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42 U. Krengel and A. Imberty 5.5. Protein or salt crystals? Obtaining protein crystals is very exciting! But are those really protein crystals? Many experimenters have been very disappointed to find out that their precious crystals turned out to be salt. Here are some simple methods to test this: (1) X-ray diffraction: If the crystal is big enough to test it in the X-ray beam, the diffraction pattern will show immediately if you crystallized protein or salt. Protein crystals exhibit many closely spaced diffraction spots, while for salt, with its small unit cell, the reflections are far apart (and much stronger!). If you do not see any spots, this can indicate a very weakly (non-)diffracting protein crystal. You should make sure, though, that you did not miss the spots by choosing a too small oscillation range for data collection (20 oscillation should do the trick). (2) Brute force: Crush the crystal – if it is salt, it will be hard to break and you will hear a distinct clicking sound when snapping the needle on the cover slip. A protein crystal is soft and easily crumbles under a needle (but watch out: never use this method if you only have one single crystal!). (3) Stains: Protein-staining dyes such as Methylene Blue (staining solutions can be purchased ready-to-use as “Izit” from Hampton Research) can be used in order to identify protein crystals in a non-destructive way. If the crystals turn blue, you can be sure that it is protein. If not, it might be salt, but you could also be dealing with very faintly staining protein crystals, especially if the crystals are small or thin. (4) Check the reservoir: If you find crystals in the reservoir, the odds are high that the crystals in the drop are not protein either. (5) Gel: Collect the crystals in an Eppendorf tube, wash them several times in mother liquor (by centrifuging the tube a couple of minutes at ca. 10,000 rpm and then removing the supernatant), then dissolve them and load them on a gel. If you see a distinct band characteristic for your protein – voilà, if not, you might either have crystallized salt, used too little material from the start or lost the protein in the procedure (usually, the method works, though). (6) Negative control: Set up experiments with the same conditions (including salts and detergents or anything else you might have added to stabilize the protein, e.g. by using the filtrate solution from the concentration step), but without protein. If you get crystals, you know it is not protein. 5.6. Seeding techniques and other tricks In some cases, it can be beneficial to induce crystal formation by introducing seeds into the crystallization solution. Three main techniques exist: (1) Macroseeding: Small crystals are introduced into a pre-equilibrated crystallization drop (e.g. with help of a loop or capillary). The crystal seeds should preferably first be washed several times in mother liquor (best in a solution with slightly lower concentration,
Crystallography and Lectin Structure Database 43 such that the surface layers start to dissolve and clean surfaces are available for crystal re-growth). The concentration of the drop, to which the seed crystal is to be added, should be in the metastable range (Fig. 8). Suitable conditions can be found either through parallel seeding experiments into drops with various concentrations or by first plotting out the phase diagram. This technique works best if perfect small crystals can be obtained, but no crystals large enough to obtain high-resolution X-ray diffraction data. (2) Streak seeding: Streak seeding works best if large crystals are available, even if they are twinned or not useful for other reasons. For capturing the seeds, a hair or a cat’s whisker is taken and gently stroked over the original crystal. Also acupuncture needles work very well for streaking. The hair or needle is then drawn through a series of preequilibrated crystallization setups. In this way, it will contain less and less seeds from one drop to the next, giving rise to streaks of crystals in the first drop(s) and single crystals in the last drop(s). This method should definitely be tried for old crystallization setups, which yielded crystals only in some of the drops. (3) Microseeding: Microseeds are preferably generated by crushing a large crystal, e.g. with a needle, but they can also be obtained by crushing (many) small crystals with a glass rod or homogenizer. The drop containing the microseeds is then diluted with the mother liquor, to final dilutions of 1:10, 1:100, 1:1,000, and 1:10,000 (always using a fresh pipette tip when preparing the next dilution in the series) and vortexed in order to obtain a homogenous mixture of the seeds. Tiny amounts of these solutions are then added one by one to pre-equilibrated crystallization drops in order to test which dilution gives the best results. The seeding solutions can sometimes be stored for several days to months. Just be careful to vortex them again before using them the next time. Good overviews over crystal seeding are also given by Bergfors [126] and Stura and Wilson [127]. Very nice illustrations of the seeding technique are shown in Ref. [128]. An interesting variation is so-called cross-seeding (seeding from one crystal form to another, which grows under a different condition) or to use other materials like porous silicon, minerals or crushed frozen hair for seeding [129, 130]. If seeding does not work, but small crystals can be obtained, one may also try to layer the experiments with various oils to slow down equilibration [131] or to transfer drops from a high to a lower precipitant concentration before crystals become visible macroscopically [132]. This method even works for very short crystallization times [133]. 5.7. Co-crystallization Co-crystallization experiments are performed in order to obtain the crystal structures of protein–ligand complexes. This method works exactly like a normal protein crystallization experiment. The only difference is that one has to add a ligand to the
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