This work was supported by grants to S.H.O. from the National Institutes of Health (R01 GM092993), the National Science Foundation (NSF CAREER Award and DBI 0964216), the Office of Naval Research (ONR) Young Investigator Program and the Minnesota Partnership Award for Biotechnology and Medical Genomics. Device fabrication was performed at the University of Minnesota Nanofabrication Center (NFC), which receives support from the NSF through the National Nanotechnology Infrastructure Network. This work was also supported by grants to M.R. from the National Institutes of Health (NS048357, R21 NS073684), the National Multiple Sclerosis Society (CA1060A11), the Applebaum, Hilton, Peterson and Sanford Foundations and the McNeilus family. The authors wish to thank Hyungsoon Im for assistance with illustrations and Shailabh Kumar for assistance with scanning electron microscopy.

1. Microfabrication of Microwell Array Substrate
<ol>
	<li>Start with a 4 inch silicon wafer with 100 nm of thermally grown oxide.</li>
	<li>Spin SPR-955 0.7 photoresist on the wafer at 4,000 rpm for 30 sec.</li>
	<li>Bake on a hotplate at 115 &#176;C for 90 sec.</li>
	<li>Expose photoresist.
	<ol>
		<li>Use a mask that will create 1 &#181;m holes arranged in a hexagonal array with a 3 &#181;m period where the array covers a 2 mm x 2 mm area.</li>
		<li>Expose wafer in an i-line stepper using a step size of 6 mm and an exposure of 200 mJ/cm2.</li>
	</ol>
	</li>
	<li>Bake on a hotplate at 115 &#176;C for 90 sec.</li>
	<li>Develop using a spin developer to deposit 2 puddles of CD-26 on the wafer for a total development time of 90 sec.</li>
	<li>Rinse thoroughly with ultrapure DI water and dry with N2.</li>
	<li>Etch the oxide in a reactive-ion etcher for 6 min with 50 sccm of Ar, 25 sccm of CF4, and 50 sccm of CHF3 flowing at a pressure of 75 mTorr.</li>
	<li>Etch silicon in an ICP deep-trench reactive-ion etcher for 2 min with 63 sccm of C4F8, 27 sccm of SF6, 40 sccm of Ar, and 10 sccm of O2 flowing at a pressure of 14 mTorr.</li>
	<li>Rinse in acetone, methanol, and isopropanol to remove resist.</li>
	<li>Place in a bath of H2SO4:H2O2 1:1 for 10 min to clean the surface.</li>
	<li>Rinse thoroughly with ultrapure DI water and dry with N2.</li>
	<li>Deposit 1,080 &#197; of Al2O3 by atomic layer deposition at 250 &#176;C.</li>
</ol>
2. Preparation of Poly(dimethylsiloxane) Squeegee
<ol>
	<li>If possible, work in a cleanroom. Otherwise work in the cleanest environment possible.</li>
	<li>Obtain a plastic Petri dish (or other clean disposable container) with &#8805;13 mm sides.</li>
	<li>In that dish, pour out 5 g PDMS curing agent, followed by 50 g PDMS base agent (or anything with a 1:10 ratio that will mostly fill the Petri dish). Mix thoroughly with a disposable plastic pipette or rod.</li>
	<li>Remove bubbles by placing the mixed PDMS in a chamber with a vacuum line, applying vacuum until bubbles come out of mixture (approximately 30 min).</li>
	<li>If not already in the Petri dish, pour PDMS in Petri dish, avoiding creation of air bubbles.</li>
	<li>Cure overnight on a hotplate at &#62;50 &#176;C (higher heat for less time also works).</li>
	<li>With new/clean razor blade, carefully cut a rectangular piece approximately 2.5 x 2.5 cm, keeping top surface as clean as possible. Peel out with a clean forceps. Trim off one edge (down to 1.5 cm) by pressing down on razor blade in one motion, to make top edge as sharp as possible (this will be the edge used in the squeegee process).</li>
	<li>Clean with acetone, methanol, isopropyl alcohol, and ultrapure DI water, and then blow dry with clean nitrogen gas. Store in a clean Petri dish.</li>
</ol>
3. Preparation of Vesicles
<ol>
	<li>Collect lipid stock solutions of 25 mg/ml egg phosphatidylcholine (PC) and 1 mg/ml 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) ammonium salt (Rho-DPPE) in chloroform and a 0.5 mg/ml stock solution of monosialoganglioside GM1 in chloroform/methanol (2:1 v/v).</li>
	<li>In a small glass vial make a mixture that results in 97 mol% PC, 2 mol% GM1 and 1 mol% Rho-DPPE by adding 18.9 &#181;l of 25 mg/ml PC, 39.6 &#181;l of 0.5 mg/ml GM1 and 7.9 &#181;l of 1 mg/ml Rho-DPPE using glass syringes.</li>
	<li>Note: The supplementary material for Nair et al.25 contains an excellent customizable Excel spreadsheet for calculating the amounts of lipids required to create vesicles of any desired composition.</li>
	<li>Place vial in a vacuum desiccator and remove the solvent by leaving the vial under vacuum for 6 hr.</li>
	<li>Make an aqueous solution of 0.1 M NaCl. Add 0.5 ml of the 0.1 M NaCl solution to the vial containing the dried lipid film.</li>
	<li>Allow the aqueous lipid mixture to rest overnight.</li>
	<li>Briefly vortex mix to create a vesicle suspension, then sonicate for 20 min in a bath sonicator at room temperature.</li>
	<li>Extrude the vesicle suspension through a polycarbonate membrane with 100 nm pore size. Pass the vesicle suspension through the filter 17x.</li>
	<li>Store the extruded vesicles in a glass vial at 4 &#176;C. </li>
</ol>
4. Formation of Spherical Supported Lipid Bilayers
<ol>
	<li>Collect 700 nm diameter SiO2 beads. The beads are supplied in distilled water with a stock concentration of 1.4 x 1011 beads/ml.</li>
	<li>In a 1.5 ml Eppendorf tube prepare a 1 ml bead suspension with a concentration of 1.5 x 1010 beads/ml by adding 107 &#181;l of bead stock solution to 893 &#181;l of 0.1 M NaCl.</li>
	<li>Vortex the suspension to ensure it is well mixed.</li>
	<li>Centrifuge the suspension at 1,700 x g for 20 min then discard the supernatant. Resuspend the beads in 1 ml of 0.1 M NaCl. Repeat this step twice more to thoroughly wash the beads.</li>
	<li>To form the SSLBs, in a 1.5 ml Eppendorf tube add 25 &#181;l of the SiO2 bead suspension to 200 &#181;l of 1 mg/ml vesicle suspension. Vortex the mixture and let stand for 1 hr at room temperature. At this stage the SSLB concentration should be 1.7 x 109 SSLBs/ml.</li>
	<li>Centrifuge at 1,700 x g for 20 min to pellet the SSLBs. The pellet should appear pink due to the rupture of vesicles with Rho-DPPE on the beads.</li>
	<li>Discard the supernatant and resuspend in 225 &#181;l phosphate buffered saline (PBS), pH = 7.4. Repeat Steps 4.6-4.7 two more times to remove any unruptured vesicles from the SSLB suspension.</li>
</ol>
5. Assembly of SSLB Array
<ol>
	<li>Cleave wafer of microwell arrays resulting in rectangular pieces with 4-6 microwell arrays per piece.</li>
	<li>Vortex mix the SSLB suspension, then place 10 &#181;l of SSLB suspension on each microwell array. Lest rest for 1 hr to allow SSLBs to settle to the surface.</li>
	<li>Gently wash the microwell array chip with PBS from a wash bottle, then submerge in a PBS bath prepared in a shallow dish.</li>
	<li>While submerged, gently place the PDMS squeegee flush on the microwell array chip and gently slide it along the surface 5x to remove SSLBs that are not immobilized in microwells.</li>
	<li>Grip the microwell array chip with tweezers and gently shake in the PBS bath to remove any excess SSLBs.</li>
	<li>Quickly remove the chip from the squeegee bath and place in a fresh PBS bath until further use. Make sure the top surface of the chip remains wet to preserve the SSLBs.</li>
</ol>
Note: The assembly method described above can be used to create arrays of natural membrane particles, like myelin particles isolated from mouse brain, or phospholipid vesicles without the SiO2 bead supports. This work is described in detail in Wittenberg et al.21 and representative results are shown below.
6. Cholera Toxin Binding Assay
<ol>
	<li>Prepare a 2 mg/ml solution of bovine serum albumin (BSA) in PBS.</li>
	<li>Prepare a solution with desired concentration of Alexa 488-conjugated cholera toxin B-subunit in PBS with 2 mg/ml BSA.</li>
	<li>Remove the SSLB array chip from the PBS bath and wick away most of the PBS solution using a laboratory wipe. Leave just enough PBS on the chip to keep the SSLB array hydrated.</li>
	<li>To prevent nonspecific binding, add 200 &#181;l of 2 mg/ml BSA solution to the SSLB array chip. Let rest for 1 hr in a humidified box.</li>
	<li>With a micropipette, remove 200 &#181;l of solution from the top of the SSLB array chip.</li>
	<li>Add 200 &#181;l of cholera toxin solution to the SSLB array chip and let rest for 1 hr in a humidified box.</li>
	<li>Gently wash the SSLB array chip with PBS from a wash bottle to remove any unbound cholera toxin.</li>
	<li>Wick away excess PBS using a laboratory wipe. Before imaging cover chip with a 24 mm x 40 mm cover slip.</li>
	<li>Image arrays with an upright microscope using filter sets appropriate for the fluorophores of interest.</li>
	<li>Analyze the fluorescence intensity of individual SSLBs in an array using the automated particle analysis function of ImageJ software.</li>
	<li>Compile mean fluorescence intensities from individual SSLBs into histograms that summarize the intensities of SSLBs found on a given array.</li>
</ol>

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H., Castillo, J., Gether, U., Hedegard, P., Stamou, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=How+Curved+Membranes+Recruit+Amphipathic+Helices+and+Protein+Anchoring+Motifs.">How Curved Membranes Recruit Amphipathic Helices and Protein Anchoring Motifs.</a> Nat. Chem. Biol. 5 (11), 835-841 (2009).</li><li>Roizard, S., Danelon, C., Hassaine, G., Piguett, J., Schulze, K., Hovius, R., Tampe, R., Vogel, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+G-Protein-Coupled+Receptors+in+Cell-Derived+Plasma+Membranes+Supported+on+Porous+Beads.">Activation of G-Protein-Coupled Receptors in Cell-Derived Plasma Membranes Supported on Porous Beads.</a> J. Am. Chem. Soc. 133 (42), 16868-16874 (2011).</li></ol>

The authors have no competing financial interests to disclose.

In this work we show that monodisperse SiO2 beads coated with supported lipid bilayers can be arrayed into microwell arrays without the need for targeting ligands on the lipid bilayers or the substrate surface, and the arrays can be used for characterizing toxin-lipid interactions. The dissociation constant we calculated for CTx/GM1 binding compares favorably, given the wide disparity of values in the literature, with a previous report by Winter et al., where colloidal assembly of lipid-coated beads was used to calculate a dissociation constant of about 30 nM. By fitting the data to the Hill-Waud binding model, we determined that the CTx/GM1 interaction is likely multivalent, which agrees with previous reports28. The B (ganglioside binding) subunit of CTx exists as a pentamer, with each of the five units capable of binding a GM1 molecule29; thus, it is not surprising that more than one subunit may be bound given the diffusive mobility of GM1 in SLBs30.
Arrays of chemically functionalized beads have been used in other analytical studies31. In these types of experiments, beads are typically immobilized in microwells created in the end of etched optical fibers. Multiplex fluorescence assays can be carried out in this configuration by mixing subpopulations of beads and immobilizing them simultaneously onto the end of the optical fiber32. In our multiplex assay (Figures 3c-3e), we immobilized optically encoded SSLBs sequentially because lipids on SSLBs free in solution could be transferred from one type of SSLB to another33. While this adds extra steps in the preparation of SSLB arrays, it not overly time consuming and is the most efficient approach for lipid-functionalized beads. We have also spatially encoded arrays by depositing different SSLB populations in adjacent microwell arrays, and there is no observable cross-talk between the arrays20. To reduce sample-preparation times, SSLB arrays can be prepared in advance and used when desired. We tested the stability of arrays over the course of one week and found no significant degradation of the arrays in terms of fluorescence intensity, which indicates that the amount of lipid material on the SSLBs does not change with time. Also, the array occupancy does not change over time, which suggests that once SSLBs are immobilized, they do not detach from the microwells (Figure 4b). 

In addition to beads coated with synthetic lipid bilayers, we have also used this method to create arrays of natural membrane particles21. In that work, we immobilized myelin particles derived from mouse brain as well as neuronal membrane particles, such as the lipid-raft fraction. We were able to use the arrays to identify cell-specific membrane surface molecules by antibody binding (Figures 6 and 7). Furthermore, when the size of the microwells was reduced to the nanoscale and the top surface of the array was coated with gold, we were able to employ surface plasmon resonance to monitor binding of antibodies to myelin membrane particles without the use of fluorescent labels. The major difference between using SSLB and vesicles or natural membrane particles to form arrays is that in the case of SSLBs the exact amount of membrane material in each microwell is known because only a single bead can fit in each well. With vesicles or natural membrane particles, there is no way to control the amount of material in each microwell, other than to adjust the starting concentration of vesicles or particles. However, the well-to-well variability of the number of vesicles is not very large compared to the distribution of vesicle diameters21.

Finally, this method can have diverse future applications. Junesch and coworkers have employed a squeegee to remove nanoparticles during colloidal lithography nanofabrication34. Membrane arrays could be formed to mimic cell-cell contacts to investigate cellular focal adhesion35 or examine the binding of membrane curvature-sensing proteins36. In addition, the use of porous beads would facilitate the inclusion of transmembrane proteins into the lipid bilayers37 or local delivery of stored cargo to cells cultured on a biomembrane array substrate. The microwell platform is fabricated by photolithography, which allows for design flexibility. Here our arrays had hexagonal geometry, but square, linear, or other geometries could be created with varying array periodicity and microwell dimensions to investigate the spatial effects of focal adhesion.

Lipid bilayer membranes are essential structures in nature. Cellular plasma membranes and organelle membranes are composed of lipid bilayers that incorporate a number of molecules that are necessary for life. Many life-sustaining processes occur on the surface of cells or are mediated by molecules associated with lipid-bilayer membranes. In fact, many pharmaceuticals target processes or molecules are found on or in membranes1,2. It is therefore necessary to analytically investigate processes, such as chemical reactions or noncovalent binding events that occur on membrane surfaces. Because natural membranes can be difficult to isolate and/or interface with sensors, many researchers employ simplified model membranes to carry out analytical studies. A number of model membrane systems are described in the literature, ranging from giant vesicles that can be tens to hundreds of microns in diameter to liposomes with nanoscale dimensions3,4. Alternatively, planar lipid bilayers deposited on solid supports, i.e., supported lipid bilayers (SLBs), can be formed on a number of different surfaces and have been widely used in biophysical, biochemical, and analytical applications5. Coupling SLBs with electrical or optical materials enables investigation of membrane biochemistry and biophysics through the use of different analytical techniques. Fluorescence microscopy6,&#160;electrochemistry7, optical spectroscopy8, scanning probe microscopy9, surface plasmon resonance10, and mass spectrometry11 have all been employed to study the structure and properties of SLBs.
SLB arrays offer additional versatility in the design of sensors for multiplex assays12,13. Other applications use SLB arrays to mimic the junction that forms between immune cells14. Preparation methods for SLB arrays have varied from microfluidic approaches15 to those that employ physical barriers between adjacent SLB patches.16 Other groups have used printing methods17, photochemical patterning18 and various nanoengineering approaches19 to create SLB arrays.
In this paper and accompanying video we demonstrate a method for forming SLB arrays by depositing SLB-coated SiO2 beads into ordered arrays of microwells20. We refer to the SLB-coated SiO2 beads as spherical supported lipid bilayers (SSLBs). This technique is an extension of earlier work that created arrays of phospholipid vesicles and biomembranes derived from natural sources21, from which we also show example results. Other methods for arraying biomembrane particles or vesicles have relied on patterns of specific targeting ligands on surfaces that associate with complementary ligands contained on the vesicle surface. Examples include biotin-avidin association22,23 and DNA hybridization schemes24. Our approach only requires a microwell array with no targeting or recognition moieties necessary. The size of SSLBs is defined by the diameter of the SiO2 bead supports, which have low poly-dispersity. By tuning the microwell diameter to just larger than the SSLB diameter, only a single SSLB settles into each microwell. A poly(dimethylsiloxane) (PDMS) squeegee then removes from the surface all SSLBs that are not immobilized in microwells. The microwells and resultant SSLB arrays have high density (~105 SSLBs/mm2) with 3 &#181;m center-to-center spacing and hexagonal periodicity. By serially depositing SSLBs with different lipid compositions, it is possible to create multicomponent arrays with randomly positioned SSLBs. To demonstrate the sensing capability of SSLB arrays, we used the interaction of cholera toxin (CTx) with a ganglioside (GM1) incorporated into the SSLBs. With natural membrane particles, we were able to detect cell-specific lipids in multicomponent arrays containing membrane material from two different cell types.&#160;

<table><tbody><tr><td>4 inch silicon wafers</td><td>University Wafer</td><td>425</td><td></td></tr><tr><td>Shipley MEGAPOSIT SPR955-CM 0.7 photoresist</td><td>MicroChem</td><td>SPR955-CM</td><td></td></tr><tr><td>Shipley MICROPOSIT CD-26 developer</td><td>MicroChem</td><td>CD-26</td><td></td></tr><tr><td>i-line stepper</td><td>Canon</td><td>2500 i3 stepper</td><td></td></tr><tr><td>Vision 320 reactive ion etcher</td><td>Advanced Vacuum</td><td>Vision 320 RIE</td><td></td></tr><tr><td>Deep trench reactive ion etcher</td><td>Plasma Therm</td><td>SLR-770</td><td></td></tr><tr><td>Atomic layer depostion system</td><td>Cambridge NanoTech</td><td>Savannah</td><td></td></tr><tr><td>Dow Corning Sylgard 184 poly(dimethylsiloxane) kit</td><td>Ellsworth Adhesives</td><td>184 SIL ELAST KIT 0.5KG</td><td></td></tr><tr><td>Egg phosphatidylcholine</td><td>Avanti Polar Lipids</td><td>840051C</td><td></td></tr><tr><td>1,2-dipalmitoyl-&lt;em&gt;sn&lt;/em&gt;-glycero-3-phosphoethanolamine-&lt;em&gt;N&lt;/em&gt;-(lissamine rhodamine B sulfonyl) ammonium salt</td><td>Avanti Polar Lipids</td><td>810158C</td><td></td></tr><tr><td>Monosialoganglioside GM1</td><td>Avanti Polar Lipids</td><td>860065P</td><td></td></tr><tr><td>Silica beads</td><td>Bangs Laboratories</td><td>SS03N/4666</td><td>Packaging on the bead container states the beads are 900 nm in diameter. However, after light-scattering and electron microscopy we determined the beads are roughly 700 nm in diameter.</td></tr><tr><td>Cholera toxin B-subunit, Alexa 488-conjugate</td><td>Molecular Probes</td><td>C-34775</td><td></td></tr><tr><td>Anti-oligodentrocyte antibody IgM O4, NorthernLights 557 conjugate</td><td>R&amp;amp;D Systems</td><td>NL1326R</td><td></td></tr><tr><td>FM1-43</td><td>Molecular Probes</td><td>T-3136</td><td></td></tr><tr><td>Eppendorf MiniSpin centrifuge</td><td>Fisher Scientific</td><td>05-401-09</td><td></td></tr></tbody></table>

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.

When SiO2 beads are mixed with a solution of vesicles composed of phospholipids, fluorescent lipids and other lipids such as gangliosides, the vesicles rupture on the SiO2 bead surfaces to form SSLBs, as shown schematically in Figure 1a. After washing the SSLBs, a drop of SSLB solution is placed on a microwell array, and the beads are allowed to settle to the surface. (Figure 1b1) This can also be done with a suspension of phospholipid vesicles or natural membrane particles, which are shown in Figure 1b2. A fluorescence image of phospholipid vesicles adsorbed to a microwell array substrate is shown in Figure 1b3. Then the PDMS squeegee is made to slide gently across the surface to remove any SSLBs (Figure 1c1), vesicles or natural membrane particles (Figure 1c2) that did not settle into microwells. This results in a SSLB array in which each microwell contains at most one SSLB (Figure 1d1) or an array where multiple vesicles or natural membrane particles are in each microwell (Figure 1d2). An image of a microarray composed of fluorescently labeled vesicles is shown in Figure 1d3.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51501/51501fig1highres.jpg" width="500" /> 
Figure 1. Formation of SSLBs and SSLB array assembly. (a) Illustration of a SSLB composed of 3 types of lipids on an SiO2 bead: egg PC (black), Rho-DPPE (red) and GM1 (blue). (b1-d1) Process flow for assembly of a SSLB array showing SSLBs scattered on a microwell array (b1), use of a PDMS squeegee to clear the surface of SSLBs not settled into microwells (c1), and the final SSLB array (d1). (b2-d2) The same process illustrated for the case of phospholipid vesicles or natural membrane particles where more than one vesicle or particle can be accommodated in each microwell. (b3) A fluorescence image of phospholipid vesicles adsorbed onto a microwell substrate corresponding to the illustration in (b2). (d3) A fluorescence image of a microarray composed of phospholipid vesicles corresponding to the illustration in (d2). Adapted from references 20 and 21 and used with permission of the American Chemical Society.
Individual SiO2 beads supports for SSLBs in microwells were imaged with scanning electron microscopy (SEM) from both the angled top-view and cross-sectional perspectives. A top view of an area approximately 15 &#181;m x 15 &#181;m is shown in Figure 2a, while Figure 2b shows a cross-sectional view of a single bead immobilized in a microwell. The lighter layer coating the microwell is 100 nm of Al2O3 deposited by atomic layer deposition to tune the well diameter so that the wells are capable of accommodating only single beads.
<img alt="Figure 2" fo:content-width="3in" src="/files/ftp_upload/51501/51501fig2highres.jpg" width="300" /> 
Figure 2. Scanning electron microscopy images of SSLB arrays. (a) An electron micrograph of a 15 mm x 15 mm area showing SSLB bead supports immobilized in microwell arrays. (b) A single 700 nm diameter SSLB in a microwell. The microwell coating is Al2O3, which is deposited to shrink the microwell so that only individual beads can fit inside. Adapted from reference 20 and used with permission of the American Chemical Society.
Once the SSLB arrays are prepared, they can be imaged by fluorescence microscopy. Figure 3a shows a 100 &#181;m &#215; 100 &#181;m area of an array with 936 SSLBs. To determine the percent of occupancy of an array, the fluorescence image can be compared with a brightfield image of the same area. Figure 3b shows an area with 550 microwells and 416 SSLBs for an occupancy rate of 76%.
Sequential deposition of different types of SSLBs was used to create arrays with more than one type of SSLB, where the identity of individual SSLBs was determined by the absence or presence of a toxin receptor (GM1) and its binding of cholera toxin (CTx). The assembly process was the same as for a single SSLB type but was carried out a second time with a different SSLB identity. The starting concentration of SSLBs was 10x lower to reduce occupancy of the first type of SSLB and to allow more vacancies to be occupied by the second type of SSLB. In the Figures 3c-3e, an SSLB array with two types of SSLBs is shown. All of the SSLBs contain red fluorescent Rho-DPPE (Figure 3c), but only some of the SSLBs contain GM1, a ganglioside that is the receptor for the B-subunit of CTx. When exposed to green fluorescent Alexa 488-conjugated CTx, only the SSLBs containing GM1 fluoresce in the green channel (Figure 3d). When the red and green images are merged, it is possible to determine which SSLBs contain GM1 (yellow) and which do not (red) (Figure 3e).
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51501/51501fig3highres.jpg" width="500" /> 
Figure 3. Fluorescence imaging of SSLB arrays. (a) A 100 &#181;m x 100 &#181;m area containing 936 Rho-DPPE-labeled egg PC SSLBs. (b) A brightfield and fluorescence overlay showing a SSLB with 76% of the microwells occupied. (c-e) A SSLB array containing SSLBs that contain Rho-DPPE (c), a fraction of which also contain GM1, which is bound by Alexa-488-conjugated CTx (d). (e) Merger of red and green channels where colocalized fluorescence indicates the presence of GM1 on the SSLB. The blue circles indicate SSLBs that only fluoresce red, i.e. lacking GM1. Adapted from reference 20 and used with permission of the American Chemical Society.
Fluorescence data from SSLB arrays was analyzed by measuring the intensity of individual SSLBs in an image. The SSLB intensity data from arrays were compiled into histograms. Figure 4a shows one such histogram from the analysis of Rho-DPPE fluorescence from a SSLB array composed solely of SSLBs containing egg PC and 1 mol% Rho-DPPE. The data in Figure 4a were acquired by analyzing the image in Figure 3a. We found the SSLB arrays to be robust and stable for at least one week when refrigerated and submerged in buffer. The arrays did not lose any fluorescence intensity, nor did the occupancy of the arrays change over the course of one week (Figure 4b).
<img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/51501/51501fig4highres.jpg" width="600" /> 
Figure 4. (a) Histogram showing the distribution of fluorescence intensities for the SSLB array shown in Figure 3a. The mean intensity for each SSLB in the array was used to compile the histogram. The solid line is a Gaussian fit to the data. (b) Mean array occupancy (black circles) and fluorescence intensity (red squares) for a SSLB array monitored over the course of one week. Adapted from reference 20 and used with permission of the American Chemical Society.
We made arrays composed of SSLBs with varying concentrations of GM1 (0, 0.5, 1 and 2 mol%) and exposed them to a fixed concentration of CTx as well as arrays with SSLBs with concentrations of GM1 and exposed them to varying concentrations of CTx. The later experiment was used to determine the equilibrium dissociation constant (Kd) for GM1/CTx binding. The SSLB arrays with varying GM1 concentration were exposed to 50 nM Alexa-488 labeled CTx for 1 hr then washed and imaged. The intensity histograms for SSLB arrays with 0.5 &#8211; 2 mol% GM1 exposed to 50 nM Alexa 488-conjugated CTx can be seen in Figure 5a. Figure 5b shows the linear dependence of mean fluorescence intensity upon GM1 concentration. When the concentration of GM1 in SSLBs is fixed at 2 mol% and the SSLB arrays are exposed to CTx ranging from 16 pM to 158 nM, the binding response follows sigmoidal curves as shown in Figure 5c. The curves fit to this data are based upon two different binding models: the Langmuir isotherm (Eq. 1) and the Hill-Waud model (Eq. 2). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51501/51501fig5highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Equation 1" fo:content-width="1in" src="/files/ftp_upload/51501/51501eq1highres.jpg" width="100" /> (1)
<img alt="Equation 2" fo:content-width="1.2in" src="/files/ftp_upload/51501/51501eq2highres.jpg" width="120" /> (2)
Where F is the fluorescence intensity at a given CTx concentration, Fmax is the fluorescence intensity when binding is saturated, [CTx] is the CTx concentration, KD and KH are the dissociation constants from the Langmuir and Hill-Waud fits, respectively, and n is cooperativity coefficient in the Hill-Waud model. The cooperativity coefficient (n) estimates the amount of multivalent binding, where an n greater than one indicates that each CTx molecule binds more than one GM1. The concentration of CTx at 0.5 x Fmax is the KD or KH for the GM1/CTx interaction. In our experiments we calculated a KD of 1.6 &#177; 0.2 nM and KH of 1.4 &#177; 0.2 nM with an n value of 1.3, indicating cooperative binding. The dissociation constants for the GM1/CTx interaction vary widely in the literature, ranging over 5 orders of magnitude from 4.5 pM to 370 nM26,27.
<img alt="Figure 5" fo:content-width="6in" src="/files/ftp_upload/51501/51501fig5highres.jpg" width="600" /> 
Figure 5. CTx/GM1 binding assays on SSLB arrays. (a) Histograms of the fluorescence intensity of individual SSLBs within SSLB arrays. The arrays contained SSLBs with 0.5, 1 or 2 mol% GM1 and were exposed to 50 nM Alexa 488-conjugated CTx. The solid lines are Gaussian fits. (b) Plot of the distribution means from (a) as a function of GM1 concentration. (c) Binding curves showing the distribution means from arrays containing SSLBs with 2 mol% GM1 after exposure to Alexa-488-conjugated CTx varying from 16 pM to 158 nM. The lines are fits to the Langmuir isotherm (red) and Hill-Waud (blue) binding models. The error bars in (b-c) represent the standard deviations of the distributions of fluorescence intensity corresponding to each data point. Adapted from reference 20 and used with permission of the American Chemical Society.
As mentioned earlier, it is also possible to form arrays with biomembrane material derived from natural sources21. The arrays are formed the same way, by dispersing membrane particles on a microwell substrate, then cleaning the top surface with the squeegee to leave behind membrane particles in the microwells. Figure 6 shows examples of myelin and neuronal lipid raft microarrays. In Figure 6a, myelin particles are labeled with the lipophilic fluorophore FM1-43. Lipid rafts are known to be enriched in cholesterol and gangliosides such as GM1; thus, Alexa 488-conjugated CTx binds strongly to lipid raft microarrays as shown in Figure 6b, while fluorescently labeled streptavidin (SAPE) does not bind to the array. (Figure 6c) We also created stripe arrays by delivering myelin and lipid raft particles to the microwell substrate with a microfluidic chip with 250 &#181;m-wide channels. After particle delivery, the microfluidic chip was detached from the microwell substrate and the squeegee process carried out as usual. The stripe arrays were then exposed to a fluorescent anti-oligodendrocyte IgM antibody (IgM O4), which binds sulfatide found in myelin. (Figure 7) The myelin and lipid raft stripe microarrays in Figure 7 are shown at increasing levels of magnification, and at the highest level of magnification, it is readily apparent that IgM O4 only binds to the myelin microarrays because sulfatide is absent from the lipid rafts.
<img alt="Figure 6" fo:content-width="5in" src="/files/ftp_upload/51501/51501fig6highres.jpg" width="500" /> 
Figure 6. Myelin particle and neuronal lipid raft particle arrays. (a) A myelin particle microarray where the myelin particles are rendered fluorescent by the lipophilic fluorophore FM1-43. The dashed line indicates the edge of the array area. (b) Lipid raft microarray showing CTx binding to the lipid raft particles. (c) Lipid raft microarray exposed to fluorescent streptavidin-phycoerythrin (SAPE) showing little to no nonspecific binding. Adapted from reference 21 and used with permission of the American Chemical Society.
<img alt="Figure 7" fo:content-width="5in" src="/files/ftp_upload/51501/51501fig7highres.jpg" width="500" /> 
Figure 7. Multicomponent arrays formed by microfluidic delivery of myelin and neuronal raft membranes. (a) Fluorescence image after myelin and rafts were deposited via microfluidic channels on the microarray substrate. The left and right stripes contain myelin and the middle stripe contains neuronal raft membranes. The membranes are stained with FM1-43, which labels all lipid material. The scale bar is 250 &#956;m. (b) Fluorescence image of the same three stripes after applying the PDMS squeegee and incubating with IgM O4. The scale bar is 250 &#956;m. (c-e) Magnified images from the three stripes showing that IgM O4 only binds to the myelin microarrays: (c) left stripe and (e) right stripe. The scale bars in panels c-e are 30 &#956;m. Adapted from reference 21 and used with permission of the American Chemical Society.

Watch this Scientific Journal Video about Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method at JoVE.com

Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method

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 ...

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Bioengineering

13.6K Views.  University of Minnesota. 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.

Video: Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method

A Guided Materials Screening Approach for Developing Quantitative Sol-gel Derived Protein Microarrays

Microarrays have found use in the development of high-throughput assays for new materials and discovery of small-molecule drug leads. Herein we describe a guided material screening approach to identify sol-gel based materials that are suitable for producing three-dimensional protein microarrays. The approach first identifies materials that can be printed as microarrays, narrows down the number of materials by identifying those that are compatible with a given enzyme assay, and then hones in on optimal materials based on retention of maximum enzyme activity. This approach is applied to develop microarrays suitable for two different enzyme assays, one using acetylcholinesterase and the other using a set of four key kinases involved in cancer. In each case, it was possible to produce microarrays that could be used for quantitative small-molecule screening assays and production of dose-dependent inhibitor response curves. Importantly, the ability to screen many materials produced information on the types of materials that best suited both microarray production and retention of enzyme activity. The materials data provide insight into basic material requirements necessary for tailoring optimal, high-density sol-gel derived microarrays.

A guided material screening approach to develop sol-gel derived protein doped microarrays using an emerging pin-printing method of fabrication is described. This methodology is demonstrated through the development of acetylcholinesterase and multikinase microarrays, which are used for cost-effective small-molecule screening.

Microarrays have found use in the development of high-throughput assays for new materials and discovery of small-molecule drug leads. Herein we describe a ...

Chemistry

Microfabrication of Chip-sized Scaffolds for Three-dimensional Cell cultivation

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell cultivation. These technologies offer a wide spectrum of advantages not only for manufacturing but also for different applications. The size and shape of formed cell clusters can be influenced by the exact and reproducible architecture of the microfabricated scaffold and, therefore, the diffusion path length of nutrients and gases can be controlled.1 This is unquestionably a useful tool to prevent apoptosis and necrosis of cells due to an insufficient nutrient and gas supply or removal of cellular metabolites.
Our polymer chip, called CellChip, has the outer dimensions of 2 x 2 cm with a central microstructured area. This area is subdivided into an array of up to 1156 microcontainers with a typical dimension of 300 m edge length for the cubic design (cp- or cf-chip) or of 300 m diameter and depth for the round design (r-chip).2
So far, hot embossing or micro injection moulding (in combination with subsequent laborious machining of the parts) was used for the fabrication of the microstructured chips. Basically, micro injection moulding is one of the only polymer based replication techniques that, up to now, is capable for mass production of polymer microstructures.3 However, both techniques have certain unwanted limitations due to the processing of a viscous polymer melt with the generation of very thin walls or integrated through holes. In case of the CellChip, thin bottom layers are necessary to perforate the polymer and provide small pores of defined size to supply cells with culture medium e.g. by microfluidic perfusion of the containers.
In order to overcome these limitations and to reduce the manufacturing costs we have developed a new microtechnical approach on the basis of a down-scaled thermoforming process. For the manufacturing of highly porous and thin walled polymer chips, we use a combination of heavy ion irradiation, microthermoforming and track etching. In this so called "SMART" process (Substrate Modification And Replication by Thermoforming) thin polymer films are irradiated with energetic heavy projectiles of several hundred MeV introducing so-called "latent tracks" Subsequently, the film in a rubber elastic state is formed into three dimensional parts without modifying or annealing the tracks. After the forming process, selective chemical etching finally converts the tracks into cylindrical pores of adjustable diameter.

We present two processes for the microfabrication of porous polymer chips for three-dimensional cell cultivation. The first one is hot embossing combined with a solvent vapour welding process. The second one uses a recently developed microthermoforming process combined with ion track technology leading to a significant simplification of manufacture.

Using microfabrication technologies is a prerequisite to create scaffolds of reproducible geometry and constant quality for three-dimensional cell ...

Biology

Antifouling Self-assembled Monolayers on Microelectrodes for Patterning Biomolecules

We present a procedure for forming a poly(ethylene glycol) (PEG) trimethoxysilane self-assembled monolayer (SAM) on a silicon substrate with gold microelectrodes. The PEG-SAM is formed in a single assembly step and prevents biofouling on silicon and gold surfaces. The SAM is used to coat microelectrodes patterned with standard, positive-tone lithography. Using the microtubule as an example, we apply a DC voltage to induce electrophoretic migration to the SAM-coated electrode in a reversible manner. A flow chamber is used for imaging the electrophoretic migration and microtubule patterning in situ using epifluorescence microscopy. This method is generally applicable to biomolecule patterning, as it employs electrophoresis to immobilize target molecules and thus does not require specific molecular interactions. Further, it avoids problems encountered when attempting to pattern the SAM molecules directly using lithographic techniques. The compatibility with electron beam lithography allows this method to be used to pattern biomolecules at the nanoscale.

We present a procedure for forming a poly(ethylene glycol) self-assembled monolayer (PEG-SAM) on a silicon substrate with gold microelectrodes. The PEG-SAM is formed in a single step and prevents biofouling on silicon and gold surfaces. Electrophoresis is then used for patterning biomolecules down to the nanoscale.

We present a procedure for forming a poly(ethylene glycol) (PEG) trimethoxysilane self-assembled monolayer (SAM) on a silicon substrate with gold ...

Biomembrane Fabrication by the Solvent-assisted Lipid Bilayer (SALB) Method

In order to mimic cell membranes, the supported lipid bilayer (SLB) is an attractive platform which enables in vitro investigation of membrane-related processes while conferring biocompatibility and biofunctionality to solid substrates. The spontaneous adsorption and rupture of phospholipid vesicles is the most commonly used method to form SLBs. However, under physiological conditions, vesicle fusion (VF) is limited to only a subset of lipid compositions and solid supports. Here, we describe a one-step general procedure called the solvent-assisted lipid bilayer (SALB) formation method in order to form SLBs which does not require vesicles. The SALB method involves the deposition of lipid molecules onto a solid surface in the presence of water-miscible organic solvents (e.g., isopropanol) and subsequent solvent-exchange with aqueous buffer solution in order to trigger SLB formation. The continuous solvent exchange step enables application of the method in a flow-through configuration suitable for monitoring bilayer formation and subsequent alterations using a wide range of surface-sensitive biosensors. The SALB method can be used to fabricate SLBs on a wide range of hydrophilic solid surfaces, including those which are intractable to vesicle fusion. In addition, it enables fabrication of SLBs composed of lipid compositions which cannot be prepared using the vesicle fusion method. Herein, we compare results obtained with the SALB and conventional vesicle fusion methods on two illustrative hydrophilic surfaces, silicon dioxide and gold. To optimize the experimental conditions for preparation of high quality bilayers prepared via the SALB method, the effect of various parameters, including the type of organic solvent in the deposition step, the rate of solvent exchange, and the lipid concentration is discussed along with troubleshooting tips. Formation of supported membranes containing high fractions of cholesterol is also demonstrated with the SALB method, highlighting the technical capabilities of the SALB technique for a wide range of membrane configurations.

Biomembrane Fabrication by the Solvent-assisted Lipid Bilayer SALB Method

We present an experimental protocol to form a supported lipid bilayer on solid substrates without using lipid vesicles. We demonstrate a one-step method to form a lipid bilayer on silicon dioxide and gold as well as supported membranes with cholesterol-enriched domain for various biological applications.

In order to mimic cell membranes, the supported lipid bilayer (SLB) is an attractive platform which enables in vitro investigation of membrane-related ...

Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, such as nucleic acids and proteins. Single molecule FRET (sm-FRET) measurements on immobilized molecules permit long observations of the system -effectively until both dyes photobleach- resulting in time-traces that report on biomolecular dynamics with a broad range of timescales from milliseconds to minutes. To facilitate the acquisition of large number of traces for statistical analyses, the process must be automated and the sample environment should be tightly controlled over the entire measurement time (~12 hours). This is accomplished using an automated scanning confocal microscope that allows the interrogation of thousands of single molecules overnight, and a microfluidic cell that permits the controlled exchange of buffer, with restricted oxygen content and maintains a constant temperature throughout the entire measuring period. Here we show how to assemble the microfluidic device and how to activate its surface for DNA immobilization. Then we explain how to prepare a buffer to maximize the photostability and lifetime of the fluorophores. Finally, we show the steps involved in preparing the setup for the automated acquisition of time-resolved single molecule FRET traces of DNA molecules.

In this article we describe how we obtain FRET traces from individual DNA molecules immobilized to a surface using an automated scanning confocal microscope.

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, ...

Patterning of Microorganisms and Microparticles through Sequential Capillarity-assisted Assembly

Controlled patterning of microorganisms into defined spatial arrangements offers unique possibilities for a broad range of biological applications, including studies of microbial physiology and interactions. At the simplest level, accurate spatial patterning of microorganisms would enable reliable, long-term imaging of large numbers of individual cells and transform the ability to quantitatively study distance-dependent microbe-microbe interactions. More uniquely, coupling accurate spatial patterning and full control over environmental conditions, as offered by microfluidic technology, would provide a powerful and versatile platform for single-cell studies in microbial ecology. 
This paper presents a microfluidic platform to produce versatile and user-defined patterns of microorganisms within a microfluidic channel, allowing complete optical access for long-term, high-throughput monitoring. This new microfluidic technology is based on capillarity-assisted particle assembly and exploits the capillary forces arising from the controlled motion of an evaporating suspension inside a microfluidic channel to deposit individual microsized objects in an array of traps microfabricated onto a polydimethylsiloxane (PDMS) substrate. Sequential depositions generate the desired spatial layout of single or multiple types of micro-sized objects, dictated solely by the geometry of the traps and the filling sequence. 
The platform has been calibrated using colloidal particles of different dimensions and materials: it has proven to be a powerful tool to generate diverse colloidal patterns and perform surface functionalization of trapped particles. Furthermore, the platform was tested on microbial cells, using Escherichia coli cells as a model bacterium. Thousands of individual cells were patterned on the surface, and their growth was monitored over time. In this platform, the coupling of single-cell deposition and microfluidic technology allows both geometric patterning of microorganisms and precise control of environmental conditions. It thus opens a window into the physiology of single microbes and the ecology of microbe-microbe interactions, as shown by preliminary experiments.

We present a technology that uses capillarity-assisted assembly in a microfluidic platform to pattern micro-sized objects suspended in a liquid, such as bacteria and colloids, into prescribed arrays on a polydimethylsiloxane substrate.

Controlled patterning of microorganisms into defined spatial arrangements offers unique possibilities for a broad range of biological applications, ...

Engineering

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

Fabrication of a Multiplexed Artificial Cellular MicroEnvironment Array

Cellular microenvironments consist of a variety of cues, such as growth factors, extracellular matrices, and intercellular interactions. These cues are well orchestrated and are crucial in regulating cell functions in a living system. Although a number of researchers have attempted to investigate the correlation between environmental factors and desired cellular functions, much remains unknown. This is largely due to the lack of a proper methodology to mimic such environmental cues in vitro, and simultaneously test different environmental cues on cells. Here, we report an integrated platform of microfluidic channels and a nanofiber array, followed by high-content single-cell analysis, to examine stem cell phenotypes altered by distinct environmental factors. To demonstrate the application of this platform, this study focuses on the phenotypes of self-renewing human pluripotent stem cells (hPSCs). Here, we present the preparation procedures for a nanofiber array and the microfluidic structure in the fabrication of a Multiplexed Artificial Cellular MicroEnvironment (MACME) array. Moreover, overall steps of the single-cell profiling, cell staining with multiple fluorescent markers, multiple fluorescence imaging, and statistical analyses, are described.

This article describes the detailed methodology to prepare a Multiplexed Artificial Cellular MicroEnvironment (MACME) array for high-throughput manipulation of physical and chemical cues mimicking in vivo cellular microenvironments and to identify the optimal cellular environment for human pluripotent stem cells (hPSCs) with single-cell profiling.

Cellular microenvironments consist of a variety of cues, such as growth factors, extracellular matrices, and intercellular interactions. These cues are ...

Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips

A significant number of lead compounds fail in the pharmaceutical pipeline because animal studies often fail to predict clinical responses in human patients. Human Organ-on-a-Chip (Organ Chip) microfluidic cell culture devices, which provide an experimental in vitro platform to assess efficacy, toxicity, and pharmacokinetic (PK) profiles in humans, may be better predictors of therapeutic efficacy and safety in the clinic compared to animal studies. These devices may be used to model the function of virtually any organ type and can be fluidically linked through common endothelium-lined microchannels to perform in vitro studies on human organ-level and whole body-level physiology without having to conduct experiments on people. These Organ Chips consist of two perfused microfluidic channels separated by a permeable elastomeric membrane with organ-specific parenchymal cells on one side and microvascular endothelium on the other, which can be cyclically stretched to provide organ-specific mechanical cues (e.g., breathing motions in lung). This protocol details the fabrication of flexible, dual channel, Organ Chips through casting of parts using 3D printed molds, enabling combination of multiple casting and post-processing steps. Porous poly (dimethyl siloxane) (PDMS) membranes are cast with micrometer sized through-holes using silicon pillar arrays under compression. Fabrication and assembly of Organ Chips involves equipment and steps that can be implemented outside of a traditional cleanroom. This protocol provides researchers with access to Organ Chip technology for in vitro organ- and body-level studies in drug discovery, safety and efficacy testing, as well as mechanistic studies of fundamental biological processes.

Here, we present a protocol that describes the fabrication of stretchable, dual channel, organ chip microfluidic cell culture devices for recapitulating organ-level functionality in vitro.

A significant number of lead compounds fail in the pharmaceutical pipeline because animal studies often fail to predict clinical responses in human ...

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.

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.