The OPPF-UK is funded by the Medical Research Council, UK (grant MR/K018779/1) and the Membrane Protein Laboratory by the Wellcome Trust (grant 099165/Z/12/Z).

1. Construction of Expression Vectors
<ol>
	<li>Use PCR Infusion cloning to construct up to 96 expression plasmids in parallel using the vectors pOPINE-3C-GFP and pOPIN-N-GFP which add either cleavable C-terminal or N-terminal GFP-His6 fusions to the sequences13&#160;respectively. 
	Note: The choice of vector is dictated by the topology of the protein since the GFP needs to be expressed in the cytosol, thus for a protein with a cytosolic N-terminus and extracellular C-terminus we would use pOPIN-N-GFP and for an extracellular N-terminus and cytosolic C-terminus in we would use pOPINE-3C-GFP. Where both termini are cytosolic we would use pOPINE-3C-GFP unless there is functional reason not to tag the C-terminus.</li>
	<li>Amplify the DNA sequences encoding the membrane proteins with primers incorporating the following extensions shown: 
	pOPINE-3C-GFP (Forward primer &#8212; AGGAGATATACCATG; Reverse primer &#8212; CAGAACTTCCAGTTT) and pOPIN-N-GFP (Forward primer &#8212; AAGTTCTGTTTCAGGGCCCG; Reverse primer &#8212; ATGGTCTAGAAAGCTTTA).</li>
	<li>Analyze the PCR reaction by agarose gel electrophoresis. Once clean PCR products have been obtained, add 5 U DpnI to each well (make up in 5 &#181;l of 1x DpnI reaction buffer). Purify the PCR products using magnetic beads in a 96-well format. 
	Note: DpnI is added to remove the template vector; this step is not necessary if genomic DNA is used as template.</li>
	<li>Add 1 &#956;l (100 ng) of the appropriate linearized vector to each well of a PCR plate.</li>
	<li>Using a multichannel pipettor add x &#956;l of purified PCR insert to the appropriate wells of the PCR plate. Ensure that all products are in the range of 10-250 ng.</li>
	<li>Add (9-x) &#956;l of water to the well and transfer the whole 10 &#956;l to a tube containing the dry-down in-fusion reagent. Leave for 30 sec.</li>
	<li>Mix contents briefly by pipetting up and down. Transfer the reactions back to the original PCR plate and seal.</li>
	<li>Incubate for 30 min at 42 &#176;C in a thermocycler.</li>
	<li>Immediately transfer the reactions to ice as soon as the reaction is complete and add 40 &#956;l of TE (10 mM Tris, pH 8.0, 1 mM EDTA).</li>
	<li>Freeze at once or transform into competent cells. For transformation, use 3 &#181;l of the diluted reaction per 50 &#956;l aliquot of high competency (&#62; 5 x 109 transformants/&#181;g) competent E. coli cells.</li>
	<li>Add 1 ml of LB agar supplemented with 50 &#181;g/ml carbenicillin per well of a 24-well tissue culture plate (for cloning, prepare wells for 2 x number of constructs).</li>
	<li>Following outgrowth, Following outgrowth plate out 25 &#956;l and 25 &#956;l of a 1 in 5 dilution of the culture onto the agar containing 24-well tissue culture plates.</li>
	<li>Spread the cells by gently tilting the plate.</li>
	<li>Dry the plate with the lid off for 10-15 min at 37 &#176;C or in a flow hood at room temperature.</li>
	<li>Replace the lid and turn the plate upside down and incubate overnight at 37 &#176;C.</li>
	<li>Pick two colonies and mini-prep the vectors.</li>
	<li>PCR verify the constructs using the construct specific reverse primer and a vector specific forward primer (e.g. pOPIN_FOR GAC CGA AAT TAA TAC GAC TCA CTA TAG GG).</li>
</ol>
2. Transformation of E. coli Expression Strains
Note: Prior to performing this protocol, E. coli competent cells are made, aliquotted and stored frozen at -80 &#176;C as described previously13. Membrane protein expression is routinely screened in two strains, Lemo21(DE3) and C41(DE3)pLysS. To aid interpretation of the results it can be helpful to include a positive control, e.g. a plasmid expressing GFP or a membrane protein fused to GFP known to express and an empty vector negative control.
<ol>
	<li>Thaw the aliquotted E. coli on ice. Add 3 &#181;l of mini-prepped expression plasmid to each aliquot of competent cells.</li>
	<li>Incubate the mix on ice for 30 min.</li>
	<li>Heat-shock the cells for 30 sec at 42 &#176;C.</li>
	<li>Allow cells to recover on ice for 2 min and then add 300 &#181;l of power broth to each tube.</li>
	<li>Incubate for 1 hr in a 37 &#176;C static incubator.</li>
	<li>Prepare the LB Agar in a 24-well tissue culture plate, supplemented with 50 &#181;g/ml carbenicillin and 35 &#181;g/ml chloramphenicol. Add rhamnose to a final concentration of 0.25 mM to the wells containing Lemo21 (DE3). 
	Note: If the product is likely to be toxic the wells containing C41(DE3) pLysS can be supplemented with 1% (w/v) glucose.</li>
	<li>Transfer 30 &#181;l of cells from the transformation mix onto the LB Agar plates. Spread and dry the plate as described in 1.13-1.15.</li>
</ol>
3. Preparation of Overnight Starter Cultures
Note: Always prepare overnight cultures the day after transformation never from an old transformation plate.
<ol>
	<li>Add 0.7 ml power broth to each well of a 96-well deep-well plate supplemented with 50 &#181;g/ml carbenicillin and 35 &#181;g/ml chloramphenicol. Add rhamnose to the wells containing Lemo21 (DE3) to a final concentration of 0.25 mM. Optionally, supplement C41(DE3) pLysS with 1% (w/v) glucose; see above (2.6 note).</li>
	<li>Pick one colony from the overnight transformation plates into each well. Seal the block with a gas-permeable seal.</li>
	<li>Shake the block at 250 rpm in an incubator with a large throw (e.g. a 25 mm orbit) or at 600 rpm in an incubator with a small throw (e.g. a 4.3 mm orbit) at 37 &#176;C overnight.</li>
</ol>
4. Growth and Induction of Cultures
<ol>
	<li>Add 3 ml of power broth supplemented with 50 &#181;g/ml carbenicillin and 35 &#181;g/ml chloramphenicol to each well of 24-well deep-well plate. Add L-rhamnose (and glucose) as described in section 3.1. Use 3 ml of culture to allow harvesting in duplicate and to allow for some evaporation of water through the gas permeable seal.</li>
	<li>Set up the expression cultures by adding 150 &#181;l of Lemo21 (DE3) or C41(DE3) pLysS overnight cultures to each well of the 24-well deep-well blocks.</li>
	<li>Shake the blocks at 37 &#176;C until the average O.D. at 595 nm is ~0.5 through the plate.</li>
	<li>Induce expression by adding IPTG to the cultures to a final concentration of 1.0 mM IPTG.</li>
	<li>Grow the cultures overnight (18-20 hr) with shaking at 20 &#176;C.</li>
</ol>
5. Analysis of Expression by In-gel Fluorescence
<ol>
	<li>Harvest in duplicate. Transfer 1 ml of culture from each well into a square well, conical bottom, 96-well deep-well block. 
	Note: Harvest in duplicate in case of breakage of the block or to allow the testing of an alternative detergent if desired.</li>
	<li>Seal the blocks and harvest the cells by centrifugation at 6,000 x g for 10 min.</li>
	<li>Invert the block and decant the media. Tap the block gently onto a paper towel to drain remaining media.</li>
	<li>Seal the plates and store at -80 &#176;C for a minimum of 20 min. Optionally, leave the experiment at this point and the block processed when convenient.</li>
	<li>De-frost the pellets for 20 min at room temperature.</li>
	<li>Use either a multi-channel pipette or an orbital shaker (1,000 rpm for 10 min) at 4 &#176;C to resuspend&#160;the cells completely in 200 &#181;l of lysis buffer (50 mM NaH2PO4,&#160;300 mM NaCl 10 mM Imidazole 1% v/v Tween 20, adjust the pH to 8.0 using NaOH).</li>
	<li>Add 20 &#181;l of the DLPI solution (20 &#181;l DNaseI (4,000 units/ml), 20 mg lysozyme, and 20 &#181;l protease inhibitor cocktail in a final volume of 2 ml of water). Incubate for 10 min with shaking (~1,000 rpm) at 4 &#176;C.</li>
	<li>Add 25 &#181;l of 10% n-Dodecyl &#946;-D-maltoside (DDM).</li>
	<li>Incubate for 60 min with shaking (~1,000 rpm) at 4 &#176;C.</li>
	<li>Centrifuge the deep-well block at 6,000 x g for 30 min at 4 &#176;C.</li>
	<li>Transfer 10 &#956;l of the cleared lysate to a microtitre plate and add 10 &#956;l of SDS PAGE gel loading buffer (100 mM Tris, pH 6.8, 4% w/v SDS, 0.2% w/v Bromophenol blue, 10% v/v &#946;-mercaptoethanol, and 20% v/v glycerol). CAUTION! DO NOT BOIL THE SAMPLE.</li>
	<li>Load 10 &#956;l of sample from step 5.11/well onto SDS PAGE gel(s) and run at 100-120 V (constant voltage) in a cold room until the dye front reaches the bottom (2-2.5 hr).</li>
	<li>Place the gel onto an imager (e.g. with a blue-light filter to detect the GFP fusion proteins). The exposure time is chosen to ensure that the brightest bands are not saturated.</li>
</ol>
6. Scale-up of Cell Cultures
<ol>
	<li>Transform an aliquot of competent E. coli as described in section 2.</li>
	<li>Pick a colony into 10 ml of power broth (supplemented as described in section 3) and grow overnight at 37 &#176;C.</li>
	<li>Dilute 5 ml of the overnight culture into 500 ml power broth.</li>
	<li>Grow with shaking at 250 rpm at 37 &#176;C until the O.D. at 595 nm is ~0.5.</li>
	<li>Induce expression by adding IPTG to a final concentration of 1.0 mM.</li>
	<li>Grow overnight with shaking at 250 rpm at 20 &#176;C.</li>
	<li>Harvest cells by centrifugation at 5,000 x g for 15 min.</li>
	<li>Discard supernatant. Cell pellets can be stored at -80 &#176;C.</li>
</ol>
7. Preparation of Membranes
<ol>
	<li>Thaw cells (if necessary) at room temperature and resuspend in 1x PBS pH 7.5 at a volume of 5ml per gram of cell pellet; Increase volume as necessary until viscosity reduces to a runny consistency.&#160;Add&#160;MgCl2 to final concentration of 1 mM, DNAse I to final concentration of 10 &#956;g/ml and one pre-packaged protease inhibitor tablet per 20 g of cell pellet. Break open cells using a chilled cell disrupter at a pressure of 30 kpsi.</li>
	<li>Pellet the unbroken cells and debris at 30,000 g for 15 min at 4 &#176;C. Transfer supernatant to an ultra-centrifugation tube.</li>
	<li>Collect the total membrane fraction by spinning down the supernatant in an ultra-centrifuge at 200,000 x g for 1 hr at 4 &#176;C.</li>
	<li>Discard the supernatant.</li>
	<li>Re-suspend the membrane pellet in ice-cold 10 ml PBS pH 7.5 using a cell homogenizer. 1 ml resuspended membranes are required for each detergent. Optionally, at this stage, flash freeze the membrane suspension in liquid nitrogen and store at -80 &#176;C.</li>
</ol>
8. Solubilization and Analysis of the Membrane Fraction
<ol>
	<li>Thaw membrane fraction (if necessary), and for each detergent to be used for solubilization, aliquot 900 &#181;l of the membrane suspension into a 1.5 ml polyallomer micro-centrifuge tube.</li>
	<li>Add 100 &#181;l of each freshly prepared detergent (10% w/v solutions of DDM, DM, Cymal-6 and LDAO) to a final concentration of 1% to the specified 1.5 ml tube. For solubilization of eukaryotic membrane proteins cholesterol hemisuccinate (0.2% final concentration) can be added to the detergents.</li>
	<li>Incubate the mixtures at 4 &#176;C for 1 hr with mild agitation.</li>
	<li>Pellet the detergent insoluble fraction with a bench top ultra-centrifuge, ensuring tubes are at least half-full, at 150,000 g at 4 &#176;C for 45 min and retain supernatant. 
	Note: The effectiveness of detergent solubilization can be estimated using a fluorescent plate reader by measuring the GFP fluorescence of the solubilized membranes with the detergent insoluble fraction resuspended in the same volume of PBS.</li>
	<li>Analyze the mono-dispersity of the different detergent : membrane protein complexes using fluorescence-detected size exclusion chromatography (FSEC).</li>
	<li>For FSEC analysis, equilibrate a size exclusion column with a broad fractionation range, e.g. 5,000 to 5,000,000 in molecular weight, with running buffer (20 mM Tris pH 7.5, 150 mM NaCl, 0.03% DDM) at a flow rate of 0.3 ml/min. Use this buffer for all samples, i.e. it is not changed for each detergent tested.</li>
	<li>Inject 100 &#181;l of solubilized membranes onto the column at a flow rate of 0.3 ml/min. Monitor elution profile using fluorescence optics (excitation at 488 nm and emission at 512 nm). If an inline fluorescence monitor is not available, collect the fractions and read the fluorescence in a plate reader.</li>
	<li>Import elution volume and fluorescence intensity data into a spreadsheet program for graphical display.</li>
	<li>Analyse remaining solubilized membrane material by in-gel fluorescence as described in section 5.</li>
</ol>

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The authors have nothing to disclose.

In the protocol described in this article, ligation independent cloning into a GFP reporter vector is combined with fluorescence detection to rapidly screen the relative expression of multiple membrane proteins in E. coli. Vectors can be made in four working days with primary expression screening taking a further week. In-gel fluorescence of detergent lysates of whole cells is used to rank expression hits for further analysis in a secondary screen. The primary expression screen as presented does not require any specialist equipment apart from a shaker incubator suitable for growing cells in deep well blocks. All other liquid handling involving 96-well SBS plates can be carried out using multi-channel pipettes. The protocol can be readily modified to include other E.coli strains grown under different expression conditions (e.g. lower temperature). In the case of the LEMO21(DE3) titrating the level of rhamnose (from 0 to 2.0 mM) added to modulate the rate of transcription can increase the level of expression of some membrane proteins7 . Detergents other than DDM could be used for extraction in the primary screen though harsher detergents may solubilize proteins from inclusion bodies and therefore give false positives. Clearly, the more elaborate the initial screen becomes, the more time-consuming the experiment.
The secondary screen involves preparing a total membrane fraction from the expression hits identified in the primary screen. This is a critical step as it will confirm the localisation of the fusion proteins to the membrane of the E. coli cells. Subsequent extraction of the GFP fusion protein in different detergents is monitored by fluorescence-detected size exclusion chromatography of solubilized material to assess the mono-dispersity of the detergent-protein complexes. This is another critical step since it identifies the most appropriate detergent for solubilization of the membrane protein for subsequent downstream processing. However, experience shows that further optimization of the choice of detergent(s) may still be required during purification and crystallization. Evidence of uniform behavior in different detergents including ones with a relatively small micelle size (e.g. LDAO) appears to be good indicator of a propensity to form well diffracting crystals at least for prokaryotic membrane proteins14. In this respect the profile shown for the membrane protein in Figure 2B appears highly favorable.
The main advantage of using a GFP reporter is that it gives a rapid read-out of expression in un-purified samples. A disadvantage is that the GFP fusion protein has to be removed during the purification of the protein for crystallization. In the case of the vectors used here, a rhinovirus 3C protease site enables cleavage of the GFP. Alternatively, it is straightforward to re-clone candidate proteins, identified in the screen, into vectors which only add a polyhistidine or other affinity tag for subsequent purification. This assumes that fusion to GFP is not necessary for expression. The main alternative to using fusion to GFP for detection of membrane protein expression and solubilization is to add only a histidine tag. However this approach typically requires starting with a membrane fraction15 which for the evaluation of many variants would become very time-consuming.
In summary, the main purpose of the protocol described in this article is to enable a large number of membrane protein sequence variants to be rapidly evaluated in terms of expression and detergent solubilization and prioritized for deeper analysis.

It has been estimated that membrane proteins account for approximately 20-30% of the genes coded in all sequenced genomes 1, including the human genome 2. Given their critical role in many biological processes, for example transport of metabolites, energy generation and as drug targets, there is intense interest in determining their three dimensional structure. However compared to soluble proteins, membrane proteins are highly under-represented in the Protein Data Bank. In fact, of the 99,624 released structures in the PDB (<a href="http://www.rcsb.org/pdb/">http://www.rcsb.org/pdb/</a>) only 471 (<a href="http://blanco.biomol.uci.edu/mpstruc/">http://blanco.biomol.uci.edu/mpstruc/</a>) are classified as membrane proteins. This reflects the technical difficulties of working with membrane proteins which are naturally expressed at relatively low levels; rhodopsin is one of a few exceptions3,4. Over-expression of recombinant versions in heterologous cells often results in misfolding and poor targeting to the membrane which limits the overall level of production. Selecting the best detergent for extraction and solubilization of a membrane protein once expressed makes subsequent purification difficult and requires careful optimization.
Escherichia coli remains the most commonly used expression host for membrane proteins accounting for 72% (<a href="http://blanco.biomol.uci.edu/mpstruc/">http://blanco.biomol.uci.edu/mpstruc/</a>) of structures solved to date which are almost exclusively from bacterial sources. Specific E. coli strains, C41(DE3) and C43(DE3), have been isolated which do not lead to the over-expression of membrane proteins and hence avoid the effects of toxicity observed for other BL21 strains5,6. Tuning the level of recombinant membrane protein expression appears to be critical for optimizing the level of production6,7.
In many cases successful production of membrane proteins for structure determination has required the evaluation of multiple versions including amino- and carboxy-terminal deletions, multiple orthologues to exploit natural sequence variation and different fusion proteins6-8. Therefore, a method for the expression screening of multiple membrane proteins in parallel is essential. One commonly used approach is to fuse the protein to green fluorescent protein (GFP) enabling expression to be tracked without the need to purify the protein. In this way, information about expression level, stability and behavior in different detergents can be assessed with small amounts of un-purified material9-12.
In this article we describe a streamlined protocol for cloning and expression screening of membrane proteins in E. coli using fusion to GFP as a reporter of production and subsequent characterization of detergent: protein complexes (Figure 1).

<table><tbody><tr><td>pOPIN-N-GFP</td><td>addgene (www.addgene.org)</td><td></td><td></td></tr><tr><td>pOPINE-3C-GFP</td><td>addgene (www.addgene.org)</td><td></td><td></td></tr><tr><td>C41(DE3)pLysS</td><td>Lucigen (http://lucigen.com/)</td><td>60444-1</td><td></td></tr><tr><td>LEMO21(DE3)</td><td>New England Biolabs</td><td>C2528H</td><td></td></tr><tr><td>In-Fusion HD EcoDry Cloning System, 96 Rxns</td><td>Clontech/Takara</td><td>639688</td><td></td></tr><tr><td>Power broth</td><td>Molecular Dimensions (http://www.moleculardimensions.com/)</td><td>MD12-106-1</td><td></td></tr><tr><td>Protease Inhibitor tablets</td><td>Roche</td><td>11697498001</td><td></td></tr><tr><td>cOmplete, Mini, EDTA-Free; Protease Inhibitor Cocktail&amp;nbsp;</td><td>Roche</td><td>11836170001</td><td></td></tr><tr><td>L-Rhamnose monohydrate</td><td>Sigma-Aldrich</td><td>83650</td><td></td></tr><tr><td>Protease Inhibitor Cocktail&amp;nbsp;</td><td>Sigma-Aldrich</td><td>P8849</td><td></td></tr><tr><td>DNAse 1</td><td>Sigma-Aldrich</td><td>DN25</td><td></td></tr><tr><td>Lysozyme</td><td>Sigma-Aldrich</td><td>L7651</td><td></td></tr><tr><td>Cholesterol hemisuccinate</td><td>Sigma-Aldrich</td><td>C6512</td><td></td></tr><tr><td>CYMAL-6 (6-Cyclohexyl-1-Hexyl-&amp;beta;-D-Maltoside)</td><td>Anatrace</td><td>C326</td><td></td></tr><tr><td>DDM (n-Dodecyl &amp;beta;-maltoside)</td><td>Generon</td><td>D97002</td><td></td></tr><tr><td>DM (n-Decyl &amp;beta;-maltoside)</td><td>Generon</td><td>D99003</td><td></td></tr><tr><td>LDAO (&lt;em&gt;N,N&lt;/em&gt;-Dimethyl-n-dodecylamine &lt;em&gt;N&lt;/em&gt;-oxide)</td><td>Generon</td><td>D71111</td><td></td></tr><tr><td>E1 ClipTip Eq384 8&amp;nbsp;channel 1-30 &amp;micro;l</td><td>Thermo Scientific&amp;nbsp;</td><td>4672030</td><td></td></tr><tr><td>Rainin Pipette Pipet-Lite XLS 6ch 100-1,200 &amp;micro;l LTS Spacer</td><td>Anachem</td><td>LA6-300XLS</td><td></td></tr><tr><td>Rainin Pipette Light XLS+ 8ch 2-20 &amp;micro;l LTS</td><td>Anachem</td><td>L8-20XLSPLUS</td><td></td></tr><tr><td>Pipette Pipet-Lite XLS 8ch 100-1,200 &amp;micro;l LTS Tip</td><td>Anachem</td><td>L8-1200XLS</td><td></td></tr><tr><td>24 well V bottom deep well blocks</td><td>Porvair sciences</td><td>360115</td><td></td></tr><tr><td>BenchMark Fluorescent Protein Standard</td><td>Life Technologies</td><td>LC5928</td><td></td></tr><tr><td>NuPAGE Novex 10% Bis-Tris Midi Protein Gels, 26 well</td><td>Life Technologies</td><td>WG1203BOX</td><td></td></tr><tr><td>XCell4 SureLock Midi-Cell</td><td>Life Technologies</td><td>WR0100</td><td></td></tr><tr><td>Vibramax 100</td><td>Heidolph</td><td>544-21200-00</td><td></td></tr><tr><td>Tension rollers attachment</td><td>Heidolph</td><td>549-81000-00</td><td></td></tr><tr><td>ptima L-100 Ultracentrifuge running 45-Ti rotor</td><td>Beckman Coulter</td><td>393253</td><td></td></tr><tr><td>Optima Max-XP ultracentrifuge running TLA-55 rotor</td><td>Beckman Coulter</td><td>393315</td><td></td></tr><tr><td>Floor standing centrifuge, Avanti J-26S XPI running JA-17 and JS-5.3 rotors&amp;nbsp;</td><td>Beckman Coulter</td><td>B14535</td><td></td></tr><tr><td>Prominence UFLC with RF-20A Fluorescence detector</td><td>Shimadzu</td><td></td><td></td></tr><tr><td>Sepharose 6 10/300 GL size exclusion column</td><td>GE Healthcare</td><td>17-5172-01&amp;nbsp;</td><td></td></tr><tr><td>SSI5R-HS SHEL LAB Floor Model Shaking Incubator, 5 Cu.Ft. (144&amp;nbsp;L), 30-850 rpm</td><td>Shel Lab&amp;nbsp;</td><td>SSI5R-HS</td><td></td></tr><tr><td>Innova44R incubator</td><td>New Brunswick /Eppendorf</td><td>Innova44R</td><td></td></tr><tr><td>TS Series 0.75 kW Disrupter 230 V 50 Hz 1&amp;nbsp;PH (40 kpsi)</td><td>Constant systems</td><td>PL083016</td><td></td></tr><tr><td>L-Rhamnose monohydrate</td><td>Sigma</td><td>83650</td><td></td></tr></tbody></table>

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.

The SEDS family of proteins (shape, elongation, division, and sporulation) is essential for bacterial cell division. These integral proteins typically have ten transmembrane domains with both amino and carboxy termini inside the cell. To exemplify the protocol described above, 47 SEDs orthologues were cloned into the vector pOPINE-3C-eGFP. A small scale expression screen of the constructs was carried out in two E .coli strains, Lemo21(DE3) grown in the presence of 0.25 mM rhamnose to block leaky expression and C41(DE3) pLysS, which are routinely used for the expression of membrane proteins5-8.
Detergent solubilized lysates of induced cultures were analyzed by SDS-polyacrylamide electrophoresis. Using in-gel fluorescence to visualize expressed GFP fusion proteins showed that between the two strains 38 out of the 47 SEDs constructs were produced. Interestingly there were differences between the two expression strains. Of the 38 expressed constructs, only 18 were produced in both strains, 14 only expressed in Lemo21(DE3) and 6 only in C41(DE3) pLysS. Overall, there was a higher success rate for the Lemo21(DE3) compared to the C41(DE3) pLysS (38 vs 24). However the observation that some proteins appeared to be strain specific means that screening in both is useful.
Representative data for 24 of the 47 SEDs constructs are shown in Figure 1. The intensity of the bands gives an indication of the level of expression, e.g. the fusion protein in well C4 in Figure 1A and 1B is expressed well in both strains however the additional lower molecular weight bands suggest that the protein is partially degraded. In many lanes there is a band which corresponds in size to GFP alone. A qualitative assessment of the integrity of the protein can be carried out by looking at the intensity of fusion protein relative to the free GFP; e.g. G4 and H4 are completely broken down to free GFP in C41(DE3) pLysS (Figure 1B) whereas in the Lemo21(DE3) there is some intact protein (Figure 1A). For 14 out of the 38 expressed proteins the intensity of the free GFP band is similar to that of the fusion protein, for 21 the intensity of the free GFP band is greater than that of the fusion protein and a minority of the fusion proteins (3/38 expressed) e.g. F6 and G6 in Figure 1A have relatively little free GFP compared to the fusion protein.
Two of the constructs that expressed well in the primary screen B5 (high intensity bands for free GFP and fusion proteins) and G6 (little free GFP) were expressed and a total membrane fraction prepared from 500 ml cell culture. Resuspended membranes were divided into aliquots and analyzed by fluorescence-detected size exclusion chromatography (FSEC). The FSEC profiles are presented in Figure 2A and 2B respectively and show that the two SEDs proteins behave differently in the four detergents tested. In one case (Figure 2A: sample B5) the protein only gave a symmetrical peak with little aggregation in DDM which would therefore be the detergent of choice for subsequent purification. By contrast the other fusion protein (Figure 2B: sample G6) gave a similar profile in all detergents tested.
<img alt="Figure 1" src="/files/ftp_upload/52357/52357fig1highres.jpg" width="700px" /> 
Figure 1: Diagram of workflow for screening membrane protein expression in E. coli using a GFP reporter.
<img alt="Figure 2" src="/files/ftp_upload/52357/52357fig2highres.jpg" /> 
Figure 2: Primary screen of membrane protein expression using in-gel fluorescence. Detergent lysates of E. coli cells were analyzed by SDS-PAGE and gels imaged using Blue Epi illumination and a 530/28 filter. The results are shown for 24 constructs (labeled A4-H6 based on their position in a 96-well plate). The positions of the bands for free GFP and GFP fusion proteins are indicated. (A) Proteins expressed in Lemo21(DE3). (B) Proteins expressed in C41(DE3) pLysS.
<img alt="Figure 3" src="/files/ftp_upload/52357/52357fig3highres.jpg" /> 
Figure 3: Secondary screen of membrane protein expression using Fluorescence-detected size exclusion chromatography (FSEC). Total membrane fractions were extracted in four different detergents and the solubilized membrane proteins analyzed by size exclusion chromatography using a FPLC system coupled to a fluorescence detector. FSEC profiles for (A) membrane protein B5 and (B) membrane protein G6. The position of the peaks corresponding to aggregated protein, solubilized GFP fusion protein and free GFP are indicated. The detergents tested are indicated below the profiles.

Watch this Scientific Journal Video about Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli at JoVE.com

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.

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 ...

green-fluorescent-protein-based-expression-screening-membrane

Research

JoVE Journal

Biology

32.7K Views.  Research Complex at Harwell.  Een gestroomlijnde benadering screening op de expressie van recombinant membraaneiwitten in Escherichia coli gebaseerd op fusie met groen fluorescent proteïne gepresenteerd. 

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

Bacterial Inner-membrane Display for Screening a Library of Antibody Fragments

Antibodies engineered for intracellular function must not only have affinity for their target antigen, but must also be soluble and correctly folded in the cytoplasm. Commonly used methods for the display and screening of recombinant antibody libraries do not incorporate intracellular protein folding quality control, and, thus, the antigen-binding capability and cytoplasmic folding and solubility of antibodies engineered using these methods often must be engineered separately. Here, we describe a protocol to screen a recombinant library of single-chain variable fragment (scFv) antibodies for antigen-binding and proper cytoplasmic folding simultaneously. The method harnesses the intrinsic intracellular folding quality control mechanism of the Escherichia coli twin-arginine translocation (Tat) pathway to display an scFv library on the E. coli inner membrane. The Tat pathway ensures that only soluble, well-folded proteins are transported out of the cytoplasm and displayed on the inner membrane, thereby eliminating poorly folded scFvs prior to interrogation for antigen-binding. Following removal of the outer membrane, the scFvs displayed on the inner membrane are panned against a target antigen immobilized on magnetic beads to isolate scFvs that bind to the target antigen. An enzyme-linked immunosorbent assay (ELISA)-based secondary screen is used to identify the most promising scFvs for additional characterization. Antigen-binding and cytoplasmic solubility can be improved with subsequent rounds of mutagenesis and screening to engineer antibodies with high affinity and high cytoplasmic solubility for intracellular applications.

We provide a method to simultaneously screen a library of antibody fragments for binding affinity and cytoplasmic solubility by using the Escherichia coli twin-arginine translocation pathway, which has an inherent quality control mechanism for intracellular protein folding, to display the antibody fragments on the inner membrane.

Antibodies engineered for intracellular function must not only have affinity for their target antigen, but must also be soluble and correctly folded in ...

Biochemistry

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

Escherichia coli-Based Cell-Free Protein Synthesis: Protocols for a robust, flexible, and accessible platform technology

Over the last 50 years, Cell-Free Protein Synthesis (CFPS) has emerged as a powerful technology to harness the transcriptional and translational capacity of cells within a test tube. By obviating the need to maintain the viability of the cell, and by eliminating the cellular barrier, CFPS has been foundational to emerging applications in biomanufacturing of traditionally challenging proteins, as well as applications in rapid prototyping for metabolic engineering, and functional genomics. Our methods for implementing an E. coli-based CFPS platform allow new users to access many of these applications. Here, we describe methods to prepare extract through the use of enriched media, baffled flasks, and a reproducible method of tunable sonication-based cell lysis. This extract can then be used for protein expression capable of producing 900 &#181;g/mL or more of super folder green fluorescent protein (sfGFP) in just 5 h from experimental setup to data analysis, given that appropriate reagent stocks have been prepared beforehand. The estimated startup cost of obtaining reagents is $4,500 which will sustain thousands of reactions at an estimated cost of $0.021 per &#181;g of protein produced or $0.019 per &#181;L of reaction. Additionally, the protein expression methods mirror the ease of the reaction setup seen in commercially available systems due to optimization of reagent pre-mixes, at a fraction of the cost. In order to enable the user to leverage the flexible nature of the CFPS platform for broad applications, we have identified a variety of aspects of the platform that can be tuned and optimized depending on the resources available and the protein expression outcomes desired.

Escherichia coli-Based Cell-Free Protein Synthesis: Protocols for a robust, flexible, and accessible platform technology

This protocol details the steps, costs, and equipment necessary to generate E. coli-based cell extracts and implement in vitro protein synthesis reactions within 4 days or less. To leverage the flexible nature of this platform for broad applications, we discuss reaction conditions that can be adapted and optimized.

Over the last 50 years, Cell-Free Protein Synthesis (CFPS) has emerged as a powerful technology to harness the transcriptional and translational capacity ...

Chemistry

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 ...

Engineering 'Golden' Fluorescence by Selective Pressure Incorporation of Non-canonical Amino Acids and Protein Analysis by Mass Spectrometry and Fluorescence

Fluorescent proteins are fundamental tools for the life sciences, in particular for fluorescence microscopy of living cells. While wild-type and engineered variants of the green fluorescent protein from Aequorea victoria (avGFP) as well as homologs from other species already cover large parts of the optical spectrum, a spectral gap remains in the near-infrared region, for which avGFP-based fluorophores are not available. Red-shifted fluorescent protein (FP) variants would substantially expand the toolkit for spectral unmixing of multiple molecular species, but the naturally occurring red-shifted FPs derived from corals or sea anemones have lower fluorescence quantum yield and inferior photo-stability compared to the avGFP variants. Further manipulation and possible expansion of the chromophore&#39;s conjugated system towards the far-red spectral region is also limited by the repertoire of 20 canonical amino acids prescribed by the genetic code. To overcome these limitations, synthetic biology can achieve further spectral red-shifting via insertion of non-canonical amino acids into the chromophore triad. We describe the application of SPI to engineer avGFP variants with novel spectral properties. Protein expression is performed in a tryptophan-auxotrophic E. coli strain and by supplementing growth media with suitable indole precursors. Inside the cells, these precursors are converted to the corresponding tryptophan analogs and incorporated into proteins by the ribosomal machinery in response to UGG codons. The replacement of Trp-66 in the enhanced &#34;cyan&#34; variant of avGFP (ECFP) by an electron-donating 4-aminotryptophan results in GdFP featuring a 108 nm Stokes shift and a strongly red-shifted emission maximum (574 nm), while being thermodynamically more stable than its predecessor ECFP. Residue-specific incorporation of the non-canonical amino acid is analyzed by mass spectrometry. The spectroscopic properties of GdFP are characterized by time-resolved fluorescence spectroscopy as one of the valuable applications of genetically encoded FPs in life sciences.

Synthetic biology enables the engineering of proteins with unprecedented properties using the co-translational insertion of non-canonical amino acids. Here, we presented how a spectrally red-shifted variant of a GFP-type fluorophore with novel fluorescence spectroscopic properties, termed &#34;gold&#34; fluorescent protein (GdFP), is produced in E. coli via selective pressure incorporation (SPI).

Fluorescent proteins are fundamental tools for the life sciences, in particular for fluorescence microscopy of living cells. While wild-type and ...

Bioengineering

High-throughput Screening and Biosensing with Fluorescent C. elegans Strains

High-throughput screening (HTS) is a powerful approach for identifying chemical modulators of biological processes. However, many compounds identified in screens using cell culture models are often found to be toxic or pharmacologically inactive in vivo1-2. Screening in whole animal models can help avoid these pitfalls and streamline the path to drug development.
C. elegans is a multicellular model organism well suited for HTS. It is small (&lt;1 mm) and can be economically cultured and dispensed in liquids. C. elegans is also one of the most experimentally tractable animal models permitting rapid and detailed identification of drug mode-of-action3.
We describe a protocol for culturing and dispensing fluorescent strains of C. elegans for high-throughput screening of chemical libraries or detection of environmental contaminants that alter the expression of a specific gene. Large numbers of developmentally synchronized worms are grown in liquid culture, harvested, washed, and suspended at a defined density. Worms are then added to black, flat-bottomed 384-well plates using a peristaltic liquid dispenser. Small molecules from a chemical library or test samples (e.g., water, food, or soil) can be added to wells with worms. In vivo, real-time fluorescence intensity is measured with a fluorescence microplate reader. This method can be adapted to any inducible gene in C. elegans for which a suitable reporter is available. Many inducible stress and developmental transcriptional pathways are well defined in C. elegans and GFP transgenic reporter strains already exist for many of them4. When combined with the appropriate transgenic reporters, our method can be used to screen for pathway modulators or to develop robust biosensor assays for environmental contaminants. 
We demonstrate our C. elegans culture and dispensing protocol with an HTS assay we developed to monitor the C. elegans cap &#x2018;n&#x2019; collar transcription factor SKN-1. SKN-1 and its mammalian homologue Nrf2 activate cytoprotective genes during oxidative and xenobiotic stress5-10. Nrf2 protects mammals from numerous age-related disorders such as cancer, neurodegeneration, and chronic inflammation and has become a major chemotherapeutic target11-13.Our assay is based on a GFP transgenic reporter for the SKN-1 target gene gst-414, which encodes a glutathione-s transferase6. The gst-4 reporter is also a biosensor for xenobiotic and oxidative chemicals that activate SKN-1 and can be used to detect low levels of contaminants such as acrylamide and methyl-mercury15-16.

High-throughput Screening and Biosensing with Fluorescent C. elegans Strains

A procedure for liquid-based culturing and dispensing of C. elegans strains expressing fluorescent reporter proteins is described that does not require expensive sorting equipment. This approach can be applied to numerous inducible C. elegans genes for drug discovery or biosensing of contaminants.

High-throughput screening (HTS) is a powerful approach for identifying chemical modulators of biological processes.  However, many compounds identified in ...

Using Enhanced Green Fluorescence Protein-expressing Escherichia Coli to Assess Mouse Peritoneal Macrophage Phagocytosis

This manuscript describes a simple and reproducible method to perform a phagocytosis assay. The first part of this method involves building a pET-SUMO-EGFP vector (SUMO = small ubiquitin-like modifier) and expressing enhanced green fluorescence protein (EGFP) in Escherichia coli (BL21DE). EGFP-expressing E. coli is coincubated with macrophages for 1 h at 37 &#176;C; the negative control group is incubated on ice for the same amount of time. Then, the macrophages are ready for assessment. The advantages of this technique include its simple and straightforward steps, and phagocytosis can be measured by both flow cytometer and fluorescence microscope. The EGFP-expressing E. coli are stable and display a strong fluorescence signal even after the macrophages are fixed with paraformaldehyde. This method is not only suitable for the assessment of macrophage cell lines or primary macrophages in vitro but also suitable for the evaluation of granulocyte and monocyte phagocytosis in peripheral blood mononuclear cells. The results show that the phagocytic capability of peritoneal macrophages from young (eight-week-old) mice is higher than that of macrophages from aged (16-month-old) mice. In summary, this method measures macrophage phagocytosis and is suitable for studying the innate immune system function.

Using Enhanced Green Fluorescence Protein-expressing Escherichia Coli to Assess Mouse Peritoneal Macrophage Phagocytosis

Here, we present a protocol to assess mouse peritoneal macrophage phagocytosis using enhanced green fluorescence protein-expressing Escherichia coli.

This manuscript describes a simple and reproducible method to perform a phagocytosis assay. The first part of this method involves building a ...

Immunology and Infection

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

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.

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.

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

Residue-specific Incorporation of Noncanonical Amino Acids into Model Proteins Using an Escherichia coli Cell-free Transcription-translation System

The canonical set of amino acids leads to an exceptionally wide range of protein functionality. Nevertheless, the set of residues still imposes limitations on potential protein applications. The incorporation of noncanonical amino acids can enlarge this scope. There are two complementary approaches for the incorporation of noncanonical amino acids. For site-specific incorporation, in addition to the endogenous canonical translational machineries, an orthogonal aminoacyl-tRNA-synthetase-tRNA pair must be provided that does not interact with the canonical ones. Consequently, a codon that is not assigned to a canonical amino acid, usually a stop codon, is also required. This genetic code expansion enables the incorporation of a noncanonical amino acid at a single, given site within the protein. The here presented work describes residue-specific incorporation where the genetic code is reassigned within the endogenous translational system. The translation machinery accepts the noncanonical amino acid as a surrogate to incorporate it at canonically prescribed locations, i.e., all occurrences of a canonical amino acid in the protein are replaced by the noncanonical one. The incorporation of noncanonical amino acids can change the protein structure, causing considerably modified physical and chemical properties. Noncanonical amino acid analogs often act as cell growth inhibitors for expression hosts since they modify endogenous proteins, limiting in vivo protein production. In vivo incorporation of toxic noncanonical amino acids into proteins remains particularly challenging. Here, a cell-free approach for a complete replacement of L-arginine by the noncanonical amino acid L-canavanine is presented. It circumvents the inherent difficulties of in vivo expression. Additionally, a protocol to prepare target proteins for mass spectral analysis is included. It is shown that L-lysine can be replaced by L-hydroxy-lysine, albeit with lower efficiency. In principle, any noncanonical amino acid analog can be incorporated using the presented method as long as the endogenous in vitro translation system recognizes it.

Residue-specific Incorporation of Noncanonical Amino Acids into Model Proteins Using an Escherichia coli Cell-free Transcription-translation System

An easy-to-use, cell-free expression protocol for the residue-specific incorporation of noncanonical amino acid analogs into proteins, including downstream analysis, is presented for medical, pharmaceutic, structural and functional studies.

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

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