Studying Left Ventricular Reverse Remodeling by Aortic Debanding in Rodents

Summary

Here we describe a step-by-step protocol of surgical aorta debanding in the well-established mice model of aortic-constriction. This procedure not only allows studying the mechanisms underlying the left ventricular reverse remodeling and regression of hypertrophy but also to test novel therapeutic options that might accelerate myocardial recovery.

Abstract

To better understand the left ventricular (LV) reverse remodeling (RR), we describe a rodent model wherein, after aortic banding-induced LV remodeling, mice undergo RR upon removal of the aortic constriction. In this paper, we describe a step-by-step procedure to perform a minimally invasive surgical aortic debanding in mice. Echocardiography was subsequently used to assess the degree of cardiac hypertrophy and dysfunction during LV remodeling and RR and to determine the best timing for aortic debanding. At the end of the protocol, terminal hemodynamic evaluation of the cardiac function was conducted, and samples were collected for histological studies. We showed that debanding is associated with surgical survival rates of 70-80%. Moreover, two weeks after debanding, the significant reduction of ventricular afterload triggers the regression of ventricular hypertrophy (~20%) and fibrosis (~26%), recovery of diastolic dysfunction as assessed by the normalization of left ventricular filling and end-diastolic pressures (E/e' and LVEDP). Aortic debanding is a useful experimental model to study LV RR in rodents. The extent of myocardial recovery is variable between subjects, therefore, mimicking the diversity of RR that occurs in the clinical context, such as aortic valve replacement. We conclude that the aortic banding/debanding model represents a valuable tool to unravel novel insights into the mechanisms of RR, namely the regression of cardiac hypertrophy and the recovery of diastolic dysfunction.

Introduction

The constriction of the transverse or ascending aorta in the mouse is a widely used experimental model for pressure overload-induced cardiac hypertrophy, diastolic and systolic dysfunction and heart failure^1,2,3,4. Aortic-constriction initially leads to compensated left ventricle (LV) concentric hypertrophy to normalize wall stress¹. However, under certain circumstances, such as prolonged cardiac overload, this hypertrophy is insufficient to decrease the wall stress, triggering diastolic and systolic dysfunction (pathological hypertrophy)⁵. In parallel, changes in extracellular matrix (ECM) lead to the collagen deposition and crosslinking in a process known as fibrosis, which can be subdivided into replacement fibrosis and reactive fibrosis. Fibrosis is, in most of the cases, irreversible and compromises myocardial recovery after overload relief^6,7. Nevertheless, recent cardiac magnetic resonance imaging studies revealed that reactive fibrosis is able to regress in the long term⁸. Altogether, fibrosis, hypertrophy and cardiac dysfunction are part of a process known as myocardial remodeling that rapidly progresses towards heart failure (HF).

Understanding the features of myocardial remodeling has become a major objective for limiting or reversing its progression, the latter known as reverse remodeling (RR). The term RR includes any myocardial alteration chronically reversed by a given intervention, such pharmacological therapy (e.g., antihypertensive medication), valve surgery (e.g., aortic stenosis) or ventricular assist devices (e.g., chronic HF). However, RR is often incomplete due to the prevailing hypertrophy or systolic/diastolic dysfunction. Thus, the clarification of RR underlying mechanisms and novel therapeutic strategies are still missing, which is mostly due to the impossibility to access and study human myocardial tissue during RR in most of these patients.

To overcome this limitation, rodent models have played a significant role in advancing our understanding of the signaling pathways involved in HF progression. Specifically, aortic debanding of mice with an aortic constriction represents a useful model to study the molecular mechanisms underlying adverse LV remodeling⁹ and RR^10,11 as it allows the collection of myocardial samples at different time points in these two phases. Moreover, it provides an excellent experimental setting to test potential novel targets that can promote/accelerate RR. For instance, in aortic stenosis context, this model might provide information about the molecular mechanisms involved in the vast diversity of myocardial response underlying the (in)completeness of the RR^6,12, as well as, the optimal timing for valve replacement, which represents a major shortcoming of the current knowledge. Indeed, the optimal timing for this intervention is a subject of debate, mainly because it is defined based on the magnitude of aortic gradients. Several studies advocate that this time point might be too late for the myocardial recovery as fibrosis and diastolic dysfunction are often already present¹².

To our knowledge, this is the only animal model that recapitulates the process of both myocardial remodeling and RR taking place in conditions such as aortic stenosis or hypertension before and after valve replacement or the onset of anti-hypertensive medication, respectively.

Seeking to address the challenges summarized above, we describe a surgical animal model that can be implemented both in mice and rats, addressing the differences between these two species. We describe the main steps and details involved when carrying out these surgeries. Finally, we report the most significant changes taking place in the LV immediately before and throughout RR.

Protocol

All animal experiments comply with the Guide for the Care and Use of Laboratory Animals (NIH Publication no. 85–23, revised 2011) and the Portuguese law on animal welfare (DL 129/92, DL 197/96; P 1131/97). The competent local authorities approved this experimental protocol (018833). Seven-week-old male C57B1/J6 mice were maintained in appropriate cages, with a regular 12/12 h light-dark cycle environment, a temperature of 22 °C and 60% humidity with access to water and a standard diet ad libitum.

1. Preparation of the surgical field

1. Disinfect the operation site with 70% alcohol and place a disposable operating room table cover over the surgical area.
2. Sterilize all the instruments before surgery.
   NOTE: This procedure requires micro surgical scissors, 2 fine curved forceps, 3 fine straight forceps, a scalpel, small forceps, an angled dissector scissor, a needle holder, an ultrafine ligation aid, 2 hemostats and, lastly, a magnetic fixator retraction system is highly recommended (Figure 1A).
3. Curve the tip of a 26 G blunted needle to 90° for an easier approach to the aorta. A 26 G needle will create a 0.45 mm diameter aortic narrowing (Figure 1B).
4. Adjust the heating pad temperature to 37 ± 0.1 °C.

2. Mice preparation and intubation

1. Anesthetize young C57B1/J6 mice (20-25 g) by inhalation of 8% sevoflurane with 0.5 - 1.0 L/min 100% O₂ in a cone tube.
2. Check the anesthesia depth using the toe pinch withdrawal reflex.
3. Place the mouse at dorsal recumbency on an inclined plate and proceed to orotracheal intubation.
4. Move the mouse to the heating pad and quickly connect the orotracheal tube to the ventilator to initiate the mechanical ventilation.
5. Adjust the ventilator parameters to a frequency of 160 breaths/min and a tidal volume of 10 mL/kg.

3. Preparation for surgery (for both banding and debanding surgeries)

1. Shave and apply the depilatory cream from the neckline to mid-chest level of the mice.
2. Apply ophthalmic gel to the animals' eyes to prevent drying out of the cornea.
3. Place a rectal probe and the oximeter at the paw or tail for monitoring temperature and blood oxygenation, and heart rate, respectively.
   NOTE: Anesthesia induces significant hypothermia, therefore, it is important to maintain normal body temperature during surgery to avoid a rapid decrease in heart rate.
4. Maintain anesthesia with sevoflurane (2.0 - 3.0%). Check the correct level of anesthesia by the lack of the toe-pinch reflex.
5. Place the mice in right-lateral decubitus on a heating pad and secure the limbs to the magnetic fixator retraction system with a tape to keep the animal in the correct position during the surgery (Figure 2, Figure 3A).
6. Disinfect the mouse chest with 70% alcohol followed by providone-iodine solution.

4. Ascending aortic banding surgery

NOTE: For a detailed protocol description, consult ^2,3,4,13.  

1. With a disposable blade, perform a small (~0.5 cm) skin incision on the left side immediately below the axilla level and dissect the skin.
2. Gently dissect and separate the pectoralis muscle and other muscle layers until the ribs become visible. Use fine forceps and avoid cutting the muscle.
3. Under a microscope, identify the intercostal spaces and open a small incision between the 2^nd and 3^rd intercostal space with fine forceps.
4. Retract the ribs by placing the chest retractor (Figure 2A).
5. Use small forceps to gently dissect and separate the thymic lobes until ascending aorta becomes visible.
   NOTE: Cotton applicators should be handy in case of bleeding. Warm sterile saline should be given subcutaneously in case of significant bleeding (e.g., the mammary artery).
6. Use small forceps to gently dissect the aorta.
   NOTE: Aorta is considered to be dissected when there are no fat or other adhesions around it and it is possible to easily encircle the vessel with a small curve forceps.
7. After aortic dissection, place a 7-0 polypropylene ligature around aorta by using ligation aid and curved forceps (Figure 2B).
8. Position the blunted 26 G needle parallel to the aorta (tip pointed towards the mice head) (Figure 2B). For mice weighing 20-25 g, this needle induces a reproducible 65-70% aortic constriction.
9. Make 2 loose knots around the aorta and the 26 G needle with the help of 2 forceps (Figure 2B).
10. Tighten the 1^st knot and, quickly after, the 2^nd knot. Briefly confirm the right positioning of the constriction and quickly remove the needle to restore aortic blood flow. Finally, make a 3^rd knot (BA group).
11. Reposition the thymus and the muscles into their initial position.
12. Perform the sham procedure identical to the constriction procedure but keeping the suture loose around the aorta (SHAM group).
13. Cut the ends of the suture and remove the chest retractor.
    NOTE: Short suture ends may increase the probability of knots loosening with aortic pressure, while long ends make the debanding procedure riskier since adhesions can occur between the suture and the left atrium.
14. Close the chest wall using 6-0 polypropylene suture with a simple interrupted or continuous suture using the lowest number of stitches possible. Tighten the last chest knot with the lungs inflated at end inspiration by pinched off the outflow of the ventilator for 2s to re-inflate the lungs.
15. Close the skin with a 6-0 silk/polypropylene suture in a continuous suture pattern. 
    NOTES: If a more recent ventilator is used, it is possible to programme it to pause in inspiration (Setup-Advanced-Pause-Inspiration)

5. Post-operative care

1. Apply providone-iodine solution to the skin suture site.
2. For proper analgesia, administer buprenorphine subcutaneously 0.1 mg/kg, twice daily, until the animal fully recovers (usually 2-3 days after surgery).
3. Inject sterile saline intraperitoneally to prevent dehydration in case of significant bleeding during the surgery.
4. Turn off the anesthesia (without deintubating the mouse) and wait until the animal recover the reflexes (whiskers movements are an awakening signal)  and starts to breathe spontaneously.
5. Remove the tracheal cannula.
6. Let the animal recover in an incubator at 37 °C.
7. Return the animal to a 12 h light/dark cycle room after full recovery.

6. Aortic debanding surgery

1. Seven weeks later, perform the debanding of the aorta in half of the BA  animals and remove the loose suture from half of the SHAM mice, giving rise to 2 new groups -- debanding (DEB) and debanding SHAMA (DESHAM), respectively. DESHAM represents the control for the DEB group (Figure 4).
2. Repeat all the steps 2.1 to 3.6 mentioned above.
3. Gently dissect the tissues, adhesions, and fibrosis around the aorta until its constriction becomes visible.
4. Carefully dissect the aorta and separate the suture from the aorta. Cut the suture with angled one-probed spring scissors (Figure 3B).
5. Close the chest wall using 6-0 polypropylene suture with a simple interrupted or continuous suture using the minimum number of stitches possible.
   NOTE: Try to tighten the last chest suture when lungs are inflated to avoid pneumothorax.
6. Close the skin with a 6-0 silk/polypropylene suture in a continuous suture pattern. 
7. Perform all post-operative care procedures as mentioned in 5.
8. Sacrifice the animals 2 weeks later.

7. Echocardiography to assess cardiac function and left ventricular hypertrophy in vivo

1. Perform the echocardiographic exam every 2-3 weeks to follow the progression of hypertrophy and cardiac function.
2. Anesthetize the animals, as described, by inhalation of 5% sevoflurane with a nose cone. Adjust the anesthesia level by decreasing it to 2.5%.
3. Shave and apply the depilatory cream from the neckline to mid-chest level.
4. Place the animal on a heating pad and place the ECG electrodes. Assure a good ECG trace and maintain heart rate between 300 and 350 beats/min.
5. Monitor the temperature (~37 °C).
6. Apply echo gel and position the animal at left lateral decubitus.
7. Start the echocardiograph and adjust the settings.
8. Position an ultrasound probe over the thorax.
9. Assess the pressure gradient across the banding at 7 and 2 weeks after banding and debanding surgery, respectively. Position the probe at the LV long axis and place the beam over aorta. Press the button PW to activate pulsed wave Doppler echocardiography. After seven weeks of banding, aortic gradients will be >25 mmHg in the banded animals.
10. Record two-dimensional guided images of aorta showing the presence or absence of the ascending aorta constriction to anatomically visualize the efficacy of the banding and debanding.
    NOTE: It is possible to visualize turbulent flow at the constriction level if the color mode is available.
11. Assess hypertrophy by positioning the probe at an LV short axis, at papillary muscles level, and press M-mode tracing to visualize LV anterior wall (LVAW), LV diameter (LVD) and LV posterior wall (LVPW) in diastole (D) and systole (S) (Figure 5).
12. Assess systolic function, calculate the ejection fraction and fractional shortening as previously described^14,15.
13. Assess diastolic function by 1) determining the peak of pulsed-wave Doppler of early and late mitral flow velocity (E and A waves, respectively) using an apical 4-chamber apical view just above the mitral leaflets; 2) recording lateral mitral annular myocardial early diastolic (E') and peak systolic (S') velocities using pulsed-TDI and apical 4-chamber apical view (Figure 5).  
14. Record at least three consecutive heartbeats to each parameter assessment. These values will be subsequently averaged.

8. Hemodynamic assessment

1. At the end of the protocol (Figure 4), perform final echocardiography, as described in 7, before the terminal hemodynamic assessment.
2. Repeat steps 2.1 to 3.6.
3. Cannulate the right jugular vein and perfuse sterile saline at 64 mL/kg/h.
4. Rotate slightly the animal to the left side and make a skin incision at the level of the xiphoid appendix.
5. Separate the skin from the muscle with forceps or with a scissor.
6. Make a lateral incision between the left ribs at the xiphoid appendix level.
7. Perform a left lateral thoracotomy to expose the heart fully.
   NOTE: To avoid bleeding and lung damage, insert a cotton swab into the thoracic cavity and push the lung gently while inserting two hemostats on the right and left side of the place to cut.
8. Pre-heat the P-V loop catheters in a water-bath at 37 °C.
9. Calibrate the catheter (setup, channel setting, chose the correct channel for pressure and volume, units).
10. Insert a catheter apically into the LV and assure the volume sensors are positioned between the aortic valve and the apex. Volumes can be assessed by echocardiography (Figure 5). Visualization of the pressure-volume loops helps to confirm the correct positioning of the catheter (Figure 6).
11. Allow the animal to stabilize 20-30 min without significant changes in the shape of the pressure-volume loops.
12. With ventilation suspended at end-expiration, acquire baseline recordings (Figure 6). Continuously acquire data at 1,000 Hz to be subsequently analyzed off-line by appropriate software.
13. Compute parallel conductance after the hypertonic saline bolus (10%, 10 µL).
14. While anesthetized, sacrifice the animal by exsanguination, collect and centrifuge the blood.
15. Lastly, excise and collect the heart. Weight the heart, the left, and the right ventricle separately and immediately store the samples in liquid nitrogen or formalin for subsequent molecular or histological studies, respectively.

9. Aortic banding/debanding procedure in rats

1. Perform aortic banding in young Wistar (70-90 g) using a 22 G needle and 6-0 polypropylene ligature to constrict the aorta.
2. Ensure a proper anesthetic and analgesic procedures with 3-4% of sevoflurane and 0.05 mg/kg of buprenorphine, respectively.
3. During echocardiography, assure a heart rate always above 300 rate / min (ideally between 300 and 350).
4. Before step 8.9, gently dissect the rat aorta, place a flow probe around it to measure cardiac output. The use of the aortic flow probe is the gold standard procedure for rats.
5. For the hemodynamic evaluation, cannulate the jugular or femoral vein for fluid administration (32 mL/kg/h).
6. Replace the pressure-volume catheter SPR-1035 by the SPR-847 or SPR-838, whose sizes better suit the rat ventricular dimensions.

Results

Post-operative and late survival
The perioperative survival of the banding procedure is 80% and the mortality during the first month is typically <20%. As previously mentioned, the success of the debanding surgery is highly dependent on how invasive the previous surgery was. After a learning curve, the mortality rate during the debanding procedures is around 25%. For this mortality accounts mostly deaths during the surgery procedure, including aorta or left atrium rupture (in rats, the survival ra...

Discussion

The model proposed herein mimics the process of LV remodeling and RR after aortic banding and debanding, respectively. Therefore, it represents an excellent experimental model to advance our knowledge on the molecular mechanisms involved in the adverse LV remodeling and to test novel therapeutic strategies able to induce myocardial recovery of these patients. This protocol details steps on how to create a rodent animal model of aortic banding and debanding with a minimally invasive and highly conservative surgical techni...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Absorption Spears	F.S.T	18105-03	To absorb fluids during the surgery
Blades	F.S.T	10011-00	To perform the skin incision
Buprenorphine	Buprelieve		Analgesia drug
Catutery	F.S.T	18010-00	To prevent exsanguination
Catutery tips	F.S.T	18010-01	To prevent exsanguination
cotton swab	Johnson's		To absorb fluids during the surgery
Depilatory cream	Veet		To delipate the animal
Disposable operating room table cover	MEDKINE	DYND4030SB	To cover the surgical area
Echo probe	Siemens	Sequoia 15L8W	Ultrasound signal aquisition
Echocardiograph	Siemens	Acuson Sequoia C512	Ultrasound signal aquisition
End-tidal CO2 monitor	Kent Scientific	CapnoStat	To control expiration gas saturation
Forcep/Tweezers	F.S.T	11255-20	To dissect the tissues and aorta
Forcep/Tweezers	F.S.T	11272-30	To dissect the tissues and aorta
Forcep/Tweezers	F.S.T	11151-10	To dissect the tissues and aorta
Forcep/Tweezers	F.S.T	11152-10	To dissect the tissues and aorta
Gas system	Penlon Sigma Delta		To anesthesia and mechanical ventilation
Hemostats	F.S.T	13010-12	To hold the suture before tight the aorta
Hemostats	F.S.T	13011-12	To hold the suture before tight the aorta
Ligation aids	F.S.T	18062-12	To place a suture around the aorta
Magnetic retractor	F.S.T	18200-20	To help keep the animal in a proper position
Needle holder	F.S.T	12503-15	To suture the animal
Needle 26G	B-BRAUN	4665457	To serve as a molde of aortic constriction diameter
Oxygen	Air Liquide		To anesthesia and mechanical ventilation
Polipropilene suture	Vycril	W8304/W8597	To suture the animal and to do the constriction
Povidone-iodine solution	Betadine®		Skin antiseptic
PowerLab	Millar instruments	ML880 PowerLab 16/30	PV loop Signal Aquisition
Pulse oximeter	Kent Scientific	MouseStat	To control heart rate and blood saturation
PVAN software	Millar Instruments		To analyse the haemodynamic data
PV loop cathether	Millar instruments	SPR-1035. 1.4 F	PV loop Signal Aquisition
Retractor	F.S.T	17000-01	To provide a better overview of the aorta
Scalpet handle	F.S.T	10003-12	To perform the skin incision
Scissors	F.S.T	15070-08	To cut the suture in debanding surgery
Scissors	F.S.T	14084-09	To cut other material during the surgery e.g. suture, papper
Sevoflurane	Baxter	533-CA2L9117
Temperature control module	Kent Scientific	RightTemp	To control animal corporal temperature
Ventilator	Kent Scientific	PhysioSuite	To ventilate the animal
Water-bath	Thermo Scientific™	TSGP02	To maintain water temperature adequate to heat the P-V loop catethers