This work was supported by the Pioneer Research Center Program (NRF-2012-0009575) and National Research Foundation Grants (NRF-2012R1A1B4002413, NRF-2014R1A1A2059341) from the National Research Foundation of Korea. This work was also partially supported by the Inha University Research Grant.

1. Solution Preparation
<ol>
	<li>Preparation of buffer solution:
	<ol>
		<li>To formulate buffer solution, dissolve 1 M KCl (Potassium chloride), 10 mM Tris-HCl (Tris-hydrochloride), and 1 mM EDTA (Ethylenediaminetetraacetic acid) in distilled water and adjust pH to 8.0.</li>
		<li>Filter the solution using a 0.20 &#181;m filter. To sterilize, autoclave the solution at 121 &#176;C for 15 min.</li>
	</ol>
	</li>
	<li>Preparation of lipid solution for pre-painting:
	<ol>
		<li>To formulate the lipid solution for pre-painting, dissolve 3% DPhPC (1,2-diphytanoyl-sn-glycero-3-phosphatidylcholine) lipid (w:v) in a mixture of 2:8 n-decane and hexadecane (v:v). Stir overnight using a rotator.</li>
	</ol>
	</li>
	<li>Preparation of lipid solution for membrane formation:
	<ol>
		<li>To formulate the lipid solution for membrane formation, dissolve 0.1% DPhPC (1, 2-diphytanoyl-sn-glycero-3-phosphatidylcholine) lipid (w:v) in a mixture of 2:8 squalene and hexadecane (v:v). Stir overnight using a rotator.</li>
	</ol>
	</li>
</ol>
2. Formation of a PDMS Thin Film
<ol>
	<li>Mix PDMS and curing agent in a 9:1 (w/w) ratio in a mixing cup to form the PDMS prepolymer. Add 5 g of PDMS prepolymer to a Petri dish to form the PDMS thin film (thickness 200 - 250 &#181;m). Spread PDMS pre-polymer using a spin coater at 800 rpm for 10 sec to form a thin film.</li>
	<li>Place the Petri dish into a vacuum desiccator at a pressure of 100 mTorr for 2 hr to remove air bubbles. To polymerize the pre-polymer thin film, bake in an oven for 5 hr at 70 &#176;C.</li>
	<li>In order to make a square PDMS thin film, cut the polymerized PDMS thin film into 2 x 2 cm2 squares. Use a 500 &#181;m micro punch to make an aperture in the center of the PDMS thin film. Pre-paint apertures with 3% DPhPC lipid solution mixed in 2:8 n-decane and hexadecane.</li>
</ol>
3. Chamber Fabrication and Assembly
<ol>
	<li>To fabricate the BLM chamber, design two symmetric blocks of the chamber using 3D drawing software with outer dimensions of 4 cm x 1.5 cm x 1 cm and inner-well dimensions of 1.5 cm x 1.3 cm x 0.8 cm17.</li>
	<li>Craft the chamber using a PTFE block with a CNC machine and follow the manufacturer&#39;s instructions.</li>
</ol>
4. Chamber Assembly
<ol>
	<li>To assemble the chamber, place the pre-painted-PDMS thin film between the two PTFE blocks such that the aperture on the PDMS thin film is aligned with the hole in the chamber.</li>
	<li>Seal the outer edges of the chamber using a cover glass with grease (facilitating optical observation). Immobilize the assembled chamber using nuts and bolts. 
	NOTE: Make sure the chamber is well-sealed so that there is no liquid leakage.</li>
</ol>
5. Formation of Membrane Precursor with Expedited Self-assembly Formation (MPES)
<ol>
	<li>Using a pipette, deposit 0.5 &#181;l of 0.1% DPhPC lipid mixed in 2:8 n-decane:hexadecane onto the aperture of the PDMS thin film assembled with the chamber.</li>
	<li>Prior to use, store the chamber in a freezer or a refrigerator below 10 &#176;C.</li>
</ol>
6. Membrane Formation and Verification
<ol>
	<li>To form a BLM with MPES, withdraw the chamber from the refrigerator and suspend 2 ml of buffer solution in each side of the chamber. Set the chamber aside for &#60; 10 min until the frozen membrane precursor thaws.</li>
	<li>Place the chamber onto a micromanipulator to precisely control the elevation with respect to the light source and the microscope. Illuminate one side of the chamber as a light source using a halogen fiber optic illuminator to brighten the aperture of the PDMS thin film for optical observation of BLM formation process.</li>
	<li>On the other side, place a digital microscope vertically with respect to the light source to observe BLM formation (magnify by 200X).</li>
	<li>To confirm BLM formation, observe the center of the aperture where the color becomes brighter than the annulus.</li>
</ol>
7. Electrical Recording
<ol>
	<li>For electrical measurement, prepare Ag/Cl electrodes using a 208 &#181;m-thick silver wire and bleach in sodium hypochlorite for &#62; 1 min. Place the Ag/Cl electrodes into each side of the chamber deep enough to be dipped into the buffer solution.</li>
	<li>Connect the electrodes to the microelectrode amplifier. Using electrophysiology software, apply a &#177;10 mV triangular waveform across the membrane to acquire a square wave. Set applying voltage by clicking the arrows indicated on V_clamp (mV).</li>
	<li>Record the electrical properties of the membrane by clicking the record button (red dot icon). Proceed with recording until a uniform square wave is observed. Quit the recording by clicking the black square icon.</li>
</ol>
8. Ion Channel Incorporation
NOTE: Gramicidin A (gA) incorporation occurs spontaneously upon formation of BLM, as gA is added directly to the lipid solution.
<ol>
	<li>To observe gA channel activities, apply 100 mV across the membrane at a sample rate of 5 kHz to measure holding potential of the membrane. Set applying voltage by clicking the arrows indicated on V_clamp (mV).</li>
	<li>Record the electrical properties of the gA incorporation by clicking the record button (red dot icon). Proceed the recording until current jumps is observed. Quit the recording by clicking the black square icon.</li>
	<li>After electrical data acquisition, filter the data with a low-pass Bessel filter at 100 Hz using an electrophysiology software.</li>
	<li>Observe current jumps in the filtered holding potential data (each current jump, ~0.15 nS, represents dimerization of a gA ion channel) to verify gA incorporation.</li>
</ol>

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The authors have nothing to disclose.

Our BLM formation technique provides a powerful tool for cell membrane and ion channel studies, in contrast to conventional techniques that have limited potential for industrial use. We developed a membrane precursor using a PDMS thin film, and devised a frozen membrane precursor with expedited self-assembly.
As opposed to conventional membrane formation methods with hydrophobic films, where membrane formation only occurs via surface interactions between the film and the lipid solution,20...

Artificial lipid bilayer membrane, or black lipid membrane (BLM), is an important tool for elucidating mechanisms of cell membranes and ion channels, as well as for understanding interactions between ion channels and ions/molecules.1-7 Although the patch-clamp method is often considered the gold standard for cell membrane studies, it is laborious and requires highly skilled operators for ion channel measurements.8 While artificially reconstituted lipid bilayer membranes have emerged as alternative tools for ion channel studies,9,10 they are also associated with laborious processes and specific expertise. Moreover, membranes are susceptible to mechanical perturbations. Hence, lipid bilayer technologies introduced to date have limited practical applications.11
In order to enhance robustness and longevity of lipid bilayer membranes, Costello et al.12, and Ide and Yanagida13 have devised a free-standing lipid bilayer supported by hydrogels. Despite enhanced longevity however (&#60; 24 hr), bilayer robustness was not improved. Jeon et al.14 devised a hydrogel encapsulated membrane (HEM) with intimate hydrogel-lipid bilayer contact, resulting in enhanced longevity (up to several days). To further increase the lifetime of the HEM, Malmstadt and Jeon et al. created a hydrogel-encapsulated membrane with hydrogel-lipid binding via in-situ covalent conjugation (cgHEM).15 In both systems, membrane lifetimes increased substantially (&#62; 10 days). However, the membrane formation systems were not sufficiently robust, and could not be stored or delivered where required to liberate expertise for use of the lipid bilayers.
The development of a lipid bilayer platform has primarily revolved around increasing robustness and longevity of BLMs. Although the longevity of BLMs has been substantially enhanced recently, their applications have been limited due to a lack of transportability and storability. To overcome these issues, Jeon et al. created a storable membrane system and introduced a membrane precursor (MP).16 To construct an MP, they prepared a mixture of n-decane and hexadecane containing 3% DPhPC (1,2-diphytanoyl-sn-glycero-3-phosphatidylcholine) to control the freezing point of the lipid solution such that it would freeze at ~14 &#176;C (below room temperature, above typical refrigerator temperature). In this experiment, the MP was spread over a small aperture on a polytetrafluoroethylene (PTFE) film and subsequently frozen in a refrigerator at 4 &#176;C. When the MP was brought to room temperature, the MP thawed and a lipid bilayer was automatically formed, eliminating the expertise typically associated with membrane formation. However, the success rate of BLM made from the MP was as low as ~27%, and membrane formation time was inconsistent (30 min to 24 hr), limiting its practical applications.
In this study, a polydimethylsiloxane (PDMS) thin film is used instead of a conventional hydrophobic thin films (PTFE, polyoxymethylene, polystyrene) to (a) control fabrication time and (b) increase the success rate of BLM formation as previously reported by Ryu et al.17 Herein, membrane formation was facilitated by extraction of solvents due to the porous nature of PDMS, and the time required for membrane formation was successfully controlled in this study. In this system, as the lipid solution was absorbed into the PDMS thin film, a consistent membrane formation time was achieved. Moreover, membrane lifetime was prolonged due to slow absorption of solvents into the PDMS thin film, a result of the addition of squalene to the lipid solution. We conducted optical and electrical measurements to verify that membranes formed using this technique are suitable for ion channels studies.

<table border="0" cellpadding="0" cellspacing="0" width="1448">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Potassium Chloride</td>
			<td style="width:172px;">Sigma-Aldrich</td>
			<td style="width:199px;">P9333</td>
			<td style="width:743px;">For buffer solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Tris-hydrochloride</td>
			<td style="width:172px;">Sigma-Aldrich</td>
			<td style="width:199px;">1185-53-1</td>
			<td>For buffer solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Ethylenediaminetetraacetic acid</td>
			<td style="width:172px;">Sigma-Aldrich</td>
			<td style="width:199px;">60-00-4</td>
			<td>For buffer solution</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:335px;">n-decane</td>
			<td style="width:172px;">Sigma-Aldrich</td>
			<td style="width:199px;">44074-U</td>
			<td>For lipid solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Hexadecane</td>
			<td style="width:172px;">Sigma-Aldrich</td>
			<td style="width:199px;">544-76-3</td>
			<td>For lipid solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Squalene</td>
			<td style="width:172px;">Sigma-Aldrich</td>
			<td style="width:199px;">S3626</td>
			<td>For lipid solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Gramicidin A</td>
			<td style="width:172px;">Sigma-Aldrich</td>
			<td style="width:199px;">11029-61-1</td>
			<td>Membrane protein</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">1,2-diphytanoyl-sn-glycero-3-phosphocholine</td>
			<td style="width:172px;">Avanti Polar Lipids, Inc.</td>
			<td style="width:199px;">850356</td>
			<td>For membrae formation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Sylgard 184a and 184b elastromer kit</td>
			<td style="width:172px;">Dow Corning Asia</td>
			<td style="width:199px;"></td>
			<td>To produce PDMS thin film</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">0.2 &#956;m filter</td>
			<td style="width:172px;">Satorius stedim</td>
			<td style="width:199px;">16534----------K</td>
			<td>To filter buffer solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Rotator</td>
			<td style="width:172px;">FinePCR</td>
			<td style="width:199px;">AG</td>
			<td>To dissolve lipid homogeneously</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Autoclave</td>
			<td style="width:172px;">Biofree</td>
			<td style="width:199px;">BF-60AC</td>
			<td>To sterilize buffer solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Spin coater</td>
			<td style="width:172px;">Shinu Mst</td>
			<td style="width:199px;">SP-60P</td>
			<td>To spread PDMS prepolymer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Vaccum dessiccator</td>
			<td style="width:172px;">Welch</td>
			<td style="width:199px;">2042-22</td>
			<td>To remove air bubble in PDMS prepolymer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">500 &#956;m&#160; punch</td>
			<td style="width:172px;">Harris Uni-Core</td>
			<td style="width:199px;">0.5</td>
			<td>To create an aperture on the PDMS thin film</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">CNC machine</td>
			<td style="width:172px;">SME trading</td>
			<td style="width:199px;">SME 2518</td>
			<td>To fabricate membrane formation chamber</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Halogen fiber optic illuminator</td>
			<td style="width:172px;">Motic</td>
			<td style="width:199px;">MLC-150C</td>
			<td>To illuminate the aperture of PDMS thin film for optical observation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Digital microscope</td>
			<td style="width:172px;">Digital blue</td>
			<td style="width:199px;">QX-5</td>
			<td>To optically observe lipid bilayer membrane formation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Electrode</td>
			<td style="width:172px;">A-M Systems</td>
			<td style="width:199px;"></td>
			<td>To electrically observe membrane formation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:335px;">Microelectrode amplifier (Axopatch amplifier)</td>
			<td style="width:172px;">Axon Instruments</td>
			<td>Axopatch 200B Amplifier</td>
			<td>To measure capacitance of the membrane (described as microelectrode amplifier in the manuscript)</td>
		</tr>
	</tbody>
</table>

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.

Optimization of MPES Solution Composition 
Different compositions of lipids and solvents were tested to successfully reconstitute lipid bilayer membranes from MPES. The MP system with a mixture of n-decane and hexadecane containing 3% DPhPC14 exhibited a low success rate of membrane formation (~27%). In addition, as the PDMS film continuously extracted lipid solution, it was necessary to optimize solvent composition to maintain an intact lipid bil...

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Automated Lipid Bilayer Membrane Formation Using a Polydimethylsiloxane Thin Film

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 ...

automated-lipid-bilayer-membrane-formation-using-polydimethylsiloxane

Research

JoVE Journal

Bioengineering

18.0K Views.  Inha University. 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.

Video: Automated Lipid Bilayer Membrane Formation Using a Polydimethylsiloxane Thin Film

A Chitosan Based, Laser Activated Thin Film Surgical Adhesive, 'SurgiLux': Preparation and Demonstration

Sutures are a 4,000 year old technology that remain the 'gold-standard' for wound closure by virtue of their repair strength (~100 KPa). However, sutures can act as a nidus for infection and in many procedures are unable to effect wound repair or interfere with functional tissue regeneration.1 Surgical glues and adhesives, such as those based on fibrin and cyanoacrylates, have been developed as alternatives to sutures for the repair of such wounds. However, current commercial adhesives also have significant disadvantages, ranging from viral and prion transfer and a lack of repair strength as with the fibrin glues, to tissue toxicity and a lack of biocompatibility for the cyanoacrylate based adhesives. Furthermore, currently available surgical adhesives tend to be gel-based and can have extended curing times which limit their application.2 Similarly, the use of UV lasers to facilitate cross-linking mechanisms in protein-based or albumin 'solders' can lead to DNA damage while laser tissue welding (LTW) predisposes thermal damage to tissues.3 Despite their disadvantages, adhesives and LTW have captured approximately 30% of the wound closure market reported to be in excess of US $5 billion per annum, a significant testament to the need for sutureless technology.4
In the pursuit of sutureless technology we have utilized chitosan as a biomaterial for the development of a flexible, thin film, laser-activated surgical adhesive termed 'SurgiLux'. This novel bioadhesive uses a unique combination of biomaterials and photonics that are FDA approved and successfully used in a variety of biomedical applications and products. SurgiLux overcomes all the disadvantages associated with sutures and current surgical adhesives (see Table 1).
In this presentation we report the relatively simple protocol for the fabrication of SurgiLux and demonstrate its laser activation and tissue weld strength. SurgiLux films adhere to collagenous tissue without chemical modification such as cross-linking and through irradiation using a comparatively low-powered (120 mW) infrared laser instead of UV light. Chitosan films have a natural but weak adhesive attraction to collagen (~3 KPa), laser activation of the chitosan based SurgiLux films emphasizes the strength of this adhesion through polymer chain interactions as a consequence of transient thermal expansion.5 Without this 'activation' process, SurgiLux films are readily removed.6-9 SurgiLux has been tested both in vitro and in vivo on a variety of tissues including nerve, intestine, dura mater and cornea. In all cases it demonstrated good biocompatibility and negligible thermal damage as a consequence of irradiation.6-10

The fabrication of a novel, flexible thin film surgical adhesive from FDA approved ingredients, chitosan and indocyanine green is described. Bonding of this adhesive to collagenous tissue through a simple activation process with a low-powered infra-red laser is demonstrated.

Sutures are a 4,000 year old technology that remain the 'gold-standard' for wound closure by virtue of their repair strength (~100 KPa). However, sutures ...

Creating Transient Cell Membrane Pores Using a Standard Inkjet Printer

Bioprinting has a wide range of applications and significance, including tissue engineering, direct cell application therapies, and biosensor microfabrication.1-10 Recently, thermal inkjet printing has also been used for gene transfection.8,9 The thermal inkjet printing process was shown to temporarily disrupt the cell membranes without affecting cell viability. The transient pores in the membrane can be used to introduce molecules, which would otherwise be too large to pass through the membrane, into the cell cytoplasm.8,9,11
The application being demonstrated here is the use of thermal inkjet printing for the incorporation of fluorescently labeled g-actin monomers into cells. The advantage of using thermal ink-jet printing to inject molecules into cells is that the technique is relatively benign to cells.8, 12 Cell viability after printing has been shown to be similar to standard cell plating methods1,8. In addition, inkjet printing can process thousands of cells in minutes, which is much faster than manual microinjection. The pores created by printing have been shown to close within about two hours. However, there is a limit to the size of the pore created (~10 nm) with this printing technique, which limits the technique to injecting cells with small proteins and/or particles. 8,9,11
A standard HP DeskJet 500 printer was modified to allow for cell printing.3, 5, 8 The cover of the printer was removed and the paper feed mechanism was bypassed using a mechanical lever. A stage was created to allow for placement of microscope slides and coverslips directly under the print head. Ink cartridges were opened, the ink was removed and they were cleaned prior to use with cells. The printing pattern was created using standard drawing software, which then controlled the printer through a simple print command. 3T3 fibroblasts were grown to confluence, trypsinized, and then resuspended into phosphate buffered saline with soluble fluorescently labeled g-actin monomers. The cell suspension was pipetted into the ink cartridge and lines of cells were printed onto glass microscope cover slips. The live cells were imaged using fluorescence microscopy and actin was found throughout the cytoplasm. Incorporation of fluorescent actin into the cell allows for imaging of short-time cytoskeletal dynamics and is useful for a wide range of applications.13-15

A description of the methods used to convert an HP DeskJet 500 printer into a bioprinter. The printer is capable of processing living cells, which causes transient pores in the membrane. These pores can be utilized to incorporate small molecules, including fluorescent G-actin, into the printed cells.

Bioprinting has a wide range of applications and significance, including tissue engineering, direct cell application therapies, and biosensor ...

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 ...

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 ...

Double Emulsion Generation Using a Polydimethylsiloxane (PDMS) Co-axial Flow Focus Device

Double emulsions are useful in a number of biological and industrial applications in which it is important to have an aqueous carrier fluid. This paper presents a polydimethylsiloxane (PDMS) microfluidic device capable of generating water/oil/water double emulsions using a coaxial flow focusing geometry that can be fabricated entirely using soft lithography. Similar to emulsion devices using glass capillaries, double emulsions can be formed in channels with uniform wettability and with dimensions much smaller than the channel sizes. Three dimensional flow focusing geometry is achieved by casting a pair of PDMS slabs using two layer soft lithography, then mating the slabs together in a clamshell configuration. Complementary locking features molded into the PDMS slabs enable the accurate registration of features on each of the slab surfaces. Device testing demonstrates formation of double emulsions from 14 &#181;m to 50 &#181;m in diameter while using large channels that are robust against fouling and clogging.

Double Emulsion Generation Using a Polydimethylsiloxane PDMS Co-axial Flow Focus Device

Microfluidic double emulsions generation typically involves devices with patterned wettability or custom-fabricated glass components. Here we describe the fabrication and testing of an all polydimethylsiloxane (PDMS) double emulsion generator that does not require surface treatment or complicated fabrication processes, and is capable of producing double emulsions down to 14 &#181;m.

Double emulsions are useful in a number of biological and industrial applications in which it is important to have an aqueous carrier fluid. This paper ...

Using Tomoauto: A Protocol for High-throughput Automated Cryo-electron Tomography

Cryo-electron tomography (Cryo-ET) is a powerful three-dimensional (3-D) imaging technique for visualizing macromolecular complexes in their native context at a molecular level. The technique involves initially preserving the sample in its native state by rapidly freezing the specimen in vitreous ice, then collecting a series of micrographs from different angles at high magnification, and finally computationally reconstructing a 3-D density map. The frozen-hydrated specimen is extremely sensitive to the electron beam and so micrographs are collected at very low electron doses to limit the radiation damage. As a result, the raw cryo-tomogram has a very low signal to noise ratio characterized by an intrinsically noisy image. To better visualize subjects of interest, conventional imaging analysis and sub-tomogram averaging in which sub-tomograms of the subject are extracted from the initial tomogram and aligned and averaged are utilized to improve both contrast and resolution. Large datasets of tilt-series are essential to understanding and resolving the complexes at different states, conditions, or mutations as well as obtaining a large enough collection of sub-tomograms for averaging and classification. Collecting and processing this data can be a major obstacle preventing further analysis. Here we describe a high-throughput cryo-ET protocol based on a computer-controlled 300kV cryo-electron microscope, a direct detection device (DDD) camera and a highly effective, semi-automated image-processing pipeline software wrapper library tomoauto developed in-house. This protocol has been effectively utilized to visualize the intact type III secretion system (T3SS) in Shigella flexneri minicells. It can be applicable to any project suitable for cryo-ET.

We present a protocol on how to utilize high-throughput cryo-electron tomography to determine high resolution in situ structures of molecular machines. The protocol permits large amounts of data to be processed, avoids common bottlenecks and reduces resource downtime, allowing the user to focus on important biological questions.

Cryo-electron tomography (Cryo-ET) is a powerful three-dimensional (3-D) imaging technique for visualizing macromolecular complexes in their native ...

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 ...

Spontaneous Formation and Rearrangement of Artificial Lipid Nanotube Networks as a Bottom-Up Model for Endoplasmic Reticulum

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense lipidic nanotubes that are, in terms of morphology and dynamics, reminiscent of ER. The networks are derived from phospholipid double bilayer membrane patches adhering to a transparent Al2O3 substrate. The adhesion is mediated by Ca2+ in the ambient buffer. Subsequent depletion of Ca2+ by means of BAPTA/EDTA causes retraction of the membrane, resulting in spontaneous lipid nanotube network formation. The method only comprises phospholipids and microfabricated surfaces for simple formation of an ER model and does not require the addition of proteins or chemical energy (e.g., GTP or ATP). In contrast to the 3D morphology of the cellular endoplasmic reticulum, the model is two-dimensional (albeit the nanotube dimensions, geometry, structure, and dynamics are maintained). This unique in vitro ER model consists of only a few components, is easy to construct, and can be observed under a light microscope. The resulting structure can be further decorated for additional functionality, such as the addition of ER-associated proteins or particles to study transport phenomena among the tubes. The artificial networks described here are suitable structural models for the cellular ER, whose unique characteristic morphology has been shown to be related to its biological function, whereas details regarding formation of the tubular domain and rearrangements within are still not completely understood. We note that this method uses Al2O3 thin-film-coated microscopy coverslips, which are commercially available but require special orders. Therefore, it is advisable to have access to a microfabrication facility for preparation.

Solid-supported, protein-free, double phospholipid bilayer membranes (DLBM) can be transformed into complex and dynamic lipid nanotube networks and can serve as 2D bottom-up models of the endoplasmic reticulum.

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense ...

Immobilization of Live Caenorhabditis elegans Individuals Using an Ultra-thin Polydimethylsiloxane Microfluidic Chip with Water Retention

Radiation is widely used for biological applications and for ion-beam breeding, and among these methods, microbeam irradiation represents a powerful means of identifying radiosensitive sites in living organisms. This paper describes a series of on-chip immobilization methods developed for the targeted microbeam irradiation of live individuals of Caenorhabditis elegans. Notably, the treatment of the polydimethylsiloxane (PDMS) microfluidic chips that we previously developed to immobilize C. elegans individuals without the need for anesthesia is explained in detail. This chip, referred to as a worm sheet, is resilient to allow the microfluidic channels to be expanded, and the elasticity allows animals to be enveloped gently. Also, owing to the self-adsorption capacity of the PDMS, animals can be sealed in the channels by covering the surface of the worm sheet with a thin cover film, in which animals are not pushed into the channels for enclosure. By turning the cover film over, we can easily collect the animals. Furthermore, the worm sheet shows water retention and allows C. elegans individuals to be subjected to microscopic observation for long periods under live conditions. In addition, the sheet is only 300 &#181;m thick, allowing heavy ions such as carbon ions to pass through the sheet enclosing the animals, thus allowing the ion particles to be detected and the applied radiation dose to be measured accurately. Because selection of the cover films used for enclosing the animals is very important for successful long-term immobilization, we conducted the selection of the suitable cover films and showed a recommended one among some films. As an application example of the chip, we introduced imaging observation of muscular activities of animals enclosing the microfluidic channel of the worm sheet, as well as the microbeam irradiation. These examples indicate that the worm sheets have greatly expanded the possibilities for biological experiments.

Immobilization of Live Caenorhabditis elegans Individuals Using an Ultra-thin Polydimethylsiloxane Microfluidic Chip with Water Retention

A series of immobilization methods has been established to allow the targeted irradiation of live Caenorhabditis elegans individuals using a recently developed ultra-thin polydimethylsiloxane microfluidic chip with water retention. This novel on-chip immobilization is also adequate for imaging observations. The detailed treatment and application examples of the chip are explained.

Radiation is widely used for biological applications and for ion-beam breeding, and among these methods, microbeam irradiation represents a powerful means ...

Material Formation of Recombinant Spider Silks through Aqueous Solvation using Heat and Pressure

Many spiders produce seven types of silks. Six of the silks are fiber in form when produced by the spiders. These fibers are not water soluble. In order to reproduce the remarkable mechanical properties of spider silks, they must be produced in heterologous hosts as spiders are both territorial and cannibalistic. The synthetic analogs of spider silk also tend to be insoluble in aqueous solutions. Thus, a large percentage of research in recombinant spider silks rely upon organic solvents that are detrimental to large scale production of materials. Our group&#8217;s method forces the solvation of these recombinant spider silks into water. Remarkably, when these proteins are prepared using this method of heat and pressure, a wide range of material forms can be prepared from the same solution of recombinant spider silk proteins (rSSp) including: films, fibers, sponge, hydrogel, lyogel, and adhesives. This article demonstrates the production of the solvated rSSp and material forms in a manner that is more easily understood than from written materials and methods alone.

Here, we present a protocol to produce water soluble recombinant spider silk protein solutions and the material forms that can be formed from those solutions.

Many spiders produce seven types of silks. Six of the silks are fiber in form when produced by the spiders. These fibers are not water soluble. In order ...