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Cancer Research

Decellularization of the Murine Cardiopulmonary Complex

doi: 10.3791/61854 Published: May 30, 2021
Alejandro E. Mayorca-Guiliani1, Maria Rafaeva1, Oliver Willacy1, Chris D. Madsen2, Raphael Reuten1, Janine T. Erler1

Abstract

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.

Introduction

The ECM is a three-dimensional network made of proteins and glycans that accommodates all cells in a multicellular organism, giving organs their shape and regulating cell behavior throughout life1. From egg fertilization onwards, cells build and remodel the ECM, and are in turn strictly controlled by it. The purpose of this protocol is to open a way to analyze and map mouse ECM, as mice are the most used model organism in mammalian pathophysiology.

The development of this method was driven by the need to characterize and isolate metastasis-associated native ECM2. As tumors lack proper anatomical vascularization and mice are relatively small organisms, microsurgical procedures were designed to retrogradely catheterize the aorta, while isolating the circulation of the major vessel leading to a tumor (e.g., the pulmonary veins), thus focusing reagent flow and allowing tumor decellularization. This method produces ECM scaffolds with a conserved structure2 that can be immunostained and imaged, allowing ECM structure mapping in submicron detail. To carry out this protocol, it is necessary to acquire surgical and microsurgical skills (dissection, microsuturing and catheterization) that may represent a potential limitation to its use. To our knowledge, this method represents the state-of-the-art for native ECM structure imaging analysis2,3.

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Protocol

All procedures included here have been reviewed and approved by the ethical committee regulating experimental medicine in the University of Copenhagen and agree with Danish and European legislation. To demonstrate this protocol, we have used female BALB/cJ mice of 8-12 weeks of age and an MMTV-PyMT female mouse of 11 weeks of age.

NOTE: Avoiding bacterial contamination of the decellularized ECM scaffold gives the best imaging outcome and allows long-term sample storage. It is therefore important to keep all the steps sterile. As such, all instruments and surgical material, including suture, micro-suture, solutions, tubing, Luer connectors and catheters, must be sterile. Surfaces, including a polystyrene tray, must be disinfected with 70% ethanol, and the perfusion should preferably be carried out under a laminar flow hood. All procedures take place at room temperature unless otherwise indicated.

1. Post-mortem microsurgery

  1. Euthanize the mouse using a CO2 chamber.
    1. Use a 4 L chamber, filled with 100% CO2, starting at 0.2 L/min for 2 min and increasing until reaching a flow of 0.8 L/min after 3 min. The mouse should fall unconscious during the first 2 min, and then respiration should cease (usually around 5 min, but flow can be maintained as necessary).
  2. Shave the thorax, abdomen and back of the mouse with the hair clipper and disinfect with 70% ethanol. Shaving greatly reduces the number of artifacts due to the presence of hair either on samples for imaging or biochemical analysis.
  3. Pin the mouse to a polystyrene tray, extending its fore- and hindlimbs, as well as its head and tail. Place it under the microsurgery microscope.
  4. Using a Mayo straight pattern scissors establish surgical access with a cutaneous incision running from the submandibular region to the lower abdomen and dissect subcutaneously to expose the thoracic wall and peritoneum.
  5. Using microsurgical scissors, cut the pectoralis major and pectoralis minor muscles along the sixth intercostal space on both sides of the thoracic wall.
  6. Using straight-pattern scissors, cut the sternum along the previous incisions, and then complete a sternotomy by cutting the sternum along its long axis, then elevate and pin both sides of the thoracic wall to expose the cardiopulmonary complex.
  7. Using round-tipped micro-forceps (or Dumont micro-forceps), excise the thymus and surrounding adipose tissue by delicately pulling them off their attachments. This will reveal the major vessels.
  8. Using the cautery, cauterize the descending cava vein and, using straight pattern scissors, cut the esophagus.
  9. Using sharp micro-forceps, separate the brachiocephalic veins and the brachiocephalic, left common carotid and left subclavian arteries from the underlying tissue to facilitate ligation and cauterization.
  10. Using micro-needle holder, sharp micro-forceps and 9-0 suture place stitches above the emergence of the brachiocephalic, left common carotid and left subclavian arteries.
  11. Cauterize the brachiocephalic veins.
  12. Separate the submandibular salivary glands along the midline to expose the neck muscles and the trachea. Separate the muscles to expose the cricothyroid ligament. Using micro-scissors, open an entrance by sectioning the ligament.
  13. Introduce a 27 G catheter in the trachea and delicately push until the trachea branches into the bronchi (i.e., until resistance to the catheter is met, then retreat 3 mm). Be careful not to disrupt the bronchi. Using a 6-0 suture, place 3 stitches around the trachea to secure the catheter.
  14. Section the mouse at the height of the 12th thoracic vertebra. The descending aorta runs anteriorly to the spine and should be sectioned here along with the spine. Set the lower half apart.
  15. Retrogradely catheterize the aorta and push the catheter until it reaches the aortic arc. Using 9-0 suture, place 4 stitches around the aorta, beginning 5 mm below the catheter tip.

2. Decellularization

  1. Connect the mouse to a pump system using silicone tubing and Luer connectors. Perfuse with deionized water at 200 µL/min for 15 min. Maintain this flow rate during decellularization.
  2. Change the perfusion agent to 0.5% sodium deoxycholate (DOC) diluted in deionized water and perfuse overnight.
  3. Change the perfusion agent to 0.1% sodium dodecyl sulphate (SDS) diluted in deionized water and perfuse for 8 hours.
  4. Perfuse with deionized water overnight to wash away SDS and DOC for 24 h.
  5. Resect the decellularized heart and lungs by sectioning its attachments to the thorax using a curved scissors and store in a sterile cryo-tube with deionized water with 1% (v/v) penicillin-streptomycin and 0.3 µM sodium azide at 4 °C. ECM scaffolds can be stored for at least for 12 weeks1. If the scaffold will be used for biochemical analysis (e.g., mass spectrometry), snap freeze in liquid nitrogen.

3. Immunostaining

  1. Plan the imaging: determine primary antibody (or antibodies) and the combination of fluorescently conjugated secondary antibodies to match each other and to fit the laser lines of the fluorescence microscope.
  2. Block the sample by immersing it in a cryotube containing 6% (v/v) donkey serum - 3% (w/v) bovine serum albumin (BSA) overnight.
  3. Incubate with primary antibody (or antibodies) in 3% donkey serum in PBS for 24 h.
  4. Wash 5 times for 1 h each time in 0.05% tween 20 in PBS (PBST).
  5. Incubate the sample with fluorescently conjugated secondary antibody (or antibodies) in 3% donkey serum in PBS for 24 h.
  6. Wash 5 times for 1 hour in 0.05% (PBST). Wait 1 h between each wash.
  7. Add deionized water and store at 4 °C away from direct light. At this point, the scaffold is ready to image.

4. Imaging

  1. Place the sample in a glass-bottomed dish and humidify it with two droplets of storing solution (PBS or deionized water).
  2. Prepare the objective. We recommend using a water immersion objective.
  3. Inspect the sample using fluorescence light.
  4. Switch to computer control. Turn on lasers and adjust laser intensity, pinhole aperture, detectors wavelengths, gain, resolution and zoom. Set the number and step size for z-stack and begin acquisition. We recommend using multiphoton laser excitation to increase tissue penetration and to minimize scattering of light, bleaching and tissue damage.

5. Hematoxylin-eosin staining

  1. Excise 1 lung lobe from a euthanized mouse.
  2. Place in a 10 mm x 10 mm x 5 mm cryomold and cover it with approximately 500 µL of OCT compound.
  3. Freeze on dry ice (-70 °C) and maintain the sample at that temperature.
  4. Excise one decellularized lung lobe from a processed mouse according to step 2.5.
  5. Place in a cryomold with the largest surface area down and cover it with OCT compound as specified in step 5.2.
  6. Freeze on dry ice (-70°C) and maintain the sample at that temperature until otherwise required. The sample can be stored for at least 12 weeks.
  7. Section frozen tissue blocks at -20 °C in a cryostat with 5 µm thickness and place sections on adhesive glass slides and store at -80 °C.
  8. Take slides to room temperature until air dried (approximately 20 min).
  9. Shortly immerse in PBS and fix by immersing the slides in 4% paraformaldehyde in PBS for 15 min. Wash once in PBS for 5 min, then twice in distilled water for 5 min.
  10. Immerse in Mayer's hematoxylin solution for 10 min. This time can be optimized according to tissue source and stain preparation.
  11. Wash in a Coplin jar under running distilled water for 10 min.
  12. Immerse in eosin solution for 7 min. This time can be optimized according to tissue source and stain preparation.
  13. Dip in 50% ethanol to remove excess Eosin and dehydrate by shortly dipping in 70% ethanol, and in 96% and 100% ethanol for 30 seconds. Dip in xylene several times.
  14. Apply few drops of DPX mounting medium and place a glass coverslip.
  15. Leave slides to dry overnight under a chemical hood.
  16. Scan slides in a slide scanner.

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Representative Results

Cardiopulmonary decellularization
After successfully completing the protocol, the heart and lungs, as well as annex tissue such as the aortic arc, will be free of cells. Decellularization can be validated by hematoxylin-eosin staining (Figure 1) of the ECM scaffolds showing removal of the nuclei comparing to the native tissue. These scaffolds retain the dimensions of fresh organs and its insoluble ECM structure is intact2. Figure 2 shows a schematic representation of the key surgical steps required to successfully perfuse the mouse cardio-pulmonary complex.

ECM Imaging
In a standard setting, secondary antibodies can be used in green, red and far-red fluorescence channels (i.e., 488 nm, 555nm/594 nm and 647 nm wavelength detection); the addition of second harmonics generation (SHG) imaging using 2-photon excitation will reveal fibrillar collagen. Laser excitation can incite tissue autofluorescence and caution must be applied when using it with green fluorescence, as it may confound imaging data. A straightforward way to validate auto-fluorescence is to image an unstained control tissue and set laser intensity and detector gain accordingly and compare this with the antibody staining. However, this autofluorescence can be used as an advantage, as it can expose elastin in lungs scaffolds.

ECM scaffolds showed increased permeability and light penetrability2. Using this protocol with a motorized microscope stage allows for three-dimensional, tiled imaging of whole-mount) samples at submicron resolution (Figure 3). In case sectioning the tissue is necessary (e.g., to image cardiac walls or deep pulmonary parenchyma) tissue should be sectioned with a sharp scalpel before staining is conducted.

Figure 1
Figure 1. Validating decellularization. Hematoxylin-Eosin staining of snap frozen samples from native and decellularized lungs and heart. Notice the absence of nuclei in decellularized samples. All scales in microns. Please click here to view a larger version of this figure.

Figure 2
Figure 2. Micro-surgery schematic showing the key steps required to decellularize the cardio-pulmonary complex. Please click here to view a larger version of this figure.

Figure 3
Figure 3. Representative multiple protein immunostaining of decellularized PyMT mouse lungs from a 11-week-old female mouse. Tile mosaic showing the maximum projection of a z-stack. Inset 1 shows the pleura. Inset 2 shows normal parenchyma ECM. Inset 3 shows a bronchiole. The colors have been made accessible for the color blind. All scales in microns. Please click here to view a larger version of this figure.

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Discussion

Decellularization techniques based on tissue agitation alter ECM structure, making them unsuitable for ECM structure analysis4. Perfusion decellularization, using an anatomical route such as the aorta of the trachea, allows to reach the capillary bed, or terminal alveoli, and facilitates the delivery of decellularizing agents throughout the organ. The use of zwitterionic, anionic and non-ionic detergents to decellularize tissue is reported4,5,6, however, sodium dodecyl sulphate (SDS, anionic) linearizes fibrillar collagen in the mouse fat pad2 but not in the lungs; this suggests the choice of detergent must be optimized, adapting to the target tissue to maintain ECM structure. Tissue clearing methods could conceivably be used for ECM analysis, although they require chemicals that can change ECM cross linking, and tissue dimensions7,8,9. While acquired tissue transparency allows enhanced microscopic imaging, the presence of cells significantly worsens antibody penetration and may cover ECM epitopes/proteins. Isolating and imaging intact ECM permits quantitative analysis of its structure with analytical tools2,10, mapping its composition2 and opens the way for further ECM biochemical examination.

The dissection and ligation of major vessels and the consequent isolation of coronary and pulmonary circulation is necessary to achieve uniform pressure of perfused solutions throughout the tissues. Therefore, this protocol is dependent on the microsurgical expertise of the main operator. It is critical to operate with precision, so as to preserve vessels, lungs and heart intact. Executing this protocol repeatedly to understand the three-dimensional anatomy of the thorax is paramount to obtain consistent results.

The surgical procedure shown here sums the basic steps to access the mouse vasculature1,2 and the organs in its territory. By changing the ligature pattern, it is possible to access the head and neck, the fore limbs and fat pads. Using the same skills, it is possible to decellularize the sub-diaphragmatic organs.

Equally as important is the careful design of the immunostaining setup. We have previously compiled a catalogue of validated antibodies against structural ECM proteins3. The standard setup can reveal up to three proteins and fibrillar collagen simultaneously, enabling cross-examination.

The significance of this method lies in the possibility of obtaining structurally and dimensionally intact ECM scaffolds. The deconstruction of a complex tissue into discreet components is one of the fundamental goals of bioengineering; while it is relatively straightforward to isolate cells, or blood, from an organ, there were no methods to obtain its ECM scaffolding. This was especially true of tumors, but the method presented here opens the way for ECM isolation in any mouse strain for anatomical and biochemical analysis of the ECM.

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Disclosures

The authors have nothing to disclose.

Acknowledgments

We thank Prof. Ivana Novak and Dr. Nynne Meyn Christensen (Centre for Advanced Bioimaging (CAB), University of Copenhagen) for providing microscope access. This work was supported by the European Research Council (ERC-2015-CoG-682881-MATRICAN; AEM-G, OW, RR and JTE); a PhD fellowship from the Lundbeck Foundation (R286-2018-621; MR); the Swedish Research Council (2017-03389; CDM); the Swedish Cancer Society, Cancerfonden (CAN 2016/783, 19 0632 Pj, and 190007; CDM); German Cancer Aid (Deutsche Krebshilfe; RR); and the Danish Cancer Society (R204-A12454; RR). 

Materials

Name Company Catalog Number Comments
MICROSURGERY
6-0 suture, triangular section needle (Vicryl) Ethicon 6301124
9-0 micro-suture (Safil) B Braun G1048611
Adson forceps Fine Science Tools 11006-12
Adson forceps with teeth Fine Science Tools 11027-12
Castroviejo microneedle holder Fine Science Tools no. 12061-01
CO2 ventilation chamber for mouse euthanasia
Deionized water (Milli-Q IQ 7000, Ultrapure lab water system)  Merck ZIQ7000T0
Disposable polystyrene tray (~30 × 50 cm)
Dissection microscope (Greenough, with two-armed gooseneck) Leica S6 D
Double-ended microspatula Fine Science Tools 10091-12
Dumont microforceps (two) Fine Science Tools 11252-20
Dumont microforceps with 45° tips (two) Fine Science Tools 11251-35
Hair clippers Oster 76998-320-051
Halsey needle holder (with tungsten carbide jaws) Fine Science Tools 12500-12
Intravenous 24-gauge catheter (Insyte) BD 381512
Intravenous 26-gauge catheter (Terumo) Surflo-W SR+DM2619WX
Mayo scissors (tough cut, straight) Fine Science Tools 14110-15
Microforceps with ringed tips (two) Aesculap FM571R
Micro-spring scissors (Vannas, curved) Fine Science Tools 15001-08
Minicutter KLS Martin 80-008-03-04
Molt Periostotome Aesculap D0543R
Needles (27 gauge; Microlance) BD 21018
Paper towel (sterile) or surgical napkin 
Serrated scissors (CeramaCut, straight) Fine Science Tools 14958-09
Spatula (Freer-Yasargil) Aesculap OL166R
Syringes (1 mL; Plastipak) BD 3021001
Syringes (10 mL; Plastipak) BD 3021110
Tendon scissors (Walton) Fine Science Tools 14077-09
IMMUNOSTAINING
Alexa Fluor 488 donkey anti-guinea pig IgG Thermo Fisher Scientific A-11055
Alexa Fluor 594 donkey anti-rabbit IgG Life Technologies A11037
BSA(albumin bovine fraction V, standard grade, lyophilized)  Serva 11930.03
Collagen IV polyclonal antibody (RRID: AB_2276457)  Millipore AB756P Host: rabbit
PBS (pH 7.4, 10×, Gibco)  Thermo Fisher Scientific 70011044 Host: goat
Periostin polyclonal antibody (a kind gift from Manuel Koch. RRID:AB2801621) Host: guinea pig
Scalpel disposable with blade no.11 (pcs. 10) VWR 233-5364)
Serum (normal donkey serum)  Jackson ImmunoResearch 017-000-121
Tween 20 Sigma-Aldrich P9416-50ML
IMAGING
 Detectors (hybrid detector (Leica, HyD S model) and photomultiplier tubes (PMTs; )  Leica
 Fluorescence light source  Leica EL6000
 Microscope (inverted multiphoton microscope)  Leica SP5-X MP
 Objective (lambda blue, 20×, 0.70 numerical aperture (NA) IMM UV)  Leica HCX PL APO
 Two-photon Ti–sapphire laser (Spectra-physics, Mai Tai DeepSee model) 
 White-light laser (WLL)  Leica
DECELLULARIZATION
70% Ethanol (absolute alcohol 99.9%); absolute alcohol must be adjusted to 70% (vol/vol) using deionized water  Plum 1680766
Deionized water (Milli-Q IQ 7000, Ultrapure lab water system)  Merck ZIQ7000T0
Luer-to-tubing male fittings (1/8 inch) World Precision Instruments 13158-100
PBS (pH 7.4, 10×, Gibco)  Thermo Fisher Scientific 70011044
Penicillin-streptomycin Gibco 15140122
Peristaltic pump (with 12 channels) Ole Dich 110AC(R)20G75
Silicone tubing (with 2-mm i.d. and 4 mm o.d.) Ole Dich 31399
Sodium Azide Sigma-Aldrich 08591-1ML-F
Sodium deoxycholate (DOC) Sigma-Aldrich D6750-100G
Sodium Dodecyl Sulphate Sigma-Aldrich L3771-500G
H&E STAINING
4% PFA Fisher Scientific 15434389
96% Ethanol Plum 201446-5L
Absolute ethanol Plum 201152-1L
Coverslips (24x50mm; 1000 pcs) Hounisen 422.245
Cryomolds Intermediate (15 x 15 x 5 mm; 100 pcs) Tissue-Tek 4566
Cryostat Leica CM3050S
DPX mounting medium Hounisen 1001.0025
Eosin Y solution alcoholic 0.5% Sigma 1024390500
Feather microtome blade stainless steel,C35 (50 pcs) Pfm medical 207500003

Fisherbrand Superfrost Plus slides (25 x 75 mm; 144 pcs)
Thermofisher 6319483
Mayers hematoxylin Sigma MHS32-1L
OCT compound VWR 361603E
Slide scanner (Nanozoomer) Hamamatsu Photonics
Xylene Sigma 534056-4L

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References

  1. Hynes, R. O. Extracellular matrix: not just pretty fibrils. Science. 326, 1216-1219 (2009).
  2. Mayorca-Guiliani, A. E., et al. ISDoT: in situ decellularization of tissues for high-resolution imaging and proteomic analysis of native extracellular matrix. Nature Medicine. 23, 890-898 (2017).
  3. Mayorca-Guiliani, A. E., et al. Decellularization and antibody staining of mouse tissues to map native extracellular matrix structures in 3D. Nature Protocols. 14, 3395-3425 (2019).
  4. White, L. J., et al. The impact of detergents on the tissue decellularization process: A TOF-sims study. Acta Biomaterialia. 50, 207-219 (2017).
  5. Ott, H. C., et al. Perfusion-decellularized matrix: using nature's platform to engineer a bioartificial heart. Nature Medicine. 14, 213-221 (2008).
  6. Uygun, B. E., et al. Organ re-engineering through development of a transplantable recellularized liver graft using decellularized liver matrix. Nature Medicine. 16, 814-820 (2010).
  7. Susaki, E. A., et al. Advanced CUBIC protocols for whole-brain and whole-body clearing and imaging. Nature Protocols. 10, 1709-1727 (2015).
  8. Tomer, R., Ye, L., Hsueh, B., Deisseroth, K. Advanced CLARITY for rapid and high-resolution imaging of intact tissues. Nature Protocols. 9, 1682-1697 (2014).
  9. Erturk, A., et al. Three-dimensional imaging of solvent-cleared organs using 3DISCO. Nature Protocols. 7, 1983-1995 (2012).
  10. Wershof, E., et al. A FIJI Macro for quantifying pattern in extracellular matrix. BioRxiv. (2019).
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Cite this Article

Mayorca-Guiliani, A. E., Rafaeva, M., Willacy, O., Madsen, C. D., Reuten, R., Erler, J. T. Decellularization of the Murine Cardiopulmonary Complex. J. Vis. Exp. (171), e61854, doi:10.3791/61854 (2021).More

Mayorca-Guiliani, A. E., Rafaeva, M., Willacy, O., Madsen, C. D., Reuten, R., Erler, J. T. Decellularization of the Murine Cardiopulmonary Complex. J. Vis. Exp. (171), e61854, doi:10.3791/61854 (2021).

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