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MINI-REVIEW
Single molecule techniques for the study
of membrane proteins
Ana J. García-Sáez & Petra Schwille
Received: 2 February 2007 / Revised: 17 April 2007 / Accepted: 17 April 2007 / Published online: 12 May 2007
#
Springer-Verlag 2007
Abstract Single molecule techniques promise novel infor-
mation about the properties and behavior of individual
particles, thus enabling access to molecular heter ogeneities
in biological systems. Their recent developments to accom-
modate membrane studies have significantly deepened the
understanding of membrane proteins. In this short review, we
will describe the basics of the three most common single-
molecule techniques used on membrane proteins: fluores-
cence correlation spectroscopy, single particle tracking, and
atomic force microscopy. We will discuss the most relevant
findings made during the recent years and their contribution
to the membrane protein field.
Keywords Membrane protein
.
Single molecule technique
.
Atomic force microscopy
.
Fluorescence correlation spectroscopy
.
Single particle tracking
.
Single molecule microscopy
Introduction
Cell membranes are noncovalent, supramolecular assemblies
of lipids and proteins that form self-contained volumes. They
define cellular compartments by acting as permeability
barriers to polar molecules. But besides this structural
function, cell membranes are involved in the transport of
energy, matter, and information into and out of the cell.
Such a diversity of functions is reflected by the complex
protein and lipid composition of natural membranes. Lipids
organize into bilayers that contain integral and associated
membrane proteins. According to genomic data, half of a
cell’s genes code for membrane proteins, which are
responsible for around one third of the dry cell-weight
(Douglass and Vale 2005; Zimmerberg and Gawrisch 2006).
This makes cellular membranes crowded environments.
For the lipids, the combination of different backbones,
head groups, acyl chains, and degree of saturation can give
rise to hundreds of different species. This allows fine-
tuning of the physical and chemical properties of mem-
branes, including lateral fluidity, permeability, rigidity, and
curvature. Indeed, this diversity is fully exploited by the
cell: Mass -spectrometry analys es of lipid composition in
cells revealed high levels of temporal and spatial variability
of membrane composition (van Meer 2005).
Besides controlling the majority of membrane properties,
lipids can affect membrane protein function through
specific interactions (Lin et al. 2005). And on the other
hand, membrane proteins can alter lipid organization and
modify the physical properties of mem branes (Douglass
and Vale 2005).
Information about how membrane proteins work, their
interactions wi th lipids and other proteins, and their
dynamics and structure, is essential for the understanding
of biological membranes and cell functioning in general.
However, the difficulties in reconstituting functional mem-
brane proteins to controllable model bilayers have delaye d
their structural and functional characterization in compari-
son to their soluble counterparts.
Recently, a number of single-molecule techniques have
been extended for the study of membrane proteins. The
applications performed to date have contributed substan-
tially to the current knowledge of membrane proteins and
membrane organization.
Appl Microbiol Biotechnol (2007) 76:257–266
DOI 10.1007/s00253-007-1007-8
A. J. García-Sáez
:
P. Schwille (*)
Biophysics Group,
Biotechnologisches Zentrum (BIOTEC) der TU Dresden,
Tatzberg 47-51, 01307 Dresden, Germany
e-mail: petra.schwille@biotec.tu-dresden.de
Biological systems are intrinsically heterogeneous, im-
plying distributions of dynamic coefficients, structural
conformations, interactions among molecules, and catalytic
rates. The main advantage of single-molecule techniques vs
averaging techniques is that individual inhomogeneities in
the system of study can be assessed (Haustein and Schwille
2004). The analysis of these distributions can provide a
deeper insight into the mechanism of action of biolo gical
molecules. For example, it can be distinguished if changes
in the measured parameter are observed for all mol ecules in
the system or only a subpopulation of them (Johnson et al.
2005). In addition, single-molecule techniques usually need
low concentrations similar to physiological conditions and
are normally applied to systems in equilibrium.
In this review, we will focus on the recent progress in the
membrane–protein field achieved with single-molecule tech-
niques. We will describe the principles of the three main single-
molecule techniques that have been applied to membrane
proteins: fluorescence correlation spectroscopy (FCS), single-
particle tracking (SPT), and atomic force microscopy/spectros-
copy (AFM), and we will discuss the main findings and their
contribution to the understanding of membrane proteins.
Fluorescence correlation spectroscopy
Fluorescence correlation spectroscopy characterizes the
fluctuations in fluorescence intensity of a system in
equilibrium (Haustein and Schwille 2004). A scheme of a
typical setup is shown in Fig. 1a. It usually contains a
microscope objective with high numerical aperture to focus
the laser and achieve efficient detection. The photons
emitted within the focal volume pass through a pinhole,
which reduces the detection volume in the z-direction. As a
result, the measurement volume is confined to dimensions in
the range of fractions of a femtoliter, which is a micrometer
cube. Then, the photons pass proper filters and are detected
by an avalanche photodiode. This configuration allows high
signal/noise ratio when using fluorophore concentrations
ranging from 10
−10
to 10
−6
M.
The diffusion of fluorophore molecules through the focal
volume or changes in their emission properties [due to
chemical reactions, association/dissociation events, photo-
dynamic processes, or conformational changes (Haustein
and Schwille 2004; Kahya 2006)] produce fluctuations in
fluorescence intensity that can be quantified and autocorre-
lated as a function of time (Fig. 1b). The autocorrel ation
function is a mathematical tool that provides a measure of
the self-similarity of the fluorescence signal over time. The
characteristic decay time of the autocorrelation function is
related to the time that the fluorophore spends within the
focal volume. It depends on the mobility of the particle:
The larger the diffusion coefficient, the faster the decay
(Fig. 1c).
FCS averages thousands of single-diffusion events,
allowing precise estimation of diffusion coefficients and
particle concentrations in relatively short acquisition times
(Schwille 2001). It thus can be used to inves tigate not only
molecule mobility, transport, and diffusion processes but
also rate constants of inter- or intramolecular reactions or
binding (Bacia et al. 2006).
As with other fluorescence techniques, the protein of
interest must be fluorescently labeled. Fusion to fluorescent
proteins and chemical labeling with organic dyes are the
most common strategies. Care must be taken that labeling
does not affect the protein behavior.
Fig. 1 Principles of fluorescence correlation spectroscopy. a The
sample is illuminated by focusing the laser beam with an objective.
The emitted photons are then spectrally filtered and detected with an
avalanche phot odiode. A pinhole before the detector limits the
detection volume in the z-direction, so that the light from upper and
lower planes is eliminated. b The emission of photons diffusing in and
out the focal volume induces fluctuations in the intensity signal. The
fluorescence trace recorded during a given peri od of time is
autocorrelated. The autocorrelation function measures the self-
similarity of the fluorescence signal with time. c The graph shows
the values obtained for the autocorrelated fluorescence signal in b
(white circles). The diffusion coefficient and the particle concentration
can be determined by fitting the experimental values to the
autocorrelation function in b (solid line) (courtesy of J. Ries)
258 Appl Microbiol Biotechnol (2007) 76:257–266
In the case of membrane measurements, some additional
considerations are required (Kahya 2006). It is critical to
correctly position the laser focus on the membrane bilayer
to avoid artifacts during the collection of autocorrelation
curves. For this purpose, it is recommended to combine the
FCS setup with an imaging device (preferably a laser scanning
microscope) and a z-stepping motor that allows measurement
of the diffusion times at different z-positions of the bilayer
(Benda et al. 2003). The measured membrane should be
planar in the focal plane, and fluorescence fluctuations due
to undulations or movement of the whole membrane should
be avoided. The analysis of aggregation processes is
restricted by sensitivity, as for freely diffusing molecules in
the two-dimensional plane of the membrane, the mass of the
monomer/oligomer should differ at least by a factor of 15–20
(Kahya 2006).
Giant unilamellar vesicles (GUVs) are an excellent
model membrane system for FCS studies: They have
similar size to eukaryotic cells, so they can be visualized
with a microscope, and the membrane area focused in the
detection volume has virtually no curvature. In addition,
they form stable structures composed of freestanding bilayers
whose lipid composition can be controlled. In spite of some
technical difficulties, membrane proteins can be reconstituted
into them. FCS measurements in GUVs have been success-
fully used during the recent years for the study of the lateral
organization and dynamics of several membrane proteins, like
an ATP-binding cassette transport system from Lactobacillus
lactis, the lactose transporter LacS from Streptococcus
thermophilus, the mechanosensitive channel MscL from
Escherichia coli (Doeven et al. 2005), and bateriorhodopsin
(Kahya et al. 2002). The possibility to produce phase-
separated membranes in GUVs has extended the use of FCS
to characterize the partition and the dynamics of membrane
proteins in liquid-disordered (L
d
) and liquid-ordered (L
o
)
raft-like phases. The SNARE proteins syntaxin 1A and
synaptobrevin 2 preferentially localized to the L
d
phase when
functionally reconstituted into domain-exhibiting GUVs
(Bacia et al. 2004b). A significantly higher partition into
the L
o
phase, around 20–30%, was found in the cases of
BACE and human placental alkaline phosphatase (Kahya et
al. 2005; Kalvodova et al. 2005). Due to the differential
diffusion properties of raft-associated cholera toxin and the
L
d
-located lipid dye DiI, FCS could be use to analyze the
relationship between domain-exhibiting GUVs and rafts in
living cells (Bacia et al. 2004a).
FCS has been employed several times to study mem-
brane receptors dynamics. By using fluorescently labeled
ligands, the ligand-binding constant to the γ-amino butyric
acid A (GABA
A
) and to the β
2
-adrenergic receptors could
be determined in living cells. Analysis of the mobility of
ligand-receptor complexes revealed two different diffusion
coefficients in both cases, which were assigned to fast and
hindered mobility (Hegener et al. 2004; Meissner and
Haberlein 2003). A similar approach was used to investigate
the interactions of T-cell receptor with CD8 during T-cell
activation (Gakamsky et al. 2005).
The interaction kinetics of fluor escently labeled IgG and
the FcγRII receptor was studied in supported planar
membranes with a combination of FCS and total internal
reflection f luorescence (TIR-FCS; Lieto et al. 2003).
Recently, the technique was extended to living cells and
applied to measure the lateral diffusion of membrane-
bound, farnesylated enhanced green fluorescent protein
(EGFP; Ohsugi et al. 2006).
The organization of the cell membrane has been investi-
gated by FCS. Diffusion measurements at different spatial
scales give rise to the so-called FCS diffusion law, which
provides information about the confinement parameters of
microdomains in cell membranes (Wawrezinieck et al.
2005). The area of measured membrane can be modified
by using laser beams of different size (Wawrezini eck et al.
2005), by performing a z-scan (Humpolickova et al. 2006),
or by means of nanometric apertures that allow to observe
areas below the diffraction limit (Wenger et al. 2007).
Interestingly, the latter approach can give an estimate of the
size of mem brane heterogeneities. The diffusion properties
of GFP proteins bound to the cell membrane through
different tags were affected by the disruption of lipid-based
or cytoskeleton-based microdomains, revealing the impor-
tance of these factors in membrane organization (Lenne et
al. 2006).
Dual-color fluorescence cross-correlation spectroscopy
(FCCS) is a powerful variation of FCS to probe interactions
between two species labeled with spectrally different dyes
(Bacia et al. 2006; Haustein and Schwille 2004). In this
mode, the two fluorophore s are excited within the same
focal volume using two lasers or two-photon excitation, and
the photons emitted are measured using spectral channels.
The analysis of the fluorescence fluctuations in the two
channels yields a cross-correlation function whose ampli-
tude is proportion al to the relative concentration of the
double-labeled particles (interacting molecules that move
together through the diffusion volume). Spectrally distinct
dyes should be used to avoid cross talk and false positive
cross-correlation.
The destiny of two cargoes along the endocytic pathway
was followed with FCCS (Bacia et al. 2002). The A and B
subunits of cholera toxin were labeled with Cy2 an d Cy5,
respectively. Positive cross-correlation could be measured
for the two subunits along the endocytic pathway, until they
were separated in the Golgi network. Larson et al. (2003)
also used a cross-correlation strategy to analyze the dynamic
interactions between labeled Lyn kinase and FcɛRI receptor
stimulated with fluorescently labeled IgE. In an elegant
approach, they were able to perform FCCS measurements in
Appl Microbiol Biotechnol (2007) 76:257–266 259
real time in living cells that revealed a decrease in Lyn diffusion
associated to FcɛRI receptor binding upon stimulation.
Scanning FCS (SFCS) extends FCS to larger diffusion
times and has thus been successfully applied to analyze
membrane dynamics (Ries and Schwille 2006). The extension
to two-focus SFCS, realized by measuring simultaneously in
two detection volumes instead of one, eliminates the need to
calibrate the detection volume, and dual-color cross-correlation
analysis is easily implemented. Accurate diffusion coefficients
could be determined on GUVs, on planar supported bilayers,
and on the membrane of yeast cells.
Single-particle tracking
SPT uses video microscopy combined with digital computer
processing to monitor the motion of single proteins or lipids
on the plane of the membrane. The molecules of interest are
often labeled with submicrometer particles (latex beads of
200–500 nm in diameter or colloidal gold of 40–100 nm in
size) coated with specific antibodies or ligands. These
particles are smaller than the wavelength of light; thus, they
scatter light, giving rise to a diffraction pattern, which is an
airy disc, and allows determination of the centroid position
with around 10 nm precision (Kusumi et al. 2005a).
Molecules can also be fluorescently labeled. The most
common fluorophores are organic dyes, quantum dots, or
fluorescent proteins. Fluorescence labels are usual ly smaller
in size than the particles mentioned above and probably less
invasive. But unlike latex or gold beads, they undergo
photobleaching, thus reducing the achiev able acquisition
time. In both cases, proper controls have to be made to
assure that labeling does not alter the behavior of the
system under study. When noncovalent labeling is used,
there is risk of label exchange among target molecules,
which increases tracking difficulties. In addition, multiva-
lent labels can promote cross-linking, modifying molecule
mobility or inducing cellular responses (Lenne et al. 2006).
Particles are usually ima ged in bright field mode with
enhanced contrast and projected onto a video camera
(Fig. 2a). TIRF microscopy can also be used for SPT, as
only a thin plane of the sample is illuminated, such that the
background fluorescence is strongly reduced and single
particles can be detected in the membrane. The video
images are processed and recorded in real time. Analysis of
the movies is performed digitally. The tracking software
used must be able to accurately track a single molecule
without mistaking it with other ones, while being flexible
enough to allow changes in the particle characteristics due
to out-of-focus movement (Lenne et al. 2006).
Analysis of a single-particle trajectory is done by
determining the x, y positions of the particle in every image
of the movie an d calculating the displacements at Δt time
intervals (Fig. 2b). Then, the mean square displacement can
be related to the diffusion coefficient through different
models, depending on the type of motion. Slow diffusing
particles are experimentally favored, and statistical quality
strongly depends on the time interval. SPT is a powerful
technique to detect deviations from Brownian diffusion,
like confined diffusion, anomalous diffusion, or directed
flow of particles. However, classification of trajectories
must be done with a proper algorithm, which can statistically
distinguish among the different ways of motion (Schmidt
et al. 1996; Schutz et al. 1997).
SPT studies on fluorescent lipid analogs and membrane
proteins led Kusumi et al. to propose a new model of
membrane organization, known as the compartmentalized
fluid model (Kusumi et al. 2005a,b; Ritchie et al. 2005;
Suzuki et al. 2005). The analysis of single-particle mobility
at high frame rates in living cells revealed short-term
confined Brownian diffusion and long-term hop diffusion
between the compartments. According to this model, the
compartment boundaries are form ed by the membrane
skeleton and the membrane proteins associated to it. This
model explains why the diffusion coefficients of lipids and
membrane proteins are larger in artifici al model membranes
than in living cell s and why their mobility is reduced after
molecular complex formation.
In accordance with the above model, Douglass and Vale
(2005) used SPT in an elegant study to show that the adaptor
protein LAT and the tyrosine kinase Lck localized to
membrane microdomains depending on protein
–protein
interactions but not on actin or lipid rafts. These protein
microdomains affected the diffusion of other membrane
proteins by partial exclusion or trapping. These clusters were
proposed to have a role in cell signaling, so that partition of
molecules in and out of the microdomains would decide the
phosphorylation/dephosporylation balance.
Relevant information about the G-protein coupled
receptors (GPCRs) mechanism of action has been obtained
with single-molecule detection and SPT studies. Single-
molecule analysis of ep idermal growth fac tor (EGF)
binding to cells showed that Ca
2+
-signaling was activated
in a sigmoidal fashion. A 50% probability of cell activation
was achieved after binding of 300 EGF molecules, a very
low number compared with the tens of thousands of
receptors expressed on the cell surface (Uyemura et al.
2005). Data analysis revealed that EGF bound preferential-
ly to pre-dimeric EGF receptor complexes, showing coop-
erativity for the binding of the second ligand. This would
allow sensitive signaling even at low EGF concentrations
(Teramura et al. 2006). In addit ion, a decrease in receptor
mobility upon ligand binding was measured for nerve
growth factor receptor, neurokinin-1 receptor, and odorant
receptor OR17-40 (Jacquier et al. 2006; Lill et al. 2005;
Shibata et al. 2006).
260 Appl Microbiol Biotechnol (2007) 76:257–266
The effect of cholesterol concentration on the mobility of
proteins in the plasma membrane has been investigated by
SPT. Interestingly, the diffusion of EGFR, HER2, and the I-
Ek proteins of the major histocompatibility complex
(MCH) decreased after cholesterol depletion in a reversible
way (Nishimura et al. 2006; Orr et al. 2005). In addition,
the mobility of EGFR and HER2, but not I-Ek proteins,
was affected by the cytoskeletal organization in cholesterol-
depleted cells.
SPT experiments were used to monitor the diffusion
properties of the G protein Ras as a function of its activation
state (Lommerse et al. 2005; Murakoshi et al. 2004). Upon
EGF or insulin activation, Ras was greatly immobilized and
confined to 20 nm domains, suggesting the formation of
Ras-signaling complexes. A sim ilar strategy was used to
investigate the diffusion properties of the heterotrimeric G
protein Gq (Perez et al. 2006). Experiments in supported
cell-membrane sheets wi th immobilized GPCRs showed
that Gq in its heterotrimeric state had a lower mobility than
in the monomeric state, suggesting a role of oligomeric
state in differential partition into signaling complexes.
Atomic force spectroscopy
AFM is a scanning probe technique that allows samples to
be imaged with nanometer to atomic level resolution
(Morris et al. 1999). As schematically shown in Fig. 3a,
the AFM microscope consists of an extremely sharp tip
mounted on the end of a cantilever, which allows the tip to
move vertically as it tracks the sample. The scanning
mechanism works via a piezoelectric transducer, which
accurately positions the tip on the sample surface. For
detection, a laser beam is focused on the cantilever and
reflected onto a photodiode. During measurements, the tip
is brought into proximity with the sample surface, which is
scanned underneath the tip. The force experienced between
the tip and samp le causes a deflection of the canti lever,
which changes the position of the refl ected beam on the
photodiode. This, in turn, can be used to provide a
topographical map of the surface (Fig. 3b).
In addition to topographical imaging, AFM can also
probe the nanomechanical properties relating to the force
experienced between the tip and sample. In these measure-
Fig. 2 Principles of single-particle tracking. a Single particles are
detected and time lapse recorded in a stack file. The raw images are
deconvoluted, filtered, and spatially aligned. Then, fluorescence
peaks are accurately localized with a two-dimensional Gaussian fit.
The peak positions are connected in the successive frames, and
particle traces are obtained. b Analysis of single-particle trajectories.
On the left, the trajectories of free-diffusing (A) and confined (B)
particles are shown. The graph on the right depicts the corresponding
mean-square-displacement vs lag time for the same particles
Appl Microbiol Biotechnol (2007) 76:257–266 261
ments, the AFM tip is brought into contact with the sample
and pulled away. Then, the deflection of the cantilever
during this approach and retraction is recorded. The plot of
cantilever deflection vs tip/sample distance is called force
curve (Fig. 3c). Small force differences can be obtained,
thanks to the low spring constant of the cantilever (Janovjak
et al. 2006; Zlatanova et al. 2000).
For sample preparation, the membrane protein of interest
is embedded in planar lipid bilayers (with a defined lipid
composition, coming from native membranes or in the form
of 2D protein-lipi d crystals) supported on a solid substrate.
Unlike with FCS and SPT, AFM does not need protein
labeling. Measurements are usually done under physiolog-
ical conditions in the presence of a bathing buffer solution
and at ambient temperature.
The AFM mode provides images of single-membrane
proteins at subnano meter resolution. The organization of
the photosynthetic machinery in several bacteria has been
visualized by AFM at high resolution, providing informa-
tion about the molecular architecture of the complexes in
native membranes (Fotiadis et al. 2002; Scheuring et al.
2001; Scheuring et al. 2003a,b; Scheuring et al. 2004a,b;
Scheuring et al. 2006). Interestingly, a comparative AFM
analysis of Rhodospirillum photometricum membranes
under low- and high-light conditions showed adaptation in
the photosynthetic complex assembly to optimize photo-
synthetic efficiency under different environmental condi-
tions (Scheuring and Sturgis 2005).
On the other hand, the single-molecule force spectros-
copy mode allows the characterization of inter- and
intramolecular interactions with high sensitivity. In unfold-
ing studies, the AFM tip is brought into contact with a
membrane protein and bound to it. Then, retraction of the
tip from the membrane stretches the polypeptide chain.
When the applied force overcomes the stability of the
protein, it unfolds and gives rise to peaks in the force curve
that correspond to elements of secondary structure in the
protein (Janovjak et al. 2006; Muller et al. 2006). The first
Fig. 3 Atomic force microscopy. a Scheme of an AFM setup. A very
sharp tip is mounted onto a cantilever and brought into proximity of the
sample, which is supported on a solid substrate. Relative positioning of
the sample and the AFM tip is performed with high accuracy , thanks to a
piezoelectric transducer . The detection module is composed of a laser
beam focused on the cantilever and reflected onto a photodiode detector
(courtesy of C. Bippes). b Three-dimensional topographical image of a
SM/DOPC/Chol supported lipid bilayer containing placental alkaline
phosphatase. The liquid-disordered and liquid-ordered phases are shown
with a height scale. Peaks of protein are depicted in white (kindly
provided by S. Chiantia). c Force curve obtained for the unfolding of a
single sodium/proton antiporter . Characteristic peaks are obtained that
correspond to the unfolding of secondary structure elements in the protein
(gently supplied by A. Kedrov)
262 Appl Microbiol Biotechnol (2007) 76:257–266
unfolding experiments exploring the stability o f the
structural elements of membrane proteins were performed
with bacteriorhodopsin (Muller et al. 2002b; Oesterhelt
et al. 2000; Scheuring et al. 2006). The peaks obtained in
the force–distance curves could be assigned to secondary
structure elements within the protein. Since then, severa l
publications have analyzed in depth how the molecular
interactions that stabilize the membrane structure of this
and other rhodopsins are affected by extra - or intracellular
unfolding, disulfide bonding, and oligomerization state
(Cisneros et al. 2005; Janovjak et al. 2004; Kessler et al.
2006; Kessler and Gaub 2006; Sapra et al. 2006; Tanuj
et al. 200 6 ). Single-molecule force spectroscopy was also
used in unfolding and refolding experiments with a
bacterial sodium/proton antiporter (Kedrov et al. 2004).
The effect of pH and of an inhibitory ligand on the molecular
interactions was investigated (Kedrov et al. 2005, 2006b).
Recently, Kedrov et al. (2006a ) reported an improved AFM
setup that enabled the study of the folding kinetics for this
antiporter with a time resolution of a few milliseconds.
Single-molecule recognition studies can be performed by
immobilizing one of the interacting partners to the AFM tip and
measuring the force necessary to break down the interacting
complex. This application allows recognition imaging, a
combination of topographical imaging and force measure-
ments.Ithasbeenusedtostudyligand/receptor interactions
(Kienberge r et al. 2006). Recently, it was also employed to
study the topology of the Na
+
/glucose co-transporter in living
cells (Puntheeranurak et al. 2006). The AFM tip was modified
with an antibody against the C-terminal loop 13 of the protein,
and single-mole cule recognition events probed that the
antigenic region was oriented towards the extracellular region.
Moreover,
D-glucose modified tips were used to investigate
the effect of the presence/absence of Na
+
on ligand binding
and to test the transporter specificity in competition experi-
ments with
L-glucose and D-galactose. Similarly, th e
appearance of HSP60 on the cell membrane of living and
fixed endothelial cells upon heat stress was confirmed by
single-molecule recognition events using an AFM tip modi-
fied with an antibody against the protein (Pfister et al. 2005).
The organization of eukaryotic membrane complexes has
been investigated by AFM. Gap junctions are channels
between neighboring cells, which are important for the flux
of ions and metabolites as well as for cell-to-ce ll signaling.
Connexon complexes that form the gap junctions were
purified and reconstituted into supported planar bilayers.
The structure and mechanical properties of the cytoplasmic
and extracellular loops were studied by imaging AFM and
single-molecule force spectroscopy (Liu et al. 2006; Muller
et al. 2002a). Aquaporins have also been studied by AFM
(Fotiadis et al. 2002; Moller et al. 2003; Schenk et al.
2005), and recently, the structural organization of mem-
brane patches containing junctional microdomains com-
posed by connexons and aquaporins was visualized by
high-resolution AFM (Buzhynskyy et al. 2007).
Recently, Gonçalves et al. (2006) reported high-resolution
AFM imaging and force spectroscopy measurements on
unsupported membranes. This two-chamber AFM setup is a
significant step towards more physiological conditions. In
addition, it allows the use of different buffer solutions
across the membranes for the study of gradient-excitable
membrane proteins.
The combination of AFM with FCS is a very promising
application that has been developed during the last couple of
years. It was initially applied to study the dynamic properties
of supported lipid bilayers exhibiting phase separation (Burns
et al. 2005; Chiantia et al. 2006a). Now, the first studies
including membrane proteins start to appear. Chiantia et al.
(2006b) used a combination of AFM and SFCS to solve the
existing controversy about the lateral organization of the
placental alkaline phosphatase, a raft associated protein, in
domain-exhibiting membranes. The dynamics and lateral
distribution of other membrane proteins, like cholera toxin,
which partitions to the liquid-ordered phase, and synapto-
brevin, found in the liquid-disordered phase, have also been
investigated (Chiantia et al. 2006b and unpublished results).
Conclusions
The use of single-molecule techniques for the study of
membrane proteins has already shown its usefulness. Among
their major achievements, single-molecule studies have
revealed a more complex organization of the cell membrane,
allowing significant extension of the fluid mosaic model.
Besides, the molecular details of signaling processes, including
receptor activation, lateral organization, and dynamics or the
formation of signaling complexes, are now better understood.
However, technical difficulties to reconstitute membrane
proteins into model membranes still are the bottleneck for the
advancement of knowledge in the field. Therefore, the
development of general, reproducible methods is highly
desirable not only for single-molecule applications but also
for averaging biochemical and biophysical techniques. Cur-
rent approaches involve detergent-mediated reconstitution of
membrane proteins into proteoliposomes (Bacia et al. 2004b;
Girard et al. 2004; Kahya et al. 2002), which can be used for
GUV or supported bilayer production. Alternatively, mem-
brane proteins can be in vitro translated and simultaneously
incorporated into supported bilayers (Robelek et al. 2007).
Alternatively to in vitro systems, single-molecule tech-
niques also offer the possibility of studying membrane
proteins in their natural environment with minor alte rations
of the biological system. Living-cell applications open a
new way to investigate the structure, molecular mecha-
nisms, and dynamics of membrane proteins.
Appl Microbiol Biotechnol (2007) 76:257–266 263
The maturation and easier accessibility of these method-
ologies for membrane research will continue to make single-
molecule analysis a major tool in the membrane protein field.
Aknowledgments We thank J. Suckale, J. Ries, and S. Chiantia for
careful reading. This work was supported by a Marie Curie Intra-
European Fellowship (FP6) and German DGF (SCHW716/4-1).
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