Waiting
Login processing...

Trial ends in Request Full Access Tell Your Colleague About Jove

Bioengineering

Decellularization for the Preparation of Highly Preserved Human Acellular Skin Matrix for Regenerative Medicine

Published: September 8, 2021 doi: 10.3791/62935
Veronica Romano*1, Immacolata Belviso*1, Domenico Cozzolino1, Anna Maria Sacco1, Fabrizio Schonauer1, Daria Nurzynska2, Franca Di Meglio1, Clotilde Castaldo1
* These authors contributed equally

Abstract

Extracellular matrix (ECM) provides biophysical and biochemical stimuli to support self-renewal, proliferation, survival, and differentiation of surrounding cells due to its content of diverse bioactive molecules. Due to these characteristics, the ECM has been recently considered a promising candidate for the creation of biological scaffolds to boost tissue regeneration. Emerging studies have demonstrated that decellularized human tissues could resemble the native ECM in their structural and biochemical profiles, preserving the three-dimensional (3D) architecture and the content of fundamental biological molecules. Hence, decellularized ECM can be employed to promote tissue remodeling, repair, and functional reconstruction of many organs. Selecting the appropriate decellularization procedure is crucial to obtain acellular tissues that retain the characteristics of the ideal microenvironment for cells.

The protocol described here provides a detailed step-by-step description of the decellularization method to obtain a reproducible and effective cell-free biological ECM. Skin fragments from patients undergoing plastic surgery were scaled down and decellularized using a combination of sodium dodecylsulfate (SDS), Triton X-100, and antibiotics. To promote the regular and homogeneous transport of the solution through the samples, they were enclosed in embedding cassettes to ensure protection from mechanical insults. After the decellularization procedure, the snow-white color of skin fragments indicated complete and successful decellularization. Additionally, decellularized samples showed an intact and well-preserved architecture. The results suggest that the proposed decellularization method was effective, fast, and reproducible and protected samples from architectural damages.

Introduction

The ECM serves as a scaffold for cells, supporting them through an intricate architecture maintained by different components, and it is one of the major factors responsible for the mechanical properties of the heart and cardiac tissue function1,2. Increasing evidence suggests that ECM plays an active role in tissue remodeling, making the conventional assumption that the ECM is a passive component obsolete3,4,5,6. The role of the ECM is to provide biophysical and biochemical cues to resident cells. It is well-established that these signals can influence many fundamental cell behaviors, impacting their contractile function, proliferation, migration, and differentiation potential7,8,9. Thus, ECM is increasingly being employed in tissue engineering and regenerative medicine as a therapeutic support tool9,10,11,12,13.

The ECM consists of several proteins such as collagen, elastin, fibronectin, proteoglycan, and laminin, along with ECM-bound growth factors, all involved in regeneration mechanisms, such as cell recruitment, migration, and differentiation, as well as cell alignment and proliferation14. The mechanical properties of the tissues also have great relevance in the physiopathology of the organs. Indeed, changes in mechanical properties are often associated with the onset and the evolution of several diseases. The reason resides in the fact that when the ECM is modified, signals coming from the environment induce changes in gene and protein expression, leading to functional impairment15,16.

Regenerative therapies for organ repair are currently focused on replicating the sophisticated microenvironment of the native tissue to heal the organ where the body fails. Despite the rapid pace of many tissue engineering approaches, the tissues still cannot be reproduced accurately in their entirety and complexity by artificial procedures. Synthetic materials have been largely employed so far, as they can be appropriately tuned to simulate the mechanical and biochemical properties of the cellular microenvironment. Nevertheless, they have limits, such as the inability to mimic the numerous interactions within the native tissue, the cost of technologies to produce them, and the fact that they are less natural and biocompatible than native tissue17,18,19. Additionally, their composition, primarily in terms of proteins and soluble factors, greatly differs from the natural one, which is extremely difficult to replicate20.

The cutting-edge approach in regenerative medicine to reduce the gap between the patients in need and organ transplants is to produce scaffolds made of decellularized extracellular matrix (d-ECM) and repopulate them with the appropriate cell types to regenerate the damaged organs. Decellularization is the process in which the ECM is isolated from its native cells and genetic material to produce a natural and biomimetic scaffold, able to avoid the immune response and rejection once implanted in the patients21,22,23. The ECM thus obtained can then be repopulated to produce functional tissue. The major issue when developing a d-ECM is the method. For any decellularization technique, the primary goal remains the preservation of the native ECM composition, stiffness, and 3D structure, and all strategies have both benefits and drawbacks. Because the elimination of cellular content and DNA from the tissue requires the use of chemical or physical agents, or the combination of both, each decellularization procedure causes, to different degrees, the disruption of the ECM. Hence, it is crucial to minimize the damage to the ECM24,25,26.

Native ECM utilization as a platform for the reconstitution of native ECM in vitro is highly desirable. For this purpose, several decellularization protocols have been applied to a wide range of tissues27,28,29,30. In fact, since the early stages of decellularization research and ECM development, several tissues, such as arteries, aortic valves, and peripheral nerves from both animals and humans, have been decellularized, and some d-ECMs are still commercially available and used for tissue replacement or wound healing31,32,33. Recently, human skin has also emerged as a suitable candidate to produce decellularized scaffolds for cardiac repair owing to its composition and mechanical properties-able to boost the regenerative potential of cardiac progenitor cells (CPCs) and adapt to cardiac contractility34. This paper describes a simple and fast protocol to produce decellularized scaffolds from adult human skin, allowing the development of a d-ECM with well-preserved architecture.

Subscription Required. Please recommend JoVE to your librarian.

Protocol

The specimens from human tissue were collected according to the principles of the Declaration of Helsinki and observing University Hospital "Federico II" guidelines. All patients involved in this study provided written consent forms.

1. Preparation of solutions

  1. Preparation of 1200 mL of 1% decellularizing solution
    1. Prepare 600 mL of 2% Triton X-100 solution by measuring 588 mL of double-distilled water in a graduated cylinder and transferring it to a 1 L beaker. Using a serological pipette, add 12 mL of Triton X-100 and a stir bar to the beaker. Place the beaker on a magnetic stirrer and mix the solution until the Triton X-100 is completely dissolved.
    2. Stop the agitation. Remove the stir bar using a stir bar retriever. Store at room temperature.
    3. Prepare 600 mL of 2% SDS solution by measuring 550 mL of double-distilled water in a graduated cylinder and transferring it to a 1 L beaker.
    4. Weigh 12 g of SDS powder in a plastic weighing boat and transfer it to the beaker containing the double-distilled water in step 1.1.3.
      NOTE: Step 1.1.4 must be performed under a chemical hood, and the user should wear personal protective equipment.
    5. Add a stir bar, place the beaker on a magnetic stirrer, and mix the solution until the SDS is completely dissolved.
    6. Stop the agitation process. Remove the stir bar using a stir bar retriever.
    7. Transfer the solution to a graduated cylinder and adjust the volume to 600 mL by adding double-distilled water.
    8. Pour 2% Triton X-100 solution prepared in step 1.1.1 into a 2 L cylinder.
    9. Pour 2% SDS solution prepared in steps 1.1.3-1.1.7 into the same 2 L cylinder to obtain a total volume of 1200 mL.
    10. Cover with parafilm and mix gently by inversion to obtain a homogeneous solution.
    11. Transfer the mixed solution to a 2 L bottle using a funnel to avoid the formation of foam. Store the solution at +4 °C.
  2. Preparation of 1x phosphate-buffered-saline (PBS)
    1. Prepare 500 mL of 1x phosphate-buffered-saline (PBS) by dissolving 0.1 g of potassium phosphate monobasic, 0.1 g of potassium chloride, 4.0 g of sodium chloride, and 0.575 g of sodium phosphate dibasic in sterile double-distilled water. Check the pH value (7.4).
    2. Store at +4 °C until use.
  3. Preparation of antibiotic solution
    1. Weigh 625 µg of amphotericin B in a plastic weighing boat.
    2. Pipette 8 mL of a penicillin and streptomycin (pen/strep) mixture into a 15 mL tube.
    3. Add the weighed amphotericin B to the pen/strep mixture, and adjust the volume to 10 mL by adding penicillin and streptomycin (pen/strep) mixture. Cap the tube and dissolve the amphotericin B by shaking vigorously.

2. Day 1 - start the decellularization procedure.

  1. Pour 400 mL of the decellularizing solution into a 500 mL beaker.
  2. Add 2.0 mL of the antibiotic solution.

3. Preparation of skin samples

  1. Wash the human skin sample from the abdomen of patients undergoing abdominoplasty in a plastic tray by shacking and turning it upside down in a 0.9% NaCl isotonic physiological solution to remove excess blood and other biological fluids.
  2. Remove hair using fine forceps and fat using large surgical scissors.
  3. Place the skin sample on the dissecting board and dissect it using a scalpel, with the graduations on the dissecting board as a reference to obtain 3 cm x 2.5 cm fragments (length by width), avoiding scars and dirty or burned areas of the tissue.
  4. Place each fragment in an embedding cassette, close it, and make sure all the cassettes are appropriately locked to avoid leakage of the sample during the decellularization procedure.
  5. Place a maximum of four embedding cassettes with samples in a beaker with the decellularizing solution. Add a stir bar. Cover with aluminum foil, writing on the top all the information to identify the sample and the starting time of the procedure.
  6. Place the beaker on the magnetic stirrer and start the agitation at a rotational velocity of 150 rpm. Stop the agitation. Remove the aluminum foil covering the beaker and take the embedding cassettes out with long forceps.
    NOTE: After stopping the agitation, check the clarity of the solution. Modify the duration of the agitation (less or more than 8 hours) based on the degree of the clouding due to the release of cellular debris.
  7. Place each cassette in a 100 mm dish. Replace the decellularizing solution by repeating steps 3.5-3.6 overnight.

4. Day 2 - check the state of the skin samples.

  1. Stop the agitation process. Remove the aluminum foil covering the beaker and take the embedding cassettes out with long forceps.
  2. Place each cassette in a 100 mm dish and open it. Check the color of the sample.
    NOTE: The samples might show a color change from the native beige to snow-white.
  3. Remove residual hair and detach the epidermis from the dermis. Replace the decellularizing solution by repeating steps 3.5-3.6.
  4. Stop the agitation after 8 h. Repeat steps 4.1-4.2 until the sample appears snow-white. Stop the decellularizing procedure.
    NOTE: The time of exposure to the action of detergents can be extended if needed.
  5. Measure 400 mL of 1x PBS in a 500 mL cylinder, pour it into a 500 mL beaker, and add 2 mL of the Pen Strep/Amphotericin B mixture. Add a stir bar.
  6. Place a maximum of four cassettes containing the samples in each beaker with the 1x PBS/antibiotics solution. Cover with aluminum foil and write all the information on the top as in step 3.5.
  7. Place the beaker on a magnetic stirrer and start the agitation at a rotational velocity of 150 rpm overnight, at room temperature.

5. Day 3 - a final wash of the samples

  1. Stop the agitation. Replace the 1x PBS solution with 400 mL of double-distilled water. Add a stir bar. Cover with aluminum foil and write all the information on top as in step 3.5.
  2. Place the beaker on a magnetic stirrer and start the agitation at a rotational velocity of 150 rpm for 30 min.
  3. Stop the agitation process. Remove the aluminum foil covering the beaker and take the embedding cassettes out with long forceps.
  4. Place each embedding cassette containing the decellularized skin sample in a 100 mm dish. Open the cassettes and gently dry the skin samples by dabbing them on laboratory wipes.
  5. Place each decellularized skin sample in a weighing boat previously tagged with the information about the sample. Cover the boat with aluminum foil and write all the information on the top to identify the sample. Store at - 80 °C.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

The aim of the protocol was to obtain a skin d-ECM sample from biological tissue, maintaining a well-organized 3D structure and well-preserved content of biological molecules (Figure 1). This method is primarily based on the constant stirring of the samples in a solution containing the combination of two detergents, Triton X-100 and SDS, thus preserving the biological and structural features typical of the native tissue and reducing the time of exposure during the decellularization process. Upon receipt, the samples were washed and prepared for the procedure, obtaining 3 cm x 2.5 cm clean skin fragments, free from hair or fat tissue (Figure 2 and Figure 3). The use of embedding cassettes allowed the minimization of tissue disruption (Figure 3), resulting in samples with a more intact architecture. The decellularized skin did not show signs of significant mechanical or chemical damage after macroscopic observation. The epidermis was detached during the decellularization procedure (Figure 4A,B), and the samples changed in color from beige, typical of the whole native tissue, to snow-white, indicating complete and successful decellularization (Figure 4C).

Figure 1
Figure 1: Protocol workflow. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Representative images of the skin sample preparation upon receipt. (A) Incoming sample. (B, C) Macroscopic evaluation of the sample placed on dissecting board. (D-F) Removal of the fat tissue using large surgical scissors. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Skin sample dissection and placement in embedding cassettes. (A) Hair removed by fine forceps. (B-E) Skin samples dissected using a scalpel to obtain 3 cm x 2.5 cm fragments to fit into the embedding cassettes. (F, G) Skin fragment placed in embedding cassette. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Checking the state of the sample during decellularization. (A, B) Detachment of the epidermis from the dermis. (C) Sample color change indicating the occurrence of decellularization. Scale bars = 3 cm. Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.

Discussion

Although the protocol described above has been optimized and improved compared to previously published protocols, it presents a few critical steps that need attention and precision. The formation of foam must be avoided during the preparation of the decellularizing solution to prevent incorrect dilution of the detergents. This could be addressed by gently pouring the solutions and making them flow along the inner side of the cylinder. Furthermore, care must be taken when manually removing fat tissue from the samples, as decellularization using detergents, such as Triton X-100 and SDS, does not eliminate fat, and lipidic remnants could negatively affect the effectiveness of the method. Another critical step is the removal of the epidermis from the samples. During the decellularization, the epidermis gradually detaches from the skin surface. In this phase, it is crucial to peel it away with forceps only when it is completely swept off, thus avoiding tearing or shredding it and causing the entrapment of debris within the samples.

This protocol focuses on reducing the impact of decellularization on the samples by two major strategies: the shortening of exposure time to the procedure by using two detergents in combination28,35,36 and the protection of the samples from mechanical insults by enclosing them in embedding cassettes37. Further, the procedure has yielded successful outcomes when applied to samples other than skin, such as human and porcine myocardium and blood vessels. The evaluation of the effectiveness of the protocol described here is primarily done by a macroscopic observation of the samples, which would show a remarkable change in color, from beige to snow-white. Certainly, some other aspects must be assessed to assess the quality of d-ECM, such as the removal of cellular debris and residual genetic material, the preservation of structural proteins and biomolecule content, and the retention of mechanical properties, as described elsewhere34,37,38. The removal of cellular debris and genetic material is crucial to reduce the immunogenicity of the construct. Hence, some criteria for assessing the efficacy of removing these components are well-established in the literature: the d-ECM must have not more than 50 ng of double-stranded DNA (dsDNA)39 per mg of ECM dry weight, and no nuclear material must be microscopically visible.

To meet these criteria, residual DNA content evaluation and staining with hematoxylin and eosin must be performed, as described by Di Meglio et al.38 to support the macroscopic data. Moreover, the evaluation of ECM preservation by detecting structural proteins, such as collagen, fibronectin, and laminin, together with glycosaminoglycans and growth factors, is also appropriate34. Finally, the mechanical properties of the d-ECM should match those of the native tissue40. Although the efficacy and feasibility of this procedure have been demonstrated to obtain desirable decellularized tissue, the method has some limitations. For example, it allows the decellularization of only a few samples at a time (4 for each beaker), leading to considerable wastage of solutions. On the one hand, the use of embedding cassettes helps protect and preserve the samples; on the other hand, it forces the operators to decellularize only samples of small sizes that fit into the cassettes. Another limitation is the possibility of microbial contamination of the d-ECM obtained. Although the procedure involves the addition of antibiotics, it is highly recommended to sterilize the d-ECM by UV prior to use. Finally, the incubations required throughout the entire procedure must be accurately calculated and organized to optimally manage the time.

This protocol was developed to address the drawbacks observed while testing other protocols in the laboratory. Existing protocols allow a varying degree of decellularization without guaranteeing adequate integrity of the tissue, thus involving a considerable loss of essential biological molecules and mechanical properties41. The preservation of the ECM structure and composition after the decellularization are fundamental aspects, as d-ECM may act as the scaffold, that once repopulated, will support cellular regenerative mechanisms within the organ. By optimizing the decellularization process, biological scaffolds derived from human skin may be utilized in regenerative medicine, helping to minimize the gap between donors and patients in need of organ transplants. In addition, advancements in recellularization methods and applications of human induced pluripotent stem cell-derived cells will improve the distribution of the proper cell types42,43 to enable the regeneration process44.

Subscription Required. Please recommend JoVE to your librarian.

Disclosures

The authors have no conflicts of interest to disclose.

Acknowledgments

None

Materials

Name Company Catalog Number Comments
0.9% NaCl isotonic Physiological solution Sigma-Aldrich S8776 0.9% in water
1 L beaker VWR 511-0318 Clean and autoclave before use
10 mL serological pipet Falcon 357551 Sterile,  polystyrene
100 mm plates Falcon 351029 Treated, sterile cell culture dish
15 mL sterile tubes Falcon 352097 Centrifuge sterile tubes, polypropylene
1 L graduated cylinder VWR 612-1524 Clean and autoclave before use
2 L bottle VWR 215-1596 Clean and autoclave before use
25 mL serological pipet Falcon 357525 Sterile,  polystyrene
2 L graduated cylinder VWR 612-3072 Clean and autoclave before use
500 mL beaker VWR 511-0317 Clean and autoclave before use
Amphotericin B Sigma-Aldrich Y0000005 Powder
Dissecting board VWR 100498-398 Made of high-density polyethylene.
Dissecting scalpel VWR 233-5526 Sterile and disposable
Embedding cassettes Diapath 070191 External dimensions: 40x26x7 mm (WxDxH)
Fine forceps VWR 232-1317 Clean and autoclave before use
Funnel VWR 221-1861 Clean and autoclave before use
Hexagonal weighing boats size M Sigma-Aldrich Z708585 Hexagonal, polystyrene, 51 mm Bottom I.D., 64 mm Top I.D.
Hexagonal weighing boats size S Sigma-Aldrich Z708577 Hexagonal, polystyrene, 25 mm Bottom I.D., 38 mm Top I.D.
Large surgical scissors VWR 233-1211 Clean and autoclave before use
Long forceps VWR 232-0096 Clean and autoclave before use
Penicillin and Streptomycin Sigma-Aldrich P4333-100ml Stabilized, with 10.000 units penicillin and 10 mg streptomycin/mL, 0.1 μm filtered. Store at -20°C.  The solution  should be aliquoted into smaller working volumes to avoid repeated freeze/thaw cycles Solution.
Pipette gun Eppendorf 613-2795 Eppendorf Easypet® 3
Plastic tray VWR BELAH162620000 Corrosion-proof polypropylene plastic tray
Potassium Chloride Sigma-Aldrich P9333 Powder
Potassium Phosphate Monobasic Sigma-Aldrich P5665 Powder
Sodium Chloride Sigma-Aldrich S7653 Powder
Sodium Dodecyl Sulfate Sigma-Aldrich 62862 Powder
Sodium Phosphate Dibasic Sigma-Aldrich 94046 Powder
Spatula VWR RSGA038.210 Clean and autoclave before use
Spoon VWR 231-1314 Clean and autoclave before use
Stir bar VWR 442-0362 Clean and autoclave before use
Stir bar retriever VWR 89026-262 Molded in pure, FDA-approved PTFE
Triton X-100 Sigma-Aldrich 9002-93-1 Liquid

DOWNLOAD MATERIALS LIST

References

  1. Daley, W. P., Peters, S. B., Larsen, M. Extracellular matrix dynamics in development and regenerative medicine. Journal of Cell Science. 121, (3), 255-264 (2008).
  2. Lu, P., Takai, K., Weaver, V. M., Werb, Z. Extracellular matrix degradation and remodeling in development and disease. Cold Spring Harbor Perspectives in Biology. 3, (12), 005058 (2011).
  3. Dobaczewski, M., Gonzalez-Quesada, C., Frangogiannis, N. G. The extracellular matrix as a modulator of the inflammatory and reparative response following myocardial infarction. Journal of Molecular and Cellular Cardiology. 48, (3), 504-511 (2010).
  4. Frangogiannis, N. G. Matricellular proteins in cardiac adaptation and disease. Physiological Reviews. 92, (2), 635-688 (2012).
  5. Etoh, T., et al. Myocardial and interstitial matrix metalloproteinase activity after acute myocardial infarction in pigs. American Journal of Physiology-Heart and Circulating Physiology. 281, (3), 987-994 (2001).
  6. Ma, Y., et al. Myofibroblasts and the extracellular matrix network in post-myocardial infarction cardiac remodeling. Pflügers Archiv: European Journal of Physiology. 466, (6), 1113-1127 (2014).
  7. Fong, A. H., et al. Three-dimensional adult cardiac extracellular matrix promotes maturation of human induced pluripotent stem cell-derived cardiomyocytes. Tissue Engineering. Part A. 22, (15-16), 1016-1025 (2016).
  8. Atance, J., Yost, M. J., Carver, W. Influence of the extracellular matrix on the regulation of cardiac fibroblast behavior by mechanical stretch. Journal of Cell Physiology. 200, (3), 377-386 (2004).
  9. Belviso, I., et al. The microenvironment of decellularized extracellular matrix from heart failure myocardium alters the balance between angiogenic and fibrotic signals from stromal primitive cells. International Journal of Molecular Sciences. 21, (21), 7903 (2020).
  10. Badylak, S. F. Regenerative medicine and developmental biology: The role of the extracellular matrix. The Anatomical Record. 287, (1), 36-41 (2005).
  11. Volpato, F. Z., Führmann, T., Migliaresi, C., Hutmacher, D. W., Dalton, P. D. Using extracellular matrix for regenerative medicine in the spinal cord Using extracellular matrix for regenerative medicine in the spinal cord. Biomaterials. 34, (21), 4945-4955 (2013).
  12. Martino, M. M., et al. Extracellular matrix and growth factor engineering for controlled angiogenesis in regenerative medicine. Frontiers in Bioengineering and Biotechnology. 3, 45 (2015).
  13. Hussey, G. S., Keane, T. J., Badylak, S. F. The extracellular matrix of the gastrointestinal tract: a regenerative medicine platform. Nature Reviews: Gastroenterology Hepathology. 14, 540-552 (2017).
  14. Sheng, Y., Fei, D., Leiiei, G., Xiaosong, G. Extracellular matrix scaffolds for tissue engineering and regenerative medicine. Current Stem Cell Research and Therapy. 12, (3), 233-246 (2017).
  15. Rozario, T., De Simone, D. W. The extracellular matrix in development and morphogenesis: A dynamic view. Developmental Biology. 341, (1), 126-140 (2010).
  16. Takawale, A., Sakamuri, S. S. V. P., Kassiri, Z. Extracellular matrix communication and turnover in cardiac physiology and pathology. Comprehensive Physiology. 5, (2), 687-719 (2015).
  17. Li, L., Zhao, Q., Kong, W. Extracellular matrix remodeling and cardiac fibrosis. Matrix Biology. 68-69, 490-506 (2018).
  18. DeQuach, J. A., et al. Simple and high yielding method for preparing tissue specific extracellular matrix coatings for cell culture. PLoS ONE. 5, (9), 13039 (2010).
  19. Saldin, L. T., Cramer, M. C., Velankar, S. S., White, L. J., Badylak, S. F. Extracellular matrix hydrogels from decellularized tissue: Structure and function. Acta Biomaterialia. 49, 1-15 (2017).
  20. Tukmachev, D., et al. Injectable extracellular matrix hydrogels as scaffolds for spinal cord injury repair. Tissue Engineering. Part A. 22, (3-4), 306-317 (2016).
  21. Shoichet, M. S. Polymer scaffolds for biomaterials applications. Macromolecules. 43, (2), 581-591 (2010).
  22. Taylor, D. A., et al. Decellularized matrices in regenerative medicine. Acta Biomaterialia. 74, 74-89 (2018).
  23. Gilpin, A., Yang, Y. Decellularization strategies for regenerative medicine: From processing techniques to applications. BioMed Research International. 2017, 9831534 (2017).
  24. Heath, D. E. A review of decellularized extracellular matrix biomaterials for regenerative engineering applications. Regenerative Engineering and Translational Medicine. 5, (2), 155-166 (2019).
  25. Hoshiba, T., et al. Decellularized extracellular matrix as an in vitro model to study the comprehensive roles of the ECM in stem cell differentiation. Stem Cell International. 2016, 6397820 (2016).
  26. Keane, T. J., Swinehart, I. T., Badylak, S. F. Methods of tissue decellularization used for preparation of biologic scaffolds and in vivo relevance. Methods. 84, 5-34 (2015).
  27. Cerbotari, S., et al. Detergent decellularization of heart valves for tissue engineering: Toxicological effects of residual detergents on human endothelial cells. Artificial Organs. 34, (3), 206-210 (2010).
  28. Ott, H. C., et al. Perfusion-decellularized matrix: using nature's platform to engineer a bioartificial heart. Nature Medicine. 14, (2), 213-221 (2008).
  29. Uygun, B. E., et al. Organ reengineering through development of a transplantable recellularized liver graft using decellularized liver matrix. Nature Medicine. 16, (7), 814-820 (2010).
  30. Petersen, T. H., et al. Tissue-engineered lungs for in vivo implantation. Science. 329, (5991), 538-541 (2010).
  31. Calle, E. A., Petersen, T. H., Niklason, L. E. Procedure for lung engineering. Journal of Visualized Experiments. (49), e2651 (2011).
  32. Zeltinger, J., Landeen, L. K., Alexander, H. G., Kidd, I. D., Sibanda, B. Development and Characterization of Tissue-Engineered Aortic Valves. Tissue engineering. Part A. 7, (1), 9-22 (2001).
  33. Dahl, S., et al. Decellularized native and engineered arterial scaffolds for transplantation. Cell Transplantation. 12, (6), 659-666 (2003).
  34. Belviso, I., et al. Decellularized human dermal matrix as a biological scaffold for cardiac repair and regeneration. Frontiers inTissue Engineering and Regenerative Medicine. 8, 229 (2020).
  35. Greco, K. V., et al. Characterisation of porcine dermis scaffolds decellularised using a novel non-enzymatic method for biomedical applications. Journal of Biomaterials Applications. 30, (2), 239-253 (2015).
  36. Brouki, M. P., et al. Decellularization and preservation of human skin: A platform for tissue engineering and reconstructive surgery. Methods. 171, 62-67 (2020).
  37. Carbonaro, D., et al. A low-cost scalable 3D-printed sample-holder for agitation-based decellularization of biological tissues. Medical Engineering & Physics. 85, 7-15 (2020).
  38. Di Meglio, F., et al. Optimization of human myocardium decellularization method for the construction of implantable patches. Tissue Engineering. Part C Methods. 23, (9), 525-539 (2017).
  39. Crapo, P. M., Gilbert, T. W., Badylak, S. F. An overview of tissue and whole organ decellularization processes. Biomaterials. 32, (12), 3233-3243 (2011).
  40. Putame, G., et al. Compact and tunable stretch bioreactor advancing tissue engineering implementation. Application to engineered cardiac constructs. Medical Engineering & Physics. 84, 1-9 (2020).
  41. Parmaksiz, M., Dogan, A., Odabas, S., Elçin, A. E., Elçin, Y. M. Clinical applications of decellularized extracellular matrices for tissue engineering and regenerative medicine. Biomedical Materials. 11, (2), 022003 (2016).
  42. Sacco, A. M., et al. Diversity of dermal fibroblasts as major determinant of variability in cell reprogramming. Journal of Cellular and Molecular Medicine. 23, (6), 4256-4268 (2019).
  43. Belviso, I., et al. Isolation of adult human dermal fibroblasts from abdominal skin and generation of induced pluripotent stem cells using a non-integrating method. Journal of Visualized Experiments: JoVE. (155), e60629 (2020).
  44. Belviso, I., Di Meglio, F., Romano, V., Montagnani, S., Castaldo, C. Non-modified RNA-based reprogramming of human dermal fibroblasts into induced pluripotent stem cells. Methods in Molecular Biology. (2021).
This article has been published
Video Coming Soon
PDF DOI DOWNLOAD MATERIALS LIST

Cite this Article

Romano, V., Belviso, I., Cozzolino, D., Sacco, A. M., Schonauer, F., Nurzynska, D., Di Meglio, F., Castaldo, C. Decellularization for the Preparation of Highly Preserved Human Acellular Skin Matrix for Regenerative Medicine. J. Vis. Exp. (175), e62935, doi:10.3791/62935 (2021).More

Romano, V., Belviso, I., Cozzolino, D., Sacco, A. M., Schonauer, F., Nurzynska, D., Di Meglio, F., Castaldo, C. Decellularization for the Preparation of Highly Preserved Human Acellular Skin Matrix for Regenerative Medicine. J. Vis. Exp. (175), e62935, doi:10.3791/62935 (2021).

Less
Copy Citation Download Citation Reprints and Permissions
View Video

Get cutting-edge science videos from JoVE sent straight to your inbox every month.

Waiting X
Simple Hit Counter