This paper was supported by the 2018 Spanish Society of Thoracic Surgery Grant to National Multicentric Study [Number 180101 awarded to Ne&#769;stor J.Marti&#769;nez-Herna&#769;ndez] and PI16-01315 [awarded to Manuel Mata-Roig] by the Instituto de Salud Carlos III. CIBERER is funded by the VI National R&#38;D&#38;I Plan 2018-2011, Iniciativa Ingenio 2010, Consolider Program, CIBER Actions, and the Instituto de Salud Carlos III, with assistance from the European Regional Development Fund.

The European directive 20170/63/EU for the care and use of laboratory animals was adhered to and the study protocol was approved by the Ethics Committee of the University of Valencia (Law 86/609/EEC and 214/1997 and Code 2018/VSC/PEA/0122 Type 2 of the Government of Valencia, Spain).
1. Tracheal decellularization
NOTE: The decellularization method has been reported elsewhere20.
<ol>
	<li>Euthanize donor male adult New Zealand white rabbits (Oryctolagus cuniculus) weighing 3.5-4.1 kg with 133 mg/kg pentobarbital sodium, using a 200 mg/mL injection through the marginal ear vein.</li>
	<li>While ensuring aseptic conditions, perform a central longitudinal cervicotomy, dissect the cervical muscles, and approach the trachea. Dissect the organ circumferentially and longitudinally. Finally, transect under the first ring and just above the carina.</li>
	<li>With a scalpel, section the tracheas into 2 cm pieces. With scissors, remove the surrounding connective tissue and inner mucosa layer6.</li>
	<li>Submerge the specimens in 12 mL of phosphate buffered saline (PBS) solution containing 2% sodium dodecyl sulfate (SDS), 5% penicillin-streptomycin, and 5% amphotericin B.</li>
	<li>Subject the tracheas to constant stirring with a magnetic stirrer at 400 rpm for 5 weeks at room temperature. Replace the decellularization solution weekly after a 2 h osmotic shock, by means of immersion of the trachea in distilled water.</li>
	<li>Cryogenize the specimens using a 12 mL mixture of 80% fetal bovine serum (FBS) and 20% dimethyl sulfoxide (DMSO) in a freezing container at -80 &#176;C.</li>
	<li>When the tracheas are going to be used (after 13-15 days), thaw them in a water bath at 37 &#176;C and wash them by submerging them in PBS after thawing is complete.</li>
</ol>
2. Sterilization
<ol>
	<li>Irradiation
	<ol>
		<li>Place batches of four tracheal pieces measuring 2 cm each in a 20 mL of methacrylate in T25 culture flask filled with PBS until a total volume of 30 mL is reached. Take care to prevent bubbles from forming, which could cause energy diffusion in the air-liquid interface.</li>
		<li>Conduct irradiation using a linear accelerator, with photons of a nominal energy of 10 MV flattening filter free beams. Apply a dose rate of 2,400 monitor units per minute at the isocenter, placing the tracheas at a source-surface distance of 100 cm to be irradiated, with a depth of field of 2.5 cm for a radiation field of 10 cm x 10 cm-so as to cover the entire container-corresponding to a dose of 24 Gy/min.</li>
		<li>Escalate the doses with each four piece batch; four pieces will be subjected to 0.5 kGy, four to 1 kGy, four to 2 kG, etc., until sterilization is reached.</li>
	</ol>
	</li>
	<li>Culture
	<ol>
		<li>Introduce the pieces into 30 mL of Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) with inactivated 10% FBS without antibiotics or antifungals.</li>
		<li>Culture them in a standard tissue incubator at 37 &#176;C and 5% CO2&#160;for 2 weeks and inspect them every 24 h. 
		NOTE: The contamination parameters are changes in the pH of the culture medium and, accordingly, changes in the color and turbidity of the medium. The tracheas were harvested from germ-free lagomorphs, which were not ill and, thus, expected to be devoid of anaerobic bacteria in their tracheas.</li>
	</ol>
	</li>
</ol>
3. Histological analysis
NOTE: Stain the pieces using hematoxylin and eosin21, Masson&#39;s trichrome, and orcein22.
<ol>
	<li>DAPI staining
	<ol>
		<li>Determine the tissue viability using DAPI (4&#8242;,6-diamidino-2-phenylindole). This blue-florescent stain binds strongly to adenine- and thymine-rich regions in DNA sequences, and therefore allows for DNA to be viewed via fluorescence microscopy.</li>
		<li>Embed the tissue samples in optimal cutting temperature (OCT) compound.</li>
		<li>Cut the samples using a cryostat.</li>
		<li>Wash the sample to be dyed three times in distilled water to remove the OCT. Place in mounting medium that includes a 30 nM solution of DAPI.</li>
		<li>Visualize fluorescence using fluorescence microscopy.</li>
	</ol>
	</li>
	<li>DNA content analysis
	<ol>
		<li>Cut segments of trachea measuring approximately 3 mm long using a scalpel.</li>
		<li>Incubate for 2 h in proteinase K (Table of Materials).</li>
		<li>Extract the DNA with a DNA extraction kit, following the manufacturer&#39;s instructions.</li>
		<li>By means of spectrophotometry, determine the concentration of DNA by measuring the absorbance at 260/280 using a spectrophotometer.</li>
		<li>Measure the size of the extracted DNA samples using capillary chromatography with a bioanalyzer.</li>
	</ol>
	</li>
</ol>
4. Biomechanical study
NOTE: Tracheal resistance to longitudinal and transverse forces is measured through axial tensile and radial compression tests23.
<ol>
	<li>Tracheal measurement
	<ol>
		<li>Measure the tracheal length, wall thickness, and external diameter using a Vernier caliper.</li>
		<li>Calculate the average values from three random measurements of each of the variables.</li>
		<li>In the radial compression tests, calculate the anteroposterior diameter by detecting the point at which the plate comes into contact with the specimen.</li>
		<li>Perform all tests at room temperature.</li>
	</ol>
	</li>
	<li>Tensile tests
	<ol>
		<li>Conduct tensile tests on a traction desktop universal testing machine (UTM) displacement control, equipped with a 100 N load (0.1 N force resolution, 0.001 mm of position, and 0.1 s). The testing machine is equipped with force and position sensors, and is connected to a computer with software specifically designed by the manufacturer23.</li>
		<li>Record data every 0.4 s and export to a spreadsheet.</li>
		<li>Construct tensile jaws adapted to the mean caliber of the rabbit tracheas from pure monolayer, non-toxic crystal polyvinyl chloride (PVC) hollow tubes with an external diameter of 1 cm and a wall thickness of 1.5 mm.</li>
		<li>Section the conducts into 3 cm long segments.</li>
		<li>Drill 12 preformed holes for the termino-terminal suture, 2 mm from the edge of the jaws and separated by a distance of 2.5 mm, to prevent bias due to the sutures.</li>
		<li>Attach the PVC glass tubes to the rabbit trachea by termino-terminal anastomosis with a continuous suture through alternate preformed holes (every 5 mm), 2 mm away from the edge of the trachea and with a 6-0 nylon monofilament suture.</li>
		<li>Stretch all the pieces at a displacement rate of 5.0 mm/min.</li>
		<li>Record the variables maximum stress (&#963;max, in N/mm2) and strain (&#949;max, with no units), together with the energy stored per unit of trachea volume (W/Vol, in mJ/mm), and Young&#39;s modulus (E, in MPa).</li>
	</ol>
	</li>
	<li>Radial compression tests
	<ol>
		<li>Perform radial compression tests on a compression desktop UTM, equipped with a 15 N load cell (force resolution 0.001 N, position 0.001 mm, and time 0.1 s) to obtain force data (N), position (mm), and time (s). Record data and export to spreadsheet at 0.5 s intervals.</li>
		<li>Place the tracheas with the membranous area resting on the lower plate. The plate gradually rises up toward the top plate at a constant speed of 5 mm/min.</li>
		<li>Calculate every unit per unit of length of the sample (f in N/mm), stiffness (R in Mpa&#183;mm), and the energy per unit of surface area (W / S in mJ/mm2) needed to completely occlude the trachea.</li>
	</ol>
	</li>
</ol>
5. Surgical technique
NOTE: The surgical technique has been widely reported elsewhere20.
<ol>
	<li>Place a sterile intraluminal PVC stent, size 14 Fr (which allows it to slide freely without compressing the walls), with a 3-4 mm margin at each end.</li>
	<li>Fix the stent with a single 6-0 nylon monofilament stitch through the intercartilaginous space of the first cartilage.</li>
	<li>Proceed to anesthetize the rabbits.
	<ol>
		<li>Pre-medicate the subjects (3.65-4.05 kg male New Zealand white rabbits) with intramuscular analgesics (35 mg/kg ketamine) with a sedative, muscle relaxant, and analgesic (2.5 mg/kg xylazine).</li>
		<li>Shave the incision zone out of the operating zone and clean the surgical area to remove the hair.</li>
		<li>Administer analgesics plus antibiotic prophylaxis: 0.05 mg/kg intramuscular buprenorphine and 10 mg/kg enrofloxacin.</li>
		<li>Put a venous catheter in the marginal ear vein of each rabbit.</li>
		<li>Induct anesthesia with an intravenous 10 mg/kg bolus of propofol.</li>
		<li>Monitor the animal&#39;s vital signs using a three-lead electrocardiogram, pulse oximetry, and noninvasive pressure measurement. Every 30 min, apply physiological serum to the eyes to prevent dryness while under anesthesia.</li>
		<li>Verify the anesthetic plane using the toe pinch method.</li>
		<li>Maintain anesthesia with inhaled isoflurane at 1.5%-2% of the minimum alveolar concentration without losing spontaneous ventilation and provide thermal support to the rabbit with a heating pad.</li>
	</ol>
	</li>
	<li>Disinfect the incision zone several times in a circular motion with an iodine-based scrub. Under aseptic conditions at all times and with sterile material, make a longitudinal 3 cm central thoracic incision, and harvest bilateral pedicled flaps composed of pectoral fascia and a muscular component.</li>
	<li>Wrap the tracheas with the flap in four rabbits, one on each hemithorax (thus, a total of eight tracheas).</li>
	<li>When the surgery is completed, reverse the anesthesia by interrupting isoflurane administration.</li>
	<li>Postsurgical period
	<ol>
		<li>Keep the animals in the operating room until they have completely recovered from anesthesia. When they have completely recovered, return them to their environment with other rabbits.</li>
		<li>Treat the rabbits with antibiotics (0.5 mL/kg of 2.5% enrofloxacin) and analgesics (5 mg/mL meloxicam; 0.05 mL/kg metacam) every 24 h for 5 days.</li>
		<li>Leave the implants in situ for the desired time.</li>
		<li>Prior to euthanasia, pre-medicate the rabbits with intramuscular analgesics (35 mg/kg ketamine) and a sedative, muscle relaxant, and analgesic (2.5 mg/kg xylazine). Then euthanize the rabbits with 133 mg/kg pentobarbital sodium using a 200 mg/mL injection through the marginal ear vein and harvest the tracheas.</li>
		<li>Perform biomechanical and histological tests on the tracheas.</li>
	</ol>
	</li>
</ol>
6. Statistical analysis
<ol>
	<li>Adjust all the models by the Bayesian method on R software, Version 3.5.3 R Core (R Foundation for Statistical Computing. 2019).</li>
	<li>Analyze the study variables, except f and R, using multiple linear regression models.</li>
	<li>For the f and R variables, apply mixed linear regression models. In these models, in addition to the variables of interest related to the treatment and condition of each trachea, introduce the percentage of occlusion as a monotonic effect and an independent term per trachea as a random factor.</li>
</ol>

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None of the authors have any conflict of interests.

There are several sterilization strategies that exist. Supercritical CO2fully penetrates tissues, acidifying the medium and deconstructing the cellular phospholipid bilayer with simple elimination by means of depressurization of the implant8,14,25. Ultraviolet radiation has also been used, and its effectiveness in rodent trachea has been published, although there are only a few reports in the literature<sup class="xre...

Sterilization of an implant is a basic requisite for its viability; in fact, prostheses that have proven to be successful are those implanted in sterile areas (blood vessels, heart, bone, etc.)1. The trachea has two surfaces: a surface in contact with the external environment, which is therefore not sterile, and a surface toward the mediastinum, which is sterile. Therefore, from the moment the trachea is extracted, it is not a sterile organ. Despite the subsequent decellularization process being carried out in maximum sterile conditions, it is not a sterilization step2. The implantation of foreign material in itself entails a risk of infection due to the probacterial microenvironment it produces3and an up to 0.014% risk of disease transmission from the donor to the recipient, even if the material has been sterilized4. To ensure correct vascularization of the trachea, in almost all experimental transplant protocols, it first undergoes heterotopic implant5,6,7&#160;to a sterile area (muscle, fascia, omentum, subcutaneous, etc.); this is because implanting a non-sterile element in this medium would lead to infection of the area3.
There are a range of possible strategies to obtain a sterile implant. Using supercritical CO2has achieved terminal sterilization8,9. Other methods, such as ultraviolet radiation or treatment with substances such as peracetic acid, ethanol, oxygen peroxide, and electrolyzed water, have obtained different success rates in sterilization, almost always depending on their dosages, but they have been shown to affect the biomechanical characteristics of implants. Indeed, some substances, such as ethylene oxide, can substantially change the structure of the implanted matrix and can even cause undesirable immunogenic effects. For this reason, many of these strategies cannot be applied to biological models2,10,11,12,13.
The most widely studied and accepted sterilization strategy is that established by the ISO 11737-1:2006 standard for the sterilization of medical devices implanted in humans, with a gamma radiation dose of 25 kGy. However, this regulation focuses only on the sterilization of inert, non-biological elements14,15. Additionally, radiotherapy doses in the radical treatment of carcinoma are three orders of magnitude lower than those used to sterilize medical devices1. With this in mind, we can conclude that said dose would not only kill the microbiota but would also destroy and radically alter the biological structure of the implant. There is also the possibility that it would generate residual lipids upon degradation, which can potentially be cytotoxic and accelerate the enzymatic degradation of the scaffold13,14,15,16,17, even when using doses as low as 1.9 kGy and with damage directly proportional to the radiation dose received17.
Thus, the objective of this paper is to try to identify the radiation dose that allows for a sterile implant to be obtained with minimal harmful effects caused by irradiation2,18,19. The strategy we followed involved the irradiation of decellularized and irradiated tracheas at different escalated doses within a range of kilograys (0.5, 1, 2, 3 kGy, etc.), until achieving a negative culture. Additional tests were carried out for those doses that achieved negative cultures, in order to confirm sterilization. After determining the minimum dose to obtain sterilization, the structural and biomechanical impact of the irradiation on the trachea were checked. All the metrics were compared with the control native rabbit tracheas. The sterilization of the construct was then tested in vivo by implanting the tracheas into New Zealand white rabbits.

<table><tbody><tr><td>6-0 nylon monofilament suture&amp;nbsp;</td><td>Monosoft. Covidien; Mansfield, MA, USA</td><td>SN-5698G</td><td></td></tr><tr><td>Amphotericin B 5%</td><td>Gibco Thermo Fisher Scientific; Waltham, MA USA</td><td>15290018</td><td></td></tr><tr><td>Bioanalyzer</td><td>Agilent, Santa Clara, CA, USA</td><td>G2939BA</td><td></td></tr><tr><td>Buprenorphine</td><td>Buprex. Reckitt Benckiser Healthcare; Hull, Reino Unido</td><td>N02AE01</td><td></td></tr><tr><td>Compression desktop UTM</td><td>Microtest, Madrid, Spain</td><td>EM1/10/FR</td><td></td></tr><tr><td>Cryostate</td><td>Leyca CM3059, Leyca Biosystems, Wetzlar, Alemania</td><td>CM3059</td><td></td></tr><tr><td>Dimethyl sulfoxide (DMSO)</td><td>Sigma-Aldrich; MO, USA</td><td>D2650</td><td></td></tr><tr><td>DMEM&amp;nbsp;</td><td>Thermo Fisher Scientific; Waltham, MA, USA</td><td>11965084</td><td></td></tr><tr><td>DNA extraction kit</td><td>DNeasy extraction kit Quiagen, Hilden, Germany</td><td>4368814</td><td></td></tr><tr><td>Enrofloxacin, 2.5%</td><td>Boehringer Ingelheim, Ingelheim am Rhein, Germany</td><td>0035-0002</td><td></td></tr><tr><td>Fetal bovine serum (FBS)</td><td>GE Healthcare Hyclone; Madrid, Spain</td><td>SH20898.03IR</td><td></td></tr><tr><td>Fluorescence microscope</td><td>Leyca DM2500 (Leica, Wetzlar, Germany)</td><td>DM2500??</td><td></td></tr><tr><td>Freezing Container&amp;nbsp;</td><td>Mr Frosty. Thermo Fisher; Madrid, Spain</td><td>&amp;nbsp;5100-0001</td><td></td></tr><tr><td>Isofluorane</td><td>Isoflo; Proyma Ganadera; Ciudad Real, Spain&amp;nbsp;</td><td>8.43603E+12</td><td></td></tr><tr><td>Ketamin</td><td>Imalgene. Merial; Toulouse, Francia</td><td>BOE127823</td><td></td></tr><tr><td>Linear accelerator&amp;nbsp;</td><td>&quot;True Beam&quot;. Varian, Palo Alto, California, USA</td><td>H191001</td><td></td></tr><tr><td>Magnetic stirrer</td><td>Orbital Shaker PSU-10i. Biosan; Riga, Letonia</td><td>BS-010144-AAN</td><td></td></tr><tr><td>Meloxicam 5 mg/ml</td><td>Boehringer Ingelheim, Ingelheim am Rhein, Germany</td><td>6283-MV</td><td></td></tr><tr><td>Penicillin-streptomycin 5%</td><td>Gibco Thermo Fisher Scientific; Waltham, MA USA</td><td>15140122</td><td></td></tr><tr><td>Pentobarbital sodium</td><td>Dolethal. Vetoquinol; Madrid, Espa&amp;ntilde;a</td><td>3.60587E+12</td><td></td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>Sigma-Aldrich; MO, USA</td><td>P2272</td><td></td></tr><tr><td>Propofol</td><td>Propofol Lipuro. B. Braun Melsungen AG; Melsungen, Alemania</td><td>G 151030</td><td></td></tr><tr><td>Proteinase K</td><td>Gibco Thermo Fisher Scientific; Waltham, Massachussetts, USA</td><td>S3020</td><td></td></tr><tr><td>PVC hollow tubes</td><td>Cristallo Extra; FITT, Sandrigo, Italy</td><td>hhdddyyZ</td><td></td></tr><tr><td>PVC stent&amp;nbsp;</td><td>ArgyleTM Medtronic; Istanbul, Turkey</td><td>019 5305 1</td><td></td></tr><tr><td>R software, Version 3.5.3 R Core</td><td>R Foundation for Statistical Computing</td><td>R 3.5.3</td><td></td></tr><tr><td>Sodium dodecyl sulfate (SDS)</td><td>Sigma-Aldrich; MO, USA</td><td>8,17,034</td><td></td></tr><tr><td>Spectrophotometer</td><td>Nanodrop, Life Technologies; Isogen Life Science. Utrech, Netherlands</td><td>ND-ONEC-W</td><td></td></tr><tr><td>Spreadsheet</td><td>Microsoft Excel for Mac, Version 16.23, Redmond, WA, USA</td><td>2864993241</td><td></td></tr><tr><td>Traction Universal Testing Machine&amp;nbsp;</td><td>Testing Machines, Veenendaal, Netherlands</td><td>84-01</td><td></td></tr><tr><td>UTM Software</td><td>TestWorks 4, MTS Systems Corporation, Eden Prairie, MN, USA&amp;nbsp;</td><td>100-093-627 F</td><td></td></tr><tr><td>VECTASHIELD Mounting Medium&amp;nbsp;</td><td>Vector Labs, Burlingame; CA; USA</td><td>H-1000-10</td><td></td></tr><tr><td>Xylacine</td><td>Xilagesic. Calier; Barcelona, Espa&amp;ntilde;a</td><td>20102-003</td><td></td></tr></tbody></table>

low-dose gamma radiation sterilization for decellularized tracheal grafts

One of the main key aspects in ensuring that a transplant evolves correctly is the sterility of the medium. Decellularized tracheal transplantation involves implanting an organ that was originally in contact with the environment, thus not being sterile from the outset. While the decellularization protocol (through detergent exposition [2% sodium dodecyl sulfate], continuous stirring, and osmotic shocks) is conducted in line with aseptic measures, it does not provide sterilization. Therefore, one of the main challenges is ensuring sterility prior to in vivo implantation. Although there are established gamma radiation sterilization protocols for inorganic materials, there are no such measures for organic materials. Additionally, the protocols in place for inorganic materials cannot be applied to organic materials, as the established radiation dose (25 kGy) would completely destroy the implant. This paper studies the effect of an escalated radiation dose in a decellularized rabbit trachea. We maintained the dose range (kGy) and tested escalated doses until finding the minimal dose at which sterilization is achieved. After determining the dose, we studied effects of it on the organ, both histologically and biomechanically. We determined that while 0.5 kGy did not achieve sterility, doses of both 1 kGy and 2 kGy did, with 1 kGy, therefore, being the minimal dose necessary to achieve sterilization. Microscopic studies showed no relevant changes compared to non-sterilized organs. Axial biomechanical characteristics were not altered at all, and only a slight reduction in the force per unit of length that the organ can radially tolerate was observed. We can therefore conclude that 1 kGy achieves complete sterilization of decellularized rabbit trachea with a minimal, if any, effects on the organ.

Decellularization 
DAPI staining shows the absence of DNA, and no DNA values higher than 50 ng were detected in any of the tracheae any by electrophoresis, with all fragments being smaller than 200 bp20.
Microbial culture 
Two of the eight pieces subjected to 0.5 kGy showed color change in less than 1 week. None of the pieces irradiated at 1 kGy and 2 kGy showed any color change (Figure 1).
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Low-Dose Gamma Radiation Sterilization for Decellularized Tracheal Grafts

Obtaining sterilization is essential for tracheal tissue transplant. Herein, we present a sterilization protocol using low-dose gamma irradiation that is fully tolerated by organs.

One of the main key aspects in ensuring that a transplant evolves correctly is the sterility of the medium. Decellularized tracheal transplantation ...

low-dose-gamma-radiation-sterilization-for-decellularized-tracheal

Research

JoVE Journal

Bioengineering

819 Views.  Hospital Universitari de la Ribera. Dit protocol biedt een volledig sterilisatieproces voor elk niet-steriel orgaan zonder de kenmerken ervan te beïnvloeden. Het belangrijkste doel van deze techniek omvat de mogelijkheid om een orgaan te steriliseren zonder de eigenschappen ervan te beïnvloeden. Hoewel deze techniek zich heeft gericht op luchtpijpen, heeft het aangetoond dat lagere gammastralingsdoses biologische materialen effectief kunnen steriliseren.Een individu kan het probleem onder ogen zien door de laagste effe...

Video: Low-Dose Gamma Radiation Sterilization for Decellularized Tracheal Grafts

Tracheotomy: A Method for Transplantation of Stem Cells to the Lung

Lung disease is a leading cause of death and likely to become an epidemic given increases in pollution and smoking worldwide. Advances in stem cell therapy may alleviate many of the symptoms associated with lung disease and induce alveolar repair in adults. Concurrent with the ongoing search for stem cells applicable for human treatment, precise delivery and homing (to the site of disease) must be reassured for successful therapy. Here, I report that stem cells can safely be instilled via the trachea opening a non-stop route to the lung. This method involves a skin incision, caudal insertion of a cannula into and along the tracheal lumen, and injection of a stem cell vehicle mixture into airways of the lung. A broad range of media solutions and stabilizers can be instilled via tracheotomy, resulting in the ability to deliver a wider range of cell types. With alveolar epithelium confining these cells to the lumen, lung expansion and negative pressure during inhalation may also assist in stem cell integration. Tracheal delivery of stem cells, with a quick uptake and the ability to handle a large range of treatments, could accelerate the development of cell-based therapies, opening new avenues for treatment of lung disease.

Significant breakthroughs in stem cell identification are continuously being made. To translate these discoveries, however, novel methods for cellular delivery must be devised. Here I report that the airways provide a safe route for stem cell transplantation to the lungs.

Lung disease is a leading cause of death and likely to become an epidemic given increases in pollution and smoking worldwide. Advances in stem cell ...

Biology

A Decellularization Methodology for the Production of a Natural Acellular Intestinal Matrix

Successful tissue engineering involves the combination of scaffolds with appropriate cells in vitro or in vivo. Scaffolds may be synthetic, naturally-derived or derived from tissues/organs. The latter are obtained using a technique called decellularization. Decellularization may involve a combination of physical, chemical, and enzymatic methods. The goal of this technique is to remove all cellular traces whilst maintaining the macro- and micro-architecture of the original tissue.
Intestinal tissue engineering has thus far used relatively simple scaffolds that do not replicate the complex architecture of the native organ. The focus of this paper is to describe an efficient decellularization technique for rat small intestine. The isolation of the small intestine so as to ensure the maintenance of a vascular connection is described. The combination of chemical and enzymatic solutions to remove the cells whilst preserving the villus-crypt axis in the luminal aspect of the scaffold is also set out. Finally, assessment of produced scaffolds for appropriate characteristics is discussed.

The article describes a methodology for the production of an acellular matrix from rat intestine. The derivation of intestinal scaffolds is important for future applications in tissue engineering, stem cell biology and drug testing.

Successful tissue engineering involves the  combination of scaffolds with appropriate cells in vitro or in vivo.  Scaffolds may be synthetic, ...

Nonhuman Primate Lung Decellularization and Recellularization Using a Specialized Large-organ Bioreactor

There are an insufficient number of lungs available to meet current and future organ transplantation needs. Bioartificial tissue regeneration is an attractive alternative to classic organ transplantation. This technology utilizes an organ&#39;s natural biological extracellular matrix (ECM) as a scaffold onto which autologous or stem/progenitor cells may be seeded and cultured in such a way that facilitates regeneration of the original tissue. The natural ECM is isolated by a process called decellularization. Decellularization is accomplished by treating tissues with a series of detergents, salts, and enzymes to achieve effective removal of cellular material while leaving the ECM intact. Studies conducted utilizing decellularization and subsequent recellularization of rodent lungs demonstrated marginal success in generating pulmonary-like tissue which is capable of gas exchange in vivo. While offering essential proof-of-concept, rodent models are not directly translatable to human use. Nonhuman primates (NHP) offer a more suitable model in which to investigate the use of bioartificial organ production for eventual clinical use.
The protocols for achieving complete decellularization of lungs acquired from the NHP rhesus macaque are presented. The resulting acellular lungs can be seeded with a variety of cells including mesenchymal stem cells and endothelial cells. The manuscript also describes the development of a bioreactor system in which cell-seeded macaque lungs can be cultured under conditions of mechanical stretch and strain provided by negative pressure ventilation as well as pulsatile perfusion through the vasculature; these forces are known to direct differentiation along pulmonary and endothelial lineages, respectively. Representative results of decellularization and cell seeding are provided.

Whole-organ decellularization produces natural biological scaffolds that may be used for regenerative medicine. The description of a nonhuman primate model of lung regeneration in which whole lungs are decellularized and then seeded with adult stem cells and endothelial cells in a bioreactor that facilitates vascular circulation and liquid media ventilation is presented.

There are an insufficient number of lungs available to meet current and future organ transplantation needs. Bioartificial tissue regeneration is an ...

Decellularization of the Murine Cardiopulmonary Complex

We present here a decellularization protocol for mouse heart and lungs. It produces structural ECM scaffolds that can be used to analyze ECM topology and composition. It is based on a microsurgical procedure designed to catheterize the trachea and aorta of a euthanized mouse to perfuse the heart and lungs with decellularizing agents. The decellularized cardiopulmonary complex can subsequently be immunostained to reveal the location of structural ECM proteins. The whole procedure can be completed in 4 days.
The ECM scaffolds resulting from this protocol are free of dimensional distortions. The absence of cells enables structural examination of ECM structures down to submicron resolution in 3D. This protocol can be applied to healthy and diseased tissue from mice as young as 4-weeks old, including mouse models of fibrosis and cancer, opening the way to determine ECM remodeling associated with cardiopulmonary disease.

This protocol aims to decellularize the heart and lungs of mice. The resulting extracellular matrix (ECM) scaffolds can be immunostained and imaged to map the location and topology of their components.

We present here a decellularization protocol for mouse heart and lungs. It produces structural ECM scaffolds that can be used to analyze ECM topology and ...

Cancer Research

Generation of ESC-derived Mouse Airway Epithelial Cells Using Decellularized Lung Scaffolds

Lung lineage differentiation requires integration of complex environmental cues that include growth factor signaling, cell-cell interactions and cell-matrix interactions. Due to this complexity, recapitulation of lung development in vitro to promote differentiation of stem cells to lung epithelial cells has been challenging. In this protocol, decellularized lung scaffolds are used to mimic the 3-dimensional environment of the lung and generate stem cell-derived airway epithelial cells. Mouse embryonic stem cell are first differentiated to the endoderm lineage using an embryoid body (EB) culture method with activin A. Endoderm cells are then seeded onto decellularized scaffolds and cultured at air-liquid interface for up to 21 days. This technique promotes differentiation of seeded cells to functional airway epithelial cells (ciliated cells, club cells, and basal cells) without additional growth factor supplementation. This culture setup is defined, serum-free, inexpensive, and reproducible. Although there is limited contamination from non-lung endoderm lineages in culture, this protocol only generates airway epithelial populations and does not give rise to alveolar epithelial cells. Airway epithelia generated with this protocol can be used to study cell-matrix interactions during lung organogenesis and for disease modeling or drug-discovery platforms of airway-related pathologies such as cystic fibrosis.

This protocol efficiently directs mouse embryonic stem cell-derived definitive endoderm to mature airway epithelial cells. This differentiation technique uses 3-dimensional decellularized lung scaffolds to direct lung lineage specification, in a defined, serum-free culture setting.

Lung lineage differentiation requires integration of complex environmental cues that include growth factor signaling, cell-cell interactions and ...

Seeding and Implantation of a Biosynthetic Tissue-engineered Tracheal Graft in a Mouse Model

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional tissue. Tissue engineering holds great promise as a potential solution with its ability to integrate cells and signaling molecules into a 3-dimensional scaffold. Recent work with tissue engineered tracheal grafts (TETGs) has seen some success but their translation has been limited by graft stenosis, graft collapse, and delayed epithelialization. In order to investigate the mechanisms driving these issues, we have developed a mouse model for tissue engineered tracheal graft implantation. TETGs were constructed using electrospun polymers polyethylene terephthalate (PET) and polyurethane (PU) in a mixture of PET and PU (20:80 percent weight). Scaffolds were then seeded using bone marrow mononuclear cells isolated from 6-8 week-old C57BL/6 mice by gradient centrifugation. Ten million cells per graft were seeded onto the lumen of the scaffold and allowed to incubate overnight before implantation between the third and seventh tracheal rings. These grafts were able to recapitulate the findings of stenosis and delayed epithelialization as demonstrated by histological analysis and lack of Keratin 5 and Keratin 14 basal epithelial cells on immunofluorescence. This model will serve as a tool for investigating cellular and molecular mechanisms involved in host remodeling.

Graft stenosis poses a critical obstacle in tissue engineered airway replacement. To investigate cellular mechanisms underlying stenosis, we utilize a murine model of tissue engineered tracheal replacement with seeded bone marrow mononuclear cells (BM-MNC). Here, we detail our protocol, including scaffold manufacturing, BM-MNC isolation, graft seeding, and implantation.

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional ...

Design of a Biocompatible Drug-Eluting Tracheal Stent in Mice with Laryngotracheal Stenosis

Laryngotracheal stenosis (LTS) is a pathologic narrowing of the subglottis and trachea leading to extrathoracic obstruction and significant shortness of breath. LTS results from mucosal injury from a foreign body in the trachea, leading to tissue damage and a local inflammatory response that goes awry, leading to the deposition of pathologic scar tissue. Treatment for LTS is surgical due to the lack of effective medical therapies. The purpose of this method is to construct a biocompatible stent that can be miniaturized to place into mice with LTS. We demonstrated that a PLLA-PCL (70% poly-L-lactide and 30% polycaprolactone) construct had optimal biomechanical strength, was biocompatible, practicable for an in vivo placement stent, and capable of eluting drug. This method provides a drug delivery system for testing various immunomodulatory agents to locally inhibit inflammation and reduce airway fibrosis. Manufacturing the stents takes 28&#8722;30 h and can be reproduced easily, allowing for experiments with large cohorts. Here we incorporated the drug rapamycin within the stent to test its effectiveness in reducing fibrosis and collagen deposition. Results revealed that PLLA-PCL tents showed reliable rapamycin release, were mechanically stable in physiological conditions, and were biocompatible, inducing little inflammatory response in the trachea. Further, the rapamycin-eluting PLLA-PCL stents reduced scar formation in the trachea in vivo.

Laryngotracheal stenosis results from pathologic scar deposition that critically narrows the tracheal airway and lacks effective medical therapies. Using a PLLA-PCL (70% poly-L-lactide and 30% polycaprolactone) stent as a local drug delivery system, potential therapies aimed at decreasing scar proliferation in the trachea can be studied.

Laryngotracheal stenosis (LTS) is a pathologic narrowing of the subglottis and trachea leading to extrathoracic obstruction and significant shortness of ...

Imaging-Guided Bioreactor for Generating Bioengineered Airway Tissue

Repeated injury to airway tissue can impair lung function and cause chronic lung disease, such as chronic obstructive pulmonary disease. Advances in regenerative medicine and bioreactor technologies offer opportunities to produce lab-grown functional tissue and organ constructs that can be used to screen drugs, model disease, and engineer tissue replacements. Here, a miniaturized bioreactor coupled with an imaging modality that allows in situ visualization of the inner lumen of explanted rat trachea during in vitro tissue manipulation and&#160;culture is described. Using this bioreactor, the protocol demonstrates imaging-guided selective removal of endogenous cellular components while preserving the intrinsic biochemical features and ultrastructure of the airway tissue matrix. Furthermore, the delivery, uniform distribution, and subsequent prolonged culture of exogenous cells on the decellularized airway lumen with optical monitoring in situ are shown. The results highlight that the imaging-guided bioreactor can potentially be used to facilitate the generation of functional in vitro airway tissues.

The protocol describes an imaging-enabled bioreactor that allows the selective removal of the endogenous epithelium from the rat trachea and homogenous distribution of exogenous cells on the lumen surface, followed by long-term in vitro culture of the cell-tissue construct.

Repeated injury to airway tissue can impair lung function and cause chronic lung disease, such as chronic obstructive pulmonary disease. Advances in ...

Transplantation of Bioengineered Lung Using Decellularized Mouse Lungs and Primary Human Endothelial Cells

Lung transplantation is a critical treatment for patients with end-stage lung diseases like idiopathic pulmonary fibrosis, but challenges such as donor shortages and posttransplant complications persist. Bioengineered lungs, integrating patient-specific cells into decellularized animal scaffolds, present a promising alternative. Despite progress in using bioengineered lungs in animal models, functionality and structure remain immature. This protocol addresses a critical barrier in organ bioengineering: the need for a cost-effective experimental platform. By using mouse models instead of larger animals like rats or swine, researchers can significantly reduce the resources required for each experiment, accelerating research progress. 
The protocol outlines a detailed procedure for lung bioengineering using mouse heart-lung blocks and human primary cells, focusing on isolation strategy for the mouse heart-lung block, decellularization, bioreactor setup, perfusion-based organ culture, and orthotopic transplantation of bioengineered lungs. This mouse-scale platform not only reduces experimental costs but also provides a viable framework for optimizing cell types and numbers for recellularization, testing different cell types using histological and molecular methods, and ensuring blood flow post-transplantation. The method holds potential for broad applications, including studying cell interactions in three-dimensional culture conditions, cell-matrix interactions, and ex vivo cancer modeling, thereby advancing the field of organ bioengineering.

This paper describes how to create bioengineered mouse lungs using decellularization and recellularization methods. It also details subsequent orthotopic lung transplantation.

Lung transplantation is a critical treatment for patients with end-stage lung diseases like idiopathic pulmonary fibrosis, but challenges such as donor ...

Exploring Obliterative Airway Disease in Lung Transplants

Murine Intrapulmonary Tracheal Transplantation: A Model for Investigating Obliterative Airway Disease After Lung Transplantation

Murine intrapulmonary tracheal transplantation (IPTT) is used as a model of obliterative airway disease (OAD) following lung transplantation. Initially reported by our team, this model has gained use in the study of OAD due to its high technical reproducibility and suitability for investigating immunological behaviors and therapeutic interventions.
In the IPTT model, a rodent tracheal graft is directly inserted into the recipient's lung through the pleura. This model is distinct from the heterotopic tracheal transplantation (HTT) model, wherein grafts are transplanted into subcutaneous or omental sites, and from the orthotopic tracheal transplantation (OTT) model in which the donor trachea replaces the recipient's trachea. 
Successful implementation of the IPTT model requires advanced anesthetic and surgical skills. Anesthetic skills include endotracheal intubation of the recipient, setting appropriate ventilatory parameters, and appropriately timed extubation after recovery from anesthesia. Surgical skills are essential for precise graft placement within the lung and for ensuring effective sealing of the visceral pleura to prevent air leakage and bleeding. In general, the learning process takes approximately 2 months.
In contrast to the HTT and OTT models, in the IPTT model, the allograft airway develops airway obliteration in the relevant lung microenvironment. This allows investigators to study lung-specific immunological and angiogenic processes involved in airway obliteration after lung transplantation. Furthermore, this model is also unique in that it exhibits tertiary lymphoid organs (TLOs), which are also seen in human lung allografts. TLOs are comprised of T and B cell populations and characterized by the presence of high endothelial venules that direct immune cell recruitment; therefore, they are likely to play a crucial role in graft acceptance and rejection. We conclude that the IPTT model is a useful tool for studying intrapulmonary immune and profibrotic pathways involved in the development of airway obliteration in the lung transplant allograft.

Author Spotlight: Investigating the Key Factors of Obliterative Bronchiolitis After Lung Transplantation

The murine intrapulmonary tracheal transplantation (IPTT) model is valuable for studying obliterative airway disease (OAD) after lung transplantation. It offers insights into lung-specific immunological and angiogenic behavior in airway obliteration after allotransplantation with high reproducibility. Here, we describe the IPTT procedure and its expected results.

Murine intrapulmonary tracheal transplantation (IPTT) is used as a model of obliterative airway disease (OAD) following lung transplantation. Initially ...

Low-Dose Gamma Radiation Sterilization for Decellularized Tracheal Grafts

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