© 1995 American Heart Association, Inc.
Articles |
Microvascular Responses to Inhibition of Nitric Oxide Production
Role of Active Oxidants
From the Departments of Physiology, Medicine, and Cellular Biology and Anatomy, Center of Excellence in Arthritis and Rheumatology, LSU Medical Center, Shreveport, La.
Correspondence to Dr D. Neil Granger, Department of Physiology and Biophysics, LSU Medical Center, 1501 Kings Hwy, PO Box 33932, Shreveport, LA 71130-3932.
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Abstract The objective of this study was to assess the potential contribution of hydrogen peroxide (H2O2) to the leukocyte–endothelial cell adhesion and increased microvascular permeability observed in rat mesenteric venules after inhibition of nitric oxide synthesis with NG-nitro-L-arginine methyl ester (L-NAME). Leukocyte adherence and emigration and leakage of fluorescein isothiocyanate–labeled albumin were monitored in postcapillary venules before and after exposure of the tissue to L-NAME. H2O2 production in mesenteric tissue was monitored by using dihydrorhodamine 123 (DHR), the H2O2-sensitive fluorochrome. L-NAME elicited a rapid increase in both the rate of albumin extravasation and oxidation of DHR, which was followed by an increased adherence and emigration of leukocytes in postcapillary venules. Treatment with either catalase or dimethylthiourea attenuated the L-NAME–induced oxidative stress, albumin leakage, and leukocyte–endothelial cell adhesion. Oxidation of DHR was enhanced in animals treated with either 3-amino-1,2,4-triazole (ATZ), an inhibitor of endogenous catalase, or a combination of ATZ and maleic acid diethyl ester, which depletes intracellular glutathione. Animals receiving a CD11/CD18-specific antibody to prevent leukocyte adhesion/emigration exhibited a reduced oxidation of DHR in response to L-NAME. These findings indicate that most of the H2O2 (and secondarily derived oxidants) generated in mesenteric tissue exposed to an inhibitor of nitric oxide production is due to accumulation of activated leukocytes.
Key Words: leukocyte–endothelial cell adhesion • vascular permeability • platelet-leukocyte aggregation • mast cell degranulation • dihydrorhodamine 123
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Introduction |
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There is a growing body of evidence that implicates nitric oxide (NO) as an important modulator of the adhesive interactions between leukocytes, platelets, and microvascular endothelium.1 This view is supported by data derived from in vitro and in vivo experiments that demonstrate leukocyte–endothelial cell adhesion,2 platelet–endothelial cell adhesion,1 and homotypic and heterotypic aggregation of leukocytes and platelets1 3 4 after inhibition of NO synthesis. Similar cell-cell interactions are observed in the microvasculature of tissues exposed to ischemia/reperfusion, a condition associated with diminished production of NO.5 Additional evidence that supports a role for NO in modulating blood cell–endothelial cell interactions is provided by the observation that NO donors (eg, spermine-NO and nitroprusside) prevent the adhesive interactions elicited by exposure of the microvasculature to either NO synthase inhibitors6 or ischemia/reperfusion.5 NO-mediated attenuation of cell-cell interactions appears to play an important role in maintaining the normal barrier function of the microvasculature, since the increased albumin leakage observed in venules exposed to NO synthase inhibitors is largely prevented when leukocyte–endothelial cell adhesion is inhibited with a monoclonal antibody directed against the leukocyte adhesion glycoprotein CD11/CD18.1
Although the cell-cell interactions and microvascular dysfunction associated with inhibition of NO synthesis have been well characterized, the biochemical mechanisms that underlie this response remain undefined. It has recently been proposed that oxygen-derived free radicals contribute to the leukocyte adherence and emigration and the subsequent endothelial cell injury elicited by inhibitors of NO synthase.7 Gaboury et al8 have recently reported that NO donors (eg, 3-morpholinosydonimine-N-ethylcarbamide) blunt the leukocyte–endothelial cell adhesion response normally observed in mesenteric venules exposed to the oxygen radical–generating system, hypoxanthine–xanthine oxidase. On the basis of these findings, the authors suggested that inhibition of NO production leads to excessive accumulation of superoxide, a free radical that is effectively scavenged by NO. Superoxide accumulation could then result in the formation of inflammatory mediators, increased expression of adhesion molecules, and increased microvascular protein leakage.9 10 11
There is also evidence that hydrogen peroxide (H2O2) may contribute to the microvascular alterations elicited by inhibition of NO synthesis. Monolayers of cultured human umbilical vein endothelial cells (HUVECs) exposed to the NO synthase inhibitor NG-nitro-L-arginine methyl ester (L-NAME) sustain an increased number of adherent neutrophils and exhibit an increased oxidation of the H2O2-sensitive probe 2,7-dichlorofluorescein diacetate.12 A platelet-activating factor (PAF)-receptor antagonist prevented the L-NAME–induced neutrophil adhesion, which is consistent with the leukocyte adhesion responses observed in postcapillary venules exposed to L-NAME13 and with the observation that PAF mediates the enhanced neutrophil adhesion to HUVEC monolayers exposed to H2O2.9 This evidence of an increased production of H2O2 by L-NAME–treated endothelial cells, coupled with the recent demonstration that NO protects cultured cells from H2O2-induced cytotoxicity,14 would suggest that H2O2 and/or secondarily derived oxidants may be a major chemical mediator of the cell-cell adhesive interactions and barrier dysfunction observed in postcapillary venules exposed to NO synthase inhibitors.
A major objective of the present study was to determine whether the cell-cell adhesive interactions and albumin leakage observed in postcapillary venules exposed to the NO synthase inhibitor are associated with an increased production of H2O2. The fluorescent probe dihydrorhodamine 123 (DHR) was used to monitor the production of H2O2 in rat mesentery. Another objective was to assess the contribution of H2O2 to the increased leukocyte–endothelial cell adhesion, platelet-leukocyte aggregation, and albumin leakage normally observed in L-NAME–treated mesenteric venules. Finally, we also assessed the contribution of adherent and emigrated leukocytes to the increased oxidation of DHR elicited by L-NAME.
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Materials and Methods |
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Surgical Procedure
Male Sprague-Dawley rats (200 to 250 g) were maintained on a purified laboratory diet and fasted for 24 hours before each experiment. The animals were initially anesthetized with pentobarbital (65 mg/kg body wt), and then a tracheotomy was performed to facilitate breathing during the experiment. The right carotid artery was cannulated, and systemic arterial pressure was measured with a Statham P23A pressure transducer connected to the carotid artery cannula. Systemic blood pressure and heart rate were continuously recorded with a Grass physiological recorder (Grass Instruments). The left jugular vein was also cannulated for drug administration.
Intravital Microscopy
Rats were placed in a supine position on an adjustable Plexiglas
microscope stage, and the mesentery was prepared for microscopic
observation as described previously.1 5 15 Briefly, the
mesentery was draped over a nonfluorescent coverslip that allowed for
observation of a 2-cm2 segment of tissue. The exposed bowel
wall was covered with Saran Wrap (Dow Chemical Co), and then the
mesentery was superfused with bicarbonate-buffered saline (BBS, 37°C,
pH 7.4) that was bubbled with a mixture of 5% CO2 and 90%
N2.
An inverted microscope (TMD-2S, Diaphoto, Nikon) with a x40 objective lens was used to observe the mesenteric microcirculation. The mesentery was transilluminated with a 12-V 100-W direct current–stabilized light source. A video camera (VK-C150, Hitachi) mounted on the microscope projected the image onto a color monitor (PVM-2030, Sony), and the images were recorded with a videocassette recorder (NV8950, Panasonic). A video time-date generator (WJ810, Panasonic) projected the time, date, and stopwatch function onto the monitor.
Single unbranched venules with diameters ranging between 25 and 35 µm and lengths >150 µm were selected for study. Venular diameter was measured either on-line or off-line by using a video caliper (Microcirculation Research Institute, Texas A&M University, College Station, Tex). Red blood cell centerline velocity was measured in venules by using an optical Doppler velocimeter (Microcirculation Research Institute, Texas A&M University). The velocimeter was calibrated against a rotating glass disk coated with red blood cells. Venular blood flow was calculated from the product of mean red blood cell velocity (Vmean=centerline velocity/1.6)16 and microvascular cross-sectional area; cylindrical geometry was assumed. Wall shear rate (SR) was calculated by the newtonian definition: SR=8(Vmean/diameter).
The number of adherent leukocytes was determined off-line during
playback of videotaped images. A leukocyte was considered to be
adherent to venular endothelium if it remained stationary for a period
30 seconds.17 Adherent cells were expressed as the
number per 100-µm length of venule. The number of emigrated
leukocytes was also determined off-line during playback of videotape
images. Any interstitial leukocytes present in the mesentery at the
onset of the experiment were subtracted from the total number of
leukocytes that accumulated during the course of the experiment.
Leukocyte emigration was expressed as the number per field of view
surrounding the venule. Platelet-leukocyte aggregates that were visible
within postcapillary venules were quantified and expressed as the
number of aggregates crossing a fixed point within the venule over a
5-minute period.1 5 15 To visualize mast cells
surrounding the mesenteric microvasculature, 0.1% of toluidine blue
(Sigma Chemical Co) was added to the mesentery at the termination of
each experiment.5 The number of intact and degranulated
mast cells was determined, and the percentage of degranulated mast
cells was calculated.
To quantify albumin leakage across mesenteric venules, 50 mg/kg of fluorescein isothiocyanate (FITC)-labeled bovine albumin (Sigma) was administered intravenously to the animals 30 minutes before each experiment.1 5 15 Fluorescence intensity (excitation wavelength, 420 to 490 nm; emission wavelength, 520 nm) was detected by using a silicon-intensified target camera (C-2400-08, Hamamatsu Photonics). The fluorescence intensity of FITC-albumin within three segments of the venule under study (Iv) and in three contiguous areas of perivenular interstitium (Ii) was measured at various times after the administration of FITC-albumin with a computer-assisted digital imaging processor (NIH Image 1.35 on a Macintosh computer). The windows to measure average fluorescence intensities within and along the venule were set at 50-µm length and 25-µm width. Over the entire L-NAME superfusion period, plasma FITC-albumin concentration and intravenular fluorescence intensity fell by <10%. An index of vascular albumin leakage was determined from the Ii-to-Iv ratio at specific intervals after L-NAME exposure, ie, 10 and 30 minutes.
The oxidant-sensitive fluorescent probe DHR (Molecular Probes, Inc)18 19 20 21 22 was added to the mesenteric superfusate (10 µmol/L) in some experiments to monitor oxidant stress. The fluorochrome was visualized by using the same microscope and image analysis system described above. DHR fluorescence intensity was monitored in a region of mesentery that was equivalent to twice the area of the venule under observation. An image processor was used to monitor fluorescence intensity just before (baseline value [It]=0) and after superfusion of the mesentery with L-NAME (It=x). The ratio of It=x to It=0 was used as an index of oxidative stress in mesenteric tissue. DHR is oxidized to fluorescent rhodamine 123 (RH 123, Sigma) by intracellular and extracellular secondary H2O2-dependent reactions.22 Since RH 123 binds to the mitochondrial inner membrane, any RH 123 formed by the oxidation of DHR is stable within endothelial cells and other cell types.22 RH 123 is known to be a mitochondrial membrane potential–sensitive fluorescent probe23 24 25 that responds to mitochondrial ATP production. Therefore, in some experiments, RH 123 rather than DHR was superfused onto the mesenteric preparation (10 µmol/L) as a control group.
Experimental Protocols
After all parameters measured on-line were in a steady state,
the mesentery was superfused (1.25 mL/min) with BBS alone for a period
of 30 minutes, with video recordings and measurements of all parameters
made at the onset and 30 minutes into the superfusion period. For an
additional 30 minutes, the mesentery was superfused with either L-NAME
(100 µmol/L) or BBS alone (control group). In some experiments,
either superoxide dismutase (SOD, 12 mg/kg IV with ligation of renal
arteries, Sigma),11 26 27 catalase (150 000 U/kg IV,
Boehringer Mannheim),11 26 27 or a monoclonal antibody
(MAb CL26, 100 µg per rat) directed against the common ß subunit
(CD18) of the leukocyte adhesion glycoprotein CD11/CD1828
was administered 30 minutes before the L-NAME superfusion, and the same
protocol was used. In another series of experiments,
N,N'-dimethylthiourea
(DMTU, Janssen Chimica), a small, permeable, and relatively nontoxic
scavenger of H2O2 and the hydroxyl radical
( · OH),29 30 31 32 was added to the superfusate to give a
final concentration of 10 mmol/L. To assess the potential involvement
of intracellular antioxidants, animals were treated either with an
inhibitor of endogenous catalase, 3-amino-1,2,4-triazole (ATZ, 1
mmol/L, Sigma),32 33 34 or with a depletor of cell
glutathione (GSH), maleic acid diethyl ester (diethyl maleate
[DMA], 100 µmol/L, Sigma) 60 minutes before L-NAME
superfusion.35 In some experiments, both ATZ and DMA were
added to the superfusate along with L-NAME, and the same protocol was
used.
In a separate group of animals (n=6), we assessed whether superfusion of rat mesentery with 100 µmol/L monomethyl-L-arginine (L-NMMA), another inhibitor of NO synthase, elicits the same changes in DHR oxidation, leukocyte–endothelial cell adhesion, and albumin leakage that were noted with L-NAME. For these studies, the same protocol was used as described above for L-NAME, except that L-NMMA was added to the superfusate, and the mesentery was exposed for a period of 30 minutes.
The activities of constitutive and inducible NO synthase (cNOS and
iNOS, respectively) were determined in rat mesenteric samples (
200
mg) harvested from three control rats. The method of Weinberg et
al36 was used for the assay. Rat mesentery was homogenized
for 30 seconds in 1.0 mL HEPES buffer (pH 7.4) containing 0.1 mmol/L
EDTA, 1.0 mmol/L dithiothreitol (DTT), 10 µmol/L tetrahydrobiopterin,
and 0.2 mmol/L phenylmethylsulfonyl fluoride. After centrifugation at
15 000g and 4°C, the supernatant was removed and mixed
with 1.0 mL of 1:1 (wt/vol) Dowex/HEPES buffer (50 mmol/L, pH 7.4) and
allowed to incubate for 5 minutes at 4°C. The samples were then
centrifuged at 400g for 5 minutes at 4°C, and the
supernants were removed and placed on ice. Total (cNOS plus iNOS)
activity was determined by using 50-µL aliquots of sample, which were
incubated in a total volume of 200 µL containing 50 mmol/L HEPES (pH
7.4), 10 µmol/L BH4, 10 µmol/L FAD, 10 µmol/L
L-[14C]arginine, 1.0 mmol/L
CaCl2, 1.0 mmol/L MgCl2, and 0.1
mmol/L NADPH. Ca2+-independent NO synthase (iNOS)
activity was assessed in the absence of CaCl2 and in the
presence of 1.0 mmol/L EGTA. After a 60-minute incubation at 37°C,
the reaction was terminated by the addition of 0.5 mL of 1:1 (wt/vol)
Dowex/HEPES mixture (50 mmol/L, pH 7.4) to remove the unreacted
L-[14C]arginine. The Dowex/sample mixture
was centrifuged at 10 000g for 5 minutes at 25°C, and the
supernatant was saved. The Dowex pellet was then washed three times by
centrifugation using 600 µL of 50 mmol/L HEPES, pH 7.4, for each
wash. All supernatants were combined and added to 15 mL of PolyFluro
scintillation cocktail (Packard Instruments). Samples were counted for
14C activity. Only L-NAME– (or L-NMMA)–inhibitable (1
mmol/L) activity was used to calculate cNOS and iNOS activities. Total
protein content for each sample was determined by the Bio-Rad protein
assay with bovine serum albumin used as a standard. cNOS activity was
calculated from the difference between total and iNOS activities and
expressed as picomoles per minute per milligram protein.
In three rat mesenteric preparations exposed to 100 µmol/L L-NAME for 30 minutes, the mesentery was perfused with 2% paraformaldehyde–2.5% glutaraldehyde in 0.1 mol/L cacodylate buffer, pH 7.4, via the superior mesenteric artery. The mesenteric tissue was excised, postfixed in 1% OsO4 in 0.1 mol/L cacodylate buffer, dehydrated in ethanol, and embedded in epoxy resins (Polybed 812/Araldite, Polysciences, Inc). Silver sections (60 to 70 nm) were cut with a diamond knife, stained with 1% uranyl acetate in 50% ethanol and lead citrate, and examined with a Philips CM10 electron microscope.
Statistics
The data were analyzed by standard statistical analysis, ie,
one-way ANOVA and Fisher's post hoc test. All values are reported as
mean±SEM from six rats, and statistical significance was set at
P<.05.
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Results |
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The total activity of NO synthase in rat mesentery was 2.03±0.22 pmol/min per milligram, with 83.7±3.8% of this activity due to the constitutive (Ca2+-dependent) form of the enzyme. L-NAME, at concentrations of 100 µmol/L and 1 mmol/L, reduced total NO synthase activity by 85% and 98%, respectively. L-NMMA, at the same concentrations, reduced mesenteric NO synthase activity by 70% and 78%, respectively. Examination of mesenteric venules exposed to 100 µmol/L L-NAME by electron microscopy revealed a modest number of adherent leukocytes and endothelial cells that appeared normal with no apparent pathological changes. The mild inflammatory response elicited by L-NAME was very similar to that previously described for mesenteric venules exposed to ischemia/reperfusion.37
Table 1
summarizes the alterations in venular
hemodynamics associated with L-NAME superfusion of rat mesenteric
venules. Within 30 minutes after beginning the L-NAME superfusion, red
blood cell velocity and wall shear rate decreased by 22% and 28%,
respectively, whereas venular diameter remained unchanged from the
control value. The reduction in erythrocyte velocity and shear rate
induced by L-NAME was significantly blunted by treatment with DMTU but
not by catalase. A further reduction in venular wall shear rate was
noted in mesenteric preparations treated with a combination of ATZ and
DMA but not with either agent alone.
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Fig 1 demonstrates the effects of exogenously
administered antioxidants on the adherence (panel A) and emigration
(panel B) of leukocytes in postcapillary venules exposed to 100
µmol/L L-NAME. In animals subjected to superfusion with BBS alone for
30 minutes, leukocyte adherence was 2.5±0.7 per 100 mm, with 1.3±0.7
emigrated leukocytes per field. Corresponding values obtained in
mesenteric preparations exposed to 100 µmol/L L-NAME were 17.2±1.7
adherent leukocytes per 100 mm and 9.0±1.2 per field. Catalase and
DMTU significantly attenuated the L-NAME–induced leukocyte adhesion to
venular endothelium by 53% and 69%, respectively (Fig 1A). The
increased number of emigrated leukocytes observed in mesenteric venules
exposed to L-NAME was also significantly attenuated by the
administration of either catalase (64%) or DMTU (58%) (Fig 1B).
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P<.05 vs L-NAME–untreated group.
Fig 2 presents the effects of inhibitors of
intracellular antioxidants on the adherence (panel A) and emigration
(panel B) of leukocytes in postcapillary venules exposed to 100
µmol/L L-NAME. Contrary to the responses observed in control
preparations, a significant increase in the number of adherent
leukocytes was observed at 10 minutes into the L-NAME superfusion
period in preparations treated with either ATZ alone or the combination
of ATZ and DMA. Treatment with a combination of ATZ and DMA also
elicited a larger increase in L-NAME–induced leukocyte adhesion than
did L-NAME alone (Fig 2A). The number of emigrated leukocytes at 10
minutes of L-NAME superfusion was also increased by treatment with
either ATZ, DMA, or a combination of ATZ and DMA, whereas L-NAME alone
did not significantly increase leukocyte emigration at 10 minutes (Fig 2B).
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Fig 3 illustrates the changes in fluorescence intensity
of the H2O2-sensitive probe DHR during
superfusion of the mesentery with L-NAME. Superfusion of the mesentery
with BBS was not associated with significant DHR activation (Fig 3, top
left panel). Within 10 minutes of L-NAME superfusion, areas of
fluorescence intensity were observed along the venule (Fig 3, top right
panel). This L-NAME–induced increase in DHR-associated fluorescence
gradually increased along the venule until the end of observation
period (Fig 3, bottom panels). Although DHR activity was largely
confined to venular endothelium, areas of fluorescence intensity were
also observed in the interstitium of rat mesentery exposed to L-NAME.
Ten percent to 20% of the mast cells in the perivenular space
(identified by toluidine blue staining) were DHR
fluorescence–positive.
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Fig 4 compares the time course of the changes in DHR
fluorescence intensity (panel A) with the simultaneously monitored
increases in the number of adherent (panel B) and emigrated (panel C)
leukocytes in single postcapillary venules exposed to L-NAME. DHR
fluorescence increased as early as 10 minutes after starting the L-NAME
superfusion (543% increase above baseline value). DHR fluorescence
intensity continued to increase until the end of the observation
period, ie, at 30 minutes (969% increase above the baseline value).
The increased DHR-fluorescence at 20 and 30 minutes of L-NAME
superfusion was associated with increased numbers of adherent and
emigrated leukocytes, as observed in other experiments (Figs 1
and 2).
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Superfusion of rat mesentery with 100 µmol/L L-NMMA elicited changes in DHR fluorescence and leukocyte–endothelial cell adhesion similar to those found with L-NAME. At 30 minutes of L-NMMA superfusion, 15.0±2.9 adherent leukocytes and 6.2±1.5 emigrated cells were noted. These changes were accompanied by a 393±75% increase in DHR fluorescence intensity. The smaller responses elicited by L-NMMA, when compared with L-NAME, presumably reflect the differences in the ability of the two agents to inhibit rat mesenteric NO synthase activity (see above).
Fig 5 summarizes the effects of different antioxidants
(panel A) and inhibitors of intracellular antioxidants (panel B) on DHR
fluorescence at 10 and 30 minutes after L-NAME superfusion. At 10
minutes, none of the agents studied had a significant effect on the
L-NAME–induced increase in DHR fluorescence. At 30 minutes, the
L-NAME–induced DHR activation was significantly attenuated by
treatment with either catalase (77%) or DMTU (66%) (Fig 5A). The
superoxide radical–scavenging enzyme SOD did not significantly alter
L-NAME–induced DHR fluorescence (data not shown). An enhancement of
L-NAME–induced DHR fluorescence was noted in mesenteric preparations
treated with ATZ (1650±387% of baseline value) when compared with
L-NAME alone (892±135%). The combination of ATZ and DMA further
increased L-NAME–induced DHR activation (2777±406%) (Fig 5B).
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Table 2 summarizes effects of treatment with a
monoclonal antibody directed against the common ß subunit (CD18) of
the leukocyte adhesion glycoprotein CD11/CD18 on L-NAME–induced
leukocyte adherence and emigration and DHR activation. At 30 minutes,
the CD18-specific monoclonal antibody significantly attenuated the
L-NAME–induced increases in the number of adherent and emigrated
leukocytes as well as the increased DHR fluorescence.
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Superfusion of the mesentery with L-NAME for 30 minutes did not alter fluorescence intensity of RH 123. After 30 minutes of L-NAME superfusion, the RH 123 fluorescence intensity was 116±5% (P>.05) of the corresponding baseline value. L-NAME plus ATZ and DMA, which induced the maximal increase in DHR fluorescence intensity, slightly (but not significantly) reduced RH 123–associated fluorescence (88±18% of the baseline value). These observations indicate that the increased DHR fluorescence intensity observed in the present study is not likely explained on the basis of changes in mitochondrial ATP production.
Fig 6 summarizes the effects of different antioxidants
(panel A) and inhibitors of intracellular antioxidants (panel B) on
L-NAME–induced albumin leakage. After 30 minutes of BBS superfusion
(control), an albumin leakage index of 4.7±1.9% was observed. A
significant increase in albumin leakage (30.7±6.4%) was noted as
early as 10 minutes after the L-NAME superfusion. This early increase
in albumin extravasation was attenuated by the antioxidants catalase
(43%) and DMTU (36%). The larger increase in albumin leakage
(52.8±4.2%) observed at 30 minutes of L-NAME superfusion was also
attenuated by administration of either catalase (56%) or DMTU (66%)
(Fig 6A). Treatment with either ATZ alone or ATZ plus DMA further
enhanced the early (10-minute) albumin leakage induced by L-NAME,
whereas these agents had no effect on albumin extravasation at 30
minutes of L-NAME superfusion (Fig 6B). In preparations superfused with
100 µmol/L L-NMMA, albumin leakage increased to 11.8±2.3% at 10
minutes and to 33.7±6.8% at 30 minutes.
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Exposure of the rat mesentery to certain stimuli is frequently
accompanied by the appearance of large platelet-leukocyte aggregates in
postcapillary venules. These aggregates have been observed in
mesenteric venules exposed to the NO synthase inhibitor
L-NAME.1 Although such aggregates are not observed during
control conditions, 6.2±0.5 aggregates per 5 minutes were observed in
venules exposed to 100 µmol/L L-NAME. Table 3
demonstrates that the L-NAME–induced formation of platelet-leukocyte
aggregates is significantly reduced in mesenteric preparations treated
with either catalase and DMTU but not in preparations treated with
inhibitors of intracellular scavengers.
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Table 3 also summarizes the responses of mast cells along the
postcapillary venules to 30 minutes of L-NAME superfusion. In the
control group, degranulated mast cells represented <3% of total
mast cell population situated along postcapillary venules. At 30
minutes after the L-NAME superfusion, degranulated mast cells increased
to 25%. Both antioxidants used in the present study (catalase
and DMTU) significantly attenuated the L-NAME–induced mast cell
degranulation. Inhibition of endogenous catalase (ATZ) as well as
depletion of GSH (DMA) had no significant effect on the L-NAME–induced
degranulation of mast cells.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Previous studies have shown that inhibition of NO synthesis with analogues of L-arginine enhances cell-cell adhesive interactions in the microcirculation and reduces the barrier function of microvascular endothelium in postcapillary venules.1 2 13 Mechanistic studies have revealed that the increased leukocyte adherence and albumin extravasation in rat mesenteric venules elicited by L-NAME involve the participation of PAF13 and a leukocyte–endothelial cell adhesive interaction that is mediated by the ß2-integrin CD11/CD18 on leukocytes and intercellular adhesion molecule-1 (ICAM-1) on endothelial cells.1 2 The results of the present study extend the growing body of literature on NO to invoke a role for H2O2 (or a secondarily derived oxidant) in the cell-cell interactions and microvascular barrier dysfunction elicited by L-NAME. The present study provides three separate lines of evidence that support the involvement of H2O2 in the microvascular responses to NO synthase inhibition: (1) Both catalase and DMTU, which effectively scavenge or detoxify H2O2, significantly attenuate the responses to L-NAME. (2) Depletion of endogenous defense mechanisms against H2O2 (catalase and GSH) exaggerates the responses to L-NAME. (3) Exposure of the mesentery to L-NAME results in a profound increase in the oxidation of DHR, which reflects an increased production of H2O2 (or a secondarily derived oxidant).
There is a large body of evidence in the literature that supports a role for H2O2 in modulating leukocyte–endothelial cell adhesion and microvascular permeability. Monolayers of cultured HUVECs,9 isolated perfused carotid arteries,38 and mesenteric venules39 all sustain a higher level of neutrophil adhesion after exposure to H2O2. Furthermore, catalase administration has been shown to reduce the leukocyte adherence in mesenteric venules elicited by ischemia/reperfusion, a condition that is associated with enhanced H2O2 production.11 26 Both in vitro and in vivo studies have implicated a role for PAF in the enhanced leukocyte–endothelial cell adhesion elicited by H2O2.9 27 40 As would be predicted for a PAF-mediated adhesion process, H2O2-induced leukocyte adhesion involves the engagement of CD11/CD18 on leukocytes with ICAM-1 on vascular endothelial cells.17 There is a similar body of evidence that implicates H2O2 as a modulator of vascular permeability. Generation of H2O2 within the microvasculature is associated with increased macromolecule extravasation.41 Similarly, catalase has been shown to be effective in attenuating the microvascular permeability responses elicited by a variety of inflammatory stimuli.27 Thus, our finding that H2O2 participates in the cell-cell adhesive interactions and microvascular barrier dysfunction associated with NO synthase inhibition is consistent with the known biological actions of this reactive oxygen species.
In an effort to monitor the oxidative stress elicited by L-NAME, we used the H2O2-sensitive fluorescent probe DHR, which has been successfully used to measure intracellular H2O2 levels in cultured endothelial cells and in other cell types.18 19 20 21 DHR exhibits a variety of physicochemical and biological properties that make it an ideal agent for detecting secondary H2O2-dependent reactions in living tissue by use of fluorescence videomicroscopy. Such properties include cell membrane permeability, minimal cellular toxicity, specificity of the nonfluorescent form (DHR) for reactive oxygen species, and intracellular sequestration of the fluorescent oxidized form of the probe.22 Royall et al22 have recently provided evidence that the oxidation of DHR is not due to H2O2 alone but appears to occur by reactions involving reactive oxygen species. Nonetheless, DHR oxidation appears to be far more sensitive to H2O2 than superoxide,22 a view that is corroborated by our ability to reduce L-NAME–induced DHR oxidation with catalase and DMTU but not SOD.
The possibility that NO synthase inhibition is associated with an increased formation of oxidants derived secondarily from H2O2 is supported by the results of a recent study wherein the hydroperoxide-sensitive fluorogenic probe carboxydichlorofluorescein (CDCF) was used in rat mesenteric preparations exposed to 100 µmol/L L-NAME. Suematsu et al42 demonstrated that the increased CDCF fluorescence elicited by L-NAME was significantly diminished by pretreatment with desferrioxamine, suggesting that the CDCF fluorescence is linked to an iron-catalyzed hydroperoxide flux.
The temporal responses of DHR oxidation in mesenteric tissue exposed to L-NAME suggest that there is an early rise in DHR oxidation that occurs within 10 minutes of NO synthase inhibition, followed by a slower more profound increase in DHR oxidation over the next 30 minutes of L-NAME exposure. There are several possible sources of the active oxygen species that are produced in tissues exposed to L-NAME. One possible source is the endothelial cell, which can produce H2O2 via the mitochondrial electron transport chain, xanthine oxidase, and/or the biosynthesis of prostaglandins.42 43 It has been proposed that the NO normally produced by endothelial cells may serve to scavenge the superoxide generated by the same cells.44 In this instance, one would expect that inhibition of NO production with L-NAME could result in an accumulation of superoxide and H2O2 within endothelial cells. An alternate source of H2O2 in L-NAME–treated tissue is the adherent and activated granulocyte. It has been reported that NO reduces the generation of superoxide by granulocytes by inhibiting the activity of the membrane-associated enzyme NADPH oxidase.45 Thus, L-NAME may promote the generation of H2O2 by acting through such a process, or it may do so indirectly by eliciting the formation of inflammatory mediators (PAF)46 that activate granulocytes and induce leukocyte–endothelial cell adhesion.47 Adherent granulocytes have been shown to produce far greater quantities of superoxide than leukocytes in suspension.48
We observed that catalase and DMTU were very effective in reducing both the leukocyte adherence and DHR oxidation in postcapillary venules that are normally elicited by L-NAME. These observations raise a question as to whether L-NAME–induced leukocyte adherence is a cause or an effect of the accompanying DHR oxidation. We addressed this question by examining the influence of a CD18-specific monoclonal antibody, which largely abolishes the leukocyte adherence induced by L-NAME, on the extent of DHR oxidation. Our findings indicate that although the initial increase in DHR oxidation is not affected by prevention of leukocyte adherence, the slower, more profound DHR oxidation observed after 30 minutes of L-NAME exposure was largely prevented by the CD18-specific antibody. These results suggest that recruited leukocytes are the major source of the H2O2 (or a secondarily derived oxidant) produced after inhibition of NO synthesis in the rat mesentery. Nonetheless, our data do not exclude a role for endothelial cells as a source of oxidants, particularly in the initial period of L-NAME exposure.
The possibility that L-NAME enhances oxidant production by venular endothelial cells is supported by the work of Suematsu et al.42 They observed that the enhanced oxidation of CDCF in rat mesenteric venules exposed to L-NAME is largely unaffected by antibodies that prevent leukocyte–endothelial cell adhesion when measured at 30 minutes of L-NAME exposure, but there is a significant diminution of CDCF fluorescence by antiadhesion interventions at 60 minutes of L-NAME superfusion. The discrepant results between that study and the present study are not readily explained; however, differences in the oxidant-sensitive fluorochromes and antiadhesion reagents used could provide an explanation. The former possibility appears tenable, inasmuch as 30 minutes was required for L-NAME to elicit significant oxidation of CDCF,42 whereas DHR oxidation was detected as early as 10 minutes after L-NAME superfusion.
The results of the present study also implicate a role for H2O2 in the mast cell degranulation and platelet-leukocyte aggregation normally associated with inhibition of NO synthesis.1 7 Both catalase and DMTU were effective in attenuating the mast cell degranulation elicited by L-NAME. Kubes et al7 have recently proposed that inhibition of NO synthesis allows for the accumulation of superoxide (due to the loss of the scavenging effect of NO), which in turn causes mast cell degranulation. They also proposed that mast cell products, released in response to superoxide exposure, promote the adhesion of leukocytes in postcapillary venules. Their hypothesis was based on experiments demonstrating a reduced L-NAME–induced mast cell degranulation and leukocyte adherence in rat mesentery treated with either SOD or ketotifen (a mast cell stabilizer). Our findings extend their observations to invoke a role for H2O2, in addition to superoxide, in the process of L-NAME–induced degranulation. Since both SOD and catalase are similarly effective in blunting L-NAME–induced mast cell degranulation, it is possible that a reactive oxygen species derived from the reaction of H2O2 with superoxide (eg, · OH) ultimately contributes to the mast cell activation and degranulation associated with the inhibition of NO synthesis. Our studies with DHR tend to support the possibility that mast cells produce reactive oxygen species after exposure to L-NAME, inasmuch as DHR oxidation was observed in areas surrounding mesenteric venules that corresponded to mast cells. Thus, the early burst of H2O2 produced by activated mast cells may contribute to the recruitment of leukocytes (eg, through PAF formation), which in turn account for the larger quantities of the oxidant detected at a later time.
Previous studies from our laboratory demonstrate that P-selectin mediates the platelet-leukocyte aggregation elicited by inactivation or inhibition of NO synthesis.1 5 Administration of a P-selectin–specific monoclonal antibody also diminishes the albumin extravasation associated with L-NAME exposure,1 suggesting that these aggregates contribute to the microvascular dysfunction elicited by L-NAME. The findings of the present study reveal that H2O2 participates in the formation of platelet-leukocyte aggregates in L-NAME–treated mesentery; ie, both catalase and DMTU dramatically blunted the appearance of aggregates in mesenteric venules. P-selectin expression on platelets is significantly increased in response to low levels of H2O2.49 50 51 Consequently, it appears tenable that H2O2 release from endothelial cells and/or activated and adherent leukocytes accounts for the P-selectin expression and resultant platelet-leukocyte aggregation that is associated with inhibition of NO synthesis.
Additional considerations that may bear on the interpretation of our findings are the changes in venular shear rate and depressed arteriolar production of NO that would accompany L-NAME treatment of the mesentery. L-NAME generally elicits about a 30% reduction in venular shear rate in rat mesenteric venules. Although such a fall in shear rate may be expected to elicit the recruitment of adherent leukocytes, Kubes et al2 have shown that a relatively small fraction of the adherent leukocytes observed in mesenteric venules exposed to L-NAME or L-NMMA can be attributed to the accompanying decline in shear rate. The changes in arteriolar tone that yield the decline in shear rate may also signal a potential contribution of a reduced influence of NO derived from arteriolar endothelium on downstream endothelial cells lining the postcapillary venules. Although such an endocrine influence appears tenable in view of the relatively long half-life and high diffusivity of NO, the fact that NO interacts avidly with hemoglobin in red blood cells would tend to argue against this interesting possibility.52 Nonetheless, the possibility that NO produced by arterioles diffuses to neighboring venules to exert a modulating influence cannot be readily dismissed.
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Acknowledgments |
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This study was supported by a grant from the National Heart, Lung, and Blood Institute (HL-26441) and the National Institutes of Diabetes and Digestive and Kidney Diseases (DK-43785).
Received March 22, 1994; accepted September 19, 1994.
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N. Okayama, H. Ichikawa, L. Coe, M. Itoh, and J. S. Alexander Exogenous NO enhances hydrogen peroxide-mediated neutrophil adherence to cultured endothelial cells Am J Physiol Lung Cell Mol Physiol, May 1, 1998; 274(5): L820 - L826. [Abstract] [Full Text] [PDF]
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N. R. Harris and K. W. Langlois Age-dependent responses of the mesenteric vasculature to ischemia-reperfusion Am J Physiol Heart Circ Physiol, May 1, 1998; 274(5): H1509 - H1515. [Abstract] [Full Text] [PDF]
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A. L. Baldwin, G. Thurston, and H. Al Naemi Inhibition of nitric oxide synthesis increases venular permeability and alters endothelial actin cytoskeleton Am J Physiol Heart Circ Physiol, May 1, 1998; 274(5): H1776 - H1784. [Abstract] [Full Text] [PDF]
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P. Kubes, D. Payne, and L. Ostrovsky Preconditioning and adenosine in I/R-induced leukocyte-endothelial cell interactions Am J Physiol Heart Circ Physiol, April 1, 1998; 274(4): H1230 - H1238. [Abstract] [Full Text] [PDF]
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D. J. Mitchell, J. Yu, and K. Tyml Local L-NAME decreases blood flow and increases leukocyte adhesion via CD18 Am J Physiol Heart Circ Physiol, April 1, 1998; 274(4): H1264 - H1268. [Abstract] [Full Text] [PDF]
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 J. S. Steingrub Novel Insights Into the Pathogenesis and Therapy of Circulatory Shock Seminars in Cardiothoracic and Vascular Anesthesia, March 1, 1998; 2(1): 31 - 45. [Abstract] [PDF]
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R. M. F. Wever, T. F. Luscher, F. Cosentino, and T. J. Rabelink Atherosclerosis and the Two Faces of Endothelial Nitric Oxide Synthase Circulation, January 13, 1998; 97(1): 108 - 112. [Full Text] [PDF]
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P. R. Kvietys and D. N. Granger Endothelial cell monolayers as a tool for studying microvascular pathophysiology Am J Physiol Gastrointest Liver Physiol, December 1, 1997; 273(6): G1189 - G1199. [Abstract] [Full Text] [PDF]
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N. R. Harris Opposing effects of L-NAME on capillary filtration rate in the presence or absence of neutrophils Am J Physiol Gastrointest Liver Physiol, December 1, 1997; 273(6): G1320 - G1325. [Abstract] [Full Text] [PDF]
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J. G. Filep, A. Delalandre, and M. Beauchamp Dual Role for Nitric Oxide in the Regulation of Plasma Volume and Albumin Escape During Endotoxin Shock in Conscious Rats Circ. Res., November 19, 1997; 81(5): 840 - 847. [Abstract] [Full Text]
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 N. Okayama, C. G. Kevil, L. Correia, D. Jourd'Heuil, M. Itoh, M. B. Grisham, and J. S. Alexander Nitric oxide enhances hydrogen peroxide-mediated endothelial permeability in vitro Am J Physiol Cell Physiol, November 1, 1997; 273(5): C1581 - C1587. [Abstract] [Full Text] [PDF]
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J. G. Filep Endogenous Endothelin Modulates Blood Pressure, Plasma Volume, and Albumin Escape After Systemic Nitric Oxide Blockade Hypertension, July 1, 1997; 30(1): 22 - 28. [Abstract] [Full Text]
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X.-F. Niu, G. Ibbotson, and P. Kubes A Balance Between Nitric Oxide and Oxidants Regulates Mast Cell–Dependent Neutrophil–Endothelial Cell Interactions Circ. Res., November 1, 1996; 79(5): 992 - 999. [Abstract] [Full Text]
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M.-L. Wu, K.-L. Tsai, S.-M. Wang, J.-C. Wu, B.-S. Wang, and Y.-T. Lee Mechanism of Hydrogen Peroxide and Hydroxyl Free Radical–Induced Intracellular Acidification in Cultured Rat Cardiac Myoblasts Circ. Res., April 1, 1996; 78(4): 564 - 572. [Abstract] [Full Text]
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J. P. Gaboury, X.-F. Niu, and P. Kubes Nitric Oxide Inhibits Numerous Features of Mast Cell–Induced Inflammation Circulation, January 15, 1996; 93(2): 318 - 326. [Abstract] [Full Text]
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