This project was funded by NIH (R21EB013717) and IUPUI OVCR (RSFG). The author thanks Ms. Han Shih for her technical assistance.

A. Macromer and Peptide Synthesis
<ol>
	<li>Synthesize 4-arm PEG-norbornene (PEG4NB) and photoinitiator Lithium arylphosphanate (LAP) using established protocols.18,19</li>
	<li>Synthesize bis-cysteine containing chymotrypsin-sensitive peptide CGGY&darr;C (arrow indicates chymotrypsin cleavage site) using standard solid phase peptide synthesis in a microwave peptide synthesizer (CEM Discover SPS).
	<ol class="lalpha">
		<li>Calculate the amount of resin (Rink-amide MBHA resin) needed based on the substitution ratio of the resin and the synthesis scale. Swell the resin in dimethylformamide (DMF) in a reaction vessel (RV) for 15 min.</li>
		<li>Remove Fmoc-protecting group from the resin using a deprotection solution containing 20% Piperidine and 0.1 M HOBt in DMF. Use the parameters listed in the table below for microwave-assisted Fmoc-deprotection (for all Fmoc-amino acids):
		<table border="1">
			<tbody>
				<tr>
					<td>Synthesis scale</td>
					<td>0.1 mmole</td>
					<td>0.2 mmole</td>
				</tr>
				<tr>
					<td>Power</td>
					<td>20 W</td>
					<td>50 W</td>
				</tr>
				<tr>
					<td>Temperature</td>
					<td>75 &deg;C</td>
					<td>75 &deg;C</td>
				</tr>
				<tr>
					<td>Time</td>
					<td>3 min</td>
					<td>3 min</td>
				</tr>
			</tbody>
		</table>
		</li>
		<li>Following deprotection, rinse the resin with DMF and perform Ninhydrin test to confirm the removal of Fmoc (exposure of N-terminal amine). Resin should turn blue after 2 min at 95 &deg;C.</li>
		<li>After successful deprotection, couple the first Fmoc-amino acid at the C-terminus (synthesis runs from C- to N-terminus) in an activator base solution (0.28 M Diisopropylethylamine (DIEA) in DMF) with the parameters listed in the table below.
		<table border="1">
			<tbody>
				<tr>
					<td>Synthesis scale</td>
					<td>0.1 mmole</td>
					<td>0.2 mmole</td>
				</tr>
				<tr>
					<td>Power</td>
					<td>20 W</td>
					<td>50 W</td>
				</tr>
				<tr>
					<td>Temperature*</td>
					<td>75 &deg;C</td>
					<td>75 &deg;C</td>
				</tr>
				<tr>
					<td>Time*</td>
					<td>5 min</td>
					<td>5 min</td>
				</tr>
			</tbody>
		</table>
		 
		* For Cyst and His, use 50 &deg;C and 10 min to reduce racemization.</li>
		<li>Rinse the resin and perform Ninhydrin test to qualitatively confirm the completion of coupling (disappearance of N-terminal amine). Resin should be clear after 5 min at 95 &deg;C. Repeat coupling step (A.2.d) if Ninhydrin test returns positive result (blue color).</li>
		<li>Repeat deprotection (step A.2.b) and coupling (step A.2.d) for additional amino acids in the sequence ending with a final deprotection step.</li>
		<li>Drain the RV, rinse resin with 25 ml of dichloromethane (DCM). This step is to rinse off DMF, which is a base and will hinder peptide cleavage.</li>
		<li>Prepare cleavage cocktail by mixing 250 mg phenol in 4.75 ml Trifluoroacetic acid (TFA), 0.125 ml Triisopropylsilane (TIS) and 0.125 ml ddH2O.</li>
		<li>Add cleavage solution into the RV and cleave for 30 min in the microwave peptide synthesizer (Power = 20 W; Temperature = 38 &deg;C).</li>
		<li>Collect the peptide solution in a 50 ml centrifuge tube using vacuum manifold.</li>
		<li>Precipitate cleaved peptide in 40 ml of ether, vortex the tube and centrifuge at 3,000 rpm for 5 min to collect the peptide.</li>
		<li>Drain the supernatant and repeat step A.2.k twice.</li>
		<li>Collect and dry peptide in vacuum.</li>
		<li>Purify the peptide using HPLC and characterize it with mass spectrometry.</li>
	</ol>
	</li>
</ol>
B. Material Preparation and Sterilization
<ol>
	<li>Prepare 20 wt% PEG4NB solution in PBS, vortex the mixture until the macromer dissolves completely, and sterilize the macromer solution using a syringe filter. Store the sterilized solution at -20 &deg;C (prepare aliquots for long-term storage).</li>
	<li>Prepare and sterilize 2 wt% photoinitiator (LAP) solution in PBS. Store the sterilized solution at room temperature protected from light.</li>
	<li>Dissolve the peptide (CGGYC) in PBS, vortex the mixture and syringe filter the solution for sterilization. Aliquot the peptide solution and store at -20 &deg;C until use (prevent freeze-thaw cycles).</li>
	<li>Determine thiol (-SH) concentration in the prepared peptide solution using Ellman's reagent (follow the manufacturer's protocol). This step will give accurate peptide concentration (i.e., [SH]/2) that is required for calculating the peptide amount to be used in thiol-ene photo-click reactions.</li>
	<li>Prepare gelation molds by cutting the top portion off of sterile 1 ml disposable syringes using a razor blade (open tip for gel removal). Sterilize the syringe molds by autoclave.</li>
	<li>Based on the desired gel formulation (thiol to ene ratio = 1) and concentrations of stock materials, calculate the required volume of polymer, peptide crosslinker, photoinitiator, and buffer solutions required.</li>
</ol>
C. Cell Preparation
<ol>
	<li>Equilibrate cell culture medium (high glucose DMEM containing 10% fetal bovine serum; Antibiotic-Antimycotic with 100 U/ml penicillin, 100 &mu;g/ml streptomycin, 250 ng/ml Fungizone; and 50 &mu;M &beta;-mercaptoethanol), Hank's balanced salt solution (HBSS, Ca2+, Mg2+ free), and Trypsin-EDTA to 37 &deg;C.</li>
	<li>Remove flasks containing MIN6 &beta;-cells from CO2 incubator and place the flasks in the laminar flow hood.</li>
	<li>Aspirate cell culture media from the flasks and rinse the cell with HBSS.</li>
	<li>Trypsinize the cells using 3 ml of 2X (0.1%) Trypsin-EDTA and incubate the flasks for 4 min at 37 &deg;C and 5% CO2.
	<ul>
		<li>Note: Use 2X trypsin solution allows complete dissociation of MIN6 &beta;-cells into single cell suspension.</li>
	</ul>
	</li>
	<li>After incubation, tap the flasks on the surface of the hood gently to completely detach the cells. Confirm the detachment and dissociation of the cells using a microscope.</li>
	<li>Add equal volume of cell culture medium to neutralize the trypsin. Mix the solution well by gentle pipetting, then transfer the mixture to a canonical tube and centrifuge for 5 min at 1,000 rpm.</li>
	<li>Aspirate the supernatant and gently re-suspend the cells in 4 - 5 ml of culture media.</li>
	<li>Determine cell density using 50% Trypan blue buffer in a hemocytometer.</li>
</ol>
D. Hydrogel Fabrication and Cell Encapsulation
<ol>
	<li>Once the cells are ready, mix (based on a 1:1 thiol to ene ratio) the required volumes of PEG4NB, CGGYC, LAP, cell solution, and HBSS in a microcentrifuge tube.</li>
	<li>Mix the solution gently using a pipette to obtain a homogeneous mixture and add 25 &mu;l of the resulting solution to a syringe mold. Expose the syringe to UV light (365 nm, 5 mW/cm2) for 2 min.</li>
	<li>Following photopolymerization, plunge gels into a sterile 24-well plate containing cell culture medium and incubate the hydrogels in 37 &deg;C and 5% CO2.</li>
	<li>Wash the cell-laden hydrogels in cell culture medium for 30 min to rinse off loosely attached cells and un-crosslinked macromers. Transfer the gels into fresh medium.</li>
	<li>Refresh cell culture media every 2 to 3 days.</li>
</ol>
E. Cell Viability Assay
Encapsulated cell morphology can be observed using light microscope (since the synthesized hydrogels are transparent). Cell viability can be visualized and measured qualitatively using Live/Dead staining and quantitatively using AlamarBlue reagent.
E.1. Live/Dead staining
<ol>
	<li>Thaw Live/Dead staining reagents at room temperature.</li>
	<li>In a canonical tube add PBS according to the number of samples (n) to be stained: 
	Total volume of PBS=n &times; 500 &mu;l</li>
	<li>Pipette 0.25 &mu;l of Calcein AM and 2 &mu;l of EthD-1 per 1 ml of PBS to the canonical tube containing PBS. Vortex the solution to mix the components.</li>
	<li>Incubate cell-laden hydrogels in Live/Dead staining solution (0.5 ml/sample) for 1 hr at room temperature on an orbital shaker.</li>
	<li>Using a pasteur pipette, remove the dye solution and rinse the sample twice with PBS.</li>
	<li>Place sample on a cover slip or glass slide. Observe and image cells using a confocal microscope.</li>
</ol>
E.2. AlamarBlue Assay
<ol>
	<li>Prepare 10 v/v% AlamarBlue solution in cell culture medium.</li>
	<li>Remove hydrogels from incubator and aspirate the media out of each well without making any contact with the hydrogel.</li>
	<li>Incubate cell-laden hydrogels and an empty well (negative control) in 500 &mu;l of 10% AlamarBlue solution for 16 hr.</li>
	<li>Following incubation, pipette 200 &mu;l of the AlamarBlue solution in a 96 well plate and measure fluorescence using a microplate reader (excitation: 560 nm, emission: 590 nm).
	<ul>
		<li>Higher cellular metabolic activity reduces the dye to give higher fluorescence reading.</li>
		<li>Be sure to include at least two wells of 10% Alamarblue solution as blanks. When analyzing the data, subtract the averaged blank fluorescence value from the fluorescence reading of the samples.</li>
	</ul>
	</li>
</ol>
F. Chymotrypsin Mediated Gel Erosion and Spheroid Recovery
<ol>
	<li>Prepare 1 mg/ml of Trypsin-free &alpha;-chymotrypsin solution in HBBS and sterile filter the solution.</li>
	<li>Incubate hydrogels in chymotrypsin solution (500 &mu;l for each gel) at room temperature with gentle shaking until complete gel erosion is achieved.
	<ul>
		<li>For a 25 &mu;l gel, the time for complete erosion is approximately 5 min.</li>
	</ul>
	</li>
	<li>Place the plate on ice for a few minutes to reduce the activity of chymotrypsin.</li>
	<li>Transfer the cell solution in 1 ml tube and centrifuge at 300 rpm, 4 &deg;C for 2 min.</li>
	<li>Carefully remove the supernatant using a pasture pipette and gently resuspend the spheroids in HBSS.</li>
	<li>Transfer the recovered spheroid solution to a 24-well plate and place the plate on ice to allow the spheroids to settle down.</li>
	<li>For size analysis, acquire phase contrast images of the settled spheroids using a light microscope. Measure recovered spheroid diameters using software such as Nikon Element software or ImageJ.</li>
</ol>
G. Functional Assay of Recovered Spheroids
G.1. Glucose stimulated insulin secretion from the recovered cell spheroids
<ol>
	<li>In a 24 well plate, incubate the gels in 500 &mu;l of 2 mM low glucose Kerbs-Ringer buffer (KRB) for 1 hr.</li>
	<li>Erode the gel and recover cell spheroids using steps F.1 to F.6.</li>
	<li>Aspirate the supernatant carefully without making any contact with the cell spheroids and resuspend the spheroids in 500 &mu;l of 2 mM low glucose KRB.</li>
	<li>Transfer 100 &mu;l of cell spheroids solution to a 0.6 ml tube and place it on ice for intracellular ATP quantification.</li>
	<li>Split the remaining cell spheroid solution in half (into two microcentrifuge tubes) and centrifuge for 2 min at 300 rpm and 4 &deg;C.</li>
	<li>Aspirate supernatant (almost to the bottom) and resuspend the first half of cell spheroids in 1 ml of 25 mM high glucose KRB and second half in 1 ml of 2 mM low glucose KRB.</li>
	<li>Gently pipette and transfer the solution containing recovered cell spheroids to a 24-well plate. Incubate the plate for 1 hr at 37 &deg;C and 5% CO2.</li>
	<li>Following incubation, transfer the solution to a microcentrifuge tube and centrifuge for 2 min at 1,000 rpm and 4 &deg;C.</li>
	<li>Transfer 500 &mu;l of the supernatant from the centrifuged tube to another microcentrifuge tube and store at 4 &deg;C for insulin ELISA.</li>
	<li>Perform insulin ELISA following manufacturer's protocol.</li>
</ol>
G.2. CellTiter Glo Assay
<ol>
	<li>ATP standards: Prepare a 10 &mu;M ATP solution and perform serial dilutions to obtain a series of 8 standard solutions.</li>
	<li>To a white 96 well plate, add 50 &mu;l of cell spheroid solution (from step G.1.6) and standards.</li>
	<li>To each well containing samples and standards, add 50 &mu;l of CellTiter Glo reagent and incubate at room temperature for 15 min.</li>
	<li>Following incubation, measure luminescence count of the samples and standards using a microplate reader and calibrate sample ATP concentrations using a linear regression curve generated by the ATP standards.</li>
	<li>Determine the total ATP concentration in sample and multiply by the sample dilution factor to get the intracellular ATP concentration in one cell-laden hydrogel.</li>
</ol>
G.3. Determine insulin secretion
<ol>
	<li>Calculate the amount of insulin secreted from the recovered spheroids by taking the ratio of the amount of insulin secreted in high glucose medium to amount of insulin secreted in low glucose medium.</li>
	<li>Normalize the quantified insulin content of cell spheroids with respective amount of intracellular ATP of the recovered cell spheroids.</li>
</ol>

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No conflicts of interest declared.

The described protocol presents details on easy encapsulation of cells in thiol-ene hydrogels formed by step-growth photopolymerization. While a stoichiometric ratio of 1:1 of norbornene to thiol functional groups was used in this protocol, the ratio can be adjusted depending on the experiments. In addition to a correct formulation, it is important to maintain homogeneity in the pre-polymer solution. In particular, use gentle pipetting to ensure that cells are well distributed in the pre-polymer solution in order t...

Hydrogels are hydrophilic crosslinked polymers with exceptional potential as scaffolding materials for repairing and regenerating tissues.1-3 The high water content of hydrogels permits easy diffusion of oxygen and exchange of nutrients and cellular metabolic products, all of which are crucial to maintaining cell viability. In addition, hydrogels are excellent carriers for controlled release and cell delivery due their high tunability.2 Synthetic hydrogels such as those prepared from poly(ethylene glycol) (PEG) are increasingly used in tissue engineering applications, largely due to their cytocompatibility, tissue-like elasticity, and high tunability in material physical and mechanical properties.4-6
Although a commonly used hydrogel platform, studies have shown that PEG diacrylate (PEGDA) hydrogels formed by chain-growth photopolymerizations have a tendency to damage encapsulated cells during network crosslinking and in situ cell encapsulation.7 The cellular damage was largely attributed to radical species generated by the photoinitiator molecules, which propagate through the vinyl groups on PEGDA to crosslink polymer chains into hydrogels. Unfortunately, these radical species also cause stresses and cellular damage during cell encapsulation, especially for radical-sensitive cells such as pancreatic &beta;-cells.8-10 In order to obtain a higher mesh size for better diffusion and cell survival, higher molecular weights PEGDA are often used for cell encapsulation. This, however, compromises polymerization kinetics and causes sub-optimal gel biophysical properties.7,11,12 In addition to the above mentioned disadvantages, it is very difficult to recover cell structures from PEGDA hydrogels due to the heterogeneity and non-degradable nature of the crosslinked networks. While protease-sensitive peptides can be incorporated into PEG macromer backbone to render the otherwise inert PEGDA hydrogels sensitive to enzymatic cleavage, the conjugation often uses expensive reagents and the resulting networks still contain high degree of heterogeneity due to the nature of chain-growth polymerization.13-15
Recently, PEG-peptide hydrogels formed via step-growth thiol-ene photopolymerization have been shown to exhibit preferential properties for cell encapsulation over hydrogels formed by chain-growth photopolymerization.7 The superior gelation kinetics of thiol-ene hydrogels is attributed to the 'click' nature of reaction between thiol and ene functionalities. Compared to chain-growth polymerization of PEGDA, thiol-ene reaction is less oxygen inhibited which results in faster gelation rate.16,17 Thiol-ene hydrogels also have higher polymerization efficiency and better gel biophysical properties compared to chain-growth PEGDA hydrogels,7,18 which results in limited cellular damage caused by radical species during photopolymerization.
Previously, thiol-ene hydrogels formed by 4-arm PEG-norbornene (PEG4NB) macromer and bis-cysteine containing peptide crosslinkers, such as protease-sensitive peptides have been utilized for cell encapsulation.7,18 High tunability of PEG hydrogel networks offers a flexible and controllable 3D microenvironment for investigating cell survival and activity, while the use of protease-sensitive peptide sequence provides a mild way for recovery of cell constructs formed naturally within hydrogels. In this protocol we utilize step-growth photopolymerized thiol-ene hydrogels fabricated using 4-arm PEG-norbornene (PEG4NB) and a chymotrypsin-sensitive peptide crosslinker (CGGY&darr;C) for the encapsulation of MIN6 &beta;-cells. This protocol systematically elaborates techniques for studying the survival, proliferation and spheroid formation of MIN6 &beta;-cells in thiol-ene hydrogels. We further provide method for &beta;-cell spheroid recovery and biological characterization of recovered spheroids.

<table border="1">
 <tbody>
 <tr>
 <td>Name</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>4-arm PEG (20kDa)</td>
 <td>Jenkem Technology USA</td>
 <td>4ARM-PEG-20K</td>
 <td></td>
 </tr>
 <tr>
 <td>Fmoc-amino acids</td>
 <td>Anaspec</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Live/Dead cell viability kit</td>
 <td>Invitrogen</td>
 <td>L3224</td>
 <td>Includes Calcein AM and Ethidium homodimer-1</td>
 </tr>
 <tr>
 <td>AlamarBlue reagent</td>
 <td>AbD Serotec</td>
 <td>BUF012</td>
 <td></td>
 </tr>
 <tr>
 <td>CellTiter Glo reagent</td>
 <td>Promega</td>
 <td>G7570</td>
 <td></td>
 </tr>
 <tr>
 <td>DPBS</td>
 <td>Lonza</td>
 <td>17-512F</td>
 <td>Without Ca+2 and Mg+2</td>
 </tr>
 <tr>
 <td>HBSS</td>
 <td>Lonza</td>
 <td>10547F</td>
 <td>Without Ca+2 and Mg+2</td>
 </tr>
 <tr>
 <td>High Glucose DMEM</td>
 <td>Hyclone</td>
 <td>SH30243.01</td>
 <td></td>
 </tr>
 <tr>
 <td>FBS</td>
 <td>Gibco</td>
 <td>16000-044</td>
 <td></td>
 </tr>
 <tr>
 <td>Antibiotic-Antimycotic</td>
 <td>Invitrogen</td>
 <td>15240-062</td>
 <td></td>
 </tr>
 <tr>
 <td>&beta;-Mercaptoethanol</td>
 <td>Sigma-Aldrich</td>
 <td>M7522-100ML</td>
 <td></td>
 </tr>
 <tr>
 <td>Trypsin-EDTA</td>
 <td>Invitrogen</td>
 <td>15400-054</td>
 <td></td>
 </tr>
 <tr>
 <td>Trypsin-free &alpha;-chymotrypsin</td>
 <td>Worthington Biochemical Corp</td>
 <td>LS001432</td>
 <td></td>
 </tr>
 <tr>
 <td>Mouse Inusin ELISA kit</td>
 <td>Mercodia</td>
 <td>10-1247-01</td>
 <td></td>
 </tr>
 <tr>
 <td>1 ml disposable syringe</td>
 <td>BD biosciences</td>
 <td></td>
 <td></td>
 </tr>
 </tbody>
</table>

generation and recovery of &beta;-cell spheroids from step-growth peg-peptide hydrogels

Hydrogels are hydrophilic crosslinked polymers that provide a three-dimensional microenvironment with tissue-like elasticity and high permeability for culturing therapeutically relevant cells or tissues. Hydrogels prepared from poly(ethylene glycol) (PEG) derivatives are increasingly used for a variety of tissue engineering applications, in part due to their tunable and cytocompatible properties. In this protocol, we utilized thiol-ene step-growth photopolymerizations to fabricate PEG-peptide hydrogels for encapsulating pancreatic MIN6 b-cells. The gels were formed by 4-arm PEG-norbornene (PEG4NB) macromer and a chymotrypsin-sensitive peptide crosslinker (CGGYC). The hydrophilic and non-fouling nature of PEG offers a cytocompatible microenvironment for cell survival and proliferation in 3D, while the use of chymotrypsin-sensitive peptide sequence (CGGY&darr;C, arrow indicates enzyme cleavage site, while terminal cysteine residues were added for thiol-ene crosslinking) permits rapid recovery of cell constructs forming within the hydrogel. The following protocol elaborates techniques for: (1) Encapsulation of MIN6 &beta;-cells in thiol-ene hydrogels; (2) Qualitative and quantitative cell viability assays to determine cell survival and proliferation; (3) Recovery of cell spheroids using chymotrypsin-mediated gel erosion; and (4) Structural and functional analysis of the recovered spheroids.

Figures 1-4 show representative results for encapsulation, survival, proliferation, spheroid formation, and spheroid recovery in thiol-ene hydrogels. Figure 1 shows the reaction schematic of (1) step-growth thiol-ene photopolymerization using PEG4NB and CGGYC, and (2) chymotrypsin mediated gel erosion which follows a surface erosion mechanism. Figures 2 and 3 present viability results obtained using Live/Dead staining and AlamarBlue assay. We observe tha...

Watch this Scientific Journal Video about Generation and Recovery of Î²-cell Spheroids From Step-growth PEG-peptide Hydrogels at JoVE.com

Generation and Recovery of &amp;#946;-cell Spheroids From Step-growth PEG-peptide Hydrogels

The following protocol provides techniques for encapsulating pancreatic &#x03B2;-cells in step-growth PEG-peptide hydrogels formed by thiol-ene photo-click reactions. This material platform not only offers a cytocompatible microenvironment for cell encapsulation, but also permits user-controlled rapid recovery of cell structures formed within the hydrogels.

Generation and Recovery of &beta;-cell Spheroids From Step-growth PEG-peptide Hydrogels

Hydrogels are hydrophilic crosslinked polymers that provide a three-dimensional  microenvironment with tissue-like elasticity and high permeability for  ...

generation-recovery-cell-spheroids-from-step-growth-peg-peptide

Research

JoVE Journal

Bioengineering

14.1K Views.  Indiana University - Purdue University at Indianapolis. The following protocol provides techniques for encapsulating pancreatic β-cells in step-growth PEG-peptide hydrogels formed by thiol-ene photo-click reactions. This material platform not only offers a cytocompatible microenvironment for cell encapsulation, but also permits user-controlled rapid recovery of cell structures formed within the hydrogels.

Video: Generation and Recovery of &#946;-cell Spheroids From Step-growth PEG-peptide Hydrogels

A Simple Hanging Drop Cell Culture Protocol for Generation of 3D Spheroids

Studies of cell-cell cohesion and cell-substratum adhesion have historically been performed on monolayer cultures adherent to rigid substrates. Cells within a tissue, however, are typically encased within a closely packed tissue mass in which cells establish intimate connections with many near-neighbors and with extracellular matrix components. Accordingly, the chemical milieu and physical forces experienced by cells within a 3D tissue are fundamentally different than those experienced by cells grown in monolayer culture. This has been shown to markedly impact cellular morphology and signaling. Several methods have been devised to generate 3D cell cultures including encapsulation of cells in collagen gels1or in biomaterial scaffolds2. Such methods, while useful, do not recapitulate the intimate direct cell-cell adhesion architecture found in normal tissues. Rather, they more closely approximate culture systems in which single cells are loosely dispersed within a 3D meshwork of ECM products. Here, we describe a simple method in which cells are placed in hanging drop culture and incubated under physiological conditions until they form true 3D spheroids in which cells are in direct contact with each other and with extracellular matrix components. The method requires no specialized equipment and can be adapted to include addition of any biological agent in very small quantities that may be of interest in elucidating effects on cell-cell or cell-ECM interaction. The method can also be used to co-culture two (or more) different cell populations so as to elucidate the role of cell-cell or cell-ECM interactions in specifying spatial relationships between cells. Cell-cell cohesion and cell-ECM adhesion are the cornerstones of studies of embryonic development, tumor-stromal cell interaction in malignant invasion, wound healing, and for applications to tissue engineering. This simple method will provide a means of generating tissue-like cellular aggregates for measurement of biomechanical properties or for molecular and biochemical analysis in a physiologically relevant model.

We describe a simple, rapid method of generating 3D tissue-like spheroids and their potential application to quantify differences in cell-cell interactions.

Studies of cell-cell cohesion and cell-substratum adhesion have historically been performed on monolayer cultures adherent to rigid substrates. Cells ...

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung tissue from porcine, rat, or mouse, the tissue is perfused and submerged in subsequent chemical detergents to remove the cellular debris. Histological comparison of the tissue before and after processing confirms removal of over 95% of double stranded DNA and alpha galactosidase staining suggests the majority of cellular debris is removed. After decellularization, the tissue is lyophilized and then cryomilled into a powder. The matrix powder is digested for 48 hr in an acidic pepsin digestion solution and then neutralized to form the pregel solution. Gelation of the pregel solution can be induced by incubation at 37 &#176;C and can be used immediately following neutralization or stored at 4 &#176;C for up to two weeks. Coatings can be formed using the pregel solution on a non-treated plate for cell attachment. Cells can be suspended in the pregel prior to self-assembly to achieve a 3D culture, plated on the surface of a formed gel from which the cells can migrate through the scaffold, or plated on the coatings. Alterations to the strategy presented can impact gelation temperature, strength, or protein fragment sizes. Beyond hydrogel formation, the hydrogel stiffness may be increased using genipin.

This is a method to create a 3-dimensional cell culture scaffold from pulmonary extracellular matrix. Intact lung is processed into hydrogels that can support the growth of cells in three-dimensions.

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung ...

Controlled Strain of 3D Hydrogels under Live Microscopy Imaging

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized in vitro stretching methods. Various available systems use 2D substrate-based stretchers, while the accessibility of 3D techniques to strain soft hydrogels, is more restricted. Here, we describe a method that allows external stretching of soft hydrogels from their circumference, using an elastic silicone strip as the sample carrier. The stretching system utilized in this protocol is constructed from 3D-printed parts and low-cost electronics, making it simple and easy to replicate in other labs. The experimental process begins with polymerizing thick (&#62;100 &#956;m) soft fibrin hydrogels (Elastic Modulus of ~100 Pa) in a cut-out at the center of a silicone strip. Silicone-gel constructs are then attached to the printed-stretching device and placed on the confocal microscope stage. Under live microscopy the stretching device is activated, and the gels are imaged at various stretch magnitudes. Image processing is then used to quantify the resulting gel deformations, demonstrating relatively homogenous strains and fiber alignment throughout the gel&#8217;s 3D thickness (Z-axis). Advantages of this method include the ability to strain extremely soft hydrogels in 3D while executing in situ microscopy, and the freedom to manipulate the geometry and size of the sample according to the user&#8217;s needs. Additionally, with proper adaptation, this method can be used to stretch other types of hydrogels (e.g., collagen, polyacrylamide or polyethylene glycol) and can allow for analysis of cells and tissue response to external forces under more biomimetic 3D conditions.

The presented method involves uniaxial stretching of 3D soft hydrogels embedded in silicone rubber while allowing live confocal microscopy. Characterization of the external and internal hydrogel strains as well as fiber alignment are demonstrated. The device and protocol developed can assess the response of cells to various strain regimes.

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized ...

Affordable Oxygen Microscopy-Assisted Biofabrication of Multicellular Spheroids

Multicellular spheroids are important tools for studying tissue and cancer physiology in 3D and are frequently used in tissue engineering as tissue assembling units for biofabrication. While the main power of the spheroid model is in mimicking physical-chemical gradients at the tissue microscale, the real physiological environment (including dynamics of metabolic activity, oxygenation, cell death, and proliferation) inside the spheroids is generally ignored. At the same time, the effects of the growth medium composition and the formation method on the resulting spheroid phenotype are well documented. Thus, characterization and standardization of spheroid phenotype are required to ensure the reproducibility and transparency of the research results. The analysis of average spheroid oxygenation and the value of O2 gradients in three dimensions (3D) can be a simple and universal way for spheroid phenotype characterization, pointing at their metabolic activity, overall viability, and potential to recapitulate in vivo tissue microenvironment. The visualization of 3D oxygenation can be easily combined with multiparametric analysis of additional physiological parameters (such as cell death, proliferation, and cell composition) and applied for continuous oxygenation monitoring and/or end-point measurements. The loading of the O2 probe is performed during the stage of spheroid formation and is compatible with various protocols of spheroid generation. The protocol includes a high-throughput method of spheroid generation with introduced red and near-infrared emitting ratiometric fluorescent O2 nanosensors and the description of multi-parameter assessment of spheroid oxygenation and cell death before and after bioprinting. The experimental examples show comparative O2 gradients analysis in homo- and hetero-cellular spheroids as well as spheroid-based bioprinted constructs. The protocol is compatible with a conventional fluorescence microscope having multiple fluorescence filters and a light-emitting diode as a light source.

The protocol describes high-throughput spheroid generation for bioprinting using the multi-parametric analysis of their oxygenation and cell death on a standard fluorescence microscope. This approach can be applied to control the spheroids viability and perform standardization, which is important in modeling 3D tissue, tumor microenvironment, and successful (micro)tissue biofabrication.

Multicellular spheroids are important tools for studying tissue and cancer physiology in 3D and are frequently used in tissue engineering as tissue ...

Fragmenting Bulk Hydrogels and Processing into Granular Hydrogels for Biomedical Applications

Granular hydrogels are jammed assemblies of hydrogel microparticles (i.e., &#34;microgels&#34;). In the field of biomaterials, granular hydrogels have many advantageous properties, including injectability, microscale porosity, and tunability by mixing multiple microgel populations. Methods to fabricate microgels often rely on water-in-oil emulsions (e.g., microfluidics, batch emulsions, electrospraying) or photolithography, which may present high demands in terms of resources and costs, and may not be compatible with many hydrogels. This work details simple yet highly effective methods to fabricate microgels using extrusion fragmentation and to process them into granular hydrogels useful for biomedical applications (e.g., 3D printing inks). First, bulk hydrogels (using photocrosslinkable hyaluronic acid (HA) as an example) are extruded through a series of needles with sequentially smaller diameters to form fragmented microgels. This microgel fabrication technique is rapid, low-cost, and highly scalable. Methods to jam microgels into granular hydrogels by centrifugation and vacuum-driven filtration are described, with optional post-crosslinking for hydrogel stabilization. Lastly, granular hydrogels fabricated from fragmented microgels are demonstrated as extrusion printing inks. While the examples described herein use photocrosslinkable HA for 3D printing, the methods are easily adaptable for a wide variety of hydrogel types and biomedical applications.

This work describes straightforward, adaptable, and low-cost methods to fabricate microgels with extrusion fragmentation, process the microgels into injectable granular hydrogels, and apply the granular hydrogels as extrusion printing inks for biomedical applications.

Granular hydrogels are jammed assemblies of hydrogel microparticles (i.e., &#34;microgels&#34;). In the field of biomaterials, granular hydrogels have ...

Human Liver Spheroids from Peripheral Blood for Liver Disease Studies

Human liver cells can form a three-dimensional (3D) structure capable of growing in culture for some weeks, preserving their functional capacity. Due to their nature to cluster in the culture dishes with low or no adhesive characteristics, they form aggregates of multiple liver cells that are called human liver spheroids. The forming of 3D liver spheroids relies on the natural tendency of hepatic cells to aggregate in the absence of an adhesive substrate. These 3D structures possess better physiological responses than cells, which are closer to an in vivo environment. Using 3D hepatocyte cultures has numerous advantages when compared with classical two-dimensional (2D) cultures, including a more biologically relevant microenvironment, architectural morphology that reassembles natural organs as well as a better prediction regarding disease state and in vivo-like responses to drugs. Various sources can be used to generate spheroids, like primary liver tissue or immortalized cell lines. The 3D liver tissue can also be engineered by using human embryonic stem cells (hESCs) or induced pluripotent stem cells (hiPSCs) to derive hepatocytes. We have obtained human liver spheroids using blood-derived pluripotent stem cells (BD-PSCs) generated from unmanipulated peripheral blood by activation of human membrane-bound GPI-linked protein and differentiated to human hepatocytes. The BD-PSCs-derived human liver cells and human liver spheroids were analyzed by light microscopy and immunophenotyping using human hepatocyte markers.

Here we present a non-genetic method to generate human autologous liver spheroids using mononuclear cells isolated from steady-state peripheral blood.

Human liver cells can form a three-dimensional (3D) structure capable of growing in culture for some weeks, preserving their functional capacity. Due to ...

Generation, High-Throughput Screening, and Biobanking of Human-Induced Pluripotent Stem Cell-Derived Cardiac Spheroids

Human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are of paramount importance for human cardiac disease modeling and therapeutics. We recently published a cost-effective strategy for the massive expansion of hiPSC-CMs in two dimensions (2D). Two major limitations are cell immaturity and a lack of three-dimensional (3D) arrangement and scalability in high-throughput screening (HTS) platforms. To overcome these limitations, the expanded cardiomyocytes form an ideal cell source for the generation of 3D cardiac cell culture and tissue engineering techniques. The latter holds great potential in the cardiovascular field, providing more advanced and physiologically relevant HTS. Here, we describe an HTS-compatible workflow with easy scalability for the generation, maintenance, and optical analysis of cardiac spheroids (CSs) in a 96-well-format. These small CSs are essential to fill the gap present in current in vitro disease models and/or generation for 3D tissue engineering platforms. The CSs present a highly structured morphology, size, and cellular composition. Furthermore, hiPSC-CMs cultured as CSs display increased maturation and several functional features of the human heart, such as spontaneous calcium handling and contractile activity. By automatization of the complete workflow, from the generation of CSs to functional analysis, we increase intra- and inter-batch reproducibility as demonstrated by high-throughput (HT) imaging and calcium handling analysis. The described protocol allows modeling of cardiac diseases and assessing drug/therapeutic effects at the single-cell level within a complex 3D cell environment in a fully automated HTS workflow. In addition, the study describes a straightforward procedure for long-term preservation and biobanking of whole-spheroids, thereby providing researchers the opportunity to create next-generation functional tissue storage. HTS combined with long-term storage will substantially contribute to translational research in a wide range of areas, including drug discovery and testing, regenerative medicine, and the development of personalized therapies.

Presented here is a set of protocols for the generation and cryopreservation of cardiac spheroids (CSs) from human-induced pluripotent stem cell-derived cardiomyocytes cultured in a high-throughput, multidimensional format. This three-dimensional model functions as a robust platform for disease modeling, high-throughput screenings, and maintains its functionality after cryopreservation.

Human-induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are of paramount importance for human cardiac disease modeling and therapeutics. We ...

Synthesis of Decellularized Cartilage Extracellular Matrix Hydrogels

Decellularized cartilage extracellular matrix (DC-ECM) hydrogels are promising biomaterials for tissue engineering and regenerative medicine due to their biocompatibility and ability to mimic natural tissue properties. This protocol aims to produce DC-ECM hydrogels that closely mimic the native ECM of cartilage tissue. The protocol involves a combination of physical and chemical disruption and enzymatic digestion to remove the cellular material while preserving the structure and composition of the ECM. The DC-ECM is cross-linked using a chemical agent to form a stable and biologically active hydrogel. The DC-ECM hydrogel has excellent biological activity, spatial structure, and biological induction function, as well as low immunogenicity. These characteristics are beneficial in promoting cell adhesion, proliferation, differentiation, and migration and for creating a superior microenvironment for cell growth. This protocol provides a valuable resource for researchers and clinicians in the field of tissue engineering. Biomimetic hydrogels can potentially enhance the development of effective tissue engineering strategies for cartilage repair and regeneration.

This paper introduces a new method for the synthesis of decellularized cartilage extracellular matrix (DC-ECM) hydrogels. DC-ECM hydrogels have excellent biocompatibility and provide a superior microenvironment for cell growth. Therefore, they can be ideal cell scaffolds and biological delivery systems.

Decellularized cartilage extracellular matrix (DC-ECM) hydrogels are promising biomaterials for tissue engineering and regenerative medicine due to their ...

Comparative Study of Decellularization Techniques for Bovine Lung ECM Hydrogels

Development and Characterization of Decellularized Lung Extracellular Matrix Hydrogels

The use of extracellular matrix (ECM)-derived hydrogels in tissue engineering has become increasingly popular, as they can mimic cells&#39; natural environment in vitro. However, maintaining the native biochemical content of the ECM, achieving mechanical stability, and comprehending the impact of the decellularization process on the mechanical properties of the ECM hydrogels are challenging. Here, a pipeline for decellularization of bovine lung tissue using two different protocols, downstream characterization of the effectiveness of decellularization, fabrication of reconstituted decellularized lung ECM hydrogels and assessment of their mechanical and cytocompatibility properties were described. Decellularization of the bovine lung was pursued using a physical (freeze-thaw cycles) or chemical (detergent-based) method. Hematoxylin and Eosin staining was performed to validate the decellularization and retention of major ECM components. For the evaluation of residual collagen and sulfated glycosaminoglycan (sGAG) content within the decellularized samples, Sirius red and Alcian blue staining techniques were employed, respectively. Mechanical properties of the decellularized lung ECM hydrogels were characterized by oscillatory rheology. The results suggest that decellularized bovine lung hydrogels can provide a reliable organotypic<span style="font-size: 11pt; font-family: Calibri, sans-serif; color: black; background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">&#160;alternative to commercial ECM products by retaining most native ECM components. Furthermore, these findings reveal that the decellularization method of choice significantly affects gelation kinetics as well as the stiffness and viscoelastic properties of resulting hydrogels.

Author Spotlight: Optimizing Bovine Lung Decellularization for Organotypic Hydrogels

The protocol elucidates two distinct decellularization methodologies applied to native bovine pulmonary tissues, providing a comprehensive account of their respective characterizations.

The use of extracellular matrix (ECM)-derived hydrogels in tissue engineering has become increasingly popular, as they can mimic cells&#39; natural ...