This work was funded by the Natural Sciences and Engineering Research Council of Canada (NSERC). Video production was carried out in collaboration with the Center for Innovation in Teaching and Learning (CITL) at Memorial University of Newfoundland.

1. Chemicals and Purified Enzymes
<ol>
	<li>Procure the following materials: PDHC and KGDHC of porcine heart origin (or another purified mitochondrial flavoenzyme); H2O2 (30% solution), pyruvate, &#945;-ketoglutarate, NAD+, NADH, CoASH, thiamine pyrophosphate (TPP), mannitol, HEPES, sucrose, EGTA, 3-methyl-2-oxo valeric acid (KMV), superoxide dismutase (SOD), horseradish peroxidase (HRP), 10-Acetyl-3,7-dihydroxyphenoxazine reagent (AUR), and CPI-613.</li>
</ol>
2. Planning the Assay and Reagent Preparation
<ol>
	<li>Set up the assay according to Table 1 in a 96-well plate. 
	NOTE: The total volume for each reaction is 200 &#181;L. Stock concentrations are adjusted such that 20 &#181;L of reagent is added to each well. This ensures rapid addition of cofactors and substrates to each reaction well prior to starting the assay.</li>
	<li>Prepare reaction buffer containing 220 mM mannitol, 70 mM sucrose, 1 mM EGTA, and 20 mM HEPES (pH 7.4 with HCl). 
	NOTE: This could change depending on the physical properties of different purified enzymes.</li>
	<li>Examine the purified KGDHC or PDHC for contamination using enzyme activity assays14,16,17 or by immunoblot or Coomassie stain5.</li>
	<li>Prepare all reagents in the reaction buffer as per the working solution concentrations in Table 1.</li>
	<li>Store all reagents as 1 mL aliquots at -20 &#176;C. Store pyruvate and &#945;-ketoglutarate at -80 &#176;C in 100-&#181;L aliquots to prevent spontaneous degradation due to auto-oxidation and repeated freeze-thaw.</li>
	<li>Prepare the AUR reagent master stock in 1 mL of DMSO and then dilute to 100 &#181;M in reaction buffer. Store protected from light. 
	NOTE: Here, the 100 &#181;M working solution is made fresh every 2 to 3 weeks and protected from light.</li>
</ol>
3. Setting Up the Assay Template
<ol>
	<li>Set up the assay procedure on a monochromatic microplate reader with dual measurement capabilities.</li>
	<li>Define the reading procedure by selecting &#34;PROTOCOL&#34;. Then click &#34;PROCEDURE&#34;.</li>
	<li>Set &#34;TEMPERATURE&#34; to 25 &#176;C (Figure 1A).</li>
	<li>Click &#34;START KINETIC&#34; to set read conditions (Figure 1A). Set the time and number of read intervals/experiments.</li>
	<li>Click &#34;READ&#34; (Figure 1B). 
	NOTE: Samples are read from TOP position with a gain of 50. Time: 5 min, read at 30-s intervals. Channel 1: Resorufin fluorescence - excitation/emission = 565 nm:600 nm, wavelength width of 13.5 nm. Channel 2: NADH autofluorescence - excitation/emission = 376 nm:450 nm, wavelength width of 20 nm.</li>
	<li>Under &#34;READ&#34;, select wells to monitor (e.g., A1-A4 and B1-B4).</li>
	<li>Select &#34;END KINETIC&#34;.</li>
</ol>
4. Standard Curves
<ol>
	<li>Thaw the reagents and store on ice until needed.</li>
	<li>NADH standard curve (Figure 2A)
	<ol>
		<li>Prepare the working solutions for NADH in the concentration range of 0.5-10 mM in reaction buffer.</li>
		<li>Add 20 &#181;L aliquots from each NADH working solution concentration to wells containing 180 &#181;L reaction buffer. The final concentration of NADH in each well is 0.05-1 mM.</li>
		<li>Click &#34;READ&#34; and set excitation/emission = 376 nm:450 nm, wavelength width of 20 nm. 
		NOTE: This is an endpoint read only - kinetic parameters are not required.</li>
	</ol>
	</li>
	<li>AUR standard curve (Figure 2B)
	<ol>
		<li>Prepare the working stocks of hydrogen peroxide in reaction buffer; stock concentrations range from 200-4,000 nM.</li>
		<li>Add 120 &#181;L of the reaction buffer to each well.</li>
		<li>Add 20 &#181;L of HRP (working stock concentration = 30 U/mL, final concentration in the well = 3 U/mL), 20 &#181;L of SOD (working stock concentration = 250 U/mL, final concentration in the well = 25 U/mL), and 20 &#181;L of AUR (working stock concentration = 100 &#181;M, final concentration in the well = 10 &#181;M) to each well containing reaction buffer.</li>
		<li>Add 20 &#181;L of each H2O2 working stock solution to each well containing buffer and assay reagents. The final reaction volume is 200 &#181;L and the final concentration of H2O2 in each well is 20-400 nM.</li>
		<li>Incubate the plates for 1 min in the plate reader at 25 &#176;C.</li>
		<li>Click &#34;READ&#34; and set excitation/emission = 565 nm/600 nm, wavelength width of 13.5 nm. Note that this is an endpoint read only - kinetic parameters are not required.</li>
	</ol>
	</li>
</ol>
5. Measuring O2 &#9679;- /H2 O2 Release and NADH Production by KGDHC and PDHC
<ol>
	<li>See Table 1 for the order of addition of the different reagents, the concentration of each working solution, and the volume added to each well.</li>
	<li>Add 40 &#181;L of reaction buffer to each well. For assays using KMV or CPI-613, add 20 &#181;L of buffer to each well.</li>
	<li>Add 20 &#181;L of PDHC or KGDHC (working stock of 1 U/mL, final concentration per well = 0.1 U/mL) to each well.</li>
	<li>Incubate PDHC and KGDHC at 25 &#176;C for 2 min.</li>
	<li>Add 20 &#181;L of KMV (working stock = 100 mM, final concentration = 10 mM) or 20 &#181;L of CPI-613 (working stock concentration = 1.5 mM, final concentration = 150 &#181;M) (Table 2). 
	NOTE: This step can be excluded if inhibitors are not being used for the assays.</li>
	<li>Incubate for 1 min at 25 &#176;C. 
	NOTE: This step can be excluded if inhibitors are not being used for the assays.</li>
	<li>Add 20 &#181;L of HRP (working stock concentration = 30 units/mL, final concentration in the well = 3 units/mL) and 20 &#181;L of SOD (working stock concentration = 250 units/mL, final concentration in the well = 25 units/mL) to each well.</li>
	<li>Add 20 &#181;L of coenzyme A (working stock concentration = 1 mM, final concentration = 0.1 mM), 20 &#181;L of TPP (working stock concentration = 3 mM, final concentration = 0.3 mM), and 20 &#181;L of NAD+ (working stock concentration = 10 mM, final concentration = 1 mM) to each well.</li>
	<li>Remove the AUR reagent from protective tinfoil covering and add 20 &#181;L to each well.</li>
	<li>Add 20 &#181;L of pyruvate (working stock concentration ranging from 1-100 mM, final concentration for assays = 0.1-10 mM) to wells containing PDHC and &#945;-ketoglutarate (working stock concentration ranging from 1-100 mM, final concentration for assays = 0.1-10 mM) to wells containing KGDHC.</li>
	<li>Click &#34;READ&#34; to start kinetic measurement.</li>
</ol>
6. Data Analysis
<ol>
	<li>Hold down the &#39;CTRL&#39; button on the keyboard and click wells to generate one graph containing all kinetic measurements corresponding to channel 1 (Figure 3A).</li>
	<li>Click &#34;data&#34; in the view icon in the upper right corner for raw values for relative fluorescence units (RFU) measured at the different time points (Figure 3B).</li>
	<li>Export the values for analysis (Table 3).</li>
	<li>Click the &#34;Graph&#34; drop down menu on top right (Figure 3B) to access RFU values associated with the fluorescence channel 2.</li>
	<li>Repeat steps 6.1 to 6.3 for channel 2.</li>
</ol>

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J., Gardiner, D., O'Brien, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=2-Oxoglutarate+dehydrogenase+is+a+more+significant+source+of+O2(.-)/H2O2+than+pyruvate+dehydrogenase+in+cardiac+and+liver+tissue.">2-Oxoglutarate dehydrogenase is a more significant source of O2(.-)/H2O2 than pyruvate dehydrogenase in cardiac and liver tissue.</a> Free Radic Biol Med. 97, 501-512 (2016).</li><li>Ambrus, A., Adam-Vizi, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+dihydrolipoamide+dehydrogenase+(E3)+deficiency:+Novel+insights+into+the+structural+basis+and+molecular+pathomechanism.">Human dihydrolipoamide dehydrogenase (E3) deficiency: Novel insights into the structural basis and molecular pathomechanism.</a> Neurochem Int. , (2017).</li><li>Young, A., Gardiner, D., Brosnan, M. E., Brosnan, J. T., Mailloux, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physiological+levels+of+formate+activate+mitochondrial+superoxide/hydrogen+peroxide+release+from+mouse+liver+mitochondria.">Physiological levels of formate activate mitochondrial superoxide/hydrogen peroxide release from mouse liver mitochondria.</a> FEBS Lett. 591 (16), 2426-2438 (2017).</li><li>Fisher-Wellman, K. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+glutathione+depletion+reveals+a+novel+role+for+the+pyruvate+dehydrogenase+complex+as+a+key+H2O2-emitting+source+under+conditions+of+nutrient+overload.">Mitochondrial glutathione depletion reveals a novel role for the pyruvate dehydrogenase complex as a key H2O2-emitting source under conditions of nutrient overload.</a> Free Radic Biol Med. 65, 1201-1208 (2013).</li><li>Kussmaul, L., Hirst, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+mechanism+of+superoxide+production+by+NADH:ubiquinone+oxidoreductase+(complex+I)+from+bovine+heart+mitochondria.">The mechanism of superoxide production by NADH:ubiquinone oxidoreductase (complex I) from bovine heart mitochondria.</a> Proc Natl Acad Sci U S A. 103 (20), 7607-7612 (2006).</li><li>Huang, L. 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There is nothing to disclose.

This protocol is advantageous since, 1) it eliminates any competing reactions that may otherwise interfere with H2O2 detection (e.g., antioxidant systems or other sources of ROS), 2) provides a direct assessment of the native rate of ROS release by a flavin-containing mitochondrial dehydrogenase, 3) allows the comparison of the native ROS release rates of two or more purified flavin-based dehydrogenases, 4) can allow for a direct comparison of the rate of ROS release and enzyme activity, an...

The ultimate goal of nutrient metabolism is to make adenosine triphosphate (ATP). In mammalian cells, this occurs in mitochondria, double-membraned organelles that convert the energy stored in carbon into ATP. The production of ATP begins when carbon is combusted by mitochondria forming two electron carriers, NADH and flavin adenine dinucleotide (FADH2)1. NADH and FADH2 are then oxidized by multi-subunit respiratory complexes I and II, respectively, and the liberated electrons are ferried to the terminal electron acceptor molecular oxygen (O2) at complex IV1. The thermodynamically favorable &#34;downhill&#34; transfer of electrons to O2 at the end of the chain is coupled to the export of protons into the intermembrane space by complexes I, III, and IV. This creates a transmembrane electrochemical gradient of protons (proton-motive force), a temporary form of Gibbs free energy that is tapped by complex V to make ATP2. Electron transfer reactions in mitochondria are not perfectly coupled to ATP production. At various points in the Krebs cycle and the respiratory chain, electrons can prematurely interact with O2 to form ROS3. The most biologically relevant ROS generated by mitochondria are O2&#9679;- and H2O2. Although O2&#9679;- is often considered the proximal ROS formed by mitochondria, it is now evident that sites of production form a mixture of O2&#9679;- and H2O2, which is associated with the free radical chemistry of flavin prosthetic groups4,5. At high levels, ROS can be dangerous, damaging biological constituents required for cell function resulting in oxidative distress6. However, at low amounts, mitochondrial ROS fulfill vital signaling functions. For instance, H2O2 release from mitochondria has been implicated in controlling T-cell activation, stress signaling (e.g., induction of Nrf2 signaling pathways), the induction of cell proliferation and differentiation, insulin signaling and release, and feeding behavior7. Considerable progress has been made in understanding the signaling function of ROS. However, important questions still remain in regard to which enzymes in mitochondria serve as the most important sources and how production is controlled.
ROS release by a site of production depends on several factors: 1) the concentration of the electron donating site, 2) the redox state of the electron donating site, 3) access to oxygen, and 4) the concentration and type of oxidation substrate3,8,9. In mitochondria, other factors like the concentration of NADH and membrane potential strength also influence ROS production8,10. For example, the rate of O2&#9679;-/H2O2 production by purified PDHC or KGDHC increases with increasing NADH availability5,11. In this scenario, electrons are flowing backwards from NADH to the FAD center in the E3 subunit of PDHC or KGDHC, the site for O2&#9679;-/H2O2 production12. Similarly, the provision of NAD+ has the opposite effect, decreasing ROS release from KGDHC12. Thus, controlling entry or exit of electrons from sites of ROS production can alter how much O2&#9679;-/H2O2 is formed. For instance, blocking the E2 subunit of PDHC or KGDHC with CPI-613, a lipoic acid analog, results in an almost 90% decrease in O2&#9679;-/H2O2 production13. Similar results can be obtained with the chemical S-glutathionylation catalysts, diamide and disulfiram, which almost abolish O2&#9679;-/H2O2 production by PDHC or KGDHC via the conjugation of glutathione to the E2 subunit14. The trade-off for blocking electron flow through the E2 subunit of PDHC and KGDHC is a decrease in NADH production which diminishes ROS formation by the electron transport chain (e.g., complex III). This can also decrease ATP output by the electron transport chain. Overall, blocking electron entry to sites of ROS production can be a highly effective means of controlling O2&#9679;-/H2O2 production.
Mitochondria can contain up to 12 potential O2&#9679;-/H2O2 sources6. Most of these sites are flavin-containing enzymes which generate a mixture of O2&#9679;- and H2O2. Use of different substrate and inhibitor combinations have allowed for the identification of which respiratory complexes and mitochondrial dehydrogenases serve as high capacity O2&#9679;-/H2O2 forming sites in different tissues3. PDHC and KGDHC have been shown to serve as high capacity O2&#9679;-/H2O2 emitting sites in muscle and liver mitochondria13,15. However, some difficulties remain in regard to the examination of the O2&#9679;-/H2O2 forming potential of individual sites in mitochondria and the impact that different substrate and inhibitor combinations have on the activities of the enzymes. This is due to the presence of unwanted side reactions (e.g., formation of O2&#9679;-/H2O2 by sites other than the enzyme of interest), contaminating endogenous nutrients (e.g.,fatty acids) or organelles (e.g., peroxisomes which also form O2&#9679;-/H2O2), and use of inhibitors that lack selectivity, and/or use of compounds that do not fully inhibit ROS production. Certain inhibitors can also alter the mitochondrial bioenergetics and direction of electron flow, and this alters ROS release from other sites of production and confounds results. Absolute rates for O2&#9679;-/H2O2 release from individual sites in mitochondria are also difficult to quantify due to the high concentration of O2&#9679;- and H2O2 eliminating enzymes in the matrix and intermembrane space. Therefore, elimination of any competing reactions that can interfere with O2&#9679;-/H2O2 release measurements can be useful when identifying high capacity O2&#9679;-/H2O2 forming sites.
Here, we present a simple method that allows for the simultaneous examination of O2&#9679;-/H2O2 production and NADH formation by purified flavin-dependent dehydrogenases. By using purified enzymes, unwanted ROS forming side reactions and ROS degrading enzymes can be eliminated allowing for a more accurate measure of native O2&#9679;-/H2O2 production rates for individual flavoenzymes. This method can be used to directly compare the O2&#9679;-/H2O2 forming capacity of different purified dehydrogenases or to screen potential site-specific inhibitors for O2&#9679;-/H2O2 release. Finally, measuring O2&#9679;-/H2O2 and NADH production simultaneously can allow for a real-time assessment of the relationship between enzyme activity and ROS release capacity.

<table><tbody><tr><td>Pyruvate dehydrogenase complex</td><td>SIGMA</td><td>P7032-10UN</td><td>purified flavoenzyme</td></tr><tr><td>alpha-ketoglutarate dehydrogenase complex</td><td>SIGMA</td><td>K1502-20UN</td><td>purified flavoenzyme</td></tr><tr><td>30% hydrogen peroxide solution</td><td>SIGMA</td><td>HX0640-5</td><td>reagent, standard curves</td></tr><tr><td>NAD+</td><td>SIGMA</td><td>N0632-1G</td><td>reagent, activity/ROS release assay</td></tr><tr><td>NADH</td><td>SIGMA</td><td>N4505-100MG</td><td>reagent, standard curves</td></tr><tr><td>pyruvate</td><td>SIGMA</td><td>P2256-5G</td><td>reagent, activity/ROS release assay</td></tr><tr><td>alpha-ketoglutarate</td><td>SIGMA</td><td>75892-25G</td><td>reagent, activity/ROS release assay</td></tr><tr><td>CoASH</td><td>SIGMA</td><td>C3019-25MG</td><td>reagent, activity/ROS release assay</td></tr><tr><td>thiamine pyrophosphate</td><td>SIGMA</td><td>C8754-1G</td><td>reagent, activity/ROS release assay</td></tr><tr><td>mannitol</td><td>SIGMA</td><td>M4125-100G</td><td>buffer component</td></tr><tr><td>Hepes</td><td>SIGMA</td><td>H3375-25G</td><td>buffer component</td></tr><tr><td>sucrose</td><td>SIGMA</td><td>S7903-250G</td><td>buffer component</td></tr><tr><td>EGTA</td><td>SIGMA</td><td>E3889-10G</td><td>buffer component</td></tr><tr><td>KMV</td><td>SIGMA</td><td>198978-5G</td><td>reagent, ROS release inhibitor</td></tr><tr><td>CPI-613</td><td>Santa Cruz</td><td>sc-482709</td><td>reagent, ROS release inhibitor</td></tr><tr><td>SOD</td><td>SIGMA</td><td>S9697-15KU</td><td>reagent, ROS release detection</td></tr><tr><td>horseradish peroxidase</td><td>SIGMA</td><td>P8375-1KU</td><td>reagent, ROS release detection</td></tr><tr><td>Amplex Ultra Red</td><td>Thermofisher</td><td>A36006</td><td>reagent, ROS release detection</td></tr><tr><td>Biotech Synergy 2 microplate reader</td><td>BioTek Instruments</td><td></td><td>microplate reader for assays</td></tr><tr><td>Gen5 software</td><td>BioTek Instruments</td><td></td><td>software, used for collection of raw RFU</td></tr><tr><td>Graphpad Prism</td><td>Graphpad software</td><td></td><td>software, data analysis</td></tr><tr><td>Microsoft EXCEL</td><td>Microsoft</td><td></td><td>software, data analysis</td></tr></tbody></table>

simultaneous measurement of superoxide/hydrogen peroxide and nadh production by flavin-containing mitochondrial dehydrogenases

It has been reported that mitochondria can contain up to 12 enzymatic sources of reactive oxygen species (ROS). A majority of these sites include flavin-dependent respiratory complexes and dehydrogenases that produce a mixture of superoxide (O2&#9679;-) and hydrogen peroxide (H2O2). Accurate quantification of the ROS-producing potential of individual sites in isolated mitochondria can be challenging due to the presence of antioxidant defense systems&#160;and side reactions that also form O2&#9679;-/H2O2. Use&#160;of nonspecific inhibitors that can disrupt mitochondrial bioenergetics can also compromise measurements by altering&#160;ROS release from other sites of production. Here, we present an easy method for the simultaneous measurement of H2O2 release and nicotinamide adenine dinucleotide (NADH) production by purified flavin-linked dehydrogenases. For our purposes here, we have used purified pyruvate dehydrogenase complex (PDHC) and &#945;-ketoglutarate dehydrogenase complex (KGDHC) of porcine heart origin as examples. This method allows for an accurate measure of native H2O2 release rates by individual sites of production by eliminating other potential sources of ROS and antioxidant systems. In addition, this method allows for a direct comparison of the relationship between H2O2 release and enzyme activity and the screening of the effectiveness and selectivity of inhibitors for ROS production. Overall, this approach can allow for the in-depth assessment of native rates of ROS release for individual enzymes prior to conducting more sophisticated experiments with isolated mitochondria or permeabilized muscle fiber.

Figure 3A provides a representative trace for the RFU collected during the simultaneous measurement of H2O2 and NADH production by purified KGDHC. The raw RFU data for each time interval is depicted in Figure 3B. The raw RFU data are then exported for analysis. By extrapolating from standard curves presented in Figure 2, the absolute amount of NADH and H2O2...

Watch this Scientific Journal Video about Simultaneous Measurement of Superoxide/Hydrogen Peroxide and NADH Production by Flavin-containing Mitochondrial Dehydrogenases at JoVE.com

Simultaneous Measurement of Superoxide/Hydrogen Peroxide and NADH Production by Flavin-containing Mitochondrial Dehydrogenases

Mitochondria contain several flavin-dependent enzymes that can produce reactive oxygen species (ROS). Monitoring ROS release from individual sites in mitochondria is challenging due to unwanted side reactions. We present an easy, inexpensive method for direct assessment of native rates for ROS release using purified flavoenzymes and microplate fluorometry.

It has been reported that mitochondria can contain up to 12 enzymatic sources of reactive oxygen species (ROS). A majority of these sites include ...

simultaneous-measurement-superoxidehydrogen-peroxide-nadh-production

Research

JoVE Journal

Biochemistry

10.0K Views.  Memorial University of Newfoundland. Mitochondria contain several flavin-dependent enzymes that can produce reactive oxygen species (ROS). Monitoring ROS release from individual sites in mitochondria is challenging due to unwanted side reactions. We present an easy, inexpensive method for direct assessment of native rates for ROS release using purified flavoenzymes and microplate fluorometry.

Video: Simultaneous Measurement of Superoxide/Hydrogen Peroxide and NADH Production by Flavin-containing Mitochondrial Dehydrogenases

Monitoring the Reductive and Oxidative Half-Reactions of a Flavin-Dependent Monooxygenase using Stopped-Flow Spectrophotometry

Aspergillus fumigatus siderophore A (SidA) is an FAD-containing monooxygenase that catalyzes the hydroxylation of ornithine in the biosynthesis of hydroxamate siderophores that are essential for virulence (e.g. ferricrocin or N',N",N'''-triacetylfusarinine C)1. The reaction catalyzed by SidA can be divided into reductive and oxidative half-reactions (Scheme 1). In the reductive half-reaction, the oxidized FAD bound to Af SidA, is reduced by NADPH2,3. In the oxidative half-reaction, the reduced cofactor reacts with molecular oxygen to form a C4a-hydroperoxyflavin intermediate, which transfers an oxygen atom to ornithine. Here, we describe a procedure to measure the rates and detect the different spectral forms of SidA using a stopped-flow instrument installed in an anaerobic glove box. In the stopped-flow instrument, small volumes of reactants are rapidly mixed, and after the flow is stopped by the stop syringe (Figure 1), the spectral changes of the solution placed in the observation cell are recorded over time. In the first part of the experiment, we show how we can use the stopped-flow instrument in single mode, where the anaerobic reduction of the flavin in Af SidA by NADPH is directly measured. We then use double mixing settings where Af SidA is first anaerobically reduced by NADPH for a designated period of time in an aging loop, and then reacted with molecular oxygen in the observation cell (Figure 1). In order to perform this experiment, anaerobic buffers are necessary because when only the reductive half-reaction is monitored, any oxygen in the solutions will react with the reduced flavin cofactor and form a C4a-hydroperoxyflavin intermediate that will ultimately decay back into the oxidized flavin. This would not allow the user to accurately measure rates of reduction since there would be complete turnover of the enzyme. When the oxidative half-reaction is being studied the enzyme must be reduced in the absence of oxygen so that just the steps between reduction and oxidation are observed. One of the buffers used in this experiment is oxygen saturated so that we can study the oxidative half-reaction at higher concentrations of oxygen. These are often the procedures carried out when studying either the reductive or oxidative half-reactions with flavin-containing monooxygenases. The time scale of the pre-steady-state experiments performed with the stopped-flow is milliseconds to seconds, which allow the determination of intrinsic rate constants and the detection and identification of intermediates in the reaction4. The procedures described here can be applied to other flavin-dependent monooxygenases.5,6

We describe the use of a stopped-flow instrument to investigate both the reductive and oxidative half-reactions of Aspergillus fumigatus siderophore A (SidA), a flavin-dependent monooxygenase. We then show the spectra corresponding to the species in the reaction of SidA and we calculate the rate constants for their formation.

Aspergillus fumigatus siderophore A (SidA) is an FAD-containing monooxygenase that catalyzes the hydroxylation of ornithine in the biosynthesis of ...

Bioengineering

Detection of Total Reactive Oxygen Species in Adherent Cells by 2',7'-Dichlorodihydrofluorescein Diacetate Staining

Oxidative stress is an important event under both physiological and pathological conditions. In this study, we demonstrate how to quantify oxidative stress by measuring total reactive oxygen species (ROS) using 2&#39;,7&#39;-dichlorodihydrofluorescein diacetate (DCFH-DA) staining in colorectal cancer cell lines as an example. This protocol describes detailed steps including preparation of DCFH-DA solution, incubation of cells with DCFH-DA solution, and measurement of normalized intensity. DCFH-DA staining is a simple and cost-effective way to detect ROS in cells. It can be used to measure ROS generation after chemical treatment or genetic modifications. Therefore, it is useful for determining cellular oxidative stress upon environment stress, providing clues to mechanistic studies.

Here, we present a protocol to detect total cellular reactive oxygen species (ROS) using 2&#39;,7&#39;-dichlorodihydrofluorescein diacetate (DCFH-DA). This method can visualize cellular ROS localization in adherent cells with a fluorescence microscope and quantify ROS intensity with a fluorescence plate reader. This protocol is simple, efficient and cost-effective.

Oxidative stress is an important event under both physiological and pathological conditions. In this study, we demonstrate how to quantify oxidative ...

Stable Isotope Tracing & LC-MS Analysis to Measure DHODH and SDH Activities

Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in mammalian cells. As oxygen (O2) is the most ubiquitous terminal electron acceptor for the mammalian ETC, the O2 consumption rate is frequently used as a proxy for mitochondrial function. However, emerging research demonstrates that this parameter is not always indicative of mitochondrial function, as fumarate can be employed as an alternative electron acceptor to sustain mitochondrial functions in hypoxia. This article compiles a series of protocols that allow researchers to measure mitochondrial function independently of the O2 consumption rate. These assays are particularly useful when studying mitochondrial function in hypoxic environments. Specifically, we describe methods to measure mitochondrial ATP production, de novo pyrimidine biosynthesis, NADH oxidation by complex I, and superoxide production. In combination with classical respirometry experiments, these orthogonal and economical assays will provide researchers with a more comprehensive assessment of mitochondrial function in their system of interest.

Author Spotlight: Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals  

Here, we present a compilation of assays to directly measure mitochondrial function in mammalian cells independently of their ability to consume molecular oxygen.

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in ...

Mitochondrial Functional Assessment in Brush Lizard Liver Mitochondria

Assessment of Mitochondrial Oxygen Consumption Using a Plate Reader-based Fluorescent Assay

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of metabolic pathways. Therefore, experimentally assessing mitochondrial functionality can provide insight into variations among different populations or disease states. Additionally, it is valuable to assess whether isolated mitochondria are healthy enough to proceed with experiments. One characteristic often used to compare mitochondrial function in different samples is the rate of oxygen consumption. Oxygen consumption and subsequent calculation of the respiratory control ratio in either intact cells or mitochondria isolated from tissue can serve all three purposes. Using mitochondria isolated from the livers of brush lizards in conjunction with a phosphorescent probe that is sensitive to the fluctuations in oxygen concentration of a solution, we measured oxygen consumption using a fluorescent plate reader. This method is not only quick and efficient but also can be conducted with a small amount of mitochondria and without the need for specialized equipment. The step-by-step protocol described here increases the accessibility of mitochondrial functional assessment to researchers.

Assessment of oxygen consumption provides integral information about mitochondrial function. Using a phosphorescent probe with a fluorescent plate reader, accurate and reproducible data can be obtained easily without specialized equipment. This assay enables any lab to measure the oxygen consumption of isolated mitochondria and calculate respiratory control ratios.

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of ...

Changing Mitochondrial Function for Therapeutic and Bioinsecticide Applications

Unveiling Xenobiotic Transport and Effects in Isolated Mitochondria: Insights from Respirometric and Enzymatic Assays

Mitochondria are often referred to as the cell's powerhouse due to their crucial role in energy production through the electron transport chain (ETC). However, their significance extends far beyond energy production. Dysregulation of mitochondrial bioenergetics can trigger intracellular cascades that impact health and cellular function. Given their physiological importance, there is growing interest in exploring the pharmacological potential of mitochondrial function for developing new therapies and bioproducts. Modulating mitochondrial bioenergetics could provide innovative approaches to address challenges such as neurodegenerative disorders, metabolic diseases, and cancer, as well as to develop bioproducts for controlling pests and vector-borne diseases.
This work presents a method for assessing the effects of xenobiotics on the electron transport chain (ETC) using various substrates, including glutamate and NADH for complex I, succinate for complex II, and cytochrome c (both oxidized and reduced) for complexes III and IV. This methodology allows the activation or inhibition of electron transport through mitochondrial complexes to be evaluated using a respirometer and spectrophotometer. The mitochondria can be sourced from isolated mitochondria, fragmented cells, or homogenized tissue from various species.
In the laboratory, mitochondrial function has been analyzed in Aedes aegypti and Wistar rats, but this method is also applicable to other species, such as Rhipicephalus microplus and Rhodnius prolixus. This approach provides the basis for theorizing about the existence of uncoupler proteins and species-specific oxidizable substrate preferences influenced by their unique energetic demands. Recent findings offer valuable insights into innovative bioinsecticide design strategies that target mitochondrial function, holding significant potential for effectively controlling vector-borne diseases and pest infestations.

This study introduces a methodology for studying xenobiotic effects on mitochondrial function via respirometric and enzymatic assays. It examines electron transport chain modulation in various species, offering insights for targeted biopesticide development against insect vector-borne diseases.

Mitochondria are often referred to as the cell's powerhouse due to their crucial role in energy production through the electron transport chain (ETC). ...

Multi-parameter Measurement of the Permeability Transition Pore Opening in Isolated Mouse Heart Mitochondria

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with a molecular mass smaller than 1.5 kDa. Although the definitive molecular identity of the pore is still under debate, proteins such as cyclophilin D, VDAC and ANT contribute to mtPTP formation. While the involvement of mtPTP opening in cell death is well established1, accumulating evidence indicates that the mtPTP serves a physiologic role during mitochondrial Ca2+ homeostasis2, bioenergetics and redox signaling 3.
mtPTP opening is triggered by matrix Ca2+ but its activity can be modulated by several other factors such as oxidative stress, adenine nucleotide depletion, high concentrations of Pi, mitochondrial membrane depolarization or uncoupling, and long chain fatty acids4. In vitro, mtPTP opening can be achieved by increasing Ca2+ concentration inside the mitochondrial matrix through exogenous additions of Ca2+ (calcium retention capacity). When Ca2+ levels inside mitochondria reach a certain threshold, the mtPTP opens and facilitates Ca2+ release, dissipation of the proton motive force, membrane potential collapse and an increase in mitochondrial matrix volume (swelling) that ultimately leads to the rupture of the outer mitochondrial membrane and irreversible loss of organelle function.
Here we describe a fluorometric assay that allows for a comprehensive characterization of mtPTP opening in isolated mouse heart mitochondria. The assay involves the simultaneous measurement of 3 mitochondrial parameters that are altered when mtPTP opening occurs: mitochondrial Ca2+ handling (uptake and release, as measured by Ca2+ concentration in the assay medium), mitochondrial membrane potential, and mitochondrial volume. The dyes employed for Ca2+ measurement in the assay medium and mitochondrial membrane potential are Fura FF, a membrane impermeant, ratiometric indicator which undergoes a shift in the excitation wavelength in the presence of Ca2+, and JC-1, a cationic, ratiometric indicator which forms green monomers or red aggregates at low and high membrane potential, respectively. Changes in mitochondrial volume are measured by recording light scattering by the mitochondrial suspension. Since high-quality, functional mitochondria are required for the mtPTP opening assay, we also describe the steps necessary to obtain intact, highly coupled and functional isolated heart mitochondria.

A spectrofluorometric protocol for the measurement of the mitochondrial permeability transition pore opening in isolated mouse heart mitochondria is presented here. The assay involves the simultaneous measurement of mitochondria Ca2+ handling, mitochondrial membrane potential and mitochondrial volume. The procedure for obtaining high-quality and functional heart mitochondria is also described.

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with ...

Biology

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...

Quantification of Reactive Oxygen Species Using 2&#8242;,7&#8242;-Dichlorofluorescein Diacetate Probe and Flow-Cytometry in M&#252;ller Glial Cells

The redox balance has an important role in maintaining cellular homeostasis. The increased generation of reactive oxygen species (ROS) promotes the modification of proteins, lipids, and DNA, which finally may lead to alteration in cellular function and cell death. Therefore, it is beneficial for cells to increase their antioxidant defense in response to detrimental insults, either by activating an antioxidant pathway like Keap1/Nrf2 or by improving redox scavengers (vitamins A, C, and E, &#946;-carotene, and polyphenols, among others). Inflammation and oxidative stress are involved in the pathogenesis and progression of retinopathies, such as diabetic retinopathy (DR) and retinopathy of prematurity (ROP). Since M&#252;ller glial cells (MGCs) play a key role in the homeostasis of neural retinal tissue, they are considered an ideal model to study these cellular protective mechanisms. In this sense, quantifying ROS levels with a reproducible and simple method is essential to assess the contribution of pathways or molecules that participate in the antioxidant cell defense mechanism. In this article, we provide a complete description of the procedures required for the measurement of ROS with DCFH-DA probe and flow cytometry in MGCs. Key steps for flow cytometry data processing with the software are provided here, so the readers will be able to measure ROS levels (geometric means of FITC) and analyze fluorescence histograms. These tools are highly helpful to evaluate not only the increase in ROS after a cellular insult but also to study the antioxidant effect of certain molecules that can provide a protective effect on the cells.

Here, we propose a systematized, accessible, and reproducible protocol to detect cellular reactive oxygen species (ROS) using 2&#8242;,7&#8242;-dichlorofluorescein diacetate probe (DCFH-DA) in M&#252;ller glial cells (MGCs). This method quantifies total cellular ROS levels with a flow cytometer. This protocol is very easy to use, suitable, and reproducible.

The redox balance has an important role in maintaining cellular homeostasis. The increased generation of reactive oxygen species (ROS) promotes the ...

Neuroscience

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

Cellular Redox Profiling Using High-content Microscopy

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, excessive ROS levels induce a state of oxidative stress, which is accompanied by irreversible oxidative damage to DNA, lipids and proteins. Thus, quantification of ROS provides a direct proxy for cellular health condition. Since mitochondria are among the major cellular sources and targets of ROS, joint analysis of mitochondrial function and ROS production in the same cells is crucial for better understanding the interconnection in pathophysiological conditions. Therefore, a high-content microscopy-based strategy was developed for simultaneous quantification of intracellular ROS levels, mitochondrial membrane potential (&#916;&#936;m) and mitochondrial morphology. It is based on automated widefield fluorescence microscopy and image analysis of living adherent cells, grown in multi-well plates, and stained with the cell-permeable fluorescent reporter molecules CM-H2DCFDA (ROS) and TMRM (&#916;&#936;m and mitochondrial morphology). In contrast with fluorimetry or flow-cytometry, this strategy allows quantification of subcellular parameters at the level of the individual cell with high spatiotemporal resolution, both before and after experimental stimulation. Importantly, the image-based nature of the method allows extracting morphological parameters in addition to signal intensities. The combined feature set is used for explorative and statistical multivariate data analysis to detect differences between subpopulations, cell types and/or treatments. Here, a detailed description of the assay is provided, along with an example experiment that proves its potential for unambiguous discrimination between cellular states after chemical perturbation.

This paper presents a high-content microscopy workflow for simultaneous quantification of intracellular ROS levels, as well as mitochondrial membrane potential and morphology &#8211; jointly referred to as mitochondrial morphofunction &#8211; in living adherent cells using the cell-permeant fluorescent reporter molecules 5-(and-6)-chloromethyl-2&#39;,7&#39;-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) and tetramethylrhodamine methylester (TMRM).

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, ...

Simultaneous Measurement of Superoxide/Hydrogen Peroxide and NADH Production by Flavin-containing Mitochondrial Dehydrogenases

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Monitoring the Reductive and Oxidative Half-Reactions of a Flavin-Dependent Monooxygenase using Stopped-Flow Spectrophotometry

Detection of Total Reactive Oxygen Species in Adherent Cells by 2',7'-Dichlorodihydrofluorescein Diacetate Staining

Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals

Assessment of Mitochondrial Oxygen Consumption Using a Plate Reader-based Fluorescent Assay

Unveiling Xenobiotic Transport and Effects in Isolated Mitochondria: Insights from Respirometric and Enzymatic Assays

Multi-parameter Measurement of the Permeability Transition Pore Opening in Isolated Mouse Heart Mitochondria

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Quantification of Reactive Oxygen Species Using 2′,7′-Dichlorofluorescein Diacetate Probe and Flow-Cytometry in Müller Glial Cells

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Cellular Redox Profiling Using High-content Microscopy