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 no...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#8211;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&#8211;70,000, 87%&#8211;90% hydrolyzed</td>
		</tr>
		<tr>
			<td>Pressure controller</td>
			<td>Elveflow&#160;</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&#160;</td>
			<td>Laurell Technologies Corporation</td>
			<td>Model WS-650MZ-23NPP</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless Steel 90&#176; 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&#34; OD x 0.02&#34; ID</td>
			<td></td>
		</tr>
		<tr>
			<td>UV laser&#160;</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)<...

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

Research

JoVE Journal

Bioengineering

2.3K Views.  Wageningen University & Research. 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.

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

Synthesis, Assembly, and Characterization of Monolayer Protected Gold Nanoparticle Films for Protein Monolayer Electrochemistry

Colloidal gold nanoparticles protected with alkanethiolate ligands called monolayer protected gold clusters (MPCs) are synthesized and subsequently incorporated into film assemblies that serve as adsorption platforms for protein monolayer electrochemistry (PME). PME is utilized as the model system for studying electrochemical properties of redox proteins by confining them to an adsorption platform at a modified electrode, which also serves as a redox partner for electron transfer (ET) reactions. Studies have shown that gold nanoparticle film assemblies of this nature provide for a more homogeneous protein adsorption environment and promote ET without distance dependence compared to the more traditional systems modified with alkanethiol self-assembled monolayers (SAM).1-3 In this paper, MPCs functionalized with hexanethiolate ligands are synthesized using a modified Brust reaction4 and characterized with ultraviolet visible (UV-Vis) spectroscopy, transmission electron microscopy (TEM), and proton (1H) nuclear magnetic resonance (NMR). MPC films are assembled on SAM modified gold electrode interfaces by using a "dip cycle" method of alternating MPC layers and dithiol linking molecules. Film growth at gold electrode is tracked electrochemically by measuring changes to the double layer charging current of the system. Analogous films assembled on silane modified glass slides allow for optical monitoring of film growth and cross-sectional TEM analysis provides an estimated film thickness. During film assembly, manipulation of the MPC ligand protection as well as the interparticle linkage mechanism allow for networked films, that are readily adaptable, to interface with redox protein having different adsorption mechanism. For example, Pseudomonas aeruginosa azurin (AZ) can be adsorbed hydrophobically to dithiol-linked films of hexanethiolate MPCs and cytochrome c (cyt c) can be immobilized electrostatically at a carboxylic acid modified MPC interfacial layer. In this report, we focus on the film protocol for the AZ system exclusively. Investigations involving the adsorption of proteins on MPC modified synthetic platforms could further the understanding of interactions between biomolecules and man-made materials, and consequently aid the development of biosensor schemes, ET modeling systems, and synthetic biocompatible materials.5-8

Alkanethiolate stabilized gold colloids known as monolayer protected clusters (MPCs) are synthesized, characterized, and assembled into thin films as an adsorption interface for protein monolayer electrochemistry of simple redox protein like Pseudomonas aeruginosa azurin (AZ) and cytochrome c (cyt c).

Colloidal gold nanoparticles protected with alkanethiolate ligands called monolayer protected gold clusters (MPCs) are synthesized and subsequently ...

On-chip Isotachophoresis for Separation of Ions and Purification of Nucleic Acids

Electrokinetic techniques are a staple of microscale applications because of their unique ability to perform a variety of fluidic and electrophoretic processes in simple, compact systems with no moving parts. Isotachophoresis (ITP) is a simple and very robust electrokinetic technique that can achieve million-fold preconcentration1,2 and efficient separation and extraction based on ionic mobility.3 For example, we have demonstrated the application of ITP to separation and sensitive detection of unlabeled ionic molecules (e.g. toxins, DNA, rRNA, miRNA) with little or no sample preparation4-8 and to extraction and purification of nucleic acids from complex matrices including cell culture, urine, and blood.9-12
ITP achieves focusing and separation using an applied electric field and two buffers within a fluidic channel system. For anionic analytes, the leading electrolyte (LE) buffer is chosen such that its anions have higher effective electrophoretic mobility than the anions of the trailing electrolyte (TE) buffer (Effective mobility describes the observable drift velocity of an ion and takes into account the ionization state of the ion, as described in detail by Persat et al.13). After establishing an interface between the TE and LE, an electric field is applied such that LE ions move away from the region occupied by TE ions. Sample ions of intermediate effective mobility race ahead of TE ions but cannot overtake LE ions, and so they focus at the LE-TE interface (hereafter called the "ITP interface"). Further, the TE and LE form regions of respectively low and high conductivity, which establish a steep electric field gradient at the ITP interface. This field gradient preconcentrates sample species as they focus. Proper choice of TE and LE results in focusing and purification of target species from other non-focused species and, eventually, separation and segregation of sample species.
We here review the physical principles underlying ITP and discuss two standard modes of operation: "peak" and "plateau" modes. In peak mode, relatively dilute sample ions focus together within overlapping narrow peaks at the ITP interface. In plateau mode, more abundant sample ions reach a steady-state concentration and segregate into adjoining plateau-like zones ordered by their effective mobility. Peak and plateau modes arise out of the same underlying physics, but represent distinct regimes differentiated by the initial analyte concentration and/or the amount of time allotted for sample accumulation.
We first describe in detail a model peak mode experiment and then demonstrate a peak mode assay for the extraction of nucleic acids from E. coli cell culture. We conclude by presenting a plateau mode assay, where we use a non-focusing tracer (NFT) species to visualize the separation and perform quantitation of amino acids.

Isotachophoresis (ITP) is a robust electrokinetic separation and preconcentration technique with applications ranging from toxin detection to sample preparation. We review the physical principles of ITP and the methodology of applying this technique to two specific example applications: separation and detection of small molecules and purification of nucleic acids from cell culture lysate.

Electrokinetic techniques are a staple of microscale applications because of their unique ability to perform a variety of fluidic and electrophoretic ...

Lensfree On-chip Tomographic Microscopy Employing Multi-angle Illumination and Pixel Super-resolution

Tomographic imaging has been a widely used tool in medicine as it can provide three-dimensional (3D) structural information regarding objects of different size scales. In micrometer and millimeter scales, optical microscopy modalities find increasing use owing to the non-ionizing nature of visible light, and the availability of a rich set of illumination sources (such as lasers and light-emitting-diodes) and detection elements (such as large format CCD and CMOS detector-arrays). Among the recently developed optical tomographic microscopy modalities, one can include optical coherence tomography, optical diffraction tomography, optical projection tomography and light-sheet microscopy. 1-6 These platforms provide sectional imaging of cells, microorganisms and model animals such as C. elegans, zebrafish and mouse embryos.
Existing 3D optical imagers generally have relatively bulky and complex architectures, limiting the availability of these equipments to advanced laboratories, and impeding their integration with lab-on-a-chip platforms and microfluidic chips. To provide an alternative tomographic microscope, we recently developed lensfree optical tomography (LOT) as a high-throughput, compact and cost-effective optical tomography modality. 7 LOT discards the use of lenses and bulky optical components, and instead relies on multi-angle illumination and digital computation to achieve depth-resolved imaging of micro-objects over a large imaging volume. LOT can image biological specimen at a spatial resolution of &lt;1 &mu;m x &lt;1 &mu;m x &lt;3 &mu;m in the x, y and z dimensions, respectively, over a large imaging volume of 15-100 mm3, and can be particularly useful for lab-on-a-chip platforms.

Lensfree optical tomography is a three-dimensional microscopy technique that offers a spatial resolution of &lt;1 &mu;m &times; &lt;1 &mu;m &times; &lt;3 &mu;m in x, y and z dimensions, respectively, over a large imaging-volume of 15-100 mm3, which can be particularly useful for integration with lab-on-a-chip platforms.

Tomographic imaging has been a widely used tool in medicine as it can provide three-dimensional (3D) structural information regarding objects of different ...

On-Chip Endothelial Inflammatory Phenotyping

Atherogenesis is potentiated by metabolic abnormalities that contribute to a heightened state of systemic inflammation resulting in endothelial dysfunction. However, early functional changes in endothelium that signify an individual's level of risk are not directly assessed clinically to help guide therapeutic strategy. Moreover, the regulation of inflammation by local hemodynamics contributes to the non-random spatial distribution of atherosclerosis, but the mechanisms are difficult to delineate in vivo. We describe a lab-on-a-chip based approach to quantitatively assay metabolic perturbation of inflammatory events in human endothelial cells (EC) and monocytes under precise flow conditions. Standard methods of soft lithography are used to microfabricate vascular mimetic microfluidic chambers (VMMC), which are bound directly to cultured EC monolayers.1 These devices have the advantage of using small volumes of reagents while providing a platform for directly imaging the inflammatory events at the membrane of EC exposed to a well-defined shear field. We have successfully applied these devices to investigate cytokine-,2 lipid-3, 4 and RAGE-induced5 inflammation in human aortic EC (HAEC). Here we document the use of the VMMC to assay monocytic cell (THP-1) rolling and arrest on HAEC monolayers that are conditioned under differential shear characteristics and activated by the inflammatory cytokine TNF-&alpha;. Studies such as these are providing mechanistic insight into atherosusceptibility under metabolic risk factors.

Microfluidic flow chambers etched by photolithography and fabricated from PDMS are applied to probe functional outcomes associated with EC dysfunction and inflammation. In a representative experiment, the ability of differential shear stress to modulate monocytic cell adhesion to cytokine activated EC monolayers is demonstrated.

Atherogenesis is potentiated by metabolic abnormalities that contribute to a heightened state of systemic inflammation resulting in endothelial ...

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

Synthesis of Cationized Magnetoferritin for Ultra-fast Magnetization of Cells

Many important biomedical applications, such as cell imaging and remote manipulation, can be achieved by labeling cells with superparamagnetic iron oxide nanoparticles (SPIONs). Achieving sufficient cellular uptake of SPIONs is a challenge that has traditionally been met by exposing cells to elevated concentrations of SPIONs or by prolonging exposure times (up to 72 hr). However, these strategies are likely to mediate toxicity. Here, we present the synthesis of the protein-based SPION magnetoferritin as well as a facile surface functionalization protocol that enables rapid cell magnetization using low exposure concentrations. The SPION core of magnetoferritin consists of cobalt-doped iron oxide with an average particle diameter of 8.2 nm mineralized inside the cavity of horse spleen apo-ferritin. Chemical cationization of magnetoferritin produced a novel, highly membrane-active SPION that magnetized human mesenchymal stem cells (hMSCs) using incubation times as short as one minute and iron concentrations as lows as 0.2 mM.

A protocol for the synthesis and cationization of cobalt-doped magnetoferritin is presented, as well as a method to rapidly magnetize stem cells with cationized magnetoferritin.

Many important biomedical applications, such as cell imaging and remote manipulation, can be achieved by labeling cells with superparamagnetic iron oxide ...

Assembly and Tracking of Microbial Community Development within a Microwell Array Platform

The development of microbial communities depends on a combination of complex deterministic and stochastic factors that can dramatically alter the spatial distribution and activities of community members. We have developed a microwell array platform that can be used to rapidly assemble and track thousands of bacterial communities in parallel. This protocol highlights the utility of the platform and describes its use for optically monitoring the development of simple, two-member communities within an ensemble of arrays within the platform. This demonstration uses two mutants of Pseudomonas aeruginosa, part of a series of mutants developed to study Type VI secretion pathogenicity. Chromosomal inserts of either mCherry or GFP genes facilitate the constitutive expression of fluorescent proteins with distinct emission wavelengths that can be used to monitor community member abundance and location within each microwell. This protocol describes a detailed method for assembling mixtures of bacteria into the wells of the array and using time-lapse fluorescence imaging and quantitative image analysis to measure the relative growth of each member population over time. The seeding and assembly of the microwell platform, the imaging procedures necessary for the quantitative analysis of microbial communities within the array, and the methods that can be used to reveal interactions between microbial species area all discussed.

The development of microbial communities depends on a combination of factors, including environmental architecture, member abundance, traits, and interactions. This protocol describes a synthetic, microfabricated environment for the simultaneous tracking of thousands of communities contained in femtoliter wells, where key factors such as niche size and confinement can be approximated.

The development of microbial communities depends on a combination of complex deterministic and stochastic factors that can dramatically alter the spatial ...

Dry Film Photoresist-based Electrochemical Microfluidic Biosensor Platform: Device Fabrication, On-chip Assay Preparation, and System Operation

In recent years, biomarker diagnostics became an indispensable tool for the diagnosis of human disease, especially for the point-of-care diagnostics. An easy-to-use and low-cost sensor platform is highly desired to measure various types of analytes (e.g., biomarkers, hormones, and drugs) quantitatively and specifically. For this reason, dry film photoresist technology - enabling cheap, facile, and high-throughput fabrication - was used to manufacture the microfluidic biosensor presented here. Depending on the bioassay used afterwards, the versatile platform is capable of detecting various types of biomolecules. For the fabrication of the device, platinum electrodes are structured on a flexible polyimide (PI) foil in the only clean-room process step. The PI foil serves as a substrate for the electrodes, which are insulated with an epoxy-based photoresist. The microfluidic channel is subsequently generated by the development and lamination of dry film photoresist (DFR) foils onto the PI wafer. By using a hydrophobic stopping barrier in the channel, the channel is separated into two specific areas: an immobilization section for the enzyme-linked assay and an electrochemical measurement cell for the amperometric signal readout.
The on-chip bioassay immobilization is performed by the adsorption of the biomolecules to the channel surface. The glucose oxidase enzyme is used as a transducer for electrochemical signal generation. In the presence of the substrate, glucose, hydrogen peroxide is produced, which is detected at the platinum working electrode. The stop-flow technique is applied to obtain signal amplification along with rapid detection. Different biomolecules can quantitatively be measured by means of the introduced microfluidic system, giving an indication of different types of diseases, or, in regard to therapeutic drug monitoring, facilitating a personalized therapy.

A microfluidic biosensor platform was designed and fabricated using low-cost dry film photoresist technology for the rapid and sensitive quantification of various analytes. This single-use system allows for the electrochemical readout of on-chip-immobilized enzyme-linked assays by means of the stop-flow technique.

In recent years, biomarker diagnostics became an indispensable tool for the diagnosis of human disease, especially for the point-of-care diagnostics. An ...

Fabrication and Validation of an Organ-on-chip System with Integrated Electrodes to Directly Quantify Transendothelial Electrical Resistance

Organs-on-chips, in vitro models involving the culture of (human) tissues inside microfluidic devices, are rapidly emerging and promise to provide useful research tools for studying human health and disease. To characterize the barrier function of cell layers cultured inside organ-on-chip devices, often transendothelial or transepithelial electrical resistance (TEER) is measured. To this end, electrodes are usually integrated into the chip by micromachining methods to provide more stable measurements than is achieved with manual insertion of electrodes into the inlets of the chip. However, these electrodes frequently hamper visual inspection of the studied cell layer or require expensive cleanroom processes for fabrication. To overcome these limitations, the device described here contains four easily integrated electrodes that are placed and fixed outside of the culture area, making visual inspection possible. Using these four electrodes the resistance of six measurement paths can be quantified, from which the TEER can be directly isolated, independent of the resistance of culture medium-filled microchannels. The blood-brain barrier was replicated in this device and its TEER was monitored to show the device applicability. This chip, the integrated electrodes and the TEER determination method are generally applicable in organs-on-chips, both to mimic other organs or to be incorporated into existing organ-on-chip systems.

This publication describes the fabrication of an organ-on-chip device with integrated electrodes for direct quantification of transendothelial electrical resistance (TEER). For validation, the blood-brain barrier was mimicked inside this microfluidic device and its barrier function was monitored. The presented methods for electrode integration and direct TEER quantification are generally applicable.

Organs-on-chips, in vitro models involving the culture of (human) tissues inside microfluidic devices, are rapidly emerging and promise to provide useful ...

Antimicrobial Characterization of Advanced Materials for Bioengineering Applications

The development of new advanced materials with enhanced properties is becoming more and more important in a wide range of bioengineering applications. Thus, many novel biomaterials are being designed to mimic specific environments required for biomedical applications such as tissue engineering and controlled drug delivery. The development of materials with improved properties for the immobilization of cells or enzymes is also a current research topic in bioprocess engineering. However, one of the most desirable properties of a material in these applications is the antimicrobial capacity to avoid any undesirable infections. For this, we present easy-to-follow protocols for the antimicrobial characterization of materials based on (i) the agar disk diffusion test (diffusion method) and (ii) the ISO 22196:2007 norm to measure the antimicrobial activity on material surfaces (contact method). This protocol must be performed using Gram-positive and Gram-negative bacteria and yeast to cover a broad range of microorganisms. As an example, 4 materials with different chemical natures are tested following this protocol against Staphylococcus aureus, Escherichia coli, and Candida albicans.The results of these tests exhibit non-antimicrobial activity for the first material and increasing antibacterial activity against Gram-positive and Gram-negative bacteria for the other 3 materials. However, none of the 4 materials are able to inhibit the growth of Candida albicans.

We present a protocol for the antimicrobial characterization of advanced materials. Here, the antimicrobial activity on material surfaces is measured by two methods that complement each other: one is based on the agar disk diffusion test, and the other is a standard procedure based on the ISO 22196:2007 norm.

The development of new advanced materials with enhanced properties is becoming more and more important in a wide range of bioengineering applications. ...