion of ANG II and interstitial fibrosis (collagen I staining). C: chronic infusion of ANG II and renal macrophage infiltration (F4/80 staining). Pos cells, positive cells; HPF, high power field.

 

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  		Table 2. Renal mRNA expression of collagen I and ADMA generating and degrading enzymes

 
Oxidative Fluorescence Microscopy

The infusion of ANG II dose dependently increased oxidative stress in WT mice as evidenced by DHE fluorescence in both the ascending and descending aorta (Fig. 5, A and B). In contrast, in TG mice only 3.0 µg·kg–1·min–1 ANG II was associated with enhanced DHE fluorescence. Overall, DHE fluorescence in mice treated with ANG II was significantly greater in WT as opposed to TG animals.


Figure 5
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  		Fig. 5. Chronic infusion of ANG II and vascular oxidative stress. A: dihydroethidium (DHE) fluorescence within the ascending and descending aorta. B: DHE fluorescence in PBS, ANG II-treated (1.0 µg·kg–1·min–1) WT, and ANG II-treated (1.0 µg·kg–1·min–1) TG animal (ascending aorta, x4 objective).

 
ANG II Infusion and Renal mRNA Expression of Collagen I and ADMA-Generating and -Degrading Enzymes

The infusion of ANG II was associated with a significant increase in renal mRNA expression of PRMT1 and DDAH2, whereas PRMT3 and mouse DDAH1 expression remained unaltered (Table 2).


   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The salient findings of the present study are that 1) the chronic infusion of ANG II over 4 wk at a dose associated with marked hypertension and target organ damage has no net effect on plasma levels of ADMA and 2) the overexpression of DDAH nevertheless protects against ANG II-induced vascular oxidative stress and renal interstitial fibrosis. The lack of effect of ANG II infusions on plasma ADMA levels observed in our studies stands in contrast to recent reports by others (13). In these experiments, Hasegawa and colleagues (13) studied the effect of ANG II infusions (1.0 µg·kg–1·min–1 delivered via osmotic minipumps over 2 wk) on plasma ADMA levels in both WT and DDAH2 TG mice. Although ADMA levels were twofold higher in ANG II- versus saline-treated WT mice (0.72 vs. 1.48 µM, P < 0.01), DDAH2 TG mice were, in part, resistant to ANG II-induced elevations of ADMA levels (0.52 vs. 0.80 µM, P < 0.05).

Notably, the results of Hasegawa et al. (13) agree with our data in several aspects. At a similar dose, these authors observed a comparable blood pressure increase regardless of whether the transgene was present or not. The reduced interstitial fibrosis in the kidney in our study is reminiscent of the effect of DDAH2 on the perivascular fibrosis reported by Hasegawa et al. (13). Similar to our study, these authors noted less vascular oxidative stress in TG versus WT mice following ANG II infusion (13). The effect of a DDAH2 transgene on vascular hypertrophy in the study by Hasegawa et al. (13) may be explained by the recent observation that the DDAH2 isoenzyme plays a greater role for ADMA metabolism in vascular tissue, whereas DDAH1 has a more pronounced effect on serum ADMA levels (39). This may explain the more pronounced effect of DDAH1 (in our study) versus DDAH2 (13) overexpression on systemic ADMA levels compared with WT littermates (37% vs. 26% decrease). In that regard, the lack of tissue ADMA measurements must be considered a limitation of our study since tissue ADMA levels might be affected by ANG II, unlike plasma ADMA.

Given the similarities, the discrepant results on the effects of ANG II on ADMA levels are hard to explain. Due to our longer duration of ANG II treatment (4 vs. 2 wk), we may have missed a transient increase of ADMA after 2 wk. A further explanation for the discrepant findings may result from different detection methods for ADMA (HPLC vs. ELISA/LC-MS), the variation of which in the literature is substantial, especially for HPLC methods (24). We took several steps to exclude other possible explanations. First, we used an adequate sample size. Second, we confirmed our initial negative observation by also studying a higher dose of ANG II, namely 3.0 µg·kg–1·min–1. Third, to corroborate our findings, ADMA levels were measured both by ELISA as well as LC-MS. Although we measured a small (~15%) increase of ADMA levels by ELISA in WT mice infused with 3.0 µg·kg–1·min–1 ANG II (and only compared with pretreatment levels), this increase was very minor and far from the 100% increase observed by Hasegawa et al. (13). In addition, we could not confirm this apparent increase with LC-MS measurements, despite good agreement between both methods in many other respects. As observed by others (22, 32), the ELISA yielded higher ADMA levels than LC-MS but showed an excellent discrimination between WT and TG mice with virtually no overlap of ADMA levels between genotypes.

Apart from the studies by Hasegawa and colleagues (13), little is known about how ANG II might affect ADMA metabolism. In vitro studies by Chen et al. (5) suggest that ANG II-induced elevations of ADMA levels are the consequence of an enhanced expression of PRMT as well as reduced activity of DDAH, but these authors used rather high doses of ANG II (1 µM) and ADMA (500 µM for DDAH activity assay). We observed a minor yet significant ANG II-induced upregulation of PRMT1 and DDAH2 gene expression in the kidney, whereas PRMT3 and endogenous mouse DDAH1 expression remained unaltered. The combined upregulation of ADMA-generating (PRMT1) and -degrading (DDAH2) enzymes may explain that no net effect of ANG II treatment on systemic ADMA levels was observed. Interestingly, enhanced mRNA expression of renal PRMT1 and DDAH2 during ANG II infusions mirrors the mRNA expression pattern observed during RAS blockade in a rat model of diabetes. Thus Onozato et al. (28) observed reduced expression of both enzymes during treatment with the ANG II receptor blocker telmisartan. Furthermore, our findings agree with earlier observations of Tojo et al. (37), who reported that DDAH protein expression was increased in rats by stimulating an endogenous ANG II production (by means of a low-salt diet) as well as by an infusion of exogenous ANG II.

Others have reported that ADMA levels may affect the expression of ACE (34) or the pressor response to ANG II (8). We did not observe any relationship between ANG II-induced hypertension and plasma ADMA levels. Evidence for an association between high blood pressure and elevated ADMA levels comes from small clinical studies (29, 35). Other reports, including several large-scale clinical studies, did not confirm such an association (25, 31). Our blood pressure results are limited because we only used a noninvasive tail-cuff method, a technique that has received criticism in the past. Although we may have missed subtle blood pressure differences between genotypes with this technique, the overall effect of ANG II treatment on blood pressure was clearly evident and within the range reported by others investigators with similar doses in which either the tail-cuff method or telemetric devices were being used (13, 41).

Despite the lack of ANG II to alter plasma levels of ADMA, we observed a functional relevance of the enzyme DDAH for ANG II-induced formation of ROS and renal interstitial fibrosis. ANG II is known to stimulate the generation of oxygen free radicals (O2–) (30), which may in turn lead to endothelial NOS (eNOS) uncoupling via depletion of the cofactor tetrahydrobiopterin (3). In such a state, NOS itself becomes a major source for ROS generation. There is growing evidence that elevated ADMA levels may facilitate eNOS uncoupling (36). Such a mechanism may serve as an explanation of why TG mice were less susceptible to ANG II-induced vascular oxidative stress. In addition, the enzyme DDAH is redox sensitive, which results from a critical sulfhydryl group within the active catalytic site of DDAH (18). Enhanced expression of DDAH may, therefore, be vasoprotective. Finally, we speculate that ADMA may play a permissive role for target organ damage. Even apparently normal levels of factors, which can contribute to vascular damage, may become deleterious in the presence of comorbid conditions such as high ANG II levels and/or hypertension.

In addition to reduced vascular oxidative stress, TG mice developed less renal interstitial fibrosis. This finding is in line with recent observations by Matsumoto and colleagues (23) in which adenovirus-mediated overexpression of DDAH1 significantly reduced glomerular and interstitial fibrosis in subtotally nephrectomized rats. The infiltration of macrophages and T lymphocytes into the interstitium is a prominent finding and precedes interstitial fibrosis in ANG II-dependent forms of hypertension (17, 21). Several strategies aimed at reducing inflammation, including the blockade of tumor necrosis factor-{alpha} (TNF-{alpha}), have been shown to reduce ANG II-dependent kidney damage (26). The potential effect of ADMA on these mechanisms will require further studies.

In conclusion, our study helps to delineate the relevance of ADMA in ANG II-induced hypertension and target organ damage in mice. Our data indicate that ADMA contributes to ANG II-induced oxidative stress and renal interstitial fibrosis. However, ANG II did not increase ADMA levels in our animals, in contrast to recent studies by others (13). The lack of an increase of ADMA may explain the unexpectedly modest role of this mediator.


   GRANTS
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 ABSTRACT
 METHODS
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This work was supported by the Interdisciplinary Center for Clinical Research at the University Hospital of the University of Erlangen-Nuremberg, project B12, and by the Deutsche Forschungsgemeinschaft (KFO 106/2-1).


   FOOTNOTES
 

Address for reprint requests and other correspondence: K. F. Hilgers, Nephrologische Forschungslabors, Loschgestrasse 8, Erlangen D-91054, Germany (e-mail: karl.hilgers{at}uk-erlangen.de)

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
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