BIOMOLECULAR NETWORKS FOR NOVEL PROTEIN-POWERED ACTIVE MATERIALS
Andy Sarles and Donald J. Leo, Ph.D.
Center for Intelligent Material Systems and Structures (CIMSS)
Dept. of Mechanical Engineering, Virginia Tech, Blacksburg, VA 24061
ABSTRACT
Biological molecules including phospholipids and
proteins offer scientists and engineers a diverse
selection of materials to develop new types of active
materials and smart systems based on ion conduction.
The inherent energy-coupling abilities of these
components create novel kinds of transduction
elements. Networks formed from droplet-interface
bilayers (DIB) are a promising construct for creating
cell mimics that allow for the assembly and study of
these active biological molecules. The current-voltage
relationship of symmetric, “lipid-in” droplet-interface
bilayers are characterized using electrical impedance
spectroscopy (EIS) and cyclic voltammetry (CV).
“Lipid-in” diphytanoyl phosphatidylcholine (DPhPC)
droplet-interface bilayers have specific resistances of
nearly 10MΩ·cm2 and rupture at applied potentials
greater than 300mV, indicating the “lipid-in” approach
produces higher quality interfacial membranes than
created using the original “lipid-out” method. The
incorporation of phospholipids into the droplet interior
allows for faster monolayer formation but does not
inhibit the self-insertion of transmembrane proteins into
bilayer interfaces that separate adjacent droplets.
Preliminary results on the encapsulation of DIB
networks within a curable matrix are also presented.
This work aims to create a durable and portable
material system that features the functionality of
biomolecules within DIB networks.
INTRODUCTION
In nature, plant and animal cells are composed of
cell membranes that define the outer boundary of the
cell and divide the inner volume of each cell into
internal compartments, called organelles.
The
composite structure of the cell membrane (featuring
both phospholipid molecules arranged in a bilayer
configuration and proteins) allows it to simultaneously
block diffusion and enable selective transport of species
into and out of the cell [1]. Selective transport of
species through this ultra-thin (5-7nm thick)
phospholipid membrane occurs via transmembrane
proteins that bind to and reside within the cell
membrane. Proteins are biomolecules that carryout
nearly all of the functions of the cell and, while most of
these functions depend on the ability of the protein to
recognize specific molecules (called ligands) through
binding [2, 3], proteins also serve as structural
elements, such as keratin in hair and fibroin in silk, they
are enzymes such ATPase that hydrolyze chemical
energy, and when residing in the cell membrane they
transport matter (pumps and ion channels) as well as
send and receive information [1, 4]. Proteins offer a
diverse selection of energy coupling abilities, including
photochemical, mechanochemical, chemomechanical,
and chemoelectrical transformation, and the behavior of
each protein is directly tied to its chemical composition.
The inherent transduction properties of biomolecules,
including proteins, enable the development of a new
class of active materials.
Making use of the inherent functions of biomolecules
hinges on the ability to host these molecules in an
environment similar to that found in the cell. A bilayer
lipid membrane (BLM), or lipid bilayer, composed of
amphiphilic phospholipid molecules mimics the
structure of a cell membrane and provides a suitable
host structure for reconstituting proteins [4-6].
Recently, interest in using biological molecules, such as
phospholipids and proteins, has led to the development
of several types of bio-inspired materials and devices:
chemical sensors [7-10], rotary motors [11-13], biotransistors [14], and chemoelectric energy-conversion
prototypes [15]. These works feature proteins as active
elements that were reconstituted into an artificial
bilayer lipid membrane (BLM) formed on either a solid
support or suspended across one or many pores in a
synthetic material. Reimhult and Kumar in their recent
review article of bilayer-based sensing platforms
formed from micro- and nanoporous substrates,
however, suggest that many of the current bilayer-based
technologies for creating functional devices are limited
by the fragility of the bilayers for long-term use, the
inability to effectively reconstitute bilayers across a
large number of pores to prevent electrical leakage
currents, and the fact that supported bilayer systems
quickly dehydrate and no longer work in air [9]. An
observation of these specific works also indicates that
the state-of-the-art device concepts contain only one
type of protein capable of usually one task. The ability
to integrate the functions of many proteins, each within
their own lipid bilayer, into one device provides
additional challenges. The droplet-interface bilayer
(DIB) method developed by researchers at Oxford and
Duke [16-18] offers several unique possibilities that are
unavailable using supported bilayers and avoids many
of the issues related to the formation of lipid bilayers on
synthetic substrates mentioned above.
contained a different type of phospholipid [18]. Our
own experience in forming droplet interface bilayers
shows that the incorporation of phospholipids as
vesicles into the water phase promotes faster lipid
monolayer formation than observed for “lipid-out”
droplet interface bilayers, in which phospholipids are
dissolved in the organic solvent [19, 20].
The aqueous droplets contain 1,2-diphytanoyl-snglycero-3-phosphocholine
(DPhPC)
phospholipid
vesicles (purchased as lyophilized powder from Avanti
Polar Lipids, Inc.) dissolved in 10mM MOPS (Sigma),
100mM NaCl (Sigma), pH7 buffer solution. The
phospholipid vesicles were prepared as described
elsewhere [18]and were stored at 3-8°C for several
weeks between tests. The formation of “lipid-in”
droplet interface bilayers begins by filling a well
machined into a poly(methyl methacrylate) (PMMA)
tray filled with hexadecane (Sigma) (Figure 2). Then,
300nl aqueous droplets containing DPhPC vesicles in
buffer solution are pipetted into the organic solvent. A
silver-silver chloride (Ag/AgCl) electrode coated with
hydrophilic agarose gel (Sigma) is inserted into each
droplet for subsequent positioning and electrical
measurements. Initially, the electrodes are positioned
such that the droplets do not touch, in order to allow for
the self-assembly of lipid monolayers at the
water/hexadecane interface surrounding each droplet.
Following 5-10 minutes for monolayer formation, the
electrodes are repositioned in order to bring the droplets
into physical contact. The spontaneous formation of a
droplet interface bilayer occurs within a few minutes
once excess solvent is removed from between the
droplets, permitting the monolayers to “zip” together.
In this report we present experimental results that
further the development of a new class of smart
material (Figure 1) that relies on the transport properties
of biomolecules as the transduction mechanism. We
build upon the methods of DIB formation [17, 18] and
characterization [19, 20] established in previous works
and show that the incorporation of biomolecules, such
as alpha-hemolysin and alamethicin proteins, into DIB
networks alters the current-voltage relationship of the
lipid membranes.
Preliminary results on the
encapsulation of droplet-interface bilayers within a
curable matrix are also presented. Together, these
ventures aim to develop portable and durable
biomolecular material systems that employ the inherent
transduction properties of biomolecules.
EXPERIMENTAL METHODS
Droplet interface bilayers (DIBs) are formed using
the procedures developed by Holden, et al [16-18]. In
this paper, “lipid-in” droplet interface bilayers are
formed by incorporating the phospholipid molecules
into the water phase that constitutes each droplet.
Hwang, et al first demonstrated this version of dropletinterface bilayers in order to create asymmetric DIBs,
where each droplet, and thus each half of the bilayer,
Inputs
Δκ, where
κ (input)pH, V, [C], etc.
force
DIB network
light
sound
heat
biomolecule A
biomolecule B
biomolecule C
species 1
species 2
i(t)
single BLM
Outputs
signal
signal
current
current
ΔV
voltage
voltage
time
time
Figure 1: Active materials that utilize the transport properties of biomolecules are constructed from networks of dropletinterface bilayers. Proteins incorporated into specific bilayer interfaces in the network determine the input/output
transduction relationship to multiple stimuli.
2009 VSGC Conference
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2
RESULTS AND DISCUSSION
Characterization of Lipid-only DIBs
3
4
step 3
step 4
800µm
Figure 2: Droplet-interface bilayers are formed when
aqueous droplets (300nl each) are pipetted (2) into a
nonpolar solvent such as hexadecane within a well (1). The
droplets are pierced with agarose-tipped Ag/AgCl
electrodes (3) and, after several minutes to allow for lipid
monolayer assembly at the oil/water interface around each
droplet, they are positioned in intimate contact (4). A
droplet-interface bilayer forms when excess organic
solvent is removed from between the droplets, allowing
the monolayers to zip together in order to stabilize the
interface and prevent droplet fusion.
Electrical impedance spectroscopy (EIS) and cyclic
voltammetry (CV) are used to characterize the electrical
properties of droplet-interface bilayers with and without
proteins.
An Autolab PGSTAT12 (Eco Chemie)
Potentiostat/Galvanostat with a FRA2 module
controlled by FRA and GPES software is used to
perform these tests.
Electrical impedance
measurements of the DIBs are performed using a 5mV
(RMS) sinusoidal potential swept from 500kHz to
10mHz. CV measurements are conducted with a scan
rate of 2.5mV/s, using voltage steps of 0.3mV. A 3axis stepper-motor controlled micromanipulator
(SM325, WPI Inc.) is used to move one of the
electrodes for precise droplet positioning throughout the
study. The entire test fixture is mounted on a Zeiss
Axiovert 40CFL inverted microscope and images of the
droplet networks on the microscope are obtained with a
Canon G6 digital camera and digital camera adapter. A
homemade Faraday cage surrounding the entire
workstation was grounded through the Autolab device
to an earth ground connection in order to minimize
electrical noise during measurements.
2009 VSGC Conference
“Lipid-in” droplet-interface bilayers are formed by
incorporating phospholipid vesicles into the interior of
the droplets. The formation of a lipid bilayer at the
interface of adjacent water droplets is confirmed using
electrical impedance spectroscopy (EIS) and cyclic
voltammetry (CV) measurements.
The electrical
resistance and capacitance of the bilayer are estimated
from EIS data by fitting an equivalent electrical model
for a single DIB consisting of a resistor in parallel with
a capacitor representing the bilayer and a series
resistance attributed to the resistance of the electrolyte
solution within each droplet. The data presented in
Figure 3 show a lipid bilayer with an initial resistance
of 2.5GΩ that increases to more than 14GΩ by the third
measurement. The capacitance of the bilayer is
estimated to be 11-12pF and, by using published values
of specific capacitance for DPhPC lipid bilayers
(~0.6μF/cm2, [21, 22]), the equivalent diameter of the
interface is computed to be 50μm.
Magnitude - 
2
10
10
10
5
10
0
2
10
Frequency - Hz
10
4
0
Phase - deg
1
-50
-100
10
0
2
10
Frequency - Hz
10
4
50
45
40
DIB rupture
35
Current - pA
3mm
30
25
20
15
10
5
0
-200
-100
0
100
Voltage - mV
200
300
400
Figure 3: Representative electrical impedance (top) and
cyclic voltammetry (bottom) data measured on a single
“lipid-in” droplet-interface bilayer.
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3
curve increases greatly. Incorporating alamethicin
proteins into both droplets produces a symmetric
voltage-dependent conductance (black trace). The
measured current traces shows that the voltagedependent change in conductance is reversible; bilayers
containing alamethicin resume a state of high resistance
for applied potentials approaching 0V.
20000
2 droplets, 1 DIB
Va
10000
5000
2009 VSGC Conference
Va
0
No ALm
-5000
EIS measurements for the three cases show a very
similar impedance signature for droplet-interface
bilayers measured using a small applied potential (5mV
RMS). Each of the three trials produced a bilayer with
an electrical resistance greater than 10GΩ and an
interface ranging from 250-350μm in diameter. Cyclic
voltammetry measurements, however, show that
differences in the current flowing through the interfaces
arise as a function of voltage. Figure 4 shows that the
conductance (measured by the slope of the currentvoltage curve) of a bilayer containing alamethicin
increases when the voltage of the droplet containing
alamethicin is at a potential approximately +60mV
relative to the opposite side. For example, when
alamethicin is incorporated into the right droplet in a
two-droplet network and the voltage in this droplet is
higher than 60mV, the slope of the current-voltage
Va
ALm in both droplets
Alamethicin Incorporated into DIBs
Alamethicin (A.G. Scientific) is stored in ethanol at
0.1% (w/v) and this stock solution is diluted further to a
concentration of 100μg/ml alamethicin in 10mM
MOPS, 100mM NaCl, pH~7 buffer solution. Droplets
containing alamethicin consist of a 50-2 (volume ratio)
vesicle solution-protein solution mixture. The first set
of experiments focuses on the current-voltage response
of alamethicin within single droplet-interface bilayers
formed from two droplets. Figure 4 shows the results
of the EIS and CV measurements on three different
single DIBs: without alamethicin in either droplet, with
alamethicin in only one droplet, and with alamethicin in
both droplets.
ALm in one droplet
15000
Current - pA
Cyclic voltammetry measurements provide an
additional tool for characterizing voltage-dependent
phenomena occurring within the bilayer. As shown in
Figure 3, the current-voltage relationship of a pure
DPhPC bilayer without proteins is highly linear for
applied potentials up to the observed failure potential at
approximately 330mV. The failure potential is marked
by a sharp, irreversible increase in current measured
when the bilayer ruptures and the droplets to coalesce.
Subsequent electrical measurements on the system
show the response of the electrolyte resistance since the
lipid membrane that previously stabilized the interface
no longer exists. Additional lipid bilayers formed at the
interface of two 300nl droplets are characterized using
EIS and CV. The measured capacitance of each
interface is approximately 360pF for an equivalent
interface diameter of 260μm. The average specific
resistance measured for lipid bilayers is 7MΩ·cm2 (n =
9), while the average failure potential recorded for a
subset of these trials is 340mV (n = 4).
-100
-50
0
50
100
Voltage - mV
150
200
250
Figure 4: Electrical impedance and cyclic voltammetry
measurements on single, “lipid-in” droplet-interface
bilayers with and without proteins. The blue (circles)
traces refer to a lipid-only DIB, the red (squares) traces
were measured for a two-droplet network that contained
alamethicin (ALm) proteins in the right droplet, and the
black (triangle) traces refer to alamethicin incorporated
into both droplets. The arrows in the sub-figures indicate
the direction of insertion.
Alamethicin proteins are also incorporated into three
droplets, two-bilayer networks. Four configurations are
tested: three droplets without alamethicin, alamethicin
incorporated into the end droplet, alamethicin added to
the central droplet, and alamethicin proteins in all three
droplets.
The incorporation of alamethicin into larger
biomolecular networks is affected by the presence of
additional resistive lipid membranes in the circuit.
Again, EIS measurements (not shown) are performed to
verify bilayer formation prior to the CV measurements.
In all four trials, the electrical impedance at low
frequency (<10mHz) approaches 100GΩ while the
high-frequency (>100kHz) asymptote of the measured
impedance signature reflects the resistance of the
electrolyte--implying that both droplet interfaces are
lipid bilayers. The current measurements shown in
Figure 5 indicate that only when alamethicin proteins
insert into all interfaces in the same direction (e.g. all
interfaces have proteins oriented the same way) does
the conductance of the membrane change significantly
Sarles
4
respectively) appear to confirm this behavior, though
this comparison does not account for unequal bilayer
resistance in the OFF state or different amounts of
protein reconstituted into each interface.
5000
4000
150
1. no ALm
100
Va
3000
Current - pA
as a function of the applied potential. The insertion of
proteins in both directions into both interfaces occurs
when alamethicin is present in all three droplets and
thus the conductance of the three-droplet network is
sensitive to both positive and negative applied
potentials. The directionality of the insertion of the
proteins is indicated in the sub-figures with red arrows.
On the other hand, the incorporation of alamethicin into
only one droplet in the network limits the number of
interfaces containing proteins and dictates the direction
of insertion.
1 DIB
2000
1000
0
-1000
2. ALm in right droplet
4. ALm in all
three droplets
3. ALm in middle droplet
2 DIB
-2000
Va
-3000
Current - pA
50
3 droplets, 2 DIBs
0
50
Voltage - mV
100
150
Encapsulating DIB Networks
-100
-150
-300
-50
Figure 6: Comparison of the voltage-dependent current
measured for a single DIB and a two-DIB network with
alamethicin present in all droplets.
0
-50
-100
-200
-100
0
Voltage - mV
100
200
300
Figure 5: Electrical impedance and cyclic voltammetry
measurements on multiple, “lipid-in” droplet-interface
bilayers with and without proteins.
Only when alamethicin inserts into both interfaces of a
three-droplet network in the same direction does a
significant change in the conductance of the network
occur. Case 4 (Figure 5) illustrates this concept and
shows a symmetric change in conductance about 0V
due to the fact that both interfaces contain proteins
inserted in both directions. The voltage-induced change
in conductance measured for this configuration arises at
larger potentials than were required to trigger
alamethicin proteins in a single DIB. Figure 6 directly
compares the currents measured for a single DIB with
alamethicin in both droplets and a two-DIB network
with alamethicin in all three droplets. Consistent with
expectations, the change in conductance for a two-DIB
network occurs at roughly twice the applied potential-indicating the two lipid membranes share the total
applied potential. The circuit analysis also suggests
that the ON conductance state of a two-DIB network
would be lower than that for a single DIB due to the
additional resistance of the second lipid bilayer in the
line of current flow. The ON resistances of the single
and two-DIB networks (approximated from the data to
be 3.7MΩ at +70mV and 5.5MΩ at +130mV,
2009 VSGC Conference
A bilayer lipid membrane is formed from two 1μl
(1.5mm in diameter) droplets of DPhPC lipid vesicle
solution injected into freshly prepared mixture of
PDMS/hexadecane while still in the liquid state.
Electrical impedance measurements of a lipid bilayer
throughout the curing of the PDMS/hexadecane mixture
indicate that this membrane survives during the curing
of the surrounding encapsulant.
Impedance
measurements on the bilayer during this time period
show that the bilayer had an initial resistance and
capacitance of 2.8MΩ and 1.83nF, respectively.
Successive EIS measurements plotted in Figure 7:
Electrical impedance measurements on the DPhPC
bilayer formed at the interface of two droplets were
taken through the 2-day PDMS/hexadecane curing
cycle. Estimates of the membrane resistance (MΩ·cm2)
and capacitance (nF) are obtained by fitting an
equivalent circuit to the successive measurements.show
that resistance then decreases before beginning to
increase several hours after initial BLM formation.
Conversely, the measured capacitance of the lipid
bilayer doubles to a value of nearly 4nF hours after the
first measurement.
The changes in the measured electrical properties
indicate that the size and the quality of the membrane
increase during cure. The measured capacitance from
EIS data and the reported specific capacitance of
DPhPC bilayers (0.6μF/cm2 [21, 22]) are used to
estimate the area and equivalent diameter of the lipid
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5
bilayer separating the two droplets. The calculations
show that the bilayer grows from approximately 600μm
in diameter initially to closer to 900μm in diameter at
the end of cure. This change in area was also
confirmed by the decrease in the resistance of the
membrane within the first few hours after bilayer
formation. Later on in the curing cycle, the size of the
bilayer stabilizes as indicated by the capacitance
measurements, but the measured electrical resistance
increases by an order of magnitude. This result shows
that throughout curing, even after bilayer formation, the
lipid vesicles within the droplets continue to supply the
bilayer with additional lipid molecules. And as more
phospholipids assemble at the interface, the electrical
barrier between the two aqueous compartments
strengthens.
Figure 8: The substantial change in the size of the bilayer
at the interface is also captured in photographs taken with
a digital camera through the eyepiece of a low-power
stereo-microscope. The picture on the left was taken
approximately 15 minutes after bilayer formation, while
the picture on the right shows the DIB after the
PDMS/hexadecane mixture fully cured.
10
Magnitude - 
10
lipid bilayer
5
10
after bilayer failure
0
10
-2
Phase - deg
10
0
2
10
10
Frequency - Hz
4
6
10
10
Figure 8 illustrates the changes that the dropletinterface bilayer underwent during the curing cycle.
The size of the bilayer increased and in doing so, the
two droplets pulled away from the two Ag/AgCl
electrodes used for initial positioning as they were
drawn together. This dramatic change in size and shape
was not seen previously in any of the experiments
forming DPhPC DIBs within pure hexadecane.
-20
-40
-60
-80
0
2
10
10
Frequency - Hz
1000
4
10
10
100
1
10
1
Measured Capacitance - nF
Membrane Resistance - MΩ∙cm2
Resistance
0.1
1
100
10000
1000000
Time - s
Figure 7: Electrical impedance measurements on the
DPhPC bilayer formed at the interface of two droplets
were taken through the 2-day PDMS/hexadecane curing
cycle. Estimates of the membrane resistance (MΩ·cm2)
and capacitance (nF) are obtained by fitting an equivalent
circuit to the successive measurements.
2009 VSGC Conference
Interactions between the hydrophobic acyl regions of
lipid membranes and the oil phase have been studied on
both the formation of lipid monolayers [23, 24] and on
supported bilayer lipid membranes [25]. Needham and
Haydon, in their measurements on the tensions and
contact angles at the Plateau-Gibbs border of supported
lipid bilayers, found that bilayers formed from lipid
solutions containing squalene exhibited lower tensions,
higher contact angles, and thinner hydrophobic regions
than when hexadecane or decane were used. The
authors attributed this trend to the size of the solvent
molecule, stating that the phospholipid chains constrain
the larger solvent molecules to lie flat in the center of
the bilayer, which is unfavorable in terms of freeenergy. Smaller solvent molecules, then, are more
likely to reside in free space between the lipid acyl
chains, where they limit the packing density of the
lipids in the monolayer and cause an increase in the
surface tension at the oil/water interface.
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The size of the solvent molecule affects the surface
tension of the lipid monolayers in the same way in the
formation of unsupported droplet-interface bilayers.
Smaller solvent molecules increase the interfacial
tension within the lipid monolayers at the oil water
interface, while larger, bulkier molecules are forced out
more effectively resulting in a lowered monolayer
tension.
   BLM   LM   LM cos  1
where, γBLM, is the surface tension of the bilayer
interface and γLM is the surface tension of the lipid
monolayer that encases both droplets (Note: It is
assumed for this analysis that γLM is constant).
R
A 
x
γLM
r
A
  LM cos   1 .
12 2
R
Fzip
d
θ
*Section view of symmetric DIB
Figure 9: The size of the interface is determined by the
amount of free-energy available to two separated droplets
“zipping” together to form a DIB pair, characterized by a
energy balance on the membrane.
A simple energy balance (per unit area) on a single
droplet-interface bilayer explains how changes in
monolayer surface tension translate into different
interfacial size. Figure 9 is an illustration of a dropletinterface bilayer formed between two identical droplets.
When symmetry is considered, the cross-sectional view
of the connected droplets at the interface can be drawn
in order to visualize the forces at play. The free-energy
of thinning of the bilayer is the decrease in Helmholtz
energy in the system as the opposing lipid monolayers
are drawn together by short-range London-van der
Waals forces [26]. Feijter and Vrij first determined that
the change in Helmholtz energy per unit area, ΔA, for a
thinning soap film manifests itself as a change in the
surface tension going from the bulk (in our case, the
lipid monolayer tension) to the thinned region (i.e. the
lipid bilayer) [27], given by
A  2 .
2009 VSGC Conference
A
12 2
(3)
where, A is the Hamaker-London-van der Waals
constant (ergs) and δ is the thickness of the
hydrophobic region of the bilayer [25, 26, 28]. Setting
Equations 2 and 3 equal to one another yields
θ
γBLM
(2)
Since the thinning is induced by London-van der Waals
forces that act across the thickness of the planar
hydrophobic region of bilayer, the change in energy per
unit area of the thinned interface can also be written as
d
y
The contact angle, θ, determines how much the lipid
monolayer surface tension and the tension in the bilayer
differ.
(1)
(4)
Equation 4 says that that resulting contact angle at
equilibrium is independent of bilayer area or droplet
size. Instead, the final dimensions of the bilayer
formed between two droplets is a function of the
surface tension of the lipid monolayer encasing each
droplet, γLM, the hydrophobic thickness of the bilayer, δ
and the Hamaker constant for the attraction of water
molecules across a hydrocarbon barrier, A. When the
surface tension of the monolayer is small, a large
contact angle is needed to balance the same energy per
unit area due to London van der Waals attractions.
Conversely, a large surface tension is accompanied by a
small contact angle. Additional geometrical analysis
omitted from this paper shows that an increase in the
contact angle results in a larger interfacial area. This
relationship confirms experimental observations that
the use of a smaller-molecule solvent results in a
smaller double angle between the droplets because of
the increased surface tension at the oil/water interface.
CONCLUSIONS
The ability to construct assemblies of biomolecules
for utilizing the inherent functions of proteins is
achieved using the droplet-interface bilayer technique.
This method relies on the self-assembly of phospholipid
molecules into lipid monolayers at the oil/water
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7
interface of water droplets immersed in a nonpolar
medium such as hexadecane. This technique stands
apart from many of the traditional lipid bilayer
formation techniques in that it allows the creation of
networks of lipid bilayers, each which can contain a
unique composition of biomolecules. Our work shows
that the current-voltage relationships of bilayers within
DIB networks can be tailored using proteins. The
incorporation of alamethicin into DIBs causes lipid
bilayers to exhibit a reversible, voltage-dependent
conductance.
The encapsulation of droplet-interface bilayers for
increasing the durability and portability of this platform
was also studied. The results presented in this paper
show that the size of the molecule of the nonpolar
medium surrounding the droplets directly affects the
surface tension of the lipid monolayer and thus
determines the size and stability of the bilayer interface
between droplets. This study helped to examine the
necessary properties that a candidate curable matrix
must require in order to create a complete material
system that feature the inherent functionality of
biomolecules in DIB networks.
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2009 VSGC Conference
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