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Am J Physiol Renal Physiol 295: F388-F396, 2008. First published April 30, 2008; doi:10.1152/ajprenal.00405.2007
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Chronic L-arginine administration increases oxidative and nitrosative stress in rat hyperoxaluric kidneys and excessive crystal deposition

Ho-Shiang Huang,1 Ming-Chieh Ma,2 and Jun Chen3

1Department of Urology, National Taiwan University Hospital, Taiwan; 2School of Medicine, Fu Jen Catholic University, Hsinchuang, Taiwan; and 3Department of Urology, National Taiwan University Hospital, Yun-Lin Branch, Taipei, Taiwan, Republic of China

Submitted 31 August 2007 ; accepted in final form 26 April 2008


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Hyperoxaluric kidneys show an impaired diuretic response to acute infusion of L-arginine. In this study, we examined the chronic effect of L-arginine supplementation on CaOx crystal formation in hyperoxaluric rat kidneys. Eight groups were tested: control (received drinking water), L group (received L-arginine, 0.6%), LN group [received NG-nitro-L-arginine methyl ester (L-NAME, 10 mg/kg)], L + LN group (received L-arginine + L-NAME), HP group [received hydroxyl-L-proline (HP, 5%) mixed with chow to induce hyperoxaluria], L + HP group (received HP + L-arginine), HP + LN group, and L + HP + LN group. The duration was 42 days, and each group had eight animals. Urinary biochemistry and renal CaOx amounts were measured, as well as renal expressions of nitric oxide synthase (NOS) isoforms and NAD(P)H oxidase. The distribution of inducible NOS (iNOS), NAD(P)H oxidase, ED1-positive cells, and nitrotyrosine was examined by immunohistochemical and immunofluorescence studies, whereas superoxide production from the kidneys was examined by fluorescence spectrometric assay. Compared with the HP group, the L + HP group had excessive CaOx crystal accumulation and enhanced endothelial NOS (eNOS), iNOS, and NAD(P)H oxidase protein expression in the kidney. Urinary excretion of nitrotyrosine was markedly increased. Increased superoxide formation in the L + HP kidney was derived from NAD(P)H oxidase and uncoupled eNOS, and increased nitrotyrosine formation might derive from iNOS and ED1-positive cells that gathered around the CaOx crystals. L-NAME cotreatment (L + HP + LN group) reduced renal oxidative nitrosative stress and tubular damage, which were induced by L + HP. The results showed that chronic L-arginine treatment to the hyperoxaluric kidney with massive CaOx crystal deposition may have a toxic effect by enhancing intrarenal oxidative and nitrosative stress.

hyperoxaluria; nephrolithiasis; NAD(P)H oxidase; nitric oxide synthase

HYPEROXALURIA IS THE RESULT of either genetic or environmental factors that can lead to urolithiasis, nephrocalcinosis, metabolic acidosis, hematuria, and renal failure (19). Because there are many similarities between experimental nephrolithiasis and human kidney stone formation, the rat model of calcium oxalate (CaOx) nephrolithiasis can be used to investigate mechanisms involved in human kidney stone formation (13–16).

Nitric oxide (NO) plays a critical role in multiple renal processes, including the regulation of renal plasma flow, glomerular filtration rate, generation of renin and ANG II, and tubular reabsorption (5). Endogenous NO is synthesized by nitric oxide synthase (NOS) using L-arginine (L-Arg) as a substrate. Three isoforms of NOS, neuronal NOS (nNOS), inducible NOS (iNOS), and endothelial NOS (eNOS), are expressed in the kidneys (1, 7). Evidence has been accumulating that, in pathological states, iNOS may be overexpressed, and both eNOS and nNOS expression can be altered (1). It has been reported that L-Arg infusion in the kidneys can stimulate intrarenal NO production by enhancing NOS activity in vivo (8, 38). Intrarenal NO production stimulated by L-Arg also increases renal sodium excretion by a direct tubular action and by regulating vascular tone in the pressure natriuretic response in the renal medulla (2, 5, 8). However, the hyperoxaluric kidney showed a poor renal excretion when responding to acute saline loading (activates endogenous NO system) or exogenous L-Arg infusion, and this was associated with less urinary NO metabolites and cGMP excretion (17). These suggested that the hyperoxaluric kidney may not properly respond to the acute increased NO formation.

Our previous study showed that CaOx crystal deposition causes a change in the balance of antioxidant-oxidant, and macrophage infiltration in the kidney may contribute to this redox imbalance (13). Nitrosative stress is also induced in kidneys with renal stones (17, 29), and the production of reactive nitrogen species may cause tubular damage and exacerbate the disease.

The symptoms in many models of experimentally induced renal disease, such as cyclosporine-induced nephropathy, diabetic nephropathy, and ureteral obstruction, are improved by chronic dietary supplementation with L-Arg (11, 31). The administration of L-Arg to injured kidneys improves excretory response and is reported to reduce lipid peroxidation (LPO) in patients with diabetes mellitus (24). Aside from its beneficial effects, excessive NO has several cytotoxic effects, including peroxynitrite formation, NO-induced nitrosylation, and the inactivation of various enzymes (7). Therefore, we aimed to study the effects of L-Arg on CaOx crystal formation after hyperoxaluria. Because hydroxy-L-proline (HP), a physiological precursor of oxalate, has been reported to induce hyperoxaluria and CaOx crystal formation but without an associated metabolic acidosis when mixed in the diet at 5% (19), we therefore conducted the HP-induced hyperoxaluric model. Our present study unexpectedly showed that L-Arg caused oxidative and nitrosative stress in the hyperoxaluric kidney; this favored more crystal formation. We therefore explored the blockade effect of NOS by NG-nitro-L-arginine methyl ester (L-NAME) to see whether NOS inhibition is beneficial to the hyperoxaluric kidney.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals. Male Wistar rats (180–220 g) were housed at a constant temperature and on a light-dark cycle (light from 0700 to 1800). Animal care and experimental protocols followed the Guide for the Care and Use of Laboratory Animals (National Academy Press, Washington DC, 1997) and were reviewed and approved by the Institutional Animal Care and Use Committee, National Taiwan University School of Medicine.

The rats were divided into eight groups that underwent treatment for 42 days. The control group received drinking water, the L group received 0.6% L-Arg (Sigma, St. Louis, MO) (6) in drinking water, LN group received the nonselective NOS inhibitor L-NAME (10 mg/kg; Sigma) (1), and the L + LN group received 0.6% L-Arg and L-NAME. The last four groups were given chow mixed with 5% HP (wt/wt HP/chow) purchased from ICN Biochemicals (Aurora, OH) (19). The HP group received only this treatment, the L + HP group received HP and 0.6% L-Arg in the drinking water, the L + HP + LN group received HP with 0.6% L-Arg and L-NAME in the drinking water, and the HP + LN group received HP and L-NAME. The drugs were used at doses indicated as effective in the literature (1, 6, 19) and which prevented toxic activity. There were eight rats (n = 8) in each group.

During the experimental period, all groups had free access to standard rat chow. The animals were placed in metabolic cages 3 days before surgery for acclimatization.

Biochemical assays. Twenty-four-hour urine samples were collected 1 day before death and were analyzed using kits for oxalate (Sigma), citrate (R-Biopharm, Darmstadt, Germany), N-acetyl-β-glucosaminidase (tubular damage marker) (13), malondialdehyde (MDA, LPO marker) (kit from Oxis Research, Portland, OR), NO metabolites (nitrate and nitrite were purchased from Cayman Chemical, Ann Arbor, MI), and cGMP (Assay Designs Ann Arbor, MI), which had been used as markers of systemic NO production, and bioactivity (17). All samples were assayed in duplicate.

Creatinine (serum and urine), urine electrolytes (calcium and magnesium), and total urine protein were measured at the Central Laboratory of the National Taiwan University Hospital. Urinary supersaturation with respect to CaOx was assessed using the index proposed previously (35). Sediments in the 24-h urine sample were collected and examined for the presence of CaOx crystals by polarized microscopy, as described below. These were then weighed, placed in an oven for 48–72 h, and weighed daily until a stable weight was achieved. Myeloperoxidase (MPO) levels, an indicator of tissue neutrophil accumulation (1) in the kidneys, were measured colorimetrically (Assay Designs).

Urinary nitrotyrosine (NTS) level, an indicator of peroxynitrite formation, was also measured colorimetrically (Kamiya Biomedical, Seattle, WA). The presence of CaOx crystals in the kidneys was examined and graded as described previously (13). Each kidney was scored semiquantitatively as one of four grades (0, I, II, and III), ranging from "no crystal deposit" to "massive crystal deposits." Reduced glutathione (GSH) and oxidized glutathione (GSSG) levels, oxidative stress markers, in the kidneys were measured colorimetrically (Oxis Research), and the GSH redox ratio was calculated using the following equation: redox ratio (%) = 2 GSSG/(GSH + 2GSSG) x 100 (12).

Luminol-dependent chemiluminescence assay. We compared chemiluminescence (CL) counts on the kidney surface in the different groups. Details of the procedures are described in our earlier publication (13). CL was expressed as a counts per 10 s.

Fluorescence spectrometric assay of O2– production from the kidneys. Fluorogenic oxidation of dihydroethidium to ethidium was used as a measure of O2– (10). The isolated kidneys were homogenized in ice-cold buffer. After centrifugation at 6,000 g for 5 min at 4°C, the supernatant containing the membrane and cytosolic components was separated. The homogenates (20 µg) were then incubated at 37°C for 30 min with dihydroethidium (0.02 mM; Sigma-Aldrich Japan), salmon testes DNA (0.5 mg/ml), and various substrates for superoxide-producing enzymes and their inhibitors in a microtiter plate placed away from direct light.

Ethidium-DNA fluorescence was measured at an excitation of 480 nm and an emission of 610 nm using a fluorescence microplate reader (Moritex Bio-Science, Tokyo, Japan). NADH (0.1 mM) was used as a substrate for NAD(P)H oxidase. Diphenylene iodonium chloride (DPI, 0.1 mM) was then added to inhibit NAD(P)H oxidase. The effect of BH4 (0.01 mM) on L-arginine-induced O2– production was examined.

Immunohistochemical staining. Details of the procedures are described in our earlier publication (14). The paraffin-embedded tissue sections were incubated for 1 h at room temperature (RT) with mouse monoclonal antibody against NTS (1:100 dilution; Cayman Chemical) or against the macrophage/monocyte antigen ED1 (1:1,000 dilution; Serotec, Oxford, UK) (13). Bound primary antibody was detected using an ABC kit with anti-mouse IgG antibody and 3,3'-diaminobenzidine as substrate. Gill's hematoxylin was used as counterstain. The method for calculating the number of ED1-positive cells in these sections has been previously described (13).

Immunofluorescence examination of iNOS and gp91phox [NAD(P)H oxidase]. Rats were anaesthetized and then perfused transcardially, as described previously (13). The perfused kidney was quickly dissected and postfixed in 4% wt/vol paraformaldehyde in PBS and then stored in 10% sucrose in 4% paraformaldehyde solution at 4°C. Kidneys embedded in optimum cutting temperature compound (Tissue-Tek; Sakura Finetek, Torrence, CA) and frozen at –20°C were used to prepare 5-µm sections on a cryostat (Microm, Heidelberg, Germany) that were thaw-mounted on coated slides.

After being rehydrated and washed with PBS, the sections were processed for indirect immunofluorescence. After being blocked with 5% skim milk in PBS for 1 h at RT, the sections were incubated overnight at 4°C with rabbit anti-iNOS antibody (Santa Cruz) or mouse anti-gp91phox antibody (Santa Cruz) diluted 500-fold in 5% normal rabbit serum in PBS or 400-fold in PBS, respectively, and then incubated for 1 h at RT with rhodamine-Red-X-conjugated donkey anti-rabbit antibody (1:100 in 5% skim milk; Jackson Immuno Research, West Grove, PA) for iNOS, or conjugated with fluorescein isothiocyanate.

Tissue sections were examined on an Olympus BX51 microscope (Tokyo, Japan) with a fluorescent image analytic system (Diagnostic Instruments) at x200 magnification. Nuclei were counterstained by 4',6'-diamidino-2-phenylindole. The specificity of the antibody was tested by preincubation of the primary antibody with blocking peptide (Santa Cruz).

Western blot. Details of the procedures of Western blot analysis are described in our earlier publication (17). In brief, the same amount of protein from each preparation was loaded on each lane and nitrocellulose membrane (Amersham Bioscience, Buckingham, UK) and then incubated overnight at 4°C with monoclonal mouse antibodies against eNOS (1:500), iNOS (1:250), nNOS (1:500), NTS (1:250), poly(ADP-ribose) polymerase (PARP, 1:250) (all from BD Biosciences), gp91phox (1:2,000) (Santa Cruz Biotechnology, Santa Cruz, CA), or actin (1:1,000; Sigma) (Cedarlane, Ontario, Canada) in the blocking solution.

The density of the band in each lane was determined semiquantitatively using a densitometer with an image analytic system (Diagnostic Instruments) and expressed as the ratio of the density units for the band divided by those for actin.

Statistical analyses. Numerical data are presented as means ± SE. Differences between the groups were analyzed using the unpaired t-test or one-way ANOVA, with a post test of Duncan's multiple-range test. Differences were regarded as significant at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Body data and urinary biochemistry. Supplementation with 0.6% L-Arg resulted in a 2.26-fold higher daily intake of L-Arg (560 ± 38 mg/day in the L group vs. 248 ± 13 mg/day in the controls) (Table 1). Food consumption (data not shown) and body weight (Table 1) did not differ among the groups. Water intake was significantly increased in all L-treated groups, whereas kidney weight was significantly increased in all HP-treated groups (Table 1). Creatinine clearance did not differ among the groups.


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  		Table 1. Body data and urinary biochemistry of the eight study groups

 
Hyperoxaluria was seen in all HP-treated groups and was associated with significantly increased excretion of tubular markers. However, increments in the L + HP + LN group were significantly lower than those of the L + HP group. L-Arg supplementation resulted in a significant increase in urinary NOx and cGMP levels in the L and L + HP groups, which returned to normal levels after cotreatment with L-NAME (L + LN and L + HP + LN groups). Changes in the urinary pH value in the experimental groups had no statistical significance when compared with the controls (data not shown).

Compared with the control group, urinary excretion of NTS increased significantly in all HP-treated groups and was significantly higher in the L + HP group than in the HP and L + HP + LN groups (Table 1). The degree of urine CaOx supersaturation was significantly higher in all HP-treated groups than in the other groups (Table 1).

L-Arg enhanced CaOx crystal deposition. In Table 1 and Fig. 1, no obvious CaOx crystal deposition was seen in the control and L, LN, and L + LN groups (data not shown). However, massive accumulation of CaOx (grade III) (Table 1) associated with a dilated tubular structure was seen in the L + HP kidney (Fig. 1c) compared with the grade II seen in the HP group (Fig. 1A) and the grades I-II seen in the HP + LN group (Fig. 1B). This returned to grade II in the L + HP + LN kidney (Fig. 1D). Retained CaOx crystals were found both in renal cortex and medulla in all HP-treated groups; however, a great amount of crystal deposits was found in the tubular segments of renal cortex in L + HP kidney.


Figure 1
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  		Fig. 1. L-Arginine enhanced calcium oxalate (CaOx) crystal deposition. Representative micrographs under a polarizing microscope show details of the kidneys in each group. CaOx crystals are indicated by black arrows. A: rat treated with hydroxyl-L-proline (HP). B: rat treated with HP and NG-nitro-L-arginine methyl ester (L-NAME). C: rat treated with L-arginine and HP. D: rat treated with L-arginine and HP + L-NAME. No obvious CaOx crystal deposition was seen in the control and L-Arg (L), L-NAME (LN), and L + LN groups (data not shown). Reduced from x40 magnification.

 
The dry weight of urinary CaOx sediment (Table 1) was significantly higher in all HP-treated groups than in the control, L, LN, and L + LN groups. However, there was no significant difference among the control, L, LN, and L + LN groups, or among the four HP-treated groups.

l-Arg induced oxidative stress. CL levels in the renal surface of the left kidney and urinary excretion of lipid peroxides (MDA) were significantly increased in all HP-treated groups compared with the control group (Table 2). Among the HP-treated groups, CL levels were higher in the L + HP group than the HP and HP + LN groups but were not significantly different among the HP, HP + LN, and L + HP + LN groups.


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  		Table 2. Indexes of oxidative stress in the eight study groups

 
Changes in intrarenal oxidative stress were indicated by the amounts of reduced and oxidized GSH (Table 2). Significant decreases in GSH were noted in both the renal cortex and medulla in the HP-treated groups, whereas increased GSSG was only seen in the renal medulla of these groups. The increase in GSH and decrease in GSSG were more significant in the L + HP group than in the HP and L + HP + LN groups. The redox ratio increased in the whole kidney in all HP-treated groups, and changes in the L + HP group were higher than the HP and L + HP + LN groups.

L-Arg enhanced nitrosative stress. Figure 2 shows representative blots of the three types of NOS from two rats in each group (top) and the statistical results for all eight rats (bottom). The expression of eNOS increased significantly in the L + HP group when compared with the control and HP groups, whereas the expression of iNOS was significantly higher in the medulla of HP-treated rats and in the cortex of HP, L + HP, and HP + LN rats when compared with the controls. The expression of eNOS was not significantly different in the HP and L + HP + LN groups when compared with the controls. There was no significant change in nNOS expression in all of the experimental groups when compared with the controls.


Figure 2
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  		Fig. 2. Nitric oxide synthase (NOS) expression. Top: representative blots of endothelial (eNOS), inducible (iNOS), and neuronal (nNOS) NOS in the renal cortex and medulla of two rats from each of the different groups. Bottom: results for all 8 rats. A1: and B1: results of C, L, L + LN, HP, L + HP, and L + HP + LN groups. A2 and B2: results of C, LN, HP, and HP + LN groups. C, control; P, positive controls as described in METHODS. P < 0.05 compared with the control group (*), compared with the HP group (#), and compared with the L + HP group ({dagger}).

 
Because iNOS(+) fluorescence signal could be found in the cortex and outer medulla of normal control rats (see GoGoGoFig. 6B), iNOS stained strongly in the medulla as a continuous glandular pattern in HP and L + HP + LN rats, whereas the signal in the cortex was the same as those in the controls. In L + HP rats, the iNOS(+) signal was stronger than the controls and formed not only a glandular pattern but also a circular pattern in both the cortex and medulla.


Figure 3
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  		Fig. 3. Changes in renal nitrotyrosine (NTS) expression in the different groups. The photographs are representative results from an untreated control rat (A), a rat treated with HP (B), and a rat treated with L + HP (C) by immunohistochemical staining. All sections were evaluated by light microscopy. NTS-positive areas were stained brown. The black arrows indicate crystal deposition in the tubular lumen. Reduced from x200. *P < 0.05 compared with the control group.

 

Figure 4
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  		Fig. 4. L-Arg induced cell infiltration. Representative micrographs show the presence of the monocyte/macrophage antigen ED1 in renal sections from each group. ED1-positive cells are indicated by arrows. A: controls. B: rat treated with HP. C: rat treated with L-arginine and HP. D: statistical results for each group. E: levels of myeloperoxidase (MPO) in the renal cortex and medulla in each group. Reduced from x200. P < 0.05 compared with the control group (*), compared with the HP group (#), and compared with the L + HP group ({dagger}).

 

Figure 5
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  		Fig. 5. L-Arg increased oxidative proteins and cell injury. Levels of gp91phox (A) and full-length (full) and cleaved poly(ADP-ribose) polymerase (PARP; B) in the renal cortex and medulla in each group. The expression of gp91phox and PARP in the LN and HP + LN groups was similar to the control and HP groups, respectively (data not shown). Representative blots for two rats and the statistical results are shown in the top and bottom. P < 0.05 compared with the control group (*), compared with the HP group (#), and compared with the L + HP group ({dagger}).

 

Figure 6
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  		Fig. 6. Immunofluorescence findings of the influence of L-Arg supplementation on the expression of gp91phox (A) and iNOS (B) in the renal cortex and medulla in each group. White arrows indicate the area surrounded by iNOS(+) signals. Representative fluorescence image micrographs of the renal cortex and medulla are shown (magnification x200).

 
Figure 3 demonstrated in situ formation of NTS in the kidney by immunohistochemical staining. Because NTS was found not only in tubular cells but also dispersed in the interstitial space in HP and L + HP kidneys, it was difficult to count the NTS-positive signals. Qualitatively, minor staining was seen in the control (Fig. 3A), L, LN, and L + LN kidneys (data not shown), but increased staining was seen in all HP-treated kidneys (Fig. 3B). In the L + HP kidneys, dense staining was seen in the tubular cells and widely dispersed in the interstitium (Fig. 3C).

L-Arg induced macrophage/monocyte infiltration. Figure 4 shows micrographs of anti-ED1 antibody staining, demonstrating infiltration of macrophages/monocytes in the kidney. Only a few ED1-positive cells were found in kidney sections of the control [0.8 ± 0.5 cells/high-power field (HPF)] (Fig. 4A), L (1.1 ± 0.6 cells/HPF), LN (0.9 ± 0.5 cells/HPF), and L + LN groups (data not shown). In the L + HP group, the ED1 cells/HPF was increased to 23.7 ± 2.1 and was mainly located in the peritubular interstitium (Fig. 4C). The number of ED1 cells/HPF was significantly reduced to 8.0 ± 1.6 in the L + HP + LN group (data not shown), similar to the 10.3 ± 2.6 cells/HPF seen in the HP group (Fig. 4B) and 9.6 ± 2.5 cells/HPF of the HP + LN group (data not shown). However, in all of the HP-treated groups, some of the ED1-positive cells gathered around the CaOx crystals and formed bigger areas of positive signals, which made cell counting more difficult.

Using the amount of MPO to represent the infiltration of neutrophils and macrophages (1), as shown in Fig. 4E, changes in MPO levels were consistent with those of ED1-positive cell infiltration.

L-Arg caused increased oxidative enzyme proteins and cell injury. Expression of gp91phox, a major component of NAD(P)H oxidase, was increased in the medulla in the HP and L + HP + LN groups and in whole kidney of the L + HP group (Fig. 5A). In the HP + LN group, the expression of gp91phox and PARP in the kidney was similar to the HP group (data no shown).

Activation of PARP is an early DNA damage response as a consequence of oxidative injury (4). Figure 5B shows that levels of full-length and cleaved PARP in the L + HP and HP groups were increased significantly compared with those of the control group, and those in the L + HP group were higher (P < 0.05).

gp91phox stained strongly in the medulla as a continuous glandular pattern in HP-treated rats, whereas the signal in the cortex was the same as in the controls (Fig. 6A). In L + HP rats, the gp91phox(+) signal was stronger than in the controls and formed not only a glandular pattern but also in the interstitium of both the renal cortex and medulla (Fig. 6A).

NAD(P)H oxidase and eNOS caused O2– superoxide production. In Fig. 7, spontaneous production of O2– was significantly higher in HP-treated rats than in control rats. Incubation with NADH resulted in a significant increase in O2– production in HP-treated rats, where the L + HP group showed much higher increases than the HP and L + HP + LN groups. Enhanced activity of NAD(P)H oxidase in HP-treated rats was confirmed by the inhibition of NADH-induced O2– overproduction by DPI.


Figure 7
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  		Fig. 7. Fluorescence spectrometric assay of superoxide (O2–) production in the kidneys. Values are means ± SE and are expressed relative to the control. –, No treatment in each group. P < 0.05 compared with the control group (*), compared with the HP group (#), and compared with the L + HP group ({dagger}).

 
BH4 is an essential cofactor of eNOS, and its deficiency is reported to decrease the bioavailability of NO (10). Incubation with BH4 reversed L-Arg-induced O2– overproduction in the HP-treated groups.


   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
As observed in hyperoxaluric rats, HP-induced hyperoxaluria and renal CaOx crystal accumulation was associated with increased LPO in the hyperoxaluric kidney and renal tubular injury. LPO may result from increased superoxide formation and decreased antioxidant system in hyperoxaluric kidneys (13, 19). CaOx crystal deposition was associated with macrophage infiltration, and MPO levels increased (19), which might have contributed to the nitrosative stress in hyperoxaluric kidney. Through cotreatment with L-Arg and HP, more NTS-, ED1-, and iNOS-positive cells gathered around the increased CaOx crystals in the kidney. Increased oxidative stress was indicated by reduced antioxidant compound (GSH), increased O2– production, and oxidative injury (MDA and PARP), which can be partially reversed by the co-administration of L-NAME. Therefore, these results clearly showed that chronic L-Arg treatment is not beneficial to hyperoxaluric kidneys with CaOx crystal deposition but instead enhances intrarenal oxidative and nitrosative stress.

However, L-Arg administration did not change urinary ion activity and excretion of urine CaOx sediments. As such, grade II CaOx crystals were still found in the kidneys of L + HP + LN rats. L-NAME administration also did not change the urinary ion activity of urine CaOx sediments, since grades I-II crystals were found in the kidneys of HP + LN rats.

Aside from phagocytic cells, the expression of NAD(P)H oxidase has also been detected in kidney cells such as proximal tubular cells, glomerular mesangial cells and cells of the thick ascending loops of Henle (36). NAD(P)H oxidase is a multicomponent enzyme complex, and both gp91phox and Rac1 are critical components of endothelial NAD(P) oxidase (37). It is the primary source of O2– in the kidneys under physiological conditions, and the induction of oxidative stress or overproduction of O2– often depends on its increased expression or activity (21).

Involvement of NAD(P)H oxidase in the production of free radicals when renal epithelial cells are exposed to oxalate ions or crystals has been reported in a cell culture study (36). In this study, increased NAD(P)H oxidase protein expression and O2– production were found in the kidneys of L + HP rats, which might have led to more oxidative injury than those treated with HP only but could still be reversed by L-NAME cotreatment. However, gp91phox protein expression showed no significant change in the kidneys of L-Arg-treated rats. We hypothesize that, to enhance NAD(P)H oxidase expression after L-Arg administration, an oxidative stress needs to be present, such as reduced GSH content. But the exact mechanisms need further evaluation.

eNOS has been found to be an important source of vascular O2– excess in adjuvant-induced arthritis (10). In certain conditions, it may cause uncoupling, which generates O2–, and uncoupled eNOS is reported to be a possible prominent source of endothelial O2– in hypertension or diabetes (27, 34). It has been reported that eNOS generates O2– rather than NO when exposed to oxidant stress, including ONOO–, or when deprived of its cofactor BH4 or substrate L-Arg (10, 22).

In the present study, L-Arg administration did not cause an increase in eNOS expression or O2– production in the kidneys of normal rats. However, L + HP rats have more eNOS protein expression in the kidneys and reduced O2– production when incubated with BH4. This suggests that BH4 may be deficient in the kidneys of L + HP rats and that its addition reverses L-Arg-induced O2– increase. Combining these results, we speculate that increased O2– production by NAD(P)H oxidase may decrease BH4 bioavailability (23) and results in eNOS uncoupling, leading to further amplification of oxidative stress in the kidneys of L + HP rats.

In addition to NAD(P)H oxidase and uncoupled eNOS, other mechanisms may contribute to oxidative stress in hyperoxaluric kidneys, such as xanthine oxidase (30) and mitochondria (25). We examined the expression of xanthine oxidase in our preliminary study and found no significant change (data not shown). Meimaridou et al. (25) had reported that mitochondria-derived superoxide was the source of oxidative stress in renal epithelial cells treated with CaOx crystals.

The additive role of oxidative and nitrosative stress in epithelial cell injury has been examined by Peresleni et al. (28). Exposure of epithelial BSC-1 cells to oxidant stress (0.5 mM H2O2) resulted in increased NO release and coincided with the appearance of immunodetectable iNOS expression. This may be the mechanism of increased iNOS protein expression in the cortex of L + HP rats. Combining the findings of immunolabeling for NTS, ED1, and iNOS, we speculate that the enhanced iNOS expression may derive from two different origins: one from the renal tubules, such as the proximal tubules and inner medullary collecting duct (9), and the other is from macrophages/monocytes, which forms a circular shape and goes to the interstitium.

NO generated by iNOS, especially its metabolite peroxynitrite (as detected with NTS antibodies), is cytotoxic to renal tubular epithelium (9). Enhanced NTS formation in the kidneys and urine without an increase in urinary NOx and cGMP excretion was observed in L + HP rats when compared with the L-Arg-treated group, and that was the possible mechanism that NO reacted with superoxide yielding peroxynitrite, which itself interacted with tyrosine residues to form NTS and then to be sequestered in tissues or excreted in the urine rather than metabolized (17).

Enhanced iNOS protein expression in the renal cortex of L + HP rats can be reversed when L-NAME is used as a cotreatment. We speculate that L-Arg administration to induce iNOS protein expression and nitrosative stress in the kidneys needs oxidative stress to exist in those kidneys already. In stress-induced gastric damage, nuclear factor-{kappa}B (NF-{kappa}B) is activated by reactive oxygen species (ROS), which then induces iNOS gene expression (18). Whether the increased iNOS protein expression after chronic L-Arg administration is through NF-{kappa}B activation needs further evaluation.

During induction of CaOx nephrolithiasis, infiltration of ED1-positive cells into the kidneys and these cells surrounding CaOx crystals have been found in animal models (13, 19). L-Arg administration reduces renal macrophage infiltration in obstructive nephropathy after a 3-day L-Arg administration (32). However, in the present study, renal ED1-positive cell infiltration was not abolished in L + HP rats after a 6-wk L-Arg administration. Instead, ED1 infiltration was partially reversed after cotreatment with L-NAME. Therefore, combined with earlier data, we hypothesize that the critical factor that determines the effect of L-Arg administration is the presence of oxidative stress.

For example, Pragasam et al. (30) reported restored antioxidant enzyme levels to normal and no stone formation in the kidneys after L-Arg cosupplementation to 0.75% ethylene glycol-treated rats for 28 days. In addition to oxalate, CaOx crystals are associated with multinucleated giant cells and macrophages that are injurious to renal epithelial cells (19). Therefore, if renal oxidative stress is not increased and no CaOx crystal accumulates in hyperoxaluric kidney, L-Arg cotreatment has an antilithic effect. We performed the same experiments on day 14 in a preliminary study (data not shown), and the results were consistent with those of Prakasam et al. (30).

Our findings are consistent with those found in the atherosclerotic model in which severity of atherosclerosis determines the L-Arg treatment to be beneficial or harmful to atherosclerosis. L-Arg (2.25%) treatment of severely atherosclerotic rabbits (induction for 24 wk) leads to increased O2– production (34), whereas reduced O2– and decreased plaque progression were found in rabbits after 12 wk induction of atherosclerosis and cotreated with L-Arg (2.25%) in Böger's study (3). The source of O2– in severely atherosclerotic aorta may be NAD(P)H oxidase, xanthine oxidase, or uncoupled eNOS (3, 34).

In summary, the present study indicates that L-Arg administration can enhance NAD(P)H oxidase, eNOS, and iNOS protein expression in the hyperoxaluric kidneys with massive CaOx crystal deposition. Increased ROS production by NAD(P)H oxidase may result in eNOS uncoupling and leads to further amplification of oxidative stress in the kidneys. Increased macrophage/monocyte infiltration and iNOS expression in the kidneys are likewise associated with increased nitrosative stress in L-Arg-treated hyperoxaluric rats. This increased renal oxidative and nitrosative stress can further reduce renal antioxidant ability and result in more tubular damage and CaOx crystal accumulation in the kidneys. Increased CaOx crystal accumulation in the kidney after long-term induction of nephrolithiasis may cause excessive ROS production, which may result in the cotreatment of L-Arg lacking an antilithic effect.


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This study was supported by grants from the National Science Council of the Republic of China (NSC 94-2320-B-030-015) and the National Taiwan University Hospital (96-N-004).


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. Chen, Dept. of Urology, National Taiwan Univ. Hospital, Yun-Lin Branch 582, Section 2, Yun-Lin Road, Douliou, Taiwan (e-mail: chenjun{at}ha.mc.ntu.edu.tw)

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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 ABSTRACT
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