Protein Crystallization Hits

Some of you may know that Emerald BioStructures, as part of the Seattle Structural Genomics Center for Infectious Disease (SSGCID) has contributed to submitting more than 444 protein structures to the PDB in the past 4 years. That's quite an achievement and my congratulations go out to the project teams that are behind these structures, most of them determined via X-ray crystallography. Some of this output, including methods used to achieve this level of productivity, are described in the  September 2011 issue of Acta Cryst F.

One of the protein production methods that has been key for several of my own 2011 protein crystallization projects: lysis scouting with the Protein Maker instrument (described in this open access article "The Protein Maker: an automated system for high-throughput parallel purification". 

Smith, E., Begley, D., Anderson, V., Raymond, A., Haffner, T., Robinson, J., Edwards, T., Duncan, N., Gerdts, C., Mixon, M., Nollert, P., Staker, B., & Stewart, L. (2011). The Protein Maker: an automated system for high-throughput parallel purification Acta Crystallographica Section F Structural Biology and Crystallization Communications, 67 (9), 1015-1021 DOI: 10.1107/S1744309111028776

What is lysis scouting?

Stated simply, lysis scouting combines the testing of a set of cell-lysis buffer conditions with IMAC (ion metal affinity chromatography) . This is done to increase the yield of proteins that appear partially soluble or insoluble under standard lysis buffer conditions.  This procedure results in a clear path forward for scaled-up production of purified protein samples for protein crystallization trials.

How is lysis scouting done?

A single batch of protein expressing E.coli cells is split into 12 pools and lysed by sonication in 12 different buffer conditions. The paper shows as an example P450 51 A1 (CYP51A1) with a 6xHis-Smt tag. This is the outline of the lysis scouting protocol:

1. Prepare 12 aliquots, each corresponding to 3 g of wet cell paste
2. Resuspend in 30 mL lysis buffer (one out of an array of 12) - see table below.

Cell lysis buffers for testing lysis conditions of recombinantly expressed fungal cytochrome P450

3. Sonicate to lyse and spin to remove cell debris

4. Clarify lysates and load on 12 x 1 mL Ni-affinity matrix column

5. Wash, elute and analyze fractions

SDS-PAGE showing that buffers 1C and 1D extract much more of the target protein CYP51A1 (red boxes). L(load), W(wash) and E(elution) fractions are shown next to MW standards.

While well expressed, CYP51A seemed insoluble using standard cell-lysis methods. The lysis-scouting procedure yielded a buffer system with a detergent (CHAPS or octyl glucoside)  in the presence of high salt concentrations (500 mM NaCl).

The utility of the Protein Maker instrument in this process is the short time it takes to run a lysis scouting experiment. Total run time is approximately 1.5 hours (excluding sample analysis). I.e. many proteins can be tested for optimal lysis conditions in a single day - and since the instrument carries out the experiment for you and in parallel, there is plenty of time to strategize the next steps of mg-scale production of the protein sample for crystallization. 

There are many protein structures that we have produced in 2011 that would not exist without Protein Maker supported lysis optimization.

A true work horse.

Peter

Imagine this: you seek out a particular target protein and succeed with expression, purification and crystallization. Only when you go about building the model you realize that it's not the target protein you intended to devode your time on, but an entirely different beast. Yes, such things happen, see for example here:

Kefala G, Ahn C, Krupa M, Esquivies L, Maslennikov I, Kwiatkowski W, Choe S.
Structures of the OmpF porin crystallized in the presence of foscholine-12.
Protein Sci. 2010

The initial goal of the project was to go after KdpD, a K+ sensor kinase, and they end up with crystal structures of OmpF, a porin. These are membrane proteins, mind you. And if I had not spent my PostDoc in Jurg Rosenbusch's lab at the Biozentrum in Basel, Switzerland, it would be difficult for me to appreciate this feat. Granted, there's a lot of OmpF in E.coli membranes. But isolating the OmpF impurity with a new detergent and determining two structures from two new crystal forms is not as embarrassing as one might think.

The ticket here was foscholine-12, a rarely used detergent, that apparently is very useful for OmpF solubilization.

Foscholine-12. A new detergent for OmpF solubilization and crystallization  

I've heard of crystallization accidents before, but have never seen a such a nice turnaround of the project.

Well done!
Peter

In an ideal world, this would be a fairly simple way to grow protein crystals for X-ray structure determination purposes:

Start out with ca. 400 ul of filtered target protein solution, in 20 mM Hepes buffer neutral pH and maybe 50 mM NaCl , freshly purified to ca. 95% purity and concentrated to 10 mg/ml.

1. Snap freeze four x 25 ul aliquots of protein samples in thin-walled PCR tubes and store at -80C. These sample will be used in a week for protein crystallization optimization.

2. Document the quality of the protein sample via SDS-PAGE and determine its concentration (ca. 5 ul). Measuring UV absorption (OD280) and calculating the protein concentration is good enough.  You'll need ca. 5 ul for such an OD measurement (depending on cuvette path length and dilution). Check this online tool to get an estimate for the extinction coefficient based on sequence.

3. Now let's get out the multichannel pipettor and set up all of the remaining 200 ul of sample solution as 1ul + 1 ul sitting drop vapor diffusion crystallization experiments. A great first pass would look like this:
Plate 1: Wizard 1 & 2 (96 drops)
Plate 2: Wizard 3 & 4 (96 drops)
By the way, Emerald offers these reagents as the Wizard Suite. This is a 192 point, non-overlapping, sparse matrix that has a proven track record to yield crystals in first-pass protein crystallization exploration trials.

And while we're at it: go with the Compact Jr. plates. These plates can be sealed with clear tape and drops form nicely on the hyrdophobic polypropylene surfaces.

4. Store the crystallization trial at room temperature, minimizing temperature fluctuations and vibration. Observe right away, after 1 day, 3 days and 1 week.

5. The initially clear drops with now contain precipitate, some clear drops and a few with clustered microcrystals or needles in them. If the corresponding well-solution does not contain similar crystals, chances are that you've grown protein crystals! Since these crystals are likely to be too small to diffract them using your home-source X-ray generator and detector system, you'll need to optimize crystallization conditions and grow larger crystals.

6. There are many different ways to optimize crystallization conditons, and depending on prior knowledge you may want to carry out seeding experiments, include additives or change the treatment of the protein sample (i.e. filtration). Here's a simple optimization schema: create a grid screen around all conditions that gave you crystals. This rational crystallization optimization schema works great since it separates the effect of pH, salt, precipitant and protein concentration). You should use the online ScreenBuilder design tool to create an optimization screen. Your fellow researchers at Emerald BioSystems  are happy to prepare and send such a customized optimization screen to you.

7. Set up 96 - follow-up optimization 1 ul + 1 ul crystallization experiments with the saved protein material that you thaw in your fingers. If everything goes according to plan, crystals of different sizes will grow.

8. Harvest a crystal, cryoprotect, diffract and determine its structure ;)

Enjoy,

Peter

What's the minimal purity level required to set up a protein crystallization trial? Simple question, no clear answer, really. Most crystallizers I asked are happy to work with protein samples that are more than 90-95% pure. With purity judged by the intensity of the Coomassie stained target band in an SDS-Polyacrylamide Gel (PAGE). Sometimes it's difficult to get to such purity levels though, and the question becomes: at what purity level should you refuse to subject protein samples to crystallization trials?
This is a question that I am very interested in, particularly for membrane proteins. Membrane proteins are difficult to purify and they tend to loose activity during the purification process (this is of course due to the detergent micelles being a poor substitute for biological membranes). So we addressed this question together with Phil Laible's group at Argonne National Lab and while we were at it, we also compared two fundamentally different protein crystallization regimes: crystallization in lipidic cubic phases (LCP) and the more regular crystallization as protein/detergent/lipid complexes. LCP won, hands down.
To our surprise, in LCP decent crystals of photosynthetic reaction center (this was the only protein we tried, I know just one datapoint) grew at up to 50% contamination levels. Check out the paper for details:

C. A. Kors, E. Wallace, D. R. Davies, L. Li, P. D. Laible and P. Nollert
Effects of impurities on membrane-protein crystallization in different systems
Acta Cryst. (2009). D65, 1062-1073

We attribute this feature to the lipid matrix. It forms a 3D network of diffusion pathways, providing diffusion channels and barriers for different molecular species.

What does it mean for the required sample purity standard?
1. Don't give up when all you've got are 'dirty' protein samples. Set them up but do check though the identity of the target crystal, you may end up with a surprise: crystals of contaminating proteins rather than your target.
2. Of course I'd like to see a similar study carried out on soluble proteins. The meshwork of the LCP should help as well (need any help setting up LCP-based crystallization trials? - send an email, I'll fill you in with details).

All the best,
Peter