The current project was funded by the University of Kansas Proof of Concept Fund. We would like to thank the nursing staff at KUMC (Kansas City, KS) for assisting us in obtaining human umbilical cords, and David Jorgensen for assisting with cochleae cultures.

All procedures, including animal euthanasia, were conducted according to the approved Institutional Animal Care and Use Committee (IACUC) protocol (ACUP #2014-2234) at the University of Kansas Medical Center (KUMC).
NOTE: HWJCs were isolated from human umbilical cords that were donated by patients that provided informed consent and specimens were used in accordance with the protocols approved by the University of Kansas Human Subjects Committee (KU-IRB #15402).
1. Temporal Bone Harvest and Cochlea Isolation
<ol>
	<li>Decapitate an appropriately euthanized, 15-week-old mouse by cutting between the skull and first cervical vertebra with a sharp pair of surgical scissors, then spray down the head with 70% ethanol to disinfect (Figure 1A).</li>
	<li>Bisect the skull in a mid-sagittal plane using the same sharp pair of surgical scissors (Figure 1B-C, dashed lines).</li>
	<li>Remove the brain tissue from the skull using a pair of forceps (Figure 1D, arrowhead).</li>
	<li>Identify the temporal bone through the presence of the auditory and vestibular nerve roots (Figure 1E-F, arrowheads) on the interior of the skull and the ear canal from the exterior (Figure 1F, arrow). Carefully cut through the skull to isolate the temporal bone from the remainder of the skull (Figure 1G). 
	NOTE: In some instances, it may be possible to delicately pry the surrounding bones of the skull away from the temporal bone without requiring cutting.</li>
	<li>Insert the fine forceps into the opening of the ear canal approximately 5 mm (Figure 1H), so the tips do not risk puncturing the cochlea. Gently break open the bone of the bulla (Figure 1H, top red shaded area) so it fractures (Figure 1I, red dotted line). 
	NOTE: If necessary, a dissection microscope may aid in this process to ensure accurate placement and manipulation of instruments.</li>
	<li>Using fine forceps, pry away the remainder of the bulla from the temporal bone, exposing the bony labyrinth of the cochlea (Figure 1J, red brackets).</li>
	<li>Place a Petri dish large enough to hold the temporal bone (e.g., 30 mm or greater) underneath the dissecting scope field of view and fill it with enough Phosphate Buffered Saline (PBS) to cover the temporal bone (e.g., 0.5 to 1 mL).</li>
	<li>Submerge the temporal bone with the bulla removed in the PBS with the oval and round windows of the cochlea facing up. 
	NOTE: Removal of the vestibular system from the cochlea is not necessary, as the vestibular system serves well as a handle for manipulating and orienting features of the cochlea (Figure 1I and Figure 2B).</li>
	<li>Using an ultra-fine pair of forceps, remove the stapedial artery, which passes through the stapes.</li>
	<li>Insert one tip of the ultra-fine forceps through the arch of the stapes where the artery passed through, and delicately lift the stapes upward. The disarticulated stapes should lift off of the oval window.</li>
	<li>Using one tip of the ultra-fine forceps, puncture the oval and round windows.</li>
	<li>Using a 1x PBS filled 28.5-gauge syringe with tubing (inner diameter 0.28 mm, outer diameter 0.61 mm) attached, position the tubing over the oval window (Figure 2A - B). 
	NOTE: The opening of the tube can be manipulated using ultra-fine forceps helping to ensure accurate placement.
	<ol>
		<li>Use a fresh syringe and tubing for each cochlea.</li>
	</ol>
	</li>
	<li>Perfuse 2 mL of 10% antibiotic-antimycotic (anti-anti) and 10% penicillin-streptomycin (pen-strep) in PBS through the cochlea over 5 min to remove the perilymph. Perfusing too fast with too much pressure will damage the fine structures within the cochlea. 
	NOTE: Manually driving the syringe by hand to perfuse the fluid through the cochlea is effective, although a perfusion pump could be used as an alternative.
	<ol>
		<li>If manually perfusing, use a pair of fine, self-closing forceps to stabilize the cochlea (Figure 2A) during perfusion by grasping the vestibular portion of the temporal bone. 
		NOTE: Although not required, this leaves both hands free to handle instruments; one hand can drive the syringe (Figure 2A) while the other can position the tubing appropriately (Figure 2B).</li>
		<li>From this step forward, take care to avoid exposing the cochlea unnecessarily to air. Introducing air bubbles into the fluid spaces will block fluid circulation, and further drying out will damage the fine structures within the cochlea.</li>
	</ol>
	</li>
	<li>Remove and discard any remaining muscle tissue and bone fragments using fine forceps.</li>
	<li>If necessary, store isolated cochleae at 4 &#176;C in 10% anti-anti and 10% pen-strep solution in PBS for up to seven days with regular changes of antibiotic solution every 48 h.</li>
</ol>
2. Cochlear Processing
<ol>
	<li>Decellularization
	<ol>
		<li>For each cochlea, fill a 20-mL glass scintillation vial with a 1% sodium dodecyl sulfate (SDS) solution in de-ionized (DI) water, which is used to decellularize the cochlea.</li>
		<li>Cut the tip off of a plastic 7-mL transfer pipette using a razor blade so that the opening is large enough for a cochlea to pass through.</li>
		<li>Using the transfer pipette, gently draw the cochlea up into the pipette so it just passes the cut-off edge by a few millimeters.</li>
		<li>Expel the cochlea into the 1% SDS solution-filled scintillation vial.</li>
		<li>Circulate 2 mL of 1% SDS in de-ionized (DI) water through the cochlea in the scintillation vial&#160;using a transfer pipette.</li>
		<li>Place the scintillation vial into a rotator, and allow the scintillation vial to rotate at 10 rpm for 72 h at room temperature.
		<ol>
			<li>Change 1% SDS solution every 24 h over a 72 h period. Take care when changing SDS to never expose the cochlea to air.</li>
		</ol>
		</li>
		<li>Wash the cochlea three times for 30 min each with DI water. Perform the washes in the same scintillation vial used for the SDS charges; at this stage, the cochlea is decellularized.</li>
	</ol>
	</li>
	<li>Decalcification
	<ol>
		<li>Remove the remaining DI water from the scintillation vial, leaving just enough to keep the cochlea submerged.</li>
		<li>To begin decalcification, add 5 mL of 10% ethylenediaminetetraacetic acid (EDTA) (0.02 M) in DI water to the scintillation vial containing the cochlea.</li>
		<li>Circulate 2 mL of 10% EDTA in DI water through the cochlea in the scintillation vial&#160;using a transfer pipette.</li>
		<li>Place the scintillation vial back in the rotator, and allow the scintillation vial to rotate at 10 rpm for 72 h at room temperature. Change the 10% EDTA solution every 24 h over a 72 h period. Take care when changing solutions so that the cochlea is not exposed to air.</li>
		<li>Rinse the cochlea 3 times for 2 h each with 1x PBS; at this stage, the cochlea is decalcified.</li>
	</ol>
	</li>
	<li>Storage
	<ol>
		<li>Store the cochleae for up to 72 h in a 10% anti-anti, 10% pen-strep solution in PBS at 4 &#176;C. 
		NOTE: Longer storage periods may be possible with regular changes of antibiotic solution; however, extended storage of processed cochleae has not been validated in this protocol.</li>
	</ol>
	</li>
</ol>
3. Procurement and Expansion of hWJCs
<ol>
	<li>Isolate hWJCs according to previously published protocols22. Briefly, segment umbilical cords into 3-cm sections.</li>
	<li>Carefully make a shallow incision with a scalpel lengthwise on each umbilical cord segment to unroll and expose the Wharton&#39;s jelly of the umbilical cord. Use forceps to remove the blood vessels.</li>
	<li>Wash umbilical cord segments twice with enough PBS (e.g., 40 mL in a 60 mm Petri dish) to submerge the tissue. Finely mince tissue segments with a scalpel. Digest tissue in a 100 mm Petri dish with 50 mL of digestion medium (0.2% type 2 collagenase, 1% penicillin-streptomycin, in low-glucose Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM)).</li>
	<li>Place tissue in digesting medium on an orbital shaker at 50 rpm overnight in a 5% CO2, 37 &#176;C incubator. The following day, dilute digestion medium at a ratio of 1:16 in 2% antibiotic-antimycotic in PBS, and pellet cells by centrifugation at 500 x g at room temperature (~ 27 &#176;C) for 10 min. Discard supernatant.</li>
	<li>Resuspend pellets in Mesenchymal Stem Cell Growth Medium (MSCGM) and recombine cells. Plate cells at a density of 7 x 103 cells/cm2 in tissue culture treated T-75 flasks. Culture HWJCs in MSCGM, and expand to passage 5 for experiments. 
	NOTE: HWJCs may be sub-cultured into larger T-flasks (e.g., T-150, T-300) when passaging to increase yield of hWJCs for experiments.</li>
</ol>
4. Infusion of hWJCs into Decellularized Cochleae
<ol>
	<li>Prepare decellularized cochleae
	<ol>
		<li>Using aseptic technique, wash stored cochleae in 1x PBS three times for 30 min each.</li>
		<li>Transfer cochleae into separate wells of a 24-well plate, and add 1 mL of 37 &#176;C pre-warmed MSCGM.</li>
		<li>Incubate cochleae in a 5% CO2, 37 &#176;C cell culture incubator for a minimum of 1 h before infusion of cells.</li>
	</ol>
	</li>
	<li>Dissociate hWJCs and resuspend
	<ol>
		<li>Wash hWJCs with 37 &#176;C pre-warmed 1x PBS twice.</li>
		<li>Add enough 37 &#176;C pre-warmed Trypsin with 0.5% EDTA to cover the cell vessel surface.</li>
		<li>Incubate hWJCs for up to 5 min in a 5% CO2, 37 &#176;C cell culture incubator.
		<ol>
			<li>Verify that 90% of cells have detached from the cell culture surface by viewing cells under an inverted microscope. 
			NOTE: Cells that move when the culture vessel is gently tapped, have detached. Cells that remain stationary, have not detached.</li>
			<li>Gently tap the sides of the flask to further dislodge attached cells.</li>
		</ol>
		</li>
		<li>Using aseptic technique, transfer hWJCs from the culture vessel to a 50-mL conical tube containing an equivalent volume of MSCGM to the volume of Trypsin used to dissociate the cells.</li>
		<li>Pellet the hWJCs by centrifuging at 500 x g at room temperature (~ 27 &#176;C) for 5 min.</li>
		<li>Aspirate supernatant, and resuspend hWJCs in MSCGM at a concentration of 500,000 cells/mL.</li>
	</ol>
	</li>
	<li>Perfuse cochleae
	<ol>
		<li>Transfer cochleae to a new 24-well plate using a transfer pipette.</li>
		<li>Add a drop of MSCGM to each cochlea to prevent any cochlea from drying out.</li>
		<li>Delicately orient the cochlea using ultra-fine forceps so that the oval and round windows are facing up. 
		NOTE: Take extreme care when directly handling each cochlea as the cochlea is easily damaged.</li>
		<li>Using a sterile 28.5-gauge insulin syringe with connected tubing, draw up 0.2 mL resuspended hWJCs (100,000 cells).</li>
		<li>Using sterilized fine forceps, position tubing over the oval window.</li>
		<li>Delicately and slowly perfuse the cochlea with 0.2 mL of resuspended hWJCs using a 28.5-gauge insulin syringe connected to polyethylene tubing (Outer Diameter: 0.61 mm, Inner Diameter: 0.28 mm) over approximately 5 min.</li>
		<li>After perfusion, add 0.8 mL of 37 &#176;C pre-warmed MSCGM to the well containing the cochlea to bring the total volume in the well to 1 mL.</li>
		<li>Repeat steps 4.3.3-4.3.7 for each cochlea, using a fresh syringe and tubing each time.</li>
		<li>Place perfused cochleae in a 5% CO2, 37 &#176;C cell culture incubator, and change media three times per week.</li>
	</ol>
	</li>
</ol>
5. Cochlea Harvest and Preservation
NOTE: Cochleae may be cultured and harvested at any time point up to 30 days post-perfusion.
<ol>
	<li>To preserve cochleae, fix in 4% paraformaldehyde in 1x PBS overnight at 4 &#176;C on a rocking platform.</li>
	<li>After fixation, wash cochleae with 1x PBS three times for 5 min each.</li>
	<li>Gradually dehydrate cochlea with ethanol and clear with an aliphatic hydrocarbon solvent before embedding in paraffin.</li>
	<li>Section samples to a thickness of 10 &#181;m using a microtome and mount on glass microscope slides. 
	NOTE: Samples are now ready for histological or immunohistochemical processing using standard protocols.</li>
</ol>

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The authors have nothing to disclose.

We have successfully demonstrated that native cochlear cells can be removed from the cochlea via a decellularization process, which allows for the use of the cochlea as an intricate, three-dimensional tissue scaffold. Santi et al.15 developed the initial method for decellularizing cochleae, and have accurately estimated the volumes of many cochlear structures through with the aid of light sheet microscopy23. Such early work served as a strong basis for the tissue e...

The cochlea is an intricate spiral structure found in the temporal bone. It is composed of an outer bony labyrinth and a concentric, inner membranous labyrinth1. The membranous labyrinth consists of three fluid spaces: Scala vestibuli, Scala media, and Scala tympani1. The scala media houses the sensory epithelium, which is composed of a multitude of cell types, but the sensory hair cells (HC), which transduce mechanical energy in sound waves to nerve impulses2, are of particular interest. Exposure to acoustic trauma3,4,5, medication6, disease7,8, and aging9 can all result in impaired auditory function via HC death. Hair cell loss in mammals is permanent, unlike avian HCs, which can regenerate after injury10.
A variety of contemporary research efforts have sought to restore lost HCs, although the specific experimental approaches vary. Manipulation of gene expression in the sensory epithelium and implantation of stem cells differentiated outside the body are dominant approaches in the field, although induction methods that seek to differentiate stem cells into cochlear organoids have been attempted11,12,13. Each of these approaches is either reliant directly on stem cells, or the developmental cues used by stem cells; however, a second shared, and potentially critical, element is the ECM of the cochlea itself14,15.
The ECM not only provides physical support for cells and tissue, which includes a surface for cell adhesion, proliferation, survival, and migration, but also plays critical roles in the development of HCs and the spiral ganglion15,16,17. Naturally occurring ECM provides inductive signals that may guide cell phenotype determination and/or cell adhesion, proliferation, and survival18. Consequently, the use of decellularized cochlea in combination with cultured hWJCs offer a unique opportunity to explore the role of the ECM and HC regeneration. HWJCs are a readily available, non-controversial cell type isolated from human umbilical cords that behave like mesenchymal stem cells19. HWJCs have shown the ability to differentiate down neurosensory cell lineages20,21. Thus, the current protocol details the isolation, decellularization, and perfusion of cochleae from C57BL mouse carcasses with hWJCs for inner ear tissue engineering.

<table>
	<tbody>
		<tr>
			<td>Allegra X-14R Centrifuge</td>
			<td>Beckman-Coulter</td>
			<td>B08861</td>
			<td></td>
		</tr>
		<tr>
			<td>Intramedic Semi-Rigid Tubing</td>
			<td>Becton Dickinson</td>
			<td>427401</td>
			<td></td>
		</tr>
		<tr>
			<td>New Brunswick Innova 2000 Orbital Shaler</td>
			<td>Eppendorf</td>
			<td>M1190-0002</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Scissors</td>
			<td>Fine Science Tools</td>
			<td>14060-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Forceps</td>
			<td>Fine Science Tools</td>
			<td>11370-40</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultra-Fine Forceps</td>
			<td>Fine Science Tools</td>
			<td>18155-13</td>
			<td></td>
		</tr>
		<tr>
			<td>50-mL Conical Tubes</td>
			<td>Fisher Scientific</td>
			<td>12565271</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri Dish</td>
			<td>Fisher Scientific</td>
			<td>FB087579B</td>
			<td></td>
		</tr>
		<tr>
			<td>U-100 Insulin Syringe</td>
			<td>Fisher Scientific</td>
			<td>14-829-1B</td>
			<td></td>
		</tr>
		<tr>
			<td>Scintillation Vial</td>
			<td>Fisher Scientific</td>
			<td>03-341-73</td>
			<td></td>
		</tr>
		<tr>
			<td>Rotator</td>
			<td>Fisher Scientific</td>
			<td>88-861-049</td>
			<td></td>
		</tr>
		<tr>
			<td>Transfer Pipette</td>
			<td>Fisher Scientific</td>
			<td>22-170-404</td>
			<td></td>
		</tr>
		<tr>
			<td>Razor Blade</td>
			<td>Fisher Scientific</td>
			<td>12-640</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic-Antimycotic</td>
			<td>Fisher Scientific</td>
			<td>15-240-062</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Fisher Scientific</td>
			<td>15-140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>24-Well Plate</td>
			<td>Fisher Scientific</td>
			<td>07-200-84</td>
			<td></td>
		</tr>
		<tr>
			<td>SuperFrost PLUS Glass Microscope Slides</td>
			<td>Fisher Scientific</td>
			<td>12-550-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Transfer Pipette</td>
			<td>Fisher Scientific</td>
			<td>22-170-404</td>
			<td></td>
		</tr>
		<tr>
			<td>ProLong Gold Antifade Mountant with DAPI</td>
			<td>Fisher Scientific</td>
			<td>P36935</td>
			<td></td>
		</tr>
		<tr>
			<td>Clear-Rite 3</td>
			<td>Fisher Scientific</td>
			<td>22-046341</td>
			<td></td>
		</tr>
		<tr>
			<td>Thermo Scientific Forma Series II 3110 Water-Jacekted CO2 Incubator</td>
			<td>Fisher Scientific</td>
			<td>13-998-078</td>
			<td></td>
		</tr>
		<tr>
			<td>Mesenchymal Stem Cell Growth Medium</td>
			<td>Lonza</td>
			<td>PT-3001</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA</td>
			<td>Lonza</td>
			<td>CC-3232</td>
			<td></td>
		</tr>
		<tr>
			<td>TPP T-75 Culture Flask</td>
			<td>MidSci</td>
			<td>TP90076</td>
			<td></td>
		</tr>
		<tr>
			<td>TPP T-150 Culture Flask</td>
			<td>MidSci</td>
			<td>TP90151</td>
			<td></td>
		</tr>
		<tr>
			<td>TPP T-300 Culture Flask</td>
			<td>MidSci</td>
			<td>TP90301</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection Microscope</td>
			<td>Nikon Instruments</td>
			<td>SMZ800</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon Eclipse Ts2R-FL Inverted Microscope</td>
			<td>Nikon Instruments</td>
			<td>MFA51010</td>
			<td></td>
		</tr>
		<tr>
			<td>NuAire Class II, Type A2 Biosafety Cabinet</td>
			<td>NuAire</td>
			<td>NU-425-600</td>
			<td></td>
		</tr>
		<tr>
			<td>1X PBS</td>
			<td>Sigma-Aldrich</td>
			<td>P5368-10PAK</td>
			<td></td>
		</tr>
		<tr>
			<td>1% SDS Solution</td>
			<td>Sigma-Aldrich</td>
			<td>436143-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>10% EDTA</td>
			<td>Sigma-Aldrich</td>
			<td>E9884-100G</td>
			<td></td>
		</tr>
	</tbody>
</table>

a protocol for decellularizing mouse cochleae for inner ear tissue engineering

In mammals, mechanosensory hair cells that facilitate hearing lack the ability to regenerate, which has limited treatments for hearing loss. Current regenerative medicine strategies have focused on transplanting stem cells or genetic manipulation of surrounding support cells in the inner ear to encourage replacement of damaged stem cells to correct hearing loss. Yet, the extracellular matrix (ECM) may play a vital role in inducing and maintaining function of hair cells, and has not been well investigated. Using the cochlear ECM as a scaffold to grow adult stem cells may provide unique insights into how the composition and architecture of the extracellular environment aids cells in sustaining hearing function. Here we present a method for isolating and decellularizing cochleae from mice to use as scaffolds accepting perfused adult stem cells. In the current protocol, cochleae are isolated from euthanized mice, decellularized, and decalcified. Afterward, human Wharton&#39;s jelly cells (hWJCs) that were isolated from the umbilical cord were carefully perfused into each cochlea. The cochleae were used as bioreactors, and cells were cultured for 30 days before undergoing processing for analysis. Decellularized cochleae retained identifiable extracellular structures, but did not reveal the presence of cells or noticeable fragments of DNA. Cells perfused into the cochlea invaded most of the interior and exterior of the cochlea and grew without incident over a duration of 30 days. Thus, the current method can be used to study how cochlear ECM affects cell development and behavior.

Using the methods presented here, successful decellularization of cochleae was assessed by examining the presence or absence of DNA through 4&#39;,6-diamidino-2-phenylindole (DAPI) staining. Cochleae were considered fully decellularized if DNA was not identified within the decellularized cochlea. A native cochlea from a previous experiment that did not undergo decellularization or decalcification was used as a positive control to illustrate the structures and cells traditionally found in ...

Watch this Scientific Journal Video about A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering at JoVE.com

A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

The goal of this protocol is to demonstrate an effective method to decellularize and decalcify mouse cochleae for utilization as scaffolds for tissue engineering applications.

In mammals, mechanosensory hair cells that facilitate hearing lack the ability to regenerate, which has limited treatments for hearing loss. Current ...

a-protocol-for-decellularizing-mouse-cochleae-for-inner-ear-tissue

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

10.7K Views.  University of Kansas Medical Center. The goal of this protocol is to demonstrate an effective method to decellularize and decalcify mouse cochleae for utilization as scaffolds for tissue engineering applications.

Video: A Protocol for Decellularizing Mouse Cochleae for Inner Ear Tissue Engineering

Fabrication of Biologically Derived Injectable Materials for Myocardial Tissue Engineering

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications. Briefly, decellularized tissue is lyophilized, milled, enzymatically digested, and then brought to physiological pH. The lyophilization removes all water content from the tissue, resulting in dry ECM that can be ground into a fine powder with a small mill. After milling, the ECM powder is digested with pepsin to form an injectable matrix. After adjustment to pH 7.4, the liquid matrix material can be injected into the myocardium. Results of previous characterization assays have shown that matrix gels produced from decellularized pericardial and myocardial tissue retain native ECM components, including diverse proteins, peptides and glycosaminoglycans. Given the use of this material for tissue engineering, in vivo characterization is especially useful; here, a method for performing an intramural injection into the left ventricular (LV) free wall is presented as a means of analyzing the host response to the matrix gel in a small animal model. Access to the chest cavity is gained through the diaphragm and the injection is made slightly above the apex in the LV free wall. The biologically derived scaffold can be visualized by biotin-labeling before injection and then staining tissue sections with a horse radish peroxidase-conjugated neutravidin and visualizing via diaminobenzidine (DAB) staining. Analysis of the injection region can also be done with histological and immunohistochemical staining. In this way, the previously examined pericardial and myocardial matrix gels were shown to form fibrous, porous networks and promote vessel formation within the injection region.

Methods for preparing an injectable matrix gel from decellularized tissue and injecting it into rat myocardium in vivo are described.

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications.  ...

Procedure for Lung Engineering

Lung tissue, including lung cancer and chronic lung diseases such as chronic obstructive pulmonary disease, cumulatively account for some 280,000 deaths annually; chronic obstructive pulmonary disease is currently the fourth leading cause of death in the United States1. Contributing to this mortality is the fact that lungs do not generally repair or regenerate beyond the microscopic, cellular level. Therefore, lung tissue that is damaged by degeneration or infection, or lung tissue that is surgically resected is not functionally replaced in vivo. To explore whether lung tissue can be generated in vitro, we treated lungs from adult rats using a procedure that removes cellular components to produce an acellular lung extracellular matrix scaffold. This scaffold retains the hierarchical branching structures of airways and vasculature, as well as a largely intact basement membrane, which comprises collagen IV, laminin, and fibronectin. The scaffold is mounted in a bioreactor designed to mimic critical aspects of lung physiology, such as negative pressure ventilation and pulsatile vascular perfusion. By culturing pulmonary epithelium and vascular endothelium within the bioreactor-mounted scaffold, we are able to generate lung tissue that is phenotypically comparable to native lung tissue and that is able to participate in gas exchange for short time intervals (45-120 minutes). These results are encouraging, and suggest that repopulation of lung matrix is a viable strategy for lung regeneration. This possibility presents an opportunity not only to work toward increasing the supply of lung tissue for transplantation, but also to study respiratory cell and molecular biology in vitro for longer time periods and in a more accurate microenvironment than has previously been possible.

We have developed a decellularized lung extracellular matrix and novel biomimetic bioreactor that can be used to generate functional lung tissue. By seeding cells into the matrix and culturing in the bioreactor, we generate tissue that demonstrates effective gas exchange when transplanted in vivo for short periods of time.

Lung tissue, including lung cancer and chronic lung diseases such as chronic obstructive pulmonary disease, cumulatively account for some 280,000 deaths ...

Elastomeric PGS Scaffolds in Arterial Tissue Engineering

Cardiovascular disease is one of the leading cause of mortality in the US and especially, coronary artery disease increases with an aging population and increasing obesity1. Currently, bypass surgery using autologous vessels, allografts, and synthetic grafts are known as a commonly used for arterial substitutes2. However, these grafts have limited applications when an inner diameter of arteries is less than 6 mm due to low availability, thrombotic complications, compliance mismatch, and late intimal hyperplasia3,4. To overcome these limitations, tissue engineering has been successfully applied as a promising alternative to develop small-diameter arterial constructs that are nonthrombogenic, robust, and compliant. Several previous studies have developed small-diameter arterial constructs with tri-lamellar structure, excellent mechanical properties and burst pressure comparable to native arteries5,6. While high tensile strength and burst pressure by increasing collagen production from a rigid material or cell sheet scaffold, these constructs still had low elastin production and compliance, which is a major problem to cause graft failure after implantation. Considering these issues, we hypothesized that an elastometric biomaterial combined with mechanical conditioning would provide elasticity and conduct mechanical signals more efficiently to vascular cells, which increase extracellular matrix production and support cellular orientation.
The objective of this report is to introduce a fabrication technique of porous tubular scaffolds and a dynamic mechanical conditioning for applying them to arterial tissue engineering. We used a biodegradable elastomer, poly (glycerol sebacate) (PGS)7 for fabricating porous tubular scaffolds from the salt fusion method. Adult primary baboon smooth muscle cells (SMCs) were seeded on the lumen of scaffolds, which cultured in our designed pulsatile flow bioreactor for 3 weeks. PGS scaffolds had consistent thickness and randomly distributed macro- and micro-pores. Mechanical conditioning from pulsatile flow bioreactor supported SMC orientation and enhanced ECM production in scaffolds. These results suggest that elastomeric scaffolds and mechanical conditioning of bioreactor culture may be a promising method for arterial tissue engineering.

Elastomeric PGS scaffolds with vascular smooth muscle cells cultured in a pulsatile flow bioreactor may lead to promising small-diameter arterial constructs with native ECM production in a relatively short culture period.

Cardiovascular disease is one of the leading cause of mortality in the US and especially, coronary artery disease increases with an aging population and ...

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 ...

Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do not have suitable autologous donor vessels due to disease progression or previous harvest. The aim of vascular tissue engineering is to develop a suitable alternative source for these bypass grafts. In addition, engineered vascular tissue may prove valuable as living vascular models to study cardiovascular diseases. Several promising approaches to engineering blood vessels have been explored, with many recent studies focusing on development and analysis of cell-based methods.2-5 Herein, we present a method to rapidly self-assemble cells into 3D tissue rings that can be used in vitro to model vascular tissues.
To do this, suspensions of smooth muscle cells are seeded into round-bottomed annular agarose wells. The non-adhesive properties of the agarose allow the cells to settle, aggregate and contract around a post at the center of the well to form a cohesive tissue ring.6,7 These rings can be cultured for several days prior to harvesting for mechanical, physiological, biochemical, or histological analysis. We have shown that these cell-derived tissue rings yield at 100-500 kPa ultimate tensile strength8 which exceeds the value reported for other tissue engineered vascular constructs cultured for similar durations (&lt;30 kPa).9,10 Our results demonstrate that robust cell-derived vascular tissue ring generation can be achieved within a short time period, and offers the opportunity for direct and quantitative assessment of the contributions of cells and cell-derived matrix (CDM) to vascular tissue structure and function.

This article outlines a versatile method to create cell-derived tissue rings by cellular self-assembly. Smooth muscle cells seeded into ring-shaped agarose wells aggregate and contract to form robust three-dimensional (3D) tissues within 7 days. Millimeter-scale tissue rings are conducive to mechanical testing and serve as building blocks for tissue assembly.

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do ...

Postproduction Processing of Electrospun Fibres for Tissue Engineering

Electrospinning is a commonly used and versatile method to produce scaffolds (often biodegradable) for 3D tissue engineering.1, 2, 3 Many tissues in vivo undergo biaxial distension to varying extents such as skin, bladder, pelvic floor and even the hard palate as children grow. In producing scaffolds for these purposes there is a need to develop scaffolds of appropriate biomechanical properties (whether achieved without or with cells) and which are sterile for clinical use. The focus of this paper is not how to establish basic electrospinning parameters (as there is extensive literature on electrospinning) but on how to modify spun scaffolds post production to make them fit for tissue engineering purposes - here thickness, mechanical properties and sterilisation (required for clinical use) are considered and we also describe how cells can be cultured on scaffolds and subjected to biaxial strain to condition them for specific applications.
Electrospinning tends to produce thin sheets; as the electrospinning collector becomes coated with insulating fibres it becomes a poor conductor such that fibres no longer deposit on it. Hence we describe approaches to produce thicker structures by heat or vapour annealing increasing the strength of scaffolds but not necessarily the elasticity. Sequential spinning of scaffolds of different polymers to achieve complex scaffolds is also described. Sterilisation methodologies can adversely affect strength and elasticity of scaffolds. We compare three methods for their effects on the biomechanical properties on electrospun scaffolds of poly lactic-co-glycolic acid (PLGA).
Imaging of cells on scaffolds and assessment of production of extracellular matrix (ECM) proteins by cells on scaffolds is described. Culturing cells on scaffolds in vitro can improve scaffold strength and elasticity but the tissue engineering literature shows that cells often fail to produce appropriate ECM when cultured under static conditions. There are few commercial systems available that allow one to culture cells on scaffolds under dynamic conditioning regimes - one example is the Bose Electroforce 3100 which can be used to exert a conditioning programme on cells in scaffolds held using mechanical grips within a media filled chamber.4 An approach to a budget cell culture bioreactor for controlled distortion in 2 dimensions is described. We show that cells can be induced to produce elastin under these conditions. Finally assessment of the biomechanical properties of processed scaffolds cultured with or without cells is described.

Electrospun scaffolds can be processed post production for tissue engineering applications. Here we describe methods for spinning complex scaffolds (by consecutive spinning), for making thicker scaffolds (by multi-layering using heat or vapour annealing), for achieving sterility (aseptic production or sterilisation post production) and for achieving appropriate biomechanical properties.

Electrospinning is a commonly used and versatile method to produce  scaffolds (often biodegradable) for 3D tissue engineering.1, 2, 3 Many tissues in vivo ...

Capillary Force Lithography for Cardiac Tissue Engineering

Cardiovascular disease remains the leading cause of death worldwide1. Cardiac tissue engineering holds much promise to deliver groundbreaking medical discoveries with the aims of developing functional tissues for cardiac regeneration as well as in vitro screening assays. However, the ability to create high-fidelity models of heart tissue has proven difficult. The heart&#8217;s extracellular matrix (ECM) is a complex structure consisting of both biochemical and biomechanical signals ranging from the micro- to the nanometer scale2. Local mechanical loading conditions and cell-ECM interactions have recently been recognized as vital components in cardiac tissue engineering3-5.
A large portion of the cardiac ECM is composed of aligned collagen fibers with nano-scale diameters that significantly influences tissue architecture and electromechanical coupling2. Unfortunately, few methods have been able to mimic the organization of ECM fibers down to the nanometer scale. Recent advancements in nanofabrication techniques, however, have enabled the design and fabrication of scalable scaffolds that mimic the in vivo structural and substrate stiffness cues of the ECM in the heart6-9.
Here we present the development of two reproducible, cost-effective, and scalable nanopatterning processes for the functional alignment of cardiac cells using the biocompatible polymer poly(lactide-co-glycolide) (PLGA)8 and a polyurethane (PU) based polymer. These anisotropically nanofabricated substrata (ANFS) mimic the underlying ECM of well-organized, aligned tissues and can be used to investigate the role of nanotopography on cell morphology and function10-14.
Using a nanopatterned (NP) silicon master as a template, a polyurethane acrylate (PUA) mold is fabricated. This PUA mold is then used to pattern the PU or PLGA hydrogel via UV-assisted or solvent-mediated capillary force lithography (CFL), respectively15,16. Briefly, PU or PLGA pre-polymer is drop dispensed onto a glass coverslip and the PUA mold is placed on top. For UV-assisted CFL, the PU is then exposed to UV radiation (&#955; = 250-400 nm) for curing. For solvent-mediated CFL, the PLGA is embossed using heat (120 &#176;C) and pressure (100 kPa). After curing, the PUA mold is peeled off, leaving behind an ANFS for cell culture. Primary cells, such as neonatal rat ventricular myocytes, as well as human pluripotent stem cell-derived cardiomyocytes, can be maintained on the ANFS2.

In this protocol, we demonstrate the fabrication of biomimetic cardiac cell culture substrata made from two distinct polymeric materials using capillary force lithography. The described methods provide a scalable, cost-effective technique to engineer the structure and function of macroscopic cardiac tissues for in vitro and in vivo applications.

Cardiovascular disease remains the leading cause of death worldwide1. Cardiac tissue engineering holds much promise to deliver groundbreaking medical ...

Protocol for Relative Hydrodynamic Assessment of Tri-leaflet Polymer Valves

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of tri-leaflet polymer valve prostheses. However, identification of a protocol for initial assessment of polymer valve hydrodynamic functionality is paramount during the early stages of the design process. Traditional in vitro pulse duplicator systems are not configured to accommodate flexible tri-leaflet materials; in addition, assessment of polymer valve functionality needs to be made in a relative context to native and prosthetic heart valves under identical test conditions so that variability in measurements from different instruments can be avoided. Accordingly, we conducted hydrodynamic assessment of i) native (n = 4, mean diameter, D = 20 mm), ii) bi-leaflet mechanical (n= 2, D = 23 mm) and iii) polymer valves (n = 5, D = 22 mm) via the use of a commercially available pulse duplicator system (ViVitro Labs Inc, Victoria, BC) that was modified to accommodate tri-leaflet valve geometries. Tri-leaflet silicone valves developed at the University of Florida comprised the polymer valve group. A mixture in the ratio of 35:65 glycerin to water was used to mimic blood physical properties. Instantaneous flow rate was measured at the interface of the left ventricle and aortic units while pressure was recorded at the ventricular and aortic positions. Bi-leaflet and native valve data from the literature was used to validate flow and pressure readings. The following hydrodynamic metrics were reported: forward flow pressure drop, aortic root mean square forward flow rate, aortic closing, leakage&#160;and regurgitant volume, transaortic closing, leakage, and total energy losses. Representative results indicated that hydrodynamic metrics from the three valve groups could be successfully obtained by incorporating a custom-built assembly into a commercially available pulse duplicator system and subsequently, objectively compared to provide insights on functional aspects of polymer valve design.

There has been renewed interest in developing polymer valves. Here, the objectives are to demonstrate the feasibility of modifying a commercial pulse duplicator to accommodate tri-leaflet geometries and to define a protocol to present polymer valve hydrodynamic data in comparison to native and prosthetic valve data collected under near-identical conditions.

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of ...

Design of a Biaxial Mechanical Loading Bioreactor for Tissue Engineering

We designed a loading device that is capable of applying uniaxial or biaxial mechanical strain to a tissue engineered biocomposites fabricated for transplantation. While the device primarily functions as a bioreactor that mimics the native mechanical strains, it is also outfitted with a load cell for providing force feedback or mechanical testing of the constructs. The device subjects engineered cartilage constructs to biaxial mechanical loading with great precision of loading dose (amplitude and frequency) and is compact enough to fit inside a standard tissue culture incubator. It loads samples directly in a tissue culture plate, and multiple plate sizes are compatible with the system. The device has been designed using components manufactured for precision-guided laser applications. Bi-axial loading is accomplished by two orthogonal stages. The stages have a 50 mm travel range and are driven independently by stepper motor actuators, controlled by a closed-loop stepper motor driver that features micro-stepping capabilities, enabling step sizes of less than 50 nm. A polysulfone loading platen is coupled to the bi-axial moving platform. Movements of the stages are controlled by Thor-labs Advanced Positioning Technology (APT) software. The stepper motor driver is used with the software to adjust load parameters of frequency and amplitude of both shear and compression independently and simultaneously. Positional feedback is provided by linear optical encoders that have a bidirectional repeatability of 0.1 &mu;m and a resolution of 20 nm, translating to a positional accuracy of less than 3 &mu;m over the full 50 mm of travel. These encoders provide the necessary position feedback to the drive electronics to ensure true nanopositioning capabilities. In order to provide the force feedback to detect contact and evaluate loading responses, a precision miniature load cell is positioned between the loading platen and the moving platform. The load cell has high accuracies of 0.15% to 0.25% full scale.

We designed a novel mechanical loading bioreactor that can apply uniaxial or biaxial mechanical strain to a cartilage biocomposite prior to transplantation into an articular cartilage defect.

We  designed a loading device that is capable of applying uniaxial or biaxial  mechanical strain to a tissue engineered biocomposites fabricated for  ...

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. Collagen has been extensively used for a wide range of biomedical applications and is considered a valid alternative to synthetic materials due to its inherent biocompatibility (i.e., low antigenicity, inflammation, and cytotoxic responses). However, the limited mechanical properties and the related low hand-ability of collagen gels have hampered their use as scaffold materials for vascular tissue engineering. Therefore, the rationale behind this work was first to engineer cellularized collagen gels into a tubular-shaped geometry and second to enhance smooth muscle cells driven reorganization of collagen matrix to obtain tissues stiff enough to be handled.
The strategy described here is based on the direct assembling of collagen and smooth muscle cells (construct) in a 3D cylindrical geometry with the use of a molding technique. This process requires a maturation period, during which the constructs are cultured in a bioreactor under static conditions (without applied external dynamic mechanical constraints) for 1 or 2 weeks. The &#8220;static bioreactor&#8221; provides a monitored and controlled sterile environment (pH, temperature, gas exchange, nutrient supply and waste removal) to the constructs. During culture period, thickness measurements were performed to evaluate the cells-driven remodeling of the collagen matrix, and glucose consumption and lactate production rates were measured to monitor the cells metabolic activity. Finally, mechanical and viscoelastic properties were assessed for the resulting tubular constructs. To this end, specific protocols and a focused know-how (manipulation, gripping, working in hydrated environment, and so on) were developed to characterize the engineered tissues.

In this work, we present a technique for the rapid fabrication of living vascular tissues by direct culturing of collagen, smooth muscle cells and endothelial cells. In addition, a new protocol for the mechanical characterization of engineered vascular tissues is described.

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. ...