The authors would like to acknowledge the CCHMC Confocal Imaging Core for their assistance and guidance in development of this model, as well as Matt Batie from Clinical Engineering for the design of all 3D printed parts. Demetria Fischesser was supported by a training grant from the National Institutes of Health, (NHLBI, T32 HL125204) and Jeffery D. Molkentin was supported by the Howard Hughes Medical Institute.

All experiments involving mice were approved by the Institutional Animal Care and Use Committee (IACUC) at Cincinnati Children&#8217;s Hospital Medical Center. The institution is also AAALAC (American Association for Accreditation of Laboratory Animal Care) certified. Mice were euthanized via cervical dislocation, and mice undergoing survival surgical procedures were given pain relief (see below). All methods used for pain management and euthanasia are based on recommendations of the Panel on Euthanasia of the American Veterinary Medical Association. All mice were housed in corn cob bedding units with water and food available at all times. Mice were housed 4 to a cage with the same sex. For surgery or uninjured tissue clearing, equal numbers of 6&#8211;8-week-old male and female mice were used.
NOTE: Sterile surgical conditions were maintained in all surgeries. The surgeon changed into clean scrubs and a sterile gown and then donned shoe covers and a hairnet. The surgeon then scrubbed their hands with chlorhexidine and donned sterile surgical gloves. The surgeon was assisted by a technician who sedated, shaved, and scrubbed the incision site 3 times each, alternating between 2% chlorohexidine gluconate and 70% isopropanol. The mice were then brought to the surgeon, and surgery was performed. Between animals, the instruments were sterilized in a bead sterilizer.
1. Cre recombination
<ol>
	<li>Cross Rosa26-loxP-eGFP mice (Rosa26-nLacZ [FVB.Cg-Gt(ROSA)26Sortm1 (CAG-lacZ,-EGFP)Glh/J]) with Tcf21-MerCreMer mice (Figure 1A)6,21.</li>
	<li>When the offspring of this cross reach 6 weeks of age, administer an intraperitoneal injection of 75 mg/kg of tamoxifen dissolved in corn oil.22 Administer this tamoxifen twice, 48 h apart (Figure 1B).</li>
	<li>After tamoxifen injections, put the mice on a diet of tamoxifen food (~0.4 g/kg tamoxifen citrate) for one week. After one week of tamoxifen chow, return the mice to a normal autoclaved chow diet for one week before performing surgery (Figure 1B). Designate an appropriate number of mice (3 per surgical condition) to be uninjured controls. Sacrifice these mice, and proceed to the clearing process delineated below following the one week of normal chow diet.</li>
	<li>Following the tamoxifen regimen and recombination, perform surgery.</li>
</ol>
2. Surgical models
<ol>
	<li>Ischemia/reperfusion (I/R)23,24
	<ol>
		<li>Anesthetize mice with 1.5% isoflurane gas in room air in a ventilated box; shave their chests and necks with electric clippers, and then scrub the shaved areas with a 2% chlorhexidine gluconate-soaked swab.</li>
		<li>Use artificial tear ointment to prevent dryness during surgery by placing ointment on the eyes of the sedated mice.</li>
		<li>Perform a mid-neck cut using surgical scissors to allow visualization of the trachea. Intubate with a 20 G catheter by placing the intubation tube into the trachea and visualize the tracheal catheter through the mid-neck incision. Place the mice on a ventilator while sedated with isoflurane gas (1.5%) in room air.</li>
		<li>Make an incision under the left front limb with surgical scissors, and cut the intercostal muscles between ribs 3 and 4. Spread the ribs open using a retractor.</li>
		<li>Place a small sponge soaked in saline between the lung and the heart to prevent damage to the lung.</li>
		<li>Using a fishhook needle (6.5 mm, 3/8 c), tie the left coronary artery with a releasable slip knot using 8-0 prolene.</li>
		<li>Remove the sponge using forceps.</li>
		<li>Suture the intercostal muscles closed using 4-0 braided silk and a continuous suture while also exteriorizing the end of the 8-0 prolene slip knot.</li>
		<li>Close the skin incisions using topical tissue adhesive with the end of the occluding slip knot protruding.</li>
		<li>After one hour of occlusion, pull the exterior end of the slip knot to internally release the coronary artery occlusion, relieving the ischemia and causing reperfusion.</li>
		<li>Administer 0.02 mL of a 1 mg/mL extended release buprenorphine via subdermal injection (72 h release) as a pain reliever, and place the mice in an oxygenated incubation chamber at 37 &#176;C, separate from other animals. Monitor the mice at least every 15 min until they have recovered from anesthesia and are able to maintain a sternal or sitting position. Return the mice to their regular housing.</li>
		<li>Assess the animals for pain and distress for 48 h following surgery, and monitor the incision site daily until fully healed.</li>
		<li>Observe the mice for hydration, nourishment, and overall well-being following surgery until sacrifice.</li>
	</ol>
	</li>
	<li>Myocardial infarction (MI)25,26,27
	<ol>
		<li>Perform steps 2.1.1&#8211;2.1.7.</li>
		<li>Use a continuous suture to close intercostal muscles using 4-0 braided silk.</li>
		<li>Close the skin incision using topical tissue adhesive.</li>
		<li>Perform steps 2.1.11&#8211;2.1.13.</li>
	</ol>
	</li>
	<li>Angiotensin II/phenylephrine micro-osmotic pump infusion
	<ol>
		<li>Prepare Prepare a 10 &#956;g/&#956;L working solution of angiotensin II and 500 &#956;g/&#956;L working solution of phenylephrine under sterile conditions (i.e., in a laminar flow hood) by adding 1 mg of angiotensin II to 100 &#956;L of sterile phosphate buffered saline (PBS) and 250 mg of phenylephrine hydrochloride into 500 &#956;L of PBS.</li>
		<li>Calculate the dilution of each working solution and final volume to dispense to the micro-osmotic pump based on mouse weight. 
		NOTE: For instance, a 20 g mouse requires 20 g x 14 days x 1.5 &#956;g/day = 420 &#956;g angiotensin, or 42 &#956;L of working solution, as well as 20 g x 14 days x 50 &#956;g/day = 14000 &#956;g phenylephrine hydrochloride, or 28 &#956;L of working solution.</li>
		<li>For the weight of each mouse, dilute the working stock of the drug in sterile PBS and fill the micro-osmotic pumps (0.25 &#956;L/h, 14 days, approximately 100 &#956;L) using a 27 G needle and 1 mL syringe. 
		NOTE: This will generate a flow rate of 1.5 &#956;g&#8729;g-1&#8729;day-1 of angiotensin II and 50 &#956;g&#8729;g-1&#8729;day-1 of phenylephrine hydrochloride once placed in the mice.</li>
		<li>Anesthetize the mice with 1.5% isoflurane inhalation (to effect) in a ventilated chamber containing room air.</li>
		<li>Place the anesthetized mice on a sterile surgical table in the opine position, and maintain anesthesia with isoflurane gas inhalation (1.5%) through a nose cone.</li>
		<li>Use artificial tear ointment on the animal&#8217;s eyes to prevent dryness during surgery.</li>
		<li>Shave the fur over the implantation area with electric hair clippers, and sterilize the area to be cut using ethanol and a sterile swab. Make a small incision (approximately 1 cm) with surgical scissors in the epidermal layer of the mouse skin on the right lateral side of the back, below the shoulder blade. Use the dull sides of a pair of surgical scissors to gently stretch the skin in and around the area of implantation to insert the minipump.</li>
		<li>Place the micro-osmotic pump within the incision, and physically maneuver it to the left of the dorsal midline of the mouse by manually massaging the pump under the skin.</li>
		<li>Close the incision via continuous suture using 4-0 silk with a taper point needle.</li>
		<li>Following pump implantation, administer slow release buprenorphine at a dose of 0.1 mg/kg body weight via subdermal injection for analgesia (72-h slow release). Place the mice in an oxygenated incubation chamber at 37 &#176;C, separate from other animals. Monitor the mice at least every 15 min until they recover from anesthesia and can maintain a sternal or sitting position.</li>
		<li>Upon recovery from anesthesia in the 37 &#176;C warming chamber, place the mice back in their standard housing units.</li>
		<li>Monitor the mice for pain and distress for 48 h following surgery, and monitor the incision site daily until fully healed.</li>
		<li>Observe the mice for hydration, nourishment, and overall well-being following surgery until sacrifice.</li>
	</ol>
	</li>
</ol>
3. Clearing adult mouse hearts using a modified active CLARITY protocol
<ol>
	<li>Prepare a clean surgical area by sterilizing the surgery surface and all surgical tools with 70% ethanol. Prepare a solution of cold 1x PBS and 4% paraformaldehyde (PFA) in PBS (100 mL each). Wear gloves, lab coat, face mask, and eye protection.</li>
	<li>Prepare heparin buffer by diluting 10 units of heparin in 0.9% w/v sodium chloride solution. Inject each mouse intraperitoneally with 60 &#956;L of heparin-NaCl using a 1 mL syringe and 27 G needle.</li>
	<li>Five minutes after the injection of heparin-NaCl solution, anesthetize the mice using 1.5% isoflurane inhalation (to effect) in a ventilated chamber containing room air.</li>
	<li>Euthanize the mice by cervical dislocation, wherein the back of the head is held with the flat, closed end of forceps, while the spinal column is dislocated by pulling the tail.</li>
	<li>Clean the ventral surface of the mouse with 70% ethanol on a cotton swab. Make a 3 cm transverse incision approximately 3 cm below the xiphoid process using surgical scissors.</li>
	<li>Separate the skin from the underlying abdominal wall tissue by degloving the abdomen up to the xiphoid process (hold the skin closest to the tail and pull the skin closest to the head up toward the ribcage). Make a 2 cm transverse incision in the subcutaneous abdominal wall tissue 3 cm below the xiphoid process using surgical scissors. Make a vertical cut from this transverse incision, up the midline and through the ribcage. Pin the ribcage back, exposing the heart.</li>
	<li>Use a 27 G needle and 10 mL syringe to inject cold PBS into the superior vena cava and aorta (any positional manipulation of the heart should be done carefully with blunt forceps to avoid puncture) to clear blood from the heart. 
	NOTE: Sufficient perfusion can be noted by tail twitch and discoloration of the lungs.</li>
	<li>Use a 27 G needle and 10 mL syringe to inject cold 4% PFA into the superior vena cava and aorta to begin the fixation process.</li>
	<li>Excise the hearts. The atria, right ventricle and septum, and left ventricle should be separated using a straight-blade scalpel. Place this tissue into a 15 mL centrifuge tube filled with cold 4% PFA, and place on a nutator at 4 &#176;C overnight.</li>
	<li>Wash the heart 3 times for 1 h each with cold 1x PBS to remove excess PFA.</li>
	<li>Prepare the hydrogel (termed A4P0 for relative acrylamide/polyacrylamide composition) by mixing the following chemicals in a 15 mL centrifuge tube: 10% of 40% acrylamide, 10% 1x PBS, 80% distilled water.</li>
	<li>Add the 0.25% solution of the photoinitiator (2,2-Azobis[2-(2-imidazolin-2yl)propane)dihydrochloride) to the hydrogel solution just prior to submerging the heart in the hydrogel.</li>
	<li>Place the heart in a conical tube containing the hydrogel. Wrap the conical tube in foil, and incubate overnight at 4 &#176;C without physical disturbance.</li>
	<li>After approximately 14 h at 4 &#176;C, move the conical tube containing the heart to a 37 &#176;C bead bath.</li>
	<li>After 2.5 h in the bead bath, remove the heart from the hydrogel carefully with forceps.</li>
	<li>Wash the hearts 3 times for 1 h each in a 15 mL conical tube containing 1x PBS at 37 &#176;C on a nutator.</li>
	<li>Place the hearts in the basket of the active electrophoresis machine using forceps, and place the lid on basket. Ensure that the lid is securely in place.</li>
	<li>Fill the active electrophoresis chamber reservoir with electrophoretic clearing solution by pouring solution into the chamber.</li>
	<li>Once the chamber is full of electrophoretic clearing solution, the basket containing the heart can be submerged. Place the cap on the electrophoresis machine securely.</li>
	<li>Run the electrophoresis machine at 1.5 A, 37 &#176;C for 1.5 h.</li>
	<li>Following 1.5 h of electrophoresis, check the hearts visually to ensure that no opaque tissue remains. If regions of the tissue are still opaque, re-submerge the heart in electrophoresis solution in the electrophoretic chamber, and continue electrophoresis for 0.5 h at a time. 
	NOTE: Electrophoresis should be discontinued at first signs of tissue damage (such as tissue fraying).</li>
	<li>Wash the hearts in 15 mL conical tube containing 1x PBS for 1 h, 3 times each, at 37 &#176;C on a nutator.</li>
	<li>Submerge the hearts in a 15 mL conical tube containing N,N,N&#8242;,N&#8242;-Tetrakis(2-Hydroxypropyl)ethylenediamine&#8212;a decolorizing agent that reduces heme autofluorescence. 
	NOTE: This solution should be changed every 24 h until the hearts are completely clear (approximately 2 days).</li>
	<li>Wash the hearts in a 15 mL conical tube with 1x PBS 3 times for 1 h each to remove excess decolorizing agent.</li>
	<li>Prepare Refractive Index Matching Solution (RIMS) by combining 30 mL of 0.02 M phosphate buffer, 40 g of 5-(N-2, 3-dihydroxypropylacetamido)-2, 4, 6-tri-iodo-N, N&#39;-bis (2, 3 dihydroxypropyl) isophthalamide with stirring (must be completely dissolved&#8212;this can take up to an hour), 5 mg of sodium azide, 50 &#181;L of Tween20, and 1 g of 1,4-diazabicyclo[2.2.2]octane, and adjust the pH to 7.5.</li>
	<li>Equilibrate the hearts in RIMS for 48 h prior to imaging.</li>
</ol>
4. Imaging cleared hearts using an upright single photon confocal microscope
NOTE: The imaging apparatus consists of the bottom half of a 10 cm glass Petri dish, a 3D printed bottom reservoir, a round glass coverslip, and a 3D printed top reservoir (Figure 1C&#8211;E). 3D printed materials were made in-house by the Cincinnati Children&#8217;s Hospital Clinical Engineering Department.
<ol>
	<li>Use vacuum grease to seal the 3D printed bottom reservoir into a glass Petri dish by putting a thin layer of vacuum grease onto the bottom of the 3D printed piece (Figure 1C). 
	NOTE: The reservoir should be the same height as the sample being imaged (i.e., the sample should not be compressed by the glass coverslip placed on top of it, and the sample also should not be able to float and move within the reservoir).</li>
	<li>Fill the reservoir with RIMS, and remove all bubbles with a pipet tip. Place the heart in the RIMS carefully with the left ventricular wall facing up, ensuring that no bubbles are introduced into the solution.</li>
	<li>Adhere a glass coverslip to the bottom surface of the 3D printed top reservoir piece (Supplemental Figure 1A) using vacuum grease (again by placing a thin layer of vacuum grease onto the 3D printed piece, and placing it onto the cover slip) (Figure 1D).</li>
	<li>Place the glass coverslip and top reservoir, coverslip side down, onto the bottom reservoir (Supplemental Figure 1B), without the introduction of bubbles. The adhesion between the RIMS and cover slip will provide a seal between the top and bottom reservoir pieces (Figure 1E).</li>
	<li>Fill the top reservoir with glycerol for use with a 10x glycerol immersion objective.</li>
	<li>Use a single photon microscope outfitted with a multiphoton confocal scan head to image the cleared hearts.</li>
	<li>Lower the stage of the single photon microscope, and place the Petri dish containing the sample in the center of the stage. Raise the stage and lower the 10x glycerol immersion objective into the glycerol in the top reservoir of the sample apparatus. Ensure that the sample is in focus by looking at the edge of the sample using the eyepieces.</li>
	<li>Switch from eyepiece view to camera view on the computer by clicking on the live button.
	<ol>
		<li>Set the X and Y parameters by first locating the widest part of the tissue sample, which should be at the bottom of the Petri dish.</li>
		<li>Click on ND Acquisition and custom multipoint. Create multipoints by first finding the middle of the tissue sample by using the joystick to sweep left to right and top to bottom.</li>
		<li>Then, based on the size of the tissue, determine the tiling size needed (typically 6 x 5 for a 6&#8211;8-week-old mouse heart). Do this by running test imaging by only capturing X and Y parameters (uncheck z under the ND Acquisition tab) to check whether the entire length and width of the tissue were captured.</li>
	</ol>
	</li>
	<li>To set z parameters, open the XYZ navigation, and click on the up and down icons until finding the top and bottom of the sample.
	<ol>
		<li>Open the Z intensity correction panel and adjust the fluorescence at the top, middle, and bottom of the tissue to correct for tissue density by increasing or decreasing the laser power at each z point. Set these by clicking on the set arrow at the right of the Z Intensity Correction panel.</li>
		<li>Set the Z step size in the ND Acquisition panel to 5 &#956;m.</li>
	</ol>
	</li>
	<li>After the X, Y, and Z parameters of the heart are delineated and z-intensity correction is set, use the automated upright microscope with a 10x glycerol immersion objective with a resonant scanner and gallium arsenide phosphide photomultiplier tubes (GaAsP PMTs) to image the cleared hearts. Ensure that the XY and Z tabs are checked under ND acquisition, and press Run Z correction.</li>
	<li>Stitch, unmix, and denoise the resulting images by opening the multipoint image in the processing software and clicking on the stitch button. Under image, click on blind unmixing, 2 channel, and then find. and Under image, click on denoise.ai, then click on the fluorescent channel of choice (i.e., GFP). 
	NOTE: This process results in a 3D image of GFP-positive fibroblasts in the entire left ventricle.</li>
	<li>Use secondary analysis software to further identify patterns and trends in the dispersion of cardiac fibroblasts. Use the Spots function by running the spots wizard program.</li>
</ol>

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H., 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=Myocardial+ischemia+and+reperfusion:+A+murine+model.">Myocardial ischemia and reperfusion: A murine model.</a> American Journal of Physiology - Heart and Circuatory Physiology. 269 (6), 2147-2154 (1995).</li><li>Ahn, D., 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=Induction+of+myocardial+infarcts+of+a+predictable+size+and+location+by+branch+pattern+probability-assisted+coronary+ligation+in+C57BL/6+mice.">Induction of myocardial infarcts of a predictable size and location by branch pattern probability-assisted coronary ligation in C57BL/6 mice.</a> American Joural of Physiology: Circulatory Physiology. 286 (3), 1201-1207 (2004).</li><li>Patten, R. D., 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=Ventricular+remodeling+in+a+mouse+model+of+myocardial+infarction.">Ventricular remodeling in a mouse model of myocardial infarction.</a> American Journal of Physiology - Heart and Circuatory Physiology. 274 (5), 1812-1820 (1998).</li><li>Gao, X. M., Dart, A. M., Dewar, E., Jennings, G., Du, X. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Serial+echocardiographic+assessment+of+left+ventricular+dimensions+and+function+after+myocardial+infarction+in+mice.">Serial echocardiographic assessment of left ventricular dimensions and function after myocardial infarction in mice.</a> Cardiovascular Research. 45 (2), 330-338 (2000).</li><li>Vagnozzi, R. 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=An+acute+immune+response+underlies+the+benefit+of+cardiac+stem+cell+therapy.">An acute immune response underlies the benefit of cardiac stem cell therapy.</a> Nature. 577, 405-409 (2020).</li><li>Sengupta, P. P., Tajik, A. J., Chandrasekaran, K., Khandheria, B. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Twist+mechanics+of+the+left+ventricle.+Principles+and+application.">Twist mechanics of the left ventricle. Principles and application.</a> Journal of the American College of Cardiology: Cardiovascular Imaging. 1 (3), 366-376 (2008).</li><li>Arts, T., 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=Macroscopic+three-dimensional+motion+patterns+of+the+left+ventricle.">Macroscopic three-dimensional motion patterns of the left ventricle.</a> Advances in Experimental Medicine and Biology. 346, 383-392 (1993).</li><li>Willems, I. E. M. G., Havenith, M. G., De Mey, J. G. R., Daemen, M. J. A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+muscle+actin-positive+cells+in+healing+human+myocardial+scars.">The muscle actin-positive cells in healing human myocardial scars.</a> American Journal of Pathology. 145 (4), 868-875 (1994).</li><li>Hashmi, S., Al-Salam, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+myocardial+infarction+and+myocardial+ischemia-reperfusion+injury:+A+comparison.">Acute myocardial infarction and myocardial ischemia-reperfusion injury: A comparison.</a> International Journal of Clinical Experimental Pathology. 8, 8786-8796 (2015).</li></ol>

The authors have no disclosures related to the content of this manuscript.

This article presents a refined method for tissue clearing that allows for visualization of cardiac fibroblasts in vivo, both at baseline and following injury, to characterize and better understand fibroblasts in the mouse heart. This enhanced protocol addresses limitations in existing tissue clearing protocols that have attempted to identify specific cell types in the adult or neonatal heart12,13,14,<sup class="xref"...

Although cardiomyocytes comprise the greatest volume fraction in the heart, cardiac fibroblasts are more plentiful and are critically involved in regulating the baseline structural and reparative features of this organ. Cardiac fibroblasts are highly mobile, mechanically responsive, and phenotypically ranging depending on the extent of their activation. Cardiac fibroblasts are necessary to maintain normal levels of extracellular matrix (ECM), and too little or too much ECM production by these cells can lead to disease1,2,3. Given their importance in disease, cardiac fibroblasts have become an increasingly important topic of investigation towards identifying novel treatment strategies, especially in attempting to limit excessive fibrosis4,5,6,7. Upon injury, fibroblasts activate and differentiate into a more synthetic cell type known as a myofibroblast, which can be proliferative and secrete abundant ECM, as well as have contractile activity that helps remodel the ventricles.
While cardiac fibroblasts have been extensively evaluated for their properties in 2-D cultures6,8,9,10, much less is understood of their properties and dynamics in the 3-D living heart, either at baseline or with disease stimulation. Here, a refined method has been described to tissue clear the adult mouse heart while maintaining the fluorescence of fibroblasts labeled with a Rosa26-loxP-eGFP x Tcf21-MerCreMer genetic reporter system. Within the heart, Tcf21 is a relatively specific marker of quiescent fibroblasts4. After tamoxifen is given to activate the inducible MerCreMer protein, essentially all quiescent fibroblasts will permanently express enhanced green fluorescent protein (eGFP) from the Rosa26 locus, which allows for their tracking in vivo.
Numerous well-established tissue clearing protocols exist, some of which have been applied to the heart11,12,13,14,15,16,17. However, many of the reagents used in different tissue clearing protocols have been found to quench endogenous fluorescence signals18. Additionally, the adult heart is difficult to clear due to abundant heme group-containing proteins that generate autofluorescence19. Therefore, the goal of this protocol was to preserve fibroblast marker fluorescence with the simultaneous inhibition of heme autofluorescence in the injured adult heart for optimal 3-D visualization in vivo12,13,14,16,17,20.
Previous studies attempting to examine the cardiac fibroblast in vivo employed perfused antibodies to label these cells, although such studies were limited by antibody penetration and cardiac vascular structure14,16,17,20. Although Salamon et al. have shown tissue clearing with maintenance of topical neuronal fluorescence in the neonatal heart, and Nehrhoff et al. have shown maintenance of fluorescence marking myeloid cells, maintenance of endogenous fluorescence through the entire ventricular wall has not yet been demonstrated, including the visualization of adult cardiac fibroblasts at baseline or following injury13,20. This tissue clearing protocol refines a mixture of previous protocols based on the CLARITY method (clear lipid-exchanged acrylamide-hybridized rigid Imaging/immunostaining/in situ-hybridization-compatible tissue hydrogel) and PEGASOS (polyethylene glycol (PEG)-associated solvent system). This refined protocol permitted a more robust examination of cardiac fibroblasts in the mouse heart at baseline and of how they respond to different types of injury. The protocol is straightforward and reproducible and will help characterize the behavior of cardiac fibroblasts in vivo.

<table>
	<tbody>
		<tr>
			<td>4-0 braided silk</td>
			<td>Ethicon</td>
			<td>K871H</td>
			<td></td>
		</tr>
		<tr>
			<td>8-0 prolene</td>
			<td>Ethicon</td>
			<td>8730H</td>
			<td></td>
		</tr>
		<tr>
			<td>40% Acrylamide Solution</td>
			<td>Bio-Rad</td>
			<td>1610140</td>
			<td></td>
		</tr>
		<tr>
			<td>Angiotensin II</td>
			<td>Sigma</td>
			<td>A9525-50G</td>
			<td></td>
		</tr>
		<tr>
			<td>Artificial Tear Ointment</td>
			<td>Covetrus</td>
			<td>048272</td>
			<td></td>
		</tr>
		<tr>
			<td>DABCO (1,4-diazabicyclo[2.2. 2]octane)</td>
			<td>Millipore Sigma</td>
			<td>D27802-25G</td>
			<td></td>
		</tr>
		<tr>
			<td>GLUture topical tissue adhesive</td>
			<td>World Precision Instruments</td>
			<td>503763</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Sigma</td>
			<td>H0777</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaris Start Analysis Software</td>
			<td>Oxford Instruments</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-osmotic pumps</td>
			<td>Alzet</td>
			<td>Model 1002</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon Elements Analysis Software</td>
			<td>Nikon</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon A1R HD upright microscope</td>
			<td>Nikon</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal autoclaved chow</td>
			<td>Labdiet</td>
			<td>5010</td>
			<td></td>
		</tr>
		<tr>
			<td>Nycodenz, 5- (N-2, 3-dihydroxypropylacetamido)-2, 4, 6-tri-iodo-N, 
			N&#39;-bis (2, 3 dihydroxypropyl) isophthalamide</td>
			<td>CosmoBio</td>
			<td>AXS-1002424</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>15710</td>
			<td></td>
		</tr>
		<tr>
			<td>Phenylephrine Hydrochloride</td>
			<td>Sigma</td>
			<td>P6126-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>Photoinitiator</td>
			<td>Wako Chemicals</td>
			<td>VA-044</td>
			<td></td>
		</tr>
		<tr>
			<td>Rosa26-nLacZ [FVB.Cg-Gt(ROSA)26Sortm1 (CAG-lacZ,-EGFP)Glh/J]</td>
			<td>Jackson Laboratories</td>
			<td>Jax Stock No:012429</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Azide</td>
			<td>Sigma Aldrich</td>
			<td>S2002-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride solution</td>
			<td>Hospira, Inc.</td>
			<td>NDC 0409-4888-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Tamoxifen</td>
			<td>Sigma Aldrich</td>
			<td>T5648</td>
			<td></td>
		</tr>
		<tr>
			<td>Tamoxifen food</td>
			<td>Envigo</td>
			<td>TD.130860</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Thermo Fisher Scientific</td>
			<td>BP337-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Quadrol, N,N,N&#8242;,N&#8242;-Tetrakis(2-Hydroxypropyl)ethylenediamine, decolorizing agent</td>
			<td>Millipore Sigma</td>
			<td>122262-1L</td>
			<td></td>
		</tr>
		<tr>
			<td>X-Clarity electrophoretic clearing chamber</td>
			<td>Logos Biosystems</td>
			<td>C30001</td>
			<td></td>
		</tr>
		<tr>
			<td>X-Clarity electrophoretic clearing solution</td>
			<td>Logos Biosystems</td>
			<td>C13001</td>
			<td></td>
		</tr>
		<tr>
			<td>X-Clarity electrophoresis tissue basket</td>
			<td>Logos Biosystems</td>
			<td>C12001</td>
			<td></td>
		</tr>
		<tr>
			<td>X-Clarity electrophoresis tissue basket holder</td>
			<td>Logos Biosystems</td>
			<td>C12002</td>
			<td></td>
		</tr>
	</tbody>
</table>

refined clarity-based tissue clearing for three-dimensional fibroblast organization in healthy and injured mouse hearts

Cardiovascular disease is the most prevalent cause of mortality worldwide and is often marked by heightened cardiac fibrosis that can lead to increased ventricular stiffness with altered cardiac function. This increase in cardiac ventricular fibrosis is due to activation of resident fibroblasts, although how these cells operate within the 3-dimensional (3-D) heart, at baseline or after activation, is not well understood. To examine how fibroblasts contribute to heart disease and their dynamics in the 3-D heart, a refined CLARITY-based tissue clearing and imaging method was developed that shows fluorescently labeled cardiac fibroblasts within the entire mouse heart. Tissue resident fibroblasts were genetically labeled using Rosa26-loxP-eGFP florescent reporter mice crossed with the cardiac fibroblast expressing Tcf21-MerCreMer knock-in line. This technique was used to observe fibroblast localization dynamics throughout the entire adult left ventricle in healthy mice and in fibrotic mouse models of heart disease. Interestingly, in one injury model, unique patterns of cardiac fibroblasts were observed in the injured mouse heart that followed bands of wrapped fibers in the contractile direction. In ischemic injury models, fibroblast death occurred, followed by repopulation from the infarct border zone. Collectively, this refined cardiac tissue clarifying technique and digitized imaging system allows for 3-D visualization of cardiac fibroblasts in the heart without the limitations of antibody penetration failure or previous issues surrounding lost fluorescence due to tissue processing.

Cardiac fibroblasts are essential for baseline function of the heart as well as for response to cardiac injury. Previous attempts to understand the arrangement and morphology of these cells have been conducted largely in 2-D settings. However, a refined cardiac tissue clearing (Figure 2) and 3-D imaging technique has been published, which allows for the advanced, more detailed visualization of cardiac fibroblasts. With this imaging technique, fibroblasts were found to be densely packed and h...

Watch this Scientific Journal Video about Refined CLARITY-Based Tissue Clearing for Three-Dimensional Fibroblast Organization in Healthy and Injured Mouse Hearts at JoVE.com

Refined CLARITY-Based Tissue Clearing for Three-Dimensional Fibroblast Organization in Healthy and Injured Mouse Hearts

A refined method of tissue clearing was developed and applied to the adult mouse heart. This method was designed to clear dense, autofluorescent cardiac tissue, while maintaining labeled fibroblast fluorescence attributed to a genetic reporter strategy.

Cardiovascular disease is the most prevalent cause of mortality worldwide and is often marked by heightened cardiac fibrosis that can lead to increased ...

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4.6K Views.  University of Cincinnati College of Medicine. A refined method of tissue clearing was developed and applied to the adult mouse heart. This method was designed to clear dense, autofluorescent cardiac tissue, while maintaining labeled fibroblast fluorescence attributed to a genetic reporter strategy.

Video: Refined CLARITY-Based Tissue Clearing for Three-Dimensional Fibroblast Organization in Healthy and Injured Mouse Hearts

Heterotypic Three-dimensional In Vitro Modeling of Stromal-Epithelial Interactions During Ovarian Cancer Initiation and Progression

Epithelial ovarian cancers (EOCs) are the leading cause of death from gynecological malignancy in Western societies. Despite advances in surgical treatments and improved platinum-based chemotherapies, there has been little improvement in EOC survival rates for more than four decades 1,2. Whilst stage I tumors have 5-year survival rates &gt;85%, survival rates for stage III/IV disease are &lt;40%. Thus, the high rates of mortality for EOC could be significantly decreased if tumors were detected at earlier, more treatable, stages 3-5. At present, the molecular genetic and biological basis of early stage disease development is poorly understood. More specifically, little is known about the role of the microenvironment during tumor initiation; but known risk factors for EOCs (e.g. age and parity) suggest that the microenvironment plays a key role in the early genesis of EOCs. We therefore developed three-dimensional heterotypic models of both the normal ovary and of early stage ovarian cancers. For the normal ovary, we co-cultured normal ovarian surface epithelial (IOSE) and normal stromal fibroblast (INOF)&nbsp;cells, immortalized by retrovrial transduction of the catalytic subunit of human telomerase holoenzyme (hTERT) to extend the lifespan of these cells in culture. To model the earliest stages of ovarian epithelial cell transformation, overexpression of the CMYC oncogene in IOSE cells, again co-cultured with INOF cells. These heterotypic models were used to investigate the effects of aging and senescence on the transformation and invasion of epithelial cells. Here we describe the methodological steps in development of these three-dimensional model; these methodologies aren't specific to the development of normal ovary and ovarian cancer tissues, and could be used to study other tissue types where stromal and epithelial cell interactions are a fundamental aspect of the tissue maintenance and disease development.

Heterotypic Three-dimensional In Vitro Modeling of Stromal-Epithelial Interactions During Ovarian Cancer Initiation and Progression

We describe methodologies for establishing in vitro heterotypic three-dimensional models comprising ovarian fibroblasts and normal ovarian surface or ovarian cancer epithelial cells. We discuss the use of these models to study stromal-epithelial interactions that occur during ovarian cancer development.

Epithelial ovarian cancers (EOCs) are the leading cause of death from gynecological malignancy in Western societies. Despite advances in surgical ...

A Three-dimensional Tissue Culture Model to Study Primary Human Bone Marrow and its Malignancies

Tissue culture has been an invaluable tool to study many aspects of cell function, from normal development to disease. Conventional cell culture methods rely on the ability of cells either to attach to a solid substratum of a tissue culture dish or to grow in suspension in liquid medium. Multiple immortal cell lines have been created and grown using such approaches, however, these methods frequently fail when primary cells need to be grown ex vivo. Such failure has been attributed to the absence of the appropriate extracellular matrix components of the tissue microenvironment from the standard systems where tissue culture plastic is used as a surface for cell growth. Extracellular matrix is an integral component of the tissue microenvironment and its presence is crucial for the maintenance of physiological functions such as cell polarization, survival, and proliferation. Here we present a 3-dimensional tissue culture method where primary bone marrow cells are grown in extracellular matrix formulated to recapitulate the microenvironment of the human bone (rBM system). Embedded in the extracellular matrix, cells are supplied with nutrients through the medium supplemented with human plasma, thus providing a comprehensive system where cell survival and proliferation can be sustained for up to 30 days while maintaining the cellular composition of the primary tissue. Using the rBM system we have successfully grown primary bone marrow cells from normal donors and patients with amyloidosis, and various hematological malignancies. The rBM system allows for direct, in-matrix real time visualization of the cell behavior and evaluation of preclinical efficacy of novel therapeutics. Moreover, cells can be isolated from the rBM and subsequently used for in vivo transplantation, cell sorting, flow cytometry, and nucleic acid and protein analysis. Taken together, the rBM method provides a reliable system for the growth of primary bone marrow cells under physiological conditions.

In standard culture methods cells are taken out of their physiological environment and grown on the plastic surface of a dish. To study the behavior of primary human bone marrow cells we created a 3-D culture system where cells are grown under conditions recapitulating the native microenvironment of the tissue.

Tissue culture has been an invaluable tool to study many aspects of cell function, from normal development to disease. Conventional cell culture methods ...

Three-dimensional Co-culture Model for Tumor-stromal Interaction

Cancer progression (initiation, growth, invasion and metastasis) occurs through interactions between malignant cells and the surrounding tumor stromal cells. The tumor microenvironment is comprised of a variety of cell types, such as fibroblasts, immune cells, vascular endothelial cells, pericytes and bone-marrow-derived cells, embedded in the extracellular matrix (ECM). Cancer-associated fibroblasts (CAFs) have a pro-tumorigenic role through the secretion of soluble factors, angiogenesis and ECM remodeling. The experimental models for cancer cell survival, proliferation, migration, and invasion have mostly relied on two-dimensional monocellular and monolayer tissue cultures or Boyden chamber assays. However, these experiments do not precisely reflect the physiological or pathological conditions in a diseased organ. To gain a better understanding of tumor stromal or tumor matrix interactions, multicellular and three-dimensional cultures provide more powerful tools for investigating intercellular communication and ECM-dependent modulation of cancer cell behavior. As a platform for this type of study, we present an experimental model in which cancer cells are cultured on collagen gels embedded with primary cultures of CAFs.

Here we present a protocol to co-culture in three-dimensions, which is useful for investigating multicellular interactions and extracellular matrix-dependent modulation of cancer cell behavior. In this experimental model, cancer cells are cultured on collagen gels embedded with human cancer-associated fibroblasts.

Cancer progression (initiation, growth, invasion and metastasis) occurs through interactions between malignant cells and the surrounding tumor stromal ...

Three-Dimensional (3D) Tumor Spheroid Invasion Assay

Invasion of surrounding normal tissues is generally considered to be a key hallmark of malignant (as opposed to benign) tumors. For some cancers in particular (e.g., brain tumors such as glioblastoma multiforme and squamous cell carcinoma of the head and neck &#8211; SCCHN) it is a cause of severe morbidity and can be life-threatening even in the absence of distant metastases. In addition, cancers which have relapsed following treatment unfortunately often present with a more aggressive phenotype. Therefore, there is an opportunity to target the process of invasion to provide novel therapies that could be complementary to standard anti-proliferative agents. Until now, this strategy has been hampered by the lack of robust, reproducible assays suitable for a detailed analysis of invasion and for drug screening. Here we provide a simple micro-plate method (based on uniform, self-assembling 3D tumor spheroids) which has great potential for such studies. We exemplify the assay platform using a human glioblastoma cell line and also an SCCHN model where the development of resistance against targeted epidermal growth factor receptor (EGFR) inhibitors is associated with enhanced matrix-invasive potential. We also provide two alternative methods of semi-automated quantification: one using an imaging cytometer and a second which simply requires standard microscopy and image capture with digital image analysis.

Three-Dimensional 3D Tumor Spheroid Invasion Assay

Invasion of surrounding normal tissues is a defining characteristic of malignant tumors. We provide here a simple, semi-automated micro-plate assay of invasion into a natural 3D biomatrix that has been exemplified with a number of models of advanced human cancers.

Invasion of surrounding normal tissues is generally considered to be a key hallmark of malignant (as opposed to benign) tumors. For some cancers in ...

Three-dimensional Alginate-bead Culture of Human Pituitary Adenoma Cells

A three-dimensional culture method is described in which primary pituitary adenoma cells are grown in alginate beads. Alginate is a polymer derived from brown sea algae. Briefly, the tumor tissue is cut into small pieces and submitted to an enzymatic digestion with collagenase and trypsin. Next, a cell suspension is obtained. The tumor cell suspension is mixed with 1.2% sodium alginate and dropped into a CaCl2 solution, and the alginate/cell suspension is gelled on contact with the CaCl2 to form spherical beads. The cells embedded in the alginate beads are supplied with nutrients provided by the culture media enriched with 20% FBS. Three-dimensional culture in alginate beads maintains the viability of adenoma cells for long periods of time, up to four months. Moreover, the cells can be liberated from the alginate by washing the beads with sodium citrate and seeded on glass coverslips for further immunocytochemical analyses. The use of a cell culture model allows for the fixation and visualization of the actin cytoskeleton with minimal disorganization. In summary, alginate beads provide a reliable culture system for the maintenance of pituitary adenoma cells.

Three-dimensional culture in alginate beads, due to its simplicity and reproducibility, was chosen to maintain the pituitary adenoma cells. The procedures included an initial enzymatic digestion and mechanical dissociation of the tumor tissue, and the subsequent cell suspension was encapsulated in alginate beads.

A three-dimensional culture method is described in which primary pituitary adenoma cells are grown in alginate beads. Alginate is a polymer derived from ...

Oral Biofilm Sampling for Microbiome Analysis in Healthy Children

Oral biofilm and its molecular analysis provide a basis for investigating various dental research and clinical questions. Knowledge of biofilm composition leads to a better understanding of cariogenic and periopathogenic mechanisms. Microbial changes taking place in the oral cavity during childhood are of interest for several reasons. The evolution of the child oral microbiota and shifts in its composition need to be analyzed further to understand and possibly prevent the onset of disease. At the same time, advanced knowledge of the natural composition of oral biofilm is needed. Early stages of caries-free permanent dentition with healthy gums provide a widely unaffected subgingival habitat that can serve as an in situ baseline for studying features of oral health and disease. Analysis of children's oral biofilm during different stages in life is thus an important theme in the field. Modern molecular analysis methods can provide comprehensive information about the bacterial diversity of such biofilms. To enable microbiota data comparison, it is important to standardize each step in the procedure for molecular data generation. This procedure spans from clinical sampling, Next Generation Sequencing (NGS), bioinformatic data processing, to taxonomic interpretation. One of the most critical factors here is biofilm sampling. Sampling in children is even more challenging in particular due to limited space in subgingival areas. We thus focus on the use of paper points for subgingival sampling. This article provides a detailed protocol for oral biofilm sampling of the subgingival sulcus, the mucosa, and saliva in children.

Changes in the oral microbiome throughout childhood are of growing interest. Comparison of different microbiome studies reveals a lack of standardized sampling protocols. Limited space makes sampling the sound subgingival sulcus of children challenging. Paper point sampling is presented here in detail as the method of choice for this area.

Oral biofilm and its molecular analysis provide a basis for investigating various dental research and clinical questions. Knowledge of biofilm composition ...

Optical Clearing and Imaging of Immunolabeled Kidney Tissue

Optical clearing techniques render tissue transparent by equilibrating the refractive index throughout a sample for subsequent three-dimensional (3-D) imaging. They have received great attention in all research areas for the potential to analyze microscopic multicellular structures that extend over macroscopic distances. Given that kidney tubules, vasculature, nerves, and glomeruli extend in many directions, which have been only partially captured by traditional two-dimensional techniques so far, tissue clearing also opened up many new areas of kidney research. The list of optical clearing methods is rapidly growing, but it remains difficult for beginners in this field to choose the best method for a given research question. Provided here is a simple method that combines antibody labeling of thick mouse kidney slices; optical clearing with cheap, non-toxic and ready-to-use chemical ethyl cinnamate; and confocal imaging. This protocol describes how to perfuse kidneys and use an antigen-retrieval step to increase antibody- binding without requiring specialized equipment. Its application is presented in imaging different multicellular structures within the kidney, and how to troubleshoot poor antibody penetration into tissue is addressed. We also discuss the potential difficulties of imaging endogenous fluorophores and acquiring very large samples and how to overcome them. This simple protocol provides an easy-to-setup and comprehensive tool to study tissue in three dimensions.

The combination of antibody labeling, optical clearing, and advanced light microscopy allows three-dimensional analysis of complete structures or organs. Described here is a simple method to combine immunolabeling of thick kidney slices, optical clearing with ethyl cinnamate, and confocal imaging that enables visualization and quantification of three-dimensional kidney structures.

Optical clearing techniques render tissue transparent by equilibrating the refractive index throughout a sample for subsequent three-dimensional (3-D) ...

On-Site Sampling and Extraction of Brain Tumors for Metabolomics and Lipidomics Analysis

Despite the variety of tools available for cancer diagnosis and classification, methods that enable fast and simple characterization of tumors are still in need. In recent years, mass spectrometry has become a method of choice for untargeted profiling of discriminatory compound as potential biomarkers of a disease. Biofluids are generally considered as preferable matrices given their accessibility and easier sample processing while direct tissue profiling provides more selective information about a given cancer. Preparation of tissues for the analysis via traditional methods is much more complex and time-consuming, and, therefore, not suitable for fast on-site analysis. The current work presents a protocol combining sample preparation and extraction of small molecules on-site, immediately after tumor resection. The sampling device, which is of the size of an acupuncture needle, can be inserted directly into the tissue and then transported to the nearby laboratory for instrumental analysis. The results of metabolomics and lipidomics analyses demonstrate the capability of the approach for the establishment of phenotypes of tumors related to the histological origin of the tumor, malignancy, and genetic mutations, as well as for the selection of discriminating compounds or potential biomarkers. The non-destructive nature of the technique permits subsequent performance of routinely used tests e.g., histological tests, on the same samples used for SPME analysis, thus enabling attainment of more comprehensive information to support personalized diagnostics.

The manuscript presents on-site sampling of human brain tumors with solid phase microextraction followed by their biochemical profiling towards the discovery of biomarkers.

Despite the variety of tools available for cancer diagnosis and classification, methods that enable fast and simple characterization of tumors are still ...

Advanced Cardiac Rhythm Management by Applying Optogenetic Multi-Site Photostimulation in Murine Hearts

Ventricular tachyarrhythmias are a major cause of mortality and morbidity worldwide. Electrical defibrillation using high-energy electric shocks is currently the only treatment for life-threatening ventricular fibrillation. However, defibrillation may have side-effects, including intolerable pain, tissue damage, and worsening of prognosis, indicating a significant medical need for the development of more gentle cardiac rhythm management strategies. Besides energy-reducing electrical approaches, cardiac optogenetics was introduced as a powerful tool to influence cardiac activity using light-sensitive membrane ion channels and light pulses. In the present study, a robust and valid method for successful photostimulation of Langendorff perfused intact murine hearts will be described based on multi-site pacing applying a 3 x 3 array of micro light-emitting diodes (micro-LED). Simultaneous optical mapping of epicardial membrane voltage waves allows the investigation of the effects of region-specific stimulation and evaluates the newly induced cardiac activity directly on-site. The obtained results show that the efficacy of defibrillation is strongly dependent on the parameters chosen for photostimulation during a cardiac arrhythmia. It will be demonstrated that the illuminated area of the heart plays a crucial role for termination success as well as how the targeted control of cardiac activity during illumination for modifying arrhythmia patterns can be achieved. In summary, this technique provides a possibility to optimize the on-site mechanism manipulation on the way to real-time feedback control of cardiac rhythm and, regarding the region specificity, new approaches in reducing the potential harm to the cardiac system compared to the usage of non-specific electrical shock applications.

This work reports a method for controlling the cardiac rhythm of intact murine hearts of transgenic channelrhodopsin-2 (ChR2) mice using local photostimulation with a micro-LED array and simultaneous optical mapping of epicardial membrane potential.

Ventricular tachyarrhythmias are a major cause of mortality and morbidity worldwide. Electrical defibrillation using high-energy electric shocks is ...

Three-Dimensional Collagen Matrix Scaffold Implantation as a Liver Regeneration Strategy

Liver diseases are the leading cause of death worldwide. Excessive alcohol consumption, a high-fat diet, and hepatitis C virus infection promote fibrosis, cirrhosis, and/or hepatocellular carcinoma. Liver transplantation is the clinically recommended procedure to improve and extend the life span of patients in advanced disease stages. However, only 10% of transplants are successful, with organ availability, presurgical and postsurgical procedures, and elevated costs directly correlated with that result. Extracellular matrix (ECM) scaffolds have emerged as an alternative for tissue restoration. Biocompatibility and graft acceptance are the main beneficial characteristics of those biomaterials. Although the capacity to restore the size and correct function of the liver has been evaluated in liver hepatectomy models, the use of scaffolds or some kind of support to replace the volume of the extirpated liver mass has not been assessed.
Partial hepatectomy was performed in a rat liver with the xenoimplantation of a collagen matrix scaffold (CMS) from a bovine condyle. Left liver lobe tissue was removed (approximately 40%), and an equal proportion of CMS was surgically implanted. Liver function tests were evaluated before and after the surgical procedure. After days 3, 14, and 21, the animals were euthanized, and macroscopic and histologic evaluations were performed. On days 3 and 14, adipose tissue was observed surrounding the CMS, with no clinical evidence of rejection or infection, as was vessel neoformation and CMS reabsorption at day 21. There was histologic evidence of an insignificant inflammation process and migration of adjacent cells to the CMS, observed with the hematoxylin and eosin (H&#38;E) and Masson&#39;s trichrome staining. The CMS was shown to perform well in liver tissue and could be a useful alternative for studying tissue regeneration and repair in chronic liver diseases.

Liver diseases are induced by many causes that promote fibrosis or cirrhosis. Transplantation is the only option for recovering health. However, given the scarcity of transplantable organs, alternatives must be explored. Our research proposes the implantation of collagen scaffolds in liver tissue from an animal model.

Liver diseases are the leading cause of death worldwide. Excessive alcohol consumption, a high-fat diet, and hepatitis C virus infection promote fibrosis, ...

Refined CLARITY-Based Tissue Clearing for Three-Dimensional Fibroblast Organization in Healthy and Injured Mouse Hearts

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On-Site Sampling and Extraction of Brain Tumors for Metabolomics and Lipidomics Analysis

Advanced Cardiac Rhythm Management by Applying Optogenetic Multi-Site Photostimulation in Murine Hearts

Three-Dimensional Collagen Matrix Scaffold Implantation as a Liver Regeneration Strategy