Name: measuring trans-plasma membrane electron transport by c2c12 myotubes
Uploaded: 2018-05-04T15:00:00.000+00:00
Duration: 10 min 27 s
Description: Watch this Scientific Journal Video about Measuring Trans-Plasma Membrane Electron Transport by C2C12 Myotubes at JoVE.com

We would like to thank Thomas Bell, Lyn Mattathil, Mark Mannino, and Neej Patel for their technical support. This work was supported by United States Public Health Service award R15DK102122 from the National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK) to Jonathan Fisher. The manuscript content is solely the responsibility of the authors and does not necessarily represent the official views of the NIDDK or the National Institutes of Health.

NOTE: See Figure 1 for a schematic overview of key steps.
1. WST-1 Reduction Assay
<ol>
	<li>Grow and differentiate C2C12 adherent cells using standard cell culture procedures7 in a 96-well plate utilizing rows A-F.
	<ol>
		<li>Use a differentiation medium consisting of Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM) supplemented with 2% horse serum, 100 U/mL penicillin, and 0.1 mg/mL streptomycin. Incubate the cells at 37 &#176;C with 5% CO2.</li>
		<li>When monitoring ascorbate involvement in tPMET, supplement differentiation media with 100 &#181;M ascorbic acid. Allow cells to incubate in differentiation media for ~24-48 h.</li>
	</ol>
	</li>
	<li>Prepare stock WST-1 and PMS solutions.
	<ol>
		<li>To make a 10 mM stock of WST-1, dissolve 0.033 g of WST-1 (Formula Weight (FW): 651.34 g/mol) in 5 mL of phosphate buffered saline (PBS). Store at 4 &#176;C.</li>
		<li>To make a 5 mM stock of PMS, dissolve 0.0023 g of PMS (FW: 306.34 g/mol) in 1.5 mL of deionized H2O (diH2O). Store at -20 &#176;C and protect from light.</li>
	</ol>
	</li>
	<li>Add 0.0108 g of glucose for a final concentration of 5 mM to 11.4 mL of PBS or HEPES buffered saline (HBS; 20 mM HEPES sodium salt, 140 mM NaCl, 5 mM KCl, 2.5 mM MgSO4, 1 mM CaCl2). Add 480 &#181;L of 10 mM WST-1 for a final concentration of 400 &#181;M and 48 &#181;L of 5 mM PMS for a final concentration of 20 &#181;M.</li>
	<li>When monitoring ascorbate involvement in tPMET, divide the solution into two 6 mL aliquots. To one aliquot, add 6 &#181;L of diH2O and to the other aliquot add 6 &#181;L of 2 kU/mL AO for a final concentration of 2 U/mL.</li>
	<li>When monitoring superoxide involvement in tPMET, divide the solution into two 6 mL aliquots. To one aliquot, add 55 &#181;L of 0.1 M KPO4 buffer and to the other aliquot add 55 &#181;L of 6.5 kU/mL SOD for a final concentration of 60 U/mL.</li>
	<li>Wash the cells with PBS. Aspirate the media and add 150 &#181;L PBS. Then aspirate the PBS. 
	NOTE: The assay is done with plated, attached cells. Thus, there is no centrifugation or use of trypsin in the washing process.</li>
	<li>Add 100 &#181;L of WST-1 solution (-) SOD or (-) AO to columns 1-6 and add 100 &#181;L of WST-1 solution (+) SOD or (+) AO to columns 7-12 in the 96-well plate. Use rows G and H as background controls (i.e., to monitor change in absorbance in the reagent alone in wells without cells).</li>
	<li>Measure the absorbance values using the spectrophotometer every 10 min for 1 h at 438 nm.</li>
	<li>After reading, aspirate the media and wash each well with 150 &#181;L of PBS. Then aspirate the PBS.</li>
	<li>Calculate the change in absorbance for each well by subtracting the initial absorbance for a well from the absorbance at any time point for that well. Correct this change for the change in absorbance (if any) observed in the background wells (i.e., wells with assay solution but no cells).</li>
	<li>For analysis, normalize the absorbance data to the 60 min control measurement or wash wells with PBS or HBS and then perform a bicinchoninic acid (BCA) protein assay.</li>
	<li>Add 2 mg/mL bovine serum albumin (BSA) standard (range from 0.5-2 &#181;L for the standard curve, depending on the degree of confluence of the cells) to the empty wells (rows G and H), then add the BCA reagent to all wells.</li>
	<li>Quantify the data via nmol of WST-1 reduction per &#181;g of protein. Use 37 mM-1 cm-1 22 as the extinction coefficient for reduced WST-1 at 438 nm.</li>
</ol>
2. DPIP Reduction Assay
<ol>
	<li>Grow and differentiate C2C12 adherent cells with the same procedure as step 1.1.</li>
	<li>Prepare stock 10 mM DPIP solution as follows. Dissolve 0.029 g of DPIP (FW: 290.08 g/mol) in 10 mL of diH2O. Confirm the concentration of DPIP by measuring absorbance at 600 nm with a spectrophotometer. Use 1 mM-1 cm-1 23 as the extinction coefficient for reduced DPIP at 600 nm. Store at 4 &#176;C.</li>
	<li>Add 0.0108 g of glucose to 11.880 mL of PBS for a final concentration of 5 mM. Add 120 &#181;L of 10 mM DPIP for a final concentration of 100 &#181;M.</li>
	<li>When monitoring ascorbate involvement in tPMET, divide the solution into two 6 mL aliquots. To one aliquot, add 6 &#181;L of diH2O and to the other aliquot add 6 &#181;L of 2 kU/mL AO for a final concentration of 2 U/mL.</li>
	<li>When monitoring superoxide involvement in tPMET, divide the solution into two 6 mL aliquots. To one aliquot, add 55 &#181;L of 0.1 M KPO4 buffer and to the other aliquot add 55 &#181;L of 6.5 kU/mL SOD for a final concentration of 60 U/mL.</li>
	<li>Aspirate the media and wash cells in 150 &#181;L of PBS. Aspirate the PBS and add 100 &#181;L of DPIP solution (-) SOD or (-) AO to columns 1-6 and add 100 &#181;L of DPIP solution (+) SOD or (+) AO to columns 7-12 in the 96-well plate. Use rows G and H will as background controls (i.e., to monitor change in absorbance in the reagent alone in wells without cells).</li>
	<li>Measure the absorbance at 600 nm using spectrophotometer every 10 min for 1 h. Quantify the change in absorbance relative to the control at 60 min similar to steps 1.9-1.13.</li>
</ol>
3. Determination of Whether Reduced Electron Acceptors Are Substrates for AO or SOD
<ol>
	<li>To 5.436 mL of PBS or HBS, add 240 &#181;L of 10 mM WST-1 for a final concentration of 400 &#181;M and 24 &#181;L of 5 mM PMS for a final concentration of 20 &#181;M.
	<ol>
		<li>For DPIP, add 60 &#181;L of 10 mM DPIP, for a final concentration of 100 &#181;M, to 5.940 mL of PBS or HBS.</li>
	</ol>
	</li>
	<li>Add 100 &#181;L of solution to each well in a flat-bottom 96-well plate in the absence of cells and measure the absorbance in a spectrophotometer at 438 nm, for WST-1, or 600 nm, for DPIP.</li>
	<li>Add 1 &#181;L of 10 mM ascorbate to half of the wells for a final concentration of 100 &#956;M. Monitor the absorbance until it stabilizes.</li>
	<li>Upon stabilization, add 1 &#181;L of 200 U/mL AO for a final concentration of 2 U/mL to each well or add 1 &#181;L of 6 kU/mL SOD for a final concentration of 60 U/mL to each well and monitor the absorbance for 1 h.</li>
</ol>

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The authors have nothing to disclose.

We have presented two methods for utilizing extracellular electron acceptors, WST-1 and DPIP, in spectrophotometric assays to monitor tPMET in C2C12 myotubes. With the growth of cell lines in standard culture procedures and a spectrophotometer plate reader, it is possible to monitor tPMET with these electron acceptors in a simple microplate assay. WST-1 reduction is reproducible from well-to-well within an assay, but there is day-to-day variability. The day-to-day coefficient of variation (CV) utilizing PBS as the buffer is 0.18 and the CV utilizing HBS as the buffer is 0.23.
From previous literature utilizing mPMS and WST-1, we have modified this protocol, specifically, for the use of PMS. Thus, troubleshooting included determining the appropriate concentrations of PMS and WST-1 for use with the myotubes described. We also determined that temperature (in a range of 23-37 &#176;C) is not a critical component for the assay, while shaking the plate prior to each absorbance reading is important. However, these issues should be investigated for other cell lines to which the assay is applied. The findings regarding the ability of SOD and AO to directly oxidize reduced WST-1 suggest that screening of new assay components for this property is a key aspect of troubleshooting.
Our data suggests that PMS should be utilized in conjunction with WST-1 in order to elicit optimal WST-1 reduction. We found that in a 1 mL volume, 100 &#181;M ascorbate reduces 322 nmol of WST-1 per minute in the presence of PMS, which is required for direct reduction of WST-1 by ascorbate. In a 1 mL volume, 2 U/mL of AO oxidizes ascorbate at a rate of 4,400 nmol/s, or 800 times faster than the reduction of WST-1 by ascorbate. Clearly, enzymatic activity of AO is much faster than direct reduction of PMS/WST-1 by ascorbate. We expect that the same would follow for superoxide and SOD, though we do not have a system for measuring the absolute rate of WST-1 reduction by superoxide.
To determine whether inclusion of PMS affects the reduction of WST-1 by superoxide, we used a xanthine oxidase (4.5 mU/mL)/hypoxanthine (0.1 mM) superoxide generating system as previously described26,27. Inclusion of PMS approximately doubles the rate of WST-1 reduction by this system. Thus, it appears that PMS mediates transfer of electrons from superoxide to WST-1.
To further evaluate the requirement of PMS for cellular WST-1 reduction, we assayed reduction in the presence or absence of PMS and WST-1, and found that PMS is required for cellular WST-1 reduction. When cells are incubated in assay reagent containing PMS alone (in the absence of WST-1), there is an increase in absorbance at 438 nm. This likely reflects an accumulation of the semiquinone form of PMS (PMS-SQ) that has a peak absorbance at 440 nm28. Accumulation of PMS-SQ suggests that PMS is being reduced faster than it can pass electrons to O2 to produce superoxide. Thus, the accumulation of PMS-SQ is consistent with the reported slow reduction of O2 by PMS-SQ compared to the rate for fully-reduced PMS28.
We have also presented a protocol to determine if the electron acceptors are substrates for the enzymes that may be utilized to monitor tPMET. By monitoring the reduction of the extracellular electron acceptors in a spectrophotometer followed by the addition of the enzyme of interest, it is possible to determine if the electron acceptor is a substrate for these enzymes. Through this method, we have shown that reduced DPIP is a substrate for AO and SOD indicating that it is not a suitable electron acceptor for monitoring ascorbate efflux and superoxide export.
The procedure for testing whether or not assay components such as SOD or AO create artifacts is a significant improvement over existing methods. For example, reduced DPIP is a substrate for SOD, suggesting that published data obtained with use of DPIP and SOD should be interpreted accordingly. Our findings support previous research conducted by Dayan and Dawson in which the absorbance of leuco 2,6-dichloroindophenol was monitored after the addition of AO: DPIP was oxidized indicating that it is a substrate for AO29. However, our findings add context to research conducted by Tan and Berridge24, as well as Bellavite et al.30, that utilized SOD to inhibit DPIP reduction. In a study conducted by Tan and Berridge comparing the electron acceptors of WST-1, DPIP, and FeCN in HeLa cells, they saw that the reduction of some of the electron acceptors was suppressed in the presence of SOD. The reduction of WST-1, in conjunction with mPMS, was inhibited ~80% in the presence of SOD while the reduction of DPIP was inhibited by 40%. The reduction of FeCN, in conjunction with an alternative intermediate electron acceptor, was not inhibited in the presence of SOD24. When evaluating the mode of electron exchange between NADPH oxidases and DPIP or cytochrome c, Bellavite et al. found that DPIP and cytochrome c are reduced by NADPH oxidases and that this reduction is significantly inhibited by SOD30. Berridge and Tan have also demonstrated that SOD can inhibit WST-1 reduction by 80% in mouse cells lines of 32Dcl23, WEHI3B, and J774 as well as human cell lines of Jurkat, MALME3M, and 143B6,31. With human primary neutrophil cells, WST-1 reduction was inhibited 95% in the presence of SOD6,32.
The data here suggest that it is important to verify that enzymes used to assess specific contributions of exported molecules to tPMET cannot re-oxidize the reduced form of the extracellular electron acceptor. We found that (data not shown) ferrous chloride and ferrous sulfate were not soluble in PBS. In HBS and phenol red-free DMEM, they were rapidly oxidized, demonstrating incompatibility with HBS or DMEM as assay media. In contrast, AO and SOD do not use potassium ferrocyanide as a substrate, suggesting that it is a suitable extracellular electron acceptor if its low extinction coefficient is not an issue.
One of the main limitations of this assay is that PMS/WST-1 and DPIP can be reduced by multiple electron donors, such as NAD(P)H, ascorbate, and flavonoids such as quercetin and myricetin19,33,34. However, this can be overcome by the addition of enzymes (e.g., SOD, AO) or inhibitors to determine specific contributors to PMS/WST-1 or DPIP reduction. Another limitation is that DPIP can act as a substrate for AO and SOD. Thus, these enzymes should not be utilized in conjunction with DPIP to monitor tPMET. An additional important limitation of this study is the ability of PMS and other redox cyclers to produce superoxide26,35,36. This is illustrated in Figure 6. On the other hand, PMS itself reportedly has no direct effect on tPMET32. Additionally, Tan and Berridge32 have shown that the NADPH oxidase (NOX) inhibitor, diphenyleneiodinium chloride (DPI), can suppress the majority of WST-1 reduction, suggesting that DPI-sensitive tPMET can be a specific measure of NOX contribution to tPMET. Another limitation is that we see small changes in absorbance when utilizing WST-1. We have found that when WST-1 is made fresh and cells are confluent, the greatest change in absorbance is seen. Previous studies conducted in P388 cell lines have shown that the amount of formazan produced positively correlates with cell number21. Therefore, the more confluent the cells are then the greater the reduction in WST-1. This can be corrected for by normalizing to microgram of protein for each well.
One caveat regarding this assay is that it is primarily designed to assess global tPMET. Although it can be manipulated to monitor multiple forms of tPMET, such as the example of ascorbate efflux, more specific assays might be more appropriate when the goal does not require measurements in the context of tPMET. Fox example, if superoxide is the main interest, there are a number of other avenues to monitor superoxide, such as use of probes including fluorescent impermeable probes, aconitase, and nitroblue tetrazolium37.
Existing probes to monitor tPMET such as DPIP, FeCN and ferricytochrome c, present disadvantages. For example, reduced DPIP is a substrate for AO and SOD, FeCN has a low molar extinction coefficient that limits its utility in real-time assays, and cytochrome c is costly. Additionally, some of these probes can produce high background leading to low sensitivity when measuring a change in absorbance32. WST-1 has been shown to not be a substrate for enzymes such as AO and SOD. Additionally, WST-1 has a high molar extinction coefficient and a low background reaction, creating a more sensitive tPMET assay. Moving forward, this assay can be utilized with a wide range of inhibitors to better understand the process of tPMET, map the path of electron transport in a given cell line, and lead to better understanding of how a cell&#39;s redox environment is maintained.
Critical steps of this protocol include determining that the cells are ready for experimentation (~70% confluent) as well as making the assay reagents, specifically WST-1 and PMS, fresh for each assay. It is also critical to wash the cells before the addition of assay reagents. The DMEM utilized in this protocol does not contain ascorbate, although we supplement differentiation medium with ascorbate. A number of available media do contain ascorbate. However, the cells are washed before the addition of the assay reagents, which are ascorbate-free, to remove any residual ascorbate from the medium. As a portion of tPMET is attributable to molecular efflux (e.g., ascorbate export), it is important to start spectrophotometric readings immediately after the addition of assay reagent so as not to miss the first moments of efflux. Also, it is critical to include background wells for each individual reagent in an experiment. The assay constituents described in this paper caused negligible background change in absorbance. However, some additives (e.g., phloretin) can promote a substantial background reaction. Thus, it is important that the background is subtracted out when the data are analyzed to eliminate any confounding effects that the reagents themselves may produce.
In summary, the protocol presented here provides a means of assessing tPMET and suggests a number of caveats and aspects of the protocol that should be taken into account when applying the assay to different cell lines and/or assessing specific contributors (ascorbate and superoxide used here) to tPMET.

The ability of purified plasma membranes to reduce electron acceptors has led to the view that the plasma membrane has an inherent redox capacity1. Previously seen in fungi, plants, and animals, tPMET is a process common to multiple organisms2,3,4,5. Specifically, this process has been demonstrated in Saccharomyces cerevisiae, carrot cells, erythrocytes, lymphocytes, osteosarcoma, melanoma, macrophages, skeletal muscle, and neutrophils2,3,4,5,6,7. In a process that transports electrons across the plasma membrane to reduce extracellular oxidants, tPMET is involved in many cellular functions including: cell growth5,8, cell viability9, iron metabolism10, cell signaling11,12,13, and protection from reactive oxygen species12,14,15. Due to tPMET&#39;s involvement in many cellular functions, an imbalance of tPMET has been hypothesized to contribute to the development of some serious health conditions, including cancer16, cardiovascular disease17, and metabolic syndrome18.
There are multiple ways to monitor the transfer of electrons across the plasma membrane, but the most widely used technique is to assess the reduction of extracellular electron acceptors through colorimetric assays. Commonly used extracellular electron acceptors are tetrazolium salts, DPIP, FeCN, and ferricytochrome c19,20. The most commonly used tetrazolium salt is a second-generation salt known as WST-119. This compound is easier to utilize in colorimetric assays compared to first generation tetrazolium salts due to two sulfonate groups, which increase its water solubility21. WST-1, in conjunction with the intermediate electron acceptor 1-methoxy-phenazine methosulfate (mPMS), is reduced in two single-electron transfer events. This reduction changes the weakly-colored oxidized form of WST-1 to a more intense, yellow formazan20,22. WST-1 has a high molar extinction coefficient of 37 x 103 M-1cm-1, leading to a high assay sensitivity21,22. DPIP is also utilized as an extracellular electron acceptor to monitor tPMET. It has been shown that DPIP can be reduced extracellularly by tPMET without the aid of intermediate electron acceptors23,24. Due to the lack of intermediate electron acceptors, DPIP can directly pick-up electrons from the plasma membrane, unlike WST-124. Similar to DPIP, FeCN has been shown to be reduced extracellularly to ferrocyanide by tPMET without the aid of intermediate electron acceptors19,24. Unlike WST-1 and DPIP, FeCN has a low molar extinction coefficient leading to a lower assay sensitivity9. Another commonly used extracellular electron acceptor to monitor tPMET is ferricytochrome c. Similar to WST-1, ferricytochrome c reduction increases with the use of intermediate electron acceptor, mPMS22. Unlike WST-1 though, the ferricytochrome c method is less sensitive due to a high background and a low molar extinction coefficient22.
Here we present a method for real-time analysis of tPMET through spectrophotometric assays. The method utilized the extracellular electron acceptors WST-1 and DPIP, as they both have a high molar extinction coefficient while being less expensive compared to the other commonly used extracellular electron acceptors such as ferricytochrome c. We utilized phenazine methosulfate (PMS) instead of mPMS they have a similar chemical makeup and PMS is far less expensive. mPMS is photochemically stable which is an important characteristic for a commercial kit that needs a long shelf life. However, we make PMS fresh for each assay, so stability should not be an issue. We also present a method to evaluate possible enzymatic interactions (see Figure 1) between the extracellular electron acceptor and enzymes that may be utilized to further characterize the process of tPMET. Specifically, the enzymes AO and SOD can be used determine which portion of tPMET is due to ascorbate transport or extracellular superoxide release, two common methods for electrons to be transported across the plasma membrane.

<table><tbody><tr><td>C2C12 myoblasts</td><td>American Type Culture Collection&amp;nbsp;</td><td>CRL-1772</td><td></td></tr><tr><td>Dulbecco&#039;s modified eagle&#039;s medium - low glucose</td><td>Sigma</td><td>D6046</td><td></td></tr><tr><td>Fetal Plex animal serum complex</td><td>Gemini Bio-Products&amp;nbsp;</td><td>100-602</td><td></td></tr><tr><td>penicillin-streptomycin</td><td>Sigma</td><td>516106</td><td></td></tr><tr><td>horse serum</td><td>Gibco Technologies</td><td>16050-130</td><td></td></tr><tr><td>Dulbecco&#039;s phosphate buffered saline</td><td>Sigma</td><td>D8537</td><td></td></tr><tr><td>trypsin-EDTA</td><td>Sigma</td><td>T4049</td><td></td></tr><tr><td>15 cm culture dishes</td><td>TPP</td><td>93150</td><td></td></tr><tr><td>96 well culture plates</td><td>TPP</td><td>92096</td><td></td></tr><tr><td>2-(4-Iodophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium Sodium Salt (WST-1)</td><td>Accela ChemBio&amp;nbsp; Inc</td><td>SY016315</td><td></td></tr><tr><td>phenazine methosulfate&amp;nbsp;</td><td>Sigma</td><td>P9625</td><td></td></tr><tr><td>L-ascorbic acid</td><td>Sigma</td><td>A5960</td><td></td></tr><tr><td>ascorbate oxidase&amp;nbsp;</td><td>Sigma</td><td>A0157</td><td></td></tr><tr><td>superoxide dismutase&amp;nbsp;</td><td>Sigma</td><td>S5395</td><td></td></tr><tr><td>2,6-dichloroindophenol sodium salt&amp;nbsp;</td><td>ICN Biomedicals</td><td>215011825</td><td></td></tr><tr><td>D-(+)-glucose</td><td>Sigma</td><td>G7528</td><td></td></tr><tr><td>HEPES sodium salt</td><td>Sigma</td><td>H3784</td><td></td></tr><tr><td>sodium chloride</td><td>Sigma</td><td>S7653</td><td></td></tr><tr><td>potassium chloride</td><td>Fisher Scientific&amp;nbsp;</td><td>BP366</td><td></td></tr><tr><td>magnesium sulfate heptahydrate</td><td>Sigma</td><td>M5921</td><td></td></tr><tr><td>calcium chloride dihydrate</td><td>Sigma</td><td>C7902</td><td></td></tr><tr><td>potassium phosphate</td><td>Fisher Scientific&amp;nbsp;</td><td>BP363</td><td></td></tr><tr><td>Pierce BCA Protein Assay Kit</td><td>Thermo Scientific</td><td>23225</td><td></td></tr><tr><td>Powerwave X-I spectrophotometer</td><td>Biotek Insturments</td><td>discontinued&amp;nbsp;</td><td></td></tr><tr><td>Spectronic Genesys 5 Spectrophotometer</td><td>Thermo Scientific</td><td>336001</td><td></td></tr><tr><td>PureGrade 96-well microplate, F-bottom, clear, untreated, non-sterile</td><td>MidSci</td><td>781602</td><td></td></tr><tr><td>Iron (II) chloride tetrahydrate</td><td>Sigma</td><td>220299</td><td></td></tr><tr><td>Iron (II) sulfate heptahydrate</td><td>Sigma</td><td>215422</td><td></td></tr><tr><td>hypoxanthine</td><td>Sigma</td><td>H9636</td><td></td></tr><tr><td>xanthine oxidase</td><td>Sigma</td><td>X4500</td><td></td></tr><tr><td>Excel</td><td>Microsoft</td><td></td><td></td></tr><tr><td>R Studio</td><td>Rstudio</td><td></td><td>https://www.rstudio.com/products/rstudio/</td></tr><tr><td>KC4</td><td>Biotek Insturments</td><td>discontinued&amp;nbsp;</td><td></td></tr></tbody></table>

measuring trans-plasma membrane electron transport by c2c12 myotubes

Trans-plasma membrane electron transport (tPMET) plays a role in protection of cells from intracellular reductive stress as well as protection from damage by extracellular oxidants. This process of transporting electrons from intracellular reductants to extracellular oxidants is not well defined. Here we present spectrophotometric assays by C2C12 myotubes to monitor tPMET utilizing the extracellular electron acceptors: water-soluble tetrazolium salt-1 (WST-1) and 2,6-dichlorophenolindophenol (DPIP or DCIP). Through reduction of these electron acceptors, we are able to monitor this process in a real-time analysis. With the addition of enzymes such as ascorbate oxidase (AO) and superoxide dismutase (SOD) to the assays, we can determine which portion of tPMET is due to ascorbate export or superoxide production, respectively. While WST-1 was shown to produce stable results with low background, DPIP was able to be re-oxidized after the addition of AO and SOD, which was demonstrated with spectrophotometric analysis. This method demonstrates a real-time, multi-well, quick spectrophotometric assay with advantages over other methods used to monitor tPMET, such as ferricyanide (FeCN) and ferricytochrome c reduction.

Statistics were performed with ANOVA with repeated measures using RStudio statistical software25. Sample sizes are indicated in the figure legends.
To monitor tPMET, C2C12 myotubes were utilized along with extracellular electron acceptors, WST-1 and DPIP. AO was used to determine which portion of WST-1 and DPIP reduction was due to ascorbate efflux and SOD was used to determine which portion of WST-1 reduction was due to extracellular superoxide release. As shown in Figure 2, C2C12 myotubes are capable of tPMET as seen by the reduction of WST-1. With the addition of AO, WST-1 reduction was suppressed by ~30% indicating that ~30% of tPMET was due to the export of ascorbate (Figure 2A). With the addition of SOD, WST-1 reduction was suppressed by ~70%, indicating that ~70% of tPMET was due extracellular superoxide release (Figure 2B). When DPIP was utilized as an extracellular electron acceptor with the addition of AO, DPIP reduction was suppressed over 100% (Figure 3). This raised questions regarding the validity of using DPIP with AO to assess contribution of ascorbate.
To look into whether reduced WST-1 is a substrate for AO or SOD, WST-1 was reduced with ascorbic acid. AO or SOD was added, and the absorbance was monitored. After 1 h, there was no change in absorbance between the reduced form of WST-1 without AO or SOD and the reduced form of WST-1 with AO or SOD (Figure 4A, B). Therefore, the reduced WST-1 is not a substrate for these enzymes, and the use of AO or SOD in conjunction with WST-1 is appropriate for assessment of roles of ascorbate or superoxide, respectively.
To determine if reduced DPIP is a substrate for AO or SOD, DPIP was reduced with ascorbic acid. After 20 min, AO was added, and the absorbance was monitored. Reduced DPIP was shown to be a substrate for AO, as seen in Figure 5A, where DPIP returns to its oxidized form within 40 min. When SOD was added, DPIP was also re-oxidized (Figure 5B). Therefore, reduced DPIP is a substrate for these enzymes and not an appropriate extracellular electron acceptor to be utilized with these enzymes.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57565/57565fig1.jpg" /> 
Figure 1: A schematic representation of the key steps of the electron transport assays and determination of enzymatic interactions with the extracellular electron acceptors. <a href="/files/ftp_upload/57565/57565fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57565/57565fig2.jpg" /> 
Figure 2: The addition of AO and SOD suppresses WST-1 reduction by C2C12 myotubes. (A) tPMET was analyzed for 60 min using 400 &#181;M WST-1 and 20 &#181;M PMS with the addition of 2 U/mL AO or diH2O (N = 18/group). (B) tPMET was analyzed for 60 min with the addition of 60 U/mL SOD or 0.1 M KPO4 buffer (N = 36/group). *p&#8805; 0.05 at the indicated time point. Error bars represent standard error of the mean and may be too small to visualize. <a href="/files/ftp_upload/57565/57565fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57565/57565fig3.jpg" /> 
Figure 3: The addition of AO suppresses DPIP reduction by C2C12 myotubes. tPMET was analyzed for 60 min using 100 &#181;M DPIP with or without the addition of 2 U/mL AO (N = 36/group). *p&#8805; 0.05 at the indicated time point. Error bars represent standard error of the mean and may be too small to visualize. <a href="/files/ftp_upload/57565/57565fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57565/57565fig4.jpg" /> 
Figure 4: Reduced WST-1 is not a substrate for AO or SOD. (A) WST-1 was reduced with ascorbate. After 45 min, AO was added, and the absorbance was monitored for 1 h. No change in absorbance was observed. (B) The procedure is the same as above, except SOD was added after 45 min. No change in absorbance was observed (N = 24/group). Error bars represent standard error of the mean and may be too small to visualize. <a href="/files/ftp_upload/57565/57565fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57565/57565fig5.jpg" /> 
Figure 5: Reduced DPIP is a substrate for AO and SOD. (A) DPIP was reduced with ascorbate. After 20 min, AO was added, and the absorbance was monitored. Within 40 min, the samples returned to their original absorbance, indicating that reduced DPIP is a substrate for AO. (B) DPIP was reduced with ascorbate. After 20 min, SOD was added, and the absorbance was monitored. Over time, the reduced DPIP was re-oxidized, indicating that reduced DPIP is a substrate for SOD (N = 24/group). Error bars represent standard error of the mean and may be too small to visualize. <a href="/files/ftp_upload/57565/57565fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/57565/57565fig6.jpg" /> 
Figure 6: Schematic depicting tPMET. In this model of tPMET, ascorbate (or another electron carrier) exported by the cell can reduce extracellular PMS. In addition, NADPH oxidases generate extracellular superoxide, which can reduce WST-1 directly or reduce it indirectly via reduction of PMS. Electrons from other plasma membrane donors can also reduce PMS, which can donate electrons to either WST-1 or O2 with subsequent WST-1 reduction. <a href="/files/ftp_upload/57565/57565fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Measuring Trans-Plasma Membrane Electron Transport by C2C12 Myotubes at JoVE.com

Measuring Trans-Plasma Membrane Electron Transport by C2C12 Myotubes

The goal of this protocol is to spectrophotometrically monitor trans-plasma membrane electron transport utilizing extracellular electron acceptors and to analyze enzymatic interactions that may occur with these extracellular electron acceptors.

Trans-plasma membrane electron transport (tPMET) plays a role in protection of cells from intracellular reductive stress as well as protection from damage ...

measuring-trans-plasma-membrane-electron-transport-by-c2c12-myotubes

Nanyang Technological University

Research

JoVE Journal

Biochemistry

6.9K Views.  Saint Louis University. Het doel van dit protocol is spectrophotometrically trans-plasma membraan elektronentransport met behulp van extracellulaire elektronen acceptoren controleren en analyseren van enzymatische interacties die bij deze extracellulaire elektronen acceptoren optreden kunnen.

Video: Measuring Trans-Plasma Membrane Electron Transport by C2C12 Myotubes

Using Isolated Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High throughput Microplate Respiratory Measurements

Skeletal muscle mitochondria play a specific role in many disease pathologies. As such, the measurement of oxygen consumption as an indicator of mitochondrial function in this tissue has become more prevalent. Although many technologies and assays exist that measure mitochondrial respiratory pathways in a variety of cells, tissue and species, there is currently a void in the literature in regards to the compilation of these assays using isolated mitochondria from mouse skeletal muscle for use in microplate based technologies. Importantly, the use of microplate based respirometric assays is growing among mitochondrial biologists as it allows for high throughput measurements using minimal quantities of isolated mitochondria. Therefore, a collection of microplate based respirometric assays were developed that are able to assess mechanistic changes/adaptations in oxygen consumption in a commonly used animal model. The methods presented herein provide step-by-step instructions to perform these assays with an optimal amount of mitochondrial protein and reagents, and high precision as evidenced by the minimal variance across the dynamic range of each assay.

The methods presented provide step-by-step instructions for the performance of a collection of microplate based respirometric assays using isolated mitochondria from minimal quantities of mouse skeletal muscle. These assays are able to measure mechanistic changes/adaptations in mitochondrial oxygen consumption in a commonly used animal model.

Skeletal muscle mitochondria play a specific role in many disease pathologies. As such, the measurement of oxygen consumption as an indicator of ...

Developmental Biology

Molecular Imaging to Target Transplanted Muscle Progenitor Cells

Duchenne muscular dystrophy (DMD) is a severe genetic neuromuscular disorder that affects 1 in 3,500 boys, and is characterized by progressive muscle degeneration1, 2. In patients, the ability of resident muscle satellite cells (SCs) to regenerate damaged myofibers becomes increasingly inefficient4. Therefore, transplantation of muscle progenitor cells (MPCs)/myoblasts from healthy subjects is a promising therapeutic approach to DMD. A major limitation to the use of stem cell therapy, however, is a lack of reliable imaging technologies for long-term monitoring of implanted cells, and for evaluating its effectiveness. Here, we describe a non-invasive, real-time approach to evaluate the success of myoblast transplantation. This method takes advantage of a unified fusion reporter gene composed of genes (firefly luciferase [fluc], monomeric red fluorescent protein [mrfp] and sr39 thymidine kinase [sr39tk]) whose expression can be imaged with different imaging modalities9, 10. A variety of imaging modalities, including positron emission tomography (PET), single-photon emission computed tomography (SPECT), magnetic resonance imaging (MRI), optical imaging, and high frequency 3D-ultrasound are now available, each with unique advantages and limitations11. Bioluminescence imaging (BLI) studies, for example, have the advantage of being relatively low cost and high-throughput. It is for this reason that, in this study, we make use of the firefly luciferase (fluc) reporter gene sequence contained within the fusion gene and bioluminescence imaging (BLI) for the short-term localization of viable C2C12 myoblasts following implantation into a mouse model of DMD (muscular dystrophy on the X chromosome [mdx] mouse)12-14. Importantly, BLI provides us with a means to examine the kinetics of labeled MPCs post-implantation, and will be useful to track cells repeatedly over time and following migration. Our reporter gene approach further allows us to merge multiple imaging modalities in a single living subject; given the tomographic nature, fine spatial resolution and ability to scale up to larger animals and humans10,11, PET will form the basis of future work that we suggest may facilitate rapid translation of methods developed in cells to preclinical models and to clinical applications.

A non-invasive means to evaluate the success of myoblast transplantation is described. The method takes advantage of a unified fusion reporter gene composed of genes whose expression can be imaged with different imaging modalities. Here, we make use of a fluc reporter gene sequence to target cells via bioluminescence imaging.

Duchenne muscular dystrophy (DMD) is a severe genetic neuromuscular disorder that affects 1 in 3,500 boys, and is characterized by progressive muscle ...

Medicine

Measuring Mitochondrial Electron Transfer Complexes in Previously Frozen Cardiac Tissue from the Offspring of Sow: A Model to Assess Exercise-Induced Mitochondrial Bioenergetics Changes

The mitochondrial electron transfer complex (ETC) profile is modified in the heart tissue of the offspring born to an exercised sow. The hypothesis proposed and tested was that a regular maternal exercise of a sow during pregnancy would increase the mitochondrial efficiency of offspring heart bioenergetics. This hypothesis was tested by isolating mitochondria using a mild-isolation procedure to assess mitochondrial ETC and supercomplex profiles. The procedure described here allowed for the processing of previously frozen archived heart tissues and eliminated the necessity of fresh mitochondria preparation for the assessment of mitochondrial ETC complexes, supercomplexes, and ETC complex activity profiles. This protocol describes the optimal ETC protein complex measurement in multiplexed antibody-based immunoblotting and super complex assessment using blue-native gel electrophoresis.

Preparation of mitochondria-enriched samples from previously frozen archived solid tissues allowed the investigators to perform both functional and analytical assessments of mitochondria in various experimental modalities. This study demonstrates how to prepare mitochondria-enriched preparations from frozen heart tissue and perform analytical assessments of mitochondria.

The mitochondrial electron transfer complex (ETC) profile is modified in the heart tissue of the offspring born to an exercised sow. The hypothesis ...

Inner Mitochondrial Membrane Sensitivity to Na+ Reveals Partially Segmented Functional CoQ Pools

Ubiquinone (CoQ) pools in the inner mitochondrial membrane (IMM) are partially segmented to either complex I or FAD-dependent enzymes. Such subdivision can be easily assessed by a comparative assay using NADH or succinate as electron donors in frozen-thawed mitochondria, in which cytochrome c (cyt c) reduction is measured. The assay relies on the effect of Na+ on the IMM, decreasing its fluidity. Here, we present a protocol to measure NADH-cyt c oxidoreductase activity and succinate-cyt c oxidoreductase activities in the presence of NaCl or KCl. The reactions, which rely on the mixture of reagents in a cuvette in a stepwise manner, are measured spectrophotometrically during 4 min in the presence of Na+ or K+. The same mixture is performed in parallel in the presence of the specific enzyme inhibitors in order to subtract the unspecific change in absorbance. NADH-cyt c oxidoreductase activity does not decrease in the presence of any of these cations. However, succinate-cyt c oxidoreductase activity decreases in the presence of NaCl. This simple experiment highlights: 1) the effect of Na+ in decreasing IMM fluidity and CoQ transfer; 2) that supercomplex I+III2 protects ubiquinone (CoQ) transfer from being affected by lowering IMM fluidity; 3) that CoQ transfer between CI and CIII is functionally different from CoQ transfer between CII and CIII. These facts support the existence of functionally differentiated CoQ pools in the IMM and show that they can be regulated by the changing Na+ environment of mitochondria.

Inner Mitochondrial Membrane Sensitivity to Na+ Reveals Partially Segmented Functional CoQ Pools

This protocol describes a comparative assay, using mitochondrial complex activities CI+CIII and CII+CIII in the presence or absence of Na+, to study the existence of partially segmented functional CoQ pools.

Ubiquinone (CoQ) pools in the inner mitochondrial membrane (IMM) are partially segmented to either complex I or FAD-dependent enzymes. Such subdivision ...

Preparation and Respirometric Assessment of Mitochondria Isolated from Skeletal Muscle Tissue Obtained by Percutaneous Needle Biopsy

Respirometric profiling of isolated mitochondria is commonly used to investigate electron transport chain function. We describe a method for obtaining samples of human Vastus lateralis, isolating mitochondria from minimal amounts of skeletal muscle tissue, and plate based respirometric profiling using an extracellular flux (XF) analyzer. Comparison of respirometric profiles obtained using 1.0, 2.5 and 5.0 &#956;g of mitochondria indicate that 1.0 &#956;g is sufficient to measure respiration and that 5.0 &#956;g provides most consistent results based on comparison of standard errors. Western blot analysis of isolated mitochondria for mitochondrial marker COX IV and non-mitochondrial tissue marker GAPDH indicate that there is limited non-mitochondrial contamination using this protocol. The ability to study mitochondrial respirometry in as little as 20 mg of muscle tissue allows users to utilize individual biopsies for multiple study endpoints in clinical research projects.

Methods for biopsy of Vastus lateralis, preparation of purified mitochondria, and respirometric profiling are described. The use of small muscle volume makes this technique suitable for clinical research applications.

Respirometric profiling of isolated mitochondria is commonly used to investigate electron transport chain function. We describe a method for obtaining ...

Bioenergetic Profile Experiment using C2C12 Myoblast Cells

The ability to measure cellular metabolism and understand mitochondrial dysfunction, has enabled scientists worldwide to advance their research in understanding the role of mitochondrial function in obesity, diabetes, aging, cancer, cardiovascular function and safety toxicity.
Cellular metabolism is the process of substrate uptake, such as oxygen, glucose, fatty acids, and glutamine, and subsequent energy conversion through a series of enzymatically controlled oxidation and reduction reactions. These intracellular biochemical reactions result in the production of ATP, the release of heat and chemical byproducts, such as lactate and CO2 into the extracellular environment.
Valuable insight into the physiological state of cells, and the alteration of the state of those cells, can be gained through measuring the rate of oxygen consumed by the cells, an indicator of mitochondrial respiration - the Oxygen Consumption Rate - or OCR. Cells also generate ATP through glycolysis, i.e.: the conversion of glucose to lactate, independent of oxygen. In cultured wells, lactate is the primary source of protons. Measuring the lactic acid produced indirectly via protons released into the extracellular medium surrounding the cells, which causes acidification of the medium provides the Extra-Cellular Acidification Rate - or ECAR.
In this experiment, C2C12 myoblast cells are seeded at a given density in Seahorse cell culture plates. The basal oxygen consumption (OCR) and extracellular acidification (ECAR) rates are measured to establish baseline rates. The cells are then metabolically perturbed by three additions of different compounds (in succession) that shift the bioenergetic profile of the cell.
This assay is derived from a classic experiment to assess mitochondria and serves as a framework with which to build more complex experiments aimed at understanding both physiologic and pathophysiologic function of mitochondria and to predict the ability of cells to respond to stress and/or insults.

A description of a method for profiling mitochondrial function in cells is provided. The mitochondrial profile generated provides four parameters of mitochondrial function that can be measured in one experiment: basal respiration rate, ATP-linked respiration, proton leak, and reserve capacity.

The ability to measure cellular metabolism and understand mitochondrial dysfunction, has enabled scientists worldwide to advance their research in ...

Biology

Modeling Myotonic Dystrophy 1 in C2C12 Myoblast Cells

Myotonic dystrophy 1 (DM1) is a common form of muscular dystrophy. Although several animal models have been established for DM1, myoblast cell models are still important because they offer an efficient cellular alternative for studying cellular and molecular events. Though C2C12 myoblast cells have been widely used to study myogenesis, resistance to gene transfection, or viral transduction, hinders research in C2C12 cells. Here, we describe an optimized protocol that includes daily maintenance, transfection and transduction procedures to introduce genes into C2C12 myoblasts and the induction of myocyte differentiation. Collectively, these procedures enable best transfection/transduction efficiencies, as well as consistent differentiation outcomes. The protocol described in establishing DM1 myoblast cell models would benefit the study of myotonic dystrophy, as well as other muscular diseases.

In this protocol, we present the procedures in establishing myotonic dystrophy 1 myoblast models, including optimized C2C12 cell maintenance, gene transfection/transduction, and myocyte differentiation.

Myotonic dystrophy 1 (DM1) is a common form of muscular dystrophy. Although several animal models have been established for DM1, myoblast cell models are ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

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 ...

Measurement and Analysis of Extracellular Acid Production to Determine Glycolytic Rate

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both mitochondrial (respiration) and non-mitochondrial (other redox) reactions consume oxygen, but these reactions can be easily distinguished by chemical inhibition of mitochondrial respiration. However, while mitochondrial oxygen consumption is an unambiguous and direct measurement of respiration rate2, the same is not true for extracellular acid production and its relationship to glycolytic rate 3-6. Extracellular acid produced by cells is derived from both lactate, produced by anaerobic glycolysis, and CO2, produced in the citric acid cycle during respiration. For glycolysis, the conversion of glucose to lactate- + H+ and the export of products into the assay medium is the source of glycolytic acidification. For respiration, the export of CO2, hydration to H2CO3 and dissociation to HCO3- + H+ is the source of respiratory acidification. The proportions of glycolytic and respiratory acidification depend on the experimental conditions, including cell type and substrate(s) provided, and can range from nearly 100% glycolytic acidification to nearly 100% respiratory acidification 6. Here, we demonstrate the data collection and calculation methods needed to determine respiratory and glycolytic contributions to total extracellular acidification by whole cells in culture using C2C12 myoblast cells as a model.

Glycolysis is a defining metabolic marker in multiple biological systems. Monitoring glycolysis by measuring the extracellular flux of H+ is common, but requires correction to be quantitative and unambiguous. Here, we demonstrate how to gather and correct extracellular flux data to distinguish between respiratory and glycolytic sources of extracellular acidification.

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both ...

Measuring Trans-Plasma Membrane Electron Transport by C2C12 Myotubes

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