What is the National Crystallization Center?
The National Crystallization 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 National 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 a recent paper describing the 20+ year history of the NCC.
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 400 μl of solution at a suggested concentration of 10 mg/ml (note this is a decrease from the previous requirement of 500 μl due to new robotics instrumentation). 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 recommend shipping samples to arrive Monday through Thursday so that any delay does not cause a shipment to be delayed over a weekend.
What’s the best way to send your sample for crystallization screening? Here’s a new paper we recently put together with colleagues from synchrotrons and cryoEM centers to provide guidance on this topic!
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 through the NIGMS National Resource.
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.
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.
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?
See Analyzing Your Results for information about viewing image data.
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.
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:
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.
How should we acknowledge using the Crystallization Center?
Citations and acknowledgment of the NIH National Resource grant help us to track publications and PDB depositions, an important metric of productivity that will help secure future funding for the Crystallization Center.