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(Circulation. 1995;92:1876-1882.)
© 1995 American Heart Association, Inc.

Articles

In Vivo Spin Trapping of Glyceryl Trinitrate–Derived Nitric Oxide in Rabbit Blood Vessels and Organs

Alexander Mülsch, PhD; Peter Mordvintcev, PhD; Eberhard Bassenge, MD; Frank Jung, PhD; Bernd Clement, PhD; Rudi Busse, MD, PhD

From the Center of Physiology, Johann-Wolfgang-Goethe University Clinic, Frankfurt (A.M., P.M., R.B.); the Department of Applied Physiology, University of Freiburg (E.B.); and the Institute of Pharmaceutical Chemistry, Christian-Albrechts-University, Kiel (F.J., B.C.), Germany.

Correspondence to Alexander Mülsch, PhD, Zentrum der Physiologie, Klinikum der Universität Frankfurt, Theodor-Stern-Kai 7, D-60590 Frankfurt, Germany.

	Abstract

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Background The objectives of this study were (1) to assess glyceryl trinitrate (GTN)–derived nitric oxide (NO) formation in vascular tissues and organs of anesthetized rabbits in vivo, (2) to establish a correlation between tissue NO levels and a biological response, and (3) to verify biotransformation of GTN to NO by cytochrome P-450.

Methods and Results NO was trapped in tissues in vivo as a stable paramagnetic mononitrosyl-iron-diethyldithiocarbamate complex [NOFe(DETC)₂]. After removal of the tissues, NO was determined by cryogenic electron spin resonance spectroscopy. NO formation in vitro was assessed by spin trapping and by activation of soluble guanylyl cyclase. The GTN-elicited decrease in coronary perfusion pressure was monitored in isolated, constant-flow perfused rabbit hearts. NO was not detected in control tissues. In GTN-treated rabbits, NO formation was higher in organs than in vascular tissues and higher in venous than in arterial vessels. In isolated hearts, ventricular NO levels and decreases in coronary perfusion pressure achieved by GTN were closely correlated. Purified cytochrome P-450 catalyzed NO formation from GTN in a P-450–NADPH reductase– and NADPH–dependent fashion.

Conclusions Since GTN-derived NO formation in myocardial tissue correlates to the GTN-elicited vasodilator response, we conclude that GTN-derived NO detected in vivo correlates with the systemic effects of GTN. Therefore, the higher rate of NO formation detected in veins compared with arteries explains the preferential venodilator activity of GTN. High NO formation in cytochrome P-450–rich organs in vivo and efficient NO formation from GTN by cytochrome P-450 in vitro highlights the importance of this pathway for NO formation from GTN in the intact organism.

Key Words: arteries • spectroscopy • glyceryl trinitrate • nitric oxide • veins

	Introduction

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GTN elicits vasodilatation by generating NO from one of its nitroester groups, which activates soluble GC and the cGMP-dependent effector cascade that finally leads to relaxation of the vascular smooth muscle.^{1 2 3 4 5} At the cellular level, indirect evidence shows that cultured vascular endothelial cells,⁶ smooth muscle cells,⁷ kidney epithelial cells,⁸ and lung fibroblasts⁹ generate NO from GTN, suggesting a widespread expression of the GTN-NO pathway in vascular and nonvascular tissues. Although this mechanism of GTN action is now generally accepted, a correlation between GTN-elicited tissue NO levels and a biological response has never been established. So far, NO formation in blood and organs of living mice could be detected only after application of toxic doses (25 and 40 mg/kg) of GTN.^{10 11} Recently, nitrosylhemoglobin was detected in blood of patients treated with therapeutic doses of GTN,¹² but other tissues were not assessed. Therefore, the efficiency of vascular and nonvascular tissues in generating NO from GTN in vivo is still unknown. For instance, venous capacitance vessels are approximately one order of magnitude more sensitive to GTN than most arterial conductance and resistance vessels.¹³ In fact, the beneficial effect of GTN in angina pectoris therapy relies on this discrepant vasodilator responsiveness: predominant venodilatation results in preload reduction^{13 14} and economizes heart work. On the other hand, the relative insensitivity of the myocardial resistance bed (small arteries and arterioles) is also beneficial, since it prevents a coronary "steal" effect.¹⁵ Since these differences are not evident with authentic NO and S-nitrosothiols,¹⁵ they are unlikely to originate at the level of activation of soluble GC and/or the cGMP effector cascade but may reflect a different efficiency of vascular tissues to generate NO from GTN.^{15 16 17}

The present study was performed (1) to assess NO formation from GTN in vascular tissues and several organs of rabbits in vivo, (2) to reveal a relation between GTN-elicited tissue NO levels and a biological response of this tissue, and (3) to assess whether tissue-specific differences in GTN-derived NO formation exist. Furthermore, since the molecular mechanism of NO release from GTN in vivo is still elusive, we analyzed the catalytic activity of cytochrome P-450 in generating NO from GTN.

	Methods

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Isolated Rabbit Heart
Beating hearts were removed from anesthetized (60 mg/kg sodium pentobarbitone) and heparinized (3000 U IV) New Zealand White rabbits (2.5 to 4 kg; SAVO) of either sex according to the Langendorff technique.¹⁸ The hearts were perfused through the aortic stump with modified Krebs-Henseleit solution (pH 7.4, 37°C, continuously gassed with 95% O₂/5% CO₂) containing (in mmol/L) NaCl 118.0, NaHCO₃ 24.0, glucose 5.0, CaCl₂ 2.5, KCl 4.0, KH₂PO₄ 1.2, MgSO₄ 1.0, and pyruvate 2.0, with perfusion rate adjusted to generate a CPP of 70 mm Hg (30 to 40 mL/min). CPP was monitored with a pressure transducer (Gould P2310) connected to the side arm of the aortic perfusion cannula. Isovolumetric left ventricular pressure was measured with a fluid-filled latex balloon connected to a second pressure transducer (Gould CP-01). The heart rate was derived from the left ventricular pressure signal.

After an equilibration period, the NO synthase inhibitor N^G-nitro-L-arginine (30 µmol/L) was continuously infused to raise resting CPP, thereby increasing the absolute value (in mm Hg) of the CPP decrease to a maximally effective dose of GTN. When CPP had stabilized (after about 15 minutes), GTN dissolved in 50 µL Krebs-Henseleit solution was injected into the perfusion line proximal to the heart as a bolus (0.1, 0.25, and 1 µmol) to elicit a transient decrease in CPP. After CPP returned to baseline, FeSO₄ (0.3 µmol/L) dissolved in Krebs-Henseleit solution was infused for 15 minutes. This procedure lowered the detection limit of NO, since exogenous iron reduced formation of the Cu(DETC)₂ complex, which interfered with ESR determination of NO (see below). Five minutes after cessation of FeSO₄ infusion, DETC (50 µmol/L) was continuously infused. After 10 minutes, a second bolus of GTN was applied during continuous infusion of N^G-nitro-L-arginine and DETC, and the perfusion was stopped 5 minutes later. Then the hearts were quickly disconnected from the perfusion line, and the ventricle was cut into small pieces and frozen in liquid nitrogen for ESR recording. CPP changes were evaluated for peak responses and for area under the curve of decrease in CPP versus time.

Animal Experiments
The following protocol was approved by the local authorities according to German regulations on experimental animal research. New Zealand White rabbits were anesthetized (Nembutal 20 mg/kg IV, left ear), and GTN (0.5 mg/kg) was infused (30 mL/h IV, right ear) for 20 minutes, concomitantly with DETC (200 mg/kg). DETC infusions were well tolerated by the animals without signs of excitation. Immediately after cessation of the infusion, the animals were killed by an overdose of Nembutal (50 mg/kg IV). The carotid artery was opened, and a sample of blood was quickly collected for later ESR analysis. Subsequently, tissues of interest, eg, abdominal and thoracic aorta, femoral arteries, vena cava, mesenteric bed, heart, lung, liver, kidney, spleen, and skeletal muscle (quadriceps), were quickly excised (within 10 minutes) and frozen for ESR analysis in liquid nitrogen, as described previously.¹⁹

Background of NO Spin Trapping In Vivo
DETC distributes freely within the organism.¹⁹ It chelates intracellular free iron ions to form a water-insoluble Fe(DETC)₂ complex that is deposited in hydrophobic cell compartments.^{11 19} The Fe(DETC)₂ complex avidly and selectively scavenges NO, generating a paramagnetic mononitrosyl-iron complex, NOFe(DETC)₂.^{11 19} This complex exhibits a characteristic anisotropic ESR signal, which is characterized by g-factors g=2.035, g||=2.02 (=perpendicular; ||=parallel), with a triplet hfs at g and a splitting constant of 1.3 mT^{11 19} (Fig 1, top).

View larger version (11K): [in this window] [in a new window]   Figure 1. Detection of GTN-derived NO formed in rabbit tissues in vivo. Representative ESR spectra of frozen pure NOFe(DETC)2 complex dissolved in DMSO (upper recording) and of frozen tissues obtained from DETC- (Control) or DETC- and GTN-treated (GTN) anesthetized rabbits (recordings below). The NOFe(DETC)2 complex exhibits an anisotropic ESR signal with g values g 2.035, g|| 2.02, and a triplet hfs at g, labeled 1, 2, and 3. The control heart exhibited ESR features of the paramagnetic copper-DETC complex only. The recordings of tissues taken from a GTN-treated rabbit show in addition the third hfs line of the NOFe(DETC)2 complex, labeled 3. The intensity of this signal was taken for calculation of tissue NO concentrations. Here, these amounted to 0.6, 0.1, 0.5, and 1.5 µmol/L in the heart, abdominal aorta (abd. aorta), mesenteric bed (mes. bed), and lung, respectively. 1 indicates the position of the first hfs line of the NOFe(DETC)2 complex, which was clearly visible in the lung. Instrument settings were as detailed in "Methods." The direction and calibration of the magnetic field are indicated by B and 2.6 mT, respectively.

Calculation of Tissue NO Concentrations
The concentration of NO trapped (nmol/g tissue) was calculated from the amplitude of the third high-field hfs line, which is not masked by other ESR signals, such as that of the copper-DETC complex (see "Results"). The concentration of the NOFe(DETC)₂ complex was calculated by comparing its ESR signal amplitude with that of the g=2.039 ESR feature of the dinitrosyl-iron-di-L-cysteine complex recorded at known concentration.¹⁹ The ratio of both amplitudes at equal concentrations is 1:1.3 (mononitrosyl versus dinitrosyl complex), as calculated previously by double integration. For samples shorter than 20 mm (blood vessels), a correction factor was applied, derived from a calibration curve of the ESR signal intensity plotted versus the sample length. This relation was obtained by measuring the signal intensity of cylindrical (4.5-mm-diameter) standards (dinitrosyl-iron-di-L-cysteine complex; 0.1 mmol/L) of different lengths (2 to 20 mm).

Recording of ESR Spectra and Estimation of the Tissue Concentration of Traps
The ESR spectra were recorded at 77 K on a Brucker 300E spectrometer at a frequency of 9.33 GHz, modulation frequency 100 kHz, modulation 0.5 or 1 mT, microwave power 20 mW, and time constant 0.05 seconds. After the first ESR analysis, the NO trapping capacity (concentration of iron-DETC complex) of selected tissues was assessed as described previously.¹⁹ In short, the tissues were thawed, exposed to gaseous NO (400 mm Hg pressure), refrozen, and analyzed again by ESR spectroscopy. The concentration of mononitrosyl-iron complex detected under these conditions is equivalent to the concentration of traps. This value also provides an estimate of the so-called labile nonheme iron pool, which interacts with DETC.¹⁹

Spin Trapping of NO in Aqueous Phase
PDTC was used to detect release of NO from GTN in aqueous solution in vitro, since the iron- and mononitrosyl-iron complexes of PDTC are soluble in water, in contrast to the DETC complexes. The ESR signals of the DETC and the PDTC mononitrosyl-iron complex are identical and exhibit the same magnetic saturation behavior (unpublished results). Therefore, the concentration of NO trapped by Fe(PDTC)₂ could be calculated from the ESR signal amplitude as described for the DETC complex.

The stock trapping solution was prepared by mixing 1 part of aqueous FeSO₄ (1 mol/L) solution and 19 parts of PDTC (0.2 mol/L) in DMSO. NO donors were incubated at 37°C in HEPES buffer (15 mmol/L, pH 7.4, 0.5 mL) containing DTT (10 mmol/L), superoxide dismutase (1 µmol/L), and 1:100 diluted stock trapping solution (final concentration, 2 mmol/L PDTC and 0.5 mmol/L Fe²⁺). Under these conditions, trapping of NO from various sources (NO gas, sodium nitroprusside, SIN-1, S-nitroso-L-cysteine) was optimal with respect to recovery of NO (approaching 100%), linearity (from 0.1 to 100 µmol/L NO), sensitivity (100 nmol/L NO), specificity (specific for NO and NO-releasing compounds), and stability of the NOFe(PDTC)₂ complex (stable for more than 2 hours) (data not shown). It should be noted that oxygen levels were strongly reduced in these incubates owing to oxygen consumption by DTT (measured by an oxygen-sensitive electrode). For assessment of cytochrome P-450–catalyzed NO formation, the trapping solution was incubated with GTN (30 µmol/L), NADPH (300 µmol/L), L--dilauroyl-phosphatidylcholine (10 µmol/L), pure cytochrome P-450 (400 pmol), and cytochrome P-450–NADPH reductase (400 pmol) for either 10 or 60 minutes at 37°C. Subsequently, the samples were frozen in liquid nitrogen and kept at -70°C until analyzed by ESR spectroscopy.

Detection of NO by GC
The activity of soluble GC purified to apparent homogeneity from bovine lung was assessed by formation of [³²P]cGMP from [³²P]GTP, as described previously.⁷ GC (4 µg protein/mL) was incubated at 37°C for 3, 10, or 30 minutes, as indicated in the legend to Fig 4, in a triethanolamine hydrochloride–buffered solution (30 mmol/L, pH 7.4) containing 200 µmol/L [³²P]GTP (0.2 µCi), 100 µmol/L unlabeled cGMP, 4 mmol/L MgCl₂, 100 nmol/L erythrocyte superoxide dismutase, 0.1 mg/mL bovine -globulin, 10 mmol/L creatine phosphate, 120 µg/mL creatine phosphokinase (1 U/mL), and 200 µmol/L NADPH, purified rabbit liver cytochrome P-450, and cytochrome P-450–NADPH reductase, as indicated. Incubations were stopped by coprecipitation of nucleotides with zinc carbonate. After centrifugation, [³²P]cGMP was isolated from the supernatant by chromatography on acid alumina and was determined by liquid scintillation counting in a ß-counter (Packard). The specific activity of GC (nanomoles of cGMP formed per milligram GC per minute incubation time) was calculated as described.⁷ In the absence of NO, soluble GC exhibited a basal activity, which was subtracted from the values shown in Fig 4.

View larger version (17K): [in this window] [in a new window]   Figure 4. Graphs showing that rabbit liver cytochrome (Cyt.) P-450 catalyzes the formation of an activator of soluble GC from GTN. GC activity (nmol cGMP formed per mg GC per minute incubation time) was determined by formation of [32P]cGMP from [32P]GTP (200 µmol/L, 0.2 µCi) at 37°C. Basal GC activity (absence of GTN and P-450) was 45±5 nmol · mg-1 · min-1. Data show the increase in GC activity above basal (mean±SEM) from three determinations performed in duplicate. SEM is indicated by error bars, which in most cases are smaller than the symbols. A, GC activation by increasing concentrations of P-450 at constant GTN (100 µmol/L). Incubations proceeded for 10 minutes. GTN already elicited an increase in GC activity (45 nmol · mg-1 · min-1) in the absence of P-450. NADPH (200 µmol/L) enhanced GC activation by P-450/GTN (, -NADPH; , +NADPH). B, Time course of GC activation by P-450 (200 nmol/L) and GTN (100 µmol/L). Incubations proceeded for 1, 3, 10, and 30 minutes, either in the absence () or in the presence () of P-450. C, GC activation by increasing concentrations of GTN in the absence of P-450 (), in the presence of P-450 (200 nmol/L; ), and in the presence of P-450 and NADPH (200 µmol/L; ). Incubations proceeded for 10 minutes.

Materials
Cytochrome P-450 and cytochrome P-450–NADPH reductase were purified to apparent homogeneity from rabbit liver.²⁰ Nembutal was from Sanofi Winthrop. PDTC was a generous gift of Dr N. Frank, German Cancer Research Center, Heidelberg. GTN was provided as a trituration in lactose (10% GTN) by Pohl-Boskamp. All other biochemicals were obtained in the highest purity available from Sigma.

Statistics
Data are presented as mean±SEM. The significance of differences between NO concentrations achieved in venous and arterial tissues was assessed by one-way ANOVA, followed by the Bonferroni t test. A value of P<.05 was considered significant.

	Results

Top Abstract Introduction Methods Results Discussion References

 
NO Formation In Vivo
Frozen tissues obtained from DETC-treated control animals exhibited ESR spectra that were dominated by the resonance of the copper-DETC complex,^{11 19} as shown for a control heart in Fig 1. The intensity of this ESR signal varied considerably between different tissue types, representing differences in the content of the Cu(DETC)₂ complex (data not shown). The ESR signal of NOFe(DETC)₂ was absent from control tissues, indicating that endogenous, L-arginine–derived NO formation⁴ was below the detection limit of our ESR installation. In contrast, tissues removed from GTN-infused rabbits exhibited ESR features of the NOFe(DETC)₂ complex. In blood vessels, spleen, and heart, only the third hfs line of this ESR signal was clearly visible (labeled 3 in Fig 1), while the first and second hfs lines, clearly discernible in the ESR spectrum of the pure NOFe(DETC)₂ complex (labeled 1 and 2 in the top recording, Fig 1), were masked by the resonance of the Cu(DETC)₂ complex. With higher NO levels, such as observed in the lung, liver, and kidney, the first hfs line of the NOFe(DETC)₂ complex also became visible (labeled 1 in Fig 1). Evaluation of all tissues (from eight rabbits) revealed significant differences in NO formation achieved by GTN treatment (Fig 2). It was significantly higher in the vena cava and the mesenteric bed than in the femoral artery and thoracic and abdominal aorta (P<.05). The concentration of trapped NO was considerably higher in liver, lung, and kidney, up to 1.65 nmol/g. GTN-derived NO was not detectable in whole blood or in skeletal muscle (quadriceps) (Fig 2).

View larger version (31K): [in this window] [in a new window]   Figure 2. Bar graphs showing GTN-derived NO formation in rabbit tissues in vivo. Tissue concentrations of NO trapped as NOFe(DETC)2 complex (mean±SEM) derived from the ESR recordings taken on eight rabbits treated with DETC and GTN are shown. For details see text. Ao. thor. indicates thoracic aorta; Ao. abd., abdominal aorta; A. fem., femoral artery; Mes. bed, mesenteric bed; and V. cava, vena cava. *Significant difference between venous and arterial tissues (P<.05).

After the first ESR analysis for GTN-derived NO, a second ESR analysis was performed for estimation of the concentration of NO traps [Fe(DETC)₂ complexes]. This concentration was significantly lower in blood vessels than in organs (Table 1, left column; P<.05). However, in all tissues, only a small fraction (0.5% to 4.5%) of the total amount of traps available was required for binding of GTN-derived NO (Table 1).

View this table: [in this window] [in a new window]   Table 1. NO Trapping Capacity [Concentration of Fe(DETC)2] in Tissues of DETC-Treated Rabbits

Correlation Between Myocardial NO Formation and Relaxation of Coronary Resistance Vessels
In isolated rabbit hearts (n=6), continuous infusion of N^G-nitro-L-arginine (30 µmol/L) increased resting CPP from 73±6 to 111±8 mm Hg. GTN applied as a bolus (0.1, 0.25, and 1 µmol) elicited a dose-dependent transient decrease in peak CPP (Fig 3), maximally 50±1% at 1 µmol. After the CPP response, FeSO₄ (0.3 µmol/L for 15 minutes) was infused to provide additional iron required for formation of NO traps [Fe(DETC)₂ complex]. FeSO₄ slightly increased CPP (by 8.5±6%). Subsequent infusion of DETC (50 µmol/L) did not alter CPP (0.5±1.3% change). After 10 minutes of DETC infusion, the decrease in CPP to a second GTN bolus was significantly attenuated compared with the decrease elicited by GTN before DETC infusion: peak responses to all doses of GTN were reduced by 5±1%, and the area under the curve (CPP decrease integrated from 0 to 5 minutes of GTN application) was reduced by 74±5%, 42±6%, and 16±4% at 0.1, 0.25, and 1 µmol GTN, respectively. As shown in Fig 3, tissue NO levels achieved during this second GTN exposure were positively correlated to the dose of GTN applied and to the GTN-elicited peak decrease in CPP. Interestingly, the NO level generated by 1 µmol of GTN in isolated hearts after 5 minutes (0.04 nmol/g) was about 10-fold lower than that detected in the same tissue in vivo after infusion of 10 µmol (2 mg per animal) of GTN during 20 minutes (0.45 nmol/g; Fig 2). This indicates a rather similar dose-effect relation in rabbit hearts in vitro and in vivo.

View larger version (12K): [in this window] [in a new window]   Figure 3. Graph showing correlation between the GTN-elicited decrease in CPP and ventricular NO levels in the isolated rabbit heart. The decrease in CPP was assessed in rabbit hearts isolated according to the Langendorff technique and continuously perfused by modified Krebs-Henseleit solution (composition as listed in "Methods"). The peak decrease in CPP (%) elicited by different doses of GTN was compared with the amount of NO (pmol/g tissue) trapped in the ventricular tissue by the Fe(DETC)2 complex. For details see text. Mean±SEM from six rabbits.

Cytochrome P-450–Catalyzed NO Formation From GTN
The high in vivo NO formation in the cytochrome P-450–rich organs liver, lung, and kidney prompted us to assess the ability of cytochrome P-450, purified from rabbit liver, to generate NO from GTN in vitro. Two different detection techniques were used that operate at reduced and at ambient oxygen concentrations: (1) NO spin trapping in aqueous solution by Fe(PDTC)₂, which unavoidably proceeds under nearly anaerobic conditions due to oxygen consumption by DTT (10 mmol/L; see "Methods"), and (2) activation of soluble GC, which is assessed at ambient oxygen levels in aqueous solution. Both techniques yielded consistent results.

ESR spectroscopy revealed that cytochrome P-450 transiently enhanced NO formation from GTN (Table 2), 26 times the control (P-450–free) value at 10 minutes and 4.1 times control at 60 minutes of incubation, in the presence of NADPH and cytochrome P-450–NADPH reductase (Table 2, line 1). Without the reductase, NO formation by cytochrome P-450 was attenuated but still significantly higher than controls (compare lines 2 and 4, Table 2). The reductase itself was not capable of generating NO from GTN (Table 2, line 3). In contrast to the transient nature of P-450–catalyzed NO formation, the low spontaneous NO formation (Table 2, line 4) resulting from reductive hydrolysis of GTN by DTT proceeded at a constant rate for up to 1 hour. The involvement of nitrite as a source of P-450–dependent NO formation was excluded, since NO formation by sodium nitrite (30 µmol/L) equaled spontaneous NO formation by GTN (30 µmol/L) (Table 2, line 5).

View this table: [in this window] [in a new window]   Table 2. NO Formation From GTN by Cytochrome P-450/Cytochrome P450–NADPH Reductase as Assessed by ESR Spectroscopy

Cytochrome P-450 also catalyzed NO formation from GTN in the presence of oxygen, as detected by activation of soluble GC (Fig 4). The enhancement of basal GC activity (45±5 nmol · mg^-1 · min^-1) increased with the concentration of GTN (Fig 4A) and cytochrome P-450 (Fig 4C), resulting in fourfold to sixfold higher GC activity at maximally effective concentrations of GTN and cytochrome P-450. The reductase was omitted in these experiments because it directly inhibited soluble GC activity in an NADPH-dependent fashion (data not shown). Cytochrome P-450–catalyzed activation of soluble GC by GTN was transient, peaking at 3 to 10 minutes of incubation (Fig 4B), thus confirming the previous findings obtained under anaerobic conditions. In the absence of cytochrome P-450, basal GC activity was slightly enhanced (0.75±0.15-fold) by higher concentrations of GTN (100 and 300 µmol/L) (Fig 4) due to a low level of NO generated by nonenzymatic release of nitrite (assessed by the Griess reaction²¹ ).

	Discussion

Top Abstract Introduction Methods Results Discussion References

 
GTN, like other organic nitrates, is believed to act as a vasodilator by releasing NO from one of its nitroester groups.^{1 2 3 4 5} However, this hypothesis still requires confirmation, since (1) no firm correlation has been established between tissue levels of NO and a pharmacological response to GTN in vascular and nonvascular tissues; (2) direct evidence for NO formation from GTN in vascular tissues in vivo is lacking; (3) the pharmacodynamics (pattern of NO formation in different tissues of the intact organism) of GTN are unknown, thus leaving the large variation in the sensitivity of different vascular beds to GTN unexplained; and (4) the molecular mechanism of biotransformation of GTN to NO in vivo is still unresolved. In an attempt to substantiate the "GTN-NO hypothesis," we now used an NO spin-trapping technique to monitor NO formation in isolated rabbit hearts and in different tissues of anesthetized rabbits exposed to GTN. We observed that (1) the GTN-elicited dilation of the coronary resistance bed (decrease in CPP) of isolated rabbit hearts was positively correlated to ventricular levels of NO; (2) GTN generated different levels of NO in several vascular tissues and organs; (3) NO levels were higher in the vena cava and the mesenteric bed compared with the aorta and femoral artery; and (4) purified cytochrome P-450 of rabbit liver catalyzed NO formation from GTN in vitro under aerobic and under unaerobic conditions.

Taken together, these findings support the hypothesis that GTN elicits vascular relaxation in vivo by releasing NO. Given that the strong correlation between tissue NO levels and vasodilation in the coronary resistance bed (Fig 2) also holds true for other vascular tissues, the finding that higher levels of NO were detected in the vena cava and the mesenteric bed than in the aorta and femoral artery (Fig 2) provides an explanation for the preferential venodilator activity of GTN.^{13 14 15} Several factors are potentially able to influence NO levels in a given tissue: tissue-specific bioavailability of GTN within the organism, relative rates of "inactivation" (nitrite formation by vascular GSH transferases²² ) and "activation" (NO formation) of GTN, and inactivation of NO (by superoxide anion radicals and hemoproteins). Limited bioavailability of GTN may be responsible for the low NO levels observed in the aorta, since extraction of infused GTN across this vessel is extremely low.^{23 24} In contrast, uptake of GTN by the vena cava, the mesenteric bed, and the femoral artery is similarly high.^{23 24} Therefore, bioavailability differences do not exist between these vascular tissues. Furthermore, according to in vitro studies with isolated vessels, overall metabolism of GTN to nitrite and glyceryldinitrates (via GSH transferase and via NO formation, respectively) would appear to be similar in the vena cava and the aorta.²⁴ Since inhibition of nitrite formation from GTN by GSH transferase inhibitors does not affect the vasodilator activity of GTN,²² it can be assumed that GTN-inactivating pathways (GSH transferases) do not significantly affect the formation of NO from GTN in vascular tissues. Finally, recent evidence indicates that inactivation of NO by reaction with superoxide anion radicals^{4 25} must be considered as a general principle controlling vascular NO levels. For example, vasodilator responses to GTN are attenuated in atherosclerotic compared with healthy blood vessels,²⁶ because the former generated higher amounts of superoxide anion radical than the nonatherosclerotic controls. This impaired vasodilator responsiveness to GTN was abolished by addition of superoxide dismutase. However, if superoxide levels in nonatherosclerotic venous and arterial tissues were different (which is unknown) and of critical importance for control of tissue NO levels, all vasodilators acting by release of NO should exhibit a similar vessel type specificity. This is clearly not the case. Sodium nitroprusside, for instance, in some vascular beds exhibits a vasodilator profile quite opposite to GTN.¹⁵ Furthermore, we analyzed vascular NO formation by the spontaneously NO-releasing nitrovasodilator SIN-1 (0.5 mg · kg^-1 · 20 min^-1 IV) infused to anesthetized rabbits. SIN-1 elicited significantly higher NO levels in arterial tissues (aorta, 0.25±0.05 nmol/g; femoral artery, 0.45±0.05 nmol/g; n=4) than an identical dose of GTN (Fig 2). In contrast, SIN-1 and GTN generated similar NO levels in the vena cava (0.45±0.05 nmol/g). This finding, too, rules out that different superoxide levels or different rates of inactivation of NO account for discrepant NO levels elicited by GTN in vascular tissues. We conclude that the rate of biotransformation of GTN to NO is the limiting factor controlling NO levels in vascular tissues.

We can exclude any influence of the trapping capacity of a given tissue on the amount of NO detected. For instance, traps [Fe(DETC)₂] were approximately equally distributed in vascular tissues, ranging from 6 (abdominal aorta) to 13 (mesenteric bed) nmol/g, but the fractional occupation of traps by GTN-derived NO varied from 0.5% (thoracic aorta) to 4.5% (vena cava) (Table 1). Similarly, the content of traps in organs was unrelated to the amount of NO detected (compare spleen and kidney, for instance). Efficient trapping of NO by iron-DETC is indicated by previous findings showing that DETC significantly attenuated the vasodilator response of isolated blood vessels²⁷ and the decrease in systemic arterial blood pressure in response to NO donors.²⁸

A surprising finding was the high rate of NO formation in several organs, equal to (heart, spleen) or even exceeding (lung, liver, and kidney) venous NO formation. Considering the reportedly low vasodilator sensitivity of the coronary and hepatic resistance bed to GTN^{15 29} and the low content of vascular tissue in the heart, liver, and kidney, nonvascular cells are likely to account for NO formation from GTN in these organs. According to in vitro studies with isolated deoxymyoglobin (Reference 30 and A. Mülsch, unpublished results), at least in the heart this reaction could be catalyzed by deoxymyoglobin present in cardiomyocytes.³¹ Furthermore, since liver, lung, and kidney are rich in cytochrome P-450, and we observed efficient NO formation from GTN by purified cytochrome P-450 from rabbit liver (Fig 4; Table 2), it is highly probable that cytochrome P-450 is a major pathway of NO formation from GTN in these organs.³² In support of this conclusion, anaerobic denitration of GTN and formation of nitrite by cytochrome P-450 in rat liver microsomes accounts for 30% to 40% of the total denitration in rat liver.³³ Therefore, this pathway may decisively contribute to the GTN-elicited vasodilatation of resistance beds in organs.^{15 29}

Furthermore, since cytochrome P-450 has been proposed as a likely candidate to catalyze formation of NO by GTN in vascular tissues,^{5 9 34 35} as well as an as yet unidentified tetrameric 200-kD protein,³⁶ this topic also remains a matter of debate.³⁷ Cytochrome P-450–like activity and immunoreactivity are present in low amounts in the vascular wall, the distribution between endothelium and smooth muscle layer depending on the vessel type and species.^{38 39} The present study demonstrates that cytochrome P-450 catalyzes NO formation both in the absence (Table 2) and presence (Fig 4) of oxygen. Although purified cytochrome P-450–NADPH reductase enhanced this NO formation, cytochrome P-450 was active per se and required NADPH only as a cofactor. Formation of NO from the nitroester moiety of GTN implies a reduction of the nitrogen atom by three electrons. This could be accomplished by direct electron transfer from the cytochrome P-450–heme. Several other reductive reactions catalyzed by cytochrome P-450 have been described.^{40 41} Furthermore, a direct, reductase-independent substrate reduction by cytochrome P-450, as observed in the present study (Fig 4), is not without precedent. The fungal cytochrome P-450 55A1 has been shown to catalyze the reduction of NO in an NADH-dependent fashion, without requiring additional proteinaceous components or redox-active mediators,⁴² and another fungal cytochrome P-450 was able to generate NO from GTN.⁴³

For the first time, we provide direct evidence that GTN generates NO in vascular tissues in vivo and that NO levels and vasodilator responses elicited by GTN in isolated perfused rabbit hearts are closely correlated. Furthermore, we demonstrate that venous tissue generates higher levels of NO from GTN than arterial tissue. This finding accounts for the prevailing venodilator activity of GTN. Surprisingly, NO formation was equal or even higher not only in the liver but also in several other organs previously not considered to express the GTN-NO pathway. Since cytochrome P-450 is abundant in these organs and we observed efficient catalysis of NO formation from GTN by reconstituted rabbit liver cytochrome P-450–NADPH reductase, this pathway probably accounts for NO formation by GTN in these organs.

	Selected Abbreviations and Acronyms

 

CPP = coronary perfusion pressure DETC = diethyldithiocarbamate ESR = electron spin resonance g = gyromagnetic constant of the free electron or radical at resonance; g=(h · )/(ß · HR), where h is the Planck constant, is the microwave frequency at resonance, ß is the Bohr magneton, and HR is the magnetic field at resonance GC = guanylyl cyclase GSH = glutathione GTN = glyceryl trinitrate hfs = hyperfine splitting NO = nitric oxide PDTC = proline dithiocarbamate SIN-1 = 3-morpholino-sydnonimine

	Acknowledgments

 
This study was supported by the Deutsche Forschungsgemeinschaft (Bu 436/4-3, BA 408/13-2, and Mü 900/4-1) and by Pohl-Boskamp, Hohenlockstedt, Germany. We thank Dr Norbert Frank, German Cancer Research Center, Heidelberg, for synthesis of PDTC and Dr Ingrid Fleming for helpful suggestions.

Received January 10, 1995; revision received February 20, 1995; accepted April 17, 1995.

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