This research was supported by startup funds at Davidson College.

1. Preparation of media and important buffers
<ol>
	<li>Prepare 500 mL of 40% galactose by adding 200 g of galactose and de-ionized water to a total volume of 500 mL. Use a hot plate or microwave to increase the rate of galactose getting into solution. Sterile filter with a vacuum filtration apparatus consisting of a 0.2 &#181;m filter top attached to a sterile glass bottle. 
	NOTE: All media solutions are prepared using de-ionized water.</li>
	<li>Prepare 500 mL of 40% glucose by adding 200 g of glucose and water to a total volume of 500 mL. Autoclave to sterilize.</li>
	<li>Into each of the four 2 L flasks, add 0.4 g of complete supplement mixture without histidine (CSM-His), 3.35 g yeast nitrogen base with ammonium sulfate (YNB + nitrogen), and 475 mL water. Autoclave. Add 25 mL of 40% glucose to achieve a final glucose concentration of 2%.</li>
	<li>Add, to a glass bottle, 50 g peptone and 25 g yeast extract and water to 375 mL. Autoclave. Add 125 mL of 40% galactose to give 5x yeast peptone (YP) solution containing 10% galactose.</li>
	<li>Add, to a 250 mL flask, 0.04 g CSM-His, 0.335 g YNB + nitrogen, and water to 47.5 mL. Autoclave. Add 2.5 mL of 40% glucose to get 50 mL of CSM-His + YNB + 2% glucose.</li>
	<li>Prepare 1 L of 1M Tris pH 7.0 by mixing 121.14 g of Tris base with 800 mL of water. Add concentrated hydrochloric acid until the pH reaches 7.0. Add water to a final volume of 1 L and pass through a 0.2 &#181;m filter.</li>
	<li>Prepare 500 mL of 0.5 M ethylenediaminetetraacetic Acid (EDTA) pH 8.0 by mixing 73.06 g of EDTA with 400 mL water. Add sodium hydroxide pellets until the EDTA has dissolved and the pH reaches 8.0. Add water to a final volume of 500 mL and pass through a 0.2 &#181;m filter.</li>
	<li>Prepare 100 mL of 100 mM phenylmethanesulfonyl fluoride (PMSF) by mixing 1.74 g of PMSF with 200 proof ethanol. Store at -20 &#176;C.</li>
	<li>Prepare 225 mL of 2x bead wash buffer consisting of 50 mM Tris pH 7.0, 1.4 M NaCl, 20% glycerol, and 1 mM EDTA pH 8.0.</li>
	<li>Prepare 100 mL of the size exclusion chromatography buffer (S200 buffer) consisting of 20 mM 4-Morpholinoethanesulfonic acid hydrate (MES) pH 6.5, 100 mM NaCl, 2% glycerol, and 0.03% n-Dodecyl-beta-D-Maltopyranoside (DDM).</li>
	<li>Prepare 100 mL of membrane resuspension buffer consisting of 50 mM Tris pH 7.0, 500 mM NaCl, and 10% glycerol.</li>
	<li>Prepare 10 mL of a 3x SDS-PAGE loading dye by mixing 3 mL 20% sodium dodecyl sulfate (SDS), 3 mL glycerol, 2.4 mL 1M Tris pH 6.8, 0.03 g bromophenol blue, and 1.6 mL 2-mercaptoethanol.</li>
</ol>
2. Overexpression of Borate Transporter in S. cerevisiae
<ol>
	<li>Inoculate three transformed yeast colonies in 50 mL of CSM-His + YNB + 2% glucose media and shake overnight at 190 rpm at 30 &#176;C.</li>
	<li>The following day use a spectrophotometer to determine the optical density at a wavelength of 600 nm (OD600) and inoculate to an OD600 of 0.01 each of four 2 L flasks that each contain 500 mL of CSM-His + YNB + 2% glucose media.</li>
	<li>Shake at 190 rpm at 30 &#176;C for 30 h to allow the cells to consume all glucose and grow to a high density. 
	NOTE: A longer growth time results in larger cell yields, larger harvested membrane yields, and ultimately larger purified protein yields.</li>
	<li>Induce expression by adding 125 mL of 5x YP media supplemented with 10% galactose for a final induction concentration of 2% galactose. Shake at 190 rpm at 30 &#176;C for 16 h.</li>
	<li>Harvest cells by spinning at 4,000 x g for 15 min. Resuspend the cells in 100 mL cold water. 
	NOTE: At this point cells may be frozen at -80 &#176;C indefinitely, or the prep may continue into cell lysis and membrane harvesting. We typically harvest 40-45 g of cells.</li>
</ol>
3. Harvesting of Yeast Membranes
<ol>
	<li>Add to the cell resuspension 11.25 mL of 1M Tris pH 7.0, 0.45 mL of 0.5 M EDTA, and 2.25 mL of 100 mM PMSF, the latter two of which act as protease inhibitors. Add water to a final volume of 225 mL, which results in a cell resuspension buffer of 50 mM Tris pH 7.0, 1 mM EDTA, and 1 mM PMSF. 
	NOTE: If using frozen cells allow cells to thaw thoroughly, about 1 h at room temperature, because if they remain partially frozen, they will resist lysis by bead beating.</li>
	<li>Add the cell resuspension into a 450 mL metal canister for bead beating. Top off the remaining volume with cold 0.5 mm glass beads.</li>
	<li>Assemble a bead-beating chamber with the rotor and immerse in an ice bath. Perform six 1 min pulses, separated by 2 min rest periods to prevent overheating the lysate.</li>
	<li>Assemble a vacuum filtration apparatus by using a plastic disposable bottle top filter screwed into a glass bottle. Remove the filtration membrane, because the remaining plastic device can capture the beads while allowing lysate to pass. Separate the beads from the lysate by pouring the contents of the bead beating chamber onto the assembly while applying the vacuum.</li>
	<li>Wash the bead beating chamber with 225 mL of a 2x wash buffer that contains 50 mM Tris pH 7.0, 1 mM EDTA, 1 mM PMSF, 1.4 M NaCl, and 20% glycerol. Empty the contents of the chamber onto the beads to wash them. 
	NOTE: The wash gives a final volume of ~450 mL lysed cells and significantly increases harvested membrane yields. The final concentrations are 50 mM Tris pH 7.0, 1 mM EDTA, 1 mM PMSF, 10% glycerol, and 700 mM NaCl, and the primary purpose of elevating the salt concentration is to help dissociate peripheral membrane proteins.</li>
	<li>Spin down the cell lysate for 15 min at 15,000 x g. Pour the supernatant into polycarbonate bottles and spin for 1 h at 135,000 x g in an ultracentrifuge to collect membranes. 
	NOTE: If an ultracentrifuge is not available, membranes may be collected by spinning for 3 h at 53,000 x g, a force that is achievable in floor model centrifuges.</li>
	<li>Discard the supernatant and weigh the bottles with the membrane pellets. Resuspend the membrane pellets in approximately 35 mL membrane resuspension buffer containing 50 mM Tris pH 7.0, 500 mM NaCl, and 10% glycerol and add to a glass douncer. Weigh the empty centrifuge tubes to determine the mass of membranes harvested. 
	NOTE: In an efficient membrane harvest, the weight of membranes will be equal to approximately 20% the weight of cells used for that membrane harvest. This protocol gives typical cell yields of 40-45 g, and thus we likewise expect a yield of 8-9 g membranes.</li>
	<li>Dounce homogenize the membranes, and aliquot into 50 mL conical tubes that may now be stored indefinitely at -80 &#176;C. 
	NOTE: For this experiment, typically two aliquots are made, to allow two separate protein purifications to be performed from one original cell preparation.</li>
</ol>
4. Solubilization and Purification of Protein
<ol>
	<li>To a beaker add a stir bar and 150 mg of n-Dodecyl-&#946;-D-Maltopyranoside (DDM) per g membrane being used. Keep protein on the ice or at 4 &#176;C at all times. 
	NOTE: DDM is hygroscopic and should be stored in a glass jar with desiccant at -20 &#176;C when not in use. Also, it is typical to use between 4-5 g of membranes in a purification to obtain an appreciable amount of pure protein.</li>
	<li>Thaw membranes and bring to a final volume of 15 mL per g membrane in membrane resuspension buffer supplemented with 1 mM PMSF and 20 mM imidazole pH 8.0. Add membranes to the beaker with DDM and stir for 1 h at 4 &#176;C. 
	NOTE: The DDM concentration will be 1% w/v, but for the solubilization of membrane proteins it is more critical to be aware of the mass of detergent added per g membrane.</li>
	<li>Spin for 25 min at 135,000 x g at 4 &#176;C to pellet non-solubilized material. Filter the supernatant through a 5 &#181;m syringe filter.</li>
	<li>Load the sample by the peristaltic pump set to a flow rate of 1 mL/min onto a 1 mL immobilized nickel affinity column equilibrated in 20 mM Tris pH 7.0, 500 mM NaCl, 10% glycerol, 20 mM imidazole pH 8.0, and 0.05% DDM. 
	NOTE: For lower expressing proteins, improved purity and yields were observed by using a 1 mL instead of 5 mL column, and nickel instead of cobalt ions for affinity chromatography.</li>
	<li>Wash the column with 10 column volumes of a wash buffer containing 20 mM Tris pH 7.0, 500 mM NaCl, 10% glycerol, 80 mM imidazole pH 8.0, and 0.05% DDM. 
	NOTE: The imidazole concentration that can be used during washes for a 10-His-tagged protein is higher than is commonly appreciated and results in improved purity.</li>
	<li>Elute the protein in the buffer containing 20 mM Tris pH 7.0, 200 mM NaCl, 10% glycerol, 300 mM imidazole pH 8.0, and 0.05% DDM. Collect the eluate in ten 1 mL fractions and run on a 4-20% Tris-glycine SDS-PAGE gel along with solubilized lysate and wash fractions.</li>
	<li>Pool peak fractions, usually about 5-6 mL, and concentrate to 500 &#181;L or less volume in a 50 kDa cutoff concentrator in a benchtop centrifuge refrigerated at 4 &#176;C. 
	NOTE: 500 &#181;L is a typical maximum volume injected into size exclusion chromatography (SEC) columns. At this point the protein may be stored indefinitely at -80 &#176;C or continue onto SEC.</li>
	<li>Filter the protein through a 0.2 &#181;m spin column filter and inject onto a size exclusion column (e.g., Superdex-200) equilibrated in S200 buffer: 20 mM Mes pH 6.5, 100 mM NaCl, 2% glycerol, and 0.03% DDM. 
	NOTE: For the subsequent glutaraldehyde cross-linking assay, the buffering agent must not contain a primary amine. SEC is an opportunity to exchange buffers if necessary, as done here when exchanging Tris for Mes.</li>
	<li>Run the peak fractions on a 4-20% Tris-glycine SDS-PAGE gel. Stain the gel, collect pure peak fractions, and concentrate in a 50 kDa-cutoff concentrator at 4 &#176;C.</li>
	<li>Determine the protein concentration by measuring the absorbance at a wavelength of 280 nm, and store indefinitely at -80 &#176;C.</li>
</ol>
5. Glutaraldehyde Cross-linking Assay
<ol>
	<li>Prepare a 10 &#181;L reaction by mixing 3 &#181;L of 0.5 mg/mL protein in S200 buffer, 5 &#181;L of S200 buffer, 1 &#181;L of either water or 20% sodium dodecyl sulfate (SDS), followed by 1 &#181;L of 1.5% glutaraldehyde. 
	NOTE: This will give a 1x reaction of 0.15 mg/mL protein and 0.15% glutaraldehyde. Samples containing a pre-treatment of SDS are important negative controls and&#160;should be mixed and incubated at room temperature for 5 min before adding glutaraldehyde.</li>
	<li>Incubate the reaction for 30 min at room temperature. Terminate the reaction by adding 5&#181;L of 3x SDS-PAGE gel loading dye, which contains an excess of Tris buffer that quenches the glutaraldehyde.</li>
	<li>Load all 15 &#181;L onto 4-20% Tris-glycine SDS-PAGE gel, which will load 1.5 &#181;g protein per lane. Run the gel at 200 V for 30 min. Stain and determine the extent of dimer cross-linking by evidence of a band that runs at twice the size of the denatured monomer. 
	NOTE: A glutaraldehyde titration and time course may be performed first to find the best conditions for a homomeric transporter.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+of+Bor1+supports+an+elevator+transport+mechanism+for+SLC4+anion+exchangers.">Structure of Bor1 supports an elevator transport mechanism for SLC4 anion exchangers.</a> Proceedings of the National Academy of Sciences of the United States of America. 113 (38), 10542-10546 (2016).</li><li>Coudray, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+of+the+SLC4+transporter+Bor1p+in+an+inward-facing+conformation.">Structure of the SLC4 transporter Bor1p in an inward-facing conformation.</a> Protein Science. 26 (1), 130-145 (2016).</li><li>Arakawa, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystal+structure+of+the+anion+exchanger+domain+of+human+erythrocyte+band+3.">Crystal structure of the anion exchanger domain of human erythrocyte band 3.</a> Science. 350 (6261), 680-684 (2015).</li><li>Huynh, K. 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The authors have nothing to disclose.

Here we have shared detailed protocols that result in the purification to homogeneity of three distinct eukaryotic borate transporters. The protocols presented here are derived from other protocols for the expression of integral membrane proteins in S. cerevisiae1,14, and our optimizations result in both improved purity and improved yields. The parameters optimized here include cell culture growth volumes and times, bead-beating lysis procedures, buffer composition during cell lysis and protein purification, amount of detergent used per gram membrane, metal ion identify in affinity purification, volume of the affinity column, and the implementation of a cross-linking assay to evaluate homolog and detergent suitability by assessing the multimeric state of oligomeric transporters. The protocols are successful for multiple SLC4 homologs from the plant and fungal species. A limitation of all strategies for the purification of integral membrane proteins is that there exists no universal membrane protein purification method that is guaranteed to work. It is possible that our protocols are more likely to be successful for proteins closer in sequence identity and structure to SLC4 transporters, and thus members of the SLC4, SLC23, and SLC26 families could be promising targets15. Likewise, the more evolutionarily distant from the borate transporters a membrane transporter might be, the more likely the protocol will have to be different, such as by varying the expression system, detergent, or other key parameters4.
The protocol takes advantage of the most commonly used affinity tag in protein purification, the His-tag. Despite the presence of impurities in the initial eluted fractions, the combinations of nickel affinity, extensive washing, and subsequent SEC purification results in the highly purified protein. A 10-His-tag allows for the more stringent imidazole washes and thus can remove more background binding proteins than can be removed in washes permitted by 8-His- or 6-His-tags. Our selections for pure fractions err on the conservative side of the most highly pure gel fractions, which correspond to the peak SEC fractions. Final protein yields can be increased by pooling and concentrating more fractions, albeit with the trade-off of slightly less pure protein. The methods presented here enabled the purification of AtBor1 in quantities that led to the determination of its crystal structure7, with purification differences consisting of cutting off the His-tag and exchanging the transporter into a different detergent to improve crystal diffraction7.
Our protocol raises important considerations for the selection of homologs and the detergent used to solubilize and purify them. DDM is a common first choice for detergent because it is relatively mild, often successful at solubilizing, and has been used in structural and functional studies of a wide variety of membrane proteins. In determining whether a detergent is a poor selection for a membrane, a common method of evaluation is whether the protein gives a single monodisperse peak on an SEC chromatogram, rather than a large peak in the void volume or a range of polydisperse peaks, which indicate misfolded or unstable protein. A more subtle consideration is raised by our purification of ScBor1. It purifies in large quantities and looks favorable and monodisperse on an SEC chromatogram, which suggests it could be an attractive target for structural studies. However, our cross-linking assay reveals that it is a monomer when purified. While it is possible that ScBor1 could natively exist as a monomer&#160;in the cell, studies indicate that SLC4 transporters and their homologs are likely to be dimers7,8,9,10. Our development of the cross-linking assay occurred after crystallizing and solving the structure of AtBor1. However, had we been able to evaluate ScBor1 in comparison with AtBor1 with the cross-linking assay, valuable time and research efforts could have been redirected to the pursuit of AtBor1 instead of ScBor1, the latter of which ultimately was not successful in crystallization and diffraction experiments. This assay can thus help researchers distinguish between either homologs or detergents to use when pursuing structural studies in order to prioritize conditions in which the protein maintains its suspected native conformation. Additionally, the assay can be used to probe which amino acids are critical for multimerization, in order to find amino acid substitutions that destabilize the multimerization interface and lead to obligate monomers. Such an approach has been used to probe the functional significance of homomeric assembly in membrane transporters16,17,18.
A general advantage of our method is that yeast has been shown to enable the expression of many challenging membrane proteins of diverse function1. Additionally, its low cost and growth times promote its accessibility for many research efforts that may be less able to use expression strategies that require tissue culture or expensive growth media. The procedures presented here are relatively inexpensive and can be performed in one week which underscores the feasibility of the approach. Implementation of these protocols can help enable structural and functional studies of other challenging membrane proteins.

The difficulty in obtaining sufficient quantities of purified membrane protein remains a critical bottleneck in the pursuit of structural and functional studies of receptors, ion channels, and transporters. Many protocols exist for&#160;moderately high throughput pipelines to screen and find candidate membrane proteins that express well enough to enable subsequent in vitro studies1,2,3,4. Typically, proteins are tagged with an N- or C-terminal green fluorescent protein (GFP), and expression levels are monitored either by in-gel fluorescence or by fluorescence-detection size exclusion chromatography (FSEC)5. Such approaches enable the triaging of membrane protein candidates into high expressing proteins, moderate or low expressing proteins, or proteins that express poorly or not at all. This approach works well when the experimental design is to investigate large numbers of candidate genes with the intention of selecting whichever protein expresses the best. However, in some cases, an experimental approach is predicated on studying a particular membrane protein, which can be challenging when that protein&#39;s expression levels are in the moderate or low range. Additionally, sometimes in these cases, expression levels can be only minimally increased by altering the construct via truncations, thermostabilizing mutations, or codon optimization. It is, therefore, sometimes necessary to optimize membrane protein expression and purification protocols for membrane proteins that express only moderately well.
The SLC4 family of transporters includes the bicarbonate transporter Anion Exchanger 1 (also known as Band 3), the most abundant membrane protein in red blood cells6, and a key driver of cellular respiration. The SLC4 family is homologous with borate transporters, which are critical in plants and have been shown to exhibit a similar structure to Anion Exchanger 1 as well as to sodium-coupled SLC4 transporters7,8,9,10. Here we report optimized protein expression and purification protocols using S. cerevisiae to purify three different borate transporters found in yeast and plants. We highlight in detail key steps we took to optimize the yield and efficiency of the purifications to homogeneity. Additionally, we report a chemical cross-linking assay using glutaraldehyde to monitor the multimeric assembly of these transporters in the context of a purified protein-detergent-lipid complex. The cross-linking experiment can help evaluate homolog and detergent suitability by assessing the multimeric state of oligomeric transporters after purification.
This protocol assumes that the desired transporter has been cloned into a 2 &#181; derived plasmid under inducible control of the GAL1 promoter for expression in S. cerevisiae. For these procedures, DNA was used encoding full-length wild-type borate transporters Saccharomyces cerevisiae Bor1 (ScBor1)11, Arabidopsis thaliana Bor1 (AtBor1)12, and Oryza sativa Bor3 (OsBor3)13. The C-terminus in each construct is appended with a 10-His-tag and a thrombin cleavage site inserted between the transporter and the 10-His-tag to enable its removal if desired. The protocol will also assume that the plasmid has already been transformed into the DSY-5 expression strain of S. cerevisiae on complete supplemental selective media (CSM) lacking histidine.

<table><tbody><tr><td>2-mercaptoethanol</td><td>Sigma-Aldrich</td><td>M3148</td><td></td></tr><tr><td>4-Morpholinoethanesulfonic acid hydrate (MES)</td><td>Acros</td><td>172591000</td><td></td></tr><tr><td>&amp;Auml;kta Pure 25 L FPLC</td><td>GE Healthcare</td><td>29018224</td><td></td></tr><tr><td>Amicon Ultra 15 mL, 50 kDa MWCO concentrator</td><td>Millipore</td><td>UFC905024</td><td>for concentrating nickel column fractions</td></tr><tr><td>Amicon Ultra 4 mL, 50 kDa MWCO concentrator</td><td>Millipore</td><td>UFC805024</td><td>for concentrating S200 fractions</td></tr><tr><td>Bacto Peptone</td><td>BD Diagnostics</td><td>211677</td><td></td></tr><tr><td>Bacto Yeast Extract</td><td>BD Diagnostics</td><td>288620</td><td></td></tr><tr><td>Glass Erlenmeyer flask, 2L</td><td>Sigma-Aldrich</td><td>CLS44442L</td><td></td></tr><tr><td>Benchtop centrifuge</td><td>Sorvall</td><td>Legend RT</td><td></td></tr><tr><td>Bottle top filter</td><td>Nalgene</td><td>595-4520</td><td>the membrane is removed and used to filter glass beads</td></tr><tr><td>Bromophenol blue</td><td>Sigma-Aldrich</td><td>114391</td><td></td></tr><tr><td>Complete supplement mixture without histidine</td><td>Sunrise Science</td><td>1006-100</td><td></td></tr><tr><td>D-Galactose</td><td>Sigma-Aldrich</td><td>G0750</td><td></td></tr><tr><td>D-Glucose</td><td>Sigma-Aldrich</td><td>RDD016</td><td></td></tr><tr><td>Ethanol, 200 proof</td><td>Pharmco-Aaper</td><td>111000200</td><td></td></tr><tr><td>Ethylenediaminetetraacetic Acid (EDTA)</td><td>Sigma-Aldrich</td><td>EDS-500G</td><td></td></tr><tr><td>Gel tank SDS-PAGE system</td><td>Bio-Rad</td><td>1658004</td><td></td></tr><tr><td>Glass bead-beating cell disruptor</td><td>BioSpec</td><td>1107900</td><td></td></tr><tr><td>Glass beads, 0.5mm</td><td>BioSpec</td><td>11079105&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;</td><td></td></tr><tr><td>Glutaraldehyde</td><td>Electron Microscopy Sciences</td><td>16019</td><td>sent as an 8% solution under nitrogen</td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>G7893</td><td></td></tr><tr><td>HiTrap IMAC FF column, 1 mL</td><td>GE Healthcare</td><td>17-0921-02</td><td>charged with nickel</td></tr><tr><td>Hydrochloric acid</td><td>Sigma-Aldrich</td><td>258148</td><td></td></tr><tr><td>Imidazole</td><td>Acros</td><td>12202</td><td></td></tr><tr><td>Instant Blue gel stain</td><td>Expedeon</td><td>ISB1L</td><td></td></tr><tr><td>JA-10 rotor</td><td>Beckman</td><td>369687</td><td></td></tr><tr><td>JA-14 rotor</td><td>Beckman</td><td>339247</td><td></td></tr><tr><td>Microcentrifuge tubes, 1.5mL</td><td>Thermo Scientific</td><td>3451</td><td></td></tr><tr><td>Microcentrifuge, refrigerated</td><td>Fisher</td><td>13-100-676</td><td></td></tr><tr><td>Mini-Protean Tris-glycine gels, 4-20%</td><td>Bio-Rad</td><td>456-1096</td><td></td></tr><tr><td>Minipuls 3 peristaltic pump</td><td>Gilson</td><td>F155005</td><td>used for loading lysate onto affinity column</td></tr><tr><td>n-Dodecyl-beta-D-Maltopyranoside (DDM)</td><td>Inalco</td><td>1758-1350</td><td></td></tr><tr><td>Nickel(II) Sulfate Hexahydrate</td><td>Sigma-Aldrich</td><td>227676</td><td></td></tr><tr><td>Orbital shaking incubator with temperature control</td><td>New Brunswick</td><td>C24</td><td></td></tr><tr><td>p423 GAL1 plasmid with borate transporter insert</td><td></td><td></td><td>available from authors</td></tr><tr><td>Phenylmethanesulfonyl fluoride (PMSF)</td><td>Acros</td><td>215740050</td><td></td></tr><tr><td>Polycarbonate ultracentrifuge tubes</td><td>Beckman</td><td>355618</td><td></td></tr><tr><td>Polypropylene bottles, 250mL</td><td>Beckman</td><td>356011</td><td></td></tr><tr><td>Polypropylene bottles, 500mL</td><td>Beckman</td><td>355607</td><td></td></tr><tr><td>Precision Plus Protein Kaleidoscope Standards</td><td>Bio-Rad</td><td>1610375</td><td></td></tr><tr><td>&lt;em&gt;S. cerevisiae&lt;/em&gt; expression strain DSY-5</td><td></td><td></td><td>available from authors</td></tr><tr><td>Sodium chloride</td><td>Sigma-Aldrich</td><td>S7653</td><td></td></tr><tr><td>Sodium dodecyl sulfate</td><td>Sigma-Aldrich</td><td>L3771</td><td></td></tr><tr><td>Sodium hydroxide</td><td>Sigma-Aldrich</td><td>S8045</td><td></td></tr><tr><td>Spin column with 0.2 &amp;micro;m filter, 0.5mL</td><td>Millipore</td><td>UFC30GV0S</td><td>for filtering protein before injecting onto S200 column</td></tr><tr><td>Sterile Falcon tubes, 15mL</td><td>Lab Depot</td><td>TLD431696</td><td></td></tr><tr><td>Sterile Falcon tubes, 50mL</td><td>Lab Depot</td><td>TLD431698</td><td></td></tr><tr><td>Superdex 200 10/300 GL column</td><td>GE Healthcare</td><td>17-5175-01</td><td>used for SEC purification</td></tr><tr><td>Syringe filter, 5&amp;micro;m</td><td>Pall</td><td>4650</td><td>for filtering lysate before loading affinity column</td></tr><tr><td>Tris-base</td><td>Sigma-Aldrich</td><td>T1503</td><td></td></tr><tr><td>Type 70 Ti rotor</td><td>Beckman</td><td>337922</td><td></td></tr><tr><td>Ultracentrifuge</td><td>Beckman</td><td>L8-70M</td><td></td></tr><tr><td>YNB+Nitrogen without amino acids</td><td>Sunrise Science</td><td>1501-500</td><td></td></tr></tbody></table>

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.

Typical gels for the eluted fractions from performing nickel affinity chromatography show all three proteins partially purified (Figure 1A). To the right of the ladder is the lysate, which in each case does not show a significant band corresponding to the borate transporter which is typical for proteins that do not overexpress well. The 80 mM wash lane shows minimal loss of 10-His-tagged protein despite the relatively high imidazole concentration. Bands corresponding to the borate transporters are readily apparent in eluted fractions, and fractions deemed to contain the bulk of the eluted protein are concentrated before injecting onto the S200 gel filtration column. At this stage, the proteins are insufficiently purified for many downstream applications, as indicated by extra bands in the concentrated sample before injecting onto the S200 column (Figure 1B). However, injecting the sample onto the size exclusion column reveals eluted fractions of high purity (Figure 1B-C). Chromatograms and their corresponding gels for each of the borate transporters are presented. Despite the moderate purity of the samples upon elution from affinity chromatography, the protein is highly pure upon eluting from the gel filtration column. Critically, the chromatograms reveal each protein to migrate primarily as a single monodisperse peak with little protein retained in the void volume. A stable and folded protein generally gives a monodisperse and symmetrical peak, while unstable, misfolded, or aggregated protein will generally give either multiple asymmetric peaks or a large peak in the void volume. It is typical to obtain a final yield of approximately 2 mg purified protein per L of yeast culture for ScBor1, and approximately 1 mg each of purified AtBor1 and OsBor3 per L culture. These numbers are the amount of protein purified from one aliquot of 4-5 g of membranes, which originates from one half of our original cell preparation.
The cross-linking experiment shows that purified homomeric transporters can have their assembly in solution readily assessed (Figure 2). The purpose of the samples containing a pre-treatment of SDS is to show that cross-linking is dependent on a folded state in solution and that cross-linking therefore does not occur when multimerization is disrupted by a harsh detergent. The results shown distinguish that one of the three borate transporters, ScBor1, is not dimerized when purified in DDM in these conditions, while the other two, AtBor1 and OsBor3, cross-link and show dimerization. It is possible to see that not all protein may cross-link. In this example, trace amounts of OsBor3 monomer are visible, while AtBor1 cross-links to completion. The extent of cross-linking will depend on the number of lysine residues in proximity to one another, and it is possible that other cross-linking reagents may efficiently cross-link different membrane proteins.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59166/59166fig1.jpg" /> 
Figure 1. Nickel affinity and size exclusion chromatography purification for three borate transporters. Each vertical panel (A), (B), and (C) contains data for ScBor1, AtBor1, and OsBor3. (A) Nickel affinity column. M is a ladder of molecular weight markers with weights in kDa specified to the left. L is the crude lysate, and W is the 80 mM imidazole wash. All other numbered lanes correspond to 1 mL fractions eluted with 300 mM imidazole. Bars above fractions indicate which were selected to be combined and concentrated for injections onto the column. (B) P indicates concentrated protein before injecting onto the S200 column. Eluted SEC fractions are to the right, with brackets used to match gel lanes with their chromatogram fractions in (C). The void volume is just after 8 mL. <a href="https://www.jove.com/files/ftp_upload/59166/59166fig1large.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/59166/59166fig2.jpg" /> 
Figure 2. Cross-linking assay assesses purified transporters for multimeric assembly. A ladder with molecular weights is indicated on the left. Above all other lanes is shown which protein is present and whether the sample has had glutaraldehyde added or a pre-treatment of SDS before glutaraldehyde addition. Arrows indicate the positions of monomer and dimer for AtBor1 and OsBor3. ScBor1 is a smaller protein than AtBor1 and OsBor3 and runs as expected. <a href="https://www.jove.com/files/ftp_upload/59166/59166fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Expression, Solubilization, and Purification of Eukaryotic Borate Transporters at JoVE.com

Expression, Solubilization, and Purification of Eukaryotic Borate Transporters

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 ...

expression-solubilization-purification-eukaryotic-borate

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Biochemistry

9.6K Views.  Davidson College. 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.

Video: Expression, Solubilization, and Purification of Eukaryotic Borate Transporters

Thermostabilization, Expression, Purification, and Crystallization of the Human Serotonin Transporter Bound to S-citalopram

The serotonin transporter is a sodium and chloride-coupled transporter that &#34;pumps&#34; extracellular serotonin into cells. S-citalopram is a drug used to treat depression and anxiety by binding to the serotonin transporter with high-affinity, blocking serotonin reuptake. Here we report an efficient procedure and a set of tools to stabilize, express, purify, and crystallize serotonin transporter-antibody complexes bound to S-citalopram and other antidepressants. Mutations which stabilize the serotonin transporter were identified using an S-citalopram binding assay. Serotonin transporter expressed in baculovirus-transduced HEK293S GnTI- cells, was reconstituted into proteoliposomes and used to raise high-affinity antibodies. We have developed a strategy to discover antibodies that are useful for structural studies. A straightforward approach for the expression of antibody fragments in Sf9 cells has also been established. Transporter-antibody complexes purified using this procedure are well-behaved and readily crystallize, producing complexes with S-citalopram that diffract X-rays to 3-4 &#197; resolution. The strategies developed here can be utilized to determine the structure of other challenging membrane proteins.

Thermostabilization, Expression, Purification, and Crystallization of the Human Serotonin Transporter Bound to S-citalopram

This manuscript describes how to screen for thermostabilizing mutations, purify the human serotonin transporter, generate high affinity antibodies, and crystallize the serotonin transporter-antibody complex bound to the antidepressant drug S-citalopram. This protocol can be adapted to the study of other challenging membrane transporters, receptors, and channels.

The serotonin transporter is a sodium and chloride-coupled transporter that &#34;pumps&#34; extracellular serotonin into cells. S-citalopram is a drug ...

Expression and Purification of Mammalian Bestrophin Ion Channels

The human genome encodes four bestrophin paralogs, namely BEST1, BEST2, BEST3, and BEST4. BEST1, encoded by the BEST1 gene, is a Ca2+-activated Cl- channel (CaCC) predominantly expressed in retinal pigment epithelium (RPE). The physiological and pathological significance of BEST1 is highlighted by the fact that over 200 distinct mutations in the BEST1 gene have been genetically linked to a spectrum of at least five retinal degenerative disorders, such as Best vitelliform macular dystrophy (Best disease). Therefore, understanding the biophysics of bestrophin channels at the single-molecule level holds tremendous significance. However, obtaining purified mammalian ion channels is often a challenging task. Here, we report a protocol for the expression of mammalian bestrophin proteins with the BacMam baculovirus gene transfer system and their purification by affinity and size-exclusion chromatography. The purified proteins have the potential to be utilized in subsequent functional and structural analyses, such as electrophysiological recording in lipid bilayers and crystallography. Importantly, this pipeline can be adapted to study the functions and structures of other ion channels.

The purification of ion channels is often challenging, but once achieved, it can potentially allow in vitro investigations of the functions and structures of the channels. Here, we describe the stepwise procedures for the expression and purification of mammalian bestrophin proteins, a family of Ca2+-activated Cl- channels.

The human genome encodes four bestrophin paralogs, namely BEST1, BEST2, BEST3, and BEST4. BEST1, encoded by the BEST1 gene, is a Ca2+-activated Cl- ...

Modifying Baculovirus Expression Vectors to Produce Secreted Plant Proteins in Insect Cells

It has been a challenge for scientists to express recombinant secretory eukaryotic proteins for structural and biochemical studies. The baculovirus-mediated insect cell expression system is one of the systems used to express recombinant eukaryotic secretory proteins with some post-translational modifications. The secretory proteins need to be routed through the secretory pathways for protein glycosylation, disulfide bonds formation, and other post-translational modifications. To improve the existing insect cell expression of secretory plant proteins, a baculovirus expression vector is modified by the addition of either a GP67 or a hemolin signal peptide sequence between the promoter and multiple-cloning sites. This newly designed modified vector system successfully produced a high yield of soluble recombinant secreted plant receptor proteins of Arabidopsis thaliana. Two of the expressed plant proteins, the extracellular domains of Arabidopsis TDR and PRK3 plasma membrane receptors, were crystallized for X-ray crystallographic studies. The modified vector system is an improved tool that can potentially be used for the expression of recombinant secretory proteins in the animal kingdom as well.

Here, we present the protocols for utilizing the insect cell and baculovirus protein expression system to produce large quantities of plant secreted proteins for protein crystallization. A baculovirus expression vector has been modified with either GP67 or insect hemolin signal peptide for plant protein secretion expression in insect cells.

It has been a challenge for scientists to express recombinant secretory eukaryotic proteins for structural and biochemical studies. The ...

Expression, Isolation, and Purification of Soluble and Insoluble Biotinylated Proteins for Nerve Tissue Regeneration

Recombinant protein engineering has utilized Escherichia coli (E. coli) expression systems for nearly 4 decades, and today E. coli is still the most widely used host organism. The flexibility of the system allows for the addition of moieties such as a biotin tag (for streptavidin interactions) and larger functional proteins like green fluorescent protein or cherry red protein. Also, the integration of unnatural amino acids like metal ion chelators, uniquely reactive functional groups, spectroscopic probes, and molecules imparting post-translational modifications has enabled better manipulation of protein properties and functionalities. As a result this technique creates customizable fusion proteins that offer significant utility for various fields of research. More specifically, the biotinylatable protein sequence has been incorporated into many target proteins because of the high affinity interaction between biotin with avidin and streptavidin. This addition has aided in enhancing detection and purification of tagged proteins as well as opening the way for secondary applications such as cell sorting. Thus, biotin-labeled molecules show an increasing and widespread influence in bioindustrial and biomedical fields. For the purpose of our research we have engineered recombinant biotinylated fusion proteins containing nerve growth factor (NGF) and semaphorin3A (Sema3A) functional regions. We have reported previously how these biotinylated fusion proteins, along with other active protein sequences, can be tethered to biomaterials for tissue engineering and regenerative purposes. This protocol outlines the basics of engineering biotinylatable proteins at the milligram scale, utilizing&#160; a T7 lac inducible vector and E. coli expression hosts, starting from transformation to scale-up and purification.

Developing biotinylatable fusion proteins has many potential applications in various fields of research. Recombinant protein engineering is a straight forward procedure that is cost-effective, providing high yields of custom-designed proteins.

Recombinant protein engineering has utilized Escherichia coli (E. coli) expression systems for nearly 4 decades, and today E. coli is still the most ...

Bioengineering

Expression, Purification, and Liposome Binding of Budding Yeast SNX-BAR Heterodimers

SNX-BAR proteins are an evolutionarily conserved class of membrane remodeling proteins that play key roles in sorting and trafficking of protein and lipids during endocytosis, sorting within the endosomal system, and autophagy. Central to SNX-BAR protein function is the ability to form homodimers or heterodimers that bind membranes using highly conserved phox-homology (PX) and BAR (Bin/Amphiphysin/Rvs) domains. In addition, oligomerization of SNX-BAR dimers on membranes can elicit the formation of membrane tubules and vesicles and this activity is thought to reflect their functions as coat proteins for endosome-derived transport carriers. Researchers have long utilized in vitro binding studies using recombinant SNX-BAR proteins on synthetic liposomes or giant unilamellar vesicles (GUVs) to reveal the precise makeup of lipids needed to drive membrane remodeling, thus revealing their mechanism of action. However, due to technical challenges with dual expression systems, toxicity of SNX-BAR protein expression in bacteria, and poor solubility of individual SNX-BAR proteins, most studies to date have examined SNX-BAR homodimers, including non-physiological dimers that form during expression in bacteria. Recently, we have optimized a protocol to overcome the major shortcomings of a typical bacterial expression system. Using this workflow, we demonstrate how to successfully express and purify large amounts of SNX-BAR heterodimers and how to reconstitute them on synthetic liposomes for binding and tubulation assays.

Here, we present a workflow for the expression, purification and liposome binding of SNX-BAR heterodimers in yeast.

SNX-BAR proteins are an evolutionarily conserved class of membrane remodeling proteins that play key roles in sorting and trafficking of protein and ...

Genetics

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.

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.

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

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

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

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

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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

Strategies for Expressing and Purifying Solute Carrier Transporters

High-Throughput Expression and Purification of Human Solute Carriers for Structural and Biochemical Studies

Solute carriers (SLCs) are membrane transporters that import and export a range of endogenous and exogenous substrates, including ions, nutrients, metabolites, neurotransmitters, and pharmaceuticals. Despite having emerged as attractive therapeutic targets and markers of disease, this group of proteins is still relatively underdrugged by current pharmaceuticals. Drug discovery projects for these transporters are impeded by limited structural, functional, and physiological knowledge, ultimately due to the difficulties in the expression and purification of this class of membrane-embedded proteins. Here, we demonstrate methods to obtain high-purity, milligram quantities of human SLC transporter proteins using&#160;codon-optimized gene sequences. In conjunction with a systematic exploration of construct design and high-throughput expression, these protocols ensure the preservation of the structural integrity and biochemical activity of the target proteins. We also highlight critical steps in the eukaryotic cell expression, affinity purification, and size-exclusion chromatography of these proteins. Ultimately, this workflow yields pure, functionally active, and stable protein preparations suitable for high-resolution structure determination, transport studies, small-molecule engagement assays, and high-throughput in vitro screening.

Author Spotlight: Expression and Purification of Human Solute Carrier Transporters Using Codon-Optimized Genes 

Structural and biochemical studies of human membrane transporters require milligram quantities of stable, intact, and homogeneous protein. Here we describe scalable methods to screen, express, and purify human solute carrier transporters using codon-optimized genes.

Solute carriers (SLCs) are membrane transporters that import and export a range of endogenous and exogenous substrates, including ions, nutrients, ...

Budding Yeast Protein Extraction and Purification for the Study of Function, Interactions, and Post-translational Modifications

Homogenization by bead beating is a fast and efficient way to release DNA, RNA, proteins, and metabolites from budding yeast cells, which are notoriously hard to disrupt. Here we describe the use of a bead mill homogenizer for the extraction of proteins into buffers optimized to maintain the functions, interactions and post-translational modifications of proteins. Logarithmically growing cells expressing the protein of interest are grown in a liquid growth media of choice. The growth media may be supplemented with reagents to induce protein expression from inducible promoters (e.g. galactose), synchronize cell cycle stage (e.g. nocodazole), or inhibit proteasome function (e.g. MG132). Cells are then pelleted and resuspended in a suitable buffer containing protease and/or phosphatase inhibitors and are either processed immediately or frozen in liquid nitrogen for later use. Homogenization is accomplished by six cycles of 20 sec&#160;bead-beating (5.5 m/sec), each followed by one minute incubation on ice. The resulting homogenate is cleared by centrifugation and small particulates can be removed by filtration. The resulting cleared whole cell extract (WCE) is precipitated using 20% TCA for direct analysis of total proteins by SDS-PAGE followed by Western blotting. Extracts are also suitable for affinity purification of specific proteins, the detection of post-translational modifications, or the analysis of co-purifying proteins. As is the case for most protein purification protocols, some enzymes and proteins may require unique conditions or buffer compositions for their purification and others may be unstable or insoluble under the conditions stated. In the latter case, the protocol presented may provide a useful starting point to empirically determine the best bead-beating strategy for protein extraction and purification. We show the extraction and purification of an epitope-tagged SUMO E3 ligase, Siz1, a cell cycle regulated protein that becomes both sumoylated and phosphorylated, as well as a SUMO-targeted ubiquitin ligase subunit, Slx5.

The preparation of high quality yeast cell extracts is a necessary first step in the analysis of individual proteins or entire proteomes. Here we describe a fast, efficient, and reliable homogenization protocol for budding yeast cells that has been optimized to preserve protein functions, interactions, and post-translational modifications.

Homogenization by bead beating is a fast and efficient way to release DNA, RNA, proteins, and metabolites from budding yeast cells, which are notoriously ...

Expression, Solubilization, and Purification of Eukaryotic Borate Transporters

Summary

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Thermostabilization, Expression, Purification, and Crystallization of the Human Serotonin Transporter Bound to S-citalopram

Expression and Purification of Mammalian Bestrophin Ion Channels

Modifying Baculovirus Expression Vectors to Produce Secreted Plant Proteins in Insect Cells

Expression, Isolation, and Purification of Soluble and Insoluble Biotinylated Proteins for Nerve Tissue Regeneration

Expression, Purification, and Liposome Binding of Budding Yeast SNX-BAR Heterodimers

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

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

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

High-Throughput Expression and Purification of Human Solute Carriers for Structural and Biochemical Studies

Budding Yeast Protein Extraction and Purification for the Study of Function, Interactions, and Post-translational Modifications