This work was supported by NIH grant GM085232 and an Irma T. Hirschl/ Monique Weill-Caulier Scholar Award (to A.A.).

1. Lipid Preparation
<ol>
	<li>Aliquot the desired amount lipids into a clear glass tube. Use E. coli polar lipid extract, but most lipid compositions can be used. If the lipids are in powder form, resuspend them in chloroform to a concentration of 20 mg/ml. Dry the lipids at RT under N2 gas until all solvent has evaporated.</li>
	<li>Resuspend the lipids in pentane and dry it again a steady stream of gas N2 to remove leftover traces of chloroform. Dry until all solvent has evaporated. While drying, gently rotate the tube so that the lipid is distributed in a uniform film covering the bottom third of the tube rather than a forming a dense mass at the bottom.</li>
	<li>Add the reconstitution buffer containing detergent (300 mM KCl, 50 mM citric acid, 25 mM K2HPO4, pH 4.5, 35 mM CHAPS) to dissolve the lipids. Use other mild detergents for this step if the protein is poorly tolerant to CHAPS.</li>
	<li>Resuspend the lipids to clarity using a water bath sonicator. Immerse the tube in the center of the water sonicator bath at maximum power in short (30 sec-1 min) pulses with short (30 sec-1 min) pauses. The solution should become clear. 
	NOTE: The final degree of clarity achieved depends on the specific lipid composition used. Some batch-to-batch variation in this step, even among nominally identical lipids, is also possible. Residual haze is due to light scattering by large particles in the suspension, which can be removed by centrifugation.</li>
	<li>Incubate the resuspended lipids at RT&#160;for 20-60 min.</li>
</ol>
2. Proteoliposome Formation
NOTE: Several strategies can be employed to insert the detergent-solubilized protein into liposomes. For CLC-ec1 dialysis works well and is therefore the method of choice 6,9,10.
<ol>
	<li>Add the purified protein to the desired protein-to-lipid (P/L) ratio (expressed in &#181;g of Protein/mg Lipid). The choice of P/L ratio depends on the purpose of the experiment.
	<ol>
		<li>If the goal is to assess the activity of the protein of interest, reconstitute at high P/L&#8217;s to maximize the signal, as each liposome contains multiple copies of the protein. To quantify the activity and/or unitary transport rate of the protein, reconstitute in a Poisson dilution regime at low P/L&#8217;s, so that each vesicle contains on average 1 protein. See Step 7 and Discussion for a more in depth analysis of when to use each regime.</li>
	</ol>
	</li>
	<li>To determine the optimal P/L values for the various regimens, perform a complete titration of the activity as a function of protein concentration 8,11-13. However, for any protein for which the molecular weight (MW, expressed in kDa) of the active unit is known the following P/L values can be used as initial guesstimates: 
	<img alt="Equation 1" fo:content-width="1.5in" src="/files/ftp_upload/52369/52369eq1.jpg" /> 
	<img alt="Equation 2" fo:content-width="1.5in" src="/files/ftp_upload/52369/52369eq2.jpg" /></li>
	<li>Load the protein and lipid mixture in a pre-wetted dialysis tube or a cassette. Place the dialysis device in a beaker containing 500-1,000x the volume of the lipid reconstitution buffer (i.e., 1 L buffer for 1 ml of lipid resuspension) under gentle stirring.
	<ol>
		<li>Change buffer 3-4 times at intervals &#62;3 hr. At every solution change, wash the cassette/tubing and the beaker with distilled water and fill the beaker with fresh reconstitution buffer. Include 1-2 O/N&#160;intervals for the solution change. The exact number and duration of the solution changes depends on the exact lipid and detergents used.</li>
	</ol>
	</li>
	<li>After the last step is complete, remove the liposomes from the dialysis vessel, aliquot them in tubes and flash freeze them in liquid N2 or freeze at -80 &#176;C.</li>
</ol>
3. Recording Set-up
NOTE: The recording set-up (Figure 2A) consists of two chambers (flat bottomed cylinders, ~3-4 ml volume), a Cl- (see below) electrode, a pH meter with an analog or digital electrical output, a digitizer, and a computer with an appropriate acquisition software.
<ol>
	<li>Connect the Cl- electrode to the pH meter, the output of the pH meter to the digitizer which is connected to the computer. Place the reference electrode in one chamber and the recoding wire in the other (Figure 2B). 
	NOTE: The polarity of the wires is correct if &#916;VCal (see Step 6.4 and 7.2) is positive.</li>
	<li>Place the chambers on a stir plate with a stir bar in the recording chamber (Figure 1B). Make sure the stir bar does not touch the electrode.</li>
	<li>Connect the chambers using an agar bridge (Figure 2B) (100 mM KCl and 2% Agarose). Store the Agar bridges in 100 mM KCl.</li>
</ol>
4. Preparation of the Cl- Electrode
<ol>
	<li>Take an old and -possibly- non-functioning pH electrode. Break the glass coating to completely reveal the silver wires.</li>
	<li>Carefully remove the coating from the silver wires using a scalpel blade. Clean the wires using copious amounts of H2O and EtOH.</li>
	<li>Place in a saturated solution of FeCl3 (or bleach) until wires are covered with a uniform dark coat of AgCl.</li>
</ol>
5. Preparation of Unilamellar Vesicles
<ol>
	<li>The night before the experiments, swell 1 g of Sephadex G-50 beads in 15 ml of external buffer (1 mM KCl, 150 mM K2SO4, 25 mM citric acid, pH 4.5) with gentle shaking (do not stir) for at least 3 hr at RT before use. Each experiment requires ~3 ml of swollen beads. Keep the swollen beads at 4 &#176;C for a few days at most.</li>
	<li>While the liposomes are thawing at RT, pour in each column ~3 ml of swollen G-50 beads. Let them dry by gravity flow at RT; this usually takes 1-2 min.</li>
	<li>Prepare unilamellar vesicles by extruding the proteoliposomes 11 times through a Mini-Extruder using a 0.4 m Teflon cutoff.</li>
	<li>Place the column in a plastic round-bottomed tube. Spin the columns to remove excess solution. Centrifuge for 20-30 sec at 1,400 x g in a clinical centrifuge.</li>
	<li>Discard flow-through, place column in a 13 x 100 mm glass tube and add 100 &#181;l of the extruded vesicles to the column. Spin column for 1 min at 500 x g in a clinical centrifuge.</li>
	<li>Collect ~200 &#181;l of flow-through that will be added to the recording chamber in step 6.5.</li>
</ol>
6. Efflux Measurement
<ol>
	<li>Place 2 ml of 100 mM KCl in the reference (ground) chamber and 1.8 ml of the external buffer (1 mM KCl, 150 mM K2SO4, 25 mM citric acid, pH 4.5) to the recording chamber. The slight osmotic imbalance between the internal and external solutions does not affect</li>
	<li>Start the acquisition program. Let the baseline equilibrate, this may take a few minutes.</li>
	<li>Once the signal reaches a stable baseline (the liposomes are ready and the columns dry), start recording (Figure 3A).</li>
	<li>Add 15 &#181;l of a 10 mM KCl solution to calibrate the system (Figure 2A).</li>
	<li>Add liposomes. A small jump might be visible due to incomplete removal of external Cl- from the proteoliposomes (Figure 3A). Wait till the baseline stabilizes.</li>
	<li>Add Valinomycin (1 &#181;l at 1 mg/ml in EtOH) to initiate efflux (Figure 3A).</li>
	<li>Let the efflux run its time course till it plateaus (Figure 3A).</li>
	<li>Add 40 &#181;l of 1.5 M &#946;-octylglucoside (&#946;-OG) prepared in the external buffer to dissolve all liposomes (Figure 3A). End the recording.</li>
</ol>
7. Data Analysis
<ol>
	<li>Export the trace file from in an ascii or text format and import it to the analysis program of choice.</li>
	<li>Measure the Voltage at the following points (Figure 3A): 
	At the beginning of the recording (V0); 
	After addition of the calibrating pulse (Vcal); 
	After addition of the liposomes (Vlipo); 
	At the end of the efflux (Vfin); 
	After addition of detergent (Vtot); 
	Baseline the voltages so that V0=0.</li>
	<li>Use the Nernst-Plank equation to determine the experimental value of <img alt="Equation 3" fo:content-width=".3in" src="/files/ftp_upload/52369/52369eq3.jpg" /> by measuring 
	<img alt="Equation 4" fo:content-width="3in" src="/files/ftp_upload/52369/52369eq4.jpg" /> 
	Where VolCal is the volume of the calibration pulse in &#181;l, 1,800 is the chamber volume at the beginning of the experiment expressed&#160;in &#181;l, [Cl]cal/in are the Cl- concentrations of the calibration pulse and of the external buffer in mM and &#916;Vcal is the jump in mV measured after the calibration pulse. 
	NOTE: This empirical determination of &#945; serves three purposes: it ensures that the system is responding properly, offers a measure of the consistency of the instrument&#8217;s response between experiments as well as to check that no mistakes were made in the solution making.</li>
	<li>Calculate the Cl concentrations after each step as follows: 
	<img alt="Equation 5" fo:content-width="3.5in" src="/files/ftp_upload/52369/52369eq5.jpg" /> 
	<img alt="Equation 6" fo:content-width="3.5in" src="/files/ftp_upload/52369/52369eq6.jpg" /> 
	<img alt="Equation 7" fo:content-width="3.5in" src="/files/ftp_upload/52369/52369eq7.jpg" /> 
	The factor 3/400 comes from the dilution of the 15 &#181;l calibration pulse into the 2,000 &#181;l final volume.</li>
	<li>Calculate the changes in [Cl] at each step, &#916;Cllipo/fin/tot.</li>
	<li>Convert V(t) in Cl(t) with 
	<img alt="Equation 8" fo:content-width="3.5in" src="/files/ftp_upload/52369/52369eq8.jpg" /> 
	Directly determine the [Cl] values at the critical points and compare them to the calculated values as an internal control.</li>
	<li>Normalize the trace so that the Clrellipo=0 and Clreltot=1 
	<img alt="Equation 9" fo:content-width="3.5in" src="/files/ftp_upload/52369/52369eq9.jpg" /></li>
	<li>Fit the efflux time course to the following equation: 
	<img alt="Equation 10" fo:content-width="3.5in" src="/files/ftp_upload/52369/52369eq10.jpg" /> 
	Where L is the rate of Cl- leak that is experimentally determined by measuring Cl- efflux from protein-free liposomes prepared in the same conditions and &#964; is the time constant of the process.</li>
	<li>Calculate v, the initial velocity: 
	<img alt="Equation 11" fo:content-width="3.5in" src="/files/ftp_upload/52369/52369eq11.jpg" /> 
	Where &#916;Clfin is expressed in millimolar and V is the volume of the chamber in &#956;l.</li>
	<li>Calculate the transport rate/conductance 
	<img alt="Equation 12" fo:content-width="3.5in" src="/files/ftp_upload/52369/52369eq12.jpg" /> 
	Where p is the P/L, MW is the molecular weight of the active complex expressed in kDa</li>
</ol>

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The authors declare no competing financial interests.

We have described a detailed protocol to measure Cl- transport mediated by purified anion-selective channels or transporters reconstituted in liposomes. The example used was the prokaryotic H+/Cl- exchanger CLC-ec1. However, the methodology can be readily adapted to study channels gated by ligands 12,13,15, voltage 11,12, or sporting different anionic selectivity 15,16 by replacing the Ag:AgCl electrode with one suitable for the ion under consideration. El...

In last two decades the ability to overexpress and purify membrane transport proteins has dramatically increased: ion channels, primary and secondary transporters are now routinely purified from heterologous expression systems as well as natural sources. New approaches to monitor expression, improve and facilitate the extraction and enhance stability of these proteins are constantly being developed 1-5. These technological breakthroughs have been instrumental in triggering the explosion of atomic-level structural information on membrane proteins which, in turn, enhanced our understanding of the structural bases of their function. In contrast, our ability to probe the functional properties of the purified proteins did not increase at the same rate, so that in some cases high resolution structural information is accompanied by qualitative functional data, thus limiting our ability to quantitatively test structure-based predictions. Hence, the development of quantitative and generalizable functional assays is a key step towards the elucidation of the mechanistic underpinnings of membrane protein function.
Here we describe an efflux assay that can be used to quantitatively determine key functional properties of purified and reconstituted Cl- channels and transporters. The principles underlying the assay can be generalized to a variety of transport systems, as well as to non ion-transporting proteins. Liposomes are reconstituted with purified Cl- channel/transporters in the presence of a large Cl- gradient (Figure 1A, B). Cl- efflux is initiated by the addition of an ionophore to allow for counter-ion flux, in our case the K+ ionophore valinomycin, which shunt the voltage established by the Cl- gradient and set the initial membrane potential to the equilibrium potential of K+6,7. Without the ionophore no significant net Cl- efflux occurs, as it is prevented by the generation of a transmembrane potential. The data is quantitatively described by two measurable parameters (Figure 1C): &#964;, the time constant of Cl- efflux, and f0, the fraction of liposomes not containing an active protein. From &#964; and f0 the unitary Cl- transport rate, the fraction of active proteins and the molecular mass of the active complex can be derived 8. The technique is illustrated here using proteoliposomes reconstituted with CLC-ec1, a well characterized CLC-type H+/Cl- exchanger of known structure and function. This assay is readily generalized to channels or transporters with different ionic selectivity or whose activity depends on the presence of voltage and/or ligands. Furthermore, this assay can be used to determine whether small molecules directly affect the reconstituted protein, to quantitate the effects of these compounds and how membrane composition or lipid-modifying reagents affect the function of the reconstituted channels and transporters.

<table border="0" cellpadding="0" cellspacing="0" width="995">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:372px;">Name of Reagent/ Equipment</td>
			<td style="width:337px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Liposomicator, Avanti Polar Lipids Inc.</td>
			<td style="width:337px;">&#160;Avanti Polar Lipids Inc.</td>
			<td>610200</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">IEC Centra CL2 Benchtop</td>
			<td style="width:337px;">Thermo Scientific</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Orion Research Model 701A Digital pH-mV meter</td>
			<td style="width:337px;"></td>
			<td style="width:119px;"></td>
			<td>These can be found on Ebay.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">&#160;Non-functional pH probe</td>
			<td style="width:337px;"></td>
			<td style="width:119px;"></td>
			<td>Any pH meter probe with silver wires will work. The glass/plastic coating needs to be removed and the wires cleaned.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">&#160; DI-710 Data Logger</td>
			<td style="width:337px;">DATAQ instruments</td>
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			<td height="21" style="height:21px;">WinDAQ acquisition software</td>
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a proteoliposome-based efflux assay to determine single-molecule properties of cl- channels and transporters

The last 15 years have been characterized by an explosion in the ability to overexpress and purify membrane proteins from prokaryotic organisms as well as from eukaryotes. This increase has been largely driven by the successful push to obtain structural information on membrane proteins. However, the ability to functionally interrogate these proteins has not advanced at the same rate and is often limited to qualitative assays of limited quantitative value, thereby limiting the mechanistic insights that they can provide. An assay to quantitatively investigate the transport activity of reconstituted Cl- channels or transporters is described. The assay is based on the measure of the efflux rate of Cl- from proteoliposomes following the addition of the K+ ionophore valinomycin to shunt the membrane potential. An ion sensitive electrode is used to follow the time-course of ion efflux from proteoliposomes reconstituted with the desired protein. The method is highly suited for mechanistic studies, as it allows for the quantitative determination of key properties of the reconstituted protein, such as its unitary transport rate, the fraction of active protein and the molecular mass of the functional unit. The assay can also be utilized to determine the effect of small molecule compounds that directly inhibit/activate the reconstituted protein, as well as to test the modulatory effects of the membrane composition or lipid-modifying reagents. Where possible, direct comparison between results obtained using this method were found to be in good agreement with those obtained using electrophysiological approaches. The technique is illustrated using CLC-ec1, a CLC-type H+/Cl- exchanger, as a model system. The efflux assay can be utilized to study any Cl- conducting channel/transporter and, with minimal changes, can be adapted to study any ion-transporting protein.

We describe a detailed and robust protocol to measure Cl- transport mediated by purified CLC-ec1, a prokaryotic CLC-type H+/Cl- exchanger, reconstituted in liposomes. A schematic representation of the experiment is shown in Figure 3. Proteoliposomes reconstituted with purified CLC-ec1 and containing high internal Cl- are immersed in a bath solution containing low Cl-. Under these conditions net Cl- efflux is prevented by the buildup of pos...

Watch this Scientific Journal Video about A Proteoliposome-Based Efflux Assay to Determine Single-molecule Properties of Cl- Channels and Transporters at JoVE.com

A Proteoliposome-Based Efflux Assay to Determine Single-molecule Properties of Cl- Channels and Transporters

Proteoliposomes are used to study purified channels and transporters reconstituted in a well-defined biochemical environment. An experimental procedure to measure efflux mediated by these proteins is illustrated. The steps to prepare proteoliposomes, perform the recordings, and analyze data to quantitatively determine the functional properties of the reconstituted protein are described.

A Proteoliposome-Based Efflux Assay to Determine Single-molecule Properties of Cl- Channels and Transporters

The last 15 years have been characterized by an explosion in the ability to overexpress and purify membrane proteins from prokaryotic organisms as well as ...

a-proteoliposome-based-efflux-assay-to-determine-single-molecule

Research

JoVE Journal

Biology

9.7K ビュー.  Weill Cornell Medical College. Proteoliposomes are used to study purified channels and transporters reconstituted in a well-defined biochemical environment. An experimental procedure to measure efflux mediated by these proteins is illustrated. The steps to prepare proteoliposomes, perform the recordings, and analyze data to quantitatively determine the functional properties of the reconstituted protein are described.

ビデオ: A Proteoliposome-Based Efflux Assay to Determine Single-molecule Properties of Cl- Channels and Transporters

Single Molecule Methods for Monitoring Changes in Bilayer Elastic Properties

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein interactions and more general lipid bilayer-protein interactions. These interactions are particularly important in pharmacological research, as many current pharmaceuticals on the market can alter the lipid bilayer material properties, which can lead to altered membrane protein function.   The formation of gramicidin channels are dependent on conformational changes in gramicidin subunits which are in turn dependent on the properties of the lipid.  Hence the gramicidin channel  current is a reporter of altered properties of the bilayer due to certain compounds.  

Membrane protein function is regulated by the cell membrane lipid composition. This video-article details how to form a patch using bilayer patch electrodes, as well as how to use gramicidin channels as reporters of altered membrane properties.

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Direct Observation of Enzymes Replicating DNA Using a Single-molecule DNA Stretching Assay

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. Linearized &lambda; DNA is modified to have a biotin on the end of one strand, and a digoxigenin moiety on the other end of the same strand. The biotinylated end is attached to a functionalized glass coverslip and the digoxigeninated end to a small bead. The assembly of these DNA-bead tethers on the surface of a flow cell allows a laminar flow to be applied to exert a drag force on the bead. As a result, the DNA is stretched close to and parallel to the surface of the coverslip at a force that is determined by the flow rate (Figure 1). The length of the DNA is measured by monitoring the position of the bead. Length differences between single- and double-stranded DNA are utilized to obtain real-time information on the activity of the replication proteins at the fork. Measuring the position of the bead allows precise determination of the rates and processivities of DNA unwinding and polymerization (Figure 2).

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system.

We describe a method for observing real time replication of individual DNA molecules mediated by proteins of the bacteriophage replication system. ...

Single-Molecule Imaging of Nuclear Transport

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the last decade. The investigation of molecular interactions with high temporal and spatial resolutions deep within cells has remained challenging due to the inherently weak signals arising from individual molecules. Recent works by Yang et al. demonstrated that narrow-field epifluorescence microscopy allows visualization of nucleocytoplasmic transport at the single molecule level. By the single molecule approach, important kinetics, such as nuclear transport time and efficiency, for signal-dependent and independent cargo molecules have been obtained. Here we described a protocol for the methodological approach with an improved spatiotemporal resolution of 0.4 ms and 12 nm. The improved resolution enabled us to capture transient active transport and passive diffusion events through the nuclear pore complexes (NPC) in semi-intact cells. We expect this method to be used in elucidating other binding and trafficking events within cells.

Single molecule microscopy approch provided novel insights into nuclear transport.

The utility of single molecule fluorescence microscopy approaches has been proven to be of a great avail in understanding biological reactions over the ...

A Fluorescent Screening Assay for Identifying Modulators of GIRK Channels

G protein-gated inward rectifier K+ (GIRK) channels function as cellular mediators of a wide range of hormones and neurotransmitters and are expressed in the brain, heart, skeletal muscle and endocrine tissue1,2. GIRK channels become activated following the binding of ligands (neurotransmitters, hormones, drugs, etc.) to their plasma membrane-bound, G protein-coupled receptors (GPCRs). This binding causes the stimulation of G proteins (Gi and Go) which subsequently bind to and activate the GIRK channel. Once opened the GIRK channel allows the movement of K+ out of the cell causing the resting membrane potential to become more negative. As a consequence, GIRK channel activation in neurons decreases spontaneous action potential formation and inhibits the release of excitatory neurotransmitters. In the heart, activation of the GIRK channel inhibits pacemaker activity thereby slowing the heart rate.
GIRK channels represent novel targets for the development of new therapeutic agents for the treatment neuropathic pain, drug addiction, cardiac arrhythmias and other disorders3. However, the pharmacology of these channels remains largely unexplored. Although a number of drugs including anti-arrhythmic agents, antipsychotic drugs and antidepressants block the GIRK channel, this inhibition is not selective and occurs at relatively high drug concentrations3.
Here, we describe a real-time screening assay for identifying new modulators of GIRK channels. In this assay, neuronal AtT20 cells, expressing GIRK channels, are loaded with membrane potential-sensitive fluorescent dyes such as bis-(1,3-dibutylbarbituric acid) trimethine oxonol [DiBAC4(3)] or HLB 021-152 (Figure 1). The dye molecules become strongly fluorescent following uptake into the cells (Figure 1). Treatment of the cells with GPCR ligands stimulates the GIRK channels to open. The resulting K+ efflux out of the cell causes the membrane potential to become more negative and the fluorescent signal to decrease (Figure 1). Thus, drugs that modulate K+ efflux through the GIRK channel can be assayed using a fluorescent plate reader. Unlike other ion channel screening assays, such atomic absorption spectrometry4 or radiotracer analysis5, the GIRK channel fluorescent assay provides a fast, real-time and inexpensive screening procedure.

A real-time screening procedure for identifying drugs that interact with G protein-gated inward rectifier K+ (GIRK) channels is described. The assay utilizes membrane potential-sensitive fluorescent dyes to measure GIRK channel activity. This technique is adaptable for use on a number of cell lines.

G protein-gated inward rectifier K+ (GIRK) channels function as cellular mediators of a wide range of hormones and neurotransmitters and are expressed in ...

Studying DNA Looping by Single-Molecule FRET

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into nucleosomes. Thermodynamics of dsDNA bending has been studied by a method called cyclization which relies on DNA ligase to covalently join short sticky ends of a dsDNA. However, ligation efficiency can be affected by many factors that are not related to dsDNA looping such as the DNA structure surrounding the joined sticky ends, and ligase can also affect the apparent looping rate through mechanisms such as nonspecific binding. Here, we show how to measure dsDNA looping kinetics without ligase by detecting transient DNA loop formation by FRET (Fluorescence Resonance Energy Transfer). dsDNA molecules are constructed using a simple PCR-based protocol with a FRET pair and a biotin linker. The looping probability density known as the J factor is extracted from the looping rate and the annealing rate between two disconnected sticky ends. By testing two dsDNAs with different intrinsic curvatures, we show that the J factor is sensitive to the intrinsic shape of the dsDNA.

This study presents a detailed experimental procedure to measure looping dynamics of double-stranded DNA using single-molecule Fluorescence Resonance Energy Transfer (FRET). The protocol also describes how to extract the looping probability density called the J factor.

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into ...

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 ...

Application of Electrophysiology Measurement to Study the Activity of Electro-Neutral Transporters

The transport of ions through cell membranes ensures the fine control of ion content within and outside the cell that is indispensable for cell survival. These transport mechanisms are mediated by the activities of specialized transporter proteins. Specifically,pH dynamics are finely controlled by plasma membrane proton (H+) extrusion systems, such as the Na+/H+ exchanger (NHE) protein family. Despite extensive efforts to study the mechanisms underlying NHE regulation, our current understanding of the biophysical and molecular properties of the NHE family is inadequate because of the limited availability of methods to effectively measure NHE activity. In this manuscript, we used H+-selective electrodes during whole-cell patch clamping recording to measure NHE-induced H+ flux. We proposed this approach to overcome some limitations of typically used methods to measure NHE activity, such as radioactive uptake and fluorescent membrane permeants. Measurement of NHE activity using the described method enables high sensitivity and time resolution and more efficient control of intracellular H+ concentrations. H+-selective electrodes are based on the fact that transporter activity creates an ion gradient in close proximity to the cell membrane. An H+-selective electrode moving up to and away from the cell membrane in a repetitive, oscillatory fashion records a voltage difference that is dependent on H+ flux. While H+-selective electrodes are used to detect H+ flux moving out of the cell, the patch clamp method in the whole-cell configuration is used to control the intracellular ion composition. Moreover, application of the giant patch clamp technique allows modification of the intracellular composition of not only ions but also lipids. The transporter activity of NHE isoform 3 (NHE3) was measured using this technical approach to study the molecular basis of NHE3 regulation by phosphoinositides.

This manuscript describes the applications of proton-selective electrodes and patch clamping methods to measure the activity of proton transport systems. These methods overcome some limitations of techniques commonly used to study proton transport activity, such as moderate sensitivity, time resolution and insufficient intracellular milieu control.

The transport of ions through cell membranes ensures the fine control of ion content within and outside the cell that is indispensable for cell survival. ...

The Lactate Dehydrogenase Sequestration Assay &#8212; A Simple and Reliable Method to Determine Bulk Autophagic Sequestration Activity in Mammalian Cells

Bulk autophagy is characterized by the sequestration of large portions of cytoplasm into double/multi-membrane structures termed autophagosomes. Here a simple protocol to monitor this process is described. Moreover, typical results and experimental validation of the method under autophagy-inducing conditions in various types of cultured mammalian cells are provided. During bulk autophagy, autophagosomes sequester cytosol, and thereby also soluble cytosolic proteins, alongside other autophagic cargo. LDH is a stable and highly abundant, soluble cytosolic enzyme that is non-selectively sequestered into autophagosomes. The amount of LDH sequestration therefore reflects the amount of bulk autophagic sequestration. To efficiently and accurately determine LDH sequestration in cells, we employ an electrodisruption-based fractionation protocol that effectively separates sedimentable from cytosolic LDH, followed by measurement of enzymatic activity in sedimentable fractions versus whole-cell samples. Autophagic sequestration is determined by subtracting the proportion of sedimentable LDH in untreated cells from that in treated cells. The advantage of the LDH sequestration assay is that it gives a quantitative measure of the autophagic sequestration of endogenous cargo, as opposed to other methods that either involve ectopic expression of sequestration probes or semi-quantitative protease protection analyses of autophagy markers or receptors.

The Lactate Dehydrogenase Sequestration Assay &amp;#8212; A Simple and Reliable Method to Determine Bulk Autophagic Sequestration Activity in Mammalian Cells

Here a simple and well-validated protocol for measuring bulk autophagic sequestration activity in mammalian cells is described. The method is based on quantifying the proportion of lactate dehydrogenase (LDH) in sedimentable cell fractions compared to total cellular LDH levels.

Bulk autophagy is characterized by the sequestration of large portions of cytoplasm into double/multi-membrane structures termed autophagosomes. Here a ...

Parallel High Throughput Single Molecule Kinetic Assay for Site-Specific DNA Cleavage

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable of measuring SSDC in many single DNA molecules simultaneously. Bead-tethered substrate DNAs, each containing a single copy of the target sequence, are prepared in a microfluidic flow channel. An external magnet applies a weak force to the paramagnetic beads. The integrity of up to 1,000 individual DNAs can be monitored by visualizing the microbeads under darkfield imaging using a wide-field, low magnification objective. Injecting of a restriction endonuclease, NdeI, initiates the cleavage reaction. Video microscopy is used to record the exact moment of each DNA cleavage by observing the frame in which the associated bead moves up and out of the focal plane of the objective. Frame-by-frame bead counting quantifies the reaction, and an exponential fit determines the reaction rate. This method allows collection of quantitative and statistically significant data on single molecule SSDC reactions in a single experiment.

A highly parallel method for measuring the site-specific cleavage of DNA at the single molecule level is described. This protocol demonstrates the technique using the restriction endonuclease NdeI. The method can easily be modified to study any process that results in site-specific DNA cleavage.

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable ...