Intramyocardial Cell Delivery: Observations in Murine Hearts

Summary

Intramyocardial cell delivery in murine models of cardiovascular diseases, such as hypertension or myocardial infarction, is widely used to test the therapeutic potential of different cell types in regenerative studies. Therefore, a detailed description and a clear visualization of this surgical procedure will help to define the limits and advantages of cardiovascular cell therapeutic analyses in small rodents.

Abstract

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue revascularization and cell survival. In addition, further observations revealed that specific stem cells, such as cardiac stem cells, mesenchymal stem cells and cardiospheres have the ability to integrate within the surrounding myocardium by differentiating into cardiomyocytes, smooth muscle cells and endothelial cells.

Here, we present the materials and methods to reliably deliver noncontractile cells into the left ventricular wall of immunodepleted mice. The salient steps of this microsurgical procedure involve anesthesia and analgesia injection, intratracheal intubation, incision to open the chest and expose the heart and delivery of cells by a sterile 30-gauge needle and a precision microliter syringe.

Tissue processing consisting of heart harvesting, embedding, sectioning and histological staining showed that intramyocardial cell injection produced a small damage in the epicardial area, as well as in the ventricular wall. Noncontractile cells were retained into the myocardial wall of immunocompromised mice and were surrounded by a layer of fibrotic tissue, likely to protect from cardiac pressure and mechanical load.

Introduction

Various cell delivery protocols have been tested in murine and rat models of cardiovascular diseases with the aim of translating the efficiency, efficacy and safety of this experimental procedure in human patients. In small rodent hearts, intramyocardial cell delivery is the most feasible method of cell delivery^1,2, whereas in the rat heart antegrade³ and retrograde⁴ intracoronary cell infusion can also be used. Both methods have limits and advantages. Cell delivery via the intracoronary route has theoretical advantages over direct intramuscular injection in promoting global cell dissemination³, but it also has the risk of causing coronary embolism^3,5. Limitations on intramyocardial delivery are associated with mechanical injury, acute inflammation, and myocardial damage^6,7. In humans, cells for cardiac repair are delivered by intramyocardial injection through an endocardial or surgical epicardial approach or by intracoronary arterial route⁸. Injection by transvascular routes is suited in patients with acute infarct and reperfused myocardium, but may not be possible in case of total occlusions or poor flow within the vessels of the effected territory⁹. Direct injection into the ventricular wall by transendocardial or transepicardial injection is technically feasible depending on the patient health status. Indeed, it has been shown that this technique is safe^10,11, although for transepicardial injection an open chest surgery is necessary and for transendocardial approaches an electrophysiological mapping for each patient is required to differentiate sites of viable ischemic or scarred myocardium⁹.

Importantly, in cell therapy studies the choice of the best cell to be transplanted is still under investigation. Short-term analyses (4 weeks) showed that injection of cardiac stem cells defined as cardiospheres¹² or side population cells from bone marrow¹³ induced cardiac functional recovery in murine¹⁴ and rat¹⁵ models of myocardial infarct by decreasing scar size and cell death. Allogeneic transplantation of cardiospheres in a rat myocardial infarct model without immunosuppression was found to be safe, promoted cardiac regeneration, and improved heart function through stimulation of endogenous repair mechanisms¹⁵. Lin-/c-kit+ adult progenitor cells in the heart were shown to be self-renewing, clonogenic, and multipotent in vitro and in vivo, and when injected into an ischemic rat heart reconstituted large portions of the injured myocardial wall¹⁶ and had the ability to form conductive and intermediate-sized coronary arteries¹⁷. These promising data fuelled phase I and II clinical trials in humans: injection of autologous and allogeneic mesenchymal stem cells (MSC)¹⁸, cardiospheres¹⁹, or c-Kit positive cardiac stem cells (CSC)20 in ischemic human hearts each showed beneficial effects in cardiac function in long term studies. Nevertheless, extensive long-term follow-up and retrospective meta-analysis demonstrated that stem cell therapy provides significant benefit to some patients, but not in others with a range of unpredictable outcomes²¹. It is possible that these limitations will necessitate the design of specific protocols of cell delivery for each individual and each disease.

In mouse and rat models, long-term studies revealed that cell injection did not further improve cardiac function (12 months). Indeed, grafts of human embryonic stem cell-derived cardiomyocytes (hESC-CM) were largely isolated from the host myocardium by a layer of fibrotic tissue^22,23. Similar results have been observed after intramyocardial transplantation of skeletal myoblasts into the heart of infarcted mice²⁴. Furthermore, the long-term ability of allogeneic MSCs to preserve function in the infarcted heart has been limited by transition from an immunoprivileged to an immunogenic state after differentiation²⁵.

Taking into consideration the challenges and prospects outlined above, we show here how to deliver cells by intramyocardial injection in mice. We observe that cells without cardiomyocyte contractile properties are not connected with the host myocardium and form a cohesive mass with a thin fibrotic barrier. Although in some cases this outcome may be advantageous, the following analysis may be useful to understand how cell engraftment may be modulated to generate functionally connected myocardial structures as well.

Protocol

All animal studies were performed in compliance with international (Directive 2010/63/EU of the European Parliament) and national (UK Home Office, Act 1986) regulations. The procedures herein described are part of our plan of work under UK license authorities and have not been undertaken for the purpose of recording.

1. Preparation of Cells

This protocol describes the preparation of a specific cell line (Human Embryonic Kidney, HEK293 cells) for demonstration purposes. Cell-specific protocols must be employed for growing and eventually differentiating pluripotent or adult stem cells into specific cell lineages.

1. Grow cells in DMEM media (high glucose, 4,500 mg/L) containing 1% sodium pyruvate, 2 mM glutamine, 1% antibiotic Pen/Strep and 10% Fetal Bovine Serum (FBS). One 10 cm plate should be normally split at 1:3 and media supplemented fresh every two days.
2. Treat cells mildly with trypsin 1x and resuspend in DMEM medium as described in 1:1.
3. Count cells in a hemocytometer chamber and collect 3 x 10⁶ cells into a separate tube.
4. Centrifuge in a swing bucket rotor at 100 g for 5 min.
5. Discard the supernatant and wash the cell pellet with sterile phosphate buffer 2x.
6. Resuspend in PBS 1x at a concentration of 10⁶/50 ul for injection into the left ventricle of immunodepleted mice.

2. Presurgical Conditions

1. Maintain immunodepleted mice (NOD.CB17-Prkdscid/JHliHSD) with sterile food pellets and water in a temperature-controlled isolator (22 °C; Table 1) before each surgery.
2. Perform surgery of immunodepleted mice under a laminar flow hood. Clean the working area with 70% Ethanol and autoclave surgical tools (forceps and scissors).
3. Turn on heating pads for the surgery and for the recovery. Heating pads are important to block the decrease of body temperature occurring during and post-surgery.
4. Place a clean and autoclaved cage on the recovery-heating pad.
5. Transport the mice in their original cages from the isolator to the surgery room into a sterile autoclaved bag.
6. Weigh the mouse and according to the body weight inject subcutaneously (s.c.) a solution containing medetomidine and ketamine hydrochloride diluted in 0.9%  sterile saline solution to anesthetize the mouse, as well as to relax the musculature (medetomidine 1 mg/kg; ketamine hydrochloride 75 mg/kg).
7. Remove the mouse from the cage when fully anesthetized (within 1-2 min, no toe-pinch reflex after stimulation)
8. Apply hair removal cream on the left side of the chest and the throat area. After an appropriate amount of time (approximately 2-5 min), wipe away the loose hairs with a water-wet tissue.
9. Place the mouse on the surgery panel in a supine position. From the surgeon's viewpoint the position is vertical, tail down-head up.
10. Inject analgesic solution at 0.1-0.2 μg/g diluted in sterile saline solution (buprenorphine) into the hind limb subcutaneously and cover the shaved area with the aseptic solution povidone iodine using a cotton tip applicator.
11. Focus the microscope (2.5X objective) on the throat area.

3. Surgical Conditions

1. With blunt scissors make a midline ventral skin incision of about 0.5-1 cm length slightly below the cricoid cartilage. Separate the skin from connective tissue to visualize the salivary glands.
2. Split the salivary glands on their natural midline division by simultaneously pulling each part sideward with forcep and magnetic chest retractor and expose the trachea.
3. Pull the tongue gently to the side to expose the larynx. Slide the intubation tube carefully into the trachea. The tube tip is visible through the displayed larynx and trachea. Importantly, if the tube is in, but not clearly visible, it is in the esophagus. Remove it and change the position of the tube tip.
4. Connect the tube to the ventilation machine. Set the stroke volume at 200 μl and 150 stroke/min. Secure the ventilation tubing on the surgery panel with a tape.
5. With blunt scissors and forceps, make a vertical tail to head 1- to 1.5-cm-long skin incision over the left thorax area.
6. Loosen the skin from the connective tissue/muscle layers and the major and minor pectoral muscles without making incisions.
7. Perform thoracotomy between the third and fourth ribs as follows:
   1. Perforate the intercostal muscle layer by pinching and scratching with small, rounded forceps.
   2. Once the intercostal muscle layer is perforated, place the chest retractor to maximally open the chest cavity. The upper and middle parts of the left ventricle (LV) with its overlying auricle (atrium) are now visible.
8. Prepare a precision syringe with cells. The syringe should be able to deliver an amount of volume of 10 ul for each injection. Cast a 30 G needle at the tip of the syringe.
9. Fix with tape a small plastic cannula (from an introcan-W 20 G) on the needle, leaving exposed only 1 mm including the open tip of the needle. This will prevent the needle from perforating the entire left ventricular wall and injecting the cells into the left ventricle cavity.
10. With the help of a 5X objective, visualize at this magnification the left ventricle and inject the cells into the left ventricular wall in 5 different locations (each injection will deliver 10 ul for a total of 106 cells injected). If the injection is successful, a white area and the absence of major backwash of the cells will be visible (if the injection was not successful, the animal would be excluded from functional analysis).
11. Remove the retractor. Close the thoracic wall by enclosing the two separated ribs with two stitches of 6-0 silk suture.
12. Moisturize the pectoral muscles and skin with a cotton tip drenched in PBS 1x and gently put the muscles back into the original position with the forceps.
13. Close the skin with 3-4 stitches of 6-0 silk suture. Suture the throat incision with 2-3 stitches to close the skin.
14. Inject atipamezole (1 mg/kg final concentration) to revert the anesthetic effects.
15. As soon as the mouse starts breathing autonomously, take out the tube from the trachea and put the mouse on its right side. The mouse is expected to recover within minutes.
16. When the mouse starts to turn and walk, place it in the warm recovery cage (small IVC cage with closed filtered top) for an hour.
17. Leave the animals overnight in a controlled heated box (37 °C).
18. Fill a dish with water and pellet food to support recovery during the first 2 days after surgery.

4. Post-surgery Analyses

Ten hearts injected with cells and 10 control not injected hearts are analyzed histologically as follows:

1. Perform analysis of the grafted cells 1 week after injection (Figures 1 and 3).
   1. After euthanasia by overdose of isoflurane, perfuse the hearts with 20 ml of 4% paraformaldehyde (PFA) to fix the tissue. Dehydrate the fixed tissue in increasing percentages of ethanol (from 70-100%) and embed in paraffin blocks.
   2. Section the hearts in paraffin with a microtome at 5 μm and stain with hematoxylin and eosin for morphological analysis.
   3. Visualize connective tissue by using Masson's Trichrome stain.
   4. Analyze the images under a slide scanner microscope (Figures 1A, 2A, 3A, 3B, and 3C) and visualize the entire heart in short axis view using a digital slide scanner microscope (Figures 1B and 1C).
2. After the above analyses, deparaffinize the heart tissues in xylene, rehydrate by submerging specimens in decreasing percentages of ethanol (from 100% to water) and process for Scanning Electron Microscopy (SEM) as follows:
   1. Incubate deparaffinized blocks with 1% tannic acid in 0.1 M phosphate buffer for 1 hr.
   2. Dehydrate specimens in increasing percentages of ethanol (from 25-100%).
   3. Dry samples with HMDS (hexamethyldisilazane).
   4. Mount the specimens on Stubbs with Acheson Silver Dag and coat them with gold-palladium.
   5. View samples in an analytical scanning electron microscope (Figure 2B).

Results

We injected HEK293 cells, which are distinguishable from the heart cells by their different morphology (Figure 1), with a cobblestone shape compared to the elongated cardiomyocytes (Figure 1A). HEK293 cells were more reactive to hematoxylin dye (blue color) compared to cardiomyocytes (pink color), likely because of their increased nuclear content (Figure 1A). To further distinguish the injected cells from the host tissue, HEK293 cells were labeled with DAPI 2 hours prior...

Discussion

In this manuscript, we have shown how to perform intramyocardial injection of cells in murine hearts. As a proof of this methodology, we have used HEK293 cells. It is important to emphasize that HEK293 cells are not used in any cell therapy study and therefore the findings of this manuscript are not appropriate for direct translation to a therapeutic approach. However, the fact that HEK293 cells are not contractile cells and do not transdifferentiate in other cell types focuses attention on technical aspects such as the ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Isolator	Pfi systems		Quotation needed
Heating Pad	Vet Tech Solutions	HE006	For small animals
Medetomidine	National Veterinary Service		Veterinary prescription is necessary
Ketamine hydrochloride	National Veterinary Service		Veterinary prescription is necessary
Atipamezole	National Veterinary Service		Veterinary prescription is necessary
Hair removal cream	commercial shops
Buprenorphine	NVS		Veterinary prescription is necessary
Leica MZFLIII microscope	Leica	Model S6E	With swing arm stand TS0
Hamamatsu Nanozoomer digital slide scanner	Hamamatsu	RS series
Scanning Electron Microscope	Jeol	JSM-6610
Blunt scissors	FST	14084-09
Minivent	Harvard Apparatus	73-0043	Including small Y adapter (73-0027) and intubation cannula (73-2844)
Forceps	FST	11052-10
Retraction system	FST	18200-20	Kit for animals up to 200 g
30 G 12 mm; ½ inch	BBraun	A210	Fine yellow
Microliter syringe	ESSLAB	81201	Also include a Hamilton repeating dispenser PB 600-1 Catalog number 83700
6-0 Silk suture	Ethicon	W1614T