Colloids and Surfaces
A: Physicochemical and Engineering Aspects 203 (2002) 273– 286
www.elsevier.com/locate/colsurfa
The effect of polar groups on structural characteristics of
phospholipid monolayers spread at the air–water interface
J. Miñones Jr a,*, J.M. Rodrı́guez Patino b, O. Conde a, C. Carrera b,
R. Seoane a
a
Department of Physical Chemistry, Faculty of Pharmacy, Uni6ersity of Santiago de Compostela, Campus Sur,
15706 Santiago de Compostela, Spain
b
Department of Chemical Engineering, Faculty of Chemistry, Uni6ersity of Se6illa, c/. Prof. Garcı́a González, s/n. 41012,
Se6illa, Spain
Received 13 September 2001; accepted 24 October 2001
Abstract
Structural characteristics (structure, morphology, and relative film thickness) of dipalmitoyl phosphatidylcholine
(DPPC), dipalmitoyl phosphatidylglycerol (DPPG) and dipalmitoyl phosphatidylserine (DPPS) monolayers were
determined at the air–water interface at 20 °C and at pH 6 by means of surface pressure (y) – area (A) isotherms
coupled with Brewster angle microscopy (BAM). At lower surface pressures, phospholipid monolayers adopted an
expanded-homogeneous structure at the air–water interface. As the surface pressure increases, in the liquid-condensed
phase (LC), phospholipid monolayers showed film anisotropy and domains with heterogeneous structures. The
homogeneous structures observed at higher surface pressures proved the existence of parallel oriented aliphatic chains
when the close-packed film molecules were in the solid state. The relative monolayer thickness increased with the
surface pressure and was at a maximum at the collapse point. The phospholipid head-group has an important role
on the structural characteristics of the monolayer at the air– water interface. © 2002 Elsevier Science B.V. All rights
reserved.
Keywords: Phospholipid monolayer; Monolayer structures; Brewster angle microscopy
1. Introduction
Biological membranes are organized assemblies
of lipids, proteins, and, to a limited extent, carbohydrates, which are vital to cell function
and development [1]. Proteins and polar lipids
* Corresponding author. Fax: + 34-9815-949-12.
E-mail address: [email protected] (J. Miñones, Jr).
account for almost all of the mass of biological
membranes with a small amount of oligosaccharides, present as part of glycoproteins or glycolipids.
Although, lipid molecules display considerable
structural diversity [2,3], those that are most important to mammalian cellular membranes are the
phospholipids, due to their amphiphatic characteristics. The head of a phospholipid molecule is a
0927-7757/02/$ - see front matter © 2002 Elsevier Science B.V. All rights reserved.
PII: S 0 9 2 7 - 7 7 5 7 ( 0 1 ) 0 1 1 0 7 - 4
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J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
negatively charged phosphate group linked to a
positively charged amino group. Phospholipid
tails can be congregated together to form a local
hydrophobic environment. This leaves the
charged phosphate groups facing out into the
hydrophilic environment.
Langmuir phospholipid monolayers serve as
useful, easy to study, models of cell membranes
[4], and a variety of monolayer properties relevant
to biological processes have been investigated.
The advantage of using a monolayer model lies in
its controllability. A monolayer’s properties can
be carefully tuned, which makes possible the
defining of molecular density by varying the area
per molecule on a Langmuir film balance. Likewise, monolayers enable us to investigate mutual
interactions between molecules in a well-defined
arrangement. However, the critical question regarding the appropriate comprehension and relationship
between
molecular
packing
in
monolayers (along with those in bilayers) still
remains unanswered.
Considerable progress in this area has been
achieved through the development of surface-sensitive optical techniques for ‘in situ’ studies. Microscopic observations of the textures of
Langmuir monolayers became possible one and a
half decades ago, first by fluorescence microscopy
[5,6], and more recently by ellipsometric microscopy [7–9] and Brewster angle microscopy
(BAM) [10,11]. These methods have been used to
identify characteristic domain structures of various phases in the monolayer. A variety of materials have been probed for morphological
information with the objective of correlating microscopic information with macroscopic film
properties.
In this work, surface pressure (y) – area (A)
isotherms coupled with BAM were applied to
analyze the effect of polar groups on the structural and morphological characteristics of phospholipid monolayers. We investigated the surface
behavior of the dipalmitoyl phosphatidylcholine
(DPPC) and dipalmitoyl phosphatidylserine
(DPPS) monolayers and then compared them
with
the
dipalmitoyl
phosphatidylglycerol
(DPPG) monolayer, whose polar group, unlike
the former, lacks the amino group.
2. Materials and methods
2.1. Materials
L-a
phosphatidyl-DL-glycerol
dipalmitoyl
(DPPG; 99% purity), L-a phosphatidylcholine
dipalmitoyl (DPPC; 99% purity) and DL-a phosphatidyl-L-serine dipalmitoyl (DPPS; 98%
purity) were purchased from Sigma. These compounds were dissolved in a chloroform:ethanol
mixture (4:1 v/v) and the solution was
spread onto the air–water interface with a Microman Gilson microsyringe, precise to 90.2 ml. In
each experiment, 4.25× 1016 molecules were deposited on the surface from the spreading
solution at a concentration of approximately 0.46
mg ml − 1. The solutions were prepared every 2
days and stored at 4 °C in a desiccator saturated
with the spreading solvent in order to maintain
the phospholipid concentration. Ultrapure
water, used as subphase, was obtained from a
Milli RO, Milli Q reverse osmosis system (Millipore Corp.) containing two carbon- and two
ion-exchange columns. Finally, the water was
purified through a 0.22 mm Zetapore filter.
The resistivity of the purified water was 18 MV
cm. Temperature and pH were maintained constant at 20 °C and 6, respectively. The subphase pH was adjusted by addition of HCl (p.a.
grade).
2.2. Surface film balance
A Langmuir –Blodgett KSV-5000 (Finland)
trough, equipped with two symmetrical compartments, 71× 12 cm2 each, was used to record the
surface pressure– area (y–A) curves. The
isotherms were obtained by the simultaneous
compression of two monolayers spread on the
water sub-phase in each compartment. Compression was carried out with two barriers moving
with the same speed from the edge of each compartment to its center, where the Wilhelmy plate,
used as a surface pressure sensor, was placed.
With this procedure, we were able to simultaneously register, under identical experimental conditions, the compression isotherms for individual
monolayers spread in each compartment. The re-
J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
producibility of the isotherms, recorded in two
successive compression experiments, was the evidence for reliable results. Each curve shown in
this paper represents the average of four independent experimental y – A isotherms. The monolayers were compressed with a barrier speed of
8.2 A, 2 per molecule per min.
2.3. Brewster angle microscopy
For microscopic observation of the monolayers
morphology, a BAM 2 (NFT, Germany),
equipped with a 30 mW laser emitting p-polarized
light at 690 nm wavelength, was reflected off the
air–water interface at approximately 53.1° (Brewster angle), as described elsewhere [12,13]. In measuring the relative reflectivity (I) of the film, a
previous camera calibration was necessary in order to determine the relationship between the gray
level (GL) and the relative reflectivity (I), according to a procedure previously described [12,13].
The intensity at each point in the BAM
image depends on the local thickness and film’s
optical properties. These parameters can be measured by determining the light intensity at the
camera, and analyzing the polarization state of
the reflected light employing the method based on
the Fresnel reflection equations [14]. At Brewster
angle
I = Rp2 = Cl 2
(1)
where I is the relative reflectivity, C a constant, l
the film thickness, and Rp is the p-component of
the light. The lateral resolution of the microscope
was 2 mm, and the images were digitized and
processed in order to obtain high quality BAM
pictures.
Combined y –A isotherms and BAM measurements on the same monolayer provide complementary structural characteristics of the
monolayer, and thus enable a global consistency
check [12,13]. Moreover, the ability of the BAM
image and relative reflectivity to gain insight
into the internal monolayer structure makes possible the study of the film-forming components’
structure separately in the mixed monolayer
[15,16].
275
3. Results and discussion
3.1. Structural characteristics of DPPC
monolayers
DPPC has frequently been the phospholipid of
choice for many monolayer studies mainly because of its phase transition properties at physiologically relevant temperatures. The y –A
isotherm exhibits a liquid-expanded to liquid-condensed phase (LE-LC) transition (Fig. 1(A)). This
transition is typical of phospholipid films at temperatures below that of the gel–liquid crystal
transition temperature. As for DPPC, the gel–liquid crystal transition temperature is 41 °C [17].
The surface pressures at the beginning and at the
end of the transition are 3.5 and 4.9 mN m − 1,
respectively. Consequently, this is not a true, firstorder phase transition due to the fact that the
surface pressure does not remain constant, nor
can it be considered to be a second order phase
transition (which is characterized by the existence
of a kink point in the y–A curve). The increase in
the surface pressure during the transition may be
due to a kinetic type of cause attributable to the
fact that the film is not compressed with sufficient
slowness. The surface pressures at the transition
agree with those found in the literature [18,19],
with yt = 3.5–3.7 mN m − 1 corresponding to the
beginning of the transition, and yt = 4.3–5.6 mN
m − 1 at the end, when the monolayer is compressed slowly (e.g. 0.8–0.86 A, 2 per molecule per
min) on water at a temperature of 18–20 °C. The
use of faster rates of compression (15 A, 2 per
molecule per min, or greater) leads to an increase
in the surface pressure during the transition, as
described elsewhere [20]. The same phenomenon
occurs, even more significantly, by increasing the
temperature [21,22], ionic strength [23,24], or pH
[25,26].
In the expanded (gaseous) phase, the relative
reflectivity (Fig. 1 (B)) of the DPPC monolayer is
small (0.65× 10 − 7), but it increases abruptly at
the end of the same. In the LE–LC transition
region, the relative reflectivity increases from
4.8×10 − 7 to 11.4× 10 − 7, which corresponds to
an increase of 1.5 times in the film’s thickness.
Finally, in the LC and solid regions, the mono-
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Fig. 1. (A) y– A isotherm for a DPPC monolayer spread on the air – water interface at pH 6 and at 20 °C and (B) time evolution
of relative reflectivity () and surface pressure () during a compression – expansion cycle (shutter speed, 125 s − 1).
J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
layer thickness increases little. It is also important
to point out that, during monolayer compression,
there are few, low intensity, reflectivity peaks due
to the fact that the LC domains are very small
and of low reflectivity (Fig. 2). Indeed, at surface
pressures lower than 4 mN m − 1, the BAM images
show the existence of a homogeneous film (Fig.
2(A)). Once the plateau surface pressure is attained, small, circular domains, that are brighter
than the surrounding area, are formed (Fig. 2(B)).
In the phase transition region, the circular domains increase in size, and adopt an irregular
shape with small intrusions and protrusions bordering their edges (Fig. 2(C)), which grow along
the plateau, subsequently giving rise to domains
with two to four lobes (Fig. 2(D)). Many of these
domains have regions with different reflectivity as
a consequence of the different tilt of the alkyl
chains with respect to the plane of incidence. The
lobed-like shape of DPPC domains was also observed by Vollhardt [27]. This is the structure
typically observed by fluorescence [28] in the LE–
LC transition region, and is caused by the chirality of the DPPC [29]. Once the LE– LC transition
region is exceeded, the domains merge together as
a result of the compression, and their lobed structure slowly blurs (Fig. 2(E)) until it disappears
completely just before the monolayer collapse,
giving a homogenous image (Fig. 2(F)). At the
end of compression, the monolayer collapse is
evidenced by the formation of bright stripes (Fig.
2(G)).
The relative reflectivity curve versus time during
monolayer expansion is symmetrical to that for
compression (Fig. 1(B)); thus, demonstrating the
existence of a reversible behavior during the compression-expansion cycle. Accordingly, the BAM
image at the end of expansion (Fig. 2(H)) is
similar to that registered at the beginning of compression (Fig. 2(A)).
Many authors have postulated that the phosphorylcholine group of DPPC is horizontally oriented at the air–water interface [30– 34]. Based on
molecular models, Vilallonga et al. [33] suggested
the possible existence of three different orientations of the polar group with respect to the long
vertical axis of the molecule: (i) a first orientation
with the trimethylammonium group extended to-
277
ward the water phase below the phosphate group
(with a parallel orientation to the hydrocarbon
chains); (ii) a second orientation with this group
pointing upward towards the air phase over the
phosphate group; and (iii) a third orientation with
both groups coexisting in a horizontal plane (with
an orientation perpendicular to the vertical axis of
the molecule). In the first two cases, an imaginary
line passing through the P and N atoms (which
would represent its dipole moment) would be
parallel to the long vertical axis, and would contribute to the total surface dipole moment of the
molecule (vn). The value of vn should decrease for
the first of the proposed orientations, and should
increase for the second one. On the other hand, if
the orientation of the PN dipole were coplanar
to the air–water interface, it would not contribute
per se to vn. Their results suggest that the most
probable configuration of the phosphorylcholine
group is the latter, in agreement with Shah and
Schulman [34]. Likewise, the molecular area values recently obtained by Pathirana et al. [24] for
DPPC monolayers are in agreement with what
would be expected from a layer-parallel arrangement of the phosphorylcholine group, which requires an area of 47–54 A, 2. This area is
significantly larger than the cross-section of two
palmitoyl chains in the condensed crystalline state
[35].
Nevertheless, this interpretation of the PN
dipole horizontal orientation is only valid as long
as the DPPC molecules are densely packed in the
monolayer, because, in the expanded state the
polar group is able to adopt more flexible orientations [36–39]. Baltes et al. state [40] that the
phosphocholine group is arranged within a plane
parallel to the surface that can be shifted along
the normal by some small distances. The possibility of a change in the orientation of this group has
also been suggested by Gally et al. [41]. These
authors point out that the coplanar conformation
is not rigid since the choline group is able to
undergo angular oscillations with respect to the
surface plane. This change in the normal position
of the terminal ammonium group was also reported by Shapovalov [23], who show that the
phosphate group and neighboring glycerol moiety
are capable of strong hydrogen bonds with water
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J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
Fig. 2. Visualization by BAM of DPPC monolayers spread on the air – water interface at pH 6 and at 20 °C. (A) BAM image at
3.5 mN m − 1; (B), (C) and (D) LC domains during the LE– LC transition phase; (E) LC domains at 12.5 mN m − 1; (F)
homogeneous structure before the collapse; (G) monolayer collapse at 65 mN m − 1; (H) expansion of the monolayer at the
maximum area (y= 0.1 mN m − 1).
J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
279
crease of approximately four carbon atoms in the
length of the hydrocarbon chains that are above
the water surface, as is schematically shown in
Fig. 3. The value of 16.5 A, for the thickness of
the DPPC monolayer in the liquid condensed
state is concordant with that obtained by Bayerl
et al. [43] for dimiristoyl phosphatidylglycerol in
the same state (14.291 A, ), taking into account
the fact that the hydrocarbon chain length of this
phospholipid is 2.54 A, shorter than that for
DPPC.
Furthermore, these relative reflectivity values
(Fig. 1(B)) can be used to approximately calculate
the tilt angle of the DPPC hydrophobic tails. In
the condensed state, this tilt angle was estimated
to be 29.6°, which agrees with the data reported
for this phospholipid by other authors [44–46].
This data also agrees with that deduced directly
from Synchroton grazing incidence X-ray diffraction for D-DPPC [47] and L and DL-DPPC [48]
monolayers in the range of 30–45 mN m − 1.
Fig. 3. Schematic representation of the DPPC polar group at
the air– water interface, as encountered in the liquid condensed
state (A) and in the liquid expanded state (B).
molecules, sinking further into the subphase. As a
result, it can be postulated that the phosphorylcholine and the glycerol polar groups are deeply
submerged into the subphase at low surface pressures, and that, as the film is compressed, they
emerge to the interface due to the expulsion of the
water solvation (Fig. 3). This change in the penetration of the polar group in the subphase can be
attributed to the appearance of the LE– LC phase
transition in DPPC monolayers. In fact, the
change in the relative film thickness in the LE–
LC transition region moves from 10.7 to 16.5 A, ,
in accordance with calculations carried out using
Eq. (1) with the data in Fig. 1(B) in combination
with the theoretical value for the length of the
hydrocarbon palmitoyl chain which was estimated
to be 19 A, — keeping in mind that the length of
the CC link is 1.53 A, and that the angle between
two neighboring CC bonds is 112° [42]. The
increase in relative thickness (5.8 A, ) along the
LE – LC phase transition corresponds to the in-
3.2. Structural characteristics of DPPG
monolayers
The time evolution of the relative reflectivity
and the surface pressure during a compression–
expansion cycle for the DPPG monolayer spread
on water (pH 6) is shown in Fig. 4(A). It can be
seen that the relative reflectivity is practically constant at the beginning of compression— as the
surface pressure is practically zero— with the
presence of reflectivity peaks as a consequence of
the coexistence of the LC domains (bright regions
in Fig. 5(A)) dispersed in the expanded (gaseous)
phase (dark zones). Upon compression, the reflectivity increases markedly, and the monolayer is
nearly covered with LC domains (Fig. 5(B) and
(C)). Under these conditions optical anisotropy is
observed by turning the analyzer angle to 50°
(Fig. 5(B%)). As the monolayer is in the LC state,
the reflectivity peaks decreases progressively as
the film is compressed, and it disappears completely at higher surface pressures as the monolayer reaches the solid (S) state (Fig. 4(A)).
At a surface pressure of 36 mN m − 1, a homogenous image sprinkled with bright nuclei of
condensation is observed (Fig. 5(D)), which de-
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J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
Fig. 4. (A) Time evolution of relative reflectivity () and surface pressure () during a compression – expansion cycle (shutter speed,
125 s − 1), and (B) y – A isotherm for a DPPG monolayer spread on the air – water interface at pH 6 and at 20 °C.
J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
Fig. 5.
281
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J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
notes the beginning of the monolayer collapse via
a nucleation mechanism. The y – A isotherm shape
(Fig. 4(B)) is characteristic of a condensed mono1
layer with a compressional modulus, C −
s , of 180
−1
−1
mN m —C s = − A(dy/dA), where dy/dA is
the slope of the y– A isotherm. At the monolayer
collapse, the nuclei of condensation merge together to form long stripes (Fig. 5(E)), demonstrating its fracture. The collapse pressure was 62
mN m − 1.
There are no reflectivity peaks in the solid
phase during monolayer expansion (Fig. 4(A)). Its
presence is observed as the film recovers the LC
state. The relative reflectivity during monolayer
expansion is higher than that during compression.
At the maximum area, as the monolayer returns
to its initial state, the relative reflectivity is of
approximately 8.1× 10 − 7. This is due to the fact
that the liquid-condensed structure is maintained
during expansion as is shown in the BAM image
at the final expansion stage (Fig. 5(F) at y= 0.1
mN m − 1). The optical anisotropy of these domains demonstrates the existence of molecules
with different orientations (Fig. 5(F) and (F%)). In
summary, the compression– expansion process is
not completely reversible within the time of the
experiment, because, during monolayer expansion
there exists a relaxation time and the monolayer
maintains the LC structure even at the end of the
expansion process. That is, at a microscopic level,
the morphological features (Fig. 5) and the reflectivity (Fig. 4(A)) of DPPG monolayers are not the
same during the compression-expansion cycle, although, the shape of the y – t isotherm is practically the same (Fig. 4(A)).
Using a similar procedure to that previously
described for DPPC, the relative thickness and tilt
angle of the molecules in the E– LC and LC states
can also be estimated. The relative reflectivity at
the beginning of compression was 4× 10 − 7 (Fig.
4(A)), which means that the relative film thickness
and tilt angle in the E– LC transition region are
11.5 A, and 52.8°, respectively. At the beginning of
the LC phase, the relative reflectivity was 1.2×
10 − 6. The film thickness and the tilt angle at the
beginning of the LC phase was calculated to be
16.5 A, and 29.6°. That is to say that, throughout
the E–LC region, the DPPG molecules in the
expanded state progressively straighten up into
vertical position. This change of orientation explains why the surface pressure does not vary
during the E–LC transition, in spite of the fact
that DPPG in this region are in physical contact,
forming LC domains that become more compact
as the film is compressed (Fig. 5(A) and (B)).
3.3. Structural characteristics of DPPS
monolayers
The time evolution of the relative reflectivity
and the surface pressure during the compression–
expansion cycle of DPPS film is shown in Fig.
6(A). At a high molecular area, and at surface
pressures close to zero, the relative reflectivity is
the same as that found for pure water until it
increases abruptly during monolayer compression,
achieving a value around 1.75× 10 − 6. This value,
observed even at a zero surface pressure, is higher
than that for DPPC and DPPG in their more
condensed structures. The change in the relative
reflectivity corresponds to an increase in the
DPPS film’s thickness of approximately four
times. Throughout this region, the DPPS expanded monolayer presents some reflectivity
peaks of pronounced intensity. This is due to the
existence of irregularly shaped solid domains of
considerable size at the interface (Fig. 7(A), at
y= 0.1 mN m − 1), which merge as the monolayer
is compressed, thereby inducing a compact structure broken up by low intensity streaks (Fig. 7(B),
at y= 0.5 mN m − 1). The monolayer islands are
isotropic, as is shown in Fig. 7(B%), which were
obtained under the same experimental conditions
Fig. 5. Visualization by BAM of DPPG monolayers spread on the air – water interface at pH 6 and at 20 °C. (A) LC domains at
0.1 mN m − 1; (B) LC domains at 1 mN m − 1 without analyzer; (B%) LC domains at 1 mN m − 1 for a position p of analyzer relative
to the plane of incidence of 50°; (C) and (D) LC domains at 7.5 mN m − 1 and homogeneous structure at 36 mN m − 1, respectively;
(E) monolayer collapse; (F) expansion of the monolayer at the maximum area (y = 0.1 mN m − 1); (F%) expansion of the monolayer
at the maximum area for a position p of analyzer of 70°.
J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
283
Fig. 6. (A) Time evolution of relative reflectivity () and surface pressure () during a compression – expansion cycle (shutter speed,
125 s − 1), and (B) y – A isotherm for a DPPS monolayer spread on the air – water interface at pH 6 and at 20 °C.
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J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
Fig. 7. Visualization by BAM of DPPS monolayers spread on the air – water interface at pH 6 and at 20 °C. (A) solid domains at
0.1 mN m − 1; (B) solid domains at 0.5 mN m − 1; (B%) solid domains at 0.5 mN m − 1 for a position p of analyzer of 60°. (C)
homogeneous structure at 19 mN m − 1; (D) monolayer collapse. (E) solid domains at the end of monolayer expansion.
J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
than that for Fig. 7(B) by turning the analyzer
angle to 60°. As the film enters into the solid
state, a homogeneous image is obtained (Fig.
7(C), at y = 19 mN m − 1) and its appearance does
not vary throughout this state until the monolayer
collapse is attained via a bulk fracturing mechanism, in which the monolayer cracks cooperatively over large length scales (Fig. 7(D)). The
collapse surface pressure is 60 mN m − 1 (Fig.
6(B)) and the molecular area at the collapse point
is 38.6 A, 2 per molecule, a value similar to that of
39 A, 2 per molecule obtained by Van Deenen et al.
[49] for DL-distearoyl phosphatidylserine spread
on a phosphate buffer of pH 7.4.
The relative reflectivity during monolayer expansion is practically symmetrical in relation to
that for compression (Fig. 6(A)), which demonstrates the reversibility of the process. Thus, the
BAM image obtained at the end of expansion
(Fig. 7(E)) is similar to that registered at the
beginning of compression (Fig. 7(B)). The reflectivity peaks observed (Fig. 6(A)) in the expanded
state during monolayer compression and, especially, during expansion, are due to solid domains
of DPPS passing through the illuminated spot.
Moreover, the abrupt increase in relative reflectivity upon compression—even at a zero surface
pressure—is due to the monolayer material collected in front of the moving barrier as it reaches
the light spot. A similar behavior was observed by
Hönig and Möbius [11] for arachidic acid.
DPPS is isoelectric at pH 1.5 [50,51]. In the pH
range between 1.5 and 5.2, the ionization of the
carboxylic group takes place [31] and an apparent
pK of 4.2 is obtained [52]. In the pH region
between 5.2 and 9.4, the three DPPS ionizable
groups are completely charged. At pH\ 9.4, the
group NH+
is discharged and the DPPS
3
molecule acquires a double negative charge. Accordingly, it can be postulated that at pH 6,
intermolecular attraction forces are established
between the carboxylic and ammonium charged
groups of the close molecules, and, as a result, the
alkyl chains exert attractive forces among themselves that are strong enough to cause the spontaneous assembly of molecules at a zero surface
pressure on water, instead of an ideal gaseous
phase where the molecules are independently
285
spread on the interface with their alkyl chains
flexing freely. This phenomenon explains the existence of irregular, isotropic domains (Fig. 7(A)),
which are different from those of DPPC, with
their circular and compact domains (Fig. 2), that
result from the lower intermolecular attractions
between the zwitterionic polar groups.
Acknowledgements
This work was supported by the Consellerı́a of
Education of the Xunta de Galicia (Spain) under
Project PGIDT99PXI20302B and by DGICYT
through grant PB97-0734.
References
[1] G. Zubay, Biochemistry, second ed., Macmillan, New
York, 1988, pp. 154 – 210.
[2] M.I. Gurr, J.L. Harwood, in: Lipid Biochemistry and
Introduction, fourth ed., London, 1991. Chapman and
Hall (Eds.).
[3] D.E. Vance, J.E. Vance, New Comprehensive Biochemistry, vol. 20, Elsevier, New York, 1991.
[4] M.C. Phillips, D. Chapman, Biochim. Biophys. Acta 163
(1968) 301.
[5] H.M. McConell, Annu. Rev. Phys. Chem. 42 (1991) 171.
[6] H. Möhwald, Annu. Rev. Phys. Chem. 41 (1990) 441.
[7] R.F. Cohn, J.W. Wagner, J. Kruger, Appl. Opt. 27 (1988)
664.
[8] D. Beaglehole, Rev. Sci. Instrum. 59 (1988) 2557.
[9] R. Reiter, H. Motschmann, H. Orendi, A. Nemetz, W.
Knoll, Langmuir 8 (1992) 1784.
[10] S. Hénon, J. Meunier, Rev. Sci. Instrum. 62 (1991) 936.
[11] D. Hönig, D. Möbius, J. Phys. Chem. 95 (1991) 4590.
[12] J.M. Rodrı́guez Patino, C.C. Sánchez, M.R. Rodrı́guez
Niño, Langmuir 15 (1999) 2484.
[13] J.M. Rodriguez Patino, C.C. Sánchez, M.R. Rodrı́guez
Niño, Food Hydrocolloids 13 (1999) 401.
[14] R.M.A. Azzam, N.M. Bashara, Ellipsometry and Polarized Light, first ed., De North-Holland, Amsterdam,
1992.
[15] J.M. Rodrı́guez Patino, C.C. Sánchez, M.R. Rodrı́guez
Niño, Langmuir 15 (1999) 4777.
[16] J.M. Rodrı́guez Patino, C.C. Sánchez, M.R. Rodrı́guez
Niño, J. Agric. Food Chem. 47 (1999) 4998.
[17] L.W. Horn, N.L. Gershfeld, Biophys. J. 18 (1977) 301.
[18] M. Matsumoto, Y. Tsujii, K.I. Nakamura, T. Yoshimoto,
Thin Solid Films 280 (1996) 238.
[19] C.W. Mc Conlogue, D. Malamud, T.K. Vanderlick,
Biochim. Biophys. Acta 1372 (1998) 124.
286
J. Miñones, Jr et al. / Colloids and Surfaces A: Physicochem. Eng. Aspects 203 (2002) 273–286
[20] V. Vié, N. Van Mau, E. Lesniewska, J.P. Goudonnet, F.
Heitz, C. Le Grimellec, Langmuir 14 (1998) 4574.
[21] P. Lucham, J. Wood, S. Frogatt, R. Swart, J. Colloid.
Interf. Sci. 156 (1993) 164.
[22] T.H. Chou, C.H. Chang, Langmuir 16 (2000) 3385.
[23] V.L. Shapovalov, Thin Solid Films 599 (1998) 327 –329.
[24] S. Pathirana, W.C. Neely, V. Vodyanoy, Langmuir 14
(1998) 679.
[25] B.M. Discher, W.R. Schief, V. Vogel, S.B. Hall, Biophys.
J. 77 (1999) 2051.
[26] U. Dahmen-Levison, G. Brezesinski, W. Möwhald, Prog.
Colloid. Polym. Sci. 110 (1998) 269.
[27] D. Vollhardt, Adv. Colloid. Interf. Sci. 64 (1996) 143.
[28] C. Naumann, C. Dietrich, J.R. Lu, R.K. Thomas, A.R.
Rennie, J. Penfold, T.M. Bayerl, Langmuir 10 (1994)
1919.
[29] R.M. Weis, H.M. Connell, Nature 310 (1984) 47.
[30] J. Seelig, Q. Rev. Biophys. 10 (1977) 353.
[31] H. Hauser, M.C. Phillips, in: J.F. Danielli, M.D. Rosenberg, D.A. Cadenhead (Eds.), Progress in Surface and
Membrane Science, vol. 13, Academic Press, New York,
1979, p. 297.
[32] M. Saundaradingham, Ann. New York Acad. Sci. 195
(1955) 324.
[33] F.A. Vilallonga, E.R. Garret, M. Cereijido, J. Pharm. Sci.
61 (1972) 1720.
[34] D.O. Shah, J.H. Schulman, J. Colloid. Interf. Sci. 25
(1967) 107.
[35] H. Hauser, I. Pascher, R.H. Pearson, S. Sundell, Biochim.
Biophys. Acta 650 (1981) 21.
[36] J. Miñones, M.I. Sández, E. Iribarnegaray, P. Sanz, Colloid. Polym. Sci. 259 (1981) 382.
[37] J. Miñones, M.I. Sández, E. Iribarnegaray, P. Sanz, Colloid. Polym. Sci. 259 (1981) 460.
[38] J. Miñones, L. Cid, O. Conde, An. Quim. 82 (1986) 340.
[39] J. Miñones, L. Cid, E. Iribarnegaray, Colloid. Polym. Sci.
266 (1988) 337.
[40] H. Baltes, M. Schwendler, C.A. Helm, H. Möhwald, J.
Colloid. Interf. Sci. 178 (1996) 35.
[41] H.V. Gally, W. Niederberger, J. Seelig, Biochem. J. 14
(1975) 3647.
[42] D. Small, in: J. Hanaban (Ed.), The Physical Chemistry
of Lipids, second ed., Plenum Press, New York, 1988, p.
22.
[43] T.M. Bayerl, R.K. Thomas, D. Penfold, A. Rennie, E.
Sackmann, Biophys. J. 57 (1990) 1095.
[44] D. Vakinin, K. Kjaer, J. Als-Nielsen, M. Lösche, Biophys. J. 59 (1991) 1325.
[45] M. Carvell, D. Hall, J.G. Lyle, G.T. Tiddy, Faraday
Discuss. Chem. Soc. 81 (1986) 223.
[46] C. Naumann, C. Dietrich, J.R. Lu, R.K. Thomas, A.R.
Rennie, J. Penfold, T.M. Bayerl, Langmuir 10 (1994)
1919.
[47] U. Dahmen-Levison, G. Brezesinski, H. Möhwald, Thin
Solid Films 616 (1998) 327 – 329.
[48] G. Brezesinski, A. Dietrich, B. Struth, C. Böhm, W.G.
Bouwman, K. Kjaer, H. Möhwald, Chem. Phys. Lipids
76 (1995) 145.
[49] L.L.M. Van Deenen, U.M.T. Houtsmuller, G.H. de
Haas, E. Mulder, J. Pharm. Pharmacol. 14 (1962) 429.
[50] M.B. Abramson, R. Katzman, H.P. Gregor, J. Biol.
Chem. 239 (1964) 70.
[51] H. Hauser, A. Darke, M.C. Phillips, Eur. J. Biochem. 62
(1976) 335.
[52] T. Seimiya, S. Ohki, Nat. New Biol. 239 (1972) 26.