The authors wish to acknowledge support from the National Research Foundation (NRF -NRFF2011-01 and NRF2015NRF-POC0001-19), the National Medical Research Council (NMRC/CBRG/0005/2012), and Nanyang Technological University to N.J.C.

1. Formation of Supported Lipid Bilayer on a Hydrophilic Solid Support
<ol>
	<li>Prepare lipid stock solutions of 10 mg/ml 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 1 mg/ml 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (Rh-PE) by dissolving the respective lipid powders (appropriately weighed beforehand with an analytical mass balance) in isopropanol solution.</li>
	<li>Dilute and mix the stock solutions in isopropanol in order to prepare the desired lipid mixture at the final concentration. For fluorescence microscopy and FRAP experiments,0.5 wt% Rh-PE should be included in the lipid mixture.</li>
	<li>Inject the lipid mixture in isopropanol into the microfluidic channel until it is filled.</li>
	<li>Incubate the lipid mixture on the glass surface for about 10 min.</li>
	<li>Gradually replace the lipid solution with water or buffer solution using a peristaltic pump at a very low flow rate (10-50 &#181;l/min). Alternatively, replace the lipid mixture by repeated pipetting.</li>
	<li>Rinse the channel thoroughly with excess buffer in order to remove the residual isopropanol.</li>
</ol>
2. Formation of Cholesterol-enriched Supported Membranes
Note: The solid surface (SiO2) supports vesicle fusion, but the membrane composition (high cholesterol) inhibits vesicle fusion because cholesterol-enriched vesicles have high bending rigidity30.
<ol>
	<li>Prepare a stock solution containing 10 mg/ml DOPC lipid, 10 mg/ml cholesterol, and 1 mg/ml Rh-PE by first dissolving the respective powders in isopropanol.</li>
	<li>Repeat steps 1.2-1.6 using the stock solutions prepared in step 2.1.</li>
</ol>
3. Membrane Fluidity Assay
<ol>
	<li>Immerse the glass slide in sodium dodecyl sulfate (SDS) solution (1%) for 10 min.</li>
	<li>Wash the slides thoroughly with deionized water and rinse with ethanol.</li>
	<li>Blow-dry slides using a gentle stream of nitrogen.</li>
	<li>Expose the slides to oxygen plasma for 30 sec at maximum radiofrequency power in the oxygen plasma chamber.</li>
	<li>Remove the protective film coating of the bottomless commercial microfluidic chamber (Figure 1A) using a tweezer and attach the glass slide onto the sticky side of the chamber (Figure 1B).</li>
	<li>Assemble the connectors and tubing into the inlet and outlet positions of the chamber and place the microfluidic channel on the microscope stage (Figure 1C).</li>
	<li>Form a fluorescently-labeled supported lipid bilayer in the microfluidic channel [repeat step 1 using the desired lipid composition (e.g., 0.5 mg/ml DOPC) including 0.5 wt% Rh-PE in organic solvent].</li>
	<li>Locate the bilayer plane with a 60X oil immersion objective (NA 1.49) in order to capture images.</li>
	<li>Take two pre-bleach images, then photo-bleach a 30 &#181;m wide circular spot with a 532 nm, 100 mW laser beam. Before photobleaching, warm up the laser until its intensity has become stable. Immediately after photobleaching, capture a series of images every 1 sec for 2 min in order to follow the recovery in fluorescence intensity at the bleached spot.</li>
</ol>
4. Cholesterol Quantification Assay
<ol>
	<li>Expose the silicon dioxide-coated quartz crystal sensor chip to oxygen plasma for 30 sec at maximum radiofrequency power in the oxygen plasma chamber. Remove the chip from the chamber and immediately mount the chip in the measurement chamber.</li>
	<li>Before experiment, first acquire the frequency and dissipation of the sensor chip in air in order to ensure proper mounting.
	<ol>
		<li>For the commercial Q-Sense E1 or E4 QCM-D instrument, run the Q-Soft software program and click &#8220;Acquisition&#8221; and select &#8220;Setup Measurement&#8221;.</li>
		<li>In the new window which appears, check the 3, 5, 7, 9, 11 and 13 overtones and then click Find and Run in order to check the resonance spectra. Set the temperature at 24 &#176;C. If one or more resonance frequencies do not agree with the expected values, check the chip mounting and re-perform this step until an agreement is reached.</li>
	</ol>
	</li>
	<li>Start the peristaltic pump and flow buffer solution (10 mM Tris, 150 mM NaCl, pH 7.5) into the measurement chamber at a flow rate of 100 &#181;l/min.</li>
	<li>In the software program, click &#8220;Acquisition&#8221; and select &#8220;Restart Measurement&#8221; in order to record the resonance frequency and energy dissipation signals. Repeat this step until a stable baseline is obtained for the frequency and dissipation shifts. Note: For steps hereafter, there will be additional frequency and dissipation shifts which can be recorded as a function of time.</li>
	<li>Inject isopropanol (without lipid) for 10 min.</li>
	<li>Inject the mixture of DOPC lipid/cholesterol at the desired molar ratio with a total lipid concentration of 0.5 mg/ml in isopropanol for 10 min.</li>
	<li>Inject buffer for 20 min at a flow rate of 100 &#181;l/min.</li>
	<li>Inject a solution of 1 mM methyl-&#946;-cyclodextrin (M&#946;CD) prepared in buffer (flow rate 100 &#181;l/min) until the frequency signal reaches a stable value.</li>
	<li>Measure the relative positive frequency shifts that is caused by the M&#946;CD treatment step and calculate the mass of cholesterol and DOPC lipids by converting the &#916;fcho and &#916;fDOPC values to mass values by using the Sauerbrey equation [&#916;m = -(C/n)&#215;&#916;f, where C=17.7 ng/cm2, n:overtone].</li>
	<li>Calculate the mole fraction of cholesterol in the SLB, taking into account the molecular weights of DOPC lipid (786.1 g/mol) and cholesterol (386.6 g/mol).</li>
</ol>

The authors have no competing financial interests to disclose.

In this work, a solvent-exchange protocol is presented in which lipids in alcohol (isopropanol, ethanol, or n-propanol) are incubated with a solid support and then the alcohol is replaced gradually with an aqueous buffer solution in order to drive a series of phase transitions eventually producing lamellar-phase lipid bilayers24. It is shown that the method enables fabrication of supported lipid bilayers on surfaces such as gold, which is intractable to the vesicle fusion method.
An optimal lipid concentration range (0.1 - 0.5 mg/ml) has been determined for complete bilayer formation in standard experimental formats tested thus far. At lipid concentrations below 0.1 mg/ml, discrete, microscopic patches of bilayers formed. On the other hand, at concentrations above 0.1 mg/ml and lower than 0.5 mg/ml, a complete and uniform bilayer is formed. At lipid concentrations above this range, a fluid bilayer was still formed as verified by FRAP analysis, however, fluorescence microscopy reveals the presence of additional lipid structures on top of the bilayer. Strikingly, the morphology of these additional lipid structures, as determined by QCM-D analysis, depended on the alcohol which was used during the incubation step. In the case of ethanol, the relatively high &#916;f and &#916;D shifts resemble the QCM-D signature obtained for an adsorbed vesicle layer. When isopropanol or n-propanol was instead used, the &#916;f was slightly higher than the value expected for a bilayer (final &#916;f between -30 to -40 Hz), while the &#916;D was appreciably higher. Such QCM-D responses would be expected for extended lipid structures (e.g., worm-like micelles) protruding outward from the membrane surface (as visible by fluorescence microscopy in some cases).
The rate of solvent exchange is another important parameter which can be critical, especially when lower lipid concentrations (e.g., 0.1 mg/ml) are used. Rapid solvent exchange at low lipid concentration can lead to the formation of incomplete bilayers. In the standard measurement chamber used for QCM-D measurements in this paper (Q-Sense E4 measurement chamber), flow rates of around 100 &#181;l/min, were suitable for highly reproducible complete bilayer formation. For flow cells with other geometries and volume, the optimal flow rate may vary and must be empirically determined based on the steps suggested herein.
In addition to form supported lipid bilayers on surfaces which are intractable to vesicle fusion, the SALB can be employed to circumvent the need for lipid vesicles which can rupture, thereby opening the door to the fabrication of supported membranes with complex compositions. As an illustrative example composition, lipid mixtures with a high fraction of cholesterol were examined. Cholesterol is an important component of mammalian cell membranes, and its fraction can approach 45&#8722;50 mol% of the membrane lipid composition (e.g., in erythrocytes). Thus, even a simple model of a lipid bilayer representing a human cell membrane should include cholesterol.
While vesicle fusion could be used to fabricate fluidic lipid bilayers containing only 10-15% cholesterol, the SALB method enables formation of fluidic lipid bilayers containing high fractions of cholesterol (up to 57 mol%, as quantified by QCM-D measurements)36. However, when the level of cholesterol was further elevated (up to 63 mol%), stripe-shape domains were observed37. The co-existing domains were liquid, reminiscent of those observed in the &#946; region in the phase diagram of the cholesterol/phospholipids monolayer at the air-water interface.
Overall, the SALB method is shown to be a simple and efficient approach to form supported lipid bilayers, especially in cases beyond the scope of the conventional vesicle fusion method. Thus far, the QCM-D technique and fluorescence microscopy were mainly used to characterize the supported lipid bilayers formed by the SALB method. Looking forward, a broad range of surface-sensitive analytical measurements techniques, including surface plasmon resonance (SPR)38, atomic force microscopy (AFM)39,40, Fourier-transform infrared spectroscopy41, X-ray42 and neutron reflectivity43, can be used to further characterize and study simple and complex bilayer configurations prepare by the SALB method. These emerging capabilities open the door to a greater number of scientists who can explore artificial cell membranes by taking advantage of a simple and robust experimental protocol.

The solid-supported lipid bilayer1 (SLB) is a versatile platform which preserves the basic characteristics of biomembranes such as the bilayer thickness, two-dimensional lipid diffusivity, and the ability to host membrane-associated biomolecules. Due to the complexity of natural cell membranes, this simple platform has been shown to function as an efficient platform for in vitro studies of membrane-related processes such as raft formation2, protein binding3, virus and virus-like particle binding4,5, and cell signaling6. Formed in close proximity to a solid support, the SLB platform is compatible with a range of surface-sensitive measurements techniques such as total internal reflection microscopy (TIRF), quartz crystal microbalance-dissipation (QCM-D), and impedance spectroscopy.
Several methods have been developed to produce different types of SLBs, including air bubble collapse7 and dip-pen nanolithography8 for submicron-sized lipid spots, spin-coating9 for bilayer stacks and Langmuir&#8722;Blodgett (LB)10 and vesicle fusion (VF)11 for full-spanning, single lipid bilayer coatings. The VF method consists of the adsorption of small unilamellar vesicles to a solid support and subsequent spontaneous rupture and fusion to form a continuous lipid bilayer. However, under physiological conditions, spontaneous vesicle rupture is mainly limited to silicon-based materials such as silicon dioxide, glass, and mica. In addition, vesicle rupture does not occur spontaneously for vesicles of complex lipid compositions such as those containing high fractions of cholesterol or negatively charged lipids. Depending on the system, vesicle rupture may be induced by further tailoring the experimental conditions such as temperature12, solution pH13, and salinity14, osmotic shock15 or pressure16, or addition of divalent ions such as Ca2+ 17 . Alternatively, the membrane-active AH peptide can be introduced in order to destabilize a layer of adsorbed vesicles, leading to vesicle rupture and bilayer formation on a range of surfaces18-22.
Moreover, successful bilayer formation requires preparation of a well-controlled population of small unilamellar vesicles that can be time consuming and difficult to achieve for certain membrane compositions. Therefore, despite its high efficiency in optimal cases (e.g., after extensive freeze-thaw pretreatment of vesicles23), the general application of vesicle fusion is limited by the scope of suitable substrates and membrane compositions .
The solvent-assisted lipid bilayer (SALB) method24-28 is an alternative fabrication technique which does not require lipid vesicles. The method is based on the deposition of lipid molecules onto a solid surface in the presence of a water miscible organic solvent followed by gradual exchange of this solvent with an aqueous buffer solution in order to trigger SLB formation. During the solvent-exchange step, the ternary mixture of lipids, organic solvent, and water undergoes a series phase transitions with increasing water fraction, which leads to the formation of lamellar phase structures in the bulk solution and an SLB on the solid substrate. Importantly, this self-assembly route bypasses the need for vesicle rupture, which is usually the limiting step for the transformation of adsorbed vesicles into an SLB. The protocol is applicable to a wide variety of surfaces including silicon dioxide, aluminum oxide, chrome, indium tin oxide, and gold. In this paper and in the accompanying video, a comparison of lipid deposition by the SALB and vesicle fusion methods is presented. In particular, the influence of experimental parameters, including lipid concentration, flow rate, and the choice of water miscible organic solvent, on the quality of the bilayer formed by the SALB method are discussed. Analytical characterization of the fabricated SLBs is performed by the QCM-D, fluorescence microscopy, and fluorescence recovery after photobleaching (FRAP) techniques. QCM-D monitoring is a surface-sensitive mass measurement technique which, since the pioneering work conducted by Keller and Kasemo29, has been extensively used to quantitatively investigate bilayer formation. Fluorescence microscopy permits inspection of membrane homogeneity as well as the visualization of membrane domains. The FRAP technique is a standard tool to determine the lateral mobility of lipid molecules in an SLB, which is an essential property of fluidic membranes.
The first part of this study involves QCM-D analysis of the SALB and vesicle fusion methods applied to attempt bilayer formation on silicon dioxide and gold. In the second part, the preparation and characterization of supported membranes containing a range of cholesterol concentrations with the SALB method are demonstrated and the results are compared with those obtained by the vesicle fusion method.

<table border="0" cellpadding="0" cellspacing="0" width="1459">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">QCM-D silicon dioxide-coated substrates</td>
			<td style="width:356px;">QSense AB,&#160; Sweden</td>
			<td style="width:217px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">QCM-D gold-coated substrates</td>
			<td style="width:356px;">QSense AB,&#160; Sweden</td>
			<td style="width:217px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Q-Sense E4 module</td>
			<td>QSense AB,&#160; Sweden</td>
			<td style="width:217px;"></td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;">Plasma Cleaner, PDC-32G</td>
			<td>Harrick Plasma, Ithaca, NY</td>
			<td>PDC-001 (115V)&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC)</td>
			<td>Avanti Polar Lipids</td>
			<td>850375P</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (ammonium salt) (Rh-PE)</td>
			<td>Avanti Polar Lipids</td>
			<td>810150P</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">cholesterol</td>
			<td>Avanti Polar Lipids</td>
			<td style="width:217px;">700000P</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Methyl-&#946;-cyclodextrin</td>
			<td style="width:356px;">Sigma&#160;</td>
			<td style="width:217px;">C4555</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isopropanol</td>
			<td style="width:356px;">Sigma&#160;</td>
			<td align="right">673773</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">Ethanol</td>
			<td style="width:356px;">Sigma&#160;</td>
			<td align="right" style="width:217px;">459844</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">n-propanol</td>
			<td style="width:356px;">Sigma&#160;</td>
			<td align="right">279544</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">Sticky-Slide I 0.1 Luer</td>
			<td style="width:356px;">IBIDI</td>
			<td align="right" style="width:217px;">81128</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">Male elbow 1/8&#8221;</td>
			<td style="width:356px;">Cole-Parmer</td>
			<td style="width:217px;">30505-70</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">Silicon tubing 1.6mm ID</td>
			<td style="width:356px;">IBIDI</td>
			<td align="right" style="width:217px;">10842</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">Glass coverslip No. 1.5H, 25 mm x 75 mm</td>
			<td style="width:356px;">IBIDI</td>
			<td align="right" style="width:217px;">10812</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">Reglo Digital M2-2/12 Peristaltic Pump</td>
			<td style="width:356px;">Ismatec</td>
			<td style="width:217px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:719px;">Sodium dodecyl sulfate</td>
			<td style="width:356px;">Sigma&#160;</td>
			<td align="right" style="width:217px;">71725</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Biomembrane Fabrication by the Solvent-assisted Lipid Bilayer SALB Method at JoVE.com

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.

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 ...

biomembrane-fabrication-solvent-assisted-lipid-bilayer-salb

Research

JoVE Journal

Bioengineering

14.5K vues.  Nanyang Technological University.  Nous présentons un protocole expérimental pour former une bicouche lipidique supporté sur des substrats solides, sans utiliser des vésicules lipidiques. Nous démontrons un procédé en une étape pour former une bicouche lipidique de dioxyde de silicium et l'or ainsi que des membranes supportées par domaine enrichi en cholestérol pour diverses applications biologiques. 

Vidéo: Biomembrane Fabrication by the Solvent-assisted Lipid Bilayer SALB Method

Constant Pressure-controlled Extrusion Method for the Preparation of Nano-sized Lipid Vesicles

Liposomes are artificially prepared vesicles consisting of natural and synthetic phospholipids that are widely used as a cell membrane mimicking platform to study protein-protein and protein-lipid interactions3, monitor drug delivery4,5, and encapsulation4. Phospholipids naturally create curved lipid bilayers, distinguishing itself from a micelle.6 Liposomes are traditionally classified by size and number of bilayers, i.e. large unilamellar vesicles (LUVs), small unilamellar vesicles (SUVs) and multilamellar vesicles (MLVs)7. In particular, the preparation of homogeneous liposomes of various sizes is important for studying membrane curvature that plays a vital role in cell signaling, endo- and exocytosis, membrane fusion, and protein trafficking8. Several groups analyze how proteins are used to modulate processes that involve membrane curvature and thus prepare liposomes of diameters &lt;100 - 400 nm to study their behavior on cell functions3. Others focus on liposome-drug encapsulation, studying liposomes as vehicles to carry and deliver a drug of interest9. Drug encapsulation can be achieved as reported during liposome formation9. Our extrusion step should not affect the encapsulated drug for two reasons, i.e. (1) drug encapsulation should be achieved prior to this step and (2) liposomes should retain their natural biophysical stability, securely carrying the drug in the aqueous core. These research goals further suggest the need for an optimized method to design stable sub-micron lipid vesicles.
Nonetheless, the current liposome preparation technologies (sonication10, freeze-and-thaw10, sedimentation) do not allow preparation of liposomes with highly curved surface (i.e. diameter &lt;100 nm) with high consistency and efficiency10,5, which limits the biophysical studies of an emerging field of membrane curvature sensing. Herein, we present a robust preparation method for a variety of biologically relevant liposomes.
Manual extrusion using gas-tight syringes and polycarbonate membranes10,5 is a common practice but heterogeneity is often observed when using pore sizes &lt;100 nm due to due to variability of manual pressure applied. We employed a constant pressure-controlled extrusion apparatus to prepare synthetic liposomes whose diameters range between 30 and 400 nm. Dynamic light scattering (DLS)10, electron microscopy11 and nanoparticle tracking analysis (NTA)12 were used to quantify the liposome sizes as described in our protocol, with commercial polystyrene (PS) beads used as a calibration standard. A near linear correlation was observed between the employed pore sizes and the experimentally determined liposomes, indicating high fidelity of our pressure-controlled liposome preparation method. Further, we have shown that this lipid vesicle preparation method is generally applicable, independent of various liposome sizes. Lastly, we have also demonstrated in a time course study that these prepared liposomes were stable for up to 16 hours. A representative nano-sized liposome preparation protocol is demonstrated below.

This protocol describes an extrusion method for preparing lipid vesicles of sub-micron sizes with a high degree of homogeneity. This method uses a pressure-controlled system with controlled nitrogen flow rates for liposome preparation. The lipid preparation1,2, liposome extrusion, and size characterization will be presented herein.

Liposomes are artificially prepared vesicles consisting of natural and synthetic phospholipids that are widely used as a cell membrane mimicking platform ...

Microfluidic Chip Fabrication and Method to Detect Influenza

Fast and effective diagnostics play an important role in controlling infectious disease by enabling effective patient management and treatment. Here, we present an integrated microfluidic thermoplastic chip with the ability to amplify influenza A virus in patient nasopharyngeal (NP) swabs and aspirates. Upon loading the patient sample, the microfluidic device sequentially carries out on-chip cell lysis, RNA purification and concentration steps within the solid phase extraction (SPE), reverse transcription (RT) and polymerase chain reaction (PCR) in RT-PCR chambers, respectively. End-point detection is performed using an off-chip Bioanalyzer (Agilent Technologies, Santa Clara, CA). For peripherals, we used a single syringe pump to drive reagent and samples, while two thin film heaters were used as the heat sources for the RT and PCR chambers. The chip is designed to be single layer and suitable for high throughput manufacturing to reduce the fabrication time and cost. The microfluidic chip provides a platform to analyze a wide variety of virus and bacteria, limited only by changes in reagent design needed to detect new pathogens of interest.

An integrated microfluidic thermoplastic chip has been developed for use as a molecular diagnostic. The chip performs nucleic acid extraction, reverse transcriptase, and PCR. Methods for fabricating and running the chip are described.

Fast  and effective diagnostics play an important role in controlling infectious  disease by enabling effective patient management and treatment. Here, we ...

Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method

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

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

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

Lipid Bilayer Vesicle Generation Using Microfluidic Jetting

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

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

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

Polymeric Microneedle Array Fabrication by Photolithography

This manuscript describes the fabrication of polymeric microneedle (MN) arrays by photolithography. It involves a simple mold-free process by using a photomask consisting of embedded micro-lenses. Embedded micro-lenses were found to influence MN geometry (sharpness). Robust MN arrays with tip diameters ranging between 41.5 &#181;m &#177; 8.4 &#181;m and 71.6 &#181;m &#177; 13.7 &#181;m, with two different lengths (1,336 &#181;m &#177; 193 &#181;m and 957 &#181;m &#177; 171 &#181;m) were fabricated. These MN arrays may provide potential applications in delivery of low molecular and macromolecular therapeutic agents through skin.

Here, we present a protocol describing a mold-free fabrication process of the polymeric microneedles by photolithography.

This manuscript describes the fabrication of polymeric microneedle (MN) arrays by photolithography. It involves a simple mold-free process by using a ...

Solid Lipid Nanoparticles (SLNs) for Intracellular Targeting Applications

Nanoparticle-based delivery vehicles have shown great promise for intracellular targeting applications, providing a mechanism to specifically alter cellular signaling and gene expression. In a previous investigation, the synthesis of ultra-small solid lipid nanoparticles (SLNs) for topical drug delivery and biomarker detection applications was demonstrated. SLNs are a well-studied example of a nanoparticle delivery system that has emerged as a promising drug delivery vehicle. In this study, SLNs were loaded with a fluorescent dye and used as a model to investigate particle-cell interactions. The phase inversion temperature (PIT) method was used for the synthesis of ultra-small populations of biocompatible nanoparticles. A 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenylphenyltetrazolium bromide (MTT) assay was utilized in order to establish appropriate dosing levels prior to the nanoparticle-cell interaction studies. Furthermore, primary human dermal fibroblasts and mouse dendritic cells were exposed to dye-loaded SLN over time and the interactions with respect to toxicity and particle uptake were characterized using fluorescence microscopy and flow cytometry. This study demonstrated that ultra-small SLNs, as a nanoparticle delivery system, are suitable for intracellular targeting of different cell types.

Solid Lipid Nanoparticles SLNs for Intracellular Targeting Applications

In this study, a method for synthesizing ultra-small populations of biocompatible nanoparticles was described, as well as several in vitro methods by which to assess their cellular interactions.

Nanoparticle-based delivery vehicles have shown great promise for intracellular targeting applications, providing a mechanism to specifically alter ...

Automated Lipid Bilayer Membrane Formation Using a Polydimethylsiloxane Thin Film

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

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

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

Ligand Nano-cluster Arrays in a Supported Lipid Bilayer

Currently there is considerable interest in creating ordered arrays of adhesive protein islands in a sea of passivated surface for cell biological studies. In the past years, it has become increasingly clear that living cells respond, not only to the biochemical nature of the molecules presented to them but also to the way these molecules are presented. Creating protein micro-patterns is therefore now standard in many biology laboratories; nano-patterns are also more accessible. However, in the context of cell-cell interactions, there is a need to pattern not only proteins but also lipid bilayers. Such dual proteo-lipidic patterning has so far not been easily accessible. We offer a facile technique to create protein nano-dots supported on glass and propose a method to backfill the inter-dot space with a supported lipid bilayer (SLB). From photo-bleaching of tracer fluorescent lipids included in the SLB, we demonstrate that the bilayer exhibits considerable in-plane fluidity. Functionalizing the protein dots with fluorescent groups allows us to image them and to show that they are ordered in a regular hexagonal lattice. The typical dot size is about 800 nm and the spacing demonstrated here is 2 microns. These substrates are expected to serve as useful platforms for cell adhesion, migration and mechano-sensing studies.

We present a protocol to functionalize glass with nanometric protein patches surrounded by a fluid lipid bilayer. These substrates are compatible with advanced optical microscopy and are expected to serve as platform for cell adhesion and migration studies.

Currently there is considerable interest in creating ordered arrays of adhesive protein islands in a sea of passivated surface for cell biological ...

A Simple and Scalable Fabrication Method for Organic Electronic Devices on Textiles

Today, wearable electronics devices combine a large variety of functional, stretchable, and flexible technologies. However, in many cases, these devices cannot be worn under everyday conditions. Therefore, textiles are commonly considered the best substrate to accommodate electronic devices in wearable use. In this paper, we describe how to selectively pattern organic electroactive materials on textiles from a solution in an easy and scalable manner. This versatile deposition technique enables the fabrication of wearable organic electronic devices on clothes.

In this paper, we present a protocol to selectively deposit organic materials on textiles, which allows for the direct integration of organic electronic devices with wearables. The fabricated devices can be fully integrated in textiles, respecting their mechanical appearance and enabling sensing capabilities.

Today, wearable electronics devices combine a large variety of functional, stretchable, and flexible technologies. However, in many cases, these devices ...

Fabrication of Decellularized Cartilage-derived Matrix Scaffolds

Osteochondral defects lack sufficient intrinsic repair capacity to regenerate functionally sound bone and cartilage tissue. To this extent, cartilage research has focused on the development of regenerative scaffolds. This article describes the development of scaffolds that are completely derived from natural cartilage extracellular matrix, coming from an equine donor. Potential applications of the scaffolds include producing allografts for cartilage repair, serving as a scaffold for osteochondral tissue engineering, and providing in vitro models to study tissue formation. By decellularizing the tissue, the donor cells are removed, but many of the natural bioactive cues are thought to be retained. The main advantage of using such a natural scaffold in comparison to a synthetically produced scaffold is that no further functionalization of polymers is required to drive osteochondral tissue regeneration. The cartilage-derived matrix scaffolds can be used for bone and cartilage tissue regeneration in both in vivo and in vitro settings.

Decellularized cartilage-derived scaffolds can be used as a scaffold to guide cartilage repair and as a means to regenerate osteochondral tissue. This paper describes the decellularization process in detail and provides suggestions to use these scaffolds in in vitro settings.

Osteochondral defects lack sufficient intrinsic repair capacity to regenerate functionally sound bone and cartilage tissue. To this extent, cartilage ...