Progress in Biophysics and Molecular Biology 88 (2005) 339–357
Available online 7 October 2004
Keywords: Membrane protein crystallization; Amphiphile; Crystal packing; Lipid; Detergent
ARTICLE IN PRESS
www.elsevier.com/locate/pbiomolbio0079-6107/$ - see front matter r 2004 Elsevier Ltd. All rights reserved.
doi:10.1016/j.pbiomolbio.2004.07.006
E-mail address: pnollert@decode.com (P. Nollert).Contents
1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340
2. Amphiphilic microenvironments conducive to crystallization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341
3. The amphiphilic microenvironment inside membrane protein crystals . . . . . . . . . . . . . . . . . . . . . . . . . . . . 346
4. Amphiphile phase transitions during crystallization. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349Abstract
Integral membrane proteins are amphiphilic molecules. In order to enable chromatographic purification
and crystallization, a complementary amphiphilic microenvironment must be created and maintained.
Various types of amphiphilic phases have been employed in crystallizations and intricate amphiphilic
microenvironmental structures have resulted from these and are found inside membrane protein crystals. In
this review the process of crystallization is put into the context of amphiphile phase transitions. Finally,
practical factors are considered and a pragmatic way is suggested to pursue membrane protein
crystallization trials.
r 2004 Elsevier Ltd. All rights reserved.Review
Membrane protein crystallization in amphiphile phases:
practical and theoretical considerations
Peter Nollert
deCODE BioStructures, 7869 NE Day Rd. W, Bainbridge Island, WA 98110, USA

5. Practical considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 351
6. Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 353
Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 354
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 354
1. Introduction
Integral membrane proteins are amphiphilic molecules, they ‘‘love both’’, water as well as oil.
This duality is rooted in the physical nature of their surfaces: they all possess two fundamentally
different types of surfaces, a hydrophobic perimeter and two hydrophilic caps (Fig. 1). While
soluble proteins interact with water molecules and ions in an aqueous medium, membrane
proteins do this only with a fraction of their surface. The hydrophobic perimeter faces alkyl chains
of lipids and hydrophobic surfaces of other integral membrane proteins. Indeed, both of these
media need to be arranged within certain dimensions into a suitable microenvironment in order to
maintain the native conformation of the protein molecule. However, having individual particles is
required to subject these proteins to chromatographic purification procedures and to arrange
them into well-ordered three-dimensional arrays, crystals suitable for X-ray diffraction
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P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357340Fig. 1. Schematic depiction of a soluble protein and a transmembrane protein in their native environment, an aqueous
solution and a membrane bilayer, respectively. Both models are based on high-resolution X-ray crystallographic
experimental structures showing protein, water and lipids. Water molecules are red and blue, the surface of the protein
is blue where there is negative charge and red where there is positive charge, hydrophobic core and lipids are colored
gray. (A) Soluble protein particles are dissolved in a homogenous medium with a dielectric constant e close to that of
distilled water. On their surface they interact with water molecules and with ions. (B) Integral protein particles are
dissolved a low dielectric amphipilic medium, the lipid bilayer membrane, contacting the hydrophobic core, while theirexperiments. These particles have to be free to translate and rotate in space to bind, aggregate,
form a crystal nucleus and associate with the faces of a growing crystal. The restrictions set byhydrophilic caps are exposed to an aqueous medium similar to that shown in A for soluble proteins.

detergent micelles of uniform size would be most suitable for crystallization experiments. At the
ARTICLE IN PRESStime the choice of the detergent for crystallization purposes was based on three factors: (i)
stabilization of the native conformation of the membrane protein in monodisperse form, (ii)
enabling protein–protein contacts in the packed crystal and, (iii) preventing detrimental phase
separations during crystal growth. This line of thinking was expanded in the recent years,
particularly with respect to lipids being recognized as beneficial and sometimes crucial
crystallization components.
Most detergents belong into one of the following categories: ionic, non-ionic or zwitterionic.
Their characteristic behavior depends on their shape, stereochemistry of the head group and tail.
According to the ‘intrinsic curvature hypothesis’ (Gruner, 1985) they form supramolecular
structures in water due to the hydrophobic effect (Tanford, 1980) and their shape (Fig. 2A). At
sufficiently high concentrations, i.e. above the critical micellar concentration (CMC), detergents
form micelles. These form roughly spherical objects in which detergent molecules are primarily
packed with their alkyl chains towards the center and their head groups towards the surface
(Rosen, 1978; Wennerstroem and Lindman, 1979). Detergent molecules in these micelles are
flexible and exhibit a high degree of mobility, allowing for dramatic fluctuations in overall
micellar shape including deformations, fusion and fission (Tieleman et al., 2000). Amphiphiles
generally display a rich phase behavior commonly described in phase diagrams (Fig. 2B).these two requirements, (i) maintaining an amphiphilic microenvironment and, (ii) allowing
essentially free diffusion of small units are severe and are at the heart of the challenge to grow
crystals of integral membrane proteins.
Nonetheless, many membrane protein crystallizations have been reported (for a current update
on published membrane protein structures visit http://www.mpibp–frankfurt.mpg.de/michel/
public/memprotstruct.html or http://blanco.biomol.uci.edu/Membrane_Proteins_xtal.html) and
it is the goal of this review to point out practical as well as theoretical considerations these
crystallizations are based on. The focus is on highlighting the delicate interplay between small
molecule amphiphiles, detergents and lipids, and integral membrane proteins before, during and
after crystallization.
The critical role of amphiphiles in crystallization cannot be fully addressed in the context
of this review and the reader is therefore directed to reviews and monographs by Garavito
and Ferguson-Miller (2001), Garavito and Picot (1990), Wiener (2001), Michel
(1983, 1991), Hunte et al. (2003) and Iwata (2003). For a fundamental introduction to the
physical chemistry of detergents consult Tanford (1980), Rosen (1978) and Wennerstroem and
Lindman (1979).
2. Amphiphilic microenvironments conducive to crystallization
The first reports of successful membrane protein crystallizations (Michel and Oesterhelt,
1980; Henderson and Shotton, 1980; Garavito and Rosenbusch, 1980) created a paradigm
for membrane protein crystallization: transfer membrane proteins from their native
environment into particulate detergent micelles in order to purify and to crystallize them
in the same way as soluble proteins. It was reasoned that homogenous, lipid-free protein
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357 341Detergents typically have consolute boundaries, separating a single-phase micellar region from a

ARTICLE IN PRESSP. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357342dual micellar phase, the latter of which consists of a detergent rich and a detergent depleted phase.
At the cloud point a clear homogenous detergent solution turns turbid upon heating. Importantly,
the addition of salt, variations of pH, etc. and in particular the introduction of additional
components may have profound but essentially unpredictable effects on amphiphile phase
behavior. The fact that a ‘simple’ ternary system consisting of oil, water and block co-polymer
amphiphile may form nine different isothermal phases spectacularly illustrates this polymorphism
that is a typical feature of amphiphiles (Alexandridis et al., 1998). Lipid polymorph phases include
fluid isotropic phases, planar, positively and negatively curved bilayer phases, rod-shaped
hexagonal phases, micellar phases and bicontinuous cubic phases (Epand, 1997) and they occur,
among other things, as a function of composition, hydration, pressure, and temperature.
Some integral membrane proteins may be introduced into detergent micelles via refolding
(Dornmair et al., 1990). Indeed, since b-barrel proteins can be expressed in Escherichia coli as
inclusion bodies, membrane proteins such as OmpA or NspA can be purified in unfolded form
and crystallized without ever having had contact with lipids (Pautsch and Schulz, 1998;
Fig. 2. Introduction to small molecule amphiphile classes and an example for amphipile polymorphism. (A) According
to the ‘intrinsic curvature hypothesis’ (Gruner, 1985) amphiphiles form supramolecular structures in water due to the
hydrophobic effect (Tanford, 1980). In this simplistic view the shape, more specifically the ratio of cross section of
hydrophilic headgroup and hydrophobic tail, determines the type of structure they self-assemble into. I.e. wedge-shaped
amphiphiles assemble into structures with positive Gaussian curvature K such as micelles, cylinder-shaped amphiphiles
assemble into structures with zero Gaussian curvature K such as planar lipid bilayers and, cone-shaped amphiphiles
assemble into structures with negative Gaussian curvature K such as bicontinuous cubic phases. (B) Amphiphiles may
adopt a multitude of such supramolecular assemblies, macroscopic phases when mixed with water. For a particular
three component system, oil, water, amphiphilic block copolymer, Alexandris et al. (1998) describe a record nine
different isothermal phases including four cubic, two hexagonal and one lamellar lyotropic liquid crystalline phase and
two micellar solutions.

ARTICLE IN PRESSFig. 3. Schematic depiction of different amphiphilic micro-environments that membrane proteins have been
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357 343Buchanan, 1999), thus ensuring that once reconstituted into detergent micelles they form true
detergent–protein micelles (Fig. 3A).
In most cases however, membrane proteins are extracted concomitantly with associated lipids
from their native environment, cellular membranes. Such mixed systems, consisting of detergent,
lipids and membrane proteins form the so-called, protein detergent complexes (PDCs) (Fig. 3B).
The phase behavior of PDCs is expectedly complex and only in some select cases portions of
phase diagrams have been mapped out. It is this state however that is usually employed in
membrane protein crystallization trials and in many cases detergents as well as lipids are present
in membrane protein crystals (Table 1). Two methodological advances have substantially aided
many membrane protein crystallizations based on PDCs: (i) the introduction of small amphiphiles
such as 1,2,3-heptanetriol to modify micelle dynamics and size (Michel, 1983) and, (ii) the increase
of the size of the hydrophilic portion by complexing with monoclonal antibodies or fragments
thereof (Iwata et al., 1995; Hunte and Michel, 2002).
The range of alternative amphiphilic vehicles for membrane proteins useful for crystallization
purposes has increased in the recent years. Besides protein–detergent micelles and PDCs,
membrane protein crystallizations were started from membraneous structures. The latter may
be obtained by adding lipids to create structures that are small in size and planar such as bicelles
(Faham and Bowie, 2002) (Fig. 3C) or that are large such as in extended planar membranes
incorporated into for crystallization purposes. Membrane proteins are shown with two hydrophilic caps (yellow)
and a hydrophobic core (gray), and lipids (orange head) and detergents (green head) provide discrete (A, B, C) or
continous (D, E, F, G) amphiphilic assemblies. (A) Protein detergent micelle; (B) Protein detergent complex (PDC); (C)
Bicelle protein complex; (D) Planar membrane bilayer with embedded protein; (E) Curved proteoliposome; (F) High-
curvature regular spherical shell assembly; (G) Membrane proteins reconstituted in bicontinuous curved membranes of
a lipidic cubic phase of the diamond type. In all of these structures the amphiphile molecules are highly mobile (Zulauf,
1991) and thus constitute a ‘fluid’ hydrophobic host medium in which the integral membrane protein is embedded.

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Table 1
Amphiphile moieties detected by X-ray diffraction in membrane protein crystals reported in the protein data bank
Protein and organism Attributed identity Reference and PDB
accession code
Photosynthetic reaction centre,
Rhodopseudomonas viridis
Lauryl dimethylamine-N-Oxide Lancaster et al. (2000)
(1DXR)
Photosynthetic reaction centre,
Rhodobacter sphaeroides
Di-Stearoyl-3-SN-Phosphatidylcholine
cardiolipin
Camara-Artigas et al. (2002)
(1M3X)
Photosynthetic reaction centre,
Thermochromatium tepidum
Di-Palmitoyl-3-SN-phosphatidylethanolamine
lauryl dimethylamine-N-oxide
Nogi et al. (2000) (1EYS)
Photosynthetic reaction centre,
Thermosynechococcus
elongates
Dodecyl-beta-D-maltoside Ferreira et al. (2004) (1S5L)
Photosystem I, Synechococcus
elongatus
1,2-Distearoyl-monogalactosyl-diglyceride 1,2-
Dipalmitoyl-phosphatidyl-glycerole
Jordan et al. (2001) (1JB0)
Bacteriorhodopsin,
Halobacterium salinarum
2,10,23-Trimethyl-tetracosane 1-[2,6,10.14-
Tetramethyl-hexadecan-16-YL]- 2-[2,10,14-
Trimethylhexadecan-16-YL] glycerol
Luecke et al. (1999) (1C3W)
Bacteriorhodopsin,
Halobacterium salinarum
O3-Sulfonylgalactose Essen et al. (1998) (1BRR)
Halorhodopsin, Halobacterium
salinarum
1-Monooleoyl-RAC-glycerol palmitic acid Kolbe et al. (2000) (1E12)
Sensory Rhodopsin II,
Natronobacterium pharaonis
complexed with transducer
fragment
B-Octylglucoside Gordeliy et al. (2002) (1H2S)
Light-harvesting complex II,
spinach chloroplasts
1,2-Dipalmitoyl-phosphatidyl-glycerole
Digalactosyl diacyl glycerol B-Nonylglucoside
Liu et al. (2004) (1RWT)
Light-harvesting complex 2,
Rhodopseudomonas
acidophila
B-Octylglucoside Papiz et al. (2003) (1NKZ)
Cytochrome C Oxidase,
Paracoccus denitrificans
Di-Stearoyl-3-SN-phosphatidylcholine Harrenga and Michel (1999)
(1QLE)
Cytochrome c oxidase,
Paracoccus denitrificans
Lauryl dimethylamine-oxide Ostermeier et al. (1997)
(1AR1)
Cytochrome C oxidase, bovine
heart mitochondria
Tristearoylglycerol decyl-beta-D-maltopyranoside Tsukihara et al. (2003)
(1V55)
Cytochrome C oxidase,
Thermus thermophilus
B-Nonylglucoside Soulimane et al. (2000)
(1EHK)
Cytochrome C oxidase,
Rhodobacter sphaeroides
Di-Stearoyl-3-SN-Phosphatidylethanolamine Svensson-Ek et al. (2002)
(1M57)
Cytochrome bc1 complex,
Saccharomyces cerevisiae
1,2-Diacyl-SN-glycero-3-phoshocholine Di-
Palmitoyl-3-SN-phosphatidylethanolamine 1,2-
diacyl-SN-glycero-3-phosphoinositol undecyl-
maltoside
Lange et al. (2001) (1KB9)
Cytochrome B6F,
Chlamydomonas reinhardtii
Eicosane 1,2-Distearoyl-monogalactosyl-
diglyceride 1,2-DI-O-acyl-3-O-[6-deoxy-6-sulfo-
alpha-D-glucopyranosyl]-SN-glycerol
Stroebel et al. (2003) (1Q90)
Fumarate reductase,
Escherichia coli
O-Dodecanyl octaethylene glycol Verson et al. (1999) (1FUM)
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357344

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Table 1 (continued )
Protein and organism Attributed identity Reference and PDB
accession code
Fumarate reductase, Wolinella
succinogenes
Dodecyl-beta-D-maltoside Lancaster et al. (2001)
(1E7P)
Succinate dehydrogenase, E.
coli
Cardiolipin L-Alpha-phosphatidyl-beta-oleoyl-
gamma-palmitoyl-phosphatidylethanolamine
Yankovskaya et al. (2003)
(1NEK)
Potassium channels KcsA,
Streptomyces lividans
Diacyl glycerol Nonan-1-ol Valiyaveetil et al. (2002)
(1K4C)
Rhodopsin, bovine rod outer
segments
B-Nonylglucoside palmitoyl Okada et al. (2002) (1L9H)
Aquaporin AQP1, bovine red
blood cells
B-Nonylglucoside Sui et al. (2001) (1J4N)
Glycerol facilitator GlpF E.
coli
B-Octylglucoside Fu et al. (2000) (1FX8)
Aquaporin Z, E. coli B-2-Octylglucoside Savage et al. (2003) (1RC2)
Chloride channels, E. coli N-Octane pentadecane Dutzler et al. (2002) (1KPL)
Formate dehydrogenase, E. coli Cardiolipin Jormakka et al. (2002)
(1KQF)
Nitrate reductase A, E. coli 1,2-Diacyl-glycerol-3-SN-phosphate Bertero et al. (2003) (1Q16)
ADP/ATP carrier, bovine heart
mitochondria
Cardiolipin di-Stearoyl-3-SN-
phosphatidylcholine 3-Laurylamido-N,N0-
dimethylpropylaminoxyde
Pebay-Peyroula et al. (2003)
(1OKC)
Porin, Rhodopseudomonas
blastica
(Hydroxyethyloxy)tri(ethyloxy)octane Kreusch et al. (1994) (1PRN)
OmpK36 (osmoporin),
Klebsiella pneumoniae
Dodecane Dutzler et al. (1999) (1OSM)
OmpA-fragment, E. coli (Hydroxyethyloxy)tri(ethyloxy)octane Pautsch and Schulz (1998)
(1BXW)
OmpX, E. coli (Hydroxyethyloxy)tri(ethyloxy)octane Vogt and Schulz (1991)
(QJ8)
NspA, Neisseria meningitidis Pentaethylene glycol monodecyl ether Vandeputte et al. (2003)
(1P4T)
FhuA, E. coli 3-Deoxy-D-manno-OCT-2-ulosonic acid Ferguson et al. (1998)
(1FCP)
2-Tridecanoyloxy-pentadecanoic acid
3-Oxo-pentadecanoic acid
FhuA, E. coli N-Octyl-2-hydroxyethyl sulfoxide Locher et al. (1998) (1BY3,
1BY5)
FhuA, E. coli Decylamine-N,N-dimethyl-N-oxide Lauric acid 3-
Hydroxy-tetradecanoic acid 3-Deoxy-D-manno-
oct-2-ulosonic acid Myristic acid
Ferguson et al. (2000)
(1QFF)
Ferric citrate transporter
(FecA) E. coli
Lauryl dimethylamine Heptane Ferguson et al. (2002)
(1KMO)
Cobalamine transporter BtuB,
E. coli
(Hydroxyethyloxy)tri(ethyloxy)octane Chimento et al. (2003)
(1NQF)
outer membrane phospholipase
A (OMPLA), E. coli
B-Octylglucoside Snijder et al. (1999) (1QD5)
protease OmpT, E. coli B-Octylglucoside 2-Methyl-2,4-pentanediol Vandeputte-Rutten et al.
(2001) (1I78)
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357 345

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document the variability and abundance of lipids and detergents found in membrane protein crystals (the list was built
from http://www.mpibp-frankfurt.mpg.de/michel/public/memprotstruct.html, version of June 29th, 2004, 75 entries).
˚(Fig. 3D) or those that exhibit positive (Figs. 3E, F), or negative curvature (Fig. 3G).
Curved membrane protein containing bilayers include proteoliposomes (Takeda et al., 1998;
Liu et al., 2004) or bincontinuous lipidic cubic phases (Landau and Rosenbusch, 1996). Of course,
the quantity and type of lipid added to PDCs determine the nature of the resulting amphiphile
phase.
For some membrane proteins it is very difficult to identify detergent-based conditions
that preserve their native conformation. This predicament has inspired several research
groups to expand the range of solubilization tools by designing new amphiphiles and to
investigate the richness of their phase behavior for the purpose of membrane
protein crystallization (Fig. 4). Among these amphiphiles are the so-called peptitergents
(Schafmeister et al., 1993), lipopeptide detergents (McGregor et al., 2003), amphiphols
The identification of the complete structure of these amphiphiles is somewhat tentative. At resolutions above 2.5A
crystallographers often resort to interpreting weak electron density maps around the hydrophobic perimeter of
membrane proteins with the crystallization condition in mind, e.g. what detergent was used. However, in several cases
the identity of amphiphiles were carefully determined independently (Belrhali et al., 1999; Essen et al., 1998).Table 1 (continued )
Protein and organism Attributed identity Reference and PDB
accession code
Outer-membrane adhesin
OpcA, Neisseria meningitidis
Pentaethylene glycol Prince et al. (2002) (1K24)
Long-chain fatty acid
transporter FadL, E. coli
(Hydroxyethyloxy)tri(ethyloxy)octane Lauryl
dimethylamine-N-oxide
van den Berg et al. (2004)
(1T16)
The proteins listed represent a subset of currently determined membrane protein structures and they were selected to
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357346(Tribet et al., 1996; Popot et al., 2003) and new detergents with reduced alkyl chain mobility
such as tripod amphiphiles (Yu et al., 2000). They all self-assemble into small micelles, can
disperse lipid membranes and are gentle, non-denaturing amphiphiles preserving the native
structure of the test protein bacteriorhodopsin and other membrane proteins in solution for
extended periods of time.
3. The amphiphilic microenvironment inside membrane protein crystals
Membrane protein crystals consist of three major molecular species: water, amphiphile and
membrane protein. Water and dissolved solutes form a fluid phase within the rigidly packed
protein network, while amphiphiles may tightly bind to the protein surface and/or form a
disordered state. The morphology and the strength of intermolecular contacts in protein crystals
were investigated by Matsuura and Chernov (2003). It was found that soluble proteins form
crystal contacts frequently involving water molecules, which form specific intermolecular
hydrogen bonds on top of non-specific attractive electrostatic interactions. Similar contacts are

ARTICLE IN PRESSpresent between hydrophilic protein surfaces of membrane proteins in crystals. In some cases
Fig. 4. Cartoons of recently developed new classes of amphiphile molecules. (A) Peptitergents are designed peptides
that form amphiphatic a-helices. Their hydrophobic faces were made to interact with the hydrophobic face of integral
membrane proteins (Schafmeister et al., 1993). (B) Lipopeptides consist of a peptide scaffold supporting two alkyl
chains each anchored at the end of an a-helix. They were designed to provide a rigid outer hydrophilic shell surrounding
an inner ‘soft’ lipidic core (McGregor, 2003). (C) Amphiphols are amphiphilic polymers with a hydrophilic backbone
and hydrophobic grafted chains. They exhibit favorable phase transitions that are expected to be useful for membrane
protein crystallization experiments (Tribet et al., 1996). (D) The design of tripod amphiphiles such as #8 in Yu et al.
(2000) is based on the rationale that detergents with diminished flexibility should be more prone to form ordered crystal
lattices. The core is a terasubstituted carbon atom that limits the flexibility of three attached hydrophobic and a
hydrophilic substitutent.
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357 347amphiphiles form interactions that are crucial to crystal packing.
The structures that amphiphiles from within membrane protein crystals are remarkably
diverse (Fig. 5) and go beyond the simple two-type classification introduced by Michel
(1983). According to these categories type I crystals consist of stacked membraneous layers
(Fig. 5B). They stick together via hydrophobic interactions in the plane of the layers, resembling
2D crystals, and polar contacts mediate interlayer interactions. Conversely, type II crystals
possess contacts involving the polar regions of membrane proteins only. The hydrophobic
perimeter is embedded in a torus of detergent molecules. Typical type I crystals have been
found in all cases where membrane proteins were crystallized with the cubic phase
method (Landau and Rosenbusch, 1996; Kolbe et al., 2000; Royant et al., 2001; Katona
et al., 2003). Subtle differences in symmetry and layer arrangement exist though. Bacteriorhopsin
packs in a unidirectional way ‘‘head to tail’’ (Belrhali et al., 1999), while halorhodopsin
packs in layers where heads interact with heads and tails with tails (Kolbe et al., 2000),
and sensory rhodopsin II packs in layers with mixed up–down arrangements (Royant et al.,
2001). Bacteriorhodopsin crystals were shown by mass spectrometry to contain native
lipids that were co-purified, namely 2,3-di-O-phytanyl derivatives of phosphatidylglycerol,
phosphatidylglycerol sulfate, phosphatidylglycerol phosphate methylester, triglycosyldiether,
sulfated triglycoside lipid and sulfated tetraglycosyldiphytanylglycerol (Belrhali et al.,
1999) when crystallized from lipidic cubic phases and they contained similar lipids
when crystallized as PDCs (Essen et al., 1998). A detailed description of the interaction of lipids
within bacteriorhodopsin crystals is provided by Belrhali et al. (1999) and by Cartailler and
Luecke (2003).

ARTICLE IN PRESSP. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357348Recently Liu et al. (2004) have described crystals of the Light harvesting chlorophyll a=b
protein complex from pea thylacoid membranes. Hollow spherical shell assemblies were produced
and packed into well-ordered three-dimensional crystals (Fig. 5C). The icosahedral spheres have a
diameter of 250 A˚ and contain 20 protein trimers and several disordered lipid molecules.
Apparently the space between the LHC-II trimers in the icosahedral structure is completely filled
with digalactosyl diacylglycerol and/or phosphatidylglycerol and/or nonylglucoside and thus
resemble small proteoliposomes.
Fig. 5. Samples of experimentally determined and deduced amphiphile arrangements in crystals of membrane proteins.
(A) ‘Type II’ discontinuous arrangement of micellar Octyl-POE around the hydrophobic perimeter of the detergent
C8E4 and OmpF porin (E. coli) R3 crystal (Pebay-Peyroula et al., 1995). Protein detergent micelle with a belt of
detergent as shown by neutron density (green). (B) ‘Type I’ continuous arrangement of stacked layers consisting of
extended two-dimensional sheets of purple membrane (Halobacterium Salinarum). Bacteriorhodopsin trimers are
separated in plane by a belt of native lipids (Belrhali et al., 1999). (C) Discontinuous icosahedral packing of spherical
LHC-II proteoliposomes (Liu et al., 2004). (D) Continuous network of b-Octylglucoside extending throughout the
entire P3121 crystal of phospholipase A OmplA (E. coli) (Snijder et al., 2003).

same time conditions need to be provided that allow the hydrophobic sections to maintain or to
ARTICLE IN PRESSrearrange into the suprastructures exemplified in Fig. 5. How is this achieved?
Crystallizations involve phase transitions: an initial homogenous medium separates into a
depleted phase and a rich in amphiphile and membrane protein, the crystal (Fig. 6). The
association of protein/detergent micelles or PDCs into a type II packed crystal can easily be
understood within the framework of present crystallization theory. The mechanism for PDC
crystallization was investigated by Marone et al. (1998) for Photosynthetic reaction centers. These
were shown to exist predominantly in the monomeric form throughout the entire crystallization
process. Small-angle neutron scattering experiments suggested that initial crystal nuclei are
kinetically transient and thermodynamically unstable intermediates and single monomer
complexes serve as crystal growth units. Comparable experiments were used to characterize the
effects of crystallization additives on the shape of pure detergent micelles (Littrell et al., 2000). It
was found that micelles were elongated and rod-like in form and that their size grew by increasing
the ionic strength and shrunk when glycerol or PEG was added. Studies like these provide a
rational basis for detergent-specific formulation screen development.
In order to form continuous structures such as layered type I crystals or crystals with aSnijder et al. (2003) describe the packing of outer membrane phospholipase A in crystals where
a crystal-penetrating continuous three-dimensional detergent network exists (Fig. 5D). They
denote this arrangement as type III. Here hydrophobic and polar contacts contribute to the
integrity of the crystal.
It is instrumental to compare these packing arrangements with those that can form by small
molecule amphiphiles themselves. Type I packing corresponds to lamellar phases, type II packing
to micellar phases. Type III and the icosahedral shell packing do not have an immediate
counterpart in amphiphile polymorph structures, however, continuous three-dimensional
networks such as those formed in bicontinuous cubic phases share some structural and
topological similarities with membrane protein crystals (Fig. 2B). This resemblance is not
unexpected since one can understand the membrane protein as a modifying component of
amphiphile polymorphism and phase behavior. Generally crystals represent lowest enthalpy states
and in the case of these membrane protein/amphiphile co-crystals it is not clear which of the two
components (membrane protein or amphiphile) dominates in providing crystal forming enthalpy.
4. Amphiphile phase transitions during crystallization
The crystallization process consists of two steps, nucleation and crystal growth. Nucleation is a
critical phenomenon and hardly anything is known specifically for membrane protein crystal-
lizations. Soluble protein crystallization growth is mainly driven by modification of the water
structure, creating conditions that allow and favor the defined association of proteins (Kam et al.,
1978; Weber, 1991). Similar conditions need to be created for the hydrophilic sections of
membrane proteins, i.e. screening of repulsive electrostatic surface charges by ions and providing
conditions where the protein is at supersaturation. The latter effect was investigated by Rosenow
et al. (2003) who concluded from biochemical studies that membrane protein crystallization is
favored with those amphiphiles that optimize the solubility of integral membrane proteins. At the
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357 349continuous network of amphiphile phase, PDCs need to fuse and allow detergent and lipid

ARTICLE IN PRESSP. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357350molecules to rearrange into the supramolecular architecture described above. Snijder et al. (2003)
point out that the continuous detergent network in their crystals and the hydrophobic crystal
contacts suggest that OmplA molecules approach each other closely and coalesce their detergent
belts. The formation of the polar contacts might actually drive crystallization and induce this
merging of micelles. They hypothesized that micelle fusion and the stabilization of a continuous
network is mediated by the organic solvent and the amphiphile 2-methyl-2,4-pentanediol. Indeed,
many of the putative type III crystal yielding conditions include the use of rather high
concentrations of organic solvents or small molecule amphiphiles such as 1,2,3-heptanetriol. The
picture emerges that this crystallization process may be driven partly or possibly be dominated by
amphiphiles undergoing a phase transition prompted by the increase in system complexity.
The formation of regular bilayer stacks from bicelles may be understood along similar lines of
reasoning (Faham and Bowie, 2002). An interesting twist though is brought about by the
incorporation of membrane proteins into membrane-like vehicles (Figs. 3D–G). Doing so reduces
two rotational degrees of freedom and one translational degree of freedom, effectively reducing
the entropic penalty for crystallization. This penalty is paid already at the step of reconstitution,
Fig. 6. Schematic representation of membrane protein crystallization concepts, routes and examples for the resulting
packing arrangements. The detergent-solubilized membrane protein may be delipidated (Pautsch and Schulz, 1998) or
may carry affine lipids throughout the purification process. Alternatively, membrane-forming lipids may be added to
form a PDC, a protein/detergent/lipid complex (Zhang et al., 2003) or bicelles (Faham and Bowie, 2002). Structured
amphiphilic phases can form when lipids with zero or negative spontaneous curvature are added to detergent solubilized
membrane protein preparations. Proteoliposomes may fuse and form layered stacks or they may directly assemble into
a regular array. Bicontinuous lipidic cubic phases can provide a matrix for embedding membrane proteins and allow
these to nucleate and form layered crystals (Nollert et al., 2001).

A kinetic barrier model was developed that together with the energetics predicts a limitation of the
ARTICLE IN PRESSlipidic cubic phase method to membrane proteins with more than five a-helices and confines a
window for the cubic lattice parameter within which crystallization may occur.
5. Practical considerations
Hunte et al. (2003) and Iwata et al. (2003) provide very useful practical guides for bench
scientists to attempt membrane protein crystallizations. Strategies for rational and random screen
design and procedures for re-lipidation and lipid supplementation are summarized in this section.
Tools are currently being developed by several groups to design screens for specific amphipile
systems. Piazza et al. (2002) aim at gaining control over the phase behavior of LDAO. They
modulate the ionization of LDAO, a pH-sensitive surfactant, to phase segregate Photosynthetic
Reaction Center by adjusting temperature, salt and pH. Tanaka et al. (2003) rationalize PEG-
based cytochrome bc1 complex crystallizations with a simple model. They use dynamic light
scattering to estimate interactions between PDCs of the bc1 complex. They control the stability of
the liquid phase of BC1 solutions via the ratio of (the range of depletion zone)/(the radius of a
BC1 particle). The subsequent crystallizations were most successful at conditions where the
stability of the liquid phase changed from stable to unstable.
Apart from such detailed characterizations, membrane protein crystallographers have
traditionally paid close attention to occurrences of phase separation (Michel, 1991; Wiener,
2001; Garavito and Ferguson-Miller, 2001). For instance, Trimeric Photosystem I from
Cyanobacterium (Synechococcus elongates) can be isolated as PDCs with stereochemically pure
b-dodecylmaltoside (Fromme and Witt, 1998) and crystals readily grow from small amphiphile-prior to the actual crystallization event and may thus positively offset enthalpic crystallization
energy contributions.
The crystallization of LHC-II proteoliposomes is a special case with a significant contribution
from the amphiphiles. The mechanism was explained as that of electrostatically driven particle
aggregation (Hino et al., 2004). Spherical shell assemblies were not visible by electron microscopy
prior to crystallization. However, when the KCl concentration was increased above 20mM,
spherical shell assemblies formed and above 30mM KCl crystallization into octahedral crystals
was observed. The authors speculate that it is not the protein but the amphiphiles that determine
the small size of the proteoliposomes, i.e. by virtue of the asymmetric charge distribution on the
protein and subsequent asymmetric association with charged lipids. Specifically, the predomi-
nantly negative charge on the stromal side faces outward and during the formation of
proteoliposomes their electrostatic repulsion is shielded and allows the membrane curvature to
increase. Ready built spheres then stack into octahedral crystals. Proteoliposomes may also fuse
to layers, possible by a mechanism resembling biological fusion (Zhang et al., 2003).
The formation of type II crystals from initial homogenously dispersed membrane proteins in
bicontinuous lipidic cubic phases was rationalized qualitatively as a phase separation process
(Nollert et al., 2001) and a quantitative theory (Grabe et al., 2003) was developed based on this
hypothesis. In short, it was shown that it is energetically favorable for membrane proteins to
cluster together in flattened regions of the otherwise highly curved membrane of the cubic phase.
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357 351rich droplets during dialysis against low salt concentrations (Fromme, 2003). Supposedly these

ARTICLE IN PRESSdroplets form a separate, amphiphile and protein-rich phase and serve as nucleation
and growth sites (P. Fromme, personal communication). It is possible that the crystallization
mechanism resembles that hypothesized for the crystallization of bacteriorhodopsin in lipidic
cubic phases.
Furthermore, crystallizers may systematically investigate amphiphile systems close to
macroscopically observable phase separations and bias crystallization screening formulations
towards conditions that produce such features (Hitscherich et al., 2001). Wiener and Snook (2001)
for instance, base their screen development on the hypothesis that the properties of the pure
detergent solutions and their phase behavior plays a significant role during the membrane protein
crystallization process. They characterize detergent phase properties and utilize dye partitioning in
detergent/solute mixtures to determine phase boundaries. Ultimately they arrive at new detergent-
specific screening formulation kits.
Until recently membrane protein crystallizers faced a paradoxical situation. On one hand it was
well established that lipids may aid the crystallization process with lipids regularly showing up in
the final crystal (Table 1), on the other hand their identity and quantity are rarely reported in the
methods section of the corresponding publications. This lack of critical information makes it very
difficult to reproduce membrane protein crystallizations. Fortunately, simple procedures to
monitor lipid and detergent in membrane protein samples have been worked out. daCosta and
Baenziger (2003) describe using Fourier-transform infrared spectroscopy for this purpose. This
method reduces the sample volume to 10ml and molar lipid:protein ratios down to 5:1 (for a
300 kDa protein) can be determined. Their ratiometric assay utilizes the intensity of the lipid ester
carbonyl band at 1740 cm1 and the protein amide I band at 1650 cm1. Detergent analysis can be
included by observation of vibrations in the 1200–1000 cm1 region and quantified using a
standard curve.
Crucially, the types of lipids that membrane proteins are associated with originate form their
host membrane. Therefore, lipid compositions in membrane protein sample preparations depend
on tissue and cell types. For the insect cell baculovirus expression system the lipid profile was
shown to vary as a function of cell line (Spodoptera frugiperda vs. Trichoplusia ni) and infection
state (Marheineke et al., 1998). Lipid supplementation and its use for the crystallization of
cytochrome b6f in defined protein–detergent–lipid complexes are described by Zhang et al. (2003).
Crystallization of the delipidated protein was not possible but when supplemented with a
synthetic, non-native lipid, dioleoyl-phosphatidylcholine, well-diffracting crystals formed.
Similarly, five to 13 lipid molecules per protein particle were required for crystal formation of
the human erythrocyte anion-exchanger membrane domain (Lemieux et al., 2002).
The advent of high-throughput crystallization techniques and microcrystallization methods
(Santarsiero, 2002; Nollert, 2002) have helped substantially to cover larger portions of the
multidimensional crystallization phase space and it is expected that membrane protein
crystallization projects may be initiated even though only small amounts of protein samples
can be produced.
Regardless of these and future latest ‘tips and tricks’—some membrane proteins crystallize
more readily than others. In this respect, bacteriorhodopsin has been named ‘the lysozyme of
membrane proteins’. It can be crystallized starting from purple membranes, from PDCs or
proteoliposomes (Fig. 3) with several crystallization methods (Fig. 7). Unfortunately, not every
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357352membrane protein is that well-behaved.

ARTICLE IN PRESS6. Conclusions
The rate at which new membrane protein structures are reported has steadily increased over
time. This progress should be attributed to a greater willingness of investigators to initiate such
projects, shifts in funding preferences and in the availability of molecular biological methods, i.e.
homology screening and expression of fragments, rather than to a thorough understanding and
application of amphiphile phase science. So does it help at all to invest resources into the
understanding of amphiphile phase behavior (Caffrey, 2003) when tackling the crystallization of a
particular membrane protein? Although one would certainly wish so, the truth is that many, if not
most successful crystallizations were initially discovered by trial and error, with little or no insight
into the phase behavior of the components. One might argue though, that these crystallizations
Fig. 7. Venn diagram schematizing crystallization method space. Depicted are four different amphiphilic micro-
environments that support membrane protein crystallization. For illustration purposes representative examples are
pointed out.
P. Nollert / Progress in Biophysics and Molecular Biology 88 (2005) 339–357 353would have been found in a more expedient way, if the phase behavior would have been known
ahead of time and properly applied. Likewise, for some of the many membrane proteins that have
failed crystallization attempts, insight into the amphiphile phase behavior might have turned these
unreported failures into success stories.
What would then constitute a sensible and pragmatic way to efficiently conduct membrane
protein crystallization trials? Along the way to overcome the barriers of expression and
purification crucial insight into the compatibility with certain detergents and biochemical
procedures is often gained. This information is very useful for crystallization trial design.
Furthermore, since the co-purified lipids play an important role in crystallization it is advisable to
monitor their type and quantities. Micro-analytical techniques such as mass spectrometry
(Belrhali et al., 1999; Cohen and Chait, 2001) and Fourier transform infrared spectroscopy
(DaCosta and Baenziger, 2003) provide means to monitor and adjust the lipid content. Such
characterizations help to assure sample consistency and increase the odds to arrive at reproducible
crystallization conditions.
Once compatible detergents are found and samples are available crystallization setups can be
set up. At this point even a rudimentary understanding of the phase behavior of the detergent of
choice can help to prevent wasting sample. For instance, if the crystallization of partially

Acknowledgements
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