JPM-06181; No of Pages 8 Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
Contents lists available at ScienceDirect
Journal of Pharmacological and Toxicological Methods journal homepage: www.elsevier.com/locate/jpharmtox
1
Review
4Q1
Lampson M. Fan b, Jian-Mei Li a,⁎ a
7
a r t i c l e
8 18 9 10 11
Article history: Received 16 January 2014 Accepted 28 March 2014 Available online xxxx
12 13 14 15 16 17
Keywords: Methods Reactive oxygen species Drug screen Cytotoxicity Cell cycle
i n f o
a b s t r a c t
E
D
P
Intracellular reactive oxygen species (ROS) production is essential to normal cell function. However, excessive ROS production causes oxidative damage and cell death. Many pharmacological compounds exert their effects on cell cycle progression by changing intracellular redox state and in many cases cause oxidative damage leading to drug cytotoxicity. Appropriate measurement of intracellular ROS levels during cell cycle progression is therefore crucial in understanding redox-regulation of cell function and drug toxicity and for the development of new drugs. However, due to the extremely short half-life of ROS, measuring the changes in intracellular ROS levels during a particular phase of cell cycle for drug intervention can be challenging. In this article, we have provided updated information on the rationale, the applications, the advantages and limitations of common methods for screening drug effects on intracellular ROS production linked to cell cycle study. Our aim is to facilitate biomedical scientists and researchers in the pharmaceutical industry in choosing or developing specific experimental regimens to suit their research needs. © 2014 Published by Elsevier Inc.
32
E
Contents
N C O
R
R
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Monitoring intracellular O2− generation by lucigenin (N,N′-dimethyl-9,9′-biacridinium dinitrate)-chemiluminescence . . . . . . . . . . . . . . 3. Detecting O 2− production by superoxide dismutase (SOD)-inhibitable cytochrome c (cyt c) reduction assay . . . . . . . . . . . . . . 4. Measuring intracellular O2− production using dihydroethidium (DHE) fluorescence high-performance liquid chromatography (HPLC) . . . . . . . . . 5. DHE fluorescence flow cytometer detection of intracellular ROS production and dual-labelling for a cell cycle marker . . . . . . . . . . . . . . . 6. DHE fluorescence plate-reader detection of intracellular ROS production in intact adherent cells cultured onto 96 well plates . . . . . . . . . . . 7. DHE fluorescence microscopy detection of intracellular ROS production in intact adherent cells . . . . . . . . . . . . . . . . . . . . . . . . . 8. Recording the intracellular ROS production by 5-(and 6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate (CM-DCF-DA) fluorescence microscopy 9. CM-DCF fluorescence flow cytometry detection of intracellular ROS production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Author contribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conflict of interest statement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Uncited reference . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
49
U
36 37 38 39 40 41 42 43 44 45 46 47 48
19 20 21 22 23 24 25 26 27 28 29
C
33 31 30 35 34
R O
Faculty of Health and Medical Sciences, University of Surrey, Guildford, UK John Radcliffe Hospital, University of Oxford, Oxford OX3 9DU, UK
T
b
O
5 6
F
3
Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies
2
50 Q2
1. Introduction
51 52
Reactive oxygen species (ROS) are a group of chemically reactive molecules and free radicals formed by incomplete single-electron Abbreviations: ROS, reactive oxygen species; DHE, dihydroethidium; 2-OH-E+, 2hydroxyethidium; SOD, superoxide dismutase; HPLC, high-performance liquid chromatography; CM-DCF-DA, 5-(and 6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate; DCF, 2′,7′-dichlorodihydrofluorescein; PCNA, proliferating cell nuclear antigen. ⁎ Corresponding author at: AY Building, Faculty of Health and Medical Sciences, University of Surrey, Guildford, Surrey GU2 7XH, UK. Tel.: +44 1483 686475; fax: +44 1483 686401. E-mail address: [email protected] (J.-M. Li).
0 0 0 0 0 0 0 0 0 0 0 0 0
reduction of oxygen as by-products of cellular aerobic metabolic processes (Valko et al., 2007). These molecules readily react with other biological products and influence the cell function in response to intracellular and extracellular stimuli such as drug intervention. ROS are produced by virtually every mammalian cell type from a variety of different sources such as mitochondrial electron transport chain, ionizing radiation, NADPH oxidase, cytochrome P450 reductase, xanthine oxidase, and nitric oxide synthase (Dröge, 2002; Finkel, 2011; Gutteridge & Halliwell, 2010; Li & Shah, 2004; Valko et al., 2007). Common ROS include superoxide (O2−), hydrogen peroxide (H2O2), peroxynitrite (ONOO−), reactive aldehydes, nitric oxide (NO) and other hydroxyl
http://dx.doi.org/10.1016/j.vascn.2014.03.173 1056-8719/© 2014 Published by Elsevier Inc.
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
53 54 55 56 57 58 59 60 61 62 63
87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 Q3 112 113 114 115 116 117 118 119 120 121 122 123 124 125 126 127 128 129
Chemiluminescence is one of the most widely used methods for detecting ROS production by mammalian cells and tissues. Several luminescence probes are available in the market and each of them has advantages and limitations depending on experimental settings. Technical details and applications of these probes can be found in previous reviews (Freitas, Porto, Lima, & Fernandes, 2009; Maghzal, Krause, Stocker, & Jaquet, 2012; Mahfouz, Sharma, Lackner, Aziz, & Agarwal, 2009; Nauseef, 2014; Tarpey & Fridovich, 2001; Tarpey et al., 2004). In this article we will focus on lucigenin-chemiluminescence for O2− detection. Lucigenin-chemiluminescence has been a popular method for measuring intracellular O2− production by non-phagocytic cells due to the alleged advantages of lucigenin i.e. great cell membrane permeability, minimal cellular toxicity, high sensitivity and specificity of reaction with O2−. The rate constant for the reaction between O2− and lucigenin is three times higher than that for O2− with cytochrome c (Afanas'ev, Ostrakhovitch, Mikhal'chik, & Korkina, 2001), which makes lucigeninchemiluminescence particularly useful in the detection of low levels of O2− production by vascular endothelial cells (Fan, Teng, & Li, 2009; Li & Shah, 2001; Li et al., 2007; Teng, Fan, Meijles, & Li, 2012), vascular smooth muscle cells (Brandes et al., 2002; Chamseddine & Miller, 2003; Menshikov et al., 2006), fibroblasts (Chamseddine & Miller, 2003; Venkatachalam et al., 2008); kidney cells (Berasi, Xiu, Yee, & Paulson, 2004; Hannken, Schroeder, Stahl, & Wolf, 1998); lymphocytes (Berasi et al., 2004) lung alveolar epithelial cells (Tickner, Fan, Du, Meijles, & Li, 2011) and human spermatozoa (Mahfouz et al., 2009). The reaction is fast and can be monitored in real-time ranging from seconds to minutes which provides a kinetic reading of O2− generation by living cells under experimental conditions. The assay can be performed in a 96 well microplate where lucigenin is injected in the dark by an auto-dispenser in the chemiluminescence microplate reader. The entire system can be pre-warmed to 37 °C, and the reading is obtained instantly and monitored for hours. A diagram of the technique is shown in Fig. 1A, and Fig. 1B shows an example of the kinetic measurement of O2− production by mouse primary coronary microvascular endothelial cells. For vascular endothelial cells, there was no difference in the basal (without adding NADPH) levels of O2− production between cells cultured under 3 different conditions. However there was a significant difference in NADPH-dependent O2− production between cells. Compared to cells cultured in 10% FCS growth medium, quiescent cells (0% FCS for 24 h) produced less ROS, and TNFα-stimulated cells produced much more ROS. It had been reported previously that lucigenin could be reduced univalently by diverse enzymes including xanthine oxidase, glucose oxidase, and might react with NADH to mediate O2− production in a system where lucigenin was mixed with high concentration of xanthine oxidase (25–100 units/mL) plus NADH (Liochev & Fridovich, 1997). However, such artificial conditions would not exist in mammalian cell biology. In fact, another report found that when lucigenin was used at
146 147
O
F
144
R O
85 86
2. Monitoring intracellular O2− generation by lucigenin (N,N′-dimethyl-9,9′-biacridinium dinitrate)-chemiluminescence
P
83 84
130 131
D
81 82
the extremely short half-life of ROS generated during each cell cycle, accurate measurement of intracellular ROS level is difficult. Additional complementary approaches of ROS measurement are necessary in order to accurately reflect the intracellular oxidative changes. In this article, we have focused on easily applicable screening techniques using commonly available research laboratory instruments for the detection of the intracellular ROS production, in particular O2− by living cells and homogenates of cells/tissue. We have provided brief descriptions for the rationale of using these methods based on our experience and illustrated with results generated using settings in our laboratories. We have also given an updated overview of the applications, advantages and limitations of these techniques in relationship to pharmacological and toxicological studies of cell cycle control, cell proliferation and cell death.
T
79 80
C
77 78
E
75 76
R
73 74
R
71 72
O
70
C
68 69
N
66 67
radicals. Among them, the O2− radical has several unique effects on other ROS i.e. rapid inactivation of NO while generating ONOO−, and it serves as the precursor of other ROS such as H2O2 and OH (Ardanaz & Pagano, 2006). In general ROS affect cellular functions mainly through (a) acting as regulatory mediators in signalling processes that lead to altered gene transcription, and protein and enzyme activities (so-called “redox signalling”) (Biswas, Chida, & Rahman, 2006; Dröge, 2002; Finkel, 2011), or (b) oxidative damage to cellular proteins, nucleic acids and lipids resulting in cell apoptosis and death (Circu & Aw, 2010; Dröge, 2002; Whiteman, Hong, Jenner, & Halliwell, 2002). Cells have also antioxidant defence systems which can be divided into non-enzymatic molecules and antioxidant enzymes. Nonenzymatic molecules are ROS scavengers such as uric acid, ascorbic acid, α-tocopherol, and sulfhydryl-containing molecules such as glutathione. Antioxidant enzymes include catalase, glutathione peroxidase and particularly superoxide dismutases (SODs) (Verbon, Post, & Boonstra, 2012; Li & Shah, 2004). The overall balance between oxidative (ROS generating) and reductive (ROS scavenging) processes in the cell constitutes the cellular “redox state”. When the redox homeostasis of a cell is interrupted and the rate of ROS formation exceeds the capacity of the antioxidant defence systems, the condition of oxidative stress occurs (Görlach et al., 2002; Vokurkova, Xu, & Touyz, 2007; Vurusaner, Poli, & Basaga, 2012). Redox homeostasis and oxidant signalling are essential elements that maintain cell function and regeneration, whereas oxidative stress or excessive production of ROS in response to environmental challenges causes genetic and epigenetic changes and may lead to cell dysfunction (Verbon et al., 2012; Vurusaner et al., 2012). The concept of redox regulation of cell cycle progression represents an important mechanism linking the oxidative metabolic processes to the cell cycle regulatory machinery (Li, Fan, George, & Brooks, 2007; Menon & Goswami, 2007; Venkatachalam et al., 2008; Vokurkova et al., 2007). ROS are short-lived and certain types of ROS i.e. NO and H2O2, can diffuse within and between cells to interact with redoxsensitive signalling molecules that in turn regulate cell cycle progression. The direction of cell cycle progression is determined by both the upstream ligand-dependent stimulation of ROS production and the downstream ROS targets (Burch & Heintz, 2005; Burhans & Heintz, 2009; Menon & Goswami, 2007). It is well known that ROS can modulate cellular signalling pathways at multiple levels from membrane receptors and ion channels to various intracellular protein kinases, phosphatases and nuclear transcription factors (Li & Shah, 2004). Many important cell signalling molecules, such as the mitogenactivated protein kinases, nuclear factor- κB and the tumour suppressor protein p53 have redox-sensitive motifs (cysteine residues or metal cofactors) (Menon & Goswami, 2007; Vurusaner et al., 2012). Several key cell cycle components such as cyclins (i.e. cyclin D1 and E), cyclin dependent kinases (i.e. CDK2 and 4) and cell cycle check point proteins (i.e. Chk1, Chk2) are also redox sensitive (Maryanovich & Gross, 2013; Verbon et al., 2012; Vokurkova et al., 2007). Their functions and activities are influenced by fluctuations in the intracellular ROS levels and which in turn direct the cell to either progress through, withdraw from the cell cycle or undergo apoptosis. There is a close link between the redox cycle and cell cycle in mammalian cells and the threshold of ROS levels required to promote or to inhibit cell cycle progression may vary according to the type of cell and the extracellular environment. There are many ways to detect ROS species generated by cells or tissues, for example electrochemical quantification (Borgmann, 2009; Dikalov, Griendling, & Harrison, 2007; Halliwell & Whiteman, 2004; Tarpey, Wink, & Grisham, 2004), fluorescent probe techniques (Dikalov et al., 2007; Halliwell & Whiteman, 2004; Kalyanaraman et al., 2012; Tarpey et al., 2004) and electron spin resonance (Dikalov et al., 2007; Halliwell & Whiteman, 2004; Tarpey et al., 2004) and their applications have been extensively reviewed previously. However, in terms of cell cycle study and screening drug compound cytotoxicity, the techniques of detecting intracellular ROS have not yet been evaluated. Moreover, due to the large variety, low quantity, high reactivity and
U
64 65
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
E
2
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
132 133 134 135 136 137 138 139 140 141 142 143
145
148 Q4 149 150 151 152 153 Q5 154 155 156 157 158 159 160 161 162 163 164 165 166 167 168 169 170 171 172 173 174 175 176 177 178 179 180 181 182 183 184 185 186 187 188 189 190 191 192 193
3
N C O
R
R
E
C
T
E
D
P
R O
O
F
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
U
Fig. 1. Lucigenin-chemiluminescence detection of cell O2− production. A) Illustration of lucigenin-chemiluminescence technique. Lucigenin buffer: Hank's balanced salt solution plus freshly added MgCl2 0.8 mM and CaCl2 1.8 mM. Lucigenin (final concentration 5 μM) and NADPH (final concentration 100 μM) were injected into the wells through the autodispensers in the dark chamber of the luminometer just before reading. The final volume was 200 μL/well. B) Representative example of kinetic reading of O2− production by lucigenin (5 μM)-chemiluminescence. Mouse primary coronary microvascular endothelial cells (CMECs) were cultured for 24 h in 10% FCS growth medium (proliferating), or stimulated with TNFα (100 U/ml), or 0% FCS medium (quiescent). Chemiluminescence was recorded every minute for 60 min. NADPH was added at 20 min and tiron (5 mM) was added at 40 min.
194 195 196 197 198 199 200 201 202 203
a lower dose of 80 μmol/L, the direct enzymatic reduction of lucigenin decreased O2− production in the system (Afanas'ev et al., 2001). Although lucigenin was reported to undergo redox cycling itself which may give false positive reading of chemiluminescence in detecting O2− production by endothelial cell homogenates, this observation was obtained under extreme experimental conditions using a very high lucigenin concentration (N 200 μmol/L) in the presence of NADH (Sohn, Keller, Gloe, Crause, & Pohl, 2000). Lucigenin had been validated against other techniques of O2− detection and it has been shown clearly that when it is used at a dose below 10 μmol/L, it does not undergo
redox cycling (Li et al., 1998). Therefore, a low concentration of 5 μmol/L of lucigenin is recommended for detecting O2− production by living mammalian cells and cell homogenates (Du, Fan, Mai, & Li, 2013; Li, Mullen, & Shah, 2001; Li et al., 2007; Teng et al., 2012). In summary, lucigenin (5 μmol/L)-chemiluminescence allows high throughput and is technically easy to perform. It generates a great amount of data in a short period, which is particularly useful for screening time- and dose-dependent effects of drug compounds on ROS production by living cells (Li & Shah, 2001; Li et al., 2001). Lucigeninchemiluminescence can be performed alongside a cell proliferation
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
204 205 206 207 208 209 210 211 212 213
222 223 224 225
3. Detecting O2− production by superoxide dismutase (SOD)-inhibitable cytochrome c (cyt c) reduction assay Reduction of ferricytochrome c can be used to measure O2− production in whole cells and tissue extracts. This reaction occurs with a rate constant of ~1.5 × 105 [mol/L] s−1 at pH 8.5 at room temperature. Fe3+ cyt c + O2− → Fe2+ cyt c + O2.
228 229
255 256
Reduction of ferricytochrome c is measured at 550 nm in a spectrophotometrical 96 well microplate reader. Due to the fact that ferricytochrome c can be reduced by other oxidants such as H2O2 and a number of cellular enzymes such as cytochrome oxidase and peroxidases, the changes detected by cytochrome c reduction assay are not specific to O2−. To overcome this, SOD is added to confirm the assay specificity for the detection of O2−. The production of O2− in nmol/L per well is calculated from the difference between samples with and without SOD (Li & Shah, 2001; Tarpey et al., 2004). SOD-inhibitable cytochrome c reduction assay has been used by a number of studies to validate quantitatively the amount of O2− detected by lucigenin-chemiluminescence using xanthine and xanthine oxidase as the O2− generation system and there is a good correlation between the quantity of O2− detected by both assays (Li & Shah, 2001; Li et al., 1998). The assay in general is easy to perform, quantitative, high throughput and is convenient for measuring O2− production by cell/tissue homogenates where a large amount of ROS generation can be detected. The limitation is that SOD and cytochrome c cannot cross the cell membrane easily and therefore it is not suitable for measuring intracellular O2− production by living cells. However, the technique can be used to measure ROS released by cells into the supernatant. Although other cell membrane permeable O2− scavengers can be used to verify the detection of O2−, the reaction can be easily disturbed by adding reagents therefore, appropriate controls should be included in the experimental design to validate the results. Compared to lucigenin (5 μM)-chemiluminescence, the SOD-inhibitable cytochrome c reduction assay is not suitable for monitoring real-time generation of O2− by living cells, and is not sensitive enough to detect low levels of O2− production by nonphagocytic cells.
257 258
4. Measuring intracellular O2− production using dihydroethidium (DHE) fluorescence high-performance liquid chromatography (HPLC)
259
DHE is a cell membrane permeable non-fluorescent compound. It is intra-cellularly oxidised and forms ethidium (a red fluorescent product) which binds to DNA and is retained inside the cells. For many years, the ethidium fluorescence (Ex 500–530 nm, Em: 590–620 nm), which can be readily detected by either fluorescence microscopy or flow cytometry, has been attributed to O2− trapping inside the cells Methods of DHE solution preparation and application have been covered comprehensively previously (Fan et al., 2009; Teng et al., 2012; Zielonka, Vasquez-Vivar, & Kalyanaraman, 2008). However, DHE oxidation has been found recently to yield at least two oxidative fluorescent products i.e. 2hydroxyethidium (2-OH-E+), (Ex 480 nm, Em 567 nm) and ethidium (Fernandes et al., 2007; Zhao et al., 2003, 2005). The 2-OH-E+ is regarded as the specific product of the reaction between O2− and DHE, whereas ethidium is the non-specific oxidation fluorescent product of DHE (Kalyanaraman et al., 2013). The fluorescence spectral overlap between 2-OH-E+ and ethidium cannot be separated by fluorescence
245 246 247 248 249 250 251 252 253 254
260 261 262 263 264 265 266 267 268 269 270 271 272 273 Q6 274
C
243 244
E
241 242
R
239 240
R
237 238
O
235 236
C
233 234
N
232
U
230 231
275 276 277 278 Q7 279 280 281 282 283 284 285 286 287 288 289 290 291 292 293 294 295 296 297 Q8 298 299
5. DHE fluorescence flow cytometer detection of intracellular ROS 300 production and dual-labelling for a cell cycle marker 301 DHE fluorescence in cell suspension can be detected by a flow cytometer, which is performed in parallel with cell cycle analysis using the same cells cultured (or treated) under the same conditions. Both the cell number and the fluorescence intensity can be counted accurately and quantified by the flow cytometry, which gives an accurate intracellular redox image correlated to the cell size and cell cycle analysis (Ahmad et al., 2008; Du et al., 2009; Fan et al., 2009; McFarlin, Williams, Venable, Dwyer, & Haviland, 2013). This is particularly useful in understanding the relationship between the redox cycle and the cell cycle, and to investigate drug effects. A major advantage of flow cytometry is the high productivity that allows simultaneous measurement of large numbers of samples under different drug treatments for a rapid and accurate evaluation of redox status related to a certain profile of cell proliferation or apoptosis. DHE fluorescence flow cytometry is sensitive enough for the detection of low levels of ROS production by a limited number of non-phagocytic cells. All of these are important features required for research of new drug development or for stem cell differentiation where limited cell number is an issue. Fig. 2 is a representative example of this technique using the same sample of cultured U937 cells for cell cycle analysis (right panel), and DHE flow cytometer detection of intracellular ROS production in the presence or absence of tiron (a superoxide scavenger) (left panel) using a BD Accuri flow cytometer. One great advantage of DHE fluorescence flow cytometry is that DHE oxidation can be stopped instantly by fixing cells in paraformaldehyde and cells after fixation can be dual immunofluorescence labelled for example FITC (Ex/Em: 495/519 nm) to detect the expression of a specific marker of cell cycle progression (such as a cyclin) or cell differentiation. Dual fluorescence can be analysed simultaneously by the flow cytometer and the results provide a vivid image of intracellular ROS production and the corresponding levels of expression of the cell cycle molecule (or differentiation marker) in the same cells. The limitation is as discussed before that DHE flow cytometry cannot separate 2-OH-E+ from ethidium, and it is therefore difficult to ascertain whether the oxidative fluorescence is from O2− or other species of ROS. To overcome this, cell membrane permeable O2− scavengers such as tiron can be applied to verify the detection of O2−. The amount of O2− detected can be calculated by the differences between samples with and without tiron.
T
226 227
F
221
O
220
R O
218 219
microscopy or flow cytometry so a HPLC method was recently developed to separate and quantify 2-OH-E+ (O2−), ethidium (other ROS) and the residual DHE which has not been oxidised inside the cells (Fernandes et al., 2007; Kalyanaraman et al., 2013; Teng et al., 2012; Zhao et al., 2005). The results can be expressed semi-quantitatively as area under the curve, or the amount of 2-OH-E+ (O2−) can be quantified against a standard curve of O2− generated using the xanthine and xanthine oxidase system detected by the same equipment. Although the DHE fluorescence HPLC technique has the advantage of separating 2-OH-E+ from ethidium, it is not convenient for cell cycle study or drug compound toxicity screening. The technique requires an HPLC instrument equipped with a variety of detection systems including both fluorescence and UV detectors, and a mass spectrometer. The column needs to be often replaced due to blockages by cell/tissue extracts. Samples for HPLC need to be specifically prepared in relatively large quantities in order to get a decent signal of 2-OH-E+ and would not be suitable for any live cell study. The assay is not high throughput, due to the fact that the equilibration of HPLC is time consuming, and the column needs to be washed thoroughly between samples. Technically it is not straightforward and requires a skilled operator to run the instrument and to oversee the experiments. In comparison to other methods of ROS detection listed in this article, DHE fluorescence HPLC is not convenient when a large number of samples need to be processed. However, DHE HPLC is particularly useful if the specificity of O2− production by cells is a crucial element for the study.
P
216 217
assay or a cytotoxicity assay using the same format of a 96 well plate under the same experimental conditions. The levels of O2− production thus measured correlate closely with the stages of cell proliferation or death in the corresponding wells (Fan et al., 2009; Li et al., 2007; Tickner et al., 2011). The amount of O2− production can be quantified against a standard curve of O2− generated using the xanthine and xanthine oxidase system and detected on the same plate.
D
214 215
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
E
4
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
302 303 304 305 306 307 308 309 310 311 312 313 314 315 316 317 318 319 320 321 322 323 324 325 326 327 328 329 330 331 332 Q9 333 334 Q10 335 336 337 338
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
5
generate fluorescence. To avoid this, use of a low concentration of DHE is recommended. The DHE reagent should be kept in the dark and its incubation period with cells should be kept as short and constant as possible between samples.
380 381 382 383
8. Recording the intracellular ROS production by 5-(and 6)- 384 chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate 385 (CM-DCF-DA) fluorescence microscopy 386
F
CM-DCF-DA is a cell membrane permeable non-fluorescent reagent (Li et al., 2007; Touyz & Schiffrin, 1999). Once it diffuses across the membrane, the acetate groups of CM-DCF-DA are cleaved by intracellular esterases and CM-DCF is trapped inside the cell, and is oxidised by intracellular ROS to form a green fluorescent product (DCF, excitation/emission, in nm: 492–495/517–527)) in the cytoplasm, which can be measured using a fluorescence or confocal microscope. Compared to DHE, CM-DCF reacts slowly with intracellular ROS, and this provides a significant advantage as the CM-DCF oxidation can be microscopically recorded in real-time for cells cultured onto a special chamber or slide (Li et al., 2007) and the technique has been applied to a single isolated muscle fibre (Palomero, Pye, Kabayo, Spiller, & Jackson, 2008). The distribution pattern of DCF fluorescence in the cytoplasm can provide additional information on ROS generation in subcellular compartments. DCF labelled cells can also be fixed and their nuclei can be labelled by propidium iodide (Ex/Em: 536/617 nm, red fluorescence) and both fluorescence of DCF and propidium iodide can be imaged under a confocal microscope, which can give information of potential DNA damage i.e. nuclear fractionation (apoptosis) or mitosis (double nuclei) in proliferating cells.
Intracellular DHE fluorescence can also be detected by a fluorometer 346 plate reader using special 96 well plates that only allows light to pass 347 through the bottom of the well where the cells are adherent and alive 348 (Nazarewicz, Bikineyeva, & Dikalov, 2013). This technique is easy to 349 use, time saving and can be performed in parallel with an assay to detect 350 Q11 cell proliferation or drug toxicity using the same format of a 96 well 351 plate. This is very useful for pharmacological and toxicological screening 352 of compounds for oxidative damage of living cells. The limitation of the 353 fluorometer plate reader is that the detecting beam passes through only 354 a few points of the well and does not cover the entire bottom area of the 355 Q12 well so that the reading/per well is only an estimate (or representative) 356 value and is not the total cell ROS production in the well. The variations 357 between wells can be large and it is not reliable for cells in suspension 358 because the cells are moving around constantly in the well and we 359 would not know if the detecting beam actually passes the cells or not 360 and how many cells it may pass in one detection.
362 363 364 365 366 367 368 369 370 371 372 373 374 375 376 377 378 379
7. DHE fluorescence microscopy detection of intracellular ROS production in intact adherent cells DHE fluorescence can be detected in adherent living cells cultured onto chamber slides and then digitally imaged under a fluorescence microscope and quantified (Tickner et al., 2011). The DHE reaction can be stopped by fixing cells using paraformaldehyde and cells can be subsequently labelled in the dark using a different fluorescence probe (such as FITC) for a cell cycle molecule of interest. The comparison between the levels of ROS production and the level and subcellular location of that particular cell cycle molecule can provide important insights into the role of redox signalling in cell cycle progression. Due to the fact that ethidium fluorescence is from DNA in the nuclei, detailed observation of nuclei under microscopy may also provide useful information on cell apoptosis (nuclear fragmentation) or mitosis. The DHE fluorescence microscopy technique can also be used for fresh tissue cryosections to detect in situ ROS production (Du et al., 2013). In summary, DHE fluorescence is a powerful technique to detect intracellular ROS production by living cells, and is relatively more specific to O2− than to other species of ROS. DHE can be oxidised by air and
U
361
N C O
R
R
E
C
R O
T
345
P
6. DHE fluorescence plate-reader detection of intracellular ROS production in intact adherent cells cultured onto 96 well plates
D
343 344
389 390 391 392 393 394 395 396 397 398 399 400 401 402 403 404 405 406
E
341 342
With adequate controls, DHE fluorescence flow cytometry provides a useful and reliable technique that links intracellular ROS production to cell cycle progression (Ahmad et al., 2008; Du et al., 2009; Fan et al., 2009; McFarlin et al., 2013).
339 340
O
Fig. 2. DHE fluorescence cytometer for intracellular ROS production. U937 cells were cultured in 10% FCS medium, harvested and separated into 2 tubes. For cell cycle analysis, the DNA contents of cells were labelled by propidium iodide (Ex/Em: 536/617 nm) and the apoptotic cells (Apo) and cells in the Go/G1, S and G2.M phases were separated by their DNA contents using a flow cytometer (right panel). The intracellular O2− production was examined by a DHE fluorescence flow cytometer in the presence or absence of tiron (10 mmol/L), an O2− scavenger (left panel).
387 388
9. CM-DCF fluorescence flow cytometry detection of intracellular 407 ROS production 408 Like DHE fluorescence, DCF fluorescence can also be applied to cells in suspension and assessed by a flow cytometer (Ahmad et al., 2008; Du et al., 2009). This can be done in parallel with cell cycle analysis of the same sample and provide information on the redox state in comparison to cell cycle progression. Fig. 3A is a representative example of DCF fluorescence flow cytometry detection of intracellular ROS production by mouse primary microvascular endothelial cells in the presence or absence of tiron. Fig. 3B is a reprehensive example of cell cycle analysis. Compared to proliferating cells cultured in 10% FCS medium, quiescent cells (0% FCS medium for 24 h) generated less ROS. TNFα (100 U/mL, 24 h) stimulation significantly increased intracellular ROS production and this was accompanied by significant increases in cell apoptosis. The limitation is that: 1) DCF can be oxidised upon exposure to light and air resulting in a false increase in fluorescence. To avoid this, low light conditions should be used during experiments and whenever possible light protected tubes of aliquot should be used for each experiment. 2) DCF can be oxidised by a host of ROS such as nitric oxide, H2O2, peroxynitrite anions, and other hydroxyl radicals so it is not specific for detecting single species of ROS. The experimental setting should be carefully considered and a series of controls should be included in the experimental protocol. Despite these limitations, DCF remains a popular global ROS probe for detecting intracellular oxidants and has been applied to vascular endothelial cells (Fan et al., 2009; Teng et al., 2012), neurons (Kaur, Schulz, Heggland, Aschner, & Syversen, 2008), muscle fibres (Palomero et al., 2008) and human spermatozoa (Mahfouz et al., 2009). In summary, we have evaluated several ROS-detecting techniques that have the potential to be used for the study of redoxregulation of cell cycle progression and for pharmacological and toxicological screening of drug-related oxidative damage of cells or for other studies where the uses of these techniques are appropriate. The molecular properties and applications of ROS-detecting reagents, or their oxidative products, discussed in this article are given in Table 1. Fig. 4 summarises the advantages and limitations
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
409 410 411 412 413 414 415 416 417 418 419 420 421 Q13 422 423 424 425 426 427 428 429 430 431 432 433 434 435 436 437 438 439 440 441
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
C
T
E
D
P
R O
O
F
6
R
R
E
Fig. 3. Representative example of ROS production measurement using a CM-DCF fluorescence flow cytometer and cell cycle analysis. Mouse primary coronary microvascular endothelial cells were cultured for 24 h in 0% FCS medium (quiescent) or in 10% FCS growth medium (proliferating) or in 10% FCS growth medium plus TNFα (100 U/ml). Cells were detached and separated into 2 portions. The intracellular ROS production was detected by a DCF fluorescence flow cytometer with or without tiron (A). For cell cycle analysis, the DNA contents of cells were labelled by propidium iodide (Ex/Em: 536/617 nm) and the apoptotic cells (Apo) and cells in the Go/G1, S and G2.M phases were separated by their DNA contents using a flow cytometer (B).
t1:1 t1:2
Table 1 Properties and applications of ROS-detecting reagents discussed in the article.
C
point when the cell cycle is analysed and the cells can be fixed and dual-labelled with a cell differentiation marker. Both DHE and DCF fluorescence can be live-imaged by fluorescence microscopy or detected by a fluorometer. However, none of these techniques can distinguish the enzymatic source of ROS generation, therefore different enzyme inhibitors or specific ROS scavengers are needed to identify
N
446
U
444 445
O
447
of these techniques. Lucigenin (5 μM)-chemiluminescence provides a kinetic analysis of real-time production of O2− and is sensitive for detecting low levels of O2− production by non-phagocytic cells. DHE fluorescence HPLC detects specifically O 2 − and can separate O 2 − from other species of ROS but is time consuming. DHE fluorescence flow cytometry gives an actual redox profile at a particular time
442 443
Mol. weight
Extinction Emission (nm)
Assay final concentration
ROS detected
Lucigenin Ferricytochrome c Dihydroethidium (DHE)
510.5 12.4 (equine) 315.4
5 μmol/L 250 μmol/L 2 μmol/L
O2− All ROS non specific All ROS preference to O2−
Ethidium (E+)
394.3
DHE oxidation product
Other ROS except O2−
2-Hydroxyethidium (2-OH-E+)
330.2
DHE oxidation product
O2−
2′,7′-Dichlorodihydrofluorescein diacetate (DCF-DA) 2′,7′-Dichlorodihydrofluorescein (DCF)
487.3 401.2
Ex: 368 nm Em: 505 nm UV absorption λmax = 550 nm (reduced form) Fluorescence Ex: 526 nm Em: 605 nm Ex: 526 nm, Em: 605 nm Ex: 396 nm Em: 579 nm Non-fluorescence Ex: 492 nm Em: 517 nm
5 μmol/L DCF-DA cleaved product
All ROS non specific All ROS non specific
t1:3
Reagent
t1:4 t1:5 t1:6 t1:7 t1:8 t1:9 t1:10 t1:11 t1:12 t1:13 t1:14 t1:15
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
448 449 450 451 452 453
7
R O
O
F
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
Fig. 4. Diagrammatic summary of ROS-detecting methods discussed in this article for drug cytotoxicity screening or investigation of the role of ROS in cell function.
458 459 460
LF: Manuscript preparation and figure generation; JML: directed, reviewed and edited the manuscript. All authors have approved the final article and its submission.
461
Conflict of interest statement There is no conflict of interest.
C
462
E
463 Q14 Uncited reference
Kalyanaraman et al., 2014
R
464
References
466 467 468 469 470 471 472 473 474 475 476 477 478 479 480 481 482 483 484 485 486 487 488 489 490 491 492 493 494 495 496 497
Afanas'ev, I. B., Ostrakhovitch, E. A., Mikhal'chik, E. V., & Korkina, L. G. (2001). Direct enzymatic reduction of lucigenin decreases lucigenin-amplified chemiluminescence produced by superoxide ion. Luminescence, 16, 305–307. Ahmad, I. M., Abdalla, M. Y., Aykin-Burns, N., Simons, A. L., Oberley, L. W., Domann, F. E., et al. (2008). 2-Deoxyglucose combined with wild-type p53 overexpression enhances cytotoxicity in human prostate cancer cells via oxidative stress. Free Radical Biology and Medicine, 44, 826–834. Ardanaz, N., & Pagano, P. J. (2006). Hydrogen peroxide as a paracrine vascular mediator: Regulation and signaling leading to dysfunction. Experimental Biology and Medicine, 231, 237–251. Berasi, S. P., Xiu, M., Yee, A. S., & Paulson, K. E. (2004). HBP1 repression of the p47phox gene: Cell cycle regulation via the NADPH oxidase. Molecular and Cellular Biology, 24, 3011–3024. Biswas, S., Chida, A. S., & Rahman, I. (2006). Redox modifications of protein-thiols: Emerging roles in cell signaling. Biochemical Pharmacology, 71, 551–564. Borgmann, S. (2009). Electrochemical quantification of reactive oxygen and nitrogen: Challenges and opportunities. Analytical and Bioanalytical Chemistry, 394, 95–105. Brandes, R. P., Miller, F. J., Beer, S., Haendeler, J., Hoffmann, J., Ha, T., et al. (2002). The vascular NADPH oxidase subunit p47phox is involved in redox-mediated gene expression. Free Radical Biology and Medicine, 32, 1116–1122. Burch, P. M., & Heintz, N. H. (2005). Redox regulation of cell-cycle re-entry: Cyclin D1 as a primary target for the mitogenic effects of reactive oxygen and nitrogen species. Antioxidants and Redox Signaling, 7, 741–751. Burhans, W. C., & Heintz, N. H. (2009). The cell cycle is a redox cycle: Linking phasespecific targets to cell fate. Free Radical Biology and Medicine, 47, 1282–1293. Chamseddine, A. H., & Miller, F. J., Jr. (2003). Gp91phox contributes to NADPH oxidase activity in aortic fibroblasts but not smooth muscle cells. American Journal of Physiology — Heart and Circulatory Physiology, 285, H2284–H2289. Circu, M. L., & Aw, T. Y. (2010). Reactive oxygen species, cellular redox systems, and apoptosis. Free Radical Biology and Medicine, 48, 749–762. Dikalov, S., Griendling, K. K., & Harrison, D. G. (2007). Measurement of reactive oxygen species in cardiovascular studies. Hypertension, 49, 717–727.
N C O
R
465
U
P
Author contribution
Dröge, W. (2002). Free radicals in the physiological control of cell function. Physiological Reviews, 82, 47–95. Du, J., Fan, L. M., Mai, A., & Li, J. -M. (2013). Crucial roles of Nox2-derived oxidative stress in deteriorating the function of insulin receptors and endothelium in dietary obesity of middle-aged mice. British Journal of Pharmacology, 170, 1064–1077. Du, C., Gao, Z., Venkatesha, V. A., Kalen, A. L., Chaudhuri, L., Spitz, D. R., et al. (2009). Mitochondrial ROS and radiation induced transformation in mouse embryonic fibroblasts. Cancer Biology and Therapy, 8, 1962–1971. Fan, L. M., Teng, L., & Li, J. -M. (2009). Knockout of p47 phox uncovers a critical role of p40 phox in reactive oxygen species production in microvascular endothelial cells. Arteriosclerosis, Thrombosis, and Vascular Biology, 29, 1651–1656. Fernandes, D. C., Wosniak, J., Jr., Pescatore, L. A., Bertoline, M. A., Liberman, M., Laurindo, F. R., et al. (2007). Analysis of DHE-derived oxidation products by HPLC in the assessment of superoxide production and NADPH oxidase activity in vascular systems. American Journal of Physiology — Cell Physiology, 292, C413–C422. Finkel, T. (2011). Signal transduction by reactive oxygen species. Journal of Cell Biology, 194, 7–15. Freitas, M., Porto, G., Lima, J. L., & Fernandes, E. (2009). Optimization of experimental settings for the analysis of human neutrophils oxidative burst in vitro. Talanta, 78, 1476–1483. Görlach, A., Kietzmann, T., & Hess, J. (2002). Redox signaling through NADPH oxidases: Involvement in vascular proliferation and coagulation. Annals of the New York Academy of Sciences, 973, 505–507. Gutteridge, J. M., & Halliwell, B. (2010). Antioxidants: Molecules, medicines, and myths. Biochemical and Biophysical Research Communications, 393, 561–564. Halliwell, B., & Whiteman, M. (2004). Measuring reactive species and oxidative damage in vivo and in cell culture: How should you do it and what do the results mean? British Journal of Pharmacology, 142, 231–255. Hannken, T., Schroeder, R., Stahl, R. A., & Wolf, G. (1998). Angiotensin II-mediated expression of p27Kip1 and induction of cellular hypertrophy in renal tubular cells depend on the generation of oxygen radicals. Kidney International, 54, 1923–1933. Kalyanaraman, B., Darley-Usmar, V., Davies, K. J., Dennery, P. A., Forman, H. J., Grisham, M. B., et al. (2012). Measuring reactive oxygen and nitrogen species with fluorescent probes: Challenges and limitations. Free Radical Biology and Medicine, 52, 1–6. Kalyanaraman, B., Dranka, B. P., Hardy, M., Michalski, R., & Zielonka, J. (2014). HPLC-based monitoring of products formed from hydroethidine-based fluorogenic probes — the ultimate approach for intra- and extracellular superoxide detection. Biochimica et Biophysica Acta, 1840, 739–744. Kaur, P., Schulz, K., Heggland, I., Aschner, M., & Syversen, T. (2008). The use of fluorescence for detecting MeHg-induced ROS in cell cultures. Toxicology In Vitro, 22, 1392–1398. Li, J. -M., Fan, L. M., George, V. T., & Brooks, G. (2007). Nox2 regulates endothelial cell cycle arrest and apoptosis via p21cip1 and p53. Free Radical Biology and Medicine, 43, 976–986. Li, J. -M., Mullen, A. M., & Shah, A. M. (2001). Phenotypic properties and characteristics of superoxide production by mouse coronary microvascular endothelial cells. Journal of Molecular and Cellular Cardiology, 33, 1119–1131. Li, J. -M., & Shah, A. M. (2001). Differential NADPH- versus NADH-dependent superoxide production by phagocyte-type endothelial cell NADPH oxidase. Cardiovascular Research, 52, 477–486. Li, J. -M., & Shah, A. M. (2004). Endothelial cell superoxide generation: Regulation and relevance for cardiovascular pathophysiology. American Journal of Physiology — Regulatory, Integrative and Comparative Physiology, 287, R1014–R1030. Li, Y., Zhu, H., Kuppusamy, P., Roubaud, V., Zweier, J. L., & Trush, M. A. (1998). Validation of lucigenin (bis-N-methylacridinium) as a chemilumigenic probe for detecting superoxide anion radical production by enzymatic and cellular systems. Journal of Biological Chemistry, 273, 2015–2023. Liochev, S. I., & Fridovich, I. (1997). Lucigenin (bis-N-methylacridinium) as a mediator of superoxide anion production. Archives of Biochemistry and Biophysics, 337, 115–120.
D
457
T
456
the source of ROS generation and the type of ROS detected. Using a combination of two or three techniques will provide accurate information on ROS generation for the study.
E
454 455
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
498 499 500 501 502 503 504 505 506 507 508 509 510 511 512 513 514 515 516 517 518 519 520 521 522 523 524 525 526 527 528 529 530 531 532 533 534 535 536 537 538 539 540 541 542 543 544 545 546 547 548 549 550 551 552 553 554 555 556
O
F
Teng, L., Fan, L. M., Meijles, D., & Li, J. -M. (2012). Divergent effects of p47(phox) phosphorylation at S303-4 or S379 on tumor necrosis factor-a signaling via TRAF4 and MAPK in endothelial cells. Arteriosclerosis, Thrombosis, and Vascular Biology, 32, 1488–1496. Tickner, J., Fan, L. M., Du, J., Meijles, D., & Li, J. -M. (2011). Nox2-derived ROS in PPARγ signaling and cell-cycle progression of lung alveolar epithelial cells. Free Radical Biology and Medicine, 51, 763–772. Touyz, R. M., & Schiffrin, E. L. (1999). Ang II-stimulated superoxide production is mediated via phospholipase D in human vascular smooth muscle cells. Hypertension, 34, 976–982. Valko, M., Leibfritz, D., Moncol, J., Cronin, M. T., Mazur, M., & Telser, J. (2007). Free radicals and antioxidants in normal physiological functions and human disease. International Journal of Biochemistry and Cell Biology, 39, 44–84. Venkatachalam, P., de Toledo, S. M., Pandey, B. N., Tephly, L. A., Carter, A. B., Little, J. B., et al. (2008). Regulation of normal cell cycle progression by flavin-containing oxidases. Oncogene, 27, 20–31. Verbon, E. H., Post, J. A., & Boonstra, J. (2012). The influence of reactive oxygen species on cell cycle progression in mammalian cells. Gene, 511, 1–6. Vokurkova, M., Xu, S., & Touyz, R. M. (2007). Reactive oxygen species, cell growth, cell cycle progression and vascular remodeling in hypertension. Future Cardiology, 3, 53–63. Vurusaner, B., Poli, G., & Basaga, H. (2012). Tumor suppressor genes and ROS: Complex networks of interactions. Free Radical Biology and Medicine, 52, 7–18. Whiteman, M., Hong, H. S., Jenner, A., & Halliwell, B. (2002). Loss of oxidized and chlorinated bases in DNA treated with reactive oxygen species: Implications for assessment of oxidative damage in vivo. Biochemical and Biophysical Research Communications, 296, 883–889. Zhao, H., Joseph, J., Fales, H. M., Sokoloski, E. A., Levine, R. L., Vasquez-Vivar, J., et al. (2005). Detection and characterization of the product of hydroethidine and intracellular superoxide by HPLC and limitations of fluorescence. PNAS, 102, 5727–5732. Zhao, H., Kalivendi, S., Zhang, H., Joseph, J., Nithipatikom, K., Vasquez-Vivar, J., et al. (2003). Superoxide reacts with hydroethidine but forms a fluorescent product that is distinctly different from ethidium: Potential implications in intracellular fluorescence detection of superoxide. Free Radical Biology and Medicine, 34, 1359–1368. Zielonka, J., Vasquez-Vivar, J., & Kalyanaraman, B. (2008). Detection of 2-hydroxyethidium in cellular systems: A unique marker product of superoxide and hydroethidine. Nature Protocols, 3, 8–21.
R O
Maghzal, G. J., Krause, K. -H., Stocker, R., & Jaquet, V. (2012). Detection of reactive oxygen species derived from the family of Nox NADPH oxidases. Free Radical Biology and Medicine, 53, 1903–1918. Mahfouz, R., Sharma, R., Lackner, J., Aziz, N., & Agarwal, A. (2009). Evaluation of chemiluminescence and flow cytometry as tools in assessing production of hydrogen peroxide and superoxide anion in human spermatozoa. Fertility and Sterility, 92, 819–827. Maryanovich, M., & Gross, A. (2013). A ROS rheostat for cell fate regulation. Trends in Cell Biology, 23, 129–134. McFarlin, B. K., Williams, R. R., Venable, A. S., Dwyer, K. C., & Haviland, D. L. (2013). Imagebased cytometry reveals three distinct subsets of activated granulocytes based on phagocytosis and oxidative burst. Cytometry. Part A, 83, 745–751. Menon, S. G., & Goswami, P. C. (2007). A redox cycle within the cell cycle: Ring in the old with the new. Oncogene, 26, 1101–1109. Menshikov, M., Plekhanova, O., Cai, H., Chalupsky, K., Parfyonova, Y., Bashtrikov, P., et al. (2006). Urokinase plasminogen activator stimulates vascular smooth muscle cell proliferation via redox-dependent pathways. Arteriosclerosis, Thrombosis, and Vascular Biology, 26, 801–807. Nauseef, W. M. (2014). Detection of superoxide anion and hydrogen peroxide production by cellular NADPH oxidase. Biochimica et Biophysica Acta, 1840, 757–767. http://dx. doi.org/10.1016/j.bbagen.2013.04.040. Nazarewicz, R. R., Bikineyeva, A., & Dikalov, S. I. (2013). Rapid and specific measurements of superoxide using fluorescence spectroscopy. Journal of Biomolecular Screening, 18, 498–503. Palomero, J., Pye, D., Kabayo, T., Spiller, D. G., & Jackson, M. J. (2008). In situ detection and measurement of intracellular reactive oxygen species in single isolated mature skeletal muscle fibers by real time fluorescence microscopy. Antioxidants and Redox Signaling, 10, 1463–1474. Sohn, H. -Y., Keller, M., Gloe, T., Crause, P., & Pohl, U. (2000). Pitfalls of using lucigenin in endothelial cells: Implications for NAD(P)H dependent superoxide formation. Free Radical Research, 32, 265–272. Tarpey, M. M., & Fridovich, I. (2001). Methods of detection of vascular reactive species. Circulation Research, 89, 224–236. Tarpey, M. M., Wink, D. A., & Grisham, M. B. (2004). Methods for detection of reactive metabolites of oxygen and nitrogen: In vitro and in vivo considerations. American Journal of Physiology — Regulatory, Integrative and Comparative Physiology, 286, R431–R444.
D
557 558 559 560 561 562 563 564 565 566 567 568 569 570 571 572 573 574 575 576 577 578 579 580 581 582 583 584 585 586 587 588 589 590 591
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
P
8
U
N
C
O
R
R
E
C
T
E
627
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
592 593 594 595 596 597 598 599 600 601 602 603 604 605 606 607 608 609 610 611 612 613 614 615 616 617 618 619 620 621 622 623 624 625 626
Contents lists available at ScienceDirect
Journal of Pharmacological and Toxicological Methods journal homepage: www.elsevier.com/locate/jpharmtox
1
Review
4Q1
Lampson M. Fan b, Jian-Mei Li a,⁎ a
7
a r t i c l e
8 18 9 10 11
Article history: Received 16 January 2014 Accepted 28 March 2014 Available online xxxx
12 13 14 15 16 17
Keywords: Methods Reactive oxygen species Drug screen Cytotoxicity Cell cycle
i n f o
a b s t r a c t
E
D
P
Intracellular reactive oxygen species (ROS) production is essential to normal cell function. However, excessive ROS production causes oxidative damage and cell death. Many pharmacological compounds exert their effects on cell cycle progression by changing intracellular redox state and in many cases cause oxidative damage leading to drug cytotoxicity. Appropriate measurement of intracellular ROS levels during cell cycle progression is therefore crucial in understanding redox-regulation of cell function and drug toxicity and for the development of new drugs. However, due to the extremely short half-life of ROS, measuring the changes in intracellular ROS levels during a particular phase of cell cycle for drug intervention can be challenging. In this article, we have provided updated information on the rationale, the applications, the advantages and limitations of common methods for screening drug effects on intracellular ROS production linked to cell cycle study. Our aim is to facilitate biomedical scientists and researchers in the pharmaceutical industry in choosing or developing specific experimental regimens to suit their research needs. © 2014 Published by Elsevier Inc.
32
E
Contents
N C O
R
R
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Monitoring intracellular O2− generation by lucigenin (N,N′-dimethyl-9,9′-biacridinium dinitrate)-chemiluminescence . . . . . . . . . . . . . . 3. Detecting O 2− production by superoxide dismutase (SOD)-inhibitable cytochrome c (cyt c) reduction assay . . . . . . . . . . . . . . 4. Measuring intracellular O2− production using dihydroethidium (DHE) fluorescence high-performance liquid chromatography (HPLC) . . . . . . . . . 5. DHE fluorescence flow cytometer detection of intracellular ROS production and dual-labelling for a cell cycle marker . . . . . . . . . . . . . . . 6. DHE fluorescence plate-reader detection of intracellular ROS production in intact adherent cells cultured onto 96 well plates . . . . . . . . . . . 7. DHE fluorescence microscopy detection of intracellular ROS production in intact adherent cells . . . . . . . . . . . . . . . . . . . . . . . . . 8. Recording the intracellular ROS production by 5-(and 6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate (CM-DCF-DA) fluorescence microscopy 9. CM-DCF fluorescence flow cytometry detection of intracellular ROS production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Author contribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conflict of interest statement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Uncited reference . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
49
U
36 37 38 39 40 41 42 43 44 45 46 47 48
19 20 21 22 23 24 25 26 27 28 29
C
33 31 30 35 34
R O
Faculty of Health and Medical Sciences, University of Surrey, Guildford, UK John Radcliffe Hospital, University of Oxford, Oxford OX3 9DU, UK
T
b
O
5 6
F
3
Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies
2
50 Q2
1. Introduction
51 52
Reactive oxygen species (ROS) are a group of chemically reactive molecules and free radicals formed by incomplete single-electron Abbreviations: ROS, reactive oxygen species; DHE, dihydroethidium; 2-OH-E+, 2hydroxyethidium; SOD, superoxide dismutase; HPLC, high-performance liquid chromatography; CM-DCF-DA, 5-(and 6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate; DCF, 2′,7′-dichlorodihydrofluorescein; PCNA, proliferating cell nuclear antigen. ⁎ Corresponding author at: AY Building, Faculty of Health and Medical Sciences, University of Surrey, Guildford, Surrey GU2 7XH, UK. Tel.: +44 1483 686475; fax: +44 1483 686401. E-mail address: [email protected] (J.-M. Li).
0 0 0 0 0 0 0 0 0 0 0 0 0
reduction of oxygen as by-products of cellular aerobic metabolic processes (Valko et al., 2007). These molecules readily react with other biological products and influence the cell function in response to intracellular and extracellular stimuli such as drug intervention. ROS are produced by virtually every mammalian cell type from a variety of different sources such as mitochondrial electron transport chain, ionizing radiation, NADPH oxidase, cytochrome P450 reductase, xanthine oxidase, and nitric oxide synthase (Dröge, 2002; Finkel, 2011; Gutteridge & Halliwell, 2010; Li & Shah, 2004; Valko et al., 2007). Common ROS include superoxide (O2−), hydrogen peroxide (H2O2), peroxynitrite (ONOO−), reactive aldehydes, nitric oxide (NO) and other hydroxyl
http://dx.doi.org/10.1016/j.vascn.2014.03.173 1056-8719/© 2014 Published by Elsevier Inc.
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
53 54 55 56 57 58 59 60 61 62 63
87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 Q3 112 113 114 115 116 117 118 119 120 121 122 123 124 125 126 127 128 129
Chemiluminescence is one of the most widely used methods for detecting ROS production by mammalian cells and tissues. Several luminescence probes are available in the market and each of them has advantages and limitations depending on experimental settings. Technical details and applications of these probes can be found in previous reviews (Freitas, Porto, Lima, & Fernandes, 2009; Maghzal, Krause, Stocker, & Jaquet, 2012; Mahfouz, Sharma, Lackner, Aziz, & Agarwal, 2009; Nauseef, 2014; Tarpey & Fridovich, 2001; Tarpey et al., 2004). In this article we will focus on lucigenin-chemiluminescence for O2− detection. Lucigenin-chemiluminescence has been a popular method for measuring intracellular O2− production by non-phagocytic cells due to the alleged advantages of lucigenin i.e. great cell membrane permeability, minimal cellular toxicity, high sensitivity and specificity of reaction with O2−. The rate constant for the reaction between O2− and lucigenin is three times higher than that for O2− with cytochrome c (Afanas'ev, Ostrakhovitch, Mikhal'chik, & Korkina, 2001), which makes lucigeninchemiluminescence particularly useful in the detection of low levels of O2− production by vascular endothelial cells (Fan, Teng, & Li, 2009; Li & Shah, 2001; Li et al., 2007; Teng, Fan, Meijles, & Li, 2012), vascular smooth muscle cells (Brandes et al., 2002; Chamseddine & Miller, 2003; Menshikov et al., 2006), fibroblasts (Chamseddine & Miller, 2003; Venkatachalam et al., 2008); kidney cells (Berasi, Xiu, Yee, & Paulson, 2004; Hannken, Schroeder, Stahl, & Wolf, 1998); lymphocytes (Berasi et al., 2004) lung alveolar epithelial cells (Tickner, Fan, Du, Meijles, & Li, 2011) and human spermatozoa (Mahfouz et al., 2009). The reaction is fast and can be monitored in real-time ranging from seconds to minutes which provides a kinetic reading of O2− generation by living cells under experimental conditions. The assay can be performed in a 96 well microplate where lucigenin is injected in the dark by an auto-dispenser in the chemiluminescence microplate reader. The entire system can be pre-warmed to 37 °C, and the reading is obtained instantly and monitored for hours. A diagram of the technique is shown in Fig. 1A, and Fig. 1B shows an example of the kinetic measurement of O2− production by mouse primary coronary microvascular endothelial cells. For vascular endothelial cells, there was no difference in the basal (without adding NADPH) levels of O2− production between cells cultured under 3 different conditions. However there was a significant difference in NADPH-dependent O2− production between cells. Compared to cells cultured in 10% FCS growth medium, quiescent cells (0% FCS for 24 h) produced less ROS, and TNFα-stimulated cells produced much more ROS. It had been reported previously that lucigenin could be reduced univalently by diverse enzymes including xanthine oxidase, glucose oxidase, and might react with NADH to mediate O2− production in a system where lucigenin was mixed with high concentration of xanthine oxidase (25–100 units/mL) plus NADH (Liochev & Fridovich, 1997). However, such artificial conditions would not exist in mammalian cell biology. In fact, another report found that when lucigenin was used at
146 147
O
F
144
R O
85 86
2. Monitoring intracellular O2− generation by lucigenin (N,N′-dimethyl-9,9′-biacridinium dinitrate)-chemiluminescence
P
83 84
130 131
D
81 82
the extremely short half-life of ROS generated during each cell cycle, accurate measurement of intracellular ROS level is difficult. Additional complementary approaches of ROS measurement are necessary in order to accurately reflect the intracellular oxidative changes. In this article, we have focused on easily applicable screening techniques using commonly available research laboratory instruments for the detection of the intracellular ROS production, in particular O2− by living cells and homogenates of cells/tissue. We have provided brief descriptions for the rationale of using these methods based on our experience and illustrated with results generated using settings in our laboratories. We have also given an updated overview of the applications, advantages and limitations of these techniques in relationship to pharmacological and toxicological studies of cell cycle control, cell proliferation and cell death.
T
79 80
C
77 78
E
75 76
R
73 74
R
71 72
O
70
C
68 69
N
66 67
radicals. Among them, the O2− radical has several unique effects on other ROS i.e. rapid inactivation of NO while generating ONOO−, and it serves as the precursor of other ROS such as H2O2 and OH (Ardanaz & Pagano, 2006). In general ROS affect cellular functions mainly through (a) acting as regulatory mediators in signalling processes that lead to altered gene transcription, and protein and enzyme activities (so-called “redox signalling”) (Biswas, Chida, & Rahman, 2006; Dröge, 2002; Finkel, 2011), or (b) oxidative damage to cellular proteins, nucleic acids and lipids resulting in cell apoptosis and death (Circu & Aw, 2010; Dröge, 2002; Whiteman, Hong, Jenner, & Halliwell, 2002). Cells have also antioxidant defence systems which can be divided into non-enzymatic molecules and antioxidant enzymes. Nonenzymatic molecules are ROS scavengers such as uric acid, ascorbic acid, α-tocopherol, and sulfhydryl-containing molecules such as glutathione. Antioxidant enzymes include catalase, glutathione peroxidase and particularly superoxide dismutases (SODs) (Verbon, Post, & Boonstra, 2012; Li & Shah, 2004). The overall balance between oxidative (ROS generating) and reductive (ROS scavenging) processes in the cell constitutes the cellular “redox state”. When the redox homeostasis of a cell is interrupted and the rate of ROS formation exceeds the capacity of the antioxidant defence systems, the condition of oxidative stress occurs (Görlach et al., 2002; Vokurkova, Xu, & Touyz, 2007; Vurusaner, Poli, & Basaga, 2012). Redox homeostasis and oxidant signalling are essential elements that maintain cell function and regeneration, whereas oxidative stress or excessive production of ROS in response to environmental challenges causes genetic and epigenetic changes and may lead to cell dysfunction (Verbon et al., 2012; Vurusaner et al., 2012). The concept of redox regulation of cell cycle progression represents an important mechanism linking the oxidative metabolic processes to the cell cycle regulatory machinery (Li, Fan, George, & Brooks, 2007; Menon & Goswami, 2007; Venkatachalam et al., 2008; Vokurkova et al., 2007). ROS are short-lived and certain types of ROS i.e. NO and H2O2, can diffuse within and between cells to interact with redoxsensitive signalling molecules that in turn regulate cell cycle progression. The direction of cell cycle progression is determined by both the upstream ligand-dependent stimulation of ROS production and the downstream ROS targets (Burch & Heintz, 2005; Burhans & Heintz, 2009; Menon & Goswami, 2007). It is well known that ROS can modulate cellular signalling pathways at multiple levels from membrane receptors and ion channels to various intracellular protein kinases, phosphatases and nuclear transcription factors (Li & Shah, 2004). Many important cell signalling molecules, such as the mitogenactivated protein kinases, nuclear factor- κB and the tumour suppressor protein p53 have redox-sensitive motifs (cysteine residues or metal cofactors) (Menon & Goswami, 2007; Vurusaner et al., 2012). Several key cell cycle components such as cyclins (i.e. cyclin D1 and E), cyclin dependent kinases (i.e. CDK2 and 4) and cell cycle check point proteins (i.e. Chk1, Chk2) are also redox sensitive (Maryanovich & Gross, 2013; Verbon et al., 2012; Vokurkova et al., 2007). Their functions and activities are influenced by fluctuations in the intracellular ROS levels and which in turn direct the cell to either progress through, withdraw from the cell cycle or undergo apoptosis. There is a close link between the redox cycle and cell cycle in mammalian cells and the threshold of ROS levels required to promote or to inhibit cell cycle progression may vary according to the type of cell and the extracellular environment. There are many ways to detect ROS species generated by cells or tissues, for example electrochemical quantification (Borgmann, 2009; Dikalov, Griendling, & Harrison, 2007; Halliwell & Whiteman, 2004; Tarpey, Wink, & Grisham, 2004), fluorescent probe techniques (Dikalov et al., 2007; Halliwell & Whiteman, 2004; Kalyanaraman et al., 2012; Tarpey et al., 2004) and electron spin resonance (Dikalov et al., 2007; Halliwell & Whiteman, 2004; Tarpey et al., 2004) and their applications have been extensively reviewed previously. However, in terms of cell cycle study and screening drug compound cytotoxicity, the techniques of detecting intracellular ROS have not yet been evaluated. Moreover, due to the large variety, low quantity, high reactivity and
U
64 65
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
E
2
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
132 133 134 135 136 137 138 139 140 141 142 143
145
148 Q4 149 150 151 152 153 Q5 154 155 156 157 158 159 160 161 162 163 164 165 166 167 168 169 170 171 172 173 174 175 176 177 178 179 180 181 182 183 184 185 186 187 188 189 190 191 192 193
3
N C O
R
R
E
C
T
E
D
P
R O
O
F
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
U
Fig. 1. Lucigenin-chemiluminescence detection of cell O2− production. A) Illustration of lucigenin-chemiluminescence technique. Lucigenin buffer: Hank's balanced salt solution plus freshly added MgCl2 0.8 mM and CaCl2 1.8 mM. Lucigenin (final concentration 5 μM) and NADPH (final concentration 100 μM) were injected into the wells through the autodispensers in the dark chamber of the luminometer just before reading. The final volume was 200 μL/well. B) Representative example of kinetic reading of O2− production by lucigenin (5 μM)-chemiluminescence. Mouse primary coronary microvascular endothelial cells (CMECs) were cultured for 24 h in 10% FCS growth medium (proliferating), or stimulated with TNFα (100 U/ml), or 0% FCS medium (quiescent). Chemiluminescence was recorded every minute for 60 min. NADPH was added at 20 min and tiron (5 mM) was added at 40 min.
194 195 196 197 198 199 200 201 202 203
a lower dose of 80 μmol/L, the direct enzymatic reduction of lucigenin decreased O2− production in the system (Afanas'ev et al., 2001). Although lucigenin was reported to undergo redox cycling itself which may give false positive reading of chemiluminescence in detecting O2− production by endothelial cell homogenates, this observation was obtained under extreme experimental conditions using a very high lucigenin concentration (N 200 μmol/L) in the presence of NADH (Sohn, Keller, Gloe, Crause, & Pohl, 2000). Lucigenin had been validated against other techniques of O2− detection and it has been shown clearly that when it is used at a dose below 10 μmol/L, it does not undergo
redox cycling (Li et al., 1998). Therefore, a low concentration of 5 μmol/L of lucigenin is recommended for detecting O2− production by living mammalian cells and cell homogenates (Du, Fan, Mai, & Li, 2013; Li, Mullen, & Shah, 2001; Li et al., 2007; Teng et al., 2012). In summary, lucigenin (5 μmol/L)-chemiluminescence allows high throughput and is technically easy to perform. It generates a great amount of data in a short period, which is particularly useful for screening time- and dose-dependent effects of drug compounds on ROS production by living cells (Li & Shah, 2001; Li et al., 2001). Lucigeninchemiluminescence can be performed alongside a cell proliferation
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
204 205 206 207 208 209 210 211 212 213
222 223 224 225
3. Detecting O2− production by superoxide dismutase (SOD)-inhibitable cytochrome c (cyt c) reduction assay Reduction of ferricytochrome c can be used to measure O2− production in whole cells and tissue extracts. This reaction occurs with a rate constant of ~1.5 × 105 [mol/L] s−1 at pH 8.5 at room temperature. Fe3+ cyt c + O2− → Fe2+ cyt c + O2.
228 229
255 256
Reduction of ferricytochrome c is measured at 550 nm in a spectrophotometrical 96 well microplate reader. Due to the fact that ferricytochrome c can be reduced by other oxidants such as H2O2 and a number of cellular enzymes such as cytochrome oxidase and peroxidases, the changes detected by cytochrome c reduction assay are not specific to O2−. To overcome this, SOD is added to confirm the assay specificity for the detection of O2−. The production of O2− in nmol/L per well is calculated from the difference between samples with and without SOD (Li & Shah, 2001; Tarpey et al., 2004). SOD-inhibitable cytochrome c reduction assay has been used by a number of studies to validate quantitatively the amount of O2− detected by lucigenin-chemiluminescence using xanthine and xanthine oxidase as the O2− generation system and there is a good correlation between the quantity of O2− detected by both assays (Li & Shah, 2001; Li et al., 1998). The assay in general is easy to perform, quantitative, high throughput and is convenient for measuring O2− production by cell/tissue homogenates where a large amount of ROS generation can be detected. The limitation is that SOD and cytochrome c cannot cross the cell membrane easily and therefore it is not suitable for measuring intracellular O2− production by living cells. However, the technique can be used to measure ROS released by cells into the supernatant. Although other cell membrane permeable O2− scavengers can be used to verify the detection of O2−, the reaction can be easily disturbed by adding reagents therefore, appropriate controls should be included in the experimental design to validate the results. Compared to lucigenin (5 μM)-chemiluminescence, the SOD-inhibitable cytochrome c reduction assay is not suitable for monitoring real-time generation of O2− by living cells, and is not sensitive enough to detect low levels of O2− production by nonphagocytic cells.
257 258
4. Measuring intracellular O2− production using dihydroethidium (DHE) fluorescence high-performance liquid chromatography (HPLC)
259
DHE is a cell membrane permeable non-fluorescent compound. It is intra-cellularly oxidised and forms ethidium (a red fluorescent product) which binds to DNA and is retained inside the cells. For many years, the ethidium fluorescence (Ex 500–530 nm, Em: 590–620 nm), which can be readily detected by either fluorescence microscopy or flow cytometry, has been attributed to O2− trapping inside the cells Methods of DHE solution preparation and application have been covered comprehensively previously (Fan et al., 2009; Teng et al., 2012; Zielonka, Vasquez-Vivar, & Kalyanaraman, 2008). However, DHE oxidation has been found recently to yield at least two oxidative fluorescent products i.e. 2hydroxyethidium (2-OH-E+), (Ex 480 nm, Em 567 nm) and ethidium (Fernandes et al., 2007; Zhao et al., 2003, 2005). The 2-OH-E+ is regarded as the specific product of the reaction between O2− and DHE, whereas ethidium is the non-specific oxidation fluorescent product of DHE (Kalyanaraman et al., 2013). The fluorescence spectral overlap between 2-OH-E+ and ethidium cannot be separated by fluorescence
245 246 247 248 249 250 251 252 253 254
260 261 262 263 264 265 266 267 268 269 270 271 272 273 Q6 274
C
243 244
E
241 242
R
239 240
R
237 238
O
235 236
C
233 234
N
232
U
230 231
275 276 277 278 Q7 279 280 281 282 283 284 285 286 287 288 289 290 291 292 293 294 295 296 297 Q8 298 299
5. DHE fluorescence flow cytometer detection of intracellular ROS 300 production and dual-labelling for a cell cycle marker 301 DHE fluorescence in cell suspension can be detected by a flow cytometer, which is performed in parallel with cell cycle analysis using the same cells cultured (or treated) under the same conditions. Both the cell number and the fluorescence intensity can be counted accurately and quantified by the flow cytometry, which gives an accurate intracellular redox image correlated to the cell size and cell cycle analysis (Ahmad et al., 2008; Du et al., 2009; Fan et al., 2009; McFarlin, Williams, Venable, Dwyer, & Haviland, 2013). This is particularly useful in understanding the relationship between the redox cycle and the cell cycle, and to investigate drug effects. A major advantage of flow cytometry is the high productivity that allows simultaneous measurement of large numbers of samples under different drug treatments for a rapid and accurate evaluation of redox status related to a certain profile of cell proliferation or apoptosis. DHE fluorescence flow cytometry is sensitive enough for the detection of low levels of ROS production by a limited number of non-phagocytic cells. All of these are important features required for research of new drug development or for stem cell differentiation where limited cell number is an issue. Fig. 2 is a representative example of this technique using the same sample of cultured U937 cells for cell cycle analysis (right panel), and DHE flow cytometer detection of intracellular ROS production in the presence or absence of tiron (a superoxide scavenger) (left panel) using a BD Accuri flow cytometer. One great advantage of DHE fluorescence flow cytometry is that DHE oxidation can be stopped instantly by fixing cells in paraformaldehyde and cells after fixation can be dual immunofluorescence labelled for example FITC (Ex/Em: 495/519 nm) to detect the expression of a specific marker of cell cycle progression (such as a cyclin) or cell differentiation. Dual fluorescence can be analysed simultaneously by the flow cytometer and the results provide a vivid image of intracellular ROS production and the corresponding levels of expression of the cell cycle molecule (or differentiation marker) in the same cells. The limitation is as discussed before that DHE flow cytometry cannot separate 2-OH-E+ from ethidium, and it is therefore difficult to ascertain whether the oxidative fluorescence is from O2− or other species of ROS. To overcome this, cell membrane permeable O2− scavengers such as tiron can be applied to verify the detection of O2−. The amount of O2− detected can be calculated by the differences between samples with and without tiron.
T
226 227
F
221
O
220
R O
218 219
microscopy or flow cytometry so a HPLC method was recently developed to separate and quantify 2-OH-E+ (O2−), ethidium (other ROS) and the residual DHE which has not been oxidised inside the cells (Fernandes et al., 2007; Kalyanaraman et al., 2013; Teng et al., 2012; Zhao et al., 2005). The results can be expressed semi-quantitatively as area under the curve, or the amount of 2-OH-E+ (O2−) can be quantified against a standard curve of O2− generated using the xanthine and xanthine oxidase system detected by the same equipment. Although the DHE fluorescence HPLC technique has the advantage of separating 2-OH-E+ from ethidium, it is not convenient for cell cycle study or drug compound toxicity screening. The technique requires an HPLC instrument equipped with a variety of detection systems including both fluorescence and UV detectors, and a mass spectrometer. The column needs to be often replaced due to blockages by cell/tissue extracts. Samples for HPLC need to be specifically prepared in relatively large quantities in order to get a decent signal of 2-OH-E+ and would not be suitable for any live cell study. The assay is not high throughput, due to the fact that the equilibration of HPLC is time consuming, and the column needs to be washed thoroughly between samples. Technically it is not straightforward and requires a skilled operator to run the instrument and to oversee the experiments. In comparison to other methods of ROS detection listed in this article, DHE fluorescence HPLC is not convenient when a large number of samples need to be processed. However, DHE HPLC is particularly useful if the specificity of O2− production by cells is a crucial element for the study.
P
216 217
assay or a cytotoxicity assay using the same format of a 96 well plate under the same experimental conditions. The levels of O2− production thus measured correlate closely with the stages of cell proliferation or death in the corresponding wells (Fan et al., 2009; Li et al., 2007; Tickner et al., 2011). The amount of O2− production can be quantified against a standard curve of O2− generated using the xanthine and xanthine oxidase system and detected on the same plate.
D
214 215
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
E
4
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
302 303 304 305 306 307 308 309 310 311 312 313 314 315 316 317 318 319 320 321 322 323 324 325 326 327 328 329 330 331 332 Q9 333 334 Q10 335 336 337 338
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
5
generate fluorescence. To avoid this, use of a low concentration of DHE is recommended. The DHE reagent should be kept in the dark and its incubation period with cells should be kept as short and constant as possible between samples.
380 381 382 383
8. Recording the intracellular ROS production by 5-(and 6)- 384 chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate 385 (CM-DCF-DA) fluorescence microscopy 386
F
CM-DCF-DA is a cell membrane permeable non-fluorescent reagent (Li et al., 2007; Touyz & Schiffrin, 1999). Once it diffuses across the membrane, the acetate groups of CM-DCF-DA are cleaved by intracellular esterases and CM-DCF is trapped inside the cell, and is oxidised by intracellular ROS to form a green fluorescent product (DCF, excitation/emission, in nm: 492–495/517–527)) in the cytoplasm, which can be measured using a fluorescence or confocal microscope. Compared to DHE, CM-DCF reacts slowly with intracellular ROS, and this provides a significant advantage as the CM-DCF oxidation can be microscopically recorded in real-time for cells cultured onto a special chamber or slide (Li et al., 2007) and the technique has been applied to a single isolated muscle fibre (Palomero, Pye, Kabayo, Spiller, & Jackson, 2008). The distribution pattern of DCF fluorescence in the cytoplasm can provide additional information on ROS generation in subcellular compartments. DCF labelled cells can also be fixed and their nuclei can be labelled by propidium iodide (Ex/Em: 536/617 nm, red fluorescence) and both fluorescence of DCF and propidium iodide can be imaged under a confocal microscope, which can give information of potential DNA damage i.e. nuclear fractionation (apoptosis) or mitosis (double nuclei) in proliferating cells.
Intracellular DHE fluorescence can also be detected by a fluorometer 346 plate reader using special 96 well plates that only allows light to pass 347 through the bottom of the well where the cells are adherent and alive 348 (Nazarewicz, Bikineyeva, & Dikalov, 2013). This technique is easy to 349 use, time saving and can be performed in parallel with an assay to detect 350 Q11 cell proliferation or drug toxicity using the same format of a 96 well 351 plate. This is very useful for pharmacological and toxicological screening 352 of compounds for oxidative damage of living cells. The limitation of the 353 fluorometer plate reader is that the detecting beam passes through only 354 a few points of the well and does not cover the entire bottom area of the 355 Q12 well so that the reading/per well is only an estimate (or representative) 356 value and is not the total cell ROS production in the well. The variations 357 between wells can be large and it is not reliable for cells in suspension 358 because the cells are moving around constantly in the well and we 359 would not know if the detecting beam actually passes the cells or not 360 and how many cells it may pass in one detection.
362 363 364 365 366 367 368 369 370 371 372 373 374 375 376 377 378 379
7. DHE fluorescence microscopy detection of intracellular ROS production in intact adherent cells DHE fluorescence can be detected in adherent living cells cultured onto chamber slides and then digitally imaged under a fluorescence microscope and quantified (Tickner et al., 2011). The DHE reaction can be stopped by fixing cells using paraformaldehyde and cells can be subsequently labelled in the dark using a different fluorescence probe (such as FITC) for a cell cycle molecule of interest. The comparison between the levels of ROS production and the level and subcellular location of that particular cell cycle molecule can provide important insights into the role of redox signalling in cell cycle progression. Due to the fact that ethidium fluorescence is from DNA in the nuclei, detailed observation of nuclei under microscopy may also provide useful information on cell apoptosis (nuclear fragmentation) or mitosis. The DHE fluorescence microscopy technique can also be used for fresh tissue cryosections to detect in situ ROS production (Du et al., 2013). In summary, DHE fluorescence is a powerful technique to detect intracellular ROS production by living cells, and is relatively more specific to O2− than to other species of ROS. DHE can be oxidised by air and
U
361
N C O
R
R
E
C
R O
T
345
P
6. DHE fluorescence plate-reader detection of intracellular ROS production in intact adherent cells cultured onto 96 well plates
D
343 344
389 390 391 392 393 394 395 396 397 398 399 400 401 402 403 404 405 406
E
341 342
With adequate controls, DHE fluorescence flow cytometry provides a useful and reliable technique that links intracellular ROS production to cell cycle progression (Ahmad et al., 2008; Du et al., 2009; Fan et al., 2009; McFarlin et al., 2013).
339 340
O
Fig. 2. DHE fluorescence cytometer for intracellular ROS production. U937 cells were cultured in 10% FCS medium, harvested and separated into 2 tubes. For cell cycle analysis, the DNA contents of cells were labelled by propidium iodide (Ex/Em: 536/617 nm) and the apoptotic cells (Apo) and cells in the Go/G1, S and G2.M phases were separated by their DNA contents using a flow cytometer (right panel). The intracellular O2− production was examined by a DHE fluorescence flow cytometer in the presence or absence of tiron (10 mmol/L), an O2− scavenger (left panel).
387 388
9. CM-DCF fluorescence flow cytometry detection of intracellular 407 ROS production 408 Like DHE fluorescence, DCF fluorescence can also be applied to cells in suspension and assessed by a flow cytometer (Ahmad et al., 2008; Du et al., 2009). This can be done in parallel with cell cycle analysis of the same sample and provide information on the redox state in comparison to cell cycle progression. Fig. 3A is a representative example of DCF fluorescence flow cytometry detection of intracellular ROS production by mouse primary microvascular endothelial cells in the presence or absence of tiron. Fig. 3B is a reprehensive example of cell cycle analysis. Compared to proliferating cells cultured in 10% FCS medium, quiescent cells (0% FCS medium for 24 h) generated less ROS. TNFα (100 U/mL, 24 h) stimulation significantly increased intracellular ROS production and this was accompanied by significant increases in cell apoptosis. The limitation is that: 1) DCF can be oxidised upon exposure to light and air resulting in a false increase in fluorescence. To avoid this, low light conditions should be used during experiments and whenever possible light protected tubes of aliquot should be used for each experiment. 2) DCF can be oxidised by a host of ROS such as nitric oxide, H2O2, peroxynitrite anions, and other hydroxyl radicals so it is not specific for detecting single species of ROS. The experimental setting should be carefully considered and a series of controls should be included in the experimental protocol. Despite these limitations, DCF remains a popular global ROS probe for detecting intracellular oxidants and has been applied to vascular endothelial cells (Fan et al., 2009; Teng et al., 2012), neurons (Kaur, Schulz, Heggland, Aschner, & Syversen, 2008), muscle fibres (Palomero et al., 2008) and human spermatozoa (Mahfouz et al., 2009). In summary, we have evaluated several ROS-detecting techniques that have the potential to be used for the study of redoxregulation of cell cycle progression and for pharmacological and toxicological screening of drug-related oxidative damage of cells or for other studies where the uses of these techniques are appropriate. The molecular properties and applications of ROS-detecting reagents, or their oxidative products, discussed in this article are given in Table 1. Fig. 4 summarises the advantages and limitations
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
409 410 411 412 413 414 415 416 417 418 419 420 421 Q13 422 423 424 425 426 427 428 429 430 431 432 433 434 435 436 437 438 439 440 441
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
C
T
E
D
P
R O
O
F
6
R
R
E
Fig. 3. Representative example of ROS production measurement using a CM-DCF fluorescence flow cytometer and cell cycle analysis. Mouse primary coronary microvascular endothelial cells were cultured for 24 h in 0% FCS medium (quiescent) or in 10% FCS growth medium (proliferating) or in 10% FCS growth medium plus TNFα (100 U/ml). Cells were detached and separated into 2 portions. The intracellular ROS production was detected by a DCF fluorescence flow cytometer with or without tiron (A). For cell cycle analysis, the DNA contents of cells were labelled by propidium iodide (Ex/Em: 536/617 nm) and the apoptotic cells (Apo) and cells in the Go/G1, S and G2.M phases were separated by their DNA contents using a flow cytometer (B).
t1:1 t1:2
Table 1 Properties and applications of ROS-detecting reagents discussed in the article.
C
point when the cell cycle is analysed and the cells can be fixed and dual-labelled with a cell differentiation marker. Both DHE and DCF fluorescence can be live-imaged by fluorescence microscopy or detected by a fluorometer. However, none of these techniques can distinguish the enzymatic source of ROS generation, therefore different enzyme inhibitors or specific ROS scavengers are needed to identify
N
446
U
444 445
O
447
of these techniques. Lucigenin (5 μM)-chemiluminescence provides a kinetic analysis of real-time production of O2− and is sensitive for detecting low levels of O2− production by non-phagocytic cells. DHE fluorescence HPLC detects specifically O 2 − and can separate O 2 − from other species of ROS but is time consuming. DHE fluorescence flow cytometry gives an actual redox profile at a particular time
442 443
Mol. weight
Extinction Emission (nm)
Assay final concentration
ROS detected
Lucigenin Ferricytochrome c Dihydroethidium (DHE)
510.5 12.4 (equine) 315.4
5 μmol/L 250 μmol/L 2 μmol/L
O2− All ROS non specific All ROS preference to O2−
Ethidium (E+)
394.3
DHE oxidation product
Other ROS except O2−
2-Hydroxyethidium (2-OH-E+)
330.2
DHE oxidation product
O2−
2′,7′-Dichlorodihydrofluorescein diacetate (DCF-DA) 2′,7′-Dichlorodihydrofluorescein (DCF)
487.3 401.2
Ex: 368 nm Em: 505 nm UV absorption λmax = 550 nm (reduced form) Fluorescence Ex: 526 nm Em: 605 nm Ex: 526 nm, Em: 605 nm Ex: 396 nm Em: 579 nm Non-fluorescence Ex: 492 nm Em: 517 nm
5 μmol/L DCF-DA cleaved product
All ROS non specific All ROS non specific
t1:3
Reagent
t1:4 t1:5 t1:6 t1:7 t1:8 t1:9 t1:10 t1:11 t1:12 t1:13 t1:14 t1:15
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
448 449 450 451 452 453
7
R O
O
F
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
Fig. 4. Diagrammatic summary of ROS-detecting methods discussed in this article for drug cytotoxicity screening or investigation of the role of ROS in cell function.
458 459 460
LF: Manuscript preparation and figure generation; JML: directed, reviewed and edited the manuscript. All authors have approved the final article and its submission.
461
Conflict of interest statement There is no conflict of interest.
C
462
E
463 Q14 Uncited reference
Kalyanaraman et al., 2014
R
464
References
466 467 468 469 470 471 472 473 474 475 476 477 478 479 480 481 482 483 484 485 486 487 488 489 490 491 492 493 494 495 496 497
Afanas'ev, I. B., Ostrakhovitch, E. A., Mikhal'chik, E. V., & Korkina, L. G. (2001). Direct enzymatic reduction of lucigenin decreases lucigenin-amplified chemiluminescence produced by superoxide ion. Luminescence, 16, 305–307. Ahmad, I. M., Abdalla, M. Y., Aykin-Burns, N., Simons, A. L., Oberley, L. W., Domann, F. E., et al. (2008). 2-Deoxyglucose combined with wild-type p53 overexpression enhances cytotoxicity in human prostate cancer cells via oxidative stress. Free Radical Biology and Medicine, 44, 826–834. Ardanaz, N., & Pagano, P. J. (2006). Hydrogen peroxide as a paracrine vascular mediator: Regulation and signaling leading to dysfunction. Experimental Biology and Medicine, 231, 237–251. Berasi, S. P., Xiu, M., Yee, A. S., & Paulson, K. E. (2004). HBP1 repression of the p47phox gene: Cell cycle regulation via the NADPH oxidase. Molecular and Cellular Biology, 24, 3011–3024. Biswas, S., Chida, A. S., & Rahman, I. (2006). Redox modifications of protein-thiols: Emerging roles in cell signaling. Biochemical Pharmacology, 71, 551–564. Borgmann, S. (2009). Electrochemical quantification of reactive oxygen and nitrogen: Challenges and opportunities. Analytical and Bioanalytical Chemistry, 394, 95–105. Brandes, R. P., Miller, F. J., Beer, S., Haendeler, J., Hoffmann, J., Ha, T., et al. (2002). The vascular NADPH oxidase subunit p47phox is involved in redox-mediated gene expression. Free Radical Biology and Medicine, 32, 1116–1122. Burch, P. M., & Heintz, N. H. (2005). Redox regulation of cell-cycle re-entry: Cyclin D1 as a primary target for the mitogenic effects of reactive oxygen and nitrogen species. Antioxidants and Redox Signaling, 7, 741–751. Burhans, W. C., & Heintz, N. H. (2009). The cell cycle is a redox cycle: Linking phasespecific targets to cell fate. Free Radical Biology and Medicine, 47, 1282–1293. Chamseddine, A. H., & Miller, F. J., Jr. (2003). Gp91phox contributes to NADPH oxidase activity in aortic fibroblasts but not smooth muscle cells. American Journal of Physiology — Heart and Circulatory Physiology, 285, H2284–H2289. Circu, M. L., & Aw, T. Y. (2010). Reactive oxygen species, cellular redox systems, and apoptosis. Free Radical Biology and Medicine, 48, 749–762. Dikalov, S., Griendling, K. K., & Harrison, D. G. (2007). Measurement of reactive oxygen species in cardiovascular studies. Hypertension, 49, 717–727.
N C O
R
465
U
P
Author contribution
Dröge, W. (2002). Free radicals in the physiological control of cell function. Physiological Reviews, 82, 47–95. Du, J., Fan, L. M., Mai, A., & Li, J. -M. (2013). Crucial roles of Nox2-derived oxidative stress in deteriorating the function of insulin receptors and endothelium in dietary obesity of middle-aged mice. British Journal of Pharmacology, 170, 1064–1077. Du, C., Gao, Z., Venkatesha, V. A., Kalen, A. L., Chaudhuri, L., Spitz, D. R., et al. (2009). Mitochondrial ROS and radiation induced transformation in mouse embryonic fibroblasts. Cancer Biology and Therapy, 8, 1962–1971. Fan, L. M., Teng, L., & Li, J. -M. (2009). Knockout of p47 phox uncovers a critical role of p40 phox in reactive oxygen species production in microvascular endothelial cells. Arteriosclerosis, Thrombosis, and Vascular Biology, 29, 1651–1656. Fernandes, D. C., Wosniak, J., Jr., Pescatore, L. A., Bertoline, M. A., Liberman, M., Laurindo, F. R., et al. (2007). Analysis of DHE-derived oxidation products by HPLC in the assessment of superoxide production and NADPH oxidase activity in vascular systems. American Journal of Physiology — Cell Physiology, 292, C413–C422. Finkel, T. (2011). Signal transduction by reactive oxygen species. Journal of Cell Biology, 194, 7–15. Freitas, M., Porto, G., Lima, J. L., & Fernandes, E. (2009). Optimization of experimental settings for the analysis of human neutrophils oxidative burst in vitro. Talanta, 78, 1476–1483. Görlach, A., Kietzmann, T., & Hess, J. (2002). Redox signaling through NADPH oxidases: Involvement in vascular proliferation and coagulation. Annals of the New York Academy of Sciences, 973, 505–507. Gutteridge, J. M., & Halliwell, B. (2010). Antioxidants: Molecules, medicines, and myths. Biochemical and Biophysical Research Communications, 393, 561–564. Halliwell, B., & Whiteman, M. (2004). Measuring reactive species and oxidative damage in vivo and in cell culture: How should you do it and what do the results mean? British Journal of Pharmacology, 142, 231–255. Hannken, T., Schroeder, R., Stahl, R. A., & Wolf, G. (1998). Angiotensin II-mediated expression of p27Kip1 and induction of cellular hypertrophy in renal tubular cells depend on the generation of oxygen radicals. Kidney International, 54, 1923–1933. Kalyanaraman, B., Darley-Usmar, V., Davies, K. J., Dennery, P. A., Forman, H. J., Grisham, M. B., et al. (2012). Measuring reactive oxygen and nitrogen species with fluorescent probes: Challenges and limitations. Free Radical Biology and Medicine, 52, 1–6. Kalyanaraman, B., Dranka, B. P., Hardy, M., Michalski, R., & Zielonka, J. (2014). HPLC-based monitoring of products formed from hydroethidine-based fluorogenic probes — the ultimate approach for intra- and extracellular superoxide detection. Biochimica et Biophysica Acta, 1840, 739–744. Kaur, P., Schulz, K., Heggland, I., Aschner, M., & Syversen, T. (2008). The use of fluorescence for detecting MeHg-induced ROS in cell cultures. Toxicology In Vitro, 22, 1392–1398. Li, J. -M., Fan, L. M., George, V. T., & Brooks, G. (2007). Nox2 regulates endothelial cell cycle arrest and apoptosis via p21cip1 and p53. Free Radical Biology and Medicine, 43, 976–986. Li, J. -M., Mullen, A. M., & Shah, A. M. (2001). Phenotypic properties and characteristics of superoxide production by mouse coronary microvascular endothelial cells. Journal of Molecular and Cellular Cardiology, 33, 1119–1131. Li, J. -M., & Shah, A. M. (2001). Differential NADPH- versus NADH-dependent superoxide production by phagocyte-type endothelial cell NADPH oxidase. Cardiovascular Research, 52, 477–486. Li, J. -M., & Shah, A. M. (2004). Endothelial cell superoxide generation: Regulation and relevance for cardiovascular pathophysiology. American Journal of Physiology — Regulatory, Integrative and Comparative Physiology, 287, R1014–R1030. Li, Y., Zhu, H., Kuppusamy, P., Roubaud, V., Zweier, J. L., & Trush, M. A. (1998). Validation of lucigenin (bis-N-methylacridinium) as a chemilumigenic probe for detecting superoxide anion radical production by enzymatic and cellular systems. Journal of Biological Chemistry, 273, 2015–2023. Liochev, S. I., & Fridovich, I. (1997). Lucigenin (bis-N-methylacridinium) as a mediator of superoxide anion production. Archives of Biochemistry and Biophysics, 337, 115–120.
D
457
T
456
the source of ROS generation and the type of ROS detected. Using a combination of two or three techniques will provide accurate information on ROS generation for the study.
E
454 455
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
498 499 500 501 502 503 504 505 506 507 508 509 510 511 512 513 514 515 516 517 518 519 520 521 522 523 524 525 526 527 528 529 530 531 532 533 534 535 536 537 538 539 540 541 542 543 544 545 546 547 548 549 550 551 552 553 554 555 556
O
F
Teng, L., Fan, L. M., Meijles, D., & Li, J. -M. (2012). Divergent effects of p47(phox) phosphorylation at S303-4 or S379 on tumor necrosis factor-a signaling via TRAF4 and MAPK in endothelial cells. Arteriosclerosis, Thrombosis, and Vascular Biology, 32, 1488–1496. Tickner, J., Fan, L. M., Du, J., Meijles, D., & Li, J. -M. (2011). Nox2-derived ROS in PPARγ signaling and cell-cycle progression of lung alveolar epithelial cells. Free Radical Biology and Medicine, 51, 763–772. Touyz, R. M., & Schiffrin, E. L. (1999). Ang II-stimulated superoxide production is mediated via phospholipase D in human vascular smooth muscle cells. Hypertension, 34, 976–982. Valko, M., Leibfritz, D., Moncol, J., Cronin, M. T., Mazur, M., & Telser, J. (2007). Free radicals and antioxidants in normal physiological functions and human disease. International Journal of Biochemistry and Cell Biology, 39, 44–84. Venkatachalam, P., de Toledo, S. M., Pandey, B. N., Tephly, L. A., Carter, A. B., Little, J. B., et al. (2008). Regulation of normal cell cycle progression by flavin-containing oxidases. Oncogene, 27, 20–31. Verbon, E. H., Post, J. A., & Boonstra, J. (2012). The influence of reactive oxygen species on cell cycle progression in mammalian cells. Gene, 511, 1–6. Vokurkova, M., Xu, S., & Touyz, R. M. (2007). Reactive oxygen species, cell growth, cell cycle progression and vascular remodeling in hypertension. Future Cardiology, 3, 53–63. Vurusaner, B., Poli, G., & Basaga, H. (2012). Tumor suppressor genes and ROS: Complex networks of interactions. Free Radical Biology and Medicine, 52, 7–18. Whiteman, M., Hong, H. S., Jenner, A., & Halliwell, B. (2002). Loss of oxidized and chlorinated bases in DNA treated with reactive oxygen species: Implications for assessment of oxidative damage in vivo. Biochemical and Biophysical Research Communications, 296, 883–889. Zhao, H., Joseph, J., Fales, H. M., Sokoloski, E. A., Levine, R. L., Vasquez-Vivar, J., et al. (2005). Detection and characterization of the product of hydroethidine and intracellular superoxide by HPLC and limitations of fluorescence. PNAS, 102, 5727–5732. Zhao, H., Kalivendi, S., Zhang, H., Joseph, J., Nithipatikom, K., Vasquez-Vivar, J., et al. (2003). Superoxide reacts with hydroethidine but forms a fluorescent product that is distinctly different from ethidium: Potential implications in intracellular fluorescence detection of superoxide. Free Radical Biology and Medicine, 34, 1359–1368. Zielonka, J., Vasquez-Vivar, J., & Kalyanaraman, B. (2008). Detection of 2-hydroxyethidium in cellular systems: A unique marker product of superoxide and hydroethidine. Nature Protocols, 3, 8–21.
R O
Maghzal, G. J., Krause, K. -H., Stocker, R., & Jaquet, V. (2012). Detection of reactive oxygen species derived from the family of Nox NADPH oxidases. Free Radical Biology and Medicine, 53, 1903–1918. Mahfouz, R., Sharma, R., Lackner, J., Aziz, N., & Agarwal, A. (2009). Evaluation of chemiluminescence and flow cytometry as tools in assessing production of hydrogen peroxide and superoxide anion in human spermatozoa. Fertility and Sterility, 92, 819–827. Maryanovich, M., & Gross, A. (2013). A ROS rheostat for cell fate regulation. Trends in Cell Biology, 23, 129–134. McFarlin, B. K., Williams, R. R., Venable, A. S., Dwyer, K. C., & Haviland, D. L. (2013). Imagebased cytometry reveals three distinct subsets of activated granulocytes based on phagocytosis and oxidative burst. Cytometry. Part A, 83, 745–751. Menon, S. G., & Goswami, P. C. (2007). A redox cycle within the cell cycle: Ring in the old with the new. Oncogene, 26, 1101–1109. Menshikov, M., Plekhanova, O., Cai, H., Chalupsky, K., Parfyonova, Y., Bashtrikov, P., et al. (2006). Urokinase plasminogen activator stimulates vascular smooth muscle cell proliferation via redox-dependent pathways. Arteriosclerosis, Thrombosis, and Vascular Biology, 26, 801–807. Nauseef, W. M. (2014). Detection of superoxide anion and hydrogen peroxide production by cellular NADPH oxidase. Biochimica et Biophysica Acta, 1840, 757–767. http://dx. doi.org/10.1016/j.bbagen.2013.04.040. Nazarewicz, R. R., Bikineyeva, A., & Dikalov, S. I. (2013). Rapid and specific measurements of superoxide using fluorescence spectroscopy. Journal of Biomolecular Screening, 18, 498–503. Palomero, J., Pye, D., Kabayo, T., Spiller, D. G., & Jackson, M. J. (2008). In situ detection and measurement of intracellular reactive oxygen species in single isolated mature skeletal muscle fibers by real time fluorescence microscopy. Antioxidants and Redox Signaling, 10, 1463–1474. Sohn, H. -Y., Keller, M., Gloe, T., Crause, P., & Pohl, U. (2000). Pitfalls of using lucigenin in endothelial cells: Implications for NAD(P)H dependent superoxide formation. Free Radical Research, 32, 265–272. Tarpey, M. M., & Fridovich, I. (2001). Methods of detection of vascular reactive species. Circulation Research, 89, 224–236. Tarpey, M. M., Wink, D. A., & Grisham, M. B. (2004). Methods for detection of reactive metabolites of oxygen and nitrogen: In vitro and in vivo considerations. American Journal of Physiology — Regulatory, Integrative and Comparative Physiology, 286, R431–R444.
D
557 558 559 560 561 562 563 564 565 566 567 568 569 570 571 572 573 574 575 576 577 578 579 580 581 582 583 584 585 586 587 588 589 590 591
L.M. Fan, J.-M. Li / Journal of Pharmacological and Toxicological Methods xxx (2014) xxx–xxx
P
8
U
N
C
O
R
R
E
C
T
E
627
Please cite this article as: Fan, L.M., & Li, J.-M., Evaluation of methods of detecting cell reactive oxygen species production for drug screening and cell cycle studies, Journal of Pharmacological and Toxicological Methods (2014), http://dx.doi.org/10.1016/j.vascn.2014.03.173
592 593 594 595 596 597 598 599 600 601 602 603 604 605 606 607 608 609 610 611 612 613 614 615 616 617 618 619 620 621 622 623 624 625 626
