Name: rapid assessment of membrane protein quality by fluorescent size exclusion chromatography
Uploaded: 2023-01-06T13:05:00.000+00:00
Duration: 6 min 26 s
Description: Watch this Scientific Journal Video about Rapid Assessment of Membrane Protein Quality by Fluorescent Size Exclusion Chromatography at JoVE.com

We would like to acknowledge the help and support of the whole Peak Proteins team. From the cell science team, we would like to acknowledge Ian Hampton for his valuable insights and guidance in insect cell expression. We would like to thank Mark Abbott for providing the resources and opportunity to pursue this project.

1. Detergent and buffer preparation for FSEC
<ol>
	<li>Prepare a detergent stock solution.
	<ol>
		<li>To prepare a 20 mL stock solution, weigh 4 g of dodecyl maltoside (DDM) powder and 0.4 g of cholesteryl hemisuccinate (CHS) powder, and make it to 20 mL with laboratory-grade distilled H2O.</li>
		<li>After adding all the components, mix with end-over-end inversion at 4 &#176;C until the components are completely solubilized. Overnight end-over-end mixing at 4 &#176;C is recommended.</li>
		<li>Aliquot and store the detergent stocks at &#8722;20 &#176;C until use. If the stock needs to be used immediately, store the detergent stock on ice. 
		NOTE: The standard detergent stock used in this work was a 20% (w/v) DDM and 2% (w/v) CHS mixture (see Table of Materials). Different detergents can be used (e.g., lauryl maltose neopentyl glycol; LMNG), or the use of detergent-free extraction with polymers such as styrene-maleic acid (SMA) can be tested. This needs to be decided when designing the experimental conditions to be tested.</li>
	</ol>
	</li>
	<li>Prepare a solubilization buffer.
	<ol>
		<li>Prepare a solubilization buffer by combining the correct weight or volume of the components to achieve a final concentration of 100 mM HEPES, 200 mM NaCl, 20 % (v/v) glycerol, and 1x protease inhibitor cocktail (see Table of Materials) in a beaker. 
		NOTE: In the present study, the preparation of 50 mL of solubilization buffer was sufficient for processing five samples.</li>
		<li>Add a 0.7 volume (e.g., 35 mL if making 50 mL of buffer) of laboratory-grade distilled H2O to the beaker.</li>
		<li>Mix on a magnetic stirrer, and using a pH meter, adjust the buffer pH to 7.5 by the dropwise addition of concentrated NaOH.</li>
		<li>Using a measuring cylinder, top up the buffer to the final required volume with laboratory-grade distilled H2O.</li>
	</ol>
	</li>
	<li>Prepare the SEC running buffer.
	<ol>
		<li>Prepare the SEC running buffer by combining the correct weight or volume of the components to achieve a final concentration of 100 mM HEPES, 150 mM NaCl, and 10% (v/v) glycerol in a beaker. 
		NOTE: In the present study, preparing 600 mL of SEC buffer was sufficient for running five samples.</li>
		<li>Add a 0.7 volume (e.g., 420 mL if making 600 mL of buffer) of laboratory-grade distilled H2O to the beaker.</li>
		<li>Mix on a magnetic stirrer, and using a pH meter, adjust the buffer pH to 7.5 by the dropwise addition of concentrated NaOH.</li>
		<li>Using a measuring cylinder, top up the buffer to the final required volume with laboratory-grade distilled H2O.</li>
		<li>Filter the SEC buffer through a bottle-top 0.45 &#181;m pore filter under a vacuum (see Table of Materials).</li>
		<li>Once the buffer has passed through the filter, degas it by leaving it under the vacuum until no more bubbles appear when shaken.</li>
		<li>Add a final concentration of 0.03% (w/v) DDM and 0.003% (w/v) CHS to the SEC buffer by adding the required volume of the detergent stock prepared in step 1 (e.g., 0.9 mL if making 600 mL of SEC buffer).</li>
		<li>Pre-chill the buffer before use. 
		&#8203;NOTE: Different buffers can be used depending on the conditions tested. For example, if testing the effect of different detergents on the protein of interest, a buffer with the same detergent as the one used to solubilize the protein would ideally need to be made. If testing detergent-free conditions with SMA, detergent should be omitted from the SEC buffer completely. However, see the discussion section for more details about protocol modifications for detergent screening.</li>
	</ol>
	</li>
</ol>
2. Sample preparation for FSEC
<ol>
	<li>Prepare the cell pellets. 
	NOTE: The starting point for the assay is harvesting the cell pellet from a suspension cell expression culture of the GFP-tagged (or other fluorescently labeled) protein of interest. The exact timings and conditions for harvest will depend on the protein being expressed, the cell line being used, the conditions that the cells that have been grown in, and the method by which protein expression has been induced. These details are beyond the scope of this protocol. In this study, 0.5-1 g of Sf21 cell pellet was used per condition to be tested, corresponding to 25-50 mL of culture 2-3 days after infection with approximately 4 x 106 viable cells/mL of a 1:20 dilution of P2 baculovirus. Please note that the protocol described here has been tested and found to work equally well with similar wet weights of cell pellets from other eukaryoticcell lines (e.g., HEK293E).
	<ol>
		<li>When the cells from the suspension cultures are ready to harvest, transfer 25-50 mL culture aliquots to 50 mL conical tubes.</li>
		<li>Counterbalance the tubes, and use a benchtop centrifuge in a swing-out bucket (see Table of Materials) at 2,000 x g for 15 min at room temperature to pellet the cells.</li>
		<li>Remove and discard the culture supernatant by gently tipping it away, or use a 50 mL pipet with a pipet filler if the cell pellet is particularly loose.</li>
		<li>If the cells will be used immediately for analysis, place the cell pellet on ice, and proceed directly to step 5. If the cells are to be saved for use at a later stage, freeze the cells by placing them at -80 &#176;C.</li>
		<li>If the cell pellet has been stored at -80 &#176;C, rapidly thaw it by incubating it at room temperature for 15 min or until the sample is no longer frozen. Move the sample immediately to ice following this step.</li>
	</ol>
	</li>
	<li>Resuspend and solubilize the sample.
	<ol>
		<li>Add 2 mL of the solubilization buffer (step 1.2) to the cell pellet.</li>
		<li>Incubate with end-over-end inversion at 4 &#176;C for 15-30 min until homogenous.</li>
		<li>Add premixed detergent stock (step 1.1) (e.g., 100 &#181;L of 20% DDM/2% CHS stock) for a final concentration of 1% DDM/0.1% CHS.</li>
		<li>Solubilize for 30 min with end-over-end inversion at 4 &#176;C. 
		&#8203;NOTE: If desirable, multiple conditions can be tested in parallel. The number of parallel samples that can be processed at once will depend on the available system for running the SEC experiment. In the setup described here, it was possible to process up to five samples at a time.</li>
	</ol>
	</li>
	<li>Perform a low-speed centrifugation step.
	<ol>
		<li>Centrifuge the sample in a pre-chilled (4 &#176;C) benchtop centrifuge in a swing-out bucket at 2,000 x g for 15 min.</li>
	</ol>
	</li>
	<li>Perform a high-speed centrifugation step.
	<ol>
		<li>Carefully transfer the supernatant from the low-speed centrifugation to ultra-centrifugation tubes (e.g., 0.5-2 mL tubes) by using a blunt-ended needle attached to a 5 mL syringe, being careful not to disturb the pellet from the low-speed spin.</li>
		<li>Balance pairs of tubes to within 0.05 g, and place them in a fixed-angle ultracentrifugation rotor (see Table of Materials).</li>
		<li>Centrifuge at 4 &#176;C for 30 min at 250,000 x g.</li>
	</ol>
	</li>
</ol>
3. Size exclusion chromatography (SEC)
<ol>
	<li>Prepare the fast protein liquid chromatography (FPLC) system, and equilibrate the column.
	<ol>
		<li>Prepare the system following the manufacturer&#39;s instructions (see Table of Materials), for example, by filling the system with SEC buffer and purging the pumps of air.</li>
		<li>Connect the SEC column to the FPLC, ensuring no air enters the column. This is accomplished by applying back pressure to the SEC column with a syringe filled with water (attached to the bottom of the column) in order to perform a drop-to-drop connection with the flow path of the FPLC system.</li>
		<li>Pre-equilibrate the SEC column by washing it first in 1.5 column volumes (36 mL for a 24 mL column) of laboratory-grade distilled and filtered H2O, followed by 1.5 column volumes of SEC buffer at the recommended flow rate and pressure for the column (see Table of Materials). 
		NOTE: The FPLC system used in this study to perform FSEC was in a cold environment and had a five-position loop valve and a six-position plate fraction collector fitted. This setup allows five samples to be loaded and run sequentially down the same column in an automated fashion without requiring manual intervention between the runs. For running the SEC experiments, a commercially available pre-packed 24 mL column was used (see Table of Materials), which contained a resin that allowed proteins in the molecular weight range of 10-600 kDa to be resolved. If the protein of interest is particularly large, an alternative column matrix could be used instead, allowing the separation of proteins up to 5,000 kDa in molecular weight. Please note the presence of detergent/lipid micelle will increase the overall size of the membrane protein by &#62;150 kDa, depending on the protein and detergent used.</li>
	</ol>
	</li>
	<li>Apply the sample to the column, and run the SEC experiment.
	<ol>
		<li>Transfer the supernatant from the high-speed centrifugation step to a 1 mL syringe using a blunt-ended needle attached to the syringe. This allows for sample recovery from the centrifuge tube without disturbing the pellet.</li>
		<li>Set the sample loop to load. Overfill a 500 &#181;L sample loop by injecting 600-700 &#181;L of the sample from the syringe into the loading port. Depending on the system used, this loading step can be programmed into the method to ensure no mistakes are made.</li>
		<li>During the method, inject the sample from the loop into the column by emptying it with 4 mL of SEC buffer at the recommended flow rate and pressure for the column (see Table of Materials).</li>
		<li>Run the column at the same flow rate until 1.5 column volumes (36 mL for a 24 mL column) of the buffer have passed down.</li>
		<li>At 0.25 of the column volume (6 mL for a 24 mL column), start collecting 0.2 mL fractions in order to collect 90 fractions. 
		&#8203;NOTE: As the void volume of the column is expected to be 0.3 column volumes, beginning the collection of fractions immediately prior to this ensures that the elution of all the protein is monitored, including any protein present in the void volume.</li>
	</ol>
	</li>
</ol>
4. Fluorescent trace collection and analysis
<ol>
	<li>Transfer the samples from the fraction collector (step 3.2.5) to a 96-well plate, and read the fluorescent signal.
	<ol>
		<li>Before taking a fluorescence reading, dilute the collected samples. Using a multi-channel pipette, transfer 90 &#181;L of laboratory-grade distilled H2O from a reservoir to each well of an opaque flat bottom 96-well plate (see Table of Materials).</li>
		<li>If fractions were collected into a 96-well block during step 3.2.5, use a multi-channel pipette to transfer 10 &#181;L of the SEC fractions from the block to the opaque flat bottom 96-well plate, and mix by pipetting up and down. Otherwise, if SEC fractions were collected into individual tubes during step 3.2.5, transfer 10 &#181;L of each fraction to the opaque flat bottom 96-well plate one by one, pipetting up and down each time to mix the samples.</li>
		<li>Place the opaque flat bottom 96-well plate into the plate reader, and measure the fluorescence. If GFP is the fluorescent label (used here), set the excitation as close as possible to 488 nm, and detect the fluorescent emission as close as possible to 507 nm. 
		NOTE: The dilution required before taking the fluorescence reading will depend on the total amount of protein of interest present in the expression culture and the sensitivity of the plate reader being used. In the examples shown in this study, the samples were diluted 10-fold in water. As a suggestion for a starting point, the fractions should be diluted 5-10-fold in water or buffer before detection. If the recorded FSEC trace signal is particularly low, smaller dilutions or even an undiluted sample can be used instead. The device used for detection in this study was a plate reader capable of excitation at 488 nm and detecting fluorescent emission at 507 nm (see Table of Materials).</li>
	</ol>
	</li>
	<li>Plot the FSEC traces.
	<ol>
		<li>Export the data from the plate reader in a raw format (raw text or a comma-separated values file). These raw data will be plotted on the Y-axis of the FSEC trace.</li>
		<li>For the X-axis, calculate the volume at which each fraction was collected. The first well needs to be the elution volume at which the first fraction was collected to account for the fractionation delay (6 mL in this example). The fractions after this must sequentially increase by the collected fraction volume (0.2 mL in this example).</li>
		<li>Once the X-axis and Y-axis data have been calculated, copy and paste the data into the graphing software (see Table of Materials) in order to plot the fluorescent signal in each well against the volume at which it eluted from the column. 
		NOTE: Figure 1 shows a schematic representation of the steps required to run an FSEC experiment.</li>
	</ol>
	</li>
	<li>Analyze the FSEC traces.
	<ol>
		<li>Assess the amount of protein elution at the void volume (approximately 8 mL for a 24 mL column), which indicates that the protein is very large in size and is likely unfolded/aggregated.</li>
		<li>Assess the amount of protein eluting in any later peaks, which indicate folded protein. This is expected to be between 10 mL and 16 mL for a 24 mL column depending on the protein size (when associated with a detergent/lipid micelle). Pay close attention to the peak shape, particularly if it is a broad or split peak spanning greater than 3-4 mL (for a 24 mL column), as this indicates a polydisperse sample.</li>
		<li>Calculate the ratio between the monomeric peak height and the void peak height by dividing the maximum signal recorded in the monomer peak by the maximum signal in the void peak. 
		NOTE: This value represents the monodispersity index and allows a quantitative measure of protein quality; larger values indicate the best possible quality, whereas values below 1 indicate problematic samples, as they have more aggregated protein than folded protein.</li>
		<li>If multiple FSEC runs are to be compared and the most important feature is the amount of monomer in each case, plot the traces as the raw signal recorded by the plate reader (e.g., RFU). 
		NOTE: If a comparison of the amount of monomer compared to the unfolded protein is more important, the signal must be normalized to a percentage of the total signal using the minimum and maximum readings in the trace, which will accentuate differences in the ratio between aggregated and monomeric protein between the traces.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64322/64322fig01.jpg" /> 
Figure 1: Schematic representation of the steps required to run an FSEC experiment. (1) Cells that express the fluorescently tagged protein of interest are solubilized. (2) The crude solubilization is clarified first with a low-speed spin, followed by (3) a high-speed spin. (4) The clarified sample supernatant is loaded and run on an appropriate SEC column, and (5) the fractions are collected. (6) Samples of the fractions are transferred to a 96-well plate, and a GFP-fluorescent signal is detected using a plate reader to (7) plot the FSEC trace. <a href="https://www.jove.com/files/ftp_upload/64322/64322fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Peak Proteins is a contract research organization that provides protein expression, purification, mass-spectrometry, and structural determination for a fee.

The generic systematic approaches for condition screening with FSEC that are presented here allow the fast optimization of solubilization and purification parameters for the production of membrane proteins. This means that stable and functionally active membrane proteins can be rapidly produced for biophysical and structural studies. Furthermore, FSEC can be run using laboratory equipment that is likely already in place in membrane protein labs, and thus, there is no requirement for the purchase of a specialist instrument for running the assays.
Critical steps 
The time taken between the point of solubilization from cells in detergent to the point at which the sample is passed down the SEC column (steps 2.1.5-3.2.5) are time critical, and there must be no pauses between these steps. All the steps should be conducted at 4 &#730;C or on ice, and the time taken to perform these steps needs to be kept to the minimum possible. These time and temperature constraints are necessary in order to record the FSEC profile for the membrane protein before any potential unfolding or degradation. After the membrane protein has been solubilized, there is a greater risk of unfolding, aggregation, and degradation, even at 4 &#730;C. Ideally, any samples for which the FSEC traces are to be compared should pass down the SEC column in the same length of time after the solubilization step. In practice, this is difficult, particularly if the samples are passed sequentially down a single column, but it is possible to collect up to five SEC traces within 3 h of one another, and in this time frame, there should not be significant degradation.
Troubleshooting 
If, on performing the FSEC experiment, there is low or no fluorescent signal, it is possible the membrane protein of interest has not expressed the chosen cell line, has very low expression of the chosen cell line, or has not been solubilized in the chosen detergent. If the samples were diluted before collecting the fluorescence signal and recording the FSEC trace, a simple first step would be to try a lower dilution or no dilution of the SEC fractions. If this still does not yield an interpretable FSEC trace, the expression and solubilization of the protein should be checked.
The analysis of protein expression can be achieved by checking the fluorescence of the sample after step 2.2.2. If there is a very low or no fluorescent signal from this sample (e.g., a signal very close to the background), there is likely an issue with the protein expression. Steps can be taken to improve the expression levels of the membrane protein, such as switching to an alternate cell line or adjusting the growth conditions, the induction of expression, and the time between the induction/infection/transfection and the harvest. However, particularly poor protein expression can indicate an unstable membrane protein and, thus, a poor construct choice.
If the expression has been checked and there is a clear fluorescent signal above the background prior to FSEC, the solubilization efficiency can be checked by measuring the remaining fluorescent signal of the sample after step 2.4.3 (soluble membrane protein) in comparison to the sample after step 2.2.2 (total protein). It is common for the solubilization efficiency to be 20%-30% and still allow for the successful analysis and purification of the membrane protein. However, if the solubilization efficiency is less than 20%, a different detergent for solubilization or different solubilization conditions may be required. If attempts to improve the solubilization are not successful, this can indicate a particularly unstable membrane protein and, thus, a poor construct choice.
If a very late eluting peak is observed in the FSEC trace (e.g., 18-24 mL), this indicates that the fluorescent protein has a protein molecular weight that is much lower than expected. This can be caused by the membrane protein of interest being degraded, resulting in &#34;free&#34; GFP. One should check if the protein is intact before and after solubilization using in-gel GFP fluorescence. If the protein of interest does appear to be degrading or being proteolyzed, the amount of protease inhibitor can be increased twofold to fourfold. However, a high sensitivity to proteases or degraded protein even before solubilization can indicate a particularly unstable protein and, thus, a poor construct choice.
Modifications and further applications of FSEC 
Commonly, the fluorescent tag that is used in FSEC is GFP or eGFP, as described in this protocol. However, many different fluorescent protein tags are available. The choice of the fluorescent tag to be used depends on having a plate reader that can achieve the correct excitation and emission parameters to record the fluorescent signal for the selected fluorescent tag and having a fluorophore with little to no change in quantum yield in different environmental conditions. Furthermore, FSEC is not restricted to fluorescent proteins but can also work equally well with a protein that has been labeled with a fluorescent dye. For example, an NTA dye could be used, which would favorably bind to histidine-tagged membrane protein constructs. Furthermore, either a fluorescently labeled antibody chemically labeled with a fluorescent dye and specific for binding the membrane protein of interest or a purification tag included in the membrane protein construct could indirectly label a target for FSEC.
When performing detergent screening using FSEC, a choice can be made regarding whether the buffer used to run the SEC column should contain the matching detergent that the protein has been solubilized in or whether a standard detergent should be used across all the runs. A more accurate representation of the behavior of the protein will be obtained if the whole experiment is performed with the matching detergent throughout. However, it can be time-consuming and wasteful of detergent if the column must be re-equilibrated in a new detergent before each run is carried out. Furthermore, as the main purpose of detergent screening is to compare traces, the trends will remain in the traces even if the conditions are not ideal. Thus, a compromise can be reached whereby the protein is solubilized in the detergent of interest but the column is run in a standard buffer with a single detergent across all the runs (e.g., DDM)11, which can save time and detergent consumables.
By modifying the FLPC equipment used, the throughput of the FSEC protocol can be significantly increased, and the sample requirement can be minimized. For example, an FPLC or HPLC system could be equipped with an autosampler, a smaller bed volume analytical column (such as a 3.2 mL analytical SEC column), and an in-line fluorescent detector for monitoring continuous FSEC traces directly from the column. The resulting setup would allow more FSEC runs to be carried out in a shorter period of time and remove the manual plotting step, thus allowing a greater number of conditions to be tested in a shorter time frame. Furthermore, the sample requirement would be further reduced, as fewer samples would have to be prepared and loaded onto the FSEC column for each run. This would open up possibilities for reducing the expression cultures to a plate-based format, as such little material would be required for the analysis.
Strengths and weaknesses of the FSEC compared to other methods 
A disadvantage of FSEC is that the membrane protein constructs need to be designed to introduce the fluorescent label, and on introduction, there is a small possibility that the placement of the label could interfere with the function or folding of the membrane protein of interest. In addition, the FSEC protocol, as described here, monitors the characteristics of a membrane protein in the presence of cell lysate, which is a crude mixture of proteins. The behavior of a membrane protein in this environment may be different than when the membrane protein of interest is subjected to a preparative SEC column at the end of purification when fully isolated from other proteins. Furthermore, FSEC provides a somewhat qualitative measure of protein quality. However, by converting the FSEC trace to a monodispersity index, as described in step 4.3.3 of the protocol, a quantitative measure of protein quality can be obtained.
FSEC is not the only method that can be used in the early analysis of membrane protein constructs, solubilization conditions, and purification buffer composition. The alternative approaches have both advantages and disadvantages over FSEC. For example, fluorophore-based thermostability assays exist, particularly the use of the dye 7-diethylamino-3-(4&#8242;-maleimidylphenyl)-4-methylcoumarin (CPM)16,17. The advantage of this method is that, unlike FSEC, which provides a qualitative measure of protein quality, thermostability assays provide a quantitative measure in the form of a relative melting temperature. Furthermore, there is no requirement to introduce a fluorescent tag on the protein construct. However, the disadvantages of thermostability assays compared with FSEC are that purified protein must be used and that the assay is not compatible with all protein constructs, as it relies on the advantageous positions of native cysteine residues in the folded protein.
Another method that has similarities to both FSEC and fluorophore-based thermostability assays is an assay that measures the temperature sensitivity of a membrane protein. In this assay, the protein is challenged with different temperatures, and the protein that remains in solution after centrifugation is detected. Detection in this method has been conducted in several ways, including measuring the fluorescence in solution18, the fluorescence of an SDS-PAGE gel band19, or the signal intensity in a western blot20. However, a significant disadvantage of these approaches is that the assay is very labor-intensive and prone to high noise in the results, as each individual temperature point must be collected independently.
Finally, several more advanced biophysical techniques can be used to assess membrane protein quality in a similar manner to FSEC, for example, flow-induced dispersion analysis21, microscale thermophoresis22, or SPR. Although very powerful approaches, the disadvantage of these methods is the requirement for highly specialized instruments to run the analyses.
In conclusion, FSEC provides an invaluable tool for use in membrane protein production campaigns, and although it is not the only option, it has several distinct advantages over other methods, as listed above. The cross-validation of the results by orthogonal assays is always recommended, and none of the methods discussed above are mutually exclusive of one another.

Size exclusion chromatography (SEC), also known as gel filtration chromatography, is commonly used in protein science1. During SEC, proteins are separated based on their hydrodynamic radius, which is a function of the protein size and shape2. In brief, this separation is achieved by applying the protein samples under flow to a packed bed of porous beads that act as a molecular sieve. The beads used are often cross-linked agarose with a defined range of pore sizes to allow proteins to either enter or be excluded from the pores of the beads3,4,5,6,7. Proteins with smaller hydrodynamic radii spend a greater proportion of time within the pores and, thus, flow through the packed bed at a slower rate, whereas larger proteins spend a greater proportion of time outside the beads (the excluded volume) and move through the packed bed at a faster rate. SEC can be used as a protein purification step when a preparative column is used1. When an analytical column is used, SEC can be used to analyze the protein quality and properties2. For example, protein aggregates that may be present in a sample and indicate poor quality protein tend to be very large, meaning they only travel in the excluded volume and, thus, are eluted from the column at the earliest point; this volume is referred to as the column void or void volume. Furthermore, molecular weight standards can be used to calibrate the column, allowing an estimated molecular weight of the protein of interest to be interpolated from a standard curve.
Typically, the protein absorbance at 280 nm is used to monitor the protein elution from a size exclusion column. This restricts the use of SEC as an analysis tool until the protein of interest is largely free from contaminating proteins, for example, at the last step of protein purification. However, fluorescent SEC (FSEC) utilizes a protein of interest that is fluorescently labeled. Therefore, a fluorescent signal can be used to specifically monitor the elution of the protein of interest in the presence of other proteins or even crude mixtures8,9. Furthermore, as fluorescent signals are highly sensitive, successful analysis can be performed on samples with extremely low protein quantities. The protein of interest is often fluorescently labeled by including a green fluorescent protein (GFP) or enhanced GFP (eGFP) tag in the expression construct. The fluorescent signal can then be monitored by excitation at 395 nm or 488 nm and detecting the fluorescent emission at 509 nm or 507 nm for GFP or eGFP, respectively10.
The benefit of using a fluorescent signal to monitor protein elution from an SEC column makes FSEC a valuable tool for analyzing membrane protein samples when the expression levels are particularly poor in comparison to soluble proteins. Crucially, the quality and properties of membrane proteins can be analyzed directly following solubilization from crude lysates without the requirement to optimize the purification process first11,12. For these reasons, FSEC can be used to rapidly analyze the membrane protein quality while exploring the different factors that may be required to improve the behavior of the membrane protein in solution. For example, it is common to trial numerous constructs containing different tags, truncations, deletions, fusion partner insertions, and stabilizing mutations to find one that is not aggregated after extraction from the membrane13,14. Furthermore, buffer screening to determine the detergent, additive, ligand, or polymer that provides the most stabilizing condition for the membrane protein can define the best buffer composition for protein purification or for providing stability for downstream uses, such as biophysical assays or structural characterization.
Thus, the overall goal of the FSEC method is to collect an SEC column elution profile for a target membrane protein of interest. Furthermore, as fluorescence is used, this SEC trace is collected at the earliest possible point in the optimization of the constructs and conditions prior to any lengthy purification. The FSEC trace can be used as a comparative tool to judge the likelihood of success of purifying a membrane protein with different buffer conditions or membrane protein constructs. In this way, the collection of FSEC profiles can be used as a quick iterative process to arrive at optimal construct design and buffer composition prior to spending effort generating the quantities of pure protein required for other analysis methods.

<table><tbody><tr><td>1 mL and 5 mL plastic syringes&amp;nbsp;</td><td>Generic&amp;nbsp;</td><td>-&amp;nbsp;</td><td>Syringes for transfer of samples&amp;nbsp;</td></tr><tr><td>10x EDTA Free Protease inhibitor cocktail</td><td>Abcam</td><td>ab201111</td><td>Protease inhibitors</td></tr><tr><td>15 mL tubes&amp;nbsp;&amp;nbsp;</td><td>Generic&amp;nbsp;</td><td>-&amp;nbsp;</td><td>15 mL tubes for pellet preparation and solubilisation&amp;nbsp;&amp;nbsp;</td></tr><tr><td>2 mL ultra-centrifuge tubes</td><td>Beckman Coulter&amp;nbsp;</td><td>344625&amp;nbsp;</td><td>Tubes for ultra-centrifuge rotor&amp;nbsp;</td></tr><tr><td>50 mL tubes</td><td>Generic&amp;nbsp;</td><td>-&amp;nbsp;</td><td>50 mL tubes for cell harvest&amp;nbsp;</td></tr><tr><td>96 deep-well blocks&amp;nbsp;</td><td>Greiner</td><td>15922302</td><td>For collecting 0.2 mL SEC fractions&amp;nbsp;</td></tr><tr><td>&amp;Auml;KTA&amp;nbsp; V9-L loop valve</td><td>Cytiva&amp;nbsp;</td><td>29011358</td><td>5 posiiton loop valve&amp;nbsp; for the &amp;Auml;KTA FPLC system</td></tr><tr><td>&amp;Auml;KTA F9-C fraction collector</td><td>Cytiva&amp;nbsp;</td><td>29027743</td><td>6 position plate fraction collector for the &amp;Auml;KTA FPLC system</td></tr><tr><td>&amp;Auml;KTA pure 25 L</td><td>Cytiva&amp;nbsp;</td><td>29018224&amp;nbsp;</td><td>FPLC system for running the experiment&amp;nbsp;</td></tr><tr><td>Benchtop centrifuge (e.g. Fisherbrand GT4 3L)&amp;nbsp;&amp;nbsp;</td><td>Fisher Scientific&amp;nbsp;</td><td>15828722&amp;nbsp;</td><td>Centrifuge for low-speed spin&amp;nbsp;</td></tr><tr><td>Blunt end filling needles&amp;nbsp;</td><td>Generic&amp;nbsp;</td><td>-&amp;nbsp;</td><td>For transfer of samples&amp;nbsp;</td></tr><tr><td>Bottle top vacuum filter&amp;nbsp;</td><td>Corning</td><td>10005490</td><td>Bottle top vacuum filter for filtering SEC buffers</td></tr><tr><td>Cholesteryl hemisuccinate (CHS)&amp;nbsp;</td><td>Generon&amp;nbsp;</td><td>CH210-5GM&amp;nbsp;</td><td>Additive for detergent solubilisation&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Disposable multichannel reseviour&amp;nbsp;</td><td>Generic&amp;nbsp;</td><td>-&amp;nbsp;</td><td>Resevior for addition of water or buffer to 96-well micro-plate</td></tr><tr><td>Dodecyl maltoside (DDM)&amp;nbsp;</td><td>Glycon&amp;nbsp;</td><td>D97002-C-25g&amp;nbsp;</td><td>Detergent for solubilisation&amp;nbsp;</td></tr><tr><td>eGFP protein standards</td><td>BioVision</td><td>K815-100</td><td>eGFP standards for fluorescent calibration curve&amp;nbsp;</td></tr><tr><td>Glycerol</td><td>Thermo Scientific&amp;nbsp;</td><td>11443297</td><td>Glycerol for buffer preparation</td></tr><tr><td>HEPES</td><td>Thermo Scientific&amp;nbsp;</td><td>10411451</td><td>HEPES for buffer preparation</td></tr><tr><td>High molecular weight SEC calibration standards kit</td><td>Cytiva&amp;nbsp;</td><td>28403842</td><td>Molecular weight calibration kit for SEC</td></tr><tr><td>Lauryl maltose neopentylglycol (LMNG)&amp;nbsp;</td><td>Generon&amp;nbsp;</td><td>NB-19-0055-5G&amp;nbsp;</td><td>Detergent for solubilisation&amp;nbsp;</td></tr><tr><td>Low molecular&amp;nbsp; weight SEC calibration standards kit</td><td>Cytiva&amp;nbsp;</td><td>28403841</td><td>Molecular weight calibration kit for SEC</td></tr><tr><td>MLA-130 ultra-centrifuge rotor</td><td>Beckman Coulter&amp;nbsp;</td><td>367114&amp;nbsp;</td><td>Rotor for ultracentrifuge that fits 2 mL capacity tubes</td></tr><tr><td>Opaque 96-well flat-bottom micro-plate&amp;nbsp;</td><td>Corning</td><td>10656853</td><td>96-well for reading fluorescent signal in plate reader&amp;nbsp;</td></tr><tr><td>Optima MAX-XP ultra-centrifuge</td><td>Beckman Coulter&amp;nbsp;</td><td>393315&amp;nbsp;</td><td>Centrifuge for high-speed spin&amp;nbsp;</td></tr><tr><td>pH meter</td><td>Generic&amp;nbsp;</td><td>-&amp;nbsp;</td><td>For adjusting the pH of buffers during preparation</td></tr><tr><td>Prism</td><td>GraphPad</td><td>-&amp;nbsp;</td><td>Graphing software for plotting traces</td></tr><tr><td>Rotary mixer&amp;nbsp;</td><td>Fisher Scientific&amp;nbsp;</td><td>12027144&amp;nbsp;</td><td>Mixer for end over end mixing in the cold&amp;nbsp;</td></tr><tr><td>Sodium chloride</td><td>Fisher Scientific&amp;nbsp;</td><td>10316943</td><td>Sodium chloride for buffer preparation</td></tr><tr><td>Sodium hydroxide</td><td>Fisher Scientific&amp;nbsp;</td><td>10488790</td><td>Sodium hydroxide for buffer preparation</td></tr><tr><td>Spectramax ID3 Plate Reader</td><td>Molecular Devices</td><td>735-0391</td><td>Micro-plate reader capable of reading fluorescence</td></tr><tr><td>Stirrer plate</td><td>Generic&amp;nbsp;</td><td>-&amp;nbsp;</td><td>For stirring buffers during preparation</td></tr><tr><td>Styrene maleic acid (SMA)&amp;nbsp;</td><td>Orbiscope&amp;nbsp;</td><td>SMALP 300&amp;nbsp;</td><td>Polymer for detergent free extraction&amp;nbsp;</td></tr><tr><td>Superdex 200 Increase 10/300 GL&amp;nbsp;</td><td>Cytiva&amp;nbsp;</td><td>28990944&amp;nbsp;</td><td>SEC column for running the experiment. The bed volume of this column is 24 mL. The recommended flow rate for this column in 0.9 ml/min (in water at 4 &amp;deg;C). The maximum pressure limit for this column is 5 MPa.</td></tr><tr><td>Vacuum pump</td><td>Sartorius</td><td>16694-2-50-06</td><td>For filtering and degassing buffers</td></tr></tbody></table>

rapid assessment of membrane protein quality by fluorescent size exclusion chromatography

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

First, the dynamic range and lower limits of eGFP detection for the plate reader used in this study were investigated. A purified eGFP standard of known concentration was diluted in a 50 &#181;L final volume to 50 ng&#183;&#181;L&#8722;1, 25 ng&#183;&#181;L&#8722;1, 12.5 ng&#183;&#181;L&#8722;1, 6.25 ng&#183;&#181;L&#8722;1, 3.125 ng&#183;&#181;L&#8722;1, 1.5625 ng&#183;&#181;L&#8722;1, 0.78125 ng&#183;&#181;L&#8722;1, and 0.390625 ng&#183;&#181;L&#8722;1, and the fluorescence was read using an excitation of 488 nm and an emission of 507 nm (Figure 2). This experiment indicated that the plate reader had a lower detection limit of 30 ng of eGFP-labeled protein per well and a dynamic range of up to 500 ng of eGFP-labeled protein per well before signal saturation. Using the value for the lower limit and assuming that the protein elution is confined to 0.33 of the column volume, as little as 1.28 &#181;g of eGFP labeled protein of interest is required for SEC column loading in order for a detectable FSEC signal to be observed.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64322/64322fig02.jpg" /> 
Figure 2: eGFP standard curve. Scatter graph of the fluorescent signal for purified eGFP standard diluted to 50 ng&#183;&#181;L&#8722;1, 25 ng&#183;&#181;L&#8722;1, 12.5 ng&#183;&#181;L&#8722;1, 6.25 ng&#183;&#181;L&#8722;1, 3.125 ng&#183;&#181;L&#8722;1, 1.5625 ng&#183;&#181;L&#8722;1, 0.78125, and 0.390625 ng&#183;&#181;L&#8722;1. (A) All dilutions are displayed, including those with the saturated signal. (B) A zoomed-in scatter graph including only the standards that fall into the dynamic range of the plate reader. <a href="https://www.jove.com/files/ftp_upload/64322/64322fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Secondly, the 24 mL column used for this study was calibrated with molecular weight standards. Using the same buffer and running conditions as used for the FSEC analysis, the molecular weight standards blue dextran (&#62;2,000 kDa), ferritin (440 kDa), aldolase (158 kDa), conalbumin (75 kDa), and ovalbumin (43 kDa) were individually injected and run through the column, and elution traces were collected at 280 nm absorption. The elution volumes recorded were 8.9 mL, 12.4 mL, 15.2 mL, 16.9 mL, and 18 mL, respectively. When these elution volumes were converted to Kav (Equation 1) and plotted against log molecular weights, a standard curve could be fit. This allowed the molecular weight of the GPCRs tested in this study to be estimated by interpolation of the standard curve (Figure 3). For example, the FSEC trace of the GPCR serotonin receptor 2A (5HT2AR) after solubilization in the detergent DDM indicated an elution volume of 13.4 mL. This 5HT2AR elution volume falls between those elution volumes recorded for ferritin and aldolase and provides an estimated molecular weight of approximately 300 kDa. The 5HT2AR construct used in this study is approximately 50 kDa (including the eGFP tag), meaning that if 5HT2AR is assumed to be monomeric, 250 kDa of molecular weight could be attributed to the DDM detergent/lipid micelle. The equation for the conversion of elution volumes is as follows (Equation 1):
<img alt="Equation 1" src="/files/ftp_upload/64322/64322eq01.jpg" style="margin: auto;" />&#160; &#160; &#160;(Equation 1)
where Ve&#160;is the elution volume, V0 is the column void volume, and Vc is the total column volume.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64322/64322fig03.jpg" /> 
Figure 3: Calibration curve of the SEC column using molecular weight standards. (A) A representative FSEC trace of 5HT2AR solubilized in DDM, with the relative elution positions of the molecular weight standards blue dextran (Void), ferritin, aldolase, conalbumin, and ovalbumin marked. Ferritin, aldolase, conalbumin, and ovalbumin are colored in green, purple, red, and cyan, respectively. (B) The molecular weight calibration curve using the elution positions of the standards after conversion to Kav (Equation 1) plotted against the log molecular weight (Mr). The Mr of 5HT2AR in DDM was interpolated from the curve using Kav and is displayed on the curve (blue square). <a href="https://www.jove.com/files/ftp_upload/64322/64322fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
FSEC was then used to assess the quality and properties of the GPCR sphingosine-1-phosphate receptor (S1PR1)15. Insect cells expressing GFP-tagged human S1PR1 were processed for FSEC as described in the protocol (Figure 1).
Firstly, the optimal membrane extraction conditions were explored by testing the detergents DDM and LMNG against detergent-free extraction with SMA (Figure 4A). The monodispersity index was used to assess the quality of the protein sample and the ratio between the protein in the void (~8 mL column retention) compared with the monodispersed sample (14-15 mL column retention). The sample solubilized in LMNG displayed a superior FSEC profile with a better monomer peak shape and a lower protein aggregate peak, indicating that solubilization and purification in LMNG were the most stabilizing conditions for this membrane protein. In contrast, the sample solubilized in DDM had a relatively poorer FSEC profile, with a larger aggregate peak and a broader monomeric peak, indicating polydispersity in the sample.
Secondly, the effect of ligand addition on the FSEC profile was investigated by adding sphingosine-1-phosphate (S1P) to the sample during solubilization. In this instance, DDM was used as the solubilization reagent, and the FSEC traces of S1PR1 in the presence and absence of S1P were compared (Figure 4B). The sample solubilized in the presence of S1P showed a superior FSEC trace with reduced aggregation. This indicated that purification in the presence of a ligand was advantageous for improving the protein sample quality by stabilizing the receptor in solution but was also advantageous also as a surrogate marker of protein activity, as the results suggested the protein was correctly folded and competent for ligand binding.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64322/64322fig04.jpg" /> 
Figure 4: FSEC using a crude extract of S1PR1 indicating optimal membrane extraction conditions and ligand binding. (A) Comparison of the FSEC traces of S1PR1 solubilized in styrene-maleic acid co-polymer (SMA; blue), lauryl maltose neopentyl glycol (LMNG; red), or dodecyl maltoside (DDM; black). (B) Comparison of the FSEC traces of S1PR1 solubilized in DDM in the presence (red) or absence (black) of the agonist sphingosine-1-phosphate (S1P). <a href="https://www.jove.com/files/ftp_upload/64322/64322fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
FSEC was also used to investigate the long-term stability of 5HT2AR under different conditions. GFP-tagged human 5HT2AR was solubilized from insect cell membranes in either detergent (DDM) or SMA polymer and analyzed by FSEC at several time points after storage at 4 &#176;C or room temperature (Figure 5). Following several days of incubation, 5HT2AR in DDM displayed a significant drop in the monomeric peak height at either temperature, and significant increases in the aggregate peak were observed. In contrast, 5HT2AR in SMA lipid particle (SMALP) did not show a significant drop in the monomeric peak height over the course of the experiment, indicating that the protein in SMALP remained stable for longer, even at unfavorable temperatures. This may be important when considering protein preparations for downstream biophysical applications such as surface plasmon resonance (SPR) experiments for which there is a requirement for the sample to be stable and active over a long time period for the successful completion of the binding experiments.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64322/64322fig05.jpg" /> 
Figure 5: Effect of time and temperature on the quality of 5HT2AR extracted from the membrane using either DDM or SMALP, as analyzed by FSEC. (A) Representative FSEC traces of 5HT2AR solubilized in DDM and stored at room temperature for 1 day (grey), 2 days (green), or 5 days (blue). (B) Histograms of the normalized monomeric peak height for DDM samples stored at 4 &#176;C (blue) or RT (green) compared to SMALP samples stored at 4 &#176;C (grey) or RT (black). Error bars are representative of the SEM. <a href="https://www.jove.com/files/ftp_upload/64322/64322fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Rapid Assessment of Membrane Protein Quality by Fluorescent Size Exclusion Chromatography at JoVE.com

Rapid Assessment of Membrane Protein Quality by Fluorescent Size Exclusion Chromatography

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

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

rapid-assessment-membrane-protein-quality-fluorescent-size-exclusion

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Biochemistry

2.8K Views.  Peak Proteins Ltd.. The present protocol describes a procedure to perform fluorescent size exclusion chromatography (FSEC) on membrane proteins to assess their quality for downstream functional and structural analysis. Representative FSEC results collected for several G-protein coupled receptors (GPCRs) under detergent-solubilized and detergent-free conditions are presented.

Video: Rapid Assessment of Membrane Protein Quality by Fluorescent Size Exclusion Chromatography

Ion Exchange Chromatography (IEX) Coupled to Multi-angle Light Scattering (MALS) for Protein Separation and Characterization

Ion-exchange chromatography with multi-angle light scattering (IEX-MALS) is a powerful method for protein separation and characterization. The combination of the high-specificity separation technique IEX with the accurate molar mass analysis achieved by MALS allows the characterization of heterogeneous protein samples, including mixtures of oligomeric forms or protein populations, even with very similar molar masses. Therefore, IEX-MALS provides an additional level of protein characterization and is complementary to the standard size-exclusion chromatography with multi-angle light scattering (SEC-MALS) technique.
Here we describe a protocol for a basic IEX-MALS experiment and demonstrate this method on bovine serum albumin (BSA). IEX separates BSA to its oligomeric forms allowing a molar mass analysis by MALS of each individual form. Optimization of an IEX-MALS experiment is also presented and demonstrated on BSA, achieving excellent separation between BSA monomers and larger oligomers. IEX-MALS is a valuable technique for protein quality assessment since it provides both fine separation and molar mass determination of multiple protein species that exist in a sample.

Ion Exchange Chromatography IEX Coupled to Multi-angle Light Scattering MALS for Protein Separation and Characterization

This protocol describes the use of high-specificity ion-exchange chromatography with multi-angle light scattering for an accurate molar mass determination of proteins, protein complexes, and peptides in a heterogeneous sample. This method is valuable for quality assessment, as well as for the characterization of native oligomers, charge variants, and mixed-protein samples.

Ion-exchange chromatography with multi-angle light scattering (IEX-MALS) is a powerful method for protein separation and characterization. The combination ...

Characterization of Proteins by Size-Exclusion Chromatography Coupled to Multi-Angle Light Scattering (SEC-MALS)

Analytical size-exclusion chromatography (SEC), commonly used for the determination of the molecular weight of proteins and protein-protein complexes in solution, is a relative technique that relies on the elution volume of the analyte to estimate molecular weight. When the protein is not globular or undergoes non-ideal column interactions, the calibration curve based on protein standards is invalid, and the molecular weight determined from elution volume is incorrect. Multi-angle light scattering (MALS) is an absolute technique that determines the molecular weight of an analyte in solution from basic physical equations. The combination of SEC for separation with MALS for analysis constitutes a versatile, reliable means for characterizing solutions of one or more protein species including monomers, native oligomers or aggregates, and heterocomplexes. Since the measurement is performed at each elution volume, SEC-MALS can determine if an eluting peak is homogeneous or heterogeneous and distinguish between a fixed molecular weight distribution versus dynamic equilibrium. Analysis of modified proteins such as glycoproteins or lipoproteins, or conjugates such as detergent-solubilized membrane proteins, is also possible. Hence, SEC-MALS is a critical tool for the protein chemist who must confirm the biophysical properties and solution behavior of molecules produced for biological or biotechnological research. This protocol for SEC-MALS analyzes the molecular weight and size of pure protein monomers and aggregates. The data acquired serve as a foundation for further SEC-MALS analyses including those of complexes, glycoproteins and surfactant-bound membrane proteins.

Characterization of Proteins by Size-Exclusion Chromatography Coupled to Multi-Angle Light Scattering SEC-MALS

This protocol describes the combination of size exclusion chromatography with multi-angle light scattering (SEC-MALS) for absolute characterization of proteins and complexes in solution. SEC-MALS determines the molecular weight and size of pure proteins, native oligomers, heterocomplexes and modified proteins such as glycoproteins.

Analytical size-exclusion chromatography (SEC), commonly used for the determination of the molecular weight of proteins and protein-protein complexes in ...

Native Cell Membrane Nanoparticles System for Membrane Protein-Protein Interaction Analysis

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal transduction; from sensing environmental signals to biological response; from metabolic regulation to developmental control. Accurate structural information of protein-protein interactions is crucial for understanding the molecular mechanisms of membrane protein complexes and for the design of highly specific molecules to modulate these proteins. Many in vivo and in vitro approaches have been developed for the detection and analysis of protein-protein interactions. Among them the structural biology approach is unique in that it can provide direct structural information of protein-protein interactions at the atomic level. However, current membrane protein structural biology is still largely limited to detergent-based methods. The major drawback of detergent-based methods is that they often dissociate or denature membrane protein complexes once their native lipid bilayer environment is removed by detergent molecules. We have been developing a&#160;native cell membrane nanoparticle system for membrane protein structural biology. Here, we demonstrate the use of this system in the analysis of protein-protein interactions on the cell membrane with a case study of the oligomeric state of AcrB.

Presented here is a protocol for the determination of oligomeric state of membrane proteins that utilizes a native cell membrane nanoparticle system in conjunction with electron microscopy.

Protein-protein interactions in cell membrane systems play crucial roles in a wide range of biological processes- from cell-to-cell interactions to signal ...

Assessing Two-dimensional Crystallization Trials of Small Membrane Proteins for Structural Biology Studies by Electron Crystallography

Electron crystallography has evolved as a method that can be used either alternatively or in combination with three-dimensional crystallization and X-ray crystallography to study structure-function questions of membrane proteins, as well as soluble proteins. Screening for two-dimensional (2D) crystals by transmission electron microscopy (EM) is the critical step in finding, optimizing, and selecting samples for high-resolution data collection by cryo-EM. Here we describe the fundamental steps in identifying both large and ordered, as well as small 2D arrays, that can potentially supply critical information for optimization of crystallization conditions.
By working with different magnifications at the EM, data on a range of critical parameters is obtained. Lower magnification supplies valuable data on the morphology and membrane size. At higher magnifications, possible order and 2D crystal dimensions are determined. In this context, it is described how CCD cameras and online-Fourier Transforms are used at higher magnifications to assess proteoliposomes for order and size.
While 2D crystals of membrane proteins are most commonly grown by reconstitution by dialysis, the screening technique is equally applicable for crystals produced with the help of monolayers, native 2D crystals, and ordered arrays of soluble proteins. In addition, the methods described here are applicable to the screening for 2D crystals of even smaller as well as larger membrane proteins, where smaller proteins require the same amount of care in identification as our examples and the lattice of larger proteins might be more easily identifiable at earlier stages of the screening.

Evaluating two-dimensional (2D) crystallization trials for the formation of ordered membrane protein arrays is a highly critical and difficult task in electron crystallography. Here we describe our approach in screening for and identifying 2D crystals of predominantly small membrane proteins in the range of 15 &#x2013; 90kDa.

Electron crystallography has evolved as a method that can be used either alternatively or in combination with three-dimensional crystallization and X-ray ...

Biology

High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in plasma membrane damage. In response to these damages, complex molecular machineries rapidly reseal the plasma membrane to restore its barrier function and maintain cell survival. Despite 60 years of research in this field, we still lack a thorough understanding of the cell resealing machinery. With the goal of identifying cellular components that control plasma membrane resealing or drugs that can improve resealing, we have developed a fluorescence-based high-throughput assay that measures the plasma membrane resealing efficiency in mammalian cells cultured in microplates. As a model system for plasma membrane damage, cells are exposed to the bacterial pore-forming toxin listeriolysin O (LLO), which forms large 30-50 nm diameter proteinaceous pores in cholesterol-containing membranes. The use of a temperature-controlled multi-mode microplate reader allows for rapid and sensitive spectrofluorometric measurements in combination with brightfield and fluorescence microscopy imaging of living cells. Kinetic analysis of the fluorescence intensity emitted by a membrane impermeant nucleic acid-binding fluorochrome reflects the extent of membrane wounding and resealing at the cell population level, allowing for the calculation of the cell resealing efficiency. Fluorescence microscopy imaging allows for the enumeration of cells, which constitutively express a fluorescent chimera of the nuclear protein histone 2B, in each well of the microplate to account for potential variations in their number and allows for eventual identification of distinct cell populations. This high-throughput assay is a powerful tool expected to expand our understanding of membrane repair mechanisms via screening for host genes or exogenously added compounds that control plasma membrane resealing.

Here we describe a high-throughput fluorescence-based assay that measures the plasma membrane resealing efficiency through fluorometric and imaging analyses in living cells. This assay can be used for screening drugs or target genes that regulate plasma membrane resealing in mammalian cells.

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in plasma membrane damage. In ...

Immunology and Infection

Size Exclusion Chromatography to Analyze Bacterial Outer Membrane Vesicle Heterogeneity

The cell wall of Gram-negative bacteria consists of an inner (cytoplasmic) and outer membrane (OM), separated by a thin peptidoglycan layer. Throughout growth, the outer membrane can bleb to form spherical outer membrane vesicles (OMVs). These OMVs are involved in numerous cellular functions including cargo delivery to host cells and communication with bacterial cells. Recently, the therapeutic potential of OMVs has begun to be explored, including their use as vaccines and drug delivery vehicles. Although OMVs are derived from the OM, it has long been appreciated that the lipid and protein cargo of the OMV differs, often significantly, from that of the OM. More recently, evidence that bacteria can release multiple types of OMVs has been discovered, and evidence exists that size can impact the mechanism of their uptake by host cells. However, studies in this area are limited by difficulties in efficiently separating the heterogeneously sized OMVs. Density gradient centrifugation (DGC) has traditionally been used for this purpose; however, this technique is time-consuming and difficult to scale-up. Size exclusion chromatography (SEC), on the other hand, is less cumbersome and lends itself to the necessary future scale-up for therapeutic use of OMVs. Here, we describe a SEC approach that enables reproducible separation of heterogeneously sized vesicles, using as a test case, OMVs produced by Aggregatibacter actinomycetemcomitans, which range in diameter from less than 150 nm to greater than 350 nm. We demonstrate separation of &#34;large&#34; (350 nm) OMVs and &#34;small&#34; (&#60;150 nm) OMVs, verified by dynamic light scattering (DLS). We recommend SEC-based techniques over DGC-based techniques for separation of heterogeneously sized vesicles due to its ease of use, reproducibility (including user-to-user), and possibility for scale-up.

Bacterial vesicles play important roles in pathogenesis and have promising biotechnological applications. The heterogeneity of vesicles complicates analysis and use; therefore, a simple, reproducible method to separate varying sizes of vesicles is necessary. Here, we demonstrate the use of size exclusion chromatography to separate heterogeneous vesicles produced by Aggregatibacter actinomycetemcomitans.

The cell wall of Gram-negative bacteria consists of an inner (cytoplasmic) and outer membrane (OM), separated by a thin peptidoglycan layer. Throughout ...

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 ...

Membrane Transport Processes Analyzed by a Highly Parallel Nanopore Chip System at Single Protein Resolution

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable efforts have been made in this field, but techniques enabling automated high-throughput transport analysis in combination with solvent-free lipid bilayer techniques required for the analysis of membrane transporters are rare. This class of transporters however is crucial in cell homeostasis and therefore a key target in drug development and methodologies to gain new insights desperately needed.
The here presented manuscript describes the establishment and handling of a novel biochip for the analysis of membrane protein mediated transport processes at single transporter resolution. The biochip is composed of microcavities enclosed by nanopores that is highly parallel in its design and can be produced in industrial grade and quantity. Protein-harboring liposomes can directly be applied to the chip surface forming self-assembled pore-spanning lipid bilayers using SSM-techniques (solid supported lipid membranes). Pore-spanning parts of the membrane are freestanding, providing the interface for substrate translocation into or out of the cavity space, which can be followed by multi-spectral fluorescent readout in real-time. The establishment of standard operating procedures (SOPs) allows the straightforward establishment of protein-harboring lipid bilayers on the chip surface of virtually every membrane protein that can be reconstituted functionally. The sole prerequisite is the establishment of a fluorescent read-out system for non-electrogenic transport substrates.
High-content screening applications are accomplishable by the use of automated inverted fluorescent microscopes recording multiple chips in parallel. Large data sets can be analyzed using the freely available custom-designed analysis software. Three-color multi spectral fluorescent read-out furthermore allows for unbiased data discrimination into different event classes, eliminating false positive results.
The chip technology is currently based on SiO2 surfaces, but further functionalization using gold-coated chip surfaces is also possible.

The presented protocol describes the analysis of membrane protein mediated transport on the single transporter level using pore-spanning solvent-free lipid bilayers. This is achieved by the creation of bulk produced nanopore array chips, combined with highly parallel data acquisition and analysis, enabling the future establishment of membrane protein effector screenings.

Membrane protein transport on the single protein level still evades detailed analysis, if the substrate translocated is non-electrogenic. Considerable ...

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 ...

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

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

Source: Bird, L. E., et al. Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli. J. Vis. Exp. (2015)
In this ...

Rapid Assessment of Membrane Protein Quality by Fluorescent Size Exclusion Chromatography

Summary

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Chapters in this video

Ion Exchange Chromatography (IEX) Coupled to Multi-angle Light Scattering (MALS) for Protein Separation and Characterization

Characterization of Proteins by Size-Exclusion Chromatography Coupled to Multi-Angle Light Scattering (SEC-MALS)

Native Cell Membrane Nanoparticles System for Membrane Protein-Protein Interaction Analysis

Assessing Two-dimensional Crystallization Trials of Small Membrane Proteins for Structural Biology Studies by Electron Crystallography

High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells

Size Exclusion Chromatography to Analyze Bacterial Outer Membrane Vesicle Heterogeneity

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

Membrane Transport Processes Analyzed by a Highly Parallel Nanopore Chip System at Single Protein Resolution

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

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