Name: procedure for lung engineering
Uploaded: 2011-03-08T12:00:00
Duration: 12 min 50 s
Description: Watch this Scientific Journal Video about Procedure for Lung Engineering at JoVE.com

We thank Maegan B. Colehour for help with bioreactor development. These studies were funded by Yale University Department of Anesthesia and by NIH grant HL 098220 (to L.E.N.). T.H.P. was supported by NIH T32 GM007171.

1. Bioreactor Assembly
 Detailed figure(s) of the bioreactor design and assembly for both decellularization and culture are provided in Figures 1 and 2, respectively. All components must be sterilized prior to assembly of the bioreactor. The following specific points are noted:
CONNECTIONS:
<ol>
 <li>The arterial cannula consists of a luer-lock fitting connected to a Y-splitter by a short segment of tubing. The luer-lock connector is attached to the perfusion tubing, and the segment of the Y that is not sutured to the pulmonary artery is connected to a one-way valve. The one-way valve is oriented such that fluid can be drawn up into the tubing (in the opposite direction as perfusion), but during organ perfusion all medium flows into the lung.</li>
 <li>The tracheal cannula also consists of a luer-lock connector linked to a Y-splitter with tubing. The luer-lock connects to a breathing loop between the trachea reservoir and the main chamber. The segment of the Y-connector that is not sutured to the trachea is also connected to a one-way valve. The one-way valve is oriented in the same fashion as the arterial cannula.</li>
</ol>
FUNCTIONS:
<ol start="3">
 <li>The one-way valves in the arterial and tracheal cannulae are utilized to clear air bubbles in the tubing by enabling reversal of flow in the tubing and therefore permitting air bubbles to be removed.</li>
 <li>The "breathing loop" includes two one-way valves, positioned such that medium follows a different path into and out of the lung. A more detailed description of this feature is provided in a prior publication from our laboratory2.</li>
 <li>Vascular perfusion is provided using a roller pump. Medium is perfused into the pulmonary artery via the attached cannula, flows through the lung vasculature, and out the pulmonary vein directly into the main bioreactor, where medium is drawn up for perfusion.</li>
</ol>
2. Organ Harvest
<ol>
 <li>Euthanize adult (3-6 month old) Fischer 344 rats by sodium pentobarbital overdose, according to guidelines set forth by the American Veterinary Medical Association (60 mg/kg IP). Note: the pentobarbital solution includes heparin at 100 units/ml for anticoagulation.</li>
 <li>Spray or wipe the chest &amp; abdomen with 70% ethanol.</li>
 <li>Open the thoracic cavity, exposing the heart and lungs, taking care to not damage the lungs. Carefully make a small window through the diaphragm into the thoracic cavity, causing the lungs to retract, and then expand this incision horizontally to expose the bases of the lungs. Make two incisions vertically through the ribs, and retract the chest cage to expose the heart and lungs.</li>
 <li>Prepare a gravity perfusion system, i.e. using a syringe with plunger removed, a 3-way stopcock, and ~20cm of tubing with a 1.5inch, 21-gauge needle. Maintain the syringe at ~20cm above the animal using a ring stand. Ensure that all air is removed from the perfusion tubing.</li>
 <li>Cut the right atrium to drain blood, preventing it from returning to the lungs. Cutting the left atrium to allow easy drainage of blood and perfusate from the lungs may also be helpful, but is not necessary.</li>
 <li>Insert the needle into the base of the right ventricle and open the stopcock to perfuse the lungs with a solution of heparin and sodium nitroprusside (50U/ml and 1ug/ml, respectively) in PBS. Confirm that the pulmonary artery fills with perfusion fluid, and that blood is clearing from lungs. If needed, refill the perfusion syringe with additional fluid. Typically only ~10ml is required.</li>
 <li>Continue perfusion until the lungs are clear of blood, then stop perfusion.</li>
 <li>Dissect the trachea free, up into the neck as far as possible. Ensure trachea is separated from the esophagus. Dissect all remaining connections to the heart, lungs and trachea, and remove en bloc.</li>
 <li>Using a scalpel or sharp scissors, cut off the apex of the heart, exposing the right and left ventricles.</li>
 <li>Cannulate the pulmonary trunk via the right ventricle, and suture in place. Remove any excess left ventricular tissue.</li>
 <li>Cannulate the trachea and suture in place. Make sure that both the tracheal and pulmonary arterial cannulae are positioned such that there is no torsional stress on the trachea, lungs, or great vessels (Figure 1).</li>
 <li>Ensure that there are no air bubbles in the arterial cannula, since air bubbles trapped in the system may prevent constant flow of through the organ. In some cases, an air bubble can completely halt fluid flow.To accomplish this, place the heart/lung bloc into a jar containing PBS. Use a syringe with needle to inject a small amount of PBS into the heart cannula to expel any bubbles.</li>
 <li>Lavage the airways with PBS 4-5 times, to remove as much air as possible from the lung.</li>
 <li>After lavage steps are complete, inflate the lung with PBS containing sodium nitroprusside (SNP) at 1ug/ml. Put a stopper on the tracheal cannula, so the solution remains in the lung. Allow the lung to incubate for up to 30 minutes, as the same solution flows through the vasculature via the pulmonary artery, to permit vasodilation.</li>
 <li>Connect heart/lungs to a bioreactor cap using the luer-lock connections linked to the Y-shaped cannulae, described in "Bioreactor Assembly" above. (Figure 1). </li>
</ol>
3. Organ Decellularization
<ol>
 <li>Connect the cap (with lungs attached) to decellularization apparatus (Figure 1). The pulmonary artery cannula should connect to the perfusion line, and the tracheal cannula should float free. Ensure all lines are clear of air. As described above, air in the lines can be a source of poor decellularization by preventing flow of decllularization fluid. Air trapped in the system can also persist into culture time, when it may have a negative impact on cell survival3.</li>
 <li>Inflate the lung with PBS/SNP until lungs are full, but not over-inflated. Immediately cap the tracheal cannula so that the lung remains inflated.</li>
 <li>Perfuse lung with PBS/SNP for at least 15 min at ~15 mmHg (20cm H2O pressure). After 15min or longer, remove the stopper from the tracheal cannula to allow the lung to deflate.</li>
 <li>Continue perfusion with PBS/SNP for 30min. If necessary, refill PBS/SNP to ensure perfusion pressure is maintained at 10-15 mmHg.</li>
 <li>Begin perfusion with decellularization solution (8mM CHAPS, 1M NaCl, 25mM EDTA in 1X PBS). Take care to ensure all lines are clear of air. A vacuum system may be helpful to provide suction.</li>
</ol>
Perfuse with decellularization solution until 500ml of solution has perfused through the lung. Optimal pressure is &lt;15 mmHg (~20 cm H2O). This typically will require 2.5 hours. Flow rates are typically very slow (0.2-0.5ml/minute) initially, and rapidly increase during the second hour to approximately 1ml/minute or greater. Periodically, remove used decellularization fluid from bioreactor, ensuring enough fluid remains to support the lung and tracheal cannula.
 4. Organ rinsing and sterilization
<ol>
 <li>Transfer lung and bioreactor to a tissue culture hood. Begin rinsing with sterile PBS by removing the 500ml jar that contained the decellularization fluid and replacing with a sterile jar containing up to 1L of sterile PBS. Use vacuum suction to ensure lines are clear of air.</li>
 <li>Perfuse PBS through the vasculature at 10-15 mmHg, in the same manner as for decellularization. Periodically, remove waste PBS from bioreactor and replace and/or refill the PBS jar with fresh, sterile PBS. It is advisable to use sterile technique.</li>
 <li>Continue rinsing until at least 2.5 L of sterile PBS have perfused the lung.</li>
 <li>Transfer lung to a new, sterile bioreactor system containing fresh PBS. Ensure that both the entire perfusion loop and the entire airway lines are filled with fluid. All subsequent steps will use a pulsatile pump to perfuse the lungs at 5ml/min.</li>
 <li>Sterilize the scaffold, either via overnight perfusion with PBS + 10% FBS + 10% pen-strep solution or for 3 hours with 0.1% peracetic acid in PBS. The latter will require rinsing the lung with 3 changes of 250ml PBS over several hours to remove residual acid. For each rinse, the lung should be ventilated as well as perfused to ensure that all parts of the tissue are rinsed thoroughly.</li>
 <li>Transfer the lung to a 37&deg;C incubator and perfuse with PBS that has 10% FBS and 10% penicillin/streptomycin for ~1hr or until temperature is equilibrated, in preparation for benzonase treatment.</li>
 <li>Treat lung with benzonase to remove remnant DNA:
 <ol>
 <li>Warm the benzonase buffer (see Table of Reagents) to 37&deg;C.</li>
 <li>For each lung, fill one 10ml syringe with benzonase buffer only and one 10ml syringe with 90U/ml benzonase in buffer.</li>
 <li>Stop perfusion of the lung.</li>
 <li>Inflate the airway with benzonase buffer.</li>
 <li>Allow the lung to deflate (for ~1 minute). Then, inflate the lung with the benzonase solution. During the inflation with benzonase buffer &amp; benzonase, try to avoid injecting any air into the lung.</li>
 <li>Allow the lung to sit without perfusion or ventilation at 37&deg;C for 1 hour after inflating with benzonase.</li>
 </ol>
 </li>
 <li>Resume perfusion with the PBS + 10% FBS, 10% penicillin/streptomycin that is already in the bioreactor, and continue overnight. The following day, the lung can either be stored at 4&deg;C (for up to 3 months) or prepared for cell seeding.</li>
 <li>To prepare the scaffold for cellular repopulation, replace the PBS/FBS/benzonase solution with ~250ml of culture medium. Perfuse for at least one hour before cells are introduced, and replace with fresh culture medium directly before seeding cells.</li>
</ol>
5. Recellularization
The choice of cell source for organ reseeding is left up to individual investigators. Many cell sources can be utilized, including commercially available populations, freshly isolated neonatal or fetal pulmonary cells, embryonic stem cells, or commercially available cell sources. Specific isolation protocols for these cell populations can be found elsewhere4,5,6. Here, we provide instructions on how to seed both endothelial and epithelial cell populations.
 Endothelial seeding:
<ol>
 <li>Prepare a suspension of desired endothelial cell population, in appropriate culture medium. Filter cell suspension through a 40um cell strainer to remove cell clumps. A typical endothelial seeding in our laboratory would utilize approximately 30 million rat lung microvascular endothelial cells in 60ml of culture medium.</li>
 <li>Dispense cell suspension into a small reservoir temporarily incorporated into the perfusion loop of the bioreactor (Figure 2). Ensure that the tubing from this reservoir and the entire perfusion loop is clear of air bubbles.</li>
 <li>Infuse cells into the pulmonary artery at 3ml/min using a roller pump.</li>
 <li>After cell infusion, continue perfusion with medium recirculated from the main bioreactor at desired rate.</li>
 <li>On a daily basis, ensure that the perfusion tubing is clear of air bubbles.</li>
 <li>Medium should be replaced with fresh medium regularly; this is often done every 3-4 days.</li>
</ol>
Epithelial seeding:
<ol>
 <li>Prepare a cell suspension of the desired epithelial cell population. In our laboratory, this typically consists of approximately 50-100 million neonatal rat pulmonary cells. Filter the population through 40um cell strainer and suspend in 15ml of culture medium in a syringe. Fill the airway reservoir with 80ml of culture medium.</li>
 <li>Place the bioreactor in a 37&deg;C tissue culture incubator, and connect the syringe pump for ventilation. Ensure all ventilation lines are clear of air.
 <ol>
 <li>Seed the cells into the lung by injecting 15ml of cell suspension as a single bolus into the tracheal cannula.</li>
 <li>Immediately begin a single, slow breath using the syringe pump. This breath is administered by withdrawing 60ml of air from the main bioreactor at 3ml/minute, thus lasting approximately 20 minutes. Ensure the air filters on the main bioreactor are capped off immediately after injecting the cell suspension and before beginning the slow breath.</li>
 </ol>
 </li>
 <li>Allow the lungs to sit statically for approximately 18hrs, and then begin slow vascular perfusion (approximately 0.5 ml/min).</li>
</ol>
6. Organ Culture
Although the details of perfusion and ventilation will vary based on experimental design, the following points are noted:
<ol>
 <li>During endothelial culture, perfusion is typically performed at 1-3 ml/min using a roller pump. During epithelial culture, ventilation is typically provided at a continuous rate of 1 breath per minute using a syringe pump. Withdrawal of 5-10ml of air from the main bioreactor is typically required to effect lung ventilation at a normal tidal volume. Due to the need to maintain the bioreactor airtight during ventilation, ventilation should be paused daily and the air in the main chamber should be exchanged. All air in the system is room air, which is approximately 21% oxygen by partial pressure. The 5-10ml withdrawal of air from the bioreactor (which induces a 5-10ml inspiration of fluid by the lung) is based on the size of the lung and how many lobes are being cultured (lobes can be tied off for analysis, while the remaining lobes continue in culture). The amount of air withdrawn by the syringe pump is chosen to approximate the "tidal volume" of the cultured lung.</li>
 <li>Medium should be changed approximately every 3-4 days during culture.</li>
 <li>During co-culture of both epithelium and endothelium, experiments in our laboratory typically first seed the epithelium for 4-8 days, during which time the engineered tissue is ventilated. The endothelium is then seeded via perfusion, after which the tissue is both perfused and ventilated.</li>
</ol>
7. Representative Results:
Decellularization
When the protocol is performed correctly, the freshly extracted lungs should hold air without leaking. Inflating them with air while submerged in liquid can check this - there should not be any bubbles indicating air leaks. The subsequent decellularization should allow ~500ml of decellularization fluid to flow through the lungs over the course of 2.5 - 3 hours at 37&deg;C, and PBS should ultimately be able to flow through the lungs at about 10 ml/min (under ~15 mm Hg of hydrostatic pressure) at the end of rinsing. After treatment with 0.1% peracetic acid and benzonase, lungs can be stored at 4&deg;C for up to 3 months, and still remain suitable for recellularization.
The final decellularized extracellular matrix should be completely devoid of cellular materials, and retain the gross, microscopic and ultrastructural characteristics of native lung. Insufficient decellularization or rinsing may result in remnant DNA "sticking" to the scaffold, which can be visualized with a standard hematoxylin and eosin stain (see Figure 3 for comparison).
Repopulation of acellular matrix and culture of lung tissue
If cells are freshly isolated from 7 day old neonatal rat pups, as described in the online supplement accompanying the work by Petersen and colleagues4, one can expect a cell yield of 120-150 millions cells per litter of 10 pups (just over 10 million cells per neonate).
Optimal conditions for cell seeding and subsequent culture of the lung in the bioreactor should yield well-distributed cells within all 5 lobes of the lung, and should provide coverage of approximately 70% of the extracellular matrix scaffold (Figure 3). The cultured cell population will be positive for key respiratory cell markers such as pro-secretory protein-C (SPC), Clara cell secretory protein (CCSP), and aquaporin-5 (AQP), in order of relative abundance (Figure 4).


<img src="/files/ftp_upload/2651/2651fig1.jpg" alt="Figure 1" /> 
Figure 1. Cannulae positions and decellularization bioreactor

<img src="/files/ftp_upload/2651/2651fig2.jpg" alt="Figure 2" /> 
Figure 2. Bioreactor used for seeding and culture of engineered lung tissue

<img src="/files/ftp_upload/2651/2651fig3.jpg" alt="Figure 3" /> 
Figure 3. Histology of native, decellularized, and repopulated lung

<img src="/files/ftp_upload/2651/2651fig4.jpg" alt="Figure 4" /> 
Figure 4. Immunofluorescence staining for key lung markers

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regeneration+and+orthotopic+transplantation+of+a+bioartificial+lung.">Regeneration and orthotopic transplantation of a bioartificial lung.</a> Nat Med. 16, 927-9233 (2010).</li><li>Cortiella, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influence+of+acellular+natural+lung+matrix+on+murine+embryonic+stem+cell+differentiation+and+tissue+formation.">Influence of acellular natural lung matrix on murine embryonic stem cell differentiation and tissue formation.</a> Tissue Eng. 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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organ+reengineering+through+development+of+a+transplantable+recellularized+liver+graft+using+decellularized+liver+matrix.">Organ reengineering through development of a transplantable recellularized liver graft using decellularized liver matrix.</a> Nat. Med. 16, 814-820 (2010).</li><li>Shannon, J. M., Mason, R. J., Jennings, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+differentiation+of+alveolar+type+II+epithelial+cells+in+vitro:+effects+of+cell+shape,+cell-matrix+interactions+and+cell-cell+interactions.">Functional differentiation of alveolar type II epithelial cells in vitro: effects of cell shape, cell-matrix interactions and cell-cell interactions.</a> Biochim. Biophys. 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L.E.N. holds stock in Humacyte, a regenerative medicine company. Humacyte did not fund these studies, and did not affect the design, interpretation, or reporting of any of the experiments or methods described. The authors (L.E.N., T.H.P., E.A.C.) and Yale University have filed a patent application related to tissue engineering of lungs.

The most critical aspects of the system presented here include maintenance of sterility, and close monitoring of pressures applied to the vascular bed throughout the process of preparing and seeding the scaffold and culturing the repopulated lung. Sterility is best maintained by autoclaving all materials prior to use, and by mounting the lung in a closed system shortly after explant and avoiding subsequent breach of this barrier. After the decellularized matrix has been thoroughly rinsed and transferred to a sterile bioreactor for culture, the silicone cap and other seals and connections should not be disturbed or removed. To keep pressures in check, flow driven by gravity is preferable whenever possible. When a pump is required for recirculation of fluid, beginning after rinsing with PBS and continuing during culture, we recommend directly measuring the pressure just before the fluid enters the pulmonary artery with a pressure transducer. The magnitude of the applied pressure should not exceed 15 mm Hg.
Recellularized lungs can be cultured for varying time periods, typically ranging from 4 days to up to 3 weeks. Vascular perfusion is typically accomplished at 1-3 ml/min during endothelial culture, while ventilation is typically applied at a rate of 1 breath/min during epithelial culture. During combined culture periods, simultaneous ventilation and perfusion is appropriate. Ventilation can be performed with either liquid medium or air.
Over the course of the last several decades, several groups have done important tissue engineering work showing the feasibility of differentiating lung epithelium in vitro and of replicating several aspects of lung microanatomy7,8,9,10. However, until recently,4,5 none of these attempts to engineer lung tissue had resulted in an implantable organ that was able to maintain separation between the blood and airway compartments and that could participate in gas exchange. Therefore, though the methods described are only an initial step toward the long-term goal of generating functional lung tissue, this work is an encouraging step toward the possibility of increasing the amount of lung tissue available for transplant. Moreover, this work elaborates work done by Ott et al. and Uygun and colleagues11,12, demonstrating the efficacy of a decellularized extracellular matrix as a scaffold for tissue engineering complex three-dimensional structures and supporting the growth and survival of various types of cells. This work is also significant for its contribution to the armamentarium of respiratory cell and molecular biologists. By providing a unique, three-dimensional environment that can also provide appropriate mechanical stimuli, and that does not share the attendant hazard of rapid de-differentiation that one might encounter when culturing rodent epithelium in the laboratory with more traditional methods13,scientists could use our system to gain new insight into cell-cell and cell-matrix interactions that play a role in cell differentiation and function. This knowledge may be particularly powerful if used as leverage to guide the fate of various stem cell populations, as Cortiella's group has demonstrated in initial studies14.

<table border="1">
<tbody>
 <tr>
 <th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 <th>Comments (optional)</th>
 </tr>
 <tr>
 <td>Euthanasia</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Heparin</td>
 <td>Sigma-Aldrich</td>
 <td>H4784</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sodium nitroprusside</td>
 <td>Fluka</td>
 <td>71778</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Euthasol euthanasia solution</td>
 <td>Virbac AH</td>
 <td>710101</td>
 <td>390 mg/ml pentobarbital sodium stock solution should be diluted appropriately for IP administration </td>
 </tr>
 <tr>
 <td colspan="4">&nbsp;</td>
 </tr>
 <tr>
 <td>Decell Solution</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>CHAPS</td>
 <td>Sigma-Aldrich</td>
 <td>C3023</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>American Bioanalytical</td>
 <td>AB01915</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>EDTA</td>
 <td>Sigma</td>
 <td>E5134</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>NaOH</td>
 <td>J.T. Baker</td>
 <td>3722-01</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>1X PBS</td>
 <td>Gibco</td>
 <td>14190</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td colspan="4">&nbsp;</td>
 </tr>
 <tr>
 <td>Bioreactor Components</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>480 ml jar </td>
 <td>Cole Parmer</td>
 <td>EW-3460560</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Silicone stopper, size 14</td>
 <td>Cole parmer</td>
 <td>EW-06298-26</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Y-connectors</td>
 <td>Cole Parmer</td>
 <td>ED-30614-08</td>
 <td>Used for arterial and tracheal cannulae</td>
 </tr>
 <tr>
 <td>Silicone tubing</td>
 <td>Masterflex</td>
 <td>96420-14; 16</td>
 <td>L/S 14; 16</td>
 </tr>
 <tr>
 <td>Pressure Transducers </td>
 <td>Edwards Lifesciences</td>
 <td>PX-212</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Check valve</td>
 <td>Cole Parmer</td>
 <td>EW-98553-20</td>
 <td>One-way valve</td>
 </tr>
 <tr>
 <td>4-way stopcocks</td>
 <td>Edwards Lifesciences</td>
 <td>594WSC</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Syringe filter</td>
 <td>Cole Parmer</td>
 <td>2915-08</td>
 <td>PTFE, 0.2&mu;m</td>
 </tr>
 <tr>
 <td colspan="4">&nbsp;</td>
 </tr>
 <tr>
 <td>Decell and rinsing apparati (additions to bioreactor)</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>500 and 1000 ml glass bottles</td>
 <td>Corning</td>
 <td>1395-500; -1L</td>
 <td>Used for decell and rinsing by gravity</td>
 </tr>
 <tr>
 <td colspan="4">&nbsp;</td>
 </tr>
 <tr>
 <td>Benzonase Treatment</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Penicillin/streptomycin</td>
 <td>Gibco</td>
 <td>15140122</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>FBS</td>
 <td>Hyclone</td>
 <td>SH30071.03</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tris-HCl</td>
 <td>American Bioanalytical</td>
 <td>AB14043</td>
 <td>1M, pH 8.0</td>
 </tr>
 <tr>
 <td>MgCl2</td>
 <td>J.T. Baker</td>
 <td>2444-01</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>BSA</td>
 <td>Sigma</td>
 <td>A9647</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Benzonase Nuclease</td>
 <td>Sigma</td>
 <td>E1014</td>
 <td>Endonuclease used to remove remnant DNA from matrix</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Procedure for Lung Engineering at JoVE.com

Procedure for Lung Engineering

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 ...

Research

JoVE Journal

Bioengineering

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 ...

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 ...

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. ...

Synthesis of Keratin-based Nanofiber for Biomedical Engineering

Electrospinning, due to its versatility and potential for applications in various fields, is being frequently used to fabricate nanofibers. Production of these porous nanofibers is of great interest due to their unique physiochemical properties. Here we elaborate on the fabrication of keratin containing poly (&#949;-caprolactone) (PCL) nanofibers (i.e., PCL/keratin composite fiber). Water soluble keratin was first extracted from human hair and mixed with PCL in different ratios. The blended solution of PCL/keratin was transformed into nanofibrous membranes using a laboratory designed electrospinning set up. Fiber morphology and mechanical properties of the obtained nanofiber were observed and measured using scanning electron microscopy and tensile tester. Furthermore, degradability and chemical properties of the nanofiber were studied by FTIR. SEM images showed uniform surface morphology for PCL/keratin fibers of different compositions. These PCL/keratin fibers also showed excellent mechanical properties such as Young&#39;s modulus and failure point. Fibroblast cells were able to attach and proliferate thus proving good cell viability. Based on the characteristics discussed above, we can strongly argue that the blended nanofibers of natural and synthetic polymers can represent an excellent development of composite materials that can be used for different biomedical applications.

Electrospun nanofibers have a high surface area to weight ratio, excellent mechanical integrity, and support cell growth and proliferation. These nanofibers have a wide range of biomedical applications. Here we fabricate keratin/ PCL nanofibers, using the electrospinning technique, and characterize the fibers for possible applications in tissue engineering.

Electrospinning, due to its versatility and potential for applications in various fields, is being frequently used to fabricate nanofibers. Production of ...

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and trade one defect for another. Tissue engineering holds the promise to address this increasing demand. However, most tissue engineering strategies fail to generate stable and functional tissue substitutes because of poor vascularization. This paper focuses on an in vivo tissue engineering chamber model of intrinsic vascularization where a perfused artery and a vein either as an arteriovenous loop or a flow-through pedicle configuration is directed inside a protected hollow chamber. In this chamber-based system angiogenic sprouting occurs from the arteriovenous vessels and this system attracts ischemic and inflammatory driven endogenous cell migration which gradually fills the chamber space with fibro-vascular tissue. Exogenous cell/matrix implantation at the time of chamber construction enhances cell survival and determines specificity of the engineered tissues which develop. Our studies have shown that this chamber model can successfully generate different tissues such as fat, cardiac muscle, liver and others. However, modifications and refinements are required to ensure target tissue formation is consistent and reproducible. This article describes a standardized protocol for the fabrication of two different vascularized tissue engineering chamber models in vivo.

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

This is a guideline for constructing in vivo vascularized tissue using a microsurgical arteriovenous loop or a flow-through pedicle configuration inside a tissue engineering chamber. The vascularized tissues generated can be employed for organ regeneration and replacement of tissue defects, as well as for drug testing and disease modeling.

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and ...

Microfluidic Buffer Exchange for Interference-free Micro/Nanoparticle Cell Engineering

Engineering cells with active-ingredient-loaded micro/nanoparticles (NPs) is becoming an increasingly popular method to enhance native therapeutic properties, enable bio imaging and control cell phenotype. A critical yet inadequately addressed issue is the significant number of particles that remain unbound after cell labeling which cannot be readily removed by conventional centrifugation. This leads to an increase in bio imaging background noise and can impart transformative effects onto neighboring non-target cells. In this protocol, we present an inertial microfluidics-based buffer exchange strategy termed as Dean Flow Fractionation (DFF) to efficiently separate labeled cells from free NPs in a high throughput manner. The developed spiral microdevice facilitates continuous collection (&#62;90% cell recovery) of purified cells (THP-1 and MSCs) suspended in new buffer solution, while achieving &#62;95% depletion of unbound fluorescent dye or dye-loaded NPs (silica or PLGA). This single-step, size-based cell purification strategy enables high cell processing throughput (106 cells/min) and is highly useful for large-volume cell purification of micro/nanoparticle engineered cells to achieve interference-free clinical application.

This protocol describes the use of an inertial microfluidics-based buffer exchange strategy to purify micro/nanoparticle engineered cells with efficient depletion of unbound particles.

Engineering cells with active-ingredient-loaded micro/nanoparticles (NPs) is becoming an increasingly popular method to enhance native therapeutic ...

Microfluidic Bioprinting for Engineering Vascularized Tissues and Organoids

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic bioprinting to generate a scaffold with multilayer interlacing hydrogel microfibers. To achieve smooth bioprinting, a core-sheath microfluidic printhead containing a composite bioink formulation extruded from the core flow and the crosslinking solution carried by the sheath flow, was designed and fitted onto the bioprinter. By blending gelatin methacryloyl (GelMA) with alginate, a polysaccharide that undergoes instantaneous ionic crosslinking in the presence of select divalent ions, followed by a secondary photocrosslinking of the GelMA component to achieve permanent stabilization, a microfibrous scaffold could be obtained using this bioprinting strategy. Importantly, the endothelial cells encapsulated inside the bioprinted microfibers can form the lumen-like structures resembling the vasculature over the course of culture for 16 days. The endothelialized microfibrous scaffold may be further used as a vascular bed to construct a vascularized tissue through subsequent seeding of the secondary cell type into the interstitial space of the microfibers. Microfluidic bioprinting provides a generalized strategy in convenient engineering of vascularized tissues at high fidelity.

We provide a generalized protocol based on a microfluidic bioprinting strategy for engineering a microfibrous vascular bed, where a secondary cell type could be further seeded into the interstitial space of this microfibrous structure to generate vascularized tissues and organoids.

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic ...

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.

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 ...

Two Methods for Decellularization of Plant Tissues for Tissue Engineering Applications

The autologous, synthetic, and animal-derived grafts currently used as scaffolds for tissue replacement have limitations due to low availability, poor biocompatibility, and cost. Plant tissues have favorable characteristics that make them uniquely suited for use as scaffolds, such as high surface area, excellent water transport and retention, interconnected porosity, preexisting vascular networks, and a wide range of mechanical properties. Two successful methods of plant decellularization for tissue engineering applications are described here. The first method is based on detergent baths to remove cellular matter, which is similar to previously established methods used to clear mammalian tissues. The second is a detergent-free method adapted from a protocol that isolates leaf vasculature and involves the use of a heated bleach and salt bath to clear the leaves and stems. Both methods yield scaffolds with comparable mechanical properties and low cellular metabolic impact, thus allowing the user to select the protocol which better suits their intended application.

Here we present, and contrast two protocols used to decellularize plant tissues: a detergent-based approach and a detergent-free approach. Both methods leave behind the extracellular matrix of the plant tissues used, which can then be utilized as scaffolds for tissue engineering applications.

The autologous, synthetic, and animal-derived grafts currently used as scaffolds for tissue replacement have limitations due to low availability, poor ...