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Am J Physiol Heart Circ Physiol 291: H1788-H1796, 2006. First published April 21, 2006; doi:10.1152/ajpheart.00066.2006
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Contribution of whole blood to the control of plasma asymmetrical dimethylarginine

Scott S. Billecke,1 Laura A. Kitzmiller,2 Joseph J. Northrup,3 Steven E. Whitesall,1 Masumi Kimoto,4 Alia V. Hinz,5 and Louis G. D'Alecy1,6,7

1Department of Molecular and Integrative Physiology, University of Michigan Medical School, Ann Arbor; 2College of Literature, Science, and The Arts and 3College of Engineering, University of Michigan, Ann Arbor, Michigan; 4Department of Nutritional Science, Faculty of Health and Welfare Science, Okayama Prefectural University, Kuboki, Soja, Okayama, Japan; 5Macromolecular Structure Facility, Michigan State University, East Lansing; 6Department of Surgery (Vascular), University of Michigan Medical School, Ann Arbor; and 7William Beaumont Hospital, Department of Surgery, Royal Oak, Michigan

Submitted 16 January 2006 ; accepted in final form 11 April 2006


   ABSTRACT
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 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The endogenous nitric oxide (NO) synthase (NOS) inhibitor asymmetrical dimethylarginine (ADMA) is elevated in many patients and may contribute to the initiation and progression of their disease. While some mechanistic pathways have been identified, tissue-specific contributions to ADMA control remain unclear. We sought to determine if whole blood (WB) could participate in ADMA control ex vivo. Anesthetized male Sprague-Dawley rats underwent exsanguinations, and WB preparations were incubated at 37°C for 5 h. ADMA and symmetrical dimethylarginine were analyzed by high-pressure liquid chromatography. Incubation of lysed red blood cell (RBC) supernatant yielded a significant decrease in ADMA that was blocked by 4124W, a synthetic inhibitor of dimethylarginine dimethylaminohydrolase, the only reported enzyme to hydrolyze ADMA. Hydrolysis of ADMA was diminished by addition of physiologically relevant concentrations of zinc (i.e., 20 µM). Conversely, when rat WB or WB supernatant was incubated at 37°C, it liberated quantities of free ADMA (1–2 µM) that in vivo would likely have pathological consequences. Addition of arginine methyltransferase inhibitors to these incubations did not reduce ADMA release, indicating no dominant role for active protein methylation during these incubations. This ADMA liberation was significantly reduced by addition of protease inhibitors, indicating a dependence on peptide bond hydrolysis. Total ADMA (protein incorporated plus free) was determined by acid hydrolysis and found to be 43.18 ± 4.79 µM in WB with ~95% of this in RBCs. These ex vivo data demonstrate the potential of blood to control the NO-NOS system by modulating free ADMA.

nitric oxide; protein arginine methyltransferase; symmetrical dimethylarginine; protease

NITRIC OXIDE (NO) is produced by NO synthase (NOS) and has a myriad of functions throughout the body, including vasodilation, signaling functions, and inhibition of platelet aggregation. Deficiency or loss of NOS activity has been termed "endothelial dysfunction" and may contribute to a spectrum of cardiovascular diseases. Asymmetrical dimethylarginine (ADMA) and NG-monomethyl-L-arginine (L-NMMA) are endogenous inhibitors of NOS, but the concentration of ADMA exceeds that of L-NMMA by ~10-fold (60) and hence is of greater potential clinical interest. ADMA and its noninhibitory regioisomer, symmetrical dimethylarginine (SDMA) (6, 12), are released into the plasma following the breakdown of proteins containing arginine residues previously dimethylated by protein arginine methyltransferases (PRMT) (37). The literature contains compelling evidence that elevated plasma ADMA exists in diabetes mellitus (37), hypertension (1, 15, 22, 45, 51, 52), hypercholesterolemia (6), hyperhomocyst(e)inemia (50), experimental hemorrhage (4), preeclampsia (16), and sickle cell anemia (47). ADMA has also been reported to increase platelet activation and atherogenesis in the cardiovascular system (33). Cooke (11, 13) and others (17, 59) give extensive reviews of much of these data and argue that ADMA plays an important role in regulation of vascular tone by acting as an endogenous inhibitor of NO synthesis. Potential pathophysiological roles for elevated ADMA have been suggested by positive correlations with chronic renal failure and end-stage renal disease (25–30, 60, 61, 63–65), the severity of peripheral arterial occlusive disease (7), and impaired flow-mediated dilation (12), a surrogate for coronary artery reserve. It is still unclear, however, if the elevated plasma ADMA levels observed in these patient populations are entirely due to decreased renal clearance of ADMA or if other disease processes elevate plasma ADMA and hence contribute to overall renal and endothelial dysfunction (9, 18, 44).

Some elements of the origin and elimination of ADMA have been described (23, 39–42, 53). It has been suggested that PRMT activity is the control point for ADMA production (8). In addition, although long assumed to be a key step in its formation (36, 56), the role of protein turnover in the release of free ADMA from intact proteins and/or peptides has not been well documented. With regard to ADMA elimination, the enzyme dimethylarginine dimethylaminohydrolase (DDAH) hydrolyzes ADMA and appears to have the greatest role in its metabolism. While these counterbalancing pathways are believed to be responsible for homeostatic control of free plasma ADMA in vivo, the location(s) and mechanism(s) of ADMA generation and elimination have not been fully described. Demonstration of these pathways in ex vivo whole blood (WB) could have profound importance due to the proximity of the blood to endothelial cells and subjacent vascular smooth muscle cells, making WB an ideal site for controlling the NO-NOS system. In this study, we examine the potential contribution of WB to the control of free plasma ADMA without the contributions from other organs.

Given that DDAH activity has been identified in WB components (24), we first set out to determine if WB and WB elements had the capacity to eliminate physiologically relevant plasma ADMA concentrations via DDAH-mediated hydrolysis. We also assessed the potential of WB to liberate ADMA. Specifically, both PRMT activity and protein turnover in ex vivo WB were assessed. We found that rat WB possesses significant DDAH activity. These analyses also led to the discovery of a large reservoir of protein-incorporated ADMA in WB in addition to the presence of the proteolytic enzymes necessary for ADMA release.

Although not necessarily indicative of the contribution of WB in vivo, these results show for the first time the presence of a large store of protein-incorporated ADMA in close proximity to the vascular endothelium. This store may be released under certain pathological conditions. These findings may have clinical implications for those diseases displaying increased red blood cell (RBC) lysis and/or blood protein turnover. Many of these diseases are characterized by hypertension and endothelial dysfunction that could easily be attributable to an increase in the endogenous NOS inhibitor ADMA. This study demonstrates that blood itself is a potential contributor to the control of plasma ADMA levels.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals and housing. All blood samples were obtained from male Sprague-Dawley rats (Charles River Laboratories) weighing 350–600 g at the time of exsanguinations. Animal care complied with American Physiological Society's Principles of Laboratory Animal Care and the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health (NIH Publication No. 85–23, Revised 1996). All animal procedures were approved by the University of Michigan Committee for the Use and Care of Animals (Approval Number 8252).

Animal blood-draw protocol. Anesthesia was induced with 5% isoflurane in a chamber for 2 min and maintained by mask as needed with isoflurane (1.5%–2.5%) throughout the blood collection. A midline incision was made on the ventral abdomen, the intestines were reflected laterally, and a 19-gauge needle was inserted into the aorta. Heparin (100 U/kg body wt) was given intra-arterially and allowed to circulate for 8–10 s before the blood was withdrawn until the heart stopped beating. In those experiments testing the effect of EDTA on ADMA, control blood was drawn directly into EDTA (Vacutainer) with final concentration of 4 mM.

Washed RBC supernatant incubation. Heparinized WB was centrifuged for 8 min at 1,800 g, and plasma and buffy coat were removed and discarded. The RBC pellet was resuspended to the original WB volume with saline containing 3.33 µM ADMA and then mixed gently by inversion. The RBC suspension was again centrifuged, separated, and resuspended in fresh ADMA-saline solution for incubations as described below for WB. The suspension was then subjected to lysis by three freeze-thaw cycles with 1 min of freezing in liquid nitrogen and 5 min of thawing in a 37°C water bath. The lysate was centrifuged (16,000 g for 8 min), and the supernatant, which we refer to as RBC supernatant, was divided into four coated (Sigmacote, SL-2, Sigma-Aldrich, St. Louis, MO) 25-ml Erlenmeyer flasks with the addition of either normal saline or 4124W (prepared by Dr. Masumi Kimoto) to make 0.00, 0.01, 0.10, or 1.00 mM solutions. These coated flasks were used for the incubation of all blood fractions analyzed in this study. Aliquots were incubated at 37°C in a shaker-water bath for 5 h, and samples (100 µl) were taken at time 0 and at 1, 3, and 5 h for analysis by high-pressure liquid chromatography (HPLC). In a separate set of experiments, incubations of RBC supernatant from heparinized WB, EDTA-treated WB (4 mM), and from heparinized WB with the addition of glucose (30 mM, approximating concentrations found in diabetic hyperglycemia) (35) were compared.

Whole blood incubation. Four-milliliter aliquots of pooled WB were placed in coated flasks and incubated at 37°C or 4°C for 5 h with samples (280 µl) taken at 0, 1, 3, and 5 h. Each sample was centrifuged for 6 min (1,800 g), and plasma drawn off for analysis of ADMA and SDMA using HPLC as described below. In a separate series of experiments, WB was subjected to lysis by the same freeze-thaw procedure described for RBC supernatant. The lysate was then centrifuged (16,000 g for 8 min), and the supernatant, which we refer to as freeze-thaw whole blood (FTWB), was removed for subsequent incubation experiments. A series of FTWB samples were treated with arginine methyltransferase inhibitors (see GoGoGoGoFig. 5) or protease inhibitors (see Table 2), each at doses well in excess of their reported IC50s, before initiating the incubation protocol. For plasma incubations, pooled and heparinized WB was centrifuged for 8 min at 1,800 g, and plasma was drawn off and then incubated at 37°C for 5 h with samples taken at 0, 1, 3, and 5 h. Samples were then analyzed for ADMA and SDMA by HPLC.


Figure 1
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  		Fig. 1. Washed red blood cell (RBC) supernatant at 37°C with or without 4124W dimethylarginine dimethylaminohydrolase (DDAH) inhibitor. The dramatic hydrolysis of asymmetrical dimethylarginine (ADMA) in a saline-treated group is inhibited in a dose-dependent manner by 4124W. The increases over baseline at 1 and 3 h in the 1.0 mM 4124W-treated group suggest ADMA release from protein exceeds DDAH-mediated hydrolysis.

 

Figure 2
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  		Fig. 2. Washed RBC supernatant at 37°C. ADMA concentrations (solid lines) decrease in all groups to <40% of baseline regardless of treatment group. Symmetrical dimethylarginine (SDMA) concentrations (dashed lines) increase in all groups. SEs (not seen) are within the symbols.

 

Figure 3
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  		Fig. 3. Effect of zinc on ADMA hydrolysis. Addition of 20 or 100 µM ZnCl2 shifts the equilibrium in RBC supernatant (A; n = 4) from hydrolysis of ADMA in favor of accumulation (P < 0.02 for control vs. 20 µM ZnCl2 at 5 h; P < 0.005 for control vs. 100 µM ZnCl2 at 5 h). In the absence of added ZnCl2, ADMA increases in freeze-thaw whole blood (FTWB) (B; n = 5) over the 5-h time course and is further increased in the presence of zinc (P = 0.03 for control vs. 100 µM ZnCl2 at 5 h). ADMA in plasma from whole blood (WB) incubations (C) also accumulated (n = 5) but was unaffected by added ZnCl2. SEs (not seen) are within the symbols.

 

Figure 4
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  		Fig. 4. ADMA concentration increases on incubation of WB or FTWB but not plasma. Samples were incubated at 37°C for 5 h and assayed for ADMA (solid lines) and SDMA (dashed lines) by HPLC. ADMA concentrations increased dramatically on removal of RBC membranes. Incubation of separated plasma does not result in an increase in ADMA or SDMA. SDMA increased significantly during the incubation of FTWB (P < 0.005). WB DMA showed no detectable change when incubated at 4°C. SEs (not seen) are within the symbols.

 

Figure 5
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  		Fig. 5. Inhibitors of protein arginine methyltransferases (PRMTs) do not attenuate increases in ADMA during incubation of FTWB. To 300 µl FTWB, either saline (control), the general methyltransferase inhibitor S-adenosyl-homocysteine (SAH) (200 µM), or the PRMT-specific inhibitor arginine methyltransferase inhibitor-1 (AMI-1) (100 µM) was added, and samples were incubated for 1 h at 37°C. The inhibitor concentrations used exceed the reported IC50s for PRMTs by at least 10-fold (8, 10, 48). No statistically significant change in ADMA release is observed as determined by ANOVA (P = 0.40).

 

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  		Table 2. Protease inhibitors attenuate ADMA release from rat FTWB

 
Incubations with zinc. In addition to the treatments described above, WB, FTWB, and RBC supernatant were treated with ZnCl2 (0, 20, or 100 µM) and incubated as described. Samples were taken at time 0 and at 1, 3, and 5 h, prepared as described for each blood preparation, and assayed for ADMA as described below.

Acid hydrolysis of blood proteins. WB, plasma, or RBCs were hydrolyzed to release any previously formed dimethylated arginines (DMAs) incorporated within intact proteins. Vapor-phase acid hydrolysis was used to completely hydrolyze the proteins in WB (14). Samples were kept on ice and diluted with an ice-cold solution containing NG-monomethyl-L-arginine (L-NMMA) (Novabiochem, San Diego, CA) in H2O as an internal standard at a final concentration of 8 µM. One-hundred microliters of 20-fold diluted WB, 2-fold diluted plasma, or 40-fold diluted RBCs were dried in a centrifugal evaporator (SpeedVac SPD 1010, ThermoSavant, Holbrook, NY). The dried samples were placed in a hydrolysis chamber (Picotag, Waters, Milford, MA) containing 6 N HCl (constant boiling) and incubated in vacuo at 150°C for 1 h and again dried completely. The resulting hydrolysate was resuspended in 100 µl of 20 mM HCl, and the supernatant was prepared for ADMA and SDMA analysis by HPLC. Subsequent concentration values were corrected for dilution, and the contribution of the sample to the total blood volume was based on hematocrit (e.g., if plasma represents one-half of WB volume, DMA content was divided by 2).

Sample preparation for ADMA/SDMA assay. Heparinized plasma, FTWB, or washed RBC supernatant (100 µl) was prepared for HPLC analysis of ADMA and SDMA by using the method described by Carello et al. (9). Briefly, samples were ultrafiltered through a 10,000 molecular weight cutoff membrane (Ultrafree MC, Millipore, Bedford, MA) at 16,000 g for 20 min at 4°C, spiked with 3 µM L-NMMA as an internal standard (20 µl filtrate plus 100 µl standard), and then subjected to solid-phase extraction by loading 60 µl onto 50 mg of conditioned (1 ml 100% methanol) and equilibrated (1 ml 2% trichloroacetic acid) cation-exchange resin (Isolute SCX2 50 mg/1cc, International Sorbent Technology, UK). Adsorbed samples were then sequentially gravity-rinsed with 0.5 ml of the equilibration solution, 2 ml of 15 mM (pH 8.00) sodium phosphate, and 0.5 ml of 100% methanol followed by elution with 1 ml of eluent (40% NH4OH ACS grade reagent containing 28–30% NH3, balance methanol). Samples were then dried, reconstituted with 80 µl of a pH 8.2 borate buffer, and derivatized with 20 µl of fluorescent derivatizing agent (AccQ*Fluor, Waters, Milford, MA). After 1-min incubation, the 100 µl was transferred to a low-volume insert and heated (55°C) for 10 min to complete reaction before injection on the column.

During the course of our investigation, Heresztyn et al. (20) described what we found to be a more efficient method for preparing samples for ADMA and SDMA analysis by HPLC. We utilized a modified version of this method for all of our freeze-thaw, PRMT inhibitor, protease inhibitor, and acid hydrolysis data, finding no significant differences in ADMA and SDMA concentrations determined by the two methods. Briefly, proteins were removed from samples (100 µl) by acid precipitation with L-NMMA added as an internal standard (40 µl 10% 5-sulfosalicylic acid, 10 µl 80 µM L-NMMA, and 320 µl water per 100 µl sample). Samples were left on ice for 10 min and then centrifuged at 9,000 g for 2 min. Supernatants were added to preconditioned columns (Bond Elut-SCX 50 mg/1cc, Varian, Palo Alto, CA) and eluted as above except that adsorbed samples were rinsed with 1.25 ml of sodium phosphate (0.1 M, pH 6) and eluted with 1.25 ml 2% triethylamine in 65% methanol-water. Eluents were dried as above and derivatized by using 90 µl of borate buffer and 10 µl of fluorescent derivatizing agent, which was then heated as above.

High-pressure liquid chromatography. ADMA and SDMA in samples were quantified by reverse-phase liquid chromatography (Breeze System, Waters) by using the method of Carello et al. (9). Separation was performed on a 4.6 mm x 150 mm, 3.5-µm column (Waters, XterraMS C18) with an identical 3.9 mm x 20 mm guard column, both controlled at 36°C. Standards, blanks, and samples (10 µl) were injected (Waters, 717 Plus Autosampler), and fluorescent peak height and area were evaluated at an excitation of 250 nm and an emission of 395 nm (Waters, 2475 Multi-wavelength Fluorescence Detector). Each sample injected onto the column, including standards, received identical treatment within each sample set. Concentrations within each sample set were determined from a standard curve generated by seven standards containing known concentrations of ADMA and SDMA (from 0 to 12 µM DMA). The average method detection limit (MDL) (38) was calculated from three sets of 10 replicates of 0.75 µM DMA standard and is 0.07 µM for SDMA and 0.13 µM for ADMA. Interassay coefficient of variation was 9%, and the intra-assay coefficient was 5%, determined by comparison of plasma quality controls (4 replicates) included within each sample set.

Statistical analysis. Measured variables are reported as means ± SE and were considered statistically significant by using two-way ANOVA followed by the paired two-tailed Student's t-test corrected, unless otherwise noted, for repeated measures.


   RESULTS
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 ABSTRACT
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 RESULTS
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DDAH inhibitor 4124W attenuates ADMA degradation in rat RBC supernatant. We first sought to replicate the findings of Kang et al. (24) in rat blood by identifying DDAH activity in RBC supernatant. Before lysis, the RBC pellets were washed and resuspended in saline containing 3.33 µM ADMA. Baseline ADMA concentrations were artificially elevated in this manner to improve resolution of ADMA changes over time and allow for the detection of concentrations well within the HPLC assay detection limits. Changes in ADMA concentration were determined over a 5-h incubation at 37°C. RBC supernatant ADMA concentration decreased during the 5-h incubation to <50% of control (Fig. 1). Addition of the DDAH-specific inhibitor 4124W at the start of incubation produced a dose-dependent inhibition of ADMA degradation with the highest dose (1.00 mM) reversing the pattern to a net increase in ADMA concentration for the 1- and 3-h time points. Incubation of RBC supernatant prepared from WB collected using the anticoagulant EDTA (4 mM), which is also a putative DDAH inhibitor (24), or glucose (30 mM) (35) failed to attenuate the degradation of ADMA with each resulting in decreased ADMA concentrations similar to control (Fig. 2; P < 0.02 over 5-h incubation). In contrast, SDMA concentration increased significantly in each group tested (P < 0.02).

Addition of zinc to rat RBC supernatant inhibits ADMA degradation and exposes ADMA generation. 4124W (1.00 mM) produced a slight increase in ADMA at 1 and 3 h, suggesting that ADMA might be released in this blood preparation. However, because we were unable to sustain ADMA accumulation with a high concentration of 4124W and did not observe any inhibitory effect with EDTA or glucose, we chose to explore possible ADMA release by inhibiting DDAH via another mechanism. Knipp et al. (32) have shown that zinc is a negative regulator of DDAH activity; thus we chose to determine the effect of ZnCl2 on ADMA concentration changes during incubation of blood at 37°C (Fig. 3). Addition of 20 or 100 µM ZnCl2 to RBC supernatant incubations produced significant increases in ADMA concentration compared with control (P < 0.02 for control vs. 20 µM ZnCl2 at 5 h; P < 0.005 for control vs. 100 µM ZnCl2 at 5 h) with 100 µM ZnCl2 reversing the equilibrium from net ADMA degradation to net ADMA accumulation over 5 h.

This increase in ADMA concentration prompted us to test ADMA changes during incubation of other blood preparation. Incubation of lysed WB (FTWB) resulted in an increase in ADMA under basal conditions that increased significantly in the presence of added zinc (P = 0.03 for control vs. 100 µM ZnCl2 over 5 h). When WB was incubated for 5 h at 37°C and plasma ADMA was measured, a small but statistically significant increase of 0.32 µM after 5 h was observed. Addition of ZnCl2 to WB did not further increase this modest accumulation of plasma ADMA over the 5-h time course.

Accumulation of ADMA in incubated rat blood fractions. To explore further the potential for rat WB and WB fractions to accumulate free ADMA and SDMA, we compared incubations of WB, FTWB, or plasma at 37°C for 5 h. Results are shown in Fig. 4. This consistent increase in ADMA when incubated at 37°C is contrasted to WB incubated at 4°C. During this cold incubation, we were unable to detect any statistical changes in either ADMA or SDMA within the 5-h incubation. Incubation of FTWB released ADMA to a greater degree (P < 0.0001 over 5 h) compared with WB (P = 0.02), while separated and incubated plasma concentrations were unchanged (P = 0.87) over the 5-h time course. We did not detect a statistically significant increase in plasma SDMA during the 5-h incubation of WB, although there was a general trend toward an increase. However, concentrations of free SDMA increased significantly during the incubation of FTWB (P < 0.005) but not plasma (P = 0.18).

PRMT inhibitors fail to attenuate ADMA release in rat FTWB. It is widely accepted that an initial step in ADMA formation is the dimethylation of arginine residues within intact proteins by the PRMT family of enzymes [reviewed by Tran et al. (56)]. To assess the possible role of PRMTs in the observed increases in free ADMA, we treated FTWB with the methylase inhibitors S-adenosyl-homocysteine (SAH; Sigma) or arginine methyltransferase inhibitor-1 (AMI-1; Calbiochem, La Jolla, CA) at 200 and 100 µM, respectively, concentrations well in excess of those reported to inhibit PRMT activity (8, 10, 48). Figure 5 shows that after 1-h incubation at 37°C, ADMA concentrations increased at least as much in the presence of either PRMT inhibitors as with saline control. A slight increase was observed but was not statistically significant.

Quantification of total (protein incorporated plus free) ADMA in rat blood fractions. Since ongoing PRMT activity did not appear necessary for the release of ADMA under the conditions employed in previous experiments, we chose to determine the amount of protein-incorporated ADMA present at baseline in WB and blood fractions. WB, plasma, and intact RBCs were diluted, dried via vacuum evaporation, hydrolyzed with HCl, and then assayed for total ADMA and SDMA (Table 1). Total ADMA concentration in WB was ~43 times that of the concentration free in plasma. Total SDMA in WB was ~25 times greater than that found free in plasma. The vast majority of DMA was located in the RBCs (~95%); thus free plasma ADMA accounts for only ~2% of the total ADMA available in WB.


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  		Table 1. DMA in WB and selected components

 
Protease inhibition reduces ADMA release in incubated rat FTWB. The large amount of DMA available from intact proteins indicates a potential role for protease activity in increasing DMA, as indicated in Figs. 3 and 4. To address this question, we treated FTWB with a commercially available protease inhibitor cocktail (Complete, mini, EDTA-free from Roche) at two concentrations (1x and 4x as described by the manufacturer) or with nonspecific cysteine- and serine-protease inhibitors. The concentrations used for these latter inhibitors were well in excess of their IC50 concentrations for most proteases. The data in Table 2 show that the presence of each protease inhibitor produced a statistically significant attenuation of ADMA release from FTWB with some variability in the effectiveness of the inhibitors. Release of SDMA was similarly attenuated (data not shown). Inhibitors reported to block both cysteine- and serine-protease activity (leupeptin and PMSF) both produced similarly potent reductions (by 55 and 57%, respectively), while the serine-protease specific inhibitor aprotinin had a less dramatic effect (34%). Each condition, however, produced a statistically significant reduction in ADMA release (P ≤ 0.05, n = 4) with the 4x inhibitor tablet producing the greatest inhibition.


   DISCUSSION
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
This study describes for the first time the potential of WB to act as an independent modulator of free plasma ADMA. This potential was documented ex vivo by identifying the presence of substantial quantities of protein-incorporated ADMA in rat blood and by demonstrating that WB possesses the molecular machinery necessary for ADMA release from this protein-incorporated store. Furthermore, this release appears to be countered by the significant DDAH-mediated, zinc-sensitive hydrolysis by blood of ADMA. Together these ex vivo observations establish the potential of WB to act as a modulator of free plasma ADMA and suggest that blood may release clinically significant quantities of ADMA when pathological hemolysis or blood protein turnover occurs.

While this ex vivo approach does not provide information on the overall in vivo capacity or contribution of WB to the systemic control of plasma ADMA, identification of the ADMA generation and elimination pathways of WB may provide valuable insights into a novel mechanism of ADMA control.

DDAH-mediated hydrolysis of ADMA in rat blood. Kang et al. (24) were the first to give a detailed description of DDAH protein and activity in diluted and sonicated supernatants from separated human WB elements (RBCs, polymorphonuclear cells, and mononuclear cells). These observations suggest a role for WB in eliminating plasma ADMA, an important finding because the rapid and continuous access of blood to the vascular endothelium makes it an ideal site for reducing free plasma ADMA. We set out to determine if WB and WB elements had the capacity to control physiologically relevant plasma ADMA concentrations via DDAH-mediated hydrolysis. Incubation of rat RBC supernatant yielded a significant reduction in ADMA over the 5-h time course (Fig. 1). This reduction was evidently not influenced by the anticoagulant used because EDTA, which is a putative DDAH inhibitor (24), did not affect ADMA reduction (Fig. 2). In Fig. 1, we show that addition of the DDAH inhibitor 4124W (2, 36, 49, 58) resulted in a dose-dependent inhibition of ADMA hydrolysis. The highest concentration tested (1.00 mM) yielded a slight increase in ADMA at the 1- and 3-h incubation times. We are not certain why 4124W does not maintain full inhibition during the entire 5-h time course, but it is possible that the 4124W is metabolized or inactivated by blood enzymes. Regardless, because hydrolysis was blocked by 4124W, it appears that the observed hydrolysis was DDAH mediated. Because plasma, white blood cells, and RBC membranes were absent in these incubations, we further conclude that these elements are not required for DDAH-mediated hydrolysis of ADMA. These RBC supernatant data do not exclude the possibility of DDAH activity in WB components other than the RBCs, but the magnitude of hydrolysis by RBC supernatant indicates substantial hydrolytic activity without the other formed elements of the blood being present.

Because of our general interest in the adverse consequences of hyperglycemia and the correlation between diabetes and elevated plasma ADMA (35, 37), we chose to test the effect of added glucose in these incubations (Fig. 2). Lin et al. (35) demonstrated that activity, but not expression, of DDAH was reduced in diabetic rats and in cultured cells treated with 25.5 mM glucose. In our experiment (Fig. 2), glucose was ineffective in attenuating the loss of ADMA. Perhaps the longer exposure (48 h) to high glucose in the study of Lin et al. is needed to inhibit DDAH. In addition, the complexity of vascular smooth muscle and cultured human endothelial cells used in their study greatly exceeds that of RBC supernatant; the extensive signaling pathways in these more complex systems may be required for glucose to inhibit DDAH. We have no other explanation why our use of 30 mM glucose did not alter the rate of disappearance of ADMA from RBC supernatant.

Addition of zinc to DDAH preparations has consistently resulted in the inhibition of ADMA hydrolysis as reported in at least three studies (24, 32, 40). However, these studies used supraphysiological zinc concentrations and/or a purified or heavily diluted source of DDAH, so we sought to test more physiologically achievable zinc concentrations. We increased the zinc concentrations by 20 or 100 µM, concentrations near those found in rat plasma [20–30 µM in rat plasma (43, 55, 62)] and which can be exceeded in rats fed a high-zinc diet [320 µM in the plasma of animals given a 100-fold increase in zinc ingestion compared with control (62)]. It should be noted that we did not measure plasma or erythrocyte zinc concentrations; determination of free vs. bound zinc was beyond the scope of this study. Our animals were, however, fed standard laboratory chow containing normal zinc levels. In rat RBC supernatant, addition of either 20 or 100 µM ZnCl2 produced a significant increase in ADMA concentration compared with control incubations (Fig. 3). The higher concentration yielded a shift from net ADMA degradation (hydrolysis) to net ADMA accumulation over 5 h (Fig. 3A), consistent with inhibition of DDAH-based ADMA hydrolysis and potential ongoing ADMA release.

The unexpected increase in ADMA, coupled with the slight increase seen at 1 and 3 h with 1.00 mM 4124W, prompted us to explore further the possible ADMA release pathways in other WB preparations. We began by first continuing our analysis of the zinc effect using FTWB supernatant and WB. Incubation of FTWB supernatant yielded a statistically significant increase in ADMA release after 5 h that was again increased in the presence of zinc (Fig. 3B, P = 0.03 for 100 µM ZnCl2). In WB incubations, plasma ADMA concentrations displayed a modest increase over the 5-h incubation and did not appear to be attenuated by addition of zinc. It is unclear why ADMA release in WB was unaffected by zinc, but it could be due to the presence of baseline zinc concentrations within RBCs that are optimally inhibitory [RBC zinc content exceeds that of plasma by ~10-fold (19)], and there could be poor transport of zinc across the RBC membrane. Conversely, we are uncertain why DDAH activity does not dominate ADMA release in WB and FTWB. What is clear is that the balance of factors governing local control of WB DDAH activity, including zinc, reactive oxygen species, and perhaps citrulline, has yet to be fully elucidated. Understanding this balance may contribute to our appreciation of the pathophysiological elevations of ADMA.

In WB the modulation of DDAH expression and/or activity may be one of the control points for plasma ADMA and hence contribute to the physiological control of the NOS-NO system for vascular smooth muscle. The inhibitory effect of zinc on DDAH activity suggests that excessive (or perhaps even normal) blood zinc concentrations might result in reduced ADMA hydrolysis and hence ADMA accumulation in vivo. Such an event would likely result in the manifestation of ADMA-related pathologies, including elevated blood pressure and increased platelet aggregation. In support of this hypothesis, Yanagisawa et al. (62) recently reported elevated systemic blood pressures in normotensive rats administered a high zinc diet. They also showed the involvement of the impaired NO-NOS pathway in facilitating this change. Determining if and how blood zinc concentrations modulate plasma ADMA levels in vivo could add to our overall understanding of the pathological conditions that negatively impact the NO-NOS system.

Storage and release of free ADMA in rat blood. In this study, we have demonstrated that WB, ex vivo, is capable of releasing enough ADMA to approach concentrations reported to result in serious endothelial dysfunction in vivo. On incubating WB at 37°C, we observed a 27% (0.32 µM) increase in plasma ADMA (Fig. 4). This increase represents a change in plasma ADMA that is over twofold higher than the 0.13 µM difference reported to have a profound impact on the incidence of cardiovascular events in patients with coronary artery disease, as described in a recent clinical study (46). The change that we observed in free plasma ADMA is temperature dependent (Fig. 4), suggesting an enzyme-mediated mechanism. At baseline, plasma concentrations of ADMA (0.94 ± 0.08 µM) were nearly identical to those found in FTWB (0.95 ± 0.08 µM), indicating that the increased plasma ADMA observed in WB incubations was not due to high intracellular concentrations of free ADMA being released into the plasma. However, disruption of the RBC membranes did appear to facilitate the release of protein-incorporated ADMA into solution, as shown by the FTWB data (Fig. 4) and even the 1-h time point of untreated RBC supernatant (Fig. 3A). Since some hemolysis occurred during the 5 h incubation of WB (data not shown), it is possible that the ADMA that accumulated in these incubations was contributed in part by release from intact RBC proteins, as suggested by acid hydrolysis data (see Table 1). If hemolysis and subsequent protein turnover do contribute to ADMA accumulation in WB, it is likely that pathological conditions characterized by RBC fragility or enhanced expression of proteolytic enzymes may facilitate increases in free plasma ADMA concentrations. The hypothesis that RBC rheology and altered protein turnover may have roles in ADMA release is supported by recent publications showing increased plasma ADMA in patients with sickle cell anemia (47), a correlation between ADMA and RBC fragility in hypertensive subjects (57), and elevated ADMA levels in the urine of patients afflicted with muscular dystrophy (21). As a further contributor to endothelial dysfunction, it is notable that RBCs contain significant quantities of arginase, an enzyme that hydrolyzes L-arginine to urea and ornithine. Arginase is released into the plasma upon RBC lysis (34) and thus, combined with the likelihood of ADMA release occurring upon hemolysis, would further reduce the L-arginine/ADMA ratio, which may be a better predictor of NO deficiency and endothelial dysfunction than plasma ADMA concentrations alone (6).

Because disruption of the RBC membranes appeared to facilitate the release of protein-incorporated ADMA, the mechanistic pathway(s) for release of ADMA were explored. It has been suggested that modulation of PRMTs, which has been reported in a wide variety of tissues and in multiple isoforms throughout the body (56), may act as a control point for plasma ADMA (8). However, even at extremely high doses, the PRMT inhibitors SAH and AMI-1 were unable to attenuate the incubation-dependent increases in free plasma ADMA concentration (Fig. 5), indicating that PRMT activity was not required for ADMA release under these conditions. This does not indicate a lack of PRMT presence or activity in the blood but rather that it is not required for the short-term, incubation-dependent release of ADMA.

Having ruled out the need for PRMT activity in increasing free ADMA during our ex vivo experiments, we sought to evaluate the possible role of protein hydrolysis, the second step in release of free ADMA from its protein-incorporated form. Recent reviews have suggested the importance of protein turnover in modulating free plasma ADMA concentrations, with some citing the potential for elevated ADMA in diseases exhibiting increased proteolysis (31, 56). In support of this, Teerlink (54) recently described the rapid release of ADMA from rat kidney homogenates. These authors presume this release to occur via a proteolytic mechanism but do not specifically identify a protease-dependent reaction as being responsible for their observations. We, however, demonstrated a significant reduction in ADMA release by treating rat FTWB with commonly used protease inhibitors or a commercially available protease inhibitor tablet. ADMA release was assessed after 1 h because these inhibitors are unstable when subjected to 37°C over longer periods of time, as indicated by their manufacturers and our own unpublished observations. Each inhibitor used in this study significantly reduced ADMA release (P ≤ 0.05). Of interest is the fact that at the doses we used, the inhibitors of both serine and cysteine proteases (leupeptin and PMSF) appeared more effective in reducing ADMA release than the serine protease-specific aprotinin. Whether this is due to the greater role of cysteine proteases in ADMA release or a reflection of suboptimal inhibitor concentration is not clear. Regardless, one can expect that any class of protease might participate in the release of free amino acids, and thus broad-spectrum protease inhibition would most likely yield the more complete inhibition, as we observed. We believe this is the first report of a protease-specific mechanism for ADMA release and that use of protease inhibitors may well be able to attenuate ADMA accumulation in other tissues in addition to blood.

Acid hydrolysis released a large quantity of DMA from WB proteins. This protein-bound DMA is not free but must be released from the intact proteins and/or peptides before it appears free in solution. We found ~95% of total WB DMA in the RBCs, and its concentration was ~46-fold (ADMA) and 25-fold (SDMA) greater than the concentration of free DMA in the plasma. Al Banchaabouchi et al. (3) observed a similar 35- to 40-fold difference between free DMA and protein-incorporated DMA throughout the various regions of the kidney. They also observed an ADMA-to-SDMA ratio similar to what we observed in WB. Given such a large store of protein-incorporated ADMA, only a small fraction (<1%) of this total would have to be released from intact WB by 37°C incubation to account for the increases we saw after 5 h. At this rate and in the absence of ongoing PRMT activity, the reservoir of protein-incorporated ADMA would be exhausted over a few days' time; thus in the intact animal we would anticipate a balance between protein-arginine methylation, protein turnover, and ultimately ADMA metabolism via DDAH and/or elimination by renal excretion. In addition, these homeostatic pathways are likely altered by our ex vivo experimental conditions. Nonetheless, it is plausible that the entire increase in plasma ADMA during WB incubation was due to release of protein-incorporated ADMA as opposed to a PRMT-controlled event. We interpret this to mean that RBCs must have been able, at some time in their life span, to either methylate their own proteins, through PRMTs or some other unknown mechanism, or sequester methylated proteins originating elsewhere in the body. In support of erythrocyte PRMT activity, PRMT1 has been recently identified in erythroid progenitor cells (5). Because demethylation of protein-incorporated DMAs has not been reported, it seems likely that ADMA-incorporated proteins would exist within RBCs until broken down by proteolytic cleavage. Determining how these ADMA-containing proteins and/or peptides accumulate in RBCs may aid in our understanding of how to manipulate free plasma ADMA concentrations with an eye toward therapeutics.

An additional question generated by these data is why DDAH activity dominates in RBC supernatant while ADMA release mechanisms are manifest in FTWB and WB incubations. ADMA degradation could become dominant in RBC supernatant preparations because of either 1) the absence in these preparations of some as-yet-undefined inhibitor of DDAH that is present in plasma and/or white blood cells, or 2) the suppression of ADMA release from proteins. SDMA concentrations in RBC supernatant preparations increased, suggesting that the release mechanism(s) for DMAs incorporated in proteins is still intact. Thus, in our RBC supernatant preparations, an enhanced DDAH activity resulting from the elimination of a DDAH inhibitor is the more likely explanation.

In conclusion, conceptually we can view WB as a well-mixed "tissue" of ~5 kg (in a 70-kg individual) circulating in intimate contact with the endothelium. Thus WB, by its accessibility to endothelial NOS and by virtue of the intrinsic enzymatic control capabilities that we have demonstrated here, is ideally suited for the ADMA-mediated control of endothelial NOS function. The general hypothesis supported by these data is that WB contains enzymatic systems that contribute to the control of plasma ADMA concentrations by acting as both a sink (via DDAH-mediated hydrolysis) and a source (via proteolytic degradation of ADMA-containing proteins) for ADMA. Dysfunction of this WB system could be acting alone or in concert with other contributors that promote endothelial cell dysfunction (e.g., reactive oxygen species, reduced tetrahydrobiopterin availability, L-arginine deficiency, etc.), resulting in diminished NO-mediated functions. This study supports the presence of enzymatic activities in WB reflective of both DDAH and protein-hydrolyzing enzymes that are in sufficient quantities to account for clinically relevant increases or decreases in plasma ADMA. Dysfunctional WB control of ADMA should thus be considered a potential contributor to increased plasma ADMA. Thus this WB system may contribute to, and provide an accessible therapeutic target for, diseases in which elevations in plasma ADMA have been demonstrated.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
We thank the Ravitz Foundation (Southfield, MI) for support of S. S. Billecke. L. A. Kitzmiller and J. J. Northrup were supported in part by National Institutes of Health T35 (2T35-HL-07690–21A1) Training Grant to the University of Michigan Medical School's Summer Biomedical Research Program.


   ACKNOWLEDGMENTS
 
We thank Katari Carello and Dr. Charles Zwemer for excellent and critical review of this manuscript.


   FOOTNOTES
 

Address for reprint requests and other correspondence: L. G. D'Alecy, 7744 Medical Sciences Bldg. II, Dept. of Molecular and Integrative Physiology, Univ. of Michigan Medical School, Ann Arbor, MI 48109-0622 (e-mail: lgdalecy{at}umich.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.


   REFERENCES
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 ABSTRACT
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