A Mouse Model of Orotracheal Intubation and Ventilated Lung Ischemia Reperfusion Surgery

Podsumowanie

A mouse surgical model to create left lung ischemia reperfusion (IR) injury while maintaining ventilation and avoiding hypoxia.

Streszczenie

Ischemia reperfusion (IR) injury frequently results from processes that involve a transient period of interrupted blood flow. In the lung, isolated IR permits the experimental study of this specific process with continued alveolar ventilation, thereby avoiding the compounding injurious processes of hypoxia and atelectasis. In the clinical context, lung ischemia reperfusion injury (also known as lung IRI or LIRI) is caused by numerous processes, including but not limited to pulmonary embolism, resuscitated hemorrhagic trauma, and lung transplantation. There are currently limited effective treatment options for LIRI. Here, we present a reversible surgical model of lung IR involving first orotracheal intubation followed by unilateral left lung ischemia and reperfusion with preserved alveolar ventilation or gas exchange. Mice undergo a left thoracotomy, through which the left pulmonary artery is exposed, visualized, isolated, and compressed using a reversible slipknot. The surgical incision is then closed during the ischemic period, and the animal is awakened and extubated. With the mouse spontaneously breathing, reperfusion is established by releasing the slipknot around the pulmonary artery. This clinically relevant survival model permits the evaluation of lung IR injury, the resolution phase, downstream effects on lung function, as well as two-hit models involving experimental pneumonia. While technically challenging, this model can be mastered over the course of a few weeks to months with an eventual survival or success rate of 80%-90%.

Wprowadzenie

Ischemia reperfusion (IR) injury can occur when blood flow is restored to an organ or tissue bed after some period of interruption. In the lung, IR can occur in isolation or in association with other injurious processes such as infection, hypoxia, atelectasis, volutrauma (from high tidal volumes during mechanical ventilation), barotrauma (high peak or sustained pressures during mechanical ventilation), or blunt (non-penetrating) lung contusion injury^1,2,3. There remain several gaps in our knowledge about the mechanisms of LIRI and the impact of concurrent processes (e.g., infection) on LIRI outcomes, and also the treatment options for LIRI are limited. An in vivo model of pure LIRI is required to identify the pathophysiology of lung IR injury in isolation and to study its contribution to any multi-hit process of which lung injury is a component.

Murine lung IR models can be used to study the lung-specific pathophysiology of multiple processes, including lung transplantation³, pulmonary embolism⁴, and lung injury following hemorrhagic trauma with resuscitation⁵. Currently used models include surgical lung transplantation⁶, hilar clamping⁷, ex vivo lung perfusion⁸, and ventilated lung IR⁹. Here, we provide a detailed protocol for a murine ventilated lung IR model of sterile lung injury. There are multiple benefits of this approach (Figure 2), including that it induces minimal hypoxia and minimal atelectasis, and it is a survival surgery model that allows for long-term studies.

Reasons to choose this model of LIRI over other models such as the hilar clamping and ex vivo perfusion models are the following: this model minimizes the inflammatory contributions of atelectasis, mechanical ventilation, and hypoxia; it preserves cyclical ventilation; it maintains an intact in vivo circulatory immune system that can respond to the IR injury; and finally, as a survival procedure, it permits the longer-term analysis of the mechanisms of secondary injury generation (2-hit models) and injury resolution. Overall, we believe this ventilated lung IR model provides the "purest" form of IR injury that can be studied experimentally.

Other publications have described the use of orotracheal intubation of mice to perform IT injections or installations^10,11, but not as the starting point for a survival surgery as it is in this model. The placement of an orotracheal tube permits the performance of lung surgery by allowing the collapse of the operative lung. It also allows for the reinflation of the lung at the end of the procedure, which is critical for the pneumothorax and for the ability of the mouse to return to spontaneous ventilation at the conclusion of the procedures. Finally, the removal of the secured orotracheal tube is a simple procedure that, unlike an invasive tracheotomy, is compatible with a survival surgery. This allows for longer term research studies focused on understanding the progression and resolution of LIRI and associated disorders, as well as the creation of chronic injury models.

Protokół

All procedures and steps described below were approved by the institutional animal care and use committee (IACUC) at the University of California San Francisco. Any mouse strain can be used, though some strains have a more robust lung IR inflammatory response compared to others¹². Mice that are approximately 12-15 weeks of age (30-40 g) or older tolerate and survive the lung IR surgery better than younger mice. Both male and female mice can be used for these surgeries.

1. Mouse Intubation Protocol

1. Anesthesia and preparation for intubation
   1. Wipe the mouse abdomen with an ethanol swab. Anesthetize the mouse with an intraperitoneal injection of tribromoethanol (250-400 mg/kg). Assess the appropriate depth of anesthesia by the lack of pedal withdrawal reflex. Place eye lubricating ointment now or later (step 2.1.4).
      NOTE: For this procedure, tribromoethanol (and etomidate as an alternative option) provides a stable anesthetic plane without affecting the hemodynamic conditions required for this surgery. This anesthetic is only used once to avoid the risk of peritoneal adhesions. Isoflurane could also be used, but we do not use it here. The practitioner is free to use whatever anesthetic recipe they see fit.
   2. Place the anesthetized mouse on an intubation stand or plastic support in a supine position, suspended by its upper incisors on looped 4-0 sutures (silk or other) across two support anchors.
   3. To keep the mouse immobilized during the intubation procedure, loosely tape the lower part of the chest (or both upper limbs) to the platform.
   4. Place the fiberoptic flexible light gently on the trachea of the mouse, slightly below the vocal cords. Adjust the level of illumination so that only a dark field is visible when looking into the mouse oropharynx except for red light emanating from below the vocal cords, demonstrating the target for the eventual placement of the endotracheal tube. Note that vocal cord movements should be visible with the naked eye or, if needed, under magnification.
2. Intubation procedure
   1. Hold the tweezers with the dominant hand and use them to gently grip and draw the tongue out of the oral cavity.
   2. Open the lower jaw using forceps held by the non-dominant hand, and then push the forceps into the larynx to gently lift the epiglottis. At this time, release the tongue from the tweezers.
   3. Look for the vocal cords. They should open and close according to each breath. Holding the cannula with the guide wire pre-loaded, insert the tip of the wire through the vocal cords.
   4. Being very careful not to move the wire by holding a portion of it that is outside the cannula but just above the vocal cords, withdraw the cannula, leaving just the wire in place with its distal end within the trachea.
   5. At this point, perform a second visualization of the vocal cords to confirm that the wire distal tip remains passed through the illuminated vocal cords and into the trachea, and is not in the unlit esophagus.
   6. Hold the wire outside the mouth with the curved forceps in the left hand, stabilized against a hard surface, and carefully advance the 20G catheter with tape wings over the wire.
   7. Once the distal end of the wire emerges from the back end of the 20G catheter or endotracheal tube, hold that end with the curved forceps and smoothly advance the 20G catheter into the trachea.
   8. Carefully remove the wire from the distal end of the 20G catheter with the curved forceps without dislodging the placement of the catheter.
   9. Briefly connect the catheter to the ventilator before securing it to confirm proper placement into the trachea and not the esophagus. Confirm tracheal placement by observation of mechanical ventilation-dependent bilateral chest wall movements and the absence of inflation of the stomach.
3. Post-intubation
   1. Disconnect the catheter from the ventilator. Fix the tape wings (attached to the catheter) through the lower lip of the mouse using a 4-0 vicryl suture to firmly secure the endotracheal tube (ETT) to the mouse during all subsequent procedures/manipulations.
      NOTE: Alternatively, silk tape or other tape can be used to secure the ETT, however care should be taken to avoid dislodgement of the ETT during movement of the animal from the intubation sled to the surgical surface.
   2. Carefully remove the mouse from the intubation sled. Briefly connect the catheter to the ventilator set at a tidal volume 0.2-0.225 mL and a respiratory rate of 120-150 breaths per min to confirm correct tracheal placement of the orotracheal tube and then disconnect with the mouse breathing spontaneously through the orotracheal tube.
   3. Do not leave the animal unattended from this point onward until it has regained sufficient consciousness to maintain sternal recumbency at the end of the procedure.

2. Lung ischemia and reperfusion (IR) surgery protocol

1. Analgesia and preparation of the surgical site
   1. Wipe the mouse abdomen with an ethanol swab and inject buprenorphine (0.05-0.1 mg/kg) intraperitoneally.
   2. Shave the hair over the left thorax area up to the left scapula. Remove excess shaved hair using alcohol swabs.
      NOTE: Steps 2.1.1 and 2.1.2 can also be performed before intubation if there concern for dislodgement of the ETT when secured with silk tape.
   3. Place the mouse on a warming pad in a left lateral or 3/4 turned position and connect the tracheal tube on the ventilator with a tidal volume of 0.2-0.225 mL (~8 mg/kg) and a respiratory rate of 120-150 breaths per min. Do not use supplemental oxygen for this procedure.
   4. Apply eye lubricant with a sterile cotton-tip swab. Turn the mouse to 3/4 left side up and immobilize all four limbs and the tail with laboratory tape.
   5. Disinfect the shaved skin area and surrounding fur with povidone-iodine and wait for the solution to dry. Then cover the surgical field with a sterile drape or clear plastic film and create a rectangular opening in the drape or plastic film for the surgical field.
2. Surgical procedure
   1. Confirm the appropriate level of anesthesia (provided by the administration of tribromoethanol and buprenorphine as described earlier) by testing response to toe pinch.
   2. Using a pair of sharp scissors and a pair of larger forceps (narrow pattern forceps or similar), make a 2 cm transverse skin incision below the inferior angle of the scapula in the left lateral thorax. Use the scissors and a finer pair of forceps (extra fine graefe forceps or similar) to cut into the muscular layer and dissect down to the ribs.
   3. Identify the second intercostal space and hold the second rib with the extra fine forceps. Pulling the rib upward, use a sterile #11 or #12 (curved) scalpel blade (no handle necessary) to enter the pleural space by separating and cutting across the 2^nd-3^rd space's intercostal muscles. Consider pausing ventilation to reduce injury to the left lung apex.
   4. Insert three sterilized retractors. Use the smallest/narrowest retractor cephalad along the orientation of the ribs, the medium size retractor to the left along the 2^nd rib, and the largest retractor to the right along the surface of the 3^rd rib.
   5. Open the chest with slow and progressive retraction using the elastic retractor cords. Expose and identify the left pulmonary artery (PA) by moving the left lung apex away with a sterile cotton-tip swab.
   6. Use the micro forceps, ultrafine forceps in the right hand and PA or vessel dilating forceps in the left hand, to gently expose and create the field in which the left PA and bronchus are both visible.
   7. Using the PA forceps, pick up the left PA and pull gently but firmly upward and cephalad to visualize the transparent bronchus below. Increase magnification on the dissection microscope (see equipment list for more details) at this point to maximum (2x).
      NOTE: Sterilize all equipment before use. Additionally, to maintain sterility, only the tips of surgical instruments should enter the sterile surgical field.
   8. While retracting the PA away from the bronchus, carefully pass the closed ultrafine forceps through the space between the left PA and bronchus. Then, use these forceps to hold and pull a 7-0 or 8-0 prolene suture through the space between the left pulmonary artery (above) and bronchus (below).
   9. Encircle the left PA by tying a slipknot to create an occlusion in the PA. Blood flow interruption is easily visualized under the microscope. This marks the initiation of the ischemic period.
   10. Externalize the free end of the knot through a different entry point in the anterior left thorax using a 24G-28G needle and secure the end of the suture with a small piece of tape for easier identification later on.
   11. Reinflate the lung to expel as much air out of the chest cavity as possible using a PEEP valve/tubing on the rodent ventilator. Then, close the ribcage with two interrupted 4-0 nylon sutures.
   12. Close the muscle and subcutaneous layer with a running 4-0 nylon suture. Then apply two or three drops of topical bupivacaine (0.5%) to the incision. Use a 4-0 nylon suture to close the skin layer with a running suture.
3. Post-operative care
   1. When spontaneous ventilation has resumed, disconnect the endotracheal tube from the ventilator and extubate the mouse.
   2. Place the mouse on the warming pad to maintain body temperature during early post-anesthesia recovery.
   3. Carefully monitor the mouse while recovering from general anesthesia. Pull the externalized slipknot gently at the end of the ischemic period (30 min or 1 h).
   4. Move the mouse from the warming pad to a cage once it has exhibited signs of recovery: self-righting and/or movement.
   5. After the period of reperfusion (1 h or 3 h), euthanize the animal and collect blood by cardiac puncture and lung tissue for further analysis. For 1 h reperfusion, collect plasma for ELISA, tissue for RNA, and protein analysis; for 3 h reperfusion, additionally collect tissue for histology.

Wyniki

Inflammation generated by unilateral ventilated sterile lung ischemia reperfusion (IR) injury: Following 1 h of ischemia, we observed increased levels of cytokines in the serum and within the lung tissue by both ELISA and qRT-PCR that peaked at 1 h following reperfusion and rapidly returned to baseline within 12-24 h after reperfusion¹³. For samples collected at 3 h following reperfusion, we observed intense neutrophil infiltration within the left lung tissue and noted that the intensity of the in...

Dyskusje

This manuscript details the steps involved in performing the ventilated lung IR model developed by Dodd-o et al.⁹. This model has helped identify molecular pathways involved in the generation and resolution of inflammation from lung IR in isolation^14,15,16,17, lung IR in combination with co-existing infection¹⁸, and lung IR in relation to the gut-...

Materiały

Name	Company	Catalog Number	Comments
Equipment
Fiber Optic Light Pipe	Cole-Parmer	UX-41720-65	Fiberoptic light pipe
Fiber Optic Light Source	AmScope	SKU: CL-HL250-B	Light source for fiberoptic lights
Germinator 500	Cell Point Scientific, Inc.	No.5-1450	Bead Sterilizer
Heating Pad	AIMS	14-370-223	Alternative option
Lithium.Ion Grooming Kits(hair clipper)	WAHL home products	SKU 09854-600B	To remove mouse hair on surgical site
Microscope	Nikon	SMZ-10	Other newer options available at the company website
MiniVent Ventilator	Havard Apparatus	Model 845	Mouse ventilator
Ultrasonic Cleaner	Cole-Parmer	UX-08895-05	Clean tools that been used in operation
Warming Pad	Kent Scientific	RT-0501	To keep mouse warm while recovering from surgery
Weighing Scale	Cole-Parmer	UX-11003-41	Weighing scale
Surgery Tools
4-0 Silk Suture	Ethicon	683G	For closing muscle layer
7-0 Prolene Suture	Ethicon Industry	EP8734H	Using for making a slip knot of left pulmonary artery
Bard-Parker (11) Scalpel (Rib-Back Carbon Steel Surgical Blade, sterile, single use)	Aspen Surgical	372611	For entering thoracic cavity (option 1)
Bard-Parker (12) Scalpel	Aspen Surgical	372612	For entering thoracic cavity (option 2)
Extra Fine Graefe Forceps	FST	11150-10	Muscle/rib holding forceps
Magnetic Fixator Retraction System	FST	1. Base Plate (Nos. 18200-03) 2. Fixators (Nos. 18200-01) 3. Retractors (Nos. 18200-05 through 18200-12) 4. Elastomer (Nos.18200-07) 5. Retractor(No.18200-08)	Small Animal Retraction System
Monoject Standard Hypodermic Needle	COVIDIEN	05-561-20	For medication delivery IP
Narrow Pattern Forceps	FST	11002-12	Skin level forceps
Needle holder/Needle driver	FST	12565-14	for holding needles
Needles	BD	305110	26 gauge needle for externalizing slipknot (24 or 26 gauge needle okay too)
PA/Vessel Dilating forceps	FST	00125-11	To hold PA; non-damaging gripper
Scissors	FST	14060-09	Used for incision and cutting into the muscular layer durging surgery
Ultra Fine Dumont micro forceps	FST	11295-10 (Dumont #5 forceps, Biology tip, tip dimension:0.05*0.02mm,11cm)	For passing through the space between the left pulmonary artery and bronchus
Reagents
0.25% Bupivacaine	Hospira, Inc.	0409-1159-02	Topical analgesic used during surgical wound closure
Avertin (2,2,2-Tribromoethanol)	Sigma-Aldrich	T48402-25G	Anesthetic, using for anesthetize the mouse for IR surgery, the concentration used in IR surgery is 250-400 mg/kg.
Buprenorphine	Covetrus North America	59122	Analgesic: concentration used for surgery is 0.05-0.1 mg/kg
Eye Lubricant	BAUSCH+LOMB	Soothe Lubricant Eye Ointment	Relieves dryness of the eye
Povidone-Iodine 10% Solution	MEDLINE INDUSTRIES INC	SKU MDS093944H (2 FL OZ, topical antiseptic)	Topical liquid applied for an effective first aid antiseptic at beginning of surgery
Materials
Alcohol Swab	BD brand	BD 326895	for sterilzing area of injection and surgery
Plastic film	KIRKLAND	Stretch-Tite premium	Alternative for covering the sterilized surgical field (more cost effective)
Rodent Surgical Drapes	Stoelting	50981	Sterile field or drape for surgical field
Sterile Cotton Tipped Application	Pwi-Wnaps	703033	used for applying eye lubricant
Top Sponges	Dukal Corporaton	Reorder # 5360	Stopping bleeding from skin/muscle