An Intact Pericardium Ischemic Rodent Model

Podsumowanie

This protocol outlines the steps for inducing myocardial infarction in mice while preserving the pericardium and its contents.

Streszczenie

This protocol has shown that the pericardium and its contents play an essential anti-fibrotic role in the ischemic rodent model (coronary ligation to induce myocardial injury). The majority of pre-clinical myocardial infarction models require the disruption of pericardial integrity with loss of the homeostatic cellular milieu. However, recently a methodology has been developed by us to induce myocardial infarction, which minimizes pericardial damage and retains the heart's resident immune cell population. An improved cardiac functional recovery in mice with an intact pericardial space following coronary ligation has been observed. This method provides an opportunity to study inflammatory responses in the pericardial space following myocardial infarction. Further development of the labeling techniques can be combined with this model to understand the fate and function of pericardial immune cells in regulating the inflammatory mechanisms that drive remodeling in the heart, including fibrosis.

Wprowadzenie

To this day, cardiovascular disease (CVD) is recognized as the leading cause of death globally, resulting in a significant financial burden and reduction in patient quality of life¹. Coronary artery disease (CAD) is a sub-type of CVD and plays an essential role in the development of myocardial infarction (MI), which is a chief contributor to mortality. By definition, MI results from irreversible injury to the myocardial tissue due to prolonged conditions of ischemia and hypoxia. Myocardial tissue lacks regeneration capacity, so injuries are permanent and result in the replacement of heart muscle with a fibrotic scar that can be initially protective but ultimately contributes to adverse cardiac remodeling and eventual heart failure².

Although the management of patients with CAD has dramatically improved over the past few decades, chronic heart failure (CHF) secondary to ischemia affects many patients worldwide. For preventing and managing this epidemic, it is necessary to understand the underlying mechanisms more extensively and develop new therapeutic approaches. Moreover, past findings highlight the limitations of systemic therapy and the necessity of developing precise alternatives. Given investigating the molecular sequelae of MI in humans is affected by the ability to access infarcted tissue, animal models that recapitulate the characteristics and development of human MI and CHF related to CVD are indispensable.

As ideal animal models closely resemble a human disorder for structural and functional characteristics, disease etiology should guide their conception. In CAD, it is the chronic atherosclerotic stenosis of coronary arteries or acute thrombotic occlusion. Different methods have been developed and applied in various species of laboratory animals to induce coronary artery narrowing or occlusion. Such strategies can be broadly classified into two groups: (1) mechanical manipulation of a coronary artery to induce an MI and (2) expediting atherosclerosis to facilitate coronary narrowing leading to an MI. The first strategy usually involves either the ligation of a coronary artery or the placement of a stent within the artery. The second approach tends to rely on modifying the animal's diet to include high fat/cholesterol food. Some of the limitations of this latter approach include the lack of control on the timing and site of coronary occlusions.

In contrast, the surgical induction of MI or ischemia in an animal model has several advantages, such as location, precise timing, and extent of the coronary event, leading to more reproducible results. The most widely used method is surgical ligation of the left anterior descending coronary artery (LAD). Such models recapitulate human responses to acute ischemic injury, as well as the progression to CHF³. Initially developed in larger animals, LAD surgery on small animals such as rodents has become more feasible with advancements in technology⁴. In establishing such models, mice have been favored for various reasons, including their relative availability, low expense in housing, and their capacity for genetic manipulation.

Contemporary surgical models of ischemic heart disease using LAD occlusion require the researcher to open the pericardium to temporarily or permanently ligate the artery⁵. Such strategies result in the disruption of the pericardial space, which plays an essentially mechanical and lubricating function to ensure proper cardiac function. Another disadvantage of opening the pericardium is losing the animal's native pericardial fluid with its various cellular and protein components^6,7. In response, a method to induce MI while keeping the pericardium intact was developed by us. In addition to minimizing the perturbation of this homeostatic environment, this approach allows for tagging and tracing specific cells after causing an MI. In addition, this approach better represents myocardial ischemic injury in the human setting.

Protokół

Male and female C57BL/6J mice between 8-14 weeks of age were used for these experiments. This protocol has received ethical approval from the Animal Care Committee at the University of Calgary and follows all animal care guidelines.

1. Mouse preparation and surgery

1. Sterilize surgical tools (via bead sterilizer or autoclave).
2. Weigh mouse for presurgical weight and analgesic dose.
3. Place the mouse in an induction box with 4% isoflurane and 800 mL/min of oxygen. Confirm the anesthetic plane by pinching the toes and observing the animal for lack of reflex.
4. Inject analgesic subcutaneously (0.1 mg/kg of Buprenorphine) (see Table of Materials).
5. Place the mouse on a heated surgical pad during surgery and maintain anesthesia with 3% isoflurane delivered via nose cone. Apply ophthalmic ointment to avoid dry eyes to each eye.
6. Shave the hair from the chest and neck surgical areas.
7. Restrain the paws of the mouse and position them on the surgical table.
8. Intubate the mouse by inserting a smooth tipped 23 G catheter into the trachea through the mouth and the pharynx.
   1. Ventilate the mouse following intubation with 2% isoflurane and 100% oxygen as a carrier gas using a commercial Ventilator (see Table of Materials) set at a rate of 110 breaths/min, a tidal volume of 250 µL, and positive end-expiratory pressure (PEEP) of 4 mmHg.
9. Roll the mouse 30% on its right side to position the left side of the chest for surgery.
10. Clean the surgical area with 3 alternating scrubs of 70% ethanol and betadine in a circular motion(seeTable of Materials). Place sterile surgical drapes around the surgical area.
11. Make a 2-3 cm lateral incision in the skin of the chest to visualize the pectoralis muscles on the left side. Cut the pectoralis major and minor using a 1 cm incision from the midline outward to visualize the intercostal muscles between the third and fourth ribs.
    NOTE: Care should be taken to avoid excess bleeding from the pectoralis muscle through cauterization of bleeding vessels.
12. Make a 2 cm incision in the left intercostal muscle to introduce air (by passive air movement) into the chest cavity to allow the heart and lungs to fall away from the surgical incision. Further, expand the opening with the help of a cautery device (see Table of Materials) to incise the intercostal and prevent bleeding
    NOTE: The ventilator should be temporarily stopped during the period of cauterization to avoid explosive reactions with oxygen. Care is taken not to damage the pericardial sac.
13. Using retractors, retract the ribs to expose the heart.
14. Observe the pericardium and the underlying heart under a stereomicroscope.
    NOTE: The mouse pericardium is thin enough to visualize the vasculature of the heart.
15. Gently place forceps on the surface of the pericardium to reduce its movement and that of the underlying heart.
16. Visually landmark the left anterior descending (LAD) coronary artery by tracing its emergence from under the left appendage.
17. Using the micro-needle driver, guide an appropriate suture (see Table of Materials) through the pericardium, under the LAD, with the suture emerging on the other side of the LAD and pericardium. Tie the suture to restrict the blood flow through the coronary artery and trim the excess suture with the help of scissors (Figure 1A).
    NOTE: Upon restricting blood flow to the coronary artery, blanching of the anterior portion of the left ventricle should be visible. This procedure represents a permanent ligation model. However, a transient ligation approach with different ischemia periods could also be applied at this stage.
18. Below the incision site within the sterile area, introduce a 24 G catheter percutaneously into the chest (remove the guide needle after entering the chest cavity). Then, close the ribs followed by muscle layers and skin using an appropriate suture (a taper needle for muscle, conventional cutting needle for skin).
19. Once the chest is closed, evacuate the remaining air from the chest cavity via the 24 G catheter using gentle suction with a 3 mL syringe and chest compressions. Once the air is removed, withdraw the 24 G catheter.
20. Reduce the isoflurane to 1%.
21. Turn off the isoflurane while maintaining ventilation with oxygen to allow the mouse to recover from anesthesia. Once the animal shows signs of breathing independently, remove the 23 G tracheal tube from the mouth and place the mouse in a recovery cage to be monitored for resumption of normal breathing.
22. Allow the mouse to recover in the cage with a portion of the cage placed on a warming pad to provide an external heat source.
23. Provide maintenance injections of analgesic (Buprenorphine 0.1 mg/kg, subcutaneously) every 12 h for 72 h post-surgery.
24. Monitor the health status of mice daily for 7 days, which includes evaluating incisions and animal discomfort.
    ​NOTE: Due to the invasiveness of this procedure (thoracotomy), the administration of antibiotics may be necessary.

2. Functional assessment of cardiac function by echocardiography (ECG)

1. Induce and maintain the mouse under general anesthesia with 1.5-2% isoflurane and 800 mL/min of oxygen.
2. Place mouse in a supine position on a heated stage platform and attach the paws to the ECG leads.
3. Shave the chest of the mouse.
4. Acquire echocardiography images using a 40 MHz linear transducer probe and contact gel and analyze with the appropriate software (see Table of Materials).
5. Turn off the isoflurane and allow the mouse to recover on the heating platform before returning the cage to an active state.
   ​NOTE: Echocardiography assessment is non-invasive and thus can be performed longitudinally throughout the experiment to determine changes before and after coronary ligation.

3. Heart tissue collection for fibrosis staining

1. Sacrifice the mice with inhalation of 100% CO₂ and carefully dissect out the heart.
   NOTE: Using scissors and forceps, this is achieved by cutting through the large vessels entering (vena cava, pulmonary vein) and exiting (pulmonary artery, aorta) the heart to release it from the circulatory system in the thoracic cavity.
2. Fix the heart in 10% formalin for at least 24 h.
3. Cut samples using a straight razor blade through the right ventricle, interventricular septum, and left ventricle, ensuring that the incision went through the center of the infarct zone. Samples are then sent to the core facility for paraffin embedding.
4. Cut tissue sections of 5 µm thickness with a microtome and place them on glass slides for staining.
5. Deparaffinize using commercial xylene and graded alcohol washes (2x 99%, 1x 95%, 1x 70%) with deionized water, then rehydrate.
6. Stain with 0.1% Sirius red in picric acid for 2 h at room temperature.
7. Wash sections with 0.5% acetic acid for 3 min and rinse with 70% ethanol for 1 min.
8. Dehydrate the sections using the reverse order of washes outlined in 3.4, with increasing and graded ethanol concentrations then xylene.
9. Mount tissue sections with a mounting solution (see Table of Materials) for microscopic evaluation.

4. Flow cytometry of heart and pericardial cavity lavage

1. Sacrifice the mice with inhalation of 100% CO₂ to effect.
2. Place the mouse on its back and fix the arms and legs to a surgical board using tape.
3. Carefully open the left side (right side from the experimenter view) of the thoracic cavity, starting with cutting the diaphragm to about the midpoint and subsequently cutting through the outside ribs towards the sternum.
   NOTE: Avoid nicking large blood vessels, particularly those that run parallel to the sternum.
4. Retract the ribs using a hemostat to expose the underlying heart and pericardium.
   NOTE: The pericardium is very fragile, so be sure not to catch it with scissors during the cutting.
5. Using a PE-10 tubing (see Table of Materials) catheter inserted into the pericardial space near the junction of the left atrium and left ventricle, inject 100 µL of sterile saline into the pericardial cavity.
   1. Allow saline to pool and collect from the posterior side of the heart, being careful not to puncture or tear the pericardium in the process. Repeat this step twice and place lavage fluid on ice while processing the heart.
6. Excise the heart by cutting the major vessels (aorta, pulmonary artery, vein, and vena cava) entering and exiting the heart. Remove the right and left atria and weigh the ventricular heart tissue.
7. Mince the tissue in small 1 mm² pieces using scissors and place in 10 mL of digestion buffer containing 450 U/mL of collagenase I, 125 U/mL of collagenase XI, 60 U/mL of DNase I, and 60 U/mL of hyaluronidase in PBS for 1 h at 37 °C on an orbital shaker.
8. Pass heart tissue homogenates through a 70 µm cell strainer (see Table of Materials) and spin down at 60 x g for 5 min at 4 °C to remove cardiac parenchymal cells.
9. Collect the supernatant, pass through a 40 µm cell strainer (see Table of Materials) for a single cell suspension, and spin down at 400 x g for 5 min at 4 °C to pellet the cells.
10. Perform fragment crystallizable (Fc) receptor blocking and staining of cellular markers on pericardial and cardiac cells as previously described⁸.
11. Run samples on a flow cytometer.

5. Labeling pericardial macrophage using the Intercostal Approach to the Pleural Space (ICAPS) method⁹

1. Sterilize surgical tools (via bead sterilizer or autoclave) and spray 70% ethanol before commencing.
2. Induce and maintain the mouse under general anesthesia with 1.5-2% isoflurane and 800 mL/min of oxygen. Confirm the anesthetic plane by pinching the toes and observing the animal for lack of reflex.
3. Inject analgesic subcutaneously (Buprenorphine 0.1 mg/Kg).
4. Place the mouse on a heated surgical pad during surgery.
5. Shave the right anterolateral thoracic area.
6. Clean the surgical area with ethanol and betadine.
7. Make a 3 cm long incision in the skin, and with forceps expose the rib cage.
8. Load 5 µL of fluorescent beads (commercially available fluorescent microspheres, 1 µm, see Table of Materials) and 45 µL of saline into a PE-10 tubing syringe catheter with a beveled tip.
9. Guide the catheter into the intercostal space as previously described⁹, slowly inject the bead solution and remove the catheter in one motion.
10. Close the skin using staples.
    NOTE: Staples are used in place of sutures to minimize potential re-opening of the incision.
11. Turn off the isoflurane, place the mouse in the recovery cage and monitor for complications over the first 24 h.

Wyniki

This modified coronary ligation model has been optimized to achieve reproducibility and animal survival. However, due to the significant injury induced in the heart, some expected intra-operative and post-operative mortality are associated with the procedure. The standard mortality is typically higher in males (~25-35%) than in females (~ 10-15%).

Successful induction of an MI with the modified coronary ligation should be evident by changes in the heart's functional parameters and structur...

Dyskusje

Inducing an MI in a closed pericardium in rodents is unique and can have potentially significant applications. The procedure relies heavily on the surgeon's familiarity with the rodent model and rodent cardiac anatomy. Success is also dependent on the care given during three critical steps: intercostal muscle incision and rib retraction (Steps 1.11-1.13), creating the infarct (Step 1.17 ), and animal recovery (Steps 1.22-1.24).

The thoracotomy must be done diligently to avoid puncturing or...

Materiały

Name	Company	Catalog Number	Comments
Steri-350 Bead Sterilizer	Inotech	NC9449759
10% Formalin	Millipore Sigma	HT501128-4L
40 µm Cell strainer	VWR	CA21008-949	Falcon, 352340
70 µm Cell strainer	VWR	CA21008-952	Falcon, 352350
ACK Lysis Buffer	Thermo Fisher	A1049201
BD Insyte-W Catheter Needle 24 G X 3/4"	CDMV Inc	108778
Betadine (10% povidone-iodine topical solution)	CDMV Inc	104826
Blunt Forceps	Fine Science Tools	FST 11000-12
BNP Ophthalmic Ointment	CDMV Inc	17909
Castroviejo Needle Driver	Fine Science Tools	FST 12061-01
Centrifuge 5810R	Eppendorf	22625101
Collagenase I	Millipore Sigma	SCR103
Collagenase XI	Millipore Sigma	C7657
Covidien 5-0 Polysorb Suture - CV-11 taper needle	Medtronic Canada	GL-890
Covidien 5-0 Polysorb Suture - PC-13 cutting needle	Medtronic Canada	SL-1659
Curved Blunt Forceps	Fine Science Tools	FST 11009-13
Dako Mounting Medium	Agilen	CS70330-2
DNase I	Millipore Sigma	11284932001
Ethanol, 100%	Millipore Sigma	MFCD00003568
Ethicon 8-0 Ethilon Suture - BV-130-4 taper needle	Johnson & Johnson Inc.	2815G
Fiber-Optic Light	Nikon	2208502
Fine Forceps	Fine Science Tools	FST 11150-10
Fluoresbrite® YG Carboxylate Microspheres 1.00 µm	Polysciences, Inc.	15702
Geiger Thermal Cautery Unit	World Precision Instruments	501293	Model 150-ST
Hyaluronidase	Millipore Sigma	H4272
Isofluorane Vaporizer	Harvard Apparatus	75-0951
Isoflurane USP, 250 mL	CDMV Inc	108737
Magnetic Fixator Retraction System	Fine Science Tools	18200-20
MX550D- 40 MHz probe	Fujifilm- Visual Sonics
Needle Driver	Fine Science Tools	FST 12002-12
PE-10 Tubing	Braintree Scienctific, Inc.	PE10 50 FT
Scissors	Fine Science Tools	FST 14184-09
SMZ-1B Stereo Microscope	Nikon	SMZ1-PS
VentElite Small Animal Ventilator	Harvard Apparatus	55-7040
Vetergesic (10 mL, 0.3mg/mL buprenorphine))	CDMV Inc	124918	controlled drug
Vevo 2100 Software	Fujifilm-Visual Sonics