Funding for this project was provided by the UW School of Medicine and Public Health from the Wisconsin Partnership Program (A.I.M.), and an American Heart Association Career Development Award 19CDA34660169 (A.I.M.).

All experiments were conducted in accordance with the Guide for the Use and Care of Laboratory Animals and in compliance with the Institutional Animal Care and Use Committee in the School of Medicine and Public Health at the University of Wisconsin&#8211;Madison. All methods were performed on wild type C57BL/6J (B6) and transgenic mouse lines obtained from Jackson Laboratories.
1. Coronary Artery Occlusion (Myocardial Infarction) Induced via Ligation of the Left Anterior Descending Artery (LAD) in 1-Day-Old Neonatal Mice6
<ol>
	<li>Separate the 1-day-old neonatal pups from the mother by placing them into a clean cage along with some of the original nesting material.</li>
	<li>Place half of the cage onto a heating pad set to medium heat. The pups should remain on the unheated side of the cage, only being placed onto the heated side after surgery.</li>
	<li>Create a sterile surgical area under an operating microscope. Gather sterilized surgical equipment (Table 1).</li>
	<li>Anesthetize the pup via hypothermia: wrap the pup in gauze to avoid direct skin contact with ice and bury it in an ice bed for approximately 3&#8211;4 min. Check hypothermia of the mice periodically by performing a toe pinch. Neonates tolerate hypothermia well, however, prolonged exposure to hypothermia may result in frostbite and subsequent mortality.</li>
	<li>Once anesthetized, place the mouse onto the surgical area in the supine position, securing the arms and legs with tape. Sterilize the surgical area of the mouse with an antiseptic solution.</li>
	<li>Locate the lower chest region and make a transverse incision in the skin with small dissecting scissors. To widen the surgical view of the ribs, separate the skin from the muscle by lifting the skin gently with a pair of dressing forceps and gently press against the intercostal muscles with the small scissors in the closed position.</li>
	<li>Locate the fourth intercostal space (Figure 1A) and make a small, superficial puncture using sharp forceps, being careful not to puncture any internal organs. Perform a blunt dissection by widening the area in between the intercostal muscles with dressing forceps. Proper anatomical positioning of the incision is essential for appropriate access to the heart.</li>
	<li>Gently guide the heart out of the chest cavity by placing a finger and applying increasing pressure on the left side of the abdomen while holding the intercostal space open with dressing forceps (Figure 1B). Once the heart is outside of the chest, remove the dressing forceps, relieve pressure, and allow the heart to rest on the intercostal muscles.</li>
	<li>Locate the LAD as the area of the heart that has less pooled blood and is in the correct anatomical position (Figure 1C). The LAD can be only seen under the microscope if the heart is accessed within a few minutes of beginning surgery.</li>
	<li>Perform LAD ligation by suturing the LAD with a 6-0 suture (Figure 1D). Tie a square knot twice to induce myocardial infarction (Figure 1E). The depth of the suture into the myocardium may vary, however, proper anatomical positioning of the LAD ligation is crucial for reproducibility. When ligating the LAD, the suture should be pulled tightly but carefully so as not to sever the LAD. Blanching at the apex of the heart will be seen immediately (Figure 1E)</li>
	<li>Allow the heart to slip back into the chest cavity; this process can be gently facilitated with dressing forceps. Suture the ribs together with a surgeon&#39;s knot followed by a square knot, using blunt forceps to lift the upper set of ribs while passing a 6-0 suture through the upper and lower set of ribs.</li>
	<li>Remove the tape that was used to secure the hind legs of the pup.</li>
	<li>Adhere the skin together by placing a small amount of skin glue on the upper abdomen. Then, grab the skin of the lower abdomen with fine forceps and cover the exposed chest region. Limit the amount of excess glue that remains on the pups, as this can increase the likelihood of rejection and cannibalism by the mother.</li>
	<li>Immediately facilitate the recovery from anesthesia by placing the pup onto a heating pad set to medium heat. Periodically switch the placement of the neonates to evenly warm all parts of the body.</li>
	<li>Allow the neonate to remain directly on the heating pad for 10&#8211;15 min. Typically, movement is regained within 5 min of being placed onto heat.</li>
	<li>Clean the residual blood and glue with an alcohol wipe.</li>
	<li>Cover foreign scents on neonates by rubbing the entire body with bedding from the original cage. Place the pup into the cage on the heated side while other surgeries are being performed.</li>
	<li>Once all surgeries are completed and pups are warm and mobile, transfer the litter along with the original nesting material to the mother&#39;s cage.</li>
	<li>Monitor the mice for 30&#8211;60 min after surgery and watch for the mother&#39;s acceptance of the pups by nesting and/or grooming.</li>
	<li>Check on the mice the morning following surgery. If mother is distressed and has not nested the pups, consider a foster mother for the pups.</li>
</ol>
2. Clearing the Mouse Heart with Passive CLARITY21,22,23
<ol>
	<li>Anesthetize the mouse with isoflurane. Perform a toe pinch to ensure the mouse is fully sedated.</li>
	<li>Place the mouse onto a clean, surgical area in the supine position, securing the arms and legs with tape.</li>
	<li>Maintain isoflurane sedation on the mouse using a nose cone until the heart is extracted.</li>
	<li>Open the lower chest by holding the fur just below the xiphoid process with tissue forceps and make an incision spanning the width of the ribcage using the large dissecting scissors.</li>
	<li>Cut alongside of the distal portions of the rib cage with surgical scissors.</li>
	<li>Expose the diaphragm muscle by grasping the xiphoid process with tissue forceps. Detach the diaphragm using curved forceps.</li>
	<li>While maintaining a grasp of the xiphoid process, pull the tissue cranially until the beating heart is accessible.</li>
	<li>Grasp the heart at the base with curved forceps and dissect the heart from the chest cavity by cutting the aorta and superior vena cava with iridectomy scissors.</li>
	<li>While the heart is still beating, place the heart into a Petri dish filled with PBS so that the heart pumps out the blood inside as it keeps beating. Myocardial infarction can be confirmed by checking that the placement of the suture is in the proper anatomical position for LAD ligation.</li>
	<li>Gently squeeze the heart with forceps to allow the heart to expel the residual blood.</li>
	<li>Transfer the mouse heart into a disposable 2.5 mL glass shell vial with 2 mL of PBS. Wash away residual blood by incubating the heart on a shaker for 10 min at room temperature (RT) several times. Change the PBS solution each time until PBS remains clear.</li>
	<li>Discard the PBS and fill the vial with 2 mL of cold 4% paraformaldehyde (PFA). Incubate for 4 hours at RT (Figure 2A).</li>
	<li>After incubation, discard the PFA and the vial with 2 mL of PBS. Wash the heart on a shaker for 10 min at RT. Repeat the washing step twice, draining and filling the vial with new PBS each time to fully wash away excess PFA.</li>
	<li>Discard PBS and fill the vial with 2 mL of 4% acrylamide and 0.5% VA-044 solution. Incubate overnight at 4 &#176;C.</li>
	<li>The next day, perform polymerization by transferring the vial to a heat block set at 37 &#176;C for 3 hours.</li>
	<li>Transfer the heart into a new glass shell vial and repeat step 2.12 (PBS wash cycle).</li>
	<li>Discard PBS and fill the vial with 2 mL of Clearing Solution (Table 2). Incubate at 37 &#176;C until the heart is cleared. Change out the solution and refill with fresh Clearing Solution every 2&#8211;3 days. The clearing process could take up to several weeks (Figure 2B-C). 
	NOTE: P1 hearts typically take around 3&#8211;5 days, whereas P21 hearts can take nearly a month before Clearing Solution incubation is complete.</li>
	<li>Discard PBS and fill the vial with 2 mL of PBS and repeat step 2.12 (PBS wash cycle). Refill the vial with fresh PBS and incubate overnight at 37 &#176;C.</li>
	<li>If immunostaining will be performed, skip steps 2.21&#8211;2.22 and proceed to Section 3 for immunostaining. If relying solely on endogenous fluorescence, proceed with steps 2.21&#8211;2.22 and skip Section 3.</li>
	<li>Discard PBS and change the solution to Refractive Index Matching Solution (RIMS) (Table 3). Incubate overnight at 37 &#176;C.</li>
	<li>After incubation, the cleared tissue can be stored in RIMS solution at RT. Tissue may appear to be white and opaque in the center when first transferred into RIMS; tissue should become transparent after being incubated in RIMS at room temperature for several weeks (Figure 2D).</li>
</ol>
3. Optional: Immunohistochemistry Staining of the Whole-Mount Mouse Heart
<ol>
	<li>Remove the cleared heart from the RIMS solution and place into a clean 2.5 mL glass vial with 2 mL of PBST (PBS with 0.1% Triton-X 100)</li>
	<li>Wash the heart in PBST 3 times on an orbital rotator with 30 min incubations at RT.</li>
	<li>Block non-specific antibody binding by immersing the heart in 2 mL of 20% blocking buffer (diluted in PBST) and incubate with rotation for 3 hours at RT. Transfer to 4 &#176;C to stain with rotation overnight. 
	NOTE: Blocking buffer is made from normal serum matching the species in which the secondary antibody was raised.</li>
	<li>Wash the heart in PBST 3 times with rotation for 5 min incubations at RT.</li>
	<li>Immerse the heart in primary antibody (diluted in 2% blocking buffer with PBST) and prevent light exposure by wrapping the glass vial in aluminum foil. Incubate with rotation overnight at RT. 
	NOTE: From this point forward, aluminum foil should be continuously used to protect the secondary from ambient light exposure.</li>
	<li>Incubate for an additional 24 hours with rotation at 4 &#176;C.</li>
	<li>Following primary staining, repeat step 3.2 (long PBST wash cycle) to remove the unbound primary antibody from the tissue. Extend the wash with an overnight incubation with rotation at RT.</li>
	<li>Working in an area with limited lighting to prevent secondary antibody light exposure, immerse the heart in secondary antibody (diluted in 2% blocking buffer with PBST) and incubate with rotation for 3 hours at RT. Transfer to 4 &#176;C to stain with rotation overnight.</li>
	<li>On the next day, repeat step 3.2 (long PBST wash cycle) to remove unbound secondary antibody.</li>
	<li>Replace the solution with 2% blocking buffer (diluted in PBST). Remove residual unbound antibody by washing overnight with rotation at RT.</li>
	<li>The next day, check that excess secondary antibody has been removed using confocal microscopy. Extend the wash as needed, replacing the 2% blocking buffer solution daily. Proceed once little to no non-specific secondary is present.</li>
	<li>Store the immunostained heart in PBS at 4 &#176;C.</li>
	<li>Directly before whole-mount microscopy, incubate the heart in RIMS solution overnight at 37 &#176;C. Extend the incubation an additional 24 hours if the tissue is still not fully cleared.</li>
	<li>Store the fully cleared and immunostained heart in RIMS at RT.</li>
</ol>
4. Visualizing Non-myocyte Populations in 3D with Single-Photon Confocal Microscopy Imaging of the Cleared Mouse Heart
NOTE: If mouse hearts are harvested embryonically, continue with section 4.1. For mouse hearts harvested postnatally, continue with section 4.2.
<ol>
	<li>Imaging the Cleared Embryonic Mouse Heart
	<ol>
		<li>Fill the microscope depression slide with the PBS solution.</li>
		<li>Carefully pick up the cleared heart with curved forceps and place the tissue onto the slide.</li>
		<li>Mount the slide with a glass coverslip. The tissue can now be imaged under a confocal microscope.</li>
	</ol>
	</li>
	<li>Imaging the Cleared Postnatal Mouse Heart
	<ol>
		<li>Fill half of the chamber of the depression slide with PBS solution. In order to create a chamber large enough to fit an adult mouse heart, a 3D-printed polypropylene depression slide was custom-made (Figure 4).</li>
		<li>Carefully pick up the cleared heart with curved forceps and place the tissue into the chamber. Fill the remaining volume of the chamber with PBS.</li>
		<li>Fill the chamber with PBS so the surface of the liquid forms a dome above the top of the chamber. Mount the cover slide. The tissue can now be imaged under an upright confocal microscope.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

Cell-cell interactions between cardiomyocytes and non-myocyte populations are a determining factor of whether the heart will undergo fibrosis or repair following injury. Discoveries have been made demonstrating that a variety of cell types, including nerves14, epicardial cells24, peritoneal macrophages25, arterioles12,13, and lymphatic endothelial cells26, all play an ...

The heart has long been considered to be a post-mitotic organ, yet recent evidence demonstrates that cardiomyocyte renewal occurs in the adult human heart at about 1% per year1. However, these low rates of cardiomyocyte turnover are insufficient to replenish the massive loss of tissue that occurs following injury. A heart that has suffered a myocardial infarction will lose around one billion cardiomyocytes, often serving as a prelude to heart failure and sudden cardiac death2,3. With over 26 million people affected by heart failure worldwide, there is an unmet need for therapeutics that can reverse the damages inflicted by heart disease4.
In order to bridge this gap in therapeutics, scientists have begun investigating evolutionarily conserved mechanisms that underlie endogenous regeneration following injury. One model for studying mammalian cardiac regeneration is the neonatal mouse. Within the week following birth, neonatal mice have a robust regenerative response following cardiac damage5. We have previously demonstrated that neonatal mice can regenerate their heart via cardiomyocyte proliferation following an apical resection5. Although this technique can evoke cardiac regeneration in the neonates, the surgery lacks clinical relevance to human heart injuries. In order to mimic a human injury in the neonatal mouse model, we have developed a technique to induce a myocardial infarction through a coronary artery occlusion6. This technique requires surgical ligation of the left anterior descending artery (LAD), which is responsible for delivering 40%&#8211;50% of the blood to the left ventricular myocardium6,7. Thus, the surgery results in an infarct that impacts a significant portion of the left ventricular wall. This damage to the myocardium will stimulate cardiomyocyte proliferation and heart regeneration in neonates5.
The coronary artery occlusion surgery provides a highly reproducible and directly translational method to uncover the inner workings of cardiac regeneration. The neonatal surgery parallels coronary artery atherosclerosis in the human heart, where accumulation of plaque within the inner walls of the arteries can cause an occlusion and subsequent myocardial infarction8. Due to a void in therapeutic treatments for heart failure patients, an occlusion in the LAD is associated with mortality rates reaching up to 26% within a year following injury9, and consequently has been termed the &#34;widow maker.&#34; Advancements in therapeutics require a model that accurately reflects the complex physiological and pathological effects of cardiac injury. Our surgical protocol for neonatal mouse cardiac injury provides a platform that allows researchers to investigate the molecular and cellular cues that signal mammalian heart regeneration after injury.
Recent research highlights the dynamic relationship between the extracellular environment and proliferating cardiomyocytes. For example, the postnatal regenerative window can be extended by decreasing the stiffness of the extracellular matrix surrounding the heart10. Biomaterials from the neonatal extracellular matrix can also promote heart regeneration in adult mammalian hearts following cardiac injury11. Also accompanying cardiomyocyte proliferation is an angiogenic response12,13; collateral artery formation unique to the regenerating heart of the neonatal mouse was shown to be essential for stimulating cardiac regeneration12. Moreover, our lab has demonstrated that nerve signaling regulates cardiomyocyte proliferation and heart regeneration via modulation of growth factor levels, as well as the inflammatory response following injury14. These findings emphasize the need to trace non-myocyte cell populations in response to cardiac injury. In order to accomplish this goal, we have taken advantage of the Cre-lox recombination system in transgenic mice lines to incorporate constitutive or conditional expression of fluorescent reporter proteins for lineage tracing. Furthermore, we can use advanced methods to determine clonal expansion patterning with the Rainbow mouse line, which relies on stochastic expression of the Cre-dependent, multi-color fluorescent reporters to determine the clonal expansion of targeted cell populations15. Employing lineage tracing with the neonatal coronary artery occlusion surgery is a powerful tool for dissecting the intricate cellular mechanisms of cardiac regeneration.
Tracking the lineage of fluorescently labeled cells with three-dimensional (3D) whole organ imaging is difficult to achieve using traditional sectioning and reconstruction technique &#8211; especially when cell populations are fragile, such as nerve fibers or blood vessels. While direct whole-mount imaging of the organ by optical sectioning can capture superficial cell populations, structures that reside deep within the tissue remain inaccessible. To circumvent these barriers, tissue clearing techniques have been developed to reduce the opacity of whole organ tissues. Recently, significant advances have been made to Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging compatible Tissue hYdrogel (CLARITY)&#8211;based methods, which clear fixed tissue via lipid extraction16. Steps are also taken to homogenize the refractive index and subsequently reduce light scattering while imaging17. One such method is active CLARITY, which expedites lipid decomposition by using electrophoresis to penetrate the detergent throughout the tissue18. Although effective, this tissue clearing method requires expensive equipment and can cause tissue damage, making the approach incompatible with fragile cell populations such as the cardiac nerves19. Thus, we employ the passive CLARITY approach, which relies on heat to gently facilitate detergent penetration, therefore aiding in the retention of intricate cell structures20,21.
Passive CLARITY is typically thought to be less efficient than active CLARITY18, as the technique is often accompanied by two major obstacles: the inability to clear the entire organ depth and the extensive amount of time required to clear adult tissues. Our passive CLARITY approach overcomes both of these barriers with an expeditated clearing process that is capable of fully clearing neonatal and adult heart tissue. Our passive CLARITY tissue clearing technique has reached an efficiency that permits the visualization of a variety of cardiac cell populations, including rare populations distributed throughout the adult heart. When the cleared heart is imaged with confocal microscopy, the architecture of cell-specific patterning during development, disease, and regeneration can be illuminated.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1-thioglycerol</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>6-0 Prolene Sutures</td>
			<td>Ethicon</td>
			<td>8889H</td>
			<td>Polypropylene Sutures</td>
		</tr>
		<tr>
			<td>Acrylamide</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Boric acid</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Curved Forceps</td>
			<td>Excelta</td>
			<td>16-050-146</td>
			<td>Half Curved, Serrated, 4 in</td>
		</tr>
		<tr>
			<td>Dressing Forceps</td>
			<td>Fisherbrand</td>
			<td>13-812-39</td>
			<td>Dissecting, 4.5 in</td>
		</tr>
		<tr>
			<td>Glass Vial</td>
			<td>Fisherbrand</td>
			<td>03-339-26A</td>
			<td>12 x 35 mm Vial with Cap</td>
		</tr>
		<tr>
			<td>Histodenz</td>
			<td>Sigma-Aldrich</td>
			<td></td>
			<td>Density gradient medium</td>
		</tr>
		<tr>
			<td>Iridectomy Scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-03</td>
			<td>2 mm Cutting Edge</td>
		</tr>
		<tr>
			<td>Large Dissecting Scissors</td>
			<td>Fisherbrand</td>
			<td>08-951-20</td>
			<td>Straight, 6 in</td>
		</tr>
		<tr>
			<td>Needle Holder</td>
			<td>Fisherbrand</td>
			<td>08-966</td>
			<td>Mayo-Hegar, 6 in</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sharp Forceps</td>
			<td>Sigma-Adrich</td>
			<td>Z168777</td>
			<td>Fine Tip, Straight, 4.25 in</td>
		</tr>
		<tr>
			<td>Small Dissecting Scissor</td>
			<td>Walter Stern Inc</td>
			<td>25870-002</td>
			<td>30 mm Cutting Edge</td>
		</tr>
		<tr>
			<td>Sodium Azide</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Dodecyl Sulfate (SDS)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Forceps</td>
			<td>Excelta</td>
			<td>16050133</td>
			<td>Medium Tissue, 1X2 Teeth</td>
		</tr>
		<tr>
			<td>VA-044</td>
			<td>Wako Chemicals</td>
			<td></td>
			<td>Water-soluble azo initiator</td>
		</tr>
	</tbody>
</table>

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.

Often the two most challenging steps are guiding the heart out of the chest cavity and ligating the LAD. To troubleshoot these steps, adjustments may be made in the placement of the initial puncture between the fourth intercostal muscles; if the puncture and blunt dissection are too close in proximity to the sternum, the heart may not be able to exit the chest cavity (Figure 1A).
Additionally, increased pressure on the left abdomen may be needed t...

Watch this Scientific Journal Video about Capturing the Cardiac Injury Response of Targeted Cell Populations via Cleared Heart Three-Dimensional Imaging at JoVE.com

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

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 ...

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Research

JoVE Journal

Developmental Biology

5.8K Views.  University of Wisconsin-Madison School of Medicine and Public Health. 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.

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

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila melanogaster, inductive microenvironments known as larval Hematopoietic Pockets (HPs) have been identified as anatomical sites for the development and regulation of blood cells (hemocytes), in particular of the self-renewing macrophage lineage.&#160;HPs are segmentally repeated pockets between the epidermis and muscle layers of the larva, which also comprise sensory neurons of the peripheral nervous system. In the larva, resident (sessile) hemocytes are exposed to anti-apoptotic, adhesive and proliferative cues from these sensory neurons and potentially other components of the HPs, such as the lining muscle and epithelial layers. During normal development, gradual release of resident hemocytes from the HPs fuels the population of circulating hemocytes, which culminates in the release of most of the resident hemocytes at the beginning of metamorphosis. Immune assaults, physical injury or mechanical disturbance trigger the premature release of resident hemocytes into circulation. The switch of larval hemocytes between resident locations and circulation raises the need for a common standard/procedure to selectively isolate and quantify these two populations of blood cells from single Drosophila larvae. Accordingly, this protocol describes an automated method to release and quantify the resident and circulating hemocytes from single larvae. The method facilitates ex vivo approaches, and may be adapted to serve a variety of developmental stages of Drosophila and other invertebrate organisms.

Assaying Blood Cell Populations of the Drosophila melanogaster Larva

Drosophila blood cells, or hemocytes, cycle between resident sites and circulation. In the larva, resident (sessile) hemocytes localize to inductive microenvironments, the Hematopoietic Pockets, while circulating hemocytes move freely in the hemolymph. The goal of this protocol is the standardized isolation and quantification of these two, behaviorally distinct but interchanging, hemocyte populations.

In vertebrates, hematopoiesis is regulated by inductive microenvironments (niches). Likewise, in the invertebrate model organism&#160;Drosophila ...

Isolation of Perivascular Multipotent Precursor Cell Populations from Human Cardiac Tissue

Multipotent mesenchymal stem/stromal cells (MSC) were conventionally isolated, through their plastic adherence, from primary tissue digests whilst their anatomical tissue location remained unclear. The recent discovery of defined perivascular and MSC cell marker expression by perivascular cells in multiple tissues by our group and other researchers has provided an opportunity to prospectively isolate and purify specific homogenous subpopulations of multipotent perivascular precursor cells. We have previously demonstrated the use of fluorescent activated cell sorting (FACS) to purify microvascular CD146+CD34- pericytes and vascular CD34+CD146- adventitial cells from human skeletal muscle. Herein we describe a method to simultaneously isolate these two perivascular cell subsets from human myocardium by FACS, based on the expression of a defined set of cell surface markers for positive and negative selections. This method thus makes available two specific subpopulations of multipotent cardiac MSC-like precursor cells for use in basic research and/or therapeutic investigations.

Human cardiac tissue harbours multipotent perivascular precursor cell populations that may be suitable for myocardial regeneration. The technique described here allows for the simultaneous isolation and purification of two multipotent stromal cell populations associated with native blood vessels, i.e. CD146+CD34- pericytes and CD34+CD146- adventitial cells, from the human myocardium.

Multipotent mesenchymal stem/stromal cells (MSC) were conventionally isolated, through their plastic adherence, from primary tissue digests whilst their ...

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, ...

Horizontal Whole Mount: A Novel Processing and Imaging Protocol for Thick, Three-dimensional Tissue Cross-sections of Skin

Processing a tissue of interest to generate a microscopic image that supports a scientific argument can be challenging. The acquisition of high-quality microscopic images is not entirely dependent upon the quality of the microscope, but also upon the methods of tissue processing, which often involve multiple critical actions or steps. Furthermore, mesenchymal cell types in the skin and other tissues represent a new challenge for tissue preparation and imaging. Here, we present a complete process, from tissue harvest to microscopy. Our technique, called &#34;horizontal whole mount,&#34; is one that novices can quickly become proficient in and that allows for antigen preservation and detection in 60-300 &#181;m-thick sections cut with a cryostat. Sections of this thickness provide enhanced visualization of tissue microarchitecture in a three-dimensional environment. In addition, the protocol preserves mesenchymal cells in a manner that enhances image quality when compared to standard cryostat or paraffin sections, thereby increasing the efficacy and reliability of immunostaining. We believe that this protocol will benefit all laboratories that visualize skin, and possibly other tissues and organs.

This work presents a novel processing and imaging protocol for thick, three-dimensional tissue cross-section analysis that enables the full exploitation of confocal imaging modalities. This protocol preserves antigenicity and represents a robust system to analyze skin histology and potentially other tissue types.

Processing a tissue of interest to generate a microscopic image that supports a scientific argument can be challenging. The acquisition of high-quality ...

Quantitative Whole-mount Immunofluorescence Analysis of Cardiac Progenitor Populations in Mouse Embryos

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development and organogenesis are two areas of research that have benefitted greatly from these advances, due to the high quality of data that can be obtained directly from imaging pre-implantation embryos or ex vivo organs. While pre-implantation embryos have yielded data with especially high spatial resolution, later stages have been less amenable to three-dimensional reconstruction. Obtaining high-quality 3D or volumetric data for known embryonic structures in combination with fate mapping or genetic lineage tracing will allow for a more comprehensive analysis of the morphogenetic events taking place during embryogenesis.
This protocol describes a whole-mount immunofluorescence approach that allows for the labeling, visualization, and quantification of progenitor cell populations within the developing cardiac crescent, a key structure formed during heart development. The approach is designed in such a way that both cell- and tissue-level information can be obtained. Using confocal microscopy and image processing, this protocol allows for three-dimensional spatial reconstruction of the cardiac crescent, thereby providing the ability to analyze the localization and organization of specific progenitor populations during this critical phase of heart development. Importantly, the use of reference antibodies allows for successive masking of the cardiac crescent and subsequent quantitative measurements of areas within the crescent. This protocol will not only enable a detailed examination of early heart development, but with adaptations should be applicable to most organ systems in the gastrula to early somite stage mouse embryo.

Here, we describe a protocol for whole mount immunofluorescence and image-based quantitative volumetric analysis of early stage mouse embryos. We present this technique as a powerful approach to qualitatively and quantitatively assess cardiac structures during development, and propose that it may be widely adaptable to other organ systems.

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development ...

Three-dimensional Rendering and Analysis of Immunolabeled, Clarified Human Placental Villous Vascular Networks

Nutrient and gas exchange between mother and fetus occurs at the interface of the maternal intervillous blood and the vast villous capillary network that makes up much of the parenchyma of the human placenta. The distal villous capillary network is the terminus of the fetal blood supply after several generations of branching of vessels extending out from the umbilical cord. This network has a contiguous cellular sheath, the syncytial trophoblast barrier layer, which prevents mixing of fetal blood and the maternal blood in which it is continuously bathed. Insults to the integrity of the placental capillary network, occurring in disorders such as maternal diabetes, hypertension and obesity, have consequences that present serious health risks for the fetus, infant, and adult. To better define the structural effects of these insults, a protocol was developed for this study that captures capillary network structure on the order of 1 - 2 mm3 wherein one can investigate its topological features in its full complexity. To accomplish this, clusters of terminal villi from placenta are dissected, and the trophoblast layer and the capillary endothelia are immunolabeled. These samples are then clarified with a new tissue clearing process which makes it possible to acquire confocal image stacks to z- depths of ~1 mm. The three-dimensional renderings of these stacks are then processed and analyzed to generate basic capillary network measures such as volume, number of capillary branches, and capillary branch end points, as validation of the suitability of this approach for capillary network characterization.

This study presents a protocol for the reversible tissue clearing, immunostaining, 3D-rendering and analysis of vascular networks in human placenta villi samples on the order of 1 - 2 mm3.

Nutrient and gas exchange between mother and fetus occurs at the interface of the maternal intervillous blood and the vast villous capillary network that ...

Three-dimensional Inflammatory Human Tissue Equivalents of Gingiva

Periodontal diseases (such as gingivitis and periodontitis) are the leading causes of tooth loss in adults. Inflammation in gingiva is the fundamental physiopathology of periodontal diseases. Current experimental models of periodontal diseases have been established in various types of animals. However, the physiopathology of animal models is different from that of humans, making it difficult to analyze cellular and molecular mechanisms and evaluate new medicines for periodontal diseases. Here, we present a detailed protocol for reconstructing human inflammatory tissue equivalents of gingiva (iGTE) in vitro. We first build human tissue equivalents of gingiva (GTE) by utilizing two types of human cells, including human gingival fibroblasts (HGF) and human skin epidermal keratinocytes (HaCaT), under three-dimensional conditions. We create a wound model by using a tissue puncher to punch a hole in the GTE. Next, human THP-1 monocytes mixed with collagen gel are injected into the hole in the GTE. By adimistration of 10 ng/mL phorbol 12-myristate 13-acetate (PMA) for 72 h, THP-1 cells differentiated into macrophages to form inflammatory foci in GTE (iGTE) (IGTE also can be stumilated with 2 &#181;g/mL of lipopolysaccharides (LPS) for 48 h to initiate inflammation). IGTE is the first in vitro model of inflammatory gingiva using human cells with a three-dimensional architecture. IGTE reflects major pathological changes (immunocytes activition, intracellular interactions among fibryoblasts, epithelial cells, monocytes and macrophages) in periodontal diseases. GTE, wounded GTE, and iGTE can be used as versatile tools to study wound healing, tissue regeneration, inflammation, cell-cell interaction, and screen potential medicines for periodontal diseases.

The goal of the protocol is to build an inflammatory human gingiva model in vitro. This tissue model co-cultivates three types of human cells, HaCaT keratinocytes, gingival fibroblasts, and THP-1 macrophages, under three-dimensional conditions. This model can be applied to investigating periodontal diseases, such as gingivitis and periodontitis.

Periodontal diseases (such as gingivitis and periodontitis) are the leading causes of tooth loss in adults. Inflammation in gingiva is the fundamental ...

Three-dimensional Organotypic Cultures of Vestibular and Auditory Sensory Organs

The sensory organs of the inner ear are challenging to study in mammals due to their inaccessibility to experimental manipulation and optical observation. Moreover, although existing culture techniques allow biochemical perturbations, these methods do not provide a means to study the effects of mechanical force and tissue stiffness during development of the inner ear sensory organs. Here we describe a method for three-dimensional organotypic culture of the intact murine utricle and cochlea that overcomes these limitations. The technique for adjustment of a three-dimensional matrix stiffness described here permits manipulation of the elastic force opposing tissue growth. This method can therefore be used to study the role of mechanical forces during inner ear development. Additionally, the cultures permit virus-mediated gene delivery, which can be used for gain- and loss-of-function experiments. This culture method preserves innate hair cells and supporting cells and serves as a potentially superior alternative to the traditional two-dimensional culture of vestibular and auditory sensory organs.

Three-dimensional organotypic cultures of the murine utricle and cochlea in optically clear collagen I gels preserve innate tissue morphology, allow for mechanical stimulation through adjustment of matrix stiffness, and permit virus-mediated gene delivery.

The sensory organs of the inner ear are challenging to study in mammals due to their inaccessibility to experimental manipulation and optical observation. ...

Generation of Scaffold-free, Three-dimensional Insulin Expressing Pancreatoids from Mouse Pancreatic Progenitors In Vitro

The pancreas is a complex organ composed of many different cell types that work together to regulate blood glucose homeostasis and digestion. These cell types include enzyme-secreting acinar cells, an arborized ductal system responsible for the transportation of enzymes to the gut, and hormone-producing endocrine cells. 
Endocrine beta-cells are the sole cell type in the body that produce insulin to lower blood glucose levels. Diabetes, a disease characterized by a loss or the dysfunction of beta-cells, is reaching epidemic proportions. Thus, it is essential to establish protocols to investigate beta-cell development that can be used for screening purposes to derive the drug and cell-based therapeutics. While the experimental investigation of mouse development is essential, in vivo studies are laborious and time-consuming. Cultured cells provide a more convenient platform for screening; however, they are unable to maintain the cellular diversity, architectural organization, and cellular interactions found in vivo. Thus, it is essential to develop new tools to investigate pancreatic organogenesis and physiology.
Pancreatic epithelial cells develop in the close association with mesenchyme from the onset of organogenesis as cells organize and differentiate into the complex, physiologically competent adult organ. The pancreatic mesenchyme provides important signals for the endocrine development, many of which are not well understood yet, thus difficult to recapitulate during the in vitro culture. Here, we describe a protocol to culture three-dimensional, cellular complex mouse organoids that retain mesenchyme, termed pancreatoids. The e10.5 murine pancreatic bud is dissected, dissociated, and cultured in a scaffold-free environment. These floating cells self-assemble with mesenchyme enveloping the developing pancreatoid and a robust number of endocrine beta-cells developing along with the acinar and the duct cells. This system can be used to study the cell fate determination, structural organization, and morphogenesis, cell-cell interactions during organogenesis, or for the drug, small molecule, or genetic screening.

Generation of Scaffold-free, Three-dimensional Insulin Expressing Pancreatoids from Mouse Pancreatic Progenitors In Vitro

Here, we present a protocol to generate insulin expressing 3D murine pancreatoids from free-floating e10.5 dissociated pancreatic progenitors and the associated mesenchyme.

The pancreas is a complex organ composed of many different cell types that work together to regulate blood glucose homeostasis and digestion. These cell ...

In Vitro Generation of Heart Field-specific Cardiac Progenitor Cells

Pluripotent stem cells offer great potential for understanding heart development and disease and for regenerative medicine. While recent advances in developmental cardiology have led to generating cardiac cells from pluripotent stem cells, it is unclear if the two cardiac fields - the first and second heart fields (FHF and SHF) &#8212; are induced in pluripotent stem cells systems. To address this, we generated a protocol for in vitro specification and isolation of heart field-specific cardiac progenitor cells. We used embryonic stem cells lines carrying Hcn4-GFP and Tbx1-Cre; Rosa-RFP reporters of the FHF and the SHF, respectively, and live cell immunostaining of the cell membrane protein Cxcr4, a SHF marker. With this approach, we generated progenitor cells which recapitulate the functional properties and transcriptome of their in vivo counterparts. Our protocol can be utilized to study early specification and segregation of the two heart fields and to generate chamber-specific cardiac cells for heart disease modelling. Since this is an in vitro organoid system, it may not provide precise anatomical information. However, this system overcomes the poor accessibility of gastrulation-stage embryos and can be upscaled for high-throughput screens.

The purpose of this method is to generate heart field-specific cardiac progenitor cells in vitro in order to study the progenitor cell specification and functional properties, and to generate chamber specific cardiac cells for heart disease modelling.

Pluripotent stem cells offer great potential for understanding heart development and disease and for regenerative medicine. While recent advances in ...