Lab on a Chip
View Article Online

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

PAPER

Cite this: Lab Chip, 2019, 19, 3472

View Journal | View Issue

Osmotic-engine-driven liposomes in microfluidic
channels†
Kan Shoji*ab and Ryuji Kawano*aSelf-propelled underwater microrobots that locomote without external sources of energy have potential
application as drug carriers and probes in narrow spaces. In this study, we focused on an osmotic engine
model, which is a migration mechanism, and applied it as a negative chemotaxis mechanism to induce
liposome displacement. First, we confirmed the osmotic flow across the lipid bilayer and calculated theosmotic flow velocity to be 8.5 fL min−1 μm−2 when a salt concentration difference was applied to the lipid
bilayer. Next, we designed and fabricated a microchannel that can trap a giant liposome and apply a salt
concentration difference to the front and rear of the liposome. Then, we demonstrated the movement of
the liposome by flowing it to the microchannel. The liposome successfully moved in the direction of thelower ion concentration at a speed of 0.6 μm min−1 owing to the osmotic pressure difference. Finally, we
Received 9th August 2019,
Accepted 2nd September 2019

visualized the inner flow in the liposome by encapsulating microbeads in the liposome and observed the
movement of the microbeads to verify that an osmotic flow was generated on the liposome. As a result,
we observed the circulation of the microbeads in the liposome when the concentration difference was

DOI: 10.1039/c9lc00788a

applied to the front and rear of the liposome, suggesting that the movement of the liposome was driven
by the osmotic flow generated by the osmotic pressure difference. These results indicate that the osmotic-

rsc.li/loc

pressure-based migration mechanism has the potential to be utilized as the actuator of molecular robots.

Introduction
Bio-integrated robots combining biological objects andmechanical components, such as bioactuators1–3 and cyborg
insects,4–8 have the exclusive abilities of chemical energy
driving, self-renewal capability, biocompatibility, and softness, and they can potentially be used as intracorporeal
microrobots and environmental monitoring robots. Since
conventional robotic systems have not yet provided these abilities, bio-integrated robots have attracted attention as novel
robots. However, differences between biological objects andmachines, such as materials, communication methods, andenergy sources, prevent the development of fully integrated
biorobots. To overcome this issue, interfaces connecting
biological and mechanical objects should be designed anddeveloped. For example, we previously reported a biofuel cellthat generates electric power from insect hemolymph for theenergy source of a mounted radio sensor.7,8aDepartment of Biotechnology and Life Science, Tokyo University of Agriculture
and Technology, 2-24-16 Naka-cho, Koganei-shi, Tokyo 184-8588, Japan.
E-mail: kan-shoji@m2.tuat.ac.jp, shojikn@ucmail.uc.edu, rjkawano@cc.tuat.ac.jpbDepartment of Chemistry, University of Cincinnati, Cincinnati, Ohio 45221, USA† Electronic supplementary information (ESI) available: Liposome formation
method, hydrodynamic simulation, and bead tracking data. See DOI: 10.1039/
c9lc00788a

3472 | Lab Chip, 2019, 19, 3472–3480

On the other hand, the concept of molecular robots
whose components are artificially designed from the molecular level and assembled by bottom-up approaches has recently been proposed,9,10 where a molecular robot is defined
as a microorganism-imitating microrobot. To realize such robots, three essential components are required: sensors,11–15
intelligence,16 and actuators,17–20 and they can be integrated
into a unitary compartment21–26 formed by a giant liposome,gel, and a DNA-based capsule. This cell-imitating microrobots would offer potential not only as a microrobot driving
inside a body for drug delivery and diagnosis but also a biomimetic model of living cells. Molecular robots which cansense chemical signals and migrate to the chemical sourceby chemotaxis would show cell-like behaviors. In this way,molecular robots constructed by synthetic biology should
also attract attention as a biomimetic application. Regarding
the development of actuators, there are two different approaches. One is the use of molecular motors. Molecular motors, such as kinesin and myosin, are capsuled in a giant liposome and deform the liposomal membrane. For example,
filopodium-like protrusions were exhibited by a giant liposome encapsulating microtubes and kinesins.27 Furthermore,a controllable and programmable actuator that responses toa specific DNA signal has been developed by linking singlestrand DNAs to a lipid membrane as an anchor of molecularmotors.19 Although these cell-imitating and molecular-motor-

This journal is © The Royal Society of Chemistry 2019

View Article Online

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

Lab on a Chip

driven actuation mechanisms have potential as biomimetic
aspects of cell migration systems in synthetic biology, it isstill challenging to completely imitate their structures andfunctions.
Another actuation mechanism is a self-propelled mechanism that induces the migration of droplets by converting
external energy to mechanical motion.28–30 These selfpropelled robots are driven by either an asymmetrical chemical reaction at the leading and ending edges of the robot or
the Marangoni effect caused by a surface tension gradient.
These driving mechanisms are straightforward and provide
stable movement compared with that for the above molecularmotor systems. However, few self-propelled mechanisms
using an osmotic pressure gap have been reported because aclosed space must be prepared to maintain a static chemical
gradient for a long time.31
In this study, we focus on the osmotic engine model32
reported by Stroka and coworkers as the cell migration
model. In this model, a polarized cell in which a spatial
gradient of ion channels and pumps in the cell membrane isestablished creates a net inflow and outflow of water andions at the cell leading and trailing edges, respectively. The
net flows create net cell displacement even when the polymerization of actin in the cell is inhibited. By applying thismodel to a liposome, we demonstrate the emergence ofchemotaxis for liposomes and propose its potential as abiomimetic model of cell migration by chemotaxis. Specifically, the semipermeability of the lipid bilayer provides net
flows of water passing through the lipid bilayers owing toosmotic pressure differences at the membranes caused by
the salt concentration differences between the inner andouter solutions. Although the salt concentration differencesat the membranes induce osmotic flows, the salt concentration differences should be generated on the front and rear of
liposomes to demonstrate the chemotaxis (the constantdisplacement of liposomes). Therefore, we utilize microfluidic techniques of cell trapping and gradient generation totrap liposomes and generate a salt concentration differenceat the front and rear of liposomes (Fig. 1). We propose thatthe osmotic-pressure-based propulsion mechanism canpotentially be applied to the engine of molecular robots bydemonstrating liposome displacement induced by an osmotic

Paperpressure difference. Furthermore, this osmotic-pressuredriven liposome would be also useful as a simple cell migration model by chemotaxis.

Experimental
Reagents and chemicals
1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC; Avanti Polar
Lipid), 1,2-dioleoyl-sn-glycero-3-phospho-L-serine (sodium salt)(DOPS;
Avanti Polar Lipid), 1,2-dioleoyl-sn-glycero-3phosphoethanolamine-N-(lissamine rhodamine B sulfonyl)(ammonium salt) (Rhod PE; Avanti Polar Lipid), cholesterol(chol; Avanti Polar Lipid) and liquid paraffin (Wako Pure
Chemical Industries, Ltd.) were used for liposome formation.
DOPC and n-decane (Sigma-Aldrich) were used for the preparation of lipid bilayers for the observation of lipid bilayer
deformation. Potassium chloride (KCl; Nacalai Tesque) and
3-morpholinopropane-1-sulfonic acid (MOPS; Nacalai Tesque)were used as aqueous solutions. The MOPS solution was
prepared using ultrapure water from a Milli-Q system (Merck
Millipore Corp.). Calcein (Tokyo Chemical Industry Co. Ltd.)was utilized as a dye for the fluorescence observation ofliposomes. Fluoresbrite plain microspheres (diameter; 1.0μm, Polysciences, Inc.) were used to observe the inner flow ofliposomes. SU-8 3025 (MicroChem Corp.) was used as aphotoresist. Polydimethylsiloxane (PDMS; Sylgard 184, Dow
Corning Toray Co., Ltd.) was used for the microchannelmaterial.
Observation of membrane deformation due to osmoticpressure differenceFirst, we observed the membrane deformation induced by
the osmotic pressure difference acting on lipid bilayers. We
prepared lipid bilayers by a droplet contact method(DCM)33–35 and observed them under a microscope. In thismethod, a lipid bilayer is formed at the interface between
two droplets in contact with a lipid/oil mixture. We prepareda chamber with the shape of an infinity symbol, in which two
circular wells with an inner diameter of 2 mm overlapped.
Lipid bilayers were prepared by inserting 2.7 μL of aqueous
solution into each well, which had previously been filled with
4 μL of DOPC/n-decane (10 mg mL−1) or DOPC : chol = 7 : 3

Fig. 1 Liposome-based osmotic-pressure-driven microrobot. The liposome trapped in the microchannel migrates in the direction of the lower salt
concentration owing to the osmotic flow generated by the osmotic pressure difference.

This journal is © The Royal Society of Chemistry 2019

Lab Chip, 2019, 19, 3472–3480 | 3473

View Article Online

Paper

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

(molar ratio)/n-decane solution. We observed and compared
the membrane deformation with and without an ion concentration difference at each droplet. The observation was
performed using an inverted microscope (AXJ-5300TPHFL,
WRAYMER Inc.) mounted with a microscope digital camera(FLOYD, WRAYMER Inc.). The water volumes passing
through the lipid bilayer were calculated by measuring thevolume of droplets by image analysis (ImageJ).
Design and fabrication of the microchannel for liposome
trapping and gradient generation
To generate an ion concentration difference at the front andrear of the liposome, we designed a microchannel satisfying
the following requirements. First, the liposome should betightly fixed in the microchannel to clear the gap between theliposome and the channel walls. Second, solutions with different ion concentrations must flow on both sides of thetrapped liposome. Therefore, we proposed a microchannel inwhich strategies of a droplet-trapping microchannel36,37 forsingle cell trapping and a concentration gradient microchannel38 are combined.
To confirm the functions of the microchannel of liposome trapping and gradient generation, hydrodynamic simulations of the flow velocity and salt concentration wereconducted with COMSOL Multiphysics 4.2a (COMSOL Inc.)
using the microfluidics module. The density and viscosityof the aqueous solution were set to 1.0 mg mL−1 and 1.0
mPa s, respectively. The diffusion coefficient was set to 1.0
× 10−9 m2 s−1.
The microchannel was fabricated by standard photolithography techniques with SU-8 3025 which was used as thevmaster photoresist. Chromium masks were fabricated with amaskless exposure system (Nano System Solutions Inc.) as
the photomasks. A mask aligner (SUSS MA6/BA6 SPEC TU,
SUSS MicroTec AG) was used for UV exposure. The microchannel consists of three different layers. The first andsecond layers were fabricated on a single silicon wafer, andthe third layer was fabricated on another silicon wafer. The
channels were fabricated by PDMS molding, where PDMS
was cured for 2 h at 95 °C. Each channel layer was bonded
using plasma etching equipment (FA-1, Samco Inc.).
Liposome preparation and trapping in the microchannel
with a salt gradient
Giant liposomes were prepared by a water-in-oil emulsion
centrifugation method.39,40 1.0 M KCl and 0.5 mg mL−1
calcein solution (10 mM MOPS, pH 7.0) were contained in
the liposomes. See the ESI† for further details of the formation of the giant liposomes. 1.0 M KCl solution (10 mM
MOPS, pH 7.0) with liposomes and 0.1 M KCl (10 mM
MOPS, pH 7.0) without liposomes flowed at 2.0 and 1.0 μL
h−1 from inlets 1 and 2, respectively, with syringe pumps(YSP-202, YMS Co., Ltd.). We observed a liposome trappedin the narrow channel every 30 s using a fluorescencemicroscope (AXJ-5300TPHFL, WRAYMER Inc.) with a3474 | Lab Chip, 2019, 19, 3472–3480

Lab on a Chip

mounted microscope digital camera (FLOYD, WRAYMERInc.). Then, the movements of the liposome membrane andthe center of mass were measured by image analysis(ImageJ).
Observation of the inner flow with liposomes including
beads
To observe the osmotic flow generated in liposomes, weprepared liposomes with fluorescent beads inside them andobserved the behavior of the beads. Polystyrene fluorescentbeads with a 1 μm diameter (Polysciences, Inc.) were
included in the liposomes at a density of 10 beads per pL.
We conducted the microfluidic experiments under the same
conditions as in the above section and tracked the bead
behavior.

Results and discussion
Concept and displacement mechanism of liposomes
The displacement of liposomes is based on the osmotic flow
across the lipid bilayer. The semipermeability of the bilayer(water can permeate the lipid bilayer, but ions cannot) causes
the osmotic flow via the osmotic pressure difference between
inside and outside of the liposome. The osmotic pressure difference, ΔΠ, is expressed as ΔΠ = Πin − Πout = RT(cin − cout),
where cin and cout are the ion concentrations inside andoutside of the liposome, respectively. When solutions withdifferent ion concentrations are separated by a single lipid
bilayer, the water flows from the lower to the higher ion
concentration owing to the osmotic pressure difference. As aresult of the volume change of the solutions, the lipid bilayer
moves toward the side with the lower ion concentration(Fig. 2a). Next, the case that solutions with different ion
concentrations are separated by the liposome with the inner
solution having an ion concentration between them is shownin Fig. 2b. Osmotic pressure is generated at the front andrear membranes of the liposome, and water flows from thelower to the higher ion concentration owing to the osmoticpressure difference at both ends of the liposome. Bothmembranes move in the direction of the lower ion concentration as a result of the volume change of the solutions.
Finally, the liposome is transported by the shear force generated by the flexure of the membrane and migrates in the
direction of the lower ion concentration.
Estimation of osmotic pressure by observing deformation of
the lipid bilayer membrane
To predict the velocity of moving liposomes, the water flux
passing through lipid bilayers was calculated from the deformation of the planar bilayer lipid membrane (pBLM) formedby the DCM (Fig. 3a). Because of the symmetric salt conditionof 0.5 M : 0.5 M KCl in each droplet, the pBLM did not bend
since it was formed with droplets of an isotonic salt solution
and the osmotic pressures from both sides were equal(Fig. 3b). By contrast, the pBLM formed with droplets having

This journal is © The Royal Society of Chemistry 2019

View Article Online

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

Lab on a Chip

Fig. 2 Mechanism of the liposome movement. a) The case that thelipid bilayer is formed in a closed chamber. An osmotic flow passing
through the pBLM from the lower to the higher salt concentration is
generated when a difference in the salt concentration is generated
across the lipid bilayer. Then, the lipid bilayer moves in the direction of
the lower salt concentration as a result of the increased volume of thesolution with the higher salt concentration. b) The case that theliposome is trapped in a closed chamber and a phased salt
concentration difference is generated between the front, the inside,and the rear of the liposome. The front and rear membranes move inthe direction of the lower salt concentration owing to the osmoticpressure difference, as in the case with a single lipid bilayer. Then,
traction force is generated at the membrane as a result of membranebending. Finally, the liposome migrates in the direction of the lower
salt concentration.

the asymmetric salt condition of 1.0 M : 0.1 M KCl moved in
the direction with the lower salt concentration because thewater flowed from the lower to the higher salt concentration
owing to the osmotic pressure difference at the pBLM
(Fig. 3c, Movie S1†). We also observed the osmotic flow when
the pBLM was formed by the lipid composition of DOPC andchol to imitate the liposomal membrane used in theliposome experiments. Although we obtained the membrane
deformation with the DOPC/chol membrane, the magnitudeof the deformation is smaller than that of the DOPC
membrane (Fig. S1†). That is because the stiffness of theliposomal membrane increased41 and the osmotic water
permeability across the lipid bilayers decreased42 by adding
chol to the lipid membrane. From these results, we calculated the time dependence of the volume of permeated water
passing through the pBLM from the volume change of thedroplets (Fig. 3d). The permeated water volumes with theasymmetric salt condition of 1.0 M : 0.1 M KCl linearly
increased with time in the DOPC and DOPC/chol
membranes, and the flow rate of water passing through thepBLMs was calculated to be 4.9 and 3.3 nL min−1,
respectively.
Next, we estimated the maximum chemotaxis speed ofliposomes by calculating the volume of permeated water per

This journal is © The Royal Society of Chemistry 2019

Paper

Fig. 3 a) pBLMs prepared by the DCM. pBLMs are formed bycontacting lipid monolayers formed around the W/O emulsions.
Microscopy images of membrane deformation b) without and c) with asalt concentration difference at each droplet. In the case of a salt
concentration difference, the pBLM moves in the direction of thelower salt concentration. In contrast, the pBLM formed with isotonic
droplets remains at the initial position. d) Time courses of thepermeated water volume calculated by image analysis. The permeatedvolumes can be linearly fitted, and the flow rate is 4.9 and 3.3 nL min−1
with DOPC and DOPC/chol membranes. e) Time courses of thepermeated water volume per unit area, which can also be linearly
fitted, and the flow rate per unit area is 12.1 and 8.5 fL μm−2 min−1,
respectively.

unit area passing through the pBLM. We assumed a rectangular shape for the pBLMs formed by the DCM.29 The heightof the pBLM was calculated to be 600 μm by the volumes of
the chamber and the aqueous solution added to the chamber. The width of the pBLM was confirmed from the microscopy image and measured by ImageJ in each experiment. As
a result, we confirmed that the permeated water volume perunit area also linearly increased with time, and the flow rate
per unit areas with the DOPC and DOPC/chol membranes
were calculated to be 12.1 and 8 fL μm−2 min−1, respectively(Fig. 3e). Assuming that a liposome is trapped in a microfluidic channel with a 10 × 10 μm2 cross-sectional surface,and a concentration difference with concentrations of 1.0
and 0.1 M KCl on the inside and outside of the liposome,respectively, the maximum displacement speeds of theliposomes exhibited by the DOPC and DOPC/chol
membranes were estimated to be 12.1 and 8.5 μm min−1,
respectively. These results indicate that osmotic-flow-driven
molecular robots can potentially migrate faster thanmolecular-motor-driven living cells, which have a migration
speed of around 1 μm min−1 even in a microchannel.43

Lab Chip, 2019, 19, 3472–3480 | 3475

View Article Online

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

Paper

Fig. 4 a) Design of the microchannel used for liposome trapping andgradient generation. The microchannel consists of two main channels
with a narrow channel connecting them. Results of the hydrodynamic
simulation of b) flow velocity and c) salt concentration. Liposomes
flowing in channel 1 enter the narrow channel because the resistanceof the narrow channel is lower than that of channel 1. After theliposome is trapped in the narrow channel, a concentration differenceis generated between the front and rear of the liposome.

Hydrodynamic simulations of the microchannel for liposome
trapping and generation of concentration difference
Details of the design and the designed values of the microchannel are shown in Fig. 4a. The microchannel is composedof long and short main channels with a narrow channel
connecting the two main channels. The narrow channel isdivided into two parts, one is a liposome-trapping part andthe other is a liposome migration part. The widths of theliposome-trapping (W1_1) and the liposome migration parts(W1_2) are designed to be 20 and 10 μm, respectively. The
length and height of the narrow channel are 100 and 10 μm,
respectively. The width and height of both main channels aredesigned to be 40 and 25 μm, and one of the main channels
is 30 mm longer than the other so that the liposomes canflow into the narrow channel by the difference in the channel
resistances between the long main channel and the narrow
channel.
To trap the liposome in the microchannel and generate aconcentration difference between the front and rear of theliposome, solutions with higher and lower salt concentrations and with and without liposomes were made to flow

3476 | Lab Chip, 2019, 19, 3472–3480

Lab on a Chipfrom inlets 1 and 2, respectively. The liposome from inlet 1
flows into the narrow channel because of the difference inthe channel resistances between the main channel and thenarrow channel. Then, the liposome is trapped in the narrow
channel because its cross-sectional area (10 × 10 μm2) ismuch smaller than the size of the liposomes (20 μm diameter). To confirm the functions of the microchannel of liposome trapping and the generation of a concentration difference between the front and rear of the trapped liposome, we
calculated the flow velocity and salt concentration in the
microchannel by hydrodynamic simulation.
The simulation results for the flow velocity in the microchannel are shown in Fig. 4b. In the simulation to obtain theflow velocity, the solutions flowed from inlets 1 and 2 at flow
rates of 2.0 and 1.0 μL s−1, respectively. The color indicatesthe flow velocity and the red arrows show the direction andmagnitude of the flow. The solution injected from inlet 1
mainly flowed to the narrow channel, indicating that theliposome should also flow to the narrow channel. The
liposome should also be trapped in the middle of the narrow
channel since its diameter is expected to be around 20 μm.
The simulation results of the salt concentration in theliposome-trapped and -untrapped microchannels are respectively shown in Fig. 4c and S2.† The salt concentrations ofsolutions injected from inlets 1 and 2 were 1.0 and 0.1 M,
respectively. When the liposome is not trapped in the narrow
channel, a linear concentration gradient is generated in the
narrow channel. By contrast, the salt concentrations at thefront and rear of the trapped liposome are almost the sameas those of the injected solutions because the narrow channel
is divided by the liposome. These results indicate that the
microchannel can trap the liposome in the narrow channel
and that a concentration difference can be generated between
the front and rear of the trapped liposome.

Liposome displacement induced by the osmotic pressure
difference
The microscopy image of the fabricated microchannel isshown in Fig. 5a. A solution of 1.0 M KCl (10 mM MOPS, pH
7.0) with liposomes and a solution of 0.1 M KCl (10 mM
MOPS, pH 7.0) without liposomes were flowed to channels 1
and 2, respectively. The liposome flowing in channel 1
entered the narrow channel owing to the difference in thechannel resistance between the narrow channel and channel
1. Then, the liposome which has a sufficiently larger size
than the migration channel (10 × 10 μm2) was trapped in the
narrow channel (Fig. 5b, Movie S2†). Specifically, liposomeswith a diameter of 15 μm or more were trapped, whereas theothers passed through the narrow channel to channel 2 orwere briefly trapped in the narrow channel before flowing tochannel 2. After the liposomes were trapped in the narrow
channel, about 50% of them ruptured or untrapped in thefirst 30 s. Although we first prepared liposomes formed by
DOPC and attempted to trap them in the microchannel, theliposomes ruptured when they were trapped in the narrow

This journal is © The Royal Society of Chemistry 2019

View Article Online

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

Lab on a Chip

Fig. 5 a) Microscopy image of the microchannel. The widths of thenarrow and main channels are 40 and 10 μm, respectively. b)
Fluorescence image of the microchannel with a liposome trapped in
the narrow channel. The liposome flows from channel 1 to the narrow
channel and clogs the narrow channel.

Paperchannel. Hence, we formed liposomes composed of DOPC,
chol, and DOPS to improve their mechanical properties (Fig.
S3†).41 Furthermore, there was no significant difference
between the success rate of liposome trapping with andwithout the osmotic pressure, indicating that osmoticpressure did not influence the rupture of liposomes (Fig.
S3†). Also, we did not observe specific membrane transformations when we applied osmotic pressure differences at thefront and rear of liposomes although lipid vesicles formed
with PC and cholesterol transform their shape from sphereto prolate, tube, pear, discocyte, and starfish shapes by applying a constant osmotic pressure difference.44 We indicatedthat is because the osmotic pressure differences were notapplied on the whole liposomes but limited membranes ofliposomes (front and rear). From these results, we concluded
that the stress by the flow pressure induced the rupture ofliposomes due to the softness of liposomes. To improve thecapture rate of liposomes, in the future, we will trap theliposomes in the microchannel with other manipulation
methods such as optical tweezers.
The expansion of the front membrane of the liposome
was clearly observed when 1.0 M and 0.1 M KCl were flowedin channels 1 and 2, respectively. (Fig. 6a, Movie S3†). On theother hand, the liposome did not move and remained at theFig. 6 Fluorescence images of a trapped liposome a) with and b) without salt concentration difference. With the salt concentration difference, theliposome moved in the direction of the lower salt concentration. Time course of the displacements of the c) front membrane, d) rear membrane,
and e) center of mass of the liposome. The displacements of the front membrane and the center of mass linearly increase with time, and their
speeds are 1.07 and 0.60 μm min−1, respectively.

This journal is © The Royal Society of Chemistry 2019

Lab Chip, 2019, 19, 3472–3480 | 3477

View Article Online

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

Paper

initial position under the isotonic salt conditions (Fig. 6b).
The measured membrane displacements at the front and therear of a liposome are shown in Fig. 6c and d, respectively.
The front membrane of the liposome linearly expanded witha constant rate of 1.0 μm min−1 when the osmotic pressure
difference was applied to the liposomal membrane. Although
the rear membrane of the liposomes also moved in the direction of the lower salt concentration, the rate of shrinkage waslower than the rate of expansion of the front membrane. We
also evaluated the displacement of the center of mass of theliposomes (Fig. 6e). The center of mass also linearly moved
in the direction of the lower salt concentration with a speedof 0.60 μm min−1. Because the initial salt concentrations of
the front, rear, and inside of the liposome were 0.1, 1.0, and
1.0 M KCl, respectively, an osmotic flow was not initiallyinduced at the rear membrane of the liposome. Thus, theosmotic flows at the front and rear of the liposome shouldhave decreased and increased with decreasing salt concentration inside the liposome, respectively. Assuming that thevolume of the liposome and the flow rate of the osmotic flow
were 4.2 pL and 0.85 pL min−1, respectively, the time required
to exchange the solution inside the liposome was calculated
to be about 4.9 min. Therefore, the velocity of the front
membrane was greater than that of the rear membrane in
the first 2.5 min. When the front membrane of the liposome
has expanded by about 3 μm (about 2.5 min after theliposome was trapped in the channel), the liposome slippedand slid back. We consider that the low adhesive force
between the liposomal membrane and the surface of the
microchannel caused the slip. Although cells generally attach
to a substrate by extracellular matrices, liposomes do nothave this function and therefore slid back as a result of the
traction force acting on the lipid bilayer. To drive liposomes
for a long time, the adhesive force between the liposomal
membrane and the substrate should be increased by electrostatic interaction or by using antibody-modified liposomes.
We visualized the internal flow of the liposomes by encapsulating fluorescent microbeads in them, and we confirmedthat they were driven by the osmotic pressure difference
since the liposomes had the potential to move owing to thepulsations generated by the syringe pumps. Although themicrobeads did not move for a few seconds after the

Lab on a Chipliposome was initially trapped in the narrow channel, several
seconds after the liposome was trapped, they started to moveand continued to circulate inside the liposome while the liposome was trapped in the narrow channel (Fig. 7 and Movie
S4†). We indicated that this circulating flow was generated as
below. First, an inflow into the liposome was induced at thefront membrane of the liposome by the osmotic pressure
difference. This inflow induced the one direction internal
flow in the liposome. Although the outflow was generated at
the rear membrane of the liposome, the flow rate was slower
than that of the inflow in the first several minutes because
the osmotic pressure difference at the rear membrane of theliposome was smaller than that of the front membrane. As aresult, the difference between the inflow and outflow induced
the circulating flow in the liposome. Also, the circulation of
the microbeads expanded in the direction of the migration
channel, indicating the expansion of the trapped liposome.
On the other hand, the microbeads did not move andremained at their initial positions in the case of no osmoticpressure difference when the liposome was trapped in the
narrow channel (Fig. S4†) or before trapping, indicating that
there is no residual flow in liposomes. In addition, since
there is no significant difference between the fluorescencedecay rates in the trapped liposomes with and without theosmotic pressure difference, there are no holes and leakage
from liposomes that might induce some flows. From theseresults of the microbead observation, we conclude that theosmotic flow is generated at the liposomal membrane by theosmotic pressure difference, and the liposome is moved by
the osmotic engine mechanism.
Next, we compared the estimated and observed speeds ofthe liposome. Although the liposome formed by the DOPC
and chol membrane has the potential to move at 8.5 μm
min−1, its actual speed was about eight times lower than theestimated value. We consider that the actual speed was
mainly influenced by the friction force between the liposome
membrane and the channel walls. Although the estimatedspeed was calculated by the water permeability passing
through the lipid bilayer prepared by the DCM, there is nofriction force between the chamber walls and the droplets
because there is an oil layer between the droplets and thechamber walls. By contrast, the liposomes completely

Fig. 7 a) Fluorescence images of a microbead in a liposome trapped in the narrow channel. b) Result of a tracking microbead in a liposome whena salt concentration difference is applied. The microbead circulates in the liposome, indicating an osmotic flow is generated in the liposome by theosmotic pressure difference.

3478 | Lab Chip, 2019, 19, 3472–3480

This journal is © The Royal Society of Chemistry 2019

View Article Online

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

Lab on a Chip

attached to the channel walls, and the friction force was
generated to the membrane. As a result, the expanding speedof the liposomal membrane became slower than theestimated value.
Although the velocities of conventional self-propelled
robots driven by the Marangoni effect or chemical engines
are faster than our osmotic-pressure-driven liposome in theorder of 1–100 μm s−1,45,46 our osmotic-pressure-driven liposome can move by simply applying a salt concentration difference between its front and rear without any chemical reactions. Furthermore, our method can be easily applied toliposome-based molecular robots owing to the semipermeability of their compartments. Also, the speed of our osmoticpressure-driven liposome is almost the same as that of living
cells, which is around 1 μm min−1,43 suggesting that ourdriving mechanism has the potential to develop cell-likemolecular robots.
To improve the speed of liposomes, in the future, we needto optimize the channel dimensions, the size of liposomes andthe adhesion force between liposomes and channel walls. The
speed of liposomes would be influenced by the channel dimensions because the traction force applied to the membrane
relates to the distance of the liposomal membrane expansion
and the cross-sectional dimensions of the migration channel.
In this experiment, however, optimizing the channel dimensions are very difficult because we cannot completely control
the size of liposomes prepared by the water-in-oil emulsion
centrifugation method, which were from 5 to 30 μm in diameter (Fig. S5†). To improve the size control of liposomes, we will
prepare liposomes with microfluidic techniques which can
produce much uniformly sized giant liposomes.47,48 In
addition, when the trapped liposome was deformed, the elastic
energy of the liposomal membrane, which tended to pull theliposome back to its initial position, increased. To overcome
the energy barrier, the adhesive force between liposomes andchannel walls should be controlled by surface modification on
the channel walls, by electrostatic interaction, or using
antibody-modified liposomes.

Conclusions
We proposed the application of an osmotic engine model tothe actuator of liposome-based molecular robots. To demonstrate osmotic-pressure-driven-liposomes, we utilized themicrofluidic technologies of particle trapping and gradient
generation to generate a salt concentration difference
between the front and rear of liposomes. As a result, we
successfully induced the displacement of liposomes by theosmotic pressure difference.
To increase the speed, working time, and displacement of
the liposome, we will increase the osmotic flow of water
passing through the liposomal membrane by the reconstitution of an aquaporin into the liposomal membrane andcontrol the interaction between the liposome and thechannel walls by surface modification in the channel orelectrostatic interaction.

This journal is © The Royal Society of Chemistry 2019Paper

For the further development of liposome-based molecularrobots, the combination of osmotic-pressure and molecularmotor-driven mechanisms will enable the realization ofmolecular robots with advanced behavior. For example, amolecular robot that can deform its shape using molecularmotors to fit a pathway and chemotactically migrate along
the pathway will be developed. In addition, the reconstitutionof membrane proteins and their functionalization13,14 will
also lead to the control of the osmotic-pressure-based actuation mechanism. We believe that the osmotic-pressure-driven
actuation mechanism will become a breakthrough in thedevelopment of liposome-based molecular robots. Moreover,
our liposome-based chemotaxis system has potential as thecell migration model to imitate the chemotaxis mechanism.

Conflicts of interest
There are no conflicts to declare.

Acknowledgements
This material is based upon work supported by the Promotionof Science (JSPS) Research Fellow and KAKENHI (Grant No.
17K19138 and 19H00901) and is partially supported by
KAKENHI (Grant No. 19K15418) from the MEXT Japan.

References
1 J. Xi, J. J. Schmidt and C. D. Montemagno, Nat. Mater.,
2005, 4, 180–184.
2 Y. Tanaka, K. Morishima, T. Shimizu, A. Kikuchi, M. Yamato,
T. Okano and T. Kitamori, Lab Chip, 2006, 6, 230–235.
3 A. W. Feinberg, A. Feigel, S. S. Shevkoplyas, S. Sheehy, G. M.
Whitesides and K. K. Parker, Science, 2007, 317, 1366–1370.
4 R. Holzer and I. Shimoyama, Proceedings of the 1997 IEEE/
RSJ International Conference on Intelligent Robot and Systems.
Innovative Robotics for Real-World Applications. IROS ‘97,
1997, vol. 3, pp. 1514–1519.
5 A. Bozkurt, J. R. F. Gilmour and A. Lal, IEEE Trans. Biomed.
Eng., 2009, 56, 2304–2307.
6 H. Sato, C. W. Berry, Y. Peeri, E. Baghoomian, B. E. Casey, G.
Lavella, J. M. VandenBrooks and M. M. Maharbiz, Front.
Integr. Neurosci., 2009, 3, 24.
7 K. Shoji, Y. Akiyama, M. Suzuki, N. Nakamura, H. Ohno and
K. Morishima, Biosens. Bioelectron., 2016, 78, 390–395.
8 K. Shoji, Y. Akiyama, N. Nakamura, H. Ohno and K.
Morishima, Proceedings of 2016 IEEE International Conference
on Mechatronics and Automation (ICMA2016), 2016, pp. 629–634.
9 M. Hagiya, A. Konagaya, S. Kobayashi, H. Saito and S.
Murata, Acc. Chem. Res., 2014, 47, 1681–1690.
10 S. Murata, A. Konagaya, S. Kobayashi, H. Saito and M.
Hagiya, New Gener. Comput., 2013, 31, 27–45.
11 Y.-R. Kim, S. Jung, H. Ryu, Y.-E. Yoo, S. M. Kim and T.-J.
Jeon, Sensors, 2012, 12, 9530–9550.
12 M. Hiratani, M. Ohara and R. Kawano, Anal. Chem.,
2017, 89, 2312–2317.

Lab Chip, 2019, 19, 3472–3480 | 3479

View Article Online

Published on 04 September 2019. Downloaded by Kyoto Daigaku on 10/18/2019 4:57:12 AM.

Paper

13 S. Mizukami, M. Kashibe, K. Matsumoto, Y. Hori and K.
Kikuchi, Chem. Sci., 2017, 8, 3047–3053.
14 M. Ohara, M. Takinoue and R. Kawano, ACS Synth. Biol.,2017, 6, 1427–1432.
15 R. Kawano, ChemPhysChem, 2018, 19, 359–366.
16 T. Masubuchi, M. Endo, R. Iizuka, A. Iguchi, D. H. Yoon, T.
Sekiguchi, H. Qi, R. Iinuma, Y. Miyazono, S. Shoji, T.
Funatsu, H. Sugiyama, Y. Harada, T. Ueda and H.
Tadakuma, Nat. Nanotechnol., 2018, 13, 933–940.
17 M. Honda, K. Takiguchi, S. Ishikawa and H. Hotani, Biophys.
J., 1999, 287, 293–300.
18 A. Roux, G. Cappello, J. Cartaud, J. Prost, B. Goud and P.
Bassereau, Proc. Natl. Acad. Sci. U. S. A., 2002, 99, 5394–5399.
19 Y. Sato, Y. Hiratsuka, I. Kawamata, S. Murata and S. M.
Nomura, Sci. Robot., 2017, 2, eaal3735.
20 S. Tanaka, K. Takiguchi and M. Hayashi, Commun. Phys.,2018, 1, 18.
21 T. Sunami, K. Hosoda, H. Suzuki, T. Matsuura and T. Yomo,
Langmuir, 2010, 26, 8544–8551.
22 S. A. Sarles and D. J. Leo, Smart Mater. Struct., 2011, 20,094018.
23 K. Kurihara, M. Tamura, K. Shohda, T. Toyota, K. Suzuki
and T. Sugawara, Nat. Chem., 2011, 3, 775–781.
24 Y. Elani, R. V. Law and O. Ces, Nat. Commun., 2014, 5, 5305.
25 L. Schoonen and J. C. M. van Hest, Adv. Mater., 2016, 28,
1109–1128.
26 C. Kurokawa, K. Fujiwara, M. Morita, I. Kawamata, Y.
Kawagishi, A. Sakai, Y. Murayama, S. M. Nomura, S. Murata,
M. Takinoue and M. Yanagisawa, Proc. Natl. Acad. Sci. U. S.
A., 2017, 114, 7228–7233.
27 F. C. Keber, E. Loiseau, T. Sanchez, S. J. DeCamp, L. Giomi,
M. J. Bowick, M. C. Marchetti, Z. Dogic and A. R. Bausch,
Science, 2014, 345, 1135–1139.
28 S. Sengupta, M. E. Ibele and A. Sen, Angew. Chem., Int. Ed.,2012, 51, 8434–8445.
29 M. Guix, C. C. Mayorga-Martinez and A. Merkoçi, Chem.
Rev., 2014, 114, 6285–6322.
30 Z. Wu, X. Lin, T. Si and Q. He, Small, 2016, 12, 3080–3093.
31 J. Nardi, R. Bruinsma and E. Sackmann, Phys. Rev. Lett.,
1999, 82, 5168–5171.

3480 | Lab Chip, 2019, 19, 3472–3480

Lab on a Chip

32 K. M. Stroka, H. Jiang, S.-H. Chen, Z. Tong, D. Wirtz, S. X.
Sun and K. Konstantopoulos, Cell, 2014, 157, 611–623.
33 K. Funakoshi, H. Suzuki and S. Takeuchi, Anal. Chem.,
2006, 78, 8169–8174.
34 H. Bayley, B. Cronin, A. Heron, M. A. Holden, W. L. Hwang,
R. Syeda, J. Thompson and M. Wallace, Mol. BioSyst.,
2008, 4, 1191–1208.
35 R. Kawano, Y. Tsuji, K. Sato, T. Osaki, K. Kamiya, M.
Hirano, T. Ide, N. Miki and S. Takeuchi, Sci. Rep., 2013, 3,
1995.
36 W.-H. Tan and S. Takeuchi, Proc. Natl. Acad. Sci. U. S. A.,
2007, 104, 1146–1151.
37 T. S. Kaminski and P. Garstecki, Chem. Soc. Rev., 2017, 46,6210–6226.
38 J. Atencia, J. Morrow and L. E. Bocaccio, Lab Chip, 2009, 9,2707–2714.
39 S. Pautot, B. J. Frisken and D. A. Weitz, Langmuir, 2003, 19,2870–2879.
40 T. Toyota, N. Ohguri, K. Maruyama, M. Fujinami, T. Sagaand I. Aoki, Anal. Chem., 2012, 84, 3952–3957.
41 X. Liang, G. Mao and K. Y. S. Ng, J. Colloid Interface Sci.,
2004, 278, 53–62.
42 D. W. Deamer and J. Bramhall, Chem. Phys. Lipids, 1986, 40,
167–188.
43 P. Maiuri, E. Terriac, P. Paul-Gilloteaux, T. Vignaud, K.
McNally, J. Onuffer, K. Thorn, P. A. Nguyen, N. Georgoulia,
D. Soong, A. Jayo, N. Beil, J. Beneke, J. C. H. Lim, C. P.-Y.
Sim, Y.-S. Chu, A. Jiménez-Dalmaroni, J.-F. Joanny, J.-P.
Thiery, H. Erfle, M. Parsons, T. J. Mitchison, W. A. Lim,
A.-M. Lennon-Duménil, M. Piel and M. Théry, Curr. Biol.,2012, 22, R673–R675.
44 H. Hotani, J. Mol. Biol., 1984, 178, 113–120.
45 T. Toyota, N. Maru, M. M. Hanczyc, T. Ikegami and T.
Sugawara, J. Am. Chem. Soc., 2009, 131, 5012–5013.
46 I. Lagzi, S. Soh, P. J. Wesson, K. P. Browne and B. A.
Grzybowski, J. Am. Chem. Soc., 2010, 132, 1198–1199.
47 S. Matosevic and B. M. Paegel, J. Am. Chem. Soc., 2011, 133,
2798–2800.
48 N.-N. Deng, M. Yelleswarapu and W. T. S. Huck, J. Am.
Chem. Soc., 2016, 138, 7584–7591.

This journal is © The Royal Society of Chemistry 2019