Name: combining wet and dry lab techniques to guide the crystallization of large coiled-coil containing proteins
Uploaded: 2017-01-06T10:30:00.000+00:00
Duration: 11 min 14 s
Description: Watch this Scientific Journal Video about Combining Wet and Dry Lab Techniques to Guide the Crystallization of Large Coiled-coil Containing Proteins at JoVE.com

This work was supported by grant NIH R01 GM097204 (APV and JDH). Funding for JHM was supplied by an HHMI Undergraduate Research Summer Fellowship.

NOTE: A diagram of the conceptual framework or workflow is described in Figure 1 for reference. The protocol can be broken down into four stages: computational or sequence based predictions, protein expression and purification, biochemical characterization, and crystallization. The examples shown analyze Shroom SD2 domains and/or Shroom-Rock complexes, but can be utilized with any protein.
1. Use Established Web-based Tools to Generate Computational Predictions of Coiled-coil Domain Boundaries
<ol>
	<li>Collect an Evolutionarily Diverse Set of Sequence Homologs.
	<ol>
		<li>Go to www.uniprot.org and type in Shroom in the upper search bar.</li>
		<li>Select sequences to download by checking the box next to their accession number. Sequences with a star indicate sequences reviewed by Uniprot and are generally more reliable. Take care to ensure that all sequences are complete and accurate.</li>
		<li>Save the selected sequences by clicking the Download button.</li>
		<li>Align the collection of Shroom sequences using Clustal-Omega21 (http://www.ebi.ac.uk/Tools/msa/clustalo/). Click the &#8220;Browse&#8221; button to select the file with the Shroom sequences and then push Submit.</li>
		<li>After the alignment is complete, click the &#34;Download Alignment File&#34; button.</li>
		<li>Open the multiple sequence alignment generated in 1.1.5 using Jalview (File|Input Alignment|From file). Within the new alignment window, color by identity (Color|Percent Identity).</li>
	</ol>
	</li>
	<li>Calculate predictions of secondary structure, predictions of flexible or disordered sequence, and predicted coiled-coil regions within the protein(s) of interest.
	<ol>
		<li>Go to http://gpcr.biocomp.unibo.it/cgi/predictors/cc/pred_cchmm.cgi, to calculate Coiled-Coil predictions. Copy and Paste the sequence into the Sequence Box and click Submit. 
		NOTE: This analysis may take a couple of hours to complete. Since Shroom sequences are quite large, the sequences may need to be split into blocks of less than 1,000 amino acids to fit within the size constraint of the webserver. When analyzing Shrm2, it is recommended that the C-terminal 1,000 amino acids are used as this section contains the Shrm SD2 domain which is known to contain coiled-coil regions. When repeating this analysis with other proteins, it is recommended to use the full-length protein sequences when possible. If that is not an option, use the largest fragment possible or dissect the sequence based on known biochemical or functional data.</li>
		<li>Go to http://gpcr.biocomp.unibo.it/cgi/predictors/cc/pred_cchmm.cgi, to calculate Coiled-Coil predictions. Copy and Paste the sequence into the Sequence Box and click Submit.</li>
	</ol>
	</li>
	<li>Combine the results of steps 1.1 to 1.2.2 into a single comprehensive annotation of the Shroom sequence. 
	NOTE: It is highly encouraged to also include any and all relevant information that can be gleaned from the literature or other sources about protein function or purification at this stage.</li>
	<li>Using this comprehensive annotated sequence, predict domain boundaries (or a series of possible boundaries) for the protein, trying to maximize conservation and predicted structural features while minimizing the amount of disordered or flexible sequence. If known, the resulting protein should retain the functional properties of interest.</li>
</ol>
2. Express and Purify Proteins with the Domain Boundaries Identified in Section 1
NOTE: The goal of this section is to use a series of quick and easily quantifiable assays to screen hypothetical domain boundaries generated in Section 1.
<ol>
	<li>Using standard molecular biology techniques, generate an expression plasmid with the desired coding sequence in frame within the His10-mRuby2-XH2 plasmid (See Figure 3 for the vector map and additional details).</li>
	<li>Test expression levels for each expression plasmid using BL21(DE3) E. coli or other suitable expression strain in autoinduction media1 at room temperature for 18-24 hr.
	<ol>
		<li>Transform expression plasmids as well as an empty plasmid control into BL21(DE3) following the instructions that came with the cells or using robust transformation protocol, such as the one here (https://www.addgene.org/plasmid-protocols/bacterial-transformation/).</li>
		<li>From the freshly transformed plate, pick a single colony and grow a 5 ml starter culture at 37 &#176;C overnight in Lysogeny Broth (LB) media with 34 &#181;g/ml kanamycin.</li>
		<li>The following day, pellet the culture and wash the pellet with fresh LB.</li>
		<li>Add the starter culture to 50 ml of autoinduction media with 34 &#181;g/ml kanamycin. Allow the culture to grow to saturation at either room temperature or 37 &#176;C. Typically, grow for ~18-24 hr.</li>
		<li>Pellet cells and freeze the resulting cell pellets at -80 &#176;C indefinitely prior to purification.</li>
		<li>Compare whole cell extracts from each expression strain and the empty vector control by SDS-PAGE to determine the strain that most efficiently produces the desired fusion protein. For His10-mRuby2-Shroom SD2 fusion proteins, run 10% SDS-PAGE gels at 180 V for ~50 min or until the dye front reaches the bottom of the gel22.</li>
		<li>Stain gel with Coomassie Blue to visualize total cellular proteins within each strain23.</li>
	</ol>
	</li>
	<li>Perform an initial affinity chromatography step on each strain that successfully expressed mRuby2-fusion protein. Typically, perform purification steps 2.3.2-2.3.7 at room temperature, unless the protein will benefit from altered conditions such as purification at 4 &#176;C.
	<ol>
		<li>Thaw cell pellets from step 2.2.5.</li>
		<li>Resuspend each cell pellet in 1.5 ml of lysis buffer (10% glycerol, 500 mM NaCl, 40 mM imidazole, 20 mM Tris pH8.0, 1 mM beta-mercaptoethanol). Supplement the lysis buffer with protease inhibitors as appropriate.</li>
		<li>Add 30 &#181;l of 10 mg/ml lysozyme and incubate at room temperature for 20 min. Sonicate following the instructions for the instrument being used.</li>
		<li>Transfer into a 1.5 ml tube and centrifuge in a table top centrifuge for 30 min at 14,000 x g to pellet insoluble material or unlysed cells. Save both supernatant and pellet for subsequent analysis via SDS-PAGE.</li>
		<li>Perform nickel affinity purification, incubating the soluble fraction with 100 &#181;l of Ni-NTA beads. Incubate the beads with soluble lysate for 5-10 min, inverting several times to mix beads.</li>
		<li>Centrifuge at 800 x g for 30 sec in a tabletop centrifuge to pellet the beads. Follow this by washing the pellet 3-5 times with lysis buffer to clean off non-specific contaminants.</li>
		<li>Elute Ruby fusion protein from the beads with lysis buffer supplemented with 1 M imidazole.</li>
		<li>Use SDS-PAGE to compare the behavior of His-mRuby2-Shroom proteins in the &#34;quick and dirty&#34; purification above.</li>
	</ol>
	</li>
</ol>
3. Characterize Protein Sample to Identify Those with Advantageous Properties
<ol>
	<li>Use a spectrophotometer set at 280 nm to measure the concentration of the Ruby fusion protein, and then assess the homogeneity of the sample by loading 1-5 &#181;g of fusion protein onto a native PAGE gel24. Run the 10% Native PAGE gel at 4 &#176;C for 140 min at 175 V.
	<ol>
		<li>Image the native PAGE using an imager equipped to visualize fluorescence, observing where the mRuby-tagged fusion migrates in this assay. 
		NOTE: Be careful to observe fusion protein that is stuck in the well as this protein is likely aggregated. If a fluorescence imager is not available, one can often visualize and image concentrated samples of Ruby tagged protein under a black light or with a UV light box.</li>
	</ol>
	</li>
	<li>Perform limited proteolysis to identify stably folded domains. Incubate 95 &#181;l of Ruby-Shrm SD2 fusion at ~1 mg/ml with 5 &#181;l of 0.025% Subtilisin A.
	<ol>
		<li>Sample the reaction at time points of 0, 0.5, 2, 5, 15, 60, and 120 min, removing 10 &#181;l from the reaction for each time point and visualizing the progress of the reaction via SDS-PAGE using a 15% acrylamide gel.</li>
		<li>Stain with Coomassie Blue as in step 2.2.7 and evaluate whether a protease resistant species can be identified.</li>
	</ol>
	</li>
	<li>Integrate data from the purification efficiency, behavior in native PAGE, and limited proteolysis on the partially purified mRuby2-Shroom fusion proteins into a comprehensive assessment of the overall behavior of these proteins in solution.
	<ol>
		<li>(Optional) If a suitable functional assay is available, check for activity at this point.</li>
	</ol>
	</li>
</ol>
4. Producing High Quality Crystals of the Coiled-coil SD2 Domain from Shroom
NOTE: All steps within section 4.1 are performed at room temperature unless the protein would benefit from purification at a different temperature, usually 4 &#176;C.
<ol>
	<li>Using the 50 ml growths as a rough estimate of expression and purification potential, perform large scale expressions of mRuby2-Shroom SD2 with the goal of achieving 5-20 mg of completely purified sample. Typically 2 L of culture provides adequate starting material. After growth pellet cells by centrifugation at 8,000 x g for 10 min.
	<ol>
		<li>Resuspend the pellet from the 2 L growth in lysis buffer as before, using ~2 ml of lysis per gram of frozen cell pellet if using lysozyme for lysis. Alternatively, resuspend at ~8 ml/g of pellet if using a homogenizer or French press. Pellet cellular debris via centrifugation at 30,000 x g for 30 min.</li>
		<li>Batch bind the soluble fraction from 4.1.1 using 10 ml of Ni-NTA resin. Pour into appropriate gravity column and allow unbound proteins to drain off.</li>
		<li>Wash resin 3-5 times with 40 ml of lysis buffer, followed by a wash with lysis buffer supplemented to 1 M NaCl.</li>
		<li>Wash resin with 40 ml of lysis buffer supplemented with 80 mM imidazole.</li>
		<li>Elute the Ruby-Shroom fusion protein with 40 ml of lysis buffer supplemented to 1 M imidazole. Manually fractionate the eluted protein into 4-10 ml fractions.</li>
		<li>Confirm fractions containing Ruby-Shroom fusion protein using 10% SDS-PAGE.</li>
		<li>Dialyze appropriate fractions (typically fractions 2-4) into 8% glycerol, 250 mM NaCl, 15 mM imidazole, 20 mM Tris pH8.0, 1 mM &#946;-ME, using 6-8 kDa MWCO dialysis tubing. Add 1 mg of TEV protease for every 25-50 mg of fusion protein. Dialyze overnight at room temperature with slow stirring.</li>
		<li>Repeat nickel affinity purification steps 4.1.2. to 4.1.6. The Shroom SD2 domain should now remain in the unbound fraction while the His10-mRuby2 will remain bound to the resin and will be found in the elution fractions. Confirm this using 12% SDS-PAGE staining with Coomassie Blue.</li>
		<li>Dialyze fractions containing Shroom SD2 domain into 8% glycerol, 100 mM NaCl, 20 mM Tris pH 8.0, 5 mM beta-mercaptoethanol buffer overnight.</li>
		<li>Perform anion exchange chromatography25 using an FPLC, and eluting with a NaCl gradient from 0.1-1.0 M NaCl. Analyze fractions using 12% SDS-PAGE.</li>
		<li>Using a spin concentrator (MWCO of 10 kDa or greater depending on the protein being studied), concentrate peak fractions to ~ 0 mg/ml, and then perform size exclusion chromatography26, analyzing peak fractions via 12% SDS-PAGE.</li>
		<li>Pool peak fractions and dialyze into 20 mM Tris pH 8.0, 50 mM NaCl, 2% glycerol, 1 mM &#946;-ME, and concentrate to 10 mg/ml prior to crystallization. 
		Optional: During this step, remove 5 &#181;l samples at concentrations of 1, 2, 5, and 10 mg/ml during the course of concentrating the sample. Run a native PAGE loading 2-5 &#181;l of each sample to look at the behavior of the sample during this step.</li>
	</ol>
	</li>
	<li>Screen a small array of 12 conditions to identify an optimal protein concentration for crystallization trials. The composition of these conditions is described in Figure 7.
	<ol>
		<li>Using 24-well sitting drop crystallization trays, perform crystallization trials using the vapor diffusion method, pipetting 500 &#181;l of each of the 12 conditions into separate wells followed by drops that initially contain 1 &#181;l of well solution and 1 &#181;l of protein sample on the coverslips. Please see27 for additional information on this technique.</li>
		<li>Quickly seal the tray with clear tape and move the tray to a suitable temperature controlled environment that is vibration free. The temperature used can vary, but 4 &#176;C, 16 &#176;C, and 20 &#176;C are quite common.</li>
		<li>Examine the drops in the trays over the next 3 days using a microscope at up to 100X magnification. At the end of the 3-day period, score each drop as containing no precipitation (clear), light precipitation, heavy precipitation (brown), or crystals. A suitable protein concentration should contain no more than 6 heavy precipitations.</li>
		<li>Using the protein concentration identified in 4.2, screen a broad range of commercially available crystallization screens. The use of a liquid handling or crystallization robot greatly speeds up this process while also minimizing error in the drops. It requires far less protein as well, allowing the user to screen more conditions with a single sample.</li>
		<li>Improve initial conditions identified from broad screens by systematically varying each of the variables within the crystallization drop using 24-well screening trays as before.</li>
	</ol>
	</li>
</ol>

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A., 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=The+JCSG+high-throughput+structural+biology+pipeline.">The JCSG high-throughput structural biology pipeline.</a> Acta Crystallogr Sect F Struct Biol Cryst Commun. 66 (Pt 10), 1137-1142 (2010).</li><li>Bach, H., 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=Escherichia+coli+maltose-binding+protein+as+a+molecular+chaperone+for+recombinant+intracellular+cytoplasmic+single-chain+antibodies.">Escherichia coli maltose-binding protein as a molecular chaperone for recombinant intracellular cytoplasmic single-chain antibodies.</a> J Mol Biol. 312 (1), 79-93 (2001).</li><li>Strong, M., 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=Toward+the+structural+genomics+of+complexes:+crystal+structure+of+a+PE/PPE+protein+complex+from+Mycobacterium+tuberculosis.">Toward the structural genomics of complexes: crystal structure of a PE/PPE protein complex from Mycobacterium tuberculosis.</a> Proc Natl Acad Sci U S A. 103 (21), 8060-8065 (2006).</li><li>Heras, B., Martin, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Post-crystallization+treatments+for+improving+diffraction+quality+of+protein+crystals.">Post-crystallization treatments for improving diffraction quality of protein crystals.</a> Acta Crystallogr D Biol Crystallogr. 61 (Pt 9), 1173-1180 (2005).</li></ol>

The authors have nothing to disclose.

The protocol described here is designed to help the user identify domain boundaries within large coiled-coil proteins to facilitate their crystallization. The protocol relies on a holistic incorporation of a variety of data from computational predictions and other sources to generate a series of potential domain boundaries. These are followed by a set of biochemical experiments which are quick and inexpensive, and are used to further refine these initial hypotheses. Using this approach, the user could quickly eliminate p...

Structure determination via X-ray crystallography has made fundamental contributions to every field of modern biology; providing an atomic view of the macromolecules that support life and how they interact with one another in a variety of contexts; allowing us to understand the mechanisms that cause disease and providing opportunities to rationally design drugs to treat disease. Crystallography has long been the dominant experimental technique for determining macromolecular structure, and currently accounts for 89.3% of the structural database (www.rcsb.org). This technique has many advantages, including the potential for very high resolution, the ability to visualize macromolecules with a broad range of sizes, relatively easy data collection, and the opportunity to visualize how the macromolecule interacts with solvent as well as ligands.
Despite numerous technological improvements in recombinant protein expression1,2, purification3, and molecular biology used to generate these systems4, the single biggest obstacle in the crystallographic process remains the ability to grow diffraction quality crystals. This has been especially true for proteins which contain large coiled-coil domains. It has been estimated that as much as 5% of all amino acids are found within coiled-coils5,6, making this a very common structural feature7, yet these proteins are often more difficult to purify and crystallize than globular proteins8-10. This is further compounded by the fact that coiled-coil domains are often found within the context of a larger protein, therefore correctly predicting the boundaries of these domains is critical to avoid the inclusion of unstructured or flexible sequence that is often detrimental for crystallization.
Here we present a conceptual framework combining web-based computational analyses with experimental data from the bench, to help guide users through the initial stages of the crystallographic process including: how to select protein fragment(s) for structural studies, and how to prepare and characterize protein samples prior to crystallization attempts. We focus our analysis on two proteins containing large coiled-coil domains, Shroom (Shrm) and Rho-kinase (Rock). These proteins were chosen as they both contain coiled-coil domains and are known to form a biologically relevant complex11-16. Shroom and Rho-kinase (Rock) are predicted to contain ~200 and 680 residues of coiled-coil respectively, many portions of which have been characterized structurally17-20. The method described here provides a streamlined workflow to quickly identify fragments of coiled-coil containing protein that will be amenable for crystallization, however, the techniques described can easily be adapted for most protein or protein-complexes or modified to incorporate high-throughput approaches as available. Lastly, these methods are generally inexpensive and can be performed by users at nearly all experience levels.

<table><tbody><tr><td>BL21(DE3) Rosetta</td><td>Emd Millipore</td><td>70954-3</td><td></td></tr><tr><td>BL21(DE3) Star</td><td>ThermoFisher Scientific</td><td>C601003</td><td></td></tr><tr><td>BL21(DE3) Codon Plus</td><td>Agilent Technologies</td><td>230245</td><td></td></tr><tr><td>Lysozyme</td><td>Spectrum Chemical Mfg Corp</td><td>L3008-5GM</td><td></td></tr><tr><td>Ni-NTA resin</td><td>Life Technologies</td><td>25216</td><td></td></tr><tr><td>SubtilisinA</td><td>Spectrum Chemical Mfg Corp</td><td>S1211-10ML</td><td></td></tr><tr><td>24 well Cryschem Plate</td><td>Hampton research</td><td>HR3-160</td><td></td></tr><tr><td>INTELLI-PLATE &amp;nbsp;96:</td><td>Art Robbins Instruments</td><td>102-0001-03</td><td></td></tr><tr><td>PEG 3350</td><td>Hampton research</td><td>HR2-591</td><td></td></tr><tr><td>PEG 8000</td><td>Hampton research</td><td>HR2-515</td><td></td></tr><tr><td>PEG 400</td><td>Hampton research</td><td>HR2-603</td><td></td></tr><tr><td>PEG 4000</td><td>Hampton research</td><td>HR2-605</td><td></td></tr><tr><td>pcDNA3.1-Clover-mRuby2</td><td>Addgene</td><td>49089</td><td></td></tr><tr><td>Overnight Express Autoinduction System 1</td><td>Emd Millipore</td><td>71300</td><td></td></tr><tr><td>Lysogeny Broth powder</td><td>ThermoFisher Scientific</td><td>12795027&amp;nbsp;</td><td></td></tr></tbody></table>

combining wet and dry lab techniques to guide the crystallization of large coiled-coil containing proteins

Obtaining crystals for structure determination can be a difficult and time consuming proposition for any protein. Coiled-coil proteins and domains are found throughout nature, however, because of their physical properties and tendency to aggregate, they are traditionally viewed as being especially difficult to crystallize. Here, we utilize a variety of quick and simple techniques designed to identify a series of possible domain boundaries for a given coiled-coil protein, and then quickly characterize the behavior of these proteins in solution. With the addition of a strongly fluorescent tag (mRuby2), protein characterization is simple and straightforward. The target protein can be readily visualized under normal lighting and can be quantified with the use of an appropriate imager. The goal is to quickly identify candidates that can be removed from the crystallization pipeline because they are unlikely to succeed, affording more time for the best candidates and fewer funds expended on proteins that do not produce crystals. This process can be iterated to incorporate information gained from initial screening efforts, can be adapted for high-throughput expression and purification procedures, and is augmented by robotic screening for crystallization.

A diagram depicting the workflow utilized in this system is shown in Figure 1 and includes three main stages. Computational analysis of the sequence is utilized to develop hypotheses about the domain boundaries of the coiled-coil protein of interest. An example of an annotated analysis of the Shrm2 SD2 domain is shown in Figure 2. In this diagram, the goal was to identify possible domain boundaries for a conserved domain at the C-terminus of the cytoskele...

Watch this Scientific Journal Video about Combining Wet and Dry Lab Techniques to Guide the Crystallization of Large Coiled-coil Containing Proteins at JoVE.com

Combining Wet and Dry Lab Techniques to Guide the Crystallization of Large Coiled-coil Containing Proteins

We describe a framework incorporating straightforward biochemical and computational analysis to guide the characterization and crystallization of large coiled-coil domains. This framework can be adapted for globular proteins or extended to incorporate a variety of high-throughput techniques.

Obtaining crystals for structure determination can be a difficult and time consuming proposition for any protein. Coiled-coil proteins and domains are ...

combining-wet-dry-lab-techniques-to-guide-crystallization-large

Research

JoVE Journal

Biochemistry

7.9K Views.  University of Pittsburgh. We describe a framework incorporating straightforward biochemical and computational analysis to guide the characterization and crystallization of large coiled-coil domains. This framework can be adapted for globular proteins or extended to incorporate a variety of high-throughput techniques.

Video: Combining Wet and Dry Lab Techniques to Guide the Crystallization of Large Coiled-coil Containing Proteins

Automated Protocols for Macromolecular Crystallization at the MRC Laboratory of Molecular Biology

When high quality crystals are obtained that diffract X-rays, the crystal structure may be solved at near atomic resolution. The conditions to crystallize proteins, DNAs, RNAs, and their complexes can however not be predicted. Employing a broad variety of conditions is a way to increase the yield of quality diffraction crystals. Two fully automated systems have been developed at the MRC Laboratory of Molecular Biology (Cambridge, England, MRC-LMB) that facilitate crystallization screening against 1,920 initial conditions by vapor diffusion in nanoliter droplets. Semi-automated protocols have also been developed to optimize conditions by changing the concentrations of reagents, the pH, or by introducing additives that potentially enhance properties of the resulting crystals. All the corresponding protocols will be described in detail and briefly discussed. Taken together, they enable convenient and highly efficient macromolecular crystallization in a multi-user facility, while giving the users control over key parameters of their experiments.

Automated systems and protocols for the routine preparation of a large number of screens and nanoliter crystallization droplets for vapor diffusion experiments are described and discussed.

When high quality crystals are obtained that diffract X-rays, the crystal structure may be solved at near atomic resolution. The conditions to crystallize ...

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

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

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

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

Chemistry

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to precisely maneuver the nucleation and crystal growth towards desired sizes of protein crystals. The controlled crystallization is based on sample investigation with in situ Dynamic Light Scattering (DLS) while all visual changes in the droplet are monitored online with the help of a microscope coupled to a CCD camera, thus enabling a full investigation of the protein droplet during all stages of crystallization. The use of in situ DLS measurements throughout the entire experiment allows a precise identification of the highly supersaturated protein solution transitioning to a new phase &#8211; the formation of crystal nuclei. By identifying the protein nucleation stage, the crystallization can be optimized from large protein crystals to the production of protein microcrystals. The experimental protocol shows an interactive crystallization approach based on precise automated steps such as precipitant addition, water evaporation for inducing high supersaturation, and sample dilution for slowing induced homogeneous nucleation or reversing phase transitions.

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

Here we present a protocol for controlled production of protein microcrystals. The process uses an automated device allowing controlled manipulation of several crystallization parameters. The protein crystallization is carried-out by controlled and automated addition of crystallization solutions while monitoring and investigating the radius distribution of particles in the crystallization droplet.

The automated crystallization device is a patented technique1 especially developed for monitoring protein crystallization experiments with the aim to ...

Characterization of Glycoproteins with the Immunoglobulin Fold by X-Ray Crystallography and Biophysical Techniques

Glycoproteins on the surface of cells play critical roles in cellular function, including signalling, adhesion and transport. On leukocytes, several of these glycoproteins possess immunoglobulin (Ig) folds and are central to immune recognition and regulation. Here, we present a platform for the design, expression and biophysical characterization of the extracellular domain of human B cell receptor CD22. We propose that these approaches are broadly applicable to the characterization of mammalian glycoprotein ectodomains containing Ig domains. Two suspension human embryonic kidney (HEK) cell lines, HEK293F and HEK293S, are used to express glycoproteins harbouring complex and high-mannose glycans, respectively. These recombinant glycoproteins with different glycoforms allow investigating the effect of glycan size and composition on ligand binding. We discuss protocols for studying the kinetics and thermodynamics of glycoprotein binding to biologically relevant ligands and therapeutic antibody candidates. Recombinant glycoproteins produced in HEK293S cells are amenable to crystallization due to glycan homogeneity, reduced flexibility and susceptibility to endoglycosidase H treatment. We present methods for soaking glycoprotein crystals with heavy atoms and small molecules for phase determination and analysis of ligand binding, respectively. The experimental protocols discussed here hold promise for the characterization of mammalian glycoproteins to give insight into their function and investigate the mechanism of action of therapeutics.

We present approaches for the biophysical and structural characterization of glycoproteins with the immunoglobulin fold by biolayer interferometry, isothermal titration calorimetry, and X-ray crystallography.

Glycoproteins on the surface of cells play critical roles in cellular function, including signalling, adhesion and transport. On leukocytes, several of ...

Multi-target Parallel Processing Approach for Gene-to-structure Determination of the Influenza Polymerase PB2 Subunit

Pandemic outbreaks of highly virulent influenza strains can cause widespread morbidity and mortality in human populations worldwide. In the United States alone, an average of 41,400 deaths and 1.86 million hospitalizations are caused by influenza virus infection each year 1. Point mutations in the polymerase basic protein 2 subunit (PB2) have been linked to the adaptation of the viral infection in humans 2. Findings from such studies have revealed the biological significance of PB2 as a virulence factor, thus highlighting its potential as an antiviral drug target.
The structural genomics program put forth by the National Institute of Allergy and Infectious Disease (NIAID) provides funding to Emerald Bio and three other Pacific Northwest institutions that together make up the Seattle Structural Genomics Center for Infectious Disease (SSGCID). The SSGCID is dedicated to providing the scientific community with three-dimensional protein structures of NIAID category A-C pathogens. Making such structural information available to the scientific community serves to accelerate structure-based drug design.
Structure-based drug design plays an important role in drug development. Pursuing multiple targets in parallel greatly increases the chance of success for new lead discovery by targeting a pathway or an entire protein family. Emerald Bio has developed a high-throughput, multi-target parallel processing pipeline (MTPP) for gene-to-structure determination to support the consortium. Here we describe the protocols used to determine the structure of the PB2 subunit from four different influenza A strains.

Structure-based drug design plays an important role in drug development. Pursuing multiple targets in parallel greatly increases the chance of success for lead discovery. The following article highlights how the Seattle Structural Genomics Center for Infectious Disease utilizes a multi-target approach for gene-to-structure determination of the PB2 influenza A subunit.

Pandemic outbreaks of highly virulent influenza strains can cause widespread morbidity and mortality in human populations worldwide. In the United States ...

Immunology and Infection

High-Throughput Protein Crystallization via Microdialysis

Understanding the structure-function relationships of macromolecules, such as proteins, at the molecular level is vital for biomedicine and modern drug discovery. To date, X-ray crystallography remains the most successful method for solving three-dimensional protein structures at atomic resolution. With recent advances in serial crystallography, either using X-ray free electron lasers (XFELs) or synchrotron light sources, protein crystallography has progressed to the next frontier, where the ability to acquire time-resolved data provides important mechanistic insights into the behavior of biological molecules at room temperature. This protocol describes a straightforward high-throughput (HTP) workflow for screening crystallization conditions through the use of a 96-well dialysis plate. These plates follow the Society for Biomolecular Screening (SBS) standard and can be easily set up using any standard crystallization laboratory. Once optimal conditions are identified, large quantities of crystals (hundreds of microcrystals) can be produced using the dialyzer. To validate the robustness and versatility of this approach, four different proteins were crystallized, including two membrane proteins.

High-Throughput Protein Crystallization via Microdialysis

The presented protocol describes a straightforward approach for screening protein crystallization conditions and crystal growth using a 96-well high-throughput dialysis plate. The use of dialyzer tubes for the large-scale growth of microcrystals is also demonstrated for serial crystallography and MicroED applications.

Understanding the structure-function relationships of macromolecules, such as proteins, at the molecular level is vital for biomedicine and modern drug ...

Protein Crystallization for X-ray Crystallography

Using the three-dimensional structure of biological macromolecules to infer how they function is one of the most important fields of modern biology. The availability of atomic resolution structures provides a deep and unique understanding of protein function, and helps to unravel the inner workings of the living cell. To date, 86% of the Protein Data Bank (rcsb-PDB) entries are macromolecular structures that were determined using X-ray crystallography.
To obtain crystals suitable for crystallographic studies, the macromolecule (e.g. protein, nucleic acid, protein-protein complex or protein-nucleic acid complex) must be purified to homogeneity, or as close as possible to homogeneity. The homogeneity of the preparation is a key factor in obtaining crystals that diffract to high resolution (Bergfors, 1999; McPherson, 1999).
Crystallization requires bringing the macromolecule to supersaturation. The sample should therefore be concentrated to the highest possible concentration without causing aggregation or precipitation of the macromolecule (usually 2-50 mg/ mL). Introducing the sample to precipitating agent can promote the nucleation of protein crystals in the solution, which can result in large three-dimensional crystals growing from the solution. There are two main techniques to obtain crystals: vapor diffusion and batch crystallization. In vapor diffusion, a drop containing a mixture of precipitant and protein solutions is sealed in a chamber with pure precipitant. Water vapor then diffuses out of the drop until the osmolarity of the drop and the precipitant are equal (Figure 1A). The dehydration of the drop causes a slow concentration of both protein and precipitant until equilibrium is achieved, ideally in the crystal nucleation zone of the phase diagram. The batch method relies on bringing the protein directly into the nucleation zone by mixing protein with the appropriate amount of precipitant (Figure 1B). This method is usually performed under a paraffin/mineral oil mixture to prevent the diffusion of water out of the drop.
Here we will demonstrate two kinds of experimental setup for vapor diffusion, hanging drop and sitting drop, in addition to batch crystallization under oil.

The 3-D structure of a molecule provides a unique understanding of how the molecule functions. The principal method for structure determination at near-atomic resolution is X-ray crystallography. Here, we demonstrate the current methods for obtaining three-dimensional crystals of any given macromolecule that are suitable for structure determination by X-ray crystallography.

Using the three-dimensional structure of biological macromolecules to infer how they function is one of the most important fields of modern biology. The ...

Biology

Crystallization of ABC Transporter Membrane Proteins in Lipidic Bicelles

ABCG5/G8 Crystallization in a Lipidic Bicelle Environment for X-Ray Crystallography

ATP-binding cassette (ABC) transporters constitute lipid-embedded membrane proteins. Extracting these membrane proteins from the lipid bilayer to an aqueous environment is typically achieved by employing detergents. These detergents disintegrate the lipid bilayer and solubilize the proteins. The intrinsic habitat of membrane proteins within the lipid bilayer poses a challenge in maintaining their stability and uniformity in solution for structural characterization. Bicelles, which comprise a blend of long and short-chain phospholipids and detergents, replicate the natural lipid structure. The utilization of lipid bicelles and detergents serves as a suitable model system for obtaining high-quality diffraction crystals, specifically to determine the high-resolution structure of membrane proteins. Through these synthetic microenvironments, membrane proteins preserve their native conformation and functionality, facilitating the formation of three-dimensional crystals. In this approach, the detergent-solubilized heterodimeric ABCG5/G8 was reintegrated into DMPC/CHAPSO bicelles, supplemented with cholesterol. This setup was employed in the vapor diffusion experimental procedure for protein crystallization.

Author Spotlight: A Bicelle Crystallization Setup for ABC Transporter Membrane Proteins to Advance Drug Development 

This protocol describes a setup for the crystallization of the sterol transporter ABCG5/G8. ABCG5/G8 is reconstituted into bicelles for hanging-drop crystallization. The protocol does not require specialized materials or substrates, making it accessible and easy to adapt in any laboratory for determining the protein structure through X-ray crystallography.

ATP-binding cassette (ABC) transporters constitute lipid-embedded membrane proteins. Extracting these membrane proteins from the lipid bilayer to an ...

Crystallization of Membrane Proteins in Lipidic Mesophases

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are implicated in a multitude of malfunctions and diseases. However, a structural and functional understanding of membrane proteins is strongly lagging behind that of their soluble partners, mainly, due to difficulties associated with their solubilization and generation of diffraction quality crystals. Crystallization in lipidic mesophases (also known as in meso or LCP crystallization) is a promising technique which was successfully applied to obtain high resolution structures of microbial rhodopsins, photosynthetic proteins, outer membrane beta barrels and G protein-coupled receptors. In meso crystallization takes advantage of a native-like membrane environment and typically produces crystals with lower solvent content and better ordering as compared to traditional crystallization from detergent solutions. The method is not difficult, but requires an understanding of lipid phase behavior and practice in handling viscous mesophase materials. Here we demonstrate a simple and efficient way of making LCP and reconstituting a membrane protein in the lipid bilayer of LCP using a syringe mixer, followed by dispensing nanoliter portions of LCP into an assay or crystallization plate, conducting pre-crystallization assays and harvesting crystals from the LCP matrix. These protocols provide a basic guide for approaching in meso crystallization trials; however, as with any crystallization experiment, extensive screening and optimization are required, and a successful outcome is not necessarily guaranteed.

The protocols describe the essential steps for obtaining diffraction quality crystals of a membrane protein starting from reconstitution of the protein in a lipidic cubic phase (LCP), finding initial conditions with LCP-FRAP pre-crystallization assays, setting up LCP crystallization trials and harvesting crystals.

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are ...

High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane bilayer that surrounds all cells and organelles. Alterations in the function of MPs result in many human diseases and disorders; thus, an intricate understanding of their structures remains a critical objective for biological research. However, structure determination of MPs remains a significant challenge often stemming from their hydrophobicity. 
MPs have substantial hydrophobic regions embedded within the bilayer. Detergents are frequently used to solubilize these proteins from the bilayer generating a protein-detergent micelle that can then be manipulated in a similar manner as soluble proteins. Traditionally, crystallization trials proceed using a protein-detergent mixture, but they often resist crystallization or produce crystals of poor quality. These problems arise due to the detergent&prime;s inability to adequately mimic the bilayer resulting in poor stability and heterogeneity. In addition, the detergent shields the hydrophobic surface of the MP reducing the surface area available for crystal contacts. To circumvent these drawbacks MPs can be crystallized in lipidic media, which more closely simulates their endogenous environment, and has recently become a de novo technique for MP crystallization.
Lipidic cubic phase (LCP) is a three-dimensional lipid bilayer penetrated by an interconnected system of aqueous channels1. Although monoolein is the lipid of choice, related lipids such as monopalmitolein and monovaccenin have also been used to make LCP2. MPs are incorporated into the LCP where they diffuse in three dimensions and feed crystal nuclei. A great advantage of the LCP is that the protein remains in a more native environment, but the method has a number of technical disadvantages including high viscosity (requiring specialized apparatuses) and difficulties in crystal visualization and manipulation3,4. Because of these technical difficulties, we utilized another lipidic medium for crystallization-bicelles5,6 (Figure 1). Bicelles are lipid/amphiphile mixtures formed by blending a phosphatidylcholine lipid (DMPC) with an amphiphile (CHAPSO) or a short-chain lipid (DHPC). Within each bicelle disc, the lipid molecules generate a bilayer while the amphiphile molecules line the apolar edges providing beneficial properties of both bilayers and detergents. Importantly, below their transition temperature, protein-bicelle mixtures have a reduced viscosity and are manipulated in a similar manner as detergent-solubilized MPs, making bicelles compatible with crystallization robots. 
Bicelles have been successfully used to crystallize several membrane proteins5,7-11 (Table 1). This growing collection of proteins demonstrates the versatility of bicelles for crystallizing both alpha helical and beta sheet MPs from prokaryotic and eukaryotic sources. Because of these successes and the simplicity of high-throughput implementation, bicelles should be part of every membrane protein crystallographer&prime;s arsenal. In this video, we describe the bicelle methodology and provide a step-by-step protocol for setting up high-throughput crystallization trials of purified MPs using standard robotics.

Bicelles are lipid/amphiphile mixtures that maintain membrane proteins (MPs) within a lipid bilayer but have unique phase behavior that facilitates high-throughput screening by crystallization robots. This technique has successfully produced a number of high-resolution structures from both prokaryotic and eukaryotic sources. This video describes protocols for generating the lipidic bicelle mixture, incorporating MPs into the bicelle mixture, setting up crystallizations trials (manually as well as robotically) and harvesting crystals from the medium.

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane ...

Combining Wet and Dry Lab Techniques to Guide the Crystallization of Large Coiled-coil Containing Proteins

Summary

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Chapters in this video

Automated Protocols for Macromolecular Crystallization at the MRC Laboratory of Molecular Biology

From Constructs to Crystals – Towards Structure Determination of β-barrel Outer Membrane Proteins

Growing Protein Crystals with Distinct Dimensions Using Automated Crystallization Coupled with In Situ Dynamic Light Scattering

Characterization of Glycoproteins with the Immunoglobulin Fold by X-Ray Crystallography and Biophysical Techniques

Multi-target Parallel Processing Approach for Gene-to-structure Determination of the Influenza Polymerase PB2 Subunit

High-Throughput Protein Crystallization via Microdialysis

Protein Crystallization for X-ray Crystallography

ABCG5/G8 Crystallization in a Lipidic Bicelle Environment for X-Ray Crystallography

Crystallization of Membrane Proteins in Lipidic Mesophases

High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method