We thank Matthew Blosser for valuable discussion and advice. This work was supported by the Office of Naval Research (N00014-16-1-2382) and the National Science Foundation (PHY-1915017).

1. Protein labeling
<ol>
	<li>Allow NHS-Rhodamine, 5-HT1A membrane fragments, and one 7 K MWCO spin desalting column to equilibrate at room temperature.</li>
	<li>Dissolve 1 mg of NHS-rhodamine in 100 &#181;L of dimethyl sulfoxide (DMSO).</li>
	<li>Add 5 &#181;L of 1 M sodium bicarbonate solution to increase the pH of 5-HT1AR solution to pH 8.</li>
	<li>Add 3.66 &#181;L of the NHS-rhodamine solution to 50 &#181;L of the 5-HT1AR solution and pipette gently up and down in a microcentrifuge tube. 
	NOTE: Ensure to have at least 10x molar excess of NHS-rhodamine.</li>
	<li>Keep the mixture protected from light and put on rotator at room temperature for 1 h.</li>
	<li>Wash a 7 k MWCO spin column with 200 &#181;L of 1x phosphate buffer saline (1x PBS) three times for 1.5 min at 1.5 RCF for each wash.</li>
	<li>Add the labeled protein to one column and balance the amount in another microcentrifuge tube.</li>
	<li>Spin down the labeled protein once for 5 min at 1.5 RCF.</li>
	<li>Take a UV-vis spectrum using a nanodrop spectrophotometer at 280 nm and 554 nm and calculate the labeling efficiency following the manufacturer&#39;s manual.</li>
	<li>Store the labeled protein covered at 5 &#176;C until further use. The solution is stable for approximately a week after labeling.</li>
</ol>
2. GUVs with membrane-incorporated 5-HT 1A
<ol>
	<li>Preparation of materials and reagents
	<ol>
		<li>Allow the protein, lipids and BSA (Bovine serum albumin) to equilibrate to room temperature.</li>
		<li>During this time, clean the coverslips by placing them in methanol and sonicating for 30 min at 40 &#176;C. Ensure that the methanol completely covers the coverslips and the water level in the water bath is above the level of the methanol in the container. 
		NOTE: Methanol is toxic and should be handled in appropriate chemical hood.</li>
		<li>Dry off the excess methanol on the coverslips with a gentle stream of air. Place the coverslip rack covered in a 40 &#176;C oven for 15 min to ensure that the excess coverslips dry off.</li>
		<li>Begin the plasma cleaning process. First, place the coverslips into the plasma cleaner and close off the air intake valve to evacuate all the air inside the chamber.</li>
		<li>Once the chamber is under vacuum, clean the coverslips for 5 min using high RF power setting and a near complete vacuum, with only a slight air intake into the vacuum chamber. To ensure the proper level of plasma, adjust the opening of the vacuum chamber such that the resultant color of the plasma is a steady, bright pink. 
		NOTE: It is crucial when using air that the plasma remains a bright pink color for the duration of the plasma treatment step, as a darker purple color indicates that there is an improper amount of air in the chamber and will result in a suboptimal plasma treatment.</li>
		<li>Once the 5 min have passed, shut off the RF power and release the vacuum. 
		NOTE: Upon removal from the plasma chamber, please ensure that the coverslips remain covered.</li>
	</ol>
	</li>
	<li>Hydrogel preparation
	<ol>
		<li>Combine 6 mg of ultra-low melting temperature agarose with 300 &#181;L of ultrapure water (i.e., 2% (w/v) agarose). 
		NOTE: 2% agarose will be used to make protein-free GUVs. Agarose solution can be kept at 45 &#176;C for two days.</li>
		<li>Combine 9 mg of ultra-low temperature agarose with 300 &#181;L of ultrapure water for 3 w/v% agarose by as prepared in step 3.1. 3% agarose will be used to make protein incorporated GUVs.</li>
		<li>Vortex the solution briefly before placing them on the 90 &#176;C heat block for 10 min. Then, vortex the tube again before transferring it to a 45 &#176;C heat block to keep it in the molten form until further use.</li>
	</ol>
	</li>
	<li>Agarose and protein mixing
	<ol>
		<li>Mix 21 &#181;L of 3% agarose with the 7 &#181;L of 5-HT1AR membrane fragments. Pipette up and down slowly many times to ensure adequate mixing. Then, incubate at 45 &#176;C for 1 min.</li>
	</ol>
	</li>
	<li>Hydrogel and lipid deposition
	<ol>
		<li>For protein-free GUVs: Make a thin film on freshly plasma-cleaned coverslips using 20 &#181;L of 2% agarose. Quickly drop another coverslip on top of the agarose droplet and gently slide the coverslips apart to make a thin film on both coverslips. 
		NOTE: This step is tricky in that the sliding of the droplet must occur while the agarose is still in the molten form.</li>
		<li>For protein-incorporated GUVs: Pipette the protein/agarose mixture up and down one more time, and then deposit 20 &#181;L of the 2% agarose on a plasma-cleaned coverslip. Follow the slip-casting directions as described above.</li>
		<li>Allow the agarose to gel protected from light for 30 min at room temperature.</li>
		<li>Deposit the lipids dropwise on top of the agarose layer. Use a total of 10 &#181;L of 2 mg/mL of 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine (POPC) with 0.4 Mol% 1,2-Dipalmitoyl-sn-glycero-3-phosphoethanolamine (DPPE) labeled with ATTO 488 (ATTO-488-DPPE) (or lipid mixture of interest) in chloroform on top of the agarose film. Deposit the droplets using a gas chromatography needle and spread one droplet at a time around via a gentle air stream. 
		NOTE: Caution is needed with this step to make a relatively uniform layer of lipids on top of the hydrogel. Also, chloroform is toxic and should be handled in appropriate chemical hood.</li>
		<li>Assemble the Sykes-Moore (S-M) chambers by placing an O-ring on top of the coverslip, and then placing the top component of the chamber on top of the O-ring. Use the key provided by the manufacturer to assemble the chamber by screwing the chamber components together to seal the chamber and prevent any leakage. 
		NOTE: The top of the chamber should be tightened on the O-ring but caution is needed to ensure the coverslip stays intact as the coverslip can crack if the O-ring doesn&#39;t sit properly in the chamber. Also, ensure that the chamber is sealed tight enough such that the chamber does not leak when the swelling solution is added. Failure to tighten the chamber enough will result in leaks and loss of sample.</li>
	</ol>
	</li>
	<li>Swelling and harvesting of vesicles
	<ol>
		<li>Hydrate the entire system by gently pipetting 450 &#181;L of 200 mM sucrose in 1x PBS and gently tapping the chambers to ensure adequate buffer coverage of the hydrogel-lipid layers. 
		NOTE: The sucrose solution can be replaced with a rehydration buffer containing biological probes of interest.</li>
		<li>Place the chambers at 45 &#176;C and cover the top part of the chamber with a coverslip to prevent evaporation. Allow the sample to swell, protected from debris and light for 1 h.</li>
		<li>Add 100 &#181;L of 1 mg/mL BSA in ultrapure water into each well of a 96-well plate intended to be used. Incubate at room temperature for 1 h.</li>
		<li>Wash three times with ultra-pure water and once with 200 mM sucrose in 1x PBS.</li>
		<li>Finally, add 200 mM of glucose in 1x PBS until the addition of the GUV sample solution. 
		NOTE: BSA was used to block GUV adsorption.</li>
		<li>After allowing the hydrogel to swell, gently shake and tap the chamber to dislodge any GUVs that may remain attached to the hydrogel surface. Then, carefully pipette up the GUV-sucrose solution. 
		NOTE: As an optional step to ensure all vesicles are detached from the surface, gently pipette some of the sucrose suspension back onto the hydrogel surface.</li>
		<li>Move the suspension into a previously prepared microcentrifuge tube containing 700 &#181;L of 200 mM glucose in 1x PBS. 
		NOTE: The density gradient will lead to settling of the vesicles to the bottom of the centrifuge tube.</li>
		<li>Allow the vesicles to settle for another hour to ensure that the vesicles can sink to the bottom of the microcentrifuge tube, allowing for optimal collection.</li>
		<li>After the settling of GUVs in glucose, transfer 300 &#181;L from the bottom of the centrifuge tube (the settled vesicles) into the previously prepared and BSA-treated 96-well plate to examine the vesicles under the confocal microscope. 
		NOTE: Be sure to avoid the very bottom of the microcentrifuge tube to minimize the amount of debris collected in the final sample.</li>
	</ol>
	</li>
	<li>Check the samples under the microscope.
	<ol>
		<li>Shine a 488 nm laser on the sample (that allows us to visualize the membrane, as the bilayer has been labeled with ATTO-488-DPPE).</li>
		<li>Shine a 561 nm laser on the sample (that allows us to visualize the protein, since it has been labeled with NHS-Rhodamine). 
		&#8203;NOTE: Caution is needed while imaging the sample as photooxidation can destabilize the vesicles. Vesicles were observed on the same day.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62830/62830fig01.jpg" /> 
Figure 1: Illustration of the detailed protocol steps. Created with BioRender.com <a href="https://www.jove.com/files/ftp_upload/62830/62830fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

The authors have no conflicts of interest to disclose.

We have identified two steps that are critical to the success of the overall protocol:&#8239; plasma treatment and lipid deposition.&#8239;Plasma cleaning of the coverslips is essential in&#8239;ensuring that there is adequate coverage and adhesion of the agarose hydrogel to the glass coverslip. Plasma cleaning accomplishes two things:&#8239;first, it removes traces of organic matter from the glass surface; second, it activates the coverslip surface, allowing for an increase in wettability as the glass surface hydrophili...

Synthetic model membranes are powerful tools in the investigation of the fundamental properties and functions of biomembranes. Giant unilamellar vesicles (GUVs) are one of the most prominent platforms to study a variety of plasma membrane properties and can be engineered to mimic different physiological conditions1,2,3,4,5,6,7,8. It is well established that the plasma membrane and its organization play a key role in a multitude of cellular processes, such as signal transduction, adhesion, endocytosis, and transport9,10,11,12,13,14,15.
GUVs have been fabricated using various methods, including gentle hydration16, hydrogel swelling17, electroformation18, microfluidic techniques19,20,21,22, jetting23, and solvent exchange24,25,26. Due to challenges in handling integral membrane proteins (IMPs), in vitro platforms to study them have been limited. GUVs present a simplified platform for studying IMPs in an environment that mimics their native environment. Although there have been several approaches for protein reconstitution in GUVs, challenges arise from incorporating proteins with the correct orientation and maintaining protein functionality27.
Most successful protein-reconstitution in GUVs requires the detergent exchange method; which involves solubilizing the proteins from their native environment by detergents, followed by protein purification, and then replacing the detergent molecules with lipids through various methods28. While detergents serve to stabilize the tertiary structure of IMPs during purification, detergent micelles are a relatively unnatural environment for these proteins, which are better stabilized, particularly for functional studies, in lipid bilayers28,29,30. Moreover, incorporating functional transmembrane proteins into the lipid bilayer using traditional GUV fabrication techniques has been difficult due to the size, the delicacy of these proteins, and the additional detergent exchange steps that would be needed27,31,32,33. The use of organic solvent to remove detergents causes protein aggregation and denaturing34. An improved detergent-mediated method has been promising, however, caution is needed for the detergent removal step and optimization might be needed for specific proteins31,35. Additionally, methods that utilize electroformation could restrict the choice of protein and may not be suitable for all lipid compositions especially charged lipids31,36,37. Another technique that has been used is peptide-induced fusion of large unilamellar vesicles (LUVs) containing the desired protein with GUVs, though it was found to be laborious and can lead to the insertion of foreign molecules-the fusogenic peptides33,38,39. Giant plasma membrane vesicles (GPMVs), which are derived from living cells, can be used to overcome some of these issues, however they allow minimal control of the resultant lipid and protein composition14,40,41. Therefore, the integration of IMPs in the bilipid layer of GUVs using our modified agarose swelling method presents a reliable method to further examine these proteins in the membrane environment42,43,44,45.
Cellular signaling and communication involves a family of proteins known as G protein-coupled receptors (GPCRs); GPCRs are among the largest family of proteins and are associated with modulating mood, appetite, blood pressure, cardiovascular function, respiration, and sleep among many other physiological functions46. In this study, we used human serotonin 1A receptor (5-HT1AR) which is a prototypical member of the GPCR family. 5-HT1AR can be found in the central nervous system (CNS) and blood vessels; it influences numerous functions such as cardiovascular, gastrointestinal, endocrine functions, as well as participating in the regulation of mood47. A large barrier to GPCR research arises from their complex amphiphilic structure, and GUVs present a promising platform for the investigation of various properties of interest, ranging from protein functionality, lipid-protein interactions, and protein-protein interactions. Various approaches have been utilized to study lipid-protein interactions such as surface plasmon resonance (SPR)48,49, nuclear magnetic resonance spectroscopy (NMR)50,51, protein lipid overlay (PLO) assay51,52,53,54, native mass spectrometry55, isothermal titration calorimetry (ITC)56,57, and liposome sedimentation assay58,59. Our lab has used the simplified GUV approach to investigate the effect of lipid-protein interactions on protein functionality by incapsulating BODIPY-GTP&#947;S, which binds with the Gi&#945; subunit in the active state of the receptor. Their binding unquenches the fluorophore producing a fluorescence signal that could be detected over time45. Moreover, various studies investigated Lipid-protein interactions and the role of proteins in sensing or stabilizing membrane curvature60,61, and utilizing a feasible GUV approach could be a key advantage.
This protocol demonstrates a straightforward method to incorporate GPCRs into the membrane of GUVs using a modified agarose hydrogel system17,42. Furthermore, based on our previous work, our method could be suitable for IMPs that can bear short-term exposure to 30-40 &#176;C. Briefly, we spread a thin film of agarose combined with membrane fragments containing the GPCR of interest. Following gelation of this layer, we deposit a lipid solution atop the agarose and allow the solvent to evaporate. Rehydration of the system was then performed with an aqueous buffer, resulting in the formation of GUVs with protein incorporated in the lipid bilayer.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC)</td>
			<td>Avanti Polar Lipids, Alabaster, AL</td>
			<td>850375C-25mg</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;TI-Eclipse inverted microscope</td>
			<td>Nikon, Melville, NY</td>
			<td>Eclipse Ti</td>
			<td></td>
		</tr>
		<tr>
			<td>1,2-dipalmitoyl-sn-glycero-3-phosphatidylcholine (DPPC)</td>
			<td>Avanti Polar Lipids, Alabaster, AL</td>
			<td>850355C-25mg</td>
			<td></td>
		</tr>
		<tr>
			<td>13/16&#8243; ID, 1&#8243; OD silicon O-rings</td>
			<td>Sterling Seal &#38; Supply, Neptune, IN</td>
			<td>5-003-8770</td>
			<td></td>
		</tr>
		<tr>
			<td>16-bit Cascade II 512 electron-multiplied charge coupled device camera</td>
			<td>Photometrics, Huntington Beach, CA</td>
			<td>&#160;Cascade II 512</td>
			<td></td>
		</tr>
		<tr>
			<td>1-palmitoyl-2-oleoyl-glycero-3-phosphocholine (POPC)</td>
			<td>Avanti Polar Lipids, Alabaster, AL</td>
			<td>850457C-25mg</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mW solid-state lasers at 488 nm and emission filter centered at 525 nm, and 561 nm with emission filter centered at 595 nm</td>
			<td>Coherent, Santa Clara, CA</td>
			<td>488/561-50-LS</td>
			<td></td>
		</tr>
		<tr>
			<td>5-HT1AR membrane fragments</td>
			<td>Perkin Elmer, Waltham, MA</td>
			<td>RBHS1AM400UA</td>
			<td></td>
		</tr>
		<tr>
			<td>ATTO-488-1,2-Dipalmitoyl-sn-glycero-3-phosphoethanolamine (DPPE)</td>
			<td>ATTO-TEC, Siegen, Germany</td>
			<td>AD 488-155</td>
			<td></td>
		</tr>
		<tr>
			<td>Bench top plasma cleaner</td>
			<td>Harrick Plasma, Ithaca, NY</td>
			<td>PDC-32G</td>
			<td></td>
		</tr>
		<tr>
			<td>bovine serum albumin (BSA)</td>
			<td>Sigma Aldrich, St. Louis, MO</td>
			<td>A9418</td>
			<td></td>
		</tr>
		<tr>
			<td>chloroform (CHCl3)</td>
			<td>Millipore Sigma, Burlington, MA</td>
			<td>CX1055</td>
			<td></td>
		</tr>
		<tr>
			<td>Cholesterol (Chol)</td>
			<td>Sigma Aldrich, St. Louis, MO</td>
			<td>C8667-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning 96-well Flat Clear Bottom</td>
			<td>Corning, Corning, NY</td>
			<td>3904</td>
			<td></td>
		</tr>
		<tr>
			<td>Elmasonic E-Series E15H Ultrasonic</td>
			<td>Elma, Singen, Germany</td>
			<td>[no longer sold on main website]</td>
			<td></td>
		</tr>
		<tr>
			<td>glucose</td>
			<td>Sigma Aldrich, St. Louis, MO</td>
			<td>G7528</td>
			<td></td>
		</tr>
		<tr>
			<td>methanol (MeOH)</td>
			<td>Millipore Sigma, Burlington, MA</td>
			<td>MX0485</td>
			<td></td>
		</tr>
		<tr>
			<td>NanoDrop ND-1000</td>
			<td>Thermo Fisher Scientific, Waltham, MA</td>
			<td>ND-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>NHS-Rhodamine</td>
			<td>Thermo Fisher Scientific, Waltham, MA</td>
			<td>46406</td>
			<td></td>
		</tr>
		<tr>
			<td>phosphate buffered saline (PBS) (10x PBS)</td>
			<td>Corning, Corning, NY</td>
			<td>21-040</td>
			<td></td>
		</tr>
		<tr>
			<td>spinning-disc CSUX confocal head</td>
			<td>Yokogawa,Tokyo, Japan</td>
			<td>CSU-X1</td>
			<td></td>
		</tr>
		<tr>
			<td>standard 25 mm no. 1 glass coverslips</td>
			<td>ChemGlass, Vineland, NJ</td>
			<td>CLS-1760</td>
			<td></td>
		</tr>
		<tr>
			<td>sucrose</td>
			<td>Sigma Aldrich, St. Louis, MO</td>
			<td>S7903</td>
			<td></td>
		</tr>
		<tr>
			<td>Sykes-Moore chambers</td>
			<td>Bellco, Vineland, NJ</td>
			<td>1943-11111</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultra-low melting temperature agarose</td>
			<td>Sigma Aldrich, St. Louis, MO</td>
			<td>A5030</td>
			<td></td>
		</tr>
		<tr>
			<td>VWR Analog Heatblock</td>
			<td>VWR International, Radnor, PA</td>
			<td>[no longer sold on main website]</td>
			<td></td>
		</tr>
		<tr>
			<td>VWR Tube Rotator</td>
			<td>VWR International, Radnor, PA</td>
			<td>10136-084</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeba Spin Desalting Columns, 7K MWCO, 0.5 mL</td>
			<td>Thermo Fisher Scientific, Waltham, MA</td>
			<td>89882</td>
			<td></td>
		</tr>
	</tbody>
</table>

construction of model lipid membranes incorporating g-protein coupled receptors (gpcrs)

Robust in vitro investigations of the structure and function of integral membrane proteins has been a challenge due to the complexities of the plasma membrane and the numerous factors that influence protein behavior in live cells. Giant unilamellar vesicles (GUVs) are a biomimetic and highly tunable in vitro model system for investigating protein-membrane interactions and probing protein behavior in a precise, stimulus-dependent manner. In this protocol, we present an inexpensive and effective method for fabricating GUVs with the human serotonin 1A receptor (5-HT1AR) stably integrated in the membrane. We fabricate GUVs using a modified hydrogel swelling method; by depositing a lipid film on top of a mixture of agarose and 5-HT1AR and then hydrating the entire system, vesicles can be formed with properly oriented and functional 5-HT1AR incorporated into the membrane. These GUVs can then be used to examine protein-membrane interactions and localization behavior via microscopy. Ultimately, this protocol can advance our understanding of the functionality of integral membrane proteins, providing profound physiological insight.

The concentration of protein was measured, and the degree of labeling was calculated as the molar ratio between the dye and the protein to be 1:1. By examining the GUVs using confocal microscopy, we were able to confirm successful formation and protein integration of the vesicles. The lipids were labeled with 0.4 mol% ATTO 488-DPPE, and the protein was covalently labeled via rhodamine NHS-ester modification of primary amines. Figure 2a and Figure 2b show a prote...

Watch this Scientific Journal Video about Construction of Model Lipid Membranes Incorporating G-protein Coupled Receptors GPCRs at JoVE.com

Construction of Model Lipid Membranes Incorporating G-protein Coupled Receptors GPCRs

This protocol utilizes agarose swelling as a powerful and generalizable technique for incorporating integral membrane proteins (IMPs) into giant unilamellar lipid vesicles (GUVs), as described here for the reconstitution of the human 1A serotonin receptor protein (5-HT1AR), one of the classes of pharmacologically important G protein-coupled receptors.

Construction of Model Lipid Membranes Incorporating G-protein Coupled Receptors (GPCRs)

Robust in vitro investigations of the structure and function of integral membrane proteins has been a challenge due to the complexities of the plasma ...

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3.4K Views.  University of Southern California. This protocol utilizes agarose swelling as a powerful and generalizable technique for incorporating integral membrane proteins (IMPs) into giant unilamellar lipid vesicles (GUVs), as described here for the reconstitution of the human 1A serotonin receptor protein (5-HT1AR), one of the classes of pharmacologically important G protein-coupled receptors.

Video: Construction of Model Lipid Membranes Incorporating G-protein Coupled Receptors GPCRs

Use of Label-free Optical Biosensors to Detect Modulation of Potassium Channels by G-protein Coupled Receptors

Ion channels control the electrical properties of neurons and other excitable cell types by selectively allowing ions to flow through the plasma membrane1. To regulate neuronal excitability, the biophysical properties of ion channels are modified by signaling proteins and molecules, which often bind to the channels themselves to form a heteromeric channel complex2,3. Traditional assays examining the interaction between channels and regulatory proteins require exogenous labels that can potentially alter the protein&#39;s behavior and decrease the physiological relevance of the target, while providing little information on the time course of interactions in living cells. Optical biosensors, such as the X-BODY Biosciences BIND Scanner system, use a novel label-free technology, resonance wavelength grating (RWG) optical biosensors, to detect changes in resonant reflected light near the biosensor. This assay allows the detection of the relative change in mass within the bottom portion of living cells adherent to the biosensor surface resulting from ligand induced changes in cell adhesion and spreading, toxicity, proliferation, and changes in protein-protein interactions near the plasma membrane. RWG optical biosensors have been used to detect changes in mass near the plasma membrane of cells following activation of G protein-coupled receptors (GPCRs), receptor tyrosine kinases, and other cell surface receptors. Ligand-induced changes in ion channel-protein interactions can also be studied using this assay. In this paper, we will describe the experimental procedure used to detect the modulation of Slack-B sodium-activated potassium (KNa) channels by GPCRs.

Optical biosensor techniques can detect changes in mass near the plasma membrane in living cells and allow one to follow cellular responses in both individual cells and populations of cells. This protocol will describe detection of the modulation of potassium channels by G-protein coupled receptors in intact cells using this approach.

Ion channels control the electrical properties of neurons and other excitable cell types by selectively allowing ions to flow through the plasma ...

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 ...

Construction and Characterization of a Novel Vocal Fold Bioreactor

In vitro engineering of mechanically active tissues requires the presentation of physiologically relevant mechanical conditions to cultured cells. To emulate the dynamic environment of vocal folds, a novel vocal fold bioreactor capable of producing vibratory stimulations at fundamental phonation frequencies is constructed and characterized. The device is composed of a function generator, a power amplifier, a speaker selector and parallel vibration chambers. Individual vibration chambers are created by sandwiching a custom-made silicone membrane between a pair of acrylic blocks. The silicone membrane not only serves as the bottom of the chamber but also provides a mechanism for securing the cell-laden scaffold. Vibration signals, generated by a speaker mounted underneath the bottom acrylic block, are transmitted to the membrane aerodynamically by the oscillating air. Eight identical vibration modules, fixed on two stationary metal bars, are housed in an anti-humidity chamber for long-term operation in a cell culture incubator. The vibration characteristics of the vocal fold bioreactor are analyzed non-destructively using a Laser Doppler Vibrometer (LDV). The utility of the dynamic culture device is demonstrated by culturing cellular constructs in the presence of 200-Hz sinusoidal vibrations with a mid-membrane displacement of 40 &#181;m. Mesenchymal stem cells cultured in the bioreactor respond to the vibratory signals by altering the synthesis and degradation of vocal fold-relevant, extracellular matrix components. The novel bioreactor system presented herein offers an excellent in vitro platform for studying vibration-induced mechanotransduction and for the engineering of functional vocal fold tissues.

A novel vocal fold bioreactor capable of delivering physiologically relevant, vibratory stimulation to cultured cells is constructed and characterized. This dynamic culture device, when combined with a fibrous poly(&#949;-caprolactone) scaffold, creates a vocal fold-mimetic environment that modulates the behaviors of mesenchymal stem cells.

In vitro engineering of mechanically active tissues requires the presentation of physiologically relevant mechanical conditions to cultured cells. To ...

Preparation of Light-responsive Membranes by a Combined Surface Grafting and Postmodification Process

In order to modify the surface tension of commercial available track-edged polymer membranes, a procedure of surface-initiated polymerization is presented. The polymerization from the membrane surface is induced by plasma treatment of the membrane, followed by reacting the membrane surface with a methanolic solution of 2-hydroxyethyl methacrylate (HEMA). Special attention is given to the process parameters for the plasma treatment prior to the polymerization on the surface. For example, the influence of the plasma-treatment on different types of membranes (e.g.&#160;polyester, polycarbonate, polyvinylidene fluoride) is studied. Furthermore, the time-dependent stability of the surface-grafted membranes is shown by contact angle measurements. When grafting poly(2-hydroxyethyl methacrylate) (PHEMA) in this way, the surface can be further modified by esterification of the alcohol moiety of the polymer with a carboxylic acid function of the desired substance. These reactions can therefore be used for the functionalization of the membrane surface. For example, the surface tension of the membrane can be changed or a desired functionality as the presented light-responsiveness can be inserted. This is demonstrated by reacting PHEMA with a carboxylic acid functionalized spirobenzopyran unit which leads to a light-responsive membrane. The choice of solvent plays a major role in the postmodification step and is discussed in more detail in this paper. The permeability measurements of such functionalized membranes are performed using a Franz cell with an external light source. By changing the wavelength of the light from the visible to the UV-range, a change of permeability of aqueous caffeine solutions is observed.

A plasma-induced polymerization procedure is described for the surface-initiated polymerization on polymer membranes. Further postmodification of the grafted polymer with photochromic substances is presented with a protocol of conducting permeability measurements of light-responsive membranes.

In order to modify the surface tension of commercial available track-edged polymer membranes, a procedure of surface-initiated polymerization is ...

Construction of Modular Hydrogel Sheets for Micropatterned Macro-scaled 3D Cellular Architecture

Hydrogels can be patterned at the micro-scale using microfluidic or micropatterning technologies to provide an in vivo-like three-dimensional (3D) tissue geometry. The resulting 3D hydrogel-based cellular constructs have been introduced as an alternative to animal experiments for advanced biological studies, pharmacological assays and organ transplant applications. Although hydrogel-based particles and fibers can be easily fabricated, it is difficult to manipulate them for tissue reconstruction. In this video, we describe a fabrication method for micropatterned alginate hydrogel sheets, together with their assembly to form a macro-scale 3D cell culture system with a controlled cellular microenvironment. Using a mist form of the calcium gelling agent, thin hydrogel sheets are easily generated with a thickness in the range of 100 - 200 &#181;m, and with precise micropatterns. Cells can then be cultured with the geometric guidance of the hydrogel sheets in freestanding conditions. Furthermore, the hydrogel sheets can be readily manipulated using a micropipette with an end-cut tip, and can be assembled into multi-layered structures by stacking them using a patterned polydimethylsiloxane (PDMS) frame. These modular hydrogel sheets, which can be fabricated using a facile process, have potential applications of in vitro drug assays and biological studies, including functional studies of micro- and macrostructure and tissue reconstruction.

We describe the fabrication of micropatterned hydrogel sheets using a simple process, which can be assembled and manipulated in a freestanding form. Using these modular hydrogel sheets, a simple macro-scaled 3D cell culture system can be generated with a controlled cellular microenvironment.

Hydrogels can be patterned at the micro-scale using microfluidic or micropatterning technologies to provide an in vivo-like three-dimensional (3D) tissue ...

Synthesis of Hydrogels with Antifouling Properties As Membranes for Water Purification

Hydrogels have been widely utilized to enhance the surface hydrophilicity of membranes for water purification, increasing the antifouling properties and thus achieving stable water permeability through membranes over time. Here, we report a facile method to prepare hydrogels based on zwitterions for membrane applications. Freestanding films can be prepared from sulfobetaine methacrylate (SBMA) with a crosslinker of poly(ethylene glycol) diacrylate (PEGDA) via photopolymerization. The hydrogels can also be prepared by impregnation into hydrophobic porous supports to enhance the mechanical strength. These films can be characterized by attenuated total reflection Fourier transform infrared spectroscopy (ATR-FTIR) to determine the degree of conversion of the (meth)acrylate groups, using goniometers for hydrophilicity and differential scanning calorimetry (DSC) for polymer chain dynamics. We also report protocols to determine the water permeability in dead-end filtration systems and the effect of foulants (bovine serum albumin, BSA) on membrane performance.

This paper reports practical methods to prepare hydrogels in freestanding films and impregnated membranes and to characterize their physical properties, including water transport properties.

Hydrogels have been widely utilized to enhance the surface hydrophilicity of membranes for water purification, increasing the antifouling properties and ...

Construction and Setup of a Bench-scale Algal Photosynthetic Bioreactor with Temperature, Light, and pH Monitoring for Kinetic Growth Tests

The optimal design and operation of photosynthetic bioreactors (PBRs) for microalgal cultivation is essential for improving the environmental and economic performance of microalgae-based biofuel production. Models that estimate microalgal growth under different conditions can help to optimize PBR design and operation. To be effective, the growth parameters used in these models must be accurately determined. Algal growth experiments are often constrained by the dynamic nature of the culture environment, and control systems are needed to accurately determine the kinetic parameters. The first step in setting up a controlled batch experiment is live data acquisition and monitoring. This protocol outlines a process for the assembly and operation of a bench-scale photosynthetic bioreactor that can be used to conduct microalgal growth experiments. This protocol describes how to size and assemble a flat-plate, bench-scale PBR from acrylic. It also details how to configure a PBR with continuous pH, light, and temperature monitoring using a data acquisition and control unit, analog sensors, and open-source data acquisition software.

This paper describes the assembly process and operation of a bench-scale photosynthetic bioreactor that can be used, in conjunction with other methods, to estimate pertinent kinetic growth parameters. This system continuously monitors the pH, light, and temperature using sensors, a data acquisition and control unit, and open-source data acquisition software.

The optimal design and operation of photosynthetic bioreactors (PBRs) for microalgal cultivation is essential for improving the environmental and economic ...

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 ...

Assembly and Characterization of Biomolecular Memristors Consisting of Ion Channel-doped Lipid Membranes

The ability to recreate synaptic functionalities in synthetic circuit elements is essential for neuromorphic computing systems that seek to emulate the cognitive powers of the brain with comparable efficiency and density. To date, silicon-based three-terminal transistors and two-terminal memristors have been widely used in neuromorphic circuits, in large part due to their ability to co-locate information processing and memory. Yet these devices cannot achieve the interconnectivity and complexity of the brain because they are power-hungry, fail to mimic key synaptic functionalities, and suffer from high noise and high switching voltages. To overcome these limitations, we have developed and characterized a biomolecular memristor that mimics the composition, structure, and switching characteristics of biological synapses. Here, we describe the process of assembling and characterizing biomolecular memristors consisting of a 5 nm-thick lipid bilayer formed between lipid-functionalized water droplets in oil and doped with voltage-activated alamethicin peptides. While similar assembly protocols have been used to investigate biophysical properties of droplet-supported lipid membranes and membrane-bound ion channels, this article focuses on key modifications of the droplet interface bilayer method essential for achieving consistent memristor performance. Specifically, we describe the liposome preparation process and the incorporation of alamethicin peptides in lipid bilayer membranes, and the appropriate concentrations of each constituent as well as their impact on the overall response of the memristors. We also detail the characterization process of biomolecular memristors, including measurement and analysis of memristive current-voltage relationships obtained via cyclic voltammetry, as well as short-term plasticity and learning in response to step-wise voltage pulse trains.

Soft, low-power, biomolecular memristors leverage similar composition, structure, and switching mechanisms of bio-synapses. Presented here is a protocol to assemble and characterize biomolecular memristors obtained from insulating lipid bilayers formed between water droplets in oil. The incorporation of voltage-activated alamethicin peptides results in memristive ionic conductance across the membrane.

The ability to recreate synaptic functionalities in synthetic circuit elements is essential for neuromorphic computing systems that seek to emulate the ...

Growth of Human and Sheep Corneal Endothelial Cell Layers on Biomaterial Membranes

Corneal endothelial cell cultures have a tendency to undergo epithelial-to-mesenchymal transition (EMT) after loss of cell-to-cell contact. EMT is deleterious for the cells as it reduces their ability to form a mature and functional layer. Here, we present a method for establishing and subculturing human and sheep corneal endothelial cell cultures that minimizes the loss of cell-to-cell contact. Explants of corneal endothelium/Descemet&#39;s membrane are taken from donor corneas and placed into tissue culture under conditions that allow the cells to collectively migrate onto the culture surface. Once a culture has been established, the explants are transferred to fresh plates to initiate new cultures. Dispase II is used to gently lift clumps of cells off tissue culture plates for subculturing. Corneal endothelial cell cultures that have been established using this protocol are suitable for transferring to biomaterial membranes to produce tissue-engineered cell layers for transplantation in animal trials. A custom-made device for supporting biomaterial membranes during tissue culture is described and an example of a tissue-engineered graft composed of a layer of corneal endothelial cells and a layer of corneal stromal cells on either side of a collagen type I membrane is presented.

This protocol describes the critical steps required to establish and grow corneal endothelial cell cultures from explants of human or sheep tissue. A method for subculturing corneal endothelial cells on membranous biomaterials is also presented.