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Grow human hepatoma HepG2 cells in 24-well plates in minimal essential media, wash with serum-free media, and further incubate in serum-free media containing 1-10 pM DCFDA or an equivalent volume of vehicle (dimethyl sulfoxide [DMSO]) for 30 min. The final concentration of DCFDA and incubation times may be cell specific and require preliminary investigations for optimization.

After this time, wash cells with warm PBS (37°C) and then incubate for 5 min in PBS.

Determine hypochlorite concentration at ^290 nm at pH 12 (e = 350/M:/cm) and store on ice diluted in ice cold in ultra high purity water (17). Immediately before use, the hypochlorite solution is diluted in warm PBS and gently poured onto the HepG2 cells. To "quench" hypochlorite, 1 mM methionine solution can be added for 1-2 min before hypochlorite addition.

4. Measure fluorescence as a function of time or hypochlorite addition.

Typical data obtained are shown in Fig. 3.2.

Untreated HOCI HOCI + methionine

Untreated HOCI HOCI + methionine

Fig. 3.2. Determination of intracellular 'ROI' produced by human hepatoma HepG2 cells exposed to the neutrophil oxidant, hypochlorite by fluorescence microscopy, and DCFDA.

2.2.2.3. Comments

The use of DCFH for the "specific" measurement of H2 O2 is associated with a number of problems (22). First, it has been reported that the H2O2-dependent oxidation of DCFH to DCF occurs slowly in the absence of ferrous iron and can be completely inhibited by deferoxamine (23). Second, because peroxidases are capable of inducing DCFH oxidation in the absence of H2 O2 (24), variations in cellular peroxidase activity may influence rates of DCF formation, and cellular fluorescence DCFH itself may also auto-oxidize to form hydrogen peroxide and the reaction of DCFH with peroxidase forms DCF radicals with the subsequent generation of O2*-, suggesting that the use of DCFH to measure ROI may be problematic (25). Third, numerous other substances are capable of directly inducing DCF formation in the absence of H2 O2, including onoo~ and HOCl . Because of the multiple pathways that can lead to DCF fluorescence, and the inherent uncertainty relating to endogenous versus artifactual oxidant generation, this assay may best be applied as a qualitative marker of cellular oxidant stress rather than a precise indicator of rates of H2O2 formation.

2.2.3. Protocol 3: Estimation of Cellular O2'- Formation With Hydroethidine and FACS Analysis: Redox Cycling With Menadione

Hydroethidine (HE) is cell permeable and reacts with O2*- to form ethidium (E), which in turn is believed to intercalate with DNA, providing nuclear fluorescence at an excitation wavelength of 520 nm and an emission wavelength of 610 nm (26). Bindokas and coworkers reported that oxidation of HE by O2*- was specific because the reaction did not occur in the presence of other ROI, including OH', H2O2, or ONOO" (27). Because of apparent selectivity, HE has frequently been used to detect intracellular o/-

2.2.3.1. Materials

1. HE (Molecular Probes).

3. FACS machine or fluorescence plate reader (excitation wavelength 520 nm and emission wavelength 610 nm).

1. Culture leukemic CEM suspension cells or HL60 suspension cells in RPMI with 10% FCS and supplements. Maintain cells in log-phase and keep them at a concentration between 2.5 and 5 x 105/mL.

2. After treatment (for example, with the redox cycling compound menadione, which generates O2*_), centrifuge cells gently to remove culture media and wash twice by centrifu-gation in PBS (1,000 rpm).

3. Suspend cells in PBS, by light vortexing, and load with 5-10 pM HE, incubating them for 15-20 min at 37°C.

4. Wash cells 2x in PBS (centrifuge for 1 min at 100^ and suspend in PBS containing 10 mM glucose).

5. Because this cell assay is a viable one, immediate FACS analysis is required with the use of appropriate software for analysis.

6. The FACS setting is FL-3 (PE) in log mode after cell debris have been electronically gated out.

2.2.4. Protocol 4: Estimation of Cellular O2'- Formation With HE and Fluorescence Microscopy: Induction by the Neutrophil Oxidant, Hypochlorous Acid

2.2.4.1. Materials

1. HE (Molecular Probes).

3. Sodium hypochlorite solution.

4. Methionine (Sigma).

5. Fluorescence microscope with rhodamine filters capable of excitation wavelength 520 nm and emission wavelength 610 nm.

1. Grow human hepatoma HepG2 cells in 24-well plates in minimal essential media and load with 5 pM dihydroergot-amine (DHE) or an equivalent volume of vehicle (DMSO) for 30 min. After this time wash cells with warm PBS (37°C) and then incubate for 5 min in PBS.

2. Determine the hypochlorite concentration at A290 nm at pH 12 (e = 350/M/cm) and store on ice diluted in ice cold in ultra high-purity water (17). Immediately before use, dilute the hypochlorite solution in warm PBS and gently pour onto the HepG2 cells. To "quench" hypochlorite, 1 mM methionine solution can be added for 1-2 min before the addition of hypochlorite.

3. Fluorescence is measured as a function of time or hypochlo-rite addition. Typical data obtained are shown in Fig. 3.3.

2.2.4.3. Comments The intracellular oxidation of HE to E by O2*_ has previously been analyzed with the use of flow cytometry and also by visualization of adherent neuronal cells and brain tissue with digital imaging microfluorometry (27). HE has also been used to study the respiratory burst in immune cells (26) and the redox state

Untreated HOCl HOCl + methionine

Untreated HOCl HOCl + methionine

Fig. 3.3. Determination of intracellular 'ROI' produced by human hepatoma HepG2 cells exposed to the neutrophil oxidant, hypochlorite by fluorescence microscopy, and DHE.

in tumor cells (28). To show the potential use of this probe, we have included an example of oxidation of HE to E by intracellular ROI generated by the neutrophil derived oxidant species, HOCl (Fig. 3.3). However, as with DCF, there are a number of problems associated with the use of HE as a quantitative marker of O2*- production (22).

First, the amount of E produced by the oxidation of HE decreases with increasing O2*- flux, which suggests to us that HE catalyses the dismutation of O2*-, which will underestimate the extent of O2*- production (29). HE also can be oxidized by a variety of heme proteins, including mitochondrial cytochromes, hemoglobin, and myoglobin (30). Most importantly, recent work has indicated that O2*- does not oxidize HE to E because it was found that O2*- generated by a variety of enzymatic and chemical systems (e.g., xanthine/xanthine oxidase, endothelial nitric oxide synthase, or potassium superoxide) oxidized HE to a fluorescent product (excitation, 480 nm; emission, 567 nm) that was different from E (31). The authors concluded that the reaction between O2*- and HE formed a fluorescent marker product that was not E. Although not entirely specific for O2*-, the method continues to be widely used for the "specific" determination of this radical species. We suggest that when HE is used for the determination of O2*- production, the combined use of pharmacological inhibitors of the mitochondrial ETC, together with the use of appropriate cell lines lacking mitochondrial DNA (i.e., without a functional ETC; generated as described in ref. 21) be used to clarify the subcellular source of the O2*- signal.

2.3. Chemiluminescence Luminol has been used as a detector of ROI formed during the

Procedures reaction of XO with hypoxanthine (HX) for several decades (32).

Light produced by luminol + XO/HX is directly proportional to the activity of XO, suggesting that this system could be used as a general test for oxidizing intermediates (32). Lucigenin (bis-N-methylacridinium) has also frequently been used for the

2.3.1. Protocols: Chemilu-mlnescence Characterization of ROI Generated by HX/XO and Phorbol Ester Stimulated White Blood Cells

2.3.1.1. Materials

2.3.1.2. Methods luminescent detection of the radical O2*- by activated phagocytes or the XO/HX system (33). More recently, several other compounds have been used for chemiluminescent detection of O2*-, including coelenterazine [2-(4-hydroxybenzyl)-6-(4-hydroxyphenyl)-8-benzyl-3,7-dihydroimidazo [1,2-a]pyrazin-3-one] and its analogs CLA (2-methyl-6-phenyl-3,7-dihydroimidazo[1,2-a]pyrazin-3-one) and MCLA [2-methyl-6-(4-methoxyphenyl)-3,7-dihydroimidazo[1,2-a]pyrazin-3-one].

Luminol and lucigenin are often used to detect the production of ROI by activated phagocytes, although they have also been used to determine ROI in other cell types (34).

2. Copper and zinc-containing superoxide dismutase (CuZn-SOD; Sigma-Aldrich, St. Louis, MO).

3. DMSO (Sigma-Aldrich).

4. Luminol (Sigma-Aldrich).

6. Ethylene diamine tetra-acetic acid (EDTA).

7. Catalase (CAT; Sigma-Aldrich).

A. Luminol-dependent characterization of ROI generated by XO/HX system

1. Add 100 pL of 1 mM HX in PBS to 50 pL of either CuZn-SOD (100 U/mL), CAT (4 x 103 U/mL), or DMSO (100 mM) and add 100 pL of chemiluminescence (CL) probe (1 mM luminol) to the test wells of a white 96-well microtiter plate (in triplicate).

2. Start the reaction by the addition of 50 pL of XO (0.25 U/ mL) using the automated luminometer dispenser.

3. Operate the luminometer in the integration mode for a time of 10 s with Ascent software (LabSystems, MN).

4. Express results as total integrated CL signal (relative light units; RLU).

5. These protocols may require substantial optimization to achieve maximum sensitivity and reliability of results.

B. Luminol-dependent characterization of ROI generated by white blood cells

1. Allow EDTA anticoagulated whole blood to sediment under gravity for 1 h at 37°C and remove the plasma and centrifuge at -400^ for 10 min to pellet out the white blood cells (WBCs).

2. Wash WBCs in PBS at 4°C and resuspend in PBS to a concentration of 2-5 x 106 cells/mL.

3. Add WBCs (100 pL) to a white 96-well microtitre plate in triplicate together with 50 pL of phorbol myristate acetate (PMA; 2 ng/mL) and 50 pL of test compounds including either CuZnSOD (100 U/mL), CAT (4 x 103 U/mL), or DMSO (100 mM).

4. Add 100 pL of 1 mM luminol giving a final total volume of 300 pL.

5. Operate the luminometer in the kinetic mode at 10-s intervals for a total of 90 min using Ascent software.

6. Use the antioxidants SOD, CAT, and DMSO to characterize relative proportions of O2*-, H2 O2 and OH 2 generated by PMA-stimulated WBCs.

7. Express the results either as the total integrated CL signal (RLU) or as RLUs per minute (RLU/min).

2.3.1.3. Comments Although luminol and lucigenin are both used for the determina tion of O2*-, the two reactions are different in that luminol requires univalent oxidation and lucigenin requires univalent reduction before they can react with O2*- and produce luminescence per se. In addition, they do not react with O'-(34, 35). A major problem with lucigenin is that the radical cation formed by its reduction can auto-oxidize and generate artifactual O2*-. Because of this reaction, it has been suggested that lucigenin can be used for assaying SOD activity but it should not be used for measuring O2*-(36). The extent to which this artifact can interfere with accurate measurement of O2*- by lucigenin is controversial (7), but it appears to be significant in some cell systems (35, 37-39). It has been suggested by some that by modulating the concentrations of lucigenin, it is possible to circumvent the problems associated with the overesti-mation of O2*-. Because it is not possible to predict the extent of redox cycling, however, this may be difficult to accurately determine (6). There are other problems associated with use of lucigenin. For example, the conversion of lucigenin to the radical by O2*- is not rapid and requires other cellular-reducing systems (e.g., XO, the mitochondrial electron transport chain, or the phagocyte NADPH oxidase), thus complicating data interpretation enormously (35, 37, 38, 40). There are similar problems with the use of luminol for the detection of radicals such as O2*- because the radical intermediate formed on oxidation also auto-oxidizes (6). Furthermore, luminol luminescence is also not a reliable indicator of O2*-, even when O2*- is involved in the reaction leading to light emission, because it can mediate O2*- formation by a variety of oxidants including ferricyanide, OCl- and XO/HX (41).

The problems of luminol and lucigenin associated with the artifactual generation of O2*- can be circumvented by the use of more specific alternative nonredox-cycling compounds for the determination of O2*- including coelenterazine, a luminophore isolated from the coelenterate Aequorea (6, 7, 37, 40), and the Cypridina luciferin analogues CLA and MCLA, of which MCLA is the most sensitive probe for O2*-. Although the intensity of light emitted from the interaction of coelenterazine with O2*- is greater than either that of lucigenin or luminol, coelenterazine-dependent chemiluminescence is not entirely specific for O2*-, because ONOO- will also cause coelenterazine to luminescence (37). A similar problem is found with MCLA which reacts with peroxyl radicals (43). Our investigations with MCLA has shown that although it is more sensitive to O2*- than luminol, it has a high level of background luminescence, is both light and temperature sensitive, and it auto-oxidizes, adding to the practical problems associated with nonspecificity. As such, it is recommended that all experiments are performed as quickly as possible in the dark and that reagents are similarly prepared in the dark.

2.4. Electron Paramagnetic Resonance Spectroscopy

The only technique that specifically detects free radicals is electron paramagnetic resonance (EPR) spectroscopy because it unequivocally measures the presence of unpaired electrons. However, unpaired electrons of species such as O2*_, ' OH, and ' NO are highly reactive radicals and generally do not accumulate to high-enough levels to be measured. One solution to the problem has been to use "spin traps" or "probes" that intercept reactive radical intermediates and form stable longer-lasting radical adducts with characteristic EPR signatures (1). These adducts accumulate to levels that can be detected with the use of EPR spectroscopy and provide information to enable the identification of the originating free radical species. A wide range of spin traps are commercially available for use both in whole animal studies and cell culture systems, including N-tertbutyl-p-phenylnitrone and 5,5-dimethyl -1-pyrroline N-oxide (DMPO) (44). Recently, newer compounds including 1,1,3-trimethyl-isoindole N-oxide (45), N-2-(2-ethoxy-carbonyl-propyl)-a-phenylnitrone (46) and 5-diethoxyphosphoryl-5-methyl-1-pyrroline N-oxide (DEPMPO) reported to be more

' specific and stable have also emerged (44, 47, 48).

2.4.1. Protocol: EPR Spectroscopy Characterization of ROI Generated by Xanthine Oxidase Using the Spin-Trap DEPMPO

2.4.1.1. Materials

1. Chelex-100.

3. DEPMPO (Oxis International Inc., Portland, OR).

6. Diethylene triamine pentaacetic acid (Sigma-Aldrich).

Fig. 3.4. Determination by electron paramagnetic resonance of O2*- generated by xantlhine/xanthine oxidase using the spin-trap DEPMPO (see text for details). DEPMPO- O2*- adduct is inhibited by the addition of SOD but not by heat inactivated SOD.

1. Pretreat PBS buffer and water with the ion-exchange resin Chelex-100 (1 g/100 mL; Bio-Rad, Hercules, CA) to remove adventitious metals in the buffer (49) and add diethylene tri-amine pentaacetic acid at a final concentration of 0.1 mM

2. Incubate hypoxanthine with xanthine oxidase (0.01 units/ mL) in the presence of the spin trap DEPMPO (10 mM).

3. Add the mixtures into an aqueous quartz flat cell (Wilmad, Buena, NJ), which should then be centred in a TE011 cavity.

4. Record EPR spectra with a Bruker 200D spectrometer operated at 9.7 GHz with a 1-GHz modulation frequency.

5. Transfer the data to a computer for simulation analysis (49). Typical data obtained under these conditions are illustrated in Fig. 3.4.

2.4.1.3. Comments

The determination of radical species such as O2*-, * OH, and * NO by EPR spectroscopy requires the use of spin-traps to generate species with longer half-lives required for their detection by EPR spectroscopy. The technique can also be used to detect more stable free radical-derived species, including ascorbyl radical, tocopheroxyl radical, and heme-nitrosyl complexes produced in vascular tissues during oxidative injury and inflammation (50). However, there are a several potential problems with the use of spin traps that require consideration (1, 44, 51). For example, one must consider whether the reaction products give the EPR signal can be rapidly removed in vivo and in cultured cells by enzymatic metabolism and also by direct reduction by agents such as ascorbate. For example, when DMPO is used to trap the *OH, ascorbate can directly reduce the DMPO-hydroxyl radical adduct to an EPR-silent species (1). DMPO can also be oxidized by ferric ions, which generates the four-line spectrum normally produced by *OH (52). Several published studies reporting on the use of spin traps to detect radical species frequently fail to show use of the proper controls (53). For example, does the added compound interfere with the radical-generating system (e.g., decomposing H2O2 or chelating iron in the Fe2 +/H2O2 system), or does it interact directly with the trap spin adduct, reducing it to an EPR-silent species?

Rizzi et al. have introduced triarylmethyl free radical, TAM OX063, as a probe for the detection of O2*- in aqueous solution (54). In this case, the O2*- reacts with the probe to cause the loss of the EPR signal. One advantage of TAM OX063 is that it is not subject to reduction by such agents as ascorbate or reduced thiols such as glutathione (GSH). Valgimigli and colleagues described an EPR method for the measurement of the oxidative stress status in biological systems (55). The method was based on the X-band EPR detection of a nitroxide generated under physiological conditions by oxidation of bis(1-hydroxy-2,2,6,6-tetramethyl-4-piperidinyl)-decandioate which is administrated as hydrochloride salt. Since the probe is reported to react rapidly with the majority of radical species involved in the oxidative stress and cross cell membranes easily it was suggested to be applicable in the clinical setting (55).

3. Measurement of ROI in the Mitochondria

3.1. Isolation of 1.

Mitochondria From

Cells and Tissue 2.

Pellet approximately 2 x 10c cells by centrifugation in a microcentrifuge tube at 1000^ for 3 min.

Carefully remove and discard the supernatant.

Wash cells in MSHE buffer (sucrose 70 mM, mannitol 220 mM, HEPES 2 mM, ethylene glycol tetraacetic acid (EGTA), bovine serum albumin (BSA) 0.5 mM @ pH 7.4) and centrifuge at 1000^ for 3 min.

Remove supernatant and suspend cells in approximately ~150 pL of fresh MSHE buffer.

5. Syringe homogenize cells through a needle (27.5 gage) until 80-90% cells are blue by trypan blue exclusion.

6. Centrifuge cells at 1000^ for 10 min at 4°C and transfer the supernatant to a new tube, which is further centrifuged at 10,000^ for 20 min at 4°C.

7. Transfer the supernatant from the 10,000^ centrifugation to a new microcentrifuge tube -this contains the cytosolic fraction. The pellet from the 10,000^ centrifugation contains the isolated mitochondrial fraction. Maintain the mitochondria pellet in MSHE buffer on ice prior to processing. Mitochondrial fractions should support coupled respiration analysed using a Clark-type electrode or equivalent.

3.1.2. Animai Tissue: Isolation of Rat Liver Mitochondria

We have found liver preparations from laboratory rodents to be suitable for large quantities of good quality and coupled mitochondria (16, 17) obtained using the following protocol.

1. Use male Sprague-Dawley rats weighing 180-220 g for isolation of liver mitochondria. Rat liver mitochondria were isolated as follows.

2. Chop fresh liver tissues finely and homogenize in ice-cold isolation medium containing 220 mM mannitol, 70 mM sucrose, 2 mM HEPES, 0.5 mM EGTA, and 0.1% BSA (fat free) (pH 7.4), using a Dounce homogenizer.

Centrifuge the homogenate at 1000^ for 10 min at 4°C in a Beckman JA20 rotor.

Transfer the supernatant into a fresh tube and centrifuge as described previously.

Centrifuge the supernatant at 10,000^ for 10 min at 4°C and retain the pellet.

Scrap the middle dark brown layer from the pellet and transfer to another fresh tube.

7. Suspend the final pellet in a small volume of the isolation buffer.

This sample may now be assayed for protein concentration.

The condition of the intact mitochondria can be tested by measuring oxygen consumption in the presence of succinate and ADP and determining respiratory control ratio (21).

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