Name: obtention of giant unilamellar hybrid vesicles by electroformation and measurement of their mechanical properties by micropipette aspiration
Uploaded: 2020-01-19T13:00:00.000+00:00
Duration: 9 min 29 s
Description: Watch this Scientific Journal Video about Obtention of Giant Unilamellar Hybrid Vesicles by Electroformation and Measurement of their Mechanical Properties by Micropipette Aspiration at JoVE.com

The authors gratefully acknowledge the ANR for financial support (ANR Sysa).

1. Fabricating micropipettes
NOTE: Here, micropipettes with an inner diameter ranging from 6 to 12 &#181;m and a taper length around 3-4 mm are necessary. A detailed method of manufacturing micropipette is described in the following.
<ol>
	<li>Place the borosilicate glass capillary in the drawbar of the puller and fix one of the ends by tightening the knob.</li>
	<li>Carefully slide the glass through the holes at the side of the heater chamber.</li>
	<li>Tighten down the clamping knob at the other end.</li>
	<li>Control the size of the tip and the taper length to achieve the desired specifications. For that, optimize technical parameters such as heating temperature, pull, velocity, delay and pressure. Here is an example of a program used: 
	Heat: 550 &#176;C 
	Pull: 10 (Range of the machine: 0-255 in arbitrary units) 
	Velocity: 30 (Range: 0-255 in arbitrary units) 
	Delay: 1 (Range: 0-255 in arbitrary units) 
	Pressure: 500 (Range: 0-999 in arbitrary units)</li>
	<li>Click on PULL to execute the events defined by the program. The capillary is then separated into two micropipettes, whose dimensions have to be adjusted using a microforge.</li>
	<li>Insert the micropipette into the metal pipette holder of the microforge (see Figure 1). By using 10x objective, adjust the microscope stage and the pipette manipulator (vertical and horizontal movement) until the pipette tip is close to the glass bead surface.</li>
	<li>Press the foot switch to melt the glass bead. Lower the tip and put it in contact with the molten glass bead. Molten glass will flow into the pipette by capillary action. Wait a few seconds until the level of the molten glass reaches a certain height as shown in the insert of the Figure 1.</li>
	<li>Stop the heating by removing the pressure on the foot switch, and quickly pull the tip away using the vertical pipette manipulator to cause a sharp break.</li>
	<li>Repeat steps 1.7 and 1.8 until the desired diameter is obtained (6 to 12 &#181;m). 
	NOTE: To improve the accuracy of the diameter measurement, during the last step, use a 32x objective equipped with a reticle.</li>
</ol>
2. Coating pipette tips with BSA (bovine serum albumin)
<ol>
	<li>To prepare a 0.1 M solution of glucose containing 1% wt. BSA in pure water.
	<ol>
		<li>Weigh 180 mg of glucose powder, place in a 15 mL polypropylene conical tube and complete with 10 mL of pure water.</li>
		<li>Add 0.1 g of BSA powder and shake gently using a test tube rotatory mixer until complete dissolution (approximately 4 h).</li>
	</ol>
	</li>
	<li>Take the solution with a 10 mL disposable Luer syringe fitted with a needle. Once filled, remove the needle and install a 0.22 &#956;m acetate cellulose filter. Fill several polypropylene micro-centrifuge tubes (1.5 mL) that will be used to immerse the tip.</li>
	<li>Place the capillaries vertically into holders. Lower the holder and immerse the tip into the glucose/BSA solution overnight. The solution should rise about 1 cm high by capillary action.</li>
	<li>Remove the pipette tip from the glucose/BSA solution. Prepare 5 mL of 0.1 M glucose solution (dilute 90 mg of glucose powder in 5 mL of pure water) and filter through a 0.22 &#956;m acetate cellulose filter.</li>
	<li>Fill the pipette with the glucose solution by using a 500 &#181;L glass syringe equipped with a flexible fused silica capillary. Then, remove all the glucose solution by sucking it back and discard it (Figure 2). Repeat this step several times to remove the unbounded BSA.</li>
</ol>
3. Formation of GUVs and GHUVs by electroformation
NOTE: Electroformation is a commonly used technic developed by Angelova9. The procedures to obtain an electroformation chamber, deposit a lipid or polymer film (or both for GHUVs (Giant Hybrid Unilamellar Vesicles)) and hydrate the film under an alternative electric field are described in the following. The procedure to collect the GUV obtained is also described.
<ol>
	<li>Amphiphile, sucrose and glucose solutions preparation
	<ol>
		<li>Prepare an amphiphile solution at a concentration of 1 mg/mL. Weigh 10 mg of amphiphile and dissolve in 10 mL of chloroform. Keep the solution in sealed vials to avoid solvent evaporation.</li>
		<li>Prepare a stock solution of 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (DOPE-Rhod) at 1 mg/mL in chloroform.</li>
		<li>Add 10 &#181;L of fluorescent lipid solution to the amphiphile solution. Keep the solutions in sealed vials to avoid solvent evaporation.</li>
		<li>Prepare sucrose and glucose solutions at a concentration of 0.1 M. Weigh 342 mg and 180 mg of sucrose and glucose, respectively, and dissolve them in 10 mL of pure water.</li>
	</ol>
	</li>
	<li>Preparation of the electroformation chamber 
	NOTE: Different conductive materials can be used to make an electroformation device (e.g., platinum wires10, stainless needles11). The electroformation chamber is composed of two ITO slides separated by an O-ring rubber spacer that has been cut on one side to create an aperture. The slides are connected to a voltage generator via two electric wires (Figure 3 and Figure 4A).
	<ol>
		<li>Clean the ITO slides with organic solvent (e.g., chloroform). Identify the conductive surface using an ohmmeter.</li>
		<li>Attach the electrical wires on the conductive side using adhesive tape.</li>
		<li>Amphiphile solution deposition
		<ol>
			<li>Dip a capillary in the solution until the level increases by capillary action and collect about 5 &#956;L of the solution.</li>
			<li>Put the capillary in contact with the center of the ITO glass plate and gently spread the solution. Wait 10 seconds to ensure complete solvent evaporation (Figure 4A).</li>
			<li>Repeat this procedure 3 times for each side.</li>
		</ol>
		</li>
		<li>Add a layer of silicon-free grease on both sides of the opened O-ring spacer. Put it around the area of deposition. Place the conductive face of the second ITO glass plate on the top of the spacer.</li>
		<li>Place the electroformation chamber under vacuum for 3 hours to remove any traces of organic solvent.</li>
	</ol>
	</li>
	<li>Electroformation procedure
	<ol>
		<li>Plug the electric wires to the generator.</li>
		<li>Use the following settings for the generator: 
		Alternative sinusoidal tension 
		Frequency: 10 Hz 
		Amplitude: 2 Vpeak-to-peak 
		NOTE: Optimum voltage frequency and duration must be found for each system.</li>
		<li>Ensure that the voltage is applied before injection of the solution in the chamber.</li>
		<li>Inject 1 mL of solution using a syringe with 0.8 mm inner diameter needle to fill the chamber. Remove eventual bubbles.</li>
		<li>Let the chamber under the applied voltage/frequency for 75 min (Figure 4B).</li>
	</ol>
	</li>
	<li>GUVs harvest
	<ol>
		<li>Switch off the generator.</li>
		<li>Using 1 mL syringe with 0.8 mm inner diameter needle, suck a small part of the solution in order to produce an air bubble inside the chamber. Slightly tilt the chamber in order to make this bubble move inside the chamber. This can help the GUVs to detach from the ITO surface (Figure 4C).</li>
		<li>Suck all the solution and transfer it into a 1 mL plastic tube.</li>
		<li>Remove the wires and clean the ITO slides with organic solvents (toluene then chloroform).</li>
	</ol>
	</li>
</ol>
4. Micromanipulation set up
NOTE: The principle of micropipette aspiration is to suck a single vesicle through a glass micropipette by applying a depression. The length of the tongue inside the pipette is measured as a function of the suction pressure. The pipette coating with BSA, described previously, is essential to prevent or minimize any adhesion between vesicles membranes and the pipette.
The protocol is illustrated below.
<ol>
	<li>Pipette and water filled reservoir connection 
	NOTE: The water filled tank and the micrometer are fixed to a sliding plate. A digital counter with a micrometer head allows a vertical displacement of the device in a range of 0 to 2.5 cm and an accuracy of 1 &#181;m. Displacement along an aluminum optical rail is possible up to 1 meter in length. A silicon tubing connects the reservoir and the capillary holder (Figure 5A).
	<ol>
		<li>Fill the tank with pure water. Connect a disposable 5 mL syringe to the capillary metal holder via silicone tubing and aspirate to create a water flow from the tank to the holder.</li>
		<li>Touch the tube slightly to eliminate air bubbles. Simultaneously, raise the water tank to create a positive pressure. The 5 mL syringe is still attached to the holder.</li>
		<li>After coating and cleaning steps described previously (see step 2), fill a capillary with glucose solution until a drop forms at the end. Remove the syringe tubing from the metal holder to create a slight water flow at its end.</li>
		<li>Turn the capillary upside down and connect the glucose drop with the water flow from the holder. Fix the capillary and the holder by screwing them together.</li>
	</ol>
	</li>
	<li>Position a pipette 
	NOTE: During this operation, the water tank is still positioned on the aluminum rail to maintain a positive pressure.
	<ol>
		<li>Take the homemade aluminum stage equipped with two glass slides (previously cleaned with ethanol) and glue them with vacuum grease on each side. Install it on the microscope stage and form a meniscus between the two slides by using a 1 mL syringe containing 0.1 M of glucose as shown in Figure 5B,C.</li>
		<li>Place the pipette and its holder on the motor unit of the micromanipulator and tighten down clamping knob.</li>
		<li>Use the control panel joystick in coarse mode to lower the micropipette near the glucose meniscus. Adjust the position of the tip to the center of the meniscus using the fine mode.</li>
		<li>Hold the tip immersed in glucose for a few minutes to clean its outer and inner surface (as a positive pressure is maintained, the water flow will rinse the inner surface of the pipette to remove uncoated BSA).</li>
		<li>Store the position of the tip on the micromanipulator keyboard and withdraw it from the meniscus.</li>
		<li>Remove the glucose meniscus and replace it by a fresh one. Suck 2 &#956;L of GUVs in 0.1 M sucrose by using a 20 &#956;L micropipette and put it in the fresh glucose meniscus. Observe in DIC mode (differential interference contrast) the GUVs located at the bottom due to the difference of density between sucrose (mainly inside the GUVs) and glucose (mainly outside the GUVs).</li>
		<li>When the vesicles are slightly deflated, reinsert the suction pipette and focus on the tip. Set the height H0 of the water tank for which pressure is almost 0. For that, take advantage of small vesicles or particles which are naturally present in solution and adjust the water tank height until the motion of these particles is stopped.</li>
		<li>At this point, surround the meniscus with mineral oil to prevent water evaporation, see Figure 5D. 
		NOTE: The room temperature should be controlled between 20-22 &#176;C by using air conditioning.</li>
	</ol>
	</li>
	<li>Micropipette aspiration experiment
	<ol>
		<li>Lower the pipette tip (6-12 &#181;m in diameter) and create a small suction (-1 cm) to aspirate a vesicle (15-30 &#181;m in diameter). The membrane of the selected vesicle should slightly fluctuate, and must not present any visible defects (no bud or filament) (Figure 6).</li>
		<li>Raise the pipette to a higher level to isolate the aspirated vesicle from the other vesicles, by using the micromanipulator and keep this position during the whole experiment.</li>
		<li>Pre-stress the vesicle by lowering the water filled tank to approximately -10 cm, then decrease the pressure to return to its initial value (-1 cm). Repeat this step several times to remove membrane excess and small defects from the membrane.</li>
		<li>From a height of -0.5 cm defined by the position of the water tank, slowly decrease the suction pressure with the micrometer to reach a regime in which the membrane fluctuates. Then increase the pressure to clearly visualize a tongue in the tip with a significant projection length (a few microns). 
		NOTE: The lowest applied pressure (P0) that allows sucking up the smallest membrane projection length (L0) will define the reference point &#945;0 (Figure 7A). All points of the curve will be measured according to this reference (&#916;L = L-L0 and &#916;P = P-P0).</li>
		<li>Increase the suction pressure with the micrometer in a stepwise manner until reaching 0.5 -0.8 mN/m. At each step, wait 5 s and take a snapshot of the tongue. This procedure at low tension enables determination of the bending modulus.</li>
		<li>Keep on increasing the suction pressure from 0.5 mN/m to the rupture tension by adjusting the height of the water filled sliding on the rail (ranging from -2 to -50 cm) (Figure 7B-D). From this experiment at high tension regime, the area compressibility modulus, lysis tension and lysis strain will be measured.</li>
		<li>Stretch about 15-20 vesicles to acquire significant statistics. Each micropipette aspiration experiment takes between 7 and 10 minutes. Perform image analysis using LASAF software for measuring the projection length of the tongue, the diameter of vesicles and the radius of the capillary.</li>
	</ol>
	</li>
	<li>Measuring bending modulus, area compressibility modulus, lysis tension and lysis strain
	<ol>
		<li>To access these parameters, use the formalism established by E. Evans12. Calculate the suction pressure applied over the membrane from equation 1: 
		&#916; P = &#961;wg (h&#8722;h0) (1) 
		where g is the gravitational acceleration (9.8 m&#8729;s&#8722;2), &#961;w is the density of the water (&#961; = 1 g&#8729;cm&#8722;3), h is the position of the water tank and h0 is the initial position where pressure is equal to zero.</li>
		<li>Calculate the membrane tension from the Laplace equation: 
		<img alt="Equation 1" src="/files/ftp_upload/60199/60199eq1.jpg" /> (2) 
		where &#916;P is the suction pressure on the micropipette, Rp and Rv are the micropipette and vesicle radii (outside the micropipette) respectively. The surface area strain (&#945;) of the membrane is defined as: 
		<img alt="Equation 2" src="/files/ftp_upload/60199/60199eq2.jpg" /> (3) 
		<img alt="Equation 3" src="/files/ftp_upload/60199/60199eq3.jpg" /> being the membrane area of the vesicle at the lower suction pressure.</li>
		<li>Calculate &#945; from the increase in projection length &#916;L of vesicle inside the capillary tip according to Equation 412: 
		<img alt="Equation 4" src="/files/ftp_upload/60199/60199eq4.jpg" /> (4)</li>
		<li>Under a very low tension regime, smoothing of thermal bending undulations dominates the apparent expansion. Plot ln(&#963;) vs &#945;. At low-&#963; values (typically 0.001&#8211;0.5 mN.m&#8722;1 13), this gives a straight line whose slope is linked to the bending modulus, Kb (first term of the equation 5)14: 
		<img alt="Equation 5" src="/files/ftp_upload/60199/60199eq5.jpg" /> (5) 
		NOTE: Under high tensions (&#62; 0.5 mN.m&#8722;1), membrane undulations are completely suppressed and membrane area increases as the result of increased spacing between molecules. In this regime, the second term of equation 5 dominates and give access to area compressibility modulus Ka (Figure 8 and Figure 9).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The coating of the micropipette is one of the key points to obtain reliable measurements. Adhesion of the vesicle to the micropipette must be prevented, and a coating is commonly used in literature17,18,19,20,21, with BSA, &#946;-casein or surfasil. Details of the coating procedure are rarely mentioned.
Dissolution of the BSA should...

Giant vesicles obtained from phospholipids (liposomes) have been widely used since the 1970s as the basic cell membrane model1. In the late 1990s, vesicular morphologies obtained from the self-assembly of copolymers, named polymersomes in reference to their lipid analogs2,3, rapidly appeared as an interesting alternative to liposomes that possess weak mechanical stability and poor modular chemical functionality. However, their cell biomimetic character is rather limited compared to liposomes since the latter are composed of phospholipids, the main component of the cell membrane. Furthermore, their low membrane permeability can be an issue in some applications like drug delivery where controlled diffusion of species through the membrane is required. Recently, the association of phospholipids with block copolymers to design hybrid polymer-lipid vesicles and membranes has been the subject of an increasing number of studies4,5. The main idea is to design entities that synergistically combine the benefits of each component (bio-functionality and permeability of lipid bilayers with the mechanical stability and chemical versatility of polymer membranes), which can be exploited in different applications: controlled and targeted drug delivery, biomolecular recognition within biosensors for diagnosis, functional membranes for artificial cells, development of bio-inspired micro-/nano-reactors.
Nowadays, different scientific communities (biochemists, chemists, biophysicists, physico-chemists, biologists) have increasing interest in development of a more advanced cell membrane model. Here, our goal is to present, as detailed as possible, existing methodologies (electroformation, micropipette aspiration) to obtain and characterize the mechanical properties of giant vesicles and the recent &#34;advanced&#34; cell membrane models that are hybrid polymer lipid giant vesicles4,5.
The purpose of these methods is to obtain reliable measurement of the area compressibility and bending moduli of the membrane as well as their lysis stress and strain. One of the most common techniques existing to measure bending rigidity of a giant vesicle is fluctuation analysis6,7, based on direct video microscope observation; but this requires large visible membrane fluctuation, and is not systematically obtained on thick membranes (e.g. polymersomes). Area compressibility modulus can be experimentally determined using the Langmuir Blodgett technique but most often on a monolayer8. The micropipette aspiration technique allows the measurement of both moduli on a bilayer forming giant unilamellar vesicle (GUV) in one experiment.
The following method is appropriate for all amphiphilic molecules or macromolecules able to form bilayers and, consequently, vesicles by electroformation. This requires a fluid character of the bilayer at the temperature of electroformation.

<table><tbody><tr><td>&lt;strong&gt;&lt;em&gt;Required equipment and materials for micropipette design&lt;/em&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Borosilicate Glass Capillaries</td><td>World Precision Instruments</td><td>1B100-4</td><td>external and internal diameter of 1mm and 0.58 mm respectively.</td></tr><tr><td>Filament installed</td><td>Sutter Instrument Co.</td><td>FB255B</td><td>2.5mm*2.5mm Box Filament</td></tr><tr><td>Flaming/Brown Micropipette Puller</td><td>Sutter Instrument Co.</td><td>Model P-97</td><td></td></tr><tr><td>Microforge</td><td>NARISHGE Co.</td><td>MF-900</td><td>fitted with two objectives (10x and 32x)</td></tr><tr><td>&lt;strong&gt;&lt;em&gt;Materials for coating pipette tips with BSA&lt;/em&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Bovine Serum Albumin Fraction V (BSA)</td><td>Sigma-Aldrich</td><td>10735078001</td><td></td></tr><tr><td>Disposable 1 ml syringe Luer Tip</td><td>Codan</td><td>62.1612</td><td></td></tr><tr><td>Disposable 10 ml syringe Luer Tip</td><td>Codan</td><td>626616</td><td></td></tr><tr><td>Disposable 5 ml syringe Luer Tip</td><td>Codan</td><td>62.5607</td><td></td></tr><tr><td>Disposable acetate cellulose filter</td><td>Cluzeau Info Labo</td><td>L5003SPA</td><td>Pore size: 0.22&amp;micro;m, diameter: 25mm</td></tr><tr><td>Flexible Fused Silica Capillary Tubing</td><td>Polymicro Technologies.</td><td>TSP530660</td><td>Inner Diameter 536&amp;micro;m, Outer Diameter 660&amp;micro;m,</td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G5767</td><td></td></tr><tr><td>Syringe 500 &amp;micro;L luer Lock GASTIGHT</td><td>Hamilton Syringe Company</td><td>1750</td><td></td></tr><tr><td>Test tube rotatory mixer</td><td>Labinco</td><td>28210109</td><td></td></tr><tr><td>&lt;strong&gt;&lt;em&gt;Micromanipulation Set up&lt;/em&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Aluminum Optical Rail, 1000 mm Length, M4 threads, X48 Series</td><td>Newport</td><td></td><td></td></tr><tr><td>Damped Optical Table</td><td>Newport</td><td></td><td>used as support of microscope to prevent external vibrations.</td></tr><tr><td>Micromanipulator</td><td>Eppendorf</td><td>Patchman NP 2</td><td>The module unit (motor unit for X, Y and Z movement) is mounted on the inverted microscope by the way of an adapter.</td></tr><tr><td>Micrometer</td><td>Mitutoyo Corporation</td><td>350-354-10</td><td>Digimatic LCD Micrometer Head 25,4 mm Range 0,001 mm</td></tr><tr><td>Plexiglass water reservoir (100 ml)</td><td>Home made</td><td></td><td></td></tr><tr><td>TCS SP5 inverted confocal microscope (DMI6000) equipped with a resonant scanner and a water immersion objective (HCX APO L 40x/0.80 WU-V-I).</td><td>Leica</td><td></td><td></td></tr><tr><td>X48 Rail Carrier 80 mm Length,with 1/4-20, 8-32 and 4-40 thread</td><td>Newport</td><td></td><td></td></tr><tr><td>&lt;strong&gt;&lt;em&gt;Materials for sucrose and amphiphile solution preparation&lt;/em&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;a href=&quot;https://www.sigmaaldrich.com/catalog/substance/2oleoyl1palmitoylsnglycero3phosphocholine760082685331611&quot; target=&quot;_blank&quot;&gt;2-Oleoyl-1-palmitoyl-sn-glycero-3-phosphocholine&lt;/a&gt;</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Chloroform</td><td>VWR</td><td>22711.244</td><td></td></tr><tr><td>L-&amp;alpha;-Phosphatidylethanolamine-N-(lissamine rhodamine B sulfonyl)</td><td>Sigma-Aldrich</td><td>810146C</td><td>Rhodamine tagged lipid</td></tr><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>S7903</td><td></td></tr><tr><td>&lt;strong&gt;&lt;em&gt;Electroformation set up&lt;/em&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>10 &amp;micro;L glass capillary ringcaps</td><td>Hirschmann</td><td>9600110</td><td></td></tr><tr><td>Disposable 1 ml syringe Luer Tip</td><td>Codan</td><td>62.1612</td><td></td></tr><tr><td>H Grease</td><td>Apiezon</td><td>Apiezon H Grease</td><td>Silicon-free grease</td></tr><tr><td>Indium tin oxide coated glass slides</td><td>Sigma-Aldrich</td><td>703184</td><td></td></tr><tr><td>Needle</td><td>Terumo</td><td>AN2138R1</td><td>0.8 x 38 mm</td></tr><tr><td>Ohmmeter (Multimeter)</td><td>Voltcraft</td><td>VC140</td><td></td></tr><tr><td>Toluene</td><td>VWR</td><td>28676.297</td><td></td></tr><tr><td>Voltage generator</td><td>Keysight</td><td>33210A</td><td></td></tr></tbody></table>

obtention of giant unilamellar hybrid vesicles by electroformation and measurement of their mechanical properties by micropipette aspiration

Giant vesicles obtained from phospholipids and copolymers can be exploited in different applications: controlled and targeted drug delivery, biomolecular recognition within biosensors for diagnosis, functional membranes for artificial cells, and development of bioinspired micro/nano-reactors. In all of these applications, the characterization of their membrane properties is of fundamental importance. Among existing characterization techniques, micropipette aspiration, pioneered by E. Evans, allows the measurement of mechanical properties of the membrane such as area compressibility modulus, bending modulus and lysis stress and strain. Here, we present all the methodologies and detailed procedures to obtain giant vesicles from the thin film of a lipid or copolymer (or both), the manufacturing and surface treatment of micropipettes, and the aspiration procedure leading to the measurement of all the parameters previously mentioned.

With the protocol aforementioned, we have studied different synthetic giant unilamellar vesicle (GUV), obtained from a phospholipid: 2-oleoyl-1-palmitoyl-sn-glycero-3-phosphocholine (POPC), a triblock copolymer: Poly(ethyleneoxide)-b-Poly(dimethylsiloxane)-b-Poly(ethyleneoxide) (PEO12-b-PDMS43-b-PEO12) synthesized in a previous study13, and a diblock copolymer Poly(dimethylsiloxane)-b...

Watch this Scientific Journal Video about Obtention of Giant Unilamellar Hybrid Vesicles by Electroformation and Measurement of their Mechanical Properties by Micropipette Aspiration at JoVE.com

Obtention of Giant Unilamellar Hybrid Vesicles by Electroformation and Measurement of their Mechanical Properties by Micropipette Aspiration

The goal of the protocol is to reliably measure membrane mechanical properties of giant vesicles by micropipette aspiration.

Giant vesicles obtained from phospholipids and copolymers can be exploited in different applications: controlled and targeted drug delivery, biomolecular ...

obtention-giant-unilamellar-hybrid-vesicles-electroformation

Research

JoVE Journal

Chemistry

8.3K Views.  CNRS, Bordeaux INP, LCPO, UMR 5629. The goal of the protocol is to reliably measure membrane mechanical properties of giant vesicles by micropipette aspiration.

Video: Obtention of Giant Unilamellar Hybrid Vesicles by Electroformation and Measurement of their Mechanical Properties by Micropipette Aspiration

Preparation of Giant Vesicles Encapsulating Microspheres by Centrifugation of a Water-in-oil Emulsion

The constructive biology and the synthetic biology approach to creating artificial life involve the bottom-up assembly of biological or nonbiological materials. Such approaches have received considerable attention in research on the boundary between living and nonliving matter and have been used to construct artificial cells over the past two decades. In particular, Giant Vesicles (GVs) have often been used as artificial cell membranes. In this paper, we describe the preparation of GVs encapsulating highly packed microspheres as a model of cells containing highly condensed biomolecules. The GVs were prepared by means of a simple water-in-oil emulsion centrifugation method. Specifically, a homogenizer was used to emulsify an aqueous solution containing the materials to be encapsulated and an oil containing dissolved phospholipids, and the resulting emulsion was layered carefully on the surface of another aqueous solution. The layered system was then centrifuged to generate the GVs. This powerful method was used to encapsulate materials ranging from small molecules to microspheres.

Giant vesicles containing highly packed micrometer-sized components are useful cell models. The water-in-oil emulsion centrifugation method is a simple, powerful tool for the preparation of giant vesicles with encapsulated materials.

The constructive biology and the synthetic biology approach to creating artificial life involve the bottom-up assembly of biological or nonbiological ...

Biochemistry

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 ...

Bioengineering

Phase Behavior of Charged Vesicles Under Symmetric and Asymmetric Solution Conditions Monitored with Fluorescence Microscopy

Phase-separated giant unilamellar vesicles (GUVs) exhibiting coexisting liquid-ordered and liquid-disordered domains are a common biophysical tool to investigate the lipid raft hypothesis. Numerous studies, however, neglect the impact of physiological solution conditions. On that account, the current work presents the effect of high-salinity buffer and trans-membrane solution asymmetry on liquid-liquid phase separation in charged GUVs grown from dioleylphosphatidylglycerol, egg sphingomyelin, and cholesterol. The effects were studied under isothermal and varying temperature conditions.
We describe equipment and experimental strategies applicable for monitoring the stability of coexisting liquid domains in charged vesicles under symmetric and asymmetric high-salinity solution conditions. This includes an approach to prepare charged multicomponent GUVs in high-salinity buffer at high temperatures. The protocol entails the option to perform a partial exchange of the external solution by a simple dilution step while minimizing the vesicle dilution. An alternative approach is presented utilizing a microfluidic device that allows for a complete external solution exchange. The solution effects on phase separation were also studied under varying temperatures. To this end, we present the basic design and utility of an in-house built temperature control chamber. Furthermore, we reflect on the assessment of the GUV phase state, pitfalls associated with it and how to circumvent them.

Experiments on phase separated giant unilamellar vesicles (GUVs) frequently neglect physiological solution conditions. This work presents approaches to study the effect of high-salinity buffer on liquid-liquid phase separation in charged multicomponent GUVs as a function of trans-membrane solution asymmetry and temperature.

Phase-separated giant unilamellar vesicles (GUVs) exhibiting coexisting liquid-ordered and liquid-disordered domains are a common biophysical tool to ...

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 ...

Membrane Remodeling of Giant Vesicles in Response to Localized Calcium Ion Gradients

In a wide variety of fundamental cell processes, such as membrane trafficking and apoptosis, cell membrane shape transitions occur concurrently with local variations in calcium ion concentration. The main molecular components involved in these processes have been identified; however, the specific interplay between calcium ion gradients and the lipids within the cell membrane is far less known, mainly due to the complex nature of biological cells and the difficultly of observation schemes. To bridge this gap, a synthetic approach is successfully implemented to reveal the localized effect of calcium ions on cell membrane mimics. Establishing a mimic to resemble the conditions within a cell is a severalfold problem. First, an adequate biomimetic model with appropriate dimensions and membrane composition is required to capture the physical properties of cells. Second, a micromanipulation setup is needed to deliver a small amount of calcium ions to a particular membrane location. Finally, an observation scheme is required to detect and record the response of the lipid membrane to the external stimulation. This article offers a detailed biomimetic approach for studying the calcium ion-membrane interaction, where a lipid vesicle system, consisting of a giant unilamellar vesicle (GUV) connected to a multilamellar vesicle (MLV), is exposed to a localized calcium gradient formed using a microinjection system. The dynamics of the ionic influence on the membrane were observed using fluorescence microscopy and recorded at video frame rates. As a result of the membrane stimulation, highly curved membrane tubular protrusions (MTPs) formed inside the GUV, oriented away from the membrane. The described approach induces the remodeling of the lipid membrane and MTP production in an entirely contactless and controlled manner. This approach introduces a means to address the details of calcium ion-membrane interactions, providing new avenues to study the mechanisms of cell membrane reshaping.

We present a technique for contactless micromanipulation of vesicles, using localized calcium ion gradients. Microinjection of a calcium ion solution, in the vicinity of a giant lipid vesicle, is utilized to remodel the lipid membrane, resulting in the production of membrane tubular protrusions.

In a wide variety of fundamental cell processes, such as membrane trafficking and apoptosis, cell membrane shape transitions occur concurrently with local ...

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

Preparation, Purification, and Use of Fatty Acid-containing Liposomes

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of these amphiphiles. In particular, fatty acid liposomes enhance dynamic character, due to the relatively high solubility of single-chain amphiphiles. Similarly, liposomes containing free fatty acids are more sensitive to salt and divalent cations, due to the strong interactions between the carboxylic acid head groups and metal ions. Here we illustrate techniques for preparation, purification, and use of liposomes comprised in part or whole of single chain amphiphiles (e.g., oleic acids).

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of single chain amphiphiles. Here we describe techniques for the preparation, purification, and use of liposomes comprised in part or whole of these amphiphiles.

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due ...

Assembly of Cell Mimicking Supported and Suspended Lipid Bilayer Models for the Study of Molecular Interactions

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial protective barrier for all cell types, separating the internal cellular components from the extracellular environment. These membranes are composed largely of a lipid bilayer, which contains outer hydrophilic head groups and inner hydrophobic tail groups, along with various proteins and cholesterol. The composition and structure of the lipids themselves play a crucial role in regulating biological function, including interactions between cells and the cellular microenvironment, which may contain pharmaceuticals, biological toxins, and environmental toxicants. In this study, methods to formulate uni-lipid and multi-lipid supported and suspended cell mimicking lipid bilayers are described. Previously, uni-lipid phosphatidylcholine (PC) lipid bilayers as well as multi-lipid placental trophoblast-inspired lipid bilayers were developed for use in understanding molecular interactions. Here, methods for achieving both types of bilayer models will be presented. For cell mimicking multi-lipid bilayers, the desired lipid composition is first determined via lipid extraction from primary cells or cell lines followed by liquid chromatography-mass spectrometry (LC-MS). Using this composition, lipid vesicles are fabricated using a thin-film hydration and extrusion method and their hydrodynamic diameter and zeta potential are characterized. Supported and suspended lipid bilayers can then be formed using quartz crystal microbalance with dissipation monitoring (QCM-D) and on a porous membrane for use in a parallel artificial membrane permeability assay (PAMPA), respectively. The representative results highlight the reproducibility and versatility of in vitro cell membrane lipid bilayer models. The methods presented can aid in rapid, facile assessment of the interaction mechanisms, such as permeation, adsorption, and embedment, of various molecules and macromolecules with a cell membrane, helping in the screening of drug candidates and prediction of potential cellular toxicity.

This protocol describes the formation of cell mimicking uni-lipid and multi-lipid vesicles, supported lipid bilayers, and suspended lipid bilayers. These in vitro models can be adapted to incorporate a variety of lipid types and can be used to investigate various molecule and macromolecule interactions.

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial ...

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

Biology

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...