Remotely Triggered LAD Occlusion Using a Balloon Catheter in Spontaneously Breathing Mice

Summary

This article presents a unique closed-chest technique for inducing myocardial ischemia-reperfusion injury (IRI) in mice. The presented method allows mice to breathe spontaneously while remotely inducing myocardial ischemia. This provides access to the animal for studying the dynamic processes of ischemia and reperfusion in situ and in real-time via noninvasive imaging.

Abstract

Acute myocardial infarction (AMI) is a prevalent and high-mortality cardiovascular condition. Despite advancements in revascularization strategies for AMI, it frequently leads to myocardial ischemia-reperfusion injury (IRI), amplifying cardiac damage. Murine models serve as vital tools for investigating both acute injury and chronic myocardial remodeling in vivo. This study presents a unique closed-chest technique for remotely inducing myocardial IRI in mice, enabling the investigation of the very early phase of occlusion and reperfusion using in-vivo imaging such as MRI or PET. The protocol utilizes a remote occlusion method, allowing precise control over ischemia initiation after chest closure. It reduces surgical trauma, enables spontaneous breathing, and enhances experimental consistency. What sets this technique apart is its potential for simultaneous noninvasive imaging, including ultrasound and magnetic resonance imaging (MRI), during occlusion and reperfusion events. It offers a unique opportunity to analyze tissue responses in almost real-time, providing critical insights into processes during ischemia and reperfusion. Extensive systematic testing of this innovative approach was conducted, measuring cardiac necrosis markers for infarction, assessing the area at risk using contrast-enhanced MRI, and staining infarcts at the scar maturation stage. Through these investigations, emphasis was placed on the value of the proposed tool in advancing research approaches to myocardial ischemia-reperfusion injury and accelerating the development of targeted interventions. Preliminary findings demonstrating the feasibility of combining the proposed innovative experimental protocol with noninvasive imaging techniques are presented herein. These initial results highlight the benefit of utilizing the purpose-built animal cradle to remotely induce myocardial ischemia while simultaneously conducting MRI scans.

Introduction

Acute myocardial infarction (AMI), a prevalent global cardiovascular condition, is associated with high mortality rates and morbidity¹. Despite technological advancements that enabled early and effective revascularization strategies for AMI patients, patients still experience myocardial ischemia-reperfusion injury (IRI) following these interventions². Therefore, understanding the fundamental mechanisms and formulating approaches to mitigate IRI is crucial. IRI represents a complex pathophysiological state involving a multitude of intricate biological processes. These encompass regulated cell death, oxidative stress responses, inflammation, wound healing, fibrosis, and ventricular remodeling. Animal models, such as mice, have been of great importance for IRI research and are extensively employed due to their cost-effectiveness, rapid breeding, and the wealth of mechanistic information from transgenic models³.

This protocol presents an innovative technique to remotely induce IRI in spontaneously breathing mice with a closed chest, more closely mimicking the human pathology. Remote occlusion was achieved by utilizing a balloon catheter positioned at a distance from the myocardium to occlude the left anterior descending artery (LAD). This approach offers continued access to the beating heart at the exact moments of occlusion and reperfusion, facilitating simultaneous imaging modalities, including ultrasound or magnetic resonance imaging (MRI). These noninvasive methods could provide crucial insights into tissue responses before, during, and after occlusion and reperfusion events.

Various surgical techniques exist to induce IRI in murine models. Surgical trauma stemming from thoracotomy in open-chest models of coronary ligation triggers an immune response impacting diverse mechanisms associated with ischemia and reperfusion. Activation of the innate immune system holds the potential to influence the extent of myocardial infarction⁴. The proposed adapted technique provides a potential approach for exploring myocardial pre- and postconditioning and possibly reducing the impact of the innate immune response to surgical trauma in murine IRI studies by minimizing the open thorax timeframe to a maximum of 5 min. Moreover, the newly developed technique might also contribute to reducing the pro-inflammatory sequelae caused by ventilator-induced lung injury⁵. However, the combined effects of this new closed chest method would require further in-depth investigation.

Thorough validation of the proposed technique was conducted by comparing it with the traditional method, which involves exposing the LAD through surgical chest dissection and ligature-induced occlusion for 30 min. Results from both techniques were compared, which included troponin measurements reflecting cardiac infarct size, assessments of the area at risk using MRI with gadolinium contrast enhancement, histological triphenyl tetrazolium chloride (TTC) staining, and the determination of final scar sizes via Sirius Red (SR) staining. The outcome demonstrates the robustness and efficacy of the proposed approach for studying ischemia-reperfusion injury in murine models.

Protocol

This animal protocol was approved by and is in accordance with the guidelines and regulations set forth by the Ethical Committee for Animal Experimentation (ECD) at The Catholic University of Leuven. All policies developed by the local ECD follow the regulations of the European Union concerning the welfare of laboratory animals as declared in Directive 2010/63/EU. All tests described below have been performed on 8-12 week-old (body weight = 20-23 g) male C57BL/6 mice. The acute experiments represent animals that were immediately sacrificed after surgery. The survival experiments represented a four-week follow-up, including troponin measurement, MRI, and SR staining.

1. Anesthesia and endotracheal intubation

1. Administer pre-anesthesia medication using a mix of ketamine (7.5 mg/kg; Nimatek 100 mg/mL), xylazine (1 mg/kg; Rompun, 20 mg/mL), and acepromazine (0.1 mg/kg; Placivet, 10 mg/mL) (KXA) (see Table of Materials) with 0.9 % saline diluent intraperitoneally. This allows a smoother and less invasive intubation.
2. Place the mouse in an induction chamber (see Table of Materials) and invoke anesthesia using 2.5% isoflurane in pure oxygen with a flow rate of 1 L/min until loss of righting and toe pinch reflex.
3. Transfer the animal to a heated pad (±38 °C) and position it supine with the head towards the surgeon. Fix the tail and front paws with tape on the pad. Ensure that the front limbs are not over-stretched, as this can compromise respiration.
4. Place a 2-0 silk suture (see Table of Materials) around the upper incisors and tape the suture on the border of the pad to pull the mouse taut. If necessary, attach a needle holder to keep tension on the suture. Maintain the animal anesthetized using 2% isoflurane in 100% oxygen with a flow of 1 L/min using a nosecone.
5. Place a light source just rostral of the sternum, remove the nose cone, and place the isoflurane supply on the ventilator. Intubate the mouse using forceps by lifting the tongue upwards and holding a self-made laryngoscope to lift the basal part of the tongue further and visualize the vocal cords.
6. Place a self-made tracheal tube using an 18 G catheter on a dull needle between the vocal cords. Move up the catheter, extract the dull needle, and connect the tube with the mouse ventilator (see Table of Materials).
7. Use a modified Y-shaped connector (see Table of Materials) to connect the intubation tube to the ventilator. The correct positioning of the tracheal tube can be confirmed by judging the symmetrical chest expansion.
8. Set the tidal volume (mL) at bodyweight (BW) (g) x 7 and ventilation rate at 53.5 x (BW (g)^-0.26) strokes per min, and adjust it to the body weight of a particular mouse if necessary⁶ (e.g., for a 25 g mouse the tidal volume is 175 mL at 140 strokes per min).
9. Reduce the isoflurane to 1% in 100% oxygen, as the animal will stay anesthetized due to the premedication. However, regularly assess the depth of anesthesia by performing a toe pinch. Adapt Isoflurane concentration based on anesthesia depth.
10. Apply a drop of ophthalmic ointment to prevent dryness while under anesthesia.

2. Installation of the IRI tool

NOTE: Remove all taping and place the animal on the base plate of the purpose-built IRI tool. Details on the properties of the IRI tool are mentioned in Supplementary File 1.

1. Fixate the base plate of the purpose-built remote IRI tool in a sideways position in front of the surgeon. Fix the tail and paws using tape. Fix the head using the 2-0 silk suture around the upper incisors to prevent accidental movement (Figure 1).
2. Insert the rectal thermal probe (see Table of Materials) to monitor the body temperature and use an infrared heating lamp above the animal to maintain the temperature around 37 °C. Pay attention to sufficient distance between the animal and the lamp to avoid overheating.
3. Position the ECG needle electrodes in all four paws to observe and record heart rate and ECG waveform. Secure the rectal probe and ECG electrodes to the platform using tape.
4. Apply hair removal cream on the left side of the thorax to remove hair and create a clear surgical field.
   NOTE: Limit the application time and pay attention to the complete removal of the cream to prevent chemical burns.

3. Thoracotomy

1. Perform aseptic preparation of the skin using Betadine Solution, afterwards place a surgical drape on the left thorax to ensure a sterile environment. Use sterilized equipment to perform the procedure. Make a vertical skin incision at the mid portion of the pectoral muscle approximately 1 cm long and 2 mm away from the left sternal border.
2. Spit the pectoral muscle to expose the ribs underneath. Avoid accidental injury to the vessel. If bleeding occurs, use cotton applicators to stop any bleeding.
3. Visualize the ribs and identify the intercostal spaces by observing the inflating lung through the thin, semitransparent chest wall. Open the chest cavity using surgical scissors by making a 6-8 mm incision in the third intercostal space. Ensure the incision is a minimum of 2 mm from the sternal border where the internal thoracic artery is located.
4. Insert a small rat eye speculum (see Table of Materials) in the intercostal space. This will function as a rib retractor (Figure 1A).
5. Gently lift the pericardium with curved forceps and pull it apart.
6. Place a small piece of cotton on top of the lung and push gently downwards.

4. LAD preparation

1. The LAD appears bright red and runs from the aorta under the left auricle towards the apex. The ideal positioning for the ligature is approximately 2 mm lower than the tip of the left auricle. Use the pulmonary trunk as a marker to help identify the left auricle and optimize the LAD visualization using light influx (Figure 2).
2. Use a tapered needle to pass a 7-0 polypropylene suture around the LAD. Do not place the needle too deep, as it can enter the left ventricle, nor too shallow, as it can damage the LAD.
3. Remove the small piece of cotton and check if the left lung is still ventilated.
4. Remove the rib retractor.
5. Ensure the suture is of a minimum length of 15 cm on both ends. Cut off the needle.
6. Place a piece of 1 mm PE-50 tubing over a piece of 15 mm PE-10 tubing (see Table of Materials). Guide both ends of the suture through the PE-10 tubing and shift the PE tubing with its thicker end against the heart.

5. Chest closure and extubating

1. Apply some fusidic acid gel on the open intercostal space and gently allow the remaining intrathoracic air to escape by briefly obstructing the outflow of the ventilator. Additional fusidic acid gel can be added to the external part of the PE tube to prevent air entry into the thorax.
2. Close the pectoral muscle with a 5-0 polypropylene X stitch above and below the level of the PE tube, and ensure the thickened part of the PE tube is under the level of the muscle.
3. Close the skin with two 5-0 polypropylene X stitches above and two below the PE tube (Figure 1B and Figure 3).
4. With a closed chest, the animal can breathe spontaneously again. Wean the animal from the ventilator by slowly reducing tidal volume and respiratory rate. When the animal starts breathing, spontaneously carefully disconnect the ventilator tube, but keep it in place until a stable breathing pattern is observed.
5. Connect the isoflurane supply with a nosecone that is placed over the nose of the animal and fixed on the base plate.

6. Assembly of the remote IR tool

1. Place the vertical side part in the base plate by inserting it in the slits.
2. Use a fine hook to retrieve the two suture ends and pull them through the central hole in the side part behind the balloon (see Table of Materials).
3. Guide one of the sutures ends above and around the balloon. Follow through with the other suture end under and around the balloon. Apply fusidic acid gel for smoother guiding of the suture.
4. Guide the suture ends through the slits on top of the side part. On the distal part of the suture, apply a weight of 2.1 g to keep each suture end in place (Figure 1C and Figure 4).
   NOTE: The configuration of appropriate weights is subject to change and must be tested beforehand. See Supplementary Figure 1 and Supplementary File 1 for details. Remote IRI tool design is provided in Supplementary Figure 2.

7. Induction of ischemia and reperfusion

1. Induce ischemia by promptly inflating the balloon using the vascular balloon pump up to 2 bar. Lock the pump. Visually check if the balloon is inflated and the weights are lifted (Figure 4).
2. Confirm the ischemia by observing the change in the ECG trace.
3. Leave the balloon inflated for the set time according to the specific study protocol (30 min in the presented experiments).
4. Stop the occlusion by simply unlocking the pump. Deflate the balloon carefully. Visually check if the balloon is deflated and weights are lowered.

8. Disassembly of the remote IR tool

1. Cut the suture and tube between the animal and the side part and carefully remove the side part.
2. Apply fusidic acid gel at the insertion site of the tubing into the thorax. Use curved forceps to gently push the skin against the tube and use another forceps to extract the tube. Cut any remaining suture close to the skin.
3. Place an extra suture of 5-0 polypropylene to close the exit site of the tube and minimize the risk of air entry into the thorax.

9. End of narcosis and recovery

1. Stop the isoflurane supply and allow the animal to wake up in a silent, warm, and oxygen-rich environment.
2. Confirm the mouse is not in any respiratory distress by observing it until full recovery. Provide 0.3 mL of sterile saline (at body temperature) by intraperitoneal injection to prevent dehydration.
3. For post-operative analgesia, administer an opioid analgesic (buprenorphine, 0.1 mg/kg) subcutaneously before the animal is ambulatory. Check the mice every 4-6 h for the next 24-48 h when buprenorphine is provided to ensure that the mice are responding to pain medications and to determine if additional pain medications are needed. Additional support should include a standard recommended supportive care diet gel or mash placed on the cage floor post-operatively.

Results

Validation of the ability to induce ischemia has been performed by four tests: Triphenyl tetrazolium chloride (TTC) and Sirius Red (SR) staining, cardiac troponin I measurement, and Late gadolinium enhancement (LGE) MR imaging. Statistical significance was evaluated using the Mann-Whitney non-parametric test, considering the limited sample sizes. Statistical significance was attributed to p < 0.05.

Acute experiments (TTC staining, n = 15) had no technical failure, and all animals w...

Discussion

The novel remote occlusion technique introduced in this study offers a unique platform to advance research in the field of ischemia-reperfusion injury modeling by avoiding the need for direct vessel manipulation during the initial surgery and allowing simultaneous multi-modal imaging of the early reperfused myocardium. Comprehensive characterization, including troponin-I measurements, LGE contrast MRI, TTC, and SR staining, shows that the proposed technique is equivalent to the current golden standard (i.e., ope...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
2-0 silk suture	Sharpoint Products	DC-2515N
5-0 polypropylene suture	Ethicon	8710H
7-0 polypropylene suture	Ethicon	8206H
ACCLARENT Balloon Inflation Device	Johnson&Johnson MedTech	BID30
Acepromazine	Kela
BD Vialon 18 G	BD	381347
BD Vialon 20 G	BD	381334
Betadine Solution	Purdue Pharma	25655-41-8
Buprenorphine (Buprenex Injectable)	Reckitt Benkiser Healthcare	NDC 12496-0757-1
Carp Zoom Styl Knijplood 2.1 g (lead fishing weights)	Visdeal.nl
Dumont #3 Forceps	Fine Science Tools	11231-30
Dumont #5 Forceps	Fine Science Tools	11251-30
Fine Scissors	Fine Science Tools	14040-10
Fucidine gel	Leo Pharma
Isoflurane	Abbott	NDC 5260-04-05
KD Mouse/Rat Eye Speculum	World Precision Instruments	501897
KD mouse/rat eye speculum	World Precision Instruments	501897
Ketamine	Dechra
Light source	Zeiss	KL 1500 LCD
MRI system	Bruker BioSpin, Ettlingen, Germany	BioSpec 70/30
NatriumChloride 0.9%	Baxter
Nocturnal Infrared Heat Lamp	Zoo Med Laboratories, Inc.	RS-75
ParaVision software	Bruker BioSpin	version 6.0.1
polyethylene tubing PE-10	SAI Infusion technologies	PE-10
polyethylene tubing PE-50	SAI Infusion technologies	PE-50
remote IRI tool (PMMA)	homemade
Rodent Surgical Monitor	Indus instruments
Segment v4.0	Medviso, segment.heiberg.se	R12067
Self-gated gradient echo sequence	Bruker BioSpin, Ettlingen, Germany	IntraGate, ParaVision 6.0.1
Slim Elongated Needle Holder	Fine Science Tools	12005-15
Sure-Seal Mouse/Rat Induction Chamber	World Precision Instruments	EZ-178
Tubing Connectors, Poly, Y Shape	Westlab	072025-0001
Ultraverse 035 PTA Dilatation Catheter: 5mm x 40mm, 17 ATM RBP balloon on 130 cm long catheter	Bard Peripheral Vascular, Inc.	00801741092671
Veet (depilatory creme)	Reckitt Benkiser Healthcare
Ventilator, MiniVent Model 845	Hugo Sachs	73-0043
Vidisic	BAUSCH & LOMB PHARMA	685313
Xylazine	Bayer