Amphipols From A to Z

Jean-Luc Popot, Thorsten Althoff, Dominique Bagnard, Jean-Louis Banères, Paola Bazzacco, Emmanuelle Billon-Denis, Laurent J Catoire, Philippe Champeil, Delphine Charvolin, Melanie J. Cocco, et al.

► To cite this version:

Jean-Luc Popot, Thorsten Althoff, Dominique Bagnard, Jean-Louis Banères, Paola Bazzacco, et al. Amphipols From A to Z. Annual Review of Biophysics and Biomolecular Structure, 2011, 40 (1), pp.379-408. 10.1146/annurev-biophys-042910-155219. hal-02327349

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<figure><img src="image_1.png" /></figure>
# Amphipols From A to Z

J.-L. Popot,1 T. Althoff, D. Bagnard, J.-L. Banères, P. Bazzacco, E. Billon-Denis, L.J. Catoire, P. Champeil, D. Charvolin, M.J. Cocco, G. Crémel, T. Dahmane, L.M. de la Maza, C. Ebel, F. Gabel, F. Giusti, Y. Gohon, E. Goormaghtigh, E. Guittet, J.H. Kleinschmidt, W. Kühlbrandt, C. Le Bon, K.L. Martinez, M. Picard, B. Pucci, J.N. Sachs, C. Tribet, C. van Heijenoort, F. Wien, F. Zito, and M. Zoonens

¹Institut de Biologie Physico-Chimique, CNRS/Université Paris-7 UMR 7099, F-75005 Paris, France; email: Jean-Luc.Popot@ibpc.fr

## Abstract

Amphipols (APols) are short amphipathic polymers that can substitute for detergents to keep integral membrane proteins (MPs) water soluble. In this review, we discuss their structure and solution behavior; the way they associate with MPs; and the structure, dynamics, and solution properties of the resulting complexes. All MPs tested to date form water-soluble complexes with APols, and their biochemical stability is in general greatly improved compared with MPs in detergent solutions. The functionality and ligand-binding properties of APol-trapped MPs are reviewed, and the mechanisms by which APols stabilize MPs are discussed. Applications of APols include MP folding and cell-free synthesis, structural studies by NMR, electron microscopy and X-ray diffraction, APol-mediated immobilization of MPs onto solid supports, proteomics, delivery of MPs to preexisting membranes, and vaccine formulation.

## Keywords

membrane proteins, amphipathic polymers, membrane biophysics, membrane biochemistry

## INTRODUCTION

Membrane proteins (MPs) are notoriously more difficult to study in vitro than their soluble counterparts, mainly because of two factors: (a) the relative scarcity of most MPs and the difficulties encountered to overexpress them under a functional form, and (b) their frequent instability outside biological membranes. The complexity of a natural membrane is such that this environment is essentially impossible to mimic, even with artificial lipid bilayers, and much less so in aqueous solutions. Detergents are traditionally used to screen the hydrophobic transmembrane (TM) surface of MPs and make them water soluble, but they tend to inactivate them. There are several reasons why detergents destabilize MPs, possibly including the loss of physical constraints provided by the

membrane environment. It is, however, our contention that a major factor is the competition of detergents with the protein-protein and protein-lipid interactions that stabilize the native structure of MPs (for a discussion, see Reference 67). This effect is compounded by the fact that MP/detergent complexes must be handled in the presence of free detergent micelles, because below the critical micellar concentration (CMC) of the detergent, entropic effects drive the aggregation of transmembrane surfaces (for a recent discussion, see Reference 89). Free micelles act as a hydrophobic sink into which lipids, cofactors, and subunits can disperse, which is a leading cause of MP inactivation. This view has been at the inception of the work summarized in the present review.

There are several possible ways to reduce or suppress the hydrophobic sink (for a discussion,

MP: membrane protein

Integral membrane protein: a protein that is in contact with the hydrophobic interior of a biological membrane to which it exposes an extensive transmembrane hydrophobic surface

Detergent: a surfactant with the ability to solubilize fats
see Reference 67). One of them is to replace detergents with compounds that would have such a high affinity for the transmembrane surface of MPs that they would remain associated to it even in the presence of vanishingly low concentrations of free micelles. This can theoretically be achieved by endowing a hydrophilic polymer with a multitude of hydrophobic chains that would act as many anchoring points. This consideration has led to the development of amphipols, a class of amphipathic polymers specially designed to complex MPs and keep them water soluble. Here the term amphipol (APol) refers to "an amphipathic polymer that is able to keep individual MPs soluble under the form of small complexes" (67).

Fifteen years have passed since the feasibility of this approach was first demonstrated (87). In the interim, much work has been devoted to learning to control the synthesis of these molecules and to understanding their physical chemical properties, to investigating the nature and properties of the complexes they form with MPs, and to developing their applications. As will be shown, the latter have extended well beyond the initial idea of enabling the biochemist and biophysicist to carry out basically the same studies as in detergent solution, but under improved conditions. Some APols are by now commercial. The thirty-odd publications that have appeared to date about the properties and uses of APols provide their potential users with a solid body of data to help them to exploit applications that have been validated and to develop novel ones. Previous reviews have discussed the advantages and disadvantages of APols versus other novel approaches to stabilizing MPs and studying them in vitro (12, 35, 67). Two other reviews have focused on APols and cover the field up to 2003 (68, 79).

Over the years, several types of APols have been designed, synthesized, and tested. Their study is at various stages of development. In the present review, we refer mainly to the following four types of APols: (a) A8-35 (Figure 1a), the first APol to be validated, whose solubility is due to the presence of carboxylates (87); (b) phosphorylcholine-

based APols (PC-APols) (Figure 1b) (25, 26); (c) glucose-based, nonionic APols (NAPols) (Figure 1c) (5, 6, 71, 83); and (d) sulfonated APols (SAPols) (Figure 1d) (23, 64). Other types of APols are described in earlier reviews (67, 68, 79).

## CHEMICAL STRUCTURE AND SOLUTION PROPERTIES OF AMPHIPOLS: A SPECIAL CLASS OF AMPHIPATHIC POLYMERS

Using APols efficiently depends on a good understanding of the way these polymers are designed and how they behave in aqueous solutions. In the present section, we summarize the chemical structure and solution behavior of the best field-tested APols and we describe the labeled or functionalized APols that have been developed and tested thus far.

### Design, Synthesis, and Solution Properties of Amphipols

APols are amphipathic polymers, carrying both many hydrophilic and many hydrophobic groups (Figure 1). Being amphipathic, they belong, along with detergents, to the general class of surfactants. APols present a distinctive feature compared to most other amphipathic polymers: They self-assemble into well-defined particles comprising a few macromolecules [e.g., four molecules, on average, for 9- to 10-kDa A8-35 (32), three for 25-kDa NAPols (6)]. This results from the favorable combination of a short backbone chain length and an appropriate number and size of hydrophobic segments (so that the equilibrium size of the hydrophobic core can be reached with the amount of hydrophobic groups carried by just a few chains, while avoiding the otherwise typical formation of large networks of interconnected polymers due to hydrophobic self-association), flexibility (allowing a small radius of curvature of the interface with the solution), and a limited polydispersity in composition (to avoid large populations of chains with too few or too many hydrophobic groups compared to the optimum). In practice, macromolecules

Critical micellar concentration (CMC): the concentration of a small surfactant at which its molecules start to organize into micelles

Amphipathic polymer: a polymer comprising both hydrophobic and hydrophilic moieties

Amphipol (APol): an amphipathic polymer that can keep membrane proteins water soluble in detergent-free solutions as small individual entities by adsorbing onto their transmembrane surface

A8-35: a particular type of anionic amphipol

PC-APol: phosphorylcholine-based APol

NAPol: nonionic amphipol

SAPol: sulfonated APol

Surfactant: a compound that adsorbs at the air/water interface and thereby lowers the surface tension of water
<figure><img src="image_2.png" /></figure>

<figure><img src="image_3.png" /></figure>

<figure><img src="image_4.png" /></figure>

<figure><img src="image_5.png" /><figcaption>Figure 1</figcaption></figure>

Chemical structures of four types of amphipols. (a) A polyacrylate-based APol, A8–35 (87); (b) a phosphorylcholine-based APol, C22–43 (26); (c) a nonionic, glucose-based APol (71); and (d) a sulfonated APol (23).

reaching or approaching such properties belong to one of the two following types: polysoaps, i.e., homopolymers made from amphipathic monomers bearing both alkyl and hydrophilic moieties, as is the case of homopolymeric NAPols (Figure 1c), or copolymers featuring a high density of hydrophobic monomers with short alkyl side groups ($C_8-C_{12}$) separated by one or a few hydrophilic monomers (all other APols described to date). Both types of APols typically contain a few tens of alkyl groups per chain.

When APols were initially devised, their expected mode of interaction with MPs was the primary concern (68, 87). In order to efficiently trap and keep water-soluble individual MPs, it was thought that polymers ought to be small

and highly flexible and to remain soluble despite the presence of multiple hydrophobic chains. The hydrophobic chains should be densely spaced (they are statistically ~1 nm apart in A8–35) to avoid the formation of extended hydrophilic loops, which would interfere with many types of biophysical approaches. Chemically, this implied that the hydrophobic chains be relatively short ($C_{18}$ chains were considered unsuitable) and that they be interspersed with highly polar groups. Polar groups are charged groups in most of the APols validated to date (23, 25, 26, 36, 59, 64, 87), hydroxyl groups in early NAPols (70), and glucose moieties in the most recently developed NAPols (5, 6, 71, 83). All the polymers that are efficient as APols self-assemble into small, compact, well-defined
globular assemblies resembling detergent micelles. This behavior is a result of the constraints initially imposed on their chemical structure.

To prepare APols, various synthetic routes can be followed, which include (a) radical copolymerization of hydrophilic and hydrophobic monomers (25, 70, 83); (b) radical polymerization of a suitable monomer (e.g., an acrylic monomer) followed by random chemical modification of the homopolymer thus obtained (6, 23, 59, 87); and (c) homopolymerization of an amphipathic monomer (71).

A8–35 (Figure 1a) is by far the most extensively studied APol to date. Its behavior in aqueous buffers has been examined by size exclusion chromatography (SEC), dynamic and static light scattering, equilibrium and sedimentation velocity analytical ultracentrifugation (AUC), small-angle neutron scattering (SANS), inelastic neutron scattering (INS), and molecular dynamics (MD) (33, 34; F. Giusti, C. Tribet, M. Tehei, G. Zaccaï & J.N. Sachs, unpublished data). From a practical point of view, the most important solution properties of A8–35 can be summarized as follows. At and above pH 7, most carboxylates are ionized in aqueous solutions, conferring A8–35 a high solubility in water (>$200$ g·liter⁻¹) (34). A8–35 molecules, whose average molecular weight varies between 9 and 10 kDa depending on the average length of the polyacrylic acid used for the synthesis, self-assemble into particles averaging ~40 kDa (33). The particles are monodisperse, much more so than the molecules that form them. Individual particles, therefore, comprise a variable number of molecules depending on the latter’s size. On average, A8–35 particles comprise approximately four molecules and 75–80 octyl chains (33).

Investigations of A8–35 and PC-APol C22–43 in water by neutron and light scattering (33) and by isothermal titration calorimetry (26) failed to detect dissociation of the particles down to 0.1 g·liter⁻¹. Down to this concentration, the hydrodynamic radius and molar mass of A8–35 particles are independent of both polymer concentration (33) and chain length polydispersity (C. Tribet &

F. Giusti, unpublished data). Mixtures of fluoroscently labeled A8–35 samples have been studied in an attempt to determine the concentration at which individual chains start associating, which gives rise to a Förster resonance energy transfer (FRET) signal. The critical aggregation concentration thus evaluated is extremely low: ~0.002 g·liter⁻¹ in 100 mM NaCl, pH 8.0 (F. Giusti & C. Tribet, unpublished data).

A8–35 particles bind a maximum of ~1.2 g water per g polymer; their radius of gyration is ~2.4 nm, and their Stokes radius is ~3.15 nm (33). Their operational specific volume ($\phi'$) in 100 mM NaCl is 0.87 mL·g⁻¹ and their contrast match point (the percentage of D₂O at which their neutron scattering length density is the same as that of the solvent and the particles become invisible to neutrons) is 23.5% D₂O (34). The shape of SANS curves indicates that the particles are globular and present a sharp interface with water (33), consistent with the results of recent MD simulations (J.N. Sachs, unpublished data). The latter approach has revealed a few interesting features that had remained inaccessible experimentally, such as the interfering of the APols chains that constitute the particles or the presence of water molecules in their center, probably a result of steric constraints that oppose the formation of a compact hydrophobic core (67; J.N. Sachs, unpublished data). The comparison of INS data collected on unlabeled versus partially deuterated A8–35 reveals backbone dynamics similar to that of CH₂ groups in lipids or free polymers (M. Tehei & G. Zaccaï, personal communication).

Because the aqueous solubility of A8–35 depends on the presence of ionized carboxylates, it is highly sensitive to their protonation, as well as to the presence of multivalent cations, in particular Ca²⁺ ions; MP/A8–35 complexes are similarly affected (19, 25, 33, 34, 64). A8–35 therefore should not be used below pH 7 (preferentially at pH ≥7.5) nor in the presence of millimolar concentrations of Ca²⁺ (Mg²⁺ is better tolerated; 64). Depending on the experiments at hand, a minor degree of aggregation may or may not be acceptable.

Critical aggregation concentration: the concentration of a polymer at which its molecules start to self-assemble
FAPolNBD: A8-35
grafted with NBD

Stability: refers either
to physical (colloidal)
stability, i.e., particles
remain water-soluble
and do not aggregate
nor precipitate, or to
biochemical stability,
i.e., MPs remain in
their native state and,
when measurable,
functional

It can, however, become critical in radiation
scattering experiments, where the presence
of even minute amounts of small oligomers
is highly detrimental (32). In NMR studies,
working above pH 7 accelerates the exchange
of solvent-exposed amide protons.

These experimental constraints have stim-
ulated the development of alternative APols,
such as PC-APols, NAPols, and SAPols
(Figure 1b-d), which are insensitive to low
pH and to the presence of Ca²⁺ ions (23, 25,
64). In aqueous buffers, SAPols and glucosy-
lated NAPols form small, compact, globular,
well-defined particles whose size and dispersity
are comparable to those of the particles formed
by A8-35 (6, 23, 83). As for A8-35, the size of
NAPol particles seems insensitive to the length
of the constitutive polymers (6).

Labeled and Functionalized
Amphipols

The chemistry of APols offers rich opportuni-
ties for labeling and functionalization. To date,
these modifications have been applied only to
A8-35 or its close congener A8-75 (which dif-
fers from A8-35 by the absence of isopropyl
grafts; 87), but they could be readily extended
to other APols. Isotopically labeled APols in-
clude ¹⁴C-labeled A8-75 and ³H-labeled A8-
35, which have been used mainly to follow
the distribution of polymers during fraction-
ation experiments and to quantify their bind-
ing to MPs (32, 68, 88). A form of A8-35
in which the polyacrylic acid chain is hydro-
genated but the isopropyl and octyl groups
are perdeuterated (34) has been heavily used
for SANS, AUC, and NMR experiments (16,
18, 32-34, 99). Four APols functionalized with
a fluorophore have been synthesized, namely
a form of A8-75 carrying a naphtyl group
(94) and three forms of A8-35 carrying a 7-
nitrobenz-2-oxa-1,3-diazol-4-yl (NBD) group
(FAPolNBD) (100), a rhodamine group, or a flu-
orescein one (F. Giusti, unpublished data), re-
spectively. They have been used particularly in
FRET experiments (100) and for observing the
distribution of APols in lipid vesicles (94), in

MP crystals (Figure 6), and in cells and whole
organisms (Figure 8). A biotinylated version
of A8-35 has been used to mediate the im-
mobilization of APol-trapped MPs onto solid
supports (21).

FORMATION, STRUCTURE,
AND PROPERTIES OF
MEMBRANE PROTEIN/
AMPHIPOL COMPLEXES

A vast body of information about MP/APol
complexes is available. In this section, we ex-
amine how these complexes are formed; what
is known of their composition, structure, and
solution behavior; what effects APols have on
the stability and functional properties of the
MPs they complex; which methodological con-
straints they impart; which opportunities they
offer; and how APol-trapped MPs can be trans-
ferred to other environments, be they detergent
micelles or lipid bilayers.

Trapping Membrane Proteins
with Amphipols

Because APols are poorly dissociating surfac-
tants, they are not efficient at solubilizing bi-
ological membranes, even though they parti-
tion into them (19, 65). As a result, APols are
generally unable to directly extract MPs from
cell membranes, even though there are a few
exceptions (68).

Usually, MPs are first extracted and puri-
fied in detergent by standard methods and then
transferred to APols (87). In a typical trapping
experiment, MPs in detergent solution are sup-
plemented with aliquots from a concentrated
stock solution of APols in water. The sample is
briefly incubated, during which time APols mix
with the detergent both in solution and at the
hydrophobic transmembrane surface of MPs
(89, 100). The MP/detergent/APol complexes
thus formed can also incorporate endogenous
lipids that had been solubilized along with MPs,
or lipids added afterward, to increase the pro-
tein's stability and/or activity (see below). It is
possible that, at this stage, lipids that had been
displaced by the detergent rebind (see below). The detergent concentration is then lowered below the CMC, usually by one of the following two methods:

* The solution is diluted under the CMC of the detergent. In this case, the detergent is not physically removed from the solution, but most of it disperses in the aqueous phase as monomers and its concentration in the surfactant layer surrounding the protein drops. This method is sufficient, for instance, to restore native-like allosteric properties to the nicotinic acetylcholine receptor (nAChR), possibly due to lipid rebinding (54). It is also useful to screen trapping conditions, by modulating the MP/APol ratio used and determining by ultracentrifugation, for each condition, which fraction of the protein remains soluble after dilution of the detergent under its CMC (see below).

* Polystyrene beads (Bio-Beads SM2) are added to the sample. Detergent molecules adsorb to the beads, which are removed by low-speed centrifugation (99). The use of Bio-Beads has the advantage of not diluting the protein, and it is usually preferred for preparative protocols. APols (at least A8-35, but this is likely to hold for all APols) do not adsorb significantly to the beads, presumably because their particles do not enter the beads' narrow anfractuosities (98).

Whichever procedure is used, residual detergent molecules, if present, can be removed by dilution/concentration cycles, dialysis, or preparative SEC using surfactant-free buffer (32, 99).

The amount of APols required for efficient trapping is an important parameter that must be established early in any study. The optimal MP/APol ratio depends on the protein of interest and can be determined by carrying out quick trapping tests using a concentration range of APols, dilution below the CMC of the detergent, and ultracentrifugation (25, 83, 87). If the concentration of APols is too low, the protein

aggregates and pellets upon ultracentrifugation [note that with low-CMC detergents such as dodecylmaltoside (DDM), precipitation can be slow and incomplete even in the absence of APols and well below the detergent's CMC]. Upon increasing the APol/protein ratio, a first threshold is reached where most of the protein is kept soluble but MP/APol complexes appear heterogeneous upon SEC or AUC. It is only above a second threshold that MP/APols complexes become homogeneous. Further increasing the APol/MP ratio is useless and can be detrimental: It may increase the viscosity of the sample after concentration (e.g., in solution NMR studies) and compromise the stability of the protein (because it recreates the hydrophobic sink that the use of APols aims to abolish). The minimal mass ratio of A8-35 to MP for a small, deeply membrane-embedded protein such as bacteriorhodopsin (BR) is ~3 g APol per g BR (32). Trapping experiments are usually carried out at a somewhat higher ratio, 5 g APol per g BR. This ratio exceeds by a factor of ~2.5 the amount of APol that actually binds to BR, which is ~2 g APol per g BR (32). The same requirement for a moderate excess of APols is observed with MPs that, like the cytochrome $b_c$1 complex, feature a much higher ratio of mass to transmembrane surface and therefore bind much less APols per g protein (D. Charvolin, unpublished data).

There are at least two rationales to explain this need for an excess of APols over that amount that actually binds to the protein. The first one, hypothetical, is that the protein may select, in a population of heterogeneous APol molecules, those that contribute to form MP/APol complexes with the lowest free-energy level. The second rationale is well documented: Because APols are not strongly dissociating, they do not prevent the formation of small MP oligomers unless present in some excess over that amount actually bound by the proteins (32, 100).

All the ~30 polytopic and bitopic MPs tested thus far, which differ widely in their origin, secondary structure, topology, size, function, and oligomeric state, form water-soluble
complexes with APols (67, 68). This reflects the
fact that the process of association is a physical
phenomenon that does not depend on details of
the protein's transmembrane surface, but sim-
ply on it exposing a large enough hydrophobic
area for APols to adsorb onto. To our knowl-
edge, no attempts have been reported yet to sta-
bilize monotopic or lipid-anchored MPs with
APols.

Composition, Structure, Dynamics,
and Solution Properties of Membrane
Protein/Amphipols Complexes:
Methodological Opportunities
and Constraints

MP/A8-35 complexes have been studied by
SEC, AUC, small-angle X-ray scattering
(SAXS), SANS, FRET, solution NMR, and
INS. Their properties have been reviewed re-
cently (67), and only the most salient facts
are recalled here. Small MPs with limited ex-
tramembrane regions such as BR (27 kDa; seven
transmembrane helices plus, in most experi-
ments, bound lipids) bind proportionately more
A8-35 (~2 g per g protein) (32) than do large
complexes with extended extramembrane do-
mains such as cytochrome *bc*₁ (490 kDa; 22
transmembrane helices; ~0.11 g A8-35 per g
protein) (D. Charvolin, unpublished data). The
amount of A8-35 bound to these two proteins
roughly corresponds to half a dozen molecules
of average molecular weight. Small transmem-
brane domains such as single α-helices can be
expected to bind approximately one particle of
APols, i.e., in the case of A8-35, ~40 kDa. The
A8-35 layer is compact (1.5 to 2 nm thick)
and does not form an extended corona (32).
In SEC, SAXS, SANS, AUC, or NMR exper-
iments, MP/A8-35 complexes indeed behave
like compact, globular particles (67). The only
MP/APol contacts detected to date are with the
transmembrane surface of the protein (18, 99).

Preparing homogeneous MP/A8-35 com-
plexes requires some care (32, 100), and it is dif-
ficult to totally avoid the presence of minor frac-
tions of small oligomers, a nuisance in radiation
scattering experiments (32). Even oligomer-

free preparations do not appear as narrowly
distributed as MP/detergent complexes, unless
some detergent is added (100). A tentative
interpretation is that the large size and small
numbers of adsorbed APol molecules make it
impossible for the system to finely tune the vol-
ume of the adsorbed layer to that corresponding
to the lowest free energy, generating differences
in the exact amount of APol bound from one
MP to the next. Detergent molecules would
provide the small change needed to optimize
the adjustment. Polydispersity is generated
when the complexes are separated from the ex-
tra, free polymer present at the end of a trapping
experiment (32, 100). The latter oligomeriza-
tion is reversible upon adding back APols (100).

Conditions that provoke the aggregation of
the polymer, such as, for A8–35, working at
pH ≤ 7 or adding Ca²⁺ ions, also cause MP/A8–
35 complexes to aggregate (17, 25, 32, 64, 99).
If either of these conditions is essential to the
experiment considered and monodispersity is
required, other APols must be used.

Upon trapping with APols a mixture of
MPs, e.g., the supernatant of solubilized mem-
branes, MPs are trapped under the association
state (e.g., monomers, oligomers, and super-
complexes) that was theirs in the detergent so-
lution, and they retain bound lipids (1, 5, 31,
32, 54, 90). APols do not interfere with most
purification techniques, such as SEC (6, 19,
32, 100), sucrose gradient fractionation (54,
87), or immobilized ligand (24) or metal (100)
affinity chromatography. In the last case, how-
ever, it seems that polyhistidine tags fused too
close to the transmembrane region may interact
less efficiently with the column than in deter-
gent solution, probably because of steric and/or
electrostatic effects (100). Ion-exchange chro-
matography should be restricted to MP/NAPol
complexes, as is isoelectrofocusing.

APols do not interfere with most light
spectroscopy techniques (for a UV-visible ab-
sorbance spectrum of A8-35, see Reference
100), including fluorescence spectroscopy (100;
M. Opačić, unpublished data), circular dichro-
ism (CD) (30, 66, 96), and synchrotron ra-
diation circular dichroism (23; T. Dahmane,
<figure><img src="image_6.png" /><figcaption>Figure 2</figcaption></figure>

Synchrotron radiation circular dichroism (SR-CD) spectra of denatured and A8-35-refolded bacteriorhodopsin (BR). The spectra correspond to the following samples: BR/OTG (octylthioglucoside), native BR (nBR) solubilized from purple membrane (PM) in OTG (along with PM lipids); nBR/A8-35, nBR transferred from OTG to A8-35 (1:5 BR/APol mass ratio); rBR/A8-35, BR refolded in A8-35 from sodium dodecylsulfate (SDS)-solubilized PM; dBO/SDS, bacterio-opsin (BO) delipidated and separated from retinal in organic solvents (dBO) and transferred to SDS; rdBO/A8-35, dBO refolded in A8-35 in the absence of retinal; rdBR/A8-35, dBO refolded in A8-35 in the presence of retinal. SR-CD spectra were recorded at 25°C on the 3m-nim Bessy and DISCO SOLEIL beamlines (29, 74). They extend down to ~175 nm, clearly resolving the 190 nm π-π* transition of the exciton split originating from peptide bond electrons. Data from Reference 23 and T. Dahmane, M. Zoonens & F. Wien, unpublished data.

M. Zoonens & F. Wien, unpublished data) (Figure 2). Attempts have been made to apply Fourier transform infrared spectroscopy to measuring the rate of ¹H/²H exchange at the amide bonds of A8-35-trapped BR. However, all current APols contain amide bonds, and the amide II region of protein spectra (1590–1510 cm⁻¹) is obliterated by APol signals (Y. Gohon & E. Goormaghtigh, unpublished data).

## Functionality of Amphipol-Trapped Membrane Proteins

Functional and/or ligand-binding studies have been carried out with a dozen different

APol-trapped MPs. These include the calcium ATPase (SERCA1a) from the sarcoplasmic reticulum (SR) (19, 64), the nAChR (21, 54), the BR (5, 23, 32, 66), the transporter enzyme II mannitol (EII<sup>mtl</sup>) (M. Opačić, J.-L. Popot & J. Broos, unpublished data) and OmpF (68), both from *Escherichia coli*, as well as half a dozen G-protein-coupled receptors (GPCRs) including rhodopsin (68), the leukotriene receptors 1 and 2 (BLT1 and BLT2), the 5HT<sub>4a</sub> serotonin receptor, the CB1 cannabinoid receptor 1, and the GHSR-1a ghrelin receptor (16, 24 and J.-L. Banères, unpublished data). The data indicate that most proteins are still

SERCA1a:
fast-twitch skeletal
muscle sarcoplasmic
reticulum
Ca²⁺-ATPase
functional after transfer to APols and that the
binding of small ligands, toxins, and antibod-
ies is unimpaired compared to that observed in
biological membranes.

BR, an archaebacterial light-driven proton
pump that accumulates in the so-called pur-
ple membrane of *Halobacterium salinarium*, ac-
complishes its entire photocycle after trapping
in A8-35 (32) or in NAPols (5). Compared
with the kinetics observed in purple membrane,
the first steps of the photocycle are acceler-
ated whether BR is solubilized in detergent or
trapped in APols. The last steps feature simi-
lar kinetics in APol and in purple membrane,
whereas in detergent solution the return to the
fundamental state is slower. The kinetics of the
photocycle is the same whether BR has been
trapped in its native state or refolded in A8-35
from sodium dodecylsulfate (SDS)-solubilized
purple membrane, i.e., in the presence of purple
membrane lipids (66). On the contrary, when
BR is refolded in A8-35 in the absence of lipids,
late kinetics is similar to that in detergent solu-
tion (23). These observations suggest that the
restoration of native-like protein-lipid interac-
tions upon transferring BR from detergent to
APol might be responsible for the recovery of
purple membrane-like kinetics (23).

The allosteric equilibrium between the rest-
ing and desensitized states of the nAChR is
strongly perturbed when receptor-rich postsy-
naptic membranes from the electric organ of
*Torpedo marmorata* are solubilized in detergent
(20). Membrane-like equilibrium and transi-
tion kinetics are recovered when the solubilized
preparation is supplemented with A8-35 and
diluted below the CMC of the detergent (54).
As for BR, one possible explanation is that trans-
fer to APols allows lipids to rebind to critical
allosteric sites at the transmembrane surface of
the receptor.

Recent studies of five GPCRs folded in
APols indicate that their affinity for their
ligands is similar in the membrane and in
APols. However, as previously observed with
rhodopsin (68), G protein activation by A8-
35-trapped BLT1 is significantly slowed down
compared to that observed in lipid/detergent

mixtures. In contrast, when folded in NAPols,
BLT1 catalyzes GDP→GTP exchange on the
Gαi subunit with kinetics similar to those in
lipid/detergent mixtures, suggesting that elec-
trostatic repulsion may account for the slowing
down observed in the presence of A8-35 (J.-L.
Banères, unpublished data).

Diacylglycerolkinase (DAGK) was fully ac-
tive when transferred to APol PMAL B-100
(59). Other, less extensive functional studies
with such MPs as the maltose transporter from
*E. coli* or photosystems I and II are reviewed
in Reference 68. The binding of antibodies by
APol-trapped MPs is discussed below.

A particularly interesting case is that of
SERCA1a, which has been studied in some
detail (19, 64) and has been reviewed previ-
ously (67, 68). In brief, several APols (A8-35,
PMAL C-12, PMALA C-12, and SAPols) both
protected SERCA1a against inactivation and
slowed down the overall turnover (ATP hydrol-
ysis) and individual steps in the catalytic cycle
(release of trapped Ca2⁺) (19, 64). Intermediate
effects are observed in mixtures of APols and
detergent. Trivial mechanisms, such as seques-
tering of Ca2⁺ or interference with ATP bind-
ing, do not account for the inhibition (19). Our
current hypothesis about the origin of this phe-
nomenon relies on the kinetics of reorganiza-
tion of APols around membrane proteins when
the latter undergo conformational changes that
affect their transmembrane surface. Small (sub-
nanometric) conformational changes, such as
those that affect or may affect BR (40) and
the nAChR (22, 39), can probably be accom-
modated by displacements of the APol's alkyl
chains, which are expected to be rapid (sub-
microsecond) and to not involve a high free
energy of activation. Larger interfacial move-
ments (nanometer), such as those undergone by
the transmembrane helix bundle of SERCA1a
upon transiting between the E1 and E2 states
(61), may cause a reorganization of the poly-
mer's backbone, which could entail a higher
free-energy penalty in APol than in detergent
and thereby slow down the enzymatic cycle.
As discussed below, this mechanism (called the
Gulliver effect; 19, 68) may well contribute
to the protective effect of APols against MP denaturation.

## Biochemical Stability of Amphipol-Trapped Membrane Proteins

As a rule, transferring a MP from a detergent solution to APols improves its stability, often dramatically (67, 68). In order to understand the underlying mechanism(s), one should pay attention to the volume of nonmonomeric surfactant to which the protein is exposed to distinguish effects that merely result from a decrease of the hydrophobic sink from those that reflect the interactions of the protein with the surfactant layer that surrounds its transmembrane region.

The effects of trapping a fixed quantity of BR with increasing amounts of APols have been examined in detail (23). Native BR was extracted from purple membrane along with

purple membrane lipids (32). Upon trapping it with A8-35 at BR/APol mass ratios ranging from 1:5 to 1:50, ternary BR/lipids/APol complexes formed. Control samples were stored in either 18-mM or 25-mM octylthioglucoside. Under the experimental conditions used, the volume of the hydrophobic sink was roughly comparable in 18- or 25-mM octylthioglucoside and at BR/A8-35 ratios of 1:10 or 1:20, respectively. Even at 40°C, BR is highly stabilized by A8-35, denaturing by <10% over a week (Figure 3). Under the same conditions, the protein in octylthioglucoside is totally inactivated in less than a day. Whereas a large excess of APols (ratio 1:50) is well tolerated at 4°C and at room temperature, at 40°C it negatively affects the stability of BR, approximately one-third of which is denatured after 6 days (Figure 3). This effect is most likely a consequence of delipidation, which is favored by increasing the volume of the hydrophobic sink (23). Inactivation is nevertheless slow compared

<figure><img src="image_7.png" /><figcaption>Figure 3</figcaption></figure>

Time stability of bacteriorhodopsin (BR) in amphipol A8-35 versus octylthioglucoside (OTG). BR was extracted with OTG from *Halobacterium halobium* purple membrane (PM), along with PM lipids, trapped in A8-35 (32) at various BR/A8-35 mass ratios, and stored in the dark either at room temperature or at 40°C in a buffer containing 100 mM NaCl and 20 mM sodium phosphate, pH 7.0 ([BR] = 0.22 g-liter⁻¹). Its absorbance at 554 nm, which is proportional to the concentration of the holoprotein, was followed as a function of time. Control samples were kept in 18- or 25-mM OTG (total OTG concentration, including bound detergent). These two concentrations correspond to roughly the same mass concentration of free detergent as that of free A8-35 particles in the samples trapped respectively at 1:10 and 1:20 BR/A8-35 mass ratios. The absence of data points in OTG at 40°C past 2 h is due to the aggregation of the protein, accompanied by complete bleaching. Adapted from Reference 23.
to that in octylthioglucoside. Thermostability
is also improved: A8-35-trapped BR stands
well being exposed at 60°C for 20 min, whereas
BR in octylthioglucoside denatures at 40°C
(23). BR/A8-35 preparations at a mass ratio of
1:5 (i.e., with ~3 g free APol per g BR) do not
exhibit any denaturation even after six months
of storage at 4°C (32). BR/A8-35 complexes
can be frozen and thawed without denaturing
the protein (32), but they cannot be lyophilized
(Y. Gohon, unpublished data).

BLT1, one of the receptors of leukotriene B₄ (LTB₄), is significantly stabilized by A8-35. After 25 days at 4°C, no loss of activity is observed whether or not lipids are present. Under the same conditions, BLT1 kept in foscholine-16/asolectin solution loses about half of its activity over the same period. In detergent/lipid mixed micelles, BLT1 denatures at ~27°C, whereas it is stable up to ~35°C when trapped in pure A8-35 and up to ~39°C in the presence of A8-35 plus lipids (24).

Data about the relative ability of various APols to stabilize MPs remain scattered, but they seem to point in at least one direction: The less charges APols bear, the more stabilizing they are. Thus, BR is more stable when trapped in A8-35 than in A8-75, which is a similar polyacrylic acid-derived APol that carries ~75% free carboxylate groups rather than ~35% (87; C. Tribet, unpublished data). Similarly, SERCA1a is more stable in A8-35 than in SAPols, which also carry ~75% of charged groups (64). Cytochrome *b*<sub>6</sub>*f*, a highly detergent-sensitive complex (13), is not very stable in A8-35 (5, 87), but much more so in NAPols (5, 70). The stability of BR in SAPols is much greater in the presence of 100 mM NaCl than in its absence (23). A simple rationale for these effects is that, as may be the case for detergents, electrostatic repulsion between net charges carried by the surfactant bound to a MP tends to favor the formation of particles with a small radius of curvature. This would drive the opening of the protein's structure or, in the case of a multisubunit assembly such as cytochrome *b*<sub>6</sub>*f*, fragmentation.

Little information is available on the stability of APol-trapped β-barrel MPs. On the one hand, A8-35-trapped OmpA, a monomeric protein, denatures at lower urea concentrations than in the detergent lauryldimethylaminoxide, perhaps a consequence of electrostatic repulsion between the protein and the APol at pH 10 (J.H. Kleinschmidt, unpublished observations). On the other hand, the major outer membrane protein (MOMP) from *Chlamydia trachomatis*, a trimeric porin, is considerably thermostabilized (by >30°C) when transferred from Zwittergent 3-14 to A8-35 (M.J. Cocco, unpublished data). Thus, whether APols stabilize β-barrel MPs seems to depend on the protein, the method of denaturation, or both.

The study of SERCA1a has yielded highly valuable information on the mechanisms underlying the protective effects of APols (19, 64). Three points are particularly worth noting. First, the mere addition of A8-35 to a solution of sarcoplasmic reticulum in C₁₂E₈ or DDM markedly improves the stability of the ATPase, even though it increases the volume of the hydrophobic sink, which should drive delipidation (19). This is yet another indication that an APol environment is intrinsically less denaturing than a detergent environment. Second, as discussed above, trapping with APols reversibly inhibits the activity of the ATPase, to an extent that depends on the nature of the APol and on the presence or absence of detergent (19, 64). Third, there is a correlation between the degree of inhibition and the degree of protection against denaturation: The more stabilizing the environment is, the more inhibitory. Thus, SERCA1a surrounded by a mixture of A8-35 and C₁₂E₈ is both partially inhibited and partially stabilized (19), as is SERCA1a trapped in SAPols rather than in A8-35 (64). These observations suggest that the same mechanism may underlie the two phenomena. The high sensitivity of calcium-free SERCA1a to detergents (19 and references therein) can probably be attributed to the detergent prying apart the transmembrane helix bundle (57), which is stabilized by Ca²⁺ ions (86). One may speculate that the Gulliver effect
suggested above slows down both the natural conformational changes that take place during the enzymatic cycle and the nonnatural conformational excursions that open the structure and initiate denaturation, which could explain the correlation between the two effects.

## Transferring MPs from Amphipols to Detergent Solutions or Membranes

APols bind to the hydrophobic surface of MPs in a noncovalent way, but, thanks to their multiple contact points, with an extremely slow dissociation rate. This makes their association with MPs permanent in the absence of a competing surfactant (68, 88, 89, 100), even at extreme dilutions (100). Nevertheless, APols do desorb from MPs in the presence of an excess of competing surfactant, be it free APols (88, 100), detergents (88, 89, 100), or lipids (59, 66).

The displacement of MP-bound APols by free APols was first evidenced upon centrifuging MPs trapped with [¹⁴C]A8-75 through a sucrose gradient containing unlabeled A8-75 (88). Subsequently, the exchange rate was determined by measuring the decrease of the FRET signal between tOmpA (the transmembrane domain of *E. coli* OmpA) and FAPolNBD upon addition of an excess of unlabeled A8-35 (100). At low ionic strength, the kinetics of exchange extends over tens of hours. In the presence of 100 mM NaCl, the exchange is essentially over after 10 min. These observations are consistent with a mechanism involving collisions between MP/APol complexes and free APol particles, probably followed by fusion, mixing, and fission. Such a mechanism is also suggested by many other observations: (a) The concentration of nonassembled A8-35 molecules is extremely low (see above), making it highly improbable that they can mediate the exchange between free and bound APol. (b) Retinal, a hydrophobic molecule that does not partition significantly in water, can be delivered from A8-35 particles to bacterio-opsin (the apoprotein) refolded beforehand in the same APol (23). (c) A8-35-trapped tOmpA reversibly oligomerizes upon elimination of free APol particles (100).

APols and detergents mix rapidly and efficiently (25, 26). Isothermal titration calorimetry measurements indicate that neutral detergents (DDM, octylthioglucoside, C₈E₄) partition in anionic (A8-35) or cationic (C22-43) APols, and their mixing is quasi-ideal (26). This result can seem surprising given that neutral and charged detergents do not mix ideally. Charged detergents are typically prone to form smaller assemblies than neutral detergents (and they facilitate the dispersion and denaturation of proteins) because of the coulombic repulsion between their ionized head groups. In ionic APols, however, the charged groups are covalently linked to one another. Covalent association balances the repulsion between vicinal ionic groups, which cancels the coulombic effects at short length scales.

When MP/FAPolNBD complexes are mixed with a nonionic detergent (DDM, C₁₂E₈) in a stopped-flow instrument, the kinetics of displacement of MP-bound APols, as followed by FRET, is extremely rapid (<1 s) (100). The thermodynamics of the exchange of A8-35 and C22-43 for octylthioglucoside or C₈E₄ has been studied in detail by isothermal titration calorimetry (89). The exchange is isoenthalpic and entropy-driven. Fluorescence and isothermal titration calorimetry data converge toward the conclusion that APols mix with MP/detergent complexes and MP-free detergent micelles nearly ideally, so that at equilibrium the layer of MP-bound surfactant and the free mixed micelles have almost the same composition (89, 100). This is an important observation because it implies that APols have no special affinity for MP transmembrane surfaces. Isothermal titration calorimetry studies also provide insights into the origin of the difference of behavior of MP/APol versus MP/detergent complexes at low surfactant concentration. Upon dilution of a solution of MP/detergent complexes below the CMC of the detergent, entropy favors the aggregation of the protein because it frees hundreds of detergent molecules as monomers. APols, on the contrary, stick to MPs even at extreme dilutions (100), because their desorption entails no

tOmpA:
transmembrane
domain of OmpA from
*Escherichia coli*
Folding: refers to a
MP acquiring the 3D
structure of the native
protein, without
having ever been
correctly folded
previously

significant entropy gain (for a discussion, see
Reference 89).

APols can also be displaced by lipids. Two
studies have described the spontaneous deliv-
ery of APol-trapped MPs to preformed lipid
vesicles and black lipid films. In the first study,
DAGK was delivered from complexes with
the APol OAPA-20 to palmitoyloleoylphos-
phatidylcholine vesicles while retaining its abil-
ity to phosphorylate dibutyrylglycerol (59).
The second study reported the transfer of
two A8-35-trapped β-barrel MPs, OmpA and
FomA, to black lipid membranes (66). After
transfer, the two proteins formed channels with
properties similar to those observed after recon-
stitution from a classical detergent solution.

At concentrations sufficient to deliver MPs,
APols are neither cytolytic nor toxic when they
are applied to living cells or injected into mice
(68, 82; D. Bagnard, G. Crémel, L.M. de la
Maza, unpublished observations). This opens
onto extremely interesting basic and biomed-
ical applications, where APols are used to de-
liver MPs, MP fragments, or other hydropho-
bic molecules to cell membranes.

There are two caveats to APol-mediated de-
ivery of MPs to preexisting membranes. First,
this type of transfer can be expected to be
a high-risk event for the protein, as some of
its hydrophilic extramembrane regions have
to cross the membrane, which necessarily ex-
erts strong distorting forces on the structure.
DAGK, FomA, and OmpA are robust pro-
teins, and in the experiments mentioned above
(59, 66), it is not known which fraction of
them actually adopt a transmembrane orienta-
tion without being denatured. This somewhat
brutal approach is unlikely to result in high
yields of functional insertion with more frag-
ile proteins, unless they refold easily. Second,
the APols with which the protein was initially
trapped can be expected to remain associated
with the target membrane while most likely
diffusing away from the protein. When A8-
35-refolded OmpA and FomA were applied to
black lipid films, the properties of the chan-
nels they formed indeed appeared perturbed
unless A8-35 was also added on the *trans* side

of the film (66). This suggests that adsorbed
APol molecules affect the electrostatics and/or
the internal pressure gradient of the lipid bi-
layer (66). Other studies have shown that APols
by themselves can form pores in natural (19,
82) or artificial (51, 82, 92, 93) membranes. As
shown below, when FAPolNBD-trapped trans-
membrane peptides are delivered to COS
cells, the polymer integrates the plasma mem-
brane along with the peptides (see **Figure 8**).
Vigilance regarding possible artefactual effects
is therefore in order.

APPLICATIONS

The original idea behind the design of APols
was that, by trapping MPs under a water-
soluble form more stable than that in detergent
solutions, APols would make it easier to pu-
rify them and to explore in vitro their structural
and functional properties. As the study of APols
progressed, however, it became apparent that
their applications extended beyond this origi-
nal concept. In the present section, we exam-
ine those applications of APols that have been
tested and either validated or found problem-
atic, along with an assessment of their prospects
and/or limitations.

Folding Membrane Proteins
to their Native State

Overexpressing MPs as inclusion bodies allows
production of large amounts of protein, which
is hard to achieve when directing them to cell
membranes. However, it is difficult to fold to
their native state the inactive MPs obtained in
this way. To date, in vitro folding of MPs in de-
tergents, detergent/lipid mixtures, or lipid vesi-
cles has been achieved for a dozen of β-barrel
MPs (14) and about as many α-helical MPs.
The latter include BR (42, 69); the major light-
harvesting complex II of higher plants (75);
DAGK (37); the small multidrug transporter
from E. coli, EmrE (58); and a few GPCRs
(2–4, 47). Conditions for efficient folding tend
to be highly idiosyncratic.
The feasibility of using APols to assist MP folding was first established using as models two β-barrel MPs, OmpA and FomA, from the eubacteria *E. coli* and *Fusobacterium nucleatum*, respectively, and an α-helical MP, BR (66). Following dilution of urea-unfolded OmpA or FomA into urea-free buffer in the presence of A8–35, both proteins developed β-sheet secondary structure, became protected from proteolysis by trypsin, and recovered the electrophoretic mobility during sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) that is characteristic of the folded β-barrel (66). The folding yields were near 100%. Upon application of the refolded MPs to black lipid films, single-channel recordings showed the formation of functional pores (66). OmpA has also been refolded in NAPols, albeit with much slower kinetics (J.H. Kleinschmidt, unpublished data). tOmpA has been renatured in SAPols, and the recovery of the native structure has been demonstrated by solution NMR (23).

When a solution of denatured BR in SDS solution is supplemented with A8–35 in various mass ratios and dodecyl sulfate precipitated as its potassium salt (PDS), the purple color of native BR starts to develop within minutes. Following removal of PDS crystals by centrifugation and overnight dialysis, UV-visible absorption spectra indicate that BR renaturation is complete at BR/A8–35 wt/wt ratios of either 1:5 or 1:10. The synchrotron radiation circular dichroism spectrum of refolded BR is indistinguishable from that of the native protein (Figure 2) (23). Upon illumination, A8–35-refolded BR undergoes its full photocycle (23, 66). Similar results have been obtained in NAPols (5).

The ability of APols to induce and stabilize the native fold of MPs has been further explored using a series of GPCRs from *E. coli* inclusion bodies purified under denaturing conditions (in SDS solutions). Conditions initially established to refold BR were applied virtually without changes to folding the LTB₄ receptors BLT1 and BLT2, serotonin receptor 5-HT₄a, cannabinoid receptor CB1 (24), and

ghrelin receptor GHSR-1a (J.-L. Banères, unpublished data). Folding yields of 40%–70% were systematically achieved, based on ligand-binding experiments. For all five receptors tested, addition of exogenous lipids (in 1:5 mass ratio to the APol) significantly increased the refolding yields (e.g., from ~50% to ~70% for BLT1). The pharmacological properties of APol-folded receptors, including the $K_d$ values for agonists, are close to those observed in either detergent solutions or membrane fractions. As observed for most APol-trapped MPs, GPCRs folded in A8–35 are significantly more stable than those kept in lipid/detergent mixtures (24). Folding and stabilization of the BLT2 receptor in A8–35 have made it possible to determine by solution NMR the structure of BLT2-bound LTB₄. The BLT1 and GHSR-1a receptors have also been folded in NAPols, with yields similar to those achieved in A8–35 (J.-L. Banères, unpublished data).

## Cell-Free Synthesis of Membrane Proteins

In vivo overexpression of MPs has to navigate the Charybdis of limited expression under a functional form and the Scylla of poor folding yields of mass-produced but inactive protein. An alternative approach that has attracted much attention over the past few years is to express target MPs in vitro, using a cell lysate that contains the machinery for transcribing and translating genes carried by appropriate plasmids (84, 101). Cell-free synthesis presents the advantages of doing away with toxicity issues and of lending itself to straightforward labeling using limited amounts of labeled amino acids (48). MPs, however, will precipitate if synthesized in the absence of any surfactant. Three main strategies have been described: (a) expression in the absence of surfactant, followed by solubilization of the precipitate with a detergent; (b) expression in the presence of a detergent; and (c) expression in the presence of a lipid bilayer, in the form of either lipid vesicles or nanodiscs (15, 44, 46). The third approach

Refolding: refers to MPs that were initially obtained under their native state and have been denatured using urea, SDS, or organic solvents, prior to being brought back to their initial 3D structure
<figure><img src="image_8.png" /><figcaption>Figure 4</figcaption></figure>

Amphipol-assisted cell-free expression of bacteriorhodopsin (BR). In vitro synthesis of polyhistidine-tagged BR in the presence of retinal and of A8-35, SAPols, or NAPols at the indicated concentrations. At the end of the synthesis, the samples were centrifuged at 16,000 × g for 20 min. The proteins present in the pellet (P) and supernatant (S) were separated by SDS-PAGE on a 12% polyacrylamide gel and detected using an anti-His-tag antibody. Far right panel: BR after purification on a nickel column. The purple color of the solution indicates the presence of native BR (representing at least two-thirds of the protein present in the purified sample). Adapted from Reference 5.

suffers from two disadvantages: The amount of protein that can be thus produced tends to be limited by the small volume of lipids that can be made available, and not all MPs insert correctly into preformed bilayers in the absence of the translocon machinery (50). The first two approaches involve the use of detergents. Resuspending precipitated MPs does not always yield a functional protein (49). Expression in the presence of detergent is a priori a hazardous, and certainly not a universal approach: Given that many, if not most, fully folded MPs become unstable upon being exposed to detergents, it is to be expected that many of them will not reach their native state when synthesized in their presence. Indeed, while many MPs can be expressed in this way (81), demonstrations of their functionality are rare (7, 45).

It is therefore of great interest to examine whether less aggressive surfactants are compatible with cell-free synthesis (11, 62, 63, 72). Both A8-35 and SAPols, however, inhibit the synthesis of MPs (**Figure 4**), without affecting that of a control soluble protein, the green fluorescent protein (63). It may be that these APols, both of which are polyanions, inhibit translation by simultaneously binding to MP hydrophobic segments as they appear out of the ribosome tunnel and interacting with positively charged patches at the surface of the ribosome. This hypothesis is supported by the observation that excellent results are obtained when BR is expressed in

vitro in the presence of NAPols: Not only is the yield of synthesis (~0.4 g·liter⁻¹ in the presence of 3 g·liter⁻¹ NAPol) higher and more reproducible (5) than that observed in the presence of DDM (63), but a majority of BR (~90%) is present in a soluble form (**Figure 4**) and remains so over several months (5), whereas in DDM it tends to precipitate (63). BR expressed in the presence of NAPols and retinal folds to a greater extent (at least two-thirds) than in detergent, as shown by the appearance of the purple color characteristic of the holoprotein (**Figure 4**) (5). Preliminary data indicate that NAPol-assisted cell-free synthesis also applies to GPCRs, as exemplified by the leukotriene receptor BLT1 (E. Billon-Denis, F. Zito, J.-L. Banères, unpublished data).

## NMR

Early NMR experiments were aimed at better characterizing MP/APol complexes and at exploring the resources and limitations of NMR to study their structure and dynamics. Three small, well-characterized β-barrel proteins from the outer membrane of either *E. coli* (tOmpA and OmpX) or *Klebsiella pneumoniae* (KpOmpA) were chosen as models. These three proteins, whose 3D structures are known, have been extensively studied by solution NMR, mostly in the presence of detergent. NMR studies of their complexes with

<footnote>KpOmpA: transmembrane domain of OmpA from *Klebsiella pneumoniae*</footnote>
APols showed them to either retain or regain their native fold when associated to A8-35 (17, 99; M. Renault & A. Milon, personal communication), to SAPols (23), or to NAPols (5). The complexes appear slightly bigger than MP/detergent complexes, based on estimates of the overall correlation time, $\tau_c$. For instance, the correlation time of A8-35-trapped OmpX (31 ns) (17) lies between that of OmpX in solutions of dihexanoylphosphatidylcholine (24 ns) (52) and that of OmpX inserted into small isotropic bicelles (35 ns) (53). The slightly larger size of MP/APol complexes compared to those formed with the most favorable detergents does not preclude obtaining high-resolution spectra nor achieving enough sensitivity to perform 3D experiments. Indeed, 3D $^{15}$N-edited [¹H,¹H]-HSQC-NOESY spectra have been collected to explore dipolar interactions between KpOmpA and A8-35 (M. Renault & A. Milon, personal communication). Comparative 2D $^{[15}$N,¹H]-TROSY spectra of BR in detergent solution, in nanodiscs, and in A8-35 have been recently published (73).

A drawback of A8-35 for NMR studies is the need to work at pH >7 in order to avoid aggregation. A high pH renders the observation of exchangeable protons more difficult, if not impossible in some cases, when the chemical exchange becomes too fast (17, 99). This has been one of the incentives leading to the development of pH-insensitive APols, such as SAPols and NAPols, both of which make it possible to record NMR spectra under acidic conditions (5, 23). These new APols, however, suffer from other disadvantages: The purification of SAPols is demanding, which currently limits the amounts that can be conveniently produced (23), and NAPols, due to their complex chemical structure (Figure 1c), cannot be easily perdeuterated.

NMR spectroscopy has been used to examine the organization of MP/A8-35 complexes, yielding an increasingly detailed view of protein-polymer interactions. A general description of the distribution of the alkyl chains was first obtained, taking advantage of the variation of dipolar environment experienced

by the amide protons of tOmpA upon trapping with either A8-35 or its deuterated homolog (99). Two further studies exploited intermolecular ¹H-¹H or ¹³C-¹H dipole-to-dipole cross-relaxation phenomena. They yielded a qualitative detection of spatial proximities between ¹³C atoms in the side chain of aromatic residues of OmpX and ¹H nuclei of A8-35 (18) and the identification of hydrophobic contacts between specific amide protons of KpOmpA and octyl and isopropyl chains of A8-35 (M. Renault & A. Milon, personal communication). The three studies showed that contacts with the alkyl chains of A8-35 are confined to the transmembrane, hydrophobic surface of the proteins. ¹H/²H exchange measurements show that some of the amide protons of the membrane-spanning region of OmpX exchange much more readily than others, which likely reflects the dynamics of the barrel (17).

Application of solution NMR to GPCRs is rare, mainly due to the difficulty of producing sufficient amounts of functional receptors (10 and references therein). APol-assisted folding of BLT2 has made it possible to solve by NMR the structure of its natural ligand, LTB₄, in its receptor-bound state. Perdeuterated BLT2 receptor was overexpressed in *E. coli* as inclusion bodies and folded to its native state (24) in deuterated A8-35. The high-resolution structure of the BLT2-bound leukotriene was derived from transferred nuclear Overhauser effect signals (16). The ligand undergoes a drastic reorganization upon binding, switching from an elongated structure with many coexisting rotamers to a highly constrained conformation (Figure 5). This information is of great interest from a pharmacological point of view, because it opens new perspectives in BLT-targeted drug design (no antagonists of this LTB₄-mediated proinflammatory pathway are available today for clinical use). It also contributes to a detailed analysis of the molecular events that lead to receptor activation. Because the ability of APols to fold and stabilize MPs seems quite general, the same strategy can probably be transposed to many other systems.
<figure><img src="image_9.png" /><figcaption>b</figcaption></figure>

<figure><img src="image_10.png" /><figcaption>c</figcaption></figure>

<figure><img src="image_11.png" /></figure>
Immobilizing Membrane Proteins
onto Solid Supports for
Ligand-Binding Studies

The development of sensitive detection methods and the implementation of high-throughput technologies increasingly resort to the immobilization of functional proteins onto surfaces. On the one hand, protein arrays combined with optical detection provide an efficient mode of high-throughput screening, with a particular potential in diagnostics (95). On the other hand, surface-sensitive techniques such as surface plasmon resonance (97) and fluorescence microscopy provide information regarding the kinetics of interaction, in addition to the classical thermodynamic parameters (91).

Because of the poor stability of purified MPs in detergent solutions, many current studies are carried out in native (41, 55) or reconstituted lipid membranes (9, 78), which entails a series of technical constraints: nonspecific binding to other proteins and/or lipids, the difficulty of working out and standardizing protocols for MP reconstitution and for the immobilization of membrane fragments or vesicles, and the impossibility to access simultaneously the extracellular and intracellular domains of MPs.

Reversible or irreversible affinity tags are
generally used to interface MPs with surfaces
without disturbing the protein's function. Tags
are typically added by fusing to the target MP a
short peptide or a cargo-protein (38, 85). This
approach is not applicable to proteins extracted
from native tissues. In that case, the protein
is chemically modified, usually on NH₂ or SH
groups.

APols provide a novel, highly versatile way
to immobilize MPs. Its advantages over the
above approaches are many:

* Trapping with functionalized APols does not alter the structure of the protein, and it leaves its N and C termini free for protein-ligand interactions or for other modifications.

* The protein can be trapped with a func-
tionalized APol after its purification, dur-
ing folding, or during cell-free synthesis.

* Although the MP/APol association is highly stable, it can be displaced by dergents or other APols. Affinity tags can thus be added or removed as desired.

* Because MPs are typically surrounded by at least five to six molecules of APols, several functional groups can be combined in a controlled stoichiometry, by the simple device of trapping the protein with a mix of appropriately functionalized APols.

* The MP benefits from the stabilization afforded by APols.

* The MP can (and, as a rule, must) be handled in surfactant-free solutions, which facilitates the handling of fluidics and detection using surface-sensitive techniques such as surface plasmon resonance.

It should be noted, however, that the loca-
tion of the functional groups in the APol belt
cannot be controlled and vary from one protein
to the next, as well as over time, which, depend-
ing on the experiment, can be considered either
a liability or an asset.

Whereas several APols carrying affinity tags
have been or are being synthesized, biotiny-
lated A8-35 (BAPol) is the best characterized to

BAPol: biotinylated
A8-35

<caption>Figure 5</caption>

Amphipol-assisted folding and stabilization of a G-protein-coupled receptor allow NMR determination of the structure of a receptor-bound ligand. The perdeuterated BLT2 receptor of proinflammatory leukotriene B4 (LTB4) was expressed as inclusion bodies in *Escherichia coli*, solubilized in SDS, and folded in deuterated A8–35 (DAPol). The structure of BLT2-bound LTB4 was determined by analyzing transferred nuclear Overhauser effect signals. (a) Chemical structure of LTB4. (b) Dipolar interactions in an LTB4/[u-2H,15N]BLT2/DAPol sample, 2D NOESY spectrum (mixing time τm = 0.5 s, vH = 600 MHz, 25°C). The corresponding 1D 1H spectrum is shown above the 2D spectrum. The 1D spectrum of free LTB4 in solution is displayed on the left side. Numbers refer to the protons annotated on the chemical structure of LTB4 in panel a. (c) Three-dimensional structure of LTB4 bound to BLT2 (hydrogen atoms are light gray, oxygen atoms are red; and carbon atoms are assigned a different color for each conformer). Adapted with permission from Reference 16. Copyright 2010 American Chemical Society.
date. As described above, trapping with APols as a rule does not interfere with ligand binding. The immobilization of BAPol-trapped MPs of various sources and sizes to streptavidin-coated chips or beads does not impair their function, e.g., the binding of a toxin and of a small acetylcholine analog to the nAChR, or the photocycle of BR, nor does it prevent the specific recognition of antibodies by tOmpA, BR, and cytochromes *bc*₁ and *b*₅f (21). The binding of antibodies and/or toxins to APol-trapped MPs has also been observed for OmpF (68) and MOMP (M.J. Cocco & L.M. de la Maza, unpublished data). APol-mediated MP immobilization thus appears as a promising system for diagnostic assays, as well as for functional investigations of ligand and protein interactions with MPs, with potential applications in drug screening. Prospects include the use of more sophisticated supports (electrodes, nanoparticles, and other nano-objects) and of other tagged APols. Histidine-tagged (F. Giusti & P. Kessler, unpublished data) and oligonucleotide-carrying APols (C. Le Bon & F. Giusti, unpublished data), for instance, would provide for the reversibility of the immobilization and, in the latter case, targeting, with a strong potential for the development of protein arrays. Tagged NAPols would reduce the probability of electrostatically driven nonspecific binding.

## X-Ray Crystallography

The application of APols to MP crystallography would provide fascinating perspectives. This goal, however, may seem a priori hard to reach, because MP/APol complexes appear more polydisperse than MP/detergent complexes. In the case of charged APols, this difficulty is compounded by the need to achieve the right balance between minimizing the electrostatic repulsion between APol layers and preventing aggregation. Over the past few years, however, considerable progress has been made toward identifying factors that favor the formation of monodisperse preparations of MP/APol particles. Furthermore, repulsive interactions between MP/A8-35 complexes can be cancelled

or even turned to attractive interactions by appropriately modulating the ionic strength of the solutions (68).

Current attempts have focused on a model MP, mitochondrial cytochrome *bc*₁, which combines the advantages of being sturdy, colored, relatively easy to crystallize, and of featuring large extramembrane domains that provide anchoring points for the formation of well-ordered 3D contacts while keeping protein-bound surfactant layers away from each other (8). A8-35 was added to purified, detergent-solubilized cytochrome *bc*₁, and the detergent adsorbed onto polystyrene beads. Crystallization trials were carried out by vapor diffusion. Under the many crystallization conditions tested, complete elimination of the detergent precluded any crystals from forming. On the contrary, ternary cytochrome *bc*₁/A8-35/detergent complexes did crystallize, in the presence of concentrations of salt and polyethylene glycol higher than is required for crystallization from detergent solution (D. Charvolin, M. Picard, E.A. Berry & L.-S. Huang, unpublished data). The red color of the crystals indicated that they do contain cytochrome *bc*₁ (Figure 6a). The presence of APols in the crystals was established by growing them in the presence of FAPolNBD (Figure 6b). A few of these crystals have been tested on a synchrotron X-ray source and proved to diffract, with unit cell dimensions consistent with their being protein crystals. The current resolution of the patterns, however, is poor (>20 Å) (Figure 6c). It is not known whether this reflects an intrinsic property of the crystals, arises from problems with handling and/or freezing them, or is due simply to the limited number of trials that could be carried out. There are several possible reasons why cytochrome *bc*₁/A8-35/detergent complexes crystallize whereas *bc*₁/A8-35 ones do not. One of them is that the presence of detergent lowers the electrostatic charge density of the surfactant layer, and it may allow for easier charge redistribution. Another is that, as observed with tOmpA (100), ternary complexes may be more closely monodisperse than binary ones.
<figure><img src="image_12.png" /><figcaption>Figure 6</figcaption></figure>

<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>

Crystals of ternary cytochrome *bc*1/A8–35/detergent complexes. (a) Crystals obtained in the presence of A8–35, observed by visible light microscopy. (b) Crystals obtained in the presence of an NBD-labeled analog of A8–35, observed by fluorescence microscopy. (c) Diffraction pattern (collected at the ESRF facility in Grenoble) of one of the crystals shown in panel b. (From D. Charvolin, M. Picard, E.A. Berry and L.-S. Huang, unpublished data). Abbreviation: NBD, 7-nitrobenz-2-oxa-1,3-diazol-4-yl.

Although these experiments were inter-
rupted, they show that crystallizing APol-
trapped MPs is indeed feasible. The advent
of NAPols opens new, exciting perspectives in
this field, because their use would eliminate the
electrostatic repulsion that is, presumably, the
major handicap in using charged APols such as
A8–35.

Electron Microscopy

APol-trapped MPs have been observed by
scanning transmission electron microscopy
(EM) (90), by transmission EM after negative
staining (1, 28, 32, 67 and references therein)
and by cryo-EM (1, 28). Because APols are
mild surfactants, they may be particularly well
suited to single-particle EM studies of fragile
MP supercomplexes. Supercomplexes of the
mitochondrial respiratory chain are stable and
soluble in A8–35 after removal of the digitonin
used for solubilization, and present NADH-
dehydrogenase and cytochrome c oxidase
activity after purification (1). Images of super-
complex B (consisting of one copy of complex I,
two of complex III, and one of complex IV)
adsorbed onto a continuous carbon support
film were recorded under cryo-conditions

(Figures 7a,b). The particles exhibited similar
views as in digitonin (80). A 3D volume
(Figure 7c) was calculated by the random
conical tilt method, revealing many details

<figure><img src="image_15.png" /><figcaption>Figure 7</figcaption></figure>

Cryo-electron microscopy (cryo-EM) reconstruction of mitochondrial supercomplex B (I₁III₂IV₁) trapped in APol A8–35. (a,b) Typical 2D projections from cryo-EM representing side (a) and top (b) views. (c) 3D reconstruction as seen slightly tilted from the matrix side. The red fringe indicates the likely distribution of A8–35. See Reference 1.
not previously visible. In keeping with NMR data, A8-35 seems to form a belt along the transmembrane region of the supercomplex (Figure 7c) (1).

## Proteomics: 2D Gels and Mass Spectrometry

2D gel electrophoresis is a standard technique in proteomics. It is used, in particular, to study protein expression in cells. Single spots can be picked out from the gels and mass spectrometry performed to determine the mass of each protein and collect sequence information. In the first dimension, proteins are usually separated according to their pI by isoelectrofocusing in a pH gradient. APols that carry a net charge cannot be used for isoelectrofocusing, but NAPols can. The feasibility of this approach has been tested using NAPols whose solubility relies on the presence of either hydroxyl groups (70) or glucose moieties (71). In the second dimension, APols are expected to be displaced by SDS. [APols indeed do not affect SDS-PAGE patterns (59, 66, 90).] In brief, the results show that these two types of NAPols can indeed be used to run 2D gels (5; Y. Gohon, unpublished data) and the spots excised and subjected to mass spectrometry analysis (Y. Gohon, P. Bazzacco, E. Billon-Denis, C. Béchara, G. Bolbach & S. Sagan, unpublished data).

Matrix-assisted laser desorption ionization (MALDI)-time of flight (TOF) measurements have been performed on several APol-trapped MPs, namely OmpX (18), tOmpA, BR, and the cytochrome *bc*₁ and *b*₆*f* complexes (P. Bazzacco, C. Béchara, G. Bolbach & S. Sagan, unpublished data). Neither NAPols nor ionic APols are detected, probably due to their high polydispersity. No significant differences are observed between MP/APols and MP/detergent complexes. Thus, mass spectrometry could be used to check on the extent of isotopic labeling of A8-35-trapped OmpX (18). In preparations of A8-35-trapped and NAPol-trapped cytochrome *b*₆*f*, all but two of the eight subunits were detected, albeit with variable yields, as well as free heme and, for NAPol-trapped com-

plexes, some lipids (P. Bazzacco, C. Béchara, G. Bolbach & S. Sagan, unpublished data).

Altogether, these results indicate that A8-35 and NAPols are compatible with MALDI-TOF mass spectrometry analyses of both intact MPs and of their digests, and that NAPols are suitable to perform isoelectrofocusing and 2D gel electrophoresis. The stabilizing properties of APols ought to facilitate analyses of fragile MPs and MP complexes by both of these techniques.

## Amphipols as Vectors for the Delivery of Membrane Proteins and Transmembrane Peptides

Upon injection of MP/A8-35 complexes into mice, no toxicity is observed. Antibodies are produced against MPs, not against APols (68). In cell cultures, A8-35 is not cytolytic, at least at the concentrations needed to deliver MPs to the plasma membrane (68). This has led to the examination of APols as vectors for immunization or other therapeutic purposes.

The transmembrane domain of bitopic receptors, which is made up of a single hydrophobic α-helix, plays an important role in their oligomerization (43, 56). Recent evidence indicates that peptides mimicking the trans-membrane domain of neuropilin-1 (NRP1), a receptor controlling various biological effects ranging from cell migration to cell proliferation and cell death (76), are able to block its biological functions (77). In addition to helping to dissect signaling mechanisms, these peptides also present a therapeutic relevance in cancer. Preclinical studies have demonstrated that blocking NRP1 using transmembrane peptides in tumor cells or in the endothelial cells establishing the neovascular network supporting tumor growth largely reduces brain tumor expansion and dissemination both in vitro and in vivo (60). However, NRP1 trans-membrane peptides are insoluble and prone to aggregation, which makes it difficult to ensure optimal distribution and efficacy in whole organisms. APols keep various hydrophobic peptides soluble (27, 30, 96), suggesting
that they could be used as solubilizing and vectorizing agents. An analysis has been undertaken of the biodistribution of APol-trapped transmembrane peptides applied to COS cells (Figure 8a). Confocal microscopy showed that the complexes can reach and enter the plasma membrane within minutes. Both peptides and APols are endocytosed after a few hours, leading to a total clearance of the plasma membrane in four days. To examine whether APols could be potentially useful delivery agents in vivo, mice bearing subcutaneous tumors were injected with FAPolNBD. Histological examination of tumor slices revealed strong staining of intratumor blood vessels (Figure 8b) (G. Crémel & D. Bagnard, unpublished data).

Another potentially promising therapeutic use of APols is the formulation of vaccines. Even though MPs from pathogenic organisms represent a priori privileged targets for the immune system, their production and handling pose formidable obstacles in the way of manufacturing affordable vaccines. The ability of APols to fold overexpressed MPs and to stabilize them may help to circumvent these difficulties. Recent data indicate that vaccines formulated with MOMP/A8-35 complexes protect mice against *C. trachomatis* infection better than vaccines containing MOMP/detergent complexes, to a level approaching that observed upon vaccination with the live organism (L.M. de la Maza, unpublished data). It remains to be established whether this improvement results from the strong stabilization of MOMP by A8-35, from a different presentation of the immunogen to cells of the immune system, or from a combination of both mechanisms. However, given how general MP stabilization by APols is, it is probably worth examining the possibility of extending this approach to MP-based vaccines against other infectious diseases, possibly also against some forms of cancer. The development of therapeutic uses of APols is still at a preliminary stage. However, the above data suggest that APols may provide an interesting alternative to the use of detergents, liposomes, and nanoparticles whenever organisms are to be injected with MPs or transmembrane fragments thereof.

<figure><img src="image_16.png" /><figcaption>Figure 8</figcaption></figure>

Using amphipols to deliver transmembrane peptides to cells and tissues. (a) Confocal microscopic analysis of the distribution of NBD-labeled A8-35 (FAPolNBD; green) and a rhodamine-labeled neuropilin-1 transmembrane peptide (red) 16 and 72 h after application of peptide/FAPolNBD complexes to COS-7 cells. Note that the distribution of the two labels does not totally overlap, indicating at least partial segregation. After 72 h, the plasma membrane has been cleared of both labels. (b) Distribution of FAPolNBD in tumor slices. Mice bearing subcutaneous tumors (C6 glioma) were injected intraperitoneally with 100 µL of 10⁻⁷ M FAPolNBD 24 h before tissue collection. Vascular and perivascular staining demonstrate tissue penetration (yellow arrow). Unpublished data from G. Crémel & D. Bagnard. Abbreviation: NBD, 7-nitrobenz-2-oxa-1,3-diazol-4-yl.

CONCLUSION

Much is now known about what can and what cannot be advantageously done by substituting detergents with APols. Some properties of MP/APol complexes, e.g., the dynamics of
APol-trapped MPs, remain to be more fully explored. Many applications have been validated but not fully exploited, whereas the difficulties met by other applications, e.g., 3D crystallization, remain to be solved. Many more applications will appear and develop as novel APols become better characterized (nonionic APols in particular), as the variety of functions grafted

onto APols increases, and as more laboratories familiarize themselves with their use. It is our hope that the work invested by so many people over so many years to turn APols into practical, reliable tools will provide the community of MP specialists with enough solid data and incentives so that their use becomes more widespread.

## SUMMARY POINTS

1. APols are a special class of amphipathic polymers designed to keep integral MPs water soluble in the absence of detergents.

2. Various types of APols have been tested and validated. They have in common the ability to assemble in aqueous solutions in the form of small, globular, well-defined particles.

3. APols make MPs water soluble by adsorbing onto their hydrophobic transmembrane surface. The APol layer is thin and remains stable even at high dilutions.

4. MP-adsorbed APols can be readily exchanged for detergents, lipids, or other APols.

5. Most MPs are much more stable as MP/APol complexes than they are in detergent solutions. Several mechanisms contribute to this effect.

6. APols, as a rule, do not interfere with ligand binding or with MP function. They may, however, slow down some transconformations or interactions.

7. APols are an excellent medium in which to fold MPs to their functional state, and some APols can be used for MP cell-free synthesis.

8. APols lend themselves to many applications, including structural studies, proteomics, MP immobilization, and delivery.

## DISCLOSURE STATEMENT

P. Bazzacco, D. Charvolin, F. Giusti, B. Pucci, C. Tribet, and J.-L. Popot are coauthors of patents on amphipols and their applications.

## ACKNOWLEDGMENTS

This review is dedicated to the memory of two colleagues without whom this work would likely not have existed: Charles Tanford (1921–2009), whose role in introducing membrane biologists to the physical chemistry of surfactants can hardly be overstated, and Roland Audebert (1939–1997), colloid and polymer chemist, with whom C.T. and J.-L.P. launched this project in 1994. Particular thanks are due to C. Béchara, E.A. Berry, G. Bolbach, J. Broos, L.-S. Huang, P. Kessler, A. Milon, M. Opačić, F. Rappaport, M. Renault, S. Sagan, M. Tehei, F. Winnik, and G. Zaccaï for permission to cite ongoing work done in collaboration with them and/or for comments on the manuscript, and to J. Barra for her invaluable help for the preparation of the figures.

Affiliations for the coauthors of this manuscript are as follows:

P. Bazzacco, E. Billon-Denis, L.J. Catoire, D. Charvolin, F. Giusti, C. Le Bon, F. Zito, M. Zoonens: Institut de Biologie Physico-Chimique, CNRS/Université Paris-7 UMR 7099, F-75005 Paris, France
T. Althoff, W. Kühlbrandt: Max-Planck-Institut für Biophysik, D-60438 Frankfurt am Main, Germany

D. Bagnard: Université de Strasbourg, INSERM U682, F-67084 Strasbourg, France

J.-L. Banères: Institut des Biomolécules Max Mousseron, CNRS/Universités Montpellier I & II UMR 5247, F-34093 Montpellier, France

P. Champeil: Institut de Biologie et de Technologies de Saclay, CEA/SB2SM and CNRS/URA 2096, F-91191 Gif-sur-Yvette Cedex, France

M.J. Cocco: Department of Molecular Biology and Biochemistry, University of California, Irvine, California 92697-3900

G. Crémel: Centre de Neurochimie, INSERM U682, F-67084 Strasbourg, France

T. Dahmane: Institut de Biologie Physico-Chimique, CNRS/Université Paris-7 UMR 7099, F-75005 Paris, France; Present address: Biochemistry and Molecular Biophysics, Columbia University, New York, New York 10032

L.M. de la Maza: Department of Pathology and Laboratory Medicine Medical Sciences I, University of California, Irvine, California 92697-4800

C. Ebel, F. Gabel: Institut de Biologie Structurale, CEA/CNRS/UJF UMR5075, F-38027 Grenoble, France

Y. Gohon: Present address: Institut de Biologie Physico-Chimique, CNRS/Université Paris-7 UMR 7099, F-75005 Paris, France; Unité de Chimie Biologique, AgroParisTech/INRA, F-78850 Thiverval-Grignon, France

E. Goormaghtigh: Center for Structural Biology and Bioinformatics, Université Libre de Bruxelles, B-10050 Bruxelles, Belgium

E. Guittet, C. van Heijenoort: Institut de Chimie des Substances Naturelles, CNRS UPR 2301, F-91198 Gif-sur-Yvette, France

J.H. Kleinschmidt: Fachbereich Biologie, Universität Konstanz, D-78457 Konstanz, Germany

K.L. Martinez: Department of Neuroscience and Pharmacology & Nanoscience Center, University of Copenhagen, DK-2100 Copenhagen, Denmark

M. Picard: Institut de Biologie Physico-Chimique, CNRS/Université Paris-7 UMR 7099, F-75005 Paris, France; Present address: Laboratoire de Cristallographie et RMN Biologiques, CNRS/Université Paris Descartes UMR 8015, F-75006 Paris

B. Pucci: Laboratoire de Chimie Bioorganique et des Systèmes Moléculaires Vectoriels, Université d'Avignon et des Pays de Vaucluse, F-84000 Avignon, France

J.N. Sachs: Department of Biomedical Engineering, University of Minnesota, Minneapolis, Minnesota 55455

C. Tribet: Département de Chimie, CNRS/ENS/Université Paris-6 UMR 8640, Ecole Normale Supérieure, F-75005 Paris

F. Wien: DISCO, Synchrotron Soleil, F-91192 Gif-sur-Yvette, France

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87.
Validates the
amphipol concept by
using A8-35 and three
of its congeners varying
in length and charge
density.

89. Provides a
thermodynamic
investigation of the
interactions of
detergents and APols
with the TM surface of
MPs.
99. Shows that MP/APol contacts are restricted to the TM surface of the protein.

100. Shows that APols bind tightly to MPs and yet are in dynamic equilibrium with the aqueous solution.

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100. Zoonens M, Giusti F, Zito F, Popot J-L. 2007. Dynamics of membrane protein/amphipol association studied by Förster resonance energy transfer. Implications for in vitro studies of amphipol-stabilized membrane proteins. *Biochemistry* 46:10392–404

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