We would like to acknowledge Dolf Weijers, Vera Gorelova, and Mark Roosjen for kindly providing us with YFP. S.D. acknowledges financial support from the Dutch Research Council (grant number: OCENW.KLEIN.465).

1. Fabricating the master wafer
<ol>
	<li>Take a 4 in (10 cm) diameter clean silicon wafer (see Table of Materials). Clean it further using pressurized air to remove any dust particles.</li>
	<li>Mount the wafer on a spin coater, and gently dispense ~5 mL of a negative photoresist (see Table of Materials) in the center of the wafer. Try to avoid air bubbles, as they might interfere with the downstream printing process of the wafer.</li>
	<li>To obtain a 10 &#181;m thick photoresist layer, spin-coat the wafer at 500 rpm for 30 s with an acceleration of 100 rpm/s for initial spreading, followed by a 60 s spin at 3,000 rpm with an acceleration of 500 rpm/s. In case a different thickness is desired, change the spinning parameters according to the manufacturer&#39;s instructions.</li>
	<li>Bake the wafer on a heating plate for 2 min at 65 &#176;C and then for 5 min at 95 &#176;C.</li>
	<li>Once the wafer cools down, mount the wafer in the printing chamber of the direct-write optical lithography machine (see Table of Materials), and feed the OLA design (Figure 2A, Supplementary Coding File 1) into the software. 
	NOTE: The OLA design essentially consists of two OA&#160;channels, two LO&#160;channels, and one IA&#160;channel, which merge to form a six-way junction that continues as a post-production channel and ends up in the experimental well (EW).</li>
	<li>Print the OLA design(s) on the coated wafer using a UV laser (see Table of Materials) with a dose of 300 mJ/cm2.</li>
	<li>Once the design is printed, bake the wafer at 65 &#176;C for 1 min, followed by 95 &#176;C for 3 min.</li>
	<li>At this point, ensure that the outline of the printed device appears on the wafer and is visible to the naked eye. To wash off the uncured photoresist, dip the wafer in a glass beaker containing the developer solution (see Table of Materials) until the uncured photoresist is fully removed. 
	NOTE: Excess developer treatment can affect the design resolution.</li>
	<li>Wash the wafer with acetone, isopropanol, and deionized (DI) water in sequence, and finally, with pressurized air/N2 using a blow gun.</li>
	<li>Hard-bake the wafer at 150 &#176;C for 30 min to ensure that the printed design is firmly attached to the wafer surface and does not come off in the downstream fabrication process. The master wafer is then ready for further use (Figure 2B).</li>
</ol>
2. Preparing the microfluidic device
<ol>
	<li>Place the master wafer on a square piece of aluminum, and wrap the aluminum foil around the wafer, forming a well-like structure (Figure 2C).</li>
	<li>Measure 40 g of polydimethylsiloxane (PDMS) and 4 g of curing agent (10:1, weight ratio, see Table of Materials) in a 50 mL centrifuge tube, and mix vigorously using a spatula or a pipette tip for 2-3 min.</li>
	<li>The vigorous mixing of the PDMS and curing agent traps air bubbles inside the mixture. Spin the tube at 100 x g for 2-3 min to remove majority of large air bubbles.</li>
	<li>Pour approximately 15-20 g of the mixture prepared in step 2.3 over the master wafer, and de-gas using a desiccator. Save the excess mixture to make PDMS-coated glass slides (step 3).</li>
	<li>Incubate the assembly in an oven at 70 &#176;C for at least 2 h.</li>
	<li>Take the master wafer out of the oven and let it cool down. To remove the solidified PDMS block, remove the aluminum foil, and carefully peel off the PDMS block from the edge of the wafer. 
	NOTE: The wafer is fragile and might break, so it is important to do this process carefully.</li>
	<li>Once the PDMS block is removed, keep the patterned structure facing upward, and using a sharp blade or a scalpel, cut individual PDMS blocks, each containing a single microfluidic device.</li>
	<li>Place the PDMS block on a dark surface, and adjust the light direction (a table lamp comes in handy for this) so that the engraved channels are shiny and, as a result, visible. Make holes in the inlets and the exit channel using biopsy punches of 0.5 mm and 3 mm diameter (see Table of Materials), respectively. 
	NOTE: Use a sharp biopsy punch to avoid cracks in the PDMS, which can cause leakage later. Gently push the biopsy punch through the PDMS block, and ensure it passes completely through it. To remove the biopsy punch, gently retract it while rotating it in alternate directions. The PDMS block with the engraved OLA design is now ready to bind to a PDMS-coated glass slide.</li>
</ol>
3. Making the PDMS-coated glass slide
<ol>
	<li>Take a transparent glass coverslip, pour approximately 0.5 mL of PDMS (prepared in excess in step 2.4) onto the center of the glass slide, and spread it across the coverslip by gently tilting the glass slide, ensuring total coverage of the glass slide with PDMS.</li>
	<li>Mount the glass slide on the spin coater, ensure its centrally placed (so that the middle of the slide overlaps with the center of the pressure shaft), and spin the glass slide at 500 rpm for 15 s (at an increment of 100 rpm/s) and then at 1,000 rpm for 30 s (at an increment of 500 rpm/s).</li>
	<li>Place the PDMS-coated glass slide (coated side facing upward) on a raised platform like a block of PDMS (1 cm x 1 cm) in a covered Petri dish, and bake it at 70 &#176;C for 2 h.</li>
</ol>
4. Bonding of the microfluidic device
<ol>
	<li>Clean the PDMS block (prepared in step 2) by sticking and removing commercially available scotch tape (see Table of Materials) twice. Ensure to keep the cleaned side facing up until use.</li>
	<li>Clean the PDMS-coated glass slide with pressurized air/N2&#160;using a blow gun, and keep the clean side facing up.</li>
	<li>Place the PDMS block (engraved channels facing up) and the PDMS-coated glass slide (coated side facing up) in the vacuum chamber of the plasma cleaner (see Table of Materials).</li>
	<li>Switch on the vacuum, and expose the contents to air plasma at a radio frequency of 12 MHz (RF mode high) for 15 s to activate the surfaces. Oxygen plasma can be seen in the form of a pinkish hue.</li>
	<li>Immediately after the plasma treatment, place the PDMS-coated glass slide on a clean surface with the PDMS side facing upward. Gently place the PDMS block with the microfluidic pattern now facing toward the PDMS-coated glass slide, allowing them to bond (Figure 2D). 
	NOTE: Removing the air trapped in the device is crucial to ensure thorough bonding.</li>
	<li>Bake the bonded devices at 70 &#176;C for 2 h.</li>
</ol>
5. Surface functionalization of the microfluidic device
NOTE: Prior to surface functionalization, it is important to calibrate the pressure pump as per the manufacturer&#39;s protocol (see Table of Materials) and assemble the tubing to connect it to the microfluidic device.
<ol>
	<li>Connecting the microfluidic device to the pressure pumps
	<ol>
		<li>Connect the pressure-driven flow controller to an external pressure source (up to 6 bars). At least four independent air pressure channels are required: three individual modules of 0-2 bar, and one module of &#8722;0.9-4 bar.</li>
		<li>Calibrate the pressure pump according to the manufacturer&#39;s protocol (see Table of Materials).</li>
		<li>Cut the tubing (1/16 in OD x 0.02 in ID) into four equal-sized pieces approximately 20 cm in length.</li>
		<li>Insert 23 G stainless steel 90&#176; bent connectors (see Table of Materials) at one end of each piece. This end is later inserted into the three inlets (OA, LO, and IA) of the microfluidic device and the fourth one is used to remove excess solution (step 5.2.5).</li>
		<li>Insert the other end of the tubing into the microfluidic reservoir stand, and seal it using microfluidic fittings, ensuring the tubing is long enough to touch the bottom of the reservoir (1.5 mL tubes, see Table of Materials). This prevents unwanted air bubbles from entering the microfluidic device during the PVA treatment/liposome production.</li>
	</ol>
	</li>
	<li>PVA treatment of the exit channel 
	NOTE: Prior to liposome generation, it is crucial to render the device partially hydrophilic downstream of the production junction. This is done by treating these channels with 5% (w/v) polyvinyl alcohol (PVA) solution. The PVA solution is prepared by dissolving the PVA powder (see Table of Materials) in water at 80 &#176;C for 3 h with constant stirring using a magnetic stirrer. Filter the solution prior to using it for surface treatment. Immediately after the bonding of the device, the microfluidic channels are hydrophilic due to the plasma-induced surface activation, and they will gradually become hydrophobic again. It is recommended to wait for 2 h after the baking (step 4.6) before starting the PVA treatment.
	<ol>
		<li>Dispense 200 &#181;L of 5% w/v PVA solution into a 1.5 mL tube, and connect it to the microfluidic reservoir stand. Insert the tubing (the one mentioned in step 5.1.3) such that one end is submerged in the PVA solution and the other end is connected to the inlet of the OA channel of the microfluidic device. 
		NOTE: A lower PVA concentration can lead to sub-standard surface functionalization.</li>
		<li>Repeat the above step without PVA (empty 1.5 mL tube), and connect it to the LO and IA channels.</li>
		<li>Increase the pressure of the OA phase to 100 mbar to flow the PVA solution in the OA channels. Increase the pressures of the IA and LO phases to 120 mbar in order to prevent the backflow of PVA solution inside these channels (Figure 2E). 
		NOTE: If needed, adjust the pressures of the individual channels in order to keep the meniscus steady at the production junction.</li>
		<li>Flow the PVA solution in this manner for approximately 5 min, ensuring complete functionalization of the exit channel.</li>
		<li>To remove the PVA solution, detach the tubing from the OA inlet, and immediately increase the pressure in the LO and IA channels to 2 bar. Simultaneously, use a tubing connected to a negative pressure channel (&#8722;900 mbar) remove excess PVA first from the exit channel and then from the OA inlet (Figure 2F).</li>
		<li>Bake the device at 120 &#176;C for 15 min, and let it cool down before use. The device can be stored under ambient conditions for at least 1 month. 
		NOTE: It is recommended to wait for 1 day (at room temperature) before proceeding with OLA to ensure the untreated PDMS regains its hydrophobicity after the plasma treatment.</li>
	</ol>
	</li>
</ol>
6.&#160;Octanol-assisted liposome assembly (OLA)
<ol>
	<li>Preparing the lipid stock 
	NOTE: Here, a mixture of 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (Liss Rhod PE) lipids (see Table of Materials) are used as an example; users should prepare the lipid composition they need in a similar manner.
	<ol>
		<li>Dispense 76 &#181;L of DOPC (25 mg/mL) and 16.6 &#181;L of Liss Rhod PE (1 mg/mL) into a round-bottom flask using separate glass syringes.</li>
		<li>Keep the round-bottom flask upright, and use a compressed N2&#160;blow gun to give a gentle stream of nitrogen to evaporate the chloroform and form a dried lipid film at the bottom of the flask.</li>
		<li>Place the flask in the desiccator for at least 2 h under a continuous vacuum to remove any remaining chloroform.</li>
		<li>Add 100 &#181;L of ethanol into the round-bottom flask, followed by gentle pipetting or shaking to ensure the lipids are dissolved to form a 10% (w/v) lipid stock. 
		NOTE: In case a particular lipid composition is not completely soluble in ethanol, use an ethanol/chloroform mixture, keeping the volume of chloroform as small as possible.</li>
		<li>Pipette the solution into a dark glass vial. Gently flush the vial with nitrogen using a blow gun to replace the air with an inert atmosphere. Seal the lid with paraffin film, and store at &#8722;20 &#176;C. 
		NOTE: The stock solution can be used for up to a few months. Each time the vial is opened, gently replace the air with nitrogen, and reseal with lid with paraffin film.</li>
	</ol>
	</li>
	<li>Preparing the OLA solutions
	<ol>
		<li>Make stock solutions of the indispensable components: 20 mM dextran (Mw 6,000); 60% (v/v) glycerol; a buffer of choice. In this case, 5x phosphate-buffered saline (0.68 M NaCl, 13.5 mM KCl, 50 mM Na2HPO4, 9 mM KH2PO4; pH 7.4) was prepared (see Table of Materials). 
		NOTE: As pure glycerol is highly viscous, it is recommended to cut out a small piece at the end of the pipette tip in a slanted manner for effective pipetting.</li>
		<li>Prepare 100 &#181;L of IA, OA, and exit solution (ES, the desired solution to fill the EW) samples. For ease of visualization, yellow fluorescent protein (YFP) was added to the IA solution in this particular case. Check the osmotic balance between the IA and OA by calculating the osmolarity of the encapsulated components and adding an appropriate amount of sugar or salt into the OA if needed. This is done in order to prevent the bursting or shrinking of the liposomes during production. 
		NOTE: The final compositions of the three phases are as follows: 
		IA: 15% glycerol, 5 mM dextran (Mw 6,000), 5.4 &#181;M YFP, 1x PBS 
		OA: 15% glycerol, 5% F68 surfactant, 1x PBS 
		&#8203;ES: 15% glycerol, 1x PBS 
		A lower concentration of F68 surfactant negatively affects the generation of double emulsions.</li>
		<li>Prepare 80 &#181;L of the LO phase by pipetting 4 &#181;L of stock lipid into 76 &#181;L of 1-octanol (see Table of Materials). The final concentration of DOPC is 5 mg/mL.</li>
		<li>Centrifuge the samples at 700 x g for 60 s to remove any large aggregates before proceeding with the microfluidic experiments. This prevents any obvious clogging factors from entering the microfluidic chip.</li>
	</ol>
	</li>
	<li>Liposome production
	<ol>
		<li>Dispense the solutions (IA, LO, OA) into three 1.5 mL tubes, assemble them (as mentioned in step 5.1), and connect them to the PVA-treated microfluidic chip (step 5.2.6).</li>
		<li>Apply a positive pressure on the three channels: ~100 mbar on the IA and LO channels and ~200 mbar on the OA channel. 
		NOTE: Since the OA pressure is higher than the others, the OA solution will be the first to arrive at EW; this is highly recommended.</li>
		<li>Once the OA solution reaches EW, decrease the OA pressure to 100 mbar and increase the LO to 200 mbar.&#160; The LO solution arriving at the production junction will likely result in air bubbles because of air being displaced from the LO channels. Once the air bubbles are dispensed into the EW, adjust the LO pressure to 100 mbar. Lastly, increase the IA pressure to 200 mbar, and wait until all the air in the IA channel is removed. The ideal sequence of arrival at the junction of the three phases is, thus, OA, LO, and IA.</li>
		<li>After removing the air from the three channels, adjust all the channel pressures to 50 mbar and pipette 10 &#181;L of ES into the exit chamber. After pipetting the ES, put a cover slide on the exit hole to avoid evaporation. In case the LO is pushed back, gradually increase the LO pressure in 1 mbar increments until it starts to flow to the junction.</li>
		<li>Once all three phases start co-flowing at the junction, ensure double emulsion production begins, and adjust according to its quality (Figure 3A-C). Change the pressures gradually (steps of 0.1-1 mbar) rather than abruptly unless the pressure is being changed to eliminate an unwanted clog in the channel. 
		NOTE: The channels to adjust depends on what is happening at the junction. For example, the IA needs to be decreased if the emulsions are too big; the OA needs to be increased if double emulsions form but burst instead of getting pinched off; and the LO needs to be decreased if the octanol pocket is too big.</li>
		<li>Check that the double-emulsion droplets flow downstream to EW. As they flow, octanol pockets become more and more prominent and finally get pinched off, forming liposomes (Figure 3D-F). Ensure that the post-production channel is long enough and that the migration of the double emulsions is slow enough for dewetting. 
		NOTE: Within minutes after decent production, liposomes and octanol droplets will exit the post-production channel and go into the EW. As a result, being less dense than water, the octanol droplets float to the surface of the ES. Due to the addition of dextran in the IA, the liposomes are heavier than their surroundings and go to the bottom of the chamber ready for observation and further manipulation.</li>
	</ol>
	</li>
</ol>

The authors declare no conflicts of interest.

Cellular complexity makes it extremely difficult to understand living cells when studied as a whole. Reducing the redundancy and interconnectivity of cells by reconstituting the key components in vitro is necessary to further our understanding of biological systems and create artificial cellular mimics for biotechnological applications22,23,24. Liposomes serve as an excellent minimal system to understand cellular phenomena. A non-exhaustive list of these phenomena includes cytoskeleton dynamics and resulting membrane deformations25,26, spatiotemporal regulation of biomolecular condensates27,28&#160;and their interaction with the membrane29, encapsulation of a wide variety of biomolecules, including cell-free transcription-translation systems30, cell-free lipid synthesis31, and evolution of proteins32,33,34. Liposomes have also been widely used as carriers for drug delivery and have been approved by the Food and Drug Administration (FDA) and European Medicines Agency (EMA) for clinical usage11. Liposomes and lipid-based nanoparticles are used as carriers for drug delivery, including mRNA vaccines like the recent COVID-19 vaccine35.
This study describes a detailed protocol on photolithography, microfluidic assembly, and surface functionalization to perform the OLA technique in order to generate on-chip liposomes (Figure 1 and Figure 3). The liposomes produced using OLA are monodispersed (coefficient of variation: 4%-11% of the mean) and in the biological cell-sized range (typically between 5-50 &#181;m in diameter), and they can be tuned using appropriate flow velocities of the inner and outer aqueous channels36. For example, the liposome population seen in Figure 1&#160;has a size distribution of 23.3 &#181;m &#177; 1.8 &#181;m (n = 50). With typical production rates of 10-30 Hz, the formed liposomes are unilamellar, as was previously confirmed by inserting a single bilayer-specific transmembrane protein, alpha-hemolysin36, into the membrane, as well as by using the dithionate-bleaching assay37. OLA is also compatible with a wide variety of lipid compositions, offering the user flexibility in the choice of lipids. Importantly, the initially used lipid composition is reflected in the final liposome composition38.
Being a relatively new technology, there is further scope for improving OLA. For instance, surface functionalization using PVA is crucial but tedious to perform. An easier and simpler way to surface-functionalize the device would significantly reduce the chip fabrication time as well as chip-to-chip variability. Pluronic F68 surfactant is an important component in the outer aqueous phase to initially stabilize the double emulsions; nonetheless, its usage can be restrictive in certain cases. Replacing Pluronic F68 with a more biocompatible surfactant or complete surfactant removal may improve the versatility of the system. During the migration of generated double emulsions in the exit channel, they can burst due to shear. Upgrading the OLA design to improve the double emulsion stability and octanol pocket separation could, thus, increase the throughput. Nonetheless, OLA has several advantages, which mainly include the efficient encapsulation, monodispersed and unilamellar liposomes, and controlled on-chip experimentation.
OLA has been employed and adapted in a diverse range of studies, including growth and division of liposomes39, studying the process of liquid-liquid phase separation27&#160;and its interaction with the membrane21, and understanding bacterial growth bioproduct formation in liposomes40. OLA-based high-throughput assays are also being used to understand ion transport across the membrane41&#160;and drug permeability across the lipid bilayer37, as a drug delivery system for therapeutic purposes42, to study the effect of antimicrobials on membrane43, and as a tool to encapsulate liquid crystals44. In addition to the wide range of applications of OLA technology, modified versions of OLA suited for particular purposes are being developed45,46,47,48,49,50. Overall, considering the pros and cons, we strongly believe that OLA is a versatile platform for synthetic biology.

All cells have a plasma membrane as their physical boundary, and this membrane is essentially a scaffold in the form of a lipid bilayer formed by the self-assembly of amphiphilic lipid molecules. Liposomes are the minimal synthetic counterparts of biological cells; they have an aqueous lumen surrounded by phospholipids, which form a lipid bilayer with the hydrophilic head groups facing the aqueous phase and the hydrophobic tails buried inward. The stability of liposomes is governed by the hydrophobic effect, as well as the hydrophilicity between the polar groups, van der Waals forces between the hydrophobic carbon tails, and the hydrogen bonding between water molecules and the hydrophilic heads1,2. Depending on the number of lipid bilayers, liposomes can be classified into two main categories, namely, unilamellar vesicles comprising a single bilayer and multilamellar vesicles constructed from multiple bilayers. Unilamellar vesicles are further classified based on their sizes. Typically spherical in shape, they can be produced in a variety of sizes, including small unilamellar vesicles (SUV, 30-100 nm diameter), large unilamellar vesicles (LUV, 100-1,000 nm diameter), and finally, giant unilamellar vesicles (GUV, &#62;1,000 nm diameter)3,4. Various techniques have been developed to produce liposomes, and these can be categorized broadly into bulk techniques5&#160;and microfluidic techniques6. Commonly practiced bulk techniques include lipid film rehydration, electroformation, inverted emulsion transfer, and extrusion7,8,9,10. These techniques are relatively simple and effective, and these are the prime reasons for their widespread usage in the synthetic biology community. However, at the same time, they suffer from major drawbacks with regard to the polydispersity in size, the lack of control over the lamellarity, and low encapsulation efficiency7,11. Techniques like continuous droplet interface crossing encapsulation (cDICE)12&#160;and droplet shooting and size filtration (DSSF)13&#160;overcome these limitations to some extent.
Microfluidic approaches have been rising to prominence over the last decade. Microfluidic technology provides a controllable environment for manipulating fluid flows within user-defined microchannels owing to the characteristic laminar flow and diffusion-dominated mass transfer. The resulting lab-on-a-chip devices offer unique possibilities for the spatiotemporal control of molecules, with significantly reduced sample volumes and multiplexing capabilities14. Numerous microfluidic methods to make liposomes have been developed, including pulsed jetting15, double emulsion templating16, transient membrane ejection17, droplet emulsion transfer18, and hydrodynamic focusing19. These techniques produce monodispersed, unilamellar, cell-sized liposomes with high encapsulation efficiency and high throughput.
This article details the procedure for octanol-assisted liposome assembly (OLA), an on-chip microfluidic method based on the hydrodynamic pinch-off and subsequent solvent dewetting mechanism20&#160;(Figure 1). One can relate the working of OLA to a bubble-blowing process. A six-way junction focuses the inner aqueous (IA) phase, two lipid-carrying organic (LO) streams, and two surfactant-containing outer aqueous (OA) streams at a single spot. This results in water-in-(lipids + octanol)-in-water double emulsion droplets. As these droplets flow downstream, interfacial energy minimization, external shear flow, and interaction with the channel walls lead to the formation of a lipid bilayer at the interface as the solvent pocket becomes detached, thus forming unilamellar liposomes. Depending on the size of the octanol pocket, the dewetting process can take tens of seconds to a couple of minutes. At the end of the exit channel, the less dense octanol droplets float to the surface, whereas the heavier liposomes (due to a denser encapsulated solution) sink to the bottom of the visualization chamber ready for experimentation. As a representative experiment, the process of liquid-liquid phase separation (LLPS) inside liposomes is demonstrated. For that, the required components are encapsulated inside liposomes at an acidic pH that prevents LLPS. By externally triggering a pH change and, thus, a transmembrane proton flux, phase-separated condensate droplets are formed inside the liposomes. This highlights the effective encapsulation and manipulation capabilities of the OLA system.

<table><tbody><tr><td>1-Octanol</td><td>Sigma-Aldrich</td><td>No. 297887</td><td></td></tr><tr><td>1.5 mL tubes</td><td>Fisher scientific</td><td>10451043</td><td>Eppendorf 3810X Polypropylene microcentrifuge tubes</td></tr><tr><td>ATP</td><td>Sigma-Aldrich</td><td>No. A2383</td><td></td></tr><tr><td>Biopsy punch</td><td>Darwin microfluidics</td><td>PT-T983-05</td><td>0.5 mm and 3 mm diameter</td></tr><tr><td>Citrate-base</td><td>Sigma-Aldrich</td><td>No. 71405</td><td></td></tr><tr><td>Dextran</td><td>Sigma-Aldrich</td><td>No. 31388</td><td>Mr~6,000</td></tr><tr><td>Direct-write optical lithography machine</td><td>Durham Magneto Optics Ltd</td><td>MicroWriter ML3 Baby</td><td>setup and software</td></tr><tr><td>DOPC lipid</td><td>Avanti</td><td>SKU:850375C</td><td></td></tr><tr><td>F68</td><td>Sigma-Aldrich</td><td>No. 24040032</td><td></td></tr><tr><td>Glass cover slip</td><td>Corning</td><td>#1, 24 x 40 mm</td><td></td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>No. G2025</td><td></td></tr><tr><td>Hydrochloric acid</td><td>Thermo Scientific Acros</td><td>No. 124630010</td><td></td></tr><tr><td>Liss Rhod PE lipid</td><td>Avanti</td><td>SKU:810150C</td><td></td></tr><tr><td>Parafilm</td><td>Sigma-Aldrich</td><td>No. P7793</td><td></td></tr><tr><td>Photoresist</td><td>Micro resist technology GmbH</td><td>EpoCore 10</td><td></td></tr><tr><td>Photoresist developer</td><td>micro resist technology GmbH</td><td>mr-Dev 600</td><td></td></tr><tr><td>Plasma cleaner</td><td>Harrick plasma</td><td>PDC-32G</td><td></td></tr><tr><td>Polydimethylsiloxane</td><td>Dow</td><td>Sylgard 184</td><td>PDMS and curing agent</td></tr><tr><td>Poly-L-lysine</td><td>Sigma-Aldrich</td><td>No. P7890</td><td></td></tr><tr><td>Poly-L-lysine&amp;ndash;FITC Labeled</td><td>Sigma-Aldrich</td><td>No. P3543</td><td></td></tr><tr><td>Polyvinyl alcohol</td><td>Sigma-Aldrich</td><td>no. P8136</td><td>molecular weight 30,000&amp;ndash;70,000, 87%&amp;ndash;90% hydrolyzed</td></tr><tr><td>Pressure controller</td><td>Elveflow&amp;nbsp;</td><td>OBK1 Mk3+</td><td>Flow controller</td></tr><tr><td>Scotch tape</td><td>Magic Tape Invisible Matt Tape</td><td></td><td></td></tr><tr><td>Silicon wafer</td><td>Silicon Materials</td><td>0620R16002</td><td></td></tr><tr><td>Spin coater&amp;nbsp;</td><td>Laurell Technologies Corporation</td><td>Model WS-650MZ-23NPP</td><td></td></tr><tr><td>Stainless Steel 90&amp;deg; Bent PDMS Couplers</td><td>Darwin microfluidics</td><td>PN-BEN-23G</td><td></td></tr><tr><td>Tris-base</td><td>Sigma-Aldrich</td><td>No. 252859</td><td></td></tr><tr><td>Tygon tubing</td><td>Darwin microfluidics</td><td>1/16&quot; OD x 0.02&quot; ID</td><td></td></tr><tr><td>UV laser&amp;nbsp;</td><td></td><td></td><td>365 nm wavelength</td></tr></tbody></table>

on-chip octanol-assisted liposome assembly for bioengineering

Microfluidics is a widely used tool to generate droplets and vesicles of various kinds in a controlled and high-throughput manner. Liposomes are simplistic cellular mimics composed of an aqueous interior surrounded by a lipid bilayer; they are valuable in designing synthetic cells and understanding the fundamentals of biological cells in an in vitro fashion and are important for applied sciences, such as cargo delivery for therapeutic applications. This article describes a detailed working protocol for an on-chip microfluidic technique, octanol-assisted liposome assembly (OLA), to produce monodispersed, micron-sized, biocompatible liposomes. OLA functions similarly to bubble blowing, where an inner aqueous (IA) phase and a surrounding lipid-carrying 1-octanol phase are pinched off by surfactant-containing outer fluid streams. This readily generates double-emulsion droplets with protruding octanol pockets. As the lipid bilayer assembles at the droplet interface, the pocket spontaneously detaches to give rise to a unilamellar liposome that is ready for further manipulation and experimentation. OLA provides several advantages, such as steady liposome generation (&#62;10 Hz), efficient encapsulation of biomaterials, and monodispersed liposome populations, and requires very small sample volumes (~50 &#181;L), which can be crucial when working with precious biologicals. The study includes details on microfabrication, soft-lithography, and surface passivation, which are needed to establish OLA technology in the lab. A proof-of-principle synthetic biology application is also shown by inducing the formation of biomolecular condensates inside the liposomes via transmembrane proton flux. It is anticipated that this accompanying video protocol will facilitate the readers to establish and troubleshoot OLA in their labs.

This study demonstrates the formation of membraneless condensates via the process of liquid-liquid phase separation (LLPS) inside liposomes as a representative experiment.
Sample preparation 
The IA, OA, ES, and feed solution (FS) are prepared as follows:
IA: 12% glycerol, 5 mM dextran, 150 mM KCl, 5 mg/mL poly-L-lysine (PLL), 0.05 mg/mL poly-L-lysine-FITC labeled (PLL-FITC), 8 mM adenosine triphosphate (ATP), 15 mM citrate-HCl (pH 4)
OA: 12% v/v glycerol, 5% w/v F68, 150 mM KCl, 15 mM citrate-HCl (pH 4)
ES: 12% glycerol, 150 mM KCl, 15 mM citrate-HCl (pH 4)
FS: 12% glycerol, 150 mM KCl, 75 mM Tris-HCl (pH 9)
Condensate formation inside liposomes 
A simple assay of pH-sensitive complex coacervation of positively charged poly-L-lysine (PLL) and negatively charged multivalent adenosine triphosphate (ATP) was selected to demonstrate the phenomena of LLPS in liposomes. To prevent phase separation of polylysine and ATP during encapsulation, the pH of the solution was maintained at 4, at which ATP is neutral. Increasing the pH of the ES by adding FS (a buffer of pH 9) eventually increased the pH inside the liposomes due to transmembrane proton flux, making ATP negatively charged and triggering its phase separation with positively charged PLL21&#160;(Figure 4A). After about 2 h of liposome generation, the IA and LO pressures were switched off. The OA channel pressure was kept at 100 mbar to flow the remaining liposomes into the observation chamber slowly. Once all the liposomes in the channel were recovered in the EW, the pressures were switched off to stop the flow and prevent the liposomes from moving. The liposomes generated at a lower pH showed a homogeneous fluorescence (from the encapsulated fluorescent PLL-FITC) in their&#160;lumen (Figure 4B,D). Octanol droplets floating at the surface were removed by carefully pipetting out 5 &#181;L of solution from the top to prevent them from affecting further pipetting steps. Subsequently, 10 &#181;L of FS buffer was added to the EW, which induced phase separation of the encapsulated PLL and ATP. The homogeneous FITC fluorescence from each liposome gradually transformed into distinct fluorescent condensate droplets. Eventually, the individual droplets merged into one bigger condensate droplet that freely diffused within the liposomes (Figure 4C,E-G).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65032/65032fig01.jpg" /> 
Figure 1: Schematic showing the assembly and working of OLA. The pressure controller is connected to the reservoirs containing outer aqueous, lipid-in-octanol, and inner aqueous solutions. The tubes inserted into the reservoirs are connected to the respective inlets of the OLA device. Appropriate flows in the three channels lead to the formation of water-in-(lipid-in-octanol)-in-water double emulsions. The formed double emulsions migrate to the exit well, during which the octanol pockets detach to form liposomes. The formed liposomes are collected at the bottom of the well for visualization and further experimentation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65032/65032fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65032/65032fig02.jpg" /> 
Figure 2: Preparation of the OLA chip (photolithography, microfabrication, surface treatment). (A) Digital design showing the key features of the OLA design, including three inlets, an outlet, and a six-way production junction. (B) A schematic of the master wafer showing multiple OLA designs produced using UV lithography. (C) A PDMS elastomer cast on the master wafer, placed in a well created out of aluminum foil, and cured by baking at 70 &#176;C for 2 h. (D) A microfluidic device bonded using oxygen plasma treatment, where the PDMS block containing the OLA design is attached to a PDMS-coated glass slide. (E) Surface functionalization of the fabricated chip to make the device partially hydrophilic. This is done by flowing 5% w/v PVA for 5 min from the outer aqueous channel toward the exit channel. Positive air pressure in the other channels prevents the PVA solution from entering these channels. (F) PVA is removed by applying a vacuum in the exit channel. The device is baked at 120 &#176;C for 15 min and is then ready to use. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65032/65032fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65032/65032fig03.jpg" /> 
Figure 3: Demonstration of OLA efficiently producing monodispersed double emulsions and eventually liposomes with excellent encapsulation. (A) A bright-field image showing rapid generation of double-emulsion droplets. (B) The fluorescent lipid channel shows the formation of an octanol pocket due to partial dewetting. The lipid-in-octanol phase contained a mixture of DOPC (5 mg/mL) and Lis Rhod PE at a 1,000:1 ratio. (C) The inner aqueous channel showing encapsulation of yellow fluorescent protein (YFP). The insets in (A-C) show representative zoomed-in views of the respective panels (scale bars = 10 &#181;m). (D-F) Dewetting of the octanol pocket in the exit channel, which forms a liposome. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65032/65032fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65032/65032fig04.jpg" /> 
Figure 4: pH-triggered liquid-liquid phase separation of pLL/ATP within liposomes. (A) Schematic of the pH-dependent transitioning of the homogenous solution of pLL and ATP encapsulated within the liposome (left) to phase-separated pLL/ATP coacervates (right). The initial acidic environment in the liposome renders the molecular charge of ATP to be neutral, inhibiting coacervation. When the pH inside the liposomes equilibrates with the externally applied pH increase, ATP gains a negative charge, triggering coacervation. (B-C) Line graphs (corresponding to the dotted lines in panels [D] and [G], respectively) showing the spatial distribution of pLL (green channel) and the membrane (red channel). (D-G) Time-lapse images showing the formation of pLL/ATP coacervates within the liposomes. The external addition of a basic buffer raises the pH level inside the liposomes over the course of minutes and initiates coacervation. t = 0 min refers to the time just before the occurrence of the first coacervation event. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65032/65032fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Coding File 1: CAD file of the OLA design. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65032/OLA device design.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about On-Chip Octanol-Assisted Liposome Assembly for Bioengineering at JoVE.com

On-Chip Octanol-Assisted Liposome Assembly for Bioengineering

The present protocol describes octanol-assisted liposome assembly (OLA), a microfluidic technique to generate biocompatible liposomes. OLA produces monodispersed, micron-sized liposomes with efficient encapsulation, allowing immediate on-chip experimentation. This protocol is anticipated to be particularly suitable for synthetic biology and synthetic cell research.

Microfluidics is a widely used tool to generate droplets and vesicles of various kinds in a controlled and high-throughput manner. Liposomes are ...

on-chip-octanol-assisted-liposome-assembly-for-bioengineering

Nanyang Technological University

Research

JoVE Journal

Bioengineering

2.5K Views.  Wageningen University & Research. OLA is een veelzijdig microfluïdisch platform dat nuttig is voor de synthetische biologiegemeenschap. in het bijzonder om kunstmatige cellen te bio-engineeren. OLA-gebaseerde liposomen kunnen ons helpen de zelforganisatieprincipes in de biologie te begrijpen en cel-nabootsende assemblages te construeren.OLA produceert monodisperse, unilaminaire, celgrote liposomen op een high-throughput manier en met een uitstekende inkapselingsefficiëntie. Het vereist zeer kleine monstervolumes, in ...

Video: On-Chip Octanol-Assisted Liposome Assembly for Bioengineering

Facile Preparation of Internally Self-assembled Lipid Particles Stabilized by Carbon Nanotubes

We present a facile method to prepare nanostructured lipid particles stabilized by carbon nanotubes (CNTs). Single-walled (pristine) and multi-walled (functionalized) CNTs are used as stabilizers to produce Pickering type oil-in-water (O/W) emulsions. Lipids namely, Dimodan U and Phytantriol are used as emulsifiers, which in excess water self-assemble into the bicontinuous cubic Pn3m phase. This highly viscous phase is fragmented into smaller particles using a probe ultrasonicator in presence of conventional surfactant stabilizers or CNTs as done here. Initially, the CNTs (powder form) are dispersed in water followed by further ultrasonication with the molten lipid to form the final emulsion. During this process the CNTs get coated with lipid molecules, which in turn are presumed to surround the lipid droplets to form a particulate emulsion that is stable for months. The average size of CNT-stabilized nanostructured lipid particles is in the submicron range, which compares well with the particles stabilized using conventional surfactants. Small angle X-ray scattering data confirms the retention of the original Pn3m cubic phase in the CNT-stabilized lipid dispersions as compared to the pure lipid phase (bulk state). Blue shift and lowering of the intensities in characteristic G and G&#39; bands of CNTs observed in Raman spectroscopy characterize the interaction between CNT surface and lipid molecules. These results suggest that the interactions between the CNTs and lipids are responsible for their mutual stabilization in aqueous solutions. As the concentrations of CNTs employed for stabilization are very low and lipid molecules are able to functionalize the CNTs, the toxicity of CNTs is expected to be insignificant while their biocompatibility is greatly enhanced. Hence the present approach finds a great potential in various biomedical applications, for instance, for developing hybrid nanocarrier systems for the delivery of multiple functional molecules as in combination therapy or polytherapy.

We report on a smart application of carbon nanotubes for kinetic stabilization of lipid particles that contain self-assembled nanostructures in their cores. The preparation of lipid particles requires rather low concentrations of carbon nanotubes permitting their use in biomedical applications such as drug delivery.

We present a facile method to prepare nanostructured lipid particles stabilized by carbon nanotubes (CNTs). Single-walled (pristine) and multi-walled ...

Chemistry

Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method

Lipid bilayer membranes form the plasma membranes of cells and define the boundaries of subcellular organelles. In nature, these membranes are heterogeneous mixtures of many types of lipids, contain membrane-bound proteins and are decorated with carbohydrates. In some experiments, it is desirable to decouple the biophysical or biochemical properties of the lipid bilayer from those of the natural membrane. Such cases call for the use of model systems such as giant vesicles, liposomes or supported lipid bilayers (SLBs). Arrays of SLBs are particularly attractive for sensing applications and mimicking cell-cell interactions. Here we describe a new method for forming SLB arrays. Submicron-diameter SiO2 beads are first coated with lipid bilayers to form spherical SLBs (SSLBs). The beads are then deposited into an array of micro-fabricated submicron-diameter microwells. The preparation technique uses a "squeegee" to clean the substrate surface, while leaving behind SSLBs that have settled into microwells. This method requires no chemical modification of the microwell substrate, nor any particular targeting ligands on the SSLB. Microwells are occupied by single beads because the well diameter is tuned to be just larger than the bead diameter. Typically, more 75% of the wells are occupied, while the rest remain empty. In buffer SSLB arrays display long-term stability of greater than one week. Multiple types of SSLBs can be placed in a single array by serial deposition, and the arrays can be used for sensing, which we demonstrate by characterizing the interaction of cholera toxin with ganglioside GM1. We also show that phospholipid vesicles without the bead supports and biomembranes from cellular sources can be arrayed with the same method and cell-specific membrane lipids can be identified.

Supported lipid bilayers and natural membrane particles are convenient systems that can approximate the properties of cell membranes and be incorporated in a variety of analytical strategies. Here we demonstrate a method for preparing microarrays composed of supported lipid bilayer-coated SiO2 beads, phospholipid vesicles or natural membrane particles.

Lipid bilayer membranes form the plasma membranes of cells and define the boundaries of subcellular organelles. In nature, these membranes are ...

Lipid Bilayer Vesicle Generation Using Microfluidic Jetting

Bottom-up synthetic biology presents a novel approach for investigating and reconstituting biochemical systems and, potentially, minimal organisms. This emerging field engages engineers, chemists, biologists, and physicists to design and assemble basic biological components into complex, functioning systems from the bottom up. Such bottom-up systems could lead to the development of artificial cells for fundamental biological inquiries and innovative therapies1,2. Giant unilamellar vesicles (GUVs) can serve as a model platform for synthetic biology due to their cell-like membrane structure and size. Microfluidic jetting, or microjetting, is a technique that allows for the generation of GUVs with controlled size, membrane composition, transmembrane protein incorporation, and encapsulation3. The basic principle of this method is the use of multiple, high-frequency fluid pulses generated by a piezo-actuated inkjet device to deform a suspended lipid bilayer into a GUV. The process is akin to blowing soap bubbles from a soap film. By varying the composition of the jetted solution, the composition of the encompassing solution, and/or the components included in the bilayer, researchers can apply this technique to create customized vesicles. This paper describes the procedure to generate simple vesicles from a droplet interface bilayer by microjetting.

Microfluidic jetting against a droplet interface lipid bilayer provides a reliable way to generate vesicles with control over membrane asymmetry, incorporation of transmembrane proteins, and encapsulation of material. This technique can be applied to study a variety of biological systems where compartmentalized biomolecules are desired.

Bottom-up synthetic biology presents a novel approach for investigating and reconstituting biochemical systems and, potentially, minimal organisms. This ...

Automated Lipid Bilayer Membrane Formation Using a Polydimethylsiloxane Thin Film

An artificial lipid bilayer, or black lipid membrane (BLM), is a powerful tool for studying ion channels and protein interactions, as well as for biosensor applications. However, conventional BLM formation techniques have several drawbacks and they often require specific expertise and laborious processes. In particular, conventional BLMs suffer from low formation success rates and inconsistent membrane formation time. Here, we demonstrate a storable and transportable BLM formation system with controlled thinning-out time and enhanced BLM formation rate by replacing conventionally used films (polytetrafluoroethylene, polyoxymethylene, polystyrene) to polydimethylsiloxane (PDMS). In this experiment, a porous-structured polymer such as PDMS thin film is used. In addition, as opposed to conventionally used solvents with low viscosity, the use of squalene permitted a controlled thinning-out time via slow solvent absorption by PDMS, prolonging membrane lifetime. In addition, by using a mixture of squalene and hexadecane, the freezing point of the lipid solution was increased (~16 &#176;C), in addition, membrane precursors were produced that can be indefinitely stored and readily transported. These membrane precursors have reduced BLM formation time of &#60; 1 hr and achieved a BLM formation rate of ~80%. Moreover, ion channel experiments with gramicidin A demonstrated the feasibility of the membrane system.

We demonstrate a storable, transportable lipid bilayer formation system. A lipid bilayer membrane can be formed within 1 hr with over 80% success rate when a frozen membrane precursor is brought to ambient temperature. This system will reduce laborious processes and expertise associated with ion channels.

An artificial lipid bilayer, or black lipid membrane (BLM), is a powerful tool for studying ion channels and protein interactions, as well as for ...

Directed Assembly of Elastin-like Proteins into defined Supramolecular Structures and Cargo Encapsulation In Vitro

Tailored proteinaceous building blocks are versatile candidates for the assembly of supramolecular structures such as minimal cells, drug delivery vehicles and enzyme scaffolds. Due to their biocompatibility and tunability on the genetic level, Elastin-like proteins (ELP) are ideal building blocks for biotechnological and biomedical applications. Nevertheless, the assembly of protein based supramolecular structures with distinct physiochemical properties and good encapsulation potential remains challenging.
Here we provide two efficient protocols for guided self-assembly of amphiphilic ELPs into supramolecular protein architectures such as spherical coacervates, fibers and stable vesicles. The presented assembly protocols generate Protein Membrane-Based Compartments (PMBCs) based on ELPs with adaptable physicochemical properties. PMBCs demonstrate phase separation behavior and reveal method dependent membrane fusion and are able to encapsulate chemically diverse fluorescent cargo molecules. The resulting PMBCs have a high application potential as a drug formulation and delivery platform, artificial cell, and compartmentalized reaction space.

At the interface of organic and aqueous solvents, tailored amphiphilic elastin-like proteins assemble into complex supramolecular structures such as vesicles, fibers and coacervates triggered by environmental parameters. The described assembly protocols yield Protein Membrane-Based Compartments (PMBCs) with tunable properties, enabling the encapsulation of various cargo.

Tailored proteinaceous building blocks are versatile candidates for the assembly of supramolecular structures such as minimal cells, drug delivery ...

Self-Assembly of Hybrid Lipid Membranes Doped with Hydrophobic Organic Molecules at the Water/Air Interface

Because of their unique properties, including an ultrathin thickness (3-4 nm), ultrahigh resistivity, fluidity and self-assembly ability, lipid bilayers can be readily functionalized and have been used in various applications such as bio-sensors and bio-devices. In this study, we introduced a planar organic molecule: copper (II) 2,9,16,23-tetra-tert-butyl-29H,31H-phthalocyanine (CuPc) to dope lipid membranes. The CuPc/lipid hybrid membrane forms at the water/air interface by self-assembly. In this membrane, the hydrophobic CuPc molecules are located between the hydrophobic tails of lipid molecules, forming a lipid/CuPc/lipid sandwich structure. Interestingly, an air-stable hybrid lipid bilayer can be readily formed by transferring the hybrid membrane onto a Si substrate. We report a straightforward method for incorporating nanomaterials into a lipid bilayer system, which represents a new methodology for the fabrication of biosensors and biodevices.

We report a protocol for producing a hybrid lipid membrane at the water/air interface by doping the lipid bilayer with copper (II) 2,9,16,23-tetra-tert-butyl-29H,31H-phthalocyanine (CuPc) molecules. The resulting hybrid lipid membrane has a lipid/CuPc/lipid sandwich structure. This protocol can also be applied to the formation of other functional nanomaterials.

Because of their unique properties, including an ultrathin thickness (3-4 nm), ultrahigh resistivity, fluidity and self-assembly ability, lipid bilayers ...

Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction with lipids employ naked bilayer membranes. Such systems lack the complexities of crowding by membrane-embedded proteins and glycans and exclude the associated volume effects encountered on cellular membrane surfaces. Also, the negatively charged glass surface onto which the lipid bilayers are formed prevents the free diffusion of transmembrane biomolecules. Here, we present a well-characterized polymer-lipid membrane as a mimic for crowded lipid membranes. This protocol utilizes polyethylene glycol (PEG)-conjugated lipids as a generalized approach for incorporating crowders into the supported lipid bilayer&#160;(SLB). First, a cleaning procedure of the microscopic slides and coverslips for performing single-molecule experiments is presented. Next, methods for characterizing the PEG-SLBs and performing single-molecule experiments of the binding, diffusion, and assembly of biomolecules using single-molecule tracking and photobleaching are discussed. Finally, this protocol demonstrates how to monitor the nanopore assembly of bacterial pore-forming toxin Cytolysin A (ClyA) on crowded lipid membranes with single-molecule photobleaching analysis. MATLAB codes with example datasets are also included to perform some of the common analyses such as particle tracking, extracting diffusive behavior, and subunit counting.

Here, a protocol to perform and analyze the binding, mobility, and assembly of single molecules on artificial crowded lipid membranes using single-molecule total internal reflection fluorescence (smTIRF) microscopy is presented.

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction ...

Biochemistry

Preparation and Characterization of Nanoliposomes for the Entrapment of Bioactive Hydrophilic Globular Proteins

Liposome nanocapsules have been applied for many purposes in the pharmaceutical, cosmetic, and food industries. Attributes of liposomes include their biocompatibility, biodegradability, non-immunogenicity, non-toxicity, and ability to entrap both hydrophilic and hydrophobic compounds. The classical hydration of thin lipid films in an organic solvent is applied herein as a technique to encapsulate tarin, a plant lectin, in nanoliposomes. Nanoliposome size, stability, entrapment efficiency, and morphological characterization are described in detail. The nanoliposomes are prepared using 1,2-dioleoyl-sn-glycerol-3-phosphoethanolamine (DOPE), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)-2000] (ammonium salt; DSPE-MPEG 2000), and cholesterylhemisuccinate (CHEMS) as the main constituents. Lipids are first dissolved in chloroform to obtain a thin lipid film that is subsequently rehydrated in ammonium sulfate solution containing the protein to be entrapped and incubated overnight. Then, sonication and extrusion techniques are applied to generate nanosized unilamellar vesicles. The size and polydispersity index of the nanovesicles are determined by dynamic light scattering, while nanovesicle morphology is assessed by scanning electron microscopy. Entrapment efficiency is determined by the ratio of the amount of unencapsulated protein to original amount of initially loaded protein. Homogeneous liposomes are obtained with an average size of 155 nm and polydispersity index value of 0.168. A high entrapment efficiency of 83% is achieved.

This study describes classical hydration using the thin lipid film method for nanoliposome preparation followed by nanoparticle characterization. A 47 kDa-hydrophilic and globular protein, tarin, is successfully encapsulated as a strategy to improve stability, avoid fast clearance, and promote controlled release. The method can be adapted to hydrophobic molecules encapsulation.

Liposome nanocapsules have been applied for many purposes in the pharmaceutical, cosmetic, and food industries. Attributes of liposomes include their ...

Microfluidic Production of Lysolipid-Containing Temperature-Sensitive Liposomes

The presented protocol enables a high-throughput continuous preparation of low temperature-sensitive liposomes (LTSLs), which are capable of loading chemotherapeutic drugs, such as doxorubicin (DOX). To achieve this, an ethanolic lipid mixture and ammonium sulfate solution are injected into a staggered herringbone micromixer (SHM) microfluidic device. The solutions are rapidly mixed by the SHM, providing a homogeneous solvent environment for liposomes self-assembly. Collected liposomes are first annealed, then dialyzed to remove residual ethanol. An ammonium sulfate pH-gradient is established through buffer exchange of the external solution by using size exclusion chromatography. DOX is then remotely loaded into the liposomes with high encapsulation efficiency (&#62; 80%). The liposomes obtained are homogenous in size with Z-average diameter of 100 nm. They are capable of temperature-triggered burst release of encapsulated DOX in the presence of mild hyperthermia (42 &#176;C). Indocyanine green (ICG) can also be co-loaded into the liposomes for near-infrared laser-triggered DOX release. The microfluidic approach ensures high-throughput, reproducible and scalable preparation of LTSLs.

The protocol presents the optimized parameters for preparing thermosensitive liposomes using the staggered herringbone micromixer microfluidics device. This also allows co-encapsulation of doxorubicin and indocyanine green into the liposomes and the photothermal-triggered release of doxorubicin for controlled/triggered drug release.

The presented protocol enables a high-throughput continuous preparation of low temperature-sensitive liposomes (LTSLs), which are capable of loading ...

Formulating and Characterizing Lipid Nanoparticles for Gene Delivery using a Microfluidic Mixing Platform

Lipid-based drug carriers have been used for clinically and commercially available delivery systems due to their small size, biocompatibility, and high encapsulation efficiency. Use of lipid nanoparticles (LNPs) to encapsulate nucleic acids is advantageous to protect the RNA or DNA from degradation, while also promoting cellular uptake. LNPs often contain multiple lipid components including an ionizable lipid, helper lipid, cholesterol, and polyethylene glycol (PEG) conjugated lipid. LNPs can readily encapsulate nucleic acids due to the ionizable lipid presence, which at low pH is cationic and allows for complexation with negatively charged RNA or DNA. Here LNPs are formed by encapsulating messenger RNA (mRNA) or plasmid DNA (pDNA) using rapid mixing of the lipid components in an organic phase and the nucleic acid component in an aqueous phase. This mixing is performed using a precise microfluidic mixing platform, allowing for nanoparticle self-assembly while maintaining laminar flow. The hydrodynamic size and polydispersity are measured using dynamic light scattering (DLS). The effective surface charge on the LNP is determined by measuring the zeta potential. The encapsulation efficiency is characterized using a fluorescent dye to quantify entrapped nucleic acid. Representative results demonstrate the reproducibility of this method and the influence that different formulation and process parameters have on the developed LNPs.

Lipid nanoparticles are developed using a microfluidic mixing platform approach for mRNA and DNA encapsulation.

Lipid-based drug carriers have been used for clinically and commercially available delivery systems due to their small size, biocompatibility, and high ...

On-Chip Octanol-Assisted Liposome Assembly for Bioengineering

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