Delayed Intramyocardial Delivery of Stem Cells after Ischemia Reperfusion Injury in a Murine Model

Summary

Stems cells are continuously investigated as potential treatments for individuals with myocardial damage, however, their decreased viability and retention within injured tissue can impact their long-term efficacy. In this manuscript we describe an alternative method for stem cell delivery in a murine model of ischemia reperfusion injury.

Abstract

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, cardiac stem cell therapy is studied by delivering SCs concurrently with the induction of myocardial injury. However, this approach presents two significant limitations: the early hostile pro-inflammatory ischemic environment may affect the survival of transplanted SCs, and it does not represent the subacute infarction scenario where SCs will likely be used. Here we describe a two-part series of surgical procedures for the induction of ischemia-reperfusion injury and delivery of mesenchymal stem cells (MSCs). This method of stem cell administration may allow for the longer viability and retention around damaged tissue by circumventing the initial immune response. A model of ischemia reperfusion injury was induced in mice accompanied by the delivery of mesenchymal stem cells (3.0 x 10⁵), stably expressing the reporter gene firefly luciferase under the constitutively expressed CMV promoter, intramyocardially 7 days later. The animals were imaged via ultrasound and bioluminescent imaging for confirmation of injury and injection of cells, respectively. Importantly, there was no added complication rate when performing this two-procedure approach for SC delivery. This method of stem cell administration, collectively with the utilization of state-of-the-art reporter genes, may allow for the in vivo study of viability and retention of transplanted SCs in a situation of chronic ischemia commonly seen clinically, while also circumventing the initial pro-inflammatory response. In summary, we established a protocol for the delayed delivery of stem cells into the myocardium, which can be used as a potential new approach in promoting regeneration of the damaged tissue.

Introduction

Cardiovascular disease remains the most common cause of morbidity and mortality worldwide. Cardiac ischemic events have been found to be detrimental to the overall function of the myocardium and surrounding cells¹. Only   ̴0.45-1.0% of cardiomyocytes will regenerate every year after myocardial damage occurs². Despite the growing demand and inherent focus on developing treatments, therapies aiding in the regeneration of injured tissue have been difficult to establish and still require further optimization^3,4,5. Stem cell therapies have been introduced as an alternative path to rejuvenate damaged tissue after an ischemic event; however, advancement of these therapies has been challenged by the limited survival and retention of the cells to an injured area⁶.

The microenvironment of the heart after an ischemic event can be characterized as hypoxic, pro-oxidant, and pro-inflammatory, presenting hostile conditions for therapeutic stem cells to adapt to for survival^7,8. As an immune response is triggered following injury, naïve lymphocytes, macrophages, neutrophils and mast cells attempt to repair the damage by removing dying cells and modulating the process for tissue remodeling^9,10,11. Within the first 3 days post-ischemia, inflammation is at its peak with the release of pro-inflammatory cytokines with high numbers of neutrophils and monocytes in the area^10,12. After 7 days, much of the inflammation has subsided and the transition to reparative cells begins, continuing until the remodeling cascade is complete, approximately 14 days in mice¹³. Our surgical method is a potential alternative approach to the introduction of biologics into the myocardium to bypass the peak innate immune response after ischemia reperfusion injury. At the same time, it will allow for the study of any treatments in a condition of subacute/chronic ischemia where there may be different variables to consider compared to acute myocardial infarction.

Protocol

The experiments were performed on female C57BL/6 mice, age 10-12 weeks and 20-25 g body weight. All animal procedures complied with the standards stated in the Guide for the Care and Use of Laboratory Animals (Institute of Laboratory Animal Resources, National Academy of Sciences, Bethesda, MD, USA) and were approved by the Mayo Clinic College of Medicine Institutional Animal Care and Use Committee (IACUC).

1. Preparation and intubation

1. Autoclave all surgical instruments before surgery. If multiple surgeries are to be performed in one session, clean the instruments after each animal and re-sterilize using a hot bead sterilizer.
2. Anesthetize the mice with 3.5-4% isoflurane at 1 L/min O₂ in an induction chamber.
3. Administer Buprenorphine SR 1 mg/kg (analgesic) subcutaneously, weigh the animal, and input the weight into the ventilator.
4. Shave the left side of the chest from the sternum to the level of the shoulder and apply depilatory cream to remove excess fur.
5. For the ischemia reperfusion procedure maintain the positive end-expiratory pressure (PEEP) on the ventilator at 2 cmH₂O. For the delayed injection of cells procedure change the PEEP to 3 cmH₂O to prevent lung collapse.
6. Intubate the animal using a 20 G endotracheal tube, transfer to a controlled heating pad to maintain a body temperature of 35-37 °C.
7. Place the mouse on a ventilator in lateral recumbency with cranial end on the left and caudal end on the right.
8. Maintain anesthesia at 2-2.5% isoflurane at 1 L/min O₂ for the remainder of the procedure.
9. Scrub the surgical area alternating between povidone-iodine and alcohol swabs three times and apply ophthalmic ointment to both eyes.

2. Ischemia reperfusion injury

1. Using a #10 blade scalpel make a vertical incision 2.5 mm to right of the leftmost nipple in the field of view.
2. Using scissors cut through the superficial muscle layers until the intercostal muscles and ribs are visible.
3. While lifting the ribs and surrounding tissue, cut through the intercostal space between the 4th and 5th ribs, then insert the eyelid retractor into the open space.
4. Retract the pericardium using curved forceps, moving the lung upwards and out of view.
5. Visualize LAD artery and, using a 9-0 nylon suture, pass through the myocardium beneath the artery 2.5 mm distal to the left auricle and tie a loose square knot.
6. Cut 1 cm of polyethylene tubing and place it within the loose knot.
7. Secure the suture around the tubing, confirm ischemia, then release after 35 min.
   NOTE: Confirm ischemia by pallor and ventricular arrhythmia.
8. After releasing the ligation and removing the tubing, wait for 5 min to confirm reperfusion of the myocardium.
9. Place a 24 G I.V. catheter tube into the thoracic cavity one intercostal space to right of the opening.
10. Close the intercostal incision with a 6-0 absorbable suture in a simple interrupted pattern.
11. Close the muscle layer with a 6-0 absorbable suture in a continuous suture pattern.
12. After closing the superficial muscle layer, remove the chest tube while withdrawing the air from thoracic cavity using a 1 mL tuberculin syringe.
13. Close the skin incision with a 6-0 absorbable suture in a continuous horizontal mattress pattern
    NOTE: Nylon sutures and a discontinuous suture pattern may also be used for the skin layer.
14. Administer 1.5 mL of warm saline subcutaneously and apply triple-antibiotic ointment to the incision site to prevent infection.
15. Turn off isoflurane and allow the animal to breathe through the ventilator on 100% O₂ until it can breathe continuously without aid.
16. Transfer the mouse to a bedding-free cage or a cage with covered bedding (paper towel or drape) on a warm pad with a temperature of 35-37 °C until fully recovered.

3. Mouse mesenchymal stem cell delivery

NOTE: The strain of mice used for the procedure are an inbred line and are deemed genetically identical. The mesenchymal stem cells were obtained from animals of the same strain and, by protocol design, immunosuppression was not induced¹.

1. Complete the preparation and intubation steps as done previously for the first procedure.
2. Remove the suture from the skin layer using scissors and forceps.
3. With a #10 scalpel, make an incision in the same location as the previous surgery.
4. Continue to use the scalpel to cut through scar tissue until muscle layer suture is visible
5. Using the scissors and forceps remove the suture and cut the muscle layer open.
6. Visualize and remove the sutures holding the ribs together and continue cutting through the intercostal muscle from the previous incision.
   NOTE: The lungs may have adhered to the chest wall, if this occurs, use blunt or curved forceps to carefully separate and release them.
7. Place the eyelid retractor into the intercostal space and locate the area of the previous ligation.
8. Load the mesenchymal stem cells (3.0 x 10⁵), suspended in 20 µL PBS, into a 30 G insulin syringe, bend the needle slightly as needed for the proper angle to inject.
   NOTE: Mesenchymal stem cells (MSCs) were isolated from the adipose tissue of 4-6-week-old C56BL/6 mice. Early passage cells (p3) were transduced with a vector expressing the firefly luciferase gene under the CMV promoter to allow in vivo cell viability monitoring. Adipose-derived mouse MSC were characterized by flow cytometry and the cells were positive for CD44, CD29, CD90 and CD105 but negative for the hematopoietic marker CD45¹⁴. Prior to the injection, MSCs were cultured for at least one passage to avoid the loss of cells from the thawing process.
9. Moving in the direction from the apex towards the base of the heart insert the syringe into the peri-infarct region until the needle opening is completely inside the myocardium.
10. Once inside slowly inject the cells into the myocardium, wait 3 s, then remove the needle.
11. Observe the heart closely for 3 min to be sure of no abnormal reactions to the cells such as ventricular fibrillation.
12. Place a 24 G IV catheter tube into the thoracic cavity one intercostal space to right of the opening.
13. Close the intercostal, muscle, and skin layers and remove the chest tube in the same method as the first procedure.
14. Administer 1.5 mL of warm saline subcutaneously and apply triple-antibiotic ointment to the incision site to prevent infection.
15. Turn off isoflurane and allow animal to breathe through the ventilator on 100% O₂ until it is able to breathe continuously without aid.
16. Transfer the mouse to a bedding-free cage or a cage with covered bedding (paper towel or drape) on a warm pad with a temperature of 35-37 °C until fully recovered.

4. Post-operative care following both procedures

1. Observe the animal continuously until spontaneous breathing, sternal recumbency and normal movement is established.
2. Continue observation every 15-30 min for at least 3 h on the day of the surgery.
3. Check the mice for wound dehiscence or abnormal pain once daily for 5 days, then 2-3 times weekly.
4. If the animal shows signs of pain (i.e. arched back, minimal movement, grimacing, or scruffy fur) after 72 h post-op, provide an additional dose of the Buprenorphine SR analgesic.

Results

Ischemia reperfusion injury was induced in mice on day 0, followed by a post-operative echocardiogram and electrocardiogram on the day preceding stem cell implantation. Ultrasound and electrocardiogram analysis confirmed infarction and decreased ventricular contractile function (Figure 1A-D). Further examination of the data showed the ejection fraction and fractional shortening were decreased in mice that received ischemic injury, while the end-diastolic and systolic volumes...

Discussion

Over 85 million people worldwide are affected by cardiovascular disease³. The high prevalence of these ischemic events warrants further development and expansion of alternative therapies for promoting the regeneration of damaged tissue. Traditional methods utilize the ischemia reperfusion procedure in an acute setting with subsequent administration of therapeutics¹. Inflammatory reactions are at its peak between 3-4 days postdating a cardiac ischemic event, with infiltratio...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.9% NaCl Irrigation, USP	Baxter	0338-0048-04
11x12" Press n' Seal surgical drape, autoclavable	SAI Infusion Technologies	PSS-SD
24G 3/4" IV catheter tube	Jelco	4053
28G x 1/2" 1mL allergy syringe	BD	305500	Injection of analgesic
30G x 1/2" 3/10cc insulin syringe	Ulticare	08222.0933.56	Injection of stem cells
6-0 S-29, 12" Vicryl suture	Ethicon	J556G	Intercostal, superficial muscle and skin layer incision closure
9-0 BV100-4, 5" Ethilon suture	Ethicon	2829G	Ligation of the LAD artery
Absorbent underpad	Thermo Fischer Scientific	14-206-64	For underneath the animal
Alcohol prep pads, 2 ply, medium	Coviden	6818
Anti-fog face mask	Halyard	49235
Bonn Strabismus scissors, curved, blunt	Fine Science Tools	14085-09
Buprenorphine HCL SR LAB 1mg/ml, 5 ml	ZooPharm Pharmacy		Buprenorphine narcotic analgesic formulated in a polymer that slows absorption extending duration of action (72 hours duration of activity).
Castroviejo needle holders, curved	Fine Science Tools	12061-01
Curity sterile gauze sponges	Coviden	397310
Delicate suture tying forceps, 45 angle bent	Fine Science Tools	11063-07
Electric Razor	Wahl		Fur removal
Isoflurane 100 ml	Cardinal Health	PI23238	Anesthetic
Lab coat
Monoject 1 mL hypodermic syringe	Coviden	8881501400
Moria iris forceps, curved, serrated (x2)	Fine Science Tools	11370-31
Moria speculum retractor	Fine Science Tools	17370-53
Mouse endotracheal intubation kit	Kent Scientific
Nair depilatory cream	Johnson & Johnson		Fur removal
Optixcare eye lube plus	Aventix		Sterile ocular lubricant
Physiosuite ventilator	Kent Scientific
PolyE Polyethylene tubing	Harvard Apparatus	72-0191	Temporary compression of LAD artery
Povidone-iodine swabs	PDI	S41125
Scalpel, 10-blade	Bard-Parker	371610
Sterile 3" cotton tipped applicators	Cardinal Health	C15055-003
Sterile 6" tapered cotton tip applicators	Puritan	25-826-5WC
Sterile gloves	Cardinal Health	N8830
Sterilization pouches	Medline	MPP100525GS
Surgery cap
Surgical Microscope	Leica	M125
Suture tying forceps, straight (x2)	Fine Science Tools	10825-10
Transpore surgical tape	3M	1527-1
Triple antibiotic ointment	G&W Laboratories	11-2683ILNC2	Topical application to prevent infection
Vannas-Tübingen Spring Scissors, curved	Fine Science Tools	15004-08
Vetflo vaporizer	Kent Scientific