To some, X-ray crystallography may be a rather dry topic.
However, the process generally entails lots of liquids. In particular, the
commonly used practice of crystal soaking entails transferring protein crystals
to a new solution containing dissolved ligands, which is both tedious and can
cause crystals to shatter or dissolve. A new paper by Jean-Francois Guiçhou at
Université de Montpellier and collaborators in Acta Cryst. D aims to streamline the process, and so lower barriers
for obtaining structural information that could guide drug design.
Rather than manually transferring crystals to new solutions,
the researchers pre-coated crystallization plates with ligands and then grew
protein crystals in them. They first dissolved the ligands, transferred these
to the wells, and allowed the solvent to evaporate. Although they tested a
variety of solvents, including acetone, tetrahydrofuran, ethanol, acetonitrile,
2-propanol, water, and DMSO, only the last two proved suitable; most of the
rest wicked up the well, spreading over too large of a surface (though methanol
has been used by Beryllium, née Emerald). DMSO is, of course, the most commonly
used solvent for storing small molecules, and so should work for most ligands.
DMSO is not very volatile, but only 1 µl was used per well, and putting the
plates in a fume hood for a week left behind dry ligand.
To make things easier still, the researchers used special
crystallization plates that could be put directly into an X-ray beam (in situ
crystallography), further diminishing the amount of manipulation required. The
technique was tested against four different proteins: the old standard hen
egg-white lysozyme and the drug targets cyclophilin D, PPARγ, and Erk-2.
For lysozyme, the water-soluble fragment benzamidine was
used, and the resulting structures showed the fragment binding in a similar
manner as previously described. So did structures of PPARγ bound with the high
affinity ligand rosiglitazone. Cyclophilin, though, was not as successful: of
nine fragments attempted, only one produced a structure. In contrast, three
fragments produced structures using conventional approaches. ‘Dry’
crystallization was more successful with two more potent (micromolar or better) cyclophilin
ligands. Interestingly, dry crystallization succeeded with one ligand that had
previously been characterized only by co-crystallization; even week-long
soaking experiments had not worked.
Finally, Erk-2 was screened against 14 ligands designed as
hinge-binders with low solubility in water. Crystals were obtained with five of
the ligands, and four were large enough to generate good-quality structures.
Overall this seems like a convenient approach, though it
does seem prone to false negatives. What do the crystallographers out there
think – is this a practical solution?
While this paper has accurately highlighted some difficulties in screening large numbers of compounds by crystallography, I think their solution has some issues and there may be more practical solutions already available.
ReplyDeleteHow do they determine if the compound is soluble in the crystallisation buffer and if it has been completely re-dissolved? This could be one source of the false negatives. Also, if the ligand induces any conformational changes to the protein it may crystallise as a different crystal form or not at all under the same crystallisation condition as previously observed, another potential source for false negatives. Even in cases where the crystallisation condition has been optimised, getting crystals in 100% of drops is rare.
A recent publication (http://pubs.acs.org/doi/abs/10.1021/cb500072z) described the identification of BRD4 inhibitors through screening by co-crystallisation. In this work they dispensed compounds directly into the crystallisation drop using a fairly standard crystallisation robot (mosquito LCP from TTP Labtech) without the need to evaporate the solvent prior to setting up the crystallisation drop. This protocol could easily be carried out using plates suitable for in situ data collection so I’m not sure what benefit pre-coating the crystallisation plate offers in comparison.
Frank von Delft et al., at Diamond Light Source have also setup a platform to enable medium-throughput fragment screening by crystal soaking that has been validated on a number of targets (http://www.diamond.ac.uk/Beamlines/Mx/Fragment-Screening.html). Rather than manually transferring hundreds of crystals to new drops or pre-coating crystallisation plates with compound, you can dispense very small volumes (2.5 nl) of fragments directly into the crystal drop using an acoustic dispenser (an ECHO). The platform at Diamond still requires manual mounting of the crystal but this has been accelerated using a robot-assisted system and data collection is completely automated. I’m not sure if plates suitable for in situ data collection are compatible with the platform but in situ data collection is not without its own problems. Also, crystal soaking does have different limitations to co-crystallisation. At Evotec we have had good experiences using the platform at Diamond and have observed hit rates >10 % when screening our own fragment library against several targets.
I'm surprised by the use of DMSO as it hygroscopic and apparently required a week under the hood to evaporate.
ReplyDeleteMy experience has been that fast evaporating solvents can be used by adding water to the solution. Put down compounds in 100% methanol and you can watch under the microscope as the drops wicks up and around where it is supposed to be. But put them down in 70% methanol and the drops just sits there and is dried after 2 minutes.
The other thing I would do that isn't mentioned in this paper but touched upon by Daren is put down different amount of compound into the different drops. False negatives are always a problem in crystallography and compound aggregates can certainly inhibit crystal formation. So by titrating the compound onto the plate you can find the sweet spot where crystals grow with maximum ligand present.
The other thing one could do to maximize crystallization is to add protein microcrystals to the drop as you set them up as this normally increase the number of successful drops in a tray. Even if the crystals are of a different crystal form thanks to the ligand this can be successful.
Hi,
ReplyDeleteIt's always nice to see these sorts of methods pop up in literature. We have used the same approach years ago at P&G and later at Pfizer, to grow ligand-loaded crystals (and also to prepare pre-coated plates for additive screening) -- but we never bothered to publish, so kudos to the authors on this one! To avoid false-negatives we used the following trick: when dissolving compounds in DMSO, AcN or other solvent we would add 0.5-1% of glycerol or sugar (e.g. glucose or sucrose) to the solvent. The resulting drop never quite dries out (but is usually quite viscous so it stays in place) and then when the protein + precipitant mix hits the drop it dissolves very nicely.
Artem
Artem's suggestion of adding glucose or sucrose to trap the ligands with a bit of water is a nice trick. It might well overcome my major concern with the 'dry solution' method that fragments are small molecules, often with quite high vapour pressures (you only need to smell your fragment collection to appreciate this!).
ReplyDeleteAstex first encountered this problem many years ago when we tried to dispense fragments by pipetting them as AcN/MeOD solutions, followed by evaporation of the solvent - and, as it turned out, many of the fragments. As we have reduced the average size of fragments in our library fragment volatility has become a more common issue. To overcome it, we have adopted quite strict procedures to ensure that pre-weighed solid samples and stock solutions (in DMSO)are stored at low temperatures to minimise the sublimation or evaporation of fragments.
Sorry, a browser problem prevented me leaving my details with the previous comment.
ReplyDeleteJoe's comment is quite good, but keep in mind that the depth and the width of the crystallization well plays a significant role as well with regard to using methanol-based fragment stock solutions. For example, we don't use methanol-based fragment stock solutions in combination with MRC crystallization plates from Hampton Research because those plates are shallow and wide, causing considerable wicking. Rather, we use the XJR plates from Rigaku Reagents for methanol-based fragment stock solution because these plates are narrow and deep and seem to have minimal spreading of the drops. It takes about 10 minutes for the XJR methanol plates to dry out in a fume hood compared with 2 minutes for the MRC plates, but the more concentrated drop is worth it.
ReplyDeleteAnother worthwhile solvent is NM2P (N-Methyl-2-pyrrolidone) if methanol or DMSO are problematic.