Frequently Asked Questions
FAQs: The Laboratory
What is the High-Throughput Crystallization Screening Center?
The High-Throughput Crystallization Screening Center 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 laboratory information management system/database to quickly set up and record the outcomes of 1,536 unique crystallization screening experiments.
How long has the laboratory been in operation?
The Crystallization Center accepted the first samples for crystallization in February 2000 and has been operating continuously since.
How many crystallization experiments have been set up in the high-throughput laboratory?
From February 2000, the laboratory has been used to set up well over 26 million crystallization experiments on more than 18,000 biological macromolecules.
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 (A deliberate approach to screening for initial crystallization conditions of biological macromolecules).
FAQs: Sample Requirements
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.
How pure should the sample be for the screening?
The sample should be monodisperse by dynamic light scattering (see the paper entitled Crystallizing proteins – a rational approach?). 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.
How much sample is required for screening?
We require 500 μl of solution at a suggested concentration of 10 mg/ml. The actual sample concentration will vary with the solubility of the individual samples. We recommend using a pre-crystallization screening test, such as the Hampton Research PCT to determine a protein-specific sample concentration.
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 dictates the chemistry of the crystallization experiment. Practically, 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 the 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 solution environment where it will be stable for several days.
How should I ship the sample?
This depends on the sample’s stability. We receive samples on both wet and dry ice, and with gel packs to control the temperature. Shipments should be made in insulated Styrofoam boxes and sent overnight. We do not recommend shipping to arrive Monday through Thursday so that any delay does not cause a shipment to be delayed over a weekend.
How do I submit a sample?
For details about how to submit a sample for academic or proprietary users, please see www.getacrystal.org, the Crystallization Center homepage.
Are there any costs?
There is a fee that covers the cost of setting up the screen for each sample. For academic, government and not-for-profit laboratories this fee is subsidized through current support from NIH and NSF.
FAQs: The Experiment
Why is it beneficial to set up crystallization experiments using high-throughput methods?
By using automated liquid-handling systems, we are able to set up crystallization experiments precisely and reproducibly using a minimum volume of macromolecular solutions. The laboratory personnel starts 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.
What is a microbatch under oil and why do you use it?
Microbatch-under-oil is a simple crystallization method developed by Naomi Chayen, Patrick Shaw-Stewart, and David Blow (See the papers entitled An automated system for micro-batch protein crystallization and screening, and Microbatch crystallization under oil — a new technique allowing many small-volume crystallization trials). The technique 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 (e.g. A novel technique to control the rate of vapour diffusion, giving larger protein crystals).
Why do you 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)
What type of oil do you use?
We use Paraffin Oil purchased from EMD Chemicals Inc. (catalog number PX0045-3).
What cocktails are used in the standard (soluble) protein screen?
There are two subgroups of cocktails for soluble protein studies: The first a PEG/Salt Buffer set with six PEG’s at two concentrations with 36 Salts at pH’s (using an incomplete factorial setup). The second is commercial screens purchased from Hampton Research which include: PEGRx HT, PEG?Ion HT, Crystal Screen HT, Index, Salt Rx HT, Silver Bullet with PEG3350 pH 6.8 precipitants, Grid Screen Ammonium Sulfate, modified Slice pH, modified Ionic Liquids, modified Polymer screen. Please Note: The Ionic liquids screen was modified with the addition of 0.09M buffer and 27% (w/v) PEG 3350. The Slice pH screen was modified with the dilution of the buffer from its initial 1.0 M to 0.5M concentration with the addition of 15%(w/v) PEG 3350 to promote supersaturation in the batch experiments. The Polymer Screen was formulated using 24 polymers 200≤Mr≤200000 at 2 concentrations all in 10% (v/v) Tacsimate pH 7.0. For membrane samples, a different cocktail approach is used.
FAQs: The Results
How do you record the experiments’ outcomes?
Outcomes of the screening experiments are recorded using Rock Imager systems with visual (brightfield), SHG, and UV-TPEF. All three imaging modes can be carried out at 23°C using the Rock Imager 1000 with SONICC and brightfield using a Rock Imager 54 in a temperature-controlled room typically operated at 14°C. These systems replace in-house developed imaging tables (as of January 2020 all imaging is performed in Formulatrix Rock Imagers). A specialized software program is used to analyze the results.
When do you 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 the addition of the protein solution: one day, one week, two weeks, three weeks, four, and six weeks. SHG and UV-TPEF images are recorded at the four week timepoint for samples incubated at 23°C and at the six week timepoint for samples incubated at 4°C or 14°C.
Why do you image the plates more than once?
The outcomes of crystallization experiments will change over time and crystallization itself is a stochastic process. Also, while the microbatch-under-oil experiments are, as the name implies, ‘batch experiments’, 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.
What types of outcomes can be expected from the screening experiments?
Each image of a crystallization well can be classified into seven predefined categories or their combinations. These categories are clear, phase separation, precipitate, skin, crystals, junk, and unsure. With the exception of “clear”, combinations (two or more) of all the other categories are allowed. Image classification into these categories has been discussed in considerable detail in two Center publications (Establishing a training set through the visual analysis of crystallization trials. Part I: approximately 150,000 images, and Establishing a training set through the visual analysis of crystallization trials. Part II: crystal examples). Detailed advice on interpretation has been given in another paper by the Center entitled “What’s in a drop? Correlating observations and outcomes to guide macromolecular crystallization experiments“.
How do you track the samples?
All of the data from every experiment is tracked through a secure, custom-designed database/LIMS. We run control experiments to track the conditions of the cocktails and the robotics so that we can perform quality control. We generally have very low experimental error (under 5% and more typically < 1%) and use the controls to rapidly identify and address any issues that arise.
How do you avoid data loss, in particular for the image data?
Image data is archived with multiple fall-over systems to avoid data loss. These systems include tape and optical disc backup of the image data to minimize any risk of data loss.
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, individual images are sent from the Rock Imager to an internal file server. The 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.
How do I view my image data?
Image data can be viewed using MacroscopeJ, a program that was developed at the Hauptman-Woodward Institute and is available free of charge. The software is java-based for multi-platform compatibility. This software has a significant number of new features from the old Macroscope program that was used in the past. Please go to the Analyzing Your Results tab to download the MacroscopeJ program and manual.
FAQs: Making Use of the Results
Can you recover crystals from the plates?
It is very difficult to recover crystals from the 1536-well experiment plates. The wells are small (~2 mm square at the top and 0.9 mm circle at the bottom with a conical interior). Manipulation of the crystals to remove them from the well often results in the destruction of the crystals. The Crystallization Center has described an approach using an inverted microscope system in a paper entitled “A new view on crystal harvesting” and has had success with capillary-based extraction but it is recommended to optimize conditions separately as described here. It is also possible to collect data directly from the plates but they are not designed with this purpose in mind.
How can I tell if it is a protein or a salt crystal?
We have integrated a Formulatrix Rock Imager 1000 with SONICC into our High Throughput Crystallization Screening Center pipeline. Using this instrument, we have imaging technologies that can be used to determine if an object is crystalline [second order non-linear imaging of chiral crystals (SONICC); Haupert LM and Simpson, (2011) Methods, 55, 379-386] and if it is protein [UV-two photon excited fluorescence (UV-TPEF); Madden JT, DeWalt EL and Simpson GJ (2011) Acta Cryst. D67, 839-846). Positive (white) signal from these independent imaging technologies can verify protein nanocrystals < 1 μm in size. This imaging takes place once, at either 4 or 6 weeks.
What crystallization cocktails are currently in use?
Please go to the Crystallization Cocktails page to download the complete list of our current and/or most recent crystallization cocktails for soluble and membrane proteins.
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 (Comparative Studies of Protein Crystallization by Vapour-Diffusion and Microbatch Techniques) and a protocol that the Center recommends here.
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, and the protein concentration was 10 mg/ml in the stock, 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.
How do you make the cocktails, and how can I reproduce them in my lab?
We purchase commercial screens directly from Hampton Research and the MemGold screen for membrane protein samples from Molecular Dimensions. All of our other cocktails are prepared in house or custom made by Molecular Dimensions by making up a concentrated stock solution 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.
Who sends the samples?
The majority of our samples come from structural biologists. Close to 2,000 structural biologists have made use of the Crystallization Center. We collaborated with structural genomics groups, including the Structural Genomics of Pathogenic Protozoa (SGPP) and the Northeast Structural Genomics Consortium (NESG), to develop and improve our technologies to determine initial crystallization conditions.
Are you planning to develop any new image viewing software?
We are continually developing the new MacroscopeJ program and expanding on the analysis features that it offers. We do have a new imaging software undergoing beta-tests. This incorporates the MARCO algorithm to automatically classify images. Notification will be sent to members of our mailing list as soon as this is available.
What information do you track for macromolecules submitted for screening?
We track the information supplied by users on the Sample Submission Form. We have collected and entered this data for more than 18,000 samples that have been set up in high-throughput crystallization screening trials. We also track citations to the Crystallization Center and are glad to promote new results and publications that benefited from the Crystallization Center on our home page.