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

Podsumowanie

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.

Streszczenie

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.

Wprowadzenie

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 death^2,3. 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%–50% of the blood to the left ventricular myocardium^6,7. 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 "widow maker." 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 response^12,13; 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 – 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)–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 structures^20,21.

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.

Access restricted. Please log in or start a trial to view this content.

Protokół

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–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

1. 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.
2. 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.
3. Create a sterile surgical area under an operating microscope. Gather sterilized surgical equipment (Table 1).
4. 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 3–4 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.
5. 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.
6. 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.
7. 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.
8. 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.
9. 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.
10. 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)
11. 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'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.
12. Remove the tape that was used to secure the hind legs of the pup.
13. 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.
14. 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.
15. Allow the neonate to remain directly on the heating pad for 10–15 min. Typically, movement is regained within 5 min of being placed onto heat.
16. Clean the residual blood and glue with an alcohol wipe.
17. 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.
18. Once all surgeries are completed and pups are warm and mobile, transfer the litter along with the original nesting material to the mother's cage.
19. Monitor the mice for 30–60 min after surgery and watch for the mother's acceptance of the pups by nesting and/or grooming.
20. 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.

2. Clearing the Mouse Heart with Passive CLARITY^21,22,23

1. Anesthetize the mouse with isoflurane. Perform a toe pinch to ensure the mouse is fully sedated.
2. Place the mouse onto a clean, surgical area in the supine position, securing the arms and legs with tape.
3. Maintain isoflurane sedation on the mouse using a nose cone until the heart is extracted.
4. 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.
5. Cut alongside of the distal portions of the rib cage with surgical scissors.
6. Expose the diaphragm muscle by grasping the xiphoid process with tissue forceps. Detach the diaphragm using curved forceps.
7. While maintaining a grasp of the xiphoid process, pull the tissue cranially until the beating heart is accessible.
8. 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.
9. 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.
10. Gently squeeze the heart with forceps to allow the heart to expel the residual blood.
11. 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.
12. Discard the PBS and fill the vial with 2 mL of cold 4% paraformaldehyde (PFA). Incubate for 4 hours at RT (Figure 2A).
13. 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.
14. Discard PBS and fill the vial with 2 mL of 4% acrylamide and 0.5% VA-044 solution. Incubate overnight at 4 °C.
15. The next day, perform polymerization by transferring the vial to a heat block set at 37 °C for 3 hours.
16. Transfer the heart into a new glass shell vial and repeat step 2.12 (PBS wash cycle).
17. Discard PBS and fill the vial with 2 mL of Clearing Solution (Table 2). Incubate at 37 °C until the heart is cleared. Change out the solution and refill with fresh Clearing Solution every 2–3 days. The clearing process could take up to several weeks (Figure 2B-C).
    NOTE: P1 hearts typically take around 3–5 days, whereas P21 hearts can take nearly a month before Clearing Solution incubation is complete.
18. 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 °C.
19. If immunostaining will be performed, skip steps 2.21–2.22 and proceed to Section 3 for immunostaining. If relying solely on endogenous fluorescence, proceed with steps 2.21–2.22 and skip Section 3.
20. Discard PBS and change the solution to Refractive Index Matching Solution (RIMS) (Table 3). Incubate overnight at 37 °C.
21. 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).

3. Optional: Immunohistochemistry Staining of the Whole-Mount Mouse Heart

1. 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)
2. Wash the heart in PBST 3 times on an orbital rotator with 30 min incubations at RT.
3. 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 °C to stain with rotation overnight.
   NOTE: Blocking buffer is made from normal serum matching the species in which the secondary antibody was raised.
4. Wash the heart in PBST 3 times with rotation for 5 min incubations at RT.
5. 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.
6. Incubate for an additional 24 hours with rotation at 4 °C.
7. 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.
8. 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 °C to stain with rotation overnight.
9. On the next day, repeat step 3.2 (long PBST wash cycle) to remove unbound secondary antibody.
10. Replace the solution with 2% blocking buffer (diluted in PBST). Remove residual unbound antibody by washing overnight with rotation at RT.
11. 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.
12. Store the immunostained heart in PBS at 4 °C.
13. Directly before whole-mount microscopy, incubate the heart in RIMS solution overnight at 37 °C. Extend the incubation an additional 24 hours if the tissue is still not fully cleared.
14. Store the fully cleared and immunostained heart in RIMS at RT.

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.

1. Imaging the Cleared Embryonic Mouse Heart
   1. Fill the microscope depression slide with the PBS solution.
   2. Carefully pick up the cleared heart with curved forceps and place the tissue onto the slide.
   3. Mount the slide with a glass coverslip. The tissue can now be imaged under a confocal microscope.
2. Imaging the Cleared Postnatal Mouse Heart
   1. 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).
   2. 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.
   3. 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.

Access restricted. Please log in or start a trial to view this content.

Wyniki

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...

Access restricted. Please log in or start a trial to view this content.

Dyskusje

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, arterioles^12,13, and lymphatic endothelial cells²⁶, all play an ...

Access restricted. Please log in or start a trial to view this content.

Materiały

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