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><tbody><tr><td>Liposomicator, Avanti Polar Lipids Inc.</td><td>&amp;nbsp;Avanti Polar Lipids Inc.</td><td>610200</td><td></td></tr><tr><td>IEC Centra CL2 Benchtop</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Orion Research Model 701A Digital pH-mV meter</td><td></td><td></td><td>These can be found on Ebay.</td></tr><tr><td>Non-functional pH probe</td><td></td><td></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><td>DI-710 Data Logger</td><td>DATAQ instruments</td><td></td><td></td></tr><tr><td>WinDAQ acquisition software</td><td>DATAQ instruments</td><td></td><td></td></tr><tr><td>Pierce Disposable Plastic Columns, Gravity-flow, 2ml</td><td>Pierce (Thermo Scientific)</td><td>29922</td><td></td></tr><tr><td>KIMAX Culture Tubes, Disposable, Borosilicate Glass</td><td>Kimble Chase</td><td>73500-13100</td><td></td></tr><tr><td>Extruder Set With Holder/Heating Block</td><td>Avanti Polar Lipids Inc.</td><td>610000</td><td></td></tr><tr><td>Computer</td><td></td><td></td><td></td></tr></tbody></table>

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

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

9.8K Views.  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.

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

Cholesterol Efflux Assay

Cholesterol content of cells must be maintained within the very tight limits, too much or too little cholesterol in a cell results in disruption of cellular membranes, apoptosis and necrosis 1. Cells can source cholesterol from intracellular synthesis and from plasma lipoproteins, both sources are sufficient to fully satisfy cells' requirements for cholesterol. The processes of cholesterol synthesis and uptake are tightly regulated and deficiencies of cholesterol are rare 2. Excessive cholesterol is more common problem 3. With the exception of hepatocytes and to some degree adrenocortical cells, cells are unable to degrade cholesterol. Cells have two options to reduce their cholesterol content: to convert cholesterol into cholesteryl esters, an option with limited capacity as overloading cells with cholesteryl esters is also toxic, and cholesterol efflux, an option with potentially unlimited capacity. Cholesterol efflux is a specific process that is regulated by a number of intracellular transporters, such as ATP binding cassette transporter proteins A1 (ABCA1) and G1 (ABCG1) and scavenger receptor type B1. The natural acceptor of cholesterol in plasma is high density lipoprotein (HDL) and apolipoprotein A-I. 
The cholesterol efflux assay is designed to quantitate the rate of cholesterol efflux from cultured cells. It measures the capacity of cells to maintain cholesterol efflux and/or the capacity of plasma acceptors to accept cholesterol released from cells. The assay consists of the following steps. Step 1: labelling cellular cholesterol by adding labelled cholesterol to serum-containing medium and incubating with cells for 24-48 h. This step may be combined with loading of cells with cholesterol. Step 2: incubation of cells in serum-free medium to equilibrate labelled cholesterol among all intracellular cholesterol pools. This stage may be combined with activation of cellular cholesterol transporters. Step 3: incubation of cells with extracellular acceptor and quantitation of movement of labelled cholesterol from cells to the acceptor. If cholesterol precursors were used to label newly synthesized cholesterol, a fourth step, purification of cholesterol, may be required.
The assay delivers the following information: (i) how a particular treatment (a mutation, a knock-down, an overexpression or a treatment) affects the capacity of cell to efflux cholesterol and (ii) how the capacity of plasma acceptors to accept cholesterol is affected by a disease or a treatment. This method is often used in context of cardiovascular research, metabolic and neurodegenerative disorders, infectious and reproductive diseases.

The cholesterol assay is designed to quantitate the rate of cholesterol efflux from cultured cells and the capacity of plasma acceptors to accept cholesterol released from cells. The assay consists of labelling cells with cholesterol, equilibration of cholesterol among intracellular pools and release of cholesterol to an extracellular acceptor.

Cholesterol content of cells must be maintained within the very tight limits, too much or too little cholesterol in a cell results in disruption of ...

Medicine

Models and Methods to Evaluate Transport of Drug Delivery Systems Across Cellular Barriers

Sub-micrometer carriers (nanocarriers; NCs) enhance efficacy of drugs by improving solubility, stability, circulation time, targeting, and release. Additionally, traversing cellular barriers in the body is crucial for both oral delivery of therapeutic NCs into the circulation and transport from the blood into tissues, where intervention is needed. NC transport across cellular barriers is achieved by: (i) the paracellular route, via transient disruption of the junctions that interlock adjacent cells, or (ii) the transcellular route, where materials are internalized by endocytosis, transported across the cell body, and secreted at the opposite cell surface (transyctosis). Delivery across cellular barriers can be facilitated by coupling therapeutics or their carriers with targeting agents that bind specifically to cell-surface markers involved in transport. Here, we provide methods to measure the extent and mechanism of NC transport across a model cell barrier, which consists of a monolayer of gastrointestinal (GI) epithelial cells grown on a porous membrane located in a transwell insert. Formation of a permeability barrier is confirmed by measuring transepithelial electrical resistance (TEER), transepithelial transport of a control substance, and immunostaining of tight junctions. As an example, ~200&#160;nm polymer NCs are used, which carry a therapeutic cargo and are coated with an antibody that targets a cell-surface determinant. The antibody or therapeutic cargo is labeled with 125I for radioisotope tracing and labeled NCs are added to the upper chamber over the cell monolayer for varying periods of time. NCs associated to the cells and/or transported to the underlying chamber can be detected. Measurement of free 125I allows subtraction of the degraded fraction. The paracellular route is assessed by determining potential changes caused by NC transport to the barrier parameters described above. Transcellular transport is determined by addressing the effect of modulating endocytosis and transcytosis pathways.

Many therapeutic applications require safe and efficient transport of drug carriers and their cargoes across cellular barriers in the body. This article describes an adaptation of established methods to evaluate the rate and mechanism of transport of drug nanocarriers (NCs) across cellular barriers, such as the gastrointestinal (GI) epithelium.

Sub-micrometer carriers (nanocarriers; NCs) enhance efficacy of drugs by improving solubility, stability, circulation time, targeting, and release. ...

Bioengineering

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used for ATP production. The amphiphilic nature of membrane proteins requires particular attention to their handling, and reconstitution into the natural lipid environment is indispensable when studying membrane transport processes like proton translocation. Here, we detail a method that has been used for the investigation of the proton-pumping mechanism of membrane redox enzymes, taking cytochrome bo3 from Escherichia coli as an example. A combination of electrochemistry and fluorescence microscopy is used to control the redox state of the quinone pool and monitor pH changes in the lumen. Due to the spatial resolution of fluorescent microscopy, hundreds of liposomes can be measured simultaneously while the enzyme content can be scaled down to a single enzyme or transporter per liposome. The respective single enzyme analysis can reveal patterns in the enzyme functional dynamics that might be otherwise hidden by the behavior of the whole population. We include a description of a script for automated image analysis.

Here, we present a protocol to study the molecular mechanism of proton translocation across lipid membranes of single liposomes, using cytochrome bo3 as an example. Combining electrochemistry and fluorescence microscopy, pH changes in the lumen of single vesicles, containing single or multiple enzyme, can be detected and analyzed individually.

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used ...

Biochemistry

Selection of Transporter-Targeted Inhibitory Nanobodies by Solid-Supported-Membrane (SSM)-Based Electrophysiology

Single domain antibodies (nanobodies) have been extensively used in mechanistic and structural studies of proteins and they pose an enormous potential as tools for developing clinical therapies, many of which depend on the inhibition of membrane proteins such as transporters. However, most of the methods used to determine the inhibition of transport activity are difficult to perform in high-throughput routines and depend on labeled substrates availability thereby complicating the screening of large nanobody libraries. Solid-supported membrane (SSM) electrophysiology is a high-throughput method, used for characterizing electrogenic transporters and measuring their transport kinetics and inhibition. Here we show the implementation of SSM-based electrophysiology to select inhibitory and non-inhibitory nanobodies targeting an electrogenic secondary transporter and to calculate nanobodies inhibitory constants. This technique may be especially useful for selecting inhibitory nanobodies targeting transporters for which labeled substrates are not available.

Selection of Transporter-Targeted Inhibitory Nanobodies by Solid-Supported-Membrane SSM-Based Electrophysiology

Nanobodies are important tools in structural biology and pose a great potential for the development of therapies. However, the selection of nanobodies with inhibitory properties can be challenging. Here we demonstrate the use of solid-supported-membrane (SSM)-based electrophysiology for the classification of inhibitory and non-inhibitory nanobodies targeting electrogenic membrane transporters.

Single domain antibodies (nanobodies) have been extensively used in mechanistic and structural studies of proteins and they pose an enormous potential as ...

A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes all liposomes of the ensemble to be identical. However, new experimental platforms able to observe liposomes at the single-particle level have made it possible to perform highly sophisticated and quantitative studies on protein-membrane interactions or drug carrier properties on individual liposomes, thus avoiding errors from ensemble averaging. Here we present a protocol for preparing, detecting, and analyzing single liposomes using a fluorescence-based microscopy assay, facilitating such single-particle measurements. The setup allows for imaging individual liposomes in a massive parallel manner and is employed to reveal intra-sample size and compositional inhomogeneities. Additionally, the protocol describes the advantages of studying liposomes at the single liposome level, the limitations of the assay, and the important features to be considered when modifying it to study other research questions.

This protocol describes the fabrication of liposomes and how these can be immobilized on a surface and imaged individually in a massive parallel manner using fluorescence microscopy. This allows for the quantification of the size and compositional inhomogeneity between single liposomes of the population.

Most research employing liposomes as membrane model systems or drug delivery carriers relies on bulk read-out techniques and thus intrinsically assumes ...

High-throughput Screening for Small-molecule Modulators of Inward Rectifier Potassium Channels

Specific members of the inward rectifier potassium (Kir) channel family are postulated drug targets for a variety of disorders, including hypertension, atrial fibrillation, and pain1,2. For the most part, however, progress toward understanding their therapeutic potential or even basic physiological functions has been slowed by the lack of good pharmacological tools. Indeed, the molecular pharmacology of the inward rectifier family has lagged far behind that of the S4 superfamily of voltage-gated potassium (Kv) channels, for which a number of nanomolar-affinity and highly selective peptide toxin modulators have been discovered3. The bee venom toxin tertiapin and its derivatives are potent inhibitors of Kir1.1 and Kir3 channels4,5, but peptides are of limited use therapeutically as well as experimentally due to their antigenic properties and poor bioavailability, metabolic stability and tissue penetrance. The development of potent and selective small-molecule probes with improved pharmacological properties will be a key to fully understanding the physiology and therapeutic potential of Kir channels.
The Molecular Libraries Probes Production Center Network (MLPCN) supported by the National Institutes of Health (NIH) Common Fund has created opportunities for academic scientists to initiate probe discovery campaigns for molecular targets and signaling pathways in need of better pharmacology6. The MLPCN provides researchers access to industry-scale screening centers and medicinal chemistry and informatics support to develop small-molecule probes to elucidate the function of genes and gene networks. The critical step in gaining entry to the MLPCN is the development of a robust target- or pathway-specific assay that is amenable for high-throughput screening (HTS).
Here, we describe how to develop a fluorescence-based thallium (Tl+) flux assay of Kir channel function for high-throughput compound screening7,8,9,10.The assay is based on the permeability of the K+ channel pore to the K+ congener Tl+. A commercially available fluorescent Tl+ reporter dye is used to detect transmembrane flux of Tl+ through the pore. There are at least three commercially available dyes that are suitable for Tl+ flux assays: BTC, FluoZin-2, and FluxOR7,8. This protocol describes assay development using FluoZin-2. Although originally developed and marketed as a zinc indicator, FluoZin-2 exhibits a robust and dose-dependent increase in fluorescence emission upon Tl+ binding. We began working with FluoZin-2 before FluxOR was available7,8 and have continued to do so9,10. However, the steps in assay development are essentially identical for all three dyes, and users should determine which dye is most appropriate for their specific needs. We also discuss the assay's performance benchmarks that must be reached to be considered for entry to the MLPCN. Since Tl+ readily permeates most K+ channels, the assay should be adaptable to most K+ channel targets.

Methods for developing and validating a quantitative fluorescence assay for measuring the activity of inward rectifier potassium (Kir) channels for high-throughput compound screening is presented.

Specific members of the inward rectifier potassium (Kir) channel family are postulated drug targets for a variety of disorders, including hypertension, ...

In vitro Investigation of the MexAB Efflux Pump From Pseudomonas aeruginosa

There is an emerging scientific need for reliable tools for monitoring membrane protein transport. We present a methodology leading to the reconstitution of efflux pumps from the Gram-negative bacteria Pseudomonas aeruginosa in a biomimetic environment that allows for an accurate investigation of their activity of transport. Three prerequisites are fulfilled: compartmentation in a lipidic environment, use of a relevant index for transport, and generation of a proton gradient. The membrane protein transporter is reconstituted into liposomes together with bacteriorhodopsin, a light-activated proton pump that generates a proton gradient that is robust as well as reversible and tunable. The activity of the protein is deduced from the pH variations occurring within the liposome, using pyranin, a pH-dependent fluorescent probe. We describe a step-by-step procedure where membrane protein purification, liposome formation, protein reconstitution, and transport analysis are addressed. Although they were specifically designed for an RND transporter, the described methods could potentially be adapted for use with any other membrane protein transporter energized by a proton gradient.

In vitro Investigation of the MexAB Efflux Pump From Pseudomonas aeruginosa

We present a protocol for the in vitro investigation of efflux pumps from Pseudomonas aeruginosa. This protocol allows for the generation of a robust, reversible, and tunable proton gradient within the liposome membrane and hence should be adaptable to any membrane protein energized by the protomotive force.

There is an emerging scientific need for reliable tools for monitoring membrane protein transport. We present a methodology leading to the reconstitution ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Membrane Transport Processes Analyzed by a Highly Parallel Nanopore Chip System at Single Protein Resolution

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable efforts have been made in this field, but techniques enabling automated high-throughput transport analysis in combination with solvent-free lipid bilayer techniques required for the analysis of membrane transporters are rare. This class of transporters however is crucial in cell homeostasis and therefore a key target in drug development and methodologies to gain new insights desperately needed.
The here presented manuscript describes the establishment and handling of a novel biochip for the analysis of membrane protein mediated transport processes at single transporter resolution. The biochip is composed of microcavities enclosed by nanopores that is highly parallel in its design and can be produced in industrial grade and quantity. Protein-harboring liposomes can directly be applied to the chip surface forming self-assembled pore-spanning lipid bilayers using SSM-techniques (solid supported lipid membranes). Pore-spanning parts of the membrane are freestanding, providing the interface for substrate translocation into or out of the cavity space, which can be followed by multi-spectral fluorescent readout in real-time. The establishment of standard operating procedures (SOPs) allows the straightforward establishment of protein-harboring lipid bilayers on the chip surface of virtually every membrane protein that can be reconstituted functionally. The sole prerequisite is the establishment of a fluorescent read-out system for non-electrogenic transport substrates.
High-content screening applications are accomplishable by the use of automated inverted fluorescent microscopes recording multiple chips in parallel. Large data sets can be analyzed using the freely available custom-designed analysis software. Three-color multi spectral fluorescent read-out furthermore allows for unbiased data discrimination into different event classes, eliminating false positive results.
The chip technology is currently based on SiO2 surfaces, but further functionalization using gold-coated chip surfaces is also possible.

The presented protocol describes the analysis of membrane protein mediated transport on the single transporter level using pore-spanning solvent-free lipid bilayers. This is achieved by the creation of bulk produced nanopore array chips, combined with highly parallel data acquisition and analysis, enabling the future establishment of membrane protein effector screenings.

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable ...

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. ...

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

Summary

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Chapters in this video

Cholesterol Efflux Assay

Models and Methods to Evaluate Transport of Drug Delivery Systems Across Cellular Barriers

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Selection of Transporter-Targeted Inhibitory Nanobodies by Solid-Supported-Membrane (SSM)-Based Electrophysiology

A Quantitative Fluorescence Microscopy-based Single Liposome Assay for Detecting the Compositional Inhomogeneity Between Individual Liposomes

High-throughput Screening for Small-molecule Modulators of Inward Rectifier Potassium Channels

In vitro Investigation of the MexAB Efflux Pump From Pseudomonas aeruginosa

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

Membrane Transport Processes Analyzed by a Highly Parallel Nanopore Chip System at Single Protein Resolution

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