In septic shock, cardiovascular collapse is caused by the release of inflammatory mediators. We previously found that lysozyme (Lzm-S), released from leukocytes, contributed to the myocardial depression and arterial vasodilation that develop in canine models of septic shock. To cause vasodilation, Lzm-S generates hydrogen peroxide (H2O2) that activates the smooth muscle soluble guanylate cyclase (sGC) pathway, although the mechanism of H2O2 generation is not known. To cause myocardial depression, Lzm-S binds to the endocardial endothelium, resulting in the formation of nitric oxide (NO) and subsequent activation of myocardial sGC, although the initial signaling event is not clear. In this study, we examined whether the myocardial depression produced by Lzm-S was also caused by the generation of H2O2 and whether Lzm-S could intrinsically generate H2O2 as has been described for other protein types. In a canine ventricular trabecular preparation, we found that the peroxidizing agent Aspergillus niger catalase, that would breakdown H2O2, prevented Lzm-S- induced decrease in contraction. We also found that compound I, a species of catalase formed during H2O2 metabolism, could contribute to the NO generation caused by Lzm-S. In tissue-free experiments, we used a fluorometric assay (Ultra Amplex red H2O2 assay) and electrochemical sensor techniques, respectively, to measure H2O2 generation. We found that Lzm-S could generate H2O2 and, furthermore, that this generation could be attenuated by the singlet oxygen quencher sodium azide. This study shows that Lzm-S, a mediator of sepsis, is able to intrinsically generate H2O2. Moreover, this generation may activate H2O2-dependent pathways leading to cardiovascular collapse in septic shock.
- reactive oxygen species
- compound I
- septic shock
- myocardial depression
wentworth et al. (40) showed that antibodies, regardless of source or antigenic specificity, generate hydrogen peroxide (H2O2) from singlet oxygen (1O2•). They further showed that this process is catalytic and that antibodies use H2O as an electron source in reactions that eventually lead to the formation of H2O2 (40, 41). Whereas antibodies may produce up to 500 mole equivalents from 1O2• without a reduction in rate, Wentworth and colleagues (40, 41) also showed that other proteins, such as chick egg ovalbumin and β-lactoglobulin, were capable of producing H2O2, although only a few of these proteins were studied, and some of these nonimmunoglobulin proteins generated only small amounts of H2O2. These investigators proposed that the nonspecific production of H2O2 by antibodies in sepsis could have a beneficial effect of killing of microorganisms, despite a lack of antigen specificity.
Of the many nonimmunoglobulin proteins that are released in septic shock and that could possibly generate H2O2, we previously found that lysozyme (Lzm-S), released from leukocytes from the spleen and other organs, contributes to the myocardial depression and the arterial vasodilation that develop in Escherichia coli canine models (17, 26–31). When we determined the pathway of vasodilation in an in vitro preparation, we were surprised to find that the initiating signaling event was the generation of H2O2 by Lzm-S, although we did not define the mechanism by which this generation occurred (28). H2O2 has been found to be an important mediator that contributes to cardiovascular signaling under many conditions. Burke and Wolin (5, 6) described a unique pathway by which H2O2 caused vasodilation in a preconstricted pulmonary artery preparation. We showed that a similar pathway participated in the vasodilatory response of Lzm-S in recent experiments (28). In a phenylephrine-constricted carotid artery preparation, we demonstrated that the generation of H2O2 by Lzm-S led to the formation of compound I, a species of catalase that is the product of the metabolism of H2O2. Compound I then caused activation of smooth muscle soluble guanylate cyclase (sGC), which subsequently resulted in an increase in guanosine 3′,5′-cyclic monophosphate (cGMP) and thereby produced vasorelaxation. We further demonstrated that ethanol, an inhibitor of compound I, the peroxidizing agent Aspergillus niger catalase, a competitor of endogenous catalase, and inhibitors of sGC-cGMP-protein kinase G (PKG) pathway all prevented the vasodilator effect of Lzm-S (28). Moreover, we showed that the vasodilatory response of Lzm-S was not related to the presence of heavy metals [as performed by Buettner (4)] or to changes in pH or calcium in the organ bath and that denatured Lzm-S did not cause vasodilation in our carotid artery preparation (28).
Compared with the mechanism of vasodilation, our knowledge of how Lzm-S causes myocardial depression is more limited. There are distinct differences between the pathways that are involved in Lzm-S-induced vasodilation and Lzm-S-induced myocardial depression. Arterial vasodilation is both independent of the intact endothelium and unrelated to nitric oxide (NO) signaling, since removal of the endothelium and administration of the nitric oxide synthase (NOS) inhibitor NG-monomethyl-l-arginine (l-NMMA), respectively, did not inhibit the effect of Lzm-S in the organ bath preparation (28). On the other hand, to cause steady myocardial depression, Lzm-S needs to bind to a glycoprotein moiety on the endocardial endothelium, particularly to the mannose-β(1-4) N-acetylglucosamine (GlcNAc)-(β1-4)GlcNAc moiety of high mannose/hybrid and tri-antennary glycan subtypes (17). This binding then initiates the generation of NO that diffuses from the endothelium to the myocyte, resulting in an increase in cGMP, activation of PKG, and, hence, myocardial depression (29). We further showed that l-NMMA and removal of the endothelium inhibited the myocardial depressant activity of Lzm-S. In other experiments, moreover, we found that Lzm-S also inhibited the myocardial response to adrenergic field stimulation by a NO-dependent mechanism (26). We found that Lzm-S increased the release of NO from the parasympathetic nerves, thereby enhancing acetylcholine release and mitigating the adrenergic response. Such a depressant effect on the adrenergic response would therefore also contribute to the cardiovascular collapse that develops in septic shock.
The initial signaling event through which Lzm-S acts to cause a decline in steady-state contraction and an inhibition of the cardiac adrenergic response is not clear. On the basis of the mechanism of vasodilation caused by Lzm-S, we hypothesized in the present study that H2O2 signaling is also a necessary part of the pathway involved. We further hypothesized that the mechanism by which Lzm-S generates H2O2 could be analogous to that described for immunoglobulins as proposed by Wentworth and colleagues (33, 40, 41). Such findings would provide evidence that a mediator of sepsis may intrinsically generate H2O2, thereby activating pathways leading to the cardiovascular collapse that develops in septic shock.
These experiments were approved by the University Animal Care Committee and conform with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (32).
The Lzm-S used in these experiments was purified from the spleens of nonseptic dogs, as described in earlier studies (17, 26–31), by ARVYS Proteins (Stamford, CT). In the purification of lysozyme, the procedure outlined by Grobler et al. (14) was essentially used, in which Lzm-S was purified from the frozen canine spleen. After the spleens were thawed in a water bath, the tissue was homogenized in a Waring blender with 1 mM EDTA and 0.1 mM PMSF. The tissue debris was then removed by centrifugation. The extract was adjusted with 10× buffer to a final concentration of 20 mM HEPES, pH 8, and 1 mM EDTA, and precipitated material was removed by centrifugation. The sample was loaded on a carboxymethyl cellulose column, washed with the same buffer containing 30 mM NaCl, and eluted with the same buffer containing 200 mM NaCl. Fractions containing Lzm-S were concentrated to 2–3 mg/ml, and final purification was achieved on a HiPrep S100 26/60 column eluted with 20 mM HEPES and 0.5M ammonium acetate, pH 8.0. Elution from this column happens at a volume much higher than expected from the molecular weight of Lzm-S due to nonspecific binding to the resin. Final material was concentrated and dialyzed in deionized water (type II) that served as the vehicle for Lzm-S.
The turbidimetric method of Shugar (37) was used to confirm the presence of Lzm-S enzymatic activity in the preparation, in which the specific activity of the sample was 37,000 U/mg, as assayed using the Micrococcus lysis assay. As previously determined by SDS-PAGE, the sample is >96.5% pure (26, 28). When the gel was stained with Coomassie blue, the results showed a major band and a minor band. Both bands were analyzed using MS/MS mass spectroscopy, performed by W. M. Keck Foundation Biotechnology Resource Laboratory, Yale University (New Haven, CT). The purity of the preparation was determined by the finding that solely sequences of dog spleen Lzm-S were present.
In the present study, to further examine for purity, the Lzm-S sample was subjected to SDS gel and then staining by the more sensitive silver stain. All bands identified on the silver stain were subjected to MS/MS spectroscopy at the Manitoba Centre for Proteomics and Systems Biology. Furthermore, both intact protein and trypsin digests of Lzm-S were additionally analyzed by matrix-assisted laser desorption/ionization (MALDI) mass spectroscopy to confirm protein identity and purity of the sample. An ∼0.6 mg/ml solution of the protein in water was reduced (10 mM dithiothreitol, 30 min, 57°C), alkylated (50 mM iodoacetamide, 30 min in the dark at room temperature), dialyzed against 100 mM NH4HCO3 (6 h, 7-kDa molecular mass cutoff; Pierce Protein Research Products), and digested overnight with (sequencing grade) modified trypsin (1:50 enzyme-substrate weight ratio, 12 h, 37°C; Promega). One microliter of the sample (trypsin digest or intact protein) was mixed 1:1 with MALDI matrix (150 mg/ml 2,5-dihydroxybenzoic acid in 1:1 water-acetonitrile) deposited on the MALDI target and air-dried. Mass spectrometry analysis was performed using the Manitoba/Sciex prototype MALDI quadrupole time-of-flight (QqTOF) mass spectrometer. This instrument normally produces mass accuracy within 10 parts per million (ppm) in the TOF spectra (23).
As an additional assessment of purity, it was also determined whether the myocardial depressant effects of Lzm-S would be eliminated when the protein was denatured. The technique of denaturation was as described by Hancock and Hsu (15), in which an irreversibly denatured but soluble form of Lzm-S was prepared in which a solution of 3.5 mM lysozyme was made 20% (wt/wt) in polyethylene glycol (PEG) 1000 and kept in a water bath of 81°C for 16 h. Thereafter, the preparation was kept frozen until use. The concentrations of denatured lysozyme used were the same as those described for the native Lzm-S molecule in the various protocols.
Right Ventricular Trabecular In Vitro Preparation
The right ventricular trabecular (RVT) steady-state preparation and field stimulation preparation used in this study were previously described (17, 21, 26, 29–31). Briefly, mongrel dogs (15–25 kg) were anesthetized with pentobarbital sodium. The hearts were removed, flushed intraluminally with 50 ml of Krebs-Henseleit solution (KH; in mM: 118 NaCl, 4.7 KCl, 2.5 CaCl2, 1.2 MgSO4, 1.4 KH2PO4, 25 NaHCO3, and 11 dextrose), and placed in ice-cold KH bubbled with a mixture of 95%O2 and 5% CO2. Three to four thin trabeculae (<1-mm diameter; 3.5- to 4-mm length) were obtained from the right ventricle and tied at each end with 6-0 silk thread. Each thin muscle was suspended in a 5-ml vertical constant temperature bath that contained KH at 37°C.
During steady-state (SS) contraction, the muscle was stimulated electrically via punctate platinum bipolar electrodes with rectangular pulses (1-ms duration) at an intensity of 50% above threshold delivered at intervals of 2,000 ms. The trabeculae were stretched to optimal length. The reason that right, rather than left, ventricular trabeculae were used was that it is possible to obtain a greater number of thin muscle strips from the right ventricle, and these responses to Lzm-S were found to be the same between ventricles in a previous study (13). Isometric contraction measured in grams was converted into units of tension (mN/mm2) in which grams per cross-sectional area were multiplied by the acceleration due to gravity (9.80 cm/s2). Cross-sectional area was determined as muscle weight/[specific gravity (1.06 g/ml)/muscle length].
The plasma concentrations of Lzm-S previously attained in our in vivo sepsis model ranged between 10−9 and 10−6 mol/l (30), and the latter 10−6 mol/l concentration was used in the steady-state experiments described below. In the steady-state protocols, measurements of isometric tension were obtained at 10 and 20 min post-Lzm-S administration, during which the results were compared at similar intervals with those found in the control and treatment groups.
Experiments Determining Whether H2O2 Signaling Plays a Role in Mediating Lzm-S-Induced Decrease in SS Contraction
To inhibit the vasodilatory effect of Lzm-S, it was previously shown that the peroxidizing enzyme A. niger catalase, a competitor of endogenous catalase, was effective in this regard (28). In the present study, to determine whether a similar finding could be observed in SS contraction, the ventricular trabeculae were preincubated with A. niger catalase (10−7 or 10−8 mol/l) for 30 min, after which Lzm-S was added to the RVT preparation. These results were compared with a non-A. niger catalase-treated group. Furthermore, in the study of Burke and Wolin (5), it was shown that that the peroxide-metabolizing enzymes all had the particular capability of blocking the effect of H2O2 on reducing vascular tone. Since glutathione peroxidase would also be expected to have an effect similar to that of A. niger catalase on SS contraction, this consideration was also examined in the present study.
The central hypothesis of this study is that the generation of H2O2 by Lzm-S elicits the production of NO that subsequently leads to myocardial depression. Based on our previous vasodilation experiments (28), human Lzm-S generates between 10−6 and 10−7 mol/l H2O2 to produce vasodilation. In the present study, we determined whether 10−10, 10−8, and 5 × 10−7 mol/l concentrations of H2O2 would produce myocardial depression in the RVT preparation. Furthermore, we determined whether the myocardial depression that occurred with H2O2 administration could be blocked by pretreatment with l-NMMA (10−6 or 10−4 mol/l).
In the pathway of Lzm-S-induced vasodilation, we (28) have previously shown that the metabolism of H2O2 by endogenous catalase led to the formation of a species of catalase, termed compound I, that in turn led to the activation of sGC (see discussion). Inhibition of compound I has been demonstrated by other investigators (6) to be prevented by small molecules, such as ethanol. In our previous study (28), the vasodilatory effect of Lzm-S was inhibited by ethanol, in which the 10−7 mol/l concentration essentially blocked vasodilation. In the present study, this concentration of ethanol was used to determine whether it also would inhibit Lzm-S-induced myocardial depression. Although it was recognized that there would be some decrease in inotropy by ethanol per se, this approach would allow us to determine whether any additional effect of Lzm-S on causing myocardial depression could be prevented by this treatment. Thus positive results would support the notion that the formation of compound I is relevant to the pathway that causes the myocardial depressant effect of Lzm-S.
Another approach to examine the role of compound I in Lzm-S-induced myocardial depression would be to use pseudosubstrates for endogenous catalase (8, 9). The utility of alkyl hydroperoxides as pseudosubstrates for catalase to form compound I was first demonstrated by Chance and colleagues (8, 9). In the reaction of catalase with alkyl hydroperoxides, the catalytic intermediate compound I is formed, although these alkyl derivatives are difficult to work with. On the other hand, Jones and Middlemiss (18) showed that acyl hydroperoxides (peroxo acids) might provide suitable compound I pseudosubstrates and that it was possible to form compound I by the reaction of peroxoacetic acid and endogenous catalase. In the present study, this approach was followed to determine whether peroxoacetic acid would produce myocardial depression in the RVT preparation in a manner similar to that previously found for Lzm-S. Stock solutions of peroxoacetic acid (5 × 10−3 mol/l) were prepared by dilution with distilled water as described by Jones and Middlemiss (18). Since the hydrolysis of peroxoacetic acid results in the gradual accumulation of H2O2, the stock solution was spiked with minute concentrations of bovine catalase (2 × 10−9 mol/l) ∼30 min before use to prevent artifacts arising from the presence of H2O2. Based on preliminary experiments, the concentrations of peroxoacetic acid used in the RVT preparation were 1.25 × 10−4 and 7.5 × 10−5 mol/l, respectively. Furthermore, as stated, a central hypothesis of this study is that the pathway by which Lzm-S leads to myocardial depression involves the generation of H2O2 that in turn forms compound I by means of endogenous catalase metabolism. This would imply that compound I is necessary for generation of NO by the endocardial endothelium. To examine whether this is the case, we also determined whether l-NMMA (10−5 mol/l) prevents the myocardial depression caused by peroxoacetic acid in the RVT preparation.
Experiments to Determine Whether A. Niger Catalase Pretreatment Can Also Prevent Lzm-S-induced Cardiac Adrenergic Dysfunction
We also examined whether generation of H2O2 by Lzm-S was important in causing the depressed cardiac adrenergic response observed with Lzm-S treatment (26). The cardiac autonomic response to field stimulation (FSR) was determined in the RVT preparation (see above and Refs. 21, 26, 30). FSR represents the net effect between the sympathetic component, which would increase cardiac inotropy, and the parasympathetic component, which would mitigate any sympathetic response that occurs. In our previous study (26), Lzm-S was found to inhibit FSR by the nonendothelial formation of NO, presumably from the parasympathetic nerves that enhanced the release of acetylcholine from these nerves. In the present study, the objective was to determine whether H2O2 also mediated the decrease in FSR caused by Lzm-S such that pretreatment with A. niger catalase (10−7 mol/l) would also prevent the adrenergic effect of Lzm-S.
The cardiac autonomic response to field stimulation was determined in the RVT preparation as previously described (21, 26). The pulse width of the electrical stimulus trains was increased from 2 to 20 ms, keeping other stimulus parameters unchanged. After the isometric contraction had plateaued, the pulse width was reduced to 2 ms to restore control responses. The increase in isometric tension observed with field stimulation (FSR) was calculated as the percent increase from SS twitch amplitude as follows: FSR = (PR − SS)/SS × 100%, where PR is the peak isometric response. In a previous study (21), it was shown that propranolol blocked the response to field stimulation in this RVT preparation. In the adrenergic experiments, two baseline measurements of FSR were obtained at 10-min intervals to ensure that the preparation was stable. Lzm-S (10−6 mol/l) was then placed into the organ bath, after which FSR was obtained 10 min later. These results determined with Lzm-S alone were compared with results with control and treatment groups.
Real-Time Measurements of Endocardial Endothelial NO Formation Induced by Lzm-S
In previous experiments, we (29) showed that Lzm-S caused myocardial depression by binding to the endocardial endothelium that resulted in the formation of NO. In these latter experiments, NO release was identified using pharmacological and dye techniques. In the present study, to further verify this finding, we examined NO formation by Lzm-S in an endocardial endothelial preparation in which real-time measurements of NO release were assessed using electrochemical probes attached to a radical analyzer (NOPF-200 sensor, Apollo 400 Radical Analyzer; Precision Instruments, Sarasota, FL) (16, 19, 24, 44). In this endocardial preparation, a small vertical slit was made with a scalpel into the endocardial surface of the ventricle, after which the endocardial endothelial tissue plane was carefully dissected from that of the ventricular myocardium. A template was used to keep the samples relatively constant at ∼5 mm2, and the sample was placed into a 2-ml vial. Although the endocardial endothelial tissue is very thin, there is a very clear tissue plane between the endocardial endothelium and the ventricular muscle. The endocardial tissue is nearly transparent in character, whereas ventricular muscle is brown, so these two tissues can be easily separated visibly. In the previous study (29), we examined this endocardial endothelial sheet under microscopy and found it to be devoid of myocytes. In that latter study, moreover, the endocardial cell suspension was additionally characterized by immunofluorescence studies and was stained for the endothelial markers von Willebrand factor and endothelial NOS (eNOS). These markers colocalized in the endocardial cell suspension, and there was no evidence that other cell types were present (see Fig. 4 in Ref. 29).
The NO electrode has a sharp tip that was used to pierce the endothelial surface of the tissue such that its sensing element was positioned just at the level of the endothelium (44). The NO electrode and tissue were placed into a 2-ml vial that was filled to the ∼1-ml mark with Hanks' balanced salts solution (pH 7.4) to which Tris·HCl (10 mM), MgCl2 (1 mM), MgSO4 (0.8 mM), and NaHCO3 were added (24). The photosensitizing agent hematoporphyrin (10−6 mol/l) (40) was placed into the bath. Low-flow oxygen (<0.5 l/min) was bubbled through a small polyethylene catheter that was also placed into the vial. The vial that contained the probe was positioned into a larger water jacket container that was maintained at ∼35°C. As instructed by the manufacturer, the NO sensor was calibrated by placement of stock solutions of S-nitroso-N-acetyl-d,l-penicillamine stabilized in EDTA mixed in a cupric sulfate solution. Furthermore, the ability of the sensor to measure NO was confirmed by exposing the sensor to various concentrations of saturated NO solution. An AeroNOx (Tofield, AB, Canada) NO delivery and detection system was used. The NO gas (INO Therapeutics, Fort Allen, LA) was passed through a 10% KOH solution to remove other oxides of nitrogen. The gas was then passed through distilled water. Aliquots of the saturated solution (1.91 mM) were then placed into a mixing chamber to calibrate the sensor.
In this protocol, the objectives were to determine whether the probe could detect NO release caused by Lzm-S and to examine whether this NO release could be eliminated by inhibitors of the pathway attributable to Lzm-S-induced myocardial depression. Measurements of NO generation were first obtained after Lzm-S instillation in l-NMMA-treated and nontreated preparations. In addition, we determined that A. niger catalase (10−5 mol/l) and ethanol (10−7 mol/l) prevented NO release in the endocardial endothelial preparation, since A. niger catalase would metabolize any H2O2 produced by Lzm-S, whereas ethanol would prevent the formation of compound I. Finally, it was also confirmed that the electrode signal measured NO release by placing another agonist, bradykinin (10−5 mol/l), into the endocardial preparation (1). NO signals were determined with bradykinin alone, bradykinin with l-NMMA pretreatment (10−4 mol/l), l-NMMA pretreatment alone, and bradykinin with d-NMMA pretreatment (10−4 mol/l). Moreover, with all of the different electrochemical sensors used in this study (see below), measurements were obtained at approximately the 1,000-s interval after Lzm-S, placebo, or treatment was instilled (40), except for the bradykinin experiment where measurements were obtained at the signal peak.
Experiments to Determine Whether Lzm-S Intrinsically Generates H2O2
Electrode measurements of H2O2 production and oxygen consumption.
Before the consideration that Lzm-S could spontaneously generate H2O2 was examined, experiments were initially performed in which real-time measurements were made by means of the H2O2 sensor (ISO-HPO-100; WPI Instruments, Sarasota, FL) in the endocardial endothelial preparation. This preparation was identical to that described for the NO measurements in which a small piece of endocardial endothelium ∼5 mm2 obtained from the left ventricle was placed into the 2-ml vial. The sharp tip on the H2O2 electrode was again used to pierce the endothelial surface of the tissue. Measurements of H2O2 generation were then obtained after Lzm-S was instilled in A. niger catalase treated and non-catalase-treated endocardial endothelial preparations. The H2O2 electrochemical probe was calibrated in which stock solutions of H2O2 were placed into a mixing chamber.
On the basis of work by Wentworth et al. (40), it was subsequently considered that Lzm-S could be one of those proteins that could spontaneously induce production of H2O2. The H2O2 probe was then immersed into the 2-ml vial without any tissue, although the preparation was otherwise identical to that described above. Measurements of H2O2 generation were first obtained after Lzm-S instillation in A. niger catalase-treated and non-catalase-treated solutions. This would allow us to determine whether A. niger catalase treatment could inhibit the H2O2 signal caused by Lzm-S. After we demonstrated that this effect occurred, denatured lysozyme and human lactalbumin were used as negative controls. Wentworth et al. (40) showed that lactalbumin did not cause H2O2 generation in their preparation. Lactalbumin is the regulatory subunit in the heterodimeric enzyme lactose synthase of the milk gland (14). Both α-lactalbumin and Lzm-S belong to the glycosyl hydrolase 22 family, and in our previous study (28), there also was a lack of effect of lactalbumin on vasodilation demonstrated in our carotid artery preparation. The concentration of α-lactalbumin used in this study was the same as that used for Lzm-S (see results). In addition, the 1O2• quencher sodium azide (NaN3) was shown by Wentworth et al. (40) to prevent the generation of H2O2 otherwise observed in their preparation. This comparable experiment also was performed in the nontissue experiment in which it was assessed whether generation of H2O2 could be prevented by pretreatment with NaN3.
Finally, in a previous study, Kramarenko et al. (19) showed in a nontissue experiment that ascorbate reacts with 1O2• to produce H2O2. In their experiment, H2O2 and oxygen consumption were measured by electrochemical sensors. In the present study, we used a very similar technique to determine whether Lzm-S could cause a reaction with singlet 1O2• in a comparable manner. An oxygen sensor identical to that used by the latter investigators (18) (ISO-OXY-2; WPI) was placed into the 2-ml vial filled to the ∼1-ml mark. The vial was sealed airtight to prevent leaks. After Lzm-S administration, the change in oxygen concentration was determined by the sensor over a 1,000-s interval. Furthermore, to assess whether singlet 1O2• could be consumed, we also examined whether the 1O2• quencher NaN3 prevented the decline in oxygen consumption caused by Lzm-S. The oxygen sensor was calibrated by exposing the sensor to various concentrations of nitrogen and oxygen. Oxygen consumption (in ml) after Lzm-S or placebo (deionized distilled water) instillation was calculated from the solubility of oxygen at 35°C, the ideal gas equation, and the fractional decrease in oxygen as measured from the oxygen sensor.
Measurement of Lzm-S-induced H2O2 generation by Ultra Amplex red assay method.
To confirm the results obtained with the H2O2 electrode probe, we used another method to assess intrinsic generation of H2O2 by Lzm-S. Dikalov et al. (12) indicated that the Amplex red H2O2 assay (Molecular Probes, Eugene, OR) is a highly specific and sensitive fluorometric assay for detection of H2O2 with low limits of detection. Amplex red agent is a colorless substrate that reacts with H2O2 with a 1:1 stoichiometry to produce a highly fluorescent product. This method for detection of H2O2 was also used by Wentworth et al. (40). For the purpose of this study, since H2O2 is generated in a cell-free system, this seemed like an excellent approach. In these confirmatory experiments, the Ultra Amplex red assay (A36006; Molecular Probes) was used, which contains an even more sensitive dye for H2O2 detection than the original Amplex red assay.
In these experiments, the concentrations of Lzm-S used were 2 × 10−7, 3.6 × 10−6, 6.5 × 10−6 mol/l, and 2 × 10−5 mol/l, and the samples were placed into the 2-ml vial (as described for the electrode measurements), filled to the ∼1-ml mark with buffering solution and/or double-deionized water, and studied at 35°C and pH ∼7.35. After ∼20-min exposure to visible light, the solution was kept frozen at −20°C and was later studied for H2O2 generation with the use of fluorescent techniques when enough samples were obtained. In a similar manner, control experiments were performed in which buffer solution or double-deionized water alone was exposed to light for ∼20 min in respective experiments. Other experimental conditions included 2.7 × 10−5 mol/l Lzm-S treated with A. niger catalase (10−5 mol/l), 6.5 × 10−6 mol/l Lzm-S instillation after pretreatment with NaN3, and 6.5 × 10−6 mol/l Lzm-S instillations subsequent to pretreatment with mannitol (10−4 mol/l) and dimethyl sulfoxide (DMSO; 6 × 10−4 mol/l), respectively. The latter experiments were examined to determine whether these hydroxyl radical scavengers would prevent the generation of H2O2 by Lzm-S as suggested by Wentworth et al. (43).
Experiments were also performed in the dark to determine whether the H2O2 generation by Lzm-S would be inhibited in the absence of light. This was the case in experiments performed by Wentworth et al. (40), in which the lack of visible light prevented the production of H2O2 by immunoglobulins. For both the electrode probe and the Ultra Amplex red assay methods, experiments were performed in the dark without tissue. The objective was to determine whether the generation of H2O2 could be prevented compared with when visible light was present.
Western Blotting to Detect Phosphorylation of eNOS by Lzm-S
To further support the notion that the generation of H2O2 by Lzm-S elicits the production of NO, we also determined whether Lzm-S could lead to the phosphorylation of endocardial eNOS and, furthermore, whether this phosphorylation could be prevented by pretreatment with A. niger catalase. In this preparation, endocardial endothelial tissue was removed from the right and left ventricles as described for the endocardial endothelial preparation (see above; Ref. 29). The tissue was pooled and divided into three portions. Each portion was placed into a water bath filled with KH that was maintained at 37°C. To one portion, the Lzm-S vehicle was added; to the second portion, Lzm-S (10−6 mol/l) was added; and to the third portion, after pretreatment with A. niger catalase (10−5 mol/l) for 30 min, Lzm-S (10−6 mol/l) was added. All portions were then incubated for 30 min in their respective organ baths and studied for phosphorylated eNOS using the Western blotting technique.
The Western blot preparation was similar to that described by Thomas et al. (38). The tissue was washed in phosphate saline buffered solution and incubated in ice-cold lysis buffer for 10–20 min containing 1.5% Nonidet P-40, 20 mM MOPS [3-(N-morpholino)propanesulfonic acid, 4-morpholinepropanesulfonic acid, pH 7.0], 50 mM MgCl2, 10% glycerol, 300 mM NaCl, 1 μg/ml leupeptin, and 1 mM PMSF, followed by brief homogenization. Tissue lysates were centrifuged at 12,000 rpm for 30 min at 4°C, and the supernatant was subjected to SDS-PAGE (8.0%) loaded with 100 μg of protein. Resolved proteins were transferred at 4°C to a nitrocellulose membrane, and the membrane was blocked with 4% nonfat dry milk in TBS-Tween buffer (20 mM Tris·HCL, pH 7.4, 135 mM NaCl, 0.1% Tween) for 30 min. Blocked membranes were incubated with primary antibody directed against phospho-eNOS (Ser1177) (1:1,000 dilution; Cell Signaling Technology, Beverly, MA) or eNOS (1:1,000; BD Biosciences) in 4% nonfat dry milk in TBS-Tween buffer at 4°C for 16 h. Membranes were washed with TBS-Tween and incubated at room temperature for 1 h with horseradish peroxidase-conjugated secondary antibody (1:7,500 dilution) in TBS-Tween containing 4% nonfat dry milk. Membranes were washed in TBS-Tween, and proteins were detected with an enhanced chemiluminescence detection kit.
Additional Experiments to Determine Whether Other Known Inhibitors of H2O2 Generation Can Prevent Lzm-S-Induced Myocardial Depression
Finally, in parallel with experiments that were performed in the RVT preparation, we also assessed whether other inhibitors of H2O2 generation could prevent Lzm-S-induced myocardial depression. We first examined whether generation of H2O2 by Lzm-S could be prevented by diethyldithiocarbamic acid (DETCA), which inhibits Zn2+,Cu2+ superoxide dismutase (SOD) (10), since superoxide generation could be important in the generation of H2O2. Moreover, because the NADPH oxidase (Nox4) has been identified as a source of H2O2 that can be inhibited by diphenylene iodonium (10−5 mol/l), this treatment was also tried (20, 25, 39). Other treatments previously administered in our vasodilation study were not tested, since they failed to prevent Lzm-S-induced vasorelaxation (28). Among others, these included lipoxygenase and cyclooxygenase pathway inhibitors (7) and the Nox3 inhibitor apocynin that may inhibit superoxide generation (39).
Differences in variables among groups were determined using two-way (between-within) and one-way analysis of variance (ANOVA). The Student-Newman-Keuls multiple comparison test was included to determine statistical differences among treatment groups when the ANOVA was used. In the design of the experiment, of the three to five ventricular trabeculae or endothelial endocardial strips that could be obtained from the ventricle, each one was used for a different subset of experiments in a specific study. Results are means (SD).
The purity of the Lzm-S preparation was further examined in this study. In the preparation of the SDS gel, Lzm-S was loaded under different denaturing conditions, after which a sensitive silver stain was applied (Fig. 1). As shown, there was a major band identified at ∼15 kDa and a group of minor bands of slightly less molecular mass that represented fragments of the Lzm-S molecule. This is similar to what was found in the Coomassie stain, but only one minor band was found in the latter study. In the present and latter studies, both the major and minor bands were subjected to MS/MS spectroscopy. No canine protein was identified in any of the bands studied other than Lzm-S. Furthermore, the trypsin digest and intact Lzm-S preparation were subjected to MALDI spectroscopy. Peptide mass fingerprint analysis of the trypsin digest showed 98% sequence coverage (1 missed cleavage, alkylation with iodoacetamide, 10 ppm mass accuracy) for lysozyme C P81709 entry in the Swiss-Prot database. The only fragment missing was T11-13 (TLK) due to the extremely low molecular mass of this peptide. For intact protein, an average mass of 14,572.5 (MH+) was found (Fig. 1B). Calculated average mass (MH+) for intact P81709 protein is 14,571.5. Deviation of ∼1 Da is possible due to the presence of deamidation sites in the sequence of the protein and the lower accuracy of mass assignment of average mass. Altogether, MALDI spectral analysis allowed us to confirm unequivocally 100% identity of the Lzm-S sample to the P81709 entry in the Swiss-Prot database. None of the spectra (digest or intact protein) showed the presence of essential protein contamination.
The Pathway of Lzm-S-Induced Formation of NO
As shown in Fig. 2, the NO probe was capable of detecting the NO signal when exposed to aqueous standards prepared with NO gas. An example of the effect of Lzm-S (1 × 10−6 mol/l) on the generation of NO obtained with and without l-NMMA pretreatment (10−5 mol/l) in the endocardial endothelial preparation is shown in Fig. 3A. Over the measurement interval, there was a steady increase in the formation of NO that occurred a few minutes after Lzm-S was added to the preparation. NO formation did not immediately increase, since Lzm-S needs to generate H2O2 (see below; also see discussion). As shown in Fig. 3A, l-NMMA prevented the increase in NO formation compared with the non-l-NMMA treated preparation. The mean findings are displayed in Fig. 3B. Additional experiments were subsequently performed to show that the NO electrodes were sensitive to a known agonist of NO formation. In these experiments, the results showed 1) that bradykinin increased NO formation from the approximately zero baseline value to 5.3 ± 1.6 × 10−6 mol/l (n = 6; P < 05 vs. l-NMMA and bradykinin and l-NMMA without bradykinin), 2) that pretreatment with l-NMMA attenuated this increase to 0.12 ± 0.7 × 10−6 mol/l (n = 5), 3) that d-NMMA had no effect on NO formation by bradykinin [NO increased over baseline to 6.7 ± 5 × 10−6 mol/l (n = 5)], and 4) that l-NMMA by itself caused no change in NO formation (−1 ± 3 × 10−7 mol/l from baseline; n = 6).
The postulated mechanism by which Lzm-S initiates the formation of NO was considered to be the production of H2O2 and its subsequent metabolism by endogenous catalase. In Fig. 4A, an example is shown in which pretreatment of the RVT preparation with the peroxidizing agent A. niger catalase, a competitor of endogenous catalase, prevented the decrease in tension otherwise observed with Lzm-S alone. The mean results illustrating this effect are shown in Fig. 4B. The 10−8 mol/l concentration of A. niger catalase seemed to have a lesser inhibitory effect than the 10−7 mol/l concentration. At a similar concentration, glutathione peroxidase also had a blocking effect on Lzm-S-induced myocardial depression, and contraction decreased significantly less compared with Lzm-S alone. With glutathione peroxidase treatment, myocardial tension was 7.2 ± 1.4 × 10−7 mN/mm2 at baseline and measured 5.8 ± 1.9 mN/mm2 at 10 min and 4.8 ± 2.4 mN/mm2 at 20 min post-Lzm-S instillation (both P < 0.05 vs. baseline and P < 0.05 vs. Lzm-S alone). Furthermore, when denatured Lzm-S (1 × 10−6 mol/l) was added to the RVT preparation, there was no difference in the reduction in tensions compared with when the PEG vehicle was used alone. In the denatured Lzm-S group (n = 6), the mean tension was 7.4 ± 0.6 mN/mm2 at baseline, 7.1 ± 1.3 mN/mm2 at 10 min postinstillation, and 5.8 ± 1.6 mN/mm2 (P < 0.05 vs. baseline; P < 0.05 vs. native Lzm-S group) at 20 min postinstillation. In the PEG alone-treated group (n = 5), the mean tension was 7.8 ± 1.0 mN/mm2 at baseline, 7.5 ± 2 mN/mm2 (P < 0.05 vs. native Lzm-S group) at 10 min postinstillation, and 5.8 ± 2.4 mN/mm2 (P < 0.05 vs. native Lzm-S group) at 20 min postinstillation.
Experiments were also performed to determine whether H2O2 per se would produce a decline in myocardial tension similar to that caused by Lzm-S. In these experiments, at 10−10 mol/l H2O2, there was no decrease in contraction, whereas at 10−8 mol/l, there was a slight decrease in tension. At a concentration of 5 × 10−7 mol/l H2O2 that was based on a previous study (28), the results show that H2O2 produced a decline in myocardial tension comparable to that found with Lzm-S; moreover, the results also demonstrate that this decline could be inhibited by pretreatment of the RVT preparation with l-NMMA (Fig. 5, A and B).
It was also postulated that during the metabolism of H2O2 by endogenous catalase, there would be the generation of a species of catalase, termed compound I, that can be inhibited by small molecules, such as ethanol (6). In Fig. 6, the results show that although ethanol per se (10−7 mol/l) caused a decrease in myocardial tension, there was no further decline when Lzm-S was added to the preparation. In addition, peroxoacetic acid was used as a pseudosubstrate of endogenous catalase to form compound I to determine whether it also produced myocardial depression. There was a dose-response effect of peroxoacetic acid on myocardial contraction. In preliminary studies, 7.5 × 10−5 mol/l peroxoacetic acid had no effect on myocardial tension, but at a higher concentration of 1.25 × 10−4 mol/l, this substance produced a decline in tension comparable to that observed with Lzm-S. In Fig. 7, A and B, it is shown that pretreatment of the RVT preparation with l-NMMA prevented the decrease in myocardial depression caused by peroxoacetic acid. Furthermore, in Fig. 3B, it also is shown that both ethanol and A. niger catalase prevented the increase in NO measured by the electrode in the endocardial endothelial preparation, with these findings again demonstrating that the H2O2 and NO generated by Lzm-S are linked to one another by means of a common pathway (see discussion).
Western blot experiments were performed to determine whether Lzm-S could lead to formation of NO by phosphorylation of eNOS in endothelial endocardial tissue. These results are delineated in Fig. 8 and show that A. niger catalase blocked the phosphorylation of eNOS caused by Lzm-S.
None of the traditional inhibitors of H2O2 production (7, 10, 25, 39), such as DETCA (10−2 mol/l) and diphenylene iodonium (10−5 mol/l), prevented the decline in isometric tension in the RVT preparation, so only a few experiments were performed. After DETCA pretreatment (n = 3), the isometric tension decreased from 5 ± 1.4 mN/mm2 at baseline to 2.9 ± 1.4 and 1.5 ± 1.1 mN/mm2 at 10 and 20 min post-Lzm-S, respectively. After diphenylene iodonium pretreatment (n = 2), the isometric tension decreased from 5.4 ± .4 mN/mm2 at baseline to 3.8 ± 1.2 and 3.1 ± 0.5 mN/mm2 at 10 and 20 min post-Lzm-S, respectively. These decreases in tension observed after treatment were not different from those in concomitant nontreatment groups, so only a few experiments were performed in each group.
A. Niger Catalase Also Prevents the Decline in Cardiac Adrenergic Response Caused by Lzm-S
The effect of A. niger catalase (10−7 mol/l) on prevention of Lzm-S (1 × 10−6 mol/l)-induced decrease in the adrenergic response in the RVT preparation was also examined in this study. Examples of FSR under the different treatment conditions are shown in Fig. 9. In this protocol, two baseline stimulations of FSR were obtained, after which Lzm-S or vehicle was added to the preparation. Two to three beats of SS contraction are shown at the beginning of the tracing. Field stimulation was initiated at the interval indicated by the arrow. The small dip in isometric tension observed at the beginning of stimulation is believed to be due to the effect of abnormal synchronization as described by Blinks (3). This effect may be due to abnormal conduction of the action potential when field stimulation is initially applied, possibly leading to an even more negative tension compared with SS contraction in some instances. At baseline, field stimulation produced an ∼25% increase in tension (distance between dashed line and peak force development). Compared with baseline measurements, Lzm-S at 10−6 mol/l eliminated the increase in inotropy observed with field stimulation, which was significantly different from that found with A. niger catalase pretreatment. The mean results are shown in Table 1.
Moreover, as previously described (see Refs. 21 and 26 for details), there was a progressive decline in SS contraction after FSR that occurred to a varying extent in both the vehicle-treated and Lzm-S-treated groups. The mechanism of this decline is multifactorial but most likely related to the negative inotropic effect of excess catecholamine release that impairs the contractile apparatus when multiple neural stimulations are performed (36), since this decline in SS was much smaller in time control groups in which field stimulation was not performed (26). Moreover, it was previously shown that any decline in SS that occurs with repeated field stimulations does not affect the magnitude of FSR that remains relatively constant over time (21, 26). In the present study, the percentage declines in SS from baseline 1 to baseline 2 and from baseline 1 to the treatment condition were slightly less in the time control group compared with the other groups. In the KH pretreatment and Lzm-S group, the respective declines were 16 ± 22 and 51 ± 18%. In the pretreatment A. niger catalase and Lzm-S group, the respective declines were 22 ± 21 and 53 ± 19%. In the pretreatment A. niger catalase and vehicle group, the respective declines were 19 ± 30 and 54 ± 24%. In the KH pretreatment and vehicle group, the respective declines were 24 ± 28 and 33 ± 39% (P < 0.05 vs. other groups).
Lzm-S Intrinsically Generates H2O2
The electrochemical probe was calibrated with different concentrations of H2O2 (Fig. 10). In initial experiments, we assessed whether Lzm-S could generate H2O2 in the endocardial endothelial preparation. An example is illustrated in Fig. 11A in which it is shown that Lzm-S produced an amount of H2O2 comparable to that found in the carotid artery preparation as determined in a previous study (28). Moreover, pretreatment of the preparation with A. niger catalase (10−5 mol/l) inhibited H2O2 production in this endocardial preparation [this higher concentration of A. niger catalase was used compared with the RVT preparation measurements to ensure that mixing occurred and was based on the work of Wentworth et al. (40)]. Lzm-S vehicle (deionized distilled water) had no effect on H2O2 generation by itself. The mean results demonstrating these findings are shown in Fig. 11B.
Comparable experiments were then performed without the presence of endocardial endothelial tissue (Fig. 12). Without tissue, Lzm-S caused an increase in production of H2O2 similar to that found in the tissue preparation, although a higher concentration of Lzm-S (2 × 10−6 vs. 1 × 10−6 mol/l) was required to produce this effect in the tissue-free experiment (see discussion). A. niger catalase (10−5 mol/l) at a concentration identical to that used for the tissue experiment also could be shown to counteract 2 × 10−6 mol/l Lzm-S in the tissue-free experiment. A dose-response effect of Lzm-S on H2O2 production was also evident, since 2.7 × 10−5 mol/l Lzm-S caused an even larger amount of H2O2 production compared with the 2 × 10−6 mol/l concentration. Moreover, without tissue, it can be seen that the generation of H2O2 by 2 × 10−6 mol/l Lzm-S also could be inhibited by pretreatment with the singlet oxygen quencher NaN3 (1 × 10−3 mol/l; Ref. 40). On the other hand, another protein that was related to Lzm-S, lactalbumin (2 × 10−6 mol/l), as well as denatured Lzm-S, all failed to generate H2O2 in the tissue-free experiment. The mean changes observed for the different conditions in the tissue-free experiment are shown in Fig. 13.
To further determine whether oxygen consumption occurred when Lzm-S (2 × 10−6 mol/l) was added to the tissue-free experiment, we used the oxygen sensor technique as described by Kramarenko et al. (19). An example is illustrated in Fig. 14A. It can be seen that Lzm-S (2 × 10−6 mol/l) resulted in oxygen consumption and that oxygen consumption also could be blocked by the 1O2• quencher NaN3 (1 × 10−3 mol/l). The mean results of oxygen consumption under the different conditions are illustrated in Fig. 14B. The fractional declines in oxygen at the 1,000-s interval as determined by the oxygen sensor were 0.0020 ± 0.002 in the Lzm-S alone-treated group (P < 0.05 vs. other groups), 0.0001 ± 0.0001 in the Lzm-S group pretreated with NaN3, and 0.0000 ± 0.0001 in the placebo-treated group.
Confirmatory Measurements of Intrinsic Generation of H2O2 by Lzm-S
As determined by the Ultra Amplex red assay, fluorescent measurements of intrinsic generation of H2O2 by Lzm-S were made to confirm the electrode results (12). The different conditions are illustrated in Fig. 15 and demonstrate that at the two lowest concentrations of Lzm-S (i.e., 2 × 10−7 and 3.6 × 10−6 mol/l), there was no increase in H2O2 detected by this assay compared with the buffer alone-treated experiments. However, at concentrations of 6.5 × 10−6 and 2.7 × 10−5 mol/l, there were large increases in H2O2 production, and there was a dose-response effect. Without any Lzm-S, the amount of background H2O2 detected in the buffer solution was not different from that found in the double-deionized water sample.
In addition, the H2O2 generated by Lzm-S at a concentration of 2.7 × 10−5 mol/l could be eliminated by A. niger catalase, whereas there was no difference in the generation of H2O2 by Lzm-S (6.5 × 10−6 mol/l) between the buffered and nonbuffered conditions (see Fig. 15). Furthermore, we also examined whether the hydroxyl radical scavengers mannitol (10−4 mol/l) or DMSO (6 × 10−4 mol/l) would affect the amount of H2O2 generated by Lzm-S. In the latter instances, there was no difference in the amount of H2O2 generated by Lzm-S at 6.5 × 10−6 mol/l, which was quite consistent among the various conditions tested (see Fig. 15).
The work by Wentworth et al. (40) also showed that light, either visible or ultraviolet, was a necessary requirement for the generation of H2O2 by immunoglobulins. To determine whether this was the case in the present study, we compared the results obtained with both the H2O2 electrode and the Ultra Amplex red assay with and without light in respective experiments. These results determined without tissue are shown in Fig. 16. In both methodologies, the absence of light significantly reduced the amount of H2O2 generated by Lzm-S to an amount comparable to background values.
The present study showed that Lzm-S, a mediator of sepsis, is intrinsically able to generate H2O2 and, moreover, that this generation is capable of activating pathways that could contribute to the cardiovascular collapse found in septic shock (2, 34). Wentworth and colleagues (33, 40, 41) demonstrated that proteins, particularly immunoglobulins, irrespective of their antigen specificity, may generate H2O2 in a complex set of reactions from 1O2• and H2O. They proposed that H2O2 generated in such a manner could serve as a mechanism for bacterial killing in sepsis. Although these investigators primarily focused on immunoglobulins as a source of H2O2, they also found that other proteins such as chick egg ovalbumin and β-lactoglobulins were capable of inducing this effect (40). In the present study, we showed that Lzm-S, a mediator of sepsis that is released from leukocytes as part of the inflammatory response, also is capable of inducing production of H2O2. However, rather than serving as a mechanism for bacterial killing, our results indicate that the H2O2 generated by Lzm-S could activate pathways leading to the myocardial depression and arterial vasodilation that are characteristic of septic shock (2).
Lzm-S is a newly discovered mediator of sepsis, and the pathways by which it causes cardiovascular collapse are just beginning to be elucidated (17, 26–31). We previously showed that to produce myocardial depression, Lzm-S needs to bind to a glycoprotein on the endocardial endothelium (17, 29), particularly to the mannose-β(1-4)GlcNAc-(β1-4)GlcNAc moiety of high-mannose/hybrid and tri-antennary glycan subtypes (17). This binding results in the release of NO that diffuses to adjacent myocytes to cause myocardial depression by activation of the sGC-cGMP-PKG pathway (22, 29). We further demonstrated that the removal of the endocardial endothelium as well as the administration of competitive inhibitors of enzymatic action of Lzm-S, such as N,N′-diacetylchitobiose and N,N′,N″-triacetylchitotriose (35), attenuates the myocardial depressant effect caused by Lzm-S in both in vitro and in vivo preparations (17, 26, 27, 29–31).
In the present study, we extended our knowledge about the pathways responsible for Lzm-S-induced myocardial depression. We showed that the formation of NO by Lzm-S is dependent on H2O2 signaling. We further demonstrated that we could block Lzm-S-induced myocardial depression in the RVT preparation by pretreatment with the peroxidizing agent A. niger catalase. In addition, we found that H2O2 by itself could also produce myocardial depression and, furthermore, that this effect of H2O2 could be inhibited by l-NMMA pretreatment. We therefore conclude that H2O2 is responsible for the NO formation caused by Lzm-S. Others also have shown that H2O2 activates eNOS to form NO. Thomas et al. (38) indicated that H2O2 mediated this effect by phosphorylation of eNOS. Our results suggest that the H2O2 generated by Lzm-S also may lead to NO by phosphorylation of eNOS in endocardial endothelial tissue (see Fig. 8).
Other investigators also have shown that in the metabolism of H2O2 by endogenous catalase, there is the formation of a species of catalase, termed compound I (5, 6). Wolin and colleagues (5, 6, 10) showed that compound I could activate sGC, leading to vasodilation in a preconstricted pulmonary artery preparation. They further showed that this effect of compound I on vasodilation could be inhibited by the peroxide-metabolizing agent A. niger catalase, as well as by small molecules such as ethanol. In our previous vasodilation study (28), we demonstrated that both A. niger catalase and ethanol inhibited Lzm-S-induced vasorelaxation in a carotid artery preparation. In the present study, in addition to finding that A. niger catalase prevented Lzm-S-induced myocardial depression in the RVT preparation, we also demonstrated that ethanol prevented this effect. These results would suggest that compound I is a necessary component for Lzm-S-induced myocardial depression. Furthermore, additional evidence supporting the role of compound I in the myocardial depression caused by Lzm-S comes from the peroxoacetic acid experiments. Peroxoacetic acid served as a pseudosubstrate for endogenous catalase in the formation of compound I (18). These results not only showed that peroxoacetic acid caused myocardial depression in the RVT preparation but also demonstrated that l-NMMA pretreatment prevented the effect of peroxoacetic acid. Thus these overall results support the notion that the important components of the pathway leading to Lzm-S-induced myocardial depression involve the generation of H2O2 that leads to the formation of compound I and the subsequent activation of eNOS (see Fig. 17 for schematic of pathway). Furthermore, in the field stimulation experiments, we found that the effect of Lzm-S on the adrenergic response also could be prevented by A. niger catalase pretreatment. These results also support a role of H2O2 production as a universal mediator of the effects of Lzm-S in causing vasodilation, depressed steady-state myocardial contraction, and cardiac adrenergic dysfunction.
We believe the mechanism by which Lzm-S generates H2O2 can be best explained in terms of the concepts described by Wentworth and colleagues (33, 40, 41). Although these investigators focused on immunoglobulins as a means of H2O2 production, they additionally showed that other proteins had the ability to generate H2O2. We propose that Lzm-S fits into the category of one of those nonimmunoglobulin entities. In our initial experiments of real-time production of H2O2, we used an endocardial endothelium preparation in which the sensor was placed through the tissue. These results showed that Lzm-S at a concentration of 1 × 10−6 mol/l caused production of H2O2 to a value equal to that found in the carotid artery preparation (28). We also found that A. niger catalase attenuated this effect. In the nontissue experiments, however, there was a larger amount of Lzm-S required to cause H2O2 production. In the endocardial endothelial preparation, we think this smaller amount was related to the presence of tissue, which served as a binding scaffold, thereby facilitating the generation of H2O2 (see below) such that a larger concentration of Lzm-S was required when tissue was not present.
We also used the Ultra Amplex red H2O2 assay to confirm the results obtained with the electrode experiments (12). Although there was a difference in the amount of H2O2 produced by Lzm-S between the two methodologies, this could be due to the different sensitivities between the two techniques to the generation of H2O2 or to differences between the two techniques in the amount of time the Lzm-S solution was exposed to light, or because when the Ultra Amplex red assay technique was used, the Lzm-S had to be initially thawed from its frozen state, after which the Lzm-S was added to the buffer, after which Lzm-S-H2O2 solution was frozen until enough samples could be obtained for fluorescence analysis, after which the solution was again thawed. These, as well as unrecognized considerations, may have been responsible for the differences between the two methods. At any rate, the significance of these Ultra Amplex red assay findings is that they confirmed that Lzm-S is capable of generating H2O2 in a manner described by Wentworth et al. (40).
In both the electrode and Ultra Amplex red assay measurements, moreover, there was a dose-response effect of Lzm-S on H2O2 production. However, a threshold concentration of Lzm-S was required before any effect could be observed. Although the generation of H2O2 by Lzm-S appears to be a physical process (see below), there would still be a threshold amount of Lzm-S required before H2O2 could be generated. This threshold response would be analogous to what is observed in any chemical-agonist receptor interaction where a certain concentration of the agonist is required before an effect can be appreciated. Wentworth et al. (40) also did not find a significant generation of H2O2 until a concentration of ∼5 × 10−6 mol/l immunoglobulin was attained. This is a value similar to what we found in the present study and suggests that an identical process is involved.
In both the electrochemical probe and Ultra Amplex red assay experiments, we found that light was required for Lzm-S to generate H2O2. When experiments were performed in the dark, the concentration of H2O2 generated by both techniques was much less than when light was present. Similar to the findings of Wentworth et al. (40), we also showed in the nontissue experiments that the singlet oxygen quencher NaN3 prevented the production of H2O2. In addition, we performed subsequent experiments without tissue in which we used an oxygen sensor to measure oxygen consumption as described by Kramarenko et al. (19). We found that the decrease in oxygen as measured by the oxygen sensor was comparable to the increase in H2O2 produced as determined by the H2O2 sensor. Moreover, when we examined whether singlet oxygen was involved in this oxygen consumption, we were able to inhibit this consumption with NaN3. All of these experiments would favor the concepts of Wentworth et al. (40) that a protein such as Lzm-S may facilitate the catalytic conversion of 1O2• and H2O to H2O2, thus allowing this reaction to take place. However, since we did not measure 1O2• directly and because NaN3 may have other effects, our conclusion with respect to 1O2• still remains speculative. Furthermore, although in the laboratory light was required to generate singlet oxygen to form H2O2, it is important to note that as pointed out by Nieva and Wentworth (33), in the septic patient, singlet oxygen would be derived from leukocytes.
In terms of a plausible mechanism by which immunoglobulins would generate H2O2, Datta et al. (11) used quantum mechanical calculations to delineate this process. They included a reaction of 1O2• with two waters to form HOOOH (plus H2O) followed by formation of HOOOH dimer, which rearranges to form HOO-HOOO + H2O, which rearranges to form 2H2O2 plus 1O2• or 3O2. Wentworth et al. (43) have shown that species such as dihydrogen trioxide, the hydrotrioxy radical HO•3, and ozone also could be found as intermediates or end products of this conversion to H2O2 by 1O2• (42, 43). We did not find that the hydroxyl radical scavengers mannitol and DMSO were able to block the generation of H2O2 by Lzm-S. Although the physical chemistry by which immunoglobulins and Lzm-S may generate H2O2 remains complex and further work needs to be performed, the present study shows that Lzm-S is capable of generating H2O2 in a manner that supports the concepts of Wentworth et al. (40).
Reversible myocardial depression and arterial vasodilation in septic shock result from the release of inflammatory mediators (2, 34). Lzm-S is a newly discovered mediator of myocardial dysfunction in septic shock (30). Lzm-S is a somewhat unique protein in that we have shown that it binds to surface glycoprotein(s) on the endocardial endothelium, after which it generates H2O2 (17, 29). Because of this binding, the distance over which H2O2 needs to diffuse for activation of cellular pathways would be short. On the other hand, without this binding, the local H2O2 concentrations achieved may be insufficient to cause myocardial depression, as was shown in a previous study where the carbohydrate moieties to which Lzm-S binds on the endocardial endothelium were removed (17). In sepsis, whether there are other proteins that have the capability of generating H2O2 is not clear, but on the basis of work by Wentworth et al. (40), this also may be the case. This study provides new evidence that a mediator of sepsis may intrinsically generate H2O2 and, furthermore, that this generation may activate pathways leading to the hemodynamic deterioration characteristic of this inflammatory process. Septic shock is an important cause of morbidity and mortality (2). Inhibitors of the effects of Lzm-S could provide for new treatments in septic shock.
This work was supported by Canadian Institutes of Health Research, Health Sciences Centre Foundation, and the Biology of Breathing Group, Manitoba Institutes of Child Health.
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