At the risk of repeating ourselves, the most important factor is your sample. If your protein is impure, poorly folded or unstable, the chances of obtaining useful crystals are close to zero.
Try to have your sample as a single band on an overloaded Coomassie stained gel. Crystallisation is a purification process, but you need to have your sample 90-99% pure to start. If crystals form in the presence of impurities, then you’ll probably end up with poorly diffracting samples. Our preference is to conduct mass spec analysis, which will quickly highlight/confirm sample impurities.
Your protein should be stable (it may take weeks for nucleation and crystal growth to occur), ideally in a formulation which has a low concentration of buffer (50 mM or less) and a low concentration of salt (200 mM or less). You may need co-factors, inhibitors, or other chemicals to ensure that your protein is stable. You may also need to add reductant (βME, DTT or TCEP). Glycerol tends to increase the solubility of proteins, but keep in mind you could be simply keeping a poorly folded protein in solution – also, it may prevent your protein from coming out of solution in a crystallisation experiments. Phosphate is a very common buffer but is problematic in crystallisation, as it
can will produce lovely salt crystals with magnesium and calcium: we discuss this in a lot more detail in the UV imaging section.
Our routine method for assessing protein stability is Thermofluor (aka Differential Scanning Fluorimetry, DSF). We test the stability of the protein as a function of temperature against a variety of different pH, chemical and salt conditions. If something appears to help increase the thermal stability, we recommend exchanging the formulation before conducting crystallisation trials.
Thermofluor can also be used to define chemical partners (specific or not) for your protein: often the protein needs a bit of conformational help from a co-factor, an inhibitor or a particular cation before it will crystallise.
Crystallisation is a precipitation event, and the protein solution has to become supersaturated for crystallisation to occur. For screening, it is best to aim for a high protein concentration, then set up different concentrations in your initial screening (in general, higher concentrations are more efficient for the entire biophysical process). If you can’t get your protein more concentrated than 1 mg/ml, it may be time to reassess the purification protocol, and your current formulation.
Keep in mind that the ideal concentration for crystallisation varies for every protein – some (rare) proteins have crystallised from solutions as dilute as 0.3 mg/ml and others require concentrations of well over 100 mg/ml. You should check that your protein is at an appropriate concentration before setting up large numbers of droplets. There are a couple of easy ways to test to see if the concentration of your protein is appropriate.
- Do a pre-crystallisation trial (PCT) – in this you set up a your protein against 2 “typical” crystallisation conditions, and see how it behaves – Hampton Research sells a kit for this.
- Run a single sparse matrix screen (e.g. Shotgun). If your protein is at a useful concentration, anywhere between 30-60% of the drops will be clear and the remainder will be precipitate.
NEXT: Crystallisation Strategy