Protein Crystallization Hits

Membrane Protein Crystallization with Additives in LCP

The question I get asked most often when the topic comes to crystallization of membrane proteins in lipidic cubic phases (LCP) is: "what screens can I use?" (for the non-specialists: LCP stands for Lipidic Cubic Phases; these are materials that have been used successfully to serve as host matrices for the crystallization of membrane proteins). The answer is: pretty much everything. After all, the precipitants that have been used successfully are not that different from other protein crystallizations (stop reading here if you're not interested in the details).

The question regarding precipitants and additives goes way back when the LCP crystallization methodology was still in its infancy: I'm talking last century here. Due to their utility in the crystallization of hepta-helical membrane proteins, specifically GPCRs, many crystallizers are interested in using additives for LCP-mediated membrane protein crystallization. Turns out that there's a substantial body of published literature that you can take as a guide through this 'lipid swamp'. To make things simple, let's divide up these additives in lipids, detergents, small molecule amphiphiles and the more traditional crystallization reagents, such as organic solvents, polymers and salt:

Lipidic cubic phases can be doped with a variety of amphiphiles, ranging from LIPIDS such as Cholesterol, DOPE, DOPC, DMPC and PLPC to DETERGENTS (dodecyl maltoside, CTAB, beta octyl glucoside, MEGA9, DTACI and Zwittergent) (see fig 1 below).

Figure 1: Lipid and detergent additives that have been added to LCP to modulate the outcome of membrane protein crystallization experiments. These tables and the one below were taken from a chapter in this book: "14. X-Ray Crystallography of Membrane Proteins: Concepts and Applications of Lipidic Mesophases to Three-Dimensional Membrane Protein Crystallization" G protein-coupled receptors, T.Haga and G.Berstein, editors, 2000, CRC press. P. p 365-388, Loewen, M., Chiu, M., Widmer, C., Landau, E.M., Rosenbusch, J.P., Nollert.

The HOST LIPID that forms a cubic phase is likewise a factor that can be varied:

Figure 2: LCP forming lipids: monopalmitolein, monoolein, monovaccenin, monoeicosenoin and a special cyclopropyl derivative of monoolein.

Indeed, the test protein bacteriorhodopsin crystallized well in a variety of lipids and lipid mixtures:

Figure 3. Crystallization in LCP consisting of different lipids and lipid mixtures, including cholesterol.

That's a lot of compounds that can be screened: LCP (consisting of a range of lipids) with and without additives, at a variety of concentrations. At the time this wealth of possibilities felt like opening "Pandora's Box": while we realized that the LCP method is useful for many membrane protein crystallization trials, the number of possible combination of crystallization components seemed daunting. Regardless, there was a lot of excitement to apply the LCP methods to other membrane proteins, such as bovine rhodopsin or the beta-2-adrenergic receptor.

There's nothing fundamental really, that has changed since these early days. The main progress we've seen in the last 10 years was miniaturization, possibly automation, and setup formats that give better visibility to the crystals.
Foremost, however are more facile protein engineering capabilities and the fact that monoolein has worked so well as the basis host lipid that have made it so simple to apply the LCP method to any membrane protein crystallization trial.

When it comes to the more conventional crystallization reagents, all standard crystallization reagents that are used for soluble proteins can be applied. Some may not be compatible with lipidic cubic phases at all, but this property may actually cause the embedded membrane protein to crystallize (think sponge phases).

 

Figure 4. Effect of water soluble compounds on cubic phase stability. Polyethylene (PEG) and modified polyethylenes have been used to grow crystals of a variety of membrane proteins. Here is an early paper documenting this:

M. L. Chiu, P. Nollert, M. C. Loewen, H. Belrhali, E. Pebay-Peyroula, J. P. Rosenbusch and E. M. Landau
Crystallization in cubo: general applicability to membrane proteins
Acta Cryst. (2000). D56, 781-784 

Again, all conventional protein crystallization screens can be (and have been) applied to LCP-based membrane protein crystallization. Alternatively, a subset of those conditions that are compatible with the existence of LCP or with protein mobility can be screened using speciality screens, such as Emerald BioSystems' Cubic Screen.

The composition of the crystallization cocktail with all the salts, buffers and additives is described in this Cubic Screen tech-sheet.
There you'll see that alcohols such as ethanol, 2-propanol, 1,4-butanediol, PEGS (400, 1000, 3000, 8000, derivatized PEG 2000 MME), salts (sodium chloride, lithium sulfate, magnesium chloride, zinc acetate, Ammonium sulfate, sodium citrate, Sodium Malonate ) and buffers (Mes, acetate, citrate, hepes, Na/K phosphate, cacodylate, citrate, imidazole) were selected from then exisiting crystallization formulations. To get a variety of concentrations of these precipitation reagents, simple dilution of the entire screen is common practice.

Customers of the Cubic LCP kit have used these components for more than 8 years to screen crystallization conditions and grow membrane protein crystals.

All the best,
Peter

Protein Crystallization phase diagrams

Most of us protein crystallizers are not physical chemists and are sometimes bewildered when looking at a protein crystallization phase diagram. The thing is, that phase diagrams serve as a useful conceptual framework and they can be used to guide the trouble-shooting process for protein crystallization trials.
So, here's it: Protein crystallization can be seen as three distinct events: nucleation, growth and stop of growth. Disregard the kinetics of these processes for simplicity sake. Crystallization  occurs only from supersaturated solutions where the protein concentration exceeds its solubility in a given solution. The state of this supersaturation depends, of course, on many factors - tweaking these factors is the game we play in protein crystallization (we win when crystal nuclei form and crystals to grow).

If you haven't done so already, please take some quality time to take in this schematic protein crystallization phase diagram:

Figure: Protein crystallization phase diagram schema. Crystal nucleation is a critical phenomenon that occurs only in an area of the supersaturation zone. Crystals grow under conditions of supersaturation only once nuclei have formed. Yellow: no crystal growth; red: no crystal growth either. Dark-blue: this is the target area to land in for crystallization screening. Light-blue: this is where you want to be for crystal growth.

Once you're familiar with the lines separating the differenct zones, consider these three pathways:

Pathway A: batch-type crystallization. By mixing protein with the precipitant solution the protein becomes supersaturated. Crystal nuclei form and crystal grow until the protein concentration in solution is saturated.
Pathway B: vapor diffusion-type crystallization. The concentration occurs during vapor diffusion, following mixing of the protein with the precipitant solution causing the protein to become supersaturated. During vapor diffusion the precipitant concentration increases and extends the crystal growth process.
Pathway C: dialysis-type crystallization. As the precipitant diffuses into the protein-holding chamber the protein supersaturates. Once nuclei have formed, protein crystals grow as long as the protein concentration remains supersaturated.

Keep in mind, that this protein crystallization phase diagram is (just) a general framework. There are not that many phase diagrams of actual proteins available, actually. A nice reality check and a thorough introduction into the topic of protein crystallization phase diagrams can be found here:

Asherie, N. (2004)
Protein Crystallization and Phase Diagrams
Methods 34, 266-272

Regards,

Peter

Protein crystal optimization with detergents

Detergents are an interesting class of molecules, mainly because of their amphipathic, or amphiphilic nature. They like both water & oil and are hence used extensively to keep hydrophobic molecular species dissolved in an aqueous environment. As a result they're used on an industrial scale to clean clothes, but they're also used in protein crystallization. Of course everybody thinks membrane proteins now, but that's not where this blog post is going.  Detergents can be used to optimize the crystallization of soluble proteins.

For example, in their 2001 paper Guan et al. describe the optimization of four soluble protein crystallizations with detergents, all resulting in improved crystal quality and crystal growth reproducibility.

Guan, R.-J., Wang, M., Liu, X.-Q, Wang, D.-C.
Optimization of soluble protein crystallization with detergents
Journal of Crystal Growth 231 (2001) 273-279

The detergents they use are rather diverse: they use Zwittergent, Mega-8, n-Octanoylsucrose, C12E8 and C-HEGA-10. The surprise here is that none of these detergents - with the exception of C12E8 - are routinely used in membrane protein crystallization trials.

Check out their nice images of protein crystals grown with and without detergent additives.

 

These dirty crystals could use some detergent scrubbing…

 

Their simple detergent additive protein crystallization trials have provided the following outcomes: 

• polycrystalline -> single crystals
• crystal clusters -> single crystals
• poor X-ray diffraction-> 2.5A diffraction
  
• non-reproducible -> reproducible

Worth a try, I'd say.

Peter

Crystallization Game Changer: Try a Different Plate

There are so many parameters to change when optimizing crystallizations and they all need to be tried out to identify the critical parameter that improves X-ray diffraction. In this series of blog posts I've mostly discussed modulating the crystallization reaction, by adding additives, changing precipitant or protein concentration etc. However, there seems to be a simple way to optimize, keeping all other parameters constant: drop volume, temperature, all concentrations. What could it be?

Try switching the crystallization plate!

Jenny Martin describes in her recent PLOS paper with statistical rigour that different 96-well crystallization plates - formats, materials - do effect the outcome of protein crystallization experiments:

King, Gordon J., Kai-En Chen, Gautier Robin, Jade K. Forwood, Begoña Heras, Anil S. Thakur, Bostjan Kobe, Simon P. Blomberg, and Jennifer L. Martin.
Interaction between Plate Make and Protein in Protein Crystallisation Screening.
PLoS ONE 4, no. 11 (November 16, 2009): e7851

In fact, she suggests to optimize protein crystal growth by matching the protein with its optimal plate make.

Figure: Compare different protein crystallization plates to grow optimized protein crystals.

So, here's a selection of 6 different Emerald BioSystems protein crystallization plates with different formats, well arrangements, volumes and materials that you may want to try to optimize protein crystal growth:

Clover 384 plate (COC material, 4 micro crystallization wells / reservoir)
Clover 384 plate (Polystyrene, 4 micro crystallization wells / reservoir)
Compact Clover plate (Polypropylene, 4 crystallization wells / reservoir)
Compact, Jr. plate (Polypropylene, 1 crystallization well / reservoir)
Combi Clover plate (Polypropylene, large wells, 4 crystallizations well / reservoir)
Combi Clover Jr. plate (Polypropylene, large wells, 1 crystallization well / reservoir)

Sometimes it's time to change the game.

Peter

PDBsum rocks for crystallization protocol

This week I came across a message from PDBsum letting us know that certain figures and captions of a paper we had published in 2008 (Gerdts et al. (2008). Acta Crystallogr D Biol Crystallogr, 64, 1116-1122.) had been included into PDBsum. I had not visited the PDBsum site before and was at first intrigued and then positively surprised about the wealth of information that was presented on a protein structure (3cxk). This is a high-density, information rich way to get a quick impression on a protein structure and other accessory information.

Figure: Screenshot of the PDBsum entry for 3cxk. The crystallization experiment is nicely referenced in PDB-sum.

What I liked in particular about this presentation is that the crystallization experiment becomes part of the story. Our paper described an earlier, beta version of the MPCS - the plug-based nanovolume microcapillary protein crystallization system. Since the publication of the paper in 2008 the technology has matured substantially (check out the New Product Award 2010 that the PlugMaker has received last week).

Any context that goes beyond just reporting the precipitation reagent helps. Having such exquisite detail available when trying to reproduce protein crystallization experiments is often necessary to build on published research.

Way to go EMBL-EBI!
Peter

Our PlugMaker wins the New Product Award at LabAutomation!

This is exciting news: our PlugMaker protein crystallization instrument won the 2010 New Product Award at LabAutomation! Of course we've always known that this instrument is a winner: 800 crystallization setups with just 3-4 uL of proteins, with crystals ready to diffract and 'simple as an iPod'. Seeing now the independent stamp of the Association for Laboratory Automation on it is very gratifying to us.

 

Here's a big THANK YOU !!! to the many people and their brilliant minds behind this instrument: the team at BioStructures and BioSystems, especially Lance and Cory, Rustem's lab and our development partners. I'm looking forward to see many protein scientists produce lots of X-ray structures with protein crystals grown within CrystalCards using the PlugMaker.

Plugs are the new drops.
Peter

Seeding with protein 'crap': Microseed Matrix Screening

What's the new thing that people are tying out these days? Lots of new methodologies, ranging from low-volume plug-based crystallization (of course) to new crystallization screening matrices for membrane proteins. I've noticed that there's a 'new' seeding method around that has come up in several conversations I've had with protein crystallizers over the past 3 years or so. It's called Microseed Matrix Seeding. Judging from people who try and stick with it, it is my impression is that this seems to be working rather well.

What is Microseed Matrix Seeding in practical terms?

You start with a 'failed' primary protein crystallization tray and:

1. Harvest some or all the precipitated drops, pool them (yes!) and call this seed stock.
2. Spike each drop of a new, secondary crystallization trial with a portion of the seed stock.
3. Obtain crystals from a protein/formulation combination that is different from that you used to create the seed stock with.

The Matrix Microseeding method and its application to yeast cytosine deaminase was first described by Gregory Ireton and Barry Stoddard in:

G. Ireton & B. Stoddard

Microseed matrix screening to improve crystals of yeast cytosine deaminase

Acta Cryst D60 (2004), 601-605.

Then Alan D'Arcy picked up on this new method and initiated a robotic application for this new protein crystallization seeding method:

A. D'Arcy, F. Villarda, M.Marsh

An automated microseed matrix-screening method for protein crystallization

Acta Cryst D63 (2007), 550-554.

Seeding with 'crap'? (mind me - not my own words, but I've heard this very question more than once)

Maybe not. What you see as precipitated material may be not properly characterized crystalline material. For all I know, there could be sub-micrometer sized microcyrstalline protein material mixed in the precipitate. And there's just no way for you to see that. Alternatively, the precipitated protein material itself may form a heterogenous nuclation surface in similar ways that seaweed or horse hair can serve as nuclei for protein crystallization.

A case for Microseed Matrix Screening(?) 

If you you've got many drops with precipitation in it (B), and no crystallization leads whatsoever - why not try give it a try?

Thanks,

Peter

Best Protein Crystallization Paper 2009

Guess what - in my opinion there was not a single paper published in 2009, covering the topic of protein crystallization review-style,  that comes close to Naomi & Emmanuel's collected wisdom published in 2008:

As much as I prefer open-source publications, this is one of the best. Nature Methods, Vol. 5 No.2, February 2008. Naomi Chayen and Emmanuel Saridakis.

Why?
Because it covers all the basics: Setting up initial protein crystallization trials and classical screening methods, deals with the question how many trials should be set up, what ideal volumes are and what to do if protein crystallization trials fail. In addition, Naomi Chayen and Emmanuel Saridakis lucidly explain the choices of available crytallization formats and go into a fairly detailed description of practical ways to influence the crystallization process to grow protien crystals that are sufficient for X-ray diffraction . Protein crysallization optimization is explained in a hands-on fashion, extensively referencing the body of published literature. A joy to read.

I've saved this paper in my archive using the file name Naomis_bible_paper.pdf

Happy new 2010!

Peter

 

Beautiful protein crystal images

Acta Cryst F is the place to go to check out images of new protein crystals. I've just recently noticed that every single crystallization report is graced with a crystal. Way to go Acta Cryst F!

Cover illustration of the Jan 2010 issue of Acta Cryst F.

While browsing the new Jan 2010 issue I saw in their Notes for Authors 2010 That Acta Crystallographica Section F requires that Validation Reports need to be submitted for publication:

11.5. Validation reports
Authors of structural papers are required to provide a validation report on submission. Authors are encouraged to presubmit their data to the PDB and obtain a validation report for their structure.

Howard Einspahr and Manfred Weiss then go on to make a point in their editorial  that this requirement will be fully enforced in 2010. I suppose that this is a direct consequence of the Krishna Murthy case and the associated recent crystal structure retractions.

Looking at the pretty pictures of protein crystals I realize that the only thing that can be possibly wrong with crystal pictures is that there's a mix-up (intentionally or not). There may be less damage with such mix-ups though, since the value of a crystal images is limited compared to all the other data (or isn't it?).

One thing is clear: each image of a new protein crystal is a gem that's worth showing off.

Happy new 2010,
Peter

Facing Protein Crystallizers' Remorse

The end of 2009 is near and it's time to clean up to make room for new and exciting projects in 2010. I'm just returning from the lab with a stack of protein crystallization trays to toss. Some of the crystallization experiments were prepared more than a year ago! There were two categories of crystallization trays that I dealt with:

1. Trays set up with protein that have never yielded any crystals at all
Should I keep them and hope that via slow desiccation or proteolytic cleavage protein crystals will eventually form? Nah! Everything that's older than 6 months must go. Gone they are. - I'm having second thoughts though, now that I'm writing this. I could open the crystallization chambers for a while and let the drops dry out just a little and then close them again. After all, protein crystallization by dehydration does work sometimes. Or move the trials to a different temperature? Or add chemotrypsin to the drops and create target fragments that crystallize.
Crystallizers' remorse setting in big time....

2. Trays set up with protein that have yielded crystals and structures
I could throw them out altogether. Crystals diffracted, structure is done. But, why not keep a few trays with crystals? You never know when a project 'comes back' - with the need to co-crystallize together with a small molecule ligand or protein partner. Even if the crystals don't diffract they may be useful and serve as seeds.

Pretty (Ammonium sulfate) crystals. These are the easy protein crystallization setups to give up on and throw out.

Phew! - that was easy, actually.

2010 here I come!
Peter