International
Tables for
Crystallography
Volume C
Mathematical, physical and chemical tables
Edited by E. Prince

International Tables for Crystallography (2006). Vol. C, ch. 3.1, pp. 148-151

Section 3.1.1. Crystallization

P. F. Lindleya

aESRF, Avenue des Martyrs, BP 220, F-38043 Grenoble CEDEX, France

3.1.1. Crystallization

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3.1.1.1. Introduction

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The preparation of single crystals probably constitutes the most important step in a crystal structure analysis, since without high-quality diffraction data many analyses will prove problematical, if not completely intractable; time and effort invested in crystallization procedures are rarely wasted. There is a wealth of literature available on the subject of growing crystals and this includes the Journal of Crystal Growth (Amsterdam: Elsevier). This section does not intend to be a comprehensive review of the subject, but rather to provide some key lines of approach with appropriate references. The field of crystallizing biological macromolecules is itself a growth area and, in consequence, has been given a special emphasis.

Useful general references for growing crystals for structure analysis include Bunn (1961[link]), Stout & Jensen (1968[link]), Blundell & Johnson (1976[link]), McPherson (1976[link], 1982[link], 1990[link]), Ducruix & Giegé (1992[link]) and Helliwell (1992[link]). Volume D50 (Part 4) of Acta Crystallographica (1994) reports the Proceedings of the Fifth International Conference on Crystallization of Biological Macromolecules (San Diego, California, 1993) and is essential reading for crystallization experiments in this area. A biological macromolecular database for crystallization conditions has also been initiated (Gilliland, Tung, Blakeslee & Ladner, 1994[link]).

3.1.1.2. Crystal growth

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Crystallization has long been used as a method of purification by chemists and biochemists, although lack of purity can severely hamper the growth of single crystals, particularly if the impurities have some structural resemblance to the molecule being crystallized (Giegé, Theobald-Dietrich & Lorber, 1993[link]; Thatcher, 1993[link]). The process of crystallization involves the ordering of ions, atoms, and molecules in the gas, liquid, or solution phases to take up regular positions in the solid state. The initial stage is nucleation, followed by deposition on the crystallite faces. The latter can be considered as a dynamic equilibrium between the fluid and the crystal, with growth occurring when the forward rate predominates. Factors that affect the equilibrium include the chemical nature of the crystal surface, the concentration of the material being crystallized, and the nature of the medium in and around the crystal. Relatively little research has been done concerning the process of nucleation, but crystal formation appears to be conditional on the appearance of nuclei of a critical size. Too small aggregates will have either a positive or an unfavourable free energy of formation, so that there is a tendency to dissolution, whilst above the critical size the intermolecular interactions will, on average, lead to an overall negative free energy of formation. The rate of nucleation will increase considerably with the degree of supersaturation, and, in order to limit the number of nuclei (and therefore number of crystals growing), the degree of supersaturation must be as low as possible. Supersaturation must be approached slowly, and, when a low degree has been achieved, it must be carefully controlled. Many factors can influence crystallization, but a conceptually simple explanation of crystal growth has been described in detail by Tipson (1956[link]) and elaborated, for example, by Ries-Kautt & Ducruix (1992[link]). These latter authors provide a useful schematic description of the two-dimensional solubility diagram relating the concentration of the molecule being crystallized to the concentration of the crystallizing agent. The presence of foreign bodies, such as dust particles, makes the nucleation process thermodynamically more favourable, and these should be removed by centrifugation and/or filtration. The addition of seed crystals can often be used to control the nucleation process (Thaller, Eichelle, Weaver, Wilson, Karlsson & Jansonius, 1985[link]). In the case of the formation of crystals of macromolecules in solution, Ferré-D'Amaré & Burley (1994[link]) have described the use of dynamic light scattering to screen crystallization conditions for monodispersity. Empirical observations suggest that macromolecules that have the same size under normal solvent conditions tend to form crystals, whereas those systems that are polydisperse, or where random aggregation occurs, rarely give rise to ordered crystals.

3.1.1.3. Methods of growing crystals

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General strategies for crystallizing low-molecular-weight organic compounds have been reported by van der Sluis, Hezemans & Kroon (1989[link]) and are listed in Table 3.1.1.1[link]. Many of these strategies are also applicable to inorganic compounds. In the case of biological macromolecules, the main methods utilize one or more of the factors described in Subsection 3.1.1.5[link] and include batch crystallization, the hot-box technique, equilibrium dialysis, and vapour diffusion (see, for example, Blundell & Johnson, 1976[link]; Helliwell, 1992[link]). The growth of macromolecular crystals in silica hydrogels minimizes convection currents, turbidity, and any strain effects due to the presence of the crystallization vessel. Heterogeneous and secondary nucleation are also reduced (Robert, Provost & Lefaucheux, 1992[link]; Cudney, Patel & McPherson, 1994[link]; García-Ruiz & Moreno, 1994[link]; Thiessen, 1994[link]; Robert, Bernard & Lefaucheux, 1994[link]; Bernard, Degoy, Lefaucheux & Robert, 1994[link]; Sica et al., 1994[link]). Various apparata have been described for use with the vapour diffusion technique (see also Subsection 3.1.1.6[link]) and include a simple capillary vapour diffusion device for preliminary screening of crystallization conditions (Luft & Cody, 1989[link]), a double-cell device that decouples the crystal nucleation from the crystal growth, facilitating the control of nucleation and growth (Przybylska, 1989[link]), microbridges for use with sitting drops in the 35–45 µl range (Harlos, 1992[link]), and diffusion cells with varying depths, in order to control the time course of the equilibration between the macromolecule and the reservoir solution (Luft et al., 1994[link]).

Table 3.1.1.1| top | pdf |
Survey of crystallization techniques suitable for the crystallization of low-molecular-weight organic compounds for X-ray crystallography (adapted from van der Sluis, Hezemans & Kroon, 1989[link])

TechniqueAdvantage(s)Limitation(s)
Evaporation from a single solvent Simple
Inexpensive
Limitation to solvents with adequate vapour pressure
Crust formation on tube walls
Crystals that are dried are less suitable as seeds, may lose included solvent and become tightly adhered to the crystallization vessel
Difficult to reproduce
Limited number of solvents give concentration 5–200 mg ml−1 for a particular compound
Evaporation from a binary mixture of solvents (volatile solvent and non-volatile precipitant) No crust formation on the tube walls
Crystals are not dried
Stringent demands on solubility, miscibility and volatility of the two solvents
Difficult to reproduce
Batch crystallization No demands on the volatility of the solvent or precipitant
Repeated seeding by thermal treatment is easy
Metastable zone with regard to supersaturation must be large
High and almost uncontrollable crystallization rate
Solvents must be miscible
Liquid–liquid diffusion Favourable change in supersaturation at the interface during crystallization
Repeated seeding by thermal treatment is easy
Density differences required for the two liquids (less stringent if capillaries are used)
Viscosity of the liquids greater than water
Solvents must be miscible
High and almost uncontrollable crystallization rate
Sitting-drop vapour-phase diffusion Crystallization rate can easily be controlled by changing the diffusion path, solvent, precipitant, or pH
Repeated seeding easily implemented
Highest number of independent variables to obtain wide variety of conditions
Solvents must be miscible
Solvent preferably less volatile than precipitant
Hanging-drop vapour-phase diffusion Crystallization rate can easily be controlled by changing solvent, precipitant, or pH
Easy examination of crystallization outcome in array-like set-up
Only applicable in case of water-based solvents
Diffusion rate is fast and difficult to control
See previous method
Temperature change Easily controllable parameter
Repeated seeding extremely easily and accurately carried out
With Dewar flask inexpensive and simple
Limited to thermally stable compounds and (pseudo)polymorphs
Gel crystallization Suited for sparingly soluble or easily nucleating compounds Limited variety of solvents possible
Sampling of crystals difficult
Laborious
Sublimation No inclusion of solvent of crystallization Limited to small hydrophobic molecules
Laborious
Solidification For liquids and gases the only applicable method Limited to thermostable compounds
High change of amorphicity
Laborious

3.1.1.4. Factors affecting the solubility of biological macromolecules

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There are many factors that influence the crystallization of macromolecules (McPherson, 1985a[link]; Giegé & Ducruix, 1992[link]; Schick & Jurnak, 1994[link]; Tissen, Fraaije, Drenth & Berendsen, 1994[link]; Carter & Yin, 1994[link]; Spangfort, Surin, Dixon & Svensson, 1994[link]; Axelrod et al., 1994[link]; Konnert, D'Antonio & Ward, 1994[link]; Forsythe, Ewing & Pusey, 1994[link]; Diller, Shaw, Stura, Vacquier & Stout, 1994[link]; Hennig & Schlesier, 1994[link]), but the following are particularly important with respect to solubility (Blundell & Johnson, 1976[link]).

Ionic strength. The solubility of macromolecules in aqueous solution depends on the ionic strength, since the presence of ions modifies the interactions of the macromolecule with the solvent. At low ion concentrations, the solubility of the macromolecule is increased, a phenomenon termed `salting-in'. As the ionic strength is increased, the ions added compete with one another and the macromolecules for the surrounding water. The resulting removal of water molecules from the solute leads to a decrease in the solubility, a phenomenon termed `salting-out'. Different ions will affect the solubility of the protein in different ways. Small highly charged ions will be more effective in the salting-out process than large low-charged ions. Commonly used ionic precipitants are listed in Table 3.1.1.2[link], column (a) (McPherson, 1985a[link]).

Table 3.1.1.2| top | pdf |
Commonly used ionic and organic precipitants, adapted from McPherson (1985a[link])

(a) Ionic compounds(b) Organic solvents
Ammonium or sodium sulfate Ethanol
Sodium or ammonium citrate Isopropanol
Sodium, potassium or ammonium chloride 2-Methyl-2,4-pentanediol (MPD)
Sodium or ammonium acetate Dioxane
Magnesium sulfate Acetone
Cetyltrimethylammonium salts Butanol
Calcium chloride Dimethyl sulfoxide
Ammonium nitrate 2,5-Hexanediol
Sodium formate Methanol
Lithium chloride 1,3-Propanediol
  1,3-Butyrolactone
  Poly(ethylene glycol) 600–20000 (PEG)
The volatility of solvents such as ethanol and acetone may cause handling problems.
Ammonium sulfate can cause problems when used as a precipitant, since pH changes occur owing to ammonium transfer following ammonium/ammonia equilibrium; this effect has been studied in detail by Mikol, Rodeau & Giegé (1989[link]). Monaco (1994[link]) has suggested that ammonium succinate is a useful substitute for ammonium sulfate.

pH and counterions. The net charge on a macromolecule in solution can be modified either by changing the pH (adding or removing protons) or by binding ions (counterions). In general terms, the protein solubility will increase with the overall net charge and will be least soluble when the net charge is zero (isoelectric point). In the latter case, the molecules can pack in the crystalline form without an overall, destabilizing accumulation of charge.

Temperature. Temperature has a marked affect on many of the factors that govern the solubility of a macromolecule. The dielectric constant decreases with increase in temperature, and the entropy terms in the free energy tend to dominate the enthalpy terms (Blundell & Johnson, 1976[link]). The temperature coefficient of solubility varies with ionic strength and the presence of organic solvents. McPherson (1985b[link]) gives a useful account of protein crystallization by variation of pH and temperature.

Organic solvents. Addition of organic solvents can produce a marked change in the solubility of a macromolecule in aqueous solution (care should be taken to avoid denaturation). This is partly due to a lowering of the dielectric constant, but may also involve specific solvation and displacement of water at the surface of the macromolecule. Generally, the solubility decreases with decrease of temperature when substantial amounts of organic solvent are present. Commonly used organic precipitants are listed in Table 3.1.1.2[link], column (b) (McPherson, 1985a[link]).

3.1.1.5. Screening procedures for the crystallization of biological macromolecules

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Optimal conditions for crystal growth are often very difficult to predict a priori, although many proteins crystallize close to their pI. In order to surmount the problem of testing a very large range of conditions, Carter & Carter (1979[link]) devised the incomplete factorial method, in which a very coarse matrix of crystallization conditions is explored initially. Finer grids are then investigated around the most promising sets of coarse conditions. This technique has been further refined to yield the sparse-matrix sampling technique described by Jancarik & Kim (1991[link]). Table 3.1.1.3[link] lists the crystallization parameters used by these authors. The 50 conditions constituting the sparse matrix are given in Table 3.1.1.4[link]. A recent update of this matrix and a set of stock solutions in the form of a crystal screen kit can be obtained commercially from Hampton Research (1994[link]). Further developments in screening methods are described in Volume D50 (Part 4) of Acta Crystallographica (1994).

Table 3.1.1.3| top | pdf |
Crystallization matrix parameters for sparse-matrix sampling, adapted from Jancarik & Kim (1991[link])

Precipitating agents
Non-volatileSaltsVolatileMixture
2-Methyl-2,4-pentanediol (MPD) Na, K tartrate 2-Propanol NH4 sulfate + PEG
Poly(ethylene glycol) (PEG) 400 NH4 phosphate   2-Propanol + PEG
PEG 4000 NH4sulfate    
PEG 8000 Na acetate    
  Li sulfate    
  Na formate    
  Na, K phosphate    
  Na citrate    
  Mg formate    
Range of pH: 4.6, 5.6, 7.5, 8.5
Salts, additives: Ca chloride, Na citrate, Mg chloride, NH4 acetate, NH4 sulfate, Mg acetate, Zn acetate, Ca acetate

Table 3.1.1.4| top | pdf |
Reservoir solutions for sparse-matrix sampling (Jancarik & Kim 1991[link])

No.SaltBufferPrecipitant
(% by mass)
1 0.02 M Ca chloride 0.1 M Acetate 30% MPD
2     0.4 M Na, K tartrate
3     0.4 M NH4 phosphate
4   0.1 M Tris 2.0 M NH4 sulfate
5 0.2 M Na citrate 0.1 M Hepes 40% MPD
6 0.2 M Mg chloride 0.1 M Tris 30% PEG 4000
7   0.1 M Cacodylate 1.4 M Na acetate
8 0.2 M Na citrate 0.1 M Cacodylate 30% 2-Propanol
9 0.2 M NH4 acetate 0.1 M Citrate 30% PEG 4000
10 0.2 M NH4 acetate 0.1 M Acetate 30% PEG 4000
11   0.1 M Citrate 1.0 M NH4 phosphate
12 0.2 M Mg chloride 0.1 M Hepes 30% 2-Propanol
13 0.2 M Na citrate 0.1 M Tris 30% PEG 400
14 0.2 M Ca chloride 0.1 M Hepes 28% PEG 400
15 0.2 M NH4 sulfate 0.1 M Cacodylate 30% PEG 8000
16   0.1 M Hepes 1.5 M Li sulfate
17 0.2 M Li sulfate 0.1 M Tris 30% PEG 4000
18 0.2 M Mg acetate 0.1 M Cacodylate 20% PEG 8000
19 0.2 M NH4 acetate 0.1 M Tris 30% 2-Propanol
20 0.2 M NH4 sulfate 0.1 M Acetate 25% PEG 4000
21 0.2 M Mg acetate 0.1 M Cacodylate 30% MPD
22 0.2 M Na acetate 0.1 M Tris 30% PEG 4000
23 0.2 M Mg chloride 0.1 M Hepes 30% PEG 400
24 0.2 M Ca chloride 0.1 M Acetate 20% 2-Propanol
25   0.1 M Imidazole 1.0 M Na acetate
26 0.2 M NH4 acetate 0.1 M Citrate 30% MPD
27 0.2 M Na citrate 0.1 M Hepes 20% 2-Propanol
28 0.2 M Na acetate 0.1 M Cacodylate 30% PEG 8000
29   0.1 M Hepes 0.8 M Na, K tartrate
30 0.2 M NH4 sulfate   30% PEG 8000
31 0.2 M NH4 sulfate   30% PEG 4000
32     2.0 M NH4 sulfate
33     4.0 M Na formate
34   0.1 M Acetate 2.0 M Na formate
35   0.1 M Hepes 1.6 M Na, K phosphate
36   0.1 M Tris 8% PEG 8000
37   0.1 M Acetate 8% PEG 4000
38   0.1 M Hepes 1.4 M Na citrate
39   0.1 M Hepes 2% PEG 400, 2.0 M Na sulfate
40   0.1 M Citrate 20% 2-Propanol + 20% PEG 4000
41   0.1 M Hepes 10% 2-Propanol + 20% PEG 4000
42 0.05 M K phosphate   20% PEG 8000
43     30% PEG 1500
44     0.2 M Mg formate
45 0.2 M Zn acetate 0.1 M Cacodylate 18% PEG 8000
46 0.2 M Ca acetate 0.1 M Cacodylate 18% PEG 8000
47   0.1 M Acetate 2.0 M NH4 sulfate
48   0.1 M Tris 2.0 M NH4 sulfate
49 1.0 M Li sulfate   2% PEG 8000
50 1.0 M Li sulfate   15% PEG 8000

Abbreviations: tris: 2-amino-2-(hydroxymethyl)-1,3-propanediol; hepes: 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid. Buffers: Na acetate buffer, pH = 4.6; Na citrate buffer, pH = 5.6; Na cacodylate buffer, pH = 6.5; Na hepes buffer, pH = 7.5; tris/HCl buffer, pH = 8.5.

3.1.1.6. Automated protein crystallization

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Several liquid-handling systems have been described that can automatically set up, reproducibly, a range of crystallization conditions (different protein concentrations, ionic strengths, amounts of organic precipitant, etc.) for the hanging-drop, sitting-drop, and microbatch methods. A useful introduction describing a system for mixing both buffered protein solutions and the corresponding reservoirs is given by Cox & Weber (1987[link]). Chayen, Shaw Stewart, Maeder & Blow (1990[link]) describe an automatic dispenser involving a bank of Hamilton syringes driven by stepper motors under computer control that can be used to set up small samples (2 µl or less) for microbatch crystallization (or hanging drops). Further systems have been described by Oldfield, Ceska & Brady (1991[link]), Eiselé (1993[link]), Soriano & Fontecilla-Camps (1993[link]), Sadaoui, Janin & Lewit-Bentley (1994[link]), and Chayen, Shaw Stewart & Baldock (1994[link]).

3.1.1.7. Membrane proteins

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Integral membrane proteins can be considered as those whose polypeptide chains span the lipid bilayer at least once. The external membrane segments exposed to an aqueous environment are hydrophilic, but it is the tight interaction of the hydrophobic segments of the chain with the quasisolid lipid bilayer that constitutes the major problem in their crystallization. Crystallization trials require disruption of the membrane, isolation of the protein, and solubilization of the resultant hydrophobic region (McDermott, 1993[link]). Organic solvents, chaotropic agents, and amphipathic detergents can be used to disrupt the membrane, but detergents such as β-octyl glucoside are most commonly used, since they minimize the loss of protein integrity. The several classes of detergent employed tend to be non-ionic or zwitterionic at the pH used, have a maximum hydrocarbon chain length of 12 carbon atoms, and possess a critical micelle concentration. The key to crystallizing membrane protein–detergent complexes appears to be the attainment of conditions in which the protein surfaces are moderately supersaturated and, in addition, the detergent micellar collar is at, or near, its solubility limit (Scarborough, 1994[link]). Most successful integral membrane protein crystallizations are near the micellar aggregation point of the detergent (Garavito & Picot, 1990[link]).

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