Intramyocardial Cell Delivery: Observations in Murine Hearts

The overall goal of this procedure is to deliver a specific amount of cells in the left ventricular myocardium of a murine heart, and to be able to visualize the I injected cells one week after injection. The first step is to prepare the selected cells by incubating them with DPI for two hours. Next, a thoracotomy through the left fourth intercostal space is performed to expose the heart.

The cells are then loaded in a syringe equipped with a small plastic cannula to avoid perforation of the ventricular wall. The final step is to inject the correct amount of cells into the wall of the left ventricle. Ultimately, histology and scanning electron microscopy images are used to show the presence of the injected cells in the left myocardium one week after injection.

The main advantage of this technique over existing methods is that you avoid injecting the cells into the left ventricular cavity and that you can trace the cells by staining them with dapi. The HC 2 93 cells used to demonstrate this procedure are grown in DMM media and are provided with fresh media every two days To facilitate cell tracking, incubate the cells with DPI for two hours before injecting immuno depleted. Mice are kept in a temperature controlled isolator with sterile food pellets and water before each surgery.

To begin position a heating pad in the work area to maintain the correct body temperature During surgery, the working area is set up under a laminar flow hood and cleaned with 70%ethanol. Arrange the autoclave sterilized surgical tools needed for the procedure. Next, prepare the recovery cage by placing a clean autoclave cage on a heating pad.

When ready, transport the mice in their original cages from the isolator to the surgery room in a sterile autoclaved bag. After administering an anesthetic, confirm sedation by the absence of the hind limb withdrawal reflex. Use hair removal cream to remove the fur from the left side of the chest and the throat area.

Next, place the mouse on the surgery panel in a supine position. After injecting an analgesic solution, subcutaneously, cover the shaved area with the aseptic solution. While viewing the area under a microscope, make a midline incision slightly below the cricoid cartilage.

Break through the connective tissue to visualize the salivary glands. Split the salivary glands at their natural midline by pulling them apart with forceps. Next, insert two chest retractors.

To expose the trachea, expose the larynx by pulling the tongue gently to one side, and then slide a 22 gauge metal intubation tube carefully into the trachea. If properly inserted, the tube tip should be visible through the trachea. Set the stroke volume at 200 microliters and the stroke per minute.

At 130, adjust tidal volume and respiratory frequency depending on the weight of the animal. Next, make an incision over the left thoracic area. Carefully separate through the connective tissue and major and minor pectoralis muscles.

To visualize the rib cage, perforate the intercostal muscle layer between the third and fourth ribs with small rounded forceps, and insert a chest retractor. To open the chest cavity. Visualize the upper and middle parts of the left ventricle with its overlying atrium.

At this point, the cells needed for injection should be prepared in a syringe with a 30 gauge needle to prevent the needle from perforating the left ventricular wall tape a small plastic cannula over the knee needle, leaving only one millimeter exposed. With the help of a five x objective, visualize the left ventricle and inject the cells into the ventricle wall. Slowly withdraw the needle to prevent the backwash of cells.

Inject cells into five different locations. If the injection is successful, a white area will be visible at the injection location. After injecting the cells, remove the retractor and close the thoracic wall with two silk sutures.

Mo the pectoral muscles and skin with a cotton tip drenched in PBS and gently put the muscles back into the original position with forceps. Next, close each of the incision sites with silk sutures. Once the wounds are closed, inject anti sedan to revert the anesthetic effects.

When the mouse starts breathing autonomously, remove the tube from the trachea and put the mouse on its right side. When the animal is ambulatory, place it in a warm recovery cage and monitor until fully recovered. One week after transplantation, heart samples were processed for hemat, toin and eoin staining.

To confirm that hec 2 9 3 cells were present in the ventricle wall, tri chrome stained tissues show that the cells were grouped in the same area where they were injected control. Sham operated mice showed no damage in the whole heart and integrity of the tissue. DPI labeling of the cells before injection helped to further distinguish between the injected cells and the host tissue.

The injected cells were surrounded by a thin layer of fibrotic tissue, which appeared as a blue area in the tri chrome. Staining cells were not connected with the host myocardial tissue as shown by energy dispersion spectrometry likely due to their non contractile function and different cellular structure. The arrows point to small areas of damage where the needle was injected.

Injected cells denoted by the green arrow were present in the proximity of the injured area marked by black arrows. In addition, DPI positive cells were visible along the site of injection. The tunnel assay did not show increased apoptotic cells at the site of injury suggesting that necrosis rather than apoptosis had occurred after injection.

After watching this video, you should have a good understanding of how to successfully inject cells into the myocardium of an anesthetized mouse without puncturing into the ventricular cavity.