This work was supported by the Howard Hughes Medical Institute (HHMI) Collaborative Innovation Award. L. A. M. was supported by the Signaling and Cellular Regulation National Institute of Health training grant (T32 GM008759) and the NIH Ruth L. Kirschstein Pre-Doctoral Fellow (CA165349-01). We would like to thank Prof. Michael Stowell (CU Boulder), Prof. Douglas Rees and Prof. Rob Phillips (Caltech) for their invaluable comments.

1. Liposome Preparation
<ol>
	<li>Retrieve a 20 mL glass vial with a Teflon-lined cap.</li>
	<li>Clean all glassware and syringes with chloroform prior to use to prevent contamination.</li>
	<li>Transfer 100 &mu;L of reagent grade chloroform to the glass vial using a 250 &mu;L air-tight glass syringe.</li>
	<li>Add 30 &mu;L of reagent grade methanol to the same glass vial using a 100 &mu;L air-tight glass syringe.</li>
	<li>To prepare a 2 mM 7:1.5:1.5 phosphatidylcholine (POPC): phosphatidylethanolamine (POPE): cholesterol lipid solution, transfer 216 &mu;L of 10 mg/ mL POPC, 42 &mu;L of 10 mg/mL POPE and 20 &mu;L of 11 mg/mL cholesterol solutions in CHCl3 to the 20 mL glass vial.</li>
	<li>Evaporate the organic solvents using slow-flow argon or nitrogen gas until a thin film of lipids is observed on the bottom of the vial.</li>
	<li>Place the uncapped glass vial in a vacuum desiccator for at least 30 minutes to remove residual solvent.</li>
	<li>Transfer 2 mL of buffer, previously passed through a 0.2 micron filter, to the glass vial to hydrate the lipids.</li>
	<li>Incubate the mixture at 4 &deg;C overnight and use within 48 hours.</li>
</ol>
2. Freeze-and-thaw: For Liposome Sizes of 30 - 100 nm Only
<ol>
	<li>Freeze liposome suspension in liquid nitrogen for 15 seconds.</li>
	<li>Thaw liposome suspension using a heating block at 42 &deg;C temperature for ~3 minutes.</li>
	<li>Repeat steps 2.1 and 2.2 for a total of 5 cycles.</li>
</ol>
3. Extrusion
Follow the Avestin instructions to correctly assemble the Liposofast LF-50 extruder, using the instrument outline in their guide book.
<ol>
	<li>Place the large hole support screen (with circular holes) in the support filter base, followed by the circular sintered dish (without holes), one drain disc (25 mm in diameter), and one polycarbonate membrane (25 mm in diameter).</li>
	<li>Place the small, black O-ring on top of the membrane to secure it to the support filter base.</li>
	<li>Attach the top filter extruder by tightening 4 screws in the 4 corresponding holes.</li>
	<li>Assemble the filter extruder unit to the underneath the large extruder barrel. Add the liposome solution to the cylinder barrel. Place the following on top of the extrusion barrel in order: large circular O-ring, narrow cap, large circular O-ring, circular cap.</li>
	<li>Attach the gas regulator to the extruder barrel top and close all valves to prevent air leakage, including the pressure relief valve. Place a 20 mL glass vial or 50 mL Erlenmeyer flask under the extruder filter unit. A safety valve is connected to the regulator and will release if pressure exceeds 600 psi. Turn on the nitrogen gas and open the gas valve connected to the extruder.</li>
	<li>Increase nitrogen pressure to 25 psi for 400 nm liposomes, 125 psi for 100 nm liposomes, and 400-500 psi for 30 nm liposomes.</li>
	<li>Watch the liposome suspension as it ejects into the container while being pushed by the nitrogen. Keep the nitrogen flowing until you no longer observe any liquid passing through the extruder filter into the glass vial.</li>
</ol>
4. Dynamic Light Scattering (DLS) Analysis
<ol>
	<li>Prepare 50 &mu;L of a 20 &mu;M liposome solution.</li>
	<li>Turn on the power source and the lamp source.</li>
	<li>Open the DynaPro software.</li>
	<li>Set the software to the MS/X algorithm model to detect liposomes of 30 nm and 100 nm and MS800 algorithm model to detect &gt;100 nm liposomes.</li>
	<li>Connect to the hardware.</li>
	<li>Pipet 14 &mu;L of liposome sample into the quartz cuvette and insert into the cell holder.</li>
	<li>Press start.</li>
	<li>Press stop after ~20 - 30 acquisitions.</li>
	<li>Analyze the average liposome diameter peaks recorded on the histogram.</li>
</ol>
5. Nanoparticle Tracking Analysis (NTA)
<ol>
	<li>Prepare a 500 &mu;L, 0.1 &mu;M liposome solution.</li>
	<li>Rinse the sample compartment with water and ethanol.</li>
	<li>Dry the sample compartment with a lint-free paper towel.</li>
	<li>Turn on the laser power source and the computer.</li>
	<li>Deliver 300 &mu;L of 0.1 &mu;M liposome solution to the sample compartment.</li>
	<li>Open the Temperature control and the Nanoparticle Tracking Analysis software.</li>
	<li>Press Capture button to turn on the laser.</li>
	<li>Use the horizontal and vertical adjustments to move the stage and adjust the microscope focus.</li>
	<li>Set the desired temperature (20 &deg;C) and recording time (at least 30 seconds).</li>
	<li>Press the Record button to take multiple picture frames of the liposome particles for a specified amount of time. Analyze the peaks corresponding to the diameter liposome sizes on the histogram as it tracks the motion of each particle.</li>
</ol>
6. Representative Results
A scheme outlining the extrusion method is presented in Figure 1. To obtain optimum results, preparation of liposomes with diameters of 30 nm requires a high pressure of ~ 500 psi and diameters of 100 nm requires a pressure of 125 psi to achieve a rapid filter rate. For diameters of 400 nm, a low pressure of ~ 25 psi is recommended to achieve a slower filter rate, which allows the vesicles to elongate and form larger, homogeneous liposomes. We conducted a series of experiments to determine the optimal pressure for producing consistent sub-micron vesicle sizes. We varied the pressure as well as the number of extrusion passes through polycarbonate filters with pore sizes of 30, 100, and 400 nm and discovered appropriate pressure for each desired size. For 30 nm pores, pressure below 500 psi will reduce the flow rate, causing elongation and thus larger vesicle sizes. For 100 nm pores, a steady flow was achieved at 125 psi. For 400 nm pores, low pressure (25 psi) allows the vesicles to elongate into larger vesicle sizes. A slow drop-wise filtration flow is optimal to create larger sub-micron vesicles13.
We performed DLS to determine the liposome sizes extruded through three different diameter sizes, i.e. 30, 100 and 400 nm. DLS is an established method that collects scattered light to determine the particle diameter. We extruded hydrated liposomes of 2 mM through a polycarbonate 30 nm membrane at 500 psi with 5 passes through the filter pore; a 100 nm polycarbonate membrane at 125 psi with 5 passes through the filter pore and a 400 nm polycarbonate membrane at 25 psi with 2 passes through the filter pore. The diameters of the liposomes and measured by DLS for 30, 100 and 400 nm pore sizes were 66 &plusmn; 28, 138 &plusmn;18 and 360 &plusmn; 25 nm (Figure 2). A suspension of 50 nm polystyrene beads was used as a calibration standard as shown in Figure 2, where it recorded a diameter of 47 &plusmn; 16 nm. The percent polydispersity shows that there is no overlap within liposome sizes. It is typical to observe diameters higher than 30 nm when using DLS analysis due to the known bias this instrument has towards larger particles12. Light scatter intensities from large and small particles are collected simultaneously from one detection method and thus harder to resolve liposomes in suspension12. Despite this instrumental limitation, the calibration curve describes a near linear correlation.
NTA is a new technology that measures the size of each particle from direct observations of diffusion in a liquid medium, independent of particle refractive index or density. This high-resolution technique can be used to supplement the measurement of liposomes with DLS. The NTA recorded diameters of 95 &plusmn; 48 and 356 &plusmn; 51 nm, two 100 nm and 400 nm polystyrene solutions were used for calibration. Lipid solutions of 30 nm and 100 nm were observed to have a more comparable diameter relative to the DLS as shown in Figure 3, producing average sizes of 29 &plusmn; 14, 95 &plusmn; 17 and 359 &plusmn; 73 nm. The NTA may be a more general characterization technique to quantify microscopic particles since its sensitivity allows it to measure particles of 50 - 1000 nm. The calibration curve shows a linear correlation between the polycarbonate membrane pores versus the recorded NTA diameter.
<table border="1">
	<tbody>
		<tr>
			<td>Liposome Sizes</td>
			<td>Detection Threshold</td>
			<td>Expected Minimum Particle Size (nm)</td>
		</tr>
		<tr>
			<td>30 nm</td>
			<td>11</td>
			<td>30</td>
		</tr>
		<tr>
			<td>100 nm</td>
			<td>11</td>
			<td>100</td>
		</tr>
		<tr>
			<td>400 nm</td>
			<td>21</td>
			<td>400</td>
		</tr>
	</tbody>
</table>
Table 1. Nanosight Parameters.
<img alt="Figure 1" src="/files/ftp_upload/4151/4151fig1.jpg" /> 
Figure 1. Flow chart of the extrusion method, describing how the pressure and nitrogen flow controls the homogeneity of different liposome diameters. Following liposome hydration, unilamellar vesicles of different sizes are extruded through different polycarbonate membrane filter pores at different pressures.
<img alt="Figure 2" src="/files/ftp_upload/4151/4151fig2.jpg" /> 
Figure 2. Dynamic Light Scattering (DLS) data describing the quantitative liposome sizes following extrusion. (a) Bar graphs are shown to describe the diameters of the three liposome samples. A calibration standard of 50 nm polystyrene (PS) beads was used as a reference. The average liposome diameters are indicated above the bar for each sample. The x-axis describes the pore sizes of which the solutions were extruded through. The y-axis describes the diameter recorded by DLS. Although the 30 nm and 100 nm sizes recorded values higher than their pore sizes while the larger 400 nm size recorded a slightly lower size, the calibration curve (b) shows a near linear correlation, where the x-axis expresses the polycarbonate membrane pore sizes, and the y-axis describes the recorded liposome diameter by DLS.
<img alt="Figure 3" src="/files/ftp_upload/4151/4151fig3.jpg" /> 
Figure 3. Nanoparticle Tracking Analysis (NTA) data describing the liposome sizes following extrusion. (a) The bar graphs represent the diameter of each liposome sample. The x-axis describes the pore sizes of which the solutions were extruded through. The y-axis describes the size diameter recorded by the NTA. The average liposome diameters are labeled above each sample. (b) The NTA calibration curve shows a more linear correlation than the DLS calibration curve between the filter membrane pore sizes versus the recorded diameter. The x-axis describes the polycarbonate membrane pore sizes. The y-axis describes the recorded liposome diameter by NTA.
<img alt="Figure 4" src="/files/ftp_upload/4151/4151fig4.jpg" /> 
Figure 4. Negative stain transmission electron microscopy (TEM) images showing each liposome size (500 &mu;M) followed by extrusion. Carbon Formvar mesh grids were negatively discharged prior to sample staining, where 1% Uranyl acetate in water was used to stain the samples prior to drying and imaging at 34,000&times; magnification. Sizes 30, 100, and 400 nm are clearly distinct from each other. The magnification was set to 25,000&times;. The scale bar represents 0.5 &mu;m.
<img alt="Figure 5" src="/files/ftp_upload/4151/4151fig5.jpg" /> 
Figure 5. A time-course experiment was conducted with three liposome sizes. Dynamic light scattering (DLS) was measured for each liposome solution immediately following extrusion. All liposome solutions were stored in 4 &deg;C overnight. Their diameters were recorded by the DLS after an overnight incubation. Little to no change was observed after a 16-hour incubation period.

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P., Yao, J., Dunning, F. M., Chapman, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptotagmin-mediated+bending+of+the+target+membrane+is+a+critical+step+in+Ca(2+)-regulated+fusion.">Synaptotagmin-mediated bending of the target membrane is a critical step in Ca(2+)-regulated fusion.</a> Cell. 138, 709-721 (2009).</li><li>Loughrey, H. C., Choi, L. S., Cullis, P. R., Bally, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimized+procedures+for+the+coupling+of+proteins+to+liposomes.">Optimized procedures for the coupling of proteins to liposomes.</a> J. Immunol. Methods. 132, 25-35 (1990).</li><li>Mui, B., Chow, L., Hope, M. 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No conflicts of interest declared.

Using the Avestin Liposofast LF-50 Extruder, we demonstrated how small-sized, synthetic liposomes are prepared through a pressure-controlled system. It is important to note that multilamellar vesicles form spontaneously following liposome hydration, which may lead to production of smaller nanoparticles. These small multilamellar vesicles will inevitably flow through the larger polycarbonate membrane pore size, causing heterogeneity in solutions of unilamellar vesicles produced by a large filter pore. Therefore, it is rec...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Chloroform</td>
 <td>Sigma-Aldrich</td>
 <td>02432-25ML</td>
 <td>95% stabilizers</td>
 </tr>
 <tr>
 <td>High Grade Methanol</td>
 <td>Sigma-Aldrich</td>
 <td>179337-4L</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Liposofast LF-50 Extruder</td>
 <td>Avestin, Inc.</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Phospholipids</td>
 <td>Avanti Polar Lipids</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Polycarbonate Pores</td>
 <td>Avestin, Inc.</td>
 <td>&nbsp;</td>
 <td>25 mm diameter</td>
 </tr>
 <tr>
 <td>Drain discs PE</td>
 <td>Avestin, Inc.</td>
 <td>230600</td>
 <td>25 mm diameter</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Constant Pressure-controlled Extrusion Method for the Preparation of Nano-sized Lipid Vesicles at JoVE.com

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

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 ...

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23.8K Views.  University of Colorado Boulder. 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.

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

Improved Method for the Preparation of a Human Cell-based, Contact Model of the Blood-Brain Barrier

The blood-brain barrier (BBB) comprises impermeable but adaptable brain capillaries which tightly control the brain environment. Failure of the BBB has been implied in the etiology of many brain pathologies, creating a need for development of human in vitro BBB models to assist in clinically-relevant research. Among the numerous BBB models thus far described, a static (without flow), contact BBB model, where astrocytes and brain endothelial cells (BECs) are cocultured on the opposite sides of a porous membrane, emerged as a simplified yet authentic system to simulate the BBB with high throughput screening capacity. Nevertheless the generation of such model presents few technical challenges. Here, we describe a protocol for preparation of a contact human BBB model utilizing a novel combination of primary human BECs and immortalized human astrocytes. Specifically, we detail an innovative method for cell-seeding on inverted inserts as well as specify insert staining techniques and exemplify how we use our model for BBB-related research.

Establishment of human models of the blood-brain barrier (BBB) can benefit research into brain conditions associated with BBB failure. We describe here an improved technique for preparation of a contact BBB model, which permits coculturing of human astrocytes and brain endothelial cells on the opposite sides of a porous membrane.

The blood-brain barrier (BBB) comprises impermeable but adaptable brain capillaries which tightly control the brain environment. Failure of the BBB has ...

An Improved Method for the Preparation of Type I Collagen From Skin

Soluble type 1 collagen (COL1) is used extensively as an adhesive substrate for cell cultures and as a cellular scaffold for regenerative applications. Clinically, this protein is widely used for cosmetic surgery, dermal injections, bone grafting, and reconstructive surgery. The sources of COL1 for these procedures are commonly nonhuman, which increases the potential for inflammation and rejection as well as xenobiotic disease transmission. In view of this, a method to efficiently and quickly purify COL1 from limited quantities of autologously-derived tissues would circumvent many of these issues; however, standard isolation protocols are lengthy and often require large quantities of collagenous tissues. Here, we demonstrate an efficient COL1 extraction method that reduces the time needed to isolate and purify this protein from about 10 days to less than 3 hr. We chose the dermis as our tissue source because of its availability during many surgical procedures. This method uses traditional extraction buffers combined with forceful agitation and centrifugal filtration to obtain highly-pure, soluble COL1 from small amounts of corium. Briefly, dermal biopsies are washed thoroughly in ice-cold dH2O after removing fat, connective tissue, and hair. The skin samples are stripped of noncollagenous proteins and polysaccharides using 0.5 M sodium acetate and a high speed bench-top homogenizer. Collagen from residual solids is subsequently extracted with a 0.075 M sodium citrate buffer using the homogenizer. These extracts are purified using 100,000 MW cut-off centrifugal filters that yield COL1 preparations of comparable or superior quality to commercial products or those obtained using traditional procedures. We anticipate this method will facilitate the utilization of autologously-derived COL1 for a multitude of research and clinical applications.

Traditional procedures for the isolation of soluble type 1 collagen (COL1) require about 10 days from start to finish because of lengthy buffer incubations and laborious resuspensions of fibrils. Here, we describe a means to purify COL1 from small dermal biopsies in less than 3 hr.

Soluble type 1 collagen (COL1) is used extensively as an adhesive substrate for cell cultures and as a cellular scaffold for regenerative applications. ...

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 ...

Preparation of Mica Supported Lipid Bilayers for High Resolution Optical Microscopy Imaging

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used. 
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.

We present a method of preparing mica supported lipid bilayers for high resolution microscopy. Mica is transparent and flat on an atomic scale, but rarely used in imaging because of handling difficulties; our preparation results in even deposition of the mica sheet, and reduces the material used in bilayer preparation.

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction ...

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 ...

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 ...

Film Extrusion of Crambe abyssinica/Wheat Gluten Blends

Crambe abyssinica is a plant with potential for use in industrial (non-food) plant oil production. The side stream from this oil production is a high-protein crambe meal that has limited value, as it is not fit for food or feed use. However, it contains proteins that could potentially make it a suitable raw material for higher-value products. The purpose of this study was to find methods of making this side stream into extruded films, showing that products with a higher value can be produced. The study mainly considered the development of material compositions and methods of preparing and extruding the material. Wheat gluten was added as a supportive protein matrix material, together with glycerol as a plasticizer and urea as a denaturant. The extrudate was evaluated with respect to mechanical (tensile testing) and oxygen barrier properties, and the extrudate structure was revealed visually and by scanning electron microscopy. A denser, more homogeneous material had a lower oxygen transmission rate, higher strength, and higher extensibility. The most homogeneous films were made at an extruder die temperature of 125-130 &#176;C. It is shown here that a film can be extruded with promising mechanical and oxygen barrier properties, the latter especially after a final compression molding step.

Film Extrusion of Crambe abyssinica/Wheat Gluten Blends

The side stream from plant oil production of Crambe abyssinica has limited value. The purpose of this study was to find methods for extruding materials based on this side stream, demonstrating that products with a higher value can be produced. The extrudates were found to have promising properties.

Crambe abyssinica is a plant with potential for use in industrial (non-food) plant oil production. The side stream from this oil production is a ...

Evaluation of the Storage Stability of Extracellular Vesicles

Extracellular vesicles (EVs) are promising targets in current research, to be used as drugs, drug-carriers, and biomarkers. For their clinical development, not only their pharmaceutical activity is important but also their production needs to be evaluated. In this context, research focuses on the isolation of EVs, their characterization, and their storage. The present manuscript aims at providing a facile procedure for the assessment of the effect of different storage conditions on EVs, without genetic manipulation or specific functional assays. This makes it possible to quickly get a first impression of the stability of EVs under a given storage condition, and EVs derived from different cell sources can be compared easily. The stability measurement is based on the physicochemical parameters of the EVs (size, particle concentration, and morphology) and the preservation of the activity of their cargo. The latter is assessed by the saponin-mediated encapsulation of the enzyme beta-glucuronidase into the EVs. Glucuronidase acts as a surrogate and allows for an easy quantification via the cleavage of a fluorescent reporter molecule. The present protocol could be a tool for researchers in the search for storage conditions that optimally retain EV properties to advance EV research toward clinical application.

Here we present a readily applicable protocol to assess the storage stability of extracellular vesicles, a group of naturally occurring nanoparticles produced by cells. The vesicles are loaded with glucuronidase as a model enzyme and stored under different conditions. After storage, their physicochemical parameters and the activity of the encapsulated enzyme are evaluated.

Extracellular vesicles (EVs) are promising targets in current research, to be used as drugs, drug-carriers, and biomarkers. For their clinical ...

Simultaneous Quantification of Selected Kynurenines Analyzed by Liquid Chromatography-Mass Spectrometry in Medium Collected from Cancer Cell Cultures

The kynurenine pathway and the tryptophan catabolites called kynurenines have received increased attention for their involvement in immune regulation and cancer biology. An in vitro cell culture assay is often used to learn about the contribution of different tryptophan catabolites in a disease mechanism and for testing therapeutic strategies. Cell culture medium that is rich in secreted metabolites and signaling molecules reflects the status of tryptophan metabolism and other cellular events. New protocols for the reliable quantification of multiple kynurenines in the complex cell culture medium are desired to allow for a reliable and quick analysis of multiple samples. This can be accomplished with liquid chromatography coupled with mass spectrometry. This powerful technique is employed in many clinical and research laboratories for the quantification of metabolites and can be used for measuring kynurenines.
Presented here is the use of liquid chromatography coupled with single quadrupole mass spectrometry (LC-SQ) for the simultaneous determination of four kynurenines, i.e., kynurenine, 3-hydroxykynurenine, 3-hydroxyanthranilic, and xanthurenic acid in the medium collected from in vitro cultured cancer cells. SQ detector is simple to use and less expensive compared to other mass spectrometers. In the SQ-MS analysis, multiple ions from the sample are generated and separated according to their specific mass-to-charge ratio (m/z), followed by the detection using a Single Ion Monitoring (SIM) mode.
This paper draws the attention on the advantages of the reported method and indicates some weak points. It is focused on critical elements of LC-SQ analysis including sample preparation along with chromatography and mass spectrometry analysis. The quality control, method calibration conditions and matrix effect issues are also discussed. We described a simple application of 3-nitrotyrosine as one analog standard for all target analytes. As confirmed by experiments with human ovary and breast cancer cells, the proposed LC-SQ method&#160;generates reliable results and can be further applied to other in vitro cellular models.

Described here is a protocol for the determination of four different tryptophan metabolites generated in the kynurenine pathway (kynurenine, 3-hydroxykynurenine, xanthurenic acid, 3-hydroxyanthranilic acid) in the medium collected from cancer cell cultures analyzed by liquid chromatography coupled with a single quadrupole mass spectrometry.

The kynurenine pathway and the tryptophan catabolites called kynurenines have received increased attention for their involvement in immune regulation and ...

Molecular Spring Constant Analysis by Biomembrane Force Probe Spectroscopy

A biomembrane force probe (BFP) has recently emerged as a native-cell-surface or in situ dynamic force spectroscopy (DFS) nanotool that can measure single-molecular binding kinetics, assess mechanical properties of ligand-receptor interactions, visualize protein dynamic conformational changes and more excitingly elucidate receptor mediated cell mechanosensing mechanisms. More recently, BFP has been used to measure the spring constant of molecular bonds. This protocol describes the step-by-step procedure to perform molecular spring constant DFS analysis. Specifically, two BFP operation modes are discussed, namely the Bead-Cell and Bead-Bead modes. This protocol focuses on deriving spring constants of the molecular bond and cell from DFS raw data.

A biomembrane force probe (BFP) is an in situ dynamic force spectroscopy (DFS) technique. BFP can be used to measure the spring constant of molecular interactions on living cells. This protocol presents spring constant analysis for molecular bonds detected by BFP.

A biomembrane force probe (BFP) has recently emerged as a native-cell-surface or in situ dynamic force spectroscopy (DFS) nanotool that can measure ...