co-crystallization

Hi ,

I am co-crystallizing a protein with compound and would like to know how much of compound to add to protein solution to start with. I know that the protein binds compound in a 1 to 1 ratio but also noticed that the compound precipitates out of solution when DMSO is diluted off. Where should I start of? A 1 protein :2 compound ratio or more? And what is the best method to determine if the binding is homogeneous (that all protein has got a compound in it)?

Any suggestions would help. Thanks

TY

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From: Shilong Fan


for me, I prefer to sock these compounds into your crystal. it will much more easy than co-crystallizaiton. But each protein should be different.

Normally when I start co-crystallization with small compound, I will set up the complex with 1:1.2 molar ratio as first trial to see what should happen.

As DMESO, you can't get rid of them. But you can find  as much lower concentraiton as you can.

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From: Annie Hassell

Hi YT—

We normally prepare our ligand stocks in DMSO and add this to the protein in 3-fold  molar excess.  The majority of our ligands are quite insoluble and precipitate when the DMSO concentration decreases upon addition to the protein……. so I am not surprised that you are seeing this.  If your compound does not bind your protein tightly, you might consider using a 5-fold molar excess of ligand. 

Some proteins crash out if the protein concentration in high when you add the ligand.  For those situations, we complex the ligand with dilute protein (1-2 mg/ml), and then concentrate this for crystallization trials.  I have had proteins where we had to complex the dilute protein with ligand, and then let it sit overnight at 4C before we concentrated the protein.  We normally incubate the protein+ligand at 4C for 1-3 hours for binding before we set up the crystallization experiments. 

Another scenario might be addition of ligand to the protein followed by incubation at room temp for ~1hr.  Then centrifuge at 4C, keep protein at 4C and set up your trays.

Hope this helps!

annie

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From: Prince, D Bryan
Date: 25 August 2011 14:38


Dear TY,

Typically between 5-10x molar concentration over the protein is enough to ensure binding when the IC50 is uM to low mM. For tighter binding compounds (nM to low uM), 2-5x is sufficient. Whatever you do, when the precipitate occurs DO NOT REMOVE it. I learned to my chagrin that you change all the dynamics of the drop when you do. I ended up with empty crystals until I left the precipitate in place. Think of it this way—

Free protein + compound ↔ protein:compound complex + precipitate (mix of protein + compound)

If you change the equilibrium by removing the precipitate, you remove the "pressure" on the P:C complex, and it will dissociate to P + C. The precipitate acts as a reserve of protein and compound, thus favoring (or stabilizing) the P:C complex. I set drops up as a slurry frequently, and if I get crystals, they always have the compound bound. Pay attention to the drops if you are screening, because it will be important to note what makes the precipitated solution better (clear drops=solubilizing) or worse (aggregated drops=decreased solubility of your complex). You can also try suspending your compound in LMW PEG's (200-400 FW) instead of DMSO. Either way, try using DMSO (~20%) or LMW PEG (~30%) depending on your crystallization conditions as a cryoprotectant. Any crystals that grow have some small amount of those agents in them already, so they should be more tolerant of them in higher concentrations.

Best of luck!

Bryan

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From: Roger Rowlett

Successful complexation depends on the concentration of protein, ligand, and the Kd of the protein-ligand complex. For Kd>>[protein], you will probably require [ligand] > 10 x Kd. As Kd approaches [protein], slightly superstoichiometric quantities will be sufficient for full occupancy. For Kd < [protein], stoichiometric quantities of ligand will suffice. Basically you need a [ligand] that puts near saturation on the binding isotherm.

Cheers.

--

---

Roger S. Rowlett
Gordon & Dorothy Kline Professor
Department of Chemistry
Colgate University
13 Oak Drive
Hamilton, NY 13346

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From: <Herman.Schreuder

If you do the calculations, you will find that you need a FREE ligand concentration of >10 * Kd to get >90% occupancy of the binding site.

If you have e.g. a ligand with a Kd of 100 nM, you would need a free ligand concentration of 1 µM. However, a solution of 10 mg/ml of a protein of 30 kDa, has a protein concentration of 333 µM, so in theory you should have a total ligand concentration (free + bound) of 334 µM. In pratice, some of the ligand may have been degraded during (prolonged) storaged, or the compound may not be as pure as the chemist would have wished it to be, so it is wise to use a safety margin of at least two. We normally use 1-2 mM compound to be on the safe side. Having too much ligand usually does not hurt, except that you use more compound, but with too little ligand you end up with an empty binding site and you will have to repeat the experiment with more ligand.

Best,

Herman