Support for this project came from the BGSU Graduate College, and the BGSU Office of Sponsored Programs and Research.

1. Generation of the E. coli Double Knock Out Mutant with P1 Transduction
<ol>
	<li>Obtain the E. coli single-knockout mutant strains &#916;PotE and &#916;CadB from the E. coli Genetic Stock Center (<a href="http://cgsc.biology.yale.edu" target="_blank">http://cgsc.biology.yale.edu</a>). 
	NOTE: The &#916;PotE strain is kanamycin resistant26 and the &#916;CadB strain is tetracycline resistant27.</li>
	<li>Construct the PotE/CadB double knockout (DKO) strain using the P1 transduction protocol28.
	<ol>
		<li>Lysate preparation
		<ol>
			<li>Dilute an overnight culture of &#916;PotE in fresh LB (1:100) supplemented with 10-25 mM MgCl2, 5 mM CaCl2, and 0.1-0.2% glucose.</li>
			<li>Grow at 37 &#176;C for 1-2 h.</li>
			<li>Add 40 &#181;L of P1 phage lysate to the culture, and continue growing at 37 &#176;C. Monitor for 1-3 h until the culture has lysed 
			NOTE: When the culture is lysed, cellular debris will be visible in the tube and the culture will have a significant loss of turbidity.</li>
			<li>Add several drops of chloroform (50-100 &#181;L) to the lysate and vortex. Centrifuge at 12,000 x g for 2 min to remove cellular debris and transfer the supernatant to a fresh tube.</li>
		</ol>
		</li>
		<li>Transduction
		<ol>
			<li>Grow &#916;CadB strain overnight in LB medium.</li>
			<li>Harvest the cells by centrifugation at 4,000 x g for 2 min and resuspend in 1/5-1/3 the harvested culture volume in fresh LB supplemented with 100 mM MgSO4 and 5 mM CaCl2.</li>
			<li>Add the &#916;CadB cell suspension to the tube with phage from step 1.2.1.4 and mix rapidly.</li>
			<li>Add 200 &#181;L of 1 M Na-Citrate (pH5.5), then add 1 mL of LB. Incubate at 37 &#176;C for 1 h to allow the expression of the antibiotic resistant marker.</li>
			<li>Spin cells at 4000 x g for 5 min.</li>
			<li>Resuspend in 100 &#181;L LB supplemeted with 100 mM Na-Citrate (pH 5.5) and vortex well to disperse cells.</li>
			<li>Select the recombinant strains on LB agar plates with 50 &#181;g/mL kanamycin and 10 &#181;g/mL tetracycline and then confirm by Polymerase Chain Reaction (PCR) with appropriate primers26,27.</li>
			<li>Verify the PCR fragments by sequencing.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Expression of the Target Gene (AtBAT1) in E. coli Mutant
<ol>
	<li>Amplify the full-length sequence of the target gene by PCR and insert into the Gateway entry vector pENTR/D-TOPO29. 
	NOTE: In this case ATBAT1.1 was amplified using forward primer 5&#39;CGGCGATCAATCCTTTGTT and reverse primer 5&#39;GCTAAGAATGTTGGAGATGG, an initial denaturation at 98 &#176;C for 2 min and then 30 cycles of denaturation at 98 &#176;C for 0.1 min, annealing at 59 &#176;C for 0.3 min, extension at 72 &#176;C for 0.40 min and final extension at 72 &#176;C for 10 min. The PCR product was purified using a PCR cleanup kit, quantified using a spectrometer. Insertion of the PCR product into the Gateway vector was done following the manufactrurers directions.
	<ol>
		<li>Transform the entry vector into competent Top10 E. coli cells by heatshock and select for successful recombinants on LB media with 50 &#181;g/mL kanamycin following standard molecular cloning protocols30.</li>
		<li>Extract plasmid DNA using a plasmid extraction kit, following the manufacturer&#39;s directions, and sequence to confirm the correct insertion.</li>
	</ol>
	</li>
	<li>Transfer the target gene into the E. coli destination vector, pBAD-DEST49 by Gateway LR cloning to create an expression vector29.
	<ol>
		<li>Transform the expression vector into competent Top10 E. coli cells by heatshock for 30 s and select successful recombinants on LB media with 100 &#181;g/mL ampicillin30.</li>
		<li>Extract plasmid DNA30 and sequence to confirm the correct insertion.</li>
	</ol>
	</li>
	<li>Transform expression vector into E. coli &#916;potE740(del)::kan, &#916;cadB2231::Tn10 and select on LB plates with ampicillin, kanamycin, and tetracycline30.</li>
	<li>In order to determine the optimal concentration of arabinose for induction, express the target gene under the control of the araBAD promoter in LB media with gradient concentrations of arabinose as described29.</li>
	<li>Collect the cell pellets and assess protein expression by SDS-PAGE and visualization of protein bands as described in the product manual29. 
	NOTE: The E. coli &#916;potE740(del)::kan, &#916;cadB2231::Tn10 was used as the control sample without BAT1 expression. E.coli double knock out mutant cells with the empty pBAD-DEST49 vector was not used as a control due to the presence of the ccdB gene in the vector.</li>
</ol>
3. Generation of Inside-out Membrane Vesicles
<ol>
	<li>Culture the E. coli mutant cells expressing the target protein by growing a single colony overnight in 5 mL of polyamine-free media (2% glycerol, 6 g of K2HPO4, 3 g of KH2PO4, 0.5 g of NaCl, NH4Cl, 50 mg of thiamine, 0.79 g yeast complete synthetic media adenine hemisulfate (10 mg/L), L-Arginine (50 mg/L), L-Aspartic acid (80 mg/L), L-Histidine hydrochloride monohydrate (20 mg/L), L-Leucine (100 mg/L), L-Lysine hydrochloride (50 mg/L), L-Methionine (20 mg/L), L-Phenylalanine (50 mg/L), L-Threonine (100 mg/L), L-Tryptophan (50 mg/L), L-Tyrosine (50 mg/L), L-Valine (140 mg/L), Uracil (20 mg/L)). Transfer this culture to 500 mL of polyamine-free media supplemented with 0.0002% arabinose and grow until the cell culture reaches an OD600 of 0.6-0.8 (usually ~5 h).</li>
	<li>Collect the cell pellet by centrifugation at 2,000 x g for 15 min at 4 &#176;C. Resuspend the pellet and wash three times with 0.1 M potassium phosphate buffer, pH 6.6, containing 10 mM EDTA (Buffer 1). The final volume of cells in Buffer 1 after the third spin should be 10 mL.</li>
	<li>Generate inside-out membrane vesicles by French Press treatment (10,000 p.s.i.) of the intact cells using a 35 mL French pressure Cell.</li>
	<li>Prior to loading the sample into the standard French pressure cell, lubricate the piston and O-rings to make it easier to insert into the cell. 
	NOTE: These instructions are specific for the use of the standard pressure cell31.
	<ol>
		<li>Carefully screw the closure plug into the lower end of the cell. Ensure that the cell is right-side up based on the lettering on the side of the cell. Insert the piston into the cell and move it into the cell, until the max fill line on the piston reach the top of the piston. Flip the unit over and place on the provided stand between the three posts.</li>
		<li>Pour the cell suspension into the body of the cell. 
		NOTE: We have been using only 10 mL, so there will be plenty of room left in the French pressure cell chamber, which has a capacity of 35 mL.</li>
		<li>Mount the standard pressure cell on the French pressure cell unit.
		<ol>
			<li>First check that the power is off. On the left side of the panel is the Ratio Selector switch. It has three positions: Down at the end of the run, Low for using a smaller pressure cell, and High for use with the standard French pressure cell. If the French press was previously used with the smaller cell the piston may not be fully retracted. Set this switch to Down.</li>
		</ol>
		</li>
		<li>Turn the machine on with the switch on the RHS at the back of the unit and turn the Pause-Run switch to the Run position. This will result in the piston being fully retracted. Move the switch back to Pause, and shut the machine off.</li>
		<li>Loosen the thumb screws and move the safety clamp that spans to two stainless steel columns to the side. It will swing out to the right. With one hand holding the base of the closure plug in place, pick the French pressure cell up with both hands, and rotate it 180&#176;, so the piston is now in the upright position. Set it onto the platform, and move the safety clamp back in place so that this clamp holds the French pressure cell in position.</li>
		<li>Note the Flow Valve assembly on the closure plug needs to be accessible to the operater and positioned so that the valve can be turned freely. Open the the flow valve assembly with a few counter clockwise turns. Position the the arms of the piston so that they are oriented in a two o&#39;clock and eight o&#39;clock position. Tighten the thumbs screws so that the standard pressure cell is held in place.</li>
		<li>Insert the Sample outlet tube into the closure plug, and connect a short piece of flexible tubing so that the broken cell debris can be collected in a falcon tube. We usually use an 300 mL ice-filled beaker to hold the falcon tube upright, and to keep the cell debris chilled.</li>
		<li>Make sure the Pause-Run switch is set to Pause, and the valve assembly is in the open position. Turn the machine back to on, adust the Ratio Selector switch to High, and turn the Pressure Increase Valve on the front panel to the right about a half turn.</li>
		<li>The piston will begin to rise from the base to displace the air in chamber. Direct the air coming out of the Tygon tubing attached to the sample outlet tube to the tube in the beaker.</li>
		<li>When the first drops of liquid come out, close the the flow valve assembly by turning it clockwise. This creates a metal to metal seal, with the pressure cell, so do not overtighten.</li>
		<li>The front of the French Press has a chart on the front panel to generate the proper pressure on the biological cells inside the pressure cell. In this case, increase the pressure by turning the Pressure Increase Valve clockwise, until the Gauge reaches 640 psi. This will create an internal pressure of 10,000 psi.</li>
		<li>Once the cell has reached target pressure open the flow valve assembly slightly by turning it counter clockwise. Adjust the the opening of the valve to allow a flow rate of only about 10 drops per min.</li>
		<li>When the stop line on the piston body reaches the top of the flow cell. Switch the Pause-Run Switch to Pause. This is critical because having the piston inserted farther into the cell will damage the unit. Open the flow valve assembly and collect the remaining drops.</li>
		<li>Turn Ratio Selector switch to Down, and then set the Pause Run switch to Run. When the bottom plate is fully retracted set the Run switch to Pause, and turn the machine off.</li>
		<li>Remove the French pressure cell from the French press, and disassemble for cleaning. Store all parts dry with a light covering of glycerol. Remove and replace any gaskets or seals with signs of wear.</li>
	</ol>
	</li>
	<li>Remove unbroken cells and cell debris of the French press eluent by centrifugation at 10,000 x g for 15 min at 4 &#176;C. Discard the pellet, and transfer supernatant to ultracentrifuge tubes.</li>
	<li>Pellet membrane vesicles from the resulting supernatant by ultracentrifugationat 150,000 g for 1 h at 4 &#176;C.</li>
	<li>Wash the membrane vesicles once, without resuspension, in 1 mM Tris-maleate, pH 5.2 containing 0.14 M KCl, 2 mM 2-mercaptoethanol and 10% glycerol and resuspend in the same buffer (Buffer 2)23 using a Dounce tissue grinder. Typically the membrane fraction from 500 mL of culture would be suspended in 5 mL of buffer, and a concentration of 5-10 mg of membrane protein/mL using the Biochinic Acid assay32.</li>
	<li>Store 100 &#181;L aliquots of membrane preparations at -80 &#176;C in 1.5 mL microcentrifuge tubes.</li>
</ol>
4. Western Blot and Orientation of Transporter Assay
<ol>
	<li>Wash and resuspend vesicles from step 3.7 in 30 mM Tris pH 7.8 + 0.1 mM CoCl2.</li>
	<li>To 100 &#181;L samples, add 1 &#181;L of 2 mg/mL carboxypeptidase A in 0.1 M NaCl, 30 mM Tris, 0.2 mM CoCl2 pH 7.8.</li>
	<li>Inculate for 20 min at 20 &#176;C. Stop digestion by adding 5 &#181;L of 0.5 M NaEDTA, 0.5 M 2-mercaptoethanol pH 7.5.</li>
	<li>To fully inactivate the enzyme, incubate the solution for 1 h at room temperature. 
	NOTE: Vesicles without the catboxypeptidase A treatment were used as a control.</li>
	<li>Analyze samples by electrophoresis in the presence of lithium dodecyl sulfate. Add 5-10 &#181;L of 150 mg/mL lithium dodecyl sulfate, 450 mg/mL glycerol, 0.1 mg/mL bromophenol blue, 0.4 M Tris pH 7.5 to 100 &#181;L of sample.</li>
	<li>Perform immunoblotting by laying a nitrocellulose filter moistened with 25 mM sodium-hydrogen phosphate pH 7.5 upon the polyacrylamide gel. Place these between two moistened cellulose filters and finally between two moistened plastic scouring pads. Place this assemblage between the electrodes of a chamber containing 25 mM sodium-hydrogen phosphate pH 7.5 at 2-4 &#176;C.</li>
	<li>Allow electrotransfer of proteins to proceed for 3 h at 20 V and 2-3 A.</li>
	<li>Block the nitrocellulose membrane overnight at 2 &#176;C in a blocking buffer of 0.15 M NaCl, 10 mM Tris, pH 7.5 and 0.5 mg/mL Tween 2033.</li>
	<li>Incubate the nitrocellulose filter for 2 h with a 1:5000 dilution of Anti-His (C terminal)-HRP antibody in the blocking buffer (20 mL) and wash twice with 50 mL of buffer for a total of 60 min.</li>
	<li>To visualize the immunoblot, dissolve 6 mg of 4-chloro-1-naphthol in 20 mL of denatured alcohol and add 80 mL of 15 mM Tris pH 7.5 and 50 &#181;L of 30% H2O2. Bathe the filter paper in the substrate solution for 20 min. The reagent will react with the antibody to form a blue precipitate on the nitrocellulose membrane. When sufficient color has developed, rinse the membrane with water, and allow to dry.</li>
</ol>
5. Transport Assay
<ol>
	<li>Incubate 100 &#956;L aliquots of membrane vesicles at 12 &#176;C for 5 min.</li>
	<li>Initiate transport by adding radiolabeled polyamines to the membrane vesicles at a final concentration of 50 &#181;M (unless otherwise stated). Make 3H-substrate (spermidine or putrescine) solutions in an assay buffer consisting of 10 mM Tris-HCl, 10 mM potassium phosphate, pH 8.0 and 0.14 M KCl23 (Buffer 3) with modifications depending on the assay.</li>
	<li>Conduct transport assays for 1 min at 12 &#176;C in 1.5 mL microfuge tubes.</li>
	<li>After 1 min, transfer the reaction mixtures to filtration manifold and filter through a 0.45 &#181;m nitrocellulose membrane filter.
	<ol>
		<li>Add 3 mL of ice-cold assay buffer containing a 10-fold higher concentration of unlabeled polyamines followed by 3 mL of assay buffer without the polyamines to reduce nonspecific binding.</li>
		<li>Transfer the washed filters to 20 mL disposable scintillation vials containing 10 mL of scintillation liquid and determine radioactivity by using a liquid scintillation counter. The scintillation counter measures the radioactivity in the sample and reports it as disintegrations per min (dpm).</li>
	</ol>
	</li>
	<li>Calculate the net polyamine uptake as the difference between the uptake at 1 min by vesicles incubated at 12 &#176;C and uptake at 0 min by vesicles incubated on ice. 
	NOTE: The mass of substrate imported into the vesicles is calculated as follows. If the sample volume in the microfuge tube is 0.2 &#181;L and the starting concentration of substrate is 100 &#181;M, then the total mass of substrate in the microfuge tube is 20 x 10-12 M. Because of the very high specific activity of commercially labelled substrates (often 2.22 x 109 dpm/mM) the mass of the isotope added can be ignored in the calculation. So if the total amount of isotope that was added to the microfuge tube was 100 x 106 dpm and the vesicles had a net uptake of 1,000 dpm, then the total mass of substrate taken up by the vesicles was 1% of the total number of moles of substrate or 2 x 10-13 M.
	<ol>
		<li>Determine Km for the substrate by measuring the uptake of 10, 25, 50, 250 and 500 &#956;M radiolabeled substrate into vesicles expressing the target protein. Calculate Michaelis-Menten kinetics using a nonlinear regression method using the Michaelis Menton model of the statistical software package34.</li>
	</ol>
	</li>
	<li>For the competition experiments, add 100 &#956;M, 500 &#956;M, 1 mM, 1.5 mM or 2 mM nonlabelled competitive substrate made in the assay buffer to the 1.5 mL microcentrifuge tube containing 100 &#181;L of vesicles at 12 &#176;C.
	<ol>
		<li>Add radiolabeled polyamine (50 &#181;M) to the microcentrifuge tube simultaneously.</li>
		<li>Measure radioactivity trapped inside vesicles as mentioned above by repeating steps 5 through 5.4.2.</li>
		<li>Determine apparent Km (Km,app) for the competitive substrates by measuring the uptake of 10, 25, 50, 250 and 500 &#956;M radiolabeled polyamines in the presence of 100 &#956;M or higher nonlabelled substrate and using a nonlinear regression method to plot the curve.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

In the present study, we outline a method for the characterization of an antiporter by first expressing the protein in E. coli and then generating membrane vesicles, so that the heterologously-expressed protein can be assayed in a cell-free system. In addition to equipment found in most molecular biology labs, this strategy requires the use of a French press, an ultracentrifuge, and access to a facility to conduct radioisotope assays.
A basic requirement of this technique is that the heterologous protein is correctly targeted to the plasma membrane of E. coli. This strategy may also be useful for functional analysis of organellar transporters since the plastid ADP glucose transporter was successfully localized to the E. coli cell membrane and functionally characterized35. The vector (pBAD-DEST49) used in these experiments contains an N-terminal thioredoxin protein to increase the solubility of the translated product. N-terminal fusions of a small B. subtilus protein mystic, have been found to enable more efficient targeting of membrane transporters to the cytoplasmic membrane36. However, misfolding events, and the failure of the proteins to be properly integrated into the cytoplasmic membrane are potential problems that preclude the use of bacterial expression systems for many types of transporters1.
Membrane vesicles have also been used to characterize plant transporters37,38. As the vesicles lack the essential energy sources such as ATP and enzymes, the interference from active transporters and other metabolic activity is minimal. Thus, this system is ideal for the analysis of passive translocations such as metabolite exchangers. The everted membrane vesicles, in particular, can be applied to the characterization of exporters and antiporters since the composition of the internal solution can be manipulated by changing the composition of buffer 1. Furthermore, using French press or ultrasound sonication is fairly efficient in generating inside-out membrane vesicles from intact E. coli cells. 95% of the vesicles generated by ultrasound sonication or French press have everted membranes39,40. PotE, the E. coli antiporter of putrescine and ornithine, was the first polyamine antiporter that was characterized using inside-out membrane vesicles23. We used P1 transduction to create a specific mutant strain for the characterization of a polyamine antiporter, and this strain may be useful for the characterization of other animal, fungal or plant polyamine exchangers. We also envision that other E. coli strains with two or more gene deletions might be useful for the characterization of other plant and animal exchange transporters using membrane vesicles.
The most critical step in this protocol is the expression of the protein in the E. coli mutant system. An E. coli expression vector with an inducible promoter is utilized to promote tight, dose dependent regulation of the heterologous gene expression. The presence of N terminal and C terminal tags such as His-patch Thioredoxin, V5 epitope or 6xHis in the vector is useful for detection and purification of the protein. In addition, the presence of a thioredoxin fusion protein which is a component of the pBAD49 vector, can increase translation efficiency and, in some cases, solubility of eukaryotic proteins expressed in E. coli41. The different codon choices in Arabidopsis and E. coli could challenge protein expression in E. coli. It is known that codon optimization can impressively increase heterozygous proteins expression in E. coli42. In the vesicle assay, codon optimized AtBAT1.2 showed a higher exchange activity than non-codon optimized AtBAT1.1 in E. coli cells (data not shown), demonstrating that codon optimization was helpful to enhance the expression and function of heterologously expressed proteins in bacterial cells. The production of membrane vesicles by careful adjustment of the valve to maintain a slow even drip of lysed cells is also a key step in the procedure. After ultracentrifugation, we have found that resuspension of membrane vesicles in a Dounce homogenizer minimizes sample to sample variation between aliquots of membrane vesicles that are prepared and subsequently stored at -80 &#176;C.
A limitation of E. coli expression systems is that they are incapable of post-translational modifications such as N-glycosylation or acetylation. Absence of these protein modifications might impact protein activity1. However, mutants capable of performing these modifications have been identified and can be used as a tool to express proteins that require such modifications43. The generation of sufficient amounts of the expressed protein could be a challenge due to unfolding and aggregation as inclusion bodies, failure of the protein to be properly integrated in to the cytoplasmic membrane, mistargeting and mis-regulation due to lack of post translational modifications.
A minor limitation of this technique is that it does not provide evidence for the natural orientation of the transporter. This can be accomplished by taking advantage of the N or C terminal tags and immunological methods. The accessibility of a particular terminus of the protein in vesicles can be achieved by the digestion of all accessible, and therefore, presumably external termini of the carrier, electrophoresis of the protein in the presence of sodium dodecyl sulfate, transfer to nitrocellulose filters and detection of the remaining, internal termini with antibodies40.

Proteins involved in the trafficking of metabolites constitute an essential level of physiological regulation, but the vast majority of plant membrane transporters have not yet been functionally characterized. Several strategies have been implemented to characterize novel transport proteins. Heterologous expression in model organisms such as E. coli and eukaryotic cells such as yeast, Xenopus oocytes, mammalian cells, insect cells and plant cells have all been used to determine their transport activity1. Eukaryotic cells are favored for the expression of eukaryotic proteins, because the basic cellular composition, signal transducing pathways, transcription and translation machineries are compatible with the native conditions.
Yeast has been an important model organism for the characterization of novel transport proteins in plants. The first plant transport protein that was successfully expressed in yeast (Saccharomyces pombe) was the hexose transporter HUP1 from Chlorella2. Since then, many plant transport proteins have been functionally characterized using a yeast expression system. These include, plant sugar transporters (SUC1 and SUC23, VfSUT1 and VfSTP14) and the auxin transporters (AUX1 and PIN5). Disadvantages of utilizing yeast to express plant proteins can include impaired activity of plastid-localized proteins because yeast lacks this organelle, mistargeting6, and formation of misfolded aggregates and activation of stress responses in yeast due to overexpression of membrane proteins7,8,9.
Heterologous expression of transport proteins in Xenopus oocytes have been widely used for the electrophysiological characterization of transporters10. The first plant transport proteins characterized using heterologous expression in Xenopus oocytes were the Arabidopsis potassium channel KAT110 and the Arabidopsis hexose transporter STP111. Since then, Xenopus oocytes have been employed to characterize many plant transport proteins such as plasma membrane transporters12, vacuolar sucrose transporter SUT413 and vacuolar malate transporter ALMT914. An important limitation of Xenopus oocytes for transport assays is that the concentration of intracellular metabolites cannot be manipulated1. Moreover, professional knowledge is required to prepare Xenopus oocytes and the variability of the oocyte batches is difficult to control.
Heterologous expression in the model organism E. coli is an ideal system in terms of characterization of novel plant transport proteins. With a fully sequenced genome15, the molecular and physiological characteristics of E. coli are well known. Molecular tools and techniques are well established16. In addition, different expression vectors, non-pathogenic strains and mutants are available17,18,19. Furthermore, E. coli has a high growth rate and can be easily grown under laboratory conditions. Many proteins can be easily expressed and purified at high amounts in E. coli9. When proteins cannot be assayed directly in cellular systems, reconstitution of proteins into liposomes has also been a successful, albeit challenging innovation for the characterization of purified membrane proteins. Functional characterization of the plant mitochondrial transport proteins including solute transporters such as phosphate transporters in soybean, maize, rice and Arabidopsis, dicarboxylate-tricarboxylate carrier in Arabidopsis have been accomplished by using this model system20,21. However, recombinant proteins of the tomato protein SICAT9 were found to be nonfunctional in reconstitution experiments, and other members of the CAT transporter family were found to be nonfunctional in Xenopus oocyte assays22. Thus, additional molecular tools are needed for the characterization of membrane transporters.
Five polyamine transport systems are found in E. coli23. They include two ABC transporters mediating the uptake of spermidine and putrescine, a putrescine/ornithine exchanger, a cadaverine/lysine exchanger, a spermidine exporter and a putrescine importer. The putrescine exchanger PotE was originally characterized using a vesicle assay, where inside out vesicles were prepared by lysing cells with a French press and measuring the uptake of radiolabeled putrescine into the vesicles in exchange for ornithine24. Vesicle assays were also used to characterize a calcium transporter, which mediated the transport of calcium in response to a proton gradient25. These experiments prompted us to develop a strategy for the characterization of other polyamine exchangers. We first created a strain of E. coli deficient in PotE and CadB exchangers. Here, we demonstrate the functional characterization of a plant polyamine antiporter by heterlogous expression in the modified E. coli strain, generation of membrane vesicles using a French press, and radiolabeled assays.

<table><tbody><tr><td>2-mercaptoethanol</td><td>Sigma-Aldrich</td><td>M6250</td><td></td></tr><tr><td>&lt;sup&gt;3&lt;/sup&gt;H-putrescine</td><td>PerkinElmer</td><td>NET185001MC</td><td></td></tr><tr><td>&lt;sup&gt;3&lt;/sup&gt;H-spermidine</td><td>PerkinElmer</td><td>NET522001MC</td><td></td></tr><tr><td>4-chloro-1-naphthol</td><td>Sigma-Aldrich</td><td>C8890</td><td></td></tr><tr><td>&lt;sup&gt;14&lt;/sup&gt;C arginine</td><td>Moravek Inc.</td><td>MC137</td><td></td></tr><tr><td>Arginine</td><td>Sigma-Aldrich</td><td>A-5006</td><td></td></tr><tr><td>Anti-His (C-term)-HRP antibody</td><td>ThermoFisher</td><td>R931-25</td><td>Detects the C-terminal polyhistidine (6xHis) tag, requires the free carboxyl&lt;br/&gt; group for detection</td></tr><tr><td>Arabinose</td><td>Sigma-Aldrich</td><td>A3256</td><td></td></tr><tr><td>BCA protein assay kit</td><td>ThermoFisher</td><td>23227</td><td>Pierce BCA protein asay kit.</td></tr><tr><td>Bromophenol blue</td><td>Bio-Rad</td><td>161-0404</td><td></td></tr><tr><td>Carboxypeptidase B</td><td>Sigma-Aldrich</td><td>C9584-1mg</td><td></td></tr><tr><td>Centrifuge</td><td>Sorvall</td><td></td><td>SS-34 fixed angle rotor and GA-6 fixed angle rotor</td></tr><tr><td>Dounce tissue grinder</td><td>LabGenome</td><td>7777-7</td><td>Corning 7777-7 pyrex homogenizer with pour spout.</td></tr><tr><td>Ecoscint-H</td><td>National Diagnostics</td><td>LS275</td><td>scintillation cocktail</td></tr><tr><td>EDTA</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Filtration manifold</td><td>Hoefer</td><td>FH225V</td><td></td></tr><tr><td>French Pressure Cell</td><td>Glen Mills</td><td>FA-080A120</td><td></td></tr><tr><td>GABA</td><td>Sigma-Aldrich</td><td>A2129</td><td></td></tr><tr><td>Glutamate</td><td>Sigma-Aldrich</td><td>G6904</td><td></td></tr><tr><td>Glycerol</td><td></td><td></td><td></td></tr><tr><td>GraphPad Prism software</td><td></td><td></td><td>&lt;a href=&quot;http://www.graphpad.com/prism/Prism.htm&quot; target=&quot;_blank&quot;&gt;http://www.graphpad.com/prism/Prism.htm&lt;/a&gt;</td></tr><tr><td>Hydrogen peroxide</td><td>KROGER</td><td></td><td></td></tr><tr><td>Potassium Chloride</td><td>J.T. Baker</td><td>3040-01</td><td></td></tr><tr><td>Liquid scintillation counter</td><td>Beckman</td><td>LS-6500</td><td></td></tr><tr><td>Maleate</td><td>Sigma-Aldrich</td><td>M0375</td><td></td></tr><tr><td>Nanodrop</td><td>ThermoFisher</td><td></td><td></td></tr><tr><td>Nitrocellulose membrane filters</td><td>Merck Millipore</td><td>hawp02500</td><td>0.45 &amp;micro;M</td></tr><tr><td>PCR clean up kit</td><td>Genscript</td><td></td><td>QuickClean II</td></tr><tr><td>Potassium Phosphate dibasic</td><td>ThermoFisher</td><td>P290-500</td><td></td></tr><tr><td>putrescine</td><td>fluka</td><td>32810</td><td></td></tr><tr><td>Potassium Phosphate monobasic</td><td>J.T.Baker</td><td>4008</td><td></td></tr><tr><td>Spermidine</td><td>Sigma-aldrich</td><td>S2501</td><td></td></tr><tr><td>Strains :E. coli &amp;Delta;potE740(del)::kan, &amp;Delta;cadB2231::Tn10</td><td>This manuscript</td><td>Available upon request.</td><td>Strain is deficient in the &lt;em&gt;PotE&lt;/em&gt; and &lt;em&gt;CadB&lt;/em&gt; polyamine exchangers.</td></tr><tr><td>Tris-base</td><td>Research Products</td><td>T60040-1000</td><td></td></tr><tr><td>Ultracentrifuge</td><td>Sorvall MTX 150</td><td>46960</td><td>Thermo Fisher S150-AT fixed angle rotor</td></tr><tr><td>Ultracentrifuge tubes</td><td>ThermoFisher</td><td>45237</td><td>Centrifuge tubes for S150-AT rotor</td></tr><tr><td>Vector: pBAD-DEST49</td><td>ThermoFisher</td><td></td><td>Gateway expression vector for &lt;em&gt;E. coli&lt;/em&gt;</td></tr></tbody></table>

characterization of membrane transporters by heterologous expression in e. coli and production of membrane vesicles

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine exchangers in plants have not been identified. Here, we outline a method to characterize polyamine antiporters using membrane vesicles generated from the lysis of Escherichia coli cells heterologously expressing a plant antiporter. First, we heterologously expressed AtBAT1 in an E. coli strain deficient in polyamine and arginine exchange transporters. Vesicles were produced using a French press, purified by ultracentrifugation and utilized in a membrane filtration assay of labeled substrates to demonstrate the substrate specificity of the transporter. These assays demonstrated that AtBAT1 is a proton-mediated transporter of arginine, &#947;-aminobutyric acid (GABA), putrescine and spermidine. The mutant strain that was developed for the assay of AtBAT1 may be useful for the functional analysis of other families of plant and animal polyamine exchangers. We also hypothesize that this approach can be used to characterize many other types of antiporters, as long as these proteins can be expressed in the bacterial cell membrane. E. coli is a good system for the characterization of novel transporters, since there are multiple methods that can be employed to mutagenize native transporters.

The major steps in this protocol are summarized pictorially in Figure 1. Briefly, E. coli cells deficient in all polyamine exchangers and expressing AtBAT1 are cultured, centrifuged, washed with a buffer and subjected to cell lysis using a French press. Lysis tends to produce vesicles that are mostly inside-out and trap the buffer outside the cells. Cell debris is removed by centrifugation, and a second ultracentifugation step is used to collect a membrane pellet. The membrane pellet is resuspended in Tris-Maleate buffer pH 5.2 and stored at -80 &#176;C. Transport assays are done at 12 &#176;C, which was found to be optimal for maintaining membrane stability. Assays are initiated by the addition of radiolabeled substrate and a shift in the pH of the buffer suspension of vesicles to pH 8.0. After 1 min, ice-cold assay buffer with unlabeled substrates is added to stop the uptake of the radiolabel into the vesicles. Radiolabelled vesicles are trapped by filtration through nitrocellulose membranes. Membranes are transferred to scintillation vials and radiolabel on the membranes is determined by liquid scintillation counting.
A western blot is used to verify that AtBAT1 is translocated to vesicles (Figure 2). Probing the blot with an Anti-His C-terminal antibody revealed a fusion construct protein of approximately 72.3 kDa (Figure 2, Lane 2). Digestion of the vesicles prior to SDS-PAGE resulted in a dimunition, but not a complete loss of the probe signal (Figure 2, lane 3). The decrease in the probe signal as a consequence of carboxypeptidase A suggests that most of the C-terminal residues are on the outside of the vesicles.
In this assay system, vesicles are suspended in a buffer at pH 5.2 so that the pH inside the vesicles equilibrates with the buffer. Transport of the radiolabeled substrate into the vesicles at pH 5.2 is initiated by suspending the vesicles in a pH 8.0 buffer, thus creating a pH gradient of pH 2.8 across the membrane. At 12 &#176;C, uptake of radiolabeled spermidine by the vesicles was highest at 1 min, and remained linear over 3 min (Figure 3A). Therefore, the incubation time for the transport assay was fixed at 1 min. To account for non-specific binding of radiolabel, the vesicles were incubated at 0 &#176;C in the presence of radiolabeled substrate for one minute, and these counts were subtracted from uptake of ladiolabel at higher temperatures.
Figure 3B shows the uptake of radiolabeled spermidine into the vesicles after one minute. There was no net uptake of isotope by membrane vesicles that were prepared and stored at pH 8.0, as there was no proton gradient across the vesicle membrane. To demonstrate the effect of dissipation of the artificial proton gradient, the membrane vesicles were incubated in pH 8.0 buffer for 10 min prior to the addition of labelled substrate25. Under these conditions, a minimal uptake of radiolabeled substrate was shown. Uptake of radiolabeled spermidine was also minimal in vesicles prepared with E. coli cells deficient in the polyamine exchangers CadB and PotE. Taken together, these results indicate that the proton driven uptake of spermidine was due to the BAT1 protein (Figure 3A,B).
To determine the substrate specificity of the protein, Km values were calculated by measuring the uptake of radiolabeled substrate at 10, 25, 50, 100, 250 and 500 &#956;M concentrations. The Km for spermidine, putrescine and arginine were 55 &#177; 12 &#956;M, 85 &#177; 20 &#956;M and 1.4 &#177; 0.5 mM, respectively, indicating that this protein is a high affinity polyamine and arginine exchanger (Figure 4).
Affinity of the transporter for a particular substrate can also be determined indirectly by using competition assays. Here, we have utilized two methods to evaluate the competition between two substrates. In the first method, the uptake of 50 &#956;M radiolabeled spermidine was measured in the presence of increasing concentrations of the nonlabelled competing substrate (Figure 5A). In the second method, the apparent Km for spermidine was calculated by measuring the uptake of increasing concentrations of radiolabeled spermidine in the presence of 100 &#956;M nonlabelled competing substrate (Figure 5B). Competition assays revealed that GABA is a competitive inhibitor of spermidine with a Km,app of 164 &#177; 15 &#956;M (Figure 5A,B). Furthermore, measuring the uptake of 50 &#956;M radiolabeled spermidine in the presence of varying concentrations of different amino acids revealed that AtBAT1 is also capable of transporting glutamate and alanine at mM concentrations (Figure 6).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60009/60009fig1a.jpg" /> 
Figure 1: Schematic representation of the method. (A) Schematic representation outlining key steps in the preparation and purification of membrane vesicles from E. coli. (B) Schematic representation outlining key steps in transport assay of membrane vesicle preparations using radiolabeled substrates. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60009/60009fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60009/60009fig2a.jpg" /> 
Figure 2: Western blot showing expression of AtBAt1 in purified vesicles. Bands were visualized using horseradish peroxidase conjugated anti-His (C-term)-HRP antibody. Lane 1, Prestained protein ladder. Lane 2, Purified vesicles expressing AtBAT1.1 showing a band of the expected size of the fusion protein. Lane 3, purified vesicles expressing AtBAT1.1 were pretreated with carboxypeptidase A prior to SDS electrophoresis and western blotting. Equivalent amounts of vesicles (protein) were added to each lane. Decreased staining indicates that the C-terminal of the protein in most vesicles is degraded by by protease digestion. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60009/60009fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60009/60009fig3a.jpg" /> 
Figure 3: Transport activity of vesicles showing the effect of BAT1 protein expression and the importance of a pH gradient. (A) Time dependent uptake of 3H labeled spermidine in vesicles expressing BAT1 with an internal pH of 5.2 and introduced to a buffer at pH 8.0. In the control assay, the vesicles were added to the assay buffer at pH 8.0, 10 min prior to the addition of 3H labeled spermidine to enable dissipation of the proton gradient. Then uptake of radiolabel into the vesicles was assessed over a 1 min interval. (B) Uptake of 3H labeled spermidine in the presence of a proton gradient (internal pH of 5.2), in the absence of a proton gradient (internal pH of 8), in vesicles added to the assay solution 10 min prior to the addition of radiolabeled spermidine and in vesicles made from E. coli mutant cells not expressing BAT1. Uptake into vesicles was monitored for 1 min. All values are presented as mean &#177; SE of five replicates. Data analysis was performed using a student&#39;s t-test and * indicates a significant difference from the control (p value &#60; 0.05). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60009/60009fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60009/60009fig4a.jpg" /> 
Figure 4: In vitro assays of polyamine and arginine transport activity of BAT1. (A) The Km values for spermidine and putrescine uptake are 55 &#177; 12 &#956;M and 85 &#177; 32 &#956;M respectively. (B)The Km for arginine uptake is 1.4 &#177; 0.5 mM. All values are presented as mean &#177; SE of five replicates. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60009/60009fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60009/60009fig5a.jpg" /> 
Figure 5: GABA is a competitive inhibitor of Spermidine transport by BAT1. (A) Uptake of 3H labeled spermidine by vesicles expressing AtBAT1.1 was significantly reduced in the presence of 100 &#956;M or 500 &#956;M GABA. (B) Apparent Km for spermidine uptake by BAT1.1 was increased to 164 &#177; 20 &#956;M in the presence of 100 &#956;M GABA. All values are presented as mean &#177; SE of five replicates. Data analysis was performed using a student&#39;s t-test and * indicates a significant difference from the control (p value &#60; 0.05). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60009/60009fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60009/60009fig6a.jpg" /> 
Figure 6: Glutamate and alanine are competitive inhibitors of spermidine transportby BAT1. Spermidine uptake was significantly reduced in the presence of 1 mM non-labeled glutamate and 1.5 mM non-labeled alanine. All values are presented as mean &#177; SE of five replicates. Data analysis was performed using a student&#39;s t-test and * indicates a significant difference from the control (p value &#60; 0.05). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60009/60009fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles at JoVE.com

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

We describe a method for the characterization of proton-driven membrane transporters in membrane vesicle preparations produced by heterologous expression in E. coli and lysis of cells using a French press.

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

Several methods have been developed to functionally characterize novel membrane transporters. Polyamines are ubiquitous in all organisms, but polyamine ...

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9.2K Views.  Bowling Green State University. Een van de belangrijkste voordelen van deze techniek voor het karakteriseren van membraantransporters is dat u de relatieve affiniteit van de transporter voor specifieke substraten bepalen en inzicht krijgt in het transportmechanisme. De karakterisering van membraaneiwitten die eiwitgradiënten gebruiken om substraattransport te stimuleren, is bijzonder geschikt voor deze binnenste vesicletest die we hier laten zien. Hoewel deze test vereist een aantal gespecialiseerde apparatuur, zoals een F...

Video: Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

An increasing interest in applying synthetic biology techniques to program outer membrane vesicles (OMV) are leading to some very interesting and unique applications for OMV where traditional nanoparticles are proving too difficult to synthesize. To date, all Gram-negative bacteria have been shown to produce OMV demonstrating packaging of a variety of cargo that includes small molecules, peptides, proteins and genetic material. Based on their diverse cargo, OMV are implicated in many biological processes ranging from cell-cell communication to gene transfer and delivery of virulence factors depending upon which bacteria are producing the OMV. Only recently have bacterial OMV become accessible for use across a wide range of applications through the development of techniques to control and direct packaging of recombinant proteins into OMV. This protocol describes a method for the production, purification, and use of enzyme packaged OMV providing for improved overall production of recombinant enzyme, increased vesiculation, and enhanced enzyme stability. Successful utilization of this protocol will result in the creation of a bacterial strain that simultaneously produces a recombinant protein and directs it for OMV encapsulation through creating a synthetic linkage between the recombinant protein and an outer membrane anchor protein. This protocol also details methods for isolating OMV from bacterial cultures as well as proper handling techniques and things to consider when adapting this protocol for use for other unique applications such as: pharmaceutical drug delivery, medical diagnostics, and environmental remediation.

Directed Protein Packaging within Outer Membrane Vesicles from Escherichia coli: Design, Production and Purification

A protocol for the production, purification, and use of enzyme packaged outer membrane vesicles (OMV) providing for enhanced enzyme stability for implementation across diverse applications is presented.

An increasing interest in applying synthetic biology techniques to program outer membrane vesicles (OMV) are leading to some very interesting and unique ...

From Constructs to Crystals &#8211; Towards Structure Determination of &#946;-barrel Outer Membrane Proteins

Membrane proteins serve important functions in cells such as nutrient transport, motility, signaling, survival and virulence, yet constitute only ~1% percent of known structures. There are two types of membrane proteins, &#945;-helical and &#946;-barrel. While &#945;-helical membrane proteins can be found in nearly all cellular membranes, &#946;-barrel membrane proteins can only be found in the outer membranes of mitochondria, chloroplasts, and Gram-negative bacteria. One common bottleneck in structural studies of membrane proteins in general is getting enough pure sample for analysis. In hopes of assisting those interested in solving the structure of their favorite &#946;-barrel outer membrane protein (OMP), general protocols are presented for the production of target &#946;-barrel OMPs at levels useful for structure determination by either X-ray crystallography and/or NMR spectroscopy. Here, we outline construct design for both native expression and for expression into inclusion bodies, purification using an affinity tag, and crystallization using detergent screening, bicelle, and lipidic cubic phase techniques. These protocols have been tested and found to work for most OMPs from Gram-negative bacteria; however, there are some targets, particularly for mitochondria and chloroplasts that may require other methods for expression and purification. As such, the methods here should be applicable for most projects that involve OMPs from Gram-negative bacteria, yet the expression levels and amount of purified sample will vary depending on the target OMP.

&#946;-barrel outer membrane proteins (OMPs) serve many functions within the outer membranes of Gram-negative bacteria, mitochondria, and chloroplasts. Here, we hope to alleviate a known bottleneck in structural studies by presenting protocols for the production of &#946;-barrel OMPs in sufficient quantities for structure determination by X-ray crystallography or NMR spectroscopy.

Membrane proteins serve important functions in cells such as nutrient transport, motility, signaling, survival and virulence, yet constitute only ~1% ...

Chemistry

Expression, Solubilization, and Purification of Eukaryotic Borate Transporters

The Solute Carrier 4 (SLC4) family of proteins is called the bicarbonate transporters and includes the archetypal protein Anion Exchanger 1 (AE1, also known as Band 3), the most abundant membrane protein in the red blood cells. The SLC4 family is homologous with borate transporters, which have been characterized in plants and fungi. It remains a significant technical challenge to express and purify membrane transport proteins to homogeneity in quantities suitable for&#160;structural or functional studies. Here we describe detailed procedures for the overexpression of borate transporters in Saccharomyces cerevisiae, isolation of yeast membranes, solubilization of protein by detergent, and purification of borate transporter homologs from S. cerevisiae, Arabidopsis thaliana, and Oryza sativa. We also detail a glutaraldehyde cross-linking experiment to assay multimerization of homomeric transporters. Our generalized procedures can be applied to all three proteins and have been optimized for efficacy. Many of the strategies developed here can be utilized for the study of other challenging membrane proteins.

Here we present a protocol to express, solubilize, and purify several eukaryotic borate transporters with homology to the SLC4 transporter family using yeast. We also describe a chemical cross-linking assay to assess the purified homomeric proteins for multimeric assembly. These protocols can be adapted for other challenging membrane proteins.

The Solute Carrier 4 (SLC4) family of proteins is called the bicarbonate transporters and includes the archetypal protein Anion Exchanger 1 (AE1, also ...

Co-Translational Insertion of Membrane Proteins into Preformed Nanodiscs

Cell-free expression systems allow the tailored design of reaction environments to support the functional folding of even complex proteins such as membrane proteins. The experimental procedures for the co-translational insertion and folding of membrane proteins into preformed and defined membranes supplied as nanodiscs are demonstrated. The protocol is completely detergent-free and can generate milligrams of purified samples within one day. The resulting membrane protein/nanodisc samples can be used for a variety of functional studies and structural applications such as crystallization, nuclear magnetic resonance, or electron microscopy. The preparation of basic key components such as cell-free lysates, nanodiscs with designed membranes, critical stock solutions as well as the assembly of two-compartment cell-free expression reactions is described. Since folding requirements of membrane proteins can be highly diverse, a major focus of this protocol is the modulation of parameters and reaction steps important for sample quality such as critical basic reaction compounds, membrane composition of nanodiscs, redox and chaperone environment, or DNA template design. The whole process is demonstrated with the synthesis of proteorhodopsin and a G-protein coupled receptor.

Co-translational insertion into pre-formed nanodiscs makes it possible to study cell-free synthesized membrane proteins in defined lipid environments without contact with detergents. This protocol describes the preparation of essential system components and the critical parameters for improving expression efficiency and sample quality.

Cell-free expression systems allow the tailored design of reaction environments to support the functional folding of even complex proteins such as ...

Cell-Free Production of Proteoliposomes for Functional Analysis and Antibody Development Targeting Membrane Proteins

Membrane proteins play essential roles in a variety of cellular processes and perform vital functions. Membrane proteins are medically important in drug discovery because they are the targets of more than half of all drugs. An obstacle to conducting biochemical, biophysical, and structural studies of membrane proteins as well as antibody development has been the difficulty in producing large amounts of high-quality membrane protein with correct conformation and activity. Here we describe a &#8220;bilayer-dialysis method&#8221; using a wheat germ cell-free system, liposomes, and dialysis cups to efficiently synthesize membrane proteins and prepare purified proteoliposomes in a short time with a high success rate. Membrane proteins can be produced as much as in several milligrams, such as GPCRs, ion channels, transporters, and tetraspanins. This cell-free method contributes to reducing the time, cost and effort for preparing high-quality proteoliposomes, and provides suitable means for functional analysis of membrane proteins, drug targets screening, and antibody development.

This protocol describes an efficient cell-free method for production of high-quality proteoliposome by bilayer-dialysis method using wheat cell-free system and liposomes. This method provides suitable means for functional analysis of membrane proteins, drug targets screening, and antibody development.

Membrane proteins play essential roles in a variety of cellular processes and perform vital functions. Membrane proteins are medically important in drug ...

Expression, Detergent Solubilization, and Purification of a Membrane Transporter, the MexB Multidrug Resistance Protein

Multidrug resistance (MDR), the ability of a cancer cell or pathogen to be resistant to a wide range of structurally and functionally unrelated anti-cancer drugs or antibiotics, is a current serious problem in public health. This multidrug resistance is largely due to energy-dependent drug efflux pumps. The pumps expel anti-cancer drugs or antibiotics into the external medium, lowering their intracellular concentration below a toxic threshold. We are studying multidrug resistance in Pseudomonas aeruginosa, an opportunistic bacterial pathogen that causes infections in patients with many types of injuries or illness, for example, burns or cystic fibrosis, and also in immuno-compromised cancer, dialysis, and transplantation patients. The major MDR efflux pumps in P. aeruginosa are tripartite complexes comprised of an inner membrane proton-drug antiporter (RND), an outer membrane channel (OMF), and a periplasmic linker protein (MFP) 1-8. The RND and OMF proteins are transmembrane proteins. Transmembrane proteins make up more than 30% of all proteins and are 65% of current drug targets. The hydrophobic transmembrane domains make the proteins insoluble in aqueous buffer. Before a transmembrane protein can be purified, it is necessary to find buffer conditions containing a mild detergent that enable the protein to be solubilized as a protein detergent complex (PDC) 9-11. In this example, we use an RND protein, the P. aeruginosa MexB transmembrane transporter, to demonstrate how to express a recombinant form of a transmembrane protein, solubilize it using detergents, and then purify the protein detergent complexes. This general method can be applied to the expression, purification, and solubilization of many other recombinantly expressed membrane proteins. The protein detergent complexes can later be used for biochemical or biophysical characterization including X-ray crystal structure determination or crosslinking studies.

In this protocol we demonstrate the expression, solubilization, and purification of a recombinantly expressed membrane protein, MexB, as a soluble protein detergent complex. MexB is a multidrug resistance membrane transporter from the opportunistic bacterial pathogen Pseudomonas aeruginosa.

Multidrug resistance (MDR), the ability of a cancer cell or pathogen to be resistant to a wide range of structurally and functionally unrelated ...

Biology

Cell-Free Expression of Membrane Proteins in Giant Unilamellar Vesicles

Reconstitution of the Bacterial Glutamate Receptor Channel by Encapsulation of a Cell-Free Expression System

Cell-free expression (CFE) systems are powerful tools in synthetic biology that allow biomimicry of cellular functions like biosensing and energy regeneration in synthetic cells. Reconstruction of a wide range of cellular processes, however, requires successful reconstitution of membrane proteins into the membrane of synthetic cells. While the expression of soluble proteins is usually successful in common CFE systems, the reconstitution of membrane proteins in lipid bilayers of synthetic cells has proven to be challenging. Here, a method for reconstitution of a model membrane protein, bacterial glutamate receptor (GluR0), in giant unilamellar vesicles (GUVs) as model synthetic cells based on encapsulation and incubation of the CFE reaction inside synthetic cells is demonstrated. Utilizing this platform, the effect of substituting the N-terminal signal peptide of GluR0 with proteorhodopsin signal peptide on successful cotranslational translocation of GluR0 into membranes of hybrid GUVs is demonstrated. This method provides a robust procedure that will allow cell-free reconstitution of various membrane proteins in synthetic cells.

Author Spotlight: Membrane Protein Reconstitution in Synthetic Cells

This protocol describes the inverted emulsion method used to encapsulate a cell-free expression (CFE) system within a giant unilamellar vesicle (GUV) for the investigation of the synthesis and incorporation of a model membrane protein into the lipid bilayer.

Cell-free expression (CFE) systems are powerful tools in synthetic biology that allow biomimicry of cellular functions like biosensing and energy ...

Bioengineering

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 ...

Small-Scale Plasma Membrane Preparation for the Analysis of Candida albicans Cdr1-mGFPHis

The successful biochemical and biophysical characterization of ABC transporters depends heavily on the choice of the heterologous expression system. Over the past two decades, we have developed a yeast membrane protein expression platform that has been used to study many important fungal membrane proteins. The expression host Saccharomyces cerevisiae AD&#916;&#916; is deleted in seven major endogenous ABC transporters and it contains the transcription factor Pdr1-3 with a gain-of-function mutation that enables the constitutive overexpression of heterologous membrane protein genes stably integrated as single copies at the genomic PDR5 locus. The creation of versatile plasmid vectors and the optimization of one-step cloning strategies enables the rapid and accurate cloning, mutagenesis, and expression of heterologous ABC transporters. Here, we describe the development and use of a novel protease-cleavable mGFPHis double tag (i.e., the monomeric yeast enhanced green fluorescent protein yEGFP3 fused to a six-histidine affinity purification tag) that was designed to avoid possible interference of the tag with the protein of interest and to increase the binding efficiency of the His tag to nickel-affinity resins. The fusion of mGFPHis to the membrane protein ORF (open reading frame) enables easy quantification of the protein by inspection of polyacrylamide gels and detection of degradation products retaining the mGFPHis tag. We demonstrate how this feature facilitates detergent screening for membrane protein solubilization. A protocol for the efficient, fast, and reliable isolation of the small-scale plasma membrane preparations of the C-terminally tagged Candida albicans multidrug efflux transporter Cdr1 overexpressed in S. cerevisiae AD&#916;&#916;, is presented. This small-scale plasma membrane isolation protocol generates high-quality plasma membranes within a single working day. The plasma membrane preparations can be used to determine the enzyme activities of Cdr1 and Cdr1 mutant variants.

Small-Scale Plasma Membrane Preparation for the Analysis of Candida albicans Cdr1-mGFPHis

This article presents a small-scale plasma membrane isolation protocol for the characterization of Candida albicans ABC (ATP-binding cassette) protein Cdr1, overexpressed in Saccharomyces cerevisiae. A protease-cleavable C-terminal mGFPHis double tag with a 16-residue linker between Cdr1 and the tag was designed to facilitate the purification and detergent-screening of Cdr1.

The successful biochemical and biophysical characterization of ABC transporters depends heavily on the choice of the heterologous expression system. Over ...

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules derived from the parental cells. EVs function as intra- and inter-species communication vectors while also contributing to the interaction between bacteria and host organisms in the context of infection and colonization. Given the multitude of functions attributed to EVs in health and disease, there is a growing interest in isolating EVs for in vitro and in vivo studies. It was hypothesized that the separation of EVs based on physical properties, namely size, would facilitate the isolation of vesicles from diverse bacterial cultures. 
The isolation workflow consists of centrifugation, filtration, ultrafiltration, and size-exclusion chromatography (SEC) for the isolation of EVs from bacterial cultures. A pump-driven tangential flow filtration (TFF) step was incorporated to enhance scalability, enabling the isolation of material from liters of starting cell culture. Escherichia coli was used as a model system expressing EV-associated nanoluciferase and non-EV-associated mCherry as reporter proteins. The nanoluciferase was targeted to the EVs by fusing its N-terminus with cytolysin A. Early chromatography fractions containing 20-100 nm EVs with associated cytolysin A - nanoLuc were distinct from the later fractions containing the free proteins. The presence of EV-associated nanoluciferase was confirmed by immunogold labeling and transmission electron microscopy. This EV isolation workflow is applicable to other human gut-associated gram-negative and gram-positive bacterial species. In conclusion, combining centrifugation, filtration, ultrafiltration/TFF, and SEC enables scalable isolation of EVs from diverse bacterial species. Employing a standardized isolation workflow will facilitate comparative studies of microbial EVs across species.

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Bacteria secrete nanometer-sized extracellular vesicles (EVs) carrying bioactive biological molecules. EV research focuses on understanding their biogenesis, role in microbe-microbe and host-microbe interactions and disease, as well as their potential therapeutic applications. A workflow for scalable isolation of EVs from various bacteria is presented to facilitate standardization of EV research.

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules ...

Characterization of Membrane Transporters by Heterologous Expression in E. coli and Production of Membrane Vesicles

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