Insight Into the Nature and Site of Oxygen

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Insight Into the Nature and Site of Oxygen-Centered Free Radical Generation by
Endothelial Cell Monolayers Using a Novel Spin Trapping Technique
By Bradley E. Britigan, Tedmund L. Roeder, and D. Michael Shasby
Spin trapping, a sensitive and specific means of detecting
free radicals, is optimally performed on cell suspensions. This
makes it unsuitable for the study of adherent endothelial cell
monolayers because disrupting the monolayer t o induce a
cell suspension could introduce confounding factors. This
problem was eliminated through the use of endothelial cells
that were grown t o confluence on microcarrier beads. Using
the spin trap 5.5-dimethyl-1-pyrroline-N-oxide (DMPO), the
nature of free radical species generated by suspensions of
microcarrier bead adherent porcine pulmonary endothelial
cells under various forms of oxidant stress was examined.
Exposure of these endothelial cells t o paraquat resulted in
the spin trapping of superoxide (.O,-). Endothelial cell incubation in the presence of either bolus or continuous fluxes of
hydrogen peroxide (H202) yielded spin trap evidence of
hydroxyl radical formation, which was preventable by pretreating the cells with deferoxamine. Chromium oxalate
which eliminates extracellular electron paramagnetic reso-
nance spectrometry (EPR) signals, prevented the detection of
DMPO spin adducts generated by paraquat but not H,O,treated endothelial cells. When endothelial cells were coincubated with PMA-stimulated monocytes evidence of both eo2and hydroxyl radical production was detected, whereas with
PMA-stimulated neutrophils only eo2-production could be
confirmed. Neutrophil elastase, cathepsin G, and the combination of PMA and A23187 have previously been suggested
t o induce endothelial cell oxy-radical generation. However,
exposure of endothelial cells t o each of these agents did not
yield DMPO spin adducts or cyanide-insensitive endothelial
cell 0, consumption. These data indicate that endothelial cell
exposure: t o paraquat induces extracellular eo2-formation;
t o H,O, leads t o intracellular hydroxyl radical production; and
t o elastase, cathepsin G, or A23187/PMA does not appear t o
cause oxy-radical generation.
o 1992by The American Society of Hematology.
HE INJURY of pulmonary endothelial cells by both
exogenously and endogenously generated oxidant species has been linked to a variety of pathologic lung processes.‘ For example, experimental data has implicated
endothelial cell injury by neutrophil-derived superoxide
(eo,-)and hydrogen peroxide (H202)in the pathogenesis of
increased vascular permeability occurring in the adult
respiratory distress ~yndr0me.I.~
Some evidence has been
obtained that hydroxyl radical, formed through the iron
catalyzed reaction of .02-and H,O, (Haber-Weiss reaction), may be the oxidant species actually responsible for
endothelial cell inj~ry.5’~
However, these conclusions are
based only on indirect evidence showing the protective
effects of “hydroxyl radical scavengers” and iron chelators
on neutrophil-mediated endothelial cell injury.
One limitation in defining the role of specific oxygencentered free radicals in various forms of endothelial cell
injury has been the lack of free radical detection systems
that are sensitive and specific for both intracellular and
extracellular free radical species.” Both 0 - and hydroxyl
radical will react with some nitrone compounds, termed
spin traps, with the resultant generation of stable nitroxide
free radicals (spin adducts) whose characteristic hyperfine
electron paramagnetic resonance spectrometry (EPR) splitting patterns allow identification of the original free radical
species present.” Spin trapping has been increasingly applied in recent years to the in vitro and in vivo study of
oxygen-centered free radical production by a variety of
cellular systems.”
One limitation to spin trapping is that, because of the
nature of EPR, the technique is most readily applicable to
studies of cells in suspension. This poses problems for
investigating cells, such as endothelial cells, which under
normal physiologic conditions exist as adherent monolayers. Spin trapping has previously been used to study free
radical production by endothelial cells, but the cells were
studied after they had been released into suspension by
Results from such studies may not be readily
applicable to adherent cells because the scraping procedure
could potentially alter important physical and biochemical
cell parameters. Consequently, we developed a new spin
trapping system that allows detection of intracellular and
extracellular free radical production by adherent endothelial cells. This report describes the use of this technique to
investigate the nature and site of free radical production
resulting from exposure of pulmonary endothelial cells to
pharmacologic agents and human phagocytes.
T
Blood, Vol79, No 3 (February 1). 1992: pp 699-707
MATERIALS AND METHODS
Reagents. 5,s-dimethyl-1-pyrroline-N-oxide
(DMPO), 2,2,6,6tetramethylpiperidine-N-oxy1(TEMPO), xanthine, catalase, CuZn
superoxide dismutase (SOD), diethylenetriaminepentaacetic acid
(DTPA), sodium cyanide (CN-),l,l’-dimethyl-4,4’-bipyridinium
From the Research Service and the Department of Intemal Medicine, VA Medical Center; and the Department of Intemal Medicine,
University of Iowa College of Medicine, Iowa City.
Submitted January 9, 1991; accepted September 27, 1991.
Supported in part through the VA Research Service, National
Institutes of Health awards HL44275, AI28412, HL33540, and
HL42385, The Pfizer Scholars Program, and The Cystic Fibrosis
Foundation. It was performed during the tenure of B.E.B. and D.M.S.
as a VA Research Associate and VA Clinical Investigator, respectively.
Portions of this work have been presented in abstractform at the fifrh
biannual meeting of the Intemational Society for Free Radical
Research at Pasadena, CA, November 14-20, 1990 (Free Radic Biol
Med 9S1:37, 1990) and at the annual meeting of the Midwest section
of The American Federation for Clinical Research at Chicago, IL,
October 31-November 2, 1990 (Clin Res 38:871A, 1990).
Address reprint requests to Bradley E. Britigan, MD, The University
of Iowa College of Medicine, Department of Intemal Medicine,
Division of Infectious Diseases, SW54, GH, Iowa City, LA 52242.
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be hereby marked
“advertisement” in accordance with 18 U.S.C. section 1734 solely to
indicate this fact.
0 1992 by The American Society of Hematology.
0006-4971I921 7903-0033$3.0010
699
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700
dichloride (paraquat), hydrogen peroxide (30%, H,O,), glucose
oxidase, and deferoxamine mesylate were purchased from Sigma
Chemical (St Louis, MO). Additional reagents were obtained as
follows: xanthine oxidase from Boehringer-Mannheim (Indianapolis, IN); dimethyl sulfoxide (DMSO) from Fisher Scientific (Fairlawn, NJ); ethanol from Aaper Alcohol and Chemical Co (Shelbeyville, KY); and human neutrophil elastase from Elastin Products
(Owensville, MO). Cathepsin G was purified from human neutrophils by previously described method^.'^ Chromium oxalate, synthesized according to the method of Bailar et a],” was kindly provided
by Dr Gerald M. Rosen (University of Maryland School of
Pharmacy, Baltimore).
Endothelial cellculture. Confluent monolayers of porcine pulmonary artery endothelial cells were cultured on microcarrier beads
(Qtodex 3; Pharmacia, Piscataway, NJ) as previously described at
our institution.16Cells were cultured for 2 to 3 days postconfluence
(as determined by experience with this system since its original
development). Multiple cell lines were used for these studies, with
cells utilized between passage 6 and 9. In each day’s experiment,
positive and negative controls were performed with that day’s cell
preparation to eliminate any contribution of variation in cell line or
passage number to the results. Before use, microcarrier beads were
washed free of cell culture media (M199 containing 10% heatinactivated fetal bovine serum; University of Iowa Cancer Center,
Iowa City) by transferring the beads to a 15-mL conical tube
(Sarstedt, Princeton, NJ) and allowing the beads to precipitate
(-5 minutes). Culture media was then removed by aspiration and
replaced with 10 mL Hanks’ balanced salt solution without phenol
red (HBSS, University of Iowa Cancer Facility). Beads and HBSS
were gently mixed and the beads allowed to precipitate again. The
HBSS was then replaced with fresh HBSS and the process
repeated three times. After the final wash the cells were resuspended in HBSS and all experiments were then performed in this
buffer system. Except where indicated, final incubation mixtures
contained 25% (vol/vol) microcarrier beads. Based on previous
work using the identical system of cell culture,’’ this 25% vol/vol
microcarrier beads would yield a final cell concentration of 0.5 to
1.0 x lo7 cells/mL. In some experiments, microcarrier beads
lacking cells, but which had been incubated in cell culture media
and in all other respects treated identically to cell-containing
beads, were used as negative controls to eliminate any potential
contribution of the beads and/or culture media to the results.
Spin trapping. Desired reaction mixtures (0.5 mL) were assembled in 12 x 75 mm glass tubes and transferred to a quartz EPR flat
cell, which was in turn placed into the cavity of the EPR
spectrometer (Varian E104A; Varian Instruments, Palo Alto, CA).
Resulting EPR spectra were recorded sequentially at 25°C. Unless
othenvise noted, spectrometer settings were: microwave power, 20
mW; modulation amplitude, 1.0 G; modulation frequency, 100
kHz; response time, 1 second; and gain 5.0 x lo4. Routinely,
reaction mixtures contained 25% (vol/vol) microcarrier beads. The
EPR spectrum of the stable nitroxide TEMPO was unchanged with
the inclusion of up to 50% (vol/vol) Cytodex 3 microcarrier beads
in the solution, with appropriate adjustment of the solution so that
the final TEMPO concentration remained lo-’ mol/L. Thus, the
presence of the microcarrier beads did not adversely effect the
sensitivity of the system.
Oxygen consumption. Oxygen consumption was measured as
the rate of oxygen depletion from a 3-mL reaction mixture located
in the chamber of a Clark oxygen electrode (YSI 53; Yellow
Springs Instrument Co, Yellow Springs, OH) and monitored
continuously via an attached strip chart recorder. Reaction mixtures routinely contained 1 mmol/L CN- to eliminate cellular 0,
consumption resulting from mitochondrial respiration. This cyanideinsensitive respiration has been previously used as a means of
BRITIGAN, ROEDER,AND SHASBY
measuring univalent reduction to 0, to .O,- induced in cellular
systems by redox active pharmacologic compounds such as paraquat.”,’’
RESULTS
Free radical generation induced by paraquat. To optimally study pulmonary endothelial cell free radical production in vitro using spin trapping techniques, it was necessary
to develop a system in which adherent monolayers could be
examined. Accordingly, the possibility that suspensions of
microcarrier bead adherent endothelial cells could be used
for such a purpose was examined. Exposure to the herbicide
paraquat induces an acute lung injury syndrome,” which
has been suggested to result from the ability of this
compound to undergo cell-mediated redox cycling with
resultant generation of .02and H202.17.19.20
Consequently,
paraquat was used as an initial means of evaluating the
feasibility of spin trapping free radicals generated by
microcarrier bead adherent endothelial cells.
Porcine pulmonary endothelial cells were grown to confluence on microcarrier beads, after which they were
washed free of cell culture media. They were then suspended in HBSS containing 100 mmol/L DMPO and 0.1
mmol/L DTPA, after which paraquat (0.1 to 10 mmol/L)
was added. The resulting EPR spectra was that of the
hydroxyl radical-derived spin adduct of DMPO, DMPO/
.OH (A, = A, = 15.0 G; Fig 1B). No spectrum above
background was observed if either paraquat (Fig 1A) or
endothelial cells (not shown) were omitted from the reaction mixture.
Zdentifcation of paraquat induced free radical. It has
been previously shown by multiple i n v e s t i g a t o r ~ that
~ ’ ~ ~in~
addition to being a product of the reaction of DMPO and
hydroxyl radical, DMPO/.OH also can result from the
decomposition of the superoxide-derived spin adduct,
DMPO/*OOH.This process is more rapid in the presence
of some cellular s y ~ t e m s . *Thus,
~ . ~ ~the detection of DMPO/
.OH is not specific for hydroxyl radical, but rather could
arise simply from the presence of 0 - alone. To differentiate these two possibilities it is necessary to determine the
effect of various oxidant scavengers on the system. Hydroxyl
radical reacts rapidly with DMSO or ethanol to generate
methyl radical (.CH,) or a-hydroxyethyl radical
(CHOHCH,), respectively.2’ In an experimental system
in which the concentration of DMSO or ethanol exceed
that of DMPO, hydroxyl radical formation results in production of the DMPO spin adducts of C H , or CHOHCH,,
respectively, with a concomitant decrease in the expected
DMPO/.OH peak amplit~des.~~.’’
Accordingly, microcarrier bead adherent endothelial cells were exposed to
paraquat in the presence of DTPA, 100 mmol/L DMPO,
and 140 mmol/L DMSO or ethanol. The magnitude of
resulting DMPO/.OH peaks was unaffected by the presence of DMSO or ethanol and no new spin adducts were
detected (Fig 1E) suggesting that the DMPO/.OH species
detected resulted from decomposition of the superoxide
derived spin adduct rather than spin trapping of hydroxyl
radical. To provide further evidence in support of this
interpretation, experiments were repeated in which reac-
’’
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ENDOTHELIAL CELL OXYGEN RADICAL GENERATION
701
10 0
EC
spectrum above background was observed with addition of
H,O, to microcarrier beads that lacked endothelial cells but
which had been maintained in cell culture media and
washed analogous to the adherent endothelial cells or used
without washing. Furthermore, even addition of H,O, to
similar unwashed “sham” microcarrier bead preparations
or a 25%/75% (vol/vol) of the endothelial cell culture
media/HBSS solution containing DTPA and DMPO did
not yield an EPR signal above background. These data
eliminate the possibility that the hydroxyl radical detected
with addition of H,O, to the endothelial cells resulted from
the presence of iron associated with the microcarrier beads
or residual cell culture media rather than the endothelial
cells themselves.
Because endothelial cells would be unlikely to be exposed in vivo to a large bolus of H,O,, the effect of a
continuous flux of H,O, generated by the oxidation of
glucose by glucose oxidase was examined. Again, an EPR
spectrum comprised of catalase-inhibitable DMPO/-OH
was detected (Fig 2, D and E).
Previous studies suggested that iron associated with the
endothelial cells was responsible for catalyzing the formation of hydroxyl radical resulting from H,O, exposure?23”
Consistent with this hypothesis, preincubation of the endothelial cell suspension for 30 minutes with deferoxamine
B
EC + PQ + CAT
EC + PQ + SOD
2
EC + PQ + DMSO
n
h
Fig 1. Free radical species resulting from exposure of endothelial
cell monolayers (EC) t o paraquat (Pa). Conditions for each EPR
spectrum shown are as follows: (A) 25% (vol/vol) microcarrier bead
adherent endothelial cells suspended in the presence of 100 mmol/L
DMPO and 0.1 mmol/L DTPA. (E) Same conditions as (A) except that
the cells were exposed t o 1 mmol/L paraquat. (C) Paraquat-treated
endothelial cells as in (E) except that catalase (CAT, 500 U/mL) was
also present. (D) Paraquat-treated endothelial cells as in (E) except
that SOD (30 U/mL) was also present. (E) Paraquat-treated endothelial cells as in (E) except that DMSO (140 mmol/L) was also present
Identical results were obtained with the use of ethanol instead of
DMSO.
tion mixtures contained SOD (30 U/mL) or catalase (500
U/mL) during endothelial cell exposure to paraquat. SOD
eliminated DMPO/.OH formation, whereas catalase was
without effect (Fig 1, C and D), consistent with 0 -rather
than hydroxyl radical induced DMPO/.OH formation.
Hydrogen peroxide associated free radical formation with
endothelial cells. Hydrogen peroxide has been shown to
cause endothelial cell i n j ~ r y . ~ , ~The
’ ” ~mechanism of H,O,
toxicity has been postulated to involve hydroxyl radical
generated by the interaction of H,O, with cellular iron
Consequently, we sought evidence for the formation of hydroxyl radical after endothelial cell exposure to
H,O,. Microcarrier bead adherent endothelial cells were
suspended in HBSS containing DTPA and DMPO, after
which H,O, was added. The resulting EPR spectrum was
that of DMPO/.OH (Fig 2A) and was detectable at H,O,
concentrations 20.1 mmol/L. In contrast to results with
paraquat, catalase totally eliminated the spectrum (Fig 2B).
Ethanol decreased DMPO/.OH peak amplitudes with a
corresponding appearance of the DMPO/CHOHCH, spin
adduct (AN= 16.0 G, AH = 23.1 G; Fig 2C). These results
are consistent with spin trapping of hydroxyl radical. No
I
10Q
EC + H202
EC + H202+ CAT
B
EC + H202+ ETOH
C
EC + G/GO + CAT
E
Fig 2. Effect of endothelial cell exposure t o H,O,. Conditions for
each EPR spectrum shown are: (A) 25% (vol/vol) microcarrier bead
adherent endothelial cells (EC) suspended in 100 mmol/L DMPO and
0.1 mmol/L DTPA after the addition of 1 mmol/L H,O,. (E) H,O,treated endothelial cells as in (A) except that 500 U/mL catalase (CAT)
was also present. (C) H,O,-treated endothelial cells as in (A) except
that 140 mmol/L ethanol (ETOH) was also present. (D) Endothelial
cells exposed t o a continuous flux of H,O, generated by the reaction of
glucose and glucose oxidase. (E) Same conditions as in (D) except that
catalase was present. Note the location of the 6-line spectrum of
DMPO/CHOHCH, in tracing C is designated by the asterisk (*).
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BRITIGAN, ROEDER, AND SHASBY
702
(0.1 mmol/L) before the addition of H 2 0 2 eliminated
detectable hydroxyl radical formation (Fig 3B). No inhibition was noted if iron-saturated deferoxamine was used
(Fig 3C). It has previously been reported that deferoxamine
may have other effects on modulating hydroxyl radical
formation beyond simple iron c h e l a t i ~ n ? To
~ - ~help
~ eliminate such an explanation for our results, endothelial cells
that had been pretreated with deferoxamine were extensively washed in deferoxamine-free HBSS before the addition of H,02. Once again, no evidence of hydroxyl radical
generation was detected (Fig 3D).
Induction of hydro& radical by a superoxide generating
system. In contrast to the above system in which H20,was
the sole oxidant species to which endothelial cells were
exposed, the respiratory burst of phagocytes generates a
continuous flux of both .O,- and H20,.36The reaction of
xanthine oxidase and (hypo) xanthine yields similar prodUCts24.37.38and has been used as a convenient model system
for the products of stimulated phago~ytes.~’,~
Therefore,
microcarrier bead adherent endothelial cells were suspended in xanthine, DMPO, ethanol, and DTPA. EPR
spectra were then obtained after the initiation of continuous 0 - /H,O, production by the addition of xanthine
oxidase. The resulting EPR spectra were a composite of
three species: the superoxide-derived spin adduct, DMPO/
*OOH (A, = 14.3 G, A, = 11.7 G, A, = 1.3 G); DMPO/
.OH (A, = A, = 15.0 G); and DMPO/CHOHCH,
(A, = 16.0 G, A, = 23.1 G, Fig 4A). The detection of
DMPO/CHOHCH, as discussed above would be expected
I
100
EC + H202+ DF
EC + H202+ Washed DF
Fig 3. Effect of deferoxamine on hydroxyl radical generated by
H,O,-treated endothelial cells. Conditions for each of the EPR spectra
shown are: (A) 25% endothelial cells exposed t o 1mmol/L H,O, in the
presence of 100 mmol/L DMPO and 0.1 mmol/L DTPA. (B) Same
conditions as in (A) except that the endothelial cells were preincubated with 0.1 mmol/L deferoxamine (DF) for 30 minutes before the
addition of H,O,. (C) Same conditions as in (B) except that iron-loaded
deferoxamine (FeDF) was used. (D) Same conditions as in (B) except
that the endothelial cells were extensively washed t o remove the
deferoxamine before the addition of HO
, .,
10 G
EC + WXO
EC + XMO + CAT
EC + WXO + SOD
C
I
l
l
OOH OH CH,
Fig 4. Detection of free radicals generated after exposure of
/HZ02generated by the
endothelial cells t o a continuous flux of 0 reaction of xanthine oxidase and xanthine. Conditions for each of the
EPR spectra shown are: (A) Endothelial cells suspended in 100
mmol/L DMPO, 0.1 mmol/L DTPA, 140 mmol/L DMSO, and 2
mmol/L xanthine, after which 0.04 U/mL xanthine oxidase was
added. (B) Same as (A) except catalase (CAT, 500 U/mL) was present.
(C) Same as (A) except that SOD (30 U/mL) was present. High and low
field peaks corresponding t o the DMPO spin adducts DMPO/*OOH,
DMPO/.OH, and DMPO/.CH, are designated OOH, OH, and CH,
respectively.
as a consequence of hydroxyl radical spin trapping. Consistent with this, the presence of DMPO/-CHOHCH, was
inhibited by both catalase (Fig 4B) and SOD (Fig 4C). In
contrast, the other two spin adducts were inhibited by only
SOD (Fig 4, B and C ) , confirming that they arose by spin
trapping .O,-, the initial product of the xanthine/xanthine
oxidase reaction. As we have previously ob~erved,‘~
for
reasons that remain unclear, in some cases, the presence of
catalase appeared to increase the relative ratio of DMPO/
.OH:DMPO/.OOH resulting from the spin trapping of 0 (Fig 4, A v B).
Site of paraquat- and hydrogen peroxide-induced free radical formation. Chromium oxalate induces a concentrationdependent broadening of the EPR spectral lines resulting
from solutions of nitroxide free radical^.^' As the concentration of chromium oxalate increases a point is reached in
which the peaks are broadened to such an extent that they
become indistinguishable from background. For example, 5
mmol/L chromium oxalate essentially eliminated the EPR
spectrum resulting from DMPO/.OH (Fig 5, A and B),
previously generated via the reaction of H,O, and Fe2+.
Because chromium oxalate is able to penetrate cell membranes to only a limited extent, its addition to cellular free
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ENDOTHELIAL CELL OXYGEN RADICAL GENERATION
703
10 0
DMPO I 'OH
A
DMPO / 'OH + CrOx
B
m
EC + PQ
C
EC + PQ + CrOX
D
sure of endothelial cells to activatedphagocytes. Experimental evidence obtained by other investigators has suggested a
role for hydroxyl radical in phagocyte-mediated endothelial
cell injury.5-'The results described above with exposure to
H,O,, glucose/glucose oxidase, and xanthine/xanthine oxidase provide support for these earlier studies. To further
examine this hypothesis, additional spin trapping studies
were performed in which endothelial cells were incubated
in the presence of PMA-stimulated human neutrophils or
peripheral blood monocytes, DMPO, DTPA, and ethanol.
EPR spectra of the reaction mixture of endothelial cells
and monocytes was that of DMPO/.OH and DMPO/
CHOHCH, (Fig 6A). Addition of catalase inhibited predominantly DMPO/CHOHCH, (Fig 6B), whereas SOD
eliminated both species (Fig 6C). These results are consistent with previous data4' showing that the spin trapping of
.02derived from stimulated mononuclear phagocytes is
manifested predominantly as DMPO/.OH rather than
DMPO/.OOH due to cell-mediated enhancement of the
rate of conversion of DMPO/.OOH to DMPO/.OH. The
detection of catalase-inhibitable DMPO/CHOHCH, as
observed here is indicative of spin trapping of hydroxyl
radical. Because formation of hydroxyl radical after stimulation of human monocytes is not seen in the absence of an
exogenous iron catalyst," it is likely that hydroxyl radical
production resulted from the presence of catalytic iron
provided by the endothelial cells.
In contrast to the results with monocytes, the EPR
spectra resulting from the exposure of endothelial cells to
stimulated neutrophils yielded a predominance of
DMPO/.OOH and DMPO/-OH, with only scant detection
of DMPO/CHOHCH, (Fig 6D). All species were inhibited
by SOD but not catalase (Fig 6, E and F), indicating that
the spectrum resulted from the spin trapping of 0 -with
subsequent decomposition of the DMPO/.OOH spin adduct to other nitroxides as has been previously noted in
neutrophil-containing ~ y s t e m s .Once
~ ~ . ~again,
~
the presence
of catalase appeared to increase the peak amplitude ratio of
DMPO/.OH:DMPO/.OOH. Inclusion of the myeloperoxidase inhibitor azide in the neutrophil reaction mixture did
not alter the EPR spectrum. Experiments with chromium
oxalate to localize the site of free radical formation were
not attempted because it had previously been shown that
the concentration of chromium oxalate necessary to eliminate extracellular EPR signals markedly inhibits phagocyte
free radical formation.#
Can elastase, cathepsin G, P M , and/or the calcium
ionophore A23187 induce endothelial cell free radical formation? The neutrophil-derived proteases elastase and cathepsin G increase permeability of endothelial cell monolayers to a l b ~ m i n ' ~ .by
~ ' ,a~protease-independent
~
mechanism
that has been suggested to involve endothelial cell free
radical
Ph4A and the calcium ionophore
A23187 have also been reported to induce endothelial cell
.O; f~rmation.~'
However, when up to 60 p,g/mL of human
elastase or cathepsin G or the combination of A23187 (0.1
mg/L) and PMA (0.2 mmol/L) were added to microcarrier
bead adherent endothelial cells in HBSS containing Caz+
(1.67 mmol/L), M$' (4.1 mmol/L), DTPA, and DMPO, no
1
E
F
Fig 5. Localization of the cellular site of paraquat and H,O,induced free radical production using the EPR line-broadening agent
chromium oxalate (CrOx) to eliminate extracellular EPR signals.
Conditions for each EPR spectrum shown are: (A) A solution containing the DMPO/-OH spin adduct generated previously by the reaction
of H,Oz and Fez+in the presence of DMPO. (6) Same as in (A) except
that 5 mmol/L chromium oxalate was added to the solution. (C)
Endothelial cells treated with 1 mmol/L paraquat (PO) in the presence
of DMPO and DTPA. (D) Same as in (C) except the reaction mixture
also contained 5 mmol/L chromium oxalate. (E) Endothelial cells
treated with 1 mmol/L HzO, in the presence of DMPO and DTPA. (F)
Same as (E) except that 5 mmol/L chromium oxalate was also
present.
radical generating systems has been used as a means of
localizing the site of free radical formation to intracellular
or extracellular location^.^^ In the presence of chromium
oxalate, extracellularly located spin adducts become undetectable whereas the EPR spectrum of intracellular species
remains detectable.
Consequently, the effect of chromium oxalate on the
EPR spectrum resulting from the addition of paraquat or
H,O, to microcarrier bead adherent endothelial cells suspended in the presence of DMPO and DTPA was examined. Chromium oxalate (5 mmol/L) eliminated the
DMPO/.OH spectrum resulting from exposure to paraquat
(Fig 5, C and D). However, it had only minimal effect on
H,O,-induced DMPO/.OH peaks (Fig 5, E and F). These
data are consistent with paraquat-mediated induction of
extracellular (or in close proximity to the extracellular
space) 0 -production. However, hydrogen peroxide exposure appears to lead to intracellular hydroxyl radical formation.
Characteristics of free radical formation induced by a p o -
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BRITIGAN, ROEDER, AND SHASBY
704
negative spin trapping data, cyanide-insensitive0,consumption was not increased with exposure of endothelial cells to
elastase (Fig 7), cathepsin G (Fig 7), or A23187/PMA (not
shown). It is worth noting that in the experiments described
earlier, DMPO/.OH detection required endothelial cell
exposure to 0.1 mmol/L paraquat, a paraquat concentration that results in consumption of 1 nmol of OJmin by
cyanide treated endothelial cells (Fig 7). Thus, the spin
trapping system should detect 0 -production in excess of 1
nmol/min.
-
DISCUSSION
Y
EC + MC + SOD
EC+PMN+SOD
,
Fig 6. Free radicals generated after endothelial cell exposure to
stimulated phagocytes. Conditions for EPR spectra shown are: (A)
Endothelial cells incubated in the presence of PMA (100 ng/mL)
stimulated peripheral blood monocytes (MC, 5 x 1O6/mL) in the
presenceof 100mmol/LDMPO, 140mmol/LethanoI,andO.l mmol/L
DTPA. (B) Same as (A) except the reaction mixture contained 500
U/mL catalase (CAT). (C) Same as (A) except 30 U/mL SOD was
present. (D) Same as (A) except that PMA-stimulated neutrophils
(PMN) were substituted for monocytes. (E) Same as (D) except
catalase was present. (F) Same as (D) but with SOD added. Note the
receiver gain in scan (F) was twice that of (D) and (E). High and low
field peaks corresponding t o DMPO/.OOH, DMPO/.OH, and DMPO/
CHOHCH, are designated OOH, OH, and CH, respectively.
EPR spectrum above background was detected over 2 6 0
minutes of observation (data not shown).
Cellular .O,- production resulting from exposure to
xenobiotics such as paraquat consumes 0, which, in contrast to that resulting from normal cellular respiration, is
not inhibited by cyanide (Fig 7).17,18
Consistent with the
Spin trapping has become an increasingly popular technique for the investigation of the role of free radical species
in human biology." However, a significant limitation to the
application of spin trapping to in vitro cellular systems, such
as endothelial cell that exist as monolayers, is that spin
trapping is most easily performed with cell suspensions.
Consequently, we set out to develop a system in which
endothelial cell monolayers could be examined intact by
conventional spin trapping techniques.
Suspensions of microcarrier beads adherent porcine
pulmonary endothelial cells did not alter the EPR spectra
of the stable nitroxide TEMPO, indicating that such cell
preparations are amenable to study by conventional spin
trapping techniques. This was confirmed by exposing such
endothelial cell monolayers to paraquat. Incubation of
microcarrier bead adherent endothelial cells with paraquat
in the presence of DMPO yielded the hydroxyl radical spin
adduct of DMPO, DMPO/-OH. However, DMPO/.OH
formation was inhibited by SOD but not catalase or the
hydroxyl radical scavengers DMSO and ethanol. This indicates that the DMPO/.OH spectrum resulted from 0 generation via the well-re~ognized~"~~
decomposition of the
superoxide spin adduct DMPO/.OOH to DMPO/.OH
rather than spin trapping of hydroxyl radical. Similar
conclusions have been reported after paraquat treatment of
other cell ~ystems.~~.~'
In some4y-51
but not all ~ y s t e m s , ~iron
~ . ' ~ chelation and
other means of limiting the potential for hydroxyl radical
formation appear to limit paraquat-mediated cytoxicity. We
ae
1
Time
Fig 7. Induction of endothelial cell cyanide-insensitive 0, consumption. 0, consumption over time by 25% microcarrier bead adherent
endothelial cells in HBSS (Control) and in the presence of 1 mmol/L
NaCN (+CN). Also shown is the increase in cyanide-insensitive
respiration after the addition t o NaCN treated endothelial cells of 1
mmol/L paraquat (+PO, +CN), 60 pg/mL elastase (+EL, +CN), or 60
pg/mL cathepsin G (+CG, +CN). Note paraquat but not elastase
increasedthe rate of cyanide-insensitive 0, consumption indicative of
'0, - formation. Time elapsed for the peroid shown was 30 minutes.
From www.bloodjournal.org by guest on January 21, 2015. For personal use only.
ENDOTHELIAL CELL OXYGEN RADICAL GENERATION
705
did not find spin trapping evidence of hydroxyl radical
generation during endothelial cell exposure to paraquat.
However, our results do not exclude a role for this free
radical species at a cellular site where the microenvironment precluded its detection.
The cellular site at which oxidants are generated is also
likely to be of considerable importance in their relative
toxicity. Addition of the EPR line broadening agent chromium oxalate during paraquat treatment of endothelial
cells eliminated the expected DMPO/.OH spectrum. Because the concentration of chromium oxalate used did not
eliminate the EPR spectra induced by H,O, exposure, these
results cannot be attributed to intracellular penetration of
chromium oxalate. Rather, they complement earlier work
suggesting that .O,- production induced by paraquat occurs
predominantly at extracellular locations?o34 Presumably
this involves the intracellular reduction of paraquat to the
paraquat free radical, which subsequently diffuses outside
the cell where it is oxidized by ambient oxygen with
regeneration of paraquat and coincident formation of
.oz-
.20.48
Exposure of endothelial cells monolayers to H,O, results
in a decrease in the capacity of the monolayer to function as
a diffusion barrier. Incubation of suspensions of adherent
endothelial cells to either a bolus or a continuous flux of
H,O, +- 0 -resulted in the generation of catalaseinhibitable DMPO/-OH whose magnitude decreased in the
presence of DMSO or ethanol with resulting appearance of
DMPO/CH, and DMPO/CHOHCH,, respectively. These
data confirm other experimental dataG9that exposure of
endothelial cells to exogenous H,O, +- -0,-results in the
generation of hydroxyl radical. The failure of chromium
oxalate to inhibit much of the DMPO/.OH spectrum after
endothelial cell incubation with H,O, suggests that a
considerable portion of the hydroxyl radical is generated
intracellularly. Preincubation of the endothelial cells with
deferoxamine prevented DMPO/.OH formation, suggesting that hydroxyl radical is most likely generated through a
Fenton reaction catalyzed by an as yet unknown intracellular iron chelate(s). Alternatively, Beckman et a135 have
recently reported that the interaction of 0 with endothelial cell-derived peroxynitrite can also generate .OH. Although not applicable to results with exposure to only H,O,,
this mechanism could contribute .OH generation observed
with endothelial cell exposure to fluxes of .O,- .
In vivo phagocytic cells would constitute one of the
principal exogenous sources of H,O, and 0 - to which
endothelial cells would likely be exposed. When endothelial
cells were exposed to PMA-stimulated peripheral blood
monocytes spin trap evidence of a small amount of hydroxyl
radical formation was detected. We have previously shown
that monocytes by themselves do not have the capacity for
hydroxyl radical g e n e r a t i ~ n . ~Thus,
~ . ’ ~ it is likely that the
source of the iron catalyst in this case was again the
endothelial cell. Unfortunately, the location (intracellular v
extracellular) of the endothelial cell-derived iron could not
be determined. Elimination of monocyte-generated H,O,
before its interaction with the endothelial cell monolayer
prevented hydroxyl radical production. However, this effect
I
of catalase would be expected regardless of whether the
iron catalyst was located within or on the external surface of
the endothelial cell. Experiments with chromium oxalate to
eliminate EPR spectra from spin trapping of extracellularly
generated hydroxyl radical could not be performed due to
the fact that chromium oxalate inhibits the phagocyte
respiratory burst. Nevertheless, because monocytes effectively serve as a source of continuous H,O, flux it seems
most likely that the source of iron responsible for hydroxyl
radical formation is inside the endothelial cell, as is the case
with hydroxyl radical formed with endothelial cells treated
with H,O, or glucose/glucose oxidase.
In contrast to the monocyte results, a reaction mixture
composed of endothelial cells and PMA-stimulated neutrophils failed to yield spin trap evidence of hydroxyl radical.
These results were somewhat surprising given considerable
circumstantial evidence obtained by o t h e d 9 that damage
to endothelial cells by neutrophil-derived oxidants is mediated by hydroxyl radical. There appear to be a number of
possible explanations for this apparent discrepancy. Release of myeloperoxidase and lactoferrin during the neutrophil respiratory burst have the capacity to inhibit the
generation of hydroxyl radical resulting from neutrophil
stimulation in the presence of exogenous catalytic iron
c h e l a t e ~ . ~In
~ -the
~ ’ present work, it seems possible that the
release of these two granule components decreased the
magnitude of hydroxyl radical generation to a level below
the limits of detection of the spin trapping system used.
However, inclusion of the myeloperoxidase inhibitor azide
failed to result in spin trap evidence of hydroxyl radical
formation. Because neutrophils contain lactoferrin and
monocytes do not, lactoferrin release would seem on the
surface to be a highly likely explanation. However, as
described above, the iron catalyst responsible for hydroxyl
radical generation on endothelial cell exposure to sources
of H,O, appears to be located intracellularly. Therefore, it
seems unlikely that lactoferrin that would remain outside
the endothelial cell could influence the result of an intracellular reaction. Alternatively, the stability of hydroxyl radicalderived DMPO spin adducts decreases in the presence of
PMA-stimulated n e u t r o p h i l ~ . ~ ’Thus,
~ ~ ~ ~ hydroxyl
’~
radical
generation could have occurred during the exposure of
endothelial cells to stimulated neutrophils, but the relative
instability of the resulting spin adducts precluded their
detection. Two new means of spin trapping hydroxyl radical
which appear to be relatively immune to this phenomenon
have recently been de~cribed’~~’~
and could help clarify the
situation.
Neutrophils possess a variety of granule proteases suggested to contribute to inflammatory tissue injury. Recently, several l a b o r a t ~ r i e s , ’ ~including
, ~ , ~ ’ our 0wn,[email protected]
have
shown that exposure of endothelial cell monolayers to two
different neutrophil granule components, elastase and cathepsin G, leads to cell injury. The ability of elastase and
cathepsin G to produce these effects appears to be independent of their enzymatic activity but related to their cationic
proper tie^'^.^' and mediated through the release of intracellular calcium.62It has been suggested that the nonenzymatic
injury of endothelial cells by elastase and/or cathepsin G
From www.bloodjournal.org by guest on January 21, 2015. For personal use only.
BRITIGAN, ROEDER, AND SHASBY
706
may result from the induction of cellular free radical
formation.46The results of the present study do not support
that hypothesis. As measured by both cyanide-insensitive
0, consumption and spin trapping, we detected no evidence
of 0 - generation (sensitivity 1 nmol/min) after exposure of microcarrier bead adherent endothelial cells to up
to 60 pg/mL elastase or cathepsin G . Similarly, in contrast
to an earlier report: no evidence of free radical formation
was observed after exposure of a similar concentration of
endothelial cells to PMA or A23187. This latter discrepancy
could relate to one or more of a variety of differences
between the earlier work of work of Matsubara and Ziff4’
and our own including the site of cell harvest and the
species used. In addition, the possibility remains that the
human umbilical vein preparations in the earlier study
could have been contaminated with tissue macrophages.
In summary, using a system that allows detection of free
radical formation by intact endothelial cell monolayers
using conventional spin trapping techniques, we have confirmed previous
that exposure of endothelial
cells to the redox active compound paraquat induces the
generation of .02at, or in close proximity to, the extracellular space. In contrast, exposure of these cells to bolus and
-
or continuous fluxes of H,O, 2
results in the intracellular generation of hydroxyl radical, catalyzed by cellular iron
chelates. These results strongly support the hyp~thesis~~~*”’
that phagocyte-mediated injury to endothelial cells may
occur as a consequence of intracellular hydroxyl radical
formation. Studies with stimulated monocytes appear to be
consistent with this hypothesis, although the exact site of
hydroxyl radical generation could not be determined. However, we were surprisingly unable to detect the generation
of hydroxyl radical during the exposure of the same endothelial cells to stimulated neutrophils. Further work is required
to delineate the reason(s) for this unexpected result.
Nevertheless, the ability to quantitate and localize free
radical formation in systems containing intact endothelial
cell monolayers will likely provided new insights into the
mechanism of oxidative injury to these cells.
ACKNOWLEDGMENT
We acknowledge the technical assistance of Kathy Lindsley in
maintaining the endothelial cell cultures, and the help of Kathy
Schmuecker and Naomi Erickson with preparation of the manuscript.
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1992 79: 699-707
Insight into the nature and site of oxygen-centered free radical
generation by endothelial cell monolayers using a novel spin trapping
technique
BE Britigan, TL Roeder and DM Shasby
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