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.

Representative 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 rate is higher in both surgical procedures).

Aortic banding and myocardial remodeling
The success of aortic constriction was verified by increased LV end-systolic pressure (LVESP) and by Doppler aortic flow velocities >2.5 m/s, which corresponds to a pressure gradient of 25 mmHg using the modified Bernoulli equation (Figure 5). Compared to SHAM mice, banding induced LV hypertrophy as assessed by increased LV mass (Table 1 and Figure 5) and impaired diastolic function evident by higher filling pressures (ratio of mitral peak velocity of early filling (E) to early diastolic mitral annular velocity (E'),  (E/e'), and left ventricular end-diastolic pressure (LVEDP) and prolonged relaxation (t, Table 1, Figure 5, and Figure 6) within 7 weeks. Ejection fraction was still preserved at this stage of the disease.

Histologically, seven-weeks of aortic banding induced significant cardiomyocyte hypertrophy and fibrosis (Figure 7).

Aortic debanding and myocardial reverse remodeling
In mice subjected to debanding, successful removal of aortic stenosis was verified by echo Doppler velocities (Table 1 and Figure 5). Overall, debanding promoted a significant decrease of afterload (decreased LVESP) and LV hypertrophy (assessed by morphometry, echocardiography, and histology). Moreover, we observed normalization of diastolic function and aortic velocities (Table 1, Figure 5, Figure 6, and Figure 7).

Table 1: Left ventricle morphofunctional changes assessed by echocardiography and by hemodynamics.

Critical steps	Advice
Invasiveness of the banding surgery	It is important to avoid: ● prolonged occlusion of the ascending aorta during the ligation, which may lead to lung edema and activation of inflammatory pathways capable of influencing the phenotype and disease severity15 ● bleeding of the mammary artery which, if not timely circumvented, can lead to decreased blood pressure and promote higher amounts of fibrosis when re-opening the thorax (debanding); ● damaging mice pleura and lungs; Mini left lateral thoracotomy for banding and debanding (same place; present study) vs left lateral thoracotomy for the banding and a sternotomy for the debanding surgery11: ● the first is less invasive and have a short-recovery time, which improves the success of the open-chest haemodynamic performed two weeks later. Neverthless, the use of same position to re-open the chest can increase the number of complications due to adhesions (around left atrium, pulmonary artery, etc). Overcome this issue by having extracarefull during banding procedure.
Suture internalization	Can be prevented by using: ● two banding sutures side-by-side16; ● silk instead of polypropylene11; ● titanium clips or a O-rings around the aorta to induce its constriction21; ● double loop-clip thecnique15; ● inflatable cuff to carry out supracoronary aortic banding22.
Physiological parameters	During surgery it is important to monitor: ● heart rate; ● blood oxygenation, keeping it above 90% (specially during aorta manupilation); ● anesthesia, keeping it at the lowest dose possible without inflicting discomfort on the animal.

Table 2: Critical steps of the protocol.

Figure 1: Ultra fine surgical instruments used for the banding and debanding procedures. (A) 2 needle-holders and a scalpel blade; 2 catheters for mice intubation and a scissor; a scalpel, 2 curved forceps, a ligation aid, a microsurgical scissor, 3 straight forceps; (B) and 26G-needle and blunted 26G-needle curved to fit the mice small thoracic opening properly. Please click here to view a larger version of this figure.

Figure 2: Aortic banding procedure. (A) The thoracic approach to the ascending aorta performed with the help of a magnetic fixator retraction system (3 retractors are visible). (B) The ascending aorta is clearly dissected and visible. The blunted needle and the polypropylene suture 6-0 are placed in the right position to perform the aortic banding. Please click here to view a larger version of this figure.

Figure 3: Aortic debanding procedure. (A) The mouse is placed in a magnetic retraction system, representing a handy tool to retract the muscles and tissues. The mouse is intubated for mechanical ventilation. A rectal probe controls temperature and an oximeter is placed on the right mice paw to monitor blood oxygenation during surgery. Fibrosis and adherent tissue is carefully removed around the aorta and suture, to be able to cut the suture (B) and (C). Please click here to view a larger version of this figure.

Figure 4: Experimental protocol design for mice. Myocardial remodeling (red) and reverse remodeling (green) are shown in the bottom together with all evaluation tasks. Of note, debanding surgery can give rise to two groups of animals with distinct degrees of reverse remodeling. Thus, we obtained DEB mouse with complete (DEB-COMP) and incomplete (DEB-INCOM) myocardial recovery. Please click here to view a larger version of this figure.

Figure 5: Echocardiographic assessment of cardiac structure and function. (A) Aortic flow velocities; (B) LV mass; (C) Ventricular dimensions (LV diameter, LVD) and wall thickness (LV posterior wall, LVPW and LV anterior wall, LVAW); (D) Transmitral flow (peak of pulse Doppler wave of late mitral flow velocity, A, and peak of pulsed Doppler wave of early mitral flow velocity, E) and (E) Myocardial velocities (late diastolic mitral annular tissue velocity, A'; early diastolic mitral annular tissue velocity, E' and systolic mitral annular tissue velocity, S'). Please click here to view a larger version of this figure.

Figure 6: Representative pressure-volume loops for SHAM, BA and DEB groups. Data were continuously acquired at 1000 Hz and subsequently analyzed off-line by PVAN software. Please click here to view a larger version of this figure.

Figure 7: Myocardial hypertrophy and fibrosis assessed histologically. (A) Left ventricle hypertrophy assessed by cardiomyocytes sectional area of hematoxylin-eosin (HE)-stained sections (5 µm) from SHAM (n = 17), BA (n = 14) and DEB group (n = 12). (B) Left Ventricular interstitial fibrosis and representative images of Red Sirius-stained sections (5 µm) from SHAM (n = 17), BA (n = 13) and DEB (n = 12). Please click here to view a larger version of this figure.

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 technique to reduce the surgical trauma.

The most critical step of the protocol is related to the degree of surgical aggression during aortic banding. The success of the subsequent aortic debanding surgery depends enormously on a minimally invasive banding procedure that avoids tissue aggression and fibrosis around the aorta and, therefore, a less-invasive approach is mandatory (Table 2). Suture internalization is associated with less LV hypertrophy and better cardiac function¹⁶ (Table 2) and makes the debanding procedure impossible to perform without causing an aortic rupture. In the present study, we tried to use silk, since it creates more scar tissue at the banding site, triggering a more stable degree of pressure overload. However, in our hands, the debanding surgery was more demanding when silk was used since it is a multifilament wire making it's total removal from the aorta more difficult. Nevertheless, these are technical issues that are widely protocol-and-operator-dependent, and these variations, type of suture, is not incompatible with good technical practices and reproductive results. Physiological parameters monitoring during banding and especially during debanding is mandatory for the success of the model implementation (Table 2).

In 1991, Rockman et al., described the transverse aorta constriction (TAC) in mouse for the first time⁴. Since then a considerable amount of papers came out providing numerous versions of this procedure with variations with respect to the animal age/size¹⁷, mice genetic background¹⁸, the diameter of the needle/constriction¹⁹, the material used for banding, the aortic location of the banding, the duration of the banding¹⁹ and debanding¹¹. All these methodological alternatives are valid as long as they fulfill the aims of each study. However, we should stress out that the progression of the disease towards heart failure is faster and thus RR is more incomplete when selecting: 1) longer banding durations, 2) heavier/older the mice²⁰ and 3) smaller needle diameter used for the aortic constriction (higher percentage of aortic constriction)¹⁶.

The duration of the banding and the debanding significantly impact the stage of the disease and, therefore, the recovery during RR. Likewise, choosing the right timing for debanding is mandatory to adjust to the severity of the disease envisaged. The results observed in our study are in accordance with pre-existence animal^11,21 and human studies²², except for cardiomyocytes hypertrophy, where some studies showed its normalization^10,21 and others its partial regression²³.  Moreover, studies have shown that, fibrosis regression can occur in the long term (70 months for human patients)²⁴. The results seem to be dependent on the technique used to address fibrosis²⁵. Recently, Treibel et al. were able to differentiate between cellular (myocytes, fibroblast, endothelial, red blood cells) and extracellular (ECM, blood plasma) compartments in patients with aortic stenosis after aortic valve replacemennt (AVR) using cardiovascular magnetic resonance with T1 mapping²². They described that regression of LV mass following AVR can be driven by 1) matrix regression alone, where the extracellular volume reduces; 2) cellular regression alone, where extracellular volume increases; 3) or by a proportional regression in cellular and matrix compartments, where the extracellular volume is unchanged²². These authors concluded that, following AVR, while diffuse fibrosis and myocardial cellular hypertrophy regress, focal fibrosis does not resolve. Thus, diffuse interstitial fibrosis, as assessed by matrix volume, is a potential therapeutic target. In our study, reduction of fibrosis seems to occur within 2 weeks of RR and before cardiomyocytes hypertrophy normalization. Also, sacrificing the animals 2 weeks after the debanding was the perfect timing to obtain ventricular diversity among DEB group, namely animals with diastolic dysfunction persistence (DEB-INCOM) and others with complete LV mass reversal and diastolic function improvement (DEB-COM). Moreover, as soon as 2 weeks after debanding, we have previously shown significant right ventricular changes in the banding group that partially recover after debanding²⁶, while Bjornstad et al. reported normalization of fetal genes expression, indicative of myocardial remodeling within the same timeframe¹¹.

The surgical procedure of banding/debanding can also be performed in rats²⁶, however, some differences should be highlighted. Due to its bigger size, rats have more muscle layers than mice which decreases aortic visualization and hinders positioning the ligature around the aorta. On the other hand, the risk of damaging adjacent tissues and organs, such as atria or lungs, are minimized. To overcome the issue of suture internalization we used a larger polypropylene ligature in rats to hold tight the aorta (6.0 instead of 7.0 polypropylene).

Due to aorta manipulation, debanding surgery might decrease cardiac output by imposing additional afterload on LV and thus impair the circulatory and respiratory system. Compared to mice, rats seem to be more resistant to more extended anesthetic period and therefore are easier to keep the physiological respiratory parameters controlled during the long debanding surgery. In rats, LV hypertrophy development is faster than mice, but it takes longer to progress to heart failure. Thus, the debanding surgery can be done between 5-9 weeks after banding procedure without compromising ejection fraction²⁶.

The major limitation of the banding/debanding animal model is the demanding microsurgical skills and technique of the operator, usually requiring a long learning curve to accomplish the debanding surgery. Another limitation is the impossibility to perform close chest hemodynamics in mouse and rat, which will be more physiologic. However, by using this method is obligatory to insert the catheter from the right carotid artery to LV which is, in this particular case not feasible since in banding animals ascending aorta is constricted before the carotid branches. Moreover, in mouse, we were not able to measure load-independent contractility (ESPVR) and diastolic parameters (slope of EDPVR) by performing vena cava occlusion maneuver, an important parameter for an adequate characterization of myocardial function. We found this maneuver difficult to perform in mice with ascending aorta constriction due to their small size (20-25g). 

Future application of the banding/debanding animal model includes the development of novel therapeutic approaches to myocardial diseases and the characterization of the pathways that underlie the process of LV remodeling and RR.

In conclusion, this clinically-relevant model allows to temporally and mechanistically characterize the progression towards HF, as well as, its recovery since it allows the collection of myocardial samples in different stages of myocardial remodeling and RR. Moreover, it proves to be a useful experimental model for testing therapeutic strategies aimed at the recovery of the failing heart.

Materials

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