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,&lt;br/&gt; N&#039;-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&amp;prime;,N&amp;prime;-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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Research

JoVE Journal

Medicine

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

Imaging Cleared Embryonic and Postnatal Hearts at Single-cell Resolution

Single clonal tracing and analysis at the whole-heart level can determine cardiac progenitor cell behavior and differentiation during cardiac development, and allow for the study of the cellular and molecular basis of normal and abnormal cardiac morphogenesis. Recent emerging technologies of retrospective single clonal analyses make the study of cardiac morphogenesis at single cell resolution feasible. However, tissue opacity and light scattering of the heart as imaging depth is increased hinder whole-heart imaging at single cell resolution. To overcome these obstacles, a whole-embryo clearing system that can render the heart highly transparent for both illumination and detection must be developed. Fortunately, in the last several years, many methodologies for whole-organism clearing systems such as CLARITY, Scale, SeeDB, ClearT, 3DISCO, CUBIC, DBE, BABB and PACT have been reported. This lab is interested in the cellular and molecular mechanisms of cardiac morphogenesis. Recently, we established single cell lineage tracing via the ROSA26-CreERT2; ROSA26-Confetti system to sparsely label cells during cardiac development. We adapted several whole embryo-clearing methodologies including Scale and CUBIC (clear, unobstructed brain imaging cocktails and computational analysis) to clear the embryo in combination with whole mount staining to image single clones inside the heart. The heart was successfully imaged at single cell resolution. We found that Scale can clear the embryonic heart, but cannot effectively clear the postnatal heart, while CUBIC can clear the postnatal heart, but damages the embryonic heart by dissolving the tissue. The methods described here will permit the study of gene function at a single clone resolution during cardiac morphogenesis, which, in turn, can reveal the cellular and molecular basis of congenital heart defects.

We describe a protocol to volumetrically image fluorescent protein labeled cells deep inside intact embryonic and postnatal hearts. Utilizing tissue-clearing methods in combination with whole mount staining, single fluorescent protein-labeled cells inside an embryonic or postnatal heart can be imaged clearly and accurately.

Single clonal tracing and analysis at the whole-heart level can determine cardiac progenitor cell behavior and differentiation during cardiac development, ...

Developmental Biology

Capturing the Cardiac Injury Response of Targeted Cell Populations via Cleared Heart Three-Dimensional Imaging

Cardiovascular disease outranks all other causes of death and is responsible for a staggering 31% of mortalities worldwide. This disease manifests in cardiac injury, primarily in the form of an acute myocardial infarction. With little resilience following injury, the once healthy cardiac tissue will be replaced by fibrous, non-contractile scar tissue and often be a prelude to heart failure. To identify novel treatment options in regenerative medicine, research has focused on vertebrates with innate regenerative capabilities. One such model organism is the neonatal mouse, which responds to cardiac injury with robust myocardial regeneration. In order to induce an injury in the neonatal mouse that is clinically relevant, we have developed a surgery to occlude the left anterior descending artery (LAD), mirroring a myocardial infarction triggered by atherosclerosis in the human heart. When matched with the technology to track changes both within cardiomyocytes and non-myocyte populations, this model provides us with a platform to identify the mechanisms that guide heart regeneration. Gaining insight into changes in cardiac cell populations following injury once relied heavily on methods such as tissue sectioning and histological examination, which are limited to two-dimensional analysis and often damage the tissue in the process. Moreover, these methods lack the ability to trace changes in cell lineages, instead providing merely a snapshot of the injury response. Here, we describe how technologically advanced methods in lineage tracing models, whole organ clearing, and three-dimensional (3D) whole-mount microscopy can be used to elucidate mechanisms of cardiac repair. With our protocol for neonatal mouse myocardial infarction surgery, tissue clearing, and 3D whole organ imaging, the complex pathways that induce cardiomyocyte proliferation can be unraveled, revealing novel therapeutic targets for cardiac regeneration.

Cardiomyocyte proliferation following injury is a dynamic process that requires a symphony of extracellular cues from non-myocyte cell populations. Utilizing lineage tracing, passive CLARITY, and three-dimensional whole-mount confocal microscopy techniques, we can analyze the influence of a variety of cell types on cardiac repair and regeneration.

Cardiovascular disease outranks all other causes of death and is responsible for a staggering 31% of mortalities worldwide. This disease manifests in ...

Minimal Invasive Surgical Procedure of Inducing Myocardial Infarction in Mice

Myocardial infarction still remains the main cause of death in western countries, despite considerable progress in the stent development area in the last decades. For clarification of the underlying mechanisms and the development of new therapeutic strategies, the availability of valid animal models are mandatory. Since we need new insights into pathomechanisms of cardiovascular diseases under in vivo conditions to combat myocardial infarction, the validity of the animal model is a crucial aspect. However, protection of animals are highly relevant in this context. Therefore, we establish a minimally invasive and simple model of myocardial infarction in mice, which assures a high reproducibility and survival rate of animals. Thus, this models fulfils the requirements of the 3R principle (Replacement, Refinement and Reduction) for animal experiments and assure the scientific information needed for further developing of therapeutical strategies for cardiovascular diseases.

A highly reproducible model for myocardial infarction in mice with minimal invasive manipulations is described. The model can be easily performed, resulting in a high reproducibility and survival rate. Thus, the described model will reduce the number of required animals as requested by the 3R principle (Replacement, Refinement and Reduction).

Myocardial infarction still remains the main cause of death in western countries, despite considerable progress in the stent development area in the last ...

Light-sheet Fluorescence Microscopy for the Study of the Murine Heart

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. Traditional light-sheet techniques involve the use of a single illumination beam directed orthogonally at sample tissue. Images of large samples that are produced using a single illumination beam contain stripes or artifacts and suffer from a reduced resolution due to the scattering and absorption of light by the tissue. This study uses a dual-sided illumination beam and a simplified CLARITY optical clearing technique for the murine heart. These techniques allow for deeper imaging by removing lipids from the heart and produce a large field of imaging, greater than 10 x 10 x 10 mm3. As a result, this strategy enables us to quantify the ventricular dimensions, track the cardiac lineage, and localize the spatial distribution of cardiac-specific proteins and ion-channels from the post-natal to adult mouse hearts with sufficient contrast and resolution.

This study uses a dual-sided illumination light-sheet fluorescence microscopy (LSFM) technique combined with optical clearing to study the murine heart.

Light-sheet fluorescence microscopy has been widely used for rapid image acquisition with a high axial resolution from micrometer to millimeter scale. ...

Bioengineering

Light-Sheet Microscopy and Tissue Clearing for Comprehensive Cardiac Imaging

Light-Sheet Imaging to Reveal Cardiac Structure in Rodent Hearts

Light-sheet microscopy (LSM) plays a pivotal role in comprehending the intricate three-dimensional (3D) structure of the heart, providing crucial insights into fundamental cardiac physiology and pathologic responses. We hereby delve into the development and implementation of the LSM technique to elucidate the micro-architecture of the heart in mouse models. The methodology integrates a customized LSM system with tissue clearing techniques, mitigating light scattering within cardiac tissues for volumetric imaging. The combination of conventional LSM with image stitching and multiview deconvolution approaches allows for the capture of the entire heart. To address the inherent trade-off between axial resolution and field of view (FOV), we further introduce an axially swept light-sheet microscopy (ASLM) method to minimize out-of-focus light and uniformly illuminate the heart across the propagation direction. In the meanwhile, tissue clearing methods such as iDISCO enhance light penetration, facilitating the visualization of deep structures and ensuring a comprehensive examination of the myocardium throughout the entire heart. The combination of the proposed LSM and tissue clearing methods presents a promising platform for researchers in resolving cardiac structures in rodent hearts, holding great potential for the understanding of cardiac morphogenesis and remodeling.

Author Spotlight: Customized Light-Sheet Imaging for Investigating Myocardial Structures in Rodent Hearts

The protocol utilizes advanced light-sheet microscopy along with adapted tissue clearing methods to investigate intricate cardiac structures in rodent hearts, holding great potential for the understanding of cardiac morphogenesis and remodeling.

Light-sheet microscopy (LSM) plays a pivotal role in comprehending the intricate three-dimensional (3D) structure of the heart, providing crucial insights ...

Ultrasound-Guided Induced Pluripotent Stem Cell-Derived Cardiomyocyte Implantation in Myocardial Infarcted Mice

The key objective of cell therapy after myocardial infarction (MI) is to effectively enhance the cell grafted rate, and human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are a promising cell source for cardiac repair after ischemic damage. However, a low grafted rate is a significant obstacle for effective cardiac tissue regeneration after transplantation. This protocol shows that multiple hiPSC-CM ultrasound-guided percutaneous injections into an MI area effectively increase cell transplantation rates. The study also describes the entire hiPSC-CM culture process, pretreatment, and ultrasound-guided percutaneous delivery methods. In addition, the use of human mitochondrial DNA help detect the absence of hiPSC-CMs in other mouse organs. Lastly, this paper describes the changes in cardiac function, angiogenesis, cell size, and apoptosis at the infarcted border zone in mice 4 weeks after cell delivery. It can be concluded that echocardiography-guided percutaneous injection of the left ventricular myocardium is a feasible, relatively invasive, satisfactory, repeatable, and effective cellular therapy.

Ultrasound-guided cell delivery around the site of myocardial infarction in mice is a safe, effective, and convenient way of cell transplantation.

The key objective of cell therapy after myocardial infarction (MI) is to effectively enhance the cell grafted rate, and human induced pluripotent stem ...

Isolation and Characterization of Adult Cardiac Fibroblasts and Myofibroblasts

Cardiac fibrosis in response to injury is a physiological response to wound healing. Efforts have been made to study and target fibroblast subtypes that mitigate fibrosis. However, fibroblast research has been hindered due to the lack of universally acceptable fibroblast markers to identify quiescent as well as activated fibroblasts. Fibroblasts are a heterogenous cell population, making them difficult to isolate and characterize. The presented protocol describes three different methods to enrich fibroblasts and myofibroblasts from uninjured and injured mouse hearts. Using a standard and reliable protocol to isolate fibroblasts will enable the study of their roles in homeostasis as well as fibrosis modulation.

Obtaining a pure population of fibroblasts is crucial to studying their role in wound repair and fibrosis. Described here is a detailed method to isolate fibroblasts and myofibroblasts from uninjured and injured mouse hearts followed by characterization of their purity and functionality by immunofluorescence, RTPCR, fluorescence-assisted cell sorting, and collagen gel contraction.

Cardiac fibrosis in response to injury is a physiological response to wound healing. Efforts have been made to study and target fibroblast subtypes that ...

Biology

An Antegrade Perfusion Method for Cardiomyocyte Isolation from Mice

In basic research using mouse heart, isolating viable individual cardiomyocytes is a crucial technical step to overcome. Traditionally, isolating cardiomyocytes from rabbits, guinea pigs or rats has been performed via retrograde perfusion of the heart with enzymes using a Langendorff apparatus. However, a high degree of skill is required when this method is used with a small mouse heart. An antegrade perfusion method that does not use a Langendorff apparatus was recently reported for the isolation of mouse cardiomyocytes. We herein report a complete protocol for the improved antegrade perfusion of the excised heart to isolate individual heart cells from adult mice (8 - 108 weeks old). Antegrade perfusion is performed by injecting perfusate near the apex of the left ventricle of the excised heart, the aorta of which was clamped, using an infusion pump. All procedures are carried out on a pre-warmed heater mat under a microscope, which allows for the injection and perfusion processes to be monitored. The results suggest that ventricular and atrial myocytes, and fibroblasts can be well isolated from a single adult mouse simultaneously.

We developed a simple method for isolating high quality individual mouse heart cells by the antegrade perfusion technique. This method is Langendorff-free and useful for isolating ventricular and atrial myocytes or interstitial cells, such as cardiac fibroblasts or progenitors.

In basic research using mouse heart, isolating viable individual cardiomyocytes is a crucial technical step to overcome. Traditionally, isolating ...

Mesoscopic Optical Imaging of Whole Mouse Heart

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition (fibrosis) and cellular misalignment, can affect electrical conduction, introduce electromechanical dysfunctions and, eventually, lead to arrhythmia. Current predictive models of these functional alterations are based on non-integrated and low-resolution structural information. Placing this framework on a different order of magnitude is challenging due to the inefficacy of standard imaging methods in performing high-resolution imaging in massive tissue. In this work, we describe a methodological framework that allows imaging of whole mouse hearts with micrometric resolution. The achievement of this goal has required a technological effort where advances in tissue transformation and imaging methods have been combined. First, we describe an optimized CLARITY protocol capable of transforming an intact heart into a nanoporous, hydrogel-hybridized, lipid-free form that allows high transparency and deep staining. Then, a fluorescence light-sheet microscope able to rapidly acquire images of a mesoscopic field of view (mm-scale) with the micron-scale resolution is described. Following the mesoSPIM project, the conceived microscope allows the reconstruction of the whole mouse heart with micrometric resolution in a single tomographic scan. We believe that this methodological framework will allow clarifying the involvement of the cytoarchitecture disarray in the electrical dysfunctions and pave the way for a comprehensive model that considers both the functional and structural data, thus enabling a unified investigation of the structural causes that lead to electrical and mechanical alterations after the tissue remodeling.

We report a method for mesoscopic reconstruction of the whole mouse heart by combining new advancements in tissue transformation and staining with the development of an axially scanned light-sheet microscope.

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition ...

Optimized Tissue-Clearing Protocols for 3D Imaging of the Murine Aorta 

Three-Dimensional Imaging of Aortic Tissues in Atherosclerosis

Recent research has advanced the understanding of atherosclerosis as a transmural chronic inflammatory disease involving all three layers of the arterial wall, including the intima plaque, the media, and the adventitia, which forms the outer connective tissue coat of arteries. Our recent studies have suggested that the adventitia is used by the peripheral nervous system as a conduit for reaching all tissue cells. We also found that the peripheral nervous system, that is, the sensory and sympathetic nervous system, undergoes major remodeling processes involving the neogenesis of axon networks adjacent to atherosclerotic plaques. In this context, understanding the structure of the neural network and its interactions with vascular components of diseased arteries holds major promises for a better understanding of cardiovascular disease pathogenesis. To achieve these objectives, methods to visualize the subcellular architecture of the intact healthy and diseased arteries together with their surrounding perivascular compartments are needed. Tissue clearing allows intact deep-tissue imaging of larger tissue compartments that are otherwise inaccessible. It allows volumetric imaging of intact arteries through the integration of labeling, clearing, advanced microscopic imaging, and image processing tools. Here, we describe two distinct but complementary passive tissue clearing approaches, that is, aqueous-based 2, 2-thiodiethanol (TDE) clearing and solvent-based immunolabeling-enabled three-dimensional imaging of solvent-cleared organ (iDISCO) clearing to image isolated aortic segments or whole aorta in-situ in the whole mouse.

Here, we present optimized tissue-clearing protocols to image the murine aorta in three dimensions (3D). We delineate state-of-the-art procedures for immunostaining, optical clearing, and imaging with the intent to define the anatomical proximity of the peripheral nervous system with atherosclerotic plaques and the adventitia in atherosclerosis.

Recent research has advanced the understanding of atherosclerosis as a transmural chronic inflammatory disease involving all three layers of the arterial ...