Microscopy: Labeling Proteins with Fluorescent Probes (Timothy Mitchison)


I thought some practical tips on protein labeling might interest people. Usually with Thiol labeling we’re aiming to get a probe in just one place on a protein. So, you may be lucky and have a protein that has a naturally reactive cysteine like like actin for example, which has one reactive near the C-terminus. It’s getting more common now for people who really want to study a protein in detail for biophysics to actually engineer out all the surface exposed cysteines and then engineer in one place a reactive cysteine. So that’s some amount of work on the cloning that can be worth it and you can actually walk a fluorochrome in two different places on the molecule by depending on where you put the cysteine. Now it’s really important to protect your cysteine until you do the labeling. So you want to include reducing agent in all the buffers until the labeling step and then remove your DTT or β-mercaptoethanol in the last step or else titrate it out with probe. Cysteine labeling is typically done at pH 7 to 8. If you go too high in a pH you risk labeling tyrosine residues. The nice thing about these cysteine probes is they are not very reactive with water so you don’t need to use very much. Just enough essentially, 1.5 to 3 equivalents is typical. And because you’re not using very much you have to react for a long time overnight or room temperature. So these are kind of cookbook recipes. With the typical lysine labeling reaction where you’re not usually aiming for site-specific, it’s the considerations are a little bit different. There’s plenty of surface lysine on most proteins so you can often want to label several of them. For example on an antibody you might want to aim for about 4 fluorescent molecules per antibody. Usually the antibody will still be fine and you get plenty of fluorescence then. These NHS ester probes, unlike the Thiol probes, they’re quite chemically reactive. It’s important to store them in a high-quality dry solvent. DMSO is the most common, or dry in the freezer. We often aliquot them in DMSO and store them at minus 20. You want to label at neutral pH or above. You want to use a non-nucleophilic buffer. We often use HEPES in my lab, bicarbonate, borate. Important not to use DTT or thiol. I mentioned they’ll decompose the probe. The issue of the pH is this: the higher your pH, the faster you’ll react with the protein but also, the faster the probe will be decomposed by hydroxide ions. And in our experience it often actually works a little better at let’s say pH 8 than it does at pH 10 because you start getting to rapid hydrolysis. And just as a rule of thumb here, these NHS ester probes, their half-life in solution will be a minute or two at pH 9 and several hours at pH 7, well about an hour or something at pH 7. So, if you do a pH 9 reaction it’s all over in an hour or something, if you do a pH 7 reaction you may want to leave it overnight, say at 4°. Now, if you want to control the stoichiometry, you don’t want too much label. If you overlabel a protein it may quench the solution or be non-fluorescent. With the Thiols, you usually do this by stoichiometry, so you add 1.5 moles per protein, or not too much anyway. With the Amine reactive probes, because they’re hydrolyzing, you’re often controlling the reaction not by stoichiometry, not by so much so many moles per mole of protein but rather by concentration. But in practice, whether it’s more stoichiometry limit, or more concentration limited, depends on the concentration of your protein and so you may have to pilot it. The good news about reacting tough proteins like antibodies is it turns out they’ll accommodate a fairly wide range of protein density so it’s actually fairly easy to get a good labeled antibody. A few other practical points. Your protein may have some essential cysteines or lysines. I have spent half my life working on the molecule tubulin and it turns out if you take tubulin dimer form in solution, it has highly reactive cysteines and lysines that are essential for its polymerization function and there are some tricks for protecting those. So, you can sometimes sterically blockade essential cysteines or lysines by having a high concentration of a physiological ligand that blocks access of the labeling reagent. So, for example in the case of tubulin, you have to label in the polymer. So, if the tubulins polymerize into a microtubule that actually protects these essential groups. And sometimes with an enzyme you can protect by having a high amount of substrate that protects the active site. I think my final practical point is separating free probes from protein. Traditionally you do this by molecular size by dialysis, which is kind of slow, or by gel permeation chromatography. With these modern dies with the sulfonate groups that works quite well. But if you have a more sticky die it often will stick to the protein and be hard to separate. And it’s useful for check how much is left by chromatography on silica gel. Increasingly, in my lab we like to use an affinity based method labeling either in solution, or first attaching the protein and labeling it on a column. So for example, with antibody labeling we’re often attaching the protein, to a protein, an antibody to a protein A column, labeling it on column, flushing away the excess and then eluting the antibody. That seems to work really well and give us very little free die.

, , , , , , ,

Post navigation

2 thoughts on “Microscopy: Labeling Proteins with Fluorescent Probes (Timothy Mitchison)

Leave a Reply

Your email address will not be published. Required fields are marked *