Encapsulation of Active Cytoskeletal Protein Networks in Cell-Sized

Jun 27, 2011 - proteins and polymers, for instance, to create polymer-reinforced liposomes for drug delivery ..... were recorded with a digital CCD ca...
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Encapsulation of Active Cytoskeletal Protein Networks in Cell-Sized Liposomes Feng-Ching Tsai, Bj€orn Stuhrmann, and Gijsje H. Koenderink* Biological Soft Matter Group, FOM Institute AMOLF, Science Park 104, 1098 XG Amsterdam, The Netherlands

bS Supporting Information ABSTRACT: We demonstrate that cytoskeletal actinmyosin networks can be encapsulated with high efficiency in giant liposomes by hydration of lipids in an agarose hydrogel. The liposomes have cell-sized diameters of 1020 μm and a uniform actin content. We show by measurements of membrane fluorescence intensity and bending rigidity that the majority of liposomes are unilamellar. We further demonstrate that the actin network can be specifically anchored to the membrane by biotinstreptavidin linkages. These protein-filled liposomes are useful model systems for quantitative studies of the physical mechanisms by which the cytoskeleton actively controls cell shape and mechanics. In a broader context, this new preparation method should be widely applicable to encapsulation of proteins and polymers, for instance, to create polymer-reinforced liposomes for drug delivery.

’ INTRODUCTION The shape of a living cell is jointly determined by the cell membrane and the cytoskeleton.1 The membrane defines the cell’s autonomy, separating the cytoplasm from the external environment, but does not provide much mechanical resistance since it is a fluid material. The cytoskeleton is a filamentous network that spans the cytoplasm and mechanically supports the membrane. The membrane is mainly supported by filamentous (F) actin, which forms a thin cortical shell.2 Actin is a globular protein that polymerizes to form double-helical filaments with two structurally distinct ends. In the presence of a chemical fuel (adenosine 50 -triphosphate, ATP), actin filaments can generate pushing forces by growing with their fast-growing plus end against the membrane.3 These pushing forces contribute to cell locomotion and the formation of membrane protrusions such as filopodia.4 The actin cortex is actively held under tension by myosin motors, which hydrolyze ATP to contract the actin network.5 When the actin cortex detaches from the membrane, the release of internal tension contributes to membrane blebbing.6 Part of the myosin-based tension in the actin cytoskeleton is transmitted to the extracellular matrix or to adjacent cells across transmembrane adhesion receptors.7 This force transmission enables cells to actively sense and respond to changes in tissue stiffness. The molecular composition of the cytoskeleton and associated cell membrane are complex and highly regulated. Therefore, it is difficult to unravel the interplay of physical and biochemical mechanisms that is responsible for the shape, mechanical properties, and mechanosensitivity of cells.8 In vitro studies of model cytoskeletal networks, reconstituted from purified actin,9 and model membranes, reconstituted from purified lipids,10 have provided many quantitative insights into the material properties r 2011 American Chemical Society

of these components in isolation. There have also been in vitro studies of interactions between actin networks and membranes. Experimentally, it is relatively straightforward to couple actin filaments to flat lipid monolayers11 or bilayers12 or onto the outer surface of liposomes.13 These model systems have enabled measurements of the viscoelastic properties of actinmembrane composites and have provided insights into the roles of actin filament growth in driving cell migration and the formation of membrane protrusions. However, these model systems do not mimic the membrane curvature and confinement that actin filaments experience in vivo and cannot be used to model transmembrane transmission of forces generated by myosin contractility. Accordingly, there have also been several efforts to encapsulate actin networks inside giant cell-sized liposomes.14 However, encapsulation of actin (or other cytoskeletal proteins, such as tubulin15,16) under physiological buffer conditions that promote actin polymerization and mechanochemical activity of motor proteins is difficult. There is currently no facile and robust method to achieve this encapsulation. There are two main factors hampering the encapsulation of active cytoskeletal systems into giant liposomes. First, the high salt levels in physiological buffers generally cause a low or even zero yield of liposomes.17,18 Second, the encapsulation procedure should preserve the biological activity of the proteins, which requires a fast procedure. The widely used electroformation method requires low salt concentrations,19 though special protocols can be used to allow formation at high salt,16,17,20 or salt can be introduced afterward by means of electroporation21 or Received: April 29, 2011 Revised: June 24, 2011 Published: June 27, 2011 10061

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Langmuir incorporation of ion channels in the membrane.22,23 However, liposome formation is slow, typically taking at least 3 h, and the electric field could potentially be harmful to the proteins that are to be encapsulated. Several alternative approaches have been developed for encapsulating high-salt solutions, including gentle hydration,25 freezethaw extrusion,26 solvent evaporation,27 inkjet or pulsed jet flow formation,2831 and an inverted emulsion method.32 Actin-filled liposomes have thus far been obtained by gentle hydration,33 inkjet formation,28a,b and the inverted emulsion method,24 yet each of these methods has its drawbacks. Gentle hydration is only successful with specific lipid compositions, for example, L-R-lysophosphatidylcholine (eggPC) or 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC). 21,33,34 The microfluidic inkjetting technique is a fast method that can produce liposomes in high yield, but it requires specialized equipment. The inverted emulsion method is fast (20 min) but requires multiple steps. Two of these steps, the creation of a water-in-oil emulsion and centrifugation of the emulsion droplets through a lipid monolayer to obtain liposomes, are particularly difficult to reproduce,35 though recent papers show that microfluidics-based methods can improve the reproducibility.36 Here we propose a new method to prepare giant liposomes filled with an active actinmyosin cytoskeleton. Our method is facile, quick, and reproducible and permits control over actin membrane anchoring. Our method is inspired by a recent study of Horger and co-workers,37 who showed that liposomes containing high-salt buffers can be obtained by gentle hydration of an agarose hydrogel film infiltrated with lipids. They proposed that the agarose hydrogel promotes multiple factors that are essential for liposome formation, including self-assembly of the dried lipids into lamellae, swelling of the lamellae, and fusion of adjacent liposomes into giant liposomes. In this paper, we show that actinfilled liposomes are obtained in high yield when hybrid agarose lipid films are hydrated in a solution of skeletal muscle actin and myosin II. The required conditions—spin-coated agaroselipid films and the presence of PEGylated (PEG = polyethylene glycol) lipids—are more restrictive than those for empty liposomes. We characterize the liposome size distribution, membrane lamellarity, and efficiency of protein encapsulation. We further demonstrate that our method affords control over the anchoring of the actin myosin network to the inner leaflet of the liposome membrane. This study is the first one where processive bipolar myosin filaments are encapsulated in liposomes. These bipolar filaments mimic minifilaments which generate actin contractility in cells.38 They are capable of generating contraction of entangled and cross-linked actin networks in vitro.39 In prior studies, only double-headed subfragments of myosin II (known as heavy meromyosin) have been encapsulated in liposomes,40 which are collectively able to cause contraction of actin networks or bundles,41 but individually are nonprocessive. The giant actomyosin-filled liposomes prepared in this study provide an experimental model system to guide future quantitative studies of physical mechanisms that determine cellular processes such as cell spreading,42 contraction and blebbing,6 and myosin-based mechanosensing.7 Ultimately, such systems may possibly evolve into artificial cells in combination with minimal gene expression systems that enable self-reproduction using synthetic genetic information.43

’ MATERIALS AND METHODS Materials. Lipids were purchased from Avanti Polar Lipids (Alabaster, AL). These included 1,2-dioleoyl-sn-glycero-3-phosphocholine

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(DOPC), 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (PEG-DOPE with a 2 kDa polyethylene glycol tail), the fluorescently labeled lipid 1,2-dioleoyl-sn-glycero-3phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (rhodamine-DOPE), and the biotinylated lipid 1,2-dipalmitoyl-sn-glycero-3phosphoethanolamine-N-(cap biotinyl) (biotin-PE). We investigated the formation of liposomes on films of type IX-A ultralow gelling agarose (gel point Tg e 17 C) from Sigma-Aldrich (St. Louis, MO). We chose this type of agarose because it has the lowest gelling temperature on the market, which we needed because we performed gentle swelling at 4 C. We also tested ultralow gelling temperature agarose from Fluka, but this gave a markedly reduced liposome yield and encapsulation efficiency. To test for incorporation of agarose in liposomes, agarose was fluorescently labeled37 with fluorescein isothiocyanate (FITC) from Sigma-Aldrich. Globular (G) actin and myosin II were purified from rabbit skeletal muscle including a gel filtration of actin on a Sephacryl S-200 highresolution column (GE Healthcare, Munich, Germany) to remove residual capping proteins, cross-linkers, and actin oligomers.44,45 G-actin was stored at 80 C in G-buffer (2 mM tris(hydroxymethyl)aminomethane (Tris)HCl, 0.2 mM Na2ATP, 0.2 mM CaCl2, 0.2 mM dithiothreitol (DTT), 0.5 mM NaN3, pH 8.0). Myosin II was stored at 20 C in a high-salt buffer with glycerol (0.6 M KCl, 25 mM KH2PO4, 10 mM EDTA, 1 mM DTT, pH 6.5, 50% (w/w) glycerol). Fresh myosin solutions were prepared daily by dialysis against AB300 buffer (300 mM KCl, 4 mM MgCl2, 1 mM DTT, 25 mM imidazole, pH 7.4). Protein concentrations were determined by absorption measurements using extinction coefficients of 0.53 cm2/mg at 280 nm for myosin and 1.1 cm2/mg at 290 nm for actin.44 Fluorescent myosin with a dye: protein ratio of 1.7 was prepared by labeling amine groups with DyLight 594 NHS ester.46 G-actin labeled with AlexaFluor 488 or AlexaFluor 568 or with biotin was purchased from Invitrogen (Breda, The Netherlands) and Tebu-bio (Heerhugowaard, The Netherlands), respectively. The polymerization activity of actin was checked by fluorescence microscopy and by measurements of rheological properties with video particle tracking microrheology and macroscopic shear rheometry.39b The motor activity of myosin was checked by standard motility assays and by bulk selforganization assays,39b and in the same way we confirmed that DyLight 594 labeling did not influence motor activity. To test the filament assembly capability of myosin II in I-buffer (25 mM imidazoleHCl (pH 7.4), 0.1 mM DTT, 0.1 mM Mg-ATP, 50 mM KCl, 2 mM MgCl2, 1 mM Tris, 280 mM sucrose),47 we took electron micrographs of myosin filaments assembled at a concentration of 0.24 μM. After incubation in I-buffer for 3 min at room temperature, 4 μL of sample was pipetted onto a glow-discharged 300 mesh carbon-coated grid (Agar Scientific, Stansted, U.K.) and negatively stained with 2% methylamine tungstate (Nanoprobes, New York). The sample was examined with a transmission electron microscope CM12 (Philips Electronic Instrument, Eindhoven, The Netherlands) operated at 120 kV. Streptavidin was obtained from Thermo Scientific (Breda, The Netherlands), and creatine phosphate and creatine kinase were obtained from Roche Diagnostics (Almere, The Netherlands). Other chemicals were purchased from Sigma-Aldrich. ATP was prepared as a 100 mM Mg-ATP stock solution using equimolar amounts of Na2ATP and MgCl2 in 10 mM imidazole HCl buffer (pH 7.4). Liposome Preparation. The liposome preparation method was adapted from a recently published procedure where liposomes that were empty or contained dextran were formed by hydrating mixed films of agarose and lipids in salt solutions.37 The main modifications are the inclusion of cytoskeletal proteins and the use of buffer conditions that promote actin polymerization and myosin activity. In addition, we found that the formation conditions were more restrictive, as explained below. Lipid mixtures of DOPC/PEG-DOPE/rhodamine-DOPE/biotin-PE with molar ratios of 94.8:5:0.2:0 and 93.8:5:0.2:1 were dissolved in 95:5 (v/v) chloroform/methanol at a total concentration of 3.75 mg/mL. 10062

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Figure 1. (A) Schematic overview of the liposome preparation procedure, which consists of three steps: (i) an ultralow gelling agarose solution is spincoated onto a glass slide and partially dried, (ii) lipids are spin-coated on top and dried under vacuum, and (iii) liposomes are formed by hydration in a solution of actin and myosin in I-buffer at 4 C. The liposomes are flown from the formation chamber into an adjacent observation chamber by injection of O-buffer. Actin polymerization is initiated by heating the sample to room temperature. (BF) Confocal fluorescence micrographs of giant actin-filled liposomes formed within 1 h by gentle hydration of agaroselipid films. Red represents rhodamine-lipid fluorescence, and green represents AlexaFluor 488-actin fluorescence (10 mol % in (B)(D) and 20 mol % in (E) and (F)). The following lipid compositions were used (mol %): (B, C, D, F) DOPC: PEG-DOPE:rhodamine-DOPE = 94.8:5:0.2, (E) DOPC:rhodamine-DOPE = 99.8:0.2. Observation conditions: (B) liposomes were diluted 5-fold with O-buffer and observed in the observation chamber; (CF) liposomes were observed inside the formation chamber. (B) Liposomes filled with actin (4 mg/mL) are obtained with high yield using hybrid films of agarose and 5 mol % PEGylated lipids. Scale bar 10 μm. (C) Liposomes filled with actin (1 mg/mL) in the formation chamber. Scale bar 50 μm. (D) In the absence of agarose, virtually no liposomes are formed. Scale bar 50 μm. (E) No liposomes are formed in the absence of PEGylated lipids. Scale bar 50 μm. (F) Liposomes formed on hybrid agaroselipid films prepared by dipcoating37 rarely contain actin, and liposomes are absent in most areas of the formation chamber (top panel) except for a few locations (bottom panel). Scale bars 30 μm. We also tested lipid solutions of 0.87 mg/mL and other chloroform: methanol ratios (100:0 and 6:4), but this gave less optimal results. Agarose was dissolved in deionized water at a concentration of 1% (w/w) and boiled for 3050 s in a microwave oven immediately before 300 μL was spin-coated onto 24  24 mm glass coverslips (precleaned by rinsing with H2O, ethanol, and methanol and drying after each rinse under a stream of nitrogen) at a speed of 1200 rpm for 30 s (DELTA 10BM, SUSS MicroTec). We also tested spin speeds of 200, 600, and 800 rpm and agarose solution volumes of 50, 100, 600, and 800 μL, but this resulted in a poorer liposome yield and quality. The spin-coating procedure was essential for high liposome yield and protein encapsulation. The originally described dip-coating procedure gave low yield and poor encapsulation efficiency (see Figure 1F). The coated slide was dried for 30 min at 37 C, yielding a partially dried agarose film that retains ∼15% water.37 Lipid solution was deposited onto this film by spin-coating 150 μL at a speed of 1200 rpm for 5 min. Less good results were obtained with a speed of 600 rpm and/or lipid solution volumes of 20, 30, 50, 100, and 200 μL. The coated slide was then placed under vacuum for 1 h to remove residual chloroform and methanol. A glass chamber was constructed by placing a glass slide on top of the coated slide using glass and Parafilm spacers sealed with vacuum grease, as sketched in Figure 1A.

I-buffer (30 μL) was injected into the formation chamber, and the chamber was incubated for 4560 min to form liposomes containing I-buffer. We performed the incubation at 4 C to prevent premature actin (and myosin) filament formation. The composition of the I-buffer was chosen to promote actin polymerization and myosin II motor activity (25 mM imidazoleHCl (pH 7.4), 0.1 DTT, 0.1 mM Mg-ATP, 50 mM KCl, 2 mM MgCl2, 1 mM Tris, 280 mM sucrose)47 and contained an enzymatic system to regenerate ATP composed of 1.33 mg/mL creatine phosphate and 2.48 mg/mL creatine phosphokinase. We used actin concentrations between 1 and 4 mg/mL, with or without myosin II (in a molar ratio of 1:200 or 1:100 or 1:50 with G-actin), biotinylated G-actin (in a molar ratio of 1:400 with G-actin), and streptavidin (in a molar ratio of 1:25 with G-actin). After incubation, the liposomes were flown from the formation chamber into the adjacent observation chamber and diluted by injecting 120300 μL of O-buffer with 500 mM glucose. The osmolarity of the O-buffer (Osmomat 030, Gonotec GmbH) was adjusted to 20 mOsmol/kg higher than that of the I-buffer to make the liposomes sufficiently floppy for contour fluctuation analysis by video microscopy.48 Sucrose and glucose provide sufficient optical contrast for visualization of the membranes by phase contrast microscopy; moreover, the slight difference in mass density 10063

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Langmuir causes liposome sedimentation to the bottom of the observation chamber. Microscopy and Image Analysis. We observed liposomes with a confocal laser scanning microscope using a Nikon C1 scanhead on a Nikon Eclipse Ti inverted microscope with 100/NA1.4 or 40/ NA1.0 Plan Apo objectives. Images were captured using EZ-C1 software (Nikon, version 3.10). AlexaFluor 488 was excited using a diodepumped solid-state laser with a wavelength of 488 nm (Melles-Griot); rhodamine and DyLight 594 were excited using a heliumneon laser with a wavelength of 543 nm (Melles-Griot). The liposome diameter, lamellarity, and encapsulation efficiency were analyzed from fluorescence images of equatorial cross sections of liposomes. The sizes of liposomes before harvesting (in the formation chamber) and after harvesting (in the observation chamber) were determined by detecting the center of mass position and diameter of each liposome. Since the liposomes were densely packed, especially in the formation chamber, we used a method based on the Hough transform, which can distinguish liposomes even when they are in contact. This method groups edge points into object candidates by performing an explicit voting procedure over a set of parametrized image objects.49,50,51 Fluorescence intensities of membranes for lamellarity analysis were measured from confocal images of liposomes in the observation chamber. For these dilute and well-separated liposomes, we used a custom-written MatLab program based on Canny edge detection.49 Two concentric circles were centered on the liposome to create a ring-shaped area enclosing the fluorescent membrane contour; the width of the ring was fixed for all liposomes. The image was thresholded to eliminate pixel intensity values below the noise level. To obtain the average pixel intensity for each membrane, the intensity values of pixels along the membrane contour were summed and normalized by the total number of pixels. To compare the F-actin content inside liposomes to the original actin content of the hydration buffer, confocal images of liposomes in the formation chamber were recorded. The average fluorescence intensity of F-actin inside a liposome (Ii) was obtained as follows. First, since F-actin was present both inside and outside liposomes, we identified the liposome boundaries from fluorescence images of rhodamine-labeled membranes. Then we summed the pixel intensities in the AlexaFluor 488-actin channel of all pixels within the boundary and finally divided the total intensity by the total number of pixels. To quantify the F-actin density outside the liposomes, we obtained the average fluorescence intensity (Io) of the entire field of view excluding areas of liposomes. The encapsulation ratio was defined as Ii/Io. To measure the variance in actin encapsulation among liposomes, we used confocal images of liposomes harvested from the formation chamber by flowing them into the observation chamber with O-buffer. We indentified liposomes by thresholding fluorescence images of AlexaFluor 488-actin. To ensure that only pixels inside liposomes were measured, each liposome image was first dilated and then eroded by a 5 pixel sized circle structure element.49 We then summed the intensities of all measured pixels and divided the total intensity by the number of pixels to obtain the average fluorescence intensity. Liposome Contour Analysis. Membrane fluctuation spectra were measured by phase contrast microscopy using a Nikon Eclipse Ti inverted microscope with a 100/NA1.4 Plan Apo objective. At least 500 consecutive images of the equatorial cross-section of a liposome were recorded with a digital CCD camera (CoolSNAP HQ2, Nikon). Membrane edge detection with subpixel resolution was achieved with a custom-written MatLab program based on a published method.48 Due to the refractive index difference between the buffers inside and outside the liposome, the liposome’s edge is silhouetted dark against the surrounding background in phase contrast, with a bright halo on the outside, as illustrated in Figure S1A, Supporting Information. We detected the edge positions of a liposome contour in four directions: horizontal, vertical, and two diagonal directions, as shown in Figure S1B.

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Figure S1C exemplifies the edge finding procedure for the horizontal direction. A gray level profile along a line of 2024 pixels perpendicular to the membrane was constructed, and a straight line was fitted to the portion of the profile having the maximum slope over a range of 812 pixels. The edge position was defined as the intersection of this line with a line corresponding to I = Iav, where Iav is the average gray level of the gray level profile. The membrane position was calculated as the average of the positions obtained from the four directions weighted by the corresponding slopes of the intensity profiles. This edge detection procedure provides a high spatial resolution of 0.1 pixel (6 nm with our setup). The detected positions in Cartesian coordinates (x and y) were transformed to polar coordinates (r and θ). The contours were projected to a Fourier series with 50 modes: rðθÞ ¼ Rð1 þ

50

∑ an cosðnθÞ þ bn sinðnθÞÞ n¼1

ð1Þ

where R is the liposome radius and n is the mode number. The power spectrum of the transverse thermal fluctuations, u, of the membrane is defined as Æjuðqx Þj2 æ ¼

πÆRæ3 ðÆjcn j2 æ  Æjcn jæ2 Þ 2

ð2Þ

where cn = (an2 + bn2)1/2, ÆRæ is the time-averaged liposome radius, and the wave vector qx = n/ÆRæ. According to the classical Helfrich model of fluctuating two-dimensional membranes,48,52 the time-averaged fluctuation spectrum decays with wave vector q as ÆjuðqÞj2 æ ¼

kB T σq2 þ kq4

ð3Þ

where q = (qx,qy), the thermal energy kBT = 4  1021 J, σ is the membrane tension, and k is the membrane bending rigidity. This general expression needs to be modified since we only observed the equatorial plane of each liposome and therefore the spectrum is one-dimensional. Moreover, we need to account for smearing of fluctuations whose lifetime is shorter than the camera integration time. Taking these two factors into account, the measured membrane fluctuation spectrum was fitted to a modified Helfrich model:48 !   Z 1 ∞ kB T τ m 2 τ τ 2 τm  1 dqy Æjuðqx Þj æ ¼ þ exp  ð4Þ π ∞ 4ηq τ2 τm τm where η is the buffer viscosity, τm is the fluctuation lifetime, τm(q)1 = (1/4ηq)(σq2 + kq4), and τ is the camera integration time (1020 ms with our camera setting). We excluded the first five modes from the fit, which was necessary due to the closed topology of the membrane, as well as modes with wave vectors above 2 μm1, since these modes have a wavelength less than the pixel size (60 nm) and a spectral density in the noise floor.48

’ RESULTS AND DISCUSSION Encapsulation of cytoskeletal proteins in giant liposomes presents a long-standing technical challenge. High liposome yield is generally incompatible with physiological salt levels, and the yield is even further reduced with inclusion of proteins. Though several strategies have been described which overcome these difficulties, there is currently no facile and robust method to achieve this encapsulation. We present a new approach based on gentle hydration of hybrid lipidagarose films in solutions of cytoskeletal proteins, inspired by a recent study of Horger and co-workers,37 where liposomes were prepared from a variety of lipids under physiological ionic conditions. Our method proceeds in three stages, as sketched in Figure 1A. First, a uniform 10064

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Langmuir layer of ultralow gelling agarose is spin-coated on a glass slide and partially dried (panel i). In contrast to standard gelling temperature agarose that forms fibrous, cross-linked gels, ultralow gelling agarose forms loose meshes of agarose molecules that partially dissolve upon hydration.37 Second, a lipid solution is spin-coated on the agarose layer and dried under vacuum (panel ii). Third, the hybrid film is hydrated in a buffer solution containing monomeric (G) actin and (optionally) myosin II (panel iii). It has been proposed in the original paper of Horger et al.37 that swelling and partial dissolution of the agarose film upon hydration promote all three stages of liposome formation. The large contact area of the lipids with the porous agarose film and the prehydration of the lipids by the residual water may facilitate lipid selfassembly into bilayers. Swelling of the agarose gel may generate forces normal to the lipid bilayers that promote bilayer separation. The dissolved agarose molecules may enhance water influx into the lipid film by generating hyperosmotic conditions. Furthermore, it was proposed that crowding of the liposomes that grow out from the porous agarose film promotes their fusion into giant liposomes. Dissolved agarose molecules were shown to associate with the lipid membranes after liposome formation, and it was proposed that they may promote liposome formation in a fashion similar to that of PEGylated lipids.25 Liposome Formation and Protein Encapsulation. Figure 1B shows giant liposomes containing F-actin that formed when films of agarose and lipids were hydrated in a physiological buffer (“I-buffer”). The I-buffer contains 4 mg/mL G-actin at a pH (7.4) and a salt level (50 mM KCl and 2 mM MgCl2) that promotes polymerization of G-actin into long filaments. The filaments formed homogeneous, entangled networks inside the liposomes as well as in the external “O-buffer” solution (the liposome suspension was diluted about 5-fold with 255 mM glucose). We could digest the external F-actin by supplementing the O-buffer with DNaseI, which complexes G-actin.53 The liposome yield was high: we typically found 50100 liposomes in the observation chamber after 4-fold dilution. Most liposomes were spherical and freely floating, while some attached to the bottom surface of the observation chamber. Strikingly, all liposomes in the observation chamber contained actin. In the presence of myosin II, 85% of the liposomes (out of 60) contained actin and myosin II. The remainder (15%) contained only actin. For comparison, we also made actin-filled liposomes with the inverted emulsion technique.24 In our hands, the liposome yield was high (in the hundreds) when we encapsulated sucrose in water, but low when we encapsulated actin in I-buffer. We found a maximum of 15 actin-filled liposomes per sample, much less than with the agarose swelling method. It is unclear what factors determine the liposome yield. There are examples of encapsulation of proteins in high-salt solutions that do give high yields.54 We also observed that liposomes obtained by the inverted emulsion technique tended to have attached oil droplets or lipid aggregates that complicate measurements of membrane fluctuations by video microscopy (Figure S2A, Supporting Information). In contrast, liposomes obtained by swelling of agaroselipid films had cleaner contours (Figure S2B). The presence of an agarose film was essential for the formation of liposomes in the presence of proteins and salts. Lipid films deposited directly on glass substrates did not form giant liposomes, as shown in Figure 1D. The few liposomes that did form were smaller than 20 μm, did not contain actin, and were predominantly multilamellar. This looks strikingly different from the formation chamber in the presence of an agarose film, where

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large liposomes containing actin are seen (Figure 1C). The reason why high salt concentrations hamper liposome formation from lipids coated on bare glass is uncertain. Although some mechanisms for liposome formation have been proposed,55 the exact molecular mechanism remains uncertain. It is thought that solutions of high ionic strength hinder in particular the first step of separation of lipid lamellae due to electrostatic screening of charges on the lipid head groups. The second step of liposome swelling is probably also influenced by the presence of salt. We doped the membranes with 5 mol % PEG-DOPE lipids bearing a 2 kDa PEG tail to prevent nonspecific membrane adsorption of actin and to reduce aggregation of liposomes.22,37 We found that PEG-DOPE was also indispensable to obtain actin-filled liposomes in high yield. In the absence of PEGylated lipids, liposomes were rarely formed and they did not contain actin, as shown in Figure 1E. This suggests that steric repulsions of the PEGylated lipids may contribute to liposome formation by promoting initial bilayer orientation and separation, similar to their role in gentle swelling on bare glass.25,56 In the study of Horger et al.,37 liposomes did form with high yield from pure POPC, but PEGylated lipids increased the yield. This difference may be caused by different buffer conditions (such as the presence of Mg2+ in the actin buffer) or by the presence of actin during formation. In addition to agarose and PEGylated lipids, the spin-coating step was crucial to obtain a good liposome yield and efficient actin encapsulation. When the agarose and lipid layers were dip-coated instead of spin-coated, most regions of the formation chamber did not have liposomes (upper panel of Figure 1F); liposomes did form in a few regions of the formation chamber, but they did not encapsulate actin (bottom panel of Figure 1F). We suspect that spin-coating improves the uniformity of the agarose and lipid films, which is apparently beneficial for liposome formation. Similar effects have been observed with lipid mixtures on indium tin oxide plates in the electroformation method.57 To test whether dissolved agarose entered the liposomes, we formed liposomes using an agarose gel containing 0.1% (w/w) FITC-labeled agarose. We measured the fluorescence intensities of FITC-agarose inside and outside liposomes right after formation using confocal microscopy (Figure S3A,D, Supporting Information). Interestingly, we observed that actin-filled liposomes did not contain agarose at a detectable level (Figure S3B), whereas liposomes devoid of actin did contain agarose (Figure S3C). Size Distribution and Membrane Lamellarity. Liposome size distributions were obtained by imaging their equatorial cross sections using confocal laser scanning microscopy, with identical laser and camera settings for all images. Liposomes adhering to the agarose film in the formation chamber displayed a wide distribution of diameters, ranging from 14 to 76 μm with an average diameter of 24 ( 10 μm, as shown in Figure 2 (open bars). Liposomes removed from the chamber and suspended in O-buffer were almost 2-fold smaller, with an average diameter of 13 ( 4 μm (gray bars in Figure 2). The larger size of the liposomes in the formation chamber may be caused by fusion of adjacent liposomes within the 4 h time span required for image recording. In addition, large liposomes are more likely to break when they are flown into the observation chamber. The liposome sizes are comparable with the sizes of actin-filled liposomes obtained with gentle hydration25 and the inverted emulsion method.24 In our hands, liposomes containing actin in I-buffer prepared with the inverted emulsion method had diameters of 10065

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Figure 2. Size distribution of giant actin-filled liposomes determined from confocal fluorescence images of equatorial cross sections. Empty bars correspond to liposomes in the formation chamber measured within 4 h after formation (right-hand y-axis, 614 liposomes counted), while gray bars correspond to liposomes harvested from the formation chamber by injection of O-buffer (left-hand y-axis, 96 liposomes counted).

approximately 10 μm, comparable to liposomes prepared with the new agarose method. However, the agarose method produced at least 10-fold more liposomes per sample and also generated actin-filled liposomes with diameters larger than 10 μm that are rarely found with the inverted emulsion method. For the membranes to serve as a faithful model of the cell membrane, it is important that the membrane be unilamellar. Liposomes containing 5 mol % PEGylated lipids are expected to be unilamellar.25 To confirm unilamellarity, we measured the mechanical properties of the membranes by video tracking of thermally driven membrane undulations. We formed liposomes with an inner solution of 0.016% (w/w) sucrose in water and diluted them in an aqueous solution of 330 mM glucose with slightly higher osmolarity (by 20 mOsmol/kg) to reduce the membrane surface tension.48 Free-floating, spherical liposomes were observed by phase contrast microscopy, and spectra of transverse bending undulations were extracted by image analysis. Figure 3A shows a typical fluctuation spectrum of a liposome encapsulating a sucrose solution (black squares). The bending rigidity and membrane tension were determined from the spectrum by fitting to a modifed Helfrich model,48,52 taking into account the effect of the finite camera integration time (black line). Figure 3B shows the bending modulus k of various liposomes with different diameters (black circles). We found bending moduli between 0.8  1019 and 2.7  1019 J with an average value of 1.7  1019 J (42.5 kBT). The large spread in the data is typical when determining bending moduli of liposomes by contour analysis.58 The average membrane tension was 3.6  107 N/m, comparable to prior reports.48 The average k value is 2-fold larger than that of liposomes composed exclusively of DOPC.59 However, that value was obtained by micropipette aspiration, which is known to yield smaller values for k than contour analysis.60 Furthermore, the PEGylated lipids used in our lipid formulation are expected to increase k compared to that of pure DOPC membranes, since they increase the membrane thickness61 and k scales as membrane thickness squared.62 Still, it is possible that the elevated moduli originate from residual agarose molecules in the lipid layer.37 To test this effect, we measured bending moduli for liposomes of the exact same lipid composition but prepared by electroformation. The electroformed liposomes had a range of k values (red diamonds in Figure 3B) and an average k value (1.1  1019 J) similar to those of the liposomes prepared by gentle swelling. The membrane tension (mean value 2  107 N/m) was also comparable to that of liposomes obtained by gentle swelling. This suggests

Figure 3. Analysis of liposome lamellarity by membrane bending modulus measurements and fluorimetric estimation. (A) Representative membrane fluctuation spectra of liposomes encapsulating a sucrose solution (black squares) or 1 mg/mL F-actin (red circles). The spectrum of the sucrose-filled liposome is fitted to the modifed Helfrich model,48 considering the influence of camera integration time to the fitting (black straight line). (B) Elastic bending modulus k of empty liposomes as a function of the liposome diameter. Every symbol corresponds to one liposome (black circles, agarose-swelling method; red diamonds, electroformation). (C) Left panel: average fluorescence intensity (F.I.) of liposome membranes as a function of the liposome diameter (65 liposomes measured). Right panel: projection of data points in the left panel on the average fluorescence intensity axis.

that residual agarose has no measurable influence on membrane stiffness or tension. We also measured membrane fluctuation spectra for a few actin-filled liposomes, as exemplified in Figure 3A by the red circles. These spectra can in principle be fitted to the Helfrich relation, but this fit returns an unphysical tension of 4  1021 N/m. This is probably a signature of the viscoelastic material properties of the encapsulated actin network, which is not captured in the Helfrich model that assumes a viscous liquid interior. In follow-up work, we plan to extend membrane fluctuation models, taking into account the viscoelastic nature of the actin network63 and the contractile activity of myosin.64 The close correspondence of the mechanical behavior of the liposomes prepared by gentle swelling to that of electroformed liposomes suggests that they are predominantly unilamellar.18 To independently assess membrane lamellarity, we analyzed the fluorescence intensity of the membranes of 10066

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Figure 4. Encapsulation efficiency of liposomes determined from confocal fluorescence images. (A) Histogram of the F-actin encapsulation ratio defined as the average fluorescence intensity inside a liposome (Ii) divided by the average fluorescence intensity outside the liposomes (Io). Filled bars: 1 mg/mL F-actin, left-hand y-axis, 76 liposomes counted. Empty bars: 0.25 mg/mL F-actin, right-hand y-axis, 35 liposomes counted. (B) Left panel: dependence of the encapsulated F-actin (1 mg/mL) density, measured as the average fluorescence intensity (F.I.), on the liposome diameter (88 liposomes counted). Right panel: histogram of the F-actin content of liposomes fitted by a Gaussian distribution (red line).

actin-filled liposomes. Figure 3C (left panel) shows the average fluorescence intensity of liposome membranes as a function of the liposome diameter, measured for 65 freely suspended liposomes in the observation chamber. The distribution of intensities is rather broad with no clear peak(s), and there is no apparent dependence on the liposome diameter. However, when we project the data to their corresponding average fluorescence intensity (Figure 3C (right panel)), there appear to be two groups of liposomes: one group with membrane intensities of 400 800 arbitrary units (63% of the liposomes) and another group with intensities of 10001200 arbitrary units (20% of the liposomes). This suggests that a small fraction of the liposomes may be multilamellar. Encapsulation Efficiency. An important application of actinfilled liposomes is as cell-mimetic systems for quantitative measurements of mechanics and force generation. To fulfill this promise, it is desirable that the encapsulated amount of actin be close to the target concentration with low variance among liposomes. To estimate the encapsulation efficiency, we measured the average fluorescence signal for F-actin inside liposomes (Ii) and outside liposomes (Io) in the formation chamber. For a bulk actin concentration of 1 mg/mL, the encapsulation ratio, defined as Ii/Io, was on average 56 ( 26%, as shown in Figure 4 (A). This is comparable with other studies65 which report typical solute encapsulation efficiencies below 50%. Some liposomes had apparent encapsulation ratios larger than 100%. This could be due to inhomogeneities in the highly viscous F-actin solution outside the liposomes. However, encapsulation efficiencies higher than 100% were also reported for encapsulation of several

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other macromolecular solutes.66,67 The underlying mechanism of this phenomenon, which has been termed “spontaneous protein crowding”,67 is unclear. It was suggested that nonspecific solutelipid interactions may increase the amount of solute entering between the lipid lamella during hydration.66 To test whether the encapsulation efficiency depends on the actin concentration, we compared encapsulation efficiencies for 1 and 0.25 mg/mL actin, as shown in Figure 4A. There was no significant concentration dependence of the efficiency. In future, the encapsulation efficiency may potentially be enhanced by using lipids with a functional group to bind the proteins65 or by adding a crowding agent.68 The encapsulation variance among liposomes was estimated by comparing the average fluorescence signal of F-actin in liposomes flown from the formation chamber into the observation chamber by injecting O-buffer. Figure 4 B shows that the average fluorescence intensity is independent of the liposome diameter (left) and well-described by a Gaussian distribution (red line) with a peak at a fluorescence intensity of 421 ( 33 arbitrary units. This narrow distribution indicates a low variance in the encapsulated amount of actin among liposomes. ActinMembrane Interaction. For the liposomes to faithfully mimic the mechanics of biological cells, it is important that the actin network can be attached to the membrane by specific anchors of controlled density. To prevent nonspecific membrane adhesion of actin, we included 5 mol % PEGylated lipids in the lipid mixture, which act as a steric brush. The membranes were indeed passivated successfully, as shown by confocal imaging (Figure 5A): while the actin network (green) is homogeneous throughout the liposome, near the membrane (red) an empty depletion zone devoid of actin is observed, as expected for rigid actin filaments near nonbinding surfaces.69 The depletion of actin close to the membrane is clearly visible in a fluorescence intensity cross-section of the liposome, as shown in Figure 5B. To create well-defined actinmembrane anchoring, we used biotinstreptavidin bonds, which are nearly permanent. When actin was doped with biotinylated G-actin in a 1:400 molar ratio (ca. 1 biotin per 1 μm length of filament) but the membrane was free of biotin, inclusion of streptavidin (1:25 molar ratio with G-actin) in the I-buffer caused the formation of actin bundles which did not adhere to the membrane, as shown in Figure 5C. However, upon inclusion of biotinylated lipids in the membrane, the biotinylated actin network adhered to the inner surface of the membrane and formed a cortexlike structure, leaving a depleted zone in the liposome center, as shown in Figure 5D. This structure is reminiscent of cortical actin structures observed in electroformed liposomes with electrostatic actinlipid attraction.53,70 It differs from actin cortices encapsulated by the inverted emulsion technique,24 where actin was grown from the membrane using actin-nucleating factors and was actively tread-milling. Encapsulation of Active Networks. The actin cytoskeleton in vivo actively exerts contractile forces on the membrane by the activity of myosin motors that assemble into bipolar filaments and slide actin filaments past each other. To the best of our knowledge, the only prior myosin encapsulation into liposomes was done with heavy meromyosin (HMM), a double-headed derivative of myosin II.40 HMM is nonprocessive with a duty ratio of only a few percent.71 Nevertheless, HMM motors can collectively generate contraction of actin filament networks and bundles.41 However, bipolar filaments are likely more effective since they are more processive, and they more closely mimic the cytoskeleton of nonmuscle cells, in which nonmuscle myosin II self-assembles into bipolar minifilaments composed of 1020 10067

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Figure 5. (A, C, D) Confocal fluorescence micrographs of giant actinfilled liposomes. Lipid membranes are labeled with rhodamine (red); actin is labeled with AlexaFluor 488 (green). The actin concentration in the I-buffer before encapsulation was 1 mg/mL. Insets indicate the nature of the actinmembrane interaction. (A) F-actin solution inside a liposome with an inert membrane (containing PEG-lipids) is homogeneous and displays a depletion zone underneath the membrane. (B) Fluorescence intensity profiles along the yellow line across the liposome in (A). Upper panel: membrane fluorescence intensity along the line scan. Bottom panel: actin fluorescence intensity along the line scan. (C) A cross-linked F-actin network inside a liposome with an inert membrane is inhomogeneous and bundled and does not attach to the membrane. (D) Liposomes which contain biotinylated lipids encapsulate networks of biotinylated actin filaments that are coupled to the membrane via biotinstreptavidin bonds. Scale bars 10 μm. Biotinylated G-actin to G-actin molar ratio 1:400.

myosins.72 We used large bipolar filaments of skeletal muscle myosin II motors with a length of around 1 μm, as shown in Figure 6A. These filaments contain about 100 myosins/ filament.39b Bulk networks of actin filaments and myosin bipolar filaments are initially homogeneous, but within 5 min the motor filaments self-assemble into large clusters39b (shown in green in Figure 6B) within the actin network (shown in red). Using the agarose-swelling method, we were able to encapsulate myosin filaments together with actin into liposomes, as shown in Figure 6C,D. The myosin:actin ratio is 1:200, and the membrane contains biotinylated lipids to bind to the biotinylated actin filaments. Similar to their organization in bulk networks, myosin formed clusters (shown in green) within the entangled F-actin network. Interestingly, there were two classes of liposomes. Half of the liposomes (out of 63) contained a cortexlike actin myosin structure (Figure 6C), whereas the other half contained a homogeneous actomyosin network (Figure 6D). We suspect that this variability is related to a variable myosin encapsulation efficiency. We were unable to quantify the encapsulation efficiency for myosin, since myosin was not homogeneously distributed in the encapsulated actin networks, but concentrated in dense foci. However, we expect that it is less efficiently encapsulated than actin, since the hydrodynamic radius of myosin (45 nm73) is much larger than that of actin (2 nm74). Prior studies showed that solutes with hydrodynamic radii

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Figure 6. (A) Typical electron micrograph of a bipolar myosin filament. Scale bar 200 nm. (BF) Confocal fluorescence micrographs of active F-actin solutions containing myosin II filaments in a bulk solution (B) (scale bar 20 μm) and inside liposomes (CF) (scale bars 10 μm). Actin (concentration 1 mg/mL) is labeled with AlexaFluor 488 (shown in red); myosin is labeled with DyLight 594 (shown in green). The F-actin network adheres to the membrane by biotinstreptavidin bonds. Biotinylated G-actin to G-actin molar ratio 1:400. Myosin II to G-actin molar ratio: (C, D) 1:200; (E) 1:100; (F) 1:50.

above 10 nm are encapsulated much less efficiently than smaller ones.66,68b The cortexlike structures appear to be more strongly anchored to the membrane than the homogeneous actomyosin networks. A possible explanation is that the homogeneous networks contain more myosin and are therefore more contractile, leading to rupture of actinlipid bonds. To test whether myosin-driven contractility can cause actin membrane rupture, we encapsulated actinmyosin networks with higher myosin:actin ratios of 1:100 (Figure 6E) and 1:50 (Figure 6F). We found that, at higher myosin densities, the myosin filaments condensed into one or a few large clusters, which contracted actin filaments into a shell-like structure around them. We did not find any cortexlike structures at these high myosin:actin ratios, indicating that motor-driven forces can indeed rupture actinmembrane attachments. The giant actomyosin-filled liposomes developed in this work can be used as a minimal model system to study the physical principles that govern the organization and mechanics of the actomyosin cytoskeleton in cells. To enhance the physiological relevance, it would be interesting to increase the actin concentration. The actin concentrations used here (∼0.6 mg/mL inside the liposomes) are at the low end of the range reported for 10068

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Langmuir mammalian, nonmuscle cells. Concentrations of actin in vivo depend on the organism and cell type, the proteins are generally inhomogeneously distributed in cells, and the distribution changes during the cell cycle. Global estimates of F-actin concentrations in nonmuscle cells range from 12 to 300 μM (from 0.5 to 12 mg/mL).75 Locally, the F-actin concentration can be higher, for instance, 500 μM (20 mg/mL) in the lamellipodium of migrating cells.76 The myosin:actin molar ratios used in our work are well within the range reported for nonmuscle cells, which is from 1:10 to 1:100 depending on the cell type.77 The coupling of the actomyosin networks to the membrane can be made more physiological by introducing proteins responsible for coupling in vivo, including PIP2-NWASP13c and spectrin/ankyrin proteins.78

’ CONCLUSIONS In the present work, we demonstrate a new preparation method for liposomes containing active actomyosin networks by gentle swelling of hybrid agaroselipid films in physiological buffers. This method is facile and fast, taking less than 1 h for liposome formation. We show that this method gives a high yield and leads to uniform encapsulation of actin among liposomes. Moreover, the encapsulated networks can be selectively anchored to the inner leaflet of the membrane via biotinstreptavidin linkages. This new method opens the way for quantitative studies of physical mechanisms by which the actin cytoskeleton and the plasma membrane jointly control cell shape and mechanics. ’ ASSOCIATED CONTENT

bS

Supporting Information. Explanation of the contour detection method, phase contrast images of liposomes encapsulating an actomyosin network obtained from the agarose-swelling method and inverted emulsion technique, and confocal fluorescence images of FITC-agarose in liposomes. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected].

’ ACKNOWLEDGMENT This work is part of the research program of the Foundation for Fundamental Research on Matter (FOM), which is financially supported by The Netherlands Organisation for Scientific Research (NWO). This work was financially supported by a VIDI grant from NWO (F.-C.T. and G.H.K.) and a Marie Curie IE Fellowship (B.S.). We thank Dr. A. Bertin (financially supported by Agence Nationale pour la Recherche Grant ANR-09-RPDOC026-01) for taking electron micrographs of myosin at the Plateforme de Microscopie Ultrastructurale of the Institut Pasteur in Paris, France. We benefited from helpful discussions with Dr. B. Alonso Latorre regarding image analysis and Dr. N. Becker, Dr. S. Semrau, Dr. J. Pecreaux, Dr. R. Rodríguez-García, Dr. M. Mell, and Dr. I. Lopez-Montero regarding analysis of liposome fluctuation spectra. Finally, we thank Dr. K. Carvalho and Dr. C. Sykes for advice on the inverted emulsion technique.

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’ REFERENCES (1) Alberts, B.; Bray, D.; Lewis, J.; Raff, M.; Roberts, K.; Watson, J. D. Molecular Biology of the Cell, 4th ed.; Garland Science: New York, 2001. (2) Morone, N.; Fujiwara, T.; Murase, K.; Kasai, R.; Ike, H.; Yuasa, S.; Usukura, J.; Kusumi, A. Three-dimensional reconstruction of the membrane skeleton at the plasma membrane interface by electron tomography. J. Cell Biol. 2006, 174 (6), 851–862. (3) Bugyi, B.; Carlier, M. Control of actin filament treadmilling in cell motility. Annu. Rev. Biophys. 2010, 39, 449–470. (4) Pollard, T.; Borisy, G. G. Cellular motility driven by assembly and disassembly of actin filaments. Cell Motil. Cytoskeleton 2003, 112, 453–465. (5) Howard, J. Mechanics of Motor Proteins and the Cytoskeleton; Sinauer: Sunderland, MA, 2001. (6) (a) Charras, G.; Yarrow, J.; Horton, M.; Mahadevan, L.; Mitchison, T. Non-equilibration of hydrostatic pressure in blebbing cells. Nature 2005, 435 (7040), 365–369. (b) Paluch, E.; Piel, M.; Prost, J.; Bornens, M.; Sykes, C. Cortical actomyosin breakage triggers shape oscillations in cells and cell fragments. Biophys. J. 2005, 89 (1), 724–733. (7) Engler, A.; Sen, S.; Sweeney, H.; Discher, D. Matrix elasticity directs stem cell lineage specification. Cell 2006, 126 (4), 677–689. (8) Doherty, G. J.; McMahon, H. T. Mediation, modulation, and consequences of membrane-cytoskeleton interactions. Annu. Rev. Biophys. 2008, 37, 65–95. (9) Liu, A.; Fletcher, D. Biology under construction: In vitro reconstitution of cellular function. Nat. Rev. Mol. Cell Biol. 2009, 10 (9), 644–650. (10) Tanaka, M.; Sackmann, E. Polymer-supported membranes as models of the cell surface. Nature 2005, 437, 656–663. (11) (a) Walder, R.; Levine, A.; Dennin, M. Rheology of twodimensional F-actin networks associated with a lipid interface. Phys. Rev. E 2008, 77 (1), 011909. (b) Sengupta, K.; Limozin, L. T., M; Haase, I.; Fischer, M.; Sackmann, E. Coupling artificial actin cortices to biofunctionalized lipid monolayers. Langmuir 2006, 22 (13), 5776–5785. (12) (a) Barfoot, R.; Sheikh, K.; Johnson, B.; Colyer, J.; Miles, R.; Jeuken, L.; Bushby, R.; Evans, S. Minimal F-actin cytoskeletal system for planar supported phospholipid bilayers. Langmuir 2008, 24 (13), 6827–6836. (b) Johnson, B.; Bushby, R.; Colyer, J.; Evans, S. Selfassembly of actin scaffolds at ponticulin-containing supported phospholipid bilayers. Biophys. J. 2006, 90 (3), L21–3. (c) Lee, K.; Gallop, J.; Rambani, K.; Kirschner, M. Self-assembly of filopodia-like structures on supported lipid bilayers. Science 2010, 329 (5997), 1341–1345. (13) (a) Isanta, S.; Espinosa, G.; Rodrıguez-Garcıa, R.; Natale, P.; Lopez-Montero, I.; Langevin, D.; Monroy, C. Active membranes with bound F-actin: Sliding vs. sticking conditions. Soft Matter 2011, 7, 3100. (b) Helfer, E.; Harlepp, S.; Bourdieu, L.; Robert, J.; MacKintosh, F.; Chatenay, D. Microrheology of biopolymer-membrane complexes. Phys. Rev. Lett. 2000, 85 (2), 457–460. (c) Liu, A.; Fletcher, D. Actin polymerization serves as a membrane domain switch in model lipid bilayers. Biophys. J. 2006, 91 (11), 4064–4070. (d) Heuvingh, J.; Franco, M.; Chavrier, P.; Sykes, C. ARF1-mediated actin polymerization produces movement of artificial vesicles. Proc. Natl. Acad. Sci. U.S.A. 2007, 104 (43), 16928–16933. (e) Giardini, P.; Fletcher, D.; Theriot, J. Compression forces generated by actin comet tails on lpid vesicles. Proc. Natl. Acad. Sci. U.S.A. 2003, 100 (11), 6493–6498. (f) Upadhyaya, A.; Chabot, J.; Andreeva, A.; Samadani, A.; vanOudenaarden, A. Probing polymerization forces by using actin-propelled lipid vesicles. Proc. Natl. Acad. Sci. U.S.A. 2003, 100 (8), 4521–4526. (g) Delatour, V.; Helfer, E.; Didry, D.; L^e, K.; Gaucher, J.; Carlier, M.; Romet-Lemonne, G. Arp2/3 controls the motile behavior of N-WASP-functionalized GUVs and modulates N-WASP surface distribution by mediating transient links with actin filaments. Biophys. J. 2008, 94 (12), 4890–4905. (14) Takiguchi, K.; Yamada, A.; Negishi, M.; Honda, M.; TanakaTakiguchi, Y.; Yoshikawa, K. Chapter 3—Construction of cell-sized liposomes encapsulating actin and actin-cross-linking proteins. Methods Enzymol. 2009, 464, 31–53. (15) (a) Elbaum, M.; Fygenson, D.; Libchaber, A. Buckling microtubules in vesicles. Phys. Rev. Lett. 1996, 76, 4078–4081. (b) Emsellem, 10069

dx.doi.org/10.1021/la201604z |Langmuir 2011, 27, 10061–10071

Langmuir V.; Cardoso, O.; Tabeling, P. Vesicle deformation by microtubules: A phase diagram. Phys. Rev. E 1998, 58, 4807–4810. (16) Shaklee, P.; Semrau, S.; Malkus, M.; Kubick, S.; Dogterom, M.; Schmidt, T. Protein incorporation in giant lipid vesicles under physiological conditions. ChemBioChem 2010, 11, 175–179. (17) Montes, L. R.; Alonso, A.; Goni, F. M.; Bagatolli, L. A. Giant unilamellar vesicles electroformed from native membranes and organic lipid mixtures under physiological conditions. Biophys. J. 2007, 93 (10), 3548–54. (18) Bagatolli, L. A.; Parasassi, T.; Gratton, E. Giant phospholipid vesicles: Comparison among the whole lipid sample characteristics using different preparation methods: A two photon fluorescence microscopy study. Chem. Phys. Lipids 2000, 105 (2), 135–47. (19) Angelova, M.; Soleau, S.; Meleard, P.; Faucon, J.; Bothorel, P. Preparation of giant vesicles by external AC fields. Kinetics and application. Prog. Colloid Polym. Sci. 1992, 89, 127–131. (20) Pott, T.; Bouvrais, H.; Meleard, P. Giant unilamellar vesicle formation under physiologically relevant conditions. Chem. Phys. Lipids 2008, 154 (2), 115–119. (21) Miyata, H.; Nishiyama, S.; Akashi, K.; Kinosita, K., Jr. Protrusive growth from giant liposomes driven by actin polymerization. Proc. Natl. Acad. Sci. U.S.A. 1999, 96 (5), 2048–2053. (22) Limozin, L.; Barmann, M.; Sackmann, E. On the organization of self-assembled actin networks in giant vesicles. Eur. Phys. J. E 2003, 10 (4), 319–330. (23) Zhang, Y.; Cheng, C.; Cusick, B.; LeDuc, P. Chemically encapsulated structural elements for probing the mechanical responses of biologically inspired systems. Langmuir 2007, 23 (15), 8129–8134. (24) Pontani, L. L.; van der Gucht, J.; Salbreux, G.; Heuvingh, J.; Joanny, J. F.; Sykes, C. Reconstitution of an actin cortex inside a liposome. Biophys. J. 2009, 96 (1), 192–198. (25) Yamashita, Y.; Oka, M.; Tanaka, T.; Yamazaki, M. A new method for the preparation of giant liposomes in high salt concentrations and growth of protein microcrystals in them. Biochim. Biophys. Acta 2002, 1561 (2), 129–134. (26) Castile, J. D.; Taylor, K. M. Factors affecting the size distribution of liposomes produced by freeze-thaw extrusion. Int. J. Pharm. 1999, 188 (1), 87–95. (27) Moscho, A.; Orwar, O.; Chiu, D. T.; Modi, B. P.; Zare, R. N. Rapid preparation of giant unilamellar vesicles. Proc. Natl. Acad. Sci. U.S.A. 1996, 93 (21), 11443–11447. (28) (a) Stachowiak, J. C.; Richmond, D. L.; Li, T. H.; BrochardWyart, F.; Fletcher, D. A. Inkjet formation of unilamellar lipid vesicles for cell-like encapsulation. Lab Chip 2009, 9 (14), 2003–2009. (b) Stachowiak, J. C.; Richmond, D. L.; Li, T. H.; Liu, A. P.; Parekh, S. H.; Fletcher, D. A. Unilamellar vesicle formation and encapsulation by microfluidic jetting. Proc. Natl. Acad. Sci. U.S.A. 2008, 105 (12), 4697–4702. (c) Ota, S.; Yoshizawa, S.; Takeuchi, S. Microfluidic formation of monodisperse, cellsized, and unilamellar vesicles. Angew. Chem., Int. Ed. 2009, 48 (35), 6553–6537. (29) Shum, H. C.; Lee, D.; Yoon, I.; Kodger, T.; Weitz, D. A. Double emulsion templated monodisperse phospholipid vesicles. Langmuir 2008, 24 (15), 7651–7653. (30) Richmond, D.; Schmid, E.; Martens, S.; Stachowiak, J.; Liska, N.; Fletcher, D. Forming giant vesicles with controlled membrane composition, asymmetry, and contents. Proc. Natl. Acad. Sci. U.S.A. 2011, 108 (23), 9431–9436. (31) Funakoshi, K.; Suzuki, H.; Takeuchi, S. Formation of giant lipid vesiclelike compartments from a planar lipid membrane by a pulsed jet flow. J. Am. Chem. Soc. 2007, 129, 12608–12609. (32) (a) Pautot, S.; Frisken, B.; Weitz, D. Production of unilamellar vesicles using an inverted emulsion. Langmuir 2003, 19 (7), 2870–2879. (b) Pautot, S.; Frisken, B.; Weitz, D. Engineering asymmetric vesicles. Proc. Natl. Acad. Sci. U.S.A. 2003, 100 (19), 10718–10721. (c) Yamada, A.; Yamanaka, T.; Hamada, T.; Hase, M.; Yoshikawa, K.; Baigl, D. Spontaneous transfer of phospholipid-coated oil-in-oil and water-in-oil micro-droplets through an oil/water interface. Langmuir 2006, 22 (24), 9824–9828.

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(33) Maemichi, H.; Shikinaka, K.; Kakugo, A.; Furukawa, H.; Osada, Y.; Gong, J. Morphogenesis of liposomes caused by polycation-induced actin assembly formation. Langmuir 2008, 24 (20), 11975–11981. (34) (a) Miyata, H.; Hotani, H. Morphological changes in liposomes caused by polymerization of encapsulated actin and spontaneous formation of actin bundles. Proc. Natl. Acad. Sci. U.S.A. 1992, 89 (23), 11547–11551. (b) Honda, M.; Takiguchi, K.; Ishikawa, S.; Hotani, H. Morphogenesis of liposomes encapsulating actin depends on the type of actin-crosslinking. J. Mol. Biol. 1999, 287 (2), 293–300. (35) Walde, P.; Cosentino, K.; Engel, H.; Stano, P. Giant vesicles: preparations and applications. ChemBioChem 2010, 11 (7), 848–65. (36) (a) Abkarian, M.; Loiseau, E.; Massiera, G. Continuous droplet interface crossing encapsulation (cDICE) for high throughput monodisperse vesicle design. Soft Matter 2011, 7, 4610. (b) Matosevic, S.; Paegel, B. Stepwise synthesis of giant unilamellar vesicles on a microfluidic assembly line. J. Am. Chem. Soc. 2011, 133, 2798–2800. (37) Horger, K. S.; Estes, D. J.; Capone, R.; Mayer, M. Films of agarose enable rapid formation of giant liposomes in solutions of physiologic ionic strength. J. Am. Chem. Soc. 2009, 131 (5), 1810–1819. (38) (a) Verkhovsky, A. B.; Borisy, G. G. Non-sarcomeric mode of myosin II organization in the fibroblast lamellum. J. Cell Biol. 1993, 123 (3), 637–652. (b) Svitkina, T. M.; Verkhovsky, A. B.; McQuade, K. M.; Borisy, G. G. Analysis of the actin-myosin II system in fish epidermal keratocytes: mechanism of cell body translocation. J. Cell Biol. 1997, 139 (2), 397–415. (39) (a) Bendix, P. M.; Koenderink, G. H.; Cuvelier, D.; Dogic, Z.; Koeleman, B. N.; Brieher, W. M.; Field, C. M.; Mahadevan, L.; Weitz, D. A. A quantitative analysis of contractility in active cytoskeletal protein networks. Biophys. J. 2008, 94 (8), 3126–3136. (b) Silva, M.; Depken, M.; Stuhrmann, B.; Korsten, M.; Mackintosh, F.; Koenderink, G. Active multistage coarsening of actin networks driven by myosin motors. Proc. Natl. Acad. Sci. U.S.A. 2011, 108 (23), 9408–9413. (40) (a) Takiguchi, K.; Negishi, M.; Tanaka-Takiguchi, Y.; Homma, M.; Yoshikawa, K. Reconstruction of motile actin networks in giant liposome. Proceedings of the 2010 International Symposium on MicroNanoMechatronics and Human Science (MHS), Nagoya, Japan, Nov 710, 2010; Institute of Electrical and Electronics Engineers: Piscataway, NJ, 2010; pp 150155. (b) Takiguchi, K.; Yamada, A.; Negishi, M.; Tanaka-Takiguchi, Y.; Yoshikawa, K. Entrapping desired amounts of actin filaments and molecular motor proteins in giant liposomes. Langmuir 2008, 24 (20), 11323–11326.(c) Takiguchi, K.; Negishi, M.; Tanaka-Takiguchi, Y.; Homma, M.; Yoshikawa, K. Real-world modeling of artificial motile cell. Proceedings of the 2009 International Symposium on Micro-NanoMechatronics and Human Science (MHS), Nagoya, Japan, Nov 911, 2009; Institute of Electrical and Electronics Engineers: Piscataway, NJ, 2009; pp 197201. (41) Tanaka-Takiguchi, Y.; Kakei, T.; Tanimura, A.; Takagi, A.; Honda, M.; Hotani, H.; Takiguchi, K. The elongation and contraction of actin bundles are induced by double-headed myosins in a motor concentration-dependent manner. J. Mol. Biol. 2004, 341 (2), 467–476. (42) Murrell, M.; Pontani, L.; Guevorkian, K.; Cuvelier, D.; Nassoy, P.; Sykes, C. Spreading dynamics of biomimetic actin cortices. Biophys. J. 2011, 100 (6), 1400–1409. (43) Noireaux, V.; Maeda, Y.; Libchaber, A. Development of an artificial cell, from self-organization to computation and self-reproduction. Proc. Natl. Acad. Sci. U.S.A. 2011, 108 (9), 3473–3480. (44) Pardee, J. D.; Spudich, J. A. Purification of muscle actin. Methods Enzymol. 1982, 85 (Pt B), 164–181. (45) Margossian, S. S.; Lowey, S. Preparation of myosin and its subfragments from rabbit skeletal muscle. Methods Enzymol. 1982, 85 (Pt B), 55–71. (46) DeBiasio, R. L.; Wang, L. L.; Fisher, G. W.; Taylor, D. L. The dynamic distribution of fluorescent analogues of actin and myosin in protrusions at the leading edge of migrating Swiss 3T3 fibroblasts. J. Cell Biol. 1988, 107 (6 Pt 2), 2631–2645. (47) Koenderink, G.; Dogic, Z.; Nakamura, F.; Bendix, P.; MacKintosh, F.; Hartwig, J.; Stossel, T.; Weitz, D. An active biopolymer network 10070

dx.doi.org/10.1021/la201604z |Langmuir 2011, 27, 10061–10071

Langmuir controlled by molecular motors. Proc. Natl. Acad. Sci. U.S.A. 2009, 106 (36), 15192–15197. (48) Pecreaux, J.; Dobereiner, H. G.; Prost, J.; Joanny, J. F.; Bassereau, P. Refined contour analysis of giant unilamellar vesicles. Eur Phys J E 2004, 13 (3), 277–290. (49) Gonzalez, R. C.; Woods, R. E.; Eddins, S. L. Digital Image Processing Using MATLAB; Pearson: London, 2008. (50) Duda, R.; Hart, P. Use of the Hough transformation to detect lines and curves in pictures. Commun. ACM 1972, 15, 11–15. (51) Ballard, D. H. Generalizing the Hough transform to detect arbitrary shapes. Pattern Recognit. 1981, 13 (2), 111–122. (52) Helfrich, W. Elastic properties of lipid bilayers—theory and possible experiments. Z. Naturforsch., C 1973, 28, 693. (53) Hackl, W.; Barmann, M.; Sackmann, E. Shape changes of selfassembled actin bilayer composite membranes. Phys. Rev. Lett. 1998, 80 (8), 1786. (54) Saito, H.; Kato, Y.; Le Berre, M.; Yamada, A.; Inoue, T.; Yosikawa, K.; Baigl, D. Time-resolved tracking of a minimum gene expression system reconstituted in giant liposomes. ChemBioChem 2009, 10 (10), 1640–1643. (55) Angelova, M.; Dimitrov, D. A mechanism of liposome electroformation. Prog. Colloid Polym. Sci. 1988, 76, 59–67. (56) Holland, J. W.; Cullis, P. R.; Madden, T. D. Poly(ethylene glycol)-lipid conjugates promote bilayer formation in mixtures of nonbilayer-forming lipids. Biochemistry 1996, 35 (8), 2610–2617. (57) Estes, D. J.; Mayer, M. Electroformation of giant liposomes from spin-coated films of lipids. Colloids Surf., B 2005, 42 (2), 115–123. (58) Faucon, J. F.; Mitov, M. D.; Meleard, P.; Bivas, I.; Bothorel, P. Bending elasticity and thermal fluctuations of lipid membranes. Theoretical and experimental requirements. J. Phys. (Paris) 1989, 50, 2389–2414. (59) Rawicz, W.; Olbrich, K. C.; McIntosh, T.; Needham, D.; Evans, E. Effect of chain length and unsaturation on elasticity of lipid bilayers. Biophys. J. 2000, 79 (1), 328–339. (60) Marsh, D. Elastic curvature constants of lipid monolayers and bilayers. Chem. Phys. Lipids 2006, 144 (2), 146–159. (61) Kaufmann, S.; Papastavrou, G.; Kumar, K.; Textor, M.; Reimhult, E. A detailed investigation of the formation kinetics and layer structure of poly(ethylene glycol) tether supported lipid bilayers. Soft Matter 2009, 5, 2804–2814. (62) Evans, E.; Rawicz, W. Elasticity of “fuzzy” biomembranes. Phys. Rev. Lett. 1997, 79, 2379–2382. (63) (a) Levine, A. J.; MacKintosh, F. C. Dynamics of viscoelastic membranes. Phys. Rev. E 2002, 66 (6), 061606. (b) Fournier, J.; Lacoste, D.; Rapha€el, E. Fluctuation spectrum of fluid membranes coupled to an elastic meshwork: Jump of the effective surface tension at the mesh size. Phys. Rev. Lett. 2004, 92 (1), 018102. (c) Gov, N.; Zilman, A.; Safran, S. Cytoskeleton confinement and tensionof red blood cell membranes. Phys. Rev. Lett. 2003, 90 (22), 228101. (64) Shlomovitz, R.; Gov, N. Membrane waves driven by actin and myosin. Phys. Rev. Lett. 2007, 98 (16), 168103. (65) Colletier, J. P.; Chaize, B.; Winterhalter, M.; Fournier, D. Protein encapsulation in liposomes: Efficiency depends on interactions between protein and phospholipid bilayer. BMC Biotechnol. 2002, 2, 9. (66) Dominak, L. M.; Keating, C. D. Polymer encapsulation within giant lipid vesicles. Langmuir 2007, 23 (13), 7148–7154. (67) Luisi, P. L.; Allegretti, M.; Pereira de Souza, T.; Steiniger, F.; Fahr, A.; Stano, P. Spontaneous protein crowding in liposomes: A new vista for the origin of cellular metabolism. ChemBioChem 2010, 11 (14), 1989–1992. (68) (a) Dominak, L. M.; Keating, C. D. Macromolecular crowding improves polymer encapsulation within giant lipid vesicles. Langmuir 2008, 24 (23), 13565–13571. (b) Dominak, L.; Omiatek, D.; Gundermann, E.; Heien, M.; Keating, C. Polymeric crowding agents improve passive biomacromolecule encapsulation in lipid vesicles. Langmuir 2010, 26 (16), 13195–13200. (69) Fisher, C.; Kuo, S. Filament rigidity causes F-actin depletion from nonbinding surfaces. Proc. Natl. Acad. Sci. U.S.A. 2009, 106 (1), 133–138.

ARTICLE

(70) Limozin, L.; Roth, A.; Sackmann, E. Microviscoelastic moduli of biomimetic cell envelopes. Phys. Rev. Lett. 2005, 95, 178101. (71) Finer, J.; Simmons, R.; Spudich, J. Single myosin molecule mechanics—Piconewton forces and nanometer steps. Nature 1994, 368 (6467), 113–119. (72) Verkhovsky, A. B.; Svitkina, T. M.; Borisy, G. G. Polarity sorting of actin filaments in cytochalasin-treated fibroblasts. J. Cell Sci. 1997, 110 (Pt 15), 1693–1704. (73) Herbert, T.; Carlson, F. Spectroscopic study of the self association of myosin. Biopolymers 1971, 10, 2231–2252. (74) Kanzaki, N.; Uyeda, T. Q.; Onuma, K. Intermolecular interaction of actin revealed by a dynamic light scattering technique. J. Phys. Chem. B 2006, 110 (6), 2881–2887. (75) Pollard, T.; Blanchoin, L.; Mullins, R. Molecular mechanisms controlling actin filament dynamics in nonmuscle cells. Annu. Rev. Biophys. Biomol. Struct. 2000, 29, 545–576. (76) Koestler, S.; Rottner, K.; Lai, F.; Block, J.; Vinzenz, M.; Small, J. F- and G-actin concentrations in lamellipodia of moving cells. PLoS ONE 2009, 4, e4810. (77) (a) Jansen, L.; Kolega, J.; Taylor, D. Modulation of contraction by gelation/solation in a reconstituted motile model. J. Cell Biol. 1991, 114, 1005–1015. (b) Pollard, T. Cytoplasmic contractile proteins. J. Cell Biol. 1981, 91, 156s–165s. (78) Merkle, D.; Kahya, N.; Schwille, P. Reconstitution and anchoring of cytoskeleton inside giant unilamellar vesicles. ChemBioChem 2008, 9 (16), 2673–2681.

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