Preparation of Protein Samples

Purification

Protein samples should be as pure as possible for successful crystallization. Protein that is >90% pure should be sufficient for commencing crystallization screens. The more homogeneous the protein, the more likely crystallization is to be successful. Purity can be evaluated by SDS-PAGE, isoelectric focusing, and/or mass spectroscopy.

Sample storage

Proteins are typically stored at 4°C or frozen at –80°C in a solution appropriate to maintain stability and activity. Proteins solutions should be as concentrated as practical to enhance stability. A stock protein concentration of 10-20 mg/mL is typical for crystallization screening. If protein is stored frozen, it should be aliquoted to minimize repeated freeze/thaw cycles that are usually deleterious to proteins. In general, protein solutions should contain the minimum concentrations of buffers, salts, and preservatives necessary for safe storage. In particular, the use of high concentrations of glycerol or other polyols in storage solutions should be avoided, as this can alter or interfere with crystallization.

Sample handling

Most proteins are sensitive to harsh handling. Unless known otherwise, proteins should always be maintained on ice when not in the refrigerator, cold room, or freezer. In addition, protein solutions should never be subjected to vortexing or vigorous mixing; the resulting foaming promotes protein denaturation. Before using proteins in crystallization trials, it is customary to remove dust and precipitated protein by centrifugation at 14000 xg for 5-10 minutes at 4 °C.

Crystallization Reagents

Crystallization solutions should be prepared from concentrated stock solutions that have been ultrafiltered through 0.2 um polyethylenesulfone (PES) membranes to remove dust and micro-particulate matter. (Nylon membranes will shrink and clog with concentrated salt solutions.) We typically prepare filtered stock solutions in 50 mL polypropylene centrifuge tubes (Steriflip) or polycarbonate 250 mL ultrafilter receivers. Stock solutions can be stored for a considerable length of time, but polypropylene and polycarbonate vessels are slightly porous, and solutions stored in them will change concentration over time. Replace solutions every few months to maintain reproducibility of crystallization conditions.

Crystallization solutions are conveniently prepared in 15 or 50 mL graduated polypropylene centrifuge tubes. The volume markings on these tubes are entirely sufficient for reproducibly diluting reagents to the appropriate final volume. Pipet the appropriate buffer, salt, and other reagent solutions into the tube, and dilute with filtered deionized water to the final volume using the markings on the side of the tube. Mix by inversion or vortexing. It is not necessary to ultrafilter this mixture if the stock solutions and deionized water are ultrafiltered.

Stock Solutions

Buffers

Buffer solutions are typically prepared and titrated to the desired pH at 1M concentration in 50 mL quantities. These solutions will be diluted 10X in a typical crystallization solution. Buffers are normally prepared from the conjugate acid and titrated with 5-10M NaOH to the desired pH prior to ultrafiltration.

Salts

Salt solutions are prepared near saturation in 200-250 mL quantities. An appropriate concentration for salt solutions can be gleaned from the Hampton Research catalog. Solutions should be ultrafiltered at 0.2 um using a PES membrane to remove dust and particulates.

Polymers, Polyols, and Solvents

Non-viscous polyols and solvents (e.g. , ethylene glycol, MPD, DMSO) and polyethylene glycols (e.g., PEG-400) can be used at 100% concentration. Viscous polyols (e.g. glycerol) can be diluted in water to 50-80% to reduce viscosity and aid in dispensing. Solid PEGs (PEG -2000 or larger) are difficult to dissolve by stirring. Instead, microwave PEGs and and water just shy of the target volume to boiling, let cool, and make up to the appropriate volume with water. The usual stock concentration of PEGs is 50% w/v. As always, filter solutions though a 0.2 um PES filter before storage and use.

Hanging Drop Crystallization

The most common method of protein crystallization is hanging drop vapor diffusion. In this method, a concentrated protein solution is combined with a solution of a precipitant and allowed to concentrate by evaporation. Under the right conditions, and with the appropriate precipitant, protein crystals will form. In hanging drop vapor diffusion, a small volume of protein sample and precipitant are combined on a glass coverslip and sealed over a well containing precipitant solution (fig 1). Because the precipitant concentration in the mixed drop of protein is lower than in the well solution, water evaporates from the drop—increasing the concentration of both protein and precipitant—until the drop is in equilibrium with the well solution. The concentration of protein and precipitant in the drop occurs slowly and gradually, favoring crystallization over precipitation.

Figure 1. Hanging drop vapor diffusion

Sitting drop crystallization is very similar to hanging drop crystallization, except that the protein sample sits in a small depression within the crystallization well, situated above or next to the reservoir solution, with which it can equilibrate. Sitting drop crystallization is more suitable for automated drop-setting.

Preparing manual crystallization trays

Crystallization trials are conveniently performed in 24-well, pre-greased crystallization trays (fig 2). Prior to setting trays, carefully organize your solutions and record in your notebook the crystallization conditions to be used in each well. The following protocol is typical:

• Obtain a pre-greased 24-well crystallization tray and a box of 22 mm siliconized cover slips. If setting trays at 4 °C, allow the tray and all solutions to equilibrate before proceeding.
• Fill the wells of the tray with the appropriate precipitant solutions.
• Remove a coverslip from the box, taking care to handle only by the edges.
• Pipet 1 μL¹ of protein² on to the cover slip, taking care not to introduce bubbles.
• Pipet an equal volume of precipitant solution (from the corresponding well) into the protein drop. Optionally, gently mix by pipetting up and down a few times. Avoiding mixing the drops may result in less nucleation and larger crystals. Experiment to see what works best.
• Immediately place the cover slip over the well, press down gently and twist 45° to ensure a good seal.
• Repeat for remaining wells.
• Immediately after preparing the plate, place it under a microscope and examine each of the drops for protein precipitation or foreign objects (glass shards, fibers, plastic bits) and make a notation of any drops that are not clear.
• Place the plate in a quiet place at the appropriate temperature and leave it undisturbed for at least 24 hours.

Figure 2. A Hampton Research 24-well VDX plate™ and siliconized coverslips

Notes

1. Up to 40 μL of protein can be used if desired.
2. Protein concentrations from 5-20 mg/mL are typical; a protein concentration of ≈10 mg/mL is a good starting point for initial screens.

Preparing crystallization trays with the Crystal Gryphon

The Crystal Gryphon is an automated drop setter with a 96-syringe head for dispensing screen solutions into 96-well plates, and a nano-needle for dispensing protein solution into crystallization wells. The Crystal Gryphon can set a 96-condition screen at two different protein concetnrations in under 2 minutes.

Materials and solutions required

The following solutions and supplies are required prior to automated dispensing

• A deep well block with screen solution, e.g. a Greiner 1.0 mL MasterBlock filled with Hampton Research Crystal Screen HT solutions¹. Verify that the deep well block has sufficient solution left to complete the necessary screens.²
• An empty crystallization plate, e.g. an Art Robbins 2-well Intelliplate
• An appropriate volume of protein solution at an appropriate concentration in a 200 μL PCR tube
• A strip of 3 inch HD clear duct tape long enough to seal a crystallization plate

Crystal Gryphon setup

• Ensure that the 10 L wash station supply carboy has an adequate supply of water.
• Ensure that the 1 L nano-needle wash reservoirs have an adequate supply of water or solvent.
• Check that all wash supply tubes are properly connected and not kinked or otherwise impinged.
• Turn on the power strip that supplies the Crystal Gryphon to power up the dispensing stage and pumps.
• Open the lid to the laptop to bring it out of hibernation and Open the Crystal Gryphon software
• Click on the Connect icon to establish communications with the Crystal Gryphon.
• Click on Tray Out to gain access to the stage.
• Place an empty crystallization plate in position 1 of the stage.
• Place a screen block in position 2 of the stage. Remove the sealing tape or cap mat before placing on teh stage!
• Place the tube with protein solution in position 10 of the stage. Open or remove the lid of the tube before placing on the stage!
• The Crystal Gryphon is ready to dispense.

Dispensing solution

• Load an appropriate protocol into the procotol window. Typical Protocols are described below.
• Important! Verify that the protocol selected is appropriate for the types of plates installed on the dispensing stage, and that the quantity of protein on hand is sufficient to carry out the protocol. Typically,
  • Stage position 1 contains an empty 2-well Intelliplate 96-well plate
  • Stage position 2 contains a Greiner 1.0 mL Masterblock with screen solutions
  • Stage position 10 contains 80-100 μL of protein solution
• Important! Verify that the cap mat has been removed from the screen block, and the lid has been opened and swung out of the way for the protein solution. Failure to do this may result in irreversible damage to the instrument.
• When you have verified that the instrument is ready to dispense, click on the GO button.

Notes:

1. Screens can be purchased in deep well blocks or can be dispensed manually from tubes into empty deep-well blocks and sealed with cap mats. A 1.0 mL deep-well block should be sufficient for 20 screens described here.
2. Screen blocks will normally be marked with the number of uses.

Gryphon Protocols

These are typical protocols used by the Crystal Gryphon to set up commonly used screens. These screens can be loaded from the menu on our instrument.

2-drop Screen

This screen will set up a 40 μL reservoir with 200 nL of screen solution at two different protein concentrations (200 nL and 400 nL protein).

• Wash 96-syringe head
  • 2 times, 50 μL, in the wash station (stage position 3)
• Wash nano-needle
  • 1 time, 3 cycles, in Stage position 12
• Aspirate 55 μL of screen from the deep well block (stage position 2)
  • Pre-dispense 5 μL of screen to compress any bubbles present
• Nano-aspirate 85μL protein from the protein tube (stage position 10)
  • 100μL of protein should be available in the 0.2 mL PCR tube
• Dispense 40 μL of screen into crystallization plate reservoir (stage position 1)
• Dispense 200 nL of screen into crystallization plate 4 μL well (stage position 1)
• Dispense 200 nL of screen into crystallziation plate 10 μL well (stage position 1)
• Nano-dispense 200 nL of protein into crystallization plate 4 μL well (stage position 1)
• Nano-dispense 400 nL of protein into crystallization plate 10 μL well (stage position 1)
• Exchange trays
  • Rremove crystalliation tray from stage position 1
  • Seal plate by applying one strip of 3 inch wide HD clear duct tape. Press tape tightly against tray by pulling a straight, hard edge across it.
• Dispense (empty) screen back into screen block (stage position 2)
• Nano-dispense (purge) 5 μL protein back into protein tube (stage position 10)
• Wash 96-syringe head
  • 2 times, 50 μL, in the wash station (stage position 3)
• Wash nano-needle
  • 1 time, 3 cycles, in Stage position 12

1-drop Screen

This screen will set up a protein-sparing 1-drop screen with 40 μL of reservoir solution and a 200+200 uL drop of well solution and protein.

• Wash 96-syringe head
  • 2 times, 50 μL, in the wash station (stage position 3)
• Wash nano-needle
  • 1 time, 3 cycles, in Stage position 12
• Aspirate 55 μL of screen from the deep well block (stage position 2)
  • Pre-dispense 5 μL to compress any bubbles present
• Nano-aspirate 45μL protein from the protein tube (stage position 10)
  • 60μL should be available in the 0.2 mL tube
• Dispense 40 μL of screen into crystallization plate reservoir (stage position 1)
• Dispense 200 nL of screen into crystallization plate 10 μL well (stage position 1)
• Nano-dispense 200 nL of protein into crystallization plate 10 μL well (stage position 1)
• Exchange trays
  • Remove crystallization tray from stage position 1
  • Seal plate by applying one strip of 3 inch wide HD clear duct tape. Press tape tightly against tray by pulling a straight, hard edge across it.
• Dispense (empty) screen back into screen block (stage position 2)
• Nano-dispense (purge) 5 μL protein back into protein tube (stage position 10)
• Wash 96-syringe head
  • 2 times, 50 μL, in the wash station (stage position 3)
• Wash nano-needle
  • 1 time, 3 cycles, in Stage position 12

Scorpion Protocols

The Scorpion robot is device that can rapidly and accurately prepare custom screening solutions from standard stock solutions of crystallization reagents.

Run Simulated

With the robot off, you can set up reagent/tube/rack definitions, rack assignments and protocols by double clicking on the Scorpion desktop icon and selecting "Run Simulated" on the bottom right hand corner of the pop-up window. To run protocols, you must have the robot turned on and select "Initialize" on the first window.

Reagents

Define a New Reagent: Setup (top tab) > Reagent Definitions (left tab) > "Add" > Type the full name of the new reagent with concentration and pH (if applicable) > input reagent specifications and select appropriate color

• PEGs are red--dark reds are high molecular weigh and pinks are low molecular weight (ie PEG-10,000)
• Solvents are blue (ie isopropanol)
• Buffers are green--dark greens are higher pH and light greens are lower pH (ie Tris)
• Salts are orange (ie AmSO4)

Defining Racks

Assigning reagents to a new rack: Prepare (top tab) > select a position by clicking a (+) > Select a rack type > "OK" > double click on a position and select the appropriate reagent >set volume > "OK" > double click on rack to edit, or cursor over the rack to remove/save.

• The "Define New Sample" option on the left-hand side of the Reagent selection window is meant for temporary reagents and will not be saved as a new reagent definition.

Selecting a saved rack:

Defining Protocols

Skeleton protocols for masterblocks such as "pH vs. [precipitant] screen" and "additive screens" can be opened from the "Scorpion Protocols" folder and edited to save time.

To set up a new protocol

1. Prepare > Select a destination plate type and location by clicking on a position (+) > drag and drop the "Build Screen" icon onto the selected plate
2. In the popup window, drag and drop your desired action from the "Available Functions" options (pH gradient, constant, or gradient) upwards into the "Applied Functions" list. Click on the action to edit it. Actions highlighted in yellow are active and the wells they are acting on are highlighted green. Click and drag to select multiple wells. When you are finished click "OK".
3. Under "Active protocol" you should see the "Build Screen" you just defined. When this is highlighted yellow, it will also highlight the destination position.

Initial Screening of Crystallization Conditions

Strategy

The determination of promising protein crystallization conditions is typically done using a sparse matrix screen, in which a protein is subjected to widely varying pH, salts and precipitants. There are excellent commercial screening kits available, making it generally unnecessary to mix your own initial screening reagents. The following commercial screens are recommended, in the order that they should be employed:

1. Hampton Research Crystal Screen. This screen contains 50 reagents. Screen conditions #25 and #27 have a historically poor record of producing crystals, and these two can be omitted in order to conduct the screen in two 24-well plates.
2. Hampton Research Crystal Screen 2. An extension of the original Hampton Research sparse matrix screen. Two conditions can be omitted for conducting a two-plate screen.

Evaluating screens

Plates should be examined under the microscope and evaluated for protein crystallization after 24 hours, and every day thereafter for a week. After one week, plates should be examined weekly. Record in your results for each drop. Suggested categories and abbreviations to use are as follows, with comments:

• Clear drop (C)—no changes in drop
• Precipitate (P)—typically light brown and granular. Crystals sometimes form from such precipitates. If the precipitate is thick and swirly, this is very bad, and unlikely to form crystals.
• Precipitate/phase separation (PP)—typically light brown and granular, with little blobs that look like oil drops. Crystals sometimes from at the edge of phase separations.
• Microcrystals (MX)—Difficult to distinguish from precipitate; however, unlike precipitated protein, microcrystals have a shiny appearance. Optimization of conditions may result in larger crystals.
• Needle cluster (NX)—Beautiful, but often useless for crystallography. Optimization of conditions may result in less needle-like forms.
• Plates (PX)—if not too thin, may be useful for crystallography. Optimization may result in better crystal form. Plate clusters may be separated in to suitable single crystals.
• Rod clusters (RX)—If separable into single crystals, may be useful for crystallography
• Single crystals (X)—the Holy Grail: large, individual crystals with blocky dimensions.

Optimizing crystallization conditions

If more than half the drops are clear, consider increasing the protein concentration and re-screening. If most of the drops have copious precipitate, consider halving the concentration of the well solutions¹ and re-screening. To save protein in the initial phase of screening, it may be advisable to run only half of a screen at a time in a single 24-well plate until you establish the appropriate protein and well concentrations for efficient screening.

Conditions from the initial screen that show the most promise for crystallization should be further optimized in order to improve crystal form and size. To find the best crystallization conditions, pH, precipitant concentration, and protein concentration should be systematically varied. This will require stock solutions of concentrates so that a variety of custom solutions can be constructed for optimization. For example, typical buffer stock solutions are 1M and pre-adjusted to the desired pH. Salt solutions are generally prepared to near saturation, 1-4 M depending on the salt. Stock solutions available from Hampton Research can be used as a guide to the proper concentration to prepare.

Crystal form can usually be further improved by exploring additives. Preformulated additive screens can be purchased (Hampton Research) or selected additive stocks at 10x the final desired concentration can be prepared and added at 10% volume to hanging drops. Mixing drops may not be beneficial for growing large crystals, because it encourages nucleation. It may be possible to grow fewer and larger crystals by simply pipetting well solution into drops without mixing.

Notes

1. Precipitant concentration can be halved by pipetting equal volumes of the standard screening solution and deionized water into each well. Mix thoroughly before setting drops.

Photodocumentation of crystals

We are currently using an Olympus SZX-12 dissecting microscope equipped with a binocular head, capable of 7-90X magnification. An Olympus 5050Z digital camera replace one of the eyepiece lenses to photograph images. The following instructions describe how to photograph and process crystal images.

Photographing images

• Ensure that the camera has a memory card inserted. The Olympus 5050Z will accept compact flash, xD, and SmartMedia memory cards.
• Loosen the set screw and remove one of the eypieces from the microsope. Set the eyepiece securely aside so that is does not become damaged or roll off the bench.
• Remove the dust cover from the camera extension tube and insert the camera/extension tube into the eypiece slot. Gently tighten the set screw to hold the camera in place on the microscope.
• Turn on the camera, and ensure that the custom mode, "My Camera" is selected.
• Reduce the magnification on the microscope and locate the drop you would like to photograph.
• Once you have found the drop you would like to photograph, return the zoom to 90x and compose your shot. (Note: It is important to set the zoom to 90x if you want to add scale bars to your image later.)
• Refocus the microscope if necessary.
• Use the remote release to fire the shutter.

Processing images

Images should be processed in an image editor to improve quality and to add scale bars. Image processing can be done in any image editing program. Instructions are given here for both Photoshop and Gimp. If images are shot at a magnification of 90x and formatted according to these instructions, you can include the scaling bars below in your images:

• 250 um scale bar
• 500 um scale bar

Photoshop instructions

• Open an image in Photoshop
• Select Image...Image size and resize the image to 900 pixels wide. (The height should automatically adjust to 675 pixels.)
• Adjust the contrast by selecting Image...Adjustments...Auto Levels
• Sharpen the image by selecting Filter...Sharpen...Unsharp Mask
  • Set amount to ≈100%
  • Set radius to ≈1.0 pixel
  • Set threshold to 0-4
  • Preview your sharpening and adjust amount as desired and click OK when satisfied
• Open the appropriate scaling bar image in Photoshop
• Select the entire image by pressing CTRL-A or Select...Select All
• Copy the image by pressing CTRL-C or Edit...Copy
• Click on the image to which you would like to add the sizing bar
• Paste the sizing bar into your image by pressing CTRL-V or Edit...Paste
• Select the Move tool and drag the sizing bar to an appropriate corner of the image.
• Combine the images by selecting Layer...Flatten Image.
• Save the image as a PNG file by selecting File...Save As... and selecting PNG as the output format.

GIMP instructions

• Open an image in GIMP
• Select Image...Scale image and resize the image to 900 pixels wide. (The height should automatically adjust to 675 pixels.)
• Adjust the contrast by selecting Color...Auto...Stretch contrast
• Sharpen the image by selecting Filters...Enhance...Unsharp Mask
  • Set amount to ≈0.5
  • Set radius to ≈5.0
  • Set threshold to 0-4
  • Preview your sharpening and adjust amount as desired and click OK when satisfied
• Open the appropriate scaling bar image in GIMP
• Select the entire image by pressing CTRL-A or Select...Select All
• Copy the image by pressing CTRL-C or Edit...Copy
• Click on the image to which you would like to add the sizing bar
• Paste the sizing bar into your image by pressing CTRL-V or Edit...Paste
• The image should appear as a floating layer (with "crawling ants")
• Select the Move tool and drag the sizing bar to an appropriate corner of the image. Grab the image by one of the dark areas to move it. When you are satisfied with the position, click anywhere outside the image to anchor it.
• Save the image as a PNG file by selecting File...Save As... and selecting PNG as the output format.