The O−2Generating NADPH Oxidase of Phagocytes: Structure and Methods of Detection

METHODS: A Companion to Methods in Enzymology 9, 563–577 (1996) Article No. 0064

The O0 2 Generating NADPH Oxidase of Phagocytes: Structure and Methods of Detection Steven W. Edwards Department of Biochemistry, University of Liverpool, P.O. Box 147, Liverpool L69 3BX, United Kingdom

The NADPH oxidase of phagocytic cells of the immune system plays an important role in the destruction of certain types of microbial pathogens during infections. Its inappropriate activation to release reactive oxidants may also contribute to host tissue damage in inflammatory diseases. In this review, the structure of the NADPH oxidase is described and many of the commonly used methods to detect its activation during the respiratory burst are listed. The advantages and disadvantages of each of these methods are critically discussed. q 1996 Academic Press, Inc.

Phagocytic cells of the immune system (neutrophils, eosinophils, monocytes, and macrophages) generate a series of reactive oxygen metabolites during a respiratory burst of nonmitochondrial O2 uptake (1, 2). These reactive oxidants play an important role in the killing of microbial pathogens (Fig. 1A) and may also be involved in the killing of certain types of tumor cells. More recently, it has been discovered that activated B cells may also possess this oxidase, giving rise to the suggestion that reactive oxidants may play as yet undefined regulatory roles in immune function.

DISCOVERY OF THE RESPIRATORY BURST The respiratory burst of phagocytes was discovered by Baldridge and Gerard (3) when they found that phagocytosis of bacteria was accompanied by a rapid ‘‘burst’’ of O2 uptake. Because O2 was known to be required for mitochondrial respiration to generate ATP, it was mistakenly believed that the respiratory burst provided the extra energy needed for the physical processes of phagocytosis. It was not until 1959 that the unusual nature of the respiratory burst was noted (4), when it was shown to be unaffected by inhibitors of mitochondrial respiration and hence unlikely to be involved in energy production. In the 1960s it was pre-

dicted and then shown experimentally that H2O2 was a product of the respiratory burst, and hence it was suggested that the burst was important for microbial killing. The link between the respiratory burst and pathogen killing was then confirmed by studies on patients with chronic granulomatous disease (CGD),1 then known as fatal granulomatosis of childhood. These patients have increased susceptibility to infections by certain types of bacteria and fungi, and in severe cases the condition is fatal unless carefully managed by prophylactic antibiotic therapy. Experiments by Holmes et al. (5, 6) showed (a) that neutrophils from CGD patients could phagocytose but not kill the microorganisms that caused infections in these patients, and (b) these neutrophils did not generate a respiratory burst during phagocytosis. Babior et al. (7) discovered that the primary product of the respiratory burst is O20, which is generated via the one-electron reduction of O2 . Because the oxidase preferentially utilizes NADPH as a source of electrons for O2 reduction, it was termed the NADPH oxidase. Thus, the unusual nature of the NADPH oxidase was appreciated and its importance in microbial killing established. However, it was not until very recently that the structure of this important enzyme was finally elucidated. In this quest to characterize its structure, the CGD neutrophil has proved to be an invaluable tool: putative components that are absent or defective in CGD neutrophils are very likely to be genuine components of the NADPH oxidase (8).

STRUCTURE OF THE NADPH OXIDASE The overall reaction carried out by the NADPH oxidase is 1 Abbreviations used: CGD, chronic granulomatous disease; O0 2, superoxide; H2O2 , hydrogen peroxide; SDS, sodium dodecyl sulfate; FAD, flavin adenine dinucleotide; PBS, phosphate-buffered saline; luminol, 5-amino-2,3-dihydro-1,4-phthalazinedione; lucigenin, bisN-methylacridinium nitrate.

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1046-2023/96 $18.00 Copyright q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

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NADPH / 2O2 r 2O20 / NADP/ / H/.

[1]

In vitro, the oxidase will utilize both NADH and NADPH but its Km values for these substrates (450 and 45 mM, respectively) indicate that in vivo the latter cofactor is the source of reductant and electrons used for O2 reduction. This NADPH is generated via the hexose monophosphate shunt (pentose phosphate pathway), the activity of which is increased during phagocytosis. The oxidase is a multicomponent complex that is dormant in nonactivated cells. All of the constituent components of the NADPH oxidase have now been identified, cloned, and characterized at the molecular level (9). Curiously, these components are present in different subcellular locations of the cell. During phagocytosis or other forms of activation, they are brought together via subcellular translocations to assemble into functional oxidase complexes. Cytochrome b0245 The first component of the NADPH oxidase to be discovered was an unusual cytochrome b (10). This cytochrome became incorporated into phagolysosomes during phagocytosis and importantly was absent in patients with X-linked CGD, the most severe form of the disease (11). This cytochrome is unusual in that it possesses a very low midpoint potential (0245 mV) compared to other b-type cytochromes (12, 13). For this reason it is often referred to as cytochrome b0245 . This midpoint potential is close to that of the O2/O20 redox couple and hence low enough to directly reduce O2 to O20. In reduced–oxidized difference spectra, it has an absorption maximum of 558 nm and so is sometimes referred to as cytochrome b558 . This cytochrome functions as the terminal component of an electron transport chain, passing electrons from NADPH to O2 . It can bind CO (albeit with low affinity), and in the presence of O2 it is reduced by NADPH at a rate almost equal to the rate of O20 formation by activated neutrophils. The first successful purification of this cytochrome b0245 was reported by Harper et al. (14). The cytochrome comprises a small (22-kDa) a-subunit and a larger bsubunit that is heavily glycosylated and runs as a diffuse 76- to 92-kDa band on SDS–PAGE. These subunits have thus been termed p22-phox and gp91-phox, respectively. The genes for both subunits have been cloned; that of gp91-phox was cloned by reverse genetics (15–18). The a and b subunits of the cytochrome possess little homology to other known b-cytochromes, but some regions of the a subunit have homology to regions of mitochondrial cytochrome oxidase complex I and to bovine chromaffin granule cytochrome b. Northern blotting and immunoassays have shown that the a subunit can be expressed in cells that do not possess spectrally identifiable cytochrome b0245 .

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In unstimulated neutrophils, the cytochrome is found on the plasma membrane, but most of the total cellular pool (approximately 90%) is present on the membranes of specific granules, gelatinase-containing granules, and secretory vesicles (19–22). After stimulation, these granules migrate to and fuse with either the plasma membrane (as is the case when the neutrophil is stimulated by soluble stimuli) or the phagosome (as is the case with stimulation by particles that are phagocytosed). Thus, the plasma membrane or the phagosome becomes enriched in cytochrome b molecules (23, 24), providing an increased supply of important oxidase constituents to the site of reactive oxidant production. This is required because once activated, oxidase complexes have a finite life and become inactivated. Sustained reactive oxidant production thus requires the recruitment, assembly, and activation of new complexes. This cytochrome is present, however, in normal amounts in neutrophils from patients with the autosomal recessive form of CGD, but it fails to become reduced upon physiological activation. These observations thus originally indicated that other NADPH oxidase components were likely to exist in normal neutrophils, but were defective or absent in autosomal recessive forms of CGD. Flavins The involvement of flavins in the NADPH oxidase has been appreciated for many years. For example, it was shown that oxidase activity in neutrophil extracts can be enhanced by the addition of flavins and that flavin analogues (e.g., 5-carbodeaza-FAD nucleotide) and flavoprotein inhibitors (e.g., diphenylene iodonium) can inhibit oxidase activity (25 – 28). Furthermore, flavins can be detected in NADPH oxidase preparations, and kinetic and thermodynamic considerations logically place flavins as intermediary electron carriers between NADPH and cytochrome b0245 . For many years, it was believed that a distinct flavoprotein component of the NADPH oxidase existed, but it has since been shown (29) that cytochrome b0245 possesses a flavin-binding site and is thus a flavocytochrome. Cytosolic Components One of the major breakthroughs in elucidating the components of the oxidase has been the development of experimental systems in which the NADPH oxidase can be activated in neutrophil homogenates prepared from unstimulated cells (30 – 34). Initial experiments showed that oxidase activity could be stimulated by the addition of anionic detergents (e.g., arachidonic acid, cis-unsaturated fatty acids, or SDS) to neutrophil homogenates. Further refinement of this system showed that the addition of cytosol to plasma membranes, in the presence of NADPH and

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arachidonic acid, could lead to the formation of O20. The plasma membrane thus contains cytochrome b0245 , and so these experiments indicated that cytosolic NADPH oxidase components also exist. This conclusion was strengthened following experiments with extracts obtained from CGD neutrophils, which failed to generate O20 in these cell-free systems. However, substituting cytosol and membranes from normal neutrophils could restore activity (35 – 37). In ex-

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tracts from neutrophils with X-linked CGD, it was shown that the membranes were defective, which was perhaps predicted because these neutrophils are defective in cytochrome b0245 . However, in autosomal recessive CGD extracts (which contain cytochrome b0245) the cytosol was defective, indicating that normal cytosol contains key oxidase components, but these are absent or defective in these patients. Purification of normal cytosol by Mono-Q anion ex-

FIG. 1. Activation of the respiratory burst and degranulation in neutrophils. (A) A bacterium (dark oval) has been phagocytosed and the NADPH oxidase has been activated to secrete reactive oxidants into the phagocytic vesicle. Concomitant with oxidase activation is the release of granule enzymes (degranulation) into the phagosome. These granule enzymes and reactive oxidants act in concert to kill and digest the pathogen. (B) Reactive oxidants and granule enzymes are released from activated neutrophils. Thus, NADPH oxidase complexes on the plasma membrane are activated and granules migrate to and fuse with the plasma membrane to release their contents. (C) Secretory vesicles, gelatinase-containing granules, and specific granules migrate to and fuse with the plasma membrane during priming. Thus, the plasma membrane becomes enriched in cytochrome b molecules and some receptors.

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change chromatography yielded three active fractions (NCF-1, -2, and -3) that were involved in oxidase activation (38). Independently of these experiments, a polyclonal antiserum was raised against neutrophil cytosolic proteins that eluted from a GTP-affinity column (39). It was somewhat fortunate that this crude protein extract, which contained a variety of cytosolic proteins, generated an antiserum (B1 antiserum) that recognized only two major cytosolic proteins of 47 and 67 kDa. NCF-1 was shown (by immunoblotting) to contain the 47-kDa protein (p47-phox) while NCF-2 contained the 67-kDa protein (p67-phox). This antiserum was then used to isolate cDNA clones from cDNA expression libraries (40, 41). Sequence analysis of these cDNA clones reveals that both p47-phox and p67-phox contain putative SH3 domains (i.e., regions of src homology) and potential SH3 binding domains (i.e., proline-rich regions). These domains are commonly found in proteins involved in cell signaling and are responsible for protein:protein interactions. Use of B1 antiserum also showed that the majority of autosomal recessive CGD patients lack p47-phox while the remainder lack p67-phox. Recombinant cytosolic proteins can restore activity to cytosol from CGD neutrophils in the cell-free system (42). Experiments using purified cytochrome b0245 , recombinant p47-phox, and p67-phox (together with NADPH and arachidonic acid or SDS) still required trace amounts of cytosol before O20 could be formed, and the ‘‘active’’ component of NCF-3 was not identified. It was also noted that during purification of the cytosolic proteins, reconstitution of oxidase activity became more and more dependent upon the addition of exogenous GTP, implicating the involvement of a G-protein. A third cytosolic component initially from guinea pig macrophage cytosol, has since been identified (43–46). This protein comprises two components, p21rac1 and GDP-dissociation inhibitor factor (GDI). In the presence of exogenous GTP in the cell-free system, only recombinant p21rac1 is required. A similar protein has been purified from human neutrophil cytosol, p21rac2, which has 92% homology to p21rac1. Thus, a ‘‘minimal’’ cell-free system for activation of the NADPH oxidase has been defined that comprises purified gp91-phox and the recombinant proteins p47-phox, p67phox, and p21rac, together with FAD, NADPH, GTP, MgCl2 , and SDS (47). It is thus unlikely that any other components are necessary for NADPH oxidase activity. However, a fourth cytosolic component, p40-phox, has been identified (48). This component is not required in the cell-free system, but is probably involved in the assembly of the oxidase components during cell activation in vivo, very likely via an interaction with p67-phox. p40phox also contains potential SH3 domains, and it probably binds via these to the proline-rich regions of p67-phox and p47-phox. The possible structural association of these

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components during assembly of active NADPH oxidase complexes is shown in Fig. 1.

REACTIVE OXYGEN METABOLITES GENERATED DURING THE RESPIRATORY BURST The primary product of the NADPH oxidase is O20 (Eq. [1]). However, O20 is a relatively unstable molecule and is capable of forming other, secondary oxidants or radicals (Fig. 2). For example, O20 can rapidly dismutate (either spontaneously or enzymically via the activity of superoxide dismutase) as follows: 2O20 / 2H/ r H2O2 / O2 .

[2]

Neutrophils also possess the hemoprotein myeloperoxidase (49, 50), which among other reactions is capable of forming HOCl via the reaction H2O2 / Cl0 / H/ r HOCl / H2O.

[3]

It has been estimated that in some circumstances between 40 and 70% of the O2 that is consumed during the respiratory burst can be detected as HOCl. HOCl can result in the formation of the long-lived oxidants, the chloramines: HOCl / R–NH2 r R–NHCl / H2O,

[4]

or in the formation of singlet oxygen, 1O2 : OCl0 / H2O2 r Cl0 / H2O / 1O2 .

[5]

Furthermore, reactions between O0 2 and H2O2 may occur in certain circumstances (such as in the presence of transition metal salt catalysts) and may lead to the formation of the hydroxyl free radical, •OH, which is the most reactive species known in biological systems (51, 52):

FIG. 2. Reactive oxidant production by activated neutrophils. The activities of the NADPH oxidase and myeloperoxidase can generate a number of reactive oxidants, as shown.

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O20 / H2O2 r O2 / 0OH / •OH.

[6]

This reaction is known as the metal-catalyzed Haber– Weiss reaction, and there is some controversy as to whether it occurs in activated neutrophils (see section on Electron Paramagnetic Resonance Spectroscopy, below). Assays for the detection of all these products of the neutrophil respiratory burst exist. For the reasons given below, none of these is, however, ideal for specifically measuring all NADPH oxidase activity that may occur both on the plasma membrane and within intracellular sites (e.g., within phagolysosomes). Because O20 is the primary product of the oxidase, it is preferable to measure this product whenever possible.

NEUTROPHIL PRIMING Until the 1970s it was believed that mature neutrophils were terminally differentiated end-of-line cells. However, experiments by McCall and colleagues (53, 54) showed that ‘‘toxic’’ neutrophils, i.e., those isolated from patients with acute bacterial infections, had higher activities than neutrophils isolated from healthy controls. Such enhanced activities in these toxic neutrophils that were observed included increased oxidative metabolism, phagocytosis, chemotaxis, and 2-deoxyglucose transport. It was later shown that toxic neutrophils did not arise in the circulation of these patients via the release from the bone marrow of a more active neutrophil subpopulation, but rather the infection somehow changed the biochemical properties of the circulating neutrophils. Thus, the concept of neutrophil ‘‘priming’’ was formulated. Exposure of blood neutrophils to substimulatory concentrations of a number of agonists (e.g., fMet-Leu-Phe, PAF, or LPS; 55) or to cytokines (such as GM-CSF, G-CSF, g-interferon, and TNF-a; 56–58) can prime the neutrophil into a state of enhanced responsiveness. Thus, after priming, neutrophil functions such as the respiratory burst, degranulation, and phagocytosis are all significantly enhanced compared to control cells, and the primed cells respond much more aggressively to stimulation. Undoubtedly, neutrophil function in vivo is also regulated by priming agents. The molecular processes that regulate priming are not fully defined. Priming can occur within 10–60 min of addition of a priming agent (depending upon the nature of the primer) and does not require de novo biosynthesis. Rather, priming induces molecular rearrangements within the neutrophil such as the mobilization of subcellular granules and secretory vesicles (59) to the plasma membrane and also the translocation of cytosolic proteins to the plasma membrane. By such processes, cytochrome b molecules and some re-

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ceptors are enriched on the plasma membrane (Fig. 1C). Apart from increases in receptor number, receptor function can also be altered during priming (60), and the signal transduction systems employed by primed cells are also modified (61). Priming induces profound changes in the molecular properties of neutrophils other than merely enhancing their activities. Within the circulation, neutrophils are only poorly responsive to certain types of stimulation, and consequently the respiratory burst cannot be activated and certain receptors are not expressed. By altering receptor number and function on the cell surface and by assembling active signal transduction systems, priming thus recruits normally nonresponsive neutrophils into cells that can mount a respiratory burst and phagocytosis upon inflammatory challenge. Neutrophil function in vivo is thus tightly controlled by a two-step process. During the first step, the nonresponsive cells are primed by cytokines or local concentrations of other proinflammatory signals; during the second step, the primed cells respond to challenge by activation of their antimicrobial systems. Such a two-step process of control probably protects against nonspecific neutrophil activation that could result in host tissue damage via the uncontrolled secretion of reactive oxidants and granule enzymes (Fig.1B). Unfortunately, many commonly used neutrophil isolation procedures also partially prime neutrophils during purification. There are several ways to overcome this problem. First, some isolation methods (see section on Centrifugation in Neutrophil Isolation Medium) are minimally perturbing and induce little or no priming. Second, some assays can measure certain neutrophil functions in whole, unfractionated blood. Third, neutrophils can be primed in vitro by pretreatment with a suitable priming agent, and the activity of the fully primed cell may then be analyzed. Finally, some agonists can activate neutrophils by bypassing both the occupancy of cell surface receptors and the generation of intracellular signaling molecules. For example, the phorbol ester PMA is an analogue of diacylglycerol, which is the physiological activator of protein kinase C. Thus, PMA treatment stimulates protein kinase Cregulated neutrophil functions, and such activation is largely (but not totally) independent of the primed status of the neutrophil.

NEUTROPHIL ISOLATION FROM WHOLE BLOOD There are many neutrophil isolation media now on the market, based largely on the density-gradient separation technique described in (62). Such media exploit the difference in density and size of neutrophils compared to other immune cells. It is not possible to give

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an exhaustive list of all of these media, but two commonly used examples are given. Both of these described methods yield highly pure preparations of polymorphonuclear leukocytes containing neutrophils, eosinophils, and basophils. Hence, the purity of the final yield of neutrophils should always be checked, usually by staining (e.g., Wright’s stain) and microscopic examination. The methods described below routinely yield preparations that are ú95% neutrophils. Note: use plastic tubes and pipettes throughout because neutrophils adhere tenaciously to glass. Dextran/Ficoll–Hypaque Technique Required Centrifuge; Phosphate-buffered saline, PBS (10 mM potassium phosphate, 0.9% NaCl, pH 7.4); Dextran T500 (Pharmacia), 6% (w/v) in PBS; Ficoll–hypaque (Flow Laboratories) (5.7 g Ficoll 400; 9.0 g sodium diatrizoate per 100 ml); Glass distilled water; 9% NaCl; RPMI 1640 medium (Flow laboratories) or a balanced salt solution (BSS, e.g., 12 mM NaCl; 1.2 mM KH2PO4 ; 4.8 mM KCl; 1.2 mM MgSO4; 1.3 mM CaCl2 ; 25 mM Hepes, pH 7.4; 0.1% BSA). Method (i) To 20 ml of whole (heparinized) blood, add 2 ml of 6% dextran solution. Mix by inversion and allow to sediment at room temperature for 30–40 min. (ii) The erythrocytes (which are permeable to dextran) sediment more rapidly than the leukocytes, and so the upper layer is transferred to plastic centrifuge tubes.

(iii) Centrifuge at 500g for 5 min at room temperature and wash the pellet in RPMI 1640 medium or BSS. (iv) Resuspend the washed pellet in 10 ml RPMI 1640 medium (or BSS) and layer onto 7 ml of Ficoll–hypaque solution (Fig. 3A). Centrifuge at 1000g for 25 min. (v) After centrifugation the neutrophils (together with contaminating erythrocytes) sediment to the bottom of the tube while the monocytes and lymphocytes band at the interface between the Ficoll and the buffer (Fig. 3B) (These can be separated further, by exploiting the fact that monocytes adhere to plastic surfaces but the lymphocytes do not.) The platelets remain largely in the upper layer. (vi) Resuspend the neutrophil/erythrocyte pellet in 9 ml of glass distilled water. Note: neutrophil pellets should be initially suspended in a small volume of medium (i.e., 0.5–1 ml) to avoid irreversible aggregation. After 20 s, isotonicity is restored by addition of 1 ml of 9% NaCl solution. Mix the tube thoroughly. (vii) Centrifuge the suspension at 500g for 5 min. The supernatant should now be pale red (containing hemoglobin released by the hypotonic lysis of the erythrocytes) while the pellet should be green (indicating the presence of the neutrophil enzyme, myeloperoxidase). (viii) The pellet should be washed in RPMI 1640 medium (or BSS) and finally suspended at 1 ml. Purity should be checked by Wright’s differential staining and viability by trypan blue exclusion. Cells can be counted after a suitable dilution using a hemocytometer slide (e.g., a Fuchs–Rosenthal chamber) or by a Coulter counter. Finally, dilute the purified neutrophil at around 1 1 107 to 2 1 107 cells ml01. This method yields approximately 2 1 107 to 4 1 107 neutrophils from 20 ml human venous blood. The neutrophils are typically ú95% pure and ú95% viable.

FIG. 3. Purification of neutrophils by centrifugation on Ficoll–hypaque. For details, see the text.

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Advantages The reagents used in the method are relatively inexpensive and it is suitable (with scale-up) for the purification of neutrophils from large volumes (up to 500 ml) of blood. If the method is scaled up, it is important not to overload the Ficoll–hypaque gradient; otherwise neutrophil purity will be compromised.

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in the final preparation should be assessed as described in step (viii) above.

Disadvantages The major drawback of this method is that it partially primes neutrophil functions such as receptor expression and NADPH oxidase activity (63).

Advantages This method typically yields 2 1 104 to 4 1 107 neutrophils from 20 ml venous blood. The preparations are routinely ú95% pure neutrophils and are ú95% viable. The method is extremely rapid, and the cells isolated show minimal priming of the NADPH oxidase and very low levels of expression of CR3. It is thus strongly recommended for the isolation of minimally perturbed neutrophils.

Centrifugation in Neutrophil Isolation Medium (NIM) Required Centrifuge; Neutrophil isolation medium (Cardinal Associates); Glass distilled water; RPMI 1640 medium or BSS.

Disadvantages The method is not really suitable for the isolation of neutrophils from large quantities of blood, mainly because of the relatively high cost of NIM. Other neutrophil isolation procedures based on centrifugation on Percoll gradients have been described (64).

Method (i) Layer 10–12 ml of heparinized blood onto 8 ml of NIM and centrifuge at 400g for 15–25 min (Fig. 4A). Usually 15 min centrifugation is sufficient, but if this is insufficient for separation, extend the time. (ii) After centrifugation, the neutrophils and monocytes/lymphocytes form discrete bands while the erythrocytes sediment to the bottom of the tube (Fig. 4B). (iii) Remove the neutrophil layer with a plastic Pasteur pipette and wash in RPMI 1640 medium (or BSS). (iv) If the neutrophil pellet appears red due to contamination by erythrocytes, perform a hypotonic lysis, as described in steps (vi) and (vii) above. (v) The purity, viability, and number of neutrophils

DETECTION OF NADPH OXIDASE ACTIVITY The Cytochrome c Reduction Assay There are many assays that can be used to measure either the products of the respiratory burst of neutrophils or the substrate for the oxidase, molecular O2 . All methods have their own particular advantages and disadvantages. Because the primary product of the NADPH oxidase is O20, this would logically appear to be the most appropriate oxidant to measure. However, it must be remembered that NADPH oxidase activity can be activated intracellularly, either within phagoly-

FIG. 4. Purification of neutrophils by centrifugation on NIM. For details, see the text.

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sosomes (Fig. 5A) or within other, as yet undefined, intracellular sites. Thus, if these latter activities are to be measured, then the probe used to measure oxidase activity must be able to gain access to these intracellular sites. Unfortunately, the most specific and commonly used probe to detect the formation of O20, namely ferri-cytochrome c, is a large molecule and is membrane impermeable. Insufficient quantities of cytochrome c are co-ingested with phagocytic particles during phagocytosis to allow for the intracellular measurement of O20 production. Thus, cytochrome c can measure only extracellular O20 production (Fig. 5B). Because O20 itself is charged and can only cross membranes when a specific carrier or transport system is present, the cytochrome c assay measures only NADPH oxidase complexes activated on the plasma membrane. Principle This assay (7) is based upon the fact that O20 is capable of reducing ferri (oxidized)-cytochrome c to the reduced form of the cytochrome, which has a characteristic absorption maximum of 550 nm (Fig. 6). The scavenging enzyme, superoxide dismutase (SOD), has a higher affinity for O20 than does cytochrome c. Thus, when SOD is added to a reaction mixture of cytochrome c and activated neutrophils, no absorption increase at 550 nm is observed because the O20 preferentially reacts with SOD rather than cytochrome c. Thus, the rate of O20 secretion by activated neutrophils is taken as the rate of SOD-inhibitable cytochrome c reduction. If the rate of O20 exceeds its rate of ‘‘trapping’’ by cytochrome c, then H2O2 can be formed (Eq. [2]) and H2O2 can

FIG. 6. The cytochrome c reduction assay for the detection of O0 2. For details, see the text.

reoxidise the reduced cytochrome c, giving an apparently low rate of cytochrome c reduction. For this reason, catalase is sometimes added to assay mixtures to degrade this H2O2 . Required Spectrophotometer (preferably double beam); 7.5 mM ferri-cytochrome c (e.g., horse heart) in RPMI 1640 medium or BSS; 6 mg ml01 superoxide dismutase; 1 mg ml01 catalase; RPMI 1640 medium or BSS. Method (i) To the sample cuvette, add 10 ml cytochrome c solution (final concentration 75 mM), 5 1 105 neutrophils, and RPMI 1640 medium (or BSS) to a total volume of 0.99 ml. If required, catalase may be added at 2 mg ml01. (ii) To the reference cuvette, add the same assay components plus 5 ml SOD suspension (final concentration 30 mg ml01). (iii) Follow the baseline rate of increase in absorption at 550 nm (should be zero in nonactivated cells). (iv) Add stimulus (total volume of 10 ml) to each cuvette, mix thoroughly, and follow the absorption increase at 550 nm. (v) Measure the rate of absorption increase per minute (DA/min). The micromolar absorption coefficient of reduced cytochrome c is 19.1, and so the rate of O20 secretion is calculated as

FIG. 5. Intracellular and extracellular generation of reactive oxidants by activated neutrophils. The NADPH oxidase may be activated on the membranes of phagocytic vesicles (in A) or on the membranes of other, as yet undefined, structures. Such intracellular activity can only be detected using probes that freely permeate the neutrophil membrane. (B) NADPH oxidase components on the plasma membrane are activated so that reactive oxygen metabolites are secreted.

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DA/min 1 1000 , 19.1 which gives a rate of O20 production by the suspension (5 1 105 cells) in micromolar per minute. Rates are usually calculated as nanomoles O20 secreted per mi-

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nute per 106 cells. A continuous trace of cytochrome c reduction can be converted to changes in the rate of O20 secretion as a function of time after stimulation. This can be achieved automatically by measuring the first derivative of the change in absorbance values or manually by measuring the rate of absorbance change at different times after stimulation. Modifications The assay can be modified for use with a single beam spectrophotometer, and the SOD sensitivity can be checked by addition of SOD prior to or following the addition of stimulus. Alternatively, a discontinuous assay may be performed. In this way, after a fixed time interval following the addition of stimulus, the cells are separated from the incubation medium by low-speed centrifugation, and the absorption of the supernatant is measured at 550 nm. For such an assay, nonstimulated controls must be used to obtain a baseline or ‘‘blank’’ absorption value, and SOD-containing controls must also be included. The basic assay is also suitable for use in a microtiter plate format. The wells should contain the same concentrations as reactants as described above in a total volume of 200 ml. This microplate assay may be discontinuous or may provide kinetic data if the plates are read at fixed time intervals after stimulation. Control wells should again be set up containing (i) no cells, (ii) no stimulus, or (iii) cells / stimulus / SOD. The plate reader must be fitted with appropriate filters. Advantages The assay is specific for the production of O20 and thus is often the most appropriate assay to measure NADPH oxidase activity. It can measure O20 secretion from intact neutrophils or NADPH oxidase reconstituted in a cell-free system. Routinely available spectrophotometers usually provide sufficient sensitivity to detect oxidase activity in 105 –106 neutrophils. It is suitable for semiautomation when used in microtiter plate formats. Disadvantages Cytochrome c cannot penetrate the neutrophil plasma membrane and is not internalized at sufficiently high concentrations during phagocytosis to measure intracellular NADPH oxidase activity. Thus, the cytochrome c reduction assay is specific for the secretion of O20 during the respiratory burst, arising from activation of NADPH oxidase molecules on the plasma membrane. It cannot measure intracellular NADPH oxidase activity. Oxygen Uptake Principle Molecular O2 (dioxygen) can be measured using conventional electrodes, such as the Clark-type electrode

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chamber. These electrodes generally utilize a platinum cathode and a silver anode, which are bathed in a solution of electrolyte such as saturated KCl solution and separated from the cell suspension via an O2-permeable membrane (e.g., Teflon). O2 may then diffuse from the cell suspension and is reduced at the cathode to generate a current. A voltage of 0.5–0.8 V is applied across the electrodes, and so the current generated at the cathode is proportional to the O2 tension in the cell suspension. The reactions at the cathode are O2 / 4H/ / 4e r 2H2O. Thus, four electrons are generated at the anode, which then reduce a molecule of O2 at the cathode. The cell suspension must be constantly stirred because O2 is consumed at the cathode; without stirring, artifactually high rates of O2 consumption will be recorded. Furthermore, because both the O2 solubility in biological solutions and the rate of O2 diffusion are temperaturedependent, the whole chamber must be temperaturecontrolled, usually by means of a water-jacket circulating water at 377C. The system is ‘‘closed’’ to the atmosphere, i.e., gaseous O2 in the atmosphere cannot diffuse into the cell suspension. Thus, as O2 is utilized by the cell suspension, the O2 tension in the suspension decreases. The rate of O2 decrease recorded at the electrodes is taken as the rate of O2 uptake by the cells. Required Clark-type O2 electrode plus potentiometric chart recorder (or other recording device); Air-saturated water at 377C; Sodium dithionite crystals; Microsyringes (Hamilton type). Method (i) Add approximately 3 ml of prewarmed, air-saturated water (which contains 210 mM O2 at 377C) to the chamber to calibrate the electrode. (Note: do not insert the stopper cap). Adjust the output to 100% O2 using the gain (sensitivity) control. (ii) Set the 0% O2 by adding a few crystals of the reductant, sodium dithionite, to the water and adjusting the zero control. At 0% O2 , there should be no current flowing through the electrodes. (iii) Aspirate the water to waste and thoroughly wash the chamber several times. (iv) Add prewarmed, aerated RPMI 1640 medium (or BSS) and neutrophil suspension to a final concentration of 1 1 106 to 2 1 106 cells ml01 in a total volume of 2 ml. (v) Insert the stopper cap, taking care to avoid trapping air bubbles in the suspension, which will give erroneous readings. Adjust the screw ring so that the suspension is clearly visible in the injection inlet.

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(vi) Measure the electrode output for 2–3 min until a steady rate of O2 uptake is recorded. In unstimulated neutrophils this should be low. (vii) Add stimulus (or inhibitor) to the suspension via the injection inlet, but avoid introducing air bubbles. Readjust the level of suspension in the injection inlet by adjusting the screw ring. (viii) Record the change in rate of O2 consumption. The electrode measures changes in O2 tension in the suspension, which are proportional to the O2 consumption of the cells. The rate of O2 consumption is the first derivative of the continuous measurement of O2 tension, which may be calculated automatically by some measuring devices or calculated manually. Thus, a maximal rate of O2 uptake or else changes in the rate of O2 consumption as a function of time after stimulation may be measured. The rate of O2 uptake is usually expressed as micromolar O2 consumed per minute per 106 cells. Advantages The technique is simple, requires fairly inexpensive apparatus, and measures a defined molecule, namely molecular O2 . Because neutrophils (unlike monocytes) possess no functional mitochondria, the assay measures the activity of the respiratory burst without interference from mitochondrial O2 uptake. O2 freely diffuses into neutrophils, and so the assay measures O2 uptake via NADPH oxidase activity that may occur either on the plasma membrane or within intracellular sites. However, at low O2 tensions, O2 diffusion into such intracellular sites (e.g., within phagolysosomes) may become limited via O2 gradients that may occur across the cell. Disadvantages The technique requires fairly large numbers of neutrophils (compared to other techniques) and is not suited to automation: the electrode must be used exclusively for the measurement of a single sample for the duration of the experiment. Some microelectrodes can utilize smaller volumes and hence fewer neutrophils, but do not overcome the time limitation. Care must be taken to avoid damage to the membrane that may occur during injection of stimuli/inhibitors. Some agents can permeate the membrane and poison the electrodes, and some agonists/inhibitors may stick to the membrane of the reaction vessel. The technique measures all O2 consumed by cells, which in the case of the neutrophil is mostly (but not exclusively) due to NADPH oxidase activity. Furthermore, some oxidative reactions (e.g., Eqs. [2], [5], and [6]) generate O2 , and these may thus result in apparently low rates of O2 consumption. H2O2 Production Principle H2O2 is generated by activated neutrophils via Eq. [2] and utilized by myeloperoxidase in Eq. [3]. Con-

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sumption of H2O2 via the metal-catalyzed Haber– Weiss reaction via Eq. [6] (if it occurs within activated neutrophils) is likely to be extremely low and probably does not affect measurements. Many assays that are specific for the detection of H2O2 exist, and these are commonly based on fluorescence assays. Thus, H2O2 generated by activated neutrophils is used (in the presence of a peroxidase) to oxidize a substrate whose fluorescence properties change upon oxidation. Here, two assays are described. The first of these uses scopoletin as the fluorescence indicator molecule (65). This molecules loses its fluorescence as it becomes oxidized, with oxidation requiring the presence of an exogenous peroxidase, horseradish peroxidase. Because myeloperoxidase consumes H2O2 (Eq. [3]), apparent rates of H2O2 secretion will be low unless a myeloperoxidase inhibitor that does not affect horseradish peroxidase (e.g., sodium azide) is used. The second method described utilizes 2*,7*-dichlorofluorescin (66–68), which exhibits increased fluorescence upon oxidation and utilizes cellular peroxidase, i.e., myeloperoxidase. (a) SCOPOLETIN ASSAY Required Spectrofluorometer, set at 350-nm excitation, 460nm emission; 1 mM scopoletin; 1 mg ml01 horseradish peroxidase. Method (i) To a fluorometer tube of 3-ml capacity add 12 ml scopoletin solution (final concentration 4 mM), 15 ml horseradish peroxidase solution (final concentration 5 mg ml01), and 3 1 106 to 6 1 106 neutrophils (final concentration 1 1 106 to 2 1 106 ml01). Add prewarmed RPMI 1640 medium (or BSS) to 2.97 ml. (ii) Adjust the output of the fluorometer to 100%. (iii) Add 30 ml stimulus and measure the decrease in fluorescence on a chart recorder. (iv) Prepare a calibration curve to allow for the conversion of these fluorescence changes to increases in H2O2 production. Prepare a tube containing all reactants plus cells but do not add stimulus. Add aliquots of known concentrations of H2O2 (or ethyl peroxide). Plot ‘‘Decrease in Fluorescence’’ on the vertical (y or ordinate) axis as a function of ‘‘H2O2 concentration’’ on the horizontal (x or abscissa) axis. (v) Convert the fluorescence decreases observed with activated neutrophils obtained in (iii) to nanomoles of H2O2 produced. Advantages The assay is fairly specific for the detection of H2O2 production and is suitable for use on microtiter plate formats using a fluorescence plate reader.

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Disadvantages Neither scopoletin nor horseradish peroxidase can penetrate the neutrophil plasma membrane, and so the assay can detect only H2O2 that is secreted from activated neutrophils. This H2O2 may arise from the activation of NADPH oxidase molecules on the plasma membrane or may arise from the diffusion of internally generated H2O2 from intracellular sites to the outside of the cell. This diffusion is considerably enhanced if neutrophil enzymes that consume H2O2 (e.g., myeloperoxidase, catalase) are poisoned by the addition of sodium azide (at 1 mM). (b) 2*,7*-DICHLOROFLUORESCIN An alternative method to detect H2O2 production by activated neutrophils is to use 2*,7*-dichlorofluorescin. This compound has low fluorescence, but when oxidized by H2O2 and a peroxidase it is converted to 2*,7*-dichlorofluorescein, which has high fluorescence. Moreover, dichlorofluorescin diacetate is membrane-permeable and freely diffuses into neutrophils. Once internalized, cytoplasmic esterase activity cleaves this molecule into 2*,7*-dichlorofluorescin, which becomes trapped within the neutrophil. Upon activation, intracellular H2O2 production, together with myeloperoxidase, then oxidizes the trapped indicator into the fluorescent molecule, 2*,7*-dichlorofluorescein. This assay thus provides a means to detect intracellularly generated H2O2 , which can be measured using a fluorometer or a FACS analyzer. Use of the latter detection system allows for the quantitation of fluorescence signals in individual cells and thus gives a measure of the heterogeneity of oxidase responses in neutrophil populations (67, 68). Alternatively, fluorescence of individual cells can be ‘‘viewed’’ using a fluorescence microscope allowing (with suitable cameras and software systems) both quantitation of signals in individual cells and subcellular localization of the fluorescence. Required Fluorometer, set at 500-nm excitation, 530-nm emission or FACS analyzer; 2*,7*-dichlorofluorescin diacetate solution (1 mM in ethanol); Fixative (such as Becton–Dickinson FACS lysing solution, which lyses erythrocytes but fixes leukocytes). Method (i) Incubate neutrophils at 2 1 106 cells ml01 (in RPMI 1640 medium of BSS) at 377C for 30 min with 5 mM 2*,7*-dichlorofluorescin diacetate solution. (ii) After loading, wash the cells three times with RPMI 1640 medium (or BSS) and resuspend at 2 1 106 cells ml01. If measuring fluorescence using a fluorometer, place the suspension in a fluorometer cuvette and

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add stimulus. Follow the increase in fluorescence using a chart recorder. (iii) If using a FACS analyzer, stimulate the cell suspension and at time intervals after stimulation fix the cells by the addition of FACS lysing solution. Analyze by FACS. Advantages The assay is fairly specific for the detection of H2O2 and can measure intracellular H2O2 production. Analysis of fluorescence by FACS can measure heterogeneity of oxidative responses in individual neutrophils, while analysis by fluorescence microscopy can also localize the fluorescent signal. Disadvantages For the intracellular detection of H2O2 , the indicator utilizes myeloperoxidase. Thus, the fluorescent signal generated may reflect the intracellular site of myeloperoxidase, rather than the site of H2O2 generation, which may diffuse through the cell. Chemiluminescence Principle In the early 1970s, it was discovered that activated neutrophils emit photons to generate light or chemiluminescence (reviewed in 69). This light emission may be detected using a scintillation counter (operating in the ‘‘out of coincidence’’ mode) or by specialized luminometers utilizing high-sensitivity photomultiplier tubes. Furthermore, this ‘‘native’’ chemiluminescence may be considerably ‘‘amplified’’ by inclusion of probes such as luminol or lucigenin (Fig. 7). The molecular species of oxidant responsible for unamplified chemiluminescence of activated neutrophils is not known with certainty, but candidate molecules include 1O2 (which may be generated by myeloperoxidase in Eq. [5]), reaction products of heme with H2O2 , and carbonyl compounds in excited states. The products of the respiratory burst can oxidize the lumigenic probes, luminol and lucigenin. Cell-free experiments show that luminol can be oxidized by mixtures of H2O2 and HOCl or by mixtures of H2O2 and peroxidase (such as myeloperoxidase). Thus, in activated neutrophils, luminol chemiluminescence measures the combined activities of the NADPH oxidase plus myeloperoxidase. Inhibitors of either the NADPH oxidase (such as diphenylene iodonium; 28) or myeloperoxidase (e.g., salicylhydroxamic acid; 70) can completely inhibit the luminol chemiluminescence of activated neutrophils. However, modification of the basic assay (as described below) can overcome this dependency on myeloperoxidase activity. Lucigenin chemiluminescence is believed to measure O20 (or H2O2) secretion (independently of the activity of myeloperoxidase), although some evidence in the literature suggests that

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the oxidant is not O20 per se but an oxidant derived from O20. Thus, lucigenin chemiluminescence is at the very least, an indirect measure of O20 secretion (71). Many phagocyte researchers do not use the chemiluminescence assay because the kinetics of luminol chemiluminescence of neutrophils activated by, e.g., fMet-Leu-Phe do not match those of O20 secretion (detected by the cytochrome c reduction assay) or H2O2 secretion (detected by the scopoletin assay) (Fig. 8). For many years this lack of correlation was something of a puzzle until it was appreciated (72–74) that luminol chemiluminescence measures both intracellular and extracellular reactive oxidant production. Luminol is a small uncharged molecule and can freely permeate the neutrophil plasma membrane; lucigenin, on the other hand, is larger and membrane impermeable and thus detects only the secretion of reactive oxidants. This initial phase of luminol chemiluminescence following fMet-Leu-Phe stimulation (and the secreted O20 and H2O2 production) is completely abolished when extracellular scavengers of reactive oxidants such as SOD, catalase, and methionine (to scavenge HOCl) are added to incubation mixtures (75). On the other hand, the later luminol chemiluminescence is unaffected by the extracellular scavengers and is intracellular in nature. The kinetics of fMet-Leu-Phe-stimulated luminol chemiluminescence is largely matched by the kinetics of activated O2 uptake, which is also capable of measuring both intracellular and extracellular oxidase activity. The luminol chemiluminescence assay is thus extremely useful for measuring reactive oxidant production in intracellular sites, e.g., in phagolysosomes.

Modification of the basic method to detect luminol chemiluminescence (76) overcomes the dependency of the assay on myeloperoxidase and thus provides a more accurate representation of NADPH oxidase activity. Furthermore, manipulation of assay conditions can clearly distinguish between intracellular and extracellular oxidant production. Because of the exquisite sensitivity of the assay, luminol chemiluminescence can be used to measure reactive oxidant production by neutrophils in small quantities (5–10 ml) of unfractionated whole blood (77). Required Luminometer (or scintillation counter operating in the ‘‘out of coincidence’’ mode); Luminol, stock solutions A (100 mM in DMSO diluted to 10 mM with RPMI 1640) and B (10 mM in DMSO diluted to 1 mM in RPMI 1640); Lucigenin, stock solution of 2.5 mM in RPMI 1640 medium or BSS. Method (a) Whole blood assay. This procedure is performed as follows: (i) Add 10 ml whole, heparinized blood to 0.97 ml of prewarmed (377C) RPMI 1640 medium. (ii) Add 10 ml of luminol solution A (final concentration 100 mM). (iii) Add 10 ml stimulus and measure chemiluminescence. Note: neutrophils in unfractionated blood generate only extremely low levels of reactive oxidants in response to agents such as fMet-Leu-Phe, and this usu-

FIG. 7. Structures of (A) luminol and (B) lucigenin.

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ally cannot be detected by this method. In order to detect luminol chemiluminescence in whole blood either stimulate the suspension with 0.1 mg/ml PMA or prime the neutrophils before the addition of 1 mM fMetLeu-Phe. To prime the cells, add 10 ml whole blood to 0.965 ml RPMI 1640 medium together with 5 ml (containing 50 U) of GM-CSF. Incubate at 377C for 1 h prior to stimulation by 1 mM fMet-Leu-Phe. (b) Purified neutrophils. (i) Add 5 1 105 neutrophils to 0.975 ml prewarmed (377C) RPMI 1640 medium. (ii) Add 10 ml of luminol solution B (final concentration 10 mM) or 10 ml of the lucigenin solution (final concentration 25 mM). (iii) Add stimulus and record chemiluminescence. Modifications Addition of 1 mM azide and horseradish peroxidase (4 U) to the suspension poisons myeloperoxidase and greatly enhances the chemiluminescence response due to H2O2 secretion. Thus, myeloperoxidase-independent secretion of oxidants is measured. Alternatively, addi-

FIG. 8. Kinetics of reactive oxidant production by activated neutrophils. In response to stimulation by fMet-Leu-Phe, the respiratory burst is biphasic. An initial phase of reactive oxidant secretion is followed by a second phase of intracellular production. These two phases of activity are detected using different assays.

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tion of catalase (2000 U) degrades extracellular H2O2 , and so only intracellular chemiluminescence is measured (76). Advantages The method is extremely sensitive and can measure reactive oxidant production by a few thousand neutrophils or by the neutrophils present in 5–10 ml of unfractionated blood. It is suitable for semiautomation, being adaptable to microtiter plate formats and measurement of light emission using a luminometric plate reader. It can measure both intracellular and extracellular reactive oxidant production and hence is one of the few methods (together with O2 uptake) that can measure oxidative events occurring within phagolysosomes. Disadvantages The luminol chemiluminescence assay measures the combined activities of the NADPH oxidase and myeloperoxidase. However, the dependency on myeloperoxidase can be overcome by modification of the basic assay, as described above. Electron Paramagnetic Resonance (EPR) Spectroscopy Principle Some of the oxidant species produced by activated neutrophils are free radicals, possessing an unpaired electron in their chemical structure. This unpaired electron often confers high chemical reactivity on the free radical, but also the property of paramagnetic resonance. Thus, free radicals can be detected by the technique of EPR spectroscopy. However, because free radicals are unstable, they must be ‘‘trapped’’ prior to detection by this technique. The reaction of free radicals with these traps generates characteristic EPR spectra that aid in the detection and quantitation of the free radical species produced. Because EPR spectroscopy can be used to detect •OH production in biological systems, it has been used to detect the formation of this important molecule by activated neutrophils (78–81). However, EPR spectroscopy is not generally used to measure the respiratory burst of neutrophils for several reasons. First, EPR spectrometers are not generally available in most biochemical laboratories. Second, the technique is relatively insensitive, and large quantities of neutrophils are required to generate reliable spectra. Third, there are some problems with the interpretation of spectra, as will be described below. A commonly used free radical trap is the molecule 5,5-dimethyl-1-pyrroline-1-oxide (DMPO), which can react with O20 to form the adduct DMPO-OOH and with • OH to form the adduct DMPO-OH. Both of these adducts have characteristic EPR spectra. However, because DMPO-OH can be formed via the decomposition of DMPO-OOH, great care must be taken with inter-

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pretation of data. Furthermore, because •OH formation via the Haber–Weiss reaction requires both O20 and H2O2 (Eq. [6]), DMPO-OH formation must be shown to be catalase-sensitive if this is the proposed route of formation of this free radical. There is some evidence in the literature in which this technique is used that •OH may be formed by activated neutrophils. Such putative •OH formation is regulated by the extent of degranulation of neutrophil enzymes. For example, myeloperoxidase release will decrease the rate of •OH formation because this enzyme utilizes H2O2 , one of the reactants of the Haber–Weiss reaction, while lactoferrin release will also decrease •OH formation because it can chelate iron, the catalyst for this formation. However, there is some doubt as to the physiological significance of these experiments because •OH formation can be detected only when neutrophil suspensions are supplemented with high concentrations of iron salts, in the form of iron–diethyletriaminepentaacetic acid. In the absence of this iron supplement, no •OH formation is detected.

CONCLUSIONS There are many methods that are capable of measuring the respiratory burst of neutrophils or other phagocytes. Where possible, it is recommended that the cytochrome c reduction assay be used, because this assay is the most clearly defined in terms of its specificity. However, this assay can detect only O20 secreted from activated cells. If intracellular reactive oxidant production is to be measured, then the luminol chemiluminescence method may be used. This assay provides excellent sensitivity (even measuring neutrophil function in whole blood), but care is required in the interpretation of results.

ACKNOWLEDGMENTS The financial support of the Arthritis and Rheumatism Council, North West Cancer Research Fund, and the Medical Research Council is gratefully acknowledged.

REFERENCES 1. Edwards, S. W. (1994) Biochemistry and Physiology of the Neutrophil, Cambridge Univ. Press, New York. 2. Edwards, S. W. (1991) in Calcium, Oxygen Radicals and Tissue Damage (C. J. Duncan, Ed.), pp. 35–76, Cambridge Univ. Press, Cambridge, UK. 3. Baldridge, C. W., and Gerard, R. W. (1933) Am. J. Physiol. 103, 235–236.

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4. Sbarra, A. J., and Karnovsky, M. L. (1959) J. Biol. Chem. 234, 1355–1362. 5. Holmes, B., Quie, P. G., Windhorst, D. B., and Good, R. A. (1966) Eur. J. Cancer 26, 1115–1118. 6. Holmes, B., Page, A. R., and Good, R. A. (1967) J. Clin. Invest. 46, 1422–1432. 7. Babior, B. M., Kipnes, R. S., and Curnutte, J. T. (1973) J. Clin. Invest. 52, 741–744. 8. Segal, A. W. (1989) Biochem. Soc. Trans. 17, 427–434. 9. Segal, A. W., and Abo, A. (1993) Trends Biochem. Sci. 18, 43– 47. 10. Segal, A. W., and Jones, O. T. G. (1978) Nature 276, 515–517. 11. Segal, A. W., Jones, O. T. G., Webster, D., and Allison, A. C. (1978) Lancet 2, 446–449. 12. Cross, A. R., Harper, A. M., and Segal, A. W. (1981) Biochem. J. 194, 599–606. 13. Cross, A. R., Higson, F. K., Jones, O. T. G., Harper, A. M., and Segal, A. W. (1982) Biochem. J. 204, 479–485. 14. Harper, A. M., Dunne, M. J., and Segal, A. W. (1984) Biochem. J. 219, 519–527. 15. Royer-Pokora, B., Kunkel, L. M., Monaco, A. P., Goff, S. C., Newburger, P. E., Baehner, R. L., Cole, F. S., Curnutte, J. T., and Orkin, S. H. (1986) Nature 322, 32–38. 16. Dinauer, M. C., Orkin, S. H., Brown, R., Jesaitis, A. J., and Parkos, C. A. (1987) Nature 327, 717–720. 17. Dinauer, M. C., Pierce, E. A., Bruns, G. A. P., Curnutte, J. T., and Orkin, S. H. (1990) J. Clin. Invest. 86, 1729–1737. 18. Parkos, C. A., Dinauer, M. C., Walker, L. E., Allen, R. A., Jesaitis, A. J., and Orkin, S. H. (1988) Proc. Natl. Acad. Sci. USA 85, 3319–3323. 19. Borregaard, N., and Tauber, A. I. (1984) J. Biol. Chem. 259, 47– 52. 20. Bjerrum, O. W., and Borregaard, N. (1989) Eur. J. Haematol. 43, 67–77. 21. Jesaitis, A. J., Buescher, E. S., Harrison, D., Quinn, M. T., Parkos, C. A., Livesey, S., and Linner, J. (1990) J. Clin. Invest. 85, 821–835. 22. Borregaard, N., Kjeldsen, L. K., Sengeløv, H., Bastholm, L., Nielson, M. H., and Bainton, D. F. (1993) Eur. J. Haematol. 51, 187– 198. 23. Garcia, R. C., and Segal, A. W. (1984) Biochem. J. 219, 233– 242. 24. Higson, F. K., Durbin, L., Pavlotsky, N., and Tauber, A. I. (1985) J. Immunol. 135, 519–524. 25. Babior, B. M., and Kipnes, R. S. (1977) Blood 50, 517–524. 26. Bellavite, P., Cross, A. R., Serra, M. C., Davoli, A., Jones, O. T. G., and Rossi, F. (1983) Biochim. Biophys. Acta 746, 40– 47. 27. Cross, A. R., Jones, O. T. G., Garcia, R., and Segal, A. W. (1982) Biochem. J. 208, 759–763. 28. Cross, A. R., and Jones, O. T. G. (1986) Biochem. J. 237, 111– 116. 29. Segal, A. W., West, I., Wientjes, F., Nugent, J. H. A., Chavan, A. J., Haley, B., Garcia, R. C., Rosen, H., and Scrace, G. (1992) Biochem. J. 284, 781–788. 30. Bromberg, Y., and Pick, E. (1984) Cell. Immunol. 88, 213–221. 31. Bromberg, Y., and Pick, E. (1985) J. Biol. Chem. 60, 13539– 13545. 32. Curnutte, J. T. (1985) J. Clin. Invest. 75, 1740–1743. 33. Heyneman, R. A., and Vercauteren, R. E. (1984) J. Leukocyte Biol. 36, 751–759. 34. Aviram, I., and Sharabani, M. (1989) Biochem. J. 261, 477–482.

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MEASUREMENT OF THE NADPH OXIDASE 35. Curnutte, J. T., Berkow, R. L., Roberts, R. L., Shurin, S. B., and Scott, P. J. (1988) J. Clin. Invest. 81, 606–610. 36. Curnutte, J. T., Scott, P. J., and Babior, B. M. (1989) J. Clin. Invest. 83, 1236–1240. 37. Clark, R. A., Malech, H. L., Gallin, J. I., Nunoi, H., Volpp, B. D., Pearson, D. W., Nauseef, W. M., and Curnutte, J. T. (1989) N. Engl. J. Med. 321, 647–652. 38. Nunoi, H., Rotrosen, D., Gallin, J. I., and Malech, H. L. (1988) Science 242, 1298–1301. 39. Volpp, B. D., Nauseef, W. M., and Clark, R. A. (1988) Science 242, 1295–1297. 40. Leto, T. L., Lomax, K. J., Volpp, B. D., Nunoi, H., Sechler, J. M. G., Nauseef, W. M., Clark, R. A., Gallin, J. I., and Malech, H. L. (1990) Science 248, 727–730. 41. Volpp, B. D., Nauseef, W. M., Donelson, J. E., Moser, D. R., and Clark, R. A. (1989) Proc. Natl. Acad. Sci. USA 86, 7195–7199. 42. Lomax, K. J., Leto, T. L., Nunoi, H., Gallin, J. I., and Malech, H. L. (1989) Science 245, 409–412. 43. Abo, A., and Pick, E. (1991) J. Biol. Chem. 266, 23577–23585. 44. Abo, A., Pick, E., Hall, A., Totty, N., Teahan, C. G., and Segal, A. W. (1991) Nature 353, 668–670. 45. Eklund, E. A., Marshall, M., Gibbs, J. B., Crean, C. D., and Gabig, T. G. (1991) J. Biol. Chem. 266, 13964–13970. 46. Knaus, U. G., Heyworth, P. G., Kinsella, T., Curnutte, J. T., and Bokoch, G. M. (1992) J. Biol. Chem. 267, 23575–23582. 47. Abo, A., Boyhan, A., West, I., Thrasher, A. J., and Segal, A. W. (1992) J. Biol. Chem. 267, 16767–16770. 48. Wientjes, F. B., Hsuan, J. J., Totty, N. F., and Segal, A. W. (1993) Biochem. J. 296, 557–561. 49. Agner, K. (1941) Acta Scand. 2(Suppl. 8) 1–62. 50. Klebanoff, S. J. (1968) J. Bacteriol. 95, 2131–2138. 51. Halliwell, B., and Gutteridge, J. M. C. (1985) Free Radicals in Biology and Medicine. p. 346, Clarendon, Oxford. 52. Halliwell, B., and Gutteridge, J. M. C. (1984) Biochem. J. 219, 1–9. 53. McCall, C. E., DeChatelet, L. R., Cooper, M. R., and Shannon, C. (1973) J. Infect. Dis. 127, 26–33. 54. McCall, C. E., Bass, D. A., DeChatelet, L. R., Link, A. S. J., and Mann, M. (1979) J. Infect. Dis. 140, 277–286. 55. Bender, J. G., McPhail, L. C., and van Epps, D. E. (1983) J. Immunol. 130, 2316–2323. 56. Edwards, S. W., Holden, C. S., Humphreys, J. M., and Hart, C. A. (1989) FEBS Lett. 256, 62–66.

/ 6707$$281v

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57. Edwards, S. W., Say, J. E., and Hughes, V. (1988) J. Gen. Microbiol. 134, 37–42. 58. Humphreys, J. M., Rugman, F., Davies, J. M., Mimnagh, P., Hart, C. A., and Edwards, S. W. (1991) J. Clin. Lab. Immunol. 34, 55–61. 59. Edwards, S. W., Watson, F., MacLeod, R., and Davies, J. M. (1990) Biosci. Rep. 10, 393–401. 60. Edwards, S. W. (1995) Trends Biochem. Sci. 20, 362–367. 61. Watson, F., Lowe, G. M., Robinson, J. J., Galvani, D. W., and Edwards, S. W. (1994) Biosci. Rep. 14, 91–102. 62. Bøyum, A. (1968) Scand. J. Clin. Lab. Invest. 97(Suppl) 77–89. 63. Watson, F., Robinson, J. J., and Edwards, S. W. (1992) Biosci. Rep. 12, 123–133. 64. Homburg, C. H. E., de Haas, M., von dem Borne, A. E. G. K., Verhoeven, A. J., Reutingsperger, C. P. M., and Roos, D. (1995) Blood 85, 532–540. 65. Root, R. K., Metcalf, J., Oshino, N., and Chance, B. (1975) J. Clin. Invest. 55, 945–955. 66. Bass, D. A., Parce, J. W., DeChalelet, L. R., Szejda, P., Seeds, M. L., and Thomas, M. (1983) J. Immunol. 130, 1910–1917. 67. Patel, A. K., Hallett, M. B., and Campbell, A. K. (1987) Biochem. J. 248, 173–180. 68. Daniels, R. H., Elmore, M. A., Hill, M. E., Shimizu, Y., Lackie, J. M., and Finnen, M. J. (1994) Immunology 82, 465–472. 69. Allen, R. C. (1986) Methods Enzymol. 133, 449–493. 70. Davies, B., and Edwards, S. W. (1989) Biochem. J. 258, 801– 806. 71. Edwards, S. W. (1987) J. Clin. Lab. Immunol. 22, 35–39. 72. Dahlgren, C. (1987) Biochim. Biophys. Acta 930, 33–38. 73. Dahlgren, C., Aniansson, H., and Magnusson, K.-E. (1985) Infect. Immun. 47, 326–328. 74. Briheim, G., Stendahl, O., and Dahlgren, C. (1984) Infect. Immun. 45, 1–5. 75. Watson, F., Robinson, J. J., and Edwards, S. W. (1991) J. Biol. Chem. 266, 7432–7439. 76. Dahlgren, C. (1991) J. Biolumin. Chemilumin. 6, 29–34. 77. Ristola, M., and Repo, H. (1989) APMIS 97, 503–512. 78. Britigan, B. E., Cohen, M. S., and Rosen, G. M. (1987) J. Leukocyte Biol. 41, 349–362. 79. Britigan, B. E., Hassett, D. J., Rosen, G. M., Hamill, D. R., and Cohen, M. S. (1989) Biochem. J. 264, 447–455. 80. Cohen, M. S., Britigan, B. E., Hassett, D. J., and Rosen, G. M. (1988) Rev. Infect. Dis. 10, 1088–1096. 81. Pou, S., Cohen, M. S., Britigan, B. E., and Rosen, G. M. (1989) J. Biol. Chem. 264, 12299–12302.

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