Protein Crystallization Hits

Most of the detergents that are in use today were available 5 or 10 years ago. Maybe the purity has increased over time, but the list of detergents extracted from successfully crystallized membrane proteins really hasn't seen any substantial additions in the past few years. In the previous blog "Applying the 80/20 rule to membrane protein crystallization pre-screening" I showed a list of detergents and referenced statistics that show that with just 8 detergents (dodecyl maltoside, decyl maltoside, nonyl glucoside, octyl glucoside, LDAO, C12E8 C12E9 and undecylmaltoside) about 80% of the membrane protein crystallization 'landscape' is captured.  If you're not working on beta barrel proteins you may as well remove the polyoxyethylene ethers and do with six detergents only, three of them being maltosides.
Only rarely new detergents are conceived, synthesized and tested with membrane proteins. Pil Seok Chae, Sam Gellman and others have done just that, and more: they've come up with a great new detergent class. This is their recent paper on their new series of detergents, called MNGs (maltose-neopentyl glycol amphiphiles):

Chae, P., Rasmussen, S., Rana, R., Gotfryd, K., Chandra, R., Goren, M., Kruse, A., Nurva, S., Loland, C., Pierre, Y., Drew, D., Popot, J., Picot, D., Fox, B., Guan, L., Gether, U., Byrne, B., Kobilka, B., & Gellman, S. (2010). Maltose–neopentyl glycol (MNG) amphiphiles for solubilization, stabilization and crystallization of membrane proteins Nature Methods, 7 (12), 1003-1008 DOI: 10.1038/nmeth.1526

The MNG molecule looks like this:

Figure: Welcome to the detergent family, MNGs:  maltose-neopentyl glycol amphiphiles. Reserving a spot in the front row on the detergent shelf in the freezer may be a good idea.

The molecule architecture, with the neopentyl in it center, was deliberately designed to provide a more rigid environment for membrane protein crystallization. Looking at the list of preferred detergents, the choice of maltosides (sic!) as a head group seems to be a non-brainer  (this is in retrospect, mind you), but the concept of a quarternary carbon is unique. The successful applications of this new detergent series range from stabilization (activity of beta 2 adrenergic receptor after detergent exchange from DDM), thermostability (of melibiose permease),  solubilization (leucine transporter, light harvesting complex and photosynthetic reaction center) and crystallization (cytochrome b6f & b2AR). While an improvement of crystal quality for cytochrome b6f was not seen, the authors mention that MNG-3 aided in improving crystals of agonist bound beta 2 adrenergic receptor. 

Overall the behavior of the MNGs seem to be on par with DDM (dodecyl maltoside), the currently highest ranked detergent in terms of use for successful crystal growth leading to high-resolution X-ray structures.

I raise may glass to Pil Seok and Sam: well done!
Peter

Protein Crystallizers know this simple rule: for crystallization to occur protein samples should be pure, homogenous and the protein be in a stable and non-aggregated state. These requirements can often be met by applying standard purification and concentration procedures to soluble protein targets. This is one of the reasons for the productivity of high-throughput structure genomics-type efforts. The buffer type, pH and salt are typically standard systems - such as 100 mM NaCl, 50 mM Tris-HCl pH7.5 - that rarely requires optimization for a particular protein target.

These requirements are also valid for membrane proteins, but achieving them often requires a lot more effort. A typical parameter to evaluate for membrane proteins is the detergent type and its concentration. Such a crystallization pre-screen can be done sometimes even before starting with the purification and, the procedure applied is called 'detergent exchange'. It turns out that there's a handful of methods available to carry out such detergent exchanges, for instance, using size exclusion chromatography, simple dilution or ultracentrifugation. The logic goes like this: once the membrane protein aggregates in the newly tested detergent solution, it can be detected as an extra peak, running with the void volume in the sizing chromatogram, as an increase in turbidity, or as a decrease in protein concentration in the supernatant of an ultracentrifuge tube.

Anybody who has concentrated a protein sample using a simple MWCO filter realizes, that filtration provides a good means to separate aggregated from non-aggregated protein. Indeed, GE Healthcare's multi trap purification filter kit utilizes  this feature for purification scouting of His-tagged proteins. Essentially, the protein is bound to a resin and eluted by filtration through a MWCO filter. The conditions that yields most protein in the filtrate wins. Michael Wiener's membrane protein group has given this concept an interesting twist. They describe in this paper:

an adaptation of the filtration concept to detergent screening and have devised a differential filtration assay that allows to identify those detergents that render a particular membrane protein target 'well behaved' and hence, more crystallizable. How does this work?

Instead of using the individual filtration devices, two 96-well filter plates of different MWCO (the use of two different MWCO sizes allows to distinguish between stability and size) are used sequentially. 94 different detergents can be used in one go. Some people may consider testing with 94 different detergents overkill, but I'm with Michael in his 'knock em dead' approach. However, using fewer detergents and spin-filters may prove more practicable in many labs. In fact, this useful review on membrane protein crystallization parameters:

Newstead, S., Ferrandon, S., & Iwata, S. (2008). Rationalizing α-helical membrane protein crystallization Protein Science, 17 (3), 466-472 DOI: 10.1110/ps.073263108

can be used to argue that by just testing 8 instead of 94 different detergents one can cover more than 80% of the 'detergent landscape'. The table below shows the ranking of detergents that have provided most membrane protein crystal structures. Note that followers of the Pareto Principle (80% of the bang for 20% of the buck) will do with a set of only 8 detergetns or filter tests.

Table: Which detergents to use for membrane proteins? Shown are detergent types and occurrences in successful membrane crystallization screens. Data from supplemental info to Newstead et al., 2008 paper, kindly provided by Simon Newstead. Note 1: Am I seeing this right that HEGA-10 is not included in the set of 94 used by Wiener?; Note 2: adhering to this table will increase investigator bias.

When I saw Michael presenting his differential filtratio assay at the NIH Roadmap Meeting in March 2009 I was particularly excited by these features:

(i) the low sample amounts required (only 10 ug protein / well to create sufficient signal in the dot-blot / western assay) and,

(ii) the type of data that can be pulled out of this assay.

It has taken me a while to appreciate the utility of the 'size stability quad plot'. The gist is that the membrane proteins can be grouped into cohorts that are formed by the quandrants as defined by stability and size. This sorting of detergents helps to predict those detergent species that should work best for crystallization.

And that's exactly the type of result you're expecting from a crystallization pre-screen.

All the best,

Peter

This is my list of indispensable online tools that I would encourage every protein crystallizer to use during different phases of the protein crystallization endeavour: 

While there's some overlap with Sean's list over at P212121.com, he has a lot of additional tools listed that relate to up- and downstream crystallization processes. Sean's blog is definitely worth a visit!

All the best,

Peter

Handling and storage of protein crystallization trials is usually carried out in a manner that minimizes exposure to vibration. I have set up crystallization trials and put them aside to a 'safe place' with the intention to let the crystallization commence without me interrupting the crystallization process. While I'm not sure if it was the lack of shaking or the temperature increase during observation caused by the microscope light, I did get larger and better diffracting crystals in at least one case. On the other hand, I've seen crystallization cabinets with hundreds of trays that got a whack every time somebody did not catch the door into the crystallization room. And this was a very productive protein structure lab.

Let's use this vibration effect and take it to the extreme: how about adding a bit more stress, such as ultrasonication? 3 seconds  of of bath-ultrasonication every 3 minutes throughout the crystallization process?

That's exactly what Crespo et al  have done and describe in their new paper:

Crespo, R., Martins, P., Gales, L., Rocha, F., & Damas, A. (2010). Potential use of ultrasound to promote protein crystallization Journal of Applied Crystallography, 43 (6) DOI: 10.1107/S0021889810040951

This is the contraption they used to compare with ultrasonication vs. without. 

Figure: Simple setup to compare the effect of micro batch-type protein crystallization with ultrasonication and without ultrasonication.

The setup is rather well controlled and was used to map the supersaturation curves for Lysozyme (I know…) in the presence and absence of the ultrasonication stress regime. The finding is that ultrasonication promotes nucleation. In doing so, it appears that crystals gown with this method are of higher quality. While the statistics on crystal quality (n=13) are not that convincing and the effect is small (0.1A average improvement), I'd do it if I'm desperate. For instance in cases where protein crystallization optimization by

• reagent screening, including additives,
• drop mixing regime
• temperature
• different formats and plates
• in-situ proteolysis

have not yielded any improvements in the protein crystal diffraction quality.

Cheers,

Peter

For an academic researcher paper writing is often the culmination of hundreds, if not many thousands of hours of work. The data is collected, a story is shaping up - how do you go about transmitting your newly found wisdom to your peers?

I have witnessed firsthand several different paper writing styles and practices and am the first to admit that there's no single best way to this. There's some good advice out there, though. For example, Bill Wells suggests in his aptly titled paper "Me Write Pretty One Day: How to Write a Good Scientific Paper"  'a few small steps" to make "...scientific writing clear, straightforward, and digestible". As I'm starting to write a paper myself I find it useful to look at this paper to remind myself of how to write clearly, to define what's my point and then adhere to the 'show, don't tell' rule. Wells guides his readers through 'The Anatomy of a Paper' and gives a lot of good advise on how to tell our story.

As a reader of a protein crystallization paper I expect to see the following:

1. The amino acid sequence of the protein that's expressed.

2. Description of a typical expression and purification experiment. What's the yield & purity? Are there functional assays that provide checkpoints for anybody who wishes to repeat your work?

3. A detailed description of what happened between eluting the protein from the last column and before setting it up for crystallization: temperature(s), storage conditions and time, dialysis (time, volume, MWCO used, volume ratio), concentrating device, centrifugation, filtration, addition of ligands.

4. The crystallization experiment/regime: composition of precipitant solution buffers, crystallization type, volumes, trays, dispensing methodology (stirred or mixed?), screen used, time it took for crystals to show up. Any 'out of the norm' observations, such as 'crystals formed at the air liquid interface'

5. Crystallization result: how many crystals, crystal habit, size, crystal quality (X-ray diffraction limit and any 'Table 1' associated data that resulted from a complete data set). Describe reproducibility (only 1 drop out of 10 produced crystals?, only 1 out of 10 crystals diffracted?) and any insight gained while handling the crystals (did the crystals bend when looping out, do they tend to break? Do they 'melt' when the temperature is changed?).

6. X-ray diffraction experiment: treatment of crystal (cryo used, orientation crystal was mounted) before and during exposure to X-rays. Diffraction equipment used, exposure time, sweep angle.

7. Two images of the crystals: close-up to see the crystal habit and an overview of the entire crystallization experiment. An X-ray image showing diffraction spots (crystal oriented along a special axis would rock).

There's really no good reason to hold back with any such details in a protein crystallization paper (I won't hold it against you when I review your paper). After all, one of the reasons you're writing the crystallization paper is that you aim to provide instructions to a crystallizer who seeks to enable repetition of your work in the future. This is a neat way you can 'pay back' to all scientists that provided you with insightful tips in their papers and that have helped you succeed with your own protein crystallization research project. Pass on the baton.

Cheers, Peter