First Department of Medicine, Tokyo Women's Medical University, Shinjuku-ku, Tokyo 162-8666, Japan
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ABSTRACT |
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Several serine proteases are directly cytotoxic. We investigated whether the cytotoxic effects of proteases are associated with increased levels of reactive oxygen species (ROS) in cells. We found that treatment of lung fibroblasts or bronchial epithelial cells with relatively high concentrations (0.1-100 U/ml) of neutrophil elastase, trypsin, and Pronase increased ROS levels in the mitochondria and cytoplasm. The protease-induced increase in ROS was associated with oxidative cellular injury as determined by generation of 8-hydroxy-2'-deoxyguanosine and malonaldehyde plus 4-hydroxyalkenal. The protease-induced increase in ROS was not merely due to cell detachment because the proteases also caused an increase in ROS in suspended cells, which precluded attachment to the extracellular matrix. The protease-induced increase in ROS appears to contribute to cytotoxicity because cell death induced by proteases was attenuated by treatment with catalase, a decomposer of H2O2, and accelerated by treatment with aminotriazole, a catalase inhibitor. These results suggest that several proteases increase oxidative stress, indicating a direct interaction between proteases and ROS in mediating cytotoxicity.
oxidants; reactive oxygen species; elastase; trypsin; Pronase
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INTRODUCTION |
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BECAUSE OF THEIR ABILITY to cause extensive tissue destruction, proteases are thought to be involved in the mechanism of a variety of lung diseases. Many proteases have been demonstrated to degrade extracellular matrix (ECM) proteins and other substrates such as complement components, proteinase inhibitors, and hemostatic proteins. In particular, the proteolytic degradation of ECM proteins has been established as a mechanism of tissue destructive disorders including pulmonary emphysema, acute respiratory distress syndrome, bronchiectasis, and pulmonary fibrosis (13, 31).
Besides their capacity to degrade noncellular components, serine proteases such as neutrophil elastase, trypsin, and bacterial Pronase are known to mediate direct cytotoxicity (2, 20, 36, 37). They primarily cause cytolysis by inducing necrosis, and some of them have been shown to induce apoptosis of endothelial cells (37), keratinocytes (18), and neutrophils (32). The mechanism of the cytotoxicity of proteases, however, is not fully understood.
Possible mechanisms of protease cytotoxicity include reactive oxygen species (ROS). ROS are produced in response to a variety of cytotoxic stimuli and have been hypothesized to be important mediators of cell death, whether by necrosis or apoptosis (12, 23). However, it is unknown whether ROS are involved in the mechanism of cytotoxicity induced by proteases.
Proteases and ROS are frequent companions at sites of inflammation, and previous studies have pointed to interactions between them. For example, ROS convert the inactive form of procollagenase to its active form (34) and protect proteases by inactivating proteinase inhibitors (6, 21, 33, 35), whereas neutrophil-derived neutral proteases stimulate ROS release from monocytes (28). Although these findings suggest that proteases and ROS may interact with each other under certain circumstances, the role of ROS in protease-induced cytotoxicity is unknown.
In this study, we investigated whether the cytotoxic effects of proteases are associated with increased intracellular levels of ROS within cells.
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METHODS |
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Materials. Cell culture reagents were obtained from GIBCO BRL (Life Technologies, Gaithersburg, MD), unless otherwise stated. 6-Carboxy-2',7'-dichlorodihydrofluorescein diacetate, di(acetoxymethyl ester) (CDCF) and reduced MitoTracker Red probe (CM-H2XRos) were obtained from Molecular Probes (Eugene, OR). Human neutrophil elastase, trypsin, Pronase, catalase, 3-amino-1,2,4-triazole, o-phenylenediamine, horseradish peroxidase, poly(2-hydroxyethyl methacrylate) (polyHEMA), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), propidium iodide, calphostin C, H-89, genistein, herbimycin A, sodium orthovanadate, antimycin A, and pertussis toxin were purchased from Sigma (St. Louis, MO). ONO-6818, a specific inhibitor of neutrophil elastase, was a kind gift from Ono Pharmaceuticals (Osaka, Japan).
Cell culture and protease treatment. Normal human fetal lung fibroblasts (IMR-90, Clonetics, San Diego, CA) were maintained in Dulbecco's modification of Eagle's MEM (DMEM) containing 10% FCS, 100 U/ml of penicillin, and 100 µg/ml of streptomycin. Cells were passaged weekly, and cells from passages 8 to 20 were used in the experiments. Normal human bronchial epithelial cells were prepared as described (25) and maintained in bronchial epithelial cell basal medium (modified LHC-9; Clonetics) containing 15 µg/ml of bovine pituitary extract, 0.25 ng/ml of hydrocortisone, 0.25 ng/ml of epidermal growth factor, 0.25 ng/ml of epinephrine, 5 µg/ml of transferrin, 2.5 ng/ml of insulin, 0.05 ng/ml of retinoic acid, 0.25 mg/ml of bovine serum albumin, 100 U/ml of penicillin, and 100 µg/ml of streptomycin. Cells from passages 2 to 4 were used in the experiments. Human lymphocytes were isolated from venous blood by dextran sedimentation and centrifugation on a Histopaque gradient (density 1.078). Before being used in the experiments, all cells were washed with PBS twice and reincubated with serum-free DMEM.
Determination of intracellular ROS levels. Intracellular ROS levels were measured with CDCF (1, 10). Briefly, the cells were loaded for 30 min with 5 µM CDCF in a 96-well tissue culture plate or a 6-well tissue culture plate. They were washed with PBS and then exposed to proteases in serum-free DMEM. For mechanical detachment, the cells in a six-well plate were detached with a rubber scraper. Fluorescence was monitored on a Cytofluor II multiplate fluorometer (Perceptive Biosystems, Framingham, MA) by using an excitation wavelength of 485 nm and an emission wavelength of 530 nm.
Determination of mitochondrial ROS levels. Mitochondrial ROS levels were determined by using CM-H2XRos (7, 10). Briefly, the cells were incubated for 30 min with 250 nM CM-H2XRos in a 96-well tissue culture plate and then exposed to proteases in serum-free DMEM. Fluorescence was monitored by excitation at 530 nm and emission at 590 nm.
Determination of H2O2 levels.
After the cells were treated with neutrophil elastase (100 U/ml) in
serum-free DMEM for 3 h, conditioned medium with 5 × 104 M ONO-6818 was added to inhibit the resultant enzyme
activity. H2O2 levels in the conditioned medium
were measured as described (4). Briefly, 50 µl of the
medium were combined with 75 µl of 100 mM Tris · HCl buffer,
pH 8.0, containing 16 mM o-phenylenediamine and 1 U/ml of
horseradish peroxidase in a 96-well microplate and incubated for 30 min
at 37°C. The reaction was quenched with 3 N sulfuric acid, and
absorbance was measured at 490 nm, with 690 nm as a reference
wavelength. H2O2 levels were estimated by
comparison with values on a reference curve generated with known
amounts of H2O2. We also determined whether
cells degrade H2O2 added externally to the
medium. Briefly, sufficient H2O2 was added to
100-mm dishes containing cells and serum-free DMEM to produce a final
concentration of 20 µM H2O2.
H2O2 levels in the medium were monitored before and after the addition of H2O2 at 5- to 10-min
intervals for 40 min.
Assay for DNA and lipid oxidation. The cells were exposed to neutrophil elastase (10 or 100 U/ml) for 3 h, and the resultant enzyme activities were inhibited as above. 8-Hydroxy-2'-deoxyguanosine (8-OHdG) levels in the conditioned medium were measured with an ELISA kit (Japan Institute for the Control of Aging, Shizuoka, Japan). Malonaldehyde (MDA) plus 4-hydroxyalkenal levels in the cell extracts were measured by using a colorimetric assay kit (BIOXYTECH LPO-586, OXIS International, Portland, OR) according to the manufacturer's instructions. The assay is based on the reaction between a chromogenic reagent, N-methyl-2-phenylindole, and MDA plus 4-hydroxyalkenal to yield a stable chromophore, with maximal absorbance at 586 nm.
Cell viability assay. The MTT assay, which estimates the number of viable cells, was performed as described (27). MTT (0.5 mg/ml) was added to the cells in serum-free DMEM in a 96-well culture plate during the final 1 h of incubation with the proteases. Cell viability was also determined by quantitating propidium iodide uptake as a measure of cell death (14). Propidium iodide (20 µg/ml) was added to the cells in a 96-well culture plate, and fluorescence was read 10 min later, with excitation at 530 nm and emission at 620 nm. The cells were lysed by a cycle of freezing and thawing, and fluorescence was read to estimate cell numbers. The percent uptake of propidium iodide was calculated according to the formula (fluorescence before freezing and thawing/fluorescence after freezing and thawing) × 100.
Statistical analysis. Data are presented as means ± SE. Comparisons were made with Student's t-test or ANOVA with Fisher's protected least significant difference as a post hoc analysis test. A value of P < 0.05 was accepted as significant.
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RESULTS |
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As observed previously (2, 20, 36, 37) and described again below, incubation with several serine proteases, i.e., neutrophil elastase, trypsin, and Streptomyces griseus Pronase, which is a mixture of protease A, protease B, and a trypsinlike enzyme, resulted in detachment and death of lung fibroblasts and bronchial epithelial cells.
We investigated whether these proteases increase intracellular ROS by
monitoring their levels with the fluorochrome CDCF. As demonstrated in
Fig. 1, ROS accumulated within cells
maintained in medium alone, suggesting that ROS are produced as
by-products of aerobic respiration as previously described
(3). When fibroblasts were exposed to neutrophil elastase
(1-100 U/ml; Fig. 1A), trypsin (0.1-10 U/ml; Fig.
1B), or Pronase E (0.1 or 10 U/ml; Fig. 1C), the
levels of CDCF fluorescence rose twofold or more compared with those in
cells maintained in medium alone. The effect of proteases on ROS levels
was not cell-type specific because bronchial epithelial cells also
showed increased levels of CDCF after exposure to neutrophil elastase
(Fig. 1D). We also determined whether these proteases
directly oxidize CDCF. Interestingly, we found that Pronase E, but not
neutrophil elastase or trypsin, directly oxidized CDCF in the absence
of cells (Fig. 1E). Although we removed CDCF from the
culture medium by rinsing the cells twice with PBS before exposure to
proteases, CDCF that leaked out of cells during exposure to Pronase E
may have been directly oxidized by Pronase E. The rise in ROS levels
after exposure to proteases appears to require their proteolytic
activity because ONO-6818, a specific inhibitor of neutrophil elastase,
abolished the elastase-induced rise in CDCF fluorescence (Fig.
2A). ONO-6818 did not inhibit
a rise in CDCF fluorescence induced by H2O2
(Fig. 2B), indicating that ONO-6818 does not directly affect
H2O2 or CDCF. CDCF fluorescence reflects various ROS including intracellular H2O2
(10, 11), which passes freely through the cell membrane
and diffuses readily out of cells. We therefore measured
H2O2 concentrations in conditioned medium with
a horseradish peroxidase assay with the substrate
o-phenylenediamine. We found that neutrophil elastase did
not directly oxidize o-phenylenediamine in the absence of
cells (data not shown). As demonstrated in Fig. 3A, incubation of fibroblasts
with 100 U/ml of neutrophil elastase in serum-free culture medium for
3 h produced a twofold increase in extracellular
H2O2 level. We also determined whether cells, which are thought to be equipped with various antioxidants, are capable
of decomposing H2O2 added externally to the
culture medium. As demonstrated in Fig. 3B, the addition of
20 µM H2O2 to dishes containing fibroblasts
bathed in serum-free culture medium produced a step increase in
H2O2 level that was sustained for at least 40 min, suggesting that cells are insufficient to decompose
H2O2. These results indicate that serine
proteases, including neutrophil elastase, trypsin, and bacterial
Pronase, increase ROS levels in cells.
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We next measured mitochondrial ROS levels in cells exposed to
neutrophil elastase. Fibroblasts and bronchial epithelial cells were
loaded with CM-H2XRos, a nonfluorescent compound that
enters cells where it can be oxidized at various sites including the mitochondria and then be sequestrated into the mitochondria by virtue
of its cationic state. As expected, incubation of fibroblasts with
antimycin A, a mitochondrial ROS generator, increased
CM-H2XRos fluorescence (Fig.
4A). Incubation with
neutrophil elastase resulted in two- to threefold increases in
CM-H2XRos fluorescence in fibroblasts (Fig. 4B)
and bronchial epithelial cells (Fig. 4C), and treatment with
rotenone, a specific inhibitor of mitochondrial electron transport, led
to significant inhibition in the rise of CDCF and CM-H2XRos
fluorescence in cells exposed to neutrophil elastase (Fig. 4,
D and E). Neutrophil elastase did not increase
CM-H2XRos fluorescence in the absence of cells (data not
shown). These results suggest that the predominant source of ROS in
protease-exposed cells is localized within the mitochondria, although
some enzymatic mechanisms such as cyclooxygenase, lipoxygenase,
monoamine oxidase, xanthine oxidase, NADPH oxidase, and nitric oxide
synthase may also be the source of ROS.
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Recent evidence (16) suggests that cell detachment from
culture plates increases intracellular levels of ROS. We therefore investigated whether the protease-induced rise in ROS levels was related to cell detachment. The results indicated that the
protease-induced rise in ROS levels was not merely due to cell
detachment from the ECM because 1) neutrophil elastase
increased ROS levels in fibroblasts suspended in polyHEMA-coated plates
that preclude cell attachment (Fig.
5A) and in lymphocytes, which
do not attach to culture plates (Fig. 5B), and
2) much more ROS were produced in response to neutrophil
elastase than to mechanical detachment of the cells (Fig.
5C).
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To investigate whether specific signaling pathways are involved in the
protease-induced rise in ROS levels, we evaluated the effect of several
inhibitors on the rise in ROS. The results showed that the rise in CDCF
fluorescence in fibroblasts exposed to neutrophil elastase (100 U/ml)
was not inhibited by prior or simultaneous exposure to the protein
kinase C inhibitor calphostin C (106 M), the protein
kinase A inhibitor H-89 (10
6 M), the broad spectrum
tyrosine kinase inhibitors genistein (10
5 M) and
herbimycin A (10
6 M), the tyrosine phosphatase inhibitor
sodium orthovanadate (10
4 M), or the Gi
protein blocker pertussis toxin (400 ng/ml) (data not shown).
Next, to determine whether the protease-induced rise in ROS causes
oxidative injury to cells, we measured the level of 8-OHdG as an
indicator of DNA oxidation (26), and the level of MDA plus
4-hydroxyalkenal as an indicator of lipid peroxidation
(8). As demonstrated in Fig.
6, incubation with neutrophil elastase resulted in two- to fourfold increases in the levels of 8-OHdG and MDA
plus 4-hydroxyalkenal, suggesting that elastase induces oxidative
injury of cells.
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Finally, we sought to determine whether the changes in intracellular
ROS levels contribute to the cytotoxicity by proteases. As demonstrated
in Fig. 7,
A-C, catalase, a decomposer of
H2O2, significantly inhibited the death of
fibroblasts as determined by both the MTT assay and the propidium
iodide exclusion test. Because H2O2 is freely
permeant in the cell membrane, and, therefore, it readily diffuses out
of cells if generated intracellularly, the reduction of extracellular
H2O2 by catalase is expected to decrease
intracellular H2O2 (1). To further
corroborate the role of ROS in protease-mediated cytotoxicity, we
evaluated the effect of aminotriazole, a catalase inhibitor, on cell
death. The results showed that prior and simultaneous exposure to
aminotriazole produced a small but significant rise in the rate of cell
death induced by Pronase (Fig. 7D). Aminotriazole did not
induce cell death in the absence of Pronase E (Fig. 7D). The
above findings suggest that ROS are, at least in part, responsible for
protease-mediated cytotoxicity.
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DISCUSSION |
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Growing evidence suggests that production of ROS is triggered by exposure to a variety of stressful and cytotoxic stimuli. In this study, we investigated the effect of several serine proteases on ROS levels of lung fibroblasts and bronchial epithelial cells. The results demonstrated that relatively high concentrations of neutrophil elastase, trypsin, and Pronase induce a rise in ROS, including H2O2. The source of the ROS appears to be the mitochondria, and the contribution of ROS to cytotoxicity has been demonstrated. To our knowledge, this study is the first to demonstrate that proteases directly increase the intracellular levels of ROS in nonphagocytic cells.
In this study, we used CDCF to monitor intracellular ROS levels. A recent study (11) demonstrated that CDCF, which was previously thought to be a specific indicator of H2O2, detects a broad range of ROS that are increased during intracellular oxidative stress. Thus the protease-induced increase in CDCF fluorescence observed in this study may reflect intracellular production of various oxidants including H2O2 and other ROS.
It is currently unknown how proteases increase the level of ROS. Many
mechanisms are possible and include detachment of the cells from the
culture plates, degradation of the ECM, activation of a
proteinase-activated receptor or an ion channel, an increase in
membrane permeability, and loss of intracellular antioxidants. A recent
report (16) noted that detachment of endothelial cells from the ECM led to a rapid rise in the intracellular level of ROS.
Another report (15) demonstrated that disruption of the actin cytoskeleton by an antibody to
5
1-integrin led to a change in cell shape
and subsequent generation of ROS in fibroblasts. However, the
protease-induced rise in ROS observed in our study was not merely due
to cell detachment because a rise in ROS induced by proteases was
observed in fibroblasts maintained in suspended conditions that
preclude cell attachment to culture plates and in lymphocytes, which do
not adhere to plates. This view is further supported by the observation
that the rise in ROS induced by proteases was much greater than the
rise induced by mechanical detachment of cells.
Alternatively, proteases may have activated specific signaling pathways either to increase ROS production or to decrease antioxidant defense levels. In our study, the protease-induced rise in ROS was unaffected by pharmacological inhibition of tyrosine kinases, tyrosine phosphatases, protein kinase C, protein kinase A, and Gi protein. However, proteases may have activated other signaling pathways such as Rac1 and ceramide signaling, both of which have recently been shown to regulate ROS levels in nonphagocytic cells (9, 15, 30). It is well established that serine proteases, including trypsin and thrombin, can stimulate cell proliferation through activation of protease-activated receptors and G proteins (5, 19). However, it is unknown whether protease-activated receptors are involved in ROS production and cell death induced by proteases. Thus whether proteases stimulate specific signaling to increase ROS levels was not confirmed in this study.
Although our results confirmed that neutrophil elastase, trypsin, and Pronase are cytotoxic, we have not yet investigated whether these proteases induce apoptosis or necrosis. Recent reports have documented that serine proteases such as neutrophil elastase and proteinase 3 are capable of inducing apoptosis in endothelial cells (37), keratinocytes (18), and neutrophils (32). Thus the protective effect of catalase on protease cytotoxicity observed in the present study may have been due to inhibition of apoptosis, and the contribution of ROS to apoptosis and necrosis in protease-exposed cells is the focus of ongoing investigations.
Proteases and ROS are frequent companions at sites of inflammation and
have been proposed as important mediators of tissue injury. Although
previous work (21, 35) has clearly established additive injurious effects of proteases and ROS during inflammation, our finding that proteases directly increase ROS points to a direct link between proteases and ROS in mediating cytotoxicity. This mechanism may be involved in a variety of inflammatory disorders, including pulmonary emphysema and acute respiratory distress syndrome, in which proteases and ROS are believed to act in concert to cause lung
injury (17, 22, 29). Besides their injurious effects, ROS
have been proposed to serve as signaling and messenger molecules involved in inflammatory processes. For example, ROS, including H2O2, regulate the activation of nuclear
factor-B and the production of proinflammatory cytokines such as
interleukins 6 and 8 and tumor necrosis factor-
(24).
Thus ROS released by protease-injured cells may also serve to propagate
inflammatory reactions by stimulating immune effector cells in the microenvironment.
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ACKNOWLEDGEMENTS |
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This work was supported by Grant-in Aid for Scientific Research 12670580 from the Ministry of Education, Science, and Culture, Japan.
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FOOTNOTES |
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Address for reprint requests and other correspondence: K. Aoshiba, First Dept. of Medicine, Tokyo Women's Medical Univ., 8-1 Kawada-cho, Shinjuku-ku, Tokyo 162-8666, Japan (E-mail: kaoshiba{at}chi.twmu.ac.jp).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 5 September 2000; accepted in final form 30 April 2001.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Aoshiba, K,
Yasui S,
Nishimura K,
and
Nagai A.
Thiol depletion induces apoptosis in cultured lung fibroblasts.
Am J Respir Cell Mol Biol
21:
54-64,
1999
2.
Ballieux, BEPB,
Hiemstra PS,
Klar-Mohamad N,
Hagen EC,
van Es LA,
van der Woude FJ,
and
Daha MR.
Detachment and cytolysis of human endothelial cells by proteinase 3.
Eur J Immunol
24:
3211-3215,
1994[ISI][Medline].
3.
Boveris, A,
and
Cadenas E.
Cellular sources and steady-state levels of reactive oxygen species.
In: Oxygen, Gene Expression, and Cellular Function, edited by Clerch LB,
and Massaro DJ.. New York: Dekker, 1997, vol. 105, p. 1-25. (Lung Biol Health Dis Ser)
4.
De Bleser, PJ,
Xu G,
Rombouts K,
Rogiers V,
and
Greerts A.
Glutathione levels discriminate between oxidative stress and transforming growth factor-signaling in activated rat hepatic stellate cells.
J Biol Chem
274:
33881-33887,
1999
5.
Déry, O,
Corvera CU,
Steinhoff M,
and
Bunnett NW.
Proteinase-activated receptors: novel mechanisms of signaling by serine proteases.
Am J Physiol Cell Physiol
274:
C1429-C1452,
1998
6.
Desrochers, PE,
and
Weiss SJ.
Proteolytic inactivation of alpha-1-proteinase inhibitor by a neutrophil metalloproteinase.
J Clin Invest
81:
1646-1650,
1988[ISI][Medline].
7.
Esposti, MD,
Hatzinisiriou I,
McLennan H,
and
Ralph S.
Bcl-2 and mitochondrial oxygen radicals.
J Biol Chem
274:
29831-29837,
1999
8.
Esterbauer, H,
Schaur RJ,
and
Zollner H.
Chemistry and biochemistry of 4-hydroxynonenal, malonaldehyde and related aldehydes.
Free Radic Biol Med
11:
81-128,
1991[ISI][Medline].
9.
García-Ruiz, C,
Colell A,
Marí M,
Morales A,
and
Fernández-Checa JC.
Direct effect of ceramide on the mitochondrial electron transport chain leads to generation of reactive oxygen species.
J Biol Chem
272:
11369-11377,
1997
10.
Haugland, RP.
Handbook of Fluorescent Probes and Research Chemicals (6th ed.). Eugene, OR: Molecular Probes, 1996.
11.
Hempel, SL,
Buettner GR,
O'Malley YQ,
Wessels DA,
and
Flaherty DM.
Dihydrofluorescein diacetate is superior for detecting intracellular oxidants: comparison with 2',7'-dichlorodihydrofluorescein diacetate, 5(and 6)-carboxy-2',7'-dichlorodihydrofluorescein diacetate, and dihydrorhodamine 123.
Free Radic Biol Med
27:
146-159,
1999[ISI][Medline].
12.
Jacobson, MD.
Reactive oxygen species and programmed cell death.
Trends Biochem Sci
21:
83-86,
1996[ISI][Medline].
13.
Janoff, A.
Elastase and emphysema: current assessment of the protease-antiprotease hypothesis.
Am Rev Respir Dis
132:
417-433,
1985[ISI][Medline].
14.
Kane, DJ,
Sarafian TA,
Anton R,
Hahn H,
Gralla EB,
Valentin JS,
Örd T,
and
Bredesen TE.
Bcl-2 inhibition of neural death: decreased generation of reactive oxygen species.
Science
262:
1274-1277,
1993[ISI][Medline].
15.
Kheradmand, F,
Werner E,
Tremble P,
Symons M,
and
Werb Z.
Role of Rac1 and oxygen radicals in collagenase-1 expression induced by cell shape change.
Science
280:
898-902,
1998
16.
Li, AE,
Ito H,
Rovira II,
Kim K,
Takeda K,
Yu Z,
Ferrans VJ,
and
Finkel T.
A role for reactive oxygen species in endothelial cell anoikisis.
Circ Res
85:
304-310,
1999
17.
MacNee, W,
and
Rahman I.
Oxidants and antioxidants as therapeutic targets in chronic obstructive pulmonary disease.
Am J Respir Crit Care Med
160:
s58-s65,
1999[ISI][Medline].
18.
Marthinuss, J,
Andrade-Gordon P,
and
Seiberg M.
A secreted serine protease can induce apoptosis in Pam keratinocytes.
Cell Growth Differ
6:
807-816,
1995[Abstract].
19.
Miyata, S,
Koshikawa N,
Yasumitsu H,
and
Miyazaki K.
Trypsin stimulates integrin 5
1-dependent adhesion to fibronectin and proliferation of human gastric carcinoma cells through activation of proteinase-activated receptor-2.
J Biol Chem
275:
4592-4598,
2000
20.
Okrent, DG,
Lichtenstein AK,
and
Ganz T.
Direct cytotoxicity of polymorphonuclear leukocyte granule proteins to human lung-derived cells and endothelial cells.
Am Rev Respir Dis
141:
179-185,
1990[ISI][Medline].
21.
Ossanna, PJ,
Test ST,
Matheson NR,
Regiani S,
and
Weiss SJ.
Oxidative regulation of neutrophil elastase-alpha-1 proteinase inhibitor interactions.
J Clin Invest
77:
1939-1951,
1986[ISI][Medline].
22.
Petty, TL.
Protease mechanisms in the pathogenesis of acute lung injury.
Ann NY Acad Sci
624:
267-277,
1991[ISI][Medline].
23.
Polyak, K,
Xia Y,
Zweier JL,
Kinzler KW,
and
Vogelstein B.
A model for p53-induced apoptosis.
Nature
389:
300-305,
1997[ISI][Medline].
24.
Remick, DG,
Villarete L,
and
DeForge LE.
Oxidant regulation of cytokine gene expression.
In: Oxygen, Gene Expression and Cellular Function, edited by Clerch LB,
and Massaro DJ.. New York: Dekker, 1997, vol. 105, p. 243-278. (Lung Biol Health Dis Ser)
25.
Romberger, DJ,
Heires P,
Rennard SI,
and
Wyatt TA.
-Adrenergic agonist modulation of monocyte adhesion to airway epithelial cells in vitro.
Am J Physiol Lung Cell Mol Physiol
278:
L139-L147,
2000
26.
Shigenaga, MK,
and
Ames BN.
Assays for 8-hydroxy-2'-deoxyguanosine, a biomarker of in vivo oxidative DNA damage.
Free Radic Biol Med
10:
211-216,
1991[ISI][Medline].
27.
Skehan, P.
Assays of cell growth and cytotoxicity.
In: Cell Growth and Apoptosis: A Practical Approach, edited by Studzinski GP.. New York: Oxford University Press, 1996, p. 169-192.
28.
Speer, CP,
Pabst MJ,
Hedgaard HB,
Rest RF,
and
Jonston RB, Jr.
Enhanced release of oxygen metabolites by monocyte-derived macrophages exposed to proteolytic enzymes: activity of neutrophil elastase and cathepsin G.
J Immunol
133:
2151-2156,
1984
29.
Stockley, RA.
Neutrophils and protease/antiprotease imbalance.
Am J Respir Crit Care Med
160:
s49-s52,
1999[ISI][Medline].
30.
Sundaresan, M,
Yu Z,
Ferrans VJ,
Sulciner DJ,
Gutkind JS,
Irani K,
Goldshmidt-Clermont PJ,
and
Finkel T.
Regulation of reactive-oxygen-species generation in fibroblasts by Rac1.
Biochem J
318:
379-382,
1996[ISI][Medline].
31.
Tetley, TD.
Proteinase imbalance: its role in lung disease.
Thorax
48:
560-565,
1993[Abstract].
32.
Trevani, AS,
Andonegui G,
Giordano M,
Nociari M,
Fontan P,
Dran G,
and
Geffner JR.
Neutrophil apoptosis induced by proteolytic enzymes.
Lab Invest
74:
711-721,
1996[ISI][Medline].
33.
Vissers, MCM,
George PM,
Bathurst IC,
Brennan SO,
and
Winterbourn CC.
Cleavage and inactivation of 1-antitrypsin by metalloproteinases released from neutrophils.
J Clin Invest
82:
706-711,
1988[ISI][Medline].
34.
Weiss, SJ,
Peppin G,
Ortiz X,
Ragsdale C,
and
Test ST.
Oxidative autoactivation of latent collagenase by human neutrophils.
Science
227:
747-749,
1985[ISI][Medline].
35.
Weiss, SJ,
and
Regiani S.
Neutrophils degrade subendothelial matrices in the presence of alpha-1-antiproteinase inhibitor: cooperative use of lysosomal proteinases and oxygen metabolites.
J Clin Invest
73:
1297-1303,
1984[ISI][Medline].
36.
Westlin, WF,
and
Gimbronem MA, Jr.
Neutrophil-mediated damage to human vascular endothelium: role of cytokine activation.
Am J Pathol
142:
117-128,
1993[Abstract].
37.
Yang, JJ,
Kettritz R,
Falk RJ,
Jennette JC,
and
Gaido ML.
Apoptosis of endothelial cells induced by the neutrophil serine proteases proteinase 3 and elastase.
Am J Pathol
149:
1617-1626,
1996[Abstract].