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><tbody><tr><td>Digitonin</td><td>Calbiochem</td><td>300410</td><td></td></tr><tr><td>Proteinase K</td><td>Sigma</td><td>P2308</td><td></td></tr><tr><td>Trypsin</td><td>Sigma</td><td>T3924</td><td></td></tr><tr><td>Cav1-GFP</td><td>Addgene</td><td>44433</td><td></td></tr><tr><td>pAcGFP-1 Golgi</td><td>Clontech</td><td>632464</td><td></td></tr><tr><td>Polylysine</td><td>Sigma</td><td>P4707</td><td></td></tr><tr><td>Mounting Media</td><td>Dako</td><td>cS704</td><td></td></tr></tbody></table>

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...

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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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Biology

17.7K 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

Using a GFP-tagged TMEM184A Construct for Confirmation of Heparin Receptor Identity

When novel proteins are identified through affinity-based isolation and bioinformatics analysis, they are often largely uncharacterized. Antibodies against specific peptides within the predicted sequence allow some localization experiments. However, other possible interactions with the antibodies often cannot be excluded. This situation provided an opportunity to develop a set of assays dependent on the protein sequence. Specifically, a construct containing the gene sequence coupled to the GFP coding sequence at the C-terminal end of the protein was obtained and employed for these purposes. Experiments to characterize localization, ligand affinity, and gain of function were originally designed and carried out to confirm the identification of TMEM184A as a heparin receptor1. In addition, the construct can be employed for studies addressing membrane topology questions and detailed protein-ligand interactions. The present report presents a range of experimental protocols based on the GFP-TMEM184A construct expressed in vascular cells that could easily be adapted for other novel proteins.

A construct encoding TMEM184A with a GFP tag at the carboxy-terminus designed for eukaryotic expression, was employed in assays designed to confirm the identification of TMEM184A as a heparin receptor in vascular cells.

When novel proteins are identified through affinity-based isolation and bioinformatics analysis, they are often largely uncharacterized. Antibodies ...

Genetics

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

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

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

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

Biochemistry

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

Membrane proteins are essential for cell viability and are therefore important therapeutic targets1-3. Since they function in complexes4, methods to identify and characterize their interactions are necessary5. To this end, we developed the Membrane Strep-protein interaction experiment, called Membrane-SPINE6. This technique combines in vivo cross-linking using the reversible cross-linker formaldehyde with affinity purification of a Strep-tagged membrane bait protein. During the procedure, cross-linked prey proteins are co-purified with the membrane bait protein and subsequently separated by boiling. Hence, two major tasks can be executed when analyzing protein-protein interactions (PPIs) of membrane proteins using Membrane-SPINE: first, the confirmation of a proposed interaction partner by immunoblotting, and second, the identification of new interaction partners by mass spectrometry analysis. Moreover, even low affinity, transient PPIs are detectable by this technique. Finally, Membrane-SPINE is adaptable to almost any cell type, making it applicable as a powerful screening tool to identify PPIs of membrane proteins.

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

A biochemical approach is described to identify in vivo protein-protein interactions (PPI) of membrane proteins. The method combines protein cross-linking, affinity purification and mass spectrometry, and is adaptable to almost any cell type or organism. With this approach, even the identification of transient PPIs becomes possible.

Membrane proteins are essential for cell viability and are therefore important therapeutic targets1-3. Since they function in complexes4, methods to ...

Bioengineering

Use of Recombinant Fusion Proteins in a Fluorescent Protease Assay Platform and Their In-gel Renaturation

Proteases are intensively studied enzymes due to their essential roles in several biological pathways of living organisms and in pathogenesis; therefore, they are important drug targets. We have developed a magnetic-agarose-bead-based assay platform for the investigation of proteolytic activity, which is based on the use of recombinant fusion protein substrates. In order to demonstrate the use of this assay system, a protocol is presented on the example of human immunodeficiency virus type 1 (HIV-1) protease. The introduced assay platform can be utilized efficiently in the biochemical characterization of proteases, including enzyme activity measurements in mutagenesis, kinetic, inhibition, or specificity studies, and it may be suitable for high-throughput substrate screening or may be adapted to other proteolytic enzymes.
In this assay system, the applied substrates contain N-terminal hexahistidine (His6) and maltose-binding protein (MBP) tags, cleavage sites for tobacco etch virus (TEV) and HIV-1 proteases, and a C-terminal fluorescent protein. The substrates can be efficiently produced in Escherichia coli cells and easily purified using nickel (Ni)-chelate-coated beads. During the assay, the proteolytic cleavage of bead-attached substrates leads to the release of fluorescent cleavage fragments, which can be measured by fluorimetry. Additionally, cleavage reactions can be analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). A protocol for the in-gel renaturation of assay components is also described, as partial renaturation of fluorescent proteins enables their detection based on molecular weight and fluorescence.

Here, we present the detailed procedure of a recently developed protease assay platform utilizing N-terminal hexahistidine/maltose-binding protein and fluorescent protein-fused recombinant substrates attached to the surface of nickel-nitrilotriacetic acid magnetic agarose beads. A subsequent in-gel analysis of the assay samples separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis is also presented.

Proteases are intensively studied enzymes due to their essential roles in several biological pathways of living organisms and in pathogenesis; therefore, ...

Rapid Assessment of Membrane Protein Quality by Fluorescent Size Exclusion Chromatography

During membrane protein structural elucidation and biophysical characterization, it is common to trial numerous protein constructs containing different tags, truncations, deletions, fusion partner insertions, and stabilizing mutations to find one that is not aggregated after extraction from the membrane. Furthermore, buffer screening to determine the detergent, additive, ligand, or polymer that provides the most stabilizing condition for the membrane protein is an important practice. The early characterization of membrane protein quality by fluorescent size exclusion chromatography provides a powerful tool to assess and rank different constructs or conditions without the requirement for protein purification, and this tool also minimizes the sample requirement. The membrane proteins must be fluorescently tagged, commonly by expressing them with a GFP tag or similar. The protein can be solubilized directly from whole cells and then crudely clarified by centrifugation; subsequently, the protein is passed down a size exclusion column, and a fluorescent trace is collected. Here, a method for running FSEC and representative FSEC data on the GPCR targets sphingosine-1-phosphate receptor (S1PR1) and serotonin receptor (5HT2AR) are presented.

The present protocol describes a procedure to perform fluorescent size exclusion chromatography (FSEC) on membrane proteins to assess their quality for downstream functional and structural analysis. Representative FSEC results collected for several G-protein coupled receptors (GPCRs) under detergent-solubilized and detergent-free conditions are presented.

During membrane protein structural elucidation and biophysical characterization, it is common to trial numerous protein constructs containing different ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

Validating interactions between different proteins is vital for investigation of their biological functions on the molecular level. There are several methods, both in vitro and in vivo, to evaluate protein binding, and at least two methods that complement the shortcomings of each other should be conducted to obtain reliable insights. 
For an in vivo assay, the bimolecular fluorescence complementation (BiFC) assay represents the most popular and least invasive approach that enables to detect protein-protein interaction within living cells, as well as identify the intracellular localization of the interacting proteins 1,2. In this assay, non-fluorescent N- and C-terminal halves of GFP or its variants are fused to tested proteins, and when the two fusion proteins are brought together due to the tested proteins&#x2019; interactions, the fluorescent signal is reconstituted3-6. Because its signal is readily detectable by epifluorescence or confocal microscopy, BiFC has emerged as a powerful tool of choice among cell biologists for studying about protein-protein interactions in living cells 3. This assay, however, can sometimes produce false positive results. For example, the fluorescent signal can be reconstituted by two GFP fragments arranged as far as 7 nm from each other due to close packing in a small subcellular compartment, rather that due to specific interactions7.
Due to these limitations, the results obtained from live cell imaging technologies should be confirmed by an independent approach based on a different principle for detecting protein interactions. Co-immunoprecipitation (Co-IP) or glutathione transferase (GST) pull-down assays represent such alternative methods that are commonly used to analyze protein-protein interactions in vitro. However, iIn these assays, however, the tested proteins must be readily soluble in the buffer that supportsused for the binding reaction. Therefore, specific interactions involving an insoluble protein cannot be assessed by these techniques.
 Here, we illustrate the protocol for the protein membrane overlay binding assay, which circumvents this difficulty. In this technique, interaction between soluble and insoluble proteins can be reliably tested because one of the proteins is immobilized on a membrane matrix. This method, in combination with in vivo experiments, such as BiFC, provides a reliable approach to investigate and characterize interactions faithfully between soluble and insoluble proteins. In this article, binding between Tobacco mosaic virus (TMV) movement protein (MP), which exerts multiple functions during viral cell-to-cell transport8-14, and a recently identified plant cellular interactor, tobacco ankyrin repeat-containing protein (ANK) 15, is demonstrated using this technique.

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

Testing protein-protein interaction is indispensable for dissection of protein functionality. Here, we introduce an in vitro protein-protein binding assay to probe a membrane-immobilized protein with a soluble protein. This assay provides a reliable method to test interaction between an insoluble protein and a protein in solution.

Validating interactions between different proteins is vital for investigation of their biological functions on the molecular level.  There are several ...

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 ...

Determination of Plasma Membrane Partitioning for Peripherally-associated Proteins

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein using the profiles of fluorescence intensity across the plasma membrane. Measured fluorescence profiles are fitted by a model for membrane and cytoplasm fluorescence distribution along a line applied perpendicularly to the cell periphery. This model is constructed from the fluorescence intensity values in reference cells expressing a fluorescently-tagged marker for cytoplasm and with FM 4-64-labeled plasma membrane. The method can be applied to various cell types and organisms; however, only plasma membranes of non-neighboring cells can be evaluated. This fast microscopy-based method is suitable for experiments, where subtle and dynamic changes of plasma membrane-associated markers are expected and need to be quantified, e.g., in the analysis of mutant versions of proteins, inhibitor treatments, and signal transduction observations. The method is implemented in a multi-platform R package that is coupled with an ImageJ macro that serves as a user-friendly interface.

Here, we present a protocol to perform a quantitative analysis of the level of plasma-membrane association for fluorescently-tagged peripherally-associated protein. The method is based on the computational decomposition of membrane and cytoplasmic component of signal observed in cells labeled with plasma membrane fluorescent marker.

This method provides a fast approach for the determination of plasma membrane partitioning of any fluorescently-tagged peripherally-associated protein ...

Fluorescence-Detection Size-Exclusion Chromatography: A Technique to Identify the Integrity of Fluorescent Membrane Proteins upon Detergent Solubilization

Source: Bird, L. E., et al. <a href="https://www.jove.com/v/52357">Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli</a>. J. Vis. Exp. (2015)
In this video, we demonstrate a fluorescence-detection size-exclusion chromatography technique to screen for green fluorescent protein (GFP)-fused membrane protein stability in Escherichia coli following detergent solubilization. Proteins exhibiting a symmetrical peak in the size-exclusion profile with little or no free GFP indicate purified proteins with minimum degradation and aggregation, which can subsequently be selected for structure-function analysis.

Chromatografia wykluczająca wielkość z detekcją fluorescencji: technika identyfikacji integralności fluorescencyjnych białek błonowych po rozpuszczaniu detergentu

Source: Bird, L. E., et al. Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli. J. Vis. Exp. (2015)
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Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

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Using a GFP-tagged TMEM184A Construct for Confirmation of Heparin Receptor Identity

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

Membrane-SPINE: A Biochemical Tool to Identify Protein-protein Interactions of Membrane Proteins In Vivo

Use of Recombinant Fusion Proteins in a Fluorescent Protease Assay Platform and Their In-gel Renaturation

Rapid Assessment of Membrane Protein Quality by Fluorescent Size Exclusion Chromatography

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Protein Membrane Overlay Assay: A Protocol to Test Interaction Between Soluble and Insoluble Proteins in vitro

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

Determination of Plasma Membrane Partitioning for Peripherally-associated Proteins

Chromatografia wykluczająca wielkość z detekcją fluorescencji: technika identyfikacji integralności fluorescencyjnych białek błonowych po rozpuszczaniu detergentu