11674
Langmuir 2004, 20, 11674-11683
Annexin A1 Interaction with a Zwitterionic Phospholipid
Monolayer: A Fluorescence Microscopy Study
J. Alfredo Freites,† Shahla Ali,† Anja Rosengarth,‡ Hartmut Luecke,‡,§,| and
Michael B. Dennin*,†
Department of Physics and Astronomy and Institute for Surface and Interface Science,
University of California, Irvine, California 92697-4575, and Department of Molecular Biology
and Biochemistry, Department of Physiology and Biophysics, and Department of Information
and Computer Science, University of California, Irvine, California 92697-3900
Received February 2, 2004. In Final Form: July 30, 2004
We present the results of a fluorescence microscopy study of the interaction of annexin A1 with
dipalmitoylphosphatidylcholine (DPPC) monolayers as a function of the lipid monolayer phase and the
pH of the aqueous subphase. We show that annexin A1-DPPC interaction depends strongly on the domain
structure of the DPPC monolayer and only weakly on the subphase pH. Annexin A1 is found to be line
active, with preferential adsorption at phase boundaries. Also, annexin A1 is found to form networks in
the presence of a domain structure in the monolayer. Our results point toward an important contribution
of the unique N-terminal domain to the organization of the protein at the interface.
Introduction
The annexins are a multigene family of proteins
characterized by their capacity of reversibly binding to
anionic phospholipids in a Ca2+-dependent manner.1,2
Their common folding motif is a disk-shaped C-terminal
core domain that contains four (eight in annexin A6)
homologous repeats of five R-helices each, with the
calcium-binding sites located on the convex side of the
disk. In contrast, the N-terminal domain is variable and
is thought to confer specific properties to each annexin.
Although no unambiguous physiological role has been
determined for this family, annexins have been associated
with various membrane-related phenomena, including
membrane organization, membrane trafficking, fusion,
and ion-channel formation.1-3
Annexin A1 is characterized by an N-terminal domain
that comprises its first 41 residues, with residues 2-17
forming an amphipathic R-helix. As is the case with other
annexins with large N-terminal domains, annexin A1
exhibits membrane aggregation properties.2 Results from
vesicle aggregation studies with annexin A1,4,5 annexin
A1-A5 chimeras,6,7 and truncated annexin A1 mutants,8-10
* Author to whom correspondence should be addressed.
† Department of Physics and Astronomy and Institute for Surface
and Interface Science.
‡ Department of Molecular Biology and Biochemistry.
§ Department of Physiology and Biophysics.
| Department of Information and Computer Science.
(1) Gerke, V.; Moss, S. E. Physiol. Rev. 2002, 82, 331-371.
(2) Gerke, V.; Moss, S. E. Biochim. Biophys. Acta 1997, 1357, 129154.
(3) Creutz, C. E. Science 1992, 258, 924-931.
(4) de la Fuente, M.; Parra, A. V. Biochemistry 1995, 34, 1039310399.
(5) Bitto, E.; Li, M.; Tikhonov, A. M.; Schlossman, M. L.; Cho, W.
Biochemistry 2000, 39, 13469-13477.
(6) Hoekstra, D.; Buist-Arkema, R.; Klappe, K.; Reutelingsperger,
C. P. M. Biochemistry 1993, 32, 14194-14202.
(7) Andree, H. A. M.; Willems, G. M.; Hauptmann, R.; Maurer-Fogy,
I.; Stuart, M. c. A.; Hermens, W. T.; Frederick, P. M.; Reutelingsperger,
C. P. M. Biochemistry 1993, 32, 4634-4640.
(8) Bitto, E.; Cho, W. Biochemistry 1998, 37, 10231-10237.
(9) Bitto, E.; Cho, W. Biochemistry 1999, 38, 14094-14100.
(10) Wang, W.; Creutz, C. E. Biochemistry 1994, 33, 275-282.
as well as structural studies,5,11,12 suggest that annexin
A1 possesses two distinct membrane binding sites. One
is the canonical calcium-dependent binding site to anionic
phospholipids that is part of the core domain. The other
one is calcium independent and nonspecific for anionic
lipids. On the basis of high-resolution structural studies
of annexin A1 in the presence12 and the absence11 of
calcium, Rosengarth and Luecke have proposed a twostep model for the annexin A1-membrane interaction
leading to membrane aggregation. Starting with the
protein in its inactive form, the first step would be the
calcium-mediated binding to anionic phospholipid headgroups of one membrane. This process involves a change
in conformation of the C-terminal core that results in the
previously buried N-terminal domain becoming solvent
accessible. The second step would be the binding of a second
membrane via hydrophobic interactions with the now
exposed amphipathic N-terminal domain.
To evaluate the hypothesis of a direct interaction
between lipid membranes and annexin A1, Rosengarth et
al.13 conducted a study of the interaction of annexin A1
with DPPC, DPPS, and DPPC-20 mol% DPPS monolayers. Tensiometry measurements were carried out both
in the presence and in the absence of calcium ions. A
monotonic increase in surface pressure as a function of
time was considered as an indication of protein penetration
into the phospholipid monolayer. It was shown in that
study that annexin A1 is capable of penetrating phospholipid monolayers in the absence of calcium and in the
absence of calcium and anionic phospholipids. The penetration process kinetics were found to be best described
as first-order in the presence of calcium and DPPS and
second-order in the absence of calcium. Similar experiments conducted with annexin A5, an annexin with a short
N-terminal domain of only 16 amino acids, did not show
any indication of penetration of this protein into any of
the monolayers. Also, no penetration into the DPPC
monolayer was found when a proteolytic fragment of
(11) Rosengarth, A.; Gerke, V.; Luecke, H. J. Mol. Biol. 2001, 306,
489-498.
(12) Rosengarth, A.; Luecke, H. J. Mol. Biol. 2003, 326, 1317-1325.
(13) Rosengarth, A.; Wintergalen, A.; Galla, H.-J.; Hinz, H.-J.; Gerke,
V. FEBS Lett. 1998, 438, 279-284.
10.1021/la049713b CCC: $27.50 © 2004 American Chemical Society
Published on Web 11/19/2004
Annexin A1 Interaction with a Phospholipid Monolayer
annexin A1 lacking the N-terminal domain was tested.
These results demonstrated the possible occurrence of a
calcium-independent hydrophobic interaction between the
annexin A1 N-terminal domain and phospholipid membranes.
The emerging model of the cell membrane14 pictures an
inhomogeneous medium that is organized into well-defined
domains. The domain structure is dependent upon local
composition and ordering. In this context, an alternative
point of view to conventional binding experiments is to
consider the influence of spatial organization on membrane-protein interaction. In the case of calcium-mediated
annexin interaction with model membranes, microscopy
studies of lipid monolayer systems at the micrometer15,16
and sub-micrometer17 scales have revealed the formation
of microstructural domains that are specific, in distribution
and morphology, to the calcium-mediated binding event.
Such behavior is manifested as formation of condensed
domains in single component fluid monolayers16 or
formation of anionic lipid-rich domains in mixed monolayers.17 A similar characterization has not been reported
for the calcium-independent annexin A1 membrane
interaction.
In this work, fluorescence microscopy results are
reported on the evolution of the interaction process of
annexin A1 with dipalmitoylphosphatidylcholine (DPPC)
monolayers as a function of the monolayer phase state
and the composition of the aqueous subphase. DPPC is
zwitterionic, and it has already been established that a
calcium-independent annexin A1 interaction with DPPC
exists.13 Central to this work is the possibility of testing
the protein behavior in a heterogeneous medium by
exploiting the well-understood phase behavior of DPPC
monolayers. Both the zwitterionic nature of DPPC and
its domain structure are potentially relevant to the
identified secondary protein membrane binding site. The
phase behavior of the annexin A1-DPPC monolayer
system is found to be consistent with a mean-field theory
proposed by Netz et al.18 Annexin A1 is line active, and
its phase behavior suggests that, upon adsorption, it
undergoes a form of self-assembly. Although DPPC is not
the best representative of a cell membrane phospholipid,
it serves as an excellent model system for probing the
calcium-independent membrane binding. The results
presented here highlight the importance of the nature of
the annexin A1 secondary binding site and the heterogeneous nature of the monolayer organization. This
provides a motivation for more-direct studies that better
resemble biological conditions.
Experimental
Dipalmitoylphosphatidylcholine (DPPC) was purchased from
Avanti Polar Lipids (Alabaster, AL). The monolayer fluorescent
probe employed was chain-labeled nitrobenzoxadiazole phosphatidylcholine (NBD-PC) from Molecular Probes (Eugene, OR).
Both lipids were specified to be more than 99% pure and were
used as received. Spreading solutions contained less than 0.7
mol% (with respect to total lipid) of fluorescent probe and were
prepared with chloroform (HPLC grade, EM Science, Gibbstown,
NJ) in concentrations of 1.4-1.5 g/L. Monolayer subphase buffers
contained 50 mM MES/NaOH, 100 mM NaCl, pH 6.0 adjusted
(14) Edidin, M. Nat. Rev. Mol. Cell Biol. 2003, 4, 414-418.
(15) Koppenol, S.; Tsao, F. H. C.; Yu, H.; Zografi, G. Biochim. Biophys.
Acta 1998, 1369, 221-232.
(16) Wu, F.; Gericke, A.; Flach, C. R.; Mealy, T. R.; Seaton, B. A.;
Mendelsohn, R. Biophys. J. 1998, 74, 3273-3281.
(17) Janshoff, A.; Ross, M.; Gerke, V.; Steinem, C. ChemBioChem
2001, 2, 587-590.
(18) Netz, R. R.; Andelman, D.; Orland, H. J. Phys. II 1996, 6, 10231047.
Langmuir, Vol. 20, No. 26, 2004 11675
by NaOH, or 50 mM Tris/HCl, 100 mM NaCl, pH 7.4 adjusted
by HCl. Additionally, buffers contained either 1 mM ethyleneglycol bis (β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA)
or 1 mM CaCl2. The water used throughout all the experiments
was filtered using a Milli-Q device (Millipore) and had a resistivity
greater than 18 MΩ.
Expression and purification of full-length porcine annexin A1
was performed according to Rosengarth et al.19 Protein fluorescent
labeling was performed by incubating a mixture of 1-5 mg of
protein dialyzed against 0.1 M sodium carbonate/bicarbonate
(pH 9.0) and 1 mg of Texas Red Sulfonyl Chloride (Pierce,
Rockford, IL) on ice for 1 h. The mixture was then dialyzed against
20 mM sodium phosphate (pH 7.5), 150 mM NaCl (three times),
and finally, against the buffer used in the monolayer experiment.
All the experiments were performed in a NIMA Technologies
(Coventry, UK) 601M Langmuir trough equipped with a PS-4
pressure sensor, and controlled by a desktop computer. Trough
temperature was controlled at 20.0 ( 0.5 C. The trough and
tensiometer were mounted on a Olympus BX-60 (Olympus
America, NY) epifluorescence microscope. This was placed on a
vibration isolation table (Newport RS 3000, Irvine, CA). Microscopy images were captured with a monochrome CCD camera
(Cohu 5515, San Diego, CA) connected to a video monitor and
a videocassette recorder. Selected video frames were digitized to
640 pixels × 480 pixels 8-bit gray scale images with a desktop
computer using a frame grabber, cropped to the desired size and,
except for Figure 2a and b, presented without further processing.
Quantitative image analysis was performed using ImageJ
(National Institute of Health, Washington, DC).
The phospholipid solution was spread with a Hamilton
microsyringe to form a monolayer at the air/buffer interface.
After the spreading, 30 min were allowed for solvent evaporation
and overall system relaxation. Isotherm experiments were
conducted with a barrier speed of 4.0 Å2/(molecule min).
Protein/lipid monolayer imaging experiments were conducted
at constant surface pressure. Initially, the monolayer was
compressed to a specific trough area, and then the motion control
of the trough barriers was set to maintain a constant surface
pressure. Once the surface pressure and the trough area had
reached stationary values, the annexin A1 solution was injected
underneath the monolayer at the bottom of the trough, using a
bent long-needle Hamilton syringe. This procedure was conducted
without perturbing the monolayer.
All the microscopy results on protein/lipid monolayer systems
presented were obtained during dual-label experiments, in which
both the protein and the phospholipid monolayer contained a
fluorescent probe. Imaging was carried out with two different
sets of excitation-observation fluorescent cubes that were
manually switched during the experiment. For NBD-PC imaging,
an Olympus U-MWB fluorescent cube (exictation filter, wideband blue 450-480 nm; long-pass barrier filter, 515 nm) was
employed, herein identified as wide-band filter. Texas Red
imaging was performed with a band-pass filter set (Omega
O-5732; excitation filter, 560 ( 20 nm; emission filter, 635 (
27.5 nm), herein identified as Texas Red filter.
DPPC monolayer imaging was conducted during isotherm
experiments (i.e., under compression). Experiments on protein
adsorption to the bare air/buffer interface were conducted at
constant trough area by injecting the annexin A1 solution at the
bottom of the trough.
Results
This section is organized as follows. We will first present
the phase behavior and kinetics of the DPPC monolayer
and annexin A1 separately. We will then focus on the
interaction between the annexin A1 and the liquid
expanded-liquid condensed (LE-LC) coexistence phase
of DPPC. This is the central result of the paper, and we
will report on the results for two different pH values. To
support our interpretation of the LE-LC results, we will
discuss separately the interaction between annexin A1
and the pure LC phase of the monolayer and the
(19) Rosengarth, A.; Rosgen, J.; Hinz, H.-J.; Gerke, V. J. Mol. Biol.
1999, 288, 1013-1025.
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Freites et al.
interaction between annexin A1 and the pure LE phase
of the DPPC monolayer.
Surface Activity of Individual Components. The
results of the phase behavior of DPPC-0.7 mol% NBDPC at 20 °C are shown in Figure 1. Both the Π vs A
isotherm and the corresponding fluorescence micrographs
are in good agreement with those reported in the literature
for DPPC monolayers.20-23 As a function of surface
pressure, DPPC monolayers exhibit three phases at room
temperature: a low-density gaslike phase (G), a liquid
isotropic phase, known as liquid expanded (LE), and a
hexatic phase with a tilted director, known as liquid
condensed (LC). NBD-PC localizes preferentially in the
LE phase, as it is excluded from the LC phase due to the
acyl chain ordering. Therefore, it only produces a significant signal in the LE phase. The onset of the monophasic
LE region occurs at values of specific area in the range
of 90-100 Å2/molecule, as indicated by the appearance of
a uniformly bright, featureless image under the fluorescent
filter. After further compression, a first-order transition
occurs from LE to LC, between 4.5 and 5.0 mN/m, as
indicated by the isotherm plateau and the observation of
dark domains on the fluorescence microscopy image (see
Figure 1a). These curved, multilobular LC domains are
characteristic of monolayers of enantiomeric phospho-
lipids.24 In phospholipid monolayers, for a given cycle of
compression and expansion, the nucleation and growth of
the LC phase depend on a series of factors: the composition
of the subphase dominates the nucleation and early stages
of growth, whereas the subsequent domain shape evolution
and growth are primarily determined by the compression
rate history.22,23,25,26 The LC monophasic region appears
in fluorescence microscopy images with the LC domain
boundaries flattened and in contact with each other (see
Figure 1b). The fluorescent probe is segregated to the
interboundary regions. As the pressure increases, the
fluorescent probe is increasingly excluded from the air/
water interface.
The overall microstructure formation of phospholipid
monolayers in a condensed biphasic state and the morphology of LC domains has been successfully described by
a simple phenomenological model,27 where mesoscopic
phenomena emerge from the interplay between electrostatic, interfacial, and chiral effects. Domain arrangement
and the lack of secondary nucleation events are considered
to be due to a long-range repulsion between collinear
effective electric dipoles of neighboring LC domains. These
electric dipoles represent the net electrostatic effect arising
from the ordering of the lipid acyl chains. Domain
morphology and growth are understood in terms of a
balance between the line tension associated with the LELC boundaries, which favors compact shapes of low
perimeter-to-area ratio, and the long-range dipolar repulsion within LC domains, which favors more-extended
morphologies. It has been shown24 that in the case of
enantiomeric lipids both aspects are governed by molecular
chirality.
The surface-active character of annexin A1 was first
reported by Rosengarth et al.13 Here, the tensiometry
characterization is complemented with fluorescence microscopy. Figure 2 shows the evolution of the surface
pressure after injection of annexin A1 into pH 6.0 buffer
solution. A nonzero surface pressure is observed after a
time lag of 1500 s. After a transient period, the surface
pressure begins to plateau around 10 mN/m 6000 s after
injection. The presence of protein at the surface was first
observed 200 s after injection. During the induction period
and during the first half of the transient period, fluorescence microscopy reveals extended condensed phase
protein domains arranged in an inhomogeneous frothlike pattern, as shown in Figure 2a and b. Eventually,
these domains coalesce, yielding a featureless, bright
image (see Figure 2c).
The adsorption of proteins to fluid interfaces reflects
the amphipathic nature of the polypeptide chain. However,
in contrast to simple amphiphilic molecules, the mechanism of adsorption is determined not only by the intrinsic
gradient of chemical potential but also by a complex
interrelation between entropic (conformational), hydrophobic, electrostatic, and van der Waals interactions. The
surface of the annexin C-terminal core is mostly hydrophilic. As a consequence, adsorption to the fluid interface
implies conformational changes that expose hydrophobic
segments to the nonpolar medium. Several studies by
neutron reflectivity on the adsorption to the air/water
interface of rigid28,29 and nonrigid30 globular proteins have
(20) Albrecht, O.; Gruler, H.; Sackmann, E. J. Phys. 1978, 39, 301313.
(21) Kane, S. A.; Compton, M.; Wilder, N. Langmuir 2000, 16, 84478455.
(22) McConlogue, C. W.; Vanderlick, T. K. Langmuir 1997, 13, 71587164.
(23) Klopfer, K. J.; Vanderlick, T. K. J. Colloid Interface Sci. 1996,
182, 220-229.
(24) Kruger, P.; Losche, M. Phys. Rev. E 2000, 62, 7031-7043.
(25) Mohwald, H. Annu. Rev. Phys. Chem. 1990, 41, 441-476.
(26) Mohwald, H. Phospholipid Monolayers. In Handbook of Biological Physics; Lipowsky, R., Sackmann, E., Eds.; Elsevier Science:
Amsterdam, 1995; Vol. 1.
(27) McConnell, H. M. Annu. Rev. Phys. Chem. 1991, 42, 171-195.
(28) Lu, J. R.; Su, T. J.; Thomas, R. K.; Penfold, J.; Webster, J. J.
Chem. Soc., Faraday Trans. 1998, 94, 3279-3287.
Figure 1. Surface pressure vs specific area isotherm for DPPC
on MES/NaOH buffer at 20 °C. Corresponding fluorescence
micrographs: (a)Π ) 4.7 mN/m, (b)Π ) 9.1 mN/m. The scale
bar is 20 µm. Imaging performed with a wide-band blue
excitation filter and a long-pass green emission filter, sensitive
to both NBD-PC and Texas Red (herein identified as wideband filter). In the micrographs, the bright regions correspond
to the monolayer LE phase and the dark domains to the LC
phase.
Annexin A1 Interaction with a Phospholipid Monolayer
Langmuir, Vol. 20, No. 26, 2004 11677
Figure 2. Surface pressure evolution after the injection of annexin A1 in MES/NaOH buffer to a final concentration of 24 nM.
Corresponding fluorescence micrographs: times after injection (a) 1120, (b) 1330, and (c) 2800 s. The scale bar is 20 µm. Imaging
performed with a narrow band-filter selective for Texas Red (herein identified as Texas red filter). The bright regions in the
micrographs correspond to annexin A1 adsorbed at the air/water interface.
revealed that most of the conformational changes associated with adsorption tend to conserve secondary structure.
This is achieved by the promotion of specific forms of
aggregation or assembly that are consistent with the
tertiary structure in solution.28,29 A recent fluorescence
microscopy study of lysozyme,31 consistent with this model,
presents a similar phase behavior to the one reported here
for annexin A1. Our fluorescence microscopy results
confirm the formation of a protein condensed phase
accompanying the surface tension relaxation, suggesting
that a similar behavior can be expected for annexin A1.
Annexin A1 interaction with DPPC Biphasic
Monolayers. To study the interaction between annexin
A1 and the DPPC monolayer, fluorescence microscopy was
conducted while the monolayer was held at stationary
values of surface pressure, as shown in Figure 3. Constant
surface pressure experiments, as opposed to constant area,
present the protein with a phospholipid monolayer that
has a stationary phase distribution. Because the LC phase
of DPPC is metastable,26 a monolayer held at fixed specific
area experiences a surface pressure relaxation and
accompanied partial dissolution of LC domains. The extent
and specific evolution of this relaxation process will depend
on the specific compression/expansion history.23 In consequence, to achieve the desired nearly constant phase
distribution, the magnitude of the surface pressure has
to be maintained stationary by continuously adjusting
the trough area.
(29) Lu, J. R.; Su, T. J.; Howlin, B. J. J. Phys. Chem. B 1999, 103,
5903-5909.
(30) Atkinson, P. J.; Dickinson, E.; Horne, D. S.; Richardson, R. M.
J. Chem. Soc., Faraday Trans. 1995, 91, 2847.
(31) Erickson, J. S.; Sundaram, S.; Stebe, K. J. Langmuir 2000, 16,
5072-5078.
Figure 3 shows the evolution of the surface pressure
and trough area throughout a complete experiment.
Region I corresponds to the initial compression to a specific
area in the LE-LC biphasic region after which the trough
barriers are controlled so as to keep a constant surface
pressure. The experiments were performed at an LC area
fraction of 33 ( 1%. Region II corresponds to a transient
stabilization period. The asterisk marks the time of
injection of annexin A1. The first indication of annexin
A1 at the air/buffer interface occurs about 800 s after
injection. During the period identified as Region III, the
presence of annexin A1 at the air/buffer interface is
observed in isolated locations. These small domains have
no measurable impact on the trough area. Only after a
uniform distribution of small protein domains at the LELC boundaries exists is a monotonic trough area increase
observed (region IV in Figure 3). This monotonic increase
in the trough area, with a corresponding constant surface
pressure, can only be explained as a displacement of the
phospholipid by the adsorbed protein due to the penetration of annexin A1 into the monolayer.
Figure 4 shows a sequence of micrographs obtained with
the Texas Red filter corresponding to the interaction
process of the protein with the phospholipid monolayer at
pH 6.0 containing EGTA in the subphase. The initial
nucleation and uniform distribution of protein domains
at the LE-LC boundaries is shown in Figure 4a (see also
Figure 1 in Supporting Information). The subsequent
growth by coalescence of the initial domains consists of
a wetting of the LE-LC boundary. Only after complete
coverage of the LE-LC boundaries are protein domains
observed in the LE phase (see Figure 4b). Occasionally,
what appear to be small protein domains can be observed
associated to the location of the monolayer LC domains
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Freites et al.
Figure 3. Surface pressure and trough area evolution for the system DPPC monolayer-annexin A1 at pH 6.0. The asterisk
indicates the time of injection of annexin A1 to a final concentration of 24 nM. Points a-d correspond to the fluorescence microscopy
images shown in Figure 4. See text for explanation of the regions labeled I-IV.
Figure 4. Fluorescence micrographs for the system DPPC monolayer-annexin A1 at pH 6.0 and with the phospholipid monolayer
in a biphasic state (LC area fraction is 33%). Imaging performed with the Texas red filter (see Figure 2 and text for more details
on the filter set). The scale bar is 20 µm. Times after protein injection are: (a) 1604, (b) 1731, (c) 1917, and (d) 2203 s. The white
regions correspond to adsorbed annexin A1.
(see Figure 2 in Supporting Information). The protein
domains at the boundaries grow toward the LE phase
keeping a circular interface with it. Micrographs (Figure
5) taken with the wide-band filter during this stage,
revealing the protein domains in light gray (red in the
visual observation), confirm the growth of the protein
domains and the wetting of the LE-LC boundary.
During the initial stages of adsorption, there are no
apparent changes in shape or size of the LC domains,
suggesting that annexin A1 has displaced the LE phospholipid phase without compressing it to form a new LC
phase, hence, the observed increase in trough area. The
next step of the interaction process is the coalescence of
protein domains located on different LE-LC boundaries.
This results in the formation of a continuous protein
network with the LC domains as nodes (Figures 4c and
5b and c). During this process, an increase of LC domain
size of 35% on average was also observed. As a consequence
Annexin A1 Interaction with a Phospholipid Monolayer
Langmuir, Vol. 20, No. 26, 2004 11679
Figure 5. Fluorescence micrographs for the system DPPC monolayer-annexin A1 at pH 6.0 and with the phospholipid monolayer
in a biphasic state (LC area fraction is 33%). Imaging was performed with the wide-band filter (see Figure 1 and text for more
details on filter set). White and gray regions appeared green and red, respectively, in the visual observation. The scale bar is 20
µm. Times after protein injection are: (a) 1626, (b) 1802, (c) 1915, and (d) 2441 s. White or light gray regions correspond to the
monolayer LE phase, gray regions correspond to adsorbed annexin A1, and dark domains correspond to the monolayer LC phase.
of this process, the LE regions are fully confined and the
protein domains appear to occupy most of the LE area.
The formation of this network and the nucleation and
growth of protein domains in the LE phase region occur
independently. The completion of the interdomain protein
network is followed by a loss of curvature and an overall
change in shape of the LC domains. It is worth noting
that this shape transition coincides with the formation of
the protein network and not with the complete wetting of
the LC domains. Notice also that, during the LC domain
shape change process, the fastest average rate of trough
area increase is about 56% of the slowest compression
rate (that of region I in Figure 3) usually employed for the
generation of surface pressure vs area isotherms. As no
morphology changes are observed due to any motion
caused by compression, it is reasonable to expect that the
LC domain shape transition and other phenomena associated with the protein-monolayer interaction are not
the result of the slower barrier motion necessary to
maintain a constant surface pressure.
In the final stage of adsorption, the fluorescence signal
originating from the annexin A1 essentially fills the
viewing field (Figure 4d). However, comparison with
Figure 5d confirms that the LE phase is still present in
the monolayer. Additionally, after the size increase
observed in the previous stage, the size of the LC domains
remains unchanged within the experimental uncertainty
(LC average domain size relative uncertainty per image
is between 15% and 16%) until the end of the protein
adsorption process. These facts suggest that the features
revealed by fluorescent microscopy during the late stages
of adsorption do not reflect a process that occurs entirely
at the air/buffer interface but immediately underneath.
Also, contributing to the features in the Texas Red images
is the larger fluorescence intensity of Texas Red. This
tends to amplify the size of the protein domains, such as
the ones observed in Figure 4. This effect was verified by
contrasting these images with those taken with the wideband filters on the same areas.
A pH of 6.0 for the aqueous subphase was selected on
the basis of previously reported results,19 indicating that
annexin A1 shows it highest thermodynamic stability
between pH 5.0 and 6.0. To investigate a potential
dependence of the protein surface activity on pH, experiments were also performed at pH 7.4 with a similar LC
area fraction. Taking the rate of change in trough area as
a qualitative measure of kinetics, the comparison of the
graph in Figure 6 with Figure 3 reveals similar penetration
kinetics at pH 6.0 and 7.4, once full coverage of LC domains
occurs. Fluorescence microscopy revealed mostly similar
microstructural features and overall interaction processes
between annexin A1 and the DPPC monolayer for pH 7.4,
as observed in the pH 6.0 experiments (see Figure 6).
Comparison of the graph in Figure 6 with Figure 3
reveals that the length of time spent in region III, the
initial adsorption of the protein into the monolayer, differs
by approximately 3000 s, with it being longer for the pH
7.4 system. However, consistent with the pH 6.0 systems,
the nonzero rate of change of area (region IV) for pH 7.4
occurs when a uniform distribution of small protein
domains covers the LE-LC boundaries (see Figure 6a
and c). At this point, the rate of trough area expansion is
very similar at both values of pH. It is also noticeable that
the time interval between the first protein adsorption
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Langmuir, Vol. 20, No. 26, 2004
Figure 6. Surface pressure and trough area evolution for the
system DPPC monolayer-annexin A1 at pH 7.4 with the
phospholipid monolayer in a biphasic state (LC area fraction
is 33%). The asterisk indicates the time of injection of annexin
A1 to a final concentration of 24 nM. Corresponding fluorescence
micrographs: times after injection (a) 3212, (b) 4280, (c) 3087,
and (d) 3917 s. (a-b) White regions correspond to adsorbed
annexin A1, imaging performed with the Texas red filter. (c-d)
White or light gray regions correspond to the monolayer LE
phase, gray regions correspond to adsorbed annexin A1, and
dark domains correspond to the monolayer LC phase, imaging
performed with the wide-band filter. The scale bar is 20 µm.
events and the full coverage of LE-LC domain boundaries
in all of these experiments is within the same time scale
(between 1500 and 2000 s) as the onset of surface pressure
increase for annexin A1 at the bare air/water interface.
For both systems at pH 6.0 and 7.4, the microstructural
organization consists of a network of protein domains.
Also, at the late stages, the LC domains undergo a shape
transition to long, skinny domains (see Figure 6d). The
only real difference is the magnitude and extent of the
domains. For pH 7.4, the growth by coalescence of protein
domains at the LE-LC boundaries does not progress to
a full extent before the experiment is terminated. Similarly, in contrast to the observations at pH 6.0, the
interdomain network formed is not uniformly extended
(Figure 6b and d). Therefore, the LE regions are not
confined by the protein network (Figure 6b). Additionally,
as a consequence of the protein domain formation, it
appears that the distribution of LC domains becomes
clustered or at least less uniform.
Given that the chosen values of pH are on opposite sides
of the protein calculated isoelectric point, these results
Freites et al.
suggest that there is not a strong pH dependence for the
protein-monolayer interaction. For both pH values, the
initial and final trough areas are approximately equal.
Therefore, the observed difference in protein coverage in
the late stages may reflect the amount of protein aggregated below the interface and may not be related to
the monolayer-protein interaction. Moreover, differences
in LC domain size and morphology were observed for
monolayers spread over the two buffer solutions in the
absence of protein. Therefore, the possibility that the
observed differences in overall microstructural organization between the two systems are due to constitutional
differences introduced by the different buffer solutions
and intrinsic to the phospholipid monolayer themselves
cannot be discarded. Further experiments are needed to
completely understand the late-time difference in network
coverage. Experiments were conducted substituting EGTA
in the subphase with CaCl2 at both pH 6.0 and 7.4 (results
not shown). No substantial or systematic differences were
found in either microstructural features or overall kinetics.
In contrast to the canonical behavior of annexins in the
presence of calcium ions and anionic phospholipids, the
results presented so far suggest that the annexin A1DPPC monolayer interaction is predominantly nonelectrostatic. The adsorption of annexin A1 to the phospholipid
monolayer is likely to be accompanied by a change in
protein conformation in a manner consistent with the
adsorption behavior of the protein at the bare air/buffer
interface. In both cases, specific domains in the protein
chain are more likely to be attracted to the lipid interface
through hydrophobic interactions. The formation of a
condensed phase in discrete domains by the adsorbed
protein suggests an aggregation processes regulated by
this change in conformation at the surface. The presence
of the LE-LC domain boundaries appears to modulate
both the nucleation and the growth of the protein domains,
as indicated by the occurrence of wetting and networking.
One consequence of this modulated growth could be
differences in protein chain packing between the domains
at the boundary and those that nucleate in the interior
of the LE phase. This would explain why the first
coalescence occurs only between domains nucleated at
the boundaries since it is the process that leads to the
network formation.
The difference in protein domain morphology between
the present case and the adsorption to the bare aqueous
interface suggests that the protein domains are insoluble
in the LE phase since the formation of circular domain
boundaries minimizes the contact between the protein
and the lipid LE phase. This idea is reinforced by the fact
that, even though the LE regions are being compressed
due to the penetration of the protein into the monolayer,
no secondary nucleation of LC domains is observed in the
interior of the LE phase. Any additional contribution to
the state of stress arising from the fine compressibility of
the LE phase is being relaxed by an increase in the trough
area. Immiscibility between the protein domains and the
LE phase could also explain why those protein domains
nucleated in the interior of LE phase seem to participate
in the coalescence process only at a late stage when their
surface coverage is high and/or the network of domains
nucleated at the LE-LC boundaries is sufficiently thick.
Annexin A1 Interaction with Monophasic Monolayers. To further confirm the previous assessment, the
specific interaction of annexin A1 with each phospholipid
monolayer phase was studied through constant-pressure
experiments conducted above the onset of the LE-LCto-LC transition at 9.0 mN/m and below the onset of the
LE-to-LE-LC at 3.5 mN/m. These results confirm the
Annexin A1 Interaction with a Phospholipid Monolayer
preference for protein adsorption at domain boundaries
(the line activity of annexin A1) and the insolubility of
annexin A1 in the LE phase.
At 9.0 mN/m, the monolayer consists of fully grown LC
domains with flattened boundaries which are in contact
with all their neighbors. In other words, the LC monophasic region is characterized by a granular texture. As
indicated before, achieving a steady state starting from
a monophasic LC state involves some relaxation. In this
case, the relaxation process introduces small isolated
domains of LE phase (see Figure 3 in Supporting
Information). Notice, however, that this microstructure
is not the same as the one for monolayers in the biphasic
region. After injection of annexin A1, an image with the
Texas red filter shows that these boundaries are fully
decorated with small protein domains (see Figure 4 in
Supporting Information) after 1000 s. As in the case of
the biphasic monolayer, the protein domains at the LCLE boundaries grow by coalescence forming a continuous
interphase among the LC domains (see Figure 7a and c).
Subsequently, the LC domains become completely isolated
from each other and the protein layer thickens. At the
same time, the LC domains are elongated until both lipid
and protein form a striped pattern (see Figure 7b and d).
Penetration kinetics are substantially slower than for the
biphasic experiments (see Figure 7). The striped microstructural pattern was maintained with minimum change
until the experiments were stopped. Similar experiments
performed with labeled protein but without fluorescent
label in the monolayer (results not shown, see Figure 5
in Supporting Information for a comparison between
experiments with and without fluorophore in the monolayer) produced a consistent behavior for the adsorption
and domain formation of the protein.
These results confirm the preferential adsorption at
monolayer domain boundaries and the line-active character of annexin A1. The lack of extended regions of LE
phase confirms that the line activity is a distinct characteristic of the quasi two-dimensional protein domains.
The complete alteration of the DPPC monolayer microstructure can only be achieved through a change of the
electric dipole field distribution over and across the
amphiphilic monolayer. This suggests again specificity in
conformation of the protein domains either at a mesoscopic
level or at the level of chain conformation.
At 3.5 mN/m, the protein penetrates the LE monolayer
forming circular domains (see Figures 8a and c), and
ultimately, an emulsion-like pattern forms between the
annexin A1 and the LE phase. The initial protein domains
nucleate uniformly in regions of about 5 µm and grow by
coalescence. This should be contrasted, particularly, with
the morphologies observed during the early stages of the
adsorption of annexin A1 to the bare air/water interface
(see Figure 2a and b). No apparent condensation of the
LE phase to LC phase was observed. At the end of the
experiment, individual protein domains were on the order
of 70-80 µm in diameter (see Figure 8b and d). At that
time, these large domains collapse onto each other to form
larger extended regions. The formation of such large
domains and extended regions is consistent with the
behavior observed for the adsorption to the bare aqueous
interface.
These observations are consistent with the results
obtained with the biphasic monolayers and confirm the
immiscibility of the protein domains in the phospholipid
LE phase. No domain networking was observed on the LE
phase in the monophasic experiment, confirming that the
growth of protein domains in the biphasic monolayer
system is modulated by the presence of the LE-LC
Langmuir, Vol. 20, No. 26, 2004 11681
Figure 7. Surface pressure and trough area evolution for the
system DPPC monolayer-annexin A1 at pH 6.0. The protein
was injected, while the monolayer was held at 9.0 mN/m with
the phospholipid monolayer in the LC phase. The asterisk
indicates the time of injection to a final concentration of 24 nM.
Corresponding fluorescence micrographs: times after injection
(a) 3683, (b) 4733, (c) 3472, and (d) 4716 s. (a-b) White or light
gray regions correspond to adsorbed annexin A1, imaging
performed with the Texas red filter. (c-d) White or light gray
regions correspond to the monolayer LE phase, gray regions
correspond to adsorbed annexin A1, and dark domains correspond to the monolayer LC phase, imaging performed with
the wide-band filter. The scale bar is 20 µm.
boundaries. In the same way, it can be asserted that
nucleation of protein domains in the interior of the LE
phase is an independent state from the preferential
adsorption to the LE-LC boundaries. (This will be
discussed in more detail in the next section in the context
of the model by Netz et al.18) Additionally, as was observed
in the biphasic monolayer systems, the protein domains
that nucleate in the LE phase reach a critical size before
starting coalescence. This behavior is consistent with a
specific pattern of aggregation for the adsorbed protein at
the scale of tertiary structure.
Discussion
We have presented fluorescence microscopy results on
the interaction of annexin A1 with DPPC monolayers as
a function of the lipid monolayer phase state. The central
features are that annexin A1 preferentially adsorbs to
LC-LE domain boundaries and that it ultimately induces
a shape change of the LC domains. Both of these results
indicate that the annexin A1 is line active, relative to
11682
Langmuir, Vol. 20, No. 26, 2004
Figure 8. Surface pressure and trough area evolution for the
system DPPC monolayer-annexin A1 at pH 6.0. The protein
was injected while the monolayer was held at 3.5 mN/m with
the phospholipid monolayer in the LE phase. The asterisk
indicates the time of injection to a final concentration of 24 nM.
Corresponding fluorescence micrographs: times after injection
(a) 748, (b) 5113, (c) 795, and (d) 5160 s. (a-b) White domains
correspond to adsorbed annexin A1, imaging performed with
the Texas red filter. (c-d) White or light gray regions correspond
to the monolayer LE phase and gray domains correspond to
adsorbed annexin A1, imaging performed with the wide-band
filter. The scale bar is 20 µm.
LC-LE domains. The adsorption in the presence of LCLE domains results in the formation of a protein network,
something that does not occur for adsorption in the absence
of the monolayer or in the LE phase. This suggests that
two different adsorbed states exist in the monolayer.
Finally, some protein fluorescent signal was also occasionally observed at the location of the LC domains.
This last feature appeared to originate from underneath
the monolayer since it could be observed even below the
monolayer focal plane. Also, the late-time protein images
suggest the existence of protein aggregates below the air/
water interface. These findings suggest that it is necessary
to consider the possibility that the complexity of the
observed adsorption process and surface behavior could
imply that the protein domains are only partially at the
surface and that the phenomena of conformational change
and aggregation have a multilayer character.
Preferential adsorption to the LE-LC domain boundaries of phospholipid monolayers has been reported for
other proteins that present interfacial activity, such as
Freites et al.
concanavalin A,25 bacterial surface layer proteins,32 fibronectin,33 and surfactant protein A.34 Netz, Andelman,
and Orland18 have developed a Flory-Huggins type meanfield theory that is able to account for this phenomenon.
According to this model, the preferential adsorption of a
protein to LE-LC domain boundaries is an entropic effect
due to the constitutional differences between the adsorbed
protein phase and the phospholipid monolayer. The model
predicts a reduction of the line tension associated with
the LE-LC boundary due to the protein adsorption. This
is consistent with the observed wetting of the LC phase
by the annexin A1 domains located at the LE-LC. The
change in shape of the LC domains can also be explained
in this context. The full coverage of the LE-LC boundary
by coalesced protein domains could screen the dipoledipole interaction between neighboring LC domains. At
the same time, a reduction of the line tension allows the
LC domain morphology to be dominated by the repulsive
dipole-dipole interaction within the domains. It has been
predicted35,36 that under these circumstances the LC
domains would assume elongated shapes, as was observed
during the last stages of the annexin A1-DPPC monolayer
interaction process. The screening of the LC interdomain
dipolar interaction by the adsorbed protein could also
explain the clustering of LC domains observed at pH 7.4.
The theory by Netz et al.18 also accounts for the observed
nucleation of new protein domains in the LE phase, as an
event that could occur due to a change in the protein
chemical potential at the surface. This is consistent with
the reported observation that the adsorption in the interior
of the LE phase occurs after the protein domains have
completely covered the LE-LC boundaries. Consequently,
nucleation of new protein domains in the interior of the
LE phase could be attributed to a critical increase of protein
surface concentration. The model of Netz et al. is based
only on pairwise interactions between the system components, which justifies the assumption that hydrophobic
interactions are dominant in the annexin A1-DPPC
monolayer system.
These ideas can be connected to the three different
penetration kinetics reported by Rosengarth et al.13 for
annexin A1 phospholipid monolayer systems: first-order
kinetics for the system containing both calcium ions and
DPPS, second-order kinetics for the systems containing
DPPS in the absence of calcium ions, and slower secondorder kinetics for the DPPC monolayer system. It can be
speculated that the occurrence of the first-order kinetics
characterizes unambiguously the canonical electrostatic
interaction between annexins and anionic phospholipids.
On the other hand, the mixed monolayer DPPC-DPPS
tested in that study has been reported to present DPPSrich domains in the absence of chelator agents,17,37 it is
then possible that the second-order kinetics correspond
to the kind of complex interfacial phenomena described
here, whereby protein aggregation and line activity play
a dominant role. The reported differences in kinetics
between the DPPC and the monolayers containing DPPS
could be attributed to the different nature of the domains
formed in these systems.
(32) Diederich, A.; Sponer, C.; Pum, D.; Sleytr, U. B.; Losche, M.
Colloids Surf., B 1996, 6, 335-346.
(33) Baneyx, G.; Vogel, V. Proc. Natl. Acad. Sci. U.S.A. 1999, 96,
12518-12523.
(34) Ruano, Miguel, L. F.; Nag, K.; Worthman, L.-A.; Casals, C.; PerezGil, J.; Keough, K. M. Biophys. J. 1998, 74, 1101-1109.
(35) Keller, D. J.; McConnell, H. M.; Moy, V. T. J. Phys. Chem. 1986,
90, 2311-2315.
(36) de Koker, R.; McConnell, H. M. J. Phys. Chem. 1993, 97, 1341913424.
(37) Ross, M.; Steinem, C.; Galla, H.-J.; Janshoff, A. Langmuir 2001,
17, 2437-2445.
Annexin A1 Interaction with a Phospholipid Monolayer
There is good evidence that the phase behavior reported
here for annexin A1 is directly linked to interactions
involving the N-terminal domain. No penetration into
DPPC monolayers was observed by Rosengarth et al.13
for a proteolytic fragment of annexin A1 lacking the
amphipathic N-terminal domain and for annexin A5,
which lacks an N-terminal domain. Further evidence for
the role of the N-terminal domain comes from considering
the association to membranes of annexin A12 and annexin
A5 in the absence of calcium. Under acidic conditions (pH
below 5.0), these annexins appear to refold and insert
into bilayers, yielding a transmembrane configuration.38-40
This phenomenon has been shown to depend on hydrophobic interactions between the protein and zwitterionic
components of the model membranes.39,40 It is highly
(38) Langen, R.; Isas, J. M.; Hubbell, Wayne, L.; Haigler, H. T. Proc.
Natl. Acad. Sci. U.S.A. 1998, 95, 14060-14065.
(39) Isas, J. M.; Cartailler, J.-P.; Sokolov, Y.; Patel, D. R.; Langen,
R.; Luecke, H.; Hall, J. E.; Haigler, H. T. Biochemistry 2000, 39, 30153022.
(40) Ladokhin, A. S.; Isas, J. M.; Haigler, H. T.; White, S. H.
Biochemistry 2002, 41, 13617-13626.
Langmuir, Vol. 20, No. 26, 2004 11683
sensitive to the protonation state of the C-terminal core.
This is in contrast to the results reported here. The fact
that the interaction of annexin A1 with zwitterionic
phospholipid monolayers presents the same microstructural features at neutral and acidic pH allows us to
speculate that this behavior is not related to the C-terminal
core conformation but rather, in accordance with the
results of Rosengarth et al.,13 is related to the amphipathic
nature of the N-terminal domain.
Acknowledgment. J.F., S.A., and M.D. thank the
Petroleum Research Fund (Grant No. 39070-AC9) for
support of this research. A.R. and H.L. thank NIH (Grant
No. GM56445) for support. The authors also thank Nathan
Benedict for his contribution to initiating this collaboration.
Supporting Information Available: Additional images are available as referred to in the text. This material is
available free of charge via the Internet at http://pubs.acs.org.
LA049713B