Name: measuring trans-plasma membrane electron transport by c2c12 myotubes
Uploaded: 2018-05-04T15:00:00
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 ...

<ol>
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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...

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 redu...

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 ...

The overall goal of this procedure is to monitor the process of trans-plasma membrane electron transport in real time through spectrophotometric assays. This method can help answer key questions in the radox biology field regarding the mechanism of trans-plasma membrane electron transport and how a cell's redox environment is maintained. The main advantage of this technique is that the assay is a real time, multi-well, quick spectrophotometric assay.To begin, seat C2C-12 adherence cells at two times 10 to the 4th cells per square centimeter in rows A through F of a 96 well plate, then cover the cells with growth media. Incubate the cells in a 37 degree Celsius incubator with 5%carbon dioxide. After 24 hours in culture, aspirate the growth media, wash with 150 microliters of PBS, replace with differentiation medium, and incubate the cells for an additional 24 to 48 hours.Next, prepare a stock WST-1 and PMS solutions. Make a 10 millimolar stock solution of WST-1 by dissolving 033 grams of WST-1 in five milliliters of PBS. Then, vortex the solution.Then, prepare the stock solution of PMS by dissolving 0023 grams of PMS in 1.5 milliliters of the ionized water. Vortex the solution. To prepare the working solution, add glucose to PBS to make 11.4 milliliters of a five millimolar glucose solution.To this solution, add 480 microliters of the WST-1 stock solution for a final concentration of 400 micromolar WST-1 and 48 microliters of the PMS stock solution for a final concentration of 20 micromolar PMS. Vortex the solutions to mix. Then divide the solution into two six milliliter aliquots.Depending on the assay, add the relevant components described here. With the solutions now prepared, aspirate the differentiation media from the cells and wash them once with 150 microliters of PBS. Remove the PBS and add 100 microliters from aliquot one into columns one through six.And add 100 microliters from aliquot two to columns seven through 12 of the 96 well plate. Place the 96 well plate into a plate reader and measure the absorbance values every 10 minutes for one hour at 438 nanometers. After the final reading, aspirate the media and wash each well with 150 microliters of PBS.Then, aspirate the PBS. For analysis, calculate the change in absorbance for each well by subtracting the initial absorbance for the well from the absorbance at any later time point for that same well. Correct this change based on the change in absorbance that is observed in the background wells.Once background corrected, normalize the absorbance data to the 60 minute control measurements. Alternatively, normalize by the protein content in each well using a standard BCA protein assay. Finally, quantify the data to obtain the amount of WST-1 reduction by microgram of protein.To begin the DPIP reduction assay, grow and differentiate C2C-12 adherence cells, as shown for the WST-1 reduction assay. Next, prepare a stock solution containing 10 millimolar DPIP by dissolving 029 grams of DPIP in 10 milliliters of the ionized water. Then vortex the solution.Confirm the concentration by measuring the solution's absorbance at 600 nanometers using a spectrophotometer. To make the working solution, add 120 microliters of the 10 millimolar DPIP stock solution to 11.88 milliliters of a five millimolar solution of glucose and PBS for a final DPIP working concentration of 100 micromolar. Then, vortex the solution.Then divide the solution into two six milliliters aliquots. Depending on the assay, add the relevant components described here. With the solutions now prepared, aspirate the differentiation media from the cells and wash them once with 150 microliters of PBS.Remove the PBS and add 100 microliters from aliquot one into columns one through six, and 100 microliters from aliquot two to columns seven through 12 of the 96 well plate. Place the 96 well plate into a plate reader and measure the absorbance values every 10 minutes for one hour at 600 nanometers and analyze the resulting values as described in the WST-1 reduction assay analysis. To begin, prepare two solutions.First, prepare six milliliters of 400 micromolar WST-1 and 20 micromolar PMS in PBS. Then prepare six milliliters of a 100 micromolar solution of DPIP and PBS. Prepare the first 96 well plate by adding 100 microliters of a WST-1/PMS solution to rows A through D in a flat bottom plate in the absence of cells, and measure the absorbance in a spectrophotometer at 438 nanometers.Then prepare a second 96 well plate by adding 100 microliters of the DPIP solution to rows A through D in a flat bottom plate in the absence of cells, and measure the absorbance in the spectrophotometer at 600 nanometers. Next, add one microliter of 10 millimolar ascorbate to half of the wells for a final concentration of 100 micromolar. Monitor the absorbance in the plate until it stabilizes.After this, repeat with DPIP. Upon stabilization, determine whether the produced electron acceptors are substrates. For ascorbate oxidase for super oxide dismutates, by adding either ascorbate oxidase or super oxide dismutates to each well and monitoring the absorbance change for one hour.To monitor transplasma membrane electron transport, ascorbate oxidase was used to block the effects of ascorbate mediated reduction and superoxide dismutase was used to block superoxide mediated reduction to determine which portion of WST-1 reduction was due to each component. With the addition of ascorbate oxidase, WST-1 reduction was suppressed by about 30%indicating that about 30%of the transplasma membrane electron transport was due to the export of ascorbate. With the addition of superoxide dismutase, WST-1 reduction was suppressed by about 70%indicating that about 70%of the transplasma membrane electron transport was due to extracellular superoxide release.To determine if reduced DPIP is a substrate for ascorbate oxidase or super oxide dismutase, DPIP levels were reduced with ascorbic acid. After 20 minutes, ascorbate oxidase was added, and the absorbance was monitored. The recovery shows that reduced DPIP is a substrate for ascorbate oxidase.When super oxide dismutase was added, DPIP was also reoxidized, albeit at a slower rate;therefore, reduced DPIP is a substrate for these enzymes and not an appropriate extracellular electron acceptor to be utilized with these enzymes. Once mastered, this technique can be completed in two hours if it is performed properly. While attempting this procedure, it's important to remember to make all reagents fresh each day.After its development, this technique paved the way for researchers in the field of redox biology to explore transplasma membrane electron transport in a variety of cell types. After watching this video, you should have a good understanding of how to measure transplasma membrane electron transport utilizing extracellular electron acceptors, as well as how to determine if the electron acceptor might be a substrate for enzymes utilized in the assay.

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

Research

JoVE Journal

Biochemistry

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 ...

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.

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

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 ...

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

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 ...

Measuring Nucleotide Binding to Intact, Functional Membrane Proteins in Real Time

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. This method combines expression of proteins tagged with the fluorescent non-canonical amino acid ANAP, and FRET between ANAP and fluorescent (trinitrophenyl) nucleotide derivatives. We present examples of nucleotide binding to ANAP-tagged KATP ion channels measured in unroofed plasma membranes and excised, inside-out membrane patches under voltage clamp. The latter allows for simultaneous measurements of ligand binding and channel current, a direct readout of protein function. Data treatment and analysis are discussed extensively, along with potential pitfalls and artefacts. This method provides rich mechanistic insights into the ligand-dependent gating of KATP channels and can readily be adapted to the study of other nucleotide-regulated proteins or any receptor for which a suitable fluorescent ligand can be identified.

This protocol presents a method for measuring adenine nucleotide binding to receptors in real time in a cellular environment. Binding is measured as F&#246;rster resonance energy transfer (FRET) between trinitrophenyl nucleotide derivatives and protein labeled with a non-canonical, fluorescent amino acid.

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. ...