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&#160;</td>
			<td>CRL-1772</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s modified eagle&#39;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&#160;</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&#39;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&#160; Inc</td>
			<td>SY016315</td>
			<td></td>
		</tr>
		<tr>
			<td>phenazine methosulfate&#160;</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&#160;</td>
			<td>Sigma</td>
			<td>A0157</td>
			<td></td>
		</tr>
		<tr>
			<td>superoxide dismutase&#160;</td>
			<td>Sigma</td>
			<td>S5395</td>
			<td></td>
		</tr>
		<tr>
			<td>2,6-dichloroindophenol sodium salt&#160;</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&#160;</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&#160;</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&#160;</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&#160;</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

6.8K Views.  Saint Louis University. 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.

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

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Measuring Protein Binding to F-actin by Co-sedimentation

Filamentous actin (F-actin) organization within cells is regulated by a large number of actin-binding proteins that control actin nucleation, growth, cross-linking and/or disassembly. This protocol describes a technique &#8211; the actin co-sedimentation, or pelleting, assay &#8211; to determine whether a protein or protein domain binds F-actin and to measure the affinity of the interaction (i.e., the dissociation equilibrium constant). In this technique, a protein of interest is first incubated with F-actin in solution. Then, differential centrifugation is used to sediment the actin filaments, and the pelleted material is analyzed by SDS-PAGE. If the protein of interest binds F-actin, it will co-sediment with the actin filaments. The products of the binding reaction (i.e., F-actin and the protein of interest) can be quantified to determine the affinity of the interaction. The actin pelleting assay is a straightforward technique for determining if a protein of interest binds F-actin and for assessing how changes to that protein, such as ligand binding, affect its interaction with F-actin.

This protocol describes a method to test the ability of a protein to co-sediment with filamentous actin (F-actin) and, if binding is observed, to measure the affinity of the interaction.

Filamentous actin (F-actin) organization within cells is regulated by a large number of actin-binding proteins that control actin nucleation, growth, ...

Analyzing Supercomplexes of the Mitochondrial Electron Transport Chain with Native Electrophoresis, In-gel Assays, and Electroelution

The mitochondrial electron transport chain (ETC) transduces the energy derived from the breakdown of various fuels into the bioenergetic currency of the cell, ATP. The ETC is composed of 5 massive protein complexes, which also assemble into supercomplexes called respirasomes (C-I, C-III, and C-IV) and synthasomes (C-V) that increase the efficiency of electron transport and ATP production. Various methods have been used for over 50 years to measure ETC function, but these protocols do not provide information on the assembly of individual complexes and supercomplexes. This protocol describes the technique of native gel polyacrylamide gel electrophoresis (PAGE), a method that was modified more than 20 years ago to study ETC complex structure. Native electrophoresis permits the separation of ETC complexes into their active forms, and these complexes can then be studied using immunoblotting, in-gel assays (IGA), and purification by electroelution. By combining the results of native gel PAGE with those of other mitochondrial assays, it is possible to obtain a completer picture of ETC activity, its dynamic assembly and disassembly, and how this regulates mitochondrial structure and function. This work will also discuss limitations of these techniques. In summary, the technique of native PAGE, followed by immunoblotting, IGA, and electroelution, presented below, is a powerful way to investigate the functionality and composition of mitochondrial ETC supercomplexes.

This protocol describes the separation of functional mitochondrial electron transport chain complexes (Cx) I-V and supercomplexes thereof using native electrophoresis to reveal information about their assembly and structure. The native gel can be subjected to immunoblotting, in-gel assays, and purification by electroelution to further characterize individual complexes.

The mitochondrial electron transport chain (ETC) transduces the energy derived from the breakdown of various fuels into the bioenergetic currency of the ...

Semi-automated Biopanning of Bacterial Display Libraries for Peptide Affinity Reagent Discovery and Analysis of Resulting Isolates

Biopanning bacterial display libraries is a proven technique for peptide affinity reagent discovery for recognition of both biotic and abiotic targets. Peptide affinity reagents can be used for similar applications to antibodies, including sensing and therapeutics, but are more robust and able to perform in more extreme environments. Specific enrichment of peptide capture agents to a protein target of interest is enhanced using semi-automated sorting methods which improve binding and wash steps and therefore decrease the occurrence of false positive binders. A semi-automated sorting method is described herein for use with a commercial automated magnetic-activated cell sorting device with an unconstrained bacterial display sorting library expressing random 15-mer peptides. With slight modifications, these methods are extendable to other automated devices, other sorting libraries, and other organisms. A primary goal of this work is to provide a comprehensive methodology and expound the thought process applied in analyzing and minimizing the resulting pool of candidates. These techniques include analysis of on-cell binding using fluorescence-activated cell sorting (FACS), to assess affinity and specificity during sorting and in comparing individual candidates, and the analysis of peptide sequences to identify trends and consensus sequences for understanding and potentially improving the affinity to and specificity for the target of interest.

Biopanning bacterial display libraries is a proven technique for discovery of peptide affinity reagents, a robust alternative to antibodies. The semi-automated sorting method herein has streamlined biopanning to decrease the occurrence of false positives. Here we illustrate the thought process and techniques applied in evaluating candidates and minimizing downstream analysis.

Biopanning bacterial display libraries is a proven technique for peptide affinity reagent discovery for recognition of both biotic and abiotic targets. ...

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

Direct electrochemical detection of c-type cytochrome complexes embedded in the bacterial outer membrane (outer membrane c-type cytochrome complexes; OM c-Cyts) has recently emerged as a novel whole-cell analytical method to characterize the bacterial electron transport from the respiratory chain to the cell exterior, referred to as the extracellular electron transport (EET). While the pathway and kinetics of the electron flow during the EET reaction have been investigated, a whole-cell electrochemical method to examine the impact of cation transport associated with EET has not yet been established. In the present study, an example of a biochemical technique to examine the deuterium kinetic isotope effect (KIE) on EET through OM c-Cyts using a model microbe, Shewanella oneidensis MR-1, is described. The KIE on the EET process can be obtained if the EET through OM c-Cyts acts as the rate-limiting step in the microbial current production. To that end, before the addition of D2O, the supernatant solution was replaced with fresh media containing a sufficient amount of the electron donor to support the rate of upstream metabolic reactions, and to remove the planktonic cells from a uniform monolayer biofilm on the working electrode. Alternative methods to confirm the rate-limiting step in microbial current production as EET through OM c-Cyts are also described. Our technique of a whole-cell electrochemical assay for investigating proton transport kinetics can be applied to other electroactive microbial strains.

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

Here we present a protocol of whole-cell electrochemical experiments to study the contribution of proton transport to the rate of extracellular electron transport via the outer-membrane cytochromes complex in Shewanella oneidensis MR-1.

Direct electrochemical detection of c-type cytochrome complexes embedded in the bacterial outer membrane (outer membrane c-type cytochrome complexes; OM ...

Analysis of &#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and structure by spectrometry and scanning electron microscopy (SEM). A&#946;, which forms neurotoxic amyloid aggregates in Alzheimer&#39;s disease (AD), has been shown to interact with fibrinogen. This A&#946;-fibrinogen interaction makes the fibrin clot structurally abnormal and resistant to fibrinolysis. A&#946;-induced abnormalities in fibrin clotting may also contribute to cerebrovascular aspects of the AD pathology such as microinfarcts, inflammation, as well as, cerebral amyloid angiopathy (CAA). Given the potentially critical role of neurovascular deficits in AD pathology, developing compounds which can inhibit or lessen the A&#946;-fibrinogen interaction has promising therapeutic value. In vitro methods by which fibrin clot formation can be easily and systematically assessed are potentially useful tools for developing therapeutic compounds. Presented here is an optimized protocol for in vitro generation of the fibrin clot, as well as analysis of the effect of A&#946; and A&#946;-fibrinogen interaction inhibitors. The clot turbidity assay is rapid, highly reproducible and can be used to test multiple conditions simultaneously, allowing for the screening of large numbers of A&#946;-fibrinogen inhibitors. Hit compounds from this screening can be further evaluated for their ability to ameliorate A&#946;-induced structural abnormalities of the fibrin clot architecture using SEM. The effectiveness of these optimized protocols is demonstrated here using TDI-2760, a recently identified A&#946;-fibrinogen interaction inhibitor.

Presented here are two methods that can be used individually or in combination to analyze the effect of beta-amyloid on fibrin clot structure. Included is a protocol for creating an in vitro fibrin clot, followed by clot turbidity and scanning electron microscopy methods.

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and ...

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

Analysis of Protein Folding, Transport, and Degradation in Living Cells by Radioactive Pulse Chase

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and the degradation of target proteins in live cells. By using short (pulse) radiolabeling times (&lt;30 min) and tightly controlled chase times, it is possible to label only a small fraction of the total protein pool and follow its folding. When combined with nonreducing/reducing SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoprecipitation with (conformation-specific) antibodies, folding processes can be examined in great detail. This system has been used to analyze the folding of proteins with a huge variation in properties such as soluble proteins, single and multi-pass transmembrane proteins, heavily N- and O-glycosylated proteins, and proteins with and without extensive disulfide bonding. Pulse-chase methods are the basis of kinetic studies into a range of additional features, including co- and posttranslational modifications, oligomerization, and polymerization, essentially allowing the analysis of a protein from birth to death. Pulse-chase studies on protein folding are complementary with other biochemical and biophysical methods for studying proteins in vitro by providing increased temporal resolution and physiological information. The methods as described within this paper are adapted easily to study the folding of almost any protein that can be expressed in mammalian or insect-cell systems.

Here we describe a protocol for a general pulse-chase method that allows the kinetic analysis of folding, transport, and degradation of proteins to be followed in live cells.

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and ...

Interactions with and Membrane Permeabilization of Brain Mitochondria by Amyloid Fibrils

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary mechanism of amyloid aggregate-induced toxicity in neurodegenerative diseases. However, most reports describing the mechanisms of membrane disruption are based on phospholipid model systems, and studies directly targeting events occurring at the level of biological membranes are rare. Described here is a model for studying the mechanisms of amyloid toxicity at the membrane level. For mitochondrial isolation, density gradient medium is used to obtain preparations with minimal myelin contamination. After mitochondrial membrane integrity confirmation, the interaction of amyloid fibrils arising from &#945;-synuclein, bovine insulin, and hen egg white lysozyme (HEWL) with rat brain mitochondria, as an in vitro biological model, is investigated. The results demonstrate that treatment of brain mitochondria with fibrillar assemblies can cause different degrees of membrane permeabilization and ROS content enhancement. This indicates structure-dependent interactions between amyloid fibrils and mitochondrial membrane. It is suggested that biophysical properties of amyloid fibrils and their specific binding to mitochondrial membranes may provide explanations for some of these observations.

Provided here is a protocol for investigating the interactions between native form, prefibrillar, and mature amyloid fibrils of different peptides and proteins with mitochondria isolated from different tissues and various areas of the brain.

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary ...

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine exchangers in plants have not been identified. Here, we outline a method to characterize polyamine antiporters using membrane vesicles generated from the lysis of Escherichia coli cells heterologously expressing a plant antiporter. First, we heterologously expressed AtBAT1 in an E. coli strain deficient in polyamine and arginine exchange transporters. Vesicles were produced using a French press, purified by ultracentrifugation and utilized in a membrane filtration assay of labeled substrates to demonstrate the substrate specificity of the transporter. These assays demonstrated that AtBAT1 is a proton-mediated transporter of arginine, &#947;-aminobutyric acid (GABA), putrescine and spermidine. The mutant strain that was developed for the assay of AtBAT1 may be useful for the functional analysis of other families of plant and animal polyamine exchangers. We also hypothesize that this approach can be used to characterize many other types of antiporters, as long as these proteins can be expressed in the bacterial cell membrane. E. coli is a good system for the characterization of novel transporters, since there are multiple methods that can be employed to mutagenize native transporters.

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

We describe a method for the characterization of proton-driven membrane transporters in membrane vesicle preparations produced by heterologous expression in E. coli and lysis of cells using a French press.

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine ...