We wish to thank the Calcium Imaging Core Facility at CMS for their help and guidance in image capture and analysis. This study was supported by the U.S. National Institutes of Health (NIH) HL102208 to N.A.B. This approach was originally pioneered by Holger Lorenz and Jennifer Lippincott-Schwartz at NIH.

1. Generation and Validation of Fluorescent Protein Chimeras
<ol>
	<li>Append green fluorescent protein (GFP) to the amino or carboxyl termini of the protein of interest using standard recombinant DNA strategies13. 
	NOTE: Although we have used GFP extensively, other variants such as cyan fluorescent protein (CFP), yellow fluorescent protein (YFP), DsRed and mCherry can be employed. Improvements in fluorophore folding efficiency, brightness and photostability are constantly being made, and investigators should avail themselves of the latest generation constructs.
	<ol>
		<li>Confirm correct DNA sequence and ensure that the GFP moiety is in frame with the protein of interest. Perform sequencing using standard recombinant DNA strategies21. 
		NOTE: Many institutions have core DNA facilities, which the investigator is encouraged to contact for additional information.</li>
		<li>For single or double pass membrane proteins, generate both amino- and carboxyl terminal versions of the fusion proteins19.</li>
		<li>For multi-spanning proteins insert GFP within putative extramembrane loops, as well as the termini. Take special care when generating transmembrane chimeras with the fluorescent fusion protein attached to the amino terminus of the target protein as signal sequences are often cleaved by proteolysis once the protein has passed through the ER translocon.</li>
		<li>In the case of extant signal sequences, insert the fluorescent fusion protein distal to the cleavage site so as to generate a mature protein containing an intact amino-terminal tagged fluorescent protein. Employ standard recombinant DNA strategies for the generation of chimeras21.</li>
	</ol>
	</li>
	<li>Determine the presence of sufficient appropriately localized fluorescent signal in the cell line of interest.
	<ol>
		<li>Observe the cells under epifluorescence microscopy using the intrinsic fluorescence of GFP under appropriate filter set conditions. For GFP filter sets that encompass 510-560 nm, confirm that the correct filters are used for the fluorophore employed. 
		NOTE: A sufficient signal is achieved when assessment by eye provides a strong enough signal that the cellular structure within which the signal resides is easily resolved above background. In addition the staining pattern of the GFP-protein should be consistent with that of markers for the appropriate organelle, and preferably identical to that of the native untagged protein.</li>
	</ol>
	</li>
	<li>Determine whether a stable or transient transfection approach will be utilized based upon the protein being evaluated. 
	NOTE: Stable transfectants generally afford lower levels of expression than transient systems. Although transient expression yields high level expression of proteins and increase signal intensity (~5-24 hr post transfection depending on the protein), it can also lead to overexpression artifacts such as protein aggregation or saturation of protein targeting pathways, leading to erroneous subcellular localization.</li>
</ol>
2. Transfection of Cells and Cell Culture
NOTE: A variety of cells are amenable to FPP analysis, including Hek293, HeLa, COS-7 and CHO cells14. The following description is based on expressing membrane proteins in Hek293 cells.
<ol>
	<li>Grow human embryonic kidney cells (Hek293) as adherent monolayers in 6 ml of Dulbecco&#8217;s Modified Eagles Medium (DMEM), supplemented with 5% fetal bovine serum (FBS), 1% sodium pyruvate, and 250 &#181;g/ml penicillin/streptomycin at 37 &#176;C with 5% CO2. Perform all work with cells in a laminar flow tissue culture hood. Grow cells in culture medium until ~80% confluent in a T-75 flask.</li>
	<li>Transfect cells using any standard high-efficiency, low toxicity transfection method, including lipid-, viral- or electroporation-based technologies to generate transient or stable expression. Typically 2 &#181;g of plasmid DNA per 1 x 106 cells gives good reproducible protein expression. Maintain cells for 5-48 hr in appropriate tissue culture media at 37 &#176;C in a CO2 capable tissue culture incubator.</li>
	<li>Monitor cells for expression of the GFP-fusion protein using epifluorescence microscopy, using the intrinsic fluorescence signal of GFP, using appropriate filter sets according to manufacturer&#8217;s protocol. Fluorescence signal usually reaches a maximum within 24-72 hr following transfection. Use cells when the fluorescence signal reaches a maximum level. Monitor protein expression levels by immunoblot analysis using antibodies directed against either the protein of interest or the GFP moiety.</li>
	<li>For experimental analysis plate cells on coated coverslips. View either under a live-cell configuration or following fixation and mount on microscopy slides. Plate cells to achieve 60%-80% confluence within 24-48 hr.
	<ol>
		<li>Coat coverslips with poly-lysine solution (0.1 mg/ml), so that the entire surface is covered. Incubate for 30 min under an ultra violet light at room temperature, followed by 60 min in a cell culture incubator at 37 &#176;C. Remove excess poly-lysine and rinse the coverslips in PBS 3 times.</li>
		<li>Alternatively, plate the cells on a chambered cover-glass consisting of cell culture plates with an optical glass bottom to allow direct imaging with oil-immersion objectives. The actual microscopy setup depends on the equipment available to the investigator, but should be capable of rapid solution changes. Depending on the cell type and cell shape, cells should be around 60-90% confluent at the time of the FPP assay.</li>
	</ol>
	</li>
</ol>
3. Fluorescence Microscopy Setup
<ol>
	<li>Use a high numerical aperture (NA) oil-immersion objective for maximal signal collection and spatial resolution, such as a 60X, 1.42 NA oil immersion objective. Use the following lasers to stimulate the fluorophores: Argon gas 488 nm (GFP and YFP) and 561 diode (RFP, mCherry).</li>
	<li>Determine the optimal exposure, gain and capture rate of images for each construct. Adjust laser intensity and exposure time to minimize photobleaching (see step 7.1.2).</li>
	<li>For studies using multiple fluorophores (e.g., a soluble marker for membrane permeabilization in concert with a membrane protein marker), choose appropriate filter sets to minimize signal cross-talk and maximize signal-to-noise ratios.</li>
</ol>
4. Establishing Optimal Conditions for Plasma Membrane Permeabilization
<ol>
	<li>Remove the cell culture medium from cells expressing only soluble fluorescent proteins (e.g. EGFP, DsRed). Wash the cells 3 times (1 min each) in KHM buffer (110 mM potassium acetate, 3 mM MgCl2, 20 mM HEPES-NaOH, pH 7.2). Perform experiments with cells and all reagents at room temperature.</li>
	<li>Place cells on coverslips (see step 2.4) in KHM buffer on a fluorescence microscope stage and collect the first image, which represents the control &#39;non-permeabilized&#39; stage.</li>
	<li>To selectively permeabilize the plasma membrane, add digitonin (30 &#181;M) in KHM buffer to the cells. Perfuse cells with buffer (buffer + digitonin) at a rate of 5 ml/min for 10-60 sec.</li>
	<li>Determine the optimal digitonin concentration by gradually increasing the concentration of digitonin. The optimal digitonin concentration is achieved when there is complete loss of fluorescence signal from soluble EGFP within 10-60 sec of digitonin application. For many cells application of 10-50 &#956;M digitonin is sufficient to permeabilize the cell membrane.
	<ol>
		<li>For all applications use the lowest possible concentration of digitonin. Collect images after digitonin permeabilization for permeabilized background signal.</li>
	</ol>
	</li>
	<li>Confirm that the GFP-protein chimera of interest is membrane integrated by confirming cellular retention of the fluorescence signal following digitonin permeabilization of the plasma membrane16.
	<ol>
		<li>Express an organelle specific fluorescent protein23. Mitochondrial proteins work well in this regard16, e.g., pDsRed2-Mito vector. Determine optimal expression conditions as described in step 2.1.</li>
		<li>Confirm that the digitonin concentration is appropriate for the cell type employed by ensuring that the morphology of intracellular organelles remains constant over prolonged (up to 60 min) exposure to digitonin. 
		NOTE: Use standard immunofluorescence microscopy protocols using antibodies against various cellular compartments, such as lysosomes (e.g., LAMP1), Golgi (e.g., TGN38), endoplasmic reticulum (e.g., calnexin), endosomes (e.g., Rab5), mitochondria (e.g., cytochrome c). If intracellular organelle morphology/integrity changes dramatically over 60 min, it is likely that the digitonin concentration was too high, and requires a reduction in digitonin concentration and/or exposure time. 
		NOTE: Digitonin is toxic by inhalation and skin contact. Wear appropriate personal protection equipment (PPE).</li>
	</ol>
	</li>
</ol>
5. Protease Treatment of Fluorescent Proteins
<ol>
	<li>Wash the cells in KHM buffer and then add protease (proteinase K; 50 &#956;g/ml in KHM buffer) directly onto the cells. Perfuse cells at a flow rate of 5 ml/min using a gravity fed perfusion system to continuously bath the preparation.
	<ol>
		<li>Store solutions in 50 ml syringes and polyethylene tubing connected independent reservoirs to a manifold with a single outlet at the bath (5 ml/min flow rate). Position a suction pipette to maintain a constant (0.5 ml) bath volume and achieve solution exchange by manually switching between perfusion reservoirs.</li>
	</ol>
	</li>
	<li>Collect images to determine whether the fluorescent signal persists or degrades. A loss of &#62;90% of the initial signal intensity over 30-60 sec is consistent with proteolytic degradation of the GFP-protein chimera (see Representative Results).</li>
	<li>If no signal degradation is observed with proteinase K, when such loss is anticipated, replace the proteinase K solution with trypsin (4-8 mM) in KHM buffer. If necessary, utilize a mixture of proteinase K and trypsin. There is no need to quench the protease activity.</li>
</ol>
6. Alternative Approach for Plasma Membrane Proteins with External Domains
<ol>
	<li>For plasma membrane proteins, perform the assay for external fluorescent proteins chimeras prior to digitonin permeabilization. Wash the cells on coverslips in KHM buffer (5 ml/min x 3 min), and take an image for pre-protease treatment and quantify pre-protease fluorescence signal as in Step 7.</li>
	<li>Expose cells to protease (proteinase K; 50 &#956;g/ml or trypsin 4-8 mM in KHM buffer) in the absence of digitonin by perfusing the cells with KHM media containing protease (flow rate 5 ml/min). Collect images of the fluorescent protein, to determine how quickly the signal disappears or how long the signal persists.</li>
	<li>Before digitonin permeabilization, quench all protease activity, by washing the cells three times (1 min each) in cell culture media containing 10% serum (FBS).</li>
	<li>Wash the cells by perfusing with KHM buffer (flow rate of 5 ml/min) for 2 min. Acquire a pre-permeabilization image. Permeabilize the cell membrane as described in Step 4.</li>
	<li>Collect images following protease addition as described in Step 5.</li>
</ol>
7. Image Analysis
<ol>
	<li>Use an inverted microscope capable of &#8220;live cell&#8221; recording, in which loss of fluorescence signal can be monitored in real-time. For continuous monitoring, employ identical parameters (exposure time, gain, laser intensity, etc.).
	<ol>
		<li>Determine experimental parameters by observing the fluorescence signal from the GFP-protein of interest under control conditions (i.e., KBH buffer alone). Adjust the gain, laser intensity and exposure time settings on the computer-driven microscope to ensure that there is no appreciable loss of signal intensity over a 10-20 min period. 
		NOTE: Signal intensity is not an absolute value, but is based on the microscope system and settings available to each PI. Monitor changes in relative fluorescence intensity.</li>
	</ol>
	</li>
	<li>Alternatively, treat cells at set time-points in digitonin followed by protease, and fix cells prior to mounting on coverslips for imaging. Determine time points empirically following procedures in steps 4 and 5. For the initial investigation, start at 0, 30, 60, 120 and 300 sec post addition of digitonin or protease.
	<ol>
		<li>Fix cells (20 min at room temperature) using a sufficient quantity of fixative (3.7% paraformaldehyde in PBS) to fully submerge the cells.</li>
		<li>Quench residual paraformaldehyde by rinsing cells with PBS containing 20 mM glycine (3 x 5 min).</li>
		<li>Mount coverslips by removing excess liquid from the coverslip and inverting the coverslip onto 50 &#181;l mounting media on a glass microscope slides for imaging. Allow slides to dry overnight before imaging.</li>
	</ol>
	</li>
	<li>Collect fluorescence microscopy images (under the appropriate filters sets compatible with the fluorophore being used) by using the video image capture function on a computer controlled fluorescence microscope.</li>
	<li>Quantify fluorescent signal intensities as a function of incubation condition. Obtain the mean pixel intensity within a region of interest (ROI) that encloses an individual cell.</li>
</ol>

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The authors declare no conflict of interest.

The correct orientation and topology of membrane proteins is essential for their proper function. Despite the importance of understanding membrane protein topology, there are many proteins for which such data is completely lacking. FPP provides an easy and efficient way of determining membrane protein topology, and one that can be performed by most laboratories. The FPP approach affords significant advantages over previous methods for determining protein topology. For example, there is no need to have a panel of multiple...

The plasma membranes, as well as the numerous intracellular membranes, serve as barriers separating two aqueous compartments. In the case of the plasma membrane, the separation is between the outside and inside of the cell; for intracellular organelles it is between the cytoplasm and the organelle lumen. For example, the endoplasmic reticulum (ER) membrane separates an oxidizing environment within the lumen of the ER from a cytosolic reducing environment1. Membrane proteins are synthesized on ER-associated ribosomes, and achieve their final topology within the ER membrane2. The acquisition of appropriate membrane orientation and topology for proteins is critical for their normal function. Correct topology allows relevant domains of membrane proteins to interact with their binding partners, it allows critical post-translational modifications to occur, and in the case of plasma membrane proteins, allows the cell to interact with and respond to its environment. To fully appreciate the function of a membrane protein, it is clearly imperative to know how that protein is oriented with respect to the membrane within which it resides, i.e., its membrane topology. In addition to acquisition of basic scientific knowledge, understanding the topology of a membrane protein and which aspects of a protein surface are exposed to different environments has marked clinical implications since membrane proteins comprise the majority of pharmacological targets3. Until recently, approaches to determining membrane protein topology have required considerable investment in time and money or have required reagents that are difficult to come by.
Both experimental and in silico approaches have been employed to determine the membrane topology of proteins residing within the plasma membrane. Since the first predictions of membrane spanning domains based on the evaluated hydrophobicity of individual amino acids3, numerous predictive algorithms are now available on the internet, and simply require knowledge of the protein&#8217;s amino acid sequence. However, assumptions are often central to such modeling programs, assumptions that can lead to incorrect assignments of topology4,5. Moreover, while these computer-based predictions can tentatively assign membrane spanning regions, they do not always determine whether the amino or carboxy termini of proteins are in the cytoplasm, organelle lumen or cell exterior. Even with increased computational power, and the use of machine-learning algorithms6, such data is still a model, and must be validated using experimentally acquired data. Direct experimental determination of membrane topology has been undertaken using panels of monoclonal antibodies with known epitopes distributed throughout the protein, where assessment of their immunoreactivity has been made before and after cell permeabilization. This approach requires a set of antibodies, which may not be available for the protein of interest.
An alternative strategy is to engineer epitope tags such as myc or hemagglutinin (HA) into various locations throughout the protein, again followed by determination of immunoreactivity before and after membrane permeabilization. In addition to immunogenic tags, enzymatic tags (including alkaline phosphatase, &#946;-galactosidase, or &#946;-lactamase) and chemical modifications such as cysteine scanning have all been employed to determine membrane protein topology7,8. An additional method employed for topological mapping of plasma membrane proteins relies on a slightly different approach to epitope tags. In this method the tag sequence is the N-linked glycosylation consensus sequence NXS/T. Since glycosylation only occurs when such a sequence is present in the lumen of the biosynthetic pathway, the presence of the tag in a luminal versus a cytosolic compartment is easily observed as a mass shift on SDS-PAGE gels. Such an approach has been applied to the multispanning ion channel CFTR9. While all these approaches have been utilized, it is clear that they all require considerable investments in molecular biology to generate and sequence the manifold constructs.
To determine topological information regarding transmembrane proteins located within intracellular organelles has proven to be more challenging. The application of fluorescence based technologies however, has made the determination of membrane topology a lot simpler. The technique of bimolecular fluorescence complementation (BiFC) relies on the interaction between two non-fluorescent fragments of a fluorescent protein, restoring the fluorescent properties of that protein10. Although initially described for determining protein-protein interactions in vivo10, this approach has also been utilized to determine membrane protein topology in plant cells11. However this approach is also time intensive as it requires the generation not only of fusion proteins with the protein of interest but also a variety of fusion proteins targeted to the cytosol or organelle lumen, and requires knowledge of which intracellular membranes the protein of interest resides in.
An alternative simpler approach to determining membrane protein topology has been described12. The assay, Fluorescence Protease Protection (FPP) requires the generation of a fusion protein between GFP and the gene of interest. The approach is based on the relative accessibility of non-specific proteases to the GFP moiety; depending on whether GFP is protected from proteolysis by residing in the lumen of intracellular organelles or is exposed to proteolysis by being present in the cytosol. Thus, if the GFP moiety on the protein of interest faces the cytoplasm, it will be exposed to protease activity and the fluorescence signal lost. Conversely, if the GFP moiety on the protein of interest faces an environment &#39;protected&#39; from the protease (such as the Golgi lumen) then the fluorescence signal will persist.
To allow proteases to enter the cell, but not enter intracellular membrane compartments the cholesterol binding drug digitonin is used. Cholesterol is the dominant sterol in vertebrates, and is specifically enriched in the plasma membrane relative to intracellular compartments. The glycoside toxin, digitonin, is extracted from the plant Digitalis purpurea. Digitonin has an affinity for cholesterol rich membranes, where it leads to selective membrane permeabiliztion14,16 (Figure 1). In addition to allowing exit of small cytosolic components, digitonin permeabilization also permits the entry of exogenous molecules, such as proteinase K or trypsin. The FPP protocol takes advantage of the fact that the plasma membrane contains up to 80% of cellular cholesterol17, whereas other organelles, such as ER, Golgi, endosomes, mitochondria, which have very low cholesterol content, remain intact14. The selective incorporation of cholesterol into the plasma membrane has been observed in many eukaryotic cells, permitting the use of digitonin-dependent plasma membrane permeabilization in such diverse eukaryotic species as S. cerevisiae to human14,18. The FPP assay provides a simple, rapid and fairly robust means of determining (a) whether a protein is membrane bound/associated or freely diffusible in the cytosol and (b) which domain of a membrane protein faces the cytosol or organelle lumen. Should a membrane protein have multiple orientations, the signal will arise from the dominant form and minor forms will not be detected. While there can be some concern that the addition of a GFP moiety to the protein on interest may affect its function and/or subcellular localization, this is actually more theoretical than actual. Indeed many studies have clearly shown that the GFP tag does not alter the properties of the protein19,20.

<table border="0" cellpadding="0" cellspacing="0" width="370">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:111px;">Name</td>
			<td style="width:80px;">Company</td>
			<td style="width:116px;">Catalgue Number</td>
			<td style="width:64px;">Comments</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Digitonin</td>
			<td>Calbiochem</td>
			<td align="right">300410</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Proteinase K</td>
			<td>Sigma</td>
			<td>P2308</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Trypsin</td>
			<td>Sigma</td>
			<td>T3924</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cav1-GFP</td>
			<td>Addgene</td>
			<td align="right">44433</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">pAcGFP-1 Golgi</td>
			<td>Clontech</td>
			<td align="right">632464</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Polylysine</td>
			<td>Sigma</td>
			<td>P4707</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mounting Media</td>
			<td>Dako</td>
			<td>cS704</td>
			<td></td>
		</tr>
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determining membrane protein topology using fluorescence protease protection (fpp)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.

Plasma Membrane Permeabilization
Efficient plasma membrane permeabilization is determined by the use of soluble fluorescent proteins (e.g., GFP, DsRed) (Step 4). These proteins, when expressed in cells, are free to diffuse in the cytosol, and are lost when the plasma membrane is permeabilized using digitonin (Figure 2). A complete disappearance of fluorescent signal should occur within 10-60 sec of digitonin application. Confirmation that the protein of inter...

Watch this Scientific Journal Video about Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP at JoVE.com

Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP.

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established ...

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17.6K Views.  Chicago Medical School. Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP. 

Video: Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect protease activity uses casein labeled with fluorescein isothiocyanate (FITC) as the substrate.
Protease activity results in the cleavage of the FITC-labeled casein substrate into smaller fragments, which do not precipitate under acidic conditions. After incubation of the protease sample and substrate, the reaction is acidified with the addition of trichloroacetic acid (TCA). The mixture is then centrifuged with the undigested substrate forming a pellet and the smaller, acid soluble fragments remaining in solution. The supernatant is neutralized and the fluorescence of the FITC-labeled fragments is measured.
The described kit procedure detects the trypsin protease control at a concentration of approximately 0.5 &mu;g/ml (5 ng of trypsin added to the assay). This sensitivity can be increased with a longer incubation time, up to 24 hours. The assay is performed in microcentrifuge tubes and procedures are provided for fluorescence detection using either cuvettes or multiwell plates.

The Protease Fluorescent Detection Kit is designed for the measurement of protease activity using fluorometry. It is also suitable for detection of trace amounts of protease contamination. The method is based on the proteolytic hydroysis of a proprietary formulation of a FITC-labeled casein substrate.

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect ...

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Using the GELFREE 8100 Fractionation System for Molecular Weight-Based Fractionation with Liquid Phase Recovery

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based fractionation. The system is comprised of single-use, 8-sample capacity cartridges and a benchtop GELFREE Fractionation Instrument. During separation, a constant voltage is applied between the anode and cathode reservoirs, and each protein mixture is electrophoretically driven from a loading chamber into a specially designed gel column gel. Proteins are concentrated into a tight band in a stacking gel, and separated based on their respective electrophoretic mobilities in a resolving gel. As proteins elute from the column, they are trapped and concentrated in liquid phase in the collection chamber, free of the gel. The instrument is then paused at specific time intervals, and fractions are collected using a pipette. This process is repeated until all desired fractions have been collected. If fewer than 8 samples are run on a cartridge, any unused chambers can be used in subsequent separations.
This novel technology facilitates the quick and simple separation of up to 8 complex protein mixtures simultaneously, and offers several advantages when compared to previously available fractionation methods. This system is capable of fractionating up to 1mg of total protein per channel, for a total of 8mg per cartridge. Intact proteins over a broad mass range are separated on the basis of molecular weight, retaining important physiochemical properties of the analyte. The liquid phase entrapment provides for high recovery while eliminating the need for band or spot cutting, making the fractionation process highly reproducible1.

The accompanying video describes the use of the GELFREE 8100 Fractionation System, which partitions complex protein samples on the basis of molecular weight and recovers the fractions in liquid phase. The video describes how the technology works, how it is used, and provides resultant data, with polyacrylamide gel electrophoresis analysis of fractionated bovine liver homogenate.

The GELFREE 8100 Fractionation System is a novel protein fractionation system designed to maximize protein recovery during molecular weight based ...

Bimolecular Fluorescence Complementation (BiFC) Assay for Protein-Protein Interaction in Onion Cells Using the Helios Gene Gun

Investigation of gene function in diverse organisms relies on knowledge of how the gene products interact with each other in their normal cellular environment. The Bimolecular Fluorescence Complementation (BiFC) Assay1 allows researchers to visualize protein-protein interactions in living cells and has become an essential research tool. This assay is based on the facilitated association of two fragments of a fluorescent protein (GFP) that are each fused to a potential interacting protein partner. The interaction of the two protein partners would facilitate the association of the N-terminal and C-terminal fragment of GFP, leading to fluorescence. For plant researchers, onion epidermal cells are an ideal experimental system for conducting the BiFC assay because of the ease in obtaining and preparing onion tissues and the direct visualization of fluorescence with minimal background fluorescence. The Helios Gene Gun (BioRad) is commonly used for bombarding plasmid DNA into onion cells. We demonstrate the use of Helios Gene Gun to introduce plasmid constructs for two interacting Arabidopsis thaliana transcription factors, SEUSS (SEU) and LEUNIG HOMOLOG (LUH)2 and the visualization of their interactions mediated by BiFC in onion epidermal cells.

Bimolecular Fluorescence Complementation BiFC Assay for Protein-Protein Interaction in Onion Cells Using the Helios Gene Gun

This article illustrates how to properly use the BioRad Helios Gene Gun to introduce plasmid DNA into onion epidermal cells and how to test for protein-protein interactions in onion cells based on the principle of Bimolecular Fluorescence Complementation (BiFC)

Investigation of gene function in diverse organisms relies on knowledge of how the gene products interact with each other in their normal cellular ...

Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter lipid bilayer properties. This is important because membrane proteins are energetically coupled to their host bilayer by hydrophobic interactions. Changes in bilayer properties thus alter membrane protein function, which provides an indirect way for amphiphiles to modulate protein function and a possible mechanism for "off-target" drug effects. We have previously developed an electrophysiological assay for detecting changes in lipid bilayer properties using linear gramicidin channels as probes 3,12. Gramicidin channels are mini-proteins formed by the transbilayer dimerization of two non-conducting subunits. They are sensitive to changes in their membrane environment, which makes them powerful probes for monitoring changes in lipid bilayer properties as sensed by bilayer spanning proteins. We now demonstrate a fluorescence assay for detecting changes in bilayer properties using the same channels as probes. The assay is based on measuring the time-course of fluorescence quenching from fluorophore-loaded large unilamellar vesicles due to the entry of a quencher through the gramicidin channels. We use the fluorescence indicator/quencher pair 8-aminonaphthalene-1,3,6-trisulfonate (ANTS)/Tl+ that has been successfully used in other fluorescence quenching assays 5,13. Tl+ permeates the lipid bilayer slowly 8 but passes readily through conducting gramicidin channels 1,14. The method is scalable and suitable for both mechanistic studies and high-throughput screening of small molecules for bilayer-perturbing, and potential "off-target", effects. We find that results using this method are in good agreement with previous electrophysiological results 12.

We introduce a fast fluorescence-based assay that monitors the rate of fluorescence quenching as a measure of gramicidin channel activity. The gramicidin channels are used as molecular force transducers to monitor changes in lipid bilayer properties as sensed by bilayer spanning proteins.

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter ...

Detection of Protein Interactions in Plant using a Gateway Compatible Bimolecular Fluorescence Complementation (BiFC) System

We have developed a BiFC technique to test the interaction between two proteins in vivo. This is accomplished by splitting a yellow fluorescent protein (YFP) into two non-overlapping fragments. Each fragment is cloned in-frame to a gene of interest. These constructs can then be co-transformed into Nicotiana benthamiana via Agrobacterium mediated transformation, allowing the transit expression of fusion proteins. The reconstitution of YFP signal only occurs when the inquest proteins interact 1-7. To test and validate the protein-protein interactions, BiFC can be used together with yeast two hybrid (Y2H) assay. This may detect indirect interactions which can be overlooked in the Y2H. Gateway technology is a universal platform that enables researchers to shuttle the gene of interest (GOI) into as many expression and functional analysis systems as possible8,9. Both the orientation and reading frame can be maintained without using restriction enzymes or ligation to make expression-ready clones. As a result, one can eliminate all the re-sequencing steps to ensure consistent results throughout the experiments. We have created a series of Gateway compatible BiFC and Y2H vectors which provide researchers with easy-to-use tools to perform both BiFC and Y2H assays10. Here, we demonstrate the ease of using our BiFC system to test protein-protein interactions in N. benthamiana plants.

Detection of Protein Interactions in Plant using a Gateway Compatible Bimolecular Fluorescence Complementation BiFC System

We have developed a technique to test protein-protein interactions in plant. A yellow fluorescent protein (YFP) is split into two non-overlapping fragments. Each fragment is cloned in-frame to a gene of interest via Gateway system, enabling expression of fusion proteins. Reconstitution of YFP signal only occurs when the inquest proteins interact.

We have developed a BiFC technique to test the interaction between two proteins in vivo. This is accomplished by splitting a yellow fluorescent protein ...

The Cell-based L-Glutathione Protection Assays to Study Endocytosis and Recycling of Plasma Membrane Proteins

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes for degradation or to the plasma membrane for recycling. The cell based L-glutathione protection assays can be used to study endocytosis and recycling of protein receptors, channels, transporters, and adhesion molecules localized at the cell surface. The endocytic assay requires labeling of cell surface proteins with a cell membrane impermeable biotin containing a disulfide bond and the N-hydroxysuccinimide (NHS) ester at 4 &#186;C -&#160;a temperature at which membrane trafficking does not occur. Endocytosis of biotinylated plasma membrane proteins is induced by incubation at 37 &#186;C. Next, the temperature is decreased again to 4 &#186;C to stop endocytic trafficking and the disulfide bond in biotin covalently attached to proteins that have remained at the plasma membrane is reduced with L-glutathione. At this point, only proteins that were endocytosed remain protected from L-glutathione and thus remain biotinylated. After cell lysis, biotinylated proteins are isolated with streptavidin agarose, eluted from agarose, and the biotinylated protein of interest is detected by western blotting. During the recycling assay, after biotinylation cells are incubated at 37 &#176;C to load endocytic vesicles with biotinylated proteins and the disulfide bond in biotin covalently attached to proteins remaining at the plasma membrane is reduced with L-glutathione at 4 &#186;C as in the endocytic assay. Next, cells are incubated again at 37 &#176;C to allow biotinylated proteins from endocytic vesicles to recycle to the plasma membrane. Cells are then incubated at 4 &#186;C, and the disulfide bond in biotin attached to proteins that recycled to the plasma membranes is reduced with L-glutathione. The biotinylated proteins protected from L-glutathione are those that did not recycle to the plasma membrane.

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes for degradation or to the plasma membrane for recycling. Methods described in this article are designed to study endocytosis and recycling of plasma membrane proteins.

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel protein-protein interaction pairs and monitoring protein interaction dynamics are of particular interest for revealing how plants respond to environmental factors and/or developmental signals. A plethora of approaches have been developed to examine protein-protein interactions, either in vitro or in vivo. Among them, the recently established luciferase complementation imaging (LCI) assay is the simplest and fastest method for demonstrating in vivo protein-protein interactions. In this assay, protein A or protein B is fused with the amino-terminal or carboxyl-terminal half of luciferase, respectively. When protein A interacts with protein B, the two halves of luciferase will be reconstituted to form a functional and active luciferase enzyme. Luciferase activity can be recorded with a luminometer or CCD-camera. Compared with other approaches, the LCI assay shows protein-protein interactions both qualitatively and quantitatively. Agrobacterium infiltration in Nicotiana benthamiana leaves is a widely used system for transient protein expression. With the combination of LCI and transient expression, these approaches show that the physical interaction between COP1 and SPA1 was gradually reduced after jasmonate treatment.

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

This manuscript describes an easy and rapid experimental procedure for determining protein-protein interactions based on the measurement of luciferase activity.

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel ...

Dosimetry for Cell Irradiation using Orthovoltage (40-300 kV) X-Ray Facilities

The importance of dosimetry protocols and standards for radiobiological studies is self-evident. Several protocols have been proposed for dose determination using low energy X-ray facilities, but depending on the irradiation configurations, samples, materials or beam quality, it is sometimes difficult to know which protocol is the most appropriate to employ. We, therefore, propose a dosimetry protocol for cell irradiations using low energy X-ray facility. The aim of this method is to perform the dose estimation at the level of the cell monolayer to make it as close as possible to real cell irradiation conditions. The different steps of the protocol are as follows: determination of the irradiation parameters (high voltage, intensity, cell container etc.), determination of the beam quality index (high voltage-half value layer couple), dose rate measurement with ionization chamber calibrated in air kerma conditions, quantification of the attenuation and scattering of the cell culture medium with EBT3 radiochromic films, and determination of the dose rate at the cellular level. This methodology must be performed for each new cell irradiation configuration as the modification of only one parameter can strongly impact the real dose deposition at the level of the cell monolayer, particularly involving low energy X-rays.

Dosimetry for Cell Irradiation using Orthovoltage 40-300 kV X-Ray Facilities

This document describes a new dosimetry protocol for cell irradiations using low energy X-ray equipment. Measurements are performed in conditions simulating real cell irradiation conditions as much as possible.