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).&#160;

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
<ol>
	<li>Euthanize the mouse using a CO2 chamber.
	<ol>
		<li>Place mouse into a 4 L chamber, and begin to&#160;fill with 100% CO2, starting at 0.2 L/min for 2 min,&#160;increasing until reaching a flow of 0.8 L/min after 3 min.&#160;The mouse should fall unconscious during the first 2 min, and then respiration&#160;should cease (usually around 5 min, but flow can be maintained as necessary). Confirm death prior to proceeding to the next step.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Using microsurgical scissors, cut the pectoralis major and pectoralis minor muscles along the sixth intercostal space on both sides of the thoracic wall.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Using the cautery, cauterize the descending cava vein and, using straight pattern scissors, cut the esophagus.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Cauterize the brachiocephalic veins.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
2. Decellularization
<ol>
	<li>Connect the mouse to a pump system using silicone tubing and Luer connectors. Perfuse with deionized water at 200 &#181;L/min for 15 min. Maintain this flow rate during decellularization.</li>
	<li>Change the perfusion agent to 0.5% sodium deoxycholate (DOC) diluted in deionized water and perfuse overnight.</li>
	<li>Change the perfusion agent to 0.1% sodium dodecyl sulphate (SDS) diluted in deionized water and perfuse for 8 hours.</li>
	<li>Perfuse with deionized water overnight to wash away SDS and DOC for 24 h.</li>
	<li>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 &#181;M sodium azide at 4 &#176;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.</li>
</ol>
3. Immunostaining
<ol>
	<li>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.</li>
	<li>Block the sample by immersing it in a cryotube containing 6% (v/v) donkey serum - 3% (w/v) bovine serum albumin (BSA) overnight.</li>
	<li>Incubate with primary antibody (or antibodies) in 3% donkey serum in PBS for 24 h.</li>
	<li>Wash 5 times for 1 h each time in 0.05% tween 20 in PBS (PBST).</li>
	<li>Incubate the sample with fluorescently conjugated secondary antibody (or antibodies) in 3% donkey serum in PBS for 24 h.</li>
	<li>Wash 5 times for 1 hour in 0.05% (PBST). Wait 1 h between each wash.</li>
	<li>Add deionized water and store at 4 &#176;C away from direct light. At this point, the scaffold is ready to image.</li>
</ol>
4. Imaging
<ol>
	<li>Place the sample in a glass-bottomed dish and humidify it with two droplets of storing solution (PBS or deionized water).</li>
	<li>Prepare the objective. We recommend using a water immersion objective.</li>
	<li>Inspect the sample using fluorescence light.</li>
	<li>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.</li>
</ol>
5. Hematoxylin-eosin staining
<ol>
	<li>Excise 1 lung lobe from a euthanized mouse.</li>
	<li>Place in a 10 mm x 10 mm x 5 mm cryomold and cover it with approximately 500 &#181;L of OCT compound.</li>
	<li>Freeze on dry ice (-70 &#176;C) and maintain the sample at that temperature.</li>
	<li>Excise one decellularized lung lobe from a processed mouse according to step 2.5.</li>
	<li>Place in a cryomold with the largest surface area down and cover it with OCT compound as specified in step 5.2.</li>
	<li>Freeze on dry ice (-70&#176;C) and maintain the sample at that temperature until otherwise required. The sample can be stored for at least 12 weeks.</li>
	<li>Section frozen tissue blocks at -20 &#176;C in a cryostat with 5 &#181;m thickness and place sections on adhesive glass slides and store at -80 &#176;C.</li>
	<li>Take slides to room temperature until air dried (approximately 20 min).</li>
	<li>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.</li>
	<li>Immerse in Mayer&#39;s hematoxylin solution for 10 min. This time can be optimized according to tissue source and stain preparation.</li>
	<li>Wash in a Coplin jar under running distilled water for 10 min.</li>
	<li>Immerse in eosin solution for 7 min. This time can be optimized according to tissue source and stain preparation.</li>
	<li>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.</li>
	<li>Apply few drops of DPX mounting medium and place a glass coverslip.</li>
	<li>Leave slides to dry overnight under a chemical hood.</li>
	<li>Scan slides in a slide scanner.</li>
</ol>

<ol>
	<li>Hynes, R. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+matrix:+not+just+pretty+fibrils.">Extracellular matrix: not just pretty fibrils.</a> Science. 326, 1216-1219 (2009).</li><li>Mayorca-Guiliani, A. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ISDoT:+in+situ+decellularization+of+tissues+for+high-resolution+imaging+and+proteomic+analysis+of+native+extracellular+matrix.">ISDoT: in situ decellularization of tissues for high-resolution imaging and proteomic analysis of native extracellular matrix.</a> Nature Medicine. 23, 890-898 (2017).</li><li>Mayorca-Guiliani, A. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Decellularization+and+antibody+staining+of+mouse+tissues+to+map+native+extracellular+matrix+structures+in+3D.">Decellularization and antibody staining of mouse tissues to map native extracellular matrix structures in 3D.</a> Nature Protocols. 14, 3395-3425 (2019).</li><li>White, L. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+impact+of+detergents+on+the+tissue+decellularization+process:+A+TOF-sims+study.">The impact of detergents on the tissue decellularization process: A TOF-sims study.</a> Acta Biomaterialia. 50, 207-219 (2017).</li><li>Ott, H. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfusion-decellularized+matrix:+using+nature's+platform+to+engineer+a+bioartificial+heart.">Perfusion-decellularized matrix: using nature's platform to engineer a bioartificial heart.</a> Nature Medicine. 14, 213-221 (2008).</li><li>Uygun, B. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organ+re-engineering+through+development+of+a+transplantable+recellularized+liver+graft+using+decellularized+liver+matrix.">Organ re-engineering through development of a transplantable recellularized liver graft using decellularized liver matrix.</a> Nature Medicine. 16, 814-820 (2010).</li><li>Susaki, E. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advanced+CUBIC+protocols+for+whole-brain+and+whole-body+clearing+and+imaging.">Advanced CUBIC protocols for whole-brain and whole-body clearing and imaging.</a> Nature Protocols. 10, 1709-1727 (2015).</li><li>Tomer, R., Ye, L., Hsueh, B., Deisseroth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advanced+CLARITY+for+rapid+and+high-resolution+imaging+of+intact+tissues.">Advanced CLARITY for rapid and high-resolution imaging of intact tissues.</a> Nature Protocols. 9, 1682-1697 (2014).</li><li>Erturk, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+imaging+of+solvent-cleared+organs+using+3DISCO.">Three-dimensional imaging of solvent-cleared organs using 3DISCO.</a> Nature Protocols. 7, 1983-1995 (2012).</li><li>Wershof, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+FIJI+Macro+for+quantifying+pattern+in+extracellular+matrix.">A FIJI Macro for quantifying pattern in extracellular matrix.</a> BioRxiv. , (2019).</li></ol>

The authors have nothing to disclose.

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.

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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>MICROSURGERY</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>6-0 suture, triangular section needle (Vicryl)</td>
			<td>Ethicon</td>
			<td>6301124</td>
			<td></td>
		</tr>
		<tr>
			<td>9-0 micro-suture (Safil)</td>
			<td>B Braun</td>
			<td>G1048611</td>
			<td></td>
		</tr>
		<tr>
			<td>Adson forceps</td>
			<td>Fine Science Tools</td>
			<td>11006-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Adson forceps with teeth</td>
			<td>Fine Science Tools</td>
			<td>11027-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Castroviejo microneedle holder</td>
			<td>Fine Science Tools</td>
			<td>no. 12061-01</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 ventilation chamber for mouse euthanasia</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Deionized water (Milli-Q IQ 7000, Ultrapure lab water system)&#160;</td>
			<td>Merck</td>
			<td>ZIQ7000T0</td>
			<td></td>
		</tr>
		<tr>
			<td>Disposable polystyrene tray (~30 &#215; 50 cm)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection microscope (Greenough, with two-armed gooseneck)</td>
			<td>Leica</td>
			<td>S6 D</td>
			<td></td>
		</tr>
		<tr>
			<td>Double-ended microspatula</td>
			<td>Fine Science Tools</td>
			<td>10091-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont microforceps (two)</td>
			<td>Fine Science Tools</td>
			<td>11252-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont microforceps with 45&#176; tips (two)</td>
			<td>Fine Science Tools</td>
			<td>11251-35</td>
			<td></td>
		</tr>
		<tr>
			<td>Hair clippers</td>
			<td>Oster</td>
			<td>76998-320-051</td>
			<td></td>
		</tr>
		<tr>
			<td>Halsey needle holder (with tungsten carbide jaws)</td>
			<td>Fine Science Tools</td>
			<td>12500-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Intravenous 24-gauge catheter (Insyte)</td>
			<td>BD</td>
			<td>381512</td>
			<td></td>
		</tr>
		<tr>
			<td>Intravenous 26-gauge catheter (Terumo)</td>
			<td>Surflo-W</td>
			<td>SR+DM2619WX</td>
			<td></td>
		</tr>
		<tr>
			<td>Mayo scissors (tough cut, straight)</td>
			<td>Fine Science Tools</td>
			<td>14110-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Microforceps with ringed tips (two)</td>
			<td>Aesculap</td>
			<td>FM571R</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-spring scissors (Vannas, curved)</td>
			<td>Fine Science Tools</td>
			<td>15001-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Minicutter</td>
			<td>KLS Martin</td>
			<td>80-008-03-04</td>
			<td></td>
		</tr>
		<tr>
			<td>Molt Periostotome</td>
			<td>Aesculap</td>
			<td>D0543R</td>
			<td></td>
		</tr>
		<tr>
			<td>Needles (27 gauge; Microlance)</td>
			<td>BD</td>
			<td>21018</td>
			<td></td>
		</tr>
		<tr>
			<td>Paper towel (sterile) or surgical napkin&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Serrated scissors (CeramaCut, straight)</td>
			<td>Fine Science Tools</td>
			<td>14958-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Spatula (Freer-Yasargil)</td>
			<td>Aesculap</td>
			<td>OL166R</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringes (1 mL; Plastipak)</td>
			<td>BD</td>
			<td>3021001</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringes (10 mL; Plastipak)</td>
			<td>BD</td>
			<td>3021110</td>
			<td></td>
		</tr>
		<tr>
			<td>Tendon scissors (Walton)</td>
			<td>Fine Science Tools</td>
			<td>14077-09</td>
			<td></td>
		</tr>
		<tr>
			<td>IMMUNOSTAINING</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 donkey anti-guinea pig IgG</td>
			<td>Thermo Fisher Scientific</td>
			<td>A-11055</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 594 donkey anti-rabbit IgG</td>
			<td>Life Technologies</td>
			<td>A11037</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA(albumin bovine fraction V, standard grade, lyophilized)&#160;</td>
			<td>Serva</td>
			<td>11930.03</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagen IV polyclonal antibody (RRID: AB_2276457)&#160;</td>
			<td>Millipore</td>
			<td>AB756P</td>
			<td>Host: rabbit</td>
		</tr>
		<tr>
			<td>PBS (pH 7.4, 10&#215;, Gibco)&#160;</td>
			<td>Thermo Fisher Scientific</td>
			<td>70011044</td>
			<td>Host: goat</td>
		</tr>
		<tr>
			<td>Periostin polyclonal antibody (a kind gift from Manuel Koch. RRID:AB2801621)</td>
			<td></td>
			<td></td>
			<td>Host: guinea pig</td>
		</tr>
		<tr>
			<td>Scalpel disposable with blade no.11 (pcs. 10)</td>
			<td>VWR</td>
			<td>233-5364)</td>
			<td></td>
		</tr>
		<tr>
			<td>Serum (normal donkey serum)&#160;</td>
			<td>Jackson ImmunoResearch</td>
			<td>017-000-121</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td>Sigma-Aldrich</td>
			<td>P9416-50ML</td>
			<td></td>
		</tr>
		<tr>
			<td>IMAGING</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Detectors (hybrid detector (Leica, HyD S model) and photomultiplier tubes (PMTs; )&#160;</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Fluorescence light source&#160;</td>
			<td>Leica</td>
			<td>EL6000</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Microscope (inverted multiphoton microscope)&#160;</td>
			<td>Leica</td>
			<td>SP5-X MP</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Objective (lambda blue, 20&#215;, 0.70 numerical aperture (NA) IMM UV)&#160;</td>
			<td>Leica</td>
			<td>HCX PL APO</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Two-photon Ti&#8211;sapphire laser (Spectra-physics, Mai Tai DeepSee model)&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;White-light laser (WLL)&#160;</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DECELLULARIZATION</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>70% Ethanol (absolute alcohol 99.9%); absolute alcohol must be adjusted to 70% (vol/vol) using deionized water&#160;</td>
			<td>Plum</td>
			<td>1680766</td>
			<td></td>
		</tr>
		<tr>
			<td>Deionized water (Milli-Q IQ 7000, Ultrapure lab water system)&#160;</td>
			<td>Merck</td>
			<td>ZIQ7000T0</td>
			<td></td>
		</tr>
		<tr>
			<td>Luer-to-tubing male fittings (1/8 inch)</td>
			<td>World Precision Instruments</td>
			<td>13158-100</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS (pH 7.4, 10&#215;, Gibco)&#160;</td>
			<td>Thermo Fisher Scientific</td>
			<td>70011044</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-streptomycin</td>
			<td>Gibco</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>Peristaltic pump (with 12 channels)</td>
			<td>Ole Dich</td>
			<td>110AC(R)20G75</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicone tubing (with 2-mm i.d. and 4 mm o.d.)</td>
			<td>Ole Dich</td>
			<td>31399</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Azide</td>
			<td>Sigma-Aldrich</td>
			<td>08591-1ML-F</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium deoxycholate (DOC)</td>
			<td>Sigma-Aldrich</td>
			<td>D6750-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Dodecyl Sulphate</td>
			<td>Sigma-Aldrich</td>
			<td>L3771-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>H&#38;E STAINING</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4% PFA</td>
			<td>Fisher Scientific</td>
			<td>15434389</td>
			<td></td>
		</tr>
		<tr>
			<td>96% Ethanol</td>
			<td>Plum</td>
			<td>201446-5L</td>
			<td></td>
		</tr>
		<tr>
			<td>Absolute ethanol</td>
			<td>Plum</td>
			<td>201152-1L</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips (24x50mm; 1000 pcs)</td>
			<td>Hounisen</td>
			<td>422.245</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryomolds Intermediate (15 x 15 x 5 mm; 100 pcs)</td>
			<td>Tissue-Tek</td>
			<td>4566</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryostat</td>
			<td>Leica</td>
			<td>CM3050S</td>
			<td></td>
		</tr>
		<tr>
			<td>DPX mounting medium</td>
			<td>Hounisen</td>
			<td>1001.0025</td>
			<td></td>
		</tr>
		<tr>
			<td>Eosin Y solution alcoholic 0.5%</td>
			<td>Sigma</td>
			<td>1024390500</td>
			<td></td>
		</tr>
		<tr>
			<td>Feather microtome blade stainless steel,C35 (50 pcs)</td>
			<td>Pfm medical</td>
			<td>207500003</td>
			<td></td>
		</tr>
		<tr>
			<td> 
			Fisherbrand Superfrost Plus slides (25 x 75 mm; 144 pcs)</td>
			<td>Thermofisher</td>
			<td>6319483</td>
			<td></td>
		</tr>
		<tr>
			<td>Mayers hematoxylin</td>
			<td>Sigma</td>
			<td>MHS32-1L</td>
			<td></td>
		</tr>
		<tr>
			<td>OCT compound</td>
			<td>VWR</td>
			<td>361603E</td>
			<td></td>
		</tr>
		<tr>
			<td>Slide scanner (Nanozoomer)</td>
			<td>Hamamatsu Photonics</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Xylene</td>
			<td>Sigma</td>
			<td>534056-4L</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

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.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61854/61854fig1v2.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61854/61854fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61854/61854fig2v2.jpg" /> 
Figure 2. Micro-surgery schematic showing the key steps required to decellularize the cardio-pulmonary complex. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61854/61854fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61854/61854fig3v3.jpg" /> 
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.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61854/61854fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Decellularization of the Murine Cardiopulmonary Complex at JoVE.com

Decellularization of the Murine Cardiopulmonary Complex

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 ...

decellularization-of-the-murine-cardiopulmonary-complex

Nanyang Technological University

Research

JoVE Journal

Cancer Research

2.7K Views.  University of Copenhagen (UCPH). 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.

Video: Decellularization of the Murine Cardiopulmonary Complex

A Mouse Model of Fatigue Induced by Peripheral Irradiation

Cancer-related fatigue (CRF) is a distressing and costly condition that often affects patients receiving cancer treatments, including radiation therapy. Here we describe a method using targeted peripheral irradiation to induce fatigue-like behavior in mice. With appropriate shielding, the irradiation targets the lower abdominal/pelvic region of the mouse, sparing the brain, in an effort to model radiation treatment received by individuals with pelvic cancers. We deliver an irradiation dose that is sufficient to induce fatigue-like behavior in mice, measured by voluntary wheel-running activity (VWRA), without causing obvious morbidity. Since wheel running is a normal, voluntary behavior in mice, its use should have little confounding effect on other behavioral tests or biological measures. Hence, wheel running can be used as a feasible outcome measure in understanding the behavioral and biological correlates of fatigue. CRF is a complex condition with frequent comorbidities, and likely has causes related both to cancer and its various treatments. The methods described in this paper are useful for investigating radiation-induced changes that contribute to the development of CRF and, more generally, to explore the biological networks that can explain the development and persistence of a peripherally-triggered but centrally-driven behavior like fatigue.

We describe a method using targeted peripheral irradiation to induce fatigue-like behavior in mice. The selected non-lethal irradiation dose leads to a week-long reduction in voluntary wheel-running activity.

Cancer-related fatigue (CRF) is a distressing and costly condition that often affects patients receiving cancer treatments, including radiation therapy. ...

Generation and Isolation of Cell Cycle-arrested Cells with Complex Karyotypes

Chromosome mis-segregation leads to aneuploidy, a condition in which cells harbor an imbalanced chromosome number. Several lines of evidence strongly indicate that aneuploidy triggers genome instability, ultimately generating cells with complex karyotypes that arrest their proliferation. Isolation and characterization of cells harboring complex karyotypes are crucial to study the impact of an imbalanced chromosome number on cell physiology. To date, no methods have been established to reliably isolate such aneuploid cells. This paper provides a protocol for the enrichment and analysis of aneuploid cells with complex karyotypes utilizing standard, inexpensive tissue culture techniques. This protocol can be used to analyze several features of aneuploid cells with complex karyotypes including their induced senescence-associated secretory phenotype, pro-inflammatory properties, and ability to interact with immune cells. Because cancer cells often harbor imbalances in chromosome number, it is crucial to decipher how aneuploidy impacts cell physiology in normal cells, with the ultimate goal of uncovering both its pro- and anti-tumorigenic effects.

Aneuploidy leads to genome instability, which eventually produces cell cycle-arrested cells with complex karyotypes. This paper provides a simple and convenient method to isolate aneuploid cells with complex karyotypes that cease to divide.

Chromosome mis-segregation leads to aneuploidy, a condition in which cells harbor an imbalanced chromosome number. Several lines of evidence strongly ...

A Bioluminescent and Fluorescent Orthotopic Syngeneic Murine Model of Androgen-dependent and Castration-resistant Prostate Cancer

Orthotopic tumor modeling is a valuable tool for pre-clinical prostate cancer research, as it has multiple advantages over both subcutaneous and transgenic genetically engineered mouse models. Unlike subcutaneous tumors, orthotopic tumors contain more clinically accurate vasculature, tumor microenvironment, and responses to multiple therapies. In contrast to genetically engineered mouse models, orthotopic models can be performed with lower cost and in less time, involve the use of highly complex and heterogeneous mouse or human cancer cell lines, rather that single genetic alterations, and these cell lines can be genetically modified, such as to express imaging agents. Here, we present a protocol to surgically injecting a luciferase- and mCherry-expressing murine prostate cancer cell line into the anterior prostate lobe of mice. These mice developed orthotopic tumors that were non-invasively monitored in vivo and further analyzed for tumor volume, weight, mouse survival, and immune infiltration. Further, orthotopic tumor-bearing mice were surgically castrated, leading to immediate tumor regression and subsequent recurrence, representing castration-resistant prostate cancer. Although technical skill is required to carry out this procedure, this syngeneic orthotopic model of both androgen-dependent and castration-resistant prostate cancer is of great use to all investigators in the field.

The goal of this protocol is to demonstrate the intra-prostatic injection of prostate cancer cells, with subsequent castration. Orthotopic pre-clinical models of androgen-dependent and castration-resistant prostate cancer are critical to study the disease in the context of a clinically relevant tumor microenvironment and an immunocompetent host.

Orthotopic tumor modeling is a valuable tool for pre-clinical prostate cancer research, as it has multiple advantages over both subcutaneous and ...

Virus Delivery of CRISPR Guides to the Murine Prostate for Gene Alteration

With an increasing incidence of prostate cancer, identification of new tumor drivers or modulators is crucial. Genetically engineered mouse models (GEMM) for prostate cancer are hampered by tumor heterogeneity and its complex microevolution dynamics. Traditional prostate cancer mouse models include, amongst others, germline and conditional knockouts, transgenic expression of oncogenes, and xenograft models. Generation of de novo mutations in these models is complex, time-consuming, and costly. In addition, most of traditional models target the majority of the prostate epithelium, whereas human prostate cancer is well known to evolve as an isolated event in only a small subset of cells. Valuable models need to simulate not only prostate cancer initiation, but also progression to advanced disease.
Here we describe a method to target a few cells in the prostate epithelium by transducing cells by viral particles. The delivery of an engineered virus to the murine prostate allows alteration of gene expression in the prostate epithelia. Virus type and quantity will hereby define the number of targeted cells for gene alteration by transducing a few cells for cancer initiation and many cells for gene therapy. Through surgery-based injection in the anterior lobe, distal from the urinary track, the tumor in this model can expand without impairing the urinary function of the animal. Furthermore, by targeting only a subset of prostate epithelial cells the technique enables clonal expansion of the tumor, and therefore mimics human tumor initiation, progression, as well as invasion through the basal membrane.
This novel technique provides a powerful prostate cancer model with improved physiological relevance. Animal suffering is limited, and since no additional breeding is required, overall animal count is reduced. At the same time, analysis of new candidate genes and pathways is accelerated, which in turn is more cost efficient.

This protocol describes a newly established method for virus delivery to the murine prostate. Using either CRISPR/Cas9 technology, gene overexpression, or Cre recombinase delivery, the technique allows orthotopic alteration of gene expression and implements a novel mouse model for prostate cancer.

With an increasing incidence of prostate cancer, identification of new tumor drivers or modulators is crucial. Genetically engineered mouse models (GEMM) ...

Enrichment and Characterization of the Tumor Immune and Non-immune Microenvironments in Established Subcutaneous Murine Tumors

The tumor immune microenvironment (TIME) has recently been recognized as a critical mediator of treatment response in solid tumors, especially for immunotherapies. Recent clinical advances in immunotherapy highlight the need for reproducible methods to accurately and thoroughly characterize the tumor and its associated immune infiltrate. Tumor enzymatic digestion and flow cytometric analysis allow broad characterization of numerous immune cell subsets and phenotypes; however, depth of analysis is often limited by fluorophore restrictions on panel design and the need to acquire large tumor samples to observe rare immune populations of interest. Thus, we have developed an effective and high throughput method for separating and enriching the tumor immune infiltrate from the non-immune tumor components. The described tumor digestion and centrifugal density-based separation technique allows separate characterization of tumor and tumor immune infiltrate fractions and preserves cellular viability, and thus, provides a broad characterization of the tumor immunologic state. This method was used to characterize the extensive spatial immune heterogeneity in solid tumors, which further demonstrates the need for consistent whole tumor immunologic profiling techniques. Overall, this method provides an effective and adaptable technique for the immunologic characterization of subcutaneous solid murine tumors; as such, this tool can be used to better characterize the tumoral immunologic features and in the preclinical evaluation of novel immunotherapeutic strategies.

Here we describe a method to separate and enrich components of the tumor immune and non-immune microenvironment in established subcutaneous tumors. This technique allows for the separate analysis of tumor immune infiltrate and non-immune tumor fractions which can permit comprehensive characterization of the tumor immune microenvironment.

The tumor immune microenvironment (TIME) has recently been recognized as a critical mediator of treatment response in solid tumors, especially for ...

Rapid Generation of Primary Murine Melanocyte and Fibroblast Cultures

Defects in fibroblast or melanocyte function are associated with skin diseases, including poor barrier function, defective wound healing, pigmentation defects and cancer. Vital to the understanding and amelioration of these diseases are experiments in primary fibroblast and melanocyte cultures. Nevertheless, current protocols for melanocyte isolation require that the epidermal and dermal layers of the skin are trypsinized and manually disassociated. This process is time consuming, technically challenging and contributes to inconsistent yields. Furthermore, methods to simultaneously generate pure fibroblast cultures from the same tissue sample are not readily available. Here, we describe an improved protocol for isolating melanocytes and fibroblasts from the skin of mice on postnatal days 0-4. In this protocol, whole skin is mechanically homogenized using a tissue chopper and then briefly digested with collagenase and trypsin. Cell populations are then isolated through selective plating followed by G418 treatment. This procedure results in consistent melanocyte and fibroblast yields from a single mouse in less than 90 min. This protocol is also easily scalable, allowing researchers to process large cohorts of animals without a significant increase in hands-on time. We show through flow cytometric assessments that cultures established using this protocol are highly enriched for melanocytes or fibroblasts.

This protocol outlines a rapid method to simultaneously generate melanocyte and fibroblast cultures from the skin of 0-4 day old mice. These primary cultures can be maintained and manipulated in vitro to study a variety of physiologically relevant processes, including skin cell biology, pigmentation, wound healing and melanoma.

Defects in fibroblast or melanocyte function are associated with skin diseases, including poor barrier function, defective wound healing, pigmentation ...

Murine Model of Metastatic Liver Tumors in the Setting of Ischemia Reperfusion Injury

Liver ischemia and reperfusion (I/R) injury, a common clinical challenge, remains an inevitable pathophysiological process that has been shown to induce multiple tissue and organ damage. Despite recent advances and therapeutic approaches, the overall morbidity has remained unsatisfactory especially in patients with underlying parenchymal abnormalities. In the context of aggressive cancer growth and metastasis, surgical I/R is suspected to be the promoter regulating tumor recurrence. This article aims to describe a clinically relevant murine model of liver I/R and colorectal liver metastasis. In doing so, we aim to assist other investigators in establishing and perfecting this model for their routine research practice to better understand the effects of liver I/R on promoting liver metastases.

We describe in detail a clinically relevant colorectal cancer liver metastases (CRLM) tumor model and the influence of liver ischemia reperfusion (I/R) in tumor growth and metastasis. This model can help to better understand the mechanisms underlying surgery-induced promotion of liver metastatic growth.

Liver ischemia and reperfusion (I/R) injury, a common clinical challenge, remains an inevitable pathophysiological process that has been shown to induce ...

In Vitro Establishment of a Genetically Engineered Murine Head and Neck Cancer Cell Line using an Adeno-Associated Virus-Cas9 System

The use of primary normal epithelial cells makes it possible to reproducibly induce genomic alterations required for cellular transformation by introducing specific mutations in oncogenes and tumor suppressor genes, using clustered regulatory interspaced short palindromic repeat (CRISPR)-based genome editing technology in mice. This technology allows us to accurately mimic the genetic changes that occur in human cancers using mice. By genetically transforming murine primary cells, we can better study cancer development, progression, treatment, and diagnosis. In this study, we used Cre-inducible Cas9 mouse tongue epithelial cells to enable genome editing using adeno-associated virus (AAV) in vitro. Specifically, by altering KRAS, p53, and APC in normal tongue epithelial cells, we generated a murine head and neck cancer (HNC) cell line in vitro,which is tumorigenic in syngeneic mice. The method presented here describes in detail how to generate HNC cell lines with specific genomic alterations and explains their suitability for predicting tumor progression in syngeneic mice. We envision that this promising method will be informative and useful to study tumor biology and therapy of HNC.

Development of murine models with specific genes mutated in head and neck cancer patients is required for understanding of neoplasia. Here, we present a protocol for in vitro transformation of primary murine tongue cells using an adeno-associated virus-Cas9 system to generate murine HNC cell lines with specific genomic alterations.

The use of primary normal epithelial cells makes it possible to reproducibly induce genomic alterations required for cellular transformation by ...

Detection of Cell-Free DNA in Blood Plasma Samples of Cancer Patients

Identifying mutations in tumors of cancer patients is a very important step in disease management. These mutations serve as biomarkers for tumor diagnosis as well as for the treatment selection and its response in cancer patients. The current gold standard method for detecting tumor mutations involves a genetic test of tumor DNA by means of tumor biopsies. However, this invasive method is difficult to be performed repeatedly as a follow-up test of the tumor mutational repertoire. Liquid biopsy is a new and emerging technique for detecting tumor mutations as an easy-to-use and non-invasive biopsy approach.
Cancer cells multiply rapidly. In parallel, numerous cancer cells undergo apoptosis. Debris from these cells are released into a patient&#8217;s circulatory system, together with finely fragmented DNA pieces, called cell-free DNA (cfDNA) fragments, which carry tumor DNA mutations. Therefore, for identifying cfDNA based biomarkers using liquid biopsy technique, blood samples are collected from the cancer patients, followed by the separation of plasma and buffy coat. Next, plasma is processed for the isolation of cfDNA, and the respective buffy coat is processed for the isolation of a patient&#39;s genomic DNA. Both nucleic acid samples are then checked for their quantity and quality; and analyzed for mutations using next-generation sequencing (NGS) techniques.
In this manuscript, we present a detailed protocol for liquid biopsy, including blood collection, plasma, and buffy coat separation, cfDNA and germline DNA extraction, quantification of cfDNA or germline DNA, and cfDNA fragment enrichment analysis.

In this paper we present a detailed protocol for non-invasive liquid biopsy technique, including blood collection, plasma and buffy coat separation, cfDNA and germline DNA extraction, quantification of cfDNA or germline DNA, and cfDNA fragment enrichment analysis.

Identifying mutations in tumors of cancer patients is a very important step in disease management. These mutations serve as biomarkers for tumor diagnosis ...

Implantation and Evaluation of Melanoma in the Murine Choroid via Optical Coherence Tomography

Establishing experimental choroidal melanoma models is challenging in terms of the ability to induce tumors at the correct localization. In addition, difficulties in observing posterior choroidal melanoma in vivo limit tumor location and growth evaluation in real-time. The approach described here optimizes techniques for establishing choroidal melanoma in mice via a multi-step sub-choroidal B16LS9 cell injection procedure. To enable precision in injecting into the small dimensions of the mouse uvea, the complete procedure is performed under a microscope. First, a conjunctival peritomy is formed in the dorsal-temporal area of the eye. Then, a tract into the sub-choroidal space is created by inserting a needle through the exposed sclera. This is followed by the insertion of a blunt needle into the tract and the injection of melanoma cells into the choroid. Immediately after injection, noninvasive optical coherence tomography (OCT) imaging is utilized to determine tumor location and progress. Retinal detachment is evaluated as a predictor of tumor site and size. The presented method enables the reproducible induction of choroid-localized melanoma in mice and the live imaging of tumor growth evaluation. As such, it provides a valuable tool for studying intraocular tumors.

Implantation and Evaluation of Melanoma in the Murine Choroid via Optical Coherence Tomography

The present protocol describes the implantation and evaluation of melanoma in the murine choroid utilizing optical coherence tomography.

Establishing experimental choroidal melanoma models is challenging in terms of the ability to induce tumors at the correct localization. In addition, ...

Decellularization of the Murine Cardiopulmonary Complex

Summary

Explore More Videos

Chapters in this video

A Mouse Model of Fatigue Induced by Peripheral Irradiation

Generation and Isolation of Cell Cycle-arrested Cells with Complex Karyotypes

A Bioluminescent and Fluorescent Orthotopic Syngeneic Murine Model of Androgen-dependent and Castration-resistant Prostate Cancer

Virus Delivery of CRISPR Guides to the Murine Prostate for Gene Alteration

Enrichment and Characterization of the Tumor Immune and Non-immune Microenvironments in Established Subcutaneous Murine Tumors

Rapid Generation of Primary Murine Melanocyte and Fibroblast Cultures

Murine Model of Metastatic Liver Tumors in the Setting of Ischemia Reperfusion Injury

In Vitro Establishment of a Genetically Engineered Murine Head and Neck Cancer Cell Line using an Adeno-Associated Virus-Cas9 System

Detection of Cell-Free DNA in Blood Plasma Samples of Cancer Patients

Implantation and Evaluation of Melanoma in the Murine Choroid via Optical Coherence Tomography