The extracellular matrix (ECM) is a major regulator of homeostasis and disease, yet the 3D structure of the ECM remains poorly understood because of limitations in ECM visualization. We recently developed an ECM-specialized method termed in situ decellularization of tissues (ISDoT) to isolate native 3D ECM scaffolds from whole organs in which ECM structure and composition are preserved. Here, we present detailed surgical instructions to facilitate decellularization of 33 different mouse tissues and details of validated antibodies that enable the visualization of 35 mouse ECM proteins. Through mapping of these ECM proteins, the structure of the ECM can be determined and tissue structures visualized in detail. In this study, perfusion decellularization is presented for bones, skeletal muscle, tongue, salivary glands, stomach, duodenum, jejunum/ileum, large intestines, mesentery, liver, gallbladder, pancreas, trachea, bronchi, lungs, kidneys, urinary bladder, ovaries, uterine horn, cervix, adrenal gland, heart, arteries, veins, capillaries, lymph nodes, spleen, peripheral nerves, eye, outer ear, mammary glands, skin, and subcutaneous tissue. Decellularization, immunostaining, and imaging take 4–5 d.
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Organ-specific extracellular matrix directs trans-differentiation of mesenchymal stem cells and formation of salivary gland-like organoids in vivo
Stem Cell Research & Therapy Open Access 15 July 2022
Journal of Biological Engineering Open Access 30 May 2022
Journal of Leather Science and Engineering Open Access 15 January 2022
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All data generated during this study are included in this article (and its Supplementary information files), except for some of the data generated during the revision process. Unpublished data that support the findings of this study are available from the corresponding authors upon request.
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We thank N. M. Christensen at the Center for Advanced Bioimaging (CAB), University of Copenhagen, for providing microscope access and training; B. Kobbe for purifying collagen XXVIII antibodies; J. Christensen for providing technical advice; and E. Schoof for running the MS/MS profile. We thank P. D. Yurchenco for providing the perlecan-specific antibody. This work was supported by the European Research Council (ERC-2015-CoG-682881-MATRICAN; A.E.M.-G., O.W., M.R., R.R., & J.T.E.), the Danish Cancer Society (R204-A12454; R.R.), German Cancer Aid (Deutsche Krebshilfe; R.R.), a Hallas Møller Stipend from the Novo Nordisk Foundation (J.T.E.), a PhD fellow grant of the Lundbeck Foundation (R286-2018-621; M.R.) the Ragnar Söderberg Foundation Sweden (N91/15; C.D.M.), the Swedish Research Council (2017-03389; C.D.M.), and Cancerfonden Sweden (CAN 2016/783; C.D.M.), the German Research Foundation (DFG) (FOR2722/B2; M.K.), (FOR2722/B1; R.W.), (FOR2722/D2; F.Z.), (FOR2722/M2 and FOR2722/C2; G.S.).
The authors declare no competing interests.
Peer review information Nature Protocols thanks Benjamin Struecker and other anonymous reviewer(s) for their contribution to the peer review of this work.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Key reference using this protocol
Mayorca-Guiliani, A. E. et al. Nat. Med. 23, 890–898 (2017) https://doi.org/10.1038/nm.4352
Integrated supplementary information
Supplementary Figure 1 LC–MS/MS chromatogram of filtered and unfiltered peptide samples obtained from decellularized tongue tissue washed with 1% Triton X-100.
a. Chromatogram of the unfiltered sample reveals poor detection of peptides and a strong peak with high intensity for Triton X-100 (red frame). The NL value (normalized level of base peak) for Triton X-100 is 1.06E9. b. The chromatogram of the filtered sample shows improved peptide detection. The NL value of Triton X-100 shows a total reduction of intensity by 3 orders of magnitude after filtration (NL: 1.25E6).
a. Microvascular clamps. b. Cauterizer. c. Applying forceps for clamps. d. Double-ended microspatula. e. Castroviejo microneedle holder. f. Spring micro-scissors. g. Dumont microforceps with 45° tips. h. Microforceps with ringed tips. i. Periostotome. j. Freer-Yasargil (Spatula). k. Halsey needle holder. l. Mayo scissors. m. Metzenbaum scissors. n. Serrated scissors. o. Tendon scissors. p. Adson forceps with teeth. q. Adson forceps. r. 9-0 Micro-suture. s. 6-0 Suture. t. 26G catheter. u. 24G catheter.
a. Euthanized mouse pinned on the surgical table. Dotted lines indicate skin incisions. b. Subcutaneous dissection reveals the neck, the thoracic, and abdominal walls. c. Section of the pectoralis muscles (blue arrowheads) and incisions through the eight intercostal spaces (blue arrows). d. Intercostal incisions are united by sectioning the sternum horizontally (dotted line). e. Sternotomy (dotted line 1) and sectioning of the ribs along the thoracic wall (dotted lines 2 and 3). f. Pinning the sectioned thorax reveals the thymus (1), heart (2), lungs (3), and cava vein (4). g. Excision of the thymus reveals the aorta (1), brachiocephalic artery (2), left common carotid artery (3), left subclavian artery (4), internal mammary arteries (5), and brachiocephalic veins (6). h. Elevating the brachiocephalic veins allows clear differentiation of the major arteries (numbering equal to g). (Scale bars, 2 mm unless otherwise stated).
a. Thoracic procedures are based on the selective clamping of major vessels to redirect flow to an area of interest. b. To perfuse the face and neck, a catheter is inserted in the emergence of the aortic arch (1) and micro-sutured (2 and 4) to deliver flow superiorly to the right subclavian artery (3). c. To perfuse the cardio-pulmonary complex it is necessary to clamp the brachiocephalic artery (1), the left common carotid artery (2) and the left subclavian artery (3), leaving the aorta permeable to retrograde perfusion (4). d. To perfuse the territory of the left subclavian artery a clamp is placed immediately after the emergence of the left common carotid artery, leaving the left subclavian artery permeable to retrograde perfusion. e. To deliver perfusion to the cardio-pulmonary complex, insert a catheter into the cava vein at the level of its emergence from the abdominal cavity (1), suture it to avoid back flow (2); the tip must be place at the entrance of the atrium (3). f. The mouse is sectioned (dotted line) above the diaphragm. g. Expose the entrance of the abdominal aorta (1) by elevating the heart and lungs as well as displacing a cava vein catheter (in case of heart decellularization). h. A catheter is inserted into the aorta (1) and sutured (2) to deliver retrograde flow to operations depicted in 2, 3 and 4. (Scale bars, 2 mm).
a. Inferior half of mouse showing the abdominal walls and the incisions to access the peritoneal cavity (dotted lines) b. The abdominal aorta catheterized to deliver anterograde flow (1). c. Sectioning and elevating the peritoneum exposes the contents of the peritoneal wall. d. Elevating these contents to the right exposes the catheter (1) and allows incision of the diaphragm (dotted line). This incision is essential to follow the progress of the catheter. Avoid tearing the aorta. The liver (3), stomach (4), spleen (5), pancreas (6), and intestines are on the right, exposing the abdominal arteries and veins to their iliac bifurcation (9). e. To perfuse the celiac trunk (blue arrow), the tip of the catheter must be located superiorly to the emergence of the superior mesenteric artery and the clamp immediately below (blue line). The catheter is sutured to avoid back flow (circle). f. Perfusion of the superior mesenteric artery (blue arrow) requires suturing the catheter below the celiac trunk (circle) and clamping the aorta immediately below (blue line). g. Perfusing the kidneys requires suturing the catheter inferiorly to the superior mesenteric artery and clamping below the left renal artery (blue line). h. To perfuse the common iliac arteries the catheter is pushed until the tip is located at the aortic bifurcation (blue arrows) and sutured (circle) to avoid back flow. (Scale bars, 2 mm unless otherwise stated).
a. Elevating the liver and the intestines reveals the portal vein (1). b. The catheter is inserted as inferiorly as possible to ensure liver perfusion (arrow) and sutured to avoid back flow (circles). (Scale bars, 2 mm).
Supplementary Figure 7 Decellularized ECM immunostaining with a single primary antibody and a 488-nm emission secondary antibody.
All antibodies shown are alternatives to the ones used against the same antigen shown in Figs. 6 and 7. These antibodies have been produced in guinea-pig (anti-periostin, RRID:AB_2801620; anti-TGFBI, RRID:AB_2801623; anti-collagen XIV, RRID:AB_2801626; anti-collagen XXVIII, RRID:AB_2801634). (Scale bars, 100 µm). All experiments were carried out under the authorization and guidance of the Danish Inspectorate for Animal Experimentation.
Supplementary Figs. 1–7
Movie of a 3D reconstruction of a z-stack from decellularized tissue adjacent to the aorta stained for laminin-γ1 (rat anti-laminin-γ1, 555 nm, cyano) and collagen IV (goat anti-collagen IV, 647 nm, magenta). All experiments were carried out under the authorization and guidance of the Danish Inspectorate for Animal Experimentation.
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Mayorca-Guiliani, A.E., Willacy, O., Madsen, C.D. et al. Decellularization and antibody staining of mouse tissues to map native extracellular matrix structures in 3D. Nat Protoc 14, 3395–3425 (2019). https://doi.org/10.1038/s41596-019-0225-8
Organ-specific extracellular matrix directs trans-differentiation of mesenchymal stem cells and formation of salivary gland-like organoids in vivo
Stem Cell Research & Therapy (2022)
Journal of Biological Engineering (2022)
Journal of Leather Science and Engineering (2022)
Cell & Bioscience (2021)
Nature Reviews Cancer (2021)