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FAQs ABOUT HIGH-THROUGHPUT CRYSTALLIZATION SCREENING home > about > HTS services > faqs
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  1. What is the ‘high-throughput’ crystallization-screening laboratory?
  2. Why is it beneficial to set up crystallization experiments using high-throughput methods, and why use 1536 cocktails?
  3. How long has the laboratory been in operation?
  4. How many crystallization experiments have been set up in the high-throughput laboratory?
  5. Who works in the crystallization screening laboratory?
  6. How do you achieve “high-throughput”?
  7. What is microbatch-under-oil?
  8. Why do we use microbatch-under-oil?
  9. What type of oil do we use?
10. How do we record the experiments’ outcomes?
11. When do we record the experiments’ outcomes?
12. Why do we image the plates more than once?
13. What types of outcomes can be expected from the screening experiments?
14. How do I translate from microbatch-under-oil to vapor diffusion?
15. What cocktails are used in the screen?
16. How do you make the cocktails, and how can I reproduce them in my lab?
17. What types of samples are accepted for screening?
18. How pure should the sample be for screening?
19. How much sample is required for screening?
20. How should I prepare the sample for screening?
21. How should I ship the sample?
22. Who sends the samples?
23. How do I submit a sample?
24. Are there any costs?
25. How do you track the samples?
26. How do you avoid data loss, in particular for the image data?
27. How do I get my image data?
28. How do I view my image data?
29. Are you planning to develop any new image viewing software?
30. What information do you track for the biological macromolecules that are submitted for high-throughput screening?
31. What happens during a typical month in the screening laboratory?
32. Can you recover crystals from the plates?
33. How can I tell if it is a protein or a salt crystal?
34.
What crystallization cocktails are currently in use?
35. Are you developing new screening cocktails?

1. What is the ‘high-throughput’ crystallization-screening laboratory?

The high-throughput screening laboratory at the Hauptman-Woodward Medical Research Institute in Buffalo, New York, is a facility that is used to identify crystallization conditions for biological macromolecules.  This facility makes use of automated liquid handling and imaging systems coordinated through a LIMS/database to quickly set up and record the outcomes of 1536 unique crystallization screening experiments.

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2. Why is it beneficial to set up crystallization experiments using high-throughput methods, and why use 1536 cocktails?

By using automated liquid-handling systems, we are able to set up crystallization experiments precisely and reproducibly using a minimum volume of macromolecular solutions that are often difficult to obtain. The laboratory personnel start with samples contained in a microcentrifuge tube and, within 10 minutes, are able to set up 1536 unique, microbatch-under-oil crystallization experiments. Setting up the same 1536 experiments manually would take a technician several weeks to complete. This speed is truly advantageous, greatly reducing the time available for sample degradation prior to the crystallization experiment. Finally, by setting up so many chemically diverse crystallization experiments, we increase the likelihood of identifying more than one crystallization condition. Crystals produced from different chemical cocktails will often have different physical properties. The ability to choose from several different initial crystallization conditions provides the researcher with multiple paths to pursue when faced with downstream bottlenecks. These downstream bottlenecks can include: ease of optimization, X-ray diffraction quality, and the ability to cryo-preserve the crystals for data collection.

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6. How do you achieve “high-throughput”?

We achieve high-throughput by aspirating solutions from source plates and delivering them, in parallel, to wells in high-density microassay plates.  Details of the screening method are available in the literature [Journal of Structural Biology, 142, 170-179 (2003)].

7. What is microbatch-under-oil?

Microbatch-under-oil is a simple crystallization method developed by Naomi Chayen, Patrick Shaw-Stewart and David Blow [J. Appl. Cryst. 23, pp 297-302 (1990).  J. Cryst. Growth, 122, 176-180 (1992)].  It uses oil to encapsulate an aqueous experiment drop to prevent rapid dehydration of the experiment.  Paraffin Oil is relatively water impermeable and reduces the dehydration rate of the aqueous experiment drop.  Silicon-based oils are more water permeable and allow the drops to dehydrate at a faster rate.  Mixtures of paraffin and silicon oil can be used to regulate the rate of dehydration.  Different types of oil can be used to regulate the rate of water loss from the experiment drop [J. Appl. Cryst. 30, 198-202 (1997)].

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Setting up a microbatch-under-oil experiment.  We use a bank of syringes to deliver oil (yellow), crystallization cocktail (red), and protein solution (blue) in parallel to the wells in a high-density microassay plate.  The cocktail and protein drops merge and are protected from rapid evaporation by the oil.  Crystals form in the aqueous drop.

8. Why do we use microbatch-under-oil?

Microbatch-under-oil was chosen as the crystallization method for the high-throughput screening laboratory because of its efficiency and amenability to automation.

  • Small-volume experiment drops (200 nanoliters protein + 200 nanoliters cocktail)
  • Only use the volume of cocktail solution (200 nanoliters) required for the crystallization experiment drop.  (i.e., No reservoir solution.
  • It is mechanically simple to set up the experiments (no seals or coverslips)
  • 9. What type of oil do we use?

    We use Paraffin Oil purchased from EMD Chemicals Inc. (catalog number PX0045-3).

    10. How do we record the experiments’ outcomes?

    Outcomes of the screening experiments are recorded using custom-built imaging systems.  We have three automated-imaging systems.  Two of them are located in a laboratory at room temperature (~ 23oC).  The third table is located in a temperature-controlled room.  Each table holds a maximum of 28 x 1536 well microassay plates and images the plates a rate of three per hour.  The latest version of the software that controls the tables was written at the Hauptman-Woodward Institute by Raymond Nagel.

11. When do we record the experiments’ outcomes?

Experiment plates are imaged immediately before adding the protein solution when they contain only the crystallization cocktail solution. This provides a control that can be used to ascertain the ‘quality’ of an initial crystallization hit.  If crystalline-like material appears in the plate prior to the addition of protein solution, it is not a hit that should be pursued.  Plates are also imaged at the following intervals after addition of the protein solution: one day, one week, two weeks, three weeks, and four weeks.

12. Why do we image the plates more than once?

The outcomes of crystallization experiments will change over time.  The microbatch-under-oil experiments are, as the name implies, ‘batch experiments’.  However, they are not ideal static batch experiments.  The experiment drops will slowly dehydrate.  As they dehydrate and the volume decreases, the relative concentration of any non-volatile solute increases.  This can decrease the solubility of the biological macromolecule.  It can drive a drop that is not sufficiently supersaturated for spontaneous, homogeneous nucleation (undersaturated, saturated, metastable) to a point where it is sufficiently supersaturated for crystallogenesis.

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13. What types of outcomes can be expected from the screening experiments?

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14. How do I translate from microbatch-under-oil to vapor diffusion?

First, ask yourself if this is really necessary.  Although you may prefer other methods, there are some real advantages to using the same crystallization method for screening and optimization.  There is information available on converting from microbatch-under-oil to vapor diffusion experiments available in the literature [Acta Cryst. D54, 8-15 (1998)].

A few basic concepts:

When you set up a batch experiment by combining equal volumes of protein and cocktail solution, you are diluting any solute not contained at the same concentration in both solutions.  For example, if the protein concentration was 10 mg/ml in the stock solution, it will be 5 mg/ml in the batch crystallization experiment.  Ideally, batch experiments will not dehydrate.  The starting and final concentrations of the protein in the batch crystallization experiment remain at 5 mg/ml, unless a phase change (e.g., crystallization, precipitation) occurs and drives some of the protein from the solution state.

In the case of a vapor diffusion experiment, this same dilution occurs when you combine the protein and cocktail solutions to prepare the experiment drop.  For example, if equal volumes of protein and cocktail solutions are combined to prepare the experiment drop, and the protein concentration was 10 mg/ml in the stock solution, it will be 5 mg/ml at the start of the vapor diffusion experiment.  The experiment drop will be left to dehydrate over a reservoir solution (typically, but not necessarily, the cocktail).  This reduces the volume of the experiment drop.  If no phase change occurs, the final concentration of protein in the experiment drop would be ~10 mg/ml after the vapor phase equilibration is completed.

15. What cocktails are used in the screen?

There are three subgroups of cocktails:

    • Salt/Buffer
      35 salts at 3 concentrations x 8 pH’s (incomplete factorial)
                                        
    • PEG/Salt/Buffer
      6 PEG’s at 2 concentrations x 36 Salts x 8 pH’s (incomplete factorial)

    • Commercial screens from Hampton Research

16. How do you make the cocktails, and how can I reproduce them in my lab?

We purchase commercial screens directly from Hampton Research (http://www.hamptonresearch.com).

All of our other cocktails are prepared by making up concentrated stock solutions of PEG, salt, and buffer.  The individual components are combined and diluted, if needed, to prepare the individual cocktail solutions.  The pH of the buffer stock is adjusted prior to combining the stock solutions to prepare the cocktail solution.

17. What types of samples are accepted for screening?

We screen an increasingly diverse set of biological macromolecules to identify crystallization conditions.  This has included both soluble and membrane proteins as well as protein complexes.

18. How pure should the sample be for screening?

The sample should be monodisperse by dynamic light scattering [Acta Cryst. D50, 469-471 (1994)].   Sample stability is as important as initial purity.  We strongly encourage verification that the sample will remain stable for a period of time after purification.  If the sample rapidly decomposes, a solution environment with the proper pH and chemical additives to stabilize the sample should be identified prior to crystallization efforts.

19. How much sample is required for screening?

We require 600 microliters of solution at a concentration of 10 mg/ml.  The sample concentration will vary with the solubility of the individual samples.  We suggest 10 mg/ml as a good starting point.

20. How should I prepare the sample for screening?

From a crystallization perspective, the solution would ideally be pure water.  This permits the cocktail to dominate the solution environment and dictate the chemistry of the crystallization experiment.  Often, it is necessary to add a low concentration of buffer, or other chemical additives to stabilize the protein for crystallization trials.  Co-factors and inhibitors can help to stabilize the three-dimensional conformation of the molecule and can be an effective way to improve crystallization of the sample.  Things to avoid are high concentrations of strong buffering agents and additives known to form insoluble compounds with the crystallization cocktails (phosphate and borate are good examples of this).  It is imperative to prepare the sample in a solution environment where it will be stable for several days.

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21. How should I ship the sample?

This depends on the sample’s stability.  We receive samples on both wet and dry ice.

22. Who sends the samples?

The majority of our samples come from structural biologists.  Over 750 structural biologists have signed up to be on our active mailing list.  We receive a significant number of samples from structural genomics groups.  For example, we screen all of the samples for the NESGC (Northeast Structural Genomics Consortium) to identify initial crystallization conditions.

23. How do I submit a sample?

Contact us by email at htslab@hwi.buffalo.edu to get further information.  Each month, a message will be sent to our user community to request samples for the screening queue.

24. Are there any costs?

There is a nominal fee ($297 USD) that does not quite cover the cost of setting up the screen for each sample.  The fee will not be charged until the final imaging of the experiment plate, four weeks after the experiment is set up.

25. How do you track the samples?

All of the data from every experiment is tracked through a secure, custom-designed database/LIMS.

26. How do you avoid data loss, in particular for the image data?

Image data is archived with multiple fail-over systems to avoid data loss.  These failover systems include dual RAID5 subsystems (the secondary subsystem snapshots the primary), tape, and optical disc backup of the image data to minimize any risk of data loss.

27. How do I get my image data?

An email is automatically generated by our database as soon as your experiment plate has been imaged and packaged to notify you that your outcomes are ready to review.  During the imaging of a microassay plate, 1536 individual TIFF images are sent from the reader table to a file server.  The image data are processed and packaged with a file that contains the chemical cocktails associated with the experiments.  The TIFF images are converted to JPEG format and placed on a secure ftp server for password-protected access by users.  The results are available to geographically distant investigators as soon as they are available in-house.  A CD containing all of the image data is sent to each investigator after the fourth week's reading is completed.

28. How do I view my image data?

Image data can be viewed using MacroScope, a program that was developed at the Hauptman-Woodward Institute.  This software is admittedly dated and is only compatible with a PC using a Windows OS.  The software is available free of charge and has been used, literally, to view millions of images.

29. Are you planning to develop any new image viewing software?

We are actively developing a new image viewing software package.  The software will be web-based and hence be multi-platform compatible, operating on any computer hardware/OS that has a web browser.  This software will have a significant number of new features.

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30. What information do you track for the biological macromolecules that are submitted for high-throughput        screening?

The information supplied by users on the sample submission form.  We have collected and entered this data for more than 8600 samples (March 2007) that have been set up in high-throughput screening trials.

High Throughput Screening Lab

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31. What happens during a typical month in the screening laboratory?

Email notification and sample tracking:
An email is sent out to the community of structural biologists who have expressed an interest in submitting a sample(s) to the screening laboratory.  This email notifies the community on a monthly basis that screening will take place so that interested researchers can enter the queue.  The users contact the laboratory to reserve a plate for screening experiments.  When the queue is full, a notification is sent out to the community to notify them of this fact.

Preparation of the experiment plates with oil and cocktail:
Meanwhile, in the laboratory, research associates use two Matrix TangoTM liquid-handling systems (equipped with a 384-syringe bank head) to aspirate USP grade mineral oil from a single-well source plate and dispense 5 microliters of the oil into each well of a 1536-well microassay plate purchased from Greiner BioOne (catalog number 790801).  This plate will contain the crystallization experiments.

Cocktail solutions are aspirated from four unique 384-well source plates, and 200 nanoliters of 1536 chemically distinct cocktails are dispensed into oil-filled 1536-well experiment plates.  Plates are stored at 4 degrees C until use, and they will have the macromolecular solution added to them within two weeks of the cocktail delivery.

Addition of macromolecules to the experiment plates:
Solutions of macromolecules sent to us for high-throughput crystallization screening trials arrive by overnight delivery.  Samples are sent to us by different investigators at room temperature, on wet ice (or more commonly ice packs), or on dry ice (this is dependent upon the researcher and the sample).  Samples are received, and information about the sample is entered into our database.  At this time, samples are either set up immediately or stored, as directed by the investigator, until screening trials can commence.

An experiment plate containing oil and cocktail is imaged immediately prior to the addition of the protein solution.  While the plate is being imaged, we prepare the macromolecular sample for delivery.  If the sample was received frozen, we will either thaw it on wet ice or do a 30o C quick thaw [Acta Cryst. D60, 203-204 (2004)] as directed by the investigator who supplied the sample.  The sample (in a 1.5 ml microcentrifuge tube) is centrifuged to pellet any precipitate that may have formed during transit.  Notes are taken on the condition of the sample (clear or precipitated, any noticeable color, or other observations) and entered into the database.

Next, the sample is manually loaded into 12 wells (one row) of a 96-well source plate.  The Matrix TangoTM liquid-handling system (equipped with a 12-syringe bank head to minimize dead volume loss) is used to aspirate protein from the source plate and to make 128 deliveries (200 nanoliters) to each well of the 1536-well experiment plate (already containing oil and cocktail).  The experiment plate is removed from the liquid-handling system and centrifuged at low speed to ensure that the cocktail and protein solutions merge.

The laboratory has the capacity to comfortably set up ~20 samples (30,720 crystallization experiments) in a single day.  We have the capacity to image 3 x 28 (84) plates (129,024 experiments) every evening.  The imaging schedule becomes complicated when each of 200 plates (many of them prepared on different days) have to be imaged on a regular basis.  The schedule is provided through the htslab database.

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32. Can you recover crystals from the plates?

It is very difficult to recover crystals from the 1536-well experiment plates.  The wells are small (~ 2mm square at the top, 0.9mm circle at the bottom) with a conical interior.  Manipulation of the crystals to remove them from the well often results in destruction of the crystals.  We have been working to develop a solution to this issue, but thus far our efforts have not been successful.

33. How can I tell if it is a protein or a salt crystal?

We strongly encourage you to set up the experiment using identical conditions to the screening experiment.  This includes the same protein solution, cocktail solution, temperature (23oC), crystallization method (microbatch-under-oil), and drop-volume ratio (2 microliters each of protein and cocktail solution are recommended).  Assuming you are able to reproduce the crystal that you observed in the images, we now recommend going through standard verification methods to make certain the crystal is composed of protein and not an inorganic compound.  These verification protocols may include:

  • Dehydration – Remove one of the crystals, and use a fiber wick to remove liquid from the crystal.  If the crystal is proteinaceous, it will dehydrate, becoming opaque and forming cracks as structurally important water evaporates from the crystal and the lattice collapses.

33A

  • Physical manipulation – Use a glass fiber and move the crystal.  See if the crystal will crush (similar to a piece of cake – evidence that it may be a macromolecular crystal) or fracture with an audible cracking sound (similar to crushing a crystal of sodium chloride – readily available for easy comparison) which is often indicative of a salt crystal.

33B

  • Examine the crystals under cross-polarization – This can in many, but not all, cases be used to determine whether or not the material you are viewing is crystalline or amorphous.  In addition, protein crystals (a) tend to be less intensely birefringent than salt crystals (b). 

33C

  • Don’t use protein in your crystallization experiment– If one of the protein buffer components is forming an insoluble salt when it is combined with the cocktail solution, you may be able to quickly and simply prove that the crystals are not protein.   Set up two crystallization experiments: (1) use the same protein solution and cocktail solution that produced the crystals in question, and (2) use protein buffer (i.e., prepare the crystallization mixture, but without the protein).   If you observe identical crystals in both of the experiments, the crystals are not protein crystals.
  • Dye absorption – A dye such as methylene blue (Hampton Research - Izit Crystal Dye, Cat # HR4-710) will sometimes become concentrated in the solvent channels of protein crystals.  There are many examples of protein crystals that do not effectively take up dye, but if the crystal does take up dye, it is highly probable that it is composed of protein and not salt.

33E

  • SDS-PAGE – Retrieve some of the crystals and place them into a microcentrifuge tube.  Centrifuge to pellet the crystals and discard the mother liquor.  Wash the pellet in a solution of concentrated cocktail.  If the crystals crack it is not important.  This is an important step to remove traces of the mother liquor (containing protein) from the crystal’s surface.  Centrifuge again to pellet the crystals and remove the wash solution.  Dissolve the crystals in SDS-PAGE sample buffer and run a gel.  If you see a band at the molecular weight of your sample, it is probable that you have protein crystals.

33F

  • X-ray diffraction experiment – A room temperature X-ray diffraction experiment can provide valuable, definitive proof of your crystal’s composition.

34. What crystallization cocktails are currently in use?

Click here to download a composite list of generation 10 of the HTS lab's 1536 crystallization cocktails.

35. Are you developing new screening cocktails?

We are actively working within CHTSB to develop improved crystallization cocktails. One effort, headed by Dr. Alexander McPherson (UC Irvine), is looking at the efficacy of small molecules to promote intra and intermolecular interactions of macromolecules.  Cocktails that focus on the small molecule crystallization promoters are under development and will be incorporated into the Hauptman-Woodward Institute's high-throughput screening laboratory.

We are also developing crystallization cocktails that are specifically designed to crystallize protein detergent complexes.  These cocktails are being developed after extensive laboratory investigations to identify the chemical phase separation points for a number of different detergents. The detergent phase separation boundary has been identified as an effective solution environment for crystallization of the protein-detergent complex.  This work is being led by Dr. Michael Malkowski.

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