APL acknowledges support by a Humboldt Research Fellowship for Experienced Researchers and from the National Science Foundation (1939310 and 1817909) and National Institutes of Health (R01 EB030031).

1. Preparation of oil-lipid-mixture
NOTE: The step needs to be performed in a fume hood following all the safety guidelines for handling chloroform.
<ol>
	<li>Take 0.5 mL of chloroform in a 15 mL glass vial. Add 88 &#181;L of 25 mg/mL dioleoyl-phosphocholine (DOPC), 9.3 &#181;L of 50 mg/mL cholesterol, and 5 &#181;L of 1 mg/mL dioleoyl-phosphoethanolamine-lissamine rhodamine B (rhodamine PE) (see Table of Materials) into the 15 ml glass vial. 
	NOTE: The final mole fractions of DOPC and cholesterol in silicone oil/mineral oil are 69.9% and 30%, respectively. It was established that 20-30 mol% cholesterol is an optimized concentration for membrane fluidity and stability of GUVs generated using the presented technique25,26. Physiologically, these values are well within the cholesterol concentration range found in mammalian cell plasma membranes6. Lipid stocks are acquired as solutions in chloroform and stored at -20 &#176;C. Lipid stock vials should be acclimated to room temperature before they are opened.</li>
	<li>Pipette 7.2 mL of silicone oil and 1.8 mL of mineral oil in a second 15 mL vial (see Table of Materials). Generally, this needs to be done in a low-humidity glove box, mainly if the mineral oil is reused.</li>
	<li>Mix the oils by vortexing at the maximum rotational speed (3200 rpm) for 10 s, add the mixture to the vial containing the lipid-in-chloroform mixture and immediately place it on the vortex mixer. Vortex for 10-15 s at the maximum rotational speed (3200 rpm). The resulting lipids-in-oil mix should be slightly cloudy, as the lipids are not fully dissolved in the oil but rather dispersed as small aggregates24.</li>
	<li>Put the lipid-in-oil dispersion in a bath sonicator (see Table of Materials) with ultrasonic power of 80 W and operating frequency of 40 kHz at room temperature for 30 min. Use the mixture immediately or store at 4 &#176;C for a maximum of 24 h.</li>
</ol>
2. Vesicle generation
<ol>
	<li>Mount the 3D-printed shaft (Supplementary File 1) made from black resin (see Table of Materials) on the benchtop stir plate and set rotational speed to 1200 rpm.</li>
	<li>Mount the 3D-printed cDICE chamber (Supplementary File 2) made from clear resin (see Table of Materials) on the shaft (Figure 1A,B).</li>
	<li>Prepare actin and actin-binding proteins (ABP) solutions separately in a total volume of 20 &#181;L.
	<ol>
		<li>Prepare 1-10 &#181;M of actin in globular actin buffer (G-buffer), including 10% ATTO 488 actin (see Table of Materials). 
		NOTE: 1x G-buffer comprises 5 mM of Tris-HCl, pH 8.0, and 0.2 mM of CaCl2.</li>
		<li>Add filamentous actin polymerization buffer (F-buffer) to begin actin polymerization on ice. Keep the solutions on ice to slow down actin polymerization before the addition of a crosslinker. 
		NOTE: The 1x F-buffer contains 50 mM of KCl, 2 mM of MgCl2, and 3 mM of ATP in 10&#160;mM of Tris, pH 7.5.</li>
		<li>Wait for 15 min to allow for initiation of actin polymerization on ice before adding crosslinkers of interest at the desired molar ratio. Keep the solution on ice until encapsulation.</li>
		<li>Prepare actin-binding proteins (ABPs) separately in a microtube (Figure 1C). 
		NOTE: This step is performed when a combination of ABPs (e.g., myosin, &#945;-actinin, fascin, Arp2/3 complex) encapsulates with actin25,26,27. In such cases, the desired amount of each ABP is drawn from its stock and added into the microtube designated for ABPs. As the total solution is to be 20 &#181;L, stock aliquots of ABPs must be prepared so that the desired amount of the total ABP mixture does not exceed 5-6 &#181;L. The only ABP used in the representative results here is fascin, the preparation of which is mentioned below.
		<ol>
			<li>Aliquot 1.57 &#181;L of fascin from a 1.75 mg/mL of fascin stock (see Table of Materials), and directly add to the actin solution in step 2.7. This corresponds to 2.5 &#181;M of fascin in the solution.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Add 7.5% of density gradient medium (see Table of Materials) into the actin solution to create a density gradient between the outer and the inner aqueous phase to facilitate GUV sedimentation.</li>
	<li>Dispense 700 &#181;L of the outer solution of 200 mM of glucose into the chamber (Figure 1D, left). 
	NOTE: The osmolarity of the inner solution determines the concentration of glucose. For these experiments, the osmolarity of the inner solution is ~200 mOsm, so a 200 mM glucose solution is used as the outer solution.</li>
	<li>Add a sufficient amount of lipid-oil mixture (&#62;3 mL based on the chamber size) into the chamber until 60%-80% of the chamber is filled (Figure 1D, right). An interface will be formed between the lipid-oil mixture and the outer solution.</li>
	<li>Transfer ABPs (prepared in step 2.3.4) into the actin solution. Using a regular 100-1000 &#181;L pipette, immediately transfer the 700 &#181;L of the lipid-oil mixture into the actin-ABP mixture (Figure 1E, left). Pipette up and down 8 times to generate cell-sized lipid-monolayer droplets with diameter in the range of 7-100 &#181;m (Figure 1E, middle). 
	NOTE: Step 2.7 needs to be completed in a few seconds to avoid any actin network assembly before encapsulation. Thus, before performing the step, ensure that pipette tips are already inserted into the pipettes and ready to transfer the mixtures.</li>
	<li>Using the same 100-1000 &#181;L pipette, immediately dispense the entire emulsion into the rotating chamber. Droplets will acquire a second leaflet of lipids by crossing the lipid monolayer at the oil-outer solution interface, thereby forming GUVs (Figure 1E, right).</li>
	<li>Remove the chamber from the stir plate and discard most of the lipid-oil mixture by tilting the chamber in the waste container so that a large portion of the lipid-oil mixture is drained from the large opening at the center of the chamber. 
	NOTE: This way, the lipid-oil mixture is drawn off the chamber, avoiding mixing of lipid-oil mixture with the outer solution in the next step.</li>
	<li>Hold the chamber with its lid facing towards the user. Open the chamber lid and slightly tilt the chamber towards the user. The interface between the outer solution containing GUVs and the lipid-oil mixture is visible from the chamber opening (where the lid is located).</li>
	<li>Using a pipette, collect enough outer solution containing GUVs and dispense 50-300 &#181;L of the outer solution into a 96-well plate to obtain an appropriate density of GUVs. 
	NOTE: Following this protocol, a total of about 2 x 105 GUVs are released in the outer solution inside the chamber. The GUV dispersity was not quantified; however, the diameter of about 90% of GUVs is in the range of 12-30&#160;&#181;m. GUVs with any diameter in the range of 7-50 &#181;m can be found in the population. Depending on the encapsulated density gradient medium, GUV size, and solution depth in the well plate, it takes 2-15 min for GUVs to settle down on the surface. The yield of GUVs with reconstituted actin bundles is about 90%.</li>
</ol>
3. Imaging and 3D image reconstruction
<ol>
	<li>Set up the 96 well plate on the stage of an inverted microscope equipped with a spinning disk (or laser scanning) confocal unit, an EMCCD or an sCMOS camera, and an oil immersion 60x objective lens (see Table of Materials).</li>
	<li>Focus on any region of interest (ROI) and take a z-stack image sequence from the ROI with a z-step interval of 0.5 &#181;m. 
	NOTE: Because GUVs can be slightly displaced on the surface over time, it is recommended to capture a multi-wavelength set of images at each z-plane if multiple fluorophores are being imaged, i.e., take a group of 561 nm and 488 nm images at each z-lane at a time to capture ATTO 488 actin and Rhod PE images.</li>
	<li>Save each z-stack image sequence in .tiff format.</li>
	<li>Open an image sequence of interest in an image processing software (ImageJ/Fiji). Identify the image with the highest intensity. Hold &#34;ctrl+shift+c&#34; to open the Brightness &#38; Contrast window and click on Reset.</li>
	<li>From the ImageJ/Fiji menu, go to Analyze &#62; Set Scale and enter the known physical distance and its unit for each image pixel.</li>
	<li>From the ImageJ/Fiji menu, go to Image &#62; Stacks &#62; 3D project to reconstruct a 3D image from the z-stack. Set &#34;Projection method&#34; as Brightness Point, &#34;Slice Spacing (&#181;m)&#34; as 0.5, and checkmark Interpolate. The default options can be used for the rest of the settings. 
	NOTE: z-intervals in some microscopes might not be calibrated, and the actual movement in z-direction might slightly differ from the entered z-interval (i.e., 0.5 &#181;m). In such cases, a 3D calibration specimen such as fluorescent microspheres with a known diameter can be used to obtain the actual z-interval. This value will thus be used as &#34;Slice Spacing (&#181;m)&#34; for 3D projection.</li>
</ol>

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The authors declare no conflicts of interest.

Different methods of generating GUVs have been explored for the creation of synthetic cells&#160;However, the complexity of the procedures, extended time to attain encapsulation, restriction of lipid types and molecular composition of the encapsulant, need for non-physiological chemicals to facilitate encapsulation, low GUV yield, and inconsistencies in encapsulation efficiency have continued to challenge researchers in this field. Considering the wide range of potential studies that can be embarked in bottom-up syntheti...

Lipid bilayer compartments are used as model synthetic cells for studying enclosed organic reactions and membrane-based processes or as carrier modules in drug delivery applications1,2. Bottom-up biology with purified components requires minimal experimental systems to explore properties and interactions between biomolecules, such as proteins and lipids3,4. However, with the advancement of the field, there is an increased need for more complex experimental systems that better imitate the conditions in biological cells. Encapsulation in GUVs is a practical approach that can offer some of these cell-like properties by providing a deformable and selectively permeable lipid bilayer and a confined reaction space. In particular, in vitro reconstitution of cytoskeletal systems, as models of synthetic cells, can benefit from encapsulation in membrane compartments5. Many cytoskeletal proteins bind and interact with the cell membrane. As most cytoskeletal assemblies form structures that span the entirety of the cell, their shape is naturally determined by cell-sized confinement6.
Different methods are used to generate GUVs, such as the swelling7,8, small vesicle fusion9,10, emulsion transfer11,12, pulsed jetting13, and other microfluidic approaches14,15. Although these methods are still utilized, each has its limitations. Thus, a robust and straightforward approach with a high yield of GUV encapsulation is highly desirable. Although techniques such as spontaneous swelling and electroswelling are widely adopted for the formation of GUVs, these methods are primarily compatible with specific lipid compositions16, low salt concentration buffers17, smaller encapsulant molecular size18, and require a high volume of the encapsulant. Fusing multiple small vesicles into a GUV is inherently energetically unfavorable, thus requiring specificity in charged lipid compositions9 and/or external fusion-inducing agents, such as peptides19&#160;or other chemicals. Emulsion transfer and microfluidic methods, on the other hand, may require droplet stabilization through surfactant and solvent removal after bilayer formation, respectively18,20. The complexity of experimental setup and device in microfluidic techniques such as pulsed jetting impose an additional challenge21. cDICE is an emulsion-based method derived from similar principles governing emulsion transfer22,23. An aqueous solution (outer solution) and a lipid-oil mixture are stratified by centrifugal forces in a rotating cylindrical chamber (cDICE chamber) forming a lipid saturated interface. Shuttling lipid monolayered aqueous droplets into the rotating cDICE chamber results in zipping of a bilayer as droplets cross the lipid-saturated interface into the outer aqueous solution22,24. The cDICE approach is a robust technique for GUV encapsulation. With the presented modified method, not only the high vesicle yield typical for cDICE with a significantly shorter encapsulation time (a few seconds) is achieved but GUV generation time that allow for the observation of time-dependent processes (e.g., actin cytoskeletal network formation) is significantly reduced. The protocol takes about 15-20 min from the start to GUV collection and imaging. Here, GUV generation is described using the modified cDICE method for encapsulating actin and actin-binding proteins (ABPs). However, the presented technique is applicable for encapsulating a wide range of biological reactions and membrane interactions, from the assembly of biopolymers to cell-free protein expression to membrane fusion-based cargo transfer.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>18:1 Liss Rhod PE lipid in chloroform</td>
			<td>Avanti Polar Lipids</td>
			<td>810150C</td>
			<td></td>
		</tr>
		<tr>
			<td>96 Well Optical Btm Pit PolymerBase</td>
			<td>ThermoFisher Scientific</td>
			<td>165305</td>
			<td></td>
		</tr>
		<tr>
			<td>Actin from rabbit skeletal muscle</td>
			<td>Cytoskeleton</td>
			<td>AKL99-A</td>
			<td></td>
		</tr>
		<tr>
			<td>ATTO 488-actin from rabbit skeletal muscle</td>
			<td>Hypermol</td>
			<td>8153-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Axygen microtubes (200 &#181;L)</td>
			<td>Fisher Scientific</td>
			<td>14-222-262</td>
			<td>for handling ABPs</td>
		</tr>
		<tr>
			<td>Black resin</td>
			<td>Formlabs</td>
			<td>RS-F2-GPBK-04</td>
			<td></td>
		</tr>
		<tr>
			<td>Cholesterol (powder)</td>
			<td>Avanti Polar Lipids</td>
			<td>700100P</td>
			<td></td>
		</tr>
		<tr>
			<td>Choloroform</td>
			<td>Sigma Aldrich</td>
			<td>67-66-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Clear resin</td>
			<td>Formlabs</td>
			<td>RS-F2-GPCL-04</td>
			<td></td>
		</tr>
		<tr>
			<td>CSU-X1 Confocal Scanner Unit</td>
			<td>YOKOGAWA</td>
			<td>CSU-X1</td>
			<td></td>
		</tr>
		<tr>
			<td>Density gradient medium (Optiprep)</td>
			<td>Sigma-Aldrich</td>
			<td>D1556</td>
			<td></td>
		</tr>
		<tr>
			<td>DOPC lipid in chloroform</td>
			<td>Avanti Polar Lipids</td>
			<td>850375C</td>
			<td></td>
		</tr>
		<tr>
			<td>Fascin</td>
			<td>homemade</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>F-buffer</td>
			<td>homemade</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand microtubes (1.5 mL)</td>
			<td>Fisher Scientific</td>
			<td>05-408-129</td>
			<td></td>
		</tr>
		<tr>
			<td>FS02 Sonicator</td>
			<td>Fischer Scientific</td>
			<td>FS20</td>
			<td></td>
		</tr>
		<tr>
			<td>G-buffer</td>
			<td>homemade</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>158968</td>
			<td></td>
		</tr>
		<tr>
			<td>iXon X3 camera</td>
			<td>Andor</td>
			<td>DU-897E-CS0</td>
			<td></td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Acros Organics</td>
			<td>8042-47-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus IX81 Inverted Microscope</td>
			<td>Olympus</td>
			<td>IX21</td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus PlanApo N 60x Oil Microscope Objective</td>
			<td>Olumpus</td>
			<td>1-U2B933</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicone oil</td>
			<td>Sigma-Aldrich</td>
			<td>317667</td>
			<td></td>
		</tr>
	</tbody>
</table>

rapid encapsulation of reconstituted cytoskeleton inside giant unilamellar vesicles

Giant unilamellar vesicles (GUVs) are frequently used as models of biological membranes and thus are a great tool to study membrane-related cellular processes in vitro. In recent years, encapsulation within GUVs has proven to be a helpful approach for reconstitution experiments in cell biology and related fields. It better mimics confinement conditions inside living cells, as opposed to conventional biochemical reconstitution. Methods for encapsulation inside GUVs are often not easy to implement, and success rates can differ significantly from lab to lab. One technique that has proven to be successful for encapsulating more complex protein systems is called continuous droplet interface crossing encapsulation (cDICE). Here, a cDICE-based method is presented for rapidly encapsulating cytoskeletal proteins in GUVs with high encapsulation efficiency. In this method, first, lipid-monolayer droplets are generated by emulsifying a protein solution of interest in a lipid/oil mixture. After being added into a rotating 3D-printed chamber, these lipid-monolayered droplets then pass through a second lipid monolayer at a water/oil interface inside the chamber to form GUVs that contain the protein system. This method simplifies the overall procedure of encapsulation within GUVs and speeds up the process, and thus allows us to confine and observe the dynamic evolution of network assembly inside lipid bilayer vesicles. This platform is handy for studying the mechanics of cytoskeleton-membrane interactions in confinement.

To demonstrate the successful generation of cytoskeletal GUVs using the current protocol, fascin-actin bundle structures in GUVs were reconstituted. Fascin is a short crosslinker of actin filaments which forms stiff parallel-aligned actin bundles and is purified from E. coli as Glutathione-S-Transferase (GST) fusion protein26. 5 &#181;M of actin was first reconstituted, including 0.53 &#181;M of ATTO488 actin in the actin polymerization buffer and 7.5% of the density gradient medium. Upon...

Watch this Scientific Journal Video about Rapid Encapsulation of Reconstituted Cytoskeleton Inside Giant Unilamellar Vesicles at JoVE.com

Rapid Encapsulation of Reconstituted Cytoskeleton Inside Giant Unilamellar Vesicles

This article introduces a simple method for expeditious production of giant unilamellar vesicles with encapsulated cytoskeletal proteins. The method proves to be useful for bottom-up reconstitution of cytoskeletal structures in confinement and cytoskeleton-membrane interactions.

Giant unilamellar vesicles (GUVs) are frequently used as models of biological membranes and thus are a great tool to study membrane-related cellular ...

rapid-encapsulation-reconstituted-cytoskeleton-inside-giant

Research

JoVE Journal

Bioengineering

4.1K Views.  University of Michigan, Ann Arbor. This article introduces a simple method for expeditious production of giant unilamellar vesicles with encapsulated cytoskeletal proteins. The method proves to be useful for bottom-up reconstitution of cytoskeletal structures in confinement and cytoskeleton-membrane interactions. 

Video: Rapid Encapsulation of Reconstituted Cytoskeleton Inside Giant Unilamellar Vesicles

Encapsulation of Cardiomyocytes in a Fibrin Hydrogel for Cardiac Tissue Engineering

Culturing cells in a three dimensional hydrogel environment is an important technique for developing constructs for tissue engineering as well as studying cellular responses under various culture conditions in vitro. The three dimensional environment more closely mimics what the cells observe in vivo due to the application of mechanical and chemical stimuli in all dimensions 1. Three-dimensional hydrogels can either be made from synthetic polymers such as PEG-DA 2 and PLGA 3 or a number of naturally occurring proteins such as collagen 4, hyaluronic acid 5 or fibrin 6,7. Hydrogels created from fibrin, a naturally occurring blood clotting protein, can polymerize to form a mesh that is part of the body's natural wound healing processes 8. Fibrin is cell-degradable and potentially autologous 9, making it an ideal temporary scaffold for tissue engineering.
Here we describe in detail the isolation of neonatal cardiomyocytes from three day old rat pups and the preparation of the cells for encapsulation in fibrin hydrogel constructs for tissue engineering. Neonatal myocytes are a common cell source used for in vitro studies in cardiac tissue formation and engineering 4. Fibrin gel is created by mixing fibrinogen with the enzyme thrombin. Thrombin cleaves fibrinopeptides FpA and FpB from fibrinogen, revealing binding sites that interact with other monomers 10. These interactions cause the monomers to self-assemble into fibers that form the hydrogel mesh. Because the timing of this enzymatic reaction can be adjusted by altering the ratio of thrombin to fibrinogen, or the ratio of calcium to thrombin, one can injection mold constructs with a number of different geometries 11,12. Further we can generate alignment of the resulting tissue by how we constrain the gel during culture 13.
After culturing the engineered cardiac tissue constructs for two weeks under static conditions, the cardiac cells have begun to remodel the construct and can generate a contraction force under electrical pacing conditions 6. As part of this protocol, we also describe methods for analyzing the tissue engineered myocardium after the culture period including functional analysis of the active force generated by the cardiac muscle construct upon electrical stimulation, as well as methods for determining final cell viability (Live-Dead assay) and immunohistological staining to examine the expression and morphology of typical proteins important for contraction (Myosin Heavy Chain or MHC) and cellular coupling (Connexin 43 or Cx43) between myocytes.

We describe the isolation of neonatal cardiomyocytes and the preparation of the cells for encapsulation in fibrin hydrogel constructs for tissue engineering. We describe methods for analyzing the tissue engineered myocardium after the culture period including active force generated upon electrical stimulation and cell viability and immunohistological staining.

Culturing cells in a three dimensional hydrogel environment is an important technique for developing constructs for tissue engineering as well as studying ...

High Throughput Single-cell and Multiple-cell Micro-encapsulation

Microfluidic encapsulation methods have been previously utilized to capture cells in picoliter-scale aqueous, monodisperse drops, providing confinement from a bulk fluid environment with applications in high throughput screening, cytometry, and mass spectrometry. We describe a method to not only encapsulate single cells, but to repeatedly capture a set number of cells (here we demonstrate one- and two-cell encapsulation) to study both isolation and the interactions between cells in groups of controlled sizes. By combining drop generation techniques with cell and particle ordering, we demonstrate controlled encapsulation of cell-sized particles for efficient, continuous encapsulation. Using an aqueous particle suspension and immiscible fluorocarbon oil, we generate aqueous drops in oil with a flow focusing nozzle. The aqueous flow rate is sufficiently high to create ordering of particles which reach the nozzle at integer multiple frequencies of the drop generation frequency, encapsulating a controlled number of cells in each drop. For representative results, 9.9 &mu;m polystyrene particles are used as cell surrogates. This study shows a single-particle encapsulation efficiency Pk=1 of 83.7% and a double-particle encapsulation efficiency Pk=2 of 79.5% as compared to their respective Poisson efficiencies of 39.3% and 33.3%, respectively. The effect of consistent cell and particle concentration is demonstrated to be of major importance for efficient encapsulation, and dripping to jetting transitions are also addressed. 
Introduction 
Continuous media aqueous cell suspensions share a common fluid environment which allows cells to interact in parallel and also homogenizes the effects of specific cells in measurements from the media. High-throughput encapsulation of cells into picoliter-scale drops confines the samples to protect drops from cross-contamination, enable a measure of cellular diversity within samples, prevent dilution of reagents and expressed biomarkers, and amplify signals from bioreactor products. Drops also provide the ability to re-merge drops into larger aqueous samples or with other drops for intercellular signaling studies.1,2 The reduction in dilution implies stronger detection signals for higher accuracy measurements as well as the ability to reduce potentially costly sample and reagent volumes.3 Encapsulation of cells in drops has been utilized to improve detection of protein expression,4 antibodies,5,6 enzymes,7 and metabolic activity8 for high throughput screening, and could be used to improve high throughput cytometry.9 Additional studies present applications in bio-electrospraying of cell containing drops for mass spectrometry10 and targeted surface cell coatings.11 Some applications, however, have been limited by the lack of ability to control the number of cells encapsulated in drops. Here we present a method of ordered encapsulation12 which increases the demonstrated encapsulation efficiencies for one and two cells and may be extrapolated for encapsulation of a larger number of cells.
To achieve monodisperse drop generation, microfluidic "flow focusing" enables the creation of controllable-size drops of one fluid (an aqueous cell mixture) within another (a continuous oil phase) by using a nozzle at which the streams converge.13 For a given nozzle geometry, the drop generation frequency f and drop size can be altered by adjusting oil and aqueous flow rates Qoil and Qaq. As the flow rates increase, the flows may transition from drop generation to unstable jetting of aqueous fluid from the nozzle.14 
When the aqueous solution contains suspended particles, particles become encapsulated and isolated from one another at the nozzle. For drop generation using a randomly distributed aqueous cell suspension, the average fraction of drops Dk containing k cells is dictated by Poisson statistics, where Dk = &lambda;k exp(-&lambda;)/(k!) and &lambda; is the average number of cells per drop. The fraction of cells which end up in the "correctly" encapsulated drops is calculated using Pk = (k x Dk)/&Sigma;(k' x Dk'). The subtle difference between the two metrics is that Dk relates to the utilization of aqueous fluid and the amount of drop sorting that must be completed following encapsulation, and Pk relates to the utilization of the cell sample. As an example, one could use a dilute cell suspension (low &lambda;) to encapsulate drops where most drops containing cells would contain just one cell. While the efficiency metric Pk would be high, the majority of drops would be empty (low Dk), thus requiring a sorting mechanism to remove empty drops, also reducing throughput.15
Combining drop generation with inertial ordering provides the ability to encapsulate drops with more predictable numbers of cells per drop and higher throughputs than random encapsulation. Inertial focusing was first discovered by Segre and Silberberg16 and refers to the tendency of finite-sized particles to migrate to lateral equilibrium positions in channel flow. Inertial ordering refers to the tendency of the particles and cells to passively organize into equally spaced, staggered, constant velocity trains. Both focusing and ordering require sufficiently high flow rates (high Reynolds number) and particle sizes (high Particle Reynolds number).17,18 Here, the Reynolds number Re =uDh/&nu; and particle Reynolds number Rep =Re(a/Dh)2, where u is a characteristic flow velocity, Dh [=2wh/(w+h)] is the hydraulic diameter, &nu; is the kinematic viscosity, a is the particle diameter, w is the channel width, and h is the channel height. Empirically, the length required to achieve fully ordered trains decreases as Re and Rep increase. Note that the high Re and Rep requirements (for this study on the order of 5 and 0.5, respectively) may conflict with the need to keep aqueous flow rates low to avoid jetting at the drop generation nozzle. Additionally, high flow rates lead to higher shear stresses on cells, which are not addressed in this protocol. The previous ordered encapsulation study demonstrated that over 90% of singly encapsulated HL60 cells under similar flow conditions to those in this study maintained cell membrane integrity.12 However, the effect of the magnitude and time scales of shear stresses will need to be carefully considered when extrapolating to different cell types and flow parameters. The overlapping of the cell ordering, drop generation, and cell viability aqueous flow rate constraints provides an ideal operational regime for controlled encapsulation of single and multiple cells.
Because very few studies address inter-particle train spacing,19,20 determining the spacing is most easily done empirically and will depend on channel geometry, flow rate, particle size, and particle concentration. Nonetheless, the equal lateral spacing between trains implies that cells arrive at predictable, consistent time intervals. When drop generation occurs at the same rate at which ordered cells arrive at the nozzle, the cells become encapsulated within the drop in a controlled manner. This technique has been utilized to encapsulate single cells with throughputs on the order of 15 kHz,12 a significant improvement over previous studies reporting encapsulation rates on the order of 60-160 Hz.4,15 In the controlled encapsulation work, over 80% of drops contained one and only one cell, a significant efficiency improvement over Poisson (random) statistics, which predicts less than 40% efficiency on average.12
In previous controlled encapsulation work,12 the average number of particles per drop &lambda; was tuned to provide single-cell encapsulation. We hypothesize that through tuning of flow rates, we can efficiently encapsulate any number of cells per drop when &lambda; is equal or close to the number of desired cells per drop. While single-cell encapsulation is valuable in determining individual cell responses from stimuli, multiple-cell encapsulation provides information relating to the interaction of controlled numbers and types of cells. Here we present a protocol, representative results using polystyrene microspheres, and discussion for controlled encapsulation of multiple cells using a passive inertial ordering channel and drop generation nozzle.

Combining monodisperse drop generation with inertial ordering of cells and particles, we describe a method to encapsulate a desired number of cells or particles in a single drop at kHz rates. We demonstrate efficiencies twice exceeding those of unordered encapsulation for single- and double-particle drops.

Microfluidic encapsulation methods have been previously utilized to capture cells in picoliter-scale aqueous, monodisperse drops, providing confinement ...

The Encapsulation of Cell-free Transcription and Translation Machinery in Vesicles for the Construction of Cellular Mimics

As interest shifts from individual molecules to systems of molecules, an increasing number of laboratories have sought to build from the bottom up cellular mimics that better represent the complexity of cellular life. To date there are a number of paths that could be taken to build compartmentalized cellular mimics, including the exploitation of water-in-oil emulsions, microfluidic devices, and vesicles. Each of the available options has specific advantages and disadvantages. For example, water-in-oil emulsions give high encapsulation efficiency but do not mimic well the permeability barrier of living cells. The primary advantage of the methods described herein is that they are all easy and cheap to implement. Transcription-translation machinery is encapsulated inside of phospholipid vesicles through a process that exploits common instrumentation, such as a centrifugal evaporator and an extruder. Reactions are monitored by fluorescence spectroscopy. The protocols can be adapted for recombinant protein expression, the construction of cellular mimics, the exploration of the minimum requirements for cellular life, or the assembly of genetic circuitry.

Herein we describe simple methods for the preparation of vesicles, the encapsulation of transcription and translation machinery, and the monitoring of protein production. The resulting cell-free systems can be used as a starting point from which to build increasingly complex cellular mimics.

As interest shifts from individual molecules to systems of molecules, an increasing number of laboratories have sought to build from the bottom up ...

Mammalian Cell Encapsulation in Alginate Beads Using a Simple Stirred Vessel

Cell encapsulation in alginate beads has been used for immobilized cell culture in vitro as well as for immunoisolation in vivo. Pancreatic islet encapsulation has been studied extensively as a means to increase islet survival in allogeneic or xenogeneic transplants. Alginate encapsulation is commonly achieved by nozzle extrusion and external gelation. Using this method, cell-containing alginate droplets formed at the tip of nozzles fall into a solution containing divalent cations that cause ionotropic alginate gelation as they diffuse into the droplets. The requirement for droplet formation at the nozzle tip limits the volumetric throughput and alginate concentration that can be achieved. This video describes a scalable emulsification method to encapsulate mammalian cells in 0.5% to 10% alginate with 70% to 90% cell survival. By this alternative method, alginate droplets containing cells and calcium carbonate are emulsified in mineral oil, followed by a decrease in pH leading to internal calcium release and ionotropic alginate gelation. The current method allows the production of alginate beads within 20 min of emulsification. The equipment required for the encapsulation step consists in simple stirred vessels available to most laboratories.

This video and manuscript describe an emulsion-based method to encapsulate mammalian cells in 0.5% to 10% alginate beads which can be produced in large batches using a simple stirred vessel. The encapsulated cells can be cultured in vitro or transplanted for cellular therapy applications.

Cell encapsulation in alginate beads has been used for immobilized cell culture in vitro as well as for immunoisolation in vivo. Pancreatic islet ...

Preparation and Characterization of Novel HDL-mimicking Nanoparticles for Nerve Growth Factor Encapsulation

The objective of this article is to introduce preparation and characterization methods for nerve growth factor (NGF)-loaded, high-density, lipoprotein (HDL)-mimicking nanoparticles (NPs). HDLs are endogenous NPs and have been explored as vehicles for the delivery of therapeutic agents. Various methods have been developed to prepare HDL-mimicking NPs. However, they are generally complicated, time consuming, and difficult for industrial scale-up. In this study, one-step homogenization was used to mix the excipients and form the prototype NPs. NGF is a water-soluble protein of 26 kDa. To facilitate the encapsulation of NGF into the lipid environment of HDL-mimicking NPs, protamine USP was used to form an ion-pair complex with NGF to neutralize the charges on the NGF surface. The NGF/protamine complex was then introduced into the prototype NPs. Apolipoprotein A-I was finally coated on the surface of the NPs. NGF HDL-mimicking NPs showed preferable properties in terms of particle size, size distribution, entrapment efficiency, in vitro release, bioactivity, and biodistribution. With the careful design and exploration of homogenization in HDL-mimicking NPs, the procedure was greatly simplified, and the NPs were made scalable. Moreover, various challenges, such as separating unloaded NGF from the NPs, conducting reliable in vitro release studies, and measuring the bioactivity of the NPs, were overcome.

Simple homogenization was used to prepare novel, high-density, lipoprotein-mimicking nanoparticles to encapsulate nerve growth factor. Challenges, detailed protocols for nanoparticle preparation, in vitro characterization, and in vivo studies are described in this article.

The objective of this article is to introduce preparation and characterization methods for nerve growth factor (NGF)-loaded, high-density, lipoprotein ...

Decellularization of Whole Human Heart Inside a Pressurized Pouch in an Inverted Orientation

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ultimately rejection can occur. Creating a functional, autologous bio-artificial heart could solve these challenges. Biofabrication of a heart comprised of scaffold and cells is one option. A natural scaffold with tissue-specific composition as well as micro- and macro-architecture can be obtained by decellularizing hearts from humans or large animals such as pigs. Decellularization involves washing out cellular debris while preserving 3D extracellular matrix and vasculature and allowing &#34;cellularization&#34; at a later timepoint. Capitalizing on our novel finding that perfusion decellularization of complex organs is possible, we developed a more &#34;physiological&#34; method to decellularize non-transplantable human hearts by placing them inside a pressurized pouch, in an inverted orientation, under controlled pressure. The purpose of using a pressurized pouch is to create pressure gradients across the aortic valve to keep it closed and improve myocardial perfusion. Simultaneous assessment of flow dynamics and cellular debris removal during decellularization allowed us to monitor both fluid inflow and debris outflow, thereby generating a scaffold that can be used either for simple cardiac repair (e.g. as a patch or valve scaffold) or as a whole-organ scaffold.

This method enables decellularization of a complex solid organ using a simple protocol based on osmotic shock and perfusion of ionic detergent with minimal organ matrix disruption. It comprises a novel decellularization technique for human hearts inside a pressurized pouch with real-time monitoring of flow dynamics and cellular debris outflow.

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ...

3D Printing of In Vitro Hydrogel Microcarriers by Alternating Viscous-Inertial Force Jetting

Microcarriers are beads with a diameter of 60-250 &#181;m and a large specific surface area, which are commonly used as carriers for large-scale cell cultures. Microcarrier culture technology has become one of the main techniques in cytological research and is commonly used in the field of large-scale cell expansion. Microcarriers have also been shown to play an increasingly important role in in vitro tissue engineering construction and clinical drug screening. Current methods for preparing microcarriers include microfluidic chips and inkjet printing, which often rely on complex flow channel design, an incompatible two-phase interface, and a fixed nozzle shape. These methods face the challenges of complex nozzle processing, inconvenient nozzle changes, and excessive extrusion forces when applied to multiple bioink. In this study, a 3D printing technique, called alternating viscous-inertial force jetting, was applied to enable the construction of hydrogel microcarriers with a diameter of 100-300 &#181;m. Cells were subsequently seeded on microcarriers to form tissue engineering modules. Compared to existing methods, this method offers a free nozzle tip diameter, flexible nozzle switching, free control of printing parameters, and mild printing conditions for a wide range of bioactive materials.

3D Printing of In Vitro Hydrogel Microcarriers by Alternating Viscous-Inertial Force Jetting

Presented here is a mild 3D printing technique driven by alternating viscous-inertial forces to enable the construction of hydrogel microcarriers. Homemade nozzles offer flexibility, allowing easy replacement for different materials and diameters. Cell binding microcarriers with a diameter of 50-500 &#181;m can be obtained and collected for further culturing.

Microcarriers are beads with a diameter of 60-250 &#181;m and a large specific surface area, which are commonly used as carriers for large-scale cell ...

Quantifying Cytoskeleton Dynamics Using Differential Dynamic Microscopy

Cells can crawl, self-heal, and tune their stiffness due to their remarkably dynamic cytoskeleton. As such, reconstituting networks of cytoskeletal biopolymers may lead to a host of active and adaptable materials. However, engineering such materials with precisely tuned properties requires measuring how the dynamics depend on the network composition and synthesis methods. Quantifying such dynamics is challenged by variations across the time, space, and formulation space of composite networks. The protocol here describes how the Fourier analysis technique, differential dynamic microscopy (DDM), can quantify the dynamics of biopolymer networks and is particularly well suited for studies of cytoskeleton networks. DDM works on time sequences of images acquired using a range of microscopy modalities, including laser-scanning confocal, widefield fluorescence, and brightfield imaging. From such image sequences, one can extract characteristic decorrelation times of density fluctuations across a span of wave vectors. A user-friendly, open-source Python package to perform DDM analysis is also developed. With this package, one can measure the dynamics of labeled cytoskeleton components or of embedded tracer particles, as demonstrated here with data of intermediate filament (vimentin) networks and active actin-microtubule networks. Users with no prior programming or image processing experience will be able to perform DDM using this software package and associated documentation.

Differential dynamic microscopy (DDM) combines features of dynamic light scattering and microscopy. Here, the process of using DDM to characterize reconstituted cytoskeleton networks by quantifying the subdiffusive and caged dynamics of particles in vimentin networks and the ballistic motion of active myosin-driven actin-microtubule composites is presented.

Cells can crawl, self-heal, and tune their stiffness due to their remarkably dynamic cytoskeleton. As such, reconstituting networks of cytoskeletal ...

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell deformation, division, migration, and adhesion. However, studying the dynamics and structure of the actin network in vivo is complicated by the biochemical and genetic regulation within live cells. To build a minimal model devoid of intracellular biochemical regulation, actin is encapsulated inside giant unilamellar vesicles (GUVs, also called liposomes). The biomimetic liposomes are cell-sized and facilitate a quantitative insight into the mechanical and dynamical properties of the cytoskeleton network, opening a viable route for bottom-up synthetic biology. To generate liposomes for encapsulation, the inverted emulsion method (also referred to as the emulsion transfer method) is utilized, which is one of the most successful techniques for encapsulating complex solutions into liposomes to prepare various cell-mimicking systems. With this method, a mixture of proteins of interest is added to the inner buffer, which is later emulsified in a phospholipid-containing mineral oil solution to form monolayer lipid droplets. The desired liposomes are generated from monolayer lipid droplets crossing a lipid/oil-water interface. This method enables the encapsulation of concentrated actin polymers into the liposomes with desired lipid components, paving the way for in vitro reconstitution of a biomimicking cytoskeleton network.

In Vitro Reconstitution of the Actin Cytoskeleton Inside Giant Unilamellar Vesicles

In this manuscript, we demonstrate the experimental techniques to encapsulate the F-actin cytoskeleton into giant unilamellar lipid vesicles (also called liposomes), and the method to form a cortex-biomimicking F-actin layer at the inner leaflet of the liposome membrane.

The actin cytoskeleton, the principal mechanical machinery in the cell, mediates numerous essential physical cellular activities, including cell ...

Light-Controlled Reversible Protein Patterning on Lipid Membranes

Dynamic Light-Induced Protein Patterns at Model Membranes

The precise localization and activation of proteins at the cell membrane at a certain time gives rise to many cellular processes, including cell polarization, migration, and division. Thus, methods to recruit proteins to model membranes with subcellular resolution and high temporal control are essential when reproducing and controlling such processes in synthetic cells. Here, a method is described for fabricating light-regulated reversible protein patterns at lipid membranes with high spatiotemporal precision. For this purpose, we immobilize the photoswitchable protein iLID (improved light-inducible dimer) on supported lipid bilayers (SLBs) and on the outer membrane of giant unilamellar vesicles (GUVs). Upon local blue light illumination, iLID binds to its partner Nano (wild-type SspB) and allows the recruitment of any protein of interest (POI) fused to Nano from the solution to the illuminated area on the membrane. This binding is reversible in the dark, which provides dynamic binding and release of the POI. Overall, this is a flexible and versatile method for regulating the localization of proteins with high precision in space and time using blue light.

Author Spotlight: Photo Switchable Protein Recruitment for Reversible Patterning in Artificial Cellular Systems

Here, a protocol is described for generating light-regulated and reversible protein patterns with high spatiotemporal precision at artificial lipid membranes. The method consists of the localized photoactivation of the protein iLID (improved light-inducible dimer) immobilized on model membranes that, under blue light, binds to its partner protein Nano (wild-type SspB).