気道抵抗測定とマウスにおける気管支肺胞洗浄液のサンプリングのためのリバーシブル、非侵襲的方法

Prior to starting this procedure, miser anesthetized with tomate injected intraperitoneal. Animals are then placed on a table and a 20 gauge catheter is inserted orally into the trachea. Intubated mice are placed on a mechanical ventilator and an intravenous line is started in the tail vein.

The mice are then enclosed in a rodent SIGGRAPH attached to two pressure transducers. The electronics enable the continuous measurement of respiratory system resistance during acetylcholine challenge. After disconnecting the ventilator and intravenous line, the tracheal catheter is advanced into the left lung, which is lavage with saline injected and withdrawn through the catheter.

Hi, I am Simone Phar from the Department of Medicine at Baylor College of Medicine. Today we are going to show you the oral intubation of mice such that airway mechanics and the broncho velar lage fluid can be collected repeatedly. We use this procedure to study experimental allergic disease, which is an experimental model of asthma, and this procedure can be adapted to other applications too.

So let's get started. To begin this procedure, the mouse is deeply anesthetized by an intraperitoneal injection of 48 milligrams per kilogram of ate. The mouse is then placed in a soft light type box to keep it calm.

A clean mouse cage covered with a cloth drape can be used for this purpose After five to 10 minutes confirmed by a toe pinch that the animal is fully anesthetized and then place it in the recumbent position. Ventral side up on a small table adjusted to a 45 degree angle. To secure the subject into place, insert a rubber band ENC circling the table behind the top row of teeth.

A heat lamp should be used to keep the mouse warm with your right hand. Use tweezers to grip, extend and lift the tongue from the mouth. Then gently insert a metal tongue depressor held in your left hand.

This will allow for an unimpeded airway and visualization of the vocal cords for intubation. Next, insert a 0.8 millimeter diameter fiber optic thread connected to a light source through the angio catheter and extend it 10 millimeters beyond the tip. While studying the depressor with your left hand, use your right hand to guide the illuminated end of the fiber optic thread through the oral cavity and pharynx until the vocal cords are visualized.

Then under direct visualization and timing it with when the cords are maximally open, pass the thread through the moving vocal cords and into the trachea. Now pass the angio catheter over the fiber optic thread into the trachea until the catheter tip lies within the mid portion of the tra trachea. For a 17 to 22 gram mouse, this corresponds to a 10 millimeter catheter segment remaining outside of the mouth.

Remove the fiber optic thread to confirm successful intubation. Observe regular deep breaths that immediately terminate following occlusion of the connector with the thumb. Lower the plethysmograph table until it is parallel with the work bench and turn the subject 180 degrees until it is facing the air ventilator port.

Turn the animal on its side before connecting to the ventilator. Secure an airtight connection and activate the ventilator. A successful intubation is further confirmed when the raso abdominal excursion is seen to pace with the ventilator later.

To prepare the needle for the intravenous line, remove a 10 millimeter 27 gauge needle from its syringe connector and bend it 90 degrees at the midpoint so that the bevel faces into the angle. Connect the non beveled end to the PE 10 tubing leading to the IV injection port. To prevent potentially fatal air embolization.

Purge the tubing and needle with 37 degrees Celsius 0.9%sodium chloride via the one milliliter syringe. The injection port consists of a 27 gauge needle pushed through a hole drilled into the cap of a 15 milliliter centrifuge tube. The cap is filled with saline such that the end of the needle is constantly submerged, thus reducing the likelihood the air will be entrained in the needle and injected intravenously.

Align the needle at the codal extreme of the tail. Parallel to and over the lateral vein. Run the needle slightly beneath the skin while directing it.

Cranial along the veins length and pushing subcutaneously to the bend. To confirm successful IV placement, pull the syringe plunger slightly. You should see blood backflow into the IV tubing.

Furthermore, there should be unimpeded flow through the IV line upon injection of 50 to 100 microliters of saline into the tail vein. After removing the heat lamp from the setup, enclos the subject in the plasmo graph and secure it as airtight with the application of four clamps. Removal of the heat lamp is important to prevent heating the air in the plasmo graph chamber and potentially altering subsequent measurements of respiratory system resistance or RRS respiratory system resistance or RRS is determined by continuous quantitation of the quotient.

Change in airway pressure over airflow or delta P over V at points of equal lung volume. Delta P is determined by using a pressure transducer connected to the tracheal angio catheter V.The differential of plasm graft volume over time is computed by the preamp module. After establishing a stable baseline, RRS inject five successive doses of increasing concentrations of acetylcholine chloride or a CH over one second via the iv.

Each subsequent dose is administered upon return of RRS to baseline until a 200%increase in airway resistance is achieved. Remove the IV from the tail vein and disconnect the mouse from the ventilator. Maintaining a patent airway by keeping the tracheal cannula in place, make sure the mouse has resumed spontaneous breathing.

If not, respiration can be encouraged by gently massaging the thorax. Once the mouse has resumed spontaneous breathing, transfer it with tracheal cannula in place to a chamber purged with 100%oxygen and maintained at 37 degrees Celsius with the heat lamp. The tracheal cannula must remain in place until the animal is awake to prevent asphyxiation related death caused by acetylcholine induced hypersalivation.

Bronchoalveolar lavage or BAL fluid can only be collected when the mouse has sufficiently recovered its gag reflex, which should occur about 20 minutes at replacement in the recovery chamber. To assess the gag reflex gently slide the angio catheter inward and outward obvious coughing or struggling indicates that the gag reflex has returned to collect BAL fluid. A metallic intubation guide wire with a continuous bend of about 30 degrees directed to the left lobe of the lung is inserted into the angio catheter.

Advance the G wire and angio catheter together into the left lobe of the lung such that the catheter hub excluded extends beyond the front teeth by only one millimeter. Make sure the tip of the G wire does not pass through the open end of the angio catheter. To avoid tracheal laceration or rupture while keeping the angio catheter in place, flush 300 microliters of sterile PBS pH 7.4 into the left lung via a one milliliter syringe immediately after while drawing up the syringe plunger.

To create negative pressure, remove the angio catheter slowly while intensely massaging the lung. A BAL return of 100 to 200 microliters is expected. Immediately return the lavage mouse to the 37 degree Celsius, 100%oxygen chamber.

Place the animal on its left side until fully recovered about 20 minutes and then place it back into its cage. We present here representative results of airway resistance measurements, airway hyperreactivity defined as RRS values that are significantly higher at any acetylcholine dose compared to naive values developed after five allergen challenges at the fourth dose. However much greater airway hyperreactivity was seen after the sixth challenge with no further increase in responsiveness.

At the seventh challenge, a CH dosing was stopped after RRS values tripled from baseline values further increases in dosing otherwise result in lethal bronchospasm. Mice repeatedly challenged with vehicle intranasally to not develop airway hyperreactivity and at all doses of a CH given RRS measurements do not significantly vary from baseline values as shown by these representative real-time RRS tracings from a naive and six x allergen challenged mouse receiving consecutive IV doses of a CH prior to the emergence of robust A HR.The dominant cell tape of the airway induced by allergen was the neutrophil similar to the trend for a HR.However, eosinophilia gradually strengthened with repeated allergen challenge and the eosinophil became the numerically dominant cell type in BAL fluid. After the sixth challenge coincident with marked decline in neutrophil numbers.

In control PBS challenged mice. Airway resistance measurements also did not vary significantly over time. Enhanced macrophage in neutrophil but not eosinophil recruitment to BAL fluid was also seen in these mice similar to those changes observed in mice receiving only repeated BAL fluid sampling.

So we have just shown you how to reversibly intubate mice orally and access the intravenous line to collect airway physiology data and bronchoalveolar large fluid. Before performing these techniques on live animals, it's essential to practice and perfect each technique separately. On cric animals.

A strict and gentle handling is essential and strict aseptic technique must be performed at all times. So that's it. Thanks for watching and good luck with your experiments.