ARCHIVES

OF BIOCHEMISTRY

AND

BIOPHYSICS

Vol. 286, No. 1, April, pp. 126-131, 1991

Kinetics of Superoxide Production by Stimulated Neutrophils Christopher D. V. Black,’ Amram Samuni,* John A. Cook, C. Murali Krishna, Dwight C. Kaufman, Harry L. Malech,? and Angelo Russo Radiation Oncology Branch, Clinical Oncology Program, Division of Cancer Treatment, National Cancer Znstitute and tBacteria1 Diseases Section, Laboratory of Clinical Investigation, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, Maryland 20892; and *Molecular Biology, School of Medicine, Hebrew University, Jerusalem, 91010, Israel

Received August 28, 1990, and in revised form November

28, 1990

Oxygen-derived active species and superoxide radical in particular are generated and excreted upon granulocyte activation and are instrumental in host defense against bacterial and fungal infections. Associated with the activation of neutrophils is an apparent transitory oxy-radical production. Evidence from independent methods has previously suggested that radical production peaks shortly following neutrophil stimulation and decays within minutes. However, since neutrophil function in the body might reasonably be expected to last beyond the few minutes following stimulation, cessation of the production of oxy-radicals is unexpected. In an attempt to reconcile this discrepancy, the formation kinetics of superoxide by stimulated human neutrophils was reinvestigated by three independent methods: electron spin resonance, chemiluminescence, and ferricytochrome c reduction. The present results demonstrate that under appropriate experimental conditions stimulated neutrophils have the capacity to produce superoxide for several hours. The reasons for the previously reported “apparent” ephemeral nature of oxy-radical formation are discussed. 0 1991 Academic Press, inc.

A burst of oxygen consumption accompanied by the release of reactive products of oxygen reduction is associated with neutrophil-mediated defense against bacterial infection. A one-electron reduction of oxygen, mediated through a membrane-associated NADPH oxidase, is the

i To whom correspondence and reprint requests should be addressed at the Radiation Oncology Branch, National Cancer Institute, Building lo/Room B3-B69, National Institutes of Health, Bethesda, MD 20892. FAX: (301) 480-5439.

primary event leading to the formation of the superoxide anion (Of)” and other oxygen-derived species such as ‘OH, H202, and HOC1 (l-lo). Evidence from many in vitro studies has suggested that cell-induced production of active species has a pronounced transient nature; i.e., stimulated cells produce radicals, but this formation is only temporary and stops within minutes. The apparent ephemeral nature of events triggered by cell stimulation has been corroborated by independent methodologies such as spectrophotometry (g-12), chemiluminescence (CL) (13-16), and electron spin resonance (ESR) (9, 17, 18). Moreover, kinetic models describing and simulating these rapid and transient events have been developed (18, 19) and several physiological mechanisms have been proposed to account for the respiratory burst termination (20,21). A priori, it would seem unreasonable under physiologic conditions to limit the function of cells so crucial to the body’s defence. Indeed, recent theories of neutrophil activation require a more prolonged period of oxidative output to inactivate key plasma components and to activate secreted enzymes (22). The importance of oxygen-derived radicals and the possible drawbacks associated with their detection called for a reevaluation of the presumed transient nature of neutrophil-mediated radical and Hz02 production. Recent findings suggest that there may be subtle and misleading experimental drawbacks associated with several methods commonly used to study cell-induced free * Abbreviations used: CL, chemiluminescence; DMPO, 5,5-dimethylI-pyrroline-N-oxide; DMPO-OH, 5,5-dimethyl-2-hydroxy-l-pyrrolidinyloxyl; Cyt-c”‘, ferricytochrome c; DMPO-OOH, 5,5-dimethyl-2-hydroperoxy-1-pyrrolidinyloxyl; Of, superoxide; ESR, electron spin resonance; DTPA, diethylenetriamine pentaacetic acid, HBSS, Hank’s balanced salt solution; HRPO, horseradish peroxidase; PBS, phosphatebuffered saline; PMA, phorbol12-myristate 13-acetate; SOD, superoxide dismutase; RPMI, Roswell Park Memorial Institute medium 1640; MPO, myeloperoxidase; PMN, polymorphonuclear leukocyte.

126 All

Copyright 0 1991 rights of reproduction

0003-9861/91 $3.00 by Academic Press, Inc. in any form reserved.

KINETICS

OF NEUTROPHIL-MEDIATED

radicals: (a) Radical-trapping by 5,5-dimethyl-l-pyrroline-N-oxide (DMPO) has been extensively used for studying stimulated neutrophils (9,17,18). The persistent spin-adducts, DMPO-OH and DMPO-CHs, have been assumed to accurately report on radical production. Contrary to previous conclusions, neither DMPO-OH nor DMPO-CHB validly monitor either ‘OH or 0; production because the spin-adducts’ signal may decrease with increasing superoxide flux (23), whereas, the DMPO-OOH adduct can be monitored directly to follow the production of 0; (24); (b) Many ESR experiments were performed in standard, flat, quartz cells in which oxygen cannot be replenished and, consequently, 0; production is halted by virtue of O2 depletion (17, 25), thus leading to an artifactual termination of neutrophil radical production. To avoid the intrinsic pitfalls associated with any single method, in the present study the time course of 0; production has been determined by several complementary and improved techniques. We find that (a) 0; formation by PMA-stimulated neutrophils is not transient but persists, and (b) the previous studies describing the apparent transient nature of oxy-radical production do not reflect actual neutropbil physiology but only experimentally imposed constraints. EXPERIMENTAL

PROCEDURES

Materials. Superoxide dismutase (EC 1.15.1.1 superoxide: superoxide oxidoreductase), horseradish peroxidase (EC 1.11.1.7 donor:hydrogen peroxide oxidoreductase), ferricytochrome c (Cyt-c”‘), diethylenetriaminopentaacetic acid, &amino-2,3-dihydro-1,4-phthalazinedione (luminol), and Ficoll/sodium diatriazoate (Histopaque 1083) were obtained from Sigma Chemical Co. Catalase was purchased from Boehringer Mannheim. 5,5-Dimethyl-1-pyrroline-N-oxide (DMPO) was purchased from Aldrich Chemical Co. Phorbol 12-myristate 13-acetate was purchased from Chemical Dynamics, Inc. DMPO was purified by vacuum distillation and checked for the absence of ESR-observable contamination. Luminol was recrystalized from KOH. All other chemicals were prepared and used without further purification. Phosphate-buffered saline, Hank’s balanced salt solution free of Ca’+ and Mg*+ without phenol red, and RPM1 1640 medium without phenol red were supplied by Biofluids Inc. Distilled-deionized water and controlled temperature were used throughout all experiments. Human mononuclear cells and granuPreparation of neutrophils. locytes were prepared from the blood of normal volunteers by sedimentation on a Ficoll/sodium diatriazoate gradient. Neutrophils were separated from erythrocytes by further sedimentation using 3% dextran at room temperature, followed by hypotonic lysis of the remaining red cells (26). Depending on the experiment, neutrophils (lOa-107/ml) were stimulated in either Hank’s balanced salt solution (HBSS), phosphate-buffered saline (PBS) with 0.1% glucose, or RPM1 1640 medium. Phenol red-free RPM1 (or HBSS) was used to avoid interference of the color with spectrophotometric and photon counting assays, although in practice the results were the same with or without phenol red in the medium. Reaction mixtures always contained 50-100 pM DTPA to minimize unwanted transition metal effects. Phorbol 12.myristate 13-acetate (PMA) was dissolved in ethyl alcohol (1 mg/ml), diluted before use in PBS, and further diluted to 0.16-l rig/ml final concentration in the media containing the cells. The stimulated PMNs were maintained separately (i.e., in the absence of cytochrome c, DMPO, or luminol) until needed. At timed intervals after stimulation the cells were gently resuspended and aliquots were taken with obvious clumps of cells being avoided.

SUPEROXIDE

PRODUCTION

127

Discontinuous sampling in this way was used so as not to limit the supply of either oxygen or the materials needed for the detection of 0;. To assess MPO activity of donors, Myeloperonidase (MPO) activity. blood smears were stained using the benzidine dihydrochloride method (27) and examined under light microscopy. None of the donors were found to be MPO-deficient. Blood donated by leukapheresis was routinely stained prior to donation to detect and reject abnormal leukocytes and MPO-deficient donors. Electron spin resonance. Neutrophils used in ESR experiments were suspended at 25°C in HBSS, PBS containing 0.1% glucose, or RPM1 (lOa-lo7 cells/ml) containing 75-100 mM DMPO, and PMA. Samples (100 @l) of stimulated neutrophils were drawn by a syringe into a gaspermeable teflon capillary tube, placed horizontally to minimize effects of cells settling in the cavity of a Varian E9 X-band spectrometer fitted with a temperature controller, and scanned for ESR signals. Unless otherwise stated, all experiments were carried out at controlled temperatures and in air. To monitor the time-dependence of the signal, either successive ESR spectra were recorded or the magnetic field was kept constant at the specified spectral line, while the intensity of the ESR signal was followed. &peroxide assay. The superoxide formation rate was determined by following superoxide dismutase (SOD)-inhibitable Cyt-c”’ reduction. Neutrophils were stimulated by PMA (0.5 pg/ml) and maintained at 25°C in HBSS, PBS containing 0.1% glucose, or RPMI. The media were generally supplemented with 50 pM DTPA and 25 mM Hepes buffer. At timed intervals an aliquot of the stimulated cells was added to each of two cuvettes (0.5-l X lo6 cells/ml). The reference cuvette contained 90 U/ml SOD. The assay was started by the addition of 100 PM Cyt-c”’ to both cuvettes and the OD change at 550 nm was followed using an SLM DW-PC dual beam spectrophotometer (27); rates of superoxide production were calculated from the initial slope of these curves. Luminol-amplified chemiluminescence (CL). Neutrophils were stimulated with PMA at 25°C in aerated RPM1 or HBSS containing 50 HIM DTPA and 50 pM luminol, with or without 10 U/ml horseradish peroxidase (HRPO) (14,15). Chemiluminescence was measured at 25°C by photon counting using an SLM 8000 spectrofluorimeter. To measure 02 only, HRPO was omitted and 65 U/ml catalase was added to eliminate H,O,-induced CL (15). Two measurement modes were used: (a) the cells were stimulated in the presence of luminol and the CL emitted from a single sample of cells was monitored continuously for a long duration; (b) aliquots from a suspension of cells stimulated and incubated in the absence of luminol were sampled at various time points, luminol was added, and the initial maximal transient CL was determined.

RESULTS

Stimulated neutrophils are usually reported to produce oxy-radicals in a single pulse-like fashion. Figure la illustrates a typical time-course of oxy-radical production by PMA-stimulated (1 pg/ml) neutrophils (10’ cells/ml) as monitored by luminol-amplified CL. Figure lb shows a similar time/intensity profile of the DMPO-OOH spinadduct obtained (full ESR spectra shown in Fig. 2) when spin-trapping ESR experiments are conducted in standard oxygen diffusion-limited flat quartz cells. From such data it might be assumed that there is a rather rapid rise and a subsequent fall in the neutrophil-mediated production of oxy-radicals. To determine the time course of oxy-radical production when oxygen is not a limiting factor, experiments were conducted under constant supply of oxygen. Time-course of 0% production by stimulated neutrophils measured by ESR. In a controlled air environment at

128

BLACK

Time (min)

FIG. 1. Apparent “transient” oxidative burst of PMA-stimulated neutrophils: Transient luminol-amplified CL and transient radical production as measured by spin-trapping and ESR detection using a standard oxygen diffusion-limited cuvette, monitored upon stimulation of human neutrophils with 1 Fg/ml PMA in RPM1 medium in air at 25Y!. (a) Time-dependence of luminol-amplified chemiluminescence, emitted by 10s PMNs/ml stimulated in the presence of 50 pM luminol, and continuously recorded using a SLM 8000 spectrofluorimeter. (b) Time-dependence of DMPO-OOH ESR signal observed using a standard, flat, quartz ESR cell containing lo7 PMNs/ml stimulated in the presence of DMPO spin trap. Experimental conditions: 10 mW microwave power, 100 kHz modulation frequency, 1 G modulation amplitude, 10 G/min sweep rate, 1 s response time, and 8 X 10’ receiver gain.

25’C, PMA-stimulated (1 pug/ml) neutrophils (lo7 cells/ ml) in RPMI, HBSS, or PBS containing 75 mM DMPO gave rise to a 12-line spectrum (uN = 14.2 G, on = 11.3 = 1.3 G) typical of DMPO-OOH accompanied by G, aHy a 4-line (uN = 14.9 G, on = 14.9 G) DMPO-OH signal, as seen in Fig. 2 (28). Since DMPO-OH reacts with 0; (23, 24), the DMPO-OH spin-adduct does not truly reflect the rate of radical generation by stimulated cells. Therefore, the rate of radical production was followed by continuously monitoring either the lowest-field or the highestfield line of the DMPO-OOH spectrum. Since the DMPOOOH spin adduct has a half-life of less than a minute (29), the signal intensity represents the rate of 0; generation, not a cumulative total, as is demonstrated below. As shown in Fig. 3 (open circles), after cell stimulation the DMPO-OOH signal intensity reached a maximum within several minutes, slowly decreased to 50-30%, and then persisted for several hours. Depending on the cell and stimulant concentrations used, the initial 0; production rate and the time to reach maximum signal intensity varied, yet the DMPO-OOH signal persisted for 5-6 h (the spectrum shown in Fig. 2 was recorded 3.5 h

ET AL.

after cell stimulation) and the cells remained trypan blue negative. DMPO-OOH dependence on cell concentration. To examine whether the DMPO-OOH signal truly represents the rate of 0: production, the concentration of the stimulated cells was varied, and the signal intensity recorded. Because of slight variations in the time to reach maximum signal intensity and in order to ensure against recording large variations solely caused by being either on or off the peak intensity, the DMPO-OOH signal was measured 30 min after stimulation. The results (Fig. 4) show that the steady state concentration of DMPO-OOH linearly increases with cell concentration. Assuming that the 0; production rate is linearly dependent on the concentration of the stimulated neutrophils, whereas the decay of DMPO-OOH is not, the linear dependence seen in Fig. 4 suggests that the [DMPO-OOHlsteadystate signal intensity reflects the rate of 0; production. Self-generated inhibition of luminol-amplified CL. In contrast with our findings with ESR and Cyt-c”’ assays (vide supra) but in agreement with the findings of others (13-15), when we performed the CL assay on PMA-stimulated PMNs at 25°C under continuous air supply in the presence of luminol and HRPO, the CL began to decay within a few minutes (Fig. la). Further addition of luminol, PMA, or HRPO did not restore CL. In similar experiments, the addition of either exogenous catalase or a combination of azide (to block MPO activity) and superoxide dismutase (to remove 0;) failed to prevent the loss of CL demonstrated in Fig. la. Assuming that luminol-amplified CL validly reflects the flux of 0; and HzOz, this rapid decay could result from cessation of radical formation by the neutrophils or from the generation of an inhibitor of light production. When neutrophils were stimulated in the absence of luminol and aliquots were removed after various incubation times and added directly to luminol, in the presence of a large excess of catalase to eliminate CL resulting from HzOz, CL was demonstrated at all time points as seen in Fig. 3. These data indicate that (a) cell-derived radical production does not cease; and (b) the continuous assay using luminol-amplified CL erroneously reports on radical production. 0; production rate studied concurrently by different methods. To assure that the persistent 0; production by stimulated cells is indeed valid, the rate of 0; production by PMA-stimulated neutrophils was investigated by three methods concurrently: Cyt-c”’ reduction, [DMPOCL. OOHl,tea, state ESR signal, and luminol-amplified Neutrophils (4 X lo7 cells/ml) in RPM1 medium were stimulated by PMA and kept at 25°C. Since the luminol reaction appeared to generate a CL inhibitor, the stimulated neutrophils were incubated without luminol, then at given times aliquots were taken and added directly to RPM1 containing luminol. Since luminol-amplified CL can originate from either HzO, or 0; and since neutrophilderived HzOz is stable and can persist whereas 0; does

KINETICS

OF NEUTROPHIL-MEDIATED

SUPEROXIDE

PRODUCTION

129

GAUSS FIG. 2. ESR spectra obtained following neutrophil stimulation in the presence of DMPO spin trap: Neutrophils (lo7 cells/ml in RPM1 medium) with continuous oxygen supply were stimulated with PMA in the presence of 75 mM DMPO and placed in a gas-permeable teflon capillary under air flow at 25°C within the ESR cavity. The ESR spectra of DMPO-OOH and DMPO-OH were recorded 3.5 h after neutrophil stimulation.

not, 65 U/ml catalase was added to the luminol CL reaction vessel to eliminate HzOz, thus ensuring that the source of CL originated exclusively from ongoing production of 0;. Concurrently, aliquots of stimulated neutrophils were diluted into RPM1 medium containing either 75 InM DMPO or 100 PM Cyt-c”’ and studied using the three different assay methods. Typical results from the cyt-c” assay method are illustrated in Fig. 5. All three methods showed similar time-dependency profiles indicating the continuation of 0; production for long duration. The rate of 0; production l-2 h post stimulation, as determined by the Cyt-c”’ assay, was about 0.5 nmol O~/min/lO’ cells, compared with the peak value of l-3 nmol O~/min/lO’ cells generally found for normal stimulated neutrophils shortly after stimulation (28). DISCUSSION Our choice of PMA in these studies is a reflection of the continuing dominant role of this stimulant in neutrophil research and, as such, these results are representative of the oxidative capacity of stimulated human neutrophils rather than an exact replication of physiological conditions. Because oxygen utilization and oxygen-derived radical production by activated neutrophils have been described as being burst-like, several physiological

mechanisms have been proposed to account for the apparent “respiratory burst termination” (20,21). Inherent, physiologically important factors responsible for the termination of the neutrophil respiratory burst have been previously invoked. Such factors have been thought to be important in limiting the neutrophil’s ability to produce superoxide, thereby protecting normal tissues from neutrophil-derived oxy-radicals. However, as Weiss has suggested (22), tissue damage is a complex series of interactions between oxy-radicals, plasma components, and neutrophil-derived enzymes; neutrophil-mediated destruction of bacteria will be enhanced and normal tissue destruction controlled by a prolongation of the oxidative burst. An interesting correlation between MPO activity and duration of radical formation by the cells has been based on the finding that zymosan-stimulated neutrophils from MPO-deficient subjects manifested a prolonged Oc-production (31). Such a difference between normal and MPO-deficient cells, however, was not detected for PMA-stimulated neutrophils, which were reported to produce 0; for much shorter periods (32,33). In the present study all human blood samples exhibited normal MPO activity. We find, however, that as long as adequate oxygen and nutrients are provided, there is no rapid termination of the production of oxygen-derived radicals. In

130

BLACK

ET AL.

Stimulated neutrophils (million cells/ml) 100

Time

after

cell stimulation

200

(min)

FIG. 3. Time dependence of the rate of 0; production by stimulated neutrophils assayed by ESR and CL: The rate of 0; production by neutrophils stimulated with 1 pg/ml PMA in RPM1 at 25°C was investigated by concurrently employing independent assay methods. All values were normalized to the peak height, which was taken as 100. (a) Time dependence of the ESR signal of [DMPO-OOH],,,,,,, obtained using lo7 PMNs/ml and 75 mM DMPO. Signal intensity was measured at various time points following cell stimulation (circles). (b) Luminolamplified CL assay performed in a “discontinuous” manner. PMAstimulated cells (4 X lo7 PMNs/ml) in RPM1 medium were stimulated and kept at 25’C in the absence of luminol. Aliquots were sampled at timed intervals from the incubation system, diluted 1:40 into RPM1 containing 50 PM luminol + 65 U/ml catalase and the transient maximal luminol-amplified CL was monitored (triangles).

fact, even several hours following potent stimulation with PMA, neutrophils are completely viable and continue to produce 0;. The present results, therefore, do not appear to be associated with decreased MPO activity. Clearly, stimulation and prolonged exposure to superoxide flux did not provoke a suicide effect on neutrophil viability as determined by trypan blue exclusion. Using a variety of physiological stimulants, Nathan (34-36) has demonstrated that neutrophils adhering to biological surfaces are able to produce hydrogen peroxide for several hours. Those findings, therefore, suggest that our data showing prolonged superoxide production by PMA-stimulated neutrophils in suspension may be a reflection of physiologic conditions. We demonstrate here that earlier failures to show persistence of superoxide production by PMAstimulated neutrophils can be due to assay systems that are either inherently incapable of accurately following the 0; production rate or that themselves perturb the system to alter the product being assayed. The short-lived DMPO-OOH spin adduct, generated from DMPO and Ot, partly decays to DMPO-OH by a poorly understood mechanism (28):

FIG. 4. The dependence of [DMPO-OOH],,,, on neutrophil concentration: The ESR signal intensity of DMPO-OOH was recorded 30 min following PMA stimulation of various concentrations of neutrophils in RPM1 with continuous air supply at 25’C.

DMPO-OH and DMPO-CH3 spin adducts do not accumulate (23, 37). The same is true for DMPO-OH and DMPO-CHB formed from stimulated neutrophils (24), where the failure of the relatively long-lived DMPO-OH and DMPO-CH3 to accumulate stems from their rapid destruction by neutrophil-produced 0; : OH + ‘0,

+ ESR-silent

products

PI

Since 0; contributes to both the formation (Reaction [l]) and the destruction of DMPO-OH (Reaction [2]), neither the DMPO-OH nor the DMPO-CH3 signal can serve as a quantitative marker of superoxide production; therefore, kinetic models analyzing time-dependencies of the DMPO-OH signal (18, 19) cannot describe the actual course of cell-induced 0; formation. By contrast, based on the linear dependence of [DMPO-OOH],,,, stateon cell concentration as seen in Fig. 4 and on the agreement found between time profiles of oxidative burst measured by independent methods (Figs. 3 and 5), we conclude that [DMP@OOHlatea,state does reflect 0; formation and can serve as an apparent measure of the rate of 0; production (23, 24).

z o.oth”““.‘I”“““‘,“‘.’ 0

Time

*OH It has been previously shown that under conditions in which high fluxes of 0; are generated enzymatically,

after

100

200

cell stimulation

(min)

FIG. 5. The kinetics of 0; production by PMA-stimulated neutrophils assayed by Cyt-c”’ reduction: The production rate of 0; was determined using the Cyt-c”’ reduction assay. PMNs were incubated in RPM1 at 25’C and stimulated by 0.5 rg/ml PMA. At various time points aliquots of 100 ~1 were sampled into 2 ml RPM1 containing 100 pM Cyt-c”’ and 65 U/ml catalase. The reference cuvette contained, in addition, 90 U/ ml SOD. Cytc”’ reduction was followed at 550 nm and rates of reduction were determined from the initial slopes.

KINETICS

OF NEUTROPHIL-MEDIATED

The standard type of ESR cuvette does not allow for oxygen replenishment as seen by comparing Fig. lb and Fig. 3 (open circles); therefore, it should not be used to study production kinetics of 0; by cells, since cellular metabolism quickly results in consumption of available oxygen. Experiments conducted with oxygen-selective electrodes are subject to similar constraints imposed by a lack of oxygen replenishment. But when ESR is used with neutrophils suspended in medium at 25’C with constant air supply, 0; generation was observed for hours (Fig. 3). Another possible source for an ephemeral radical production by PMA-stimulated neutrophils is the use of buffer (11, 13-15, 17, 18, 25) instead of nutrient medium such as RPMI. We have shown that PMNs stimulated at 37°C in buffer shortly stop producing 0: (38). In contrast, cells incubated in RPM1 at 25°C continued to generate superoxide as seen in Figs. 3 and 5. Because, in these cytochrome c experiments, the rate of 0; production was not measured continuously but rather at timed intervals after stimulation, the data are not limited by the availability of oxygen or substrate. In the present study the use of luminol-amplified chemiluminescent probing of 0; production by stimulated neutrophils is shown to be problematic, since the assay system itself generates an as yet undefined material that obliterates the CL (Fig. la). We are currently attempting to isolate and determine the structure of the compound(s) responsible for the inhibition. However, even without knowing the identity of this inhibitor, the present results clearly show that assays employing continuous luminolamplified CL cannot be used to determine the kinetics of 0; and H202 formation. When taken as a whole, our results show that previous failures to monitor persistent production of oxy-radicals in vitro may reflect artifacts and do not necessarily imply that in uiuo neutrophils rapidly lose their ability to produce 0:. REFERENCES 1. Babior, B. M., Kipnes, R. S., and Curnutte, J. T. (1973) J. Clin. Invest. 52, 741.-744. 2. Johnston, R. B. Jr., Keele, B. B. Jr., Misra, H. P., Lehmeyer, J. E., Webb, L. S., Baehner, R. L., and Rajagopaian, K. V. (1975) J. Clin. Invest. 55, 1357-1372. 3. Weiss, S. J., King, G. W., and LoBuglio, A. F. (1977) J. Clin. Inoest. 60,370-373. 4. Rosen, H., and Klebanoff, S. J. (1977) J. Bio2. Chem. 252, 48034810. 5. Babior, B. M. (1978) N. Engl. J. Med. 298, 659-668 and 721-725. 6. Nathan, C., and Cohn, Z. (1980) J. Exp. Med. 152, 198-208. 7. Weiss, S. J., and Slivka, A. (1982) J. Clin. Inuest. 69, 255-262. 8. Cross, A. R., Parkinson, J. F., and Jones, 0. T. G. (1984) Biochem. J.. 223. 337-344.

SUPEROXIDE

131

PRODUCTION

9. Hawley, D. A., Kleinhans, F. W., and Biesecker, J. L. (1983) Amer. J. Clin. Pathol. 79, 673-677. 10 Bass, D. A., McPhail, L. C., Schmitt, J. D., Morris-Natschke, S., McCall, C. E., and Wykle, R. L. (1988) J. Biol. Chem. 263, 19,61019,617. 11. Simchowitz, L. (1985) J. Clin. Inuest. 76, 1079-1089. 12. Shurin, S. B., Cohen, H. J., Whitin, J. C., and Newburger, P. E. (1983) Blood 62, 564-571. 13. Bender, J. G., and Van Epps, D. E. (1983) Infect Zmmun. 41, 10621070. 14. Dahlgren, C., and Stendahl, 0. (1983) Znfecr. Immun. 39,736-741. 15. Wymann, M. P., von Tscharner, V., Deranleau, D. A., and Baggiolini, M. (1987) Anal. Biochem. 165, 371-378. 16. Allen, R. C., Mead, M. E., and Kelly, J. L. (1985) in CRC Handbook of Methods for Oxygen Radical Research, (Greenwald, R. A. Ed.), pp. 343-351. CRC Press, Boca Raton, FL. 17. Cheung, K., Lark, J., Robinson, M. F., Pomery, P. J., and Hunter, S. (1986) Aust. J. Enp. Biol. Med. Sci. 64, 157-164. 18. Kleinhans, F. W., and Barefoot, S. T. (1987) J. Biol. Chem. 262, 12,452-12,457. 19. Dahinden, C. A., Fehr, J., and Hugli, T. E. (1983) J. Clin. Inuest. 72, 113-121. 20. Whitin, J. C., and Cohen, H. J. (1988) Hematol. Oncol. Clin. North Amer. 2, 289-299. 21. Jandl, R. C., Andre-Schwartz, J., Borges-Dubois, L., Kipness, R. S., McMurrieh, J., and Babior, B. M. (1978) J. Clin. Znuest. 61, 1176-1185. 22. Weiss, S. J. (1989) N. Engl. J. Med. 320, 356-376. 23. Samuni, A., Krishna, C. M., Riesz, P., Finkelstein, E., and Russo, A. (1989) Free Radical Biol. Med. 6, 141-148. 24. Samuni, A., Black, C. D. V., Krishna, C. M., Malech, H. L., Bernstein, E. F., and Russo, A. (1988) J. Biol. Chem. 263, 13,797-13,801. 25. Rosen, G. M., Britigan, B. E., Cohen, M. S., Ellington, S. P., and Barber, M. J. (1988) Biochim. Biophys. Acta 969,236-241. 26. Boyum, A. (1968) Scan. J. Clin. Lab. Invest. 2l(SuppI. 97), 77-89. 37 Kaplow, L. S. (1965) Blood 26, 215-219. -.. 28. Markert, M., Andrews, P. C., and Babior, B. M. (1984) in Methods in Enzymology (Packer, L., Ed.), Vol. 105, pp 358-365, Academic Press, San Diego. 29. Mottley, C., Connor, H. D., and Mason, R. P. (1986) Biochem. Biophys. Res. Commun. 141,622-628. 30. Finkelstein, Pharmacol.

E., Rosen, G. M., and Rauckman, 2 1, 262-265.

E. J. (1982) Mol.

31. Rosen, H., and Klebanoff,

S. J. (1976) J. Clin. Znuest. 58, 50-60.

32. Nauseef, W. M., Metcalf, 483-492.

J. A., and Root, R. K. (1983) Blood 61,

33. Dri, P., Soranzo, M. R., Cramer, R., Menegazzi, Patriarca, P. (1985) ZnfZammation 9, 21-31. 34. Nathan, 35. Nathan,

R., Miotti,

V., and

C. F. (1987) J. Clin. Znuest. 80, 155061560. C. F. (1989) Blood 73,301-306.

36. Nathan, C., Srimal, S., Farber, C., Sanchez, E., Kabbash, L., Asch, A., Gailit, J., Wright, S. D. (1989). J. Cell. Biol. 109, 1341-1349. 37. Pou, S., Cohen, M. S., Britigan, B. E., and Rosen, G. M. (1989) J. Biol. Chem. 264, 12,299-12,302. 38. Black, C. D. V., Cook, J. A., Russo, A., and Samuni, A. (1991). Free Radical Res. Commun., in press.

Kinetics of superoxide production by stimulated neutrophils.

Oxygen-derived active species and superoxide radical in particular are generated and excreted upon granulocyte activation and are instrumental in host...
728KB Sizes 0 Downloads 0 Views