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J Appl Physiol 100: 1267-1277, 2006. First published December 8, 2005; doi:10.1152/japplphysiol.01059.2005
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Brisk production of nitric oxide and associated formation of S-nitrosothiols in early hemorrhage

James L. Atkins,1,* Billy W. Day,2 Michael T. Handrigan,1 Zhe Zhang,2 Motilal B. Pamnani,3 and Nikolai V. Gorbunov1,*

1Division of Military Casualty Research, Walter Reed Army Institute of Research, Silver Spring, Maryland; 2Department of Pharmaceutical Sciences, University of Pittsburgh, Pittsburgh, Pennsylvania; and 3Department of Biochemistry and Physiology, Uniformed Services University of the Health Sciences, Bethesda, Maryland

Submitted 30 August 2005 ; accepted in final form 5 December 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
The results of previous inhibitor studies suggest that there is some increase in nitric oxide (NO) production from constitutive NO synthase in early hemorrhage (H), but the magnitude of NO production early after H has not been previously assessed. It is generally believed that only modest production rates are possible from the constitutively expressed NO synthases. To study this, anesthetized male Sprague-Dawley rats were subjected to 90 min of isobaric (40 mmHg) H. During this period of time, the dynamics of accumulation of NO intermediates in the arterial blood was assessed using electron paramagnetic resonance spectroscopy, chemiluminescence, fluorescence imaging, and mass spectrometry. Electron paramagnetic resonance-detectable NO adducts were also measured with spin traps in blood plasma and red blood cells. H led to an increase in the concentration of hemoglobin-NO from 0.9 ± 0.2 to 4.8 ± 0.7 µM. This accumulation was attenuated by a nonselective inhibitor of NO synthase, NG-nitro-L-argininemethyl ester (L-NAME), but not by NG-nitro-D-argininemethyl ester (D-NAME) or 1400W. Administration of L-NAME (but not 1400W or D-NAME) during H produced a short-term increase in mean arterial pressure (~90%). In H, the level of N oxides in red blood cells increased sevenfold. S-nitrosylation of plasma proteins was revealed with "biotin switch" techniques. The results provide compelling evidence that there is brisk production of NO in early H. The results indicate that the initial compensatory response to H is more complicated than previously realized, and it involves an orchestrated balance between intense vasoconstrictor and vasodilatory components.

blood cells; shock; physiology

THE EARLY PHYSIOLOGICAL EVENTS following hemorrhage (H) involve a "compensatory" response that results in an increase in the total peripheral resistance (61) and a redistribution of blood flow. This well-described response serves to maintain sufficient blood pressure to ensure tissue perfusion and generates a salutary redistribution of blood flow, preserving flow to critical vascular beds such as the coronaries at the expense of others such as the splanchnic bed (54). Increased sympathetic tone and the release of vasoconstrictors have well-defined roles in the early compensatory response (45); however, the potential role for vasodilators has not been well delineated. Theoretically, the dynamic regulation of blood flow distribution is best accomplished by spatially differential release of both vasoconstrictors and vasodilators. Because of its pivotal role in control of tissue perfusion, nitric oxide (NO) is a prime candidate as the postulated vasodilator, but the data to date supporting increased NO production in early H are inferential at best.

Nonselective inhibition NO synthase (NOS) in early H increases tissue injury (20), indicating that at least a basal production rate of NO is required to preserve tissue perfusion during this early timeframe. The most compelling evidence of increased NO production comes from experiments demonstrating the early loss of responsiveness to the exogenous vasoconstrictors. The loss of responsiveness to norepinephrine and angiotensin II is NO dependent (12, 35, 46, 47, 59). Studies on animals resuscitated as early as 1 h after the onset of hypotension have shown loss of responsiveness in some vascular beds (35, 46) and in isolated vessels (59). The effect persists for 4 h after resuscitation, and it can be blocked by infusion of nonselective inhibitors of NOS given in vivo (35, 46) near the time of resuscitation or in vitro (59). These findings suggest that NO production is increased either during H or immediately after resuscitation. More direct evidence of increased NO production could be obtained by measuring the accumulation of relatively stable NO reaction products in plasma and red blood cells (RBCs) during H-induced hypotension.

It has been shown recently that vasoactive NO compounds exist in three distinct redox forms: nitroxyl anion (NO–), NO radical, and the nitrosonium cation (NO+) (23, 57). Although it is generally accepted that NO radical is produced through a 5-electron oxidation of L-arginine, it has recently been suggested that NOS carries out only a four-electron oxidation, producing NO– (13, 53). The final step of oxidation of NO– to NO radical is accomplished by superoxide dismutase or other Cu-containing proteins [e.g., ceruloplasmin (CP)] (37, 42, 53). This raises the possibility that the plasma CP may be involved in formation of NO or NO adducts (2, 18, 25, 42, 60).

Based on this information, we hypothesized that the "fingerprints" of increased NO production should include changes in the redox state of plasma CP in addition to increases in the concentrations of NO adducts and the appearance of S-nitrosylated proteins. In the present study, we measured NO production in early H induced in rats by examining the formation of NO adducts with hemoglobin (Hb) and iron-N-methyl-D-glucamine dithiocarbamate {(MGD)2[Fe2+]} and iron-diethyldithiocarbamate {(DETC)2[Fe2+]} complexes by electron paramagnetic resonance (EPR) spectroscopy. Since high production rates for NO should result in the formation of S-nitrosothiols and S-nitrosylated proteins, we also looked for evidence of these compounds in the RBCs and blood plasma.

The results demonstrate large increases in the concentration of HbNO and the appearance of all the expected fingerprints of increased NO production. The study provides compelling evidence of brisk production of NO early after H. The results also indicate that circulating RBCs may serve as "chariots" of NO-like products.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Animal treatment.   Animal handling and treatments were conducted in compliance with the Animal Welfare Act and other Federal statutes and regulations related to animals, and experiments involving animals adhere to principles stated in the Guide to the Care and Use of Laboratory Animals, National Research Council. The facilities are fully accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International.

Thirty male Sprague-Dawley rats (Charles River Laboratories, Wilmington, MA) weighing 280–320 g received a standard diet and water ad libitum. All animals were anesthetized with ketamine (60 mg/kg ip) (NLS Animal Health, Owings Mills, MD), and anesthesia was maintained by supplementary injection of ketamine as required. The right femoral artery and vein were catheterized (Renepulse, Braintree Scientific, Braintree, MA). At the end of surgery, 0.1 ml of 1,000 IU/ml heparin was injected into each cannula after placement. The venous catheter was used for experimental induction of H, as well as for the administration of drugs. The induction of H was conducted by using a computer feedback and control program written in LabVIEW (National Instruments, Austin, TX), as previously described (19). Briefly, the closed-loop, computer-assisted system controls a low-flow peristaltic pump (model P720, Instech, Plymouth Meeting, PA). Blood withdrawn via the venous cannula was pumped into a heparinized reservoir placed on a balance in direct response to feedback from the blood pressure transducer (BPA-400, Micro-Med, Louisville, KY) connected to the arterial catheter for the measurement of phasic arterial blood pressure, mean arterial pressure (MAP), and heart rate. This system allowed for continuous acquisition of hemodynamic and shed blood data, along with continuous direct feedback control of the peristaltic pump, giving a tightly controlled and reproducible H insult. The arterial catheter was also used for blood sampling. Cardiovascular parameters were allowed to stabilize for 30 min. At the end of the stabilization period, the baseline blood sample (time point = 0 min) was taken from each rat from the catheter placed in the femoral artery. Blood was then withdrawn via the venous catheter to achieve a fall in MAP to 40 mmHg over 15 min. Thereafter, MAP was maintained at 40 mmHg for a total period of 90 min by either withdrawal (during the compensation period) or reinjection of blood. In some experiments assessing the blood pressure response to NOS inhibition, the feedback control of blood pressure was stopped just before drug administration. A typical profile of the MAP level during a H experiment is shown in Fig. 1B (compared with the sham treatment in Fig. 1A). The average peak shed blood volume of all H animals was 23.41 ± 1.76 ml/kg (n = 10). Blood samples were collected at 30, 60, and 90 min after the onset of H (time point = 0 min).


Figure 1
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  		Fig. 1. Graphs illustrating a typical time course of mean arterial blood pressure (MAP) during sham (A) and hemorrhage (B) experiments.

 
Experimental design.   Rats were randomly assigned to H or sham groups (n = 5 per group). Blood samples were taken at 0, 30, 60, and 90 min after H (0.7 ml of blood was removed, and this volume was replaced with an equal volume of normal saline).

Statistical analysis.   Results are given as means ± SE. Differences were evaluated by using one-way ANOVA with Tukey's post hoc multiple comparisons test. Differences were considered significant at P < 0.05.

Drug sources and timing of administration.   Saline (1 ml/kg) was administered as a vehicle control to designated H and sham animals immediately at the completion of surgical preparation before the 30-min stabilization period. 1400W dihydrochloride (Alexis Biochemical, San Diego, CA, 10 mg/kg in 1 ml) was administered to designated H and sham animals as described above. (MGD)2[Fe2+] [Alexis Biochemical, San Diego, CA; 326 mg/kg of MGD and 34 mg/kg FeSO4 as recommended (31)] was administered in 1 ml at the 90-min point of the experiment, and blood samples were collected 5 min later. Drug was administered to both sham and H animals. L-NAME or D-NAME (Alexis Biochemical, San Diego, CA) (15 mg/kg in 1 ml) was administered at 70 min into H, and blood samples were collected at 100 min. (DETC)2[Fe2+] (5 mM) and 0.5 mM Fe2SO4 (Sigma-Aldrich, St. Louis, MO) were added to red cells in vitro that were taken from the animal at 90 min into H.

EPR detection of HEME-iron nitrosyl complexes in blood.   Low-temperature EPR spectroscopy of blood was conducted as described previously (16). The frozen samples were placed in an EPR nitrogen Dewar flask (Wilmad Glass, Buena, NJ), and low-temperature EPR spectra were recorded with X-band EMX EPR spectrometer (Bruker Instruments, Rheinstetten, Germany). The recorded EPR spectra were analyzed using the WINEPR program package (Bruker Instruments). CP has a singlet signal with g = 2.030, trough and HbNO complexes have a multiplet signal characterized by a three-line hyperfine structure with Az (14N) = 17.2 G (where Az is the z axis anistrophic hyperfine coupling constant) and a trough at g =1.98 (22) (Fig. 2. ). The EPR signals of CP were measured in plasma samples. The EPR signals of HbNO complexes were obtained by subtraction of EPR signals of plasma CP and free radicals from the recorded blood EPR spectra using recently described protocols (17, 27) (Fig. 2). These calculated signals were not different from the EPR signals recorded from separated fractions of RBCs from the same experiments (data not shown).


Figure 2
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  		Fig. 2. Representative low-temperature electron paramagnetic resonance (EPR) spectra of blood collected from a sham-treated animal and an animal subjected to hemorrhage. A: samples from sham-treated animal. B: samples from hemorrhaged animal. 1–4, Spectra of blood samples at 0 min, 30 min, 60 min, and 90 min time points, respectively, 5–8, spectra of plasma samples demonstrating the EPR signals of ceruloplasmin (g = 2.030, trough); 9–12, spectra of HbNO in red blood cells (RBCs) derived from blood spectra 1–4, respectively. Note that the inverted free signal (g =2.004) is a calibration signal derived from the Dewar flask. In B [hemorrhage (5–8)], note the progressive decrease in ceruloplasmin trough (g = 2.03). Also note (9–12) the progressive increase in trough of HbNO (g = 1.97) and the characteristic triplet of this compound (12).

 
The second integral values were used to calculate amounts of HbNO in the peripheral blood samples and standard samples of HEME-nitrosylated Hb in RBCs. The standards for quantitative calculations of HbNO concentration were prepared and monitored in anaerobic spectrophotometric cuvette using freshly prepared RBCs and NOC-7, a donor of NO. Concentration of HbNO in erythrocytes was calculated using {epsilon} = 13 mM–1 cm–1 (3).

EPR spin trapping of NO products with MGD-iron complexes.   For in vivo detection of RNS compounds (i.e., NO, NO–, and NO+) in blood plasma, the water-soluble (MGD)2[Fe2+] was used. EPR analysis of both HbNO and (MGD)2[Fe2+]-NO complexes in blood was conducted in liquid nitrogen. Analysis of MGD2[Fe2+]-NO complexes in blood plasma was conducted with both low-temperature and room-temperature EPR techniques. The iron-nitrosyl complexes (MGD)2[Fe2+]-NO and (DETC)2[Fe2+]-NO, respectively, are characterized by EPR features of g{perp} = 2.039 and g|| = 2.017, and NA = 12.5 G (1, 63) where g{perp} and g|| are the axial symmetry components of the tensor of the gyromagnetic factor and NA is the nitrogen hyperfine coupling constant.

For ex vivo detection of RNS in RBCs, the lipid-soluble NO-trapping complex (DETC)2[Fe2+] was applied. RBCs were resuspended in an equal volume of saline, pH 7.4, incubated with 5 mM DETC and 0.5 mM Fe2+ for 5 min, frozen in liquid nitrogen, and then subjected to low-temperature EPR spectroscopy, as described above, detecting the characteristic signal of (DETC)2[Fe2+]-NO, with features g{perp} = 2.039 and g|| = 2.017.

Detection of S-nitrosylated proteins in RBCs.   RBCs diluted 1:10 in PBS were put onto slides using a Cytospin then washed with PBS and fixed for 30 min at 4°C in 4% paraformaldehyde in PBS, pH 7.4. The cells were again washed, then permeabilized with 0.2% Triton X-100 for 5 min. Free thiols in the cells were then rendered unreactive by methylthiolation with methylmethanothiosulfonate (200 mM for 30 min). After washing with PBS, the specimens were incubated in a solution of PBS containing 0.1 µM 2-[(5(6)-tetramethylrhodamine)amino]ethyl methanesulfonate and 1 mM ascorbic acid for 1 h. S-nitrosothiols, after being selectively reduced to thiols by ascorbic acid, were labeled with a rhodamine derivative of methanethiosulfonate. The labeled specimens were rinsed and mounted in Gelvatol (Monsanto, St. Louis, MO) under a coverslip for fluorescence microscopy. The specimens were analyzed with an Olympus AX 80 microscope equipped with a Hamamatsu digital camera. RBCs (50 cells/image) were randomly selected for assessment of fluorescence. Processing and analysis of digital images were conducted using SimplePCI high-performance imaging software (Compix, Cranberry, PA).

Biotin switch method for the detection of S-nitrosylated proteins in blood plasma.   The biotin switch method was adopted from the original protocol developed for the detection of S-nitrosylated proteins (26). Biotinylation of S-nitrosylated proteins was conducted as recommended. An aliquot of the purified biotinylated proteins was segregated by one-dimensional SDS-PAGE and analyzed by Western blot using an anti-biotin antibody. Another aliquot was examined by one-dimensional SDS-PAGE and SyproRuby staining and subsequently after destaining and Coomassie blue staining. Remaining samples were segregated by two-dimensional gel electrophoresis using immobilized pH gradient strips and PROTEAN II gels (Bio-Rad) and stained with SyproRuby. Gel bands/spots were visualized by visual and/or fluorescence image acquisition on a custom-built instrument with a high-resolution cooled Prometrix charge-coupled device camera (CH350 model, 16-bit chip, Photometrics, Munich, Germany) and analyzed with ImageJ software. Gel plugs were picked with with a robotic system on the imaging instrument, placed into Eppendorf tubes, washed to remove the dye, dehydrated with acetonitrile, lyophilized, and reswelled with a solution of sequencing grade trypsin (Promega, Madison, WI) in 25 mM ammonium carbonate.

Mass spectrometry.   Alterations in plasma proteins due to H were determined with mass spectrometry (MS) using previously developed protocol (4). Briefly, following trypsin digestion, the peptides derived from each protein gel spot were suspended in 1:1 water-acetonitrile containing 0.1% trifluoroacetic acid, mixed with a saturated solution of {alpha}-cyano-4-hydroxycinnamic acid in the same solvent, and 0.5-µl aliquots were spotted onto stainless steel plates. The spots on the plates were analyzed in duplicate by matrix-assisted laser desorption ionization-time of flight-MS in a 4700 Proteomics Analyzer with time of flight/time of flight optics (Applied Biosystems, Foster City, CA). Analysis of data with GPS Explorer Protein Analysis Software on a remote client workstation (Applied Biosystems) provided semiautomated acquisition of optimized mass spectra and the derivation of monoisotopic peptide mass fingerprint information. Searches of the National Center for Biotechnology Information nonredundant database, based on the peptide mass results using MASCOT (Matrix Science, Boston, MA) via the GPS Explorer workstation and MS-Fit (University of California San Francisco MS Facility) via the internet identified proteins with matching peptide mass fingerprints. The MASCOT parameters were set to include Rattus, tolerance of four missed trypsin cleavages per protein, allowance of protein modification by methionine oxidation, peptide tolerance of 30 parts/million (i.e., an example mass accuracy of 1,000 ± 0.03 Da), and restriction of peptides to the 700- to 4,200-Da range. The MS-Fit parameters included tolerance of four missed trypsin cleavage per protein, potential cysteine modification by acrylamide and oxidation of methionine, and changes at the amino termini, such as conversion of glutamine to pyro-glutamate and acetylation. Protein identifications were accepted when the observed and predicted pI and Mr were consistent, when scores indicated nonrandom identifications at a significance level of P < 0.05 and were corroborated by acquisition of MS/MS data that were consistent with the predicted b, y, and immonium ion series of the peptide.

Immunoblot analysis.   Alterations in the amounts of CP in blood plasma were assessed using immunoblot techniques followed by protein separation in polyacrylamide gels, as described previously (15). The primary antibodies used for hybridization were rabbit anti-CP polyclonal IgG (Innovex Biosciences, Richmond, CA) at 1:1,000 dilution. The washed membranes were subsequently probed with secondary antibody for 1 h at 25°C. The secondary antibody was goat anti-rabbit IgG (Upstate Biotechnology, Lake Placid, NY) used at a dilution of 1:3,000. The membranes were then washed three times in Tween-TBS for 5 min each, followed by development using an enhanced chemiluminescence detection kit (Amersham Biosciences, Piscataway, NJ) and subsequent exposure to Kodak Biomax MR-2 photographic film (Sigma-Aldrich, St. Louis, MO). Protein bands were identified by comparison with a molecular weight marker (Bio-Rad). beta-Actin was probed to monitor equal loading. Semi-quantitative assessment of immunoblots was conducted using ImageJ image processing software.

Chemiluminescence analysis of N-oxides.   Analysis of NO-derived products (N-oxides) in the RBCs and plasma was conducted with a NO analyzer NOA 280 (Sivers Instruments, Boulder, CO). The assay is based on detection of chemiluminescence response of ozone reaction with NO released from adducts present in RBCs and blood plasma and catalytically produced from NOFormula and NOFormula, the end products of NO oxidation. Analysis of the lable NO compounds, i.e., NO, NO–, and NO+ and NOFormula in RBCs was conducted at room temperature. The analyses of NOFormula and NOFormula in blood plasma were conducted at 25 and 90°C, respectively, following protein elimination on 5-kDa molecular mass cutoff filters within 1 h after blood sampling. The recorded chemiluminescence signals were analyzed using NO Analysis Software (Sivers Instruments).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Is there brisk production of NO in early H?   This study was undertaken to examine the production of NO in early H. We have extended previous studies that provided indirect evidence of increased NO production in early H (5, 35, 46, 47, 59) by demonstrating the accumulation of large amounts of HbNO in RBCs and the accumulation of multiple reaction products that are expected during such brisk production of NO. Additionally, we have confirmed the H-induced elevated production of NO through the use of NO spin traps.

Because NO binds avidly to hemoproteins, we were able to measure the systemic release of NO products by accumulation of HbNO in circulating RBCs. Increased HbNO was identified as early as 30 min into H (Fig. 3) (1.9 ± 0.2 vs. 1.2 ± 0.2 µM in sham treatment; P ≤ 0.017, n = 5), and a progressive increase in HbNO levels was seen throughout the entire observation period, reaching ~5 µM at 90 min (4.8 ± 0.7 vs. 1.4 ± 0.3 µM in sham; P ≤ 0.01, n = 5). Increased levels of HbNO have been detected previously by Kelly et al. (28) at 3 h into H. The characteristic EPR signal was consistent with a substantial level of HbNO at that time and a more intense signal was seen at 5 h of H. This suggests that the accumulation of HbNO continues into late H. Recently, Davies et al. (10) have examined a hypotension effect at 60 mmHg and shown a significant increase in HbNO at 3 h after H. In the present study, we have also demonstrated a sevenfold increase in N oxides within the RBCs as measured by chemiluminescence (19 ± 14 vs. 1.7 ± 0.8 µM; P ≤ 0.01, n = 5) (Fig. 4). EPR spin trapping was used to confirm both the high production rates in vivo and ex vivo and the presence of large amounts of reactive N species within the RBCs. The in vivo addition of (MGD)2[Fe2+] resulted in a brisk increase in arterial blood pressure from 40.2 ± 0.3 to 76.1 ± 5.5 mmHg (vs. 107.1 ± 4.4 mmHg in control period; n = 5, P < 0.001) and heart rate from 178.6 ± 7.0 to 351.6 ± 31 beats/min (vs. 302.5 ± 16 beats/min in control period; n = 5, P < 0.001) (Pearson correlation of the two parameters was r = 0.6; P < 0.03, n = 5), which was consistent with the systemic inactivation of vasoactive RNS entrapped by circulating (MGD)2[Fe2+].


Figure 3
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  		Fig. 3. Graphs illustrating dynamics of formation of heme-iron adducts with NO in blood of sham-treated animals (line 1) and animals subjected to hemorrhage (line 2). Blood samples were collected from each individual sham-treated and hemorrhaged rats at different time points (i.e., 0, 30, 60, and 90 min). The amount of HbNO was calculated based on EPR measurements as described in MATERIALS AND METHODS.

 

Figure 4
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  		Fig. 4. Alterations in chemiluminescence of RBCs and plasma associated with accumulation of NO products during hemmorhage. A: RBCs before hemmorhage. B: RBCs after 90 min of hemmorhage. C: plasma after hemmorhage. D: standard –25 µM sodium nitrite in PBS. RBCs were prepared as described in MATERIALS AND METHODS. After dilution with PBS (1:1), 3 µl of RBCs were introduced into the reaction vessel and incubated with the catalytic reagent at room temperature. The presented data are representative of 5 separate experiments.

 
The presence of these compounds was confirmed by a strong EPR signal of (MGD)2[Fe2+]-NO adducts in the H group but not the sham-treated animals (Fig. 5). This signal was not generated by extracting NO from HbNO in the RBC since the signal of HbNO did not change with the addition of (MGD)2[Fe2+]. The presence of large amounts of NO adducts within the RBCs was confirmed by separate experiments using the cell membrane permeable NO/trap-(DETC)2[Fe2+].


Figure 5
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  		Fig. 5. Representative low-temperature (A) and room-temperature (B) EPR spectra of spin trapping of NO-derived adducts in blood plasma from animals subjected to sham and hemorrhage treatments. 1, 90-min time point, sham treatment; 2, 90-min time point, hemorrhage; 3, standard solution of 10 mM (MGD)2[Fe2+] and 100 µM SNAP, an NO donor. Characteristic signals of (MGD)2[Fe2+] with features g{perp} = 2.039 and g|| = 2.017, and NA = 12.5 G (see MATERIALS AND METHODS for definitions) were detected in blood plasma after 90 min of hemorrhage. The presented diagrams are representative of 5 experiments.

 
Together, the data provide compelling evidence of brisk production of NO in early H. The data suggest that, in the face of intense release of vasoconstrictors previously demonstrated in early H, blood flow may be preserved at least in some critical beds by the increased production of NO. This raises the possibility that the redistribution of cardiac output and the relative hypoperfusion of some vascular beds results from regional differences in NO production. The data also demonstrate that RBCs carry substantial amounts of NO adducts. If this pool of NO adducts results in the release of vasodilators, then RBCs may serve as "chariots" for the "long-distance" delivery of NO-like vasodilatory products. One specific group of NO adducts, the S-nitrosothiols, have been proposed to serve this role.

Do S-nirosothiols increase in the RBCs?   We have demonstrated that the NO adducts in RBCs include S-nitrosothiols as detected by fluorescence image analysis (the RSNO-associated RBC fluorescence was 138 ± 56 AU in H vs. 28.7 ± 13.1 AU in sham; P ≤ 0.01, n = 150 cells, 3 animals) (Fig. 6). Rassaf et al. (49) have postulated that S-nitrosothiol production may be larger in the RBCs of rodents than in primates because of the higher thiol reactivity of the Cysbeta-125 residue of rat Hb, a residue not present in human Hb. However, even lower rates of S-nitrosothiol production may be important.


Figure 6
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  		Fig. 6. Fluorescence analysis of deposition of S-nitrosothiols in RBC proteins. Representative images of fluorescent adducts of S-nitrosothiols in RBCs obtained from sham-treated (A) and hemorrhaged (B) animals. Relative fluorescence intensity of A and B are shown in C and D, respectively. Fluorescence intensity values and RBC counts were estimated as described in the MATERIALS AND METHODS.

 
Stamler and colleagues (55, 56) have postulated that HbNO can donate NO to the Cysbeta-93 residue of Hb, resulting in the formation of S-nitrosothiols within the RBCs. These can subsequently exit the RBCs when the Hb is in the T state, resulting in the "targeted" delivery of S-nitrosothiols and vasodilation in tissues with low oxygen tension. In H, this mechanism could potentially serve as a "safety valve," ensuring that tissues did not become ischemic during the intense vasoconstriction by delivering vasodilators to the tissues with the lowest oxygen tension. Although the current data indicate that S-nitrosothiols increased in RBC, we have no direct evidence that the exit of these compounds from the RBC is dependent on the redox state of Hb, and this question is beyond the scope of the present study.

What are the mechanisms of protein S-nitrosylation in H?   In the presence of brisk NO production and the formation of S-nitrosothiols, it is expected that proteins with a favorable structure will be S-nitrosylated. This weak covalent bond has proven to be an important reversible posttranslational modification of some proteins and an important regulator of several signal transduction pathways (9). We have demonstrated S-nitrosylation of several plasma proteins by biotin switch two-dimensional gels with subsequent identification by mass spectrometry (Fig. 7). To date, not all of the S-nitrosylated proteins have been identified. The analysis continues, but one of the identified proteins deserves special mention, since previous work on transferrin suggested that all of its cysteines are involved in disulfide bonds (64). However, we have confirmed in vitro that transferrin can be S-nitrosylated, and we have confirmed its identification by Western blotting. Because of the finding of S-nitrosylated proteins in plasma, we examined postulated mechanisms for the formation of S-nitrosothiols within blood.


Figure 7
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  		Fig. 7. Biotin switch detection of S-nitrosylated proteins. Region of the 2-dimensional gel showing the presence of proteins trapped by the biotin switch. Those proteins identified conclusively by matrix-assisted laser desorption ionization-time of flight-mass spectrometry and mass spectrometry/mass spectrometry are indicated. Note that the keratins were also present in the S-nitrosylated form in the plasma from animals receiving sham treatment.

 
As previously described, HbNO can serve as a source for the formation of S-nitrosothiols in RBCs. Additionally, Inoue et al. (25) have shown in vitro that CP can catalyze the formation of S-nitrosothiols from NO, resulting in the "silencing" of the EPR signal of CP. Consistent with the occurrence of this reaction in vivo, in H we have demonstrated that the EPR signal of CP progressively decreased and was half of the original signal at 90 min (Fig. 2B). This occurred without any change in protein concentration of CP as measured by immunoblotting. The changes in CP inversely correlated with the increase in HbNO. This suggests that there are two mechanisms for the formation of S-nitrosothiols, one in the RBCs and one in plasma, providing potential mechanisms for the delivery of S-nitrosothiols both in the immediate vicinity of high NO production (CP mechanism) and also to distant sites (RBC mechanism).

How does an increase in NO production result in the loss of responsiveness to vasoconstrictors?   The loss of vascular response to epinephrine and angiotensin II is caused by increased NO production (12, 59). The previous finding of decreased responsiveness to these pressors early in H in some vascular beds (35, 46, 47) is consistent with the current finding of increased NO production in early H. The link between increased NO production and loss of responsiveness has not been established, but one possible explanation involves protein S-nitrosylation. The loss of responsiveness in H is thought to result from opening of the K+-ATP channels in vascular smooth muscle, and it is reversed by inhibitors of this channel (41, 68). Recently, Lin et al. (34) showed that S-nitrosylation of the RAS/mitogen-activated protein kinase in neural tissue results in the opening of these channels. However, many other factors have been implicated in the opening of the K+-ATP channels, including intercellular acidosis, and it is likely that the complexity of this response has only been partially revealed (33).

What is the source of NO in early hemorrhagic hypotension?   NO is normally produced by a family of NOS. NOS I and NOS III are constitutively expressed. Their NO production rates are normally low, and changes in production result from changes in intracellular calcium concentrations and posttranslational modifications of the NOSs. NOS II (iNOS) normally has low levels of expression. NO production from iNOS can be prodigious, and it is independent of changes in intracellular calcium. Although some posttranslational modifications have been described, large increases in NO production from iNOS normally result from the expression of new protein. The expression of new iNOS is a relatively late event in H. To our knowledge, the earliest demonstrated increase in mRNA for iNOS is at 45 min after H. The results came from our laboratory using the same model that was used in the present study (29). These data suggest that new iNOS expression occurs too late to contribute significantly to the increased NO production seen in the present study. We examined this question further by measuring the effect of NOS inhibitors on the production of HbNO.

The administration of a selective iNOS inhibitor has little cardiovascular effect in early H, and it can be administered without disruption of the normal hemodynamic responses to H. We administered a specific inhibitor of iNOS, 1400W, before H, and it did not change levels of HbNO in H animals (at 90 min 4.1 ± 0.3 µM in H animals and 4.8 ± 0.7 µM in H + 1400W; P < 0.6, n = 5). There was no effect of 1400W on HbNO levels in sham-treated animals either. The use of nonselective NOS inhibitors is more complicated, since they have significant cardiovascular effects. To minimize these consequences, we administered L-NAME at 70 min into H. Although there was the expected rise in MAP, the pressure was returned to 40 mmHg within 10 min (Fig. 8). The levels of HbNO fell by half in 30 min instead of the continued rise seen in animals administered D-NAME, an inactive stereoisomer of L-NAME, (Fig. 9). These data are consistent with the idea that the source of NO in early H is one of the constitutive NOSs.


Figure 8
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  		Fig. 8. Diagrams of the response of mean arterial blood pressure to administration of NO synthase inhibitors during hemorrhage experiments. Feedback control of blood pressure was stopped just before drug administration. Administrations of 10 mg/kg NG-nitro-L-argininemethyl ester (L-NAME) (A), 1 mg/kg 1400W (B), and 10 mg/kg NG-nitro-D-argininemethyl ester (D-NAME) (C), as recommended Garvey et al. (14), are indicated with arrows. *Increase in MAP following L-NAME administration. A–C are representative of 6 experiments.

 

Figure 9
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  		Fig. 9. Representative low-temperature EPR spectra of arterial blood demonstrating alterations in the intensities of signals of heme-iron nitrosyl complexes followed administration of L-NAME to animal subjected to hemorrhage. A: EPR signal of blood sample before hemorrhage. B: EPR signal of blood sample obtained at 70 min into hemorrhage. C: same as B at 30 min after administration of L-NAME. D: decrement of EPR signal of HbNO followed administration of L-NAME (calculated by subtraction of C from B). EPR components of HbNO in the presented spectra are indicated with arrows.

 

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Two mechanisms have been described for the production of NO from nitrite. Neither is likely to be the primary source of NO in this study, since plasma nitrite concentration is much smaller than the final concentrations of HbNO. However, they could potentially contribute to the response. Plasma nitrite levels are expected to increase as a result of large increases in NO production; however, there was no increase in nitrite levels in the present study. Indeed, NOFormula was <1.0 µM in both sham and H animals, whereas NOFormula increased from 19 ± 1 to 25 ± 1 µM in H animals (P ≤ 0.022, n = 5). This could be explained if one of the proposed mechanisms participated in the "recapture" of nitrite into NO. Under this paradigm, some NO metabolism proceeds to the formation of nitrate and nitrite in well-oxygenated tissues. Since this blood circulates into poorly oxygenated tissues, some of the nitrite would be converted back to NO. As a net result, nitrite levels would remain constant and nitrate levels would increase as seen in the present study. We cannot exclude the participation of one or both of these mechanisms of nitrite conversion.

Both mechanisms of nitrite conversion to NO function in tissues with decreased oxygen tension. Zweier et al. (70) have described the formation of NO from nitrite-mediated through xanthine oxidase. This mechanism requires ischemia and profound tissue acidosis, and it is normally thought to function in ischemia/reperfusion or cardiac arrest. We have administered a xanthine oxidase inhibitor and not seen any change in the production of HbNO (data not shown). The second mechanism, described by Cosby et al. (8) involves the conversion of NOFormula to NO through the mediation of deoxygenated Hb. Decreased Hb oxygenation has been documented in the mesenteric circulation in early H (44). Either mechanism could explain the lack of increase in nitrite in the present studies, but further studies are required to resolve this question.Our tentative conclusion is that a constitutive NOS is responsible for the large increase in NO production and that nitrite reclamation may amplify the effect (Fig. 10).


Figure 10
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  		Fig. 10. Schematic representation of model of NO production in early hemorrhagic shock, the effects of inhibitors and the likely mediators of vasodilation. Hemorrhage leads to a large increase in NO production, as evidenced by increased concentrations of NO adducts in RBCs to include HbNO, S-nitrosothiols, and other NO adducts. There are also increased plasma levels of nitrate and S-nitrosylated proteins (not shown). The effects of NO synthase (NOS) inhibitors are indicated. Dotted lines indicate speculated pathways.

 
Did the use of ketamine anesthesia and/or heparin affect the results of this study?   Ketamine has been shown to reduce NO synthesis in umbilical veins, so it is possible that even higher rates of NO synthesis would be found after H in a conscious animal (7, 67). It is difficult to predict how any form of anesthesia may indirectly affect systemic response to H through changes in the neurohumoral response (38, 39).

Heparin may indirectly increase NO production by preventing endothelial dysfunction(65). Savoye et al. (51) have recently demonstrated some endothelial dysfunction during H alone, especially in the mesenteric circulation. Heparin and heparin-like compounds prevent or reverse the endothelial dysfunction that is seen after resuscitation (48, 65, 66), and presumably they would have a similar effect during H.

What is the potential significance of increased NO production and the formation of NO adducts in early H?   Brisk production of NO during early H and the resultant vasodilation is normally beneficial and presumably ensures continued perfusion of at least some vascular beds during the intense early vasoconstrictor response to H. However, it could contribute to vascular collapse if there is an abrupt withdrawal of vasoconstrictor activity. This can occur very early in brisk H. In conscious animals, the initial compensatory response to H maintains normal blood pressure by increased sympathetic tone; however, with continued bleeding, H-induced sympathoinhibition results in the abrupt fall in blood pressure. Koch et al. (30) have shown that this fall in blood pressure also involves NO and can be slowed by NOS inhibition.

Other mechanisms may be involved in vasodilation in H. Numerous studies in the early 1980s demonstrated a large increase in interstitial potassium during H (11, 24, 36, 52, 62). Our laboratory recently confirmed these findings using modern microdialysis techniques that correct for in vivo diffusional constraints (43). Extracellular potassium in the range of 10–15 mM can induce vasodilation by opening the inward rectified potassium channel in vascular smooth muscle and by changes in sodium-potassium ATPase (6). Recent studies have also demonstrated increased production of carbon monoxide in H (32, 50, 69) and decreased levels of vasopressin in late H (40).

We speculate that NO adducts have an important role in early resuscitation from H. The brisk NO production demonstrated in the present study is expected to abate if resuscitation successfully restores normal blood pressure. However, the oxygen-dependent conversion of the excess of NO to reactive nitrogen species during resuscitation could potentially provide targeting of nitrosative damage and would predict, for instance, that proteins, which are S-nitrosylated during H, may be particularly susceptible to oxidative damage and nitration at the time of resuscitation.

In summary, we have shown that H leads to the rapid accumulation of large amounts of HbNO and other NO adducts including S-nitrosothiols within the RBCs. We have also demonstrated the S-nitrosylation of some plasma proteins early after H. These findings along with the results using NO spin traps provide compelling evidence of brisk NO production in early hemorrhagic shock. The results indicate that the initial compensatory response to H is more complicated than previously realized. It involves an orchestrated balance between intense vasoconstrictor and vasodilatory components. The redistribution of cardiac output in early hemorrhagic shock likely results from local regional differences in this balance. In the later phases of H, NO is already known to be an important mediator (21, 58), and the present results indicate that these processes may begin very early in the initial stages after H.


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 ABSTRACT
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This work was supported by the Department of the Army Peer Reviewed Medical Research Program no. PR033201.

The views, opinions, and/or findings contained herein are those of the authors and should not be construed as an official Department of the Army position, policy, or decision.


   ACKNOWLEDGMENTS
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We thank Jim Schooley, Matthew Bara, Dara Wolfe, Sara Smith, and Drs. Raghavan Balachandran and Ashraf Elamin for excellent technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. Atkins, Division of Military Casualty Research, Walter Reed Army Institute of Research, Bldg. 503, Rm. 1N80, 503 Robert Grant Ave., Silver Spring, MD 20910-7500 (e-mail: james.atkins{at}na.amedd.army.mil)

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.

* N. V. Gorbunov and J. L. Atkins contributed equally to the development of the presented model. Back


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